*Bacillus thuringiensis* **Toxins: Functional Characterization and Mechanism of Action**

Editors

**Yolanda Bel Juan Ferr´e Patricia Hern´andez-Mart´ınez**

MDPI • Basel • Beijing • Wuhan • Barcelona • Belgrade • Manchester • Tokyo • Cluj • Tianjin

*Editors* Yolanda Bel Universitat de Valencia ` Spain

Juan Ferre´ Universitat de Valencia ` Spain

Patricia Hernandez-Mart ´ ´ınez Universitat de Valencia ` Spain

*Editorial Office* MDPI St. Alban-Anlage 66 4052 Basel, Switzerland

This is a reprint of articles from the Special Issue published online in the open access journal *Toxins* (ISSN 2072-6651) (available at: https://www.mdpi.com/journal/toxins/special issues/Bt toxins).

For citation purposes, cite each article independently as indicated on the article page online and as indicated below:

LastName, A.A.; LastName, B.B.; LastName, C.C. Article Title. *Journal Name* **Year**, *Volume Number*, Page Range.

**ISBN 978-3-0365-2049-0 (Hbk) ISBN 978-3-0365-2050-6 (PDF)**

Cover image courtesy of Yolanda Bel, Juan Ferre and Patricia Hern ´ andez-Mart ´ ´ınez.

© 2021 by the authors. Articles in this book are Open Access and distributed under the Creative Commons Attribution (CC BY) license, which allows users to download, copy and build upon published articles, as long as the author and publisher are properly credited, which ensures maximum dissemination and a wider impact of our publications.

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### **Contents**




Reprinted from: *Toxins* **2020**, *12*, 116, doi:10.3390/toxins12020116 .................. **313**

### **About the Editors**

**Yolanda Bel** (Dr.) studied Biology (major in Biochemistry) and received her Ph.D. in Biology from the University of Valencia (UV), Spain. Her Ph.D. was undertaken in the Department of Genetics of the UV and in the Biology Division of the Oak Ridge National Laboratory, Oak Ridge (Tennessee, USA). She undertook her postdoctoral studies (1992–1993) in Sandoz, Basel, Switzerland. After a contract at the University of Valencia (1993–1996) to work on *Bacillus thuringiensis* (Bt), she completed a "Master in Industrial Wastewater Treatment", and worked in water microbiology in the private industry. She returned to the UV in 2003, where she currently works as a Research Associate in the Institute of Biotechnology and Biomedicine. Her current research interests include the screening and study of new Bt pesticidal proteins, the binding of Bt toxins to pests' midguts, study of their mode of action, and the biochemical and genetic bases of insect resistance.

**Juan Ferr´e** (Prof.) received his Ph.D. degree in Chemistry from the University of Valencia (UV), Spain, with a study which he carried out in both the Department of Genetics of the UV and the Biology Division of the Oak Ridge National Laboratory, Oak Ridge (Tennessee, USA). He conducted his postdoctoral studies in the Department of Reproductive Genetics of the Magee Womens Hospital (Pittsburgh, Pennsylvania, USA). He became a Professor of Genetics in 2000, and served as the Head of the Department of Genetics of the UV for 7 years. He is currently Director of the Institute of Biotechnology and Biomedicine of the UV (Biotecmed). His current research interests, ongoing since 1990, are (i) to understand the biochemical and genetic bases of insect resistance to *Bacillus thuringiensis* (Bt) toxins; (ii) to study the mode of action of Bt toxins; and (iii) to find novel Bt strains and insecticidal protein genes for the development of Bt-based insecticides to control agricultural insect pests.

**Patricia Hern´andez-Mart´ınez** (Dr.) studied Biology (1998–2003) at the University of Valencia (UV), Spain. She received her Ph.D. in Biology from the UV in 2009, with the work "Study of susceptibility and resistance towards *Bacillus thuringiensis* proteins on different *Spodoptera exigua* colonies", which she conducted in the Department of Genetics of the UV. After her Ph.D., her research career continued at the Institute of Biotechnology and Biomedicine (Biotecmed), where she still works as a Research Fellow. Her current research interests are (i) to study the mode of action of *B. thuringiensis* (Bt) proteins; (ii) to study the response mechanisms of different hosts after Bt protein challenge; and (iii) to understand the bases of insect resistance to Bt proteins.

### *Editorial Bacillus thuringiensis* **Toxins: Functional Characterization and Mechanism of Action**

**Yolanda Bel 1,2,\*, Juan Ferré 1,2,\* and Patricia Hernández-Martínez 1,2,\***


Received: 30 November 2020; Accepted: 8 December 2020; Published: 10 December 2020

*Bacillus thuringiensis* (*Bt*)-based products are the most successful microbial insecticides to date. This entomopathogenic bacterium produces different kinds of proteins whose specific toxicity has been shown against a wide range of insect orders, nematodes, mites, protozoa, and human cancer cells. Some of these proteins are accumulated in parasporal crystals during the sporulation phase (Cry and Cyt proteins), whereas other proteins are secreted in the vegetative phase of growth (Vip and Sip toxins). Currently, insecticidal proteins belonging to different groups (Cry and Vip3 proteins) are widely used to control insect pests and vectors both in formulated sprays and in transgenic crops (the so-called Bt crops). Despite the extensive use of these proteins in insect pest control, especially Cry and Vip3, their mode of action is not completely understood.

The aim of this Special Issue was to gather information that could summarize (in the form of review papers) or expand (research papers) the knowledge of the structure and function of Bt proteins, as well as shed light on their mode of action, especially regarding the insect receptors. This subject has generated great interest, and this interest has been materialized into the 18 papers published in this issue.

This Special Issue, "*Bacillus thuringiensis* Toxins: Functional Characterization and Mechanism of Action", includes five review papers and 13 research papers. The review papers bring up to date important aspects of Bt pathogenicity, such as its interaction with the intestinal microbiota and the immune system of the insect [1]. The current knowledge about Vip proteins has also been reviewed [2], as has the contribution that the use of toxin mutants has made to the knowledge of the mode of action of the three-domain Cry proteins [3]. On the other hand, two more review papers recapitulate the information on the cytocidal activity of Bt proteins [4] or the insecticidal activity of Bt proteins against coleopteran pests [5]. All these review papers are of high value, allowing readers to stay updated on the different aspects of the Bt field described here.

The Special Issue also gathers information that could expand the knowledge of the structure and function of Bt proteins and sheds light on their mode of action, especially regarding the insect receptors. Publishing papers focusing on the steps that remain blurred within the mode of action of all Bt insecticidal proteins, including the three-domain Cry proteins, was one of the main goals. The role of receptors such as cadherin, ABCC2, and ABCA2 on the toxicity of Bt proteins in different lepidopterans has been investigated in three different papers [6–8]. In addition, other steps in the mode of action (that comprises protein solubilization, activation, binding, oligomerization, and pore formation) have also been addressed. Examples of these steps include the involvement of a novel trypsin protein for toxin activation in *Plutella xylostella*, discovered after studying a Cry1Ac resistant strain [9], and the promotion of oligomerization of the activated Cry1Ia with insect brush border midgut vesicles, in vitro [10]. The toxicity-promoting effect of a Bt chitin-binding protein that binds to the insect peritrophic matrix has also been studied [11]. Moreover, the Special Issue includes a paper highlighting the synergistic mosquitocidal activity of the parasporal Cry and Cyt proteins present in *B. thuringiensis* ser. *israelensis* [12], and it also includes a manuscript focused on deciphering the amino

acid residues important for the interaction of Cyt2A protein with membrane lipids, a binding step necessary to exert its cytolytic action [13].

The vegetative insecticidal proteins (Vip3) secreted by *Bacillus thuringiensis* are nowadays considered as the new generation of insecticidal Bt toxins because of their different structural and molecular properties regarding the classical Bt 3-D Cry proteins. Vip3 toxins have been already introduced in Bt-crops to control lepidopteran pests. However, little is known about their mode of action. In the Special Issue, five papers analyze different aspects of its biology. They cover aspects ranging from its crystal structure [14] and structural–functional domain analyses [15] to different aspects in the mode of action, such as a study of a possible receptor (the alkaline phosphatase) in a resistant strain [16], the role of oligomerization in toxicity [17], and the study of intracellular events promoted by Vip3A intoxication in *Spodoptera frugiperda* Sf9 cells [18].

In summary, the Special Issue brings together papers of important scientific value in the field of Bt. The review and research papers included will help keep readers up to date on the topic and, at the same time, will contribute to increasing the vast knowledge of Bt and its insecticidal proteins. These studies will help to provide useful information for the development of new strategies to fight against pest insects, in the least aggressive and harmful but better environmental scenario.

**Funding:** This research received no external funding.

**Acknowledgments:** The editors are grateful to all of the authors who contributed to this Special Issue "*Bacillus thuringiensis* Toxins: Functional Characterization and Mechanism of Action". Special thanks to the peer reviewers for their expertise evaluations, which have contributed to increasing the quality of the research works and reviews compiled in this Special Issue. Finally, we thank the MDPI management team and staff for their valuable contributions, organization, and editorial support.

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**


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© 2020 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).

### *Obituary* **A Tribute to a** *Bacillus thuringiensis* **Master: Professor David J. Ellar**

#### **Susana Vílchez**

Institute of Biotechnology, Department of Biochemistry and Molecular Biology I, Faculty of Science, University of Granada, 18071 Granada, Spain; svt@ugr.es

Received: 27 November 2020; Accepted: 29 November 2020; Published: 3 December 2020

This Special Issue, on *Bacillus thuringiensis* and its toxins, seems to be the right place to pay tribute to one of the most influential scientists in the field of research into this peculiar bacterium. Professor David J. Ellar passed away at his home in Cambridge, UK, on 21 May 2020, aged 80, as a result of pneumonia, after a long and debilitating illness. Everyone who knew him and had the opportunity to work hand in hand with him remain devastated by his loss.

Professor Ellar developed a very relevant scientific career in the Department of Biochemistry at the University of Cambridge. Through his laboratory, commonly known as Skylab, passed literally hundreds of postdoctoral researchers, visiting professors, PhD students, master's students, and undergraduate students. We all received the best training possible on *B. thuringiensis*, Cry toxins, their receptors, and above all on how cutting-edge and quality science is conducted. Professor Ellar has been a referent in the field of *B. thuringiensis* for all researchers working with this special bacterium and its entomopathogenic toxins. Thanks to his more than 160 research papers, published in the best scientific journals, we all know today a little bit more about this microorganism, which has proved to be extremely useful in the area of biotechnology.

Professor Ellar was a pioneer in *B. thuringiensis* research. Thanks to him, we were able to "see" for the first time the three-dimensional structure of a Cry toxin [1]. He was also the first to show us what a Cyt toxin, another entomotoxin produced by *B. thuringiensis*, looked like [2]. Thanks to his research we learned about the existence of the first Cry toxin receptor described [3], present on the enterocyte membrane of an insect, further identified as the well-known APN receptor [4]. This created an opportunity to begin understanding the mechanism of action of Cry toxins. In addition, he was the first to relate the Domain II Loops of a Cry toxin to specificity [5], explaining why some Cry toxins are active against some insects and not against others. He was the first person who managed to successfully display a functional Cry toxin on the surface of a phage [6], opening the possibility of using phage display technology for the in vitro evolution of Cry toxins.

However, David not only stood out for being a brilliant professional. On a personal level, he was simply a great person. He was always willing to help to anyone in need. He was extremely generous, and he had an outstanding sense of humour. He also had the remarkable ability to keep a huge research group motivated, in which each of its members worked with the precision of a Swiss watch.

I have the pleasure to say that David was a real master for me: the person that I have as a model in life and the person I want to become when I grow older. With this humble letter, and on behalf of the *B. thuringiensis* research community, I would like to give you the most sincere thanks for all your work and knowledge, and for your human greatness. Rest in peace.

**Funding:** This research received no external funding.

**Acknowledgments:** I would like to thank the Guest Editors Yolanda Bel, Juan Ferré and Patricia Hernández-Martínez for letting me include this tribute in this Special Issue.

**Conflicts of Interest:** The author declares no conflict of interest.

#### **References**


**Publisher's Note:** MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

© 2020 by the author. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).

## **The Tripartite Interaction of Host Immunity–***Bacillus thuringiensis* **Infection–Gut Microbiota**

#### **Shuzhong Li, Surajit De Mandal, Xiaoxia Xu and Fengliang Jin \***

Laboratory of Bio-Pesticide Innovation and Application of Guangdong Province, College of Agriculture, South China Agricultural University, Guangzhou 510642, China; shuzhongli@stu.scau.edu.cn (S.L.); surajit\_micro@scau.edu.cn (S.D.M.); xuxiaoxia111@scau.edu.cn (X.X.) **\*** Correspondence: jflbang@scau.edu.cn; Tel.: +86-20-85280203; Fax: +86-20-85280293

Received: 28 June 2020; Accepted: 7 August 2020; Published: 12 August 2020

**Abstract:** *Bacillus thuringiensis* (Bt) is an important cosmopolitan bacterial entomopathogen, which produces various protein toxins that have been expressed in transgenic crops. The evolved molecular interaction between the insect immune system and gut microbiota is changed during the Bt infection process. The host immune response, such as the expression of induced antimicrobial peptides (AMPs), the melanization response, and the production of reactive oxygen species (ROS), varies with different doses of Bt infection. Moreover, *B. thuringiensis* infection changes the abundance and structural composition of the intestinal bacteria community. The activated immune response, together with dysbiosis of the gut microbiota, also has an important effect on Bt pathogenicity and insect resistance to Bt. In this review, we attempt to clarify this tripartite interaction of host immunity, Bt infection, and gut microbiota, especially the important role of key immune regulators and symbiotic bacteria in the Bt killing activity. Increasing the effectiveness of biocontrol agents by interfering with insect resistance and controlling symbiotic bacteria can be important steps for the successful application of microbial biopesticides.

**Keywords:** *Bacillus thuringiensis*; antimicrobial peptide; gut microbiota

**Key Contribution:** This review focused on describing the tripartite interaction of host immunity, Bt infection, and gut microbiota.

#### **1. Introduction**

The Gram-positive bacterium *Bacillus thuringiensis* (Bt) and its toxins are used to control several orders of insects, including agricultural pests and pathogen vectors [1,2]. Due to their selective insecticidal activity, *B. thuringiensis* toxins have become the most widely used commercial biopesticide worldwide [3,4]. Besides, the isolated Bt toxin genes have also been expressed in several transgenic Bt crops, and these strategies have reduced reliance on chemical pesticides [5–7]. The most common virulence factors of Bt are the crystal (Cry) toxin proteins produced during the sporulation phase of its growth cycle when ingested by susceptible insect larvae. The Cry toxins solubilize in the gut and are further activated by the host gut protease. The active fragments cross the peritrophic membrane and bind to the protein receptor located on the brush border membrane of midgut epithelial cells and create pores that induce osmotic cell lysis and subsequent death [8–10].

The widespread use of Bt spray products in high-value horticulture and the large-scale cultivation of Bt transgenic cotton and maize has resulted in cases of field resistance in several lepidopteran pest species and the western corn rootworm, *Diabrotica virgifera virgifera* [11,12]. The most common factors associated with Bt resistance are alteration of the Bt toxin receptors' binding site, mutations, and altered expressions of the midgut receptor genes [12–14]. Several Bt Cry toxin receptors, such as aminopeptidase-N (APN), alkaline phosphatase (ALP), cadherin, and ATP-binding cassette transporter (ABC transporter), have been identified and characterized in the midgut membrane of the insects [9,15,16]. *B. thuringiensis* resistance has also been linked to several other factors, such as inactivation of the midgut protease required for processing the Bt protoxins [17], gut stem cell proliferation, and differentiation [18]. However, the host immune response and the function of the gut microbiota during Bt infection, which are important aspects of Bt research, has still been inadequately studied and remains controversial [19–22].

The insect's innate immune system consists of both humoral and cellular immune responses, which depend on the non-self recognition of microbes and the subsequent production of immune effectors [23]. The humoral immune response of insects includes the induction of antimicrobial peptides (AMPs), lysozymes, and the rapidly activated phenoloxidase (PO) cascade-mediated melanization [24]. AMPs are produced by two major immune pathways, Toll and IMD (immune deficiency). These pathways produce and regulate the expression of AMPs that are specific to either Gram-positive bacterial/fungal and Gram-negative bacterial infection, respectively [25,26]. The insect cellular immune process consists of encapsulation, nodulation, and phagocytosis, which is primarily driven by the hemocyte [27]. Recognition by pattern recognition receptors (PRRs) triggers immune signal transduction, and results in the activation of the Toll, Imd, Janus kinase/signal transducer and activator of transcription (JAK/STAT), c-Jun N-terminal kinase (JNK), and prophenoloxidase (PPO) pathways [28–30]. Furthermore, insects possess both midgut-specific and systemic immune responses to combat the infection, and reactive oxygen species (ROS) production mediated by dual oxidase (DUOX) is another inducible immune defense mechanism of insects [23].

The insect gut microbiota includes not only the bacterial community but also fungi, protists, and archaea, although the bacterial species dominate in the gut microbial community [31], and plays a vital role in insect development, nutrition, immunity, metabolism, and colonization resistance against pathogens [32–37]. Several factors, such as environmental habitat, host, developmental stage, and diet, play a significant role in the structure and function of the insect gut microbiota [38,39]. It has also been reported that Bt toxicity is often associated with the abundance of the gut microbiota. The lepidopteran pest *Spodoptera exigua* can tolerate the action of Bt toxin when it contains an increased midgut microbiota load [40]. Native gut microbiota can also stimulate the host immune system. It has been reported that the native gut microbiota of bee is associated with the upregulated expression of AMPs, such as apidaecin and hymenoptaecin [41]. This indicates that gut microbiota could help the host to maintain an appropriate immune level, and play a vital role in the survival against Bt toxicity. In this review, we describe the tripartite interaction between host immunity, Bt infection, and gut microbiota (Figure 1). We discuss the effects of Bt infection on the host immune response and intestinal microbes, and how gut microbiota responds to Bt toxicity or causes Bt resistance, as well as the mechanism used by the host to limit Bt infection while maintaining intestinal homeostasis.

**Figure 1.** The tripartite interaction model between host immunity, Bt (*Bacillus thuringiensis*) infection, and gut microbiota.

#### **2. The Interaction between Host Immunity and Gut Microbiota**

#### *2.1. Native Gut Microbiota-Induced Host Immune Response*

It is well known that the host insect immune system is stimulated upon immune challenge by invading pathogens [23]. The local gut immunity plays a vital role in maintaining gut homeostasis by inhibiting or removing invading pathogens and limiting the growth of the symbionts [42]. Native

gut microbiota participates in various symbiotic interactions and also affects host immunity, but the relationship between native gut microbiota and host immune function has so far been less well studied. Kwong et al. (2007) reported that the expression of the antimicrobial peptides (AMPs) is highly upregulated in gut tissue having normal gut microbiota in comparison with the gut tissues deficient in the microbiota. They suggest that the native microbiota may induce an immune response in the host [41]. Intriguingly, colonization of one specific intestinal bacteria, *Snodgrassella alvi,* to the host alone does not induce AMPs expression. However, colonization of another gut symbiont *Frischella perrara* results in a strong host immune response involving the upregulation of AMPs and the genes associated with the melanization cascade [43]. It suggests that different microbial species may have a different regulatory function in the host. Similar to the above results, the gut commensal microbiota of Red palm weevil (RPW), *Rhynchophorus ferrugineus* Olivier can help to protect against pathogenic infection by priming the immune system [44], and the colonization of gut commensal microbiota could enhance the immunocompetence of the host. Futo et al. (2016) reported that *Tribolium castaneum* larvae with less microbiota load showed a decrease in survival rate upon immune challenge by Bt [45], which indicates that gut microbiota is essential for immune priming.

Another important aspect is the messengers of immune priming between hosts and microbes, which explains why different gut commensal bacteria showed a different effect on host immunity and physiology. It has been reported that peptidoglycan and uracil, which are released from intestinal commensal bacteria, can induce AMPs gene expression and ROS production to maintain the gut homeostasis [46–49]. Growing evidence revealed that messenger molecules not only involve peptidoglycan and uracil but also contain numerous bioactive compounds, such as short-chain fatty acids (SCFAs), choline metabolites, and lipids [50–52]. Furthermore, it has been confirmed that the axenic population of *D. melanogaster* has altered lipid metabolism and insulin signaling, but the host physiology can be restored after the administration of the gut microbial metabolite acetate [53]. A recent study found that gut microbiota may also affect the systemic immune response apart from gut immunity in Red palm weevil (RPW) *Rhynchophorus ferrugineus* Olivier larvae [44]. They might derive some metabolites, which can cross the gut epithelium and enter the host hemolymph. However, more studies are required to decipher this mechanism.

In addition to immune priming to defend microbial pathogen infection, gut commensal bacteria-mediated immune responses are also crucial for efficient arboviral acquisition in mosquitoes. Intestinal symbiotic bacterium *Serratia* Y1 has been shown to inhibit successful establishment of the *Plasmodium* through direct activation of the mosquito immune response [54], and gut microbiota could elicit a protective immune response against the *Plasmodium* transmission [55]. In contrast, *Serratia* J1, another *Serratia* strain isolated from field-caught mosquito, has no impact on *Plasmodium* development [54]. Likewise, different strains of the same bacterial species have a different effect on *Plasmodium* infections in the *Anopheles* mosquito midgut [56]. It further reminds us that the interaction between host and gut commensal bacteria is complex and may involve strain-specific outcomes according to the corresponding metabolites.

#### *2.2. Multiple Immune Reactions Help to Maintain Gut Homeostasis*

The insect immune system not only protects the host against pathogen infection but also regulates the colonization of symbiotic microorganisms in the gut to maintain host homeostasis [57]. Several interesting mechanisms contribute to the proper maintenance of the microbiota by balancing the complex interaction between the host and the microbiota, which is mainly under the control of Toll and IMD pathways, and dual oxidase (DUOX) pathways, respectively (Figure 2) [23,58]. However, the functions of the Toll pathway are not consistent in different insect species, e.g., the Toll pathway is not found to be associated with the regulation of local gut immunity in *D. melanogaster* [59]. Recent reports by Abrar et al. found that the Spatzle-mediated Toll-like signaling pathway could regulate the homeostasis of gut microbiota by mediating the synthesis of AMPs in Red palm weevil, *Rhynchophorus ferrugineus* Olivier [60]. Royet et al. (2011) reported that the Toll signaling pathway could also be

activated in the midgut of *P. xylostella* larvae by oral ingestion of pathogenic microbes. They also found that several essential elements for the Toll signaling pathway, including Spatzle, Toll receptor, tube, pelle, cactus, and dorsal, were expressed in *P. xylostella* midgut after the infection (Figure 2) [61]. Both Toll and IMD pathways can be activated following the detection of peptidoglycan (PGN) released from bacteria by different peptidoglycan recognition proteins (PGRPs). The family of PGRP is one of the key modulators in this process, which coordinates between the host immune response with the gut commensal bacteria. Similar to invading pathogens, gut commensal microorganisms can produce many immune-activating compounds (such as peptidoglycan) during growth and proliferation. A total of seven PGRPs were identified in *D. melanogaster* that can degrade peptidoglycan into non-immunostimulatory muropeptides. With the help of amidase activity, the peptidoglycan released from intestinal bacteria was maintained at a low basal level, so that the host can avoid the overactivation of the Toll and IMD pathway by gut microbiota (Figure 2) [62]. It has also been revealed that the low levels of peptidoglycan were limited by the PGRPs with amidase activity to transfer across the epithelial barrier and reach into hemolymph to stimulate the systematic immune response [63].

**Figure 2.** Insect gut immunity protects against infections and maintains gut microbiota homeostasis. DAP-type peptidoglycan (PGN) from intestinal bacteria is sensed by PGRP-LC, which triggers the IMD-dependent MEKK1-MKK3-p38 DUOX-expression pathway. Uracil also activates MEKK1-MKK3 p38 in a PLCβ-dependent manner; the activation of p38 enhances the transactivating function of ATF, which in turn activates the transcription of dual oxidase (DUOX). On the other hand, PLCβ-calcium signaling is responsible for the induction of DUOX enzymatic activity. Both contribute to the production of reactive oxygen species (ROS) in the gut lumen, where they control endogenous and infectious bacteria [64]. DAP-type PGN recognition by PGRP-LC also triggers the IMD pathway through the translocation of the nuclear factor-κB (NF-κB) family member Relish, which then induces increased transcription of antimicrobial peptides (AMPs) genes [23]. Besides, the IMD pathway has established a negative feedback loop to prevent overactivation. One is the members of the PGRP family gene (PGRP-LB or PGRP-SC) with amidase activity can cleave PGN and therefore blocks the activation of the

IMD pathway. Another is Pirk, which interferes with the plasma membrane localization of PGRP-LC [62]. In some insect species, the Toll signaling pathway is activated with the Lys-type PGN recognition by PGRP-SA or PGRP-SD after microbial infection. This initiates a proteolytic cascade that ultimately cleaves pro-Spatzle into an active ligand for Toll, leading to the activation of the NF-κB-like transcription factors dorsal and then translocation into the nucleus to induce increased transcription of the AMP gene. Finally, these immune regulatory networks cooperatively help to maintain gut homeostasis.

Similarly, the PGRP-LB homolog with amidase activity also acts as a negative modulator in the immunity of Red palm weevil, *Rhynchophorus ferrugineus* Olivier, in which abnormal expression alters the abundance and community structure of gut microbiota [65]. The intracellular protein Pirk can prevent PGRP-LC from recognizing extracellular peptidoglycan, thereby preventing hyperactivation of the gut immune response in flies [66]. Besides, it has been shown that peritrophic membrane (PM) integrity is related to the gut microbiota homeostasis in *A. stephensi* [67] and that PGRP-LD can help the PM to maintain structural integrity by preventing overactivation of the gut immune response, in turn limiting *P. berghei* infection. The knockdown of PGRP-LD can increase gut immunity and alters the gut microbial spatial distribution, which results in the dysbiosis of the gut microbiota. It suggests that PGRP-LD acts as a negative regulator of the immune signal pathway.

Research was also conducted to study other immune pathway regulators for maintaining gut homeostasis. Relish is an important regulator gene of the IMD pathway. Silencing the expression of Relish in the model insect *G. mellonella* results in a significant increase in the concentration of gut bacteria and decreases in the expression of AMPs [68]. Similar findings were also reported in Red palm weevil, which showed a compromised ability of pathogen clearance and increased gut bacterial load after silencing the Relish expression [69]. Moreover, a change in the gut commensal microbiota was observed after the inhibition of Caudal, a transcription repressor of NF-κB-mediated expression of AMPs [70]. The elimination of gut microbiota through antibiotics results in the downregulation of the IMD pathway and AMP gene expression [68]. Collectively, these results indicate that the IMD pathway plays a vital role in maintaining gut microbiota homeostasis.

The production of reactive oxygen species (ROS) is another inducible defense mechanism in the gut in addition to AMPs production (Figure 2) [23]. ROS are produced by the DUOX protein with an N-terminal extracellular peroxidase domain, which can convert H2O2 into HOCl in the presence of chloride, and thereby are detoxified in the presence of IRC catalase [58,71]. Unlike the gut IMD pathway, it is the uracil nucleobase, not peptidoglycan (PGN), that acts as an agonist to induce DUOX-dependent ROS production [48]. However, DUOX cannot be activated by most of the symbiotic bacteria under natural conditions, which suggest that symbiotic bacteria may block their uracil secretion pathway under natural conditions, and initiate it under specific dysregulated gut environments [48]. DUOX is also involved in the regulation of gut permeability in *Anopheles gambiae* [72]. The knockdown of DUOX increases the overall bacterial load in the oriental fruit fly *Bactrocera dorsalis;* however, the relative abundance of the bacterial symbionts *Enterobacteriaceae* is decreased in the gut [73].

#### **3. The Host Immune System in Response to Bt Infection**

Insects can initiate humoral and cellular immune responses to reduce the damage caused by Bt infection [74–80]. It has been reported that Bt tolerance in the flour moth, *Ephestia kuehniella*, can be achieved by the preexposure of low-concentrated Bt endotoxins (Syngenta, North Ryde, NSW, Australia) [81]. This phenomenon is mostly denoted as immune priming, which implies that the primary exposure to pathogen activates the basic immune response result in an improved immune response upon second exposure [82–84]. Therefore, the mechanism of Bt toxicity does not only depend on the host receptor but is also associated with the elevated immune response of the host. It has also been reported that Bt endotoxin-tolerant *E. kuehniella* larvae can increase the lipid carrier lipophorin in the gut lumen, which inactivates Bt toxins through the aggregation of lipophorin particles to break down toxins into coagulation products [85]. A soluble toxin-binding glycoprotein is also found in the intestinal lumen of the Bt (Cry1Ac)-resistant larvae of the lepidopteran pest *Helicoverpa armigera*, which can bind to Cry1Ac and GalNAc-specific lectins and forms an insoluble aggregate [78]. An LC5 dose of Bt ssp *galleria* strain 69-6 can trigger phagocytic activity in the larvae of *Galleria melonella*, whereas an LC15 dose of Bt increases the encapsulation rate in the hemolymph during infection [86]. Similarly, both the LC15 dose and LC50 dose of Bt resulted in elevated hemolymph phenoloxidase, and lysozyme-like activity in Bt-infected *Galleria mellonella* larvae. However, the difference is that low doses of Bt can increase the humoral and cellular immune response, involve an increased encapsulation response, and enhance the phagocytic activity of hemocytes. However, a higher dose decreases cellular reactions, involving the coagulation index and activity of phenoloxidase in hemocytes [77]. The host's immune response to Bt is likely dose dependent, as a sublethal dose of Bt damages the mid-intestinal epithelial cells, but it can be repaired by stem cell proliferation, and the enhanced immune response of the hemocytes can help to limit further infection and prevent septicemia [87]. At the LC50 dose for Bt, the situation is generally different, as the symbiotic bacteria and destroyed intestinal cells lead to dysfunctional humoral and cellular immune reactions. This indicates that Bt infection not only stimulates the local immune response in the gut but also induces the systematic immune response, where the radiation of immune pathway activation begins at the site of initial infection and radiates out, reaching the hemolymph. We found the LC50 dose of Bt infection could suppress the humoral immune response in the third instar larvae of *Plutella xylostella* [80]. Growing evidence suggests that Bt-induced immunity is a dose-dependent effect [88–91].

Comparatively, less information is available on the intestinal melanization response during Bt infection. It can be assumed that hemocytes can be recruited to seal perforations in the site of intestinal damage, and melanization may play a key role in this process. It has been reported that plasma phenoloxidase (PO) activity can be induced by both low and high concentrations of Bt in *G. melonella* and *E. kuehniella* larvae [77,81]. The prophenoloxidase (PPO) of insects comes from different sources and performs diverse functions, such as wound repairing, protection against pathogens, catalyzing, and detoxifying phenolics in the diet [92–94]. Several studies have shown that the PO may come from the hemolymph of the adult mosquito midguts [95,96]. However, other studies have shown that PPO is secreted into the foregut and can be activated by gut proteinase to detoxify phenolic present in the diet of Lepidoptera [92,93]. A recent study revealed that the PPO cascade is triggered after the infection of the Bt strain (Bt8010) in the midgut of *P. xylostella* larvae, which involves pattern recognition receptors (PRRs) and genes encoding proteases and protease inhibitors in the PPO cascade [94]. PPO can also be secreted into the hindgut to clean fecal bacteria by induced melanization of feces [92]. Similarly, the melanization response was reported in the hindgut of *Drosophila* mutant species [97]. PO-mediated melanization in the midgut might prevent symbiotic bacteria from escaping into the hemocoel through damaged midgut epithelial cells. However, despite various scientific investigations, the origin of the PO remains controversial in insects [98].

*B. thuringiensis* also synthesize another insecticidal protein (Vip) during the vegetative growth phase [99], and Vip3A, Cry1, and Cry2 genes have pyramided in cotton and maize to control lepidopteran insects [100]. The resistance to Vip3A has been selected in several lepidopteran species under laboratory conditions [101,102], while little is known of the biochemical mechanisms of resistance to Vip3A. Studies have shown this toxin does not share binding sites with Cry1 or Cry2 toxins [103,104], and in a laboratory-selected population of *Heliothis virescens*, resistance to Vip3A was shown to confer little cross-resistance to Cry1Ab and no cross-resistance to Cry1Ac [101]. The genome-wide analysis showed that most of the immune response genes, including AMPs, were upregulated, and genes involved in the metabolism and digestion process were downregulated in *Spodoptera exigua* larvae in response to Vip3 insecticidal challenge [105,106]. Similar to Bt and Bt Cry toxins, Vip3A toxin also triggers the PPO cascade and upregulates most of the genes involved in the midgut melanization process of *S. litura* and *S. exigua* [74,105]. Vip proteins also have a dose-dependent effect on the host. An increasing concentration increases the number of upregulated genes involved in the immune system and hormone modulation, and the downregulated genes involved in peritrophic membrane stability and the digestion process [106]. The genome-wide microarray analysis of Vip3Aa toxin-treated beet armyworm, *Spodoptera*

*exigua*, showed that the upregulated enriched genes are involved in innate immune response, such as AMPs and *repat* genes [105]. This information helps to understand the host insect immune response after Bt Vip protein toxin challenge.

It is well known that the interaction between Cry toxin and toxin receptors from the host midgut brush border membrane vesicles (BBMVs) is the initial step in the insecticidal activity of the Cry protein toxins [107–110]. Several researchers also showed the interaction between midgut immune-related proteins and Cry toxin [111–113]. There is evidence showing that immune-related protein like Dorsal and peroxidase C in the midgut juice of *P. xylostella* and *S. exigua* can bind to the Cry1Ab1 protein toxin [111]. The protein Dorsal plays a significant role in the insect immune system, especially in the Toll pathway; therefore, a possible insecticidal mechanism of Bt Cry1Ab1 mediated by the midgut immune-related protein can be proposed.

Similarly, it has been reported that C-type lectin-20 (CTL-20) in *Aesdes aegyptii* has the potential to bind to both toxin receptors and Cry toxins to affect the interactions between Cry toxins and toxin receptors to reduce Cry toxicity in *Aedes aegypti* [112]. Similarly, another study on immune-related peptidoglycan recognition protein (PGRP) gene expression and PO activities in Cry1Ac-susceptible and -resistant *P. xylostella* found that the resistant strain of *P. xylostella* had higher PO activity compared with the susceptible strain [114]. Moreover, among three different *P. xylostella* strains, a Cry1Ac-susceptible, a Cry1Ac–resistant strain, and a field strain, both PGRP1 (belong to PGRP-SA family) and PGRP3 (PGRP-LF) showed higher expression levels in the gut of susceptible strains compared to the resistant strain and field strains, and PGRP2 (PGRP-LB) showed the highest expression levels in the gut of resistant strains [114]. It has been found that Cry1Ah toxins can bind directly to the PPO proteins in *Ostrinia furnacalis*[115]. The interaction between Cry protein toxins and the host midgut immune-related proteins requires further investigation as the study progresses.

The above studies have greatly enriched our knowledge of the host immune response after Bt or Bt toxin infection, and it is now confirmed that Bt or Bt toxin protein can affect the host's immune system in a dose-dependent manner. It is known that the insect gut harbors a diverse indigenous microbiota, and the host immune system plays an important role in maintaining the gut homeostasis [42]. In the next section, we discuss the intestinal microbiota and its functions when the immune system of the insect host has been compromised by Bt or Bt toxins.

#### **4. The Interaction between Bt and Host Gut Microbiota**

#### *4.1. B. thuringiensis Infection Altered Host Insect Gut Microbiota*

In general, insects maintain a balanced local intestinal microbial community that plays a vital role in their host, including host development, nutrition, and tolerance against pathogens [116,117]. It has been shown that gut microbiota is also associated with the resistance against Bt SV2 in mosquito and Bt HD-1 in Indian meal moth, *Plodia interpunctella* (Hübner) [118,119]. The diversity and richness of gut microbiota are changed by pathogen infection. However, relatively few studies have been published on the effects of Bt or Bt toxin infection on host gut microbiota. An investigation into mosquito larvae exposed to time increasing doses of Bt showed that the lowest diversity of gut microbiota comes from the most tolerant mosquito larvae [88]. Interestingly, the same study also found that the most tolerant larvae had the highest inter-individual difference. Similarly, *B. thuringiensis* infection significantly reduced the diversity and abundance of the gut microbiota in the Bt-resistant line of *G. mellonella* [120]. However, honeybees feeding on transgenic Cry1Ah maize pollen did not result in significant changes in the gut microbiota community composition under laboratory conditions [121]. Intestinal epithelial cells act as a barrier to separate the microbiota of midgut and hemocoel, and the microbial composition differs between these two tissues under normal conditions. However, the bacterial profile between the gut and hemocoel has been reported to be similar following treatment of *Spodoptera littoralis* larvae with an LC50 dose of Bt Cry1Ca toxin [122]. This indicates that gut bacteria cross the intestinal barrier to

hemolymph as a result of the Bt toxin infection and then reproduce in the insect hemolymph. However, further research is needed to reveal the action mechanism of Bt on host gut microbiota.

#### *4.2. The Function of Gut Microbiota in Response to Bt Infection*

The effect of insect gut microbiota in Bt toxicity has long been controversial. In 2006, Broderick et al. reported that midgut microbiota is required for Bt subspecies *kurstaki* insecticidal activity in the larvae of the gypsy moth, *Lymantria dispar*. They also found that Bt *kurstaki* was unable to multiply in insect hemolymph in vitro, indicating that intestinal bacteria cause septicemia and contribute to Bt toxicity, but without Bt, intestinal bacteria cannot induce death [21]. However, several studies showed contrasting results. In 2009, Johnston reported that intestinal bacteria were not responsible for Bt HD73 strain toxicity in the tobacco hornworm, *Manduca Sexta* [22]. Interestingly, Bt HD73 Cry− cells can grow rapidly in plasma after intra-hemocoelic inoculation in many species. The same year, another study also confirmed that midgut microbiota is not required for the pathogenicity of Bt HD-73 and Bt HD-1 strains in the larvae of *P. xylostella* [123]. Work on the same insect host found that inoculation with one isolated gut bacteria *Enterobacter* sp. Mn2 has a different effect on Bt HD-1 and Bt HD-73 strain pathogenicity [123]. The contrasting results of different Bt strain pathogenicity after inoculation with the same gut bacteria to host insects indicate that we need to have an in-depth knowledge of different Bt strains before designing the experiment. From such studies, it is clear we need to pay attention to the host insect gut bacterial community, whether different diets and environments cause different gut bacteria communities between species or diverse populations within one species, and how it influences the interaction between Bt and host gut microbiota.

The interaction between Bt and gut microbiota can be competitive. *B. thuringiensis* can produce bacteriocin to inhibit the growth of gut bacteria [124]; on the other hand, insect gut microbiota can inhibit Bt multiplication, growth, and alteration of its toxins [118,125,126]: This is a kind of competition relationship. Conversely, Bt and host gut microbiota also show beneficial interactions to a certain extent; for example, some intestinal bacteria species can produce proteases that help solubilize Bt protoxins to their active form [127]. Furthermore, *B. thuringiensis* infection can promote translocation of gut-opportunistic pathogenic bacteria to hemocoel, which relies on gut epithelial damage caused by Bt toxins or some other factors, and then rapidly reproduce in the hemocoel and participate in host septicemia, finally leading to the death of the host [20,128]. One study showed that hemolymph microbiota are changed dramatically and the change is dominated by *Serratia* and *Clostridium* species upon Bt infection in *Spodoptera littoralis* larvae, which switch from asymptomatic gut symbionts to hemocoelic pathogens [122]. This translocation phenomenon agrees with the hypothesis discussed earlier in Section 4.1 of the present review.

Many gut symbiotic bacteria have been isolated and characterized; some of them showed a beneficial effect on the host and can be called probiotic. Such bacteria are widely used as animal feed additives in food production [129]. *Enterococcus mundtii* bacteria isolated from the feces of *Ephestia kuehniella* have the function of protecting the flour beetle, *Tribolium castaneum*, against Bt infection [130]. The surface properties test showed that this isolate has intense levels of auto-aggregation, which is related to the formation of colonies in the host insect's gut [131]. Moreover, bacteria cell wall compounds, such as lipopolysaccharide (LPS) and peptidoglycan (PGN), have been well studied, and can stimulate the host immune response [132,133]. However, *Tribolium castaneum* larvae exposed to the corresponding supernatant can also increase the resistance to Bt infection [130]. This perhaps suggests that the protective function of probiotic bacteria is based on the secreted proteins or some small peptides, which may act directly against Bt infection or through the triggering of immune priming.

Similarly, another *E. mundtii* strain isolated from *S. littoralis* also showed a protective function for the host insect, which can directly inhibit competitors and suppress pathogens' growth through its antimicrobial activity [134]. Previous studies confirmed that *E. mundtii* cells accumulate on the surface of the intestinal epithelium and form a biofilm-like structure, which helps to keep its predominant colonization status in the host insect's gut [135]. Additionally, after removing the dominant bacteria from the gut, this resulted in increased susceptibility of the spruce budworm larvae to Bt infection [20]. Although growing evidence confirmed that intestinal bacteria play important roles in the host defense response [136,137], the molecular function mechanism requires more research. It has been reported that *E. mundtii* SL can secrete a kind of bacteriocin, which strongly inhibits some of the competing organisms and can impair pathogen colonization in vivo [134]. Through this, we can speculate that Bt may also shown inhibited growth and has limited activity in the insect gut lumen, and the battle between Bt and probiotic *E. mundtii* may depend on the dose effect. It is also noteworthy that the secreted bacteriocin showed a selective antibacterial activity and has no influence on other intestinal bacteria, and as a result, the gut microbiota can develop normally. There is another study report that normal gut microbiota mediated pathogen clearance from the gut lumen [138]; this suggests that the gut microbiota can act as another form of protection response in organisms, or at least an important complement to host gut immune protection.

Insect gut microbiota also play an important role in Bt infection indirectly through intestinal epithelium cell regeneration. The Lepidoptera larvae intestinal epithelium mainly includes four kinds of cell types: Columnar cells, goblet cells, enteroendocrine cells, and intestinal stem cells. Each cell type has a specific role and helps maintain normal gut functions. The intestinal stem cell is the only type capable of division, which mediates epithelial renewal and the healing response [139]. *B. thuringiensis* infection can disrupt the gut epithelial cells by producing toxins, and insects mount a series of defensive responses, which involves melanization, AMP-mediated antimicrobial activity, and gut stem cell proliferation and differentiation in response to gut damage [18,94]. It has been shown that REPAT and MAPK p38 signaling pathways may be involved in the regulation of the gut defensive response to Bt toxins [18,140]. The REPAT gene was also predicted to be associated with a regenerative response in Bt-resistant insect species and showed constitutively increased expression in a Bt-resistant *S. exigua* [141]. Moreover, it showed that Cry1Ac resistance is related to an enhanced midgut healing response in the tobacco budworm [142,143]. These results suggest that intestinal stem cell activity is associated with Bt resistance because bacteria cannot get through the healthy midgut epithelial cells. This allows the host to quickly repair the damage, resulting in a limited number of invaders into the hemocoel.

It has been reported that indigenous gut microbiota can modulate the activity of intestinal stem cells in *Drosophila*, which correlates with the activated JAK/STAT pathway and epidermal growth factor receptor (EGFR) pathways [33,144]. Both the rate of epithelium renewal and the number of dividing intestinal stem cells were reduced after removing all the intestinal bacteria. The abundance of gut microbiota can also be increased after host immune suppression [145]. Interestingly, the number of mitotic intestinal stem cells is increased after blocking the *Drosophila* IMD pathway, which caused an abnormal intestinal microbiota, and many genes related to stem cell proliferation and differentiation were also upregulated by the induced gut microbiota [144]. It indicated that the intestinal stem cell proliferation could be stimulated by increased gut microbiota. In summary, these results demonstrate that insect gut microbiota can affect Bt resistance by mediated intestinal stem cell activity.

#### **5. Conclusions**

*B. thuringiensis* infection induces a variety of host immune responses, and interferes with the gut microbiota of the host. The resulting dysbiosis, in turn, stimulates both the expression of AMPs and the production of ROS by different ligand molecules. DUOX also plays a key role in regulating the gut microbial homeostasis. The interaction between the host immune system and gut symbionts is more cooperative rather than antagonistic. However, *B. thuringiensis* or other pathogenic infections can cause dysregulated gut environments in insects, which makes it possible to convert some symbiotic bacteria into pathobiont, known as an opportunistic pathogen. The above interaction relationship has an important effect on Bt pathogenicity or toxicity. Most studies have focused on the interaction between Bt and the host immune system or the interaction between Bt and microbiota, which significantly expands our knowledge about the dynamic Bt infection process. However, a few important aspects are

still unanswered and need to be explored, i.e., (i) How does Bt trigger the immune signaling pathway? (ii) Do the membranes of the intestinal lumen or intestinal epithelial cells have any toxin recognition receptors attached to the Toll and IMD pathways? (iii) Why do different intestinal bacteria have a different effect on the host during Bt infection? and (iv) which bacterial metabolite plays a significant role in Bt toxicity and host immunocompetence?

**Author Contributions:** Conceptualization, F.J. and S.L.; software, S.L.; writing—original draft preparation, S.L.; S.D.M.; F.J. and X.X.; writing—review and editing, S.L.; F.J.; S.D.M. and X.X.; supervision, F.J.; funding acquisition, F.J. All authors have read and agreed to the published version of the manuscript.

**Funding:** This work was supported by a grant from the National Natural Science Foundation of China (31972345), Natural Science Foundation of Guangdong, China (2019A1515011221,2020A1515010300), Provincial Agricultural Science and technology innovation and Extension project of Guangdong Province (2019KJ147).

**Acknowledgments:** We would like to thank anonymous referees for their valuable comments and suggestion.

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**


© 2020 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).

## **Current Insights on Vegetative Insecticidal Proteins (Vip) as Next Generation Pest Killers**

#### **Tahira Syed** †**, Muhammad Askari** †**, Zhigang Meng, Yanyan Li, Muhammad Ali Abid, Yunxiao Wei, Sandui Guo, Chengzhen Liang \* and Rui Zhang \***

Biotechnology Research Institute, Chinese Academy of Agricultural Sciences, Beijing 100081, China; syedtahira98@gmail.com (T.S.); 2017Y90100082@caas.cn (M.A.); mengzhigang@caas.cn (Z.M.); liyanyan01@caas.cn (Y.L.); abid@caas.cn (M.A.A.); weiyunxiao@caas.cn (Y.W.); guosandui@caas.cn (S.G.)

**\*** Correspondence: liangchengzhen@caas.cn (C.L.); zhangrui@caas.cn (R.Z.); Tel.: +86-10-82106127 (R.Z.)

† These authors contributed equally.

Received: 7 July 2020; Accepted: 11 August 2020; Published: 14 August 2020

**Abstract:** *Bacillus thuringiensis* (Bt) is a Gram positive soil bacterium. This bacterium secretes various proteins during different growth phases with an insecticidal potential against many economically important crop pests. One of the important families of Bt proteins is vegetative insecticidal proteins (Vip), which are secreted into the growth medium during vegetative growth. There are three subfamilies of Vip proteins. Vip1 and Vip2 heterodimer toxins have an insecticidal activity against many Coleopteran and Hemipteran pests. Vip3, the most extensively studied family of Vip toxins, is effective against Lepidopteron. Vip proteins do not share homology in sequence and binding sites with Cry proteins, but share similarities at some points in their mechanism of action. Vip3 proteins are expressed as pyramids alongside Cry proteins in crops like maize and cotton, so as to control resistant pests and delay the evolution of resistance. Biotechnological- and in silico-based analyses are promising for the generation of mutant Vip proteins with an enhanced insecticidal activity and broader spectrum of target insects.

**Keywords:** *Bacillus thuringiensis*; vegetative insecticidal proteins; insecticidal activity; resistance; pyramids

**Key Contribution:** This review addressed the recent advances in the structure, function, and mode of action of vegetative insecticidal proteins (Vip), and explored the strategies applied so far to modify the Vip proteins for an enhanced insecticidal activity.

#### **1. Introduction**

Common soil bacterium *Bacillus thuringiensis* (Bt) is a Gram positive motile bacterium that gained significant popularity over the past decades because of its role as an invertebrate pest killer [1]. *B. thuringiensis* has been extensively studied for its ability to produce immense arsenal of toxins with an insecticidal potential against insect vectors of human diseases, agricultural pests, nematodes, fungi, gastropods, and protozoans [1–5]. Bt proteins, as an active ingredient of biopesticides, are a valuable eco-friendly approach to replacing chemical insecticides. Bt genes are engineered in many crops like maize, cotton, soybean, and rice, offering a sustainable solution to control insect pests [6]. A Bt-transformed cotton plant expressing Cry1Ac was first introduced in 1996 in Australia and the United States of America [7]. The agriculture land covered by transgenic Bt crops reached 98.5 million hectors in 2016 [8]. Nevertheless, the rise of insect resistance is becoming a major hurdle in the commercialization of Bt transgenic crops [9].

*B. thuringiensis* produce Crystal (Cry) and Cytolytic (Cyt) toxins during sporulation, which are stored in parasporal crystalline inclusions and released after the disintegration of the cell wall in a culture medium. However, vegetative cells produce non-crystalline toxins, such as vegetative insecticidal proteins (Vip) and secreted insecticidal proteins (Sip), secreted as soluble proteins in a growth medium. Vip proteinaceous toxins were isolated from the culture medium of both *Bascillus cereus* and *B. thuringiensis* after screening [10,11]. The Vip toxin family is classified into four subfamilies, namely Vip1, Vip2, Vip3, and Vip4, based on their amino acid similarity, which also guides Vip proteins' nomenclature [10,12]. The Bt Toxin Nomenclature Committee assigned a four-rank name to each toxin—primary rank (two toxins with 45% similarity), denoted by an Arabic number; secondary rank (<78% similarity), denoted by an uppercase letter; tertiary rank (95% similarity), denoted by a lower case letter; and quaternary rank (>95% similarity), denoted by the final number (Figure 1).

To date, 15 Vip1 proteins, 20 Vip2 proteins, 111 Vip3 proteins, and 5 Vip4 proteins have been reported and named [13]. (Vip1 and Vip2 heterodimer toxins are effective against insects from Coleopteran and Hemipteran orders [14]. The largest family, Vip3, is effective against many species of Lepidoptera, and crops like cotton and maize have been successfully transformed with various Vip3 toxins [15]. Interestingly, Vip proteins have no sequence homology with Cry proteins, and do not share common binding sites in target insects [16–19]. This makes them ideal toxins to be used in combination with Cry proteins in insect resistance management (IRM) programs.

Significant knowledge about the structure and mode of action is available for Cry proteins, but this information is still rudimentary for Vip toxins. This review discusses recent insights on the structure and mechanism of action of Vip toxins. Detailed knowledge about the structure and functional characterization of Vip toxins will lead to the development of new strategies for designing improved toxins against insects that have developed resistance. The Vip family of Bt toxins is a potential candidate against these resistant pests. This study also focused on the natural and in vitro evolution of Vip toxins, and the strategies developed so far that improve the insecticidal activity of these toxins at a molecular level.

**Figure 1.** Schematic representation of vegetative insecticidal proteins (Vip) proteins' nomenclature system. Each protein is assigned a four ranked name—primary rank, given to proteins sharing less than 45% homology in amino acid sequences; secondary and tertiary ranks, with less than 78% and 95% similarity, respectively; and finally, the quaternary rank is given with more than 95% identical proteins.

#### **2. Structure and Function of Vip Toxins**

Vip proteins are widely distributed among Bacillus species. These proteins are not produced in parasporal crystalline inclusions, but are instead secreted into the culture medium. Like other Bt toxins, Vip proteins are also inactive in their native form, and are activated after being secreted in the membranes of insect midgut cells through the action of enzymes [20]. Detailed structural information of the Vip proteins is not yet elucidated; therefore, the current structural and functional predictions are based on in silico and mutagenic studies. A comparison between the Vip family of proteins is summarized in Table 1.


**Table 1.** Comparison of Vip family proteins.

#### *2.1. Structure of Vip1*/*Vip2 Binary Toxins*

Vip1 and Vip2 act as binary toxins possessing an ADP ribosyltransferase activity [21]. The genes of the Vip1 and Vip2 proteins are located in a ~4 kb single operon with different reading frames [12,22]. To date, 35 *Vip1* and *Vip2* genes have been listed in the Bt nomenclature database. Sequence analysis found that Vip1 is synthesized as a protoxin of 100 kDa, with an N terminal signal peptide sequence of 35 amino acids. Similarly, Vip2 proteins in protoxin form are 52 kDa in size, with an N terminal signal peptide of 50 amino acids [22,23]. After their modification at the N terminal signal peptide, Vip1 and Vip2 are transformed into a mature proteins of 80 kDa and 45 kDa, respectively [24]. The structural analysis of Vip2 has confirmed two domains, N-terminal (Nt) and an NAD-binding (Nicotinamide adenine dinucleotide) C-terminal (Ct) domain [15,25]. Moreover, X-ray crystallography revealed homology between the N and C terminal domains of the Vip2 protein, and both domains are formed by the perpendicular packing of five mixed β sheets, with one flanking α helix and three anti parallel β sheets with three flanking α helices [25].

Research based on their sequence homology suggests that Vip1 and Vip2 act as binary toxins of A + B type, with similarities to many mammalian toxins. Vip1 has very little structural similarity with the *Clostridium spiroforme* toxin, protective antigen of *Bacillus anthracis*, and CdtB toxin of *Clostridium dificile*. Vip2 has a structural similarity to the active domain of CdtA toxin produced by *C. difcile* [26]. Both Vip1 and Vip2 toxins have a similarity to the C2 toxin of *Clostridium botulinum* and the domain Ia of the iota toxin produced by *Clostridium perfringens* [27]. Altogether, this predicts Vip2 as an ADP-ribosyltransferase toxin inhibiting the polymerization of actin filaments, causing cytoskeleton abnormalities and insect cell death [28]. Vip1 is inferred to act as a B toxin (Binding domain) responsible for the translocation of Vip2 inside insect midgut cells. Vip2 is a cytotoxic A toxin with a binary toxin response, showing no toxicity to insects when applied alone [28].

#### *2.2. Structure and Function of Vip3 Proteins*

Vip3 toxin is also produced during the vegetative growth phase of Bt, and Vip3A is the most widely studied Vip toxin so far. Vip3 is a diverse family of toxins with 95% similarity between its members, and it shares no primary sequence homology to any other Vip families or Bt toxins [29]. These proteins show a strong inhibition of insect larval growth at a low concentration [29,30], and

the structural differences among the Vip3 members predict a broader mode of action against a wide spectrum of insects.

The Vip3 gene encodes a protein of 89 kDa having 787 to 789 amino acids [29]. Functional characterization of the *vip3Aa16* gene revealed that its −35 and −10 promoter regions have homology to the *Bacillus subtilis* promoters, which suggests that the *Vip3* gene promoter is under the control of the σ<sup>35</sup> holoenzyme [31]. It shares no sequence homology with any other toxins produced by *B. thuringiensis* [1]. Phylogenetic analysis showed that the Vip3 protein belongs to distant clade rather than to Cry toxins.

A comprehensive structure of Vip3A has not yet been resolved, and is only derived through in silico modeling [32,33]. Notably, the Vip3A signal peptide located at the N-terminal is responsible for the translocation of protein. The N-terminal region is highly conserved and performs important regulatory insecticidal functions [34], while the C-terminal region undergoes various modifications and has the ability to target insect specificity. At present, the role of both domains in insecticidal activity is mainly perceived by mutagenic studies. The deletion or addition of amino acids at the Nand C-terminals greatly affects the entomocidal property of a particular protein. For example, the deletion of amino acids at the Nt region causes a negative effect on the insecticidal activity of the Vip3A protein [35,36]. Substitutions at position T167 or G168 at the N-terminal of Vip3Af with alanine leads to a decreased insecticidal activity [32]. In contrast, the deletion of 200 amino acids from the N-terminal enhanced the toxicity of Vip3Aa against various Lepidoptera pests, and the deletion of 200 amino acids from the C-terminal region abolished the insecticidal activity of Vip3BR. Also, the deletion of 127 amino acids at the C-terminal maintained a low level of insecticidal activity against *Agrotis ipsilon* (Lepidoptera: Noctuidae) and *Helicoverpa armigera* (Lepidoptera: Noctuidae) [37], which suggests the important roles of the N- and C-terminal parts in the insecticidal activity of Vip3 proteins. It is also possible that the interactions of other plant or insect proteins with Vip3 C- and N-terminal domains could enhance their insecticidal activity. Hence, further studies are needed to explore the exact mechanism of action.

Each C-terminal amino acid plays an important role in the target specificity and toxicity against many Lepidopteron pests. Meanwhile, Vip3A11 mutants are generated after replacing nine residues at the C-terminus with Vip3A39 residues by site-targeted mutagenesis. Here, the cysteine residue CYS784 of the C-terminal region is found to be a crucial trypsin cleave site for bioactivity and toxicity [38]. However, the Vip3 C-terminal region alone does not possess an insecticidal activity, as the expression and purification of the C-terminal region of Vip3Ab1 and Vip3Bc1 cause no harm to the insects. Therefore, in contrast with the Cry proteins, both the C- and N-terminal regions are important for oligomerization and proteolytic stability, with a significant contribution to the toxicity of the Vip3 proteins [39].

Site directed mutagenesis anticipates putative trypsin cleave sites Lys195, Lys197, and Lys198 inside Vip3Aa. The mutants generated by replacing these three Lysine residues with alanine lose sensitivity to trypsin or midgut juices (MJ), and also show toxicity against *Spodoptera exigua* (Lepidoptera: Noctuidae) [40]. In the same manner, the substitution of cysteine with serine at the C-terminal also reduces the Vip3A7 protein insecticidal activity against *Plutella xylostella* (Lepidoptera: Plutellidae), possibly due to the disruption of disulfide bonds between cysteine residues [41]. An alanine scanning analysis of 588 residues unveiled a five-domain structure of the Vip3Af1 protein and its role in toxicity. This approach revealed 50 residues with a significant impact on Vip3Aa structural conformation and toxicity. Among them, two clusters of 19 substitutions, located near the N-terminus region between Leu167–Tyr27 or on the C-terminus between Gly689–Phe741, abolished toxicity to *Agrotis segetum* (Lepidoptera: Noctuidae). Another 19 substitutions also reduced toxicity to *Spodoptera frugiperda* (Lepidoptera: Noctuidae). Hence, it is evident that each amino acid within the Vip3 protein plays a diverse role in protein stability and toxicity [32]. Transmission electron microscopy and single particle analysis of Vip3Ag4 have revealed the surface topology of its tetramers. After trypsin treatment, the

protein forms an octamer containing tetramers of 65 kDa and 22 kDa fragments. In addition, the tetrameric form and main topology are retained even after trypsin treatment [42].

Although no clear evidence about the Vip3 3D structure is available, in silico analyses point to the presence of five domains in the Vip3Af protein. The trypsin fragmentation of alanine mutants depicted five domains in the Vip3Af proteins structure. Domain I spans from amino acids 12 to 198, domain II 199 to 313, domain III 314 to 526, domain IV 527 to 668, and domain V 669 to 788 amino acids. Domains I to III are necessary for tetramerization, however not domain V. In addition, the role of domain IV remains unclear [43]. This evidence is further supported by the 2D structure of the Vip3B protein predicted by X-ray diffraction studies. According to this model, Vip3B is composed of five domains; two domains carrying α helices (DI and DII) at the N-terminus and three β sheet containing domains (DIII, DIV, and DV) on the C-terminal region. Domain III shows a slight resemblance to domain II of the Cry4Aa and Cry1Ac proteins. Domain IV and V share homology with carbohydrate binding modules (CBM), indicative of glycosylated receptor-binding inside the midgut [44]. A carbohydrate binding motif (CBM; CBM\_4\_9) has also been identified at the C-terminal region in all of the Vip3 protein members, except for Vip3Ba. It is then possible that the C-terminal region plays a crucial role in recognition and binding to midgut receptors [45].

A recent report sheds more light on the structure of all five domains of Vip3A and their related function. Domain II has two highly conserved hydrophobic α helices, predicting their role in membrane insertion and pore formation inside the insect midgut. Domain III comprises three β sheets potent for cell binding, along with domain II, persistent with previous results. In Vip3A, two CBM domains with different glycan binding pockets are found in C-terminal region, which forms domain IV and domain V [46]. This indicates a specificity in their binding capacity with glycan on the targeted cell surface. The cryo-EM (cryogenic electron microscopy) structural analysis solves the structure of Vip3A and showed how the toxin forms pores in the insect midgut. The protein architecture has five distinct domains in protoxin form, with domains I (coiled α 1–α 4) ending at the primary protease cleavage site, and domain II having five α helices mainly producing the core before trypsin digestion. Antiparallel β sheets of domain III form a β prism fold analogous to the Cry toxins. In the Cry toxin, this fold assists in receptor recognition. Similar to previous results, two CBM folds form the last two domains. After trypsinization, all four monomers stay connected, and no conformational change takes place in domains II to V. Three N-terminal α helices form a parallel four helix coiled coil, which forms a long dipole to lodge the ions in its cavity. Its dimension has the ability to form pores in the lipid bilayer [47].

#### *2.3. Vip4 Toxins*

Vip4 is the least characterized toxin of the Vpb class. Only five Vip4 proteins have been identified to date. The first reported Vip4 toxin was Vip4Aa1 (now named Vpb4Aa1), isolated from Bt strain Sbt009, with no insecticidal activity against any pests. Its molecular mass is ~108 kDa and it possesses 965 amino acids [13]. The main region spanning from 47 to 77 amino acids is a PA14 domain, and the region from 218–631 residues is named the bacterial Binary\_ToxB domain. This novel protein shares 34% identity with the Vip1Aa1 protein and 65% with the Ia domain of the Iota toxin of *B. cereus*, specifically to the B component of the binary toxin [26].

#### **3. Modern Classification of Vip Proteins**

Recently, the classification and names of the Vip family of toxins were modified by Crickmore et al. [48]. Accordingly, all Vip toxins are placed in three different classes, namely Vip3, Vpa, and Vpb. The Vip3 family mnemonic remains the same. In class Vpa, all of the Vip2 proteins are placed. Class Vpb contains a binary toxin component of Vip2, such as Vip1, and its structural analogues, previously known as Vip4 [13].

#### **4. Mechanism of Action**

The mode of action of the Vip toxins inside insect guts is not yet clear and needs further investigation. Differences in the structure of the Vip and Cry toxins determine the different target sites in the insect midgut, making them suitable candidates for insects-resistant control [16].

#### *4.1. Mechanism of Action of Vip1*/*Vip2 Binary Toxin*

There is no clear mechanism of action for the Vip1/Vip2 binary toxin. Each protein alone is not enough to cause toxicity, but rather they act in combination as A + B binary toxins [28]. This multistep process begins with toxin entry at the insect midgut and ends with larvae death. There are many proposed models on this mechanism. Toxins first enter the midgut and are digested by trypsin-like proteases. Enzymatic action by trypsin or midgut juices (MJ) cleaves Vip1Ac into its activated form before entering into the brush border membrane (BBM). After enzymatic activation, Vip1Ac oligomerizes to form seven monomers containing a multimeric structure [49], which binds the BBMVs of cotton aphids with a high target specificity [14]. Similar results were observed with the activated form of the Vip1Ad and Vip2Ag binary toxin activity. Vip2Ag binding, insecticidal activity, and toxicity is increased tremendously in the presence of Vip1Ad in *Holotrichia parallela* (Coleoptera: Scarabaeidae). This shows strong evidence for Vip1 as a potential receptor of Vip2 after trypsin activation [50].

After internalization, Vip2 transfers the ADP ribose group to the actin filaments, inhibiting its polymerization. This ultimately leads to abnormal microfilament formation and cell death [15,23]. The insecticidal activity of Vip2Ag and Vip1Ad is characterized by the vacuolization and destruction of BBMVs and microvilli depletion, similar to what has been observed in *H. armigera* fed on Cry2Ab and Vip3AcAa/Cry1Ac binary toxins [51,52]. In a nutshell, the histopathological effects of Vip1/Vip2 binary toxins are comparable to those of Cry proteins.

#### *4.2. Mechanism of Action of Vip3 Toxin*

The Vip3 toxins mechanism of action has some similarity to the Cry toxin-like protease activation, binding with midgut cells and pore formation. The complex multistep process is not fully elucidated yet. At first, all Vip3 toxins are activated by midgut juices. Proteolytic analysis of Vip3A toxin has revealed several fragments (62–66 kDa, 45 kDa, 33 kDa, and 19–22 kDa) with a similar pattern after trypsin treatment or insect midgut proteases. The main cleavage product from the C-terminus region is a 62–66 kDa core peptide, generally considered to be the main part of the activated toxin [19,37,53,54]. However, a 19–22 kDa fragment comprising the 199 amino acids at the N-terminus, together with a 62–66 kDa fragment, are crucial for lethality, as shown by bioactivity assays after purification [43]. Furthermore, 45 kDa and 33 kDa fragments are formed after cleavage of a 62–66 kDa fragment [53]. Other research on protease cleavage inferred that these smaller fragments formed after the digestion of 65 kDa fragment could be the result of denaturing conditions of SDS-Page, as the C-terminal (65 kDa fragment) domain remains intact under native conditions [55]. In contrast, the proteolytic digestion of Vip3Ca produces a 70 kDa fragment [56]. Variations in the insecticidal activity of the Vip3 toxin seem to depend on the hydrolysis pattern inside insects' midgut [54], however, the proteolytic digestion pattern of Vip3Ab1 and Vip3Bc1 in a non-susceptible insect, *Ostrinia nubilalis* (Lepidoptera: Crambidae), was identical to the susceptible insects, and shed light on the fact that resistance might be unrelated to protein cleavage [39].

After the trypsin processing of Vip3A, two of its fragments, 19–22 kDa of the N-terminal region and 62–66 kDa fragment of the C-terminal region join to form a ~360 kDa homo-tetramer, which cannot be degraded by proteases [57]. Moreover, another study proposed the formation of a >240 kDa complex and identified a novel site, S164, crucial for the formation and stability of this complex. Mutations at this site lead to a loss of insecticidal activity [58]. In addition, the two fragments are eluted together in gel permeation chromatography, emphasizing the fact that they may remain together

after cleavage [55]. The interaction between the 22 kDa and 65 kDa fragment is necessary for their stability and toxicity [39].

The next phase is the binding of the activated toxin with the BBMVs inside the midgut of susceptible insects. The Vip3 protein binding sites do not overlap with the Cry proteins. A clear mechanism regarding the binding of the Vip3 toxin is still not available, and only a few studies have addressed the recognition of binding molecules inside the insect midgut cells. So far, only a few Vip3 binding proteins have been identified, one of which is ribosome S2 protein, identified by yeast hybrid assay, and confirmed by vitro pull-down assays in sf21 cells of *S. frugiperda*. A knock down of the ribosomal S2 gene in the Sf21 cells and the larvae of *Spodoptera litura* (Lepidoptera: Noctuidae) resulted in a reduced larvicidal activity, considering that S2 is one of the proteins involved in the Vip3 insecticidal mechanism [59]. Another potential protein is a 48 kDa tenascin-like glycoprotein, which strongly binds to Vip3Aa in BBMVs from black cutworm [36,60].

Many current studies are focused on the recognition of Vip3 potent receptors in order to understand the mechanism of action. A novel receptor, fibroblast growth factor receptor-like protein (Sf-FGFR), has been identified on the membrane of Sf9 cells, as a result of its binding affinity toVip3, confirmed by in vitro analysis. Silencing of the *Sf-FGFR* gene resulted in a reduced toxicity of Vip3Aa to Sf9 cells. The localization of Sf-FGFR and Vip3Aa on the surface and then inside the cytoplasm suggests that binding takes place on the surface, leading to internalization [61]. Similarly, Vip3Aa has shown a strong interaction with scavenger receptor class C like protein (Sf-SR-C) in both in vivo and in vitro analysis with Sf9 cells of *S. frugiperda*. Knocking down the expression of these receptor genes results in a reduced mortality of Sf9 cells and *S. exigua* larvae to Vip3Aa [62]. Further studies are required to fully clarify the specificities of the Vip3 toxin receptor binding.

Proteolytic activation and receptor mediated binding of Vip3 toxins leads to pore formation and cell death. After feeding Vip3 toxins, the insect midgut is damaged, which is proposed as the main target site for the Vip3 toxin. Histopathological analyses have revealed similar symptoms to Cry toxins, like swollen or lysed midguts and pore formation [19,63]. However, clear mechanisms on toxin intercellular localization and pore formation in BBM are not available. The localization mechanism of active the Vip3Aa protein inside Sf9 cells, by laser scanning confocal microscopy with fluorescently labeled Vip3Aa (Alexa488-actVip3Aa), has demonstrated that Vip3Aa is not internalized by the endocytic- or clathrin-dependent endocytic pathway. Instead, this seems to happen through receptor mediated endocytosis, after which the Vip3Aa protein interacts with various cytosolic proteins (e.g., ribosomal S2 protein) [64].

Voltage clamping and planar lipid bilayer experiments predict the ability of a toxin to form discrete ion channels without involving receptors, which differs from Cry1Ab [16]. Pore formation in lipid bilayers by an activated toxin inside the midgut has been reported in H. armigera using the florescent quenching method [65]. Additionally, the maximum potential of the activated Vip3Aa toxin to form pores has been seen at specific pHs during in vitro analyses, showing that pore formation only happens at acidic or neutral pH [57]. The mechanism of Vip3 pore formation and virulence is generally the most accepted (Figure 2).

Contrary to the pore formation model, is another mechanism in which the Vip3 toxin induces apoptotic cell death in insects. The intercellular localization of Vip3 causes abnormalities of cell division and leads to the apoptosis of insect midgut cells. Vip3A treated Sf9 cells undergo arrest at the G2/M phase and the disruption of the mitochondrial membrane potential (ΔΨm), leading to apoptotic cell death via the sf-caspase-I mediated pathway [62]. Another study has evidenced the involvement of regulatory proteins and lysosomes in apoptosis. Furthermore, symptoms of apoptosis and mitochondrial collapse are prevalent in sf9 cells when administered Vip3Aa, such as the accumulation of reactive oxygen species (ROS), caspases (caspase 3,9), and cytochrome c [66] (Figure 3). Because of the lack of sound information, more investigation is needed to clarify the downstream mechanism of Vip3 induced apoptosis and cell death.

**Figure 2.** Proposed mechanism of pore formation by the Vip3 toxin. The Vip3A toxin is activated by proteolysis inside the insect midgut. In the next step, activated toxins, including 22 kDa and 65 kDa fragments, bind with receptors, leading to pore formation in the insect midgut cells and, ultimately, to the death of the insects.

To understand the mechanism of insect response to toxins, the transcriptomic and proteomic characterization of genes and proteins is of great interest. The gene expression profiles of toxic-dose-treated larvae of *S. exigua* and *S. litura* have been analyzed in two independent studies. From the analysis of the transcriptome profile in *S. exigua* larvae, >29,000 unigenes were obtained, in which the S2 and tenascin-like protein gene expression, was stable. The up regulated genes were mostly related to immune reposes and defense mechanism while down regulated genes were mainly metabolic ones [67]. Similarly, the genes coding for lysosomes and antimicrobial peptides have been found to be up-regulated in *S. exigua* [68]. In the gene expression profile of the Vip3 toxin treated larvae of *S. exigua*, immune response genes are up-regulated and the genes involved in the digestion process are down regulated. The up regulation of initiator and effector caspases genes and antimicrobial effectors provides strong evidence for the apoptosis of insect cells, similar to previous reports [69]. In another analysis, 56,498 unigenes were identified in *S. litura* larvae. The transcription levels of the trypsin related genes increased in this case after toxin induction, which supports the role of trypsin in the metabolism of the Vip3Aa toxin [70]. However, further investigation is necessary to elucidate how Vip3 toxins cause apoptotic cell death.

**Figure 3.** Schematic representation of the mechanism of Vip3 toxin induced apoptotic cell death of insect midgut cells. Vip3A protoxin binds with receptors, and the receptor mediated internalization of toxin takes place. Toxin internalization leads to changes like DNA damage, mitochondrial membrane disruption, and the activation of caspases (caspase 3 or 9), in turn promoting apoptotic cell death.

#### **5. Insecticidal Activity of Vip Proteins**

After research on various insect species, it has been found that Vip1/Vip2 has an insecticidal activity against some pests of Coleopteran and Hemipteran orders [14]. Vip1 and Vip2 act in combination, and none of the toxins have an insecticidal activity when administered alone. The combination of Vip1Aa/Vip2Aa or Vip2Ab has been found to be affective against Diabrotica spp [12]. The Vip1Ad/Vip2Ag toxins, when combined and expressed as a binary toxin, show toxicity against *H. parallela*, *H. oblita,* and *Anomala corpulenta* (Coleoptera: Scarabaeidae) [24].

The most extensively studied Vip3 protein family is Vip3Aa, which is widely known for their insecticidal activity against many species of Lepidopteron and pests like *S. exigua*, *H. armigera*, *S. frugiperda*, *Heliothis virescens* (Lepidoptera: Noctuidae), *Helicoverpa zea* (Lepidoptera: Noctuidae), and *A. ipsilon*. Although they have very minor differences in their sequences, Vip proteins exhibit great variability in their targeted insects. For instance, one of the most recently discovered members of the Vip3 family protein, Vip3Ca, has 70% homology with Vip3Aa and has been found to be toxic against *Chrysodeixis chalcites* (Lepidoptera: Noctuidae), *Mamestra brassicae* (Lepidoptera: Noctuidae), and *Trichoplusia ni* (Lepidoptera: Noctuidae). However, the Vip3Ca toxin shows a moderate insecticidal activity against *Cydia pomonella* (Lepidoptera: Tortricidae; non-susceptible to the Cry toxin) and *O. nubilalis* (susceptible to the Cry toxin) [71]. Vip3Ca is the most potent toxin against *Mythimna separate* (Lepidoptera: Noctuidae), with an LC50 value 3.4 μg/g. This toxin could be used in future maize crop protection to control Oriental armyworm [72]. Vip3Ca is also more toxic to *Ostrinia furnacalis* (Lepidoptera: Crambidae), which is more similar to Cry1Ab than Vip3A, and can be an effective candidate against Cry1Ab-resistant colonies of *O. furnacalis* [73].

Likewise, Vip3Ae and Vip3Af are toxic to *Spodoptera littoralis*(Lepidoptera: Noctuidae), *M. brassicae,* and *Lobesia botrana* (Lepidoptera: Tortricidae), and Vip3Ab is lethal against *A. ipsilon*. Nonetheless, Vip3Ad exerts no toxicity to insects like *H. armigera, M. brassicae, S. frugiperda, S. exigua, S. littoralis,* and *A. ipsilon* [29]. The insecticidal activity of Vip3Aa59 is significantly higher than Vip3Aa58 towards

*Dendrolimus pini* (Lepidoptera: Lasiocampidae) [20], and Vip3Aa45 shows a 40-fold higher toxicity than Cry proteins against *S. exigua* [26].

In some reports, the toxicity of the Vip3 protoxin was found to be more than active toxin, e.g., Vip3Ae protoxin is more insecticidal than the active toxin when compared with Vip3Aa [54]. These could result from differences in the protocols for protein isolation and purification, bioassay conditions, and quantification methods. For example, metal chelate chromatography clearly affects the insecticidal activity of Vip3 [20,54].

#### **6. Evolution of Resistance and Cross Resistance to Vip3 Toxins**

Since the application of Vip3 transformed Bt crops, only few cases of practical resistance have been reported. The reported cases of resistance are from both laboratory and field selected insects. For example, in laboratory conditions, insects of *S. litura* show a 280-fold resistant to Vip3A after 12 generations of selection. This is probably due to the lower protease activity in those insects [74]. Similarly, the laboratory selection of *H. virescens* with Vip3Aa for 12 generations results in a >2040-fold resistance. This resistance is polygenic and decreases after 13 to 28 generations, without toxin administration. A lack of cross resistance is also seen against Cry toxins [75].

In a laboratory evolved resistance study, conducted with transgenic maize expressing Vip3Aa20 in Brazil, the target pest, *S. frugiperda,* showed a >3200-fold resistance. The pattern of resistance inheritance was autosomal recessive and monogenic, with a very low frequency of resistant alleles (0.00009 estimated by the F2 screening method) [76]. A Vip3A resistant strain of *S. frugiperda* evolved >632-fold resistance to Vip3A, with minor cross resistance to Cry1F, Cry2Ab2, or Cry2Ae toxins [77].

Despite the fact that Vip3A expressing crops are not yet commercialized in Australia, a high frequency of resistant alleles has been observed in various studies conducted on *H. armigera* and *Helicoverpa punctigera* (Lepidoptera: Noctuidae). In a study screening for resistance alleles in *H. armigera* and *H. punctigera* using the F2 method, a natural polymorphism and very high baseline frequency of 0.027 and 0.008, respectively, were observed, with no cross resistance to Cry1Ac or Cry2Ab. The presence of both resistance alleles on the same locus confirmed resistance to be recessive [78]. Another biochemical study also found a high frequency of resistant alleles in the same insect, *H. armigera,* with no significant change in the binding of Vip3 toxins to BBMVs compared with susceptible insects. Instead, a low proteolytic activity was found in resistant insects. [79].

Despite the intrinsic specificities of Vip3 toxicity, no cross resistance to Vip3C has been observed in insects of different species, previously found to be resistant to Cry1A, Cry2Ab, Dipel (Mixture of Cry1 and Cry2). Vip3C shows cross resistance to *H. armigera* colonies resistant to Vip3Aa or Vip3Aa/Cry2Ab, and toxicity against *O. furnacalis,* which is nonsusceptible to Vip3A [73]. The biochemical basis of resistance could not be established by the down regulation of membrane bound alkaline phosphatase (mALP) isoform HvmALP1, observed in Vip3 resistant insects and results does not support it to be the functional receptor of Vip3. In addition, mALP can be used as a marker for the detection of Vip3A resistance [80]. Moreover, Cry1F and Vip3A do not share common binding sites in *S. frugiperda* [17], and also lack cross resistance [54].

#### **7. Identification of Bt Isolates Containing** *Vip Genes*

Over the past few decades, researchers have been trying to find new *B. thuringiensis* isolates from different geographical regions and diverse environments, to develop new toxins with a high insecticidal potential and to cope with resistance. The discovery of new isolates not only helped in the production of new pesticides of a wide insecticidal spectrum, but also in overcoming insecticidal resistance [81]. *B. thuringiensis* strains were isolated from diverse habitats, like milk and mossy pine cone [82], soil, leaf [83] and insect cadavers [84], and goat gut [85].

After the characterization of native *B. thuringiensis* isolates isolated from soil, and fig leaves and fruits from a Turkish collection, a new *B. thuringiensis* isolate, 6A, was identified carrying a high expression of Vip3Aa. The identified protein, named Vip3Aa65, has a similar insecticidal activity against *Grapholita molesta* (Lepidoptera: Tortricidae) and *H. armigera,* but is less toxic to Spodoptera spp. compared with Vip3Aa16 [86]. In another study, two Bt strains, BnBt and MnD, were found to produce Vip proteins in isolates of great potential with a high toxicity (LC50 = 41.860 ng/μL of BnBt and 55.154 ng/μL of MnD) in the second instar larvae of *S. littoralis* [87]. Lone and co-workers isolated and expressed a novel Vip3Aa61 gene in *Escherichia coli* from isolate *B. thuringiensis* JK37. Nucleotide analysis found differences in many amino acids compared with Vip3A. Because of its high insecticidal activity (LC50 = 169.63 ng/cm2) against second instar larvae of *H. armigera*, the Vip3Aa61 toxin is a potential candidate for transgenic crop production and pest protection [88].

Bacterial 16S ribosomal RNA (rRNA) sequencing is frequently used to characterize microbes at a species rank. In the past, this method was not successful because of the close resemblance of *B. thuringiensis* to other strains, making it ambiguous for phylogenetic and diversity analysis [89]. A new and more reliable method of phylogenetic analysis is multiple locus sequence typing (MLST). MLST-based analysis of *B. thuringiensis* kurstaki isolates from Assam, India, confirmed the presence of Vip1 (53.3%), Vip2 (46.6%), and Vip3 (40%) genes [90]. The results were in contrast to a previous study, where the Vip3 gene was more abundant than Vip1 and Vip2 [91]. Recently, the characterization of an indigenous Bt strain, found Vip3 gene in this strain, and that the spore crystal mixture of this isolate had a high mortality rate against *S. frugiperda* [92].

#### **8. Transgenic Crops Expressing Vip Proteins**

Despite the high toxicity of Vip1/Vip2 toxins against corn rootworms, Bt maize crops containing these binary toxins cannot be developed because of the cytotoxicity of Vip2 proteins [93].

Vip3 proteins are introduced in crops like cotton and maize. The *Vip3Aa19* gene was first introduced into Bollgord cotton (COT102), expressed as a single insecticidal protein (VIPCOT commercialized in 2008 in Unite States of America). This toxin provided protection against three major cotton pests, *H. virescens*, cotton bollworm *H. zea*, and *Pectinophora gossypiella* (Lepidoptera: Gelechiidae). Later, it was pyramided with Cry genes (modified Cry1Ab) for insect resistance management. Vip3Aa20 was introduced (MIR162) in corn and was commercialized in 2009 in the United States. Later, Vip3 proteins were pyramided with Cry genes (Cry1Ab + Vip3Aa20) and (Cry1Ab, Vip3Aa20, and mCry3A) in corn.

Other than commercialized crops, *Vip3* genes are successfully transformed and the proteins have been expressed in transgenic crops in laboratory-based studies. Transgenic sugar cane lines expressing Vip3A showed a high mortality rate against sugar borer *Chilo infuscatellus* (Lepidoptera: Pyraloidea) [94]. Cow pea is an important food crop in many African countries, and is harmed by Lepidopteron pest, *Maruca vitrata* (Family: Crambidae). The *Vip3Ba1* gene, isolated from Australian Bt isolates, was transformed and expressed in cowpea to provide protection against legume pod borer (*M. vitrata*), by strongly inhibiting larvae growth [95]. Similarly, a transgenic corn event (C008 and C010) expressing Vip3Aa19 has been found to be highly toxic against black cut worm [96]. When a tobacco plant was transformed with the N-terminal deletion mutant of the Vip3BR protein (Ndv200), it acquired full protection against *S. littoralis*, *A. ipsilon,* and *H. armigera* [97]. These results are helpful for future Bt-derived mutant protein transformation in crops.

#### **9. Biotechnological Strategies to Improve Toxicity and Insecticidal Spectrum of Vip3 Proteins**

In vitro directed evolution to increase the insecticidal potential of Vip proteins can be employed with success. The fact that Vip3 toxins share no sequence homology with Cry toxins makes them an ideal candidate for insect resistance management (IRM) programs. Vip3 proteins are found to be toxic against insects that are less susceptible to Cry toxins, such as Lepidopteron [10]. Various strategies can be employed to increase the toxicity and insecticidal spectrum of Vip proteins. For example, sequence or domain swapping to form chimeric Vip toxins successfully enhances toxicity against susceptible and resistant insects.

#### *Genetic Engineering of Vip3A Genes to Form Chimeric Proteins*

To increase the insecticidal spectrum and activity of the Vip3 proteins, these were genetically modified. In this process, the genes were swapped for the construction of protein chimeras, where one protein expresses the sequence or domains of another protein. These novel chimeras have shown great toxicity against resistant and non-susceptible insect pests. The successful implication of a domain swapping method to generate chimeric Cry1 proteins (Cry1Ba/Cry1Ia hybrid) with enhanced toxicity against Colorado potato beetle has already been reported [98]. However, fewer reports on Vip3 chimera formation are available. A chimera of two Vip3 proteins, Vip3AcAa generated by sequence shuffling of Vip3Aa and Vip3Ac, not only had enhanced toxicity against the fall armyworm, but also to European corn borer, against which Vip3Ac was not toxic, even at high concentrations. These new chimeric proteins caused growth retardation in a Vip3A non-susceptible insect *O. nubilalis* [99].

Using domain shuffling, six chimeric proteins were generated by joining fragments of N-terminal, C-terminal, and the central part of the core protein from Vip3Aa and Vip3Ca. Two of these chimeras, in which only Nt domain (Vip3C having Nt domain of Vip3Aa) was shuffled, had shown no effect over stability and solubility. The exchange of the Ct domain in four chimeric proteins resulted in proteins that were insoluble and unstable to trypsin, except for one soluble and stable chimera. Compared with the parental proteins, one chimeric protein formed by shuffling the Nt domain of Vip3C with Vip3Aa showed an enhanced insecticidal activity against *S. frugiperda* (Table 2). Another chimeric protein was highly unstable and formed after shuffling Ct domain of Vip3Aa with Vip3Ca *O. furnacalis*, which is susceptible to Vip3C and non-susceptible to Vip3Aa, and also showed vulnerability against those chimeras containing the Ct domain of Vip3C. Considering the above mentioned observations, it is evident that the Ct domain is involved in the specificity of Vip3 proteins for targeted insects [100].

The administration of a dual toxin with non-homologous mechanisms of action is an effective way to circumvent resistance. For this purpose, chimeric proteins were formed by fusing *Vip* and *Cry* genes sequences. By combining the sequence of the full-length Vip3Aa16 toxin gene with the Nt region of the Cry1Ac activated core, a Vip3A16-Cry1Ac chimeric protein of 150 kDa was generated. The resulting fusion protein toxicity was triggered against the first-instar larva of *Ephestia kuehniella* (Lepidoptera: Pyralidae) in contrast with parental toxin Vip3Aa [33]. Similarly, another successful chimeric protein was made by fusing the nucleotide sequence of Vip3Aa7 and the Nt region of a synthetic toxin Cry9Ca, with an enhanced insecticidal activity (compared with the single parent proteins or a mixture of both) against *P. xylostella* [41].

Even if domain shuffling and sequence swapping are successfully implemented to form new toxin combination with improved insecticidal activity against new or resistant pests, site directed mutations and in silico analyses still provide crucial information necessary to understand protein toxicity. With the help of ever-evolving bioinformatics, it will be possible to better understand the effect of mutations on the mechanism of action of particular toxins against target insects (Table 2).


**2.** Chimeric Vip3 proteins and their toxicity profiles.

**Table**

#### **10. In Silico Analyses for Generation of Mutagenic Vip3 Proteins**

Over the past two decades, researchers have been trying to produce mutagenic proteins with an enhanced insecticidal activity against specific pests. By using computational methodology the analysis of the chimeric protein, Vip3Aa-Cry1Ac (formed by the fusion of the functional regions of Cry1Ac and Vip3Aa), unveiled its enhanced toxicity and broad-spectrum insect control. Molecular docking analysis was performed with five Lepidopteron insect receptors, forming a strong interaction. This new protein is proposed to be the potential toxin for future crop protection against Lepidopteron pests [103].

#### *E*ff*ect of Amino Acid Modifications on Toxicity of Mutant Vip3A*

Mutagenic analyses have been widely utilized to explore the amino acids present at specific sites critical for toxicity. For this purpose, Vip3A11 mutants were generated by replacing nine residues at N-terminus with Vip3A39 residues, using site targeted mutagenesis. An approximately two-fold increase in toxicity for three mutants (S9N, S193T, and S194L) was seen against *H. armigera* larvae compared with Vip3A11. Furthermore, the N-terminal amino acids also played a great role in toxicity and insect specificity against Lepidopteron pests [104].

Similarly, the docking and binding site prediction analysis identified the amino acids Y616, H618, Y619, W552, K557, E627, and Q652 in the Ct region as crucial sites for Vip3Aa toxin binding and insecticidal activity. The insecticidal activity of only one mutant, Y619A, was increased against *H. armigera* and *S. exigua* [38]. In another case, the Vip3Aa protein substitutions at site S164 with alanine or proline resulted in a loss of oligomer formation, and an ablation of the insecticidal potential against *S. litura*. Notably, substitution with threonine resulted in only a 35% reduction in toxicity [58].

A cysteine residue at the C-terminal region, CYS784, is a crucial site for trypsin cleavage and for the formation of the active core for toxicity. Hence, both the C- and N-terminal regions are necessary elements for toxicity. Cysteine to serine substitutions at the C-terminal also reduced the Vip3A7 protein insecticidal activity against *P. xylostella,* likely due to the disruption of the disulfide bonds between the cysteine residues [41]. A modified Vip3Ca protein, ARP150v02, with amendments at eight locations near the N-terminus region, was expressed and purified in *E. coli*. The ARP150v02 protein showed an insecticidal activity against many insects, but a high insecticidal effect against *S. frugiperda* (LC50 = 450 ng/cm2). In contrast, this protein was ineffective against *H. armigera,* even at a high dose. The binding assays revealed that the ARP150v02 protein competes for binding with Vip3Aa in *S. frugiperda* [105]. More studies based on site directed mutagenesis are necessary in order to overcome pest resistance.

#### **11. Synergism and Antagonism in Vip3 and Cry Proteins**

Various studies reported the presence of synergism in Vip3 and other Bt toxin (Cry and Cyt). The coexpression of Vip3Aa and Cyt2Aa in *E.coli* lead to synergism in *S. exigua* and *Chilo suppressalis* (Lepidoptera: Crambidae) [106]. Similar synergism was observed between Vip3A and Cry1Ia in *S. frugiperda*, *Spodoptera albula*, and *Spodoptera cosmioides* (Lepidoptera: Noctuidae) [107]. Another synergistic combination was found between Cry9Aa and Vip3Aa, possibly due to the binding mechanism between two toxins with BBMVs in *C. suppressalis* and *O. furnacalis.* Interestingly, the synergism between Vip3Aa and Cry9Aa mutants was disturbed moderately in *C. suppressalis,* and severely in and *O. furnacalis.* Synergism resulted in an improved toxicity of Vip3Aa and Cry9Aa in *C. suppressalis,* which is a great threat to rice crops in China [108]. Strong synergism was also seen in the Cry1Ab/Vip3Ca protein combination. These can be useful in *M. separate* and *O. furnacalis* control in future pyramided gene stacking [72]. A high rate of synergism was identified in the Vip3Aa and Cry1Ab combination against the neonatal larvae of *S. frugiperda,* without competing for binding sites [109]. In the same study, several other Bt protein combinations, like Vip3Aa/Cry2Ab, Cry1Ab/Cry2Ab, Cry1Ab/Cry2Ab/Vip3Aa, Cry1Ea/Cry1Ca, and Vip3Ca/Cry1Ea, also showed synergism against *S. frugiperda*.

Finally, it is worth mentioning that some toxin combinations show antagonism as well. For example, Vip3Aa showed slight antagonism with Cyt2Aa in *Culex quinquefasciatus* (Diptera: Culicidae) [106], and with Cry1Ia in *Spodoptera eridania* (Lepidoptera: Noctuidae) [107]. These antagonisms could result from direct competition between the CRY and Vip3A toxin for the same binding sites in insect BBMVs [110]. Another study identified many antagonist combinations in various Cry and Vip3 protein pairs, such as Vip3A/Cry1A or Cry1Ca, Cry1Ca/Vip3Aa, Cry1Ca/Vip3Ae, Cry1Ca/Vip3Af, Vip3Af/Cry1Aa, or Cry1A. Vip3A and Cry1Ca showed more antagonism in *S. frugiperda* at LC90 [111]. This knowledge can be helpful in the future for stacking genes in pyramids in order to broaden the insect spectrum, and for managing the evolution of insect resistance.

#### *E*ffi*cacy of Pyramided Vip3 and Cry Proteins*

The past decade marked an increased use of Vip3A with Cry proteins in pyramided crops for broader insecticidal activity and in insect resistance management [93]. The pyramiding of Cry1A and Cry2A with Vip3A is a promising strategy in IRM programs. No cases of cross resistance have been reported yet in pyramided Bt crops. Meanwhile, registered varieties of pyramided Bt cotton and maize containing Vip3Aa19 and Vip3Aa20 are commercialized worldwide [112].

In order to form pyramided Bt rice, a fusion gene (C1V3) was formed by combining truncated Cry1Ab and the full-length Vip3A by a linker, to generate a chimeric protein that could be digested efficiently by trypsin. After digestion into activated fragments, both toxins function just like an individual toxin of Cry1Ab and Vip3A. Transgenic rice with the fusion gene (C1V3) showed insecticidal activity against two major rice pests, *C. suppressalis* and *Cnaphalocrocis medinalis* (Lepidoptera: Crambidae). A high toxin content was seen after two generations in fields, along with disease spots. No difference in phenotypes was seen in the transgenic (A1L3) and control rice plants. Further investigations will clarify the implications of this strategy [113]. Cry1Ac and Vip3Aa are potential candidates for sugarcane protection against *Diatraea flavipennella* (Lepidoptera: Crambidae) and *Elasmopalpus lignosellus* (Lepidoptera: Pyralidae), through pyramided transgenic Bt sugarcane production and commercialization [114].

The main threat to the commercialization of these crops is the existence of resistance to Cry1A or Cry2A, which may spark the evolution of resistance to these pyramided crops [112]. To overcome this barrier, other strategies could be implemented, alone or by pyramiding them with Bt toxins, in order to control resistant pests. For this purpose, post transcriptional gene silencing of insects specific genes involved in various physiological functions could be effective to inhibit insect growth and development. In this method, double stranded RNAs (dsRNA) are designed to target essential insect genes, disrupting their expression by RNA interference (RNAi). Short sequences of dsRNA are incorporated into insects through diet and are also transformed in plants [115,116]. The chances of cross resistance are very low in this case, as both pathways have diverse and independent mechanisms of action.

Pyramided Cry toxins and RNAi corn plants targeting *D. v. virgifera* have already been developed [117]. Another pyramid formed by the combination of Bt toxins, Cry1Ac and dsRNA, designed to target the metabolism of juvenile hormone (JH) in *H. armigera*, was introduced in cotton. Two types of cotton plants, JHA (targeting JH acid methyltransferase) and JHB (JH transporter protein), showed a high activity against resistant *H. armigera* [118]. For the safety and high efficiency of this strategy, dsRNA can be transformed into plastids to be expressed with plastid genomes rather than a nuclear genome. The chloroplast transformation of dsRNA also increases the protein content in the cell, as RNAi machinery is absent in the chloroplast compartment. Introducing dsRNA into potato plastids targeted the β-actin gene of the deadly potato pest, Colorado potato beetle, and protected the crop against this notorious pest [119] (Table 3).


**Table 3.**


**Table 3.** *Cont.*

#### **12. Future Perspectives**

There is an increasing need for new IRM strategies. The effective control of resistant pests and the delay of adaptive evolution to resistance in insects will not be achieved solely with pyramid strategies. In depth knowledge of the mechanism of action of Bt proteins and the mechanisms of insect resistance is crucial for prolonged benefits of Bt toxins in pest control. For effective resistance management, it is necessary to develop novel toxins, and to combine more than one strategy in pest control. Vip toxins are a promising new generation of insecticides to be used in spray formulation and transgenic crops, because of their broad spectrum of insect targets. Researchers are focusing on the structure and function of Vip proteins and are attempting to find new Vip proteins from already identified and novel Bt strains. Finding new proteins could be a strategy of choice to manage resistance to Bt toxins. Next generation sequencing (NGS) can accelerate the discovery of novel proteins through the complete sequencing of novel genomes and already known Bt strains.

**Author Contributions:** Conceptualization, T.S., C.L., and R.Z.; writing (original draft preparation), T.S., M.A., Z.M., M.A.A., Y.W., Y.L., S.G., C.L., and R.Z. All authors have read and agreed to the published version of the manuscript.

**Funding:** This work was supported by grants from the Ministry of Agriculture of China (grant nos. 2016ZX08005004 and 2016ZX08009003-003-004).

**Acknowledgments:** We would like to apologize to all the investigators whose research could not be cited appropriately, owing to space limitations.

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**


© 2020 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).

### *Correction* **Correction: Syed, T., et al. Current Insights on Vegetative Insecticidal Proteins (Vip) as Next Generation Pest Killers.** *Toxins* **2020,** *12***, 522**

**Tahira Syed †, Muhammad Askari †, Zhigang Meng, Yanyan Li, Muhammad Ali Abid, Yunxiao Wei, Sandui Guo, Chengzhen Liang \* and Rui Zhang \***

> Biotechnology Research Institute, Chinese Academy of Agricultural Sciences, Beijing 100081, China; syedtahira98@gmail.com (T.S.); 2017Y90100082@caas.cn (M.A.); mengzhigang@caas.cn (Z.M.); liyanyan01@caas.cn (Y.L.); abid@caas.cn (M.A.A.); weiyunxiao@caas.cn (Y.W.); guosandui@caas.cn (S.G.) **\*** Correspondence: liangchengzhen@caas.cn (C.L.); zhangrui@caas.cn (R.Z.); Tel.: +86-10-82106127 (R.Z.)

† These authors contributed equally.

The authors wish to make the following corrections to their paper [1]: In the abstract and introduction section, the words "gram negative" should be replaced with "Gram positive" in first sentence.

In Section 6, the first line of the second paragraph, the second word should be replaced with "laboratory evolved resistance". In the third sentence of same paragraph, reference 72 in the original paper should be replaced with reference [2] in this correction.

In the third paragraph of this section, sentence five should be replaced with "Instead, a low proteolytic activity was found in resistant insects". The last reference in this paragraph (reference 78 in the original paper) should be replaced by reference [3] in this correction.

In the fourth paragraph of this section, the first line should be replaced with "no cross resistance to Vip3C has been observed in insects of different species, previously found to be resistant to Cry1A, Cry2Ab, Dipel (Mixture of Cry1 and Cry2)". The third sentence of this paragraph should be replaced with "The biochemical basis of resistance could not be established by the down regulation of membrane bound alkaline phosphatase (mALP) isoform HvmALP1, observed in Vip3 resistant insects, and results does not support it to be the functional receptor of Vip3".

The reference in the second paragraph of Section 9.1 and the seventh row of Table 2 (reference 98 in the original paper) should be replaced by reference [4] in this correction.

We apologize for any inconvenience caused to readers of *Toxins* by this change. The manuscript will be updated and the original will remain online on the article webpage. We have also rearranged all references and citations according to the correct order. We apologize for any inconvenience caused to our readers.

#### **References**


**Citation:** Syed, T.; Askari, M.; Meng, Z.; Li, Y.; Abid, M.A.; Wei, Y.; Guo, S.; Liang, C.; Zhang, R. Correction: Syed, T., et al. Current Insights on Vegetative Insecticidal Proteins (Vip) as Next Generation Pest Killers. *Toxins* 2020, *12*, 522. *Toxins* **2021**, *13*, 200. https://doi.org/10.3390/ toxins13030200

Received: 23 February 2021 Accepted: 2 March 2021 Published: 11 March 2021

**Publisher's Note:** MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

**Copyright:** © 2021 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https:// creativecommons.org/licenses/by/ 4.0/).

### *Review* **Making 3D-Cry Toxin Mutants: Much More Than a Tool of Understanding Toxins Mechanism of Action**

#### **Susana Vílchez**

Institute of Biotechnology, Department of Biochemistry and Molecular Biology I, Faculty of Science, University of Granada, 18071 Granada, Spain; svt@ugr.es; Tel.: +34-958-240071

Received: 15 July 2020; Accepted: 20 August 2020; Published: 16 September 2020

**Abstract:** 3D-Cry toxins, produced by the entomopathogenic bacterium *Bacillus thuringiensis*, have been extensively mutated in order to elucidate their elegant and complex mechanism of action necessary to kill susceptible insects. Together with the study of the resistant insects, 3D-Cry toxin mutants represent one of the pillars to understanding how these toxins exert their activity on their host. The principle is simple, if an amino acid is involved and essential in the mechanism of action, when substituted, the activity of the toxin will be diminished. However, some of the constructed 3D-Cry toxin mutants have shown an enhanced activity against their target insects compared to the parental toxins, suggesting that it is possible to produce novel versions of the natural toxins with an improved performance in the laboratory. In this report, all mutants with an enhanced activity obtained by accident in mutagenesis studies, together with all the variants obtained by rational design or by directed mutagenesis, were compiled. A description of the improved mutants was made considering their historical context and the parallel development of the protein engineering techniques that have been used to obtain them. This report demonstrates that artificial 3D-Cry toxins made in laboratories are a real alternative to natural toxins.

**Keywords:** 3D-Cry toxins; in vitro evolution; rational design; *Bacillus thuringiensis*; toxin enhancement

**Key Contribution:** Compilation of all of 3D-Cry toxin mutants with enhanced activity made with different molecular techniques.

#### **1. Introduction**

Sporulating *Bacillus thurigiensis* produces four non-phylogenetically related insecticidal protein families, the three domain Cry toxins or 3D-Cry toxins, the mosquitocidal Mtx, the binary-like (Bin), and Cyt toxins. All of these proteins form crystals (with the exception of Cry1Ia toxin [1]) concomitantly with the sporulation process. Since the discovery of these crystals in 1953 by Hannay [2], and the demonstration one year later [3] that they were responsible for the already-described entomopathogenic activity of *B. thuringiensis* [4], the study of these toxins has not stopped.

All known Cry toxins (3D-Cry, Mtx like, Bin and Cyt toxins) have been compiled in a brand new database [5], maintained by a commission of experts in charge of assigning a name when a novel Cry protein is described, using the recently proposed structure-based nomenclature rules [6] but with the same basic principles of the rules that were established in 1998 [7]. Most of the 3D-Cry toxins are active against insects from different orders, mainly Lepidoptera, Diptera, Coleoptera, Hemiptera (low toxicity for some aphids), and Hymenoptera, but some of them have other targets such as nematodes, snails, and even cancer cells [8]. Recently, a toxin active against Orthopteran insects has been described [9].

3D-Cry toxins are the best-characterized among the Cry proteins. Among them, Lepidoptera-active toxins are the best known from a mechanistical point of view. 3D-Cry toxins, synthesized as inactive protoxins by *B. thuringiensis*, have to undergo a proteolytic activation process in the guts of the susceptible insects to become active. Since the elucidation of the first three-dimensional structure of the active part of the Cry3A toxin by Li et al. in 1991 [10], the structure of nine other members of the family have been elucidated (Cry1Aa [11], Cry3Bb1 [12], Cry1Ac [13], Cry2Aa [14], Cry4Ba [15], Cry4Aa [16], Cry8Ea1 [17], Cry5B [18], and Cry7Ca1 [9]). Recently, the 3D structure of the 120 KDa Cry1Ac1 prototoxin has also been described [19], showing seven different domains (DI–DVII). Although very different at the amino acid sequence level, the structural disposition of 3D-toxins is very conserved. Active toxins present three very distinct structural domains (hence their name of 3D-toxins), each of them with a specific function. Domain I, at the N terminus end of the protein, is comprised of a bundle of seven α-helices and is responsible for the formation of a pore in the midgut cells of susceptible insects. Domain II, the middle domain, formed by three antiparallel β-sheets, plays an important role in receptor recognition. Domain III, a two antiparallel β-sheet sandwich, is thought to be involved in receptor binding and pore formation [20]. Protoxin Domains IV and VI are α-bundles similar to domains present in other proteins such as spectrin or the bacterial fibrinogen-binding complement inhibitor. On the other hand, Domains V and VII are β-rolls resembling carbohydrate-binding proteins such as sugar hydrolases [19]. Although the functions of DIV–DVII are not known, they have been related with crystal formation, toxin stability, and selective solubilization in the insect gut. In addition, it has recently been suggested that Domains V and VII could also be interacting domains with proteins present in gut membranes, and hence be involved in the recognition of receptors of the full toxin [21].

3D-Cry toxins have been exploited in insect pest management since the late 1930s [22] in agriculture and against health-related insect populations. 3D-Cry toxins have been used in agriculture, not only as spray formulations, but also in plant transgenesis to protect plants such as maize, cotton, soybean, potato, and tomato from insects [23]. 3D-Cry toxins are extremely efficient and their main characteristic is the narrow spectrum of action that each toxin shows. Their specificity is due to a complex mechanism of action, and although it is not completely understood and many questions remain unanswered, it is known to involve several steps. Currently, two models to explain the mechanism of 3D-Cry toxins have been proposed: the sequential binding model and the signaling pathway model. The sequential binding model involves crystal solubilization at a specific pH, proteolytic activation by gut digestive enzymes, receptor recognition at the membrane cells of susceptible insects, helix 1 proteolysis, conformational changes of the molecule, polymerization, and finally, membrane insertion. The signaling pathway model shares the steps of crystal solubilization, proteolytic activation, and receptor binding, but death of the cell is not explained by toxin insertion in the membrane but by the activation of cell apoptosis. Both models [22] have in common the need of 3D-Cry toxins to bind to a specific receptor at the enterocytes and possibly many other interacting receptors are required for the toxin mode of action. Among them, aminopeptidase N (APN), cadherin-like proteins (BT-R1, BtR175, HevCaLP), alkaline phosphatase (ALP), and ABC transporters have been described, with the first two receptors being the best characterized.

Everything that we know today about the mechanism of action of 3D-toxins has been mainly obtained from two sources of information: the construction of 3D-Cry mutants and the study of the resistance phenomena that insects have shown to the action of 3D-Cry toxins [24,25]. The study of mutant proteins is one of the pillars for the elucidation of any mechanism of action of any protein. The principle is simple: if one amino acid of the protein is essential for its mechanism (or the structure), when changed for another amino acid, the functionality of the protein is modified. This is a consequence of the biological principle that the structure and function of any protein are always linked. In the case of 3D-Cry toxins, thousands of mutants have been constructed to elucidate which amino acids are important for maintaining the three-dimensional structure of 3D-toxins, which are responsible for the binding to their receptors, and which are relevant for the solubilization or activation of the toxins. The information provided by the behavior of these mutants has allowed researchers to propose models that explain the mechanism of action of these natural machines, specialized in killing insects.

However, constructing mutant proteins also presents the possibility of obtaining functional variant proteins with different behaviors, even with improved activity. By manipulating the DNA sequence that codifies the 3D-toxins, versions of toxins completely novel in nature with an enhanced activity toward a particular insect, with a broader insect target, or with a novel activity against a non-susceptible insect can be obtained. The more we understand 3D-Cry toxins, the more creative we can be in the generation of artificial toxins. The fact that a high number of molecular techniques for DNA manipulation is available ensures that in the generation of 3D-Cry toxins variants, only our imagination is the limit.

The techniques available for DNA manipulation and generation of novel mutants or variants can be separated in two main groups: (i) those where mutations are generated randomly and afterward, there is a screening process and the desired mutant is selected, and (ii) those where there is a rational design behind the mutant construction. A deep understanding of protein structure and function is needed in order to use rational design, as we must decide which amino acid (or amino acids) is (or are) going to be changed and which amino acid is going to be substituted for. In contrast, random mutation can be generated anywhere in the protein sequence without previous knowledge, and selection of the suitable variant is carried out later on. The latest group of techniques is known as directed evolution or in vitro evolution of proteins in protein engineering. It is a process that simulates natural evolution, introducing a mutation and selecting it if it represents an advantage, but carried out in a laboratory context and with human intervention. For this review, I would like to use the term "in vitro evolution" in a much broader sense. I would like to include those mutants obtained by rational design and those obtained when investigating the function of the toxin, but instead of getting an impaired mutant with no function, a better 3D-Cry toxin is generated. If the reader grants me this license, then the purpose of this review makes much more sense as the intention is to compile all of the mutants generated in 3D-Cry toxins, independently of the objective of the work and the technique used. These man-made (and woman-made) mutants presented here collectively represent the "in vitro evolution" that 3D-Cry toxins have experienced thanks to the work of hundreds of researchers worldwide in a relatively short period of time. Although it must sound highly pretentious comparing human work that of Mother Nature, the fact is that some of the mutants obtained in a laboratory sometimes show better characteristics than natural toxins, at least from a practical point of view.

Previous reports have reviewed the enhancement of 3D-Cry toxin activity [26,27], but this time I would like to give a historical perspective of what the methodological context of protein engineering was like when the enhanced toxins were obtained. Therefore, apart from the objective of updating the information to the present time, this review has the purpose of describing the parallel development of molecular techniques that were used for constructing the improved versions of 3D-Cry toxins. The mutants reviewed here represent all successful variations of the 3D-Cry toxins that have been experienced in a laboratory, collectively, through many different techniques, even if the intention of the researchers was not to obtain an improved mutant. In this report are compiled all the enhanced mutants constructed through the history of 3D-Cry toxin research (Table 1; shown at the end of section two), together with all the relevant positions in the toxin molecules. The sequences of all these mutants are detailed in Table S1. I believe that this is a valuable source of information that I hope will contribute to the production of even better molecules in the future.

#### **2. "In Vitro Evolution" of 3D-Cry Toxins: An Historical Perspective**

Although Cry toxins are very efficient molecules and only very minute quantities are required for toxicity, the obsession to improve their efficiency has led to the development of diverse strategies [28]. The developed strategies include (i) the combination of several Cry toxins to increase efficiency toward a specific target [29]; (ii) co-expression with other *B. thuringiensis* proteins such as the P20 protein to provide protection in the larval gut environment [29,30], or chitinases for peritrophic membrane degradation [31]; (iii) combination with chemical compounds such as calcofluor for peritrophic membrane digestion [32], or coating with Mg(OH)2 to increase their resistance to UV light [33]; (iv) combination with other insecticidal toxins such as Cyt toxins [34–37], VIP toxins [38], Bin toxins [39], Metalloproteinase Bmp1 [40], or insect-specific scorpion toxins [41] that synergize their

effect; (v) combination with insect chaperones such as Hsp90 chaperone [42]; or (vi) expression of Cry toxins in other backgrounds such as baculovirus [43]. Other strategies developed to increase potency include the fusion of 3D-Cry to other toxins such as neurotoxins (Huwentoxin-I [44], ω-ACTX-Hv1a [45], or huwentoxin XI [46]) present in spiders venom, VIP proteins [47], the N-terminal region of PirB toxin from *Photorhabdus luminescens* [48], or the fusion to other proteins that provide interesting domains such as garlic lectin [49,50], cellulase-binding peptides [51], or *Escherichia coli* maltose binding protein (MBP) [52], thus rendering chimeras with improved activity. Lately, the combination or co-expression of Cry toxins with peptides with sequences similar to natural receptors, or other proteins present in the gut cells, is attracting the attention of researchers. Among them, peptides such as HcAPN3E, derived from an APN receptor in *Hyphantria cunea* [53], or cadhering fragments [54–59] can be mentioned.

Some of the mechanisms for enhancing 3D-Cry toxin toxicity above-mentioned are common strategies in the field of protein engineering (like the fusion of 3D-Cry toxin with other proteins), but are out of the scope of this review as the intention here is to describe specific changes carried out in the amino acid sequence of 3D-Cry toxins that are responsible for the improvement of toxicity.

The starting point in the history of the 3D-Cry toxin "in vitro evolution" could be set at the beginning of the 1980s. At that time, researchers started to understand that the differences in activity of the different *B. thuringiensis* isolated strains were due to the expression of different Cry toxin variants. In 1981, the first *cry* gene was cloned and expressed in a heterologous system [60], a milestone that represented the beginning of molecular biology for Cry toxins. A few years later, the sequence of the first *cry* gene and its deduced amino acid sequence were determined [61]. When the sequence of several other 3D-Cry toxins was available, researchers realized that areas with conserved and variable sequences were present in all of the Cry toxins, so conserved and variable blocks were established. Pretty soon, "variable regions" were related to the specificity observed in 3D-Cry toxins and represented a good starting point for the manipulation of the molecules in order to increase the activity toward insects or expand their target insects. In this sense, the patent number EP0228838 was filed in 1986 by the Mycogen Corporation at the European Patent Office [62] to commercially protect the idea that activity of 3D-Cry toxins could be modified and improved by exchanging specific "variable" regions at their sequence, and a novel method with which to do it. Since then, the history of Cry toxin "in vitro evolution" has not stopped and continues until the present day. Cry toxin history is long and exciting, and has been possible thanks to the development of the molecular tools for DNA manipulation. A description of all the molecular techniques used for the improvement of 3D-Cry toxins will also be made.

#### *2.1. Evolution by Chemical Mutagenesis and Homologue Scanning Mutagenesis, the First Molecular Techniques Used for Cry-Toxins "In Vitro Evolution"*

In the 80s, tools for molecular biology were extremely limited. To provide the reader with some background context, restriction enzymes, essential for DNA manipulation nowadays, had only just been recently discovered [63] and the number available was very low. The Sanger method for determining DNA nucleotide sequences had just been developed [64,65] and chemical mutagenesis was pretty much the only tool available for the "in vitro evolution" of Cry toxins. The technique consisted in subjecting *cry* genes to the action of mutagenic substances such as bisulfite or formic acid to obtain random mutations. Bisulfite, a single DNA strand mutagen, converts cytosines into uracils by deamination [66], rendering a transition from cytosine to thiamine or guanine to adenine, depending on the mutagenized strand (sense or antisense strand). Formic acid depurinates DNA by hydrolyzing the N-glycosyl bond between the ribose and purines [67], and when polymerization takes place using this mutated DNA as a template, a transversion is produced.

Chemical mutagenesis has been used to improve the activity of 3D-Cry toxins [68], specifically on CryIA(b) (the original names of the Cry toxins are maintained in this review). After cloning the *cry1A(b)* gene in the M13 phage, in order to obtain single stranded DNA, and setting mutagenesis conditions to obtain 2–3 mutations per gene (by limiting the exposure time of the DNA to the mutagen), the authors obtained eleven mutant toxins that were 3–5 times more toxic toward *Heliothis virescens* than the parental toxin. When the DNA sequence of these clones was determined, a wide range of substitutions was observed, all of them at Domain I of the toxin, although the distribution in the molecule was not known at that time, as none of the 3D-Cry toxin three-dimensional structure had been elucidated yet.

Another option that researchers had available at that time was the technique called homologuescanning mutagenesis. The technique consisted in exchanging equivalent regions of two 3D-Cry toxins in order to create hybrid molecules. The fragments exchanged were limited by the restriction enzymes that these *cry* genes had in common. Without a doubt, these kinds of exchanges would have been done by PCR nowadays, but this technique was not developed until 1988 [69], and it took time until the research community understood its potential in protein engineering. The main objective of using homologue-scanning mutagenesis was to understand the remarkable specificity that Cry toxins had against the same insects (at the beginning most of them Lepidoptera), but soon researchers understood the potential that this knowledge also had from a practical point of view; identifying regions responsible for the specificity was of paramount importance as they could be manipulated and modified to extend or change the specificity toward other insects.

One of the firsts reports using homolog-scanning mutagenesis in a Cry toxin was published in 1989 [70]. Here, the authors used two *cry* genes, called at the time *icpA1* and *icpC73*, as the first systematic nomenclature for Cry toxins had not been implemented [71] (in fact, the first proposed nomenclature was published the same month as this report). The authors exchanged several fragments between ICPA1, which is highly toxic toward *Bombyx mori*, and the non-toxic ICPC73 by making use of the restriction sites that these two *cry* genes have in common. After the resulting novel toxins were bioassayed, the authors observed that when conserved blocks were exchanged, the activity of the hybrid toxins did not change. However, when variable regions were exchanged, the activity of the toxins dramatically changed and could be redirected toward other insects. In other words, ICPC73 became toxic toward *B. mori* when certain regions from ICPA1 were substituted in its sequence. The region responsible for *B. mori* specificity in the ICPA1 toxin was even narrowed down to the region between residues 332 and 450, a region that we know today includes loop 1, loop 2, and loop 3 in most of the 3D-Cry toxins and has been proven to be involved in receptor recognition and specificity. These authors suggested that if these regions were indeed responsible for specificity in other "IPC" toxins, that they would be an excellent area for mutation in order to redirect activity toward other insects.

The same strategy and the same toxins (now called CryIA(a) and CryIA(c)) were tested in other insects by Schnepf et al. [72]. Both toxins showed a similar activity against *Manduca sexta*, but very different activities against *H. virescens* (as CryIA(c) is 50 times more potent than CryIA(a) for this insect). With these models, the specificity determinant regions for Lepidoptera were determined and it was demonstrated that Cry toxins could be "in vitro evolved" and their specificity could be changed completely. The same year, the specificity-determining region for a Dipteran toxin was determined [73] using homologue-scanning mutagenesis with the CryIIA toxin (active against Diptera and Lepidoptera) and CryIIB (active against Lepidoptera only). They determined that when a 241 nt segment from *cryIIA* was inserted on the *cryIIB* gene, the lepidopteran toxin showed a broader insect spectrum, also becoming active against Diptera. They even narrowed down the region responsible for the specificity of CryIIA protein toward mosquitoes to 76 amino acids.

A following work [74], done by the pioneers in the use of homologue-scanning mutagenesis in a 3D-Cry toxin, demonstrated the same effect in the CryIA(c) toxin and two other economically important pests (*H. virescens* and *Trichoplusia ni*). They defined the minimal region responsible for the toxicity of CryIA(c) as the region between amino acid 332 and 450, an equivalent region described in CryIA(a). Surprisingly, one of the hybrids obtained (hybrid 4109) showed an enhanced activity compared to the parental toxins, being 30 times more toxic than the most potent natural toxin known for *H. virescens*. This represented the first proof that by changing specific areas in the sequence, not

only was it a means of modifying the specificity of the 3D-Cry toxins, but also a way of increasing their activity.

At this point in 3D-Cry toxin history, researchers started to have the overall view that 3D-Cry toxins were modular structures and that their function could easily be manipulated by exchanging parts of the molecules.

#### *2.2. Evolution by Domain Swapping*

With the elucidation of the first three-dimensional structure of a Cry toxin (Cry3A; PDB: 1DLC) by Li et al. [10], researchers had the opportunity to "see" the spatial distribution of amino acids in a 3D-Cry molecule and to verify that active 3D-Cry toxins showed three very well defined domains (hence their name). Through comparison with other proteins, hypothetical functions for some domains were described. For example, Domain I, with seven long α-helices, long enough to span the lipid bilayer of the cell the membrane, was associated with the pore formation function. Domain II, with three loops at the apical part showing highly variable amino acid sequences, was suggested to be responsible for specificity.

Once the structure of Cry toxins was elucidated and the concept that each domain had a function was established, the idea of improving and redirecting the toxicity of a Cry toxin by exchanging complete domains was reinforced. Many reports of what was called domain swapping were produced using molecular strategies such as in vivo intramolecular recombination, cloning, or overlapping PCR.

The technique known as in vivo intramolecular recombination [75] is based on the construction of a "tandem plasmid" where two truncated proteins, in direct repeated orientations, are cloned (Figure 1). The truncated genes (one lacking the 3 end of the gene and the other the 5 end) only overlap in an area where the recombination is intended. Tandem plasmid contains an enzyme restriction site to further discriminate between recombinant plasmids (where the restriction site is lost) and the parental one. Once the tandem plasmid is introduced in a recombinase positive *E. coli* strain (Rec+), random recombination takes place at the homologue regions and novel hybrids or chimeras are produced.

One of the first works using in vivo intramolecular recombination to obtain 3D-Cry toxin hybrids was reported by Caramori et al. [76]. They cloned two truncated toxin genes (for CryIA(a) and CryIA(c)) with overlapping variable regions (63% identity). After the hybrid toxins were bioassayed against several Lepidoptera, it was determined that hybrid 32 (pHy32) and hybrid 45 (pHy45) were more active toward *T. ni* and *Heliotis* sp., respectively, than any of the parental toxins. Two of the hybrid toxins (pHy104 and pHy122) with the same amino acidic sequence even gained a novel activity against *Spodoptera littoralis* as none of the parental toxins were active against this insect.

In vivo intramolecular recombination has been used for the combination of other 3D-Cry toxins and some of them rendered enhanced toxins. This is the case for the combination of CryIC and CryIE toxins [77], both active against Lepidoptera, but with different specificities. CryIC is particularly active against *Spodoptera exigua* (LC50 26 ng/cm2) and *Mamestra brassicae* (LC50 8 ng/cm2) while CryIE is not (LC50 > 1000 ng/cm2). The authors constructed two tandem plasmids with truncated genes overlapping at Domain II and III of the toxins to construct CryIC-CryIE and CryIE-CryIC hybrid proteins. One of the resulting hybrids, named G27 and containing Domain I and II from CryIE and Domain III from CryIC, was toxic to *S. exigua* (LC50 2 ng/cm2). The reverse hybrid (DI and DII from CryIC and DIII from CryIE) was not toxic at all, meaning that the CryIC Domain III was involved in the specificity against this insect.

Using in vivo intramolecular recombination, de Maagd et al. [78] demonstrated that the moderate toxicity of CryIA(b) toward *S. exigua* could be enhanced by constructing a hybrid between DI and DII from CryIA(b) and DIII from a highly active toxin (CryIC). Hybrid H04 showed an increase in toxicity of more than 66-fold (CL50 from > 100 μg of toxin/g of diet to 1.66 μg/g) compared to Cry1Ab. The activity of the novel hybrid was even better than the parental CryIC toxin (LC50 11 μg/g), showing a 6.6-fold increase in toxicity. The authors also determined that binding sites of CryIA(b) and CryIC to

*S. exigua* Brush Border Membrane Vesicles (BBMVs) were different for both toxins and argued that in the case of insect resistance, hybrid H04 could be of great use.

**Figure 1.** In vivo intramolecular recombination. A tandem plasmid is constructed by the cloning of two truncated genes, one truncated at the 3 end (blue gene) and the other one at the 5 end (green gene), leaving an overlapping region were recombination is desired. The plasmid contains a restriction site to further discriminate if recombination took place. Plasmid is then introduced in a *recA*<sup>+</sup> *E. coli* strain and DNA recombination is allowed. After plasmid extraction from the *E. coli recA*<sup>+</sup> strain, the pool of plasmids are digested with a restriction enzyme and selected in a *recA*- *E. coli* strain. Each clone represents a recombination event where the two toxins genes have been fused [77].

Other hybrid toxins obtained by in vivo recombination from the less studied Cry1Ba, Cry1Da, and Cry1Fa toxins (Cry toxins nomenclature updated following [7] rules) were reported to have an improved activity [79]. This was the case of hybrid BBC13 (DI-DII from Cry1Ba and DIII from Cry1Ca) that showed an 11.8-fold increase in toxicity toward *M. sexta*, or hybrid BBC15 (DI-DII of Cry1Ba and DIII of Cry1Ca), which showed 8.3-fold and 7.8-fold increases in toxicity against *S. exigua* and *M. sexta*, respectively, or hybrid FFC1 (DI-DII from Cry1Fa and DIII from Cry1Ca) that showed a 5.5-fold increase toward *S. exigua*.

The latest example of hybrids made by in vivo recombination were made from Cry1Ca and Cry1Fb toxins, and DIII from Cry1Ac [80]. Hybrids RK15 (Cry1Ca/Cry1Ca/Cry1Ac) and RK12 (Cry1Fb/Cry1Fb/Cry1Ac) showed an increase in activity, compared to wild type toxins Cry1Ca and Cry1Fb, of more than 172 and 69 times toward *H. virescens*, respectively.

Another way of performing domain swapping and constructing hybrid toxins has been through standard cloning using restriction enzymes (already present or expressly created). This is the case of the hybrid Cry1C/Ab [81], constructed with the first 2194 nucleotides from *cry1C* (731 aa) and the last 1295 nucleotides (432 aa) of the 3 end of the *cry1Ab* gene. The fusion was possible thanks to the presence of a unique *Kpn*I site in a conserved region of the two genes. The Cry1C/Ab hybrid was 3, 4, and 35 times more active against *S. littoralis*, *Ostrinia nubilalis*, and *Plutella xylostella*, respectively, than the parental Cry1C toxin.

Domain swapping of Coleoptera active toxins has also been successfully accomplished by standard cloning [82], although in this case, it was necessary to introduce restriction sites (*Rsr*II) to obtain the hybrids. Hybrid 1Ia/1Ia/1Ba (DI-DII from Cry1Ia and DIII from Cry1Ba; LC50 22.4 μg/mL) was 2.5 and 7.5 times more toxic than their parental toxins, respectively. Hybrid 1Ba/1Ia/1Ba (DI from Cry1Ba, DII from Cry1Ia, and DIII from Cry1Ba, LC50 7.94 μg/mL) increased its activity even further, showing 17.9 times more potency than the parental Cry1Ba toxin. The latest hybrid toxin was almost as toxic as the Cry3Aa toxin, the most active natural protein for the Colorado potato beetle.

More recently, and thanks to the determination of the Cry1Ac1 full-length toxin 3D structure ([19]; PDB 4W8J), Zghal et al. [21] constructed a 116 KDa chimeric toxin called Cry(4Ba-1Ac) by fusing DI–DIV from Cry4Ba to DV–DVII from Cry1Ac1, using PCR amplification and cloning techniques. This represents a unique case in which other domains, apart from Cry toxin toxic domains, have been swapped. The chimeric toxin showed low toxicity toward *Culex pipiens* when expressed in an acrystalipherous *B. thuringiensis* strain (HD1 CryBpHcry(4Ba-1Ac)), but when co-expressed in a Cry2Aa producing strain (BNS3pHTcry4BLB), the activity increased from 10% mortality to 100% mortality at 200 μg/L. The LC50 of the strain bearing only Cry2Aa switched from >>200 μg/L to 0.84 μg/L when co-expressed with the chimeric toxin, meaning that an increase in toxicity of more than 238-fold was produced. This synergy was also observed when Cry2Aa and the chimeric toxin Cry(4Ba-1Ac) were bioassayed in combination. The authors suggested that the increased toxicity could be explained by a better solubilization of the crystals and also proved the importance of the protoxin domains (DIV–DVII) in the stability and the activity of the Cry toxins, a fact that will possibly be exploited in the future.

3D-Cry toxin domain swapping has also been obtained by overlapping PCR [83]. In this case, a hybrid using toxins from different classes, one coleopteran (mCry3A) and one lepidopteran specific (Cry1Ab), was obtained. DNA regions codifying for DI and DII from mCry3A [84] and DIII from Cry1Ab were amplified, containing an overlapping region at the 3 and 5 ends, respectively. Amplicons were used as a template in an overlapping PCR with flanking primers. The resulting chimera (called eCry3.1Ab; GenBank GU327680) was highly active (93% mortality) against *Diabrotica virgifera virgifera* when bioassayed with a toxin concentration ranging from 5 to 10 μg of toxin per mL of diet. Although the authors did not determine the LC50 of eCry3.1Ab, the toxicity observed was much higher than the previously reported parental mCry3A toxin (LC50 65 μg/mL; [84]).

A more recent example of constructing improved mutants by domain swapping using overlapping PCR [85] was the hybrid toxin Cry1Ac-Cry9Aa containing DI from Cry1Ac and DII and DIII from Cry9Aa. The hybrid showed 4.9 times more activity against *Helicoverpa armigera* than the wild type Cry9Aa. In addition, a Cry1Ac-Cry9AaMod toxin, where helix 1 was proteolytically removed, showed a 5.1-fold increase in toxicity.

#### *2.3. Evolution by Site-Directed Mutagenesis*

Site directed mutagenesis is a molecular strategy used to create specific changes in the amino acid sequence of a protein in order to evaluate its role in the molecule. If an amino acid is involved in the mechanism of action of a protein, when changed, the function of the protein is modified and normally abolished. However, the technique could also be used as a means of obtaining novel proteins with improved characteristics.

Site-directed mutagenesis consists of the in vitro synthesis of the codifying DNA of a protein in which one or several nucleotides are changed in a specific site in order to produce a mutant protein. The changed nucleotide is normally introduced using a mutant primer that is in vitro extended thanks to a DNA polymerase (the *E. coli* DNA polymerase Klenow fragment in the early days, and *Taq* polymerase lately when PCR [69] and site-directed mutagenesis by PCR [86] were developed). The in vitro synthesized DNA (mutant DNA) is counter selected from the wild-type DNA, and the mutant protein is expressed and tested for activity.

The elucidation of the three-dimensional structure of a 3D-Cry toxin facilitated the design of mutants for site-directed mutagenesis as the position of the amino acid to be changed was localized in the space together with its interactions with other amino acids in the molecule. Hundreds of site-directed mutagenesis studies in many Cry toxins have been performed, allowing the elucidation of the function of each domain of the protein. Although most of the mutants obtained by site-directed mutagenesis showed an impaired or diminished toxicity, in some cases, the activity of the resulting mutants was higher than the parental toxins. These mutants were not as useful as the impaired mutant to elucidate the mechanism of action of 3D-Cry toxins, but served to settle the concept that activity improvement was possible with only the change of a single amino acid.

This was the case in the work reported by Wu et al. [87] in which the authors constructed 31 mutants at two conserved regions at CryIA(c) Domain I (residues from 84 to 93 and from 160 to 177). Although most of the mutant toxins showed no toxicity, or no change in toxicity at all, one of them, the mutant H168R, localized at the hydrophobic face of the amphypatic α-helix 5, showed a 3–5-fold increase in toxicity toward *M. sexta* compared to the wild type toxin. Another example of Domain I improved mutants was obtained by investigating the role of nine tryptophan residues in the toxicity of Cry1Ab toward *M. sexta* [88]. These authors found that a conservative change to phenylalanine (W73F, W210F, W219F) produced mutant toxins 3.3, 1.5, and 2.3 times more toxic than the parental toxin. In a similar study [89], two α-helix 5 Cry1Ab mutants (V171C and L157C) with a 25-fold and 4-fold increase in toxicity toward *Lymantria dispar*, respectively, were reported.

Site directed mutagenesis studies carried out in 3D-Cry toxins showed that Domain II was particularly sensitive to amino acid changes [90,91]. Some of the mutations performed in this domain were very successful in improving the toxicity against certain insects, and represented an excellent place for redesigning the activity of 3D-Cry toxins. This was the case reported by Rajamohan in 1996 [92] where two single mutants N372A, N372G, and a triple mutant DF-1 (N372A, A282G, L283S) of the Cry1Ab toxin were constructed by site-directed mutagenesis. These residues were localized between the Cry1Ab α-helix 8a and α-helix 8b at Domain II. When bioassayed, N372A and N372G were, respectively, 8.5 and 8.3 times more potent for neonates of *L. dispar* than the parental Cry1Ab toxin, and 9.61 and 9.51 times more potent for fourth instar larvae, respectively. The DF-1 mutant showed an increase in activity of 36- and 17-fold toward neonates, and fourth instar larvae, respectively. The triple mutant was even more toxic (4-fold) than Cry1Aa, the most potent natural toxin active against *L. dispar*.

Loop 1 from Domain II has also been a successful place for mutagenesis, rendering improved mutants in the coleopteran active Cry3A toxin [93]. Site-directed mutagenesis at positions R345, Y350, and Y351 rendered eight single mutants, four double mutants, and three triple mutants, two of them showing an enhanced activity against *Tenebrio molitor*. Mutant A1 (R345A, Y350F, Y351F) and mutant A2 (R345A, ΔY350, ΔY351) were 11.4 and 2.7 times more active than wild type Cry3A, respectively. In addition, an enhanced activity of these two mutants against two other Coleoptera species, *Leptinotarsa decemlineata* and *Chrysomela scripta*, was also observed. Although changes introduced in these mutants were not very drastic (as Y and F differ only by an OH group), differences in toxicity were remarkable. In addition, loop 3 of the Cry3A toxin also proved to be relevant for toxicity enhancement [91]. The triple mutant S484A, R485A, G486A, showed a 2.4-fold increase in toxicity toward *T. molitor* compared to the wild type toxin.

Another place in Domain II where the substitutions introduced by site-directed mutagenesis rendered an enhanced toxin was the area known as D block or Dipteran specific block [73] in the dual toxin Cry2A (dual as it is also active against Lepidoptera). Cry2Ab is not a very potent toxin toward *Anopheles gambiae* (LC50 540 ng/mL), but when residues from 307–337 were mutated, three mutants (N309S, F311I, and A334S) with enhanced toxicity (1.17, 3.17, and 6.75-fold, respectively) were obtained [94]. One of the mutants, A334S, was even more toxic than the highly active natural toxin Cry2Aa [95].

Recently, 3D-Cry toxin Domain III has also been targeted for mutation by site-directed mutagenesis, and mutants with improved characteristics have been found. This is the case of the work reported by Lv et al. [96] in Cry1Ac5. Although the structure of this 3D-Cry toxin is not elucidated, 3D structure modeling using known structures (Cry1Aa (PDB:1CIY), Cry2Aa (PDB:1I5P), Cry3Aa (PDB:1DLC), Cry3Bb1 (PDB:1JI6), and Cry4Ba (PDB:1W99) was useful in localizing the loop sequence between β-20 and β-21 (576NFTSSLGNIV586). Two mutants obtained by site directed mutagenesis, S581A and I585A, showed 1.72- and 1.89-fold increases in toxicity toward *H. armigera*.

Another area that has recently been demonstrated to be susceptible to improvement by site directed mutagenesis is the β sheet 16 in Domain III. A study of alanine substitution in Cry1Ab [97] performed on this area rendered mutants S509A, V513A, and N514A, which showed an increase in toxicity toward *Spodoptera frugiperda* of more than 9.5-, 12.7-, and 51-fold, respectively. As N514 was the most relevant position experiencing toxicity enhancement in β-16, saturation mutagenesis was performed at this position (N was changed for any of the other 19 amino acids). Some of the obtained mutants, N514F, N514H, N514K, N514L, N514Q, N514S, and N514V showed a 44-, 16-, 7-, 9-, 26-, 23-, and 9-fold increase in toxicity, respectively, when compared to the wild type. An equivalent mutation in Cry1Fa (N504A), a more potent toxin than Cry1Ab for *S. frugiperda*, rendered a mutant 6–11 times more toxic than the wild type Cry1Fa in different populations of *S. frugiperda* from different countries. The authors suggested that the increase in toxicity correlated with an increase in the stability of the mutants toward gut proteases and an increase in BBMV binding.

The fact that two other mutants in β-16, this time in Cry1C [98], showed a slight increase in toxicity compared to the wild type (mutant V509A was 1.6 times more toxic than Cry1C for *M. sexta* and mutant N510A was 1.5 times more toxic toward *S. frugiperda*) suggest that β-16 is a recently discovered site for toxicity improvement.

The loop sequence between β-18 and β-19 in Domain III also seems to be a relevant region for toxicity enhancement. Mutant N546A [99] showed a slight increase in toxicity (1.8-fold) in Cry1Ac toxin, which correlated with a binding increase toward *H. armigera* BBMVs [100].

Single mutations in Domain III have also been described to be useful for toxicity enhancement. This was demonstrated in the nematocidal toxin Cry5Ba [101]. Investigating the role of the 3 asparagines present in block 3 of the toxin found that alanine substitution, by site directed mutagenesis, rendered a mutant (N586A) with a 9-fold increase in toxicity (GIC50 from 42.11 to 4.75 ng/mL) toward *Caenorhabditis elegans*. Mutant N586A was surprisingly soluble in a wide range of pHs (from pH 5 to pH 12), which correlated with the observed increase in toxicity.

#### *2.4. Evolution by Rational Design*

Rational design is a particular case of site-directed mutagenesis performed only when the knowledge of the structure and the function of the protein under study are very deep. In rational design, a hypothesis is formulated and proven by the construction of mutants with single amino acids changes, deletions, or insertions, normally performed by site-directed mutagenesis. Several works have described successful evolution of 3D-Cry toxins by rational design, although some of these were found by accident trying to prove a different hypothesis. This was the case in the work reported by Angsuthanasombat et al. in 1993 [102] where the authors were interested in demonstrating that R203 in Cry4B toxin was essential for proteolytic processing and the α-helices mobility of Domain I. For that, they replaced the R (proteolytic site) with an A, expecting that toxicity would be completely lost as a consequence of the impossibility of the helices to move properly. The effect was completely the opposite as the R203A mutant was 2.8 times more toxic to *Aedes aegypti* than the wild type.

An example of rational design where the researcher did prove their hypothesis was that reported for the evolution of Cry4Ba toxicity [103]. The Cry4Aa toxin is highly active against four species of mosquito (*Ae. aegypti*, *Anopheles quadrimaculatus*, *Culex quinquefasciatus*, and *C. pipiens*), however, the closely related Cry4Ba toxin (the one to be evolved) showed toxicity toward *Ae. aegypti* and *An. quadrimaculatus*, but not to *C. quinquefasciatus* and *C. pipiens*. Through site-directed mutagenesis, the authors delimited the putative loop 3 in Cry4Ba (VIDYNS) and in Cry4Aa (IPATYK), as the Cry4Ba and Cry4Aa 3D structures were not determined until 2005 and 2006, respectively [15,16]. The authors mutated the Cry4Ba loop 3 by replacing D454 with a P and inserting AT after position 454 to yield a novel toxin (named 4BL3PAT) with a novel loop 3 sequence (VIPATYNS). This small change increased Cry4Ba activity 700-fold toward *C. quinquefasciatus*, and by 285-fold to *C. pipiens*. Other versions of the novel toxin were also constructed in loop 3 by substitution of the PAT motive to other motives (4BL3AAT, 4BL3GAT, 4BL3GAV, 4BL3PAA, and 4BL3AAA), showing an activity gain

toward *C. quinquefasciatus* and *C. pipiens* in almost all variants (with the exception of 4BL3AAA toward *C. quinquefasctiatus*). The study of the mechanism of the 4BL3PAT mutant, compared to the wild type toxin, showed that both toxins had little difference in the ability of reversible binding to *Culex* BBMVs, a similar capability of irreversible binding, but 4BL3PAT showed a higher pore-forming ability than the Cry4Ba parental toxin. The authors identified two novel proteins in the BBMVs, which are 35 and 36 KDa in size, that the novel mutant bonds to instead of the parental toxin, proposing that these two proteins could be functional receptors and explain why the 4BL3PAT variant is toxic to the *Culex* species (although it was not proven). One year later [104], the same authors reported the evolution of the mosquitocidal Cry19Aa toxin by rational design. The Cry19Aa toxin, active against mosquito species such as *An. quadrimaculatus* (LC50 3 ng/mL) and *C. pipiens* (LC50 6 ng/mL), showed low activity against *Ae. aegypti* (LC50 1.4x10<sup>5</sup> ng/mL). After in silico modeling of the Cry4Ba structure and Cry19Aa using Cry3Aa and Cry4Aa as templates, the following changes in Cry19Aa toxin were introduced: (i) Cry19Aa Loop 1 (355SYWT358) was substituted by the Cry4Ba loop 1 (332YQDLR336), and (ii) Cry19Aa loop 2 ( 414YPWGD418) was completely deleted to mimic the length present in the Cry4Ba toxin. The resulting mutant 19AL1L2 was >42,000 times more toxic to *Ae. aegypti* (LC50 3.3 ng/mL) than the parental toxin Cry19Aa, being one of the highest activity enhancements in the history of 3D-Cry toxin evolution. The rationale behind these substitutions was to test the hypothesis that changing loop sequences involved in receptor binding could be useful to enhance toxicity by increasing the affinity of the toxin to its receptor. Unfortunately, no differences between Cry19Aa and 19AL1L2 were detected in either reversible or irreversible binding to BBMVs, so the hypothesis was not correct, but it was useful to demonstrate that "in vitro evolution" of Cry-toxins could be efficiently performed by rational design.

However, the greatest achievement in 3D-Cry toxin evolution by rational design was that reported by Liu and Dean in 2006 [105]. These authors were able to redirect the toxicity of a lepidopteran active toxin toward an insect from a different order, the dipteran *C. pipiens*. The mutant was constructed using the lepidopteran active Cry1Aa toxin as a scaffold, and changing loops 1 and 2 of the molecule. Loop 1 from Cry1Aa (311RG312) was enlarged and substituted by the Loop 1 (332YQDL335) from the mosquito active toxin Cry4Ba (although not active against *C. pipiens*). In addition, part of loop 2 in Cry1Aa (LY367RRIILGSGPNNQ378) was deleted (LY365RRIIL), and the 376NNQ378 sequence was replaced by a single G, resulting in the mutant 1AaMosq (L1:311YQDL314; L2:367GSGPG371), which gained activity against *C. pipiens* while the activity against Lepidoptera *M. sexta* was abolished. The novel toxin 1AaMosq showed an LC50 of 45.73 μg/mL when bioassayed against *C. pipiens*, while the parental toxin Cry1Aa was not toxic to this mosquito at toxin concentrations of 100 μg/mL.

A similar rational design performed in the loops of the Domain II has recently been reported in another 3D-Cry toxin [106]. Cry1Ah and Cry1Ai show high sequence similarity (77% identity at amino acid level) but very different specificity. Cry1Ah is toxic to *H. armigera* but non-toxic to *B. mori* and conversely, Cry1Ai is highly toxic to *B. mori* but has no activity against *H. armigera*. As loops in Domain II of the 3D-Cry toxins are involved in specificity, the authors exchanged loops in the two toxins by reverse PCR in order to evolve the activity of Cry1Ai toward *H. armigera*. One of the obtained mutants, Cry1Ai-h-loop2 (Cry1Ai toxin with loop 2 from Cry1Ah toxin), showed a change of specificity. Toxicity against *H. armigera* increased more than 7.8-fold (from LC50 > 500 μg/g to 64.23 μg/g). When the exchange was done in loop 2 and loop 3, the resulting Cry1Ai-h-loop2&3 mutant showed an increase in toxicity even higher, around 58 times (from LC50 > 500 μg/g to 8.61 μg/g).

Domain I has also been the subject of evolution by rational design [107]. Through bioinformatic analysis, it was found that the first 42 amino acids of Cry2A toxins interacted with a predicted transmembrane (TM) domain (amino acids 51–62) in helix 2 of the toxin. In addition, it was observed that the predicted TM in Cry2A was shorter than the equivalent TMs domains found in other Cry toxins, as a consequence of the presence of two lysines at positions 63 and 64. They predicted that this interaction could be the reason for the Cry2A toxin being less active against lepidopteran pests compared to Cry1A type toxins, so they made the hypothesis that by removing these 42 amino acids from the molecule, a Cry2A variant with increased activity could be obtained. The deleted mutant, D42, showed an increase in activity ranging from 2–3-fold toward Lepidoptera *S. littoralis*, *H. armigera*, and *Agrotis ipsilon*. In addition, when lysines 63 and 64 were substituted by a conserved amino acid present in other toxins (F and P), making the TM domain as long as the one present in other Cry toxins, the activity increased even further. Lysine 63 and 64 in the deleted mutant D42 were replaced by phenylalanine/proline by site-directed mutagenesis, and the mutant toxins D42/K63F, D42/K64F, D42/K63F/K64F, and D42/K63F/K64P were obtained. Single mutant toxins showed the same toxicity as D42, but double mutants increased their toxicity toward the tested Lepidoptera between 1.3 and 2.3 times compared to D42 toxicity, and between 4 and 6.5 times compared to the Cry2A wild type, as predicted.

Another successful example of rational design was performed on the Cry3A toxin [84]. In this work, a chymotrypsin/cathepsin G site (AAPF) was introduced into the loop region between α-helix 3 and α-helix 4 in Cry3A Domain I in order to increase the proteolytic efficiency and hence toxicity. The resulting mutant, mCry3, with a loop sequence 153NPAAPFRN160, was active against *D. virgifera virgifera* larvae (LC50 65 μg/mL) compared to the residual activity of the parental toxin Cry3A (LC50 >> 100 μg/mL). The authors determined that the increase in activity was due to several factors such as a higher solubility at neutral pH, an increase in the efficiency of the proteolytic process, and an increase of specific membrane binding. The introduced mutation did not alter the activity against the Colorado potato beetle larvae, a susceptible insect for Cry3A, but extended activity toward other coleopteran (*D. virgifera virgifera*).

#### *2.5. Evolution by Random Mutagenesis*

Random mutagenesis is one of the strategies that molecular biologists have available to obtain protein variants. In opposition to site-directed mutagenesis, the position of the mutation in random mutagenesis is not controlled. It is frequently used to discover relevant amino acids in a protein when not enough information on the function of the protein is available [90,108], or to create novel variants of a protein with novel functions. There are several techniques that have been used to perform random mutagenesis in 3D-Cry toxins such as in vitro DNA amplification with degenerated primers and error-prone PCR.

The development of the synthesis of degenerated oligonucleotides in 1988 [109] allowed researchers to introduce random mutations in a specific region of the DNA. In 1999, Kumar et al. [110] used a mixture of degenerated primers for the random mutation of a 3D-Cry toxin using a M13mp19 system that provided ssDNA. The researchers' objective was to introduce variability at α-helix 4, and at the loop between α-helix 4 and α-helix 5 of the Cry1Ac toxin. For that, a mix of primers with the wild-type sequence (97% of the primers) and degenerated oligonucleotides (3% of the primers) was used to ensure only one mutation per cycle of amplification. Using this technique, the authors obtained the mutant F134L with a 3-fold enhanced toxicity toward *M. sexta* and *H. virescens*.

Error-prone PCR is an in vitro evolution technique that generates variants using the property of the *Taq* polymerase of introducing substitutions in the DNA amplification process when subjected to certain conditions. Error-prone PCR is the most common method for creating combinatorial libraries based on a single gene. Since its development in 1989 [111], it has been used in many applications, not only to mutate DNA codifying for proteins, but also non-coding DNA regions such as promoter regions [112]. The technique is simple and only requires a PCR mixture slightly different from a standard PCR. The gene subjected to mutagenesis is amplified by upstream and downstream primers in a reaction mix containing Mn2<sup>+</sup> ions, an unbalanced ratio of dNTPs, and/or a higher concentration of Mg2<sup>+</sup>. Depending on the conditions, the overall mutagenesis rate can be controlled. Error-prone PCR mutagenesis has not been extensively used in obtaining Cry toxin variants. Only a few examples of using this technique are found in the bibliography, even though the technique has rendered improved versions of a 3D-Cry toxin. This was the case of the work reported by Shu et al. [113] made in Cry8Ca2 toxin (accession number AY518201), which was active against *Anomala corpulenta*. Two mutants (M100 and M102) showed 5- and 4.4-fold increases in toxicity, respectively, against the larva of this Coleoptera

(LC50 0.23 <sup>×</sup> <sup>10</sup><sup>8</sup> CFU/g and 0.26 <sup>×</sup> <sup>10</sup><sup>8</sup> CFU/g, respectively, compared to LC50 1.15 <sup>×</sup> <sup>10</sup><sup>8</sup> CFU/g of the parental toxin). The sequence analysis of the novel variants showed that only a single mutation in each mutant (E642G in M100 and Q439P in M102) was enough to enhance toxicity.

A more recent example of the use of error-prone PCR, although combined with other techniques, is the random evolution of Cry1Ac5, rendering the T525N mutant with a slight increase in toxicity (1.5-fold) toward *S. exigua* [114].

#### *2.6. Evolution by Mixing Cry Genes: DNA Shu*ffl*ing, In Vitro Recombination, and StEP (Staggered Extension Process)*

DNA shuffling is a powerful in vitro evolution tool for generating artificially and highly diversified sequences by homologous gene recombination (Figure 2a). Although this technique is normally used in proteins of unknown three-dimensional structure, it can be used for any protein. The technique involves random DNA fragmentation of two or more homologous genes with DNAse I, and fragment reassembly in a primer-less PCR. After the generation of a variant library, a screening and selection process of functional variants is conducted.

Since the development of this technique [115,116], DNA shuffling has been used to evolve thousands of proteins, mainly enzymes, to modify their function or activity. The convenience of this powerful technique has also been applied for the in vitro evolution of several 3D-Cry toxins.

Although in vitro evolution by DNA shuffling has not been always successful [117], and sometimes no improved 3D-Cry toxins have been obtained, in several other cases, it has been a suitable technique for obtaining variants with higher activity. For example, DNA shuffling of *cry11Aa*, *cry11Ba*, and *cry11Bb* genes, codifying for toxins active against *Ae. aegypti* and *C. quinquefasciatus*, rendered a mutant (Variant 8) 3.8 times more toxic toward *Ae. aegypti* than the parental toxin Cry11Bb (LC50 22.9 ng/mL), and 6.09 times more toxic than the Cry11Aa toxin (LC50 36.9 ng/mL) [118]. DNA sequence analysis of Variant 8 showed that the mutant contained a deletion of 219 nucleotides (73 aa) at the N-terminal end of the molecule (Domain I), and 6 and 13 nucleotide substitutions in Domain II and III, respectively. The comparative analysis at the protein level between Variant 8 and its parental toxins (Cry11Aa and Cry11Bb) showed 13 amino acid substitutions (GenBank access number MH068787).

Very recently, DNA shuffling has been used to increase the toxicity of an already improved mutant derived from Cry3Aa toxin [52]. The mutant IP3-1, engineered by rational design, contained 15 mutations over the three domains of the toxin (W106L, M117I, V140F, I186V, F206L, K230H, S258T, P292S, E294G, F346L, G468A, L491F, M503T, R531G, and I593M) and showed a higher activity toward *D. virgifera* (LC50 214 ppm) than the parental toxin. To further increase its activity, in vitro evolution by DNA shuffling was carried out and six novel mutant toxins (IP3-2, IP3-3, IP3-4, IP3-5, IP3-6, IP3-7) showed more activity than the parental toxin IP3-1 (LC50 19, 14.7, 13.7, 11.3, 11.6, 7.3 ppm, respectively). An analysis of their sequences showed that mutant toxins contained between six and nine additional mutations. From the six mutants obtained, the IP3-7 variant showed the best increase in toxicity of all (LC50 from 214 to 7.33 ppm). Most of the mutations obtained after DNA shuffling resulted in a reduction of positively charged residues such as lysine and arginine, making novel toxins more acidic and more soluble at neutral pH (*D. virgifera* gut juice is weakly acidic) and hence more active. In addition, all the selected variants showed a similar mutation at two different positions (K152E and R158E), located in the α-helices 3 and 4 loop. According to the authors, these mutations made the loops more resistant to *D. virgifera* gut proteases, contributing to the increase in toxicity compared to the parental toxin IP3-1.

A more sophisticated way of obtaining chimeras is by in vitro recombination using the technique called in vitro template-change PCR. With this strategy, a library of recombinant toxins made from the lepidopteran active toxin Cry2Aa (active toward *Ostrinia furnacalis*, *P. xylostella*, *Chilo suppressalis*, and *H. armigera*) and the low toxicity Cry2Ac [119] was made. The strategy involved four steps: (i) ssDNA amplification of the complementary strand of both genes by asymmetric PCR (amplification using a single reverse primer); (ii) synthesis of the coding strand using the ssDNA from gene 1 as a template in the presence of ddATP, which avoids further extension once it is incorporated in the polymerized strand; (iii) DNA extension of the randomly truncated library using gene 2 as a template, which is achieved thanks to the use of the KOD DNA polymerase, shows 3- –5 exonuclease proofreading activity, and is able to remove the ddATP from the truncated molecule and carry on with the extension of the DNA strand; and (iv) amplification of the full toxin fragment with flanking primers, cloning, and expression in a heterologous system. With this strategy, the authors obtained 37 chimeras (named R1–R37) showing recombination events at 37 different regions of the toxin. When recombination occurred at Domain I or Domain III, no change in specificity was observed. However, when recombination took place at Domain II, toxin specificity drastically changed. The Cry2Ad toxin gained toxicity toward *O. furnacalis* when recombination was in the 416NY417 region (recombinant R24), toward *P. xylostella* when it occurred at 440RPL442 (recombinant R26), and toward *C. suppressalis* and *H. armigera* when it took place at 455GTPGGA460 (recombinant R27).

**Figure 2.** In vitro recombination techniques. (**a**) In DNA shuffling [116], two or more homologous genes are randomly digested with DNAse I (only one strand is represented for simplification purposes). The resulting fragments are extended in a primer-less PCR using homologue fragments as templates. Finally, a PCR using flanking primers is performed in order to obtain a full size library of chimeras, after few rounds of primer-less extension. (**b**) In the staggered extension process or StEP [120], two homologous genes are PCR amplified under restricted conditions (short extension times, and low extension temperature). In cycle 1, a short fragment is extended from a primer in both genes (only one strand is represented). After denaturing, the generated fragments can anneal in the opposite homologous gene, and be extended in cycle 2. After cycle n, a library of recombined chimeras is generated.

The technique, called the staggered extension process or StEP [120], has the same objective as DNA shuffling of producing an in vitro recombination of two or more genes, but with a slightly different methodology. In this technique (Figure 2b), two (or more) homologous genes are denatured and extended from the same primer, using a thermo-cycling program in which the extension step is highly limited in time (few seconds (5 s)) and temperature (extension is carried out at 55 ◦C, temperature in

which *Taq* polymerase has a low extension rate). After 70–80 cycles of denaturing and priming-extension, a library with full-length recombined genes that is cloned and screened can be obtained. This technique has been used for in vitro evolution of the active part (DI, DII, and DIII) of the Cry1Ac5 toxin (GenBank acc. Number M73248) in combination with error-prone PCR. The variants were cloned into a plasmid containing the pro-toxin C-terminal end by Red/ET homologous recombination [121,122]. From the 57 variants obtained, only one was expressed as a full-length 130 kDa toxin containing the mutation T524N in the β-16 and β-17 loops in Domain III. The variant produced more crystals than the wild type, but slightly smaller. When bioassayed toward *S. exigua*, a toxicity 1.5 times higher (LC50 9.6 μg/mL compared to 14.1 μg/mL of the wild type) was observed.

#### *2.7. Evolution by Phage Display*

As has been reviewed thus far, generation of 3D-Cry toxin variants does not represent a big challenge nowadays as many molecular techniques for constructing libraries with a high number of mutants are available. The real challenge is to find which of the variants in the library is useful and has the desired properties. Therefore, the key question of in vitro evolution of a protein is the library screening. In the particular case of 3D-Cry toxins, this screening is labor-intensive as every single mutant has to be expressed, solubilized, and bioassayed, representing a time consuming task. As a consequence, only a reduced number of variants from the library are normally tested, and on many occasions, no improved mutants are found. To overcome this problem, approaches such as phage display have been explored to provide a means for the selection of potentially useful 3D-Cry mutants. Phage display is a molecular tool for the screening of library variants with a specific binding characteristic. A phage displayed protein (or a mutant library) consists of its expression on the surface of a bacteriophage in such a way that the protein is available to interact with other proteins, while it is bound to the virus. Display is achieved thanks to the fusion of the protein of interest (or library) to one of the proteins on the surface of the phage. When the phage replicates and viral particles assemble, the protein of interest is also assembled on the surface, being available to interact with other proteins. Displayed libraries can be screened by a process called biopanning, a methodology that allows for the selection of those phages with the desired binding properties (Figure 3). As the phenotype of the selected variant and the genotype in a phage display system are linked, once a phage is selected, the coding DNA for the protein variant can be obtained from the phage genome. Given that one of the premises for 3D-Cry toxin toxicity is to bind to a receptor, this makes phage display a suitable molecular tool for the screening of variants with novel binding characteristics.

Since the invention of the phage display technique [123] and its use for the selection of a peptide from a library with an antibody, many other applications have been developed [124–127]. 3D-Cry toxins have also benefited from the advantages of phage display technology, although several technical limitations had to be overcome before the methodology could be used for big proteins such as 3D-Cry toxins. The first attempt at displaying a 3D-Cry toxin on the surface of a phage was reported by Marzari et al. [128] using the Cry1Aa toxin and M13 phage. Unfortunately, the toxin was not properly displayed and deletions on the fusion protein were observed. One year later, Kasman et al. [129] reported the successful display of the Cry1Ac toxin on the surface of the M13 phage, although the toxin was unable to bind to the APN receptor in in vitro experiments. Later on, other display systems based on the λ and T7 phage, which are assembled in the cytoplasm of *E. coli* and released after lysis instead of being secreted through the bacterial membrane as in M13, were proven to be more appropriate for 3D-Cry toxins. The first successful phage display system, in which the 3D-Cry toxin was able to bind to natural receptors, was described by Vilchez et al. [130]. In this study, the Cry1Ac1 toxin was fused to the gpD protein, an auxiliary protein that represents one of the major components of the λ phage capsid. The displayed toxin was able to selectively recognize and bind proteins present in *M. sexta* BBMVs. Later on, other display systems using the T7 phage [131] and M13 [132] were described. Once the problem of displaying a 3D-Cry toxin was overcome, library variants were developed.

**Figure 3.** Biopanning of a phage display toxin library. The phage displayed toxin library is biopanned against a specific insect receptor (1). Those phage displaying toxins with affinity to the insect receptor will be retained and those without affinity will be washed out (2). Bond phage will be recovered (3) and amplified (4) by a susceptible *E. coli* strain, making possible to repeat the process (5) in order to obtain higher affinity toxins from the selected pool.

Mutant libraries have been constructed in specific areas of the 3D-Cry toxin using several molecular approaches such as degenerated primers [133–135], DNA shuffling [136,137], or using a previously constructed antibodies library [138]. All these libraries were screened for variants showing high binding affinity toward two of the most well-known receptors (cadherin like receptor and APN), and although an increase of binding affinity is not a guarantee for increased toxicity [134,135], some authors have managed to obtain enhanced 3D-Cry toxin variants compared to the parental toxin.

The first successful report describing the use of a phage display library for the evolution of a 3D-Cry toxin was made by Ishikawa et al. [133] using the T7 phage. They constructed a library of Cry1Aa1 variants at the loop 2 of Domain II, one of the main determinants for specificity in 3D-Cry toxins. Loop 2 variants were constructed by PCR with the degenerated primer Aa369(IILGSGP)375-degenerate-sense (5- TTATATAGAAGANNNNNNNNNNNNNNNNNNNNNAATAATCAGGAACTGTTTG3- ), which could theoretically introduce 1.28 <sup>×</sup> 109 possible combinations on the seven amino acid residues of the loop. However, in practice, the library contained only 5.0 <sup>×</sup> 105 variants, less than 0.04% of the possible mutations. Despite the reduced number of variants, the authors managed to select a toxin mutant (R5–51) with strong binding affinity to the bead-immobilized cadherin-like protein BtR175. The selected variant, R5–51, was four times more toxic (LC50 1.6 μg/g diet) than the Cry1Aa1 wild type toxin (LC50 6.3 μg/g diet) toward *B. mori*.

Another case of success in the quest of improving the toxicity of a natural 3D-Cry toxin by phage display technology was the in vitro evolution performed on the moderately active Cry8Ka1 toxin toward the Coleoptera *Anthonomus grandis* [137]. This time, variability was obtained by *cry8Ka1* gene shuffling, which was cloned in the pComb3X phagemid [139] fused to the pIII protein in a M13 system. The resulting library, pCOMBcry8Ka1var, containing 1.0 <sup>×</sup> <sup>10</sup><sup>5</sup> cfu/mL variants, was screened toward *A. grandis* BBMVs. Biopanning rendered one variant (Cry8Ka5 mutant) that showed a 3-fold increase in toxicity. Sequence analysis of the Cry8Ka5 variant demonstrated that mutations were randomly introduced at different positions in Domain I (R82Q), Domain II (Y260C, P321A), and Domain III (R508G, K538E, E594N) of the toxin. In addition, a deletion of 16 residues at the N-terminal end was observed.

Non-natural 3D-Cry toxins have also been evolved by DNA shuffling and phage display technology [136]. This is the case of the Cry1Ia12synth toxin (NCBI gene bank accession number FJ938022), a synthetically derived toxin from Cry1Ia12 with a modified codon usage for plant expression optimization. Cyr1Ia12synth is toxic for *S. frugiperda*, but not for the sugarcane giant borer *Telchin licus licus*. From the 30 variants selected by phage display using *T. l. licus* BBMVs, four of them showed higher activity toward *T. l. licus* compared to the wild type toxin. Variant 1 (D233N, E639G), variant 2 (D233N), variant 3 (I116T, L266F, K580R), and variant 4 (M45V, D233N) showed 61%, 75% 56%, and 58% mortality, respectively, higher than the wild type and the negative control (25% mortality). This represents an example of the in vitro evolution of a 3D-Cry toxin in order to be active toward another Lepidoptera specie.

The latest report of a 3D-Cry toxin in vitro evolution using phage display technology was by Dominguez-Flores et al. [140]. In this case, a library of "Crybodies" was displayed on a λ phage system similar to the one reported by Vilchez et al. [130]. Crybodies are molecules derived from the lepidopteran active Cry1Aa13 toxin where loop 2 of Domain II has been replaced by the hypervariable region contained at the complementary determinant region 3 (CDR-H3) of a human antibody library [138]. The Crybody library was biopanned to *Ae. aegypti* larvae guts homogenates, and the selected phage, with high affinity toward gut proteins, was used to obtain the novel Crybodies. Crybodies Cry1Aa13-A8 (L2:367GAREGSSSAYDYW379) and Cry1Aa13-A12, (L2:367GARGDPDFDHSTSYYLDYC385) showed significant mortality (around 90%) after 120 h at 20 μg/mL, while no toxicity was observed in the parental Cry1Aa13 toxin. Concomitantly, both variants showed a 50% decrease in toxicity toward their natural lepidopteran target (*B. mori*). In this case, phage display was proven to be useful not only for improving toxicity against an insect or related species, but also to select variants active against insects from a different order.

Another example that used phage display technology in the field of 3D-Cry toxin evolution is the work reported by Shao et al. [141]. This work describes the construction of six 3D-toxin mutants, obtained by replacing loop 1, loop 2, and loop 3 in Domain II of the Cry1Ab toxin with what they called "gut-binding peptides" or GBPs. These peptides were obtained from a random peptide library displayed on a phage that was biopanned against BBMVs obtained from the hemipteran *Nilaparvata lugens* [141]. P2S (CLMSSQAAC) and P1Z (CHLMSSQAAC) were introduced by overlapping PCR in substitution of loop 1 (278RG279), loop 2 (335RRPFNIGINNQ345), and loop 3 (401SMFRSGFSNSSVS413) in the Cry1Ab toxin. *N. lugens* nymph bioassays showed increased toxicity in five of the six variants selected. Only mutant L3-P1Z was less toxic than the wild type (with an LC50 of 189.83 μg/mL). The rest of the mutant toxins (L1-P2S, L2-P2S, L3-P2S, L1-P1Z, and L2-P17) were 5, 9, 5, 1.4, and 2.5 times more toxic, respectively, than the parental toxin. Substitution of loops 1, 2, and 3 was concomitant with a loss in toxicity of Cry1Ab toward *P. xylostella*. This work demonstrated that the in vitro evolution of Cry toxins is not only restricted to the selection of variants with an improved binding to natural receptors, but also evolution can be directed to bind other molecules in the insect guts.

#### *2.8. Evolution by PACE (Phage-Assisted Continuous Evolution)*

Phage-assisted continuous evolution is a technique developed at Harvard University by David L. Liu´s research group [142]. It is one of the latest techniques developed in the field of the in vitro evolution of proteins. It is a complex technique, that, as its name implies, is performed with the assistance of a phage. Strictly speaking, it is not an in vitro technique, as evolution is performed inside of a highly engineered *E. coli* strain, but as it is performed in a laboratory, it is considered to be an in vitro evolution technique. The evolution is carried out in what is called "the lagoon" (Figure 4a), an *E. coli* culture with a constant inflow and outflow of growing media. The flow is set up at the appropriate speed to serve as the selection process for the mutants generated in the lagoon, as only the suitable and fast growing mutants stay in the lagoon, while the non-useful mutants are washed out from the growing flask. The average residence time of the cells is less than the *E. coli* replication time. The *E. coli* strain where the evolution takes place contains three plasmids (Figure 4b): (i) an arabinose-inducible mutagenic plasmid (MP), that contains proteins that disrupt the proofreading activity of DNA polymerases, so increasing the error rate in replication; (ii) a selection plasmid (SP), that contains the protein to be evolved and all the genes necessary for M13 phage replication except for *gen III*, essential for host infection; and (iii) an accessory plasmid (AP) where the essential gen III for M13 phage replication is expressed, but only if the right mutant is generated. With this system, the mutation process and the selection process are coupled as only the desired mutants allowing the expression of protein III are replicated. Non-useful mutations produce non-infective phage, so they will be unable to reproduce and will be washed out from the lagoon. The system solves the cumbersome need to screen the entire library in each round of evolution and, given the life cycle of M13 is just 10 min, a high number of rounds of protein evolution could be conducted in a single week. PACE has been used to evolve proteins such as polymerases [143], proteases [144], and genome-editing proteins [145], obtaining variants with completely novel activities and specificities.

**Figure 4.** Phage assisted continuous evolution (PACE). In the PACE technique, the evolution occurs in a continuous culture of a highly engineered *E. coli* strain called the lagoon (**a**). The *E. coli* strain contains three plasmids (**b**), the mutagenic plasmid (MP), containing mutagenic proteins induced by arabinose, the selection plasmid (SP), which contains all the M13 genes for phage replication, except for *gene III*, and a transcriptional fusion of the evolving *cry* gene toxin and the *rpoZ* gene, codifying for the omega sub-unit of the RNA polymerase, and the accessory plasmid (AP). AP plasmid contains the M13 *gene III* downstream of a cI site and a promoter. The fusion protein between cI and a fragment from *T. ni* cadherin like receptor (TnTBR3-F3) binds to the cI site. Only Cry toxin variants interacting with TnTBR3-F3 will bind to the proximity of the promoter site so the *gen III* expression will be possible, rendering viable infecting M13 particles. However, if Cry toxins with no affinity toward the TnTBR3-F3 fragment are produced, *gene III* will not be expressed and no infecting M13 particles will be generated. Reproduced from [146] Copyright 2016, Springer Nature.




**Table 1.** *Cont.*




**Table 1.** *Cont.*






1 the time.

PACE has recently been adapted for the evolution of a 3D-Cry toxin, specifically the lepidopteran Cry1Ac toxin [146]. The cabbage looper *T. ni*, naturally susceptible to Cry1Ac, has developed resistance to the toxin, a fact that has been associated with the mutation of the *ABCC2* transporter gene [147] and the downregulation of the expression of *APN1* [148]. There is no evidence that *T. ni* uses any cadherin-like receptors for its function, so the objective of this work was to evolve a Cry1Ac toxin to specifically bind to a cadherin-like protein (TnCAD), present in the insect cell membrane of *T. ni*, in order to be used as a toxin receptor. For that, the SP plasmid contained, apart from all phage genes (except for *gen III*), a transcriptional fusion of *cry1Ac* with the *rpo*Z gene codifying for the omega sub-unit of the RNA polymerase (Figure 4b). The omega sub-unit is essential for the activity of the RNA polymerase and unless it is present in the RNA polymerase enzymatic complex, the transcription is not possible. The AP plasmid contains M13 *gen III* with an upstream promoter region for the RNA polymerase, and the binding site for the cI protein, a phage repressor protein. In addition, a transcriptional fusion between the cI protein and a fragment of the TnCAD cadherin-like protein, called TnTBR3-F3, was included in the AP plasmid. This fusion protein is able to recognize the cI binding site, allowing the TnTBR3-F3 receptor to interact and bind to other proteins.

In this system, if a Cry toxin variant has the ability of interacting with the TnTBR3-F3 receptor as a consequence of the generated mutations, then it will bind at the promoter region of the M13 *gen III* through the omega sub-unit of the RNA polymerase, allowing the expression of the essential *gen III* for phage replication. If the introduced mutation is not suitable for TnTBR3-F3 binding, then the mutant will not be present in the promoter region, the protein III will not be produced, and the resulting phage will not be infective and it will be lost. The PACE system adapted to the evolution of a Cry1Ac toxin, rendered A01s, C03s, and C05s variants with high binding affinities to the membrane protein TnCAD, in opposition to the wild type toxin Cry1Ac. In addition, when these toxins were bioassayed against *T. ni*, they were 2.2, 1.1, and 1.8 times more active compared to the wild type Cry1Ac, respectively, indicating that toxin-receptor evolution had been taking place. Furthermore, when A01s, C03s and C05s were bioassayed against Cry1Ac resistant *T. ni*, toxin variants showed an increase in toxicity of 334-, 27.8-, and 26.4-fold compared to the wild type Cry1Ac toxin. This work represents a proof of concept that evolution of 3D-Cry toxins to bind novel receptors is possible through PACE and that the technique could be useful in cases where insects have developed resistance to natural toxins.

#### **3. Concluding Remarks**

The use of several molecular techniques has allowed researchers to obtain 3D-Cry toxin mutants with improved activities compared to natural toxins. Although in many cases the reason behind this enhancement is not known, the reality is that molecular techniques have been proven to be useful to develop artificial variants. From a practical point of view, these variants represent a real alternative to (i) the intrinsic limitation that 3D-Cry toxins show, as they are only active against a narrow range of insects, and (ii) the resistance phenomena that insects have experienced as a consequence of the extensive use of natural 3D-Cry toxins. This report is proof that minimal changes in the amino acid sequence of a 3D-Cry toxin can lead to a great improvement in toxicity, and that protein engineering, rational design, and in vitro evolution are powerful tools to develop artificial 3D-Cry toxins with surprising and novel activities. The compilation of all of these successful examples and the description of all the sensitive positions that have been used to obtain 3D-Cry toxin variants represents a valuable source of information for the further manipulation of natural toxins.

**Supplementary Materials:** The following are available online at http://www.mdpi.com/2072-6651/12/9/600/s1, Table S1: Sequence of some mutant toxins mentioned in the review.

**Funding:** This research was funded by the Andalusian Operative Program, Grant No. B-BIO-081-UGR18.

**Acknowledgments:** I would like to thank Guest Editors Juan Ferré, Yolanda Bel, and Patricia Hernandez-Martínez from the University of Valencia, Spain for the invitation to write this review for the Special Issue "*Bacillus thuringiensis* Toxins: Functional Characterization and Mechanism of Action". I wish to thank my mentor Prof. David Ellar from the University of Cambridge, UK, who introduced me to the exciting field of Cry toxin research. **Conflicts of Interest:** The author declares no conflict of interest.

#### **References**


© 2020 by the author. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).

### *Review* **The Cytocidal Spectrum of** *Bacillus thuringiensis* **Toxins: From Insects to Human Cancer Cells**

**Gretel Mendoza-Almanza 1, Edgar L. Esparza-Ibarra 2, Jorge L. Ayala-Luján 3, Marisa Mercado-Reyes 2, Susana Godina-González 3, Marisa Hernández-Barrales <sup>3</sup> and Jorge Olmos-Soto 4,\***


Received: 25 March 2020; Accepted: 2 May 2020; Published: 6 May 2020

**Abstract:** *Bacillus thuringiensis* (Bt) is a ubiquitous bacterium in soils, insect cadavers, phylloplane, water, and stored grain, that produces several proteins, each one toxic to different biological targets such as insects, nematodes, mites, protozoa, and mammalian cells. Most Bt toxins identify their particular target through the recognition of specific cell membrane receptors. Cry proteins are the best-known toxins from Bt and a great amount of research has been published. Cry are cytotoxic to insect larvae that affect important crops recognizing specific cell membrane receptors such as cadherin, aminopeptidase-N, and alkaline phosphatase. Furthermore, some Cry toxins such as Cry4A, Cry4B, and Cry11A act synergistically with Cyt toxins against dipteran larvae vectors of human disease. Research developed with Cry proteins revealed that these toxins also could kill human cancer cells through the interaction with specific receptors. Parasporins are a small group of patented toxins that may or may not have insecticidal activity. These proteins could kill a wide variety of mammalian cancer cells by recognizing specific membrane receptors, just like Cry toxins do. Surface layer proteins (SLP), unlike the other proteins produced by Bt, are also produced by most bacteria and archaebacteria. It was recently demonstrated that SLP produced by Bt could interact with membrane receptors of insect and human cancer cells to kill them. Cyt toxins have a structure that is mostly unrelated to Cry toxins; thereby, other mechanisms of action have been reported to them. These toxins affect mainly mosquitoes that are vectors of human diseases like *Anopheles spp* (malaria), *Aedes spp* (dengue, zika, and chikungunya), and *Culex spp* (Nile fever and Rift Valley fever), respectively. In addition to the Cry, Cyt, and parasporins toxins produced during spore formation as inclusion bodies, Bt strains also produce Vip (Vegetative insecticidal toxins) and Sip (Secreted insecticidal proteins) toxins with insecticidal activity during their vegetative growth phase.

**Keywords:** *Bacillus thuringiensis*; Cry; Cyt; parasporins; S-layer proteins; Vip; Sip; membrane receptors; insecticidal activity; anticancer activity

**Key Contribution:** This review focused on describing *Bacillus thuringiensis* proteins and their toxic and cytotoxic activity.

#### **1. Introduction**

*Bacillus thuringiensis* is a Gram-positive and sporulated bacterium that is widely distributed in soils, plants, and insects around the world [1,2]. Bt is well known because it produces a great variety of useful proteins for pest control in agriculture (Cry, Vip, Sip) [3,4], minimizes diseases transmitted by mosquitoes (Cyt) [5], inhibits pathogens development in animals [6], and because it induces cytotoxicity in human cancer cells (PS, SLP, and Cry) [7,8].

In 1901, Ishiwata found Bt for the first time in *Bombyx mori* and called it *Bacillus sotto*. In 1915, in Thuringia, Berliner isolated this bacterium from the moth *Ephestia kuehniella* and called it *Bacillus thuringiensis* [4].

During the sporulation process, Bt produces inclusion bodies (IB) in parasporal position with cubic, bipyramidal, spherical, oval, and irregular shapes that can be distinguished by scanning electron microscopy (SEM) (Figure 1) [9–11]. The IB are formed by a conglomeration of delta-endotoxin monomers classified according to the sequence similarities between two significant families; Cry and Cyt toxins [10–12]. In addition, Bt produces other important toxins such as parasporins, S-layer, Vip, and Sip proteins that will be discussed in this review.

**Figure 1.** Different morphologies of *Bacillus thuringiensis* crystals; (**A**) image observed at 40× in an optical microscope, the crystal was stained with malachite green. (**B**) Image observed at SEM with different magnifications from the left to the right (1) 20,000×, (2) 15,000×, (3) 15,000×, and (4) 50,000×. VC: vegetative cell; CC: cubic crystal; S: spore; BC: bipyramidal crystal; OC: ovoid crystal; SC: spherical crystal.

Cry and Cyt proteins received their current nomenclature after creation of the *Bacillus thuringiensis* Toxin Nomenclature Committee [13]. This Committee proposed a classification system of four hierarchical ranks based on the place each toxin occupies in the phylogenetic tree. Cry and Cyt delta-endotoxins with less than 45% sequence identity differ in primary level and are classified as Cyt1, Cry1, Cry2, etc. Cry and Cyt delta-endotoxins with 78% sequence identity differ in secondary rank and a capital letter is added to their name, e.g., Cyt1A, Cry1A, Cry2A. Toxins with 95% identity constitute

the border for a tertiary rank and small letters differentiate these proteins from each other, e.g., Cyt1Aa, Cry1Aa, Cry1Ab, Cry1Ac [13–15].

Parasporins have less than 25% amino acid sequence homology with Cry toxins [7]. However, their mechanism of action is very similar; both families recognize specific membrane receptors on cancer cells to trigger cell death [6]. Parasporins do not induce hemolytic activity but may or may not have insecticidal activity, nevertheless, they show preferential cytotoxicity against human cancer cells instead of healthy human cells in vitro [16].

Surface layer proteins (SLP) are embedded into cell membranes of many Gram-negative and Gram-positive bacteria; they are commonly associated with polysaccharides and peptidoglycans, respectively [9]. Main functions of SLP proteins are: (1) interaction with extracellular proteins, (2) protection against pathogens, (3) phagocytosis, (4) stabilization of membranes, and (5) adhesion, among others [17]. Unlike *cry* genes, which are expressed during the sporulation process, *s-layer* genes are constitutively expressed throughout the entire cell life cycle. According to previous reports, S-layer proteins from Bt are associated with toxicity against *Epilachna varivestis* [9,18]. Furthermore, there is a report from an S-layer protein with selective cytotoxic activity against MDA-MB-231 breast cancer cells line [8].

Bt synthesizes and secretes to the medium Sip (secreted insecticidal protein) and Vip (vegetative insecticidal protein) proteins during the exponential growth phase. There are reports about the insecticidal activity of these proteins against some coleopterans and lepidopterans [19–21].

Bt toxins have been isolated and classified into at least 78 Cry [13], 3 Cyt [13], 6 parasporins [7], 1 SLP [18], 1 Sip [11], and 4 Vip families [11].

#### **2. Leading Toxic Proteins of** *Bacillus thuringiensis* **and their Mechanism of Action**

#### *2.1. Cry Toxins*

Cry proteins are widely known by their toxic activity against insects belonging to orders such as *Hymenoptera*, *Coleoptera*, *Homoptera*, *Orthoptera*, and *Mallophaga*, as well as nematodes, mites, and protozoa [2,11,22,23]. Their toxic activity against insect larvae has allowed these toxins to be used for biological control of pests through spray formulations and transgenic crops that incorporate Cry proteins or some active fragment [22,24–28]. Tobacco was the first Bt crop produced by "Plant Genetic Systems" in Belgium in 1985 [29]. Since then, other crops, such as corn, cotton, potato, rice, brinjal, and soybean, have been genetically modified with Bt toxins to resist insect pests [30,31].

Cry toxins are highly specific to their target organisms. It is unusual to find a Cry toxin that targets more than one insect order, as is the case of Cry1Ba which is toxic to moths, flies, and beetles larvae [32]. *Cry* genes reside on plasmids that are naturally transferred from one Bt strain to another by conjugation or recombination [33,34]. This transfer of information plays an essential role in the biodiversity of Bt strains [34]. The final composition of *cry* genes in a strain determines the specificity and toxicity against biological targets, including human cells [3,15,34]. More than 700 *cry* genes have been classified into groups and subgroups, according to their amino acid sequence similarity [11,13].

X-ray crystallography of Cry proteins has evidenced three structural domains; hence, Cry toxins are also known as 3d-Cry toxins. The N-terminal Domain I is formed by seven α-helices, with a conserved hydrophobic helix α-5 in the center, which is related to oligomerization of the toxin [3,4,35]. Helix α-5 is also responsible for pore-formation in the membrane of susceptible cells, and for toxin insertion into the cell. Given these characteristics, Cry proteins are classified as pore-forming toxins (PFT). In this sense, Domain I is the most conserved among all Cry toxins, sharing some structural similarity with Colicin Ia, another PFT [3,4,36,37].

Domain II is composed of three antiparallel β-sheets that form a hydrophobic core, with highly variable exposed loop regions. This domain is responsible for toxin specificity; therefore, indicates the binding sites into receptors in susceptible larvae [3,38–40].

Domain III is composed of antiparallel β sheets that form a β sandwich structure. This domain is also involved in receptor binding specificity; additionally, it is also associated with pore formation in cell membranes [3,38–40].

Cry toxins belong to the PFT family due to their mechanism of action, in which Domain I is inserted into the membrane of target cells, creating a trans-membrane ion channel and triggering the host's death [2,3,41]. The PFT family comprises different types of toxins, including, among others, Colicin family, produced by Escherichia coli [42]. The ClyA family is produced by *Escherichia coli* and *Salmonella* enteric strains [43]. The Actinoporin family is produced by sea anemones [44]. The Haemolysin family, produced by *Staphylococcus aureus* [45]. The Aerolysin family, produced by *Aeromonas hydrophila* [46]. The CDC family, produced by pathogenic Gram-positive bacteria such as *Clostridium perfringens*, *Bacillus anthracis*, and *Streptococcus pneumoniae* [47]. The toxins from the PFT family have several characteristics in common: (1) The way they fold, which suggests all share a similar mechanism of action [48–51]; (2) All recognize specific receptors on cell membranes [48–51]; (3) Promote oligomerization at the interaction site after receptor recognition [48–51].

PFT are classified into two main groups according to their secondary structures, which are responsible to the formation of pores: toxins from α-helical group includes Colicin, Exotoxin A, Diphtheria, and Cry toxins. In this group, the α-helix region is responsible for the trans-membrane ion channel formation [48–50]. On the other hand, β-barrel toxins include Aerolysin [50], Hemolysin [45], Perfringolysin O [52], and Cyt toxins [53]. These toxins insert themselves into the cell membrane, forming a barrel composed of β sheets hairpin monomers [48–50].

#### Mechanism of Action from Cry Toxins

3d-Cry proteins are produced as large protoxins with a molecular weight around 130 kDa, such as Cry1Aa protein [3,4,11,14], or short protoxins between 65 and 70 kDa, such as Cry11Aa protein [4,14]. The large protoxins are processed by insect midgut proteases at C-terminal and N-terminal ends [3,4,11,14], while short protoxins are processed only at the N-terminal end [4,14]. In both cases, protease-resistant core results in an active Cry toxin of 60 and 70 kDa approximately which retains the 3d structure [1,4,11,14]. The resistant fragment is responsible for cytotoxicity against larvae insects, nematodes, protozoans, and human cancer cells [1,4,11,14]. However, incorrect or deficient protoxin activation, and/or rapid degradation of toxins by other proteases, could reduce the toxicity of Cry proteins against their target [4,14].

#### (a) The Pore-Forming Model

The most accepted mechanism of action of Cry toxins against insect larvae is the pore-forming model [33,37,54,55], which is summarized in Figure 2A. Once larvae ingest toxin crystals, these solubilized at extreme pH (acid or alkaline, depending on the Cry toxin) and proteolyzed by proteases under suitable physicochemical conditions on the midgut. The activated toxins can reach the apical brush border membrane (microvilli) of the insect's midgut by crossing the peritrophic matrix. Cry toxins must recognize receptors in brush-border membranes to form pores; therefore, specificity is crucial for Cry proteins toxicity [3,55,56]. In this sense, Domain I needs to be inserted into the cell membrane through its hydrophobic helical hairpin [55,57]. The amphipathic helices attach to the surface of membranes using hydrophobic helices α-4 and α-5 to enter into the phospholipid bilayer [55,57]. In addition, highly variable and exposed loops from Domain II also participate in binding to receptors, a process that apparently involves two steps. The first step consists of recognition of aminopeptidase N (APN) and alkaline phosphatase (ALP) receptors and the formation of a weak binding with Cry toxins [11,55–59], which produces a reversible reaction [10,55].

The second step consist in the formation of an irreversible binding (Kd 19 nM) through recognition of a 12 amino acid ectodomain region (EC12) from the cadherin receptor (BT-R1) [10,60–62]. Conserved sequence motifs near the N and C ends of EC12 have been reported to be crucial for binding of toxins in insect cells [10,61,62]. Bt-R1 is a highly specific and selective binding receptor to Cry1Ab toxin, it was identified for the first time in the midgut of *Manduca sexta* larvae; this receptor is also responsible for Cry1Ab oligomerization [60–62].

BT-R1 is a protein of 210 kDa composed of four domains: (1) an ectodomain with twelve modules (EC1-EC12) composed of β-barrel cadherin repeats; (2) a membrane-proximal extracellular domain; (3) a transmembrane receptor; (4) a cytoplasmic domain [10,60,63]. The interaction between Cry1A toxin and BT-R1 facilitate the proteolytic cleavage of helix α1 from Domain I, which is located at the N-terminal end, resulting in the formation of a pre-pore oligomer structure that increases the affinity between oligomer and membrane receptors APN and ALP [35,62,64]. The oligomer inserted into the cell membrane creates an ionic pore that leads to osmotic failure, followed by septicemia and insect death [2,12,23,41]. Other intracellular molecules, such as actin, flotillin, prohibitin, and V-ATPase, have been found to participate in the binding to Cry toxins [11,65].

Domain III is a key structure in toxin stability, it binds to N-acetylgalactosamine (GalNAc) in the APN receptor [55,66,67]. APN has been identified as a binding receptor for Cry1A toxins in *M. sexta*, *H. dispar* [55,66,67], and *B. mori* [55]. ATP-binding cassette transporters (ABC proteins) are also involved in Cry toxicity, especially member 2 of subfamily C (ABCC2). These proteins may help Cry1A toxins carry out their primary task: binding to receptors and inserting oligomers into the cell membrane [68,69].

**Figure 2.** *Cont*.

**Figure 2.** Mechanism of action of Cry proteins. (**a**) Pore-forming model, once larvae ingest crystals, these are solubilized and proteolyzed in larvae midgut. Cry toxins recognize APN, ALP, and EC12/BT-R1 membrane receptors. Cry toxins suffer a proteolytic cleavage on helix α1, resulting in formation of a pre-pore oligomer structure. Posteriorly, oligomer structure is inserted into the cell membrane and creates an ionic pore that leads to osmotic failure, followed by septicemia and insect death. (**b**) Signaling pathway model, once Cry toxins recognize and bind to a cadherin receptor, induces activation of adenylyl cyclase that triggers an increase in cAMP and activates protein kinase A (PKA). This activation will induce a cascade of events that results in an ion channel formation in the membrane, cytoskeleton destabilization, and programmed cell death. A, Solubilization. B, Activation by proteolysis. C, Recognition of membrane receptor. Created with Biorender.com.

However, recently published evidence has shown that toxicity of Cry1AbMod and Cry1AcMod (Cry toxins whose structure has been modified by deleting helix α1) against *M. sexta* that does not involve the expression of the cadherin receptor and still can form toxic oligomeric structures [70–72]. This finding is of great importance for development of strategies to counteract resistance in transgenic crops, and to increase our knowledge of the mechanism of action of Cry toxins.

#### (b) The Signaling Pathway Model

Cell culture toxicity assays developed to elucidate the action mechanism used by Cry toxins in the signaling pathway model have been carried out in High Five (H5) cell line; which involves expression of a cadherin receptor from *Manduca sexta* [73,74] in a Sf9 cell line from lepidopterans [74,75]. The signaling pathway model postulates that Cry proteins cytotoxicity is mediated by recognizing and binding to a cadherin receptor, which activates an Mg2<sup>+</sup>- dependent cellular signal cascade pathway that leads to cell death [74]. The binding between Cry toxin and cadherin receptor induces the activation of adenylyl cyclase; which triggers an increase in cAMP and activates protein kinase

A (PKA). These events trigger a cascade that results in an ion channel formation on the membrane, cytoskeleton destabilization, and programmed cell death (Figure 2B) [76].

#### *2.2. Cyt Family*

Cyt toxins size ranges from 25 to 28 kDa; their three-dimensional structure shows Cyt proteins have a single α-β domain with low sequence homology to Cry toxins [77]. Cyt toxins affect mainly mosquitoes that are vectors of human diseases; *Anopheles spp* (malaria), *Aedes spp* (dengue, zika, and chikungunya), and *Culex spp* (Nile fever and Rift Valley fever) [78]. Bt subsp. israelensis (Bti) is the most commonly used worldwide to control these vectors [3,78], because it produces Cry4Aa, Cry4Ba, Cry10Aa, Cry11Aa, Cyt1Aa, Cyt2Ba, and Cyt1Ca toxins that together act synergistically to kill mosquito larvae of Culex, Aedes, and Anopheles [79,80] (Table 1).

**Table 1.** Cyt toxins could synergize with several Cry toxins to act against mosquitoes.


Furthermore, Cyt proteins have other important biologic targets, among others, mammalian and erythrocyte cells [81]; and the pea aphid (*Acyrthosiphon pisum*) and certain weevils (*Diaprepes abbreviates*) [82], both pest insects where Cyt toxins could be used as biocontrol.

Three subfamilies of Cyt toxins have been described so far, all of them sharing a high level of sequence identity: Cyt1 (1Aa, 1Ab, 1Ab, 1Ac, and 1Ad), Cyt2 (2Aa, 2Ba, 2Bb, 2Bc, and 2Ca), and Cyt3Aa1 [10]. Cyt1Aa shows significant similarity with volvatoxin 2 (VVA2), a cardiotoxin isolated from the mushroom *V. volvacea* [83]. VVA1 and VVA2 form the VVA toxin family, a PFT family that has hemolytic and cytotoxic activity in human red blood cells and tumor cells, respectively [84]. Cyt1Ca is different from other members of the Cyt family, since they have an extra domain in the C-terminal end with homology to a carbohydrate-binding domain of ricin. However, there are no reports of larvicidal or hemolytic activity for Cyt1Ca [11,85].

The overall structure of Cyt toxins is formed by a β sheet consisting of six antiparallel strands flanked by a α helix layer and an extra strand β0 at the N-terminal end, which could be involved in dimerization and proteolytic activation [85]. Notoriously, an extra strand at the N-terminal end has not been reported in Cyt2 toxins.

#### Cyt Proteins Mechanism of Action

Cyt proteins are synthesized as short protoxins [3,86]. The proteolytic cleavage sites in Cyt toxins are found at N-terminal and C-terminal ends [77]. It is widely recognized that loops of the helices are involved in membrane–cell interaction and intermolecular assembly [80]. However, the action mechanism followed by Cyt toxins is still not clear; moreover, it is unknown whether there are specific receptors through which Cyt toxins recognize their target cells. Nevertheless, there are currently two main models of the cytotoxicity mechanism carried out by Cyt toxins.

#### (a) The Pore-Formation Model

The pore-formation model describes binding of Cyt toxins in their monomer form to specific receptors in the membrane surface, similar to the Cry toxin mechanism. In this case, Cyt toxins interact directly with saturated membrane lipids such as phosphatidylcholine, phosphatidylethanolamine, and sphingomyelin [80,81]. Cyt toxins undergo a conformational change that helps to recruit six monomers of Cyt and assemble them into an open-umbrella structure in which the strands span the lipid bilayer transversely, while alpha helices rest on the membrane surface. The result is a pore-forming model with membrane permeabilization (Figure 3a) [80].

#### (b) The Detergent-Effect Model

The detergent-effect model suggest that Cyt toxins kill target cells through a solubilization effect on their membrane. In this model, Cyt toxins concentrate on the cell membrane surface and destroy the lipid bilayer in a detergent-like manner (Figure 3b) [80].

Both the pore-formation model and detergent model are not mutually exclusive, as it is thought that, depending on toxin concentration, one or both may act on susceptible cells [80]. Cyt oligomerization and pore formation could be carried out at low Cyt concentration [87], while the detergent effect could be induced only at high toxin concentration [87]. Therefore, the cell membrane from target cells is unable to assemble oligomers at high toxin concentration; instead, it forms a toxin–lipid complex in which the integrity of the membrane is completely lost [80].

Another mechanism of action of Cyt toxins involves a synergistic activity between different members of the family. Thus, when Cyt1Ab and Cyt2Ba act together, they enhance the insecticidal activity against *Aedes aegypti* larvae and resistant *Culex quinquefasciatus* larvae [88]. It has also been found that two known epitopes of Cyt1Aa (196EIKVSAVKE204 and 220NIQSLKFAQ228) binds to Cry4B and Cry11Aa toxins to enhance their toxic effect against the mosquito *Anopheles albimanus* and *Culex quinquefasciatus* [15,84,89]. Epitopes of Cyt1Aa play a receptors role to Cry4B and Cry11Aa, similar to the cadherin receptor of *Manduca sexta*. When Cyt1Aa binds to membrane cell receptors, Cry11Aa or Cry4B binds to this toxin, increasing oligomerization and pore formation effects [3,89,90]. Another synergistic effect occurs when Cyt toxins act together with Mtx1 (another toxin produced by Bt) against *Culex quinquefasciatus* larvae [91].

**Figure 3.** Mechanism of action of Cyt proteins. (**a**) Pore-forming model; once Cyt toxins interact with phosphatidylcholine, phosphatidylethanolamine, and sphingomyelin, they undergo a conformational change that helps recruit six Cyt monomers and assemble them into an open-umbrella structure, which results in a pore-formation and subsequent membrane permeabilization and larva death. (**b**) Detergent effect model; high Cyt toxin concentration binds to the lipid bilayer on cell membrane surface and destroys it through a detergent-effect. Both, the pore-formation model and detergent model are not mutually exclusive, because, the detergent effect acts at high Cyt toxins concentrations, while the pore-forming model acts at low Cyt toxins concentrations. A, Solubilization. B, Activation by proteolysis. C, Recognition of membrane receptor. Created with Biorender.com.

In recent years, several studies have been published related to Cyt proteins modification throughout genetic engineering to produce chimeric toxins [82,92]. These modifications take advantage of small Cyt toxin sizes and low degree complexity of their quaternary structure, as well as their high toxic capacity. The advantages of creating chimeras is to diversify their targets and to increase Cyt toxin potency.

The first successful report of a Cyt chimeric creation describes the insertion of a pea aphid gut-binding peptide GBP3.1, into the amino acid sequence of Cyt2Aa toxin. This peptide prevents the uptake of a plant virus by its vector, the pea aphid. It comprises 12 amino acids (TCSKKYPRSPCM), which bind to alanyl aminopeptidase-N on membrane surface of the aphid gut epithelium. Naturally, Cyt2Aa has low toxicity against the pea aphid, however, Chogule and coworkers succeeded in enhancing the binding of Cyt toxin and increasing its toxicity against aphids, by turning it into a chimeric protein [82].

In another study, Torres-Quintero et al. [92] modified Cyt1Aa by inserting the amino acid sequence of loop3 from Domain II of Cry1Ab (FRSGFSNSSVSI), which induces binding affinity of Cyt1Aa toxin to APN and CAD receptors of *Manduca sexta* [91]. Naturally, Cyt1Aa is not toxic to *M. sexta*, however, chimeric toxin had more significant toxicity to *M. sexta* and *Plutella xylostella* [92]. These results open new possibilities to the application of delta-endotoxins from *Bacillus thuringiensis*, to a new target pest.

#### *2.3. Parasporins*

Extensive screening analysis to find new possible targets to Bt strains has shown that non-insecticidal Cry proteins are more widely distributed in nature than insecticidal Cry proteins [16,93,94]. This fact has led researchers to inquire about the possible biological activities or targets of non-insecticidal and non-hemolytic Cry proteins. In this sense, Mizuki et al., in 1999, was the first group to report delta-endotoxins from Bt with selective cytotoxicity against leukemia cells, after a large-scale screening analysis involving protease-digested parasporal proteins from 1744 Bt strains. 1700 strains were isolated in Japan, while 44 were obtained from the Pasteur Institute in Paris [95]. From the isolated strains, 60 presented hemolysis activity and were eliminated by containing *cyt* genes, the rest of strains were tested in vitro for cytocidal activity against MOLT-4 cells and insecticidal activity [95]. At the end of the screening, authors selected only two strains (A1190 and A1462), because they produced toxins that selectively killed leukemic cells instead of normal T-cells [95]. These results inspired an intense and extensive screening of non-insecticidal and non-hemolytic toxins with cytotoxicity against cancer cells throughout the world, which led to the classification of a new type of Cry proteins called parasporins (PS) [96]. At first, this new classification included all non-insecticidal and non-hemolytic Cry toxins with selective cytotoxic activity against cancer cells [7,96]. Time later, it was accepted that non-hemolytic but insecticidal activity could also be part of this classification [96]. Bt strains that produce PS have been isolated from soils of various ecosystems in several countries such as Japan [95,97–120], Vietnam [121,122], India [94,123], Malaysia [124], China [125], Iran [126], and Saudi Arabia [127]. Canada [128] and Caribbean [129] have contributed with new parasporins.

In nature, there are several toxins produced by bacteria capable of killing mammalian cells through pores formation in cell membranes and/or by apoptosis activation [51]. Such is the case of aerolysin from *Aeromonas hydrophila* and alpha-toxin of *Clostridium perfringens*, which are both PFTs that recognize GPI-anchored proteins in the membrane of susceptible cells [51]. Some parasporins share structural homology with both toxins, therefore, it is assumed that PS proteins contain an action mechanism similar to aerolysin and alpha-toxin [130]. So far, 19 parasporins have been identified and organized in six families (Table 2) [7,96].

#### 2.3.1. Parasporin Classification

#### PS1 Family

PS1 family has cytotoxic effects against certain cancer cell lines such as HeLa (cervix cancer) [99], HL60 (leukemia) [109], Jurkat (leukemia), and HepG2 (liver cancer) [128]. Findings suggest PS1 toxins

recognize a common receptor contained between these cell lines, identified as beclin-1 [131]. In healthy cells, beclin-1 exists intracellularly and is involved in autophagia and apoptosis processes, however, in susceptible cell, beclin-1 exists extracellularly and acts as a PS1 receptor [132].

The PS1 family includes, PS1Aa1 (Cry31Aa1), PS1Aa2 (Cry31Aa2), PS1Aa3 (Cry31Aa3), PS1Aa4 (Cry31Aa4), PS1Aa5 (Cry31Aa5), PS1Aa6 (Cry31Aa6), PS1Ab1 (Cry31Ab1), PS1Ab2 (Cry1Ab2), PS1Ac1 (Cry31Ac1), PS1Ac2 (Cry31Ac2) [96].

#### PS2 Family

PS2Aa1 (Cry46Aa1), PS2Aa2 (Cry46Aa2), and PS2Ab2 (Cry46Ab1) proteins constitute the PS2 family [96]. It is reported that the Bt A1547 strain produce PS2Aa1 and PS2Aa2 [95]. Parasporins from this family are produced as small toxins of around 30 kDa that are cytotoxic to cancer cell lines like HepG2 (liver cancer), Sawano (endometrial cancer), HL60, CaCo-2 (colon cancer), Jurkat (leukemia), and MOLT-4 [133,134].

A peculiarity of PS2 family toxins is that they do not have a typical 3d structure; instead, they are very similar to PFT-aerolysin which is formed mainly by elongated β sheets [133,134]. In 2009, Akiba et al. [134] proposed that PS2 toxins were able to produce oligomers that induce membrane pores and cell death by bind to lipid rafts. In 2017, Abe et al. reported that GPI was an essential co-receptor to PS2 parasporins toxic activity [135]. The action mechanism of PS2 family members is apparent by activating apoptosis, which is associated to an increase in the tumor suppressor gene PAR-4 expression and through inhibition of the PI3K/AKT pathway [136].

#### PS3 Family

PS3Aa1 (Cry41Aa1) is the single member of this family [96]. This parasporin is the only one with a ricin domain that plays a role in stabilization of the interaction between toxins and carbohydrate residues of the membrane [137]. In 2018, a research group led by Crickmore studied the PS3 protein and observed that it is structurally related to insecticidal toxins, except for the ricin domain. Using site-directed mutagenesis, they concluded that ricin domain is not associated with PS3 selective cytotoxic activity against the HepG2 cancer cell line [138].

In contrast to the mechanism of action used by PS2, it was proposed that PS3 induced cell death by necrosis throughout a pore formation in cancer cell membranes, as was evidenced by an increase in lactate dehydrogenase (LDH) release; mainly in HL60 and HepG2 cancer cell lines [138].

#### PS4 Family

PS4 is similar in size and structure to PS2 family members, its active form is around 27 kDa and presents a selective cytotoxicity against MOLT-4, CaCo-2, HL60, U937, HepG2, Sawano, DE-4 (leukemia), TS (uterine cancer), and TCS (cervical cancer) cancer cell lines [114,115]. A peculiarity of PS4 toxin is that it can be activated in both alkaline and acid pH [129], while most of parasporins are solubilized in alkaline pH. Actually, acid pH increases their cytotoxic effect against several cancer cell lines [115].

Regarding the mechanism of action of PS4, the evidence found (non-specific binding to the membrane, release of LDH and entrance into the cell of dextrans with different molecular weights) suggests that cell death occurs by necrosis [114,115,130]. The PS4 family includes PS4Aa1 (Cry45Aa1) as the only member [96].

#### PS5 Family

PS5Aa1 (Cry64Aa1) is the only member of the PS5 family; this protein has been isolated from the *Bt tohokuensis* A1100 strain [96]. The active form of PS5Aa1 has an approximate size of 30 kDa and is selectively toxic to MOLT-4 [120]. Concerning sequence similarity, PS5 shows higher similarity to PFT aerolysin than the other parasporins [120,139]. It has been reported that PS5 is toxic to MOLT-4, Jurkat, HL-60, HepG2, HeLa, Sawano, TCS, CaCo-2, and K562 cancer cell lines, but it also shows potent

activity against healthy tissue cells such as UtSMC (normal uterus) and MRC-5 (normal lung) [120]. However, there is still no evidence about its mechanism of action.

#### PS6 Family

PS6 is closely related to PS1, sharing a conserved sequence of fifty amino acids. PS6 is selectively toxic to HepG2 and HeLa [117], but its mechanism of action is still unknown. The PS6 family includes only PS6Aa1 (Cry63Aa1), which has been isolated from Bt M019 [96,117] and 64-1-94 [129].

#### 2.3.2. Mechanism of Action of Parasporins

The cytocidal activity of PS against cancer cell lines ranges from EC50 0.0017 μg/mL of PS2 against Sawano to 3.0 μg/mL of PS1 against HepG2. Table 2 shows the reported EC50 to PS in cancer cell lines.

In management of parasporins, different methods of crystals solubilization and activation should be tried, since effectiveness of their cytocidal activity against cancer cells depends on this. Similar to what happens on insects, a correct activation of protoxins is essential for cell membrane receptors recognition and subsequent triggering of cancer cell death.

As an example, there is a particular case of PS2Aa1 not showing toxic activity when activated using trypsin, but, when activated with proteinase K, it shows activity against human cancer cells [137]. In this sense, it is crucial to know that sites of cleavage to trypsin and proteinase K are different.

The mechanism of action of parasporins against target cancer cells is poorly understood, however, available information has shown that parasporins exhibit several mechanisms of action to kill cancer cells. These proteins act in a similar way to Cry toxins because they are highly specific to a cell type, nevertheless, it is well known that Cry toxins specificity depends on cell membrane receptors (cadherin, aminopeptidase-N, and alkaline phosphatase) recognition. On the other hand, PS interaction with cell membrane receptors is still being investigated, several molecules that act as parasporin receptors have been reported and patented. In this sense, Beclin-1 acts as PSAa1 receptor [131]. Glycosylphosphatidylinositol (GPI)-anchored protein is involved in efficient cytocidal action of PS2Aa1 [140]. GADPH from CEM-SS leukemic cell line acts as receptor from a PS found in Malaysia [124]. One of the most important characteristics of parasporins is their ability to discriminate cancer cells from non-cancer cells, which is directly related to cell membrane receptors recognition.



NP- not published. ND- not determined.

#### *Toxins* **2020**, *12*, 301

#### *2.4. S-Layer Proteins*

Surface layer proteins (SLP) are widely represented, both in Gram-negative and Gram-positive bacteria, including *Bacillus* [17]. Similar to delta-endotoxins, SLP are assembled into parasporal positions with several shapes (oblique, square, or hexagonal) [8]. They have a molecular mass between 40 and 170 kDa [141,142], and are involved mainly in growth, survival, and maintenance of cell integrity [9]. There are also reports of their antiviral and antibacterial activity, as well as of their anti-inflammatory effects [141,142].

SLP toxins activity is still unclear; it has been suggested that SLP have a similar insecticidal activity to Cry proteins but with a different mechanism [141,142]. The SLP obtained from GP1 Bt strain is the only one that has been reported to have pesticidal activity against *Epilachna varivestis* [18].

A recent study reported an SLP from Bt with high selective cytotoxic activity in vitro against the MDA-MB-231 breast cancer cell line. Authors suggest that cadherin-11 receptor present in cancer cells seems to be involved in SLP recognition; however, the mechanism of action is still under study [8].

#### *2.5. Toxins Secreted by Bt*

In addition to Cry, Cyt, PS, and SLP toxins produced in parasporal bodies during sporulation, Bt secretes during vegetative growth phase other toxins with insecticidal activity [11]. There are two main families of secreted insecticidal proteins, one is known as vegetative insecticidal proteins (Vip) [20,21] and the other as secreted insecticidal proteins (Sip) [19]. These proteins contain a signal peptide sequence in the N-terminal end that is cleaved after the secretion process is completed [143,144].

#### 2.5.1. Vip Family

Vip toxins are not shaped as parasporal inclusion bodies, instead they are produced and secreted during the vegetative growth phase and their expression ends before the sporulation stage begins [11]. These insecticidal toxins have been characterized as; Vip1, Vip2, Vip3, and Vip4; however, their mechanism of action against insects is not entirely understood yet [20,28].

Vip1 is synthesized as a protoxin of 100 kDa and after secretion; a mature toxin of 80 kDa is produced, additionally, Vip2 releases a trypsin-resistant fragment of 50 kDa [144]. Together, Vip1 and Vip2 produce a Vip binary toxin [145], with synergistic insecticidal activity against some coleopteran pests and the sap-sucking insect pest *Aphis gossypii (Hemiptera)* [146].

Vip2 is similar in structure and behavior to the CdtA toxin from *Clostridium di*ffi*cile*; this toxin presents an ADP-ribosyltransferase activity and its principal target is the actin protein, therefore, could induce cytoskeletal disruption and cell death when it is activated [147].

In monomer form, Vip1 binds to its receptors and a conformational change is produced because of this interaction; thus, more Vip1 toxins are attracted to form a heptamer that translocates Vip2 into the cytoplasm through acid endosomes [148]. Once inside the cell, Vip2 destroys actin filaments that disrupt the cytoskeleton and eventually induce cell death [148].

Due to its similarity with other A+B binary toxins, it has been concluded that Vip2 is responsible for most of the cytotoxic activity, while Vip1 is responsible for binding to membrane receptors in susceptible insects [149].

Vip3 is a single-chain protein that is toxic to a wide variety of lepidopterans and other insects, such as *Agrotis ipsilon*, *Spodoptera exigua*, and *S. frugiperda*, which are less susceptible to Cry1A toxins [21]. Vip3 are proteins of 88 kDa approximately without homology to any other known insecticidal protein [86,150]. In contrast to Vip1 and Vip2, signal peptide sequences in Vip3 are not processed during secretion and are present in the mature secreted peptide, suggesting they play an important role in protein structure and insecticidal activity. However, cleavage of the N-terminal end activates the protoxin; the 66 kDa active toxin is fragmented from the 22-kDa N-terminal portion [21]. The mechanism of action is still unclear and it has been suggested that Vip3 proteins act in a similar

way to PFT, but their membrane receptors are still unknown. In vitro experiments have shown that Vip3 does not compete for binding sites of Cry1A in *Manduca sexta* nor *S. frugiperda* [151].

Vip 4 is the most recently discovered member of the Vip family. It has a molecular mass of ~108 kDa and a 34% identity to Vip1Aa1 protein, specifically to the B component of the binary toxin. For this reason, it has been suggested that Vip4 might interact with unknown A component to produce toxicity; therefore, such information is needed to understand its action mechanism [20].

The Vip proteins that have been reported so far are 15 Vip1 proteins, 20 Vip2 proteins, 101 Vip3 proteins, and 1 Vip4 [13].

#### 2.5.2. Sip Toxins

Secreted insecticidal proteins (Sip) are toxins produced by Bt with an approximate size of 41 kDa. Similarly to Vip proteins, Sip toxins are synthesized containing a signal peptide sequence of 30 amino acids, which is processed by proteases [19] and an active protein is released [11]. It is known that Sip proteins have insecticidal activity against Coleopterans such as *Leptinotarsa decemlineata*, *Diabrotica undecimpunctata howardi*, and *Diabrotica virgifera virgifera* [19]. However, their mechanism of action is still unknown.

#### **3. Conclusions**

The horizontal transfer of genetic information through the conjugation of plasmids in *Bacillus thuringiensis* opens up a world of possibilities for the discovery of new toxins, new structures, new targets, and even new classification.

Understanding the structural characteristics of Bt toxins and their mechanism of action will allow us to develop new products for improving pest management and human health. In this sense, new combinations of insecticidal Cry proteins have been recently found, opening new possibilities to pest control without genetic and neither molecular manipulation [152]. Additionally, in 2019, Mendoza and coworkers for first time reported that Cry1A toxins from Bt presented a highly specific anticancer activity in HeLa cells and also against insects. Authors suggested that in both cases, a specific interaction between Cry toxins and cell membrane receptors could be initiating toxicity on insects and in human cancer cells [153].

SLP proteins are other Bt toxins with both pesticide and anticancer activity. Recently studies have shown that these proteins also could be recognizing specific cell membrane receptors in cancer cells line [8], as Cry toxins do [153]. In addition to cytotoxic activity on insects and human cancer cell lines, SLP carry out structural and protection activities in several microorganisms. Furthermore, these proteins could be found in Gram-positive, Gram-negative, and archaebacteria, not only in Bt.

Parasporins also present specific anticancer activity in vitro; these proteins are found in non-insecticidal and non-hemolytic strains, opening the possibility to develop anticancer agents with therapeutic potential and without secondary effects in patients. However, very little is known about their mechanism of action and the receptors recognized to carry out their cytotoxicity. PS1 and PS2 are the most extensively studied parasporin families, therefore, have been used as a model to answers several questions regarding preferential activity of these toxins against cancer cells.

Finally, it is important to mention that more research is needed to understand the mechanisms of action used by Bt toxins. According to reported studies, it seems that most of them recognize specific cell membrane receptors in susceptible cells; however, what is happening inside cells once the interaction has begun still is a mystery. Thus, it is essential to know and understand the signaling pathways involved in toxicity to be capable of developing new anticancer compounds and to improve pest control including resistance developed to Bt toxins.

**Author Contributions:** G.M.-A., E.L.E.-I., J.L.A.-L., M.M.-R., S.G.-G., M.H.-B., and J.O.-S., were responsible for cited literature and writing the manuscript. G.M.-A. and J.O.S. Writing Original Draft Preparation. G.M.-A. was responsible of the Conceptualization. All authors read and approved the final manuscript.

**Funding:** This research received no external funding.

**Acknowledgments:** G.M.-A. thank National Council for Science and Technology, Mexico, for the program Cátedras CONACYT.

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**


© 2020 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).

#### *Review*

## **Insecticidal Activity of** *Bacillus thuringiensis* **Proteins against Coleopteran Pests**

#### **Mikel Domínguez-Arrizabalaga 1, Maite Villanueva 1,2, Baltasar Escriche 3, Carmen Ancín-Azpilicueta <sup>4</sup> and Primitivo Caballero 1,\***


Received: 4 June 2020; Accepted: 25 June 2020; Published: 29 June 2020

**Abstract:** *Bacillus thuringiensis* is the most successful microbial insecticide agent and its proteins have been studied for many years due to its toxicity against insects mainly belonging to the orders Lepidoptera, Diptera and Coleoptera, which are pests of agro-forestry and medical-veterinary interest. However, studies on the interactions between this bacterium and the insect species classified in the order Coleoptera are more limited when compared to other insect orders. To date, 45 Cry proteins, 2 Cyt proteins, 11 Vip proteins, and 2 Sip proteins have been reported with activity against coleopteran species. A number of these proteins have been successfully used in some insecticidal formulations and in the construction of transgenic crops to provide protection against main beetle pests. In this review, we provide an update on the activity of Bt toxins against coleopteran insects, as well as specific information about the structure and mode of action of coleopteran Bt proteins.

**Keywords:** *Bacillus thuringiensis* proteins; coleopteran pests; insecticidal activity; structure; mode of action

**Key Contribution:** This contribution provide an update on the activity of Bt toxins against coleopteran insects.

#### **1. Introduction**

The use of entomopathogenic microorganisms as biological control agents has become one of the most effective alternatives to chemical pest control. Among all, the Gram-positive bacterium *Bacillus thuringiensis* (Bt) is the most important entomopathogenic microorganism used to date in crop protection. This bacterium is widely distributed in various ecological niches, such as water, soil, insects, and plants [1]. The feature that distinguishes *B. thuringiensis* from other members of the *Bacillus* group is the capacity to produce parasporal crystalline inclusions. These crystals are composed of proteins (Cry and Cyt) which are toxic against an increasing number of insect species from the orders Lepidoptera, Diptera, Coleoptera, Hymenoptera, and Hemiptera, among others, as well as against other organisms such as mites [2] and nematodes [3]. Bt also synthesizes insecticidal toxins associated with the vegetative growth phase, named Vip (vegetative insecticidal protein) and Sip (secreted insecticidal protein), which are secreted into the growth medium [4]. These toxins are uniquely specific, safe, and completely biodegradable, and have been used for more than 60 years as an alternative to chemical insecticides [5]. Products based on Bt isolates are the most successful microbial insecticides, with current worldwide benefits estimated at \$8 billion annually [6]. However, not all Bt proteins are designated as toxins, for example, some parasporins do not have known insect targets, although they are toxic to human cancer cells [7]. The insecticidal activity of Bt toxins has also been transferred to crop plants through genetic engineering, providing very high protection levels against injurious pests and decreasing the use of chemical insecticides in many instances [8,9]. The success of these insecticidal proteins has fuelled the search for new Bt isolates and proteins that can render novel insecticidal agents with different specificities.

Since Schnepf and Whiteley cloned the first *cry* gene in the early 1980- s [10], many others have been described and are now classified according to Bt Toxin Nomenclature, that consists of four ranks based on amino acid sequence identity [11]. To date, the Bt Toxin Nomenclature Committee [12] has reported at least 78 Cry protein groups, from Cry1 to Cry 78, divided into at least three phylogenetically non-related protein subfamilies that may have different modes of action: the three-domain Cry toxins (3-domain), the mosquitocidal Cry toxins (Etx\_Mtx2), Toxin\_10 proteins, and alpha-helical toxins (reviewed in [13,14]).

The largest group, with more than 53 Cry toxin subgroups, is the 3-domain Cry toxin group. Even though the sequence identity among these proteins is low, the overall structure of the three domains is quite similar, providing proteins with different specificities but with quite similar modes of action [15]. Thus, proteins such as Cry1Aa (lepidopteran specific) and Cry3Aa (coleopteran specific) have a 32.5% identity but a structural similarity as high as 98% [16]. Phylogenetic analysis shows that the great variability in the insecticidal activity of this 3-domain group has resulted from the independent evolution of the three structural domains as well as from the swapping of domain III between different toxins [15].

Due to their feeding habits, many species of coleoptera cause serious damage to both cultivated plants and stored products, leading to significant economic losses in all regions of the world [17,18]. Both larvae and adults have strong jaws, which enable them to feed on a wide variety of plant substrates, such as roots, stems, leaves, grains or wood [19]. Beetles represent the order of the Insecta class that includes the largest number of species. However, the studies carried out to identify toxins of *B. thuringiensis* active against beetles are far from being equal to those carried out in the order Lepidoptera. Thus, 45 Cry proteins, 2 Cyt proteins, 11 Vip proteins, and 2 Sip proteins have been reported with activity against coleopteran insects to date, of which the toxins of the Cry3 and Cry8 families have the largest host spectrum (Figure 1). In this review, we provide an update on the activity of Bt toxins against coleopteran pests.

**Figure 1.** Number of susceptible coleopteran insects to Bt (*Bacillus thuringiensis*) proteins, grouped into protein families.

#### **2. The Crystal Coleopteran-Active Proteins**

*Bt* crystal proteins (δ-endotoxins) are produced during the stationary growth phase and have been isolated from a wide range of insect pests. These crystal inclusions are mainly formed by Cry and Cyt proteins that are toxic to a wide variety of insect species. Most of the information on the insecticidal properties has been obtained for the Cry3 family, and only a few data come from other Cry families. The Cyt proteins constitute a smaller group, mainly active against dipterans, although some Cyt proteins are toxic to coleopteran pests and increase the potential of certain Cry toxins [20].

#### *2.1. Protein Structure*

As mentioned above, Bt Cry proteins can be basically subdivided into three different groups according to their homology and molecular structure: the 3-domain group, Etx\_Mtx2 proteins, Toxin\_10 proteins, and alpha-helical toxins. The 3-domain Cry proteins constitute the largest and best-studied group, although there is increasing information on the 'non-3-domain' and Cyt proteins.

#### 2.1.1. The 3-Domain Group Toxins

All 3-domain Cry proteins are produced as protoxins of two main sizes, a ~130 kDa protoxin and shorter one of approximately 70 kDa [16] (Figure 2). The 130 kDa proteins share a highly conserved C terminus containing 15-17 cysteine residues, which is dispensable for toxicity but necessary for the formation of intermolecular disulphide bonds during crystal formation [15,21]. This group has been mainly studied on lepidopteran toxins such as Cry1A, but also includes some coleopteran active toxins such as Cry7A and Cry8. The structure of the small protoxins is quite similar to the N-terminal half of the large toxin group. Since these do not contain the C-terminal extension, they require, in some cases, the presence of accessory proteins for crystallization [22,23]. This second group includes Cry2A, Cry11A, and some toxins active against Coleoptera, such as Cry3A or Cry3B. Proteolytic cleavage of the N-terminal peptide and the C-terminal extension (mainly in the long Cry protoxins) yields active ~60 kDa protease-resistant fragments [24]. The first crystal structure solved by X-ray crystallography was the coleopteran-specific Cry3Aa [25]. Since then, the tertiary structure of other six 3-domain Cry active proteins, including Cry1Aa, Cry2Aa, Cry3Bb, Cry4Aa, Cry4Aa, Cry4Ba and Cry8Ea, has been determined [26–31]. Among all, Cry3Aa, Cry3Bb and Cry8Ea have been defined as coleopteran-active proteins (Figure 3A,B). Using the FATCAT server [32], the structural alignment between these anti-coleopteran proteins is significantly similar, despite their low sequence identity. Pardo-López et al. [16] analyzed the structural similarity between Cry1Aa and the other 3-domain Cry proteins aforementioned, indicating the same structural likeness. The marked similarity in terms of the structure of the 3-domain Cry proteins, despite the low sequence identity and the differences in specificity, has rendered different proteins with similar modes of action.

**Figure 2.** Relative length of 3-domain Cry proteins of *B. thuringiensis*, representing both main sizes of approximately 130 and 70 kDa. Dashed parts represent the activated toxin, while the white boxes represent the amino- and carboxy-terminal parts. Adapted from Bravo et al., 2007 [33].

Domain I consists of six α-helix surrounding a hydrophobic helix-α5. This domain, which shares strong similarities with the structure of the pore-forming domain of α+PFTs colicin A, might be responsible for membrane penetration and pore formation [23]. The binding domain II is constituted by three antiparallel β-sheets packing together and has an important role in receptor binding affinity. Finally, domain III is a two-twisted anti-parallel β-sheet and is also involved in receptor binding and pore formation [24,34]. Although it has been demonstrated that domains I and II have co-evolved over the years, swapping by homologous recombination of domain III has also been reported [15,35]. Local alignment of coleopteran-active Cry3, Cry7, and Cry8 showed that domain I was strongly conserved while domains II and III diversified [35]. Bt might use this mechanism to get adapted to a new insect host, which may explain the great variability in the biocidal activity of the 3-domain Cry proteins.

#### 2.1.2. Non-3-Domain Cry Toxins

In addition to the 3-domain Cry proteins, some unrelated Cry proteins are also designated by the Cry nomenclature: Etx\_Mtx2 proteins, Toxin\_10 proteins and alpha-helical toxins [4]. The structure and function of Etx\_Mtx2 proteins remains unclear, although the similarities with the *Clostridium perfringens* epsilon toxin (closely related to aerolysin) seem to indicate that they may have a β-sheet-based structure and a pore-forming activity [36]. It is important to notice that, while most of them have activity by themselves, some toxins are proposed as protein complexes to induce mortality, such as the Etx\_Mtx2 protein Cry23 and the Cry37 protein [37]. The crystal structure of Cry23Aa reveals a single β-stranded domain protein, with structural similarity to several β-pore forming toxins as proaerolysins, produced by other bacterial species [38]. Cry37Aa conforms to a C2 β-sandwich fold, similar to the calcium phospholipid-binding domain observed in human cytosolic phospholipase A2 (Figure 3C) [38]. Moreover, the toxins Cry34 and Cry35 have been reported to have binary activity against coleopteran insects [39,40]. Crystal structures of Cry34Ab and Cry35Ab have been published (Figure 3D). Cry35Ab, a member of Toxin\_10 proteins, shows an aerolysin-like fold, containing a β-trefoil N-terminal domain similar to the carbohydrate-binding domain in Mtx1. Cry34Ab is also a member of the aerolysin family with a β-sandwich fold, common among other cytolytic proteins [41].

#### 2.1.3. Cyt Proteins

Similar to the Cry proteins, Cyt proteins are produced as protoxins with a proteolytically activated size of around 25 kDa [20]. As with some Cry proteins, the tertiary structure of some Cyt proteins has already been solved. Cyt1Aa [42], Cyt2Aa [43] and Cyt2Ba [44] show a similar structure composed of a single α−β-domain, with two outer layers of α-helix wrapped around a β-sheet (Figure 3E). Studies performed with peptides of Cyt1A show that α-helix peptides are major structural elements involved in membrane interaction [45] and also in the oligomerization process [46], while the β-strand forms an oligomeric pore with a β-barrel structure into the membrane [43].

#### *2.2. Insecticidal Activity*

The vast majority of Cry proteins described to date are toxic to lepidopteran pests, but there are also a few crystal proteins toxic to either coleopteran or dipteran insects, and a small number are toxic to nematodes [47]. Currently, 45 Bt crystal proteins, including Cry, Cyt or binary proteins, have been tested against different coleopteran insects (Table 1).

#### 2.2.1. Host Range

Cry proteins are toxic to a large number of beetle pests. Mainly, the Cry3 group, the best-studied one, has been described with activity against most of the coleopteran species assayed. These Cry proteins, encoded by *cry3* genes, were first discovered in the subspecies *tenebrionis* [48] and *san diego* [49] although, years later, both strains turned out to be the same subsp. [50]. Since then, more isolates like Bt subsps. *tolworthi, kumamotoensis,* or *kurstaki* have been reported to encode a *cry3* gene [51,52]. Owing to the well-known activity in important coleopteran pests, such as *Leptinotarsa decemlineata* (Coleoptera: Chrysomelidae) or *Diabrotica* spp. (Coleoptera: Chrysomelidae), some of these isolates have been

developed as bioinsecticides for beetle control [47]. Cry3Aa, Cry3Ba, Cry3Bb and Cry3Ca proteins have shown activity against most major coleopteran families, including *Chrysomelidae*, *Curculionidae*, *Scarabaeidae,* and *Tenebrionidae*, among others (Table 1). Although Cry3 proteins are the most effective Bt toxins against chrysomelid beetles, the widespread use of Cry3-based insecticides and Bt crops carries the risk of selecting insect biotypes tolerant to that proteins. The appearance of resistant populations of the chrysomelids *L. decemlineata*, *Chrysomela scripta* under laboratory conditions or *Diabrotica* spp. to Bt maize have been reported [53–55].

**Figure 3.** *Bacillus thuringiensis* proteins, with particular activity against coleopteran pests, for which three-dimensional structure has been predicted. (**A**) Cry3Aa (PBD accession number 4QX1); (**B**) Cry8Ea (PBD accession number 3EB7); (**C**) Protein complex Cry23Aa/Cry37Aa (PBD accession number 4RHZ); (**D**) Binary proteins Cry34Ab and Cry35Ab (PBD accession number 4JOX and 4JPO); (**E**) Cyt1Aa (PBD accession number 3RON); (**F**) Secretable protein Vip2Aa with a NAD complex (PBD accession number 1QS2).

Cry7 and Cry8 groups are comparatively less active on chrysomelids, but they represent a serious alternative to Cry3 proteins. Cry7Aa, formerly known as CryIIIC, is very toxic to Cylas species (Coleoptera: Brentidae) [56], even more than Cry3 protein, but it has no negative effects against *Anthonomus grandis* (Coleoptera: Curculionidae) or *D. undecimpuntata* [52]. Moreover, toxicity to Colorado potato beetle has been reported, but only after in vitro solubilization [52], which was countered by a recent report of a Cry7Aa-type protoxin which is active against *L. decemlineata* without any previous solubilization step [57]. Solubilized Cry7Ab is active against *Henosepilachna vigintiomaculata* (Coleoptera: Coccinellidae) and *Acanthoscelides obtectus* (Coleoptera: Chrysomelidae), but not against *Anomala corpulenta* (Coleoptera: Scarabaeidae) or *Pyrrhalta aenescens* (Coleoptera: Chrysomelidae) [57,58]. Cry8-type proteins are toxic to a large number of coleopteran pests, particularly against species in the Scarabaeidae family [59–61]. Furthermore, Cry8A and Cry8B proteins have shown activity against the chrysomelids *L. decemlineata* and *Diabrotica* spp., Cry8Ca against the tenebrionid *Alphitobius diaperinus* (Coleoptera: Tenebrionidae) [62] and Cry8Ka against the curculionid *A. grandis* [63]. Moreover, some Cry8 proteins, such as Cry8Ea, Cry8Ga or Cry8Na, are very specific, showing different activities against very closely related host species [64,65]. Cry6Aa and Cry6Ba are active against the curculionid beetles *Hypera postica* and *Hypera brunipennis*, two of the more important pests in alfalfa [66,67], as well as *D. virgifera*, which is susceptible to the activated toxin. Cry22 proteins also have activity to a wide spectrum of coleopteran insects. In particular, Cry22A

and Cry22B proteins are toxic to coleopterans of the Brentidae, Chrysomelidae and Curculionidae families [56,68,69].

Generally, Bt protein groups are particularly toxic to a certain insect order. However, some proteins may be active against different orders [70]. Mainly lepidopteran proteins Cry1Ba and Cry1Ia have shown activity against the key coleopteran pests *A. grandis*, *A. obctetus*, *C. scripta* and *L. decemlineata* [71–76]. Dual activity against Lepidoptera-Coleoptera has also been demonstrated by Cry9-type proteins. Cry9 toxins exhibit strong activity against main lepidopteran pests, but Cry9Da is also toxic against the scarab *Anomala cuprea* [77]. Other example of cross-order toxicity is depicted by the dipteran toxin Cry10Aa, which can kill the Cotton boll weevil (*A. grandis*) [78]. Additionally, Cry51Aa is toxic against *Lygus* spp. (Hemiptera) and *L. decemlineata* [79] and Cry55Aa, a typical nematicidal protein, has been reported as toxic to the chrysomelid *Phyllotreta cruciferae* [80].

Binary toxins, structurally different from classical 3-domain Cry proteins [25], used to be considered as single toxins because both proteins are required to kill their target. To date, two binary complex toxins have been proposed to have activity against beetles. The coleopteran specific Cry23Aa has been assayed together with Cry37Aa protein to kill *Popillia japonica* (Coleoptera: Sacarabaeidae) and *Tribolium castaneum* (Coleoptera: Tenebrionidae) [37]. Furthermore, this protein mixture has been found to be active against *Cylas* spp. (Coleoptera: Brentidae) and *A. obtectus* [56,75]. On the other hand, Cry34 protein is only active in association with Cry35 protein [17]. Cry34 and Cry35 are closely related and are often encoded in the same operon, with coordinated function and appearance in crystals [40,81]. The Cry34/Cry35 binary proteins are mainly active against corn rootworms and have been developed for in-plant control in Bt maize [40,82].

*B. thuringiensis* Cyt proteins have an in vitro cytolytic (hemolytic) activity, hence their name, and show predominant dipteran specificity [24]. However, some of them are also toxic to coleopteran pests, such as Cyt1Aa to *C. scripta* [72] or Cyt2Ca to the chrysomelids *L. decemlineata* and *Diabrotica* spp. [83] and the curculionid *Diaprepes abbreviates* (Coleoptera: Curculionidae) [84,85]. Besides, Cyt proteins improve the activity of Cry proteins. For instance, Cyt1Aa is able to overcome high levels of resistance to Cry3Aa by *C. Scripta*, playing an important role in resistance management [72].




**Table 1.** *Cont.*


**Table 1.** *Cont.*


**Table 1.** *Cont.*


**Table 1.** *Cont.*



(a) The parameter is mortality. A = active; N = not active; LA = low activity, with significant inhibition of growth; (b) LC50 = lethal concentration that causes 50% mortality of the insects. Data are expressed in μg/mL, unless otherwise stated. "//" separate two different values of the LC50.

#### 2.2.2. Genetically Engineered Cry Genes

Recent advances in next generation sequencing and genetic engineering technologies allow the construction of new synthetic *cry* genes that increase or amplify their toxicity. The domain regions of some lepidopteran-specific proteins have been modified in an attempt to improve their specific activity or broaden their host range [15,153]. The first coleopteran hybrid protein was made by fusing the sequences located in domain III of the *cry3A* and *cry1Aa* genes, although unfortunately, it caused the loss of activity against *L. decemlineata* [154]. Nonetheless, substituting domain III of Cry3Aa with the same domain from Cry1Ab induced activity against WCR (Western corn rootworm) larvae [155]. On a different approach, a *cry3Bb1* gene was engineered with five amino acid substitutions to produce the new Cry3Bb1.11098 protein, which increased the activity of the natural protein against WCR [156]. Similarly, a Cry3A variant (eCry3.1Ab) was designed to confer novel activity against rootworms by creating a cathepsin G protease recognition site [157]. This technology has been introduced successfully in the development of transgenic plants, mainly to overcome the appearance of resistance by WCR populations [158].

#### *2.3. Mode of Action*

The mode of action has been mostly studied in lepidopteran insects, although it is believed to be similar between different insect orders, with some peculiarities [8]. Briefly, it is widely accepted that the process begins once the target insect ingests the protein and reaches the insect midgut, where it is solubilized and proteolytically activated. Such an activation allows toxins to first bind to their specific receptors in the host cell membrane, then to their oligomerization and, eventually, to the formation of pores in the cell membrane (Figure 4). In this multi-step mode of action, several factors may contribute to protein specificity [159].

#### 2.3.1. Solubilization and Proteolytic Processing

Once proteins reach the host midgut, they are released from their crystal package to initiate the pathogenic process. The crystals are stabilized by disulfide bridges among the C-terminal ends of the protoxins. More recently, the occurrence of 20 kbp DNA fragments with protoxins and 100–300 pb DNA fragments with in vitro proteolytic activated toxins has been established [160]. These DNA fragments have been observed to be associated with different Bt-toxins as Cry1A, Cry2A, etc., however, they have been more extensively studied on Cry8 toxins [21]. The sequence of the DNA fragments is not specific and they are located in plasmids and chromosomes [161]. Bioinformatics modelling suggests that two protoxin regions bind to major grooves and another one, combined with phosphoric acid, binds to the minor groove [162]. The associated DNA should be eliminated by the DNAses in the insect gut for the correct protein activation. In fact, DNA-protein association impairs the specific binding [163].

**Figure 4.** Schematic representation of the particularities in the mechanism of action of crystal proteins against coleopteran pests. (**1**) Crystal solubilizes in the acidic conditions of the coleopteran midgut lumen and (**2**) activates into toxin by proteolytic processing of the protoxin by the specific digestive enzymes, specially cysteine and aspartic proteases. (**3**) Toxins are able to bind to a first receptor (CADR), (**4**) oligomerizate and (**5**) form an oligomeric pre-pore structure that (**6**) is able to bind to a second specific receptor (ADAM metalloproteases/GPI-anchored alkaline phosphatases/sodium solute symporters). This event induces the insertion into the membrane, leading to (**7**) pore formation and finally to cell lysis.

It is well accepted that solubilization processes are due to the environmental conditions in the susceptible insect midgut, mainly to pH values. Of note, unlike the alkaline midgut of lepidopteran and dipteran insects, beetles have an acidic midgut, suggesting that different solubilization conditions are needed for each protein [164]. For instance, the midgut fluids of *L. decemlineata* and *D. virgifera* larvae do not seem to solubilize Cry1B and Cry7Aa1, and only after a previous in vitro solubilization, these proteins become active [52,71]. However, recent reports show that Cry7Ab2 and Cry7Aa2 proteins solubilize into midgut fluids of *H. vigintioctomaculata* and *L. decemlineata* larvae, respectively, suggesting that the lack of solubilization involves more factors than pH [57,58]. Cyt proteins dissolve readily under alkaline conditions, especially at pH 8 or higher, and they are harder to solubilize in neutral or slightly acidic pHs, which occurs in coleopteran midguts [72]. Another example of the importance of crystal solubilization was published by Galitsky et al. [28]. They related that differences in toxin solubility, oligomerization and binding for the Cry3-type toxins, in addition to differences in domain III, might explain the different specificities of Cry3A and Cry3B (e.g., WCR is susceptible to Cry3Bb1 but not to Cry3A). Solubilized proteins are proteolytically activated by gut proteases, which generate the toxic three-domain fragment of about 65 kDa [33]. In Lepidoptera and Diptera species, the main proteases present in the alkaline midgut juices are serine proteases, especially trypsin and chymotrypsin proteases [165]. However, the coleopteran species use digestive proteases belonging to cysteine and aspartic proteases and serine proteases are only present in some cases [166]. The presence of different proteases may be an important factor in toxin activation specificity, and improper processing of Bt toxins can involve the development of insect resistances. It has been reported that the combination of Cry3Aa protein and certain protease inhibitors enhances the toxicity against *Rhyzopertha dominica* (Coleoptera: Bostrichidae) larvae, evidencing that protease inhibitors may play an important role in resistant pests management [110]. Moreover, the relevance of a nicking in the N-terminal end, in the alpha 1–3 of Domain I in the activated Cry3A and Cry8Da toxins, has been shown, which rendered an 8 kDa fragment to obtain a functional 54 kDa toxin for receptor binding [167].

#### 2.3.2. Binding to the Larval Epithelium

The activated toxin is able to bind to specific receptors located in the midgut epithelial cells to form an oligomeric pre-pore structure and alterations in the midgut receptors is a critical step for insect resistance appearance [159]. It has been demonstrated that Cry3Ba protein shares a binding receptor with Cry3Aa and Cry3Ca proteins, although heterologous-competition experiments show that both proteins may have other binding sites and only share one with Cry3Ba3 [168]. It has also been shown that Cry3Bb, Cry3Ca and Cry7Aa proteins competed for the same binding sites in *C. puncticollis*, so a mutation in the midgut receptor could render all three proteins ineffective [169]. To date, several specific coleopteran binding proteins have been identified. It has been shown that an ADAM metalloprotease can be considered as a Cry3Aa receptor in *L. decemlineata*, and this binding interaction improves Cry3Aa pore-formation [170]. GPI-anchored alkaline phosphatases (ALP) are also important for the Cry3Aa binding to *Tenebrio molitor* brush border membrane vesicles (BBMV) and are highly expressed when larvae are exposed to Cry3Aa [171]. In the same way, the Cry1Ba toxin binds to ALPs from *A. grandis* midgut cells [74]. Although some putative cadherines have been previously described [172,173], Fabrick et al. [127] were the first authors reporting a cadherin protein (TmCad1), cloned from *T. molitor* larval midgut as a Cry3Aa binding receptor. Furthermore, injection of *TmCad1* dsRNA into *T. molitor* larvae conferred resistance to Cry3Aa. Another truncated cadherin protein (DvCad1-CR8–10), isolated from the WCR, binds to activated Cry3Aa, Cry3Bb [118] and also Cry8Ca [62], enhancing the activity of *L. decemlineata*, *Diabrotica* spp. and *A. diaperinus*. Finally, in *T. castaeneum* larvae, a cadherine (TcCad1) and a sodium solute symporter (TcSSS) have been identified as putative Cry3Ba functional receptors, determinant for the specific Cry protein toxicity against coleopterans [174].

Studied Cry8-binding proteins revealed a difference from those confirmed previously as receptors for Cry1A or Cry3A proteins in lepidopteran and coleopteran insect species, such as aminopeptidases, cadherins or ABCC transporters [175,176]. A Cry8-like toxin without the C-terminal end has been described, which completely shared binding sites with Cry8Ga, despite only sharing 30% of the sequence, in *Holotrichia oblita*. Cry8Da tested on *Popillia japonica* BBMV, bound specifically with a 150 kDa membrane protein which shared homology with coleopteran β-glucosidases [177]. Cry8E and Cry8-like toxins showed, in *H. parallela* and *H. oblita,* binding to several different proteins. The most relevant for both insect species and Cry8 proteins were serine proteases, sodium/potassium-transporting proteins, and a transferrin-like protein [177,178].

There is evidence that some proteins work together to cause mortality in certain coleopteran species, although the mechanism of interaction between them remains unclear. In this way, it is hypothesized that Cry37 protein may facilitate linkage of channel-forming Cry23 toxin, given their homology to other binding proteins [24]. Moreover, the fact that Cry34Ab has some activity against the Western corn rootworm (WCR) on its own [150] seems to indicate that Cry35 has the role as a receptor of Cry34, which is mainly responsible for toxicity. Cyt proteins enhancing the insecticide potency of certain Cry toxins has been also observed. The Cyt1Aa protein, from Bt sub. *israelensis,* increases the activity of Cry11Aa toxin by acting as a membrane receptor [178]. Cyt1A also helps to overcome high levels of Cry3A resistance against *C. scripta* larvae [72]. Although this mechanism of action has not yet been elucidated, Cyt1A may act as a receptor of Cry3A to enhance the binding of this protein. This synergism between Cry and Cyt toxins is an excellent strategy to decrease the appearance of resistance to Cry proteins.

#### 2.3.3. Oligomerization and Pore Formation

Although it remains unclear, some studies suggest that activated toxins need to form an oligomeric structure before insertion to the membrane as a result of binding to specific receptors [16]. In fact, Cry proteins that form oligomeric structures are related to a high pore activity [33]. Oligomerization of 3-domain Cry proteins has been described for toxins active against different insect orders, such as Cry3 proteins in coleopteran larvae. In the brush border membrane of *L. decemlineata*, Cry3A, Cry3B

and Cry3C form an oligomer prior to membrane insertion, generating a pre-pore structure that can be inserted into the membrane [168]. Cry3Aa oligomeric structures have also been reported after incubation of Cry3Aa protoxin with *T. molitor* BBMV [127]. The oligomeric structure eventually leads to the lytic pore formation that disrupts the midgut insect cell by osmotic shock. However, oligomerization studies of Cry1Ab and Cry1Ia proteins incubated with lepidopteran and coleopteran BBMV, as well as culture insect cells, showed that Cry1Ia oligomerization may not be a requirement for toxicity [179]. Besides, the appearance of Cry1Ab oligomers when incubated with coleopteran BBMV could be due to an improper insertion of oligomers into the membrane or the inability to induce the post-pore events in the cells [179]. Either way, susceptible insects can withstand minor damage, but greater damage destroys the epithelium of the midgut, leading to a disruption in feeding and subsequent starvation death. Additional to the toxin action, spores may pass through the channel, to colonize and germinate in the hemolymph and contribute to insect death by septicemia [1].

#### **3. The Secretable Coleopteran-Active Proteins**

In addition to the δ-endotoxins produced during the stationary phase, other protein compounds have been found in the culture supernatant of certain entomopathogenic *Bacillus*isolates. These proteins, produced during the vegetative growth stage of the bacterium, were designated as vegetative insecticidal proteins (Vip) [180] and secreted insecticidal proteins (Sip) [181]. Within the Vip family, *vip1* and *vip2* genes are co-transcript in a single 4 kbp operon, which render proteins of about 100 kDa (Vip1) and 50 kDa (Vip2) [171]. The absence of toxicity of the proteins alone suggests that it is a binary toxin for some members of the coleopteran [180] and hemipteran [182] orders. In contrast, Vip3 proteins are single-chain toxins with insecticidal activity against a wide range of lepidopteran species [183]. While *B. thuringiensis* is a good source of Vip proteins, these proteins have also been found in other closely related bacteria, such as *Bacillus cereus*, *Lysinibacillus sphaericus,* or *Brevibacillus leterosporus*. Currently, two Sip proteins have been described, both active against several coleopteran pests. The fact that strains harboring *sip1Aa* and *sip1Ab* genes also contain *cry3* and *cry8* genes, respectively, suggests that Sip1 proteins may have a role in the insecticidal mechanism against coleopteran insects [184].

#### *3.1. Protein Structure*

Vip1 and Vip2 proteins are found in the culture supernatant before cell lysis due to specific secretion [181,185]. Both proteins have an N-terminal signal peptide for secretion, commonly cleaved after the secretion process is completed [24,181]. The Vip1/Vip2 homology with other bacterial binary toxins and the fact that these proteins are codified by two genes encoded in a single operon, suggest the presence of a typical "A+B" binary toxin [24,185]. It has been proposed that Vip1, with moderate sequence identity (30%) and structural similarity with the binding C2-II *Clostridium botulinum* toxin and the toxin "B" of *Clostridium di*ffi*cile*, is the binding domain that translocates Vip2, with homology to the Rho-ADP-ribosylatin exotoxin C3 of *Clostridium* spp, to the host cell [186,187]. As occurs with other related "B" compounds, Vip1 is formed by four domains involved in docking to enzymatic components, binding to specific cell surface receptors, oligomerization, and channel formation in lipid membranes [188]. Coleopteran active Sip1Aa protein contains a predicted Gram-positive consensus secretion signal [4] and exhibits 46% similarity with Mtx3 mosquitocidal toxin of *Lysinibacillus sphaericus* [184]. This homology may indicate that Sip1Aa toxicity should be caused by pore formation.

#### *3.2. Insecticidal Activity*

The activity of the Bt secretable toxins against coleopterans is depicted in Table 2. Currently, four Vip protein families have been identified, but only Vip1/Vip2 showed activity against coleopteran pests [189]. Vip1/Vip2 proteins have been tested against different coleopteran families but they have shown active only against the Chrysomelidae, Curculionidae, and Scarabeidae families, being particularly toxic to corn rootworms. Single Vip1 or Vip2 showed no mortality, confirming that

these proteins must act together to be toxic [185]. Vip1Aa was highly toxic against *Diabrotica* spp. when combined with Vip2Aa or Vip2Ab, but Vip1Ab/Vip2Ab (co-expressed in the same operon) and Vip1Ab/Vip2Aa were not active [185]. These data show the specificity of these proteins and suggest that the absence of toxicity is due to Vip1Ab. Moreover, Vip1Ba/Vip2Ba and Vip1Bb/Vip1Ba were toxic against *Diabrotica virgifera virgifera* [190] and binary Vip1Da/Vip1Ad had activity against the curculionid *A. grandis* and the chrysomelids *Diabrotica* spp and *L. decemlineata* [191]. These are the only Vip proteins active against the Colorado potato beetle. Vip1Ad/Vip2Ag binary proteins were the first report of demonstrated toxicity against any Scarabaeoidea larvae, being active against *Holotrichia parallela*, *H. oblita* and *Anomala corpulenta* [192]. Sip1Aa and Sip1Ab proteins have specific activity against coleopteran pests. Sip1Aa caused lethal toxicity for *L. decemlineata* larvae and stunting in *D. virgifera* and *D. undecimpunctata* larvae [181]. Sip1Ab was also toxic to *Colaphellus bowringi* Baly (Coleoptera: Chrysomelidae) but it did not harm *Hloltrichia diomphalia* (Coleoptera: Scarabaeidae) larvae [184], suggesting specific chrysomelid activity, although further studies are needed to determine its host range.

#### *3.3. Mode of Action*

The mode of action of coleopteran-specific Bt secretable proteins is poorly understood, but some information is available for this binary mechanism of action. The proposed multistep process begins with the ingestion of the two toxins by the susceptible larvae. Though the two encoded proteins are synthesized together, they are thought not to get associated in solution and reach the insect midgut as single proteins [188]. Then, the proteolytic processing by the trypsin-like proteases of the insect midgut juice of Vip1 allows the cell-bound "B" to bind to a specific membrane receptor, followed by the formation of oligomers containing seven Vip1 molecules [193]. It is at this stage when the docking between Vip1 and Vip2 translocates the toxic component (Vip2) into the cytoplasm though the "B" (Vip1) channel [188]. Recent studies in BBMVs of *H. parallela* evidenced that although Vip2Ag showed a low degree of binding on its own, the degree of binding increased when Vip1Ad was added, showing that Vip1Ad acted as a receptor to help Vip2 bind to BBMVs [194]. Once inside the cytosol, Vip2 destroys filamentous actin by blocking its polymerization and leading to cell death [195].

Sip1 proteins have no homology with Vip proteins, but Sip1A exhibits limited sequence similarity with the 36-kDa mosquitocidal Mtx3 protein of *B. sphaericus*, suggesting that toxicity is related with pore formation [181].


**Table 2.** Insecticidal activity of Vip and Sip proteins against coleopteran pests.


**Table 2.** *Cont.*

(a) The parameter is mortality. A = active; N = not active; (b) LC50 = lethal concentration that causes 50% mortality of the insects. data are expressed in μg/mL, unless otherwise stated. "//" separate two different values of the LC50.

#### **4. Bt Based Insecticides**

In 1938, the first insecticide based on *B. thuringiensis* was produced and marketed under the name *Sporéine*for the control of lepidopteran insect pests [47]. Since then, sporulated cultures of *B. thuringiensis* have been used widely as foliar sprays to protect crops from insect damage. Since *B. thuringiensis* subsp. *tenebrionis* was discovered [48], it was rapidly formulated as a bioinsecticide and commercialized against the Colorado potato beetle. Bt-based insecticides to control coleopteran pests are mainly developed against chrysomelid beetles [198]. Novodor® (Kenogard) uses the NB-176 strain of Bt subsp. *tenebrionis* as the active ingredient and is widely used for the control of *L. decemlineata*. However, the toxicity of this commercial product has been verified for other species of beetles, such as the chrysomelids *Chrysophtharta bimaculata, C. agricola* and *C. scripta* [199,200] under laboratory conditions. Furthermore, this product has been shown to be effective against *C. scripta* in field conditions [200], while the use of Novodor did not exert good control of the populations of *Lissorhoptrus oryzophilus* (Coleoptera: Curculionidae) [201].

To date, most of the Bt-based bioinsecticide products effectively use natural Bt strains for the control of foliar-feeding pests. However, several factors have limited their use. Usually, Bt strains have a narrow insecticidal spectrum compared with other insecticides, even when insects are closely related [202]. Advances in genetic manipulation technologies offer improvements in the efficiency of Bt-based formulates and reductions in their production costs. The development of new strains by conjugation or transduction has been used to confer natural strains with new insecticidal properties [203]. The natural Bt subsp. *kurstaki*, for example, has been modified to express several *cry3* genes and extend its host range to both lepidopteran and coleopteran pests [202]. The active ingredient in Foil® is the Bt strain EG2424,

expressing both Cry1Ac and Cry3A proteins, the latter of which was transferred from a Cry3Aa-encoding plasmid belonging to the Bt subsp. *morrisoni* [204]. Similarly, the Cry3-overproducing strain, EG7673, was obtained by transforming a natural strain with a recombinant plasmid containing a *cry3Bb1* gene. A formulation with this strain as the active ingredient was commercialized as Raven® and was four-fold more active than the parental strain [205].

#### **5. Bt-Crops**

By expressing one or more Bt toxic genes in a target plant tissue transgenic insect-resistant crops, Bt crops, can be produced. Such cultivars need no further pest control measures. To date, the Bt crops extension has increased worldwide, particularly that of Bt cotton, Bt rice and Bt corn [9]. Bt plants have been created for the control of several insect pests, among others, Colorado potato beetle (*L. decemlineata*) and corn rootworms (*Diabrotica* spp.). The first human-modified pesticide-producing crop was potato, which expressed the *cry3A* gene from *B. thuringiensis* subsp. *tenebrionis* in their leaves [206]. The transgenic gene expression confers potato plants protection against the Colorado potato beetle and allows reducing insecticide applications [207]. A few years later, this Bt crop was complemented with another gene expression cassette that also provided protection against the Potato leafroll virus [208]. However, genetically modified potatoes were commercialized from 1995 to 2001, and eventually removed from the marketplace due to social concern for genetically modified crops [209].

A coleopteran-active Bt maize was designed for the control of corn rootworms, expressing a variant of the wild-type *cry3Bb1* gene from Bt subsp. *kumamotoensis* in the root tissue [210]. Currently, Bt maize hybrids express four different crystal proteins (Cry3Bb, mCry3A, Cry34Ab/35Ab and eCry3.1Ab), individually or co-expressing two toxins [211,212]. Vip1 and Vip2 proteins were also candidates to be expressed in maize plants, mainly due to the great toxicity against rootworms. However, the cytotoxic activity of the Vip2 protein has prevented the development of a Bt plant expressing this binary toxins [189]. The opportunity of expressing the toxin in a specific tissue allows minimization of the exposure of non-target fauna while increasing the control of tunneling and root pests, which are otherwise difficult to manage. However, Western corn rootworm has developed field resistance to all four currently available Bt toxins [212–214] as did *D. virgifera* in 2009 against Bt corn [55]. These facts show that although Bt crops have the potential to increase productivity while conserving biodiversity, resistance management programs and a better use of integrated pest management are necessary to delay resistance development as much as possible [215].

#### **6. Resistance and Cross-Resistance**

The widespread use of *B. thuringiensis* biopesticides, as well as the planting of millions of hectares of Bt plants to protect crops from pests, carry the risk of selecting insect biotypes that are tolerant or resistance to Bt toxins. The appearance of resistance may be due to alterations in any step involved their mode of action, from the solubilization and activation steps to the capacity of pore formation [159]. It is established that the lack of solubilization is favored by the physicochemical conditions of the midgut fluids, particularly the pH. The acidic midgut of the coleopteran insects seemed to be a limiting factor in the solubilization of Cry proteins, such as Cry1B and Cry7Aa [52,71], although recent reports seem to indicate that more factors are involved as Cry7Aa proteins are dissolved in *L. decemlineata* and *H. vigintioctomaculata* midgut fluids [57,58]. Once the Cry toxin is solubilized in the midgut, protoxins are proteolytically cleaved to activated toxins. This toxin processing depends on the presence of the right digestive enzymes in the host midgut fluid. As an example, it was observed in *D. virgifera* larvae that the Cry3Aa protein was poorly processed by its own proteases, which leads to low activity of Cry3Aa against rootworms [157]. Introduction of a chymotrypsin/cathepsin recognition site in domain I of Cry3A has been shown to enhance the bioactivity of this toxin against the western corn rootworm larvae [157].

Molecularly, the insect resistance basis is a modification or loss of the specific midgut cell membrane receptors or some mediator, which eliminates or reduces the capacity of the toxin to initiate a lethal pathway [216]. Cross-resistance between Cry toxins is often associated with sequence similarities in domains II and III, related to specific protein binding [217]. Under laboratory conditions, populations of *L. decemlineata* and *C. scripta* resistant to Cry3Aa have been described [53,54]. To date, the appearance of field resistance is still relatively low despite the extensive use of products based on the same protein, which increases the probability of resistance development.

Conversely, rootworm populations have developed resistance to all proteins used in transgenic corn. The intense selection pressure posed by the continuous exposure of insects to Bt toxins has increased the emergence of pest resistance. Since the first case of resistance to Cry3Bb1 Bt-maize in 2009, *Diabrotica* has developed resistance to Cry3Aa and Cry34/35Ab binary protein [211]. New strategies are being carried out to try to delay resistance, including a combined use of several proteins in the same Bt plant [218]. Pyramiding of two Bt proteins can delay resistance to those proteins because when insects become tolerant to one toxin, most will still be susceptible to the other toxin [211]. However, there is already evidence of cross-resistance to Cry3 proteins and even to Cry34/35, which may invalidate, in the long run, the use of all these proteins [212].

**Author Contributions:** Conceptualization, M.D.-A., C.A.-A. and P.C.; resources, C.A.-A., B.E., and P.C.; writing—original draft, M.D.-A.; writing and editing, M.D.-A., M.V., B.E., and P.C.; review and supervision, M.V. and P.C. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research was funded by the Spanish Ministry of Science and Innovation (RTI2018-095204-B-C22).

**Acknowledgments:** M.D.-A. received a doctoral grant from Universidad Pública de Navarra, Pamplona, Spain.

**Conflicts of Interest:** The authors declare no conflict of interest. The funders had no role in the design of the study; in the collection, analyses, or interpretation of data; in the writing of the manuscript, or in the decision to publish the results.

#### **References**


© 2020 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).

## **The Cadherin Protein Is Not Involved in Susceptibility to** *Bacillus thuringiensis* **Cry1Ab or Cry1Fa Toxins in** *Spodoptera frugiperda*

**Jianfeng Zhang 1,**†**, Minghui Jin 2,**†**, Yanchao Yang 1, Leilei Liu 1, Yongbo Yang 1, Isabel Gómez 3, Alejandra Bravo 3, Mario Soberón 3, Yutao Xiao <sup>2</sup> and Kaiyu Liu 1,\***


Received: 4 May 2020; Accepted: 3 June 2020; Published: 6 June 2020

**Abstract:** It is well known that insect larval midgut cadherin protein serves as a receptor of *Bacillus thuringiensis* (Bt) crystal Cry1Ac or Cry1Ab toxins, since structural mutations and downregulation of *cad* gene expression are linked with resistance to Cry1Ac toxin in several lepidopteran insects. However, the role of *Spodoptera frugiperda* cadherin protein (SfCad) in the mode of action of Bt toxins remains elusive. Here, we investigated whether SfCad is involved in susceptibility to Cry1Ab or Cry1Fa toxins. *In vivo*, knockout of the *SfCad* gene by CRISPR/Cas 9 did not increase tolerance to either of these toxins in *S. frugiperda* larvae. *In vitro* cytotoxicity assays demonstrated that cultured insect TnHi5 cells expressing GFP-tagged SfCad did not increase susceptibility to activated Cry1Ab or Cry1Fa toxins. In contrast, expression of another well recognized Cry1A receptor in this cell line, the ABCC2 transporter, increased the toxicity of both Cry1Ab and Cry1Fa toxins, suggesting that SfABCC2 functions as a receptor of these toxins. Finally, we showed that the toxin-binding region of SfCad did not bind to activated Cry1Ab, Cry1Ac, nor Cry1Fa. All these results support that SfCad is not involved in the mode of action of Cry1Ab or Cry1Fa toxins in *S. frugiperda*.

**Keywords:** *Bacillus thuringiensis*; *Spodoptera frugiperda*; cadherin; Cry1Ab; Cry1Fa; mode of action of Cry toxin

**Key Contribution:** The CRISPR/Cas 9 gene editing, cytotoxicity assessment and biochemical analysis demonstrate the mutations of *cadherin* gene does not result in the development of resistance to Cry1Ab or Cry1Fa toxins in *S. frugiperda*.

#### **1. Introduction**

The crystal (Cry) toxins and vegetative insecticidal proteins (Vip) produced by *Bacillus thuringiensis* (Bt) bacteria are important biological tools for the control of insect pests and provide good protection for plants growth [1]. During sporulation, Bt bacteria accumulate Cry toxins in crystal inclusion bodies inside the mother cell, while the Vip proteins are secreted in the vegetative phase of growth [2,3]. The Bt toxin receptors, located on the larval midgut cells, play important roles in the toxicity of these Bt toxins. After ingestion of Bt crystal inclusions or Vip protein by the larvae, these proteins are dissolved under

the alkaline conditions of the gut lumen, releasing protoxins that are activated by midgut proteases. The activated toxins bind to receptors, forming oligomers that insert into the cell membrane leading to pore formation, which results in death of the larvae [2,3]. The mode of action of Vip3Aa might be different from crystal toxins, since receptors for Vip3Aa are not shared with the Cry toxins [4–7].

In several lepidopteran insects, mutations in the *cadherin* gene (*cad*) are associated with resistance to Cry1Ac or Cry1Ab toxins [8–12]. The Cry1Ac toxin-binding region of *Helicoverpa armigera* cadherin (HaCad) and the membrane-proximal region of HaCad are required for Cry1Ac toxicity [13,14]. The downregulated expression of the *cadherin* gene has also been associated with resistance against the Bt Cry1Ac toxin in *Pectinophora gossypiella* [15]. Besides cadherin, the ATP-binding cassette sub-family C member 2 (ABCC2) is also recognized as an important insect molecule involved in the mode of action of Cry1A toxins [16]. Furthermore, it is known that HaCad and *Heliothis virescens* cadherin (HvCad) have a synergistic effect with ABCC2 on toxicity of Cry1A in cultured insect cells, since co-expression of cadherin receptors or the toxin-binding region of HaCad with the ABCC2 protein induced a synergistic effect on the cytotoxicity of Cry1Ac [14,16].

Even though cadherin has been shown to be an important Cry1A receptor in different Lepidopteran species, this is not always the case for some other lepidopteran insects. For instance, it has been reported that the cadherin from *Plutella xylostella* (PxCad) is not associated with resistance in *P. xylostella* to Cry1Ac [17]. However, other reports suggest that PxCad is a functional receptor of Cry1Ac, since PxCad can increase cytotoxicity of Cry1Ac when expressed in the Sf9 cell line [18–20]. In addition, we reported that *Spodoptera litura* cadherin (SlCad), in contrast to HaCad, cannot increase cytotoxicity of Cry1Ac when expressed in Hi5 cells, suggesting that SlCad is not a functional receptor of Cry1Ac in *S. litura* [14].

Although *S. frugiperda* is susceptible to Cry1Ab, Cry2Ab, Cry1Fa, and Vip3Aa toxins [21–30], there are no reports regarding whether *S. frugiperda* cadherin (SfCad) is involved in the mode of action of these Bt toxins. It has been shown that resistance to Cry1Fa in *S. frugiperda* is linked to different ABCC2 mutant alleles [27,28,31]. In addition, most *S. frugiperda* populations show low susceptibility to Cry1Ab or Cry1Ac toxins, in contrast to Cry1Fa that is highly active against this pest [21,32]. Here, we investigated whether SfCad is involved in the toxicity of Cry1Ab and Cry1Fa using both CRISPR/Cas 9 genome editing technology and cytotoxicity assays of Bt toxins in an insect cell line expressing SfCad. Our results suggest that *S. frugiperda* cadherin is not involved in the mode of action of Cry1Ab or Cry1Fa toxins.

#### **2. Results**

#### *2.1. Construction of SfCad Gene Deleted Mutant by CRISPR*/*Cas 9 Genome Editing*

To construct an *S. frugiperda cadherin* gene knockout mutant strain, we made use of the CRISPR/Cas 9 system to produce a large fragment deletion by designing two sgRNAs targeting different exons of the *SfCad* gene (Figure 1A). Freshly laid eggs were co-injected with the two *in vitro* transcribed sgRNAs, that are complementary to 20 bp DNA sequences from the fourth or fifth exons of *SfCad*, respectively, along with Cas 9 protein (Figure 1A). The results show that 22.5% (45/200) of the injected eggs hatched, and 71.1% (32) of the 45 neonates, raised in diet, survived into adults (F0). The F0 male and female moths were mass backcrossed with the DH19 strain to produce the next generation in single pair matings (F1).

**Figure 1.** The knock-out of the *SfCad* gene. (**A**). Deleted fragment of the *SfCad* gene by the CRISPR/Cas 9 system between the two red arrow heads. (**B**). Sequencing and agarose gel electrophoresis of DNA confirming the knock-out of the *SfCad* gene.

After enough eggs were collected, genomic DNA from individual F1 moths was prepared. Deletion events were detected by PCR using primers across the two target site regions (Figure 1A). Fragment deletions were initially screened by agarose gel electrophoresis and then those samples that showed multiple bands were cloned using T vector, and the DNA was sequenced to identify their mutations. We found that 25% (8/32) of the examined individuals showed deletions in the *cad* gene.

From the detected mutations, we selected a 382-bp deletion to generate a homozygous knockout strain (Figure 1B). The F1 larvae (progeny crosses of the 382-bp deletion F0 moth and strain DH19) were reared to pupation, and 96 exuviates of the final instar larvae were used to prepare genomic DNA. The DNA fragments flanking the two target sites were amplified by PCR, which were 515 bp in the wild type and 133 bp in the mutant (Figure 1B).

Among the 96 pupae screened, 30 carried the 382-bp deletion allele. Adults from these pupae were mass-crossed to obtain the F2 generation. The genotypes of more than 100 F2 individuals were visualized by agarose gel electrophoresis using gDNA samples from randomly selected exuviates of final-instar larvae. The agarose gel electrophoresis results showed that 21.4% (30/140) were homozygous for the 382-bp deletion. The 30 individuals were further sequenced and verified to be homozygotes. Finally, these homozygous individuals were pooled and mass-crossed to establish the *SfCad* knockout strain (Cad-KO).

#### *2.2. Susceptibility of Cad-KO Strain to the Bt Toxins*

To determine whether *SfCad* is involved in Cry1Ab or Cry1Fa resistance in *S. frugiperda*, we performed bioassays using the Cad-KO (knockout strain) and the progenitor DH19 *S. frugiperda* strains. Bioassay results showed that the knockout strain Cad-KO did not decrease susceptibility to these two Bt toxins (Figure 2). The LC50 values of Cry1Ab and Cry1Fa to the Cad-KO strain were not significantly different from the control DH19 strain because their 95% fiducial limit (FL) values overlapped (Table 1), suggesting that SfCad is not a functional receptor of Cry1Ab and Cry1Fa. The bioassay data also showed that Cry1Ab was at least 20- to 40-fold less toxic to both *S. frugiperda* strains analyzed compared to Cry1Fa toxin.

**Figure 2.** Influences of the knockout of the *SfCad* gene on susceptibility of the first instar *S. frugiperda* larvae to Cry1Ab, Vip3Aa, and Cry1Fa, respectively. Cad-KO (knockout strain), *S. frugiperda* larvae with the knockout of the *SfCad* gene. DH19-S, Bt toxin-susceptible *S. frugiperda* without knockout of the *SfCad* gene.

**Table 1.** Comparison of the susceptibility of first instar *S. frugiperda* larvae from the Cad-KO and DH19-S strains to different Bt toxins.


As an additional control, we also tested toxicity of the Vip3Aa protein; we found that the knockout strain Cad-KO did not decrease susceptibility to the Vip3Aa toxin (Table 1 and Figure 2), indicating that SfCad does not participate in Vip3Aa toxicity.

A total of 24 larvae in each group were tested with the indicated concentrations of Bt toxins, and the values of LC50 were calculated after day 7 of oral feeding. Assays were done in triplicate. The 95% fiducial limits (FL) values, shown inside the parenthesis, indicate that there are no significant differences between Cad-KO and DH19-S strains in each column, since these values overlap. Cad-KO are *S. frugiperda* larvae with the knockout of the *SfCad* gene. DH19-S are Bt toxin-susceptible *S. frugiperda* larvae without knockout of the *SfCad* gene.

#### *2.3. SfCAD Expression Did Not Increase Susceptibility of Hi5 Insect Cells to Cry1Ab or Cry1Fa Toxins*

The plasmid pIE2-SfCad-GFP was used to transiently express the fusion protein SfCad-GFP in Hi5 cells. As a control, Hi5 cells were also transfected with pIE2-SfABCC2-GFP that was previously shown to confer susceptibility to Hi5 cells to Cry1Ac toxin [33]. After transfection, cells were observed under the confocal fluorescent microscope, and the results revealed that SfCad-GFP was mainly localized on the cell membrane, suggesting proper expression and folding of the recombinant protein (Figure 3).

The transfection efficiency of the plasmid pIE2-SfCad-GFP was around 45–50% in Hi5 cells. Bioassay data showed that Hi5 cells expressing SfCad-GFP were still tolerant to Cry1Ab or Cry1Fa toxins, since the toxin-treated cells did not swell even at the highest concentration, 20 μg/mL, of these toxins (Figure 4 and Table 2). In contrast, Hi5 cells that were transfected with plasmid pIE2-SfABCC2 were susceptible to Cry1Ab or Cry1Fa toxins (Table 2 and Figure 4). The EC50 values of Bt toxins mediated by SfCad could not be calculated because there were no swollen cells after treatment with the Bt toxins for 1 h. These results also confirmed that SfCad could not mediate cytotoxicity of Cry1Ab and Cry1Fa in Hi5 cells.

**Figure 3.** Subcellular localization of SfCad-GFP in *Trichoplusia ni* Hi5 cells. Green, SfCad-GFP (GFP tag); red, endoplasmic reticulum marker (ER marker); blue, nucleus (Hoechst). Scale bar, 50 μm.

**Figure 4.** Susceptibility of Hi5 cells expressing SfCad-GFP to activated Cry1Ab and Cry1Fa toxins. The cells were transfected with plasmids pIE2-SfCad-GFP or pIE2-GFP (empty vector), respectively, and cultured for 24 h. Then, they were treated with activated toxins at 20 μg/mL for 1 h. A negative control of PBS-treated cells, treated with buffer, is included in the figure. A positive control of cells expressing SfABCC2-GFP is also shown in the figure. The susceptible cells pointed by arrow heads would become swollen, as shown in the positive control. Cells expressing SfCad-GFP or transfected with empty vector showed no swelling of the cells, similar to the negative control. Scale bar, 50 μm.


**Table 2.** Effect of SfCad or SfABCC2 on the cytotoxicity of activated Cry1Ab and Cry1Fa toxins in Hi5 cells.

\* indicates that the cells expressing the putative receptors are not susceptible to the indicated toxins; \*\* the different lowercase letters indicate that there are significant differences between EC50 values of Cry1Ab and Cry1Fa in the same column.

#### *2.4. Cry1Ac, Cry1Ab, and Cry1Fa Did Not Bind to the Toxin-Binding Region (TBR) of SfCad*

The phylogenetic tree constructed with cadherin protein sequences from different Lepidopteran insects showed that the cadherin proteins of three *Spodoptera* species (*S. frugiperda*, *S. exigua,* and *S. litura*) cluster together and share high amino acid sequence identities (around 84%). The *Spodoptera* cadherin cluster is far away from *Helicoverpa armigera* cadherin (Figure 5). It is known that HaCad can mediate toxicity of Cry1Ac in larvae and also induces susceptibility to Cry1Ac when expressed in Hi5 cells [13,14,34].

**Figure 5.** Phylogenetic analysis of cadherin protein in Lepidoptera insects. Harm, *Helicoverpa armigera*; Hzea, *Helicoverpa zea*; Hpun, *Helicoverpa punctigera*; Hvir, *Heliothis virescens*; Msex, *Manduca sexta*; Bman, *Bombyx mandarina*; Bmor, *Bombyx mori*; Msep, *Mythimna separata*; Sinf, *Sesamia inferens*; Snon, *Sesamia nonagrioides*; Sexi, *S. exigua*; Slit, *S. litura*. The Genbank accession numbers of the Cad proteins sequences used in this phylogenetic analysis are indicated in the graph.

Finally, we performed ligand blot binding assays confirming that the toxin-binding regions (TBR) of SfCad, SeCad, and SlCad did not bind to the activated Cry1Ac, Cry1Ab, and Cry1Fa toxins, in contrast to the positive control HaCad that clearly bound to Cry1Ac- and Cry1Ab-activated toxins (Figure 6). Cry1Fa did not bind to any of the TBR regions analyzed, including the TBR from the HaCad protein (Figure 6).

**Figure 6.** Ligand blot analysis of His-tagged toxin-binding regions (TBRs) binding to activated Cry1Ac, Cry1Ab, or Cry1Fa. The His-tagged TBRs of HaCad, SlCad, SeCad, and SlCad were used at different concentrations (0.125, 0.250, and 0.50 μg/mL) with one lane for each His-TBR. Binding of all toxins was assayed at 10 nM. Bound toxin was revealed with the corresponding polyclonal antibody (rabbit anti-Cry1Ac, rabbit anti-Cry1Ab, or rabbit anti-Cry1Fa antibody) as indicated in the Materials and Methods Section.

#### **3. Discussion**

The cadherin proteins from some Lepidopteran insects are involved in susceptibility of those larvae to Cry1A toxins. Mutations or reduced expression of *cadherin* genes in *H. virescens*, *H. armigera,* or *P. gossypiella* are associated with resistance to Cry1Ac [8,11,13,35]. In addition, *Bombyx mori* cadherin was shown to be involved in toxicity of Cry1Aa and Cry1Ab toxins [36,37]. However, *S. litura* and *Trichoplusia ni* cadherins do not function as Cry1Ac receptors [14,38]. A previous study demonstrated that cadherin protein from *H. virescens* functions as a receptor for Cry1A toxins, but not for Cry1Fa, when expressed in *Drosophila* S2 cells, suggesting that Cry1A and Cry1Fa toxins may rely on different receptor molecules [39]. In the present study, both the knockout in *S. frugiperda* insect larvae and over-expression of the *SfCad* gene in cultured Hi5 insect cells indicated that SfCad is not involved in toxicity of Cry1Ab or Cry1Fa in *S. frugiperda*. As described above, it has been shown that Vip3Aa does not share receptors with Cry1A or Cry1Fa toxins [4–7]. Thus, we also performed bioassays of the Cad-KO and DH19 *S. frugiperda* strains with Vip3Aa and showed that there was also no difference in the toxicity of Vip3Aa in the two *S. frugiperda* strains (Figure 2 and Table 1). These results also show that SfCad is not a functional receptor of Vip3Aa in *S. frugiperda*.

Interestingly, we showed that expression of the *SfABCC2* transporter gene in Hi5 cells greatly increased the susceptibility to Cry1Ab or Cry1Fa toxins (Table 2), supporting that ABCC2 is a functional receptor for both Cry1Ab and Cry1Fa toxins in *S. frugiperda,* as previously reported [27]. In the case of Cry1Fa, our results agree with the fact that resistance to Cry1Fa in different populations is linked to mutant alleles of *ABCC2* [28,31]. However, the toxicity of Cry1Ab to Hi5 cells expressing SfABCC2 was 3.5-fold higher than that of Cry1Fa. These results do not correlate with the toxicities of both toxins to wild type DH19 *S. frugiperda* larvae, where Cry1Fa showed 20- to 40-fold higher toxicity than Cry1Ab (Figure 2). These results indicate that Cry1Ab toxicity is limited by some additional mechanisms, other than receptor binding, in the wild type DH19 larvae. It was reported that the lack of toxicity of Cry1Ab to an *S. frugiperda* population from México correlated with enhanced toxin degradation by midgut proteases and also with reduced receptor binding [32]. In addition, it is still possible that an additional Cry1Ab receptor is expressed in the Hi5 cells but not in *S. frugiperda* larvae. Thus, different toxin susceptibility to midgut proteases or lower binding to brush border membrane vesicles (BBMV) could explain the differences in the larval susceptibility to Cry1Fa and Cry1Ab. These hypotheses remain to be analyzed.

As mentioned above, *S. frugiperda ABCC2* mutations are linked with resistance to Cry1Fa [27,28,31]. Interestingly, some Cry1Fa-resistant *S. frugiperda* strains showed cross-resistance to Cry1Ab or Cry1Ac toxins but not to Cry2Ab or Vip3Aa toxins [20,23]. In the present study, the knockout and over-expression of the *SfCad* gene revealed that SfCad is not involved in susceptibility of *S. frugiperda* to Cry1Ab,

Cry1Fa, nor Vip3Aa toxins. Nevertheless, RNAi silencing experiments of *SeCad* showed that cadherin from *S. exigua* might be involved in the toxicity of Cry1Ac and Cry2Aa to some degree [34]. Previously, we reported that SlCad did not increase the toxicity of Cry1Ac when expressed in Hi5 cells, indicating that SlCad is not a functional receptor of Cry1Ac [14]. These data agree with the lack of binding of the TBR from SlCad, SeCad, or SfCad to the Cry1Ac toxin (Figure 6) and support that cadherin proteins from the *Spodoptera* cluster species are not involved in the toxicity of Cry1Ab or Cry1Ac toxins. In the future, we will investigate whether SfCad is involved in Cry2A toxicity in *S. frugiperda*.

Even though some cadherins are not functional receptors of Cry1Ab or Cry1Ac toxins in different insect species, the cadherin protein could be a target for the evolution of Cry1Ab or Cry1Ac variants that bind to this receptor and increase their toxicity to susceptible or resistant insects where cadherin is not a functional receptor of the wild type of Cry1Ab or Cry1Ac. In the case of *T. ni,* Cry1Ac variants that could bind to the TnCad protein were selected by continuous evolution, and it was found that the Cry1Ac variants that were able to bind to TnCad were also able to counter resistance of *T. ni* insects linked to *ABCC2* mutations [40]. In addition, Cry1Ab domain III mutants were shown to increase the toxicity of Cry1Ab to different *S. frugiperda* strains, which was correlated with their increased binding to SfCad receptor [32]. Overall our results show that *S. frugiperda* cadherin is not a functional receptor of Cry1Fa and Cry1Ab toxins. Defining the structural basis for the lack of binding between Cry1Ab or Cry1Fa with SfCad could provide strategies for improving binding and toxicity of these Cry proteins to this invasive pest.

#### **4. Materials and Methods**

#### *4.1. S. frugiperda Strain and Insect Cell Cultures*

The *S. frugiperda* strain DH19 was established from individual moths collected from Dehong, Yunnan Province of China in January 2019 and reared in laboratory conditions on artificial diet without exposure to any insecticide or Bt toxin. Insects were reared at 27 ± 2 ◦C and 75% ± 10% relative humidity (RH) with a photoperiod of 14L:10D. For adults, 10% sugar solution was supplied as a food source.

The *Trichoplusia ni* BTI-Tn-5B1 cell line (Hi5) was established from insect ovaries [41] and kept in our laboratory. The cell line was cultured in Grace's insect cell culture medium (Life Technologies Co., Gand Island, NY, USA) supplemented with 10% fetal bovine serum (Life Technologies Inc.), 100 U/mL penicillin and 100 μg/mL streptomycin (Life technologies Inc.) at 28 ºC under normal atmospheric conditions.

#### *4.2. Preparation of sgRNAs*

A pair of sgRNAs against the *S. frugiperda cadherin* gene (*SfCad*) (Genbank accession no.: AX147205.1) was designed using the sgRNAcas9 design tool [42]. The sgRNA1 target sequence (5- -ATC CTG ACG CAA CTG GAG ACT GG-3- ) and sgRNA2 target sequence (5- -AGG CCA GTC GCT GGT TGT AAC GG-3- ) were selected in exons 4 and 5 of the *SfCad* gene, respectively, (Figure 1A). The selected sgRNAs were analyzed in the *S. frugiperda* genome (https://bipaa.genouest.org), and no potential off-target sites were found. The DNA template for *in vitro* transcription of these sgRNAs was constructed by using PCR-based fusion of two oligonucleotides with a T7 promoter (Target F: TAA TAC GAC TCA CTA TAG + the target sequence; Target R: TTC TAG CTC TAA AAC + the reverse complementary sequence of the target). The PCR conditions were as reported by Jin et al. [43]. The sgRNAs were synthesized using an *in vitro* transcription GeneArt Precision gRNA Synthesis Kit (Thermo Fisher Scientific, Shanghai, China), according to the manufacturer's instructions.

#### *4.3. Cas 9 Protein*

Cas 9 protein (GeneArt Platinum Cas 9 Nuclease) was purchased from Thermo Fisher Scientific (Shanghai, China).

#### *4.4. Egg Collection and Microinjection*

Freshly laid eggs (within 2 h of oviposition) were washed with distilled water. Then, the eggs were placed on a microscope slide and fixed with double-sided adhesive tape. We injected each egg with 1–2 nL of a mixture solution containing two sgRNAs (150 ng/μL for each) and Cas 9 protein (50 ng/μL) using Nanoject III (Drummond, Broomall, PA, USA). The injected eggs were incubated at 25 ◦C and 65% RH for hatching.

#### *4.5. Identification of SfCad Mutations Mediated by CRISPR*/*Cas 9 System*

To identify the mutations, a specific pair of primers (Cad-F: CCT CCT CAA ATA AGA TTA CC; Cad-R: ATG ATG GGC GCA TTG TCG T) were designed that flanked the target sites, and genomic DNA samples of individual insects were used as the template. The genomic DNA of the larvae was extracted using a Multisource Genomic DNA Miniprep Kit (Axygen, New York, NY, USA) according to the manufacturer's instructions. The PCR conditions were as reported by Jin et al. [42]. Then, 10 μL PCR products were analyzed by agarose gel electrophoresis. Multiple bands indicated that double nicking had occurred. To analyze the exact type of mutation (insertion or deletion), the bands were recovered, cloned, and sequenced by Sangon Biotech (Shanghai, China).

#### *4.6. Bt Toxins and Bioassay*

The activated Cry1Ab toxin and Vip3Aa protoxins used in the *in vivo* bioassay were provided by the institute of Plant Protection, Chinese Academy of Agricultural Science (CAAS), Beijing, China. The other purified activated and lyophilized Cry1Ac, Cry1Ab, and Cry1Fa toxins were kindly donated by Dr. Marianne Pusztai-Carey from Case Western Reserve University, USA. Toxicity of each Bt toxin to DH19 and SfCad knockout strain was determined with diet overlay bioassays. Gradient concentrations of Bt toxin solution were prepared by diluting the stock suspensions in PBS (pH 7.0) solution. Artificial diet (900 μL) was dispensed into a 24-well plate (surface area per well = 2 cm2) and after the diet cooled down, 50 μL Bt toxin solution was applied on the surface in each well. A single 1st-instar larva was put in each well after the toxin solution was dried at room temperature, and mortality was recorded after 7 days. The LC50 (median lethal concentration that killed 50% of the tested larvae) and the corresponding 95% fiducial limits were calculated through Probit analysis of the mortality data using SPSS. Control wells were treated with buffer solution.

#### *4.7. Plasmids, Transfection, and Fluorescence Observation*

The *SfCad* gene was synthesized by Genescript Company (Nanjing, China) and inserted into a pGEM-T easy vector (Promega Inc., Madison, WI, USA). Then, the gene was amplified by PCR and inserted into pIE2-GFP-N1, and the new construct was named as pIE2-SfCad-GFP [14]. The plasmid purified from the transformed *E. coli* DH5α was transfected into Hi5 cells as previously reported [14]. Briefly, Hi5 cells were grown overnight in 48-well cell culture plates (Corning Inc., New York, NY, USA) at 1.2 <sup>×</sup> 10<sup>5</sup> cells/well. Then, the transfection was performed using the mixture of the plasmid (250 ng/well) with a transfection kit Genefusion HD (1 μL/well) (Promega Inc., Madison, WI, USA). Plasmids pIE2-GFP-N1, pIE2-SfCad-GFP, pIE2-SfABCC2-GFP, and pIE2-dsRED-ER were previously reported [33,44]. At 24 h post transfection, the cells were fixed using 4% paraformaldehyde for 10 min, and stained by Hoechst 33,342 (1 μg/mL) for 10 min. Then, the cells were observed and photographed under a laser confocal scanning microscope (Carl Zeiss, Jena Deutschland, Germany). The transfection efficiency was calculated: A = the number of cells emitting green fluorescence (SfCad-GFP) divided by the number of cells emitting blue fluorescence (nucleus stained by Hoechst 33,342) × 100%. Three biological replicates were performed.

#### *4.8. Cytotoxicity Assay*

Hi5 cells were transfected using single plasmids (pIE2-SfCad-GFP or pIE2-SfABCC2-GFP) as described above. At 24 h post transfection, the cells were treated with the indicated toxin concentrations (at least five different concentrations, two-fold serial dilution) of activated Bt toxins (Cry1Ab or Cry1Fa) for 1 h, and they were photographed under an inverted confocal microscope (Nikon, Tokyo, Japan). The cells transfected with the empty vector (pIE2-GFP-N1) were used as a control group and were also treated with the Cry toxins. An additional negative control of cells treated with phosphate buffer solution (PBS) was included in these assays. The percentage of swollen cells resulting from these toxin treatments was calculated as follows: B = the number of the swollen cells divided by the number of the total cells × 100%. The percentage of the swollen cells expressing SfCad-GFP or SfABCC2-GFP was calculated as follows: C = B/A × 100%. The transfection efficiency (A) was described above in Section 4.7. The effective concentration inducing 50% mortality value (EC50) was obtained by regression analysis using SPPS 16.0 software. For two particular populations, the EC50 values were considered as significantly different if their 95% fiducial limits (FL) did not overlap [45].

#### *4.9. Construction of a Lepidoptera Cadherin Phylogenetic Tree*

The sequences of Lepidopteran insect cadherin proteins were selected for constructing a cadherin evolutionary tree by analyzing their phylogeny. GenBank accession numbers of the sequences of these cadherin proteins are as follow. Harm: *Helicoverpa armigera* cadherin, AFB74174.1; Hzea: *Helicoverpa zea* cadherin, AKH49609.1; Hpun: *Helicoverpa punctigera* cadherin, AVE17268.1; Hvir: *Heliothis virescens* cadherin, AAV80768.1; Msex: *Manduca sexta* cadherin, AAG37912.1; Bman: *Bombyx mandarina* cadherin, XP\_028026250.1; Bmor: *Bombyx mori* cadherin, BAA99404.1; Msep: *Mythimna separata* cadherin, AEI61920.1; Sinf: *Sesamia inferens* cadherin, AEL22856.1; Snon: *Sesamia nonagrioides* cadherin, ABV74206.1; Sexi: *S. exigua* cadherin, AFH96949.1; Slit: *S. litura* cadherin, XP\_022826291.1. The phylogeny of these sequences was analyzed using the neighbor-joining tree method with MEGA 5.0 software (https://mega.software.informer.com/5.0/).

#### *4.10. Purification of Proteins Expressed in Bacteria*

The coding DNA of toxin-binding regions of *SfCad*, *SlCad*, *SeCad,* and *HaCad* were amplified by PCR from the corresponding plasmids containing these genes or the cDNA obtained from midgut tissue of these insects [14]. The primers are listed in Table 3. The amplified fragments were purified and digested with restriction nucleases and cloned into the cleaved plasmids listed in Table 3. The constructs were transformed into *Escherichia coli* BL21 cells and the His-tagged proteins were purified with Ni-NTA affinity column (GE Healthcare Bioscience, Piscataway, NJ, USA) according to the manufacturer's manual. Detailed protocols were described previously [14]. All the purified proteins were stored at −80 ◦C until use.


**Table 3.** Primers used for expression of the different fragments of proteins.

The underlined letters indicate restriction sites of endonucleases.

#### *4.11. Ligand Blot Assays*

The 6 × His-tagged HaCad, SfCad, SlCad, or SeCad TBRs were separated on 12% SDS-PAGE gel and transferred onto Polyvinylidene fluoride (PVDF) membrane (three different membranes were prepared). The loading of proteins on these membranes was checked by Ponceau S staining 0.2% (*w*/*v*) in 3% (*v*/*v*) acetic acid and followed by complete destaining by washing with water. These membranes were blocked with 2% BSA in PBS-Tween (0.2%) for 3 h, then each membrane was incubated with a different activated Cry toxin (Cry1Ab, Cry1Ac or Cry1Fa toxin) at 10 nM for 2 h. After washing three times with PBS-Tween (0.2%), the membranes were further incubated with the corresponding polyclonal antibody (rabbit anti-Cry1Ac, rabbit anti-Cry1Ab, or rabbit anti-Cry1Fa antibody) diluted in PBS-Tween (0.2%) at 1:1000 dilution for 3 h. Then, the membranes were incubated with horseradish peroxidase (HRP)-conjugated goat anti-rabbit secondary antibody (Abbkine) diluted in PBS at 1:10,000. Finally, the membranes were incubated with the enhanced chemiluminescence (ECL) reagent (GE Healthcare Biosciences, Piscataway, NJ, USA) and then covered with X-ray film for exposure for a few minutes, and the film was developed and fixed as previously described [46].

**Author Contributions:** Conceptualization, K.L. and A.B.; Formal analysis, J.Z., M.J., Y.Y. (Yanchao Yang), Y.X., L.L., I.G., and M.S.; Funding acquisition, K.L.; Investigation, J.Z., M.J., Y.Y. (Yangchao Yang), and Y.Y. (Yongbo Yang); Writing original draft, J.Z. and M.J.; Writing review and editing, K.L., A.B., and M.S.; Supervision, K.L. All authors have read and agreed to the published version of the manuscript.

**Funding:** This work was supported by the National Key R&D Program of China (grant number: 2017YFD0200400).

**Conflicts of Interest:** M.S. and A.B. are coauthors of a patent on modified Bt toxins. "Suppression of resistance in insects to *Bacillus thuringiensis* Cry toxins, using toxins that do not require the cadherin receptor" (patent numbers: CA2690188A1, CN101730712A, EP2184293A2, EP2184293A4, EP2184293B1, WO2008150150A2, WO2008150150A3).

#### **References**


© 2020 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).

*Article*

### **CRISPR-Mediated Knockout of the** *ABCC2* **Gene in** *Ostrinia furnacalis* **Confers High-Level Resistance to the** *Bacillus thuringiensis* **Cry1Fa Toxin**

#### **Xingliang Wang, Yanjun Xu, Jianlei Huang, Wenzhong Jin, Yihua Yang and Yidong Wu \***

College of Plant Protection, Nanjing Agricultural University, Nanjing 210095, China; wxl@njau.edu.cn (X.W.); 2016102098@njau.edu.cn (Y.X.); 2016202024@njau.edu.cn (J.H.); 2018102105@njau.edu.cn (W.J.); yhyang@njau.edu.cn (Y.Y.)

**\*** Correspondence: wyd@njau.edu.cn; Tel.: +86-25-8439-6062

Received: 16 March 2020; Accepted: 9 April 2020; Published: 11 April 2020

**Abstract:** The adoption of transgenic crops expressing *Bacillus thuringiensis* (Bt) insecticidal crystalline (Cry) proteins has reduced insecticide application, increased yields, and contributed to food safety worldwide. However, the efficacy of transgenic Bt crops is put at risk by the adaptive resistance evolution of target pests. Previous studies indicate that resistance to *Bacillus thuringiensis* Cry1A and Cry1F toxins was genetically linked with mutations of ATP-binding cassette (ABC) transporter subfamily C gene *ABCC2* in at least seven lepidopteran insects. Several strains selected in the laboratory of the Asian corn borer, *Ostrinia furnacalis*, a destructive pest of corn in Asian Western Pacific countries, developed high levels of resistance to Cry1A and Cry1F toxins. The causality between the *O. furnacalis ABCC2* (*OfABCC2*) gene and resistance to Cry1A and Cry1F toxins remains unknown. Here, we successfully generated a homozygous strain (OfC2-KO) of *O. furnacalis* with an 8-bp deletion mutation of *ABCC2* by the CRISPR/Cas9 approach. The 8-bp deletion mutation results in a frame shift in the open reading frame of transcripts, which produced a predicted protein truncated in the TM4-TM5 loop region. The knockout strain OfC2-KO showed much more than a 300-fold resistance to Cry1Fa, and low levels of resistance to Cry1Ab and Cry1Ac (<10-fold), but no significant effects on the toxicities of Cry1Aa and two chemical insecticides (abamectin and chlorantraniliprole), compared to the background NJ-S strain. Furthermore, we found that the Cry1Fa resistance was autosomal, recessive, and significantly linked with the 8-bp deletion mutation of *OfABCC2* in the OfC2-KO strain. In conclusion, *in vivo* functional investigation demonstrates the causality of the *OfABCC2* truncating mutation with high-level resistance to the Cry1Fa toxin in *O. furnacalis*. Our results suggest that the *OfABCC2* protein might be a functional receptor for Cry1Fa and reinforces the association of this gene to the mode of action of the Cry1Fa toxin.

**Keywords:** Asian corn borer; *ABCC2*; CRISPR/Cas9; Cry1Fa; resistance

**Key Contribution:** In vivo functional investigation demonstrates the causality of the *ABCC2* truncating mutation with high level of resistance to the Cry1Fa toxin in *Ostrinia furnacalis*. Our results suggest that *O. furnacalis* ABCC2 might be a functional receptor for Cry1Fa and reinforces the association of this gene to the mode of action of the Cry1Fa toxin.

#### **1. Introduction**

Transgenic crops expressing *Bacillus thuringiensis* (Bt) insecticidal crystalline (Cry) proteins have been commercialized worldwide since 1996. The global planting area of Bt crops reached 104 million hectares in 2018 [1]. The widespread Bt crop adoption has suppressed pest populations, reduced insecticide usage, promoted biocontrol services, and economically benefited growers [2]. However, the efficacy of Bt crops is put at risk from the adaptive evolution of resistance by the target pests, and practical resistance to Bt crops has been documented at least in nine pest species in six countries [3–5].

The European corn borer *Ostrinia nubilalis* (Hübner) and the Asian corn borer *Ostrinia furnacalis* (Guenée) are two sibling species, both of which are economically important insect pests of corn, *Zea mays* (L.) [6]. *O. nubilalis* is present in Europe, North Africa, Central Asia, and North America [7], while *O. furnacalis* is distributed widely in East and Southeast Asia, Australia, and the Western Pacific Islands [8]. Bt corn expressing Cry1Ab has been widely planted for the control of some lepidopteran pests, including *O. nubilalis*, in North America since 1996, resulting in the suppression of target pest populations and reduced insecticide applications in both Bt and non-Bt corn [9,10]. No practical resistance to Cry1Ab has been identified in *O. nubilalis* field populations from North America for more than 20 years [5]. Bt corn expressing Cry1F has been used commercially in North America since 2003, and the frequency of alleles conferring Cry1F resistance did not increase in field populations of *O. nubilalis* sampled during 2003 to 2009 from the US corn belt [11]. However, practical resistance to Cry1F was discovered in 2018 from *O. nubilalis* populations from Nova Scotia of Canada [4]. It indicates that Bt resistane has already become a real threat to the long-term effectiveness of Bt corn for the control of *O. nubilalis*.

China is a major corn producer in the world and its corn acreage was 41.5 million hectares in 2018 [12]. *O. furnacalis* is the domiant pest and widely distributed in most of the corn-growing regions of China, while *O. nubilalis* is limited to some regions of northwestern China [13]. Although the commercial planting of Bt corn has not yet been approved in China, two Bt corn events (DBN9936 and Shuangkang12-5) obtained biosafety certificates in 2019 (MARA, 2020) [14], which is considered a prerequisite and landmark for commercial production. To be prepared for the switch to the adoption of Bt corn in the near future, a number of investigations have been conducted in China to assess resistance risk and cross-resistance by laboratory selection of *O. furnacalis* with Bt proteins. Under laboratory selection conditions, *O. furnacalis* developed high levels of resistance to various Cry1 toxins, including Cry1Ab, Cry1Ac, Cry1Ah, Cry1F, and Cry1Ie [15–18], proving its potential to develop Bt resistance in the field. Symmetrical cross-resistance was found among Cry1Ab, Cry1Ac, Cry1Ah, and Cry1F [15–18]. Asymmetrical cross-resistance was observed between Cry1Ie and other Cry1 toxins. Selection with Cry1Ab, Cry1Ac, Cry1Ah, or Cry1F did not confer cross-resistance to Cry1Ie, but selection with Cry1Ie resulted in high-level cross-resistance to Cry1F [15–19]. These results are valuable for the future designing of resistance management strategies for Bt corn in China. However, the resistance mechanisms underlying Bt resistance of *O. furnacalis* remain elusive.

Several proteins have been identified and characterized as receptors for Cry toxins, including cadherins, aminopeptidase *N* (APN), alkaline phosphatases (ALP), and ATP-binding cassette (ABC) transporters [20]. One of the major mechanisms of resistance to Cry toxin is reduced toxin binding to their specific larval midgut receptors through the disruption of the receptor genes [21]. Since the disruption of the ABC transporter subfamily C2 (*ABCC2*) gene was first identified to confer Cry1Ac resistance in *Heliothis virescens* [22], mutations of the homologous *ABCC2* genes associated with Cry1A and/or Cry1F resistance have been found in several lepidopteran insects, including *Plutella xylostella*, *Trichoplusia ni* [23], *Bombyx mori* [24], *Helicoverpa armigera* [25], *Spodoptera exigua* [26], and *Spodoptera frugiperda* [27–29]. Recently, the CRISPR/Cas9 system has been applied to investigate the *in vivo* role of insect *ABCC2* in the mode of action and resistance mechanisms of Bt toxins. The causal relationship between *ABCC2* knockout and Cry1A/Cry1F resistance has been confirmed in *P. xylostella* [30], *S. frugiperda* [31], and *S. exigua* [32]. Interestingly, the knockout of either *ABCC2* or *ABCC3* of *H. armigera* did not confer Cry1Ac resistance, whereas the knockout of *ABCC2* and *ABCC3* together resulted in extremely high levels of resistance to Cry1Ac [33]. However, until now, whether or not the *ABCC2* gene of *O. furnacalis* (*OfABCC2*) is involved in Bt resistance development remains unknown.

In this study, we knocked out the *OfABCC2* employing the CRISPR/Cas9 system and constructed a homozygous mutant strain (OfC2-KO). Next, we performed toxicity bioassays and found that the *OfABCC2* knockout obtained a resistance to Cry1Fa greater than 300-fold compared to the wild-type control strain. Finally, we accessed the inheritance mode of the acquired resistance and confirmed the linkage between manipulated gene deletion and high-level resistance to Cry1Fa in the OfC2-KO strain.

#### **2. Results**

#### *2.1. CRISPR*/*Cas9-Mediated Mutagenesis of OfABCC2 in O. Furnacalis*

A total of 572 newly laid eggs (< 2 h) were injected with a mixture of the synthesized sgRNA and Cas9 protein. A total of 150 injected embryos (~26%) hatched and developed to 5th instar larvae. In order to obtain individuals with edited genomes as quickly as possible, the genomic DNA of 90 exuviates of the final instar larvae were isolated, and *OfABCC2* genotypes were identified by the direct sequencing of PCR products flanking the target site. Sequencing chromatograms revealed that 7.8% (7/90) of the examined G0 individuals were mutagenized at the target site with a stretch of double peaks. Then, the seven chimeras (six females and one male) were single crossed with the wild-type NJ-S moth, respectively (G0, Figure 1).

After G0 oviposited, the genomic DNA of randomly selected eight–nine exuviates from each single-pair progeny were prepared, and their *OfABCC2* genotype was identified as described above. Among the 60 exuviates genotyped, 46 samples were wild-type homozygotes, seven individuals were heterozygotes harboring a wild-type allele and an 8-bp deletion allele, three samples were heterozygotes carrying a wild-type allele and a 1-bp insertion allele, and the genotype of the rest of the four individuals could not be identified by visual checks based on the sequencing chromatogram. We therefore confirmed efficient mutagenesis induced by CRISPR/Cas9 system had occurred in *OfABCC2* and the genome-edited alleles were transmitted to G1.

**Figure 1.** Diagram of the crossing strategy used to obtain the knockout strain homozygous for the 8-bp deletion mutation in exon 4 of *OfABCC2*. (+/-) means heterozygote (0/-8), (-/-) means mutant homozygote (-8/-8).

#### *2.2. Construction of a Homozygous Strain with OfABCC2 Knocked Out*

The mass mating was made among the above seven heterozygotes (three males and four females) that were harboring a wild-type allele and an 8-bp deletion allele (+/-) in G1 (Figure 1). The genomic DNA of 30 exuviates from G2 progeny were isolated and the genotype of *OfABCC2* was screened, and 21, five, and four samples were respectively identified as wild-type homozygotes (+/+), heterozygotes (+/-), and mutant homozygotes (-/-). The four moths (three females and one male) harboring both the 8-bp deletion alleles were mass crossed and their progeny (G3) was reared to form a homozygous knockout strain named OfC2-KO (Figure 1). Subsequently, the TA-clone sequencing of the PCR products using both gDNA and cDNA from the OfC2-KO larvae were performed, and confirmed the *OfABCC2* carrying the 8-bp deletion at the desired site (data not shown).

*Toxins* **2020**, *12*, 246

Based on the deduced peptide sequences, the 8-bp deletion at exon 4 caused a pre-mature stop codon (Figure 2a,b). The consequence of this 8-bp deletion is predicted to lose TM5-TM12 transmembrane segments and two nucleotide-binding domains (NBDs) (Figure 2c). In view of the absence of about 70% of the protein structure, the *ABCC2* gene in the OfC2-KO strain is predicted to be defective and most likely non-functional.

**Figure 2.** CRISPR/Cas9-mediated editing of the *OfABCC2* gene. (**a**) A diagram of the *OfABCC2* gene and sgRNA targeting site. The white boxes represent predicted exons through sequence alignment with *ABCC2*s from *Heliothis virescens* and *Plutella xylostella*. The sgRNA targeting site was located at exon 4, containing a proto spacer and a protospacer adjacent motif (PAM) sequence (TGG, in red). (**b**) The deduced peptide sequences from partial exon 4 to exon 6 of *OfABCC2*. The stop code is shown by a red asterisk. (**c**) A schematic diagram of the 12 transmembrane domains (TM1–TM12). The cleaved site induced by CRISPR/Cas9 is located at TM4, resulting in a frame shift of the transcript. The predicted protein produced from this mutant allele would be truncated in the intracellular TM4–TM5 loop of OfABCC2.

#### *2.3. Impact of OfABCC2 Disruption on the Susceptibility of O. Furnacalis to Bt Toxins and Chemical Insecticides*

Toxicity assays with larvae from the mutagenesis OfC2-KO strain and the background NJ-S strain against four Bt Cry toxins and two insecticides were carried out with the aim of assessing the impact of disrupted *OfABCC2* on larvae's susceptibility. The bioassay results indicate that the OfC2-KO strain showed low levels of resistance to Cry1Ac (8.1-fold) and Cry1Ab (3.6-fold), but no significant resistance to Cry1Aa (1.4-fold) compared to the NJ-S strain based on LC50 values (Table 1). However, because the susceptibility of the OfC2-KO strain to Cry1Fa was reduced to a large extent, the LC50 cannot be obtained by bioassay. The mortality of the OfC2-KO larvae was only 4% when treated by 120 μg/g Cry1Fa, and the estimated resistance ratio was much more than 300-fold. In contrast, the two strains exhibited approximately equal susceptibility to two chemical insecticides (abamectin and chlorantraniliprole) with 0.6- and 1.3-fold difference of LC50s. Our findings provide strong reverse genetics evidence for *OfABCC2* involved in the toxicity and mode of action of Cry1Fa.


**Table 1.** Toxicity response to four Bt toxins and two chemical insecticides of larvae from the original NJ-S and OfC2-KO strains of *O. furnacalis*.

<sup>1</sup> Numbers of larvae used in bioassay; <sup>2</sup> RR (resistance ratio) = LC50 (OfC2-KO)/LC50 (NJ-S); <sup>3</sup> LC50 for OfC2-KO could not be determined because of an insufficient dose response (only 4% mortality at 120 μg/g of Cry1Fa treatment).

#### *2.4. Dominance of Cry1Fa and Cry1Ac Resistance in the OfC2-KO Strain*

To investigate the inheritance of different levels of resistance to Cry1Fa (high) and Cry1Ac (low) in the OfC2-KO strain, it was crossed with the susceptible NJ-S strain, and the responses of the two strains and their F1 progeny were determined at a diagnostic concentration of Cry1Fa (2 μg/g) and Cry1Ac (1 μg/g), respectively (Table 2). For Cry1Fa, the F1a and F1b progeny had a high mortality (100% and 98.3%) at 2 μg/g, and the dominance parameters (*h*) were 0 and 0.02. Similarly, for Cry1Ac, the corresponding mortality was 100%, and both of the two *h* values were 0. The results indicated that either a high level of resistance to Cry1Fa or a low level of resistance to Cry1Ac in OfC2-KO strain was inherited as a recessive mode.

**Table 2.** Mortality and dominance of the susceptible NJ-S strain, OfC2-KO strain, and their F1 progeny from reciprocal crosses to the diagnostic concentration of Cry1Fa and Cry1Ac, respectively.


<sup>1</sup> Numbers of larvae measured at the Cry1Fa (2 μg/g) or Cry1Ac (1 μg/g) diagnostic concentration; <sup>2</sup> The degree of dominance (*h*) = (survival of F1 - survival of NJ-S)/(survival of OfC2-KO - survival of NJ-S). *h* = 0, completely recessive; *h* = 1, completely dominant.

#### *2.5. Genetic Association between the 8-bp Deletion of OfABCC2 and Cry1Fa Resistance*

To clarify the causal relationship of the 8-bp deletion in exon 4 of *OfABCC2* with high levels of Cry1Fa resistance, a set of genetic crosses was performed (Figure 3a). By using direct-sequencing analysis of the target PCR products (see typical chromatogram in Figure 3b), the genotype of 25 individuals from NJ-S were homozygous for the wild-type (*ss*) and that of 30 individuals from the OfC2-KO strain were homozygous for the 8-bp deletion of *OfABCC2* (*rr*) (Table 3). When treated with 2 μg/g of Cry1Fa in F2 progeny, 22.6% (38/168) of the larvae survived after 7 days of treatment. All the detected 21 survivors randomly selected from the F2-treated group were homozygous for the 8-bp deletion of *OfABCC2* (*rr*), and the F2-untreated individuals were separated into wild-type homozygous (*ss*: 7), heterozygous (*rs*: *13*), and 8-bp deletion homozygous (*rr*: 9). Our results clearly demonstrated that the 8-bp deletion of *OfABCC2* is significantly linked (Fisher's exact test, *p* < 0.0001) with Cry1Fa resistance in the manipulated OfC2-KO strain.

**Figure 3.** Linkage analysis of Cry1Fa resistance in the OfC2-KO strain of *O. furnacalis*. *OfABCC2* genotypes: *ss* = wild type; *rs* = heterozygous mutant; *rr* = homozygous mutant (8-bp deletion). (**a**) The crossing design used to generate F2 progeny (1*ss*: 2*rs*:1*rr*). (**b**) The direct sequencing chromatograms of PCR products amplified from a fragment of gDNA flanking the 8-bp deletion site (red box) of *OfABCC2*.

**Table 3.** Genetic linkage between the 8-bp deletion of *OfABCC2* and resistance to Cry1Fa in *O. furnacalis*.


<sup>1</sup> F1 progeny between the susceptible NJ-S and Cry1Fa-resistant *OfABCC2* strains were crossed to produce F2 progeny. 168 larvae from the F2 progeny were treated with 2 μg/g of Cry1Fa toxin. 21 of 38 survivors and 30 untreated larvae were genotyped individually by direct sequencing of the PCR products; <sup>2</sup> *ss* represent homozygous for the wild type *OfABCC2*, while *rs* means heterozygous for the 8-bp deletion allele of *OfABCC2*, and *rr* represent homozygous for the 8-bp deletion allele of *OfABCC2*.

#### **3. Discussion**

In the current study, we successfully induced a deletion mutation of 8-bp into the *OfABCC2* gene of *O. furnacalis* by the CRISPR/Cas9 genome editing system, and characterized Bt resistance properties of the knockout OfC2-KO strain. We found that the OfC2-KO strain acquired a high level of resistance to Cry1Fa (>300-fold) and low levels of resistance to Cry1Ab and Cry1Ac (< 10-fold). We also confirmed the genetic association between the 8-bp deletion of *OfABCC2* and the obtained resistance to Cry1Fa in the knockout strain. These findings provide strong evidence that OfABCC2 plays a major role in meditating the toxicity of Cry1Fa in *O. furnacalis*. Moreover, the cross-resistance and inheritance pattern results provide helpful information for designing of resistance management strategies for future adoption of Bt corn in China. Furthermore, the OfC2-KO strain can be employed in an F1 screen program to investigate the diversity and frequency of the *OfABCC2* mutant alleles in field populations of *O. furnacalis*.

ABCC2 proteins have been identified as receptors for Bt toxins Cry1A and/or Cry1F in a number of lepidopteran insects, but they have differential contributions to the toxicities for individual Cry1 toxins. The CRISPR-mediated knockout of *P. xylostella ABCC2* conferred high levels of resistance to Cry1Aa, Cry1Ab, and Cry1Ac [30]. The double knockout of *ABCC2* and *ABCC3* of *H. armigera* confers a >15,000-fold resistance to Cry1Ac [33]. A point mutation in the *ABCC2* of *B. mori* resulted in high levels of resistance to Cry1Ab and Cry1Ac, but not to Cry1Aa [24]. The CRISPR-mediated knockout of

*S. frugiperda ABCC2* conferred a 118-fold resistance to Cry1F [31], and the knockout of *S. exigua ABCC2* resulted in high levels of resistance to both Cry1Fa and Cry1Ac [32]. In our study, the knockout of *ABCC2* in *O. furnacalis* produced high-level resistance to Cry1Fa, low levels of resistance to Cry1Ab and Cry1Ac, and no resistance to Cry1Aa. The present study provides a new case for the investigation of the interaction between lepidopteran ABCC2 receptors and Bt Cry1 toxins.

A laboratory-selected strain of *O. nubilalis* developed a >1200-fold resistance to Cry1F, and the Cry1F resistance trait is controlled by a single quantitative trait locus (QTL) on linkage group 12 [34]. The subsequent fine mapping of the Cry1F resistance QTL identified a genomic region containing the *ABCC2* locus tightly linked with Cry1F resistance [35]. Practical resistance to Cry1F was recently documented in some field populations of Canadian *O. nubilalis* [4]. It will be interesting to check whether there are mutations in the *ABCC2* gene in both the laboratory-selected strain and field-derived resistant populations of *O. nubilalis*. The identification of the specific gene for Cry1F resistance of *O. nubilalis* is urgently needed for developing molecular tools to monitor the spreading of the practical resistance in the field.

Several studies reported the potential mechanisms of Cry1Ab and Cry1Ac resistance in the laboratory-selected strains of *O. furnacalis*, such as the up-regulation of the V-ATPase catalytic subunit A and heat shock 70 kDa proteins [36], the down-regulation and mutation of a cadherin gene [37], the differential expression of the miRNAs targeting potential Bt receptors [38], and the reduced expression of APN and ABC subfamily G transcripts [39]. The CRISPR-mediated knockout approach established for *O. furnacalis* in the present study can be used to evaluate the functional role of the candidate genes relating to Bt resistance.

The characterization of the inheritance of Bt resistance will provide important information for evaluating the risks of evolution of resistance and will make it possible to formulate effective resistance management strategies. Based on previous reports, resistance to Cry1-type toxins mediated by *ABCC2* mutations was recessive or incompletely recessive [22–24,27,28,30,32,33]. Consistent with these results, both the high-level resistance to Cry1Fa (>300-fold) and low-level resistance to Cry1Ac (~8-fold) were inherited as a recessive mode in the knockout OfC2-KO strain of *O. furnacalis*.

In the present work, the obtained Cry1Fa resistance was confirmed to be genetically associated with the 8-bp deletion of *OfABCC2*, which excludes the CRISPR-mediated off-target effects on resistance phenotype. We analyzed 18 research cases that employed the CRISPR/Cas9 system to manipulate the resistance genes to Bt toxins or insecticides, and found that only five of them performed linkage analysis between acquired resistance and the introduced mutation, including the knockout of the cadherin gene in *H. armigera* [40], nicotinic acetylcholine receptor α6 subunit in *P. xylostella* and *S. exigua* [41,42], the ryanodine receptor G4946E mutation in *Drosophila melanogaster* [43], and a *CYP9M10* gene in *Culex quinquefasciatus* [44]. We therefore recommend that when CRISPR-based gene editing is conducted to verify the function of a candidate gene, it is necessary to perform a genetic linkage analysis in order to clarify whether there are off-target effects.

#### **4. Materials and Methods**

#### *4.1. Insect Strains and Rearing*

The susceptible NJ-S strain was originally collected from Nanjing, China, in May 2010, and has been maintained in the laboratory without exposure to any insecticides or Bt toxins. By using the CRISPR/Cas9 genome-editing system, the *OfABCC2* gene in the background strain NJ-S was knocked out to construct a manipulated strain denoted as OfC2-KO. The genome-edited OfC2-KO strain is homozygous for the 8-bp deletion in exon 4 of *OfABCC2*, which was predicted to produce a truncated and loss-of-function protein.

The larvae of *O. furnacalis* were reared on an artificial diet with corn and soybean powder as major ingredients at 27 ± 1 ◦C, 80% relative humidity (RH), and a photoperiod of 16 h light:8 h dark. The pupae were transferred to mating cages with more than 80% RH and a photoperiod of 16:8 h (L: D). Adults were supplied with 10% sugar solution to replenish energy. About 5–6 pieces of waxed papers, as substrate for oviposition, were placed on the top of the cage, and the bottom sheet was collected daily. Egg masses were incubated in plastic boxes lined with moistened filter paper until hatching.

#### *4.2. Diet Bioassay*

The activated Cry1Aa, Cry1Ab, Cry1Ac, and Cry1Fa toxins were purchased from Marianne Pusztai-Carey (Case Western Reserve University, Cleveland, OH, USA). Abamectin (20 g/L EC) was obtained from the Institute of Plant Protection, Guangdong Academy of Agricultural Sciences, Guangzhou, Guangdong Province, China. Chlorantraniliprole (200 g/L SC) was purchased from DuPont Agricultural Chemicals Ltd. (Wilmington, DE, USA).

We used the diet incorporation method to evaluate the toxicity of Cry toxin or chemical insecticide to *O. furnacalis*. Briefly, 5 to 7 concentrations of Bt or insecticide test solutions were first diluted in distilled water. Then, we added the solution (or distilled water for control) to a proper amount diet in a clean mixing bowl and thoroughly mixed all the ingredients together until a soft, smooth dough was obtained. Next, we dispensed the toxin-incorporated diet into each well of a 24-well plate. After the diet cooled and solidified, one *O. furnacalis* larva (neonate for Cry toxin susceptibility test and 2nd instar larva for chemical insecticide bioassay) was placed in each well. All the plates were kept at an illumination incubator set at 27 ± 1 ◦C, 80% RH, and a photoperiod of 16:8 h (L:D). For Cry toxin, the mortality was recorded after 7 days of treatment, and the larvae were considered dead if they died or weighed less than 5 mg. For abamectin and chlorantraniliprole, the mortality was recorded after 2 days of treatment, and the larvae were considered dead if they did not move after gentle prodding with a brush. The data were analyzed with PoloPlus (LeOra Software) [45] to estimate the LC50 with 95% fiducial limits (FL), as well as the slopes of the concentration–mortality lines. Resistance ratios (RR) were calculated by dividing the LC50 for a particular strain by the LC50 for the susceptible NJ-S strain.

#### *4.3. Design and Preparation of sgRNA*

In our previous work, the full-length sequences of *OfABCC2* mRNA (GenBank no. MN783372) had been obtained from the susceptible NJ-S strain of *O. furnacalis*. By scanning the GN19NGG motifs, we identified a sgRNA target site (5- -GCACCTTTCGTTGGACTTTTTGG-3- ) in predicted exon 4 of *OfABCC2* (Figure 2a). A PCR-based approach was employed to prepare sgRNA according to the instructions [46]. In brief, a forward oligonucleotide encoding a T7 polymerase-binding site and the sgRNA target sequences GN19 (OfC2\_sgF, Table 4) and a universal oligonucleotide encoding the remaining sgRNA sequences (OfC2\_sgR, Table 4) were designed at first. The OfC2\_sgF and OfC2\_sgR were fused by PCR to generate a sgRNA DNA template. The PCR reaction mixture (50 μL) contained 10 μL of 5 × PCR buffer, 4 μL of 2.5 mM dNTP, 4 μL of 10 μM OfC2\_sgF, 4 μL of 10 μM OfC2\_sgR, 0.5 μL of PrimeSTAR polymerase (TaKaRa, Dalian, China), and 27.5 μL of Nuclease-free water. PCR was performed at 98 °C for 30 s, 30 cycles (98 °C 5 s, 60 °C 30 s, 72 °C 15 s), 72 °C for 10 min, and holding at 4 °C. After identification by electrophoresis, the PCR products were purified by a QIAprep® Spin Miniprep Kit (QIAGEN, Hilden, Germany). A MEGAshortscript™ T7 High Yield Transcription Kit (Ambion, Foster City, CA, USA) was used for sgRNA in vitro transcription according to the manufacturer's protocol.


#### *4.4. Egg Collection and Microinjection*

Mated female moths of *O. furnacalis* were allowed to lay egg masses on pieces of wax paper previously placed on the top of the mating cage. Fresh egg masses (within 2 h after oviposition) were immediately collected by cutting the wax paper. Then, the eggs were lined on double-sided adhesive tape on a microscope slide. About 1 nL of mixture containing 150 ng/μL of Cas9 protein (GeneArt™ Platinum™ Cas9 Nuclease, Thermo Fisher Scientific, Shanghai, China) and 300 ng/μL of sgRNA were injected into each egg using a FemtoJet and InjectMan NI 2 microinjection system (Eppendorf, Hamburg, Germany). After injection, the embryos were incubated in an incubator as described above. The hatched larvae were fed an artificial diet without any toxin.

#### *4.5. Generation of OfABCC2 Mutation Mediated by CRISPR*/*Cas9*

After embryo injection, the genomic DNAs of exuviate from 90 5th-instar larva were isolated individually using an AxyPrep Multisource Genomic DNA Miniprep Kit (Axygen, Hangzhou, China) following the manufacturer's instruction. To identify the indel mutations at predicted exon 4 of *OfABCC2*, the intron 4 sequences was first amplified by a specific pair of primers (4Ex\_F and 5Ex\_R, Table 4) and then by using the primers of 4Ex\_F and 4In\_R (Table 4) to amplify a 280-bp region flanking the desired cleavage site. The second PCR reaction mixture contained 1 μL of template, 1 μL of each of the 4Ex\_F or 4In\_R primer, 12.5 μL of 2× Taq Master Mix (TaKaRa, Dalian, China), and 9.5 μL of PCR-grade water in a final volume of 25 μL. PCR was performed at 95 °C 3 min, 35 cycles (95 °C 30 s, 55°C 30 s, 72 °C 1 min), 72 °C for 10 min, and 4 °C forever, and then the PCR products were directly sequenced with 4Ex\_F (sequencing primer) by TSINGKE Biological Technology (Nanjing, China). Direct sequencing chromatograms of mutant *OfABCC2* have double peaks around the cutting site at G0 generation. To detect the detailed deletion information of G2 genomic DNAs, the 280-bp PCR products were subcloned into pTOPO-T vector (Aidlab Biotechnologies, Beijing, China) and sequenced by TSINGKE Biological Technology. The acquired 8-bp deletion in *OfABCC2* was reconfirmed by clone sequencing using genomic DNA and mRNA templates from the knockout strain OfC2-KO.

#### *4.6. Inheritance Model Determination and Genetic Association Analysis*

The sex of *O. furnacalis* was visually determined based on the bottom characters of the pupa. Male adults (30 moths) from the original strain NJ-S were mass crossed with virgin female adults (30 moths) of the knockout strain OfC2-KO and vice versa. The degree of dominance (*h*) was estimated using the formula: *h* = (S*rs* − S*ss*)/(S*rr* − S*ss*), where S*rs*, S*ss*, and S*rr* are the survival rate for F1 hybrids, the NJ-S strain, and the OfC2-KO strains, respectively. The *h* varies from 0 for completely recessive resistance to 1 for completely dominant resistance [47].

For genetic association analysis between the 8-bp deletion of *OfABCC2* and Cry1Fa resistance phenotype, the F1 progeny from the reciprocal crosses were pooled and mass crossed to produce F2 progeny (Figure 3a). A total of 168 newly hatched larvae of the F2 progeny were treated with 2 μg/g of Cry1Fa toxin. The survivors (F2-treated) were collected after 7 days of treatment. The DNA from random selected parents (NJ-S and OfC2-KO), F2-treated survivors, and F2-untreated individuals were extracted for *OfABCC2* genotyping.

**Author Contributions:** Conceptualization, Y.W. and Y.Y.; methodology, X.W., Y.Y. and Y.W.; Investigation, X.W., Y.X., J.H. and W.J.; visualization, X.W., Y.X. and Y.Y. Funding acquisition, Y.W.; writing-original draft preparation, X.W.; writing-review and editing, Y.W. All authors have read and agreed to the published version of the manuscript.

**Funding:** This work was supported by grants from National Science and Technology Major Project of China (2019ZX08012004-005) and Fundamental Research Funds for the Central Universities of China (KYT201803).

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**


© 2020 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).

*Article*

## **ATP-Binding Cassette Subfamily a Member 2 Is a Functional Receptor for** *Bacillus thuringiensis* **Cry2A Toxins in** *Bombyx mori***, But Not for Cry1A, Cry1C, Cry1D, Cry1F, or Cry9A Toxins**

**Xiaoyi Li 1, Kazuhisa Miyamoto 2, Yoko Takasu 2, Sanae Wada 2, Tetsuya Iizuka 2, Satomi Adegawa 1, Ryoichi Sato 1,\* and Kenji Watanabe 2,\***


Received: 29 December 2019; Accepted: 3 February 2020; Published: 6 February 2020

**Abstract:** Cry toxins are insecticidal proteins produced by *Bacillus thuringiensis* (Bt). They are used commercially to control insect pests since they are very active in specific insects and are harmless to the environment and human health. The gene encoding ATP-binding cassette subfamily A member 2 (ABCA2) was identified in an analysis of Cry2A toxin resistance genes. However, we do not have direct evidence for the role of ABCA2 for Cry2A toxins or why Cry2A toxin resistance does not cross to other Cry toxins. Therefore, we performed two experiments. First, we edited the *ABCA2* sequence in *Bombyx mori* using transcription activator-like effector-nucleases (TALENs) and confirmed the susceptibility-determining ability in a diet overlay bioassay. Strains with C-terminal half-deleted BmABCA2 showed strong and specific resistance to Cry2A toxins; even strains carrying a deletion of 1 to 3 amino acids showed resistance. However, the C-terminal half-deleted strains did not show cross-resistance to other toxins. Second, we conducted a cell swelling assay and confirmed the specific ability of BmABCA2 to Cry2A toxins in HEK239T cells. Those demonstrated that BmABCA2 is a functional receptor for Cry2A toxins and that BmABCA2 deficiency-dependent Cry2A resistance does not confer cross-resistance to Cry1A, Cry1Ca, Cry1Da, Cry1Fa or Cry9Aa toxins.

**Keywords:** Cry2Ab toxin; *Bombyx mori*; ATP-binding cassette subfamily a member 2 (ABCA2); genome editing; transcription activator-like effector-nucleases (TALENs); HEK293T cell; functional receptor

**Key Contribution:** To know Cry2A-susceptibility determining role of BmABCA2, *B. mori* strains were created by genome editing using TALEN. Larvae of genome edited strains showed high level resistance to both Cry2Aa and Cry2Ab but not to Cry1A, Cry1Ca, Cry1Da, Cry1Fa, and Cry9Aa, indicating that BmABCA2 is a susceptibility determinant specific to Cry2A toxins. The heterologous expressing of BmABCA2 in HEK293T cell showed high susceptibility to Cry2Ab toxin, but not to Cry1A and Cry9Aa toxins, suggesting that BmABCA2 is a functional receptor specific to Cry2A toxins.

#### **1. Introduction**

Cry toxins are insecticidal crystal proteins and pore-forming toxins produced by *Bacillus thuringiensis* (Bt) [1,2]. Proteases activate Cry toxins in the midgut of the host insect; the toxins

then interact with specific receptors on the columnar cell membrane [3]. This interaction drives the toxins to insert partial structures into the membrane, forming ion channels [4]. A cation influx triggers the influx of water [5], resulting in cell swelling and lysis [2,6]. Given their strong toxicity in specific species and inability to harm the environment and human health, Cry toxins are used widely in pest control [7]. As a gene source, Bt toxin genes have been used efficiently to make transgenic crops (Bt crops) that resist pests [8]. However, resistant insect strains have been found in these crops [9].

To delay the selection and evolution of resistance in exposed insect populations, current commercial insecticidal systems combine two or more Cry toxins that bind to different receptors in the target pests [10]. Nevertheless, in traditional insecticidal systems that use a single Cry toxin, the generation of resistant insects and cross-resistance to other Cry toxins are still problems, reducing the value of commercial Bt crops. Regarding cross-resistance, in *Ostrinia nubilalis* and *Spodoptera frugiperda*, Cry1Ab, Cry1Ac, and Cry1Fa compete for the same binding sites with high affinity; however, Cry2Ab does not compete for the binding sites of Cry1 proteins [10]. Heterologous competition binding assays in *Helicoverpa armigera* and *Helicoverpa zea* midguts showed a common binding site for three Cry2A toxins (Cry2Aa, Cry2Ab, and Cry2Ae), but this binding site was not shared with Cry1Ac [11]. Cry 1Ab-resistant *Pectinophora gossypiella* was also reported to have strong cross-resistance to Cry1Aa, but little or no cross-resistance to Cry1Ca, Cry1Da, Cry2Aa, and Cry9Aa [12]. In the diamondback moth (*Plutella xylostella*), the Cry1C-resistant strain had strong cross-resistance to Cry1Ab, Cry1Ac, and Cry1F, but low cross-resistance to Cry1Aa and no cross-resistance to Cry2Aa [13]. Therefore, it is necessary to clarify the reason for cross-resistance or the receptors used by each Cry toxin to devise new strategies to defeat cross-resistance.

ATP-binding cassette (ABC) transporters, a class of transmembrane proteins, are found widely in organisms and are involved in Bt toxin activities [14]. Many studies seeking to identify functional receptors of target Cry toxins have focused on ABC transporters [15]. When BmABCC2 was expressed in *S. frugiperda* (Sf9) cells and *Drosophila* tissues that were originally insensitive, they became sensitive to Cry1 toxins, and when BmABCC2 was expressed in *Xenopus* oocytes the Cry toxins made pores in the membrane and cations flowed into the cells through these pores [16–18]. These results indicate that BmABCC2 is a receptor for Cry1A toxins. Using heterologous expression in Sf9 cells, ABCB1 was found to be a functional Cry3Aa receptor [19]. In comparison, an *H*. *armigera* strain with 6000-fold resistance to Cry2Ab [15] had a mutation in ABC subfamily A member 2 (HaABCA2), suggesting that ABCA2 is linked to Cry2Ab resistance [20]. This was confirmed using CRISPR/Cas9-mediated genome editing, leading to the conclusion that HaABCA2 determines the susceptibility of *H*. *armigera* to Cry2Aa and Cry2Ab [21]. The knockout of ABCA2 using CRISPR/Cas9 conferred resistance to Cry2Ab on *Trichoplusia ni* [22]. Furthermore, in Cry2Ab-resistant *P*. *gossypiella* strains, PgABCA2 was disrupted in several ways, indicating that the Cry2Ab-resistance of *P*. *gossypiella* is associated with an ABCA2 deficiency [23]. Therefore, ABCA2s seem to cause Cry2 resistance in most insects. In addition, since ABCC2 and ABCB1 function as receptors for Cry1A and Cry3A toxins, respectively, ABCA2s likely function as receptors for Cry2A toxins. However, to confirm whether ABCAs are really functional receptors for Cry2 toxins, it is necessary to show the receptor function of ABCAs using a heterologous expression system.

In this study, we used transcription activator-like effector nucleases (TALEN) to edit the genome at the *BmABCA2* locus and created mutant strains of *Bombyx mori* as a model system to clarify susceptibility determining activity. Then, we performed Cry2A toxin-contaminated leaf disc feeding assays using these mutant strains to determine whether BmABCA2 is really a susceptibility determinant for Cry2A toxins in *B*. *mori*. Using HEK293T cells expressing BmABCA2, we conducted cell swelling assays to demonstrate the Cry2A toxin-specific receptor function of BmABCA2.

#### **2. Results**

#### *2.1. Creation of Silkworm Strains with C-Terminal Half-Deleted BmABCA2s and Mutants with C-Terminal Deletions in TM7 by TALEN-Mediated Mutagenesis*

First, 200 eggs were injected individually with TALEN mRNAs and a donor oligonucleotide prepared to mutate the C-terminus of transmembrane domain 7 (TM7) in exon 15 (Figure 1A,B). The 14 neonates that hatched were allowed to develop (G0). Two female moths survived and were crossed singly with the wild-type strain to produce the next generation (G1). The G1 larvae were allowed to develop into adults. Genomic DNA samples of individual G1 moths were prepared, and insertion/deletion mutations at TM7 in exon 15 of *BmABCA2* were identified. One mutant allele (named A2T01) was presumably derived from homology-directed repair (HDR), and 12 mutant alleles (named A2T03–A2T14) obtained via non-homologous end joining (NHEJ) were detected (Figure 1C). The mutations in A2T01, A2T05, A2T06, A2T09, A2T10, A2T12, A2T13, and A2T14 led to truncation of the C-terminus of TM7 in BmABCA2. The mutations in A2T03, A2T04, A2T07, A2T08, and A2T11 were predicted to cause 3, 4, 1, 1, and 2 amino-acid deletions from the C-terminus of TM7, respectively.

**Figure 1.** TALEN-induced mutations generated at the C-terminus of TM7 in BmABCA2. (**A**) Schematic structure of BmABCA2 and the mutation sites created with the TALEN system in the wild-type (Ringetsu) *B. mori* strain. The transmembrane topology of BmABCA2 was predicted by Phobius (http://phobius.sbc.su.se/). The approximate position of the mutation is indicated by the red rectangle located at the C-terminus of TM7 in BmABCA2. (**B**) The TALEN-binding site to TM7 in exon 15 used to introduce an insertion/deletion mutation in BmABCA2 and donor oligonucleotides for HDR. (**C**) The detected G1 mutant alleles of the mutant strains. A2T01 was obtained as a result of HDR, while A2T03–A2T14 were obtained as the result of NHEJ.

To construct homozygous strains, heterozygous G1 strains were selected after confirming the PCR products with direct sequencing and were mated with individuals of the same genotype. The homozygous individuals of the next generation (G2) were screened by genotyping, resulting in the establishment of three strains with C-terminal half-deleted BmABCA2 (A2T01, A2T06, and A2T14) and three strains (A2T03, A2T08, and A2T11) with amino acids deleted from the C-terminus of TM7 (Figure S1).

#### *2.2. BmABCA2 Activity against Cry2A Toxins*

To test the susceptibility of the homozygous BmABCA2 mutant strains to Cry2Ab, larvae were reared on leaf disks contaminated with Cry2Ab for 2 days, and then on leaf disks without toxin for an additional 2 days. Larvae of all of the strains with C-terminal half-deleted BmABCA2s (A2T01, A2T06, and A2T14; Figure 1) showed strong resistance to Cry2Ab (Figure 2A). The median lethal dose (LC50) of Cry2Ab on A2T14 was >9990-fold higher than that of the wild-type strain (Table 1). Surprisingly, the larvae of strains with BmABCA2s in which 1–3 amino acids had been deleted from the C-terminus of TM7 (A2T03, A2T11, and A2T08; Figure S1), were also resistant to Cry2Ab (Figure 2B,C). However, in association with the Cry2Ab concentration, those strains were slightly susceptible to Cry2Ab, i.e., the resistance of those strains was not as high as that of strains with C-terminal half-deleted BmABCA2s.

**Figure 2.** Toxin feeding assay to evaluate the susceptibility of the larvae with mutations in BmABCA2 to Cry2Ab. (**A**) Evaluation of strains with C-terminal half-deleted BmABCA2s (A2T01, A2T06, and A2T14; indicated in Figure 1C. The wild-type strain (Ringetsu), which is susceptible to Cry2A toxins, was used as a control. (**B**,**C**) The susceptibility of strains with BmABCA2s carrying 1–3 amino acid deletions at the C-terminus of TM7 [A2T03, A2T11 (**B**), and A2T08 (**C**); indicated in Figure 1A,C] to Cry2Ab. The wild-type strain was used as a control. Cry2Ab was spread on the leaf disk at 10 μL/cm2. The toxin feeding assays were performed as described in the Materials and Methods.


**Table 1.** Responses of the knock-out (A2T14) and wild-type strains to Cry toxins.

<sup>1</sup> Number of larvae tested. <sup>2</sup> Concentration of toxins killing 50% of larvae and its 95% confidence interval (CI). <sup>3</sup> Slope of the concentration-mortality line. <sup>4</sup> Resistance ratio (RR) <sup>=</sup> LC50 of knock-out strain divided by LC50 of the same toxin for wild-type.

The susceptibility of *B*. *mori* strains carrying mutant BmABCA2 to Cry2Aa was also investigated. All of the strains with C-terminal half-deleted BmABCA2s (A2T01 and A2T14) and the strains with BmABCA2s in which 1 or 2 amino acids were deleted from the C-terminus of TM7 (A2T08 and A2T11) were resistant to Cry2Aa (Figure 3). With increasing Cry2Aa concentrations, A2T08 showed slight susceptibility to Cry2Aa (Figure 3B).

**Figure 3.** Toxin feeding assay to evaluate the susceptibility of larvae with mutations in BmABCA2 to Cry2Aa. Strains with C-terminal half-deleted BmABCA2s [A2T01 (**A**) and A2T14 (**C**)] and strains with BmABCA2s carrying 1 or 2 amino acid deletions at the C-terminal end of TM7 [see Figure 1A,C; A2T08 (**B**) and A2T11 (**A**)] were evaluated using the toxin feeding assay as described in Figure 2 and the Materials and Methods. The wild-type strain (Ringetsu) was used as a control.

The susceptibility of a *B*. *mori* strain with C-terminal half-deleted BmABCA2s (A2T14) to Cry1A, Cry1Ca, Cry1Da, Cry1Fa, and Cry9Aa was investigated further. We found that the susceptibility of strain A2T14 to Cry1Aa, Cry1Ab, Cry1Ac, Cry1Ca, Cry1Da, Cry1Fa, and Cry9Aa was similar to that of the wild-type strain (Figure 4). Moreover, the LC50 with the 95% confidence interval of each toxin was calculated (Table 1). The LC50 of every toxin did not differ between the wild-type and A2T14 strains, but A2T14 strains had no or very limited resistance to Cry1Aa (<2-fold), Cry1Ac (<5-fold) and Cry1Fa (<3-fold), indicating that the knockout of BmABCA2 did not affect susceptibility to Cry1A, Cry1Ca, Cry1Da, Cry1Fa, or Cry9Aa.

**Figure 4.** Toxin feeding assay Cry toxins. The strain with C-terminal half-deleted BmABCA2 (A2T14) and the wild-type strain (Ringetsu) were fed leaf disks contaminated with Cry1Aa (**A**), Cry1Ab (**B**), Cry1Ac (**C**), Cry1Ca (**D**), Cry1Da (**E**), Cry1Fa (**F**), or Cry9Aa (**G**) as described in the Materials and Methods.

#### *2.3. BmABCA2-Dependent Cry2A Toxins Induce Cell Swelling*

To examine whether BmABCA2 is a functional Cry2A toxin receptor, we used cell swelling assays. Enhanced green fluorescent protein (EGFP) cDNA was equipped with BmABCA2 cDNA in the same vector and transiently expressed in HEK293T cells showing green fluorescence on transfection. Cry2Ab was administered to those transient expression cells. Only the EGFP-positive cells swelled when they were treated with more than 40 nM Cry2Ab (Figure 5). The cells swelled in a Cry2Ab toxin concentration-dependent manner (Figure 5). By contrast, even 1.1 μM Cry2Ab did not induce swelling in cells not transfected with the BmABCA2 expression vector (Figure 5). Furthermore, cells that were transfected with the BmABCA2 expression vector did not swell when they were incubated in buffer lacking Cry2Ab.

**Figure 5.** Cell swelling assay against Cry2Ab using BmABCA2-expressing HEK292T cells. The cells were attached to coverslips set on the six plates. BmABCA2 was transiently expressed on the surface of HEK293T cells via the transfection of an expression vector with attached EGFP. The coverslips were set on the wells of glass slides filled with the Cry2Ab test solutions, incubated for 1 h, and observed under a microscope. Arrowheads indicate swollen cells. (Scale bar = 50 μm).

To examine whether BmABCA2 acts as a functional receptor for other Cry toxins, BmABCA2-expressing cells were treated with Cry1Aa, Cry1Ac, and Cry9Aa. However, no cells were swollen after treatment with up to 1.5 μM Cry1Aa, 1.1 μM Cry1Ac, and 3.3 μM Cry9Aa (Figure 6).

To clarify whether the BmABCC2 receptor for Cry1A toxins can function as a Cry2Ab receptor, BmABCC2-expressing cells were administered Cry2Ab. However, no cells swelled with up to 1.1 μM Cry2Ab (Figure 7). By contrast, Cry 1Aa induced swelling in BmABCC2-expressing cells at 15 nM, indicating that the level of BmABCC2 expression was sufficient to assess Cry2Ab receptor function (Figure 7). Cry1Ac induced swelling of the BmABCC2-expressing cells at 500 nM (Figure 7).

**Figure 6.** Cell swelling assay against Cry1A toxins and Cry9Aa using BmABCA2-expressing HEK293T cells. The cells were attached to coverslips set on the six plates. BmABCA2 was transiently expressed on the surface of HEK293T cells via the transfection of an expression vector with attached EGFP. Then, the cells were incubated for 1 h with Cry toxins on glass slides and observed under a microscope, as described in Figure 5. (Scale bar = 50 μm).

**Figure 7.** Cell swelling assay against Cry1A toxins and Cry2Ab using BmABCC2-expressing HEK293T cells. The cells were attached to coverslips set on the six plates. BmABCC2 co-expressed with EGFP was transiently expressed on the surface of HEK293T cells via transfection of the expression vector. Then, the cells were incubated for 1 h with Cry toxins on glass slides and observed under a microscope, as described in Figure 5. Arrowheads indicate swollen cells. (Scale bar = 50 μm).

#### **3. Discussion**

Toxicity tests of the strains with C-terminal half-deleted BmABCA2 (A2T01, A2T06, and A2T14) showed that they were highly resistant to Cry2Ab (Figure 2), indicating that BmABCA2 plays an essential role in determining the susceptibility of *B*. *mori* to Cry2Ab. ABCA2 was first suggested to be linked to Cry2Ab resistance in *H*. *armigera* [20]. This was confirmed by generating an ABCA2 knockout strain via CRISPR/Cas9 mutagenesis in which the resistance of *H*. *armigera* and *T*. *ni* larvae to Cry2Ab was linked to defects in ABCA2 [21,22], indicating that ABCA2 plays an important role in the mode of action of Cry2A toxins. Although the resistance was slightly lower than that of the C-terminal half-deleted strains (Figure 2B,C), toxicity testing of the strains with BmABCA2s that had 1–3 amino acids deleted from the C-terminal end of TM7 (A2T03, A2T08, and A2T11) showed that they were also resistant to Cry2Ab (Figure 2B,C). Thus, BmABCA2 also plays an essential role in determining the susceptibility to Cry2Ab in *B*. *mori*. In *B*. *mori*, even the partial deletion and alanine replacement of several amino acid residues at the N-terminus of extracellular loop 4 (ECL4) decreased the susceptibility-conferring activity of BmABCC2 to Cry1Aa [17]. This suggests that ECL4 of BmABCC2 is part of the site that interacts with Cry1Aa. ECL4 of BmABCA2 might also play a role in the interaction with Cry2Ab (Figure 1A), and the 1–3 amino acids that were deleted from the C-terminal end of TM7 might affect the interaction of BmABCA2 with Cry2Ab by changing the three-dimensional structure of the binding epitopes.

It is still not known how ABCA2 is involved in the susceptibility of larvae to Cry2A toxins. ABCA2 is a membrane protein and the ABC transporter ABCC2 is a functional Cry1A toxin receptor [5,16,18,24,25]. Therefore, we examined whether BmABCA2 transiently expressed in HEK293T cells (Figure 5) could function as a Cry2Ab receptor and whether HEK293T cells would swell like the columnar cells in a Cry toxin-intoxicated midgut. The BmABCA2-expressing cells started swelling on exposure to 40 nM Cry2Ab (Figure 5). When BmABCC2, a highly functional receptor for a single Cry1Aa molecule [16], was expressed in HEK293T cells using a very similar expression system, the HEK293T cells started swelling in response to 1 nM Cry1Aa [26]. By contrast, when BmABCC3, a less functional receptor for a single Cry1Aa molecule, was expressed in HEK293T cells, the cells started to swell in response to 100 nM Cry1Aa [26]. Based on our cell swelling assays using HEK293T cells, BmABCA2 has sufficient functional receptor activity for Cry2Ab. This is consistent with the strong resistance to Cry2Ab that was generated by the TALEN-induced BmABCA2 mutation (Figure 2).

In *H*. *armigera*, an HaABCA2 knockout strain showed resistance to Cry2Ab, but not to Cry1Ac [21], suggesting that this receptor is highly tuned to Cry2A toxins. Regarding Cry1Aa, the BmABCC2 binding site was thought to be the pocket made by loops 2 and 3 [27]. However, we could not find any amino acids near loops 2 and 3 that were conserved in Cry1Aa and Cry2Ab. This might explain the receptor specificity difference between Cry1Aa and Cry2Ab. The BmABCA2 knockout strains were susceptible to Cry1Aa (Figure 4). In addition, the BmABCA2 knockout strains were susceptible to all of the Cry1 and Cry9 toxins tested (Figure 4). This implies that Cry2A toxin-resistant insects lack cross-resistance to Cry1Ca, Cry1Da, Cry1Fa, or Cry9Aa. Actually, a Cry1Ca-resistant diamondback moth was reported to lack cross-resistance to Cry2Aa [13]. Furthermore, BmABCA2-expressing HEK293T cells were susceptible to Cry2Ab, but not to Cry1A or Cry9A toxins (Figures 5 and 6). Therefore, our results suggest that the specificity of BmABCA2 as a Cry toxin receptor is narrowly tuned to Cry2A toxins. By contrast, BmABCC2-expressing HEK293T cells were susceptible to Cry1A toxins, but not to Cry2Ab (Figure 7). This also suggests that ABC transporters are highly tuned to a narrow group of Cry toxins.

#### **4. Materials and Methods**

#### *4.1. Silkworm Strains and Rearing*

The wild-type silkworm strain was distributed from the Genetics Resources Center, National Agriculture and Food Research Organization (NARO) and was reared on mulberry leaves or artificial diet (Nihon Nosan Kogyo, Yokohama, Japan) at 25 ◦C.

#### *4.2. DNA Target Site Selection and Preparation of TALEN mRNA*

The DNA target site was selected in the fifteenth exon of the BmABCA2 (KP219767) gene that encodes the extracellular region between the 7th and 8th transmembrane regions. Two TALEN half sites were designed, as shown in Figure 1. TALEN-encoding genes were constructed by Golden Gate assembly, as described previously [28]. To prepare mRNAs for microinjection, TALEN-encoding plasmids were linearized by *Xba*I (TaKaRa Bio, Kusatsu, Japan) and transcribed using an mMESSAGE mMACHINE T7 transcription kit (Thermo Fisher Scientific, Waltham, MA, USA) according to the protocols of manufacturer.

#### *4.3. Egg Microinjection*

The poly(A)-tailed TALEN mRNAs (0.2 μg/μL) were dissolved in injection buffer (5 mM KCl, 0.5 mM phosphate buffer, pH 7.0) together with donor oligonucleotides (0.2–0.4 μg/μL), as described previously [29], and injected to the silkworm eggs at the syncytial preblastodermal stage [30]. The embryos were incubated at 25 ◦C in a humidified atmosphere.

#### *4.4. Identification of BmABCA2 Mutation Induced by TALENs*

To extract genomic DNA, a leg of each G1 moth was homogenized in 50 μL of DNAzol® Direct (Molecular Research Center, Cincinnati, OH, USA). After 10 min of incubation at room temperature, the homogenate was mixed vigorously and separated by centrifugation. The supernatant containing genomic DNA was used as a template for PCR. The target region of the BmABCA2 gene was amplified using a specific primer set (forward: 5- -GTGTCAGGAGCAAGTCTGGTC-3- , reverse: 5- -AGACGTGTTAAATATCTCGTCTCG-3- ). Direct sequencing of the PCR products was performed using the reverse primer as a sequencing primer. Mutations induced by TALENs were identified according to the sequencing results.

#### *4.5. Cry Toxins Preparation*

The DNAs of Cry2Ab (AAA22342), and Cry1Fa (AAA22348) genes which were optimized for expression of heterologous proteins in *Escherichia coli* was synthesized by Strings DNA Fragments service (Thermo Fisher Scientific) and subcloned into between *Bam*HI and *Xho*I sites of pGEX6P-3 (GE healthcare lifesciences, Amersham, UK) using In-fusion HD Cloning kit (TaKaRa Bio, Kusatsu, Japan). The DNA clones were used to produce the Cry2Ab, and Cry1Fa toxins. For the production of the Cry1Aa, Cry1Ab, Cry1Ac, Cry1Da, and Cry9Aa toxins, the genes of these toxins were subcloned into pGEX4T-3 and then *E. coli* cells were transformed with those as described previously [16]. The transformed cells were cultured in LB liquid medium with ampicillin at 37 ◦C and gene expression was induced by isopropyl thio-b-d-galactoside. The inclusion bodies were harvested and washed as describe as previously [31]. Inclusion bodies of Cry2Aa toxin was prepared as described elsewhere [32]. The Cry1Ca toxin was produced by a *B. thuringiensis* recombinant stain, and inclusion bodies of Cry1Ca were washed as described elsewhere [33]. The Cry1Aa, Cry1Ac, and Cry9Aa were solubilized and activated as described previously [27]. The inclusion bodies of Cry2Ab was activated with a High-Performance Liquid Chromatography method (HPLC) that the Cry2Ab was solubilized as same as described above. After the solution, the pro-toxin of Cry2Ab was dialyzed with 20 mM Tris-HCl pH 9 and passed through a 0.45 μM filter (Millipore Millex-HP Hydrophilic PVDF, Millipore,

Burlington, MA, USA) to remove bacteria. Then, the pro-toxin was applied to an HPLC system equipped with a Shodex IEC DEAE-825 column (0.8 × 7–5cm, Showa Denko Co.) and equilibrated with 250 mM Tris-HCl, pH 9. The pro-toxin was bound with the column, and non-absorbent pellets were washing by 100 mM Tris-HCl pH 9. Then, the bound pro-toxin was treated with 0.0625 mg/mL Trypsin (Sigma-Aldrich, St. Louis, MO, USA) for 40 min at 37 ◦C. The treatment Cry2Ab was eluted by Elix water 5 min later, with a linear gradient of 0~250 mM Tris-HCl pH 9 buffer eluted over 55 min with a 0.5 mL/min flow rate, and the activated toxin was dialyzed by phosphate-buffered saline (PBS, 137 mM sodium chloride, 2.7 mM potassium chloride, 10 mM sodium phosphate dibasic, 1.8 mM potassium dihydrogen phosphate, pH 7.4) for cell swelling assay.

#### *4.6. Diet Overlay Bioassays*

Susceptibility to Cry toxins of the BmABCA2 genome edited mutants and wild-type strains were evaluated with diet overlay bioassays. To make leaf disks, sixth open leaves from the top of each branch of the mulberry leaves were picked up. Cry toxins solutions were diluted with Silwet® L-77 (Momentive Performance Materials, Waterford, NY, USA), and the suspensions were spread to be 10 μL/cm2 on the leaf disks. After Cry toxins suspension were dried at room temperature, two leaf disks were put in each petri dish with 10 larvae of 2nd instar of the genome edited, and wild-type strains were respectively reared on each disk for 2 days. After 2 days, the larvae were moved to non-toxins leaf disks and mortality was recorded at 4 days after feeding initiation.

The median lethal dose (LC50) values and the 95% confidence interval were calculated based on Probit Analysis.

#### *4.7. cDNA Cloning of BmABCA2 and Construction of Expression Vectors for HEK293T Cells*

Total RNA was isolated from midgut tissue of 5th instar larvae of the wild-type silkworm strain, using TRIzol (Thermo Fisher Scientific, Waltham, MA, USA) according to the protocols of manufacturer, and used for cDNA synthesis as a template. The BmABCA2 cDNA was amplified by one-step RT-PCR using PrimeScript High Fidelity RT-PCR Kit (TaKaRa Bio). The amplified cDNA using primers (forward: 5- -CCACCCGGATCCGATATGAGACCTCAGAGAAAAGAAGCC-3- , reverse: 5- - GTCTTTGTAGTCGATCAAGCCTTCCCTTTGATATTTCGT-3- ) was cloned into *EcoR*V site of the pcDNA3.1 (Thermo Fisher Scientific, Waltham, MA, USA) using In-Fusion® HD Cloning Kit (TaKaRa Bio, Kusatsu, Japan). The neomycin resistance gene of pcDNA3.1 was replaced by Enhanced green fluorescent protein (EGFP) - *Streptoalloteichus hindustanus* ble (Sh ble) fusion gene by In-fusion cloning method described below. The linearized vector was generated from pcDNA3.1 plasmid as a template by PCR using primers (forward: 5- -GCCCTTGCTCACCATGCGAACGATCCTCATCCTGTC-3- , reverse: 5- - GAGGAGCAGGACTGAGCGGGACTCTGGGGTTCG-3- ). The EGFP-ble fusion gene was amplified using primers (forward: 5- -ATGGTGAGCAAGGGCGAGGAG-3- , reverse: 5- - TCAGTCCTGCTCCTCGGCCAC-3- ) and cloned into the linearized vector.

#### *4.8. Expression of BmABCA2 and BmABCC2 in HEK293T Cells and Cell Swelling Assay with Cry Toxins*

HEK293T cells were cultured and transfected, as described previously [5]. The HEK293T cells were cultured on the cover glass in a 6-well plate (Truesline; Nippon Genetics, Tokyo, Japan). Until the HEK293T cells grow up to 70 ~ 80% confluence, expression vectors for BmABCA2 and BmABCC2 were transfected in Opti-MEM® (Thermo Fisher Scientific, Waltham, MA, USA) with polyethylenimine (PEI Max, Polysciences, Warrington, PA, USA) for 2 h incubation at 37 ◦C. Then, the media were changed to a fresh Dulbecco's modified Eagle medium (D-MEM) and incubated at 37 ◦C for 48 h in a CO2 incubator. After that, the cover glass was taken out and covered onto Cry toxins solutions in a hole of the 2-hole glass slide. Cry toxins were diluted with PBS (pH 7.4). After 1 h incubation at 37 ◦C, the cells were observed by phase-contrast and fluorescent microscopy.

*Toxins* **2020**, *12*, 104

**Supplementary Materials:** The following are available online at http://www.mdpi.com/2072-6651/12/2/104/s1, Figure S1: Mutations in BmABCA2 in mutant strains produced by genome-editing method using TALENs.

**Author Contributions:** K.W. and R.S. conceived and designed the experiments; X.L., K.W., S.A., and K.M. performed the experiments and analyzed the data; Y.T., S.W. and T.I. helped to prepare the mutant silkworm strains used in the study; X.L., K.W. and R.S. wrote the manuscript. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research was partially supported by a Grant-in-Aid for Scientific Research (B) (18H03397) from the Ministry of Education, Culture, Sports, Science, and Technology of Japan.

**Acknowledgments:** We thank Ritsuko Murakami for rearing the silkworms and providing technical assistance. We also thank Shin-ichiro Asano for providing the *Bacillus thuringiensis* strain Bt51(pHS2). The silkworm strain "Ringetsu" was provided from The Genetic Resources Center, NARO. The work was funded by a Grant-in-Aid for Scientific Research (B) (18H03397) from the Ministry of Education, Culture, Sports, Science, and Technology of Japan.

**Conflicts of Interest:** We declare no conflict of interest, and the funders had no role in the design of the study.

#### **References**


© 2020 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).

*Article*

### **Reduced Expression of a Novel Midgut Trypsin Gene Involved in Protoxin Activation Correlates with Cry1Ac Resistance in a Laboratory-Selected Strain of** *Plutella xylostella* **(L.)**

**Lijun Gong 1,2, Shi Kang 2, Junlei Zhou 2, Dan Sun 2, Le Guo 1,2, Jianying Qin 2, Liuhong Zhu 2, Yang Bai 2, Fan Ye 1,2, Mazarin Akami 2, Qingjun Wu 2, Shaoli Wang 2, Baoyun Xu 2, Zhongxia Yang 1, Alejandra Bravo 3, Mario Soberón 3, Zhaojiang Guo 2,\*, Lizhang Wen 1,\* and Youjun Zhang 2,\***


Received: 7 January 2020; Accepted: 21 January 2020; Published: 23 January 2020

**Abstract:** *Bacillus thuringiensis* (Bt) produce diverse insecticidal proteins to kill insect pests. Nevertheless, evolution of resistance to Bt toxins hampers the sustainable use of this technology. Previously, we identified down-regulation of a trypsin-like serine protease gene *PxTryp\_SPc1* in the midgut transcriptome and RNA-Seq data of a laboratory-selected Cry1Ac-resistant *Plutella xylostella* strain, SZ-R. We show here that reduced *PxTryp\_SPc1* expression significantly reduced caseinolytic and trypsin protease activities affecting Cry1Ac protoxin activation, thereby conferring higher resistance to Cry1Ac protoxin than activated toxin in SZ-R strain. Herein, the full-length cDNA sequence of *PxTryp\_SPc1* gene was cloned, and we found that it was mainly expressed in midgut tissue in all larval instars. Subsequently, we confirmed that the *PxTryp\_SPc1* gene was significantly decreased in SZ-R larval midgut and was further reduced when selected with high dose of Cry1Ac protoxin. Moreover, down-regulation of the *PxTryp\_SPc1* gene was genetically linked to resistance to Cry1Ac in the SZ-R strain. Finally, RNAi-mediated silencing of *PxTryp\_SPc1* gene expression decreased larval susceptibility to Cry1Ac protoxin in the susceptible DBM1Ac-S strain, supporting that low expression of *PxTryp\_SPc1* gene is involved in Cry1Ac resistance in *P. xylostella*. These findings contribute to understanding the role of midgut proteases in the mechanisms underlying insect resistance to Bt toxins.

**Keywords:** *Bacillus thuringiensis*; *Plutella xylostella*; Cry1Ac resistance; trypsin-like midgut protease; protoxin activation

**Key Contribution:** Low expression of a novel midgut trypsin-like protease gene *PxTryp\_SPc1* involved in protoxin activation contributes to Cry1Ac resistance in a laboratory-selected *P. xylostella* strain.

#### **1. Introduction**

*Bacillus thuringiensis* (Bt) are gram-positive entomopathogenic bacteria most widely used as a biopesticide worldwide, and transgenic crops expressing insecticidal toxins produced by these bacteria (transgenic Bt crops) have been planted in 104 million hectares globally in 2018, which has a central role in pest control and global food security [1,2]. However, field-evolved resistance to Bt crops soared from three cases in 2005 to 16 in 2016, documenting an accelerated evolution of practical resistance [3]. Due to the commercial application of Bt proteins, such as Cry proteins for the control of insect pests, it is necessary to probe the resistance mechanism to Cry proteins in order to propose effective strategies to delay the resistance evolution.

Bt Cry proteins are produced as inactive and insoluble crystals formed by protoxins [4]. Cry protoxins are solubilized in the alkaline environment of the midgut and are further processed into activated toxins by midgut proteases when ingested by susceptible larvae [5]. Activated toxins then interact with specific midgut receptor proteins, such as aminopeptidase N (APN), alkaline phosphatase (ALP), cadherin (CAD) and ABC transporters, located in the brush border membrane (BBM) of the midgut epithelium cells from the larvae [6,7] Receptor binding leads to the formation of lytic pores in the membrane that burst cells and finally kill the insects [8]. However, it has been shown that protoxins also bind to specific receptors, and then they are activated by midgut proteases inducing also toxin oligomerization and pore-formation [8,9]. Whether protoxins are activated before or after receptor binding is an important step, transforming the 130 kDa protoxin into a 55–65 kDa activated toxin [4,7]. Trypsin proteases are important midgut proteinases, which participate in Bt Cry protein degradation and protoxin activation [10,11]. It has been reported that alteration of the midgut trypsin genes or trypsin proteolytic activities are linked to Bt resistance in *Plodia interpunctella* (Hübner) (Lepidoptera: Pyralidae) [10], *Ostrinia nubilalis* (Hübner) (Lepidoptera: Pyralidae) [12–14], *Spodoptera frugiperda* (JE Smith) (Lepidoptera: Noctuidae) [15], *Helicoverpa armigera* (Hübner) (Lepidoptera: Noctuidae) [16,17], *Aedes aegypti*(L.) (Diptera: Culicidae) [18], *Mythimna unipuncta* (Haworth) (Lepidoptera: Noctuidae) [19], and *Helicoverpa zea* (Boddie) (Lepidoptera: Noctuidae) [20].

The diamondback moth, *Plutella xylostella* (L.), is a cosmopolitan insect pest of cruciferous crops that was the first example of resistance to Bt sprays in the field [21]. The economic damage produced by *P. xylostella* was estimated to be up to USD 5 billion every year [22]. Moreover, since *P. xylostella* was the first documented insect developing field-evolved Bt resistance, it is a good model to understand insect resistance mechanisms to Bt toxins. Previous studies showed that in *P. xylostella*, resistance to the Cry1Ac toxin was not associated with alterations of the *PxABCH1* and *PxCAD* genes [23–25]. In contrast, Cry1Ac resistance rather correlated with a mutation in the *PxABCC2* gene [26] or with the differential expression of the *PxmALP*, *PxABCB1*, *PxABCC1*, *PxABCC2*, *PxABCC3,* and *PxABCG1* genes, which were shown to be down-regulated by an enhanced MAPK signaling pathway [27–29]. In the MAPK-mediated *trans*-regulatory mechanism, we reported that over-expression of the *PxMAP4K4* gene resulted in down-regulation of diverse midgut genes, thereby conferring a Cry1Ac resistance phenotype. Different *P. xylostella* strains that are resistant to Cry1Ac showed different induction levels of *PxMAP4K4*, resulting in different resistance levels. For example, the SZ-R resistant strain showed moderate resistance levels (662-fold to Cry1Ac) in contrast with the near-isogenic strain (NIL-R) that was highly resistant to Cry1Ac (>3900-fold). The relative expression of *PxMAP4K4* in SZ-R is slightly higher than that of the susceptible DBM1Ac-S strain; thus, expression of some Cry toxin receptors are also down-regulated in the SZ-R strain although at a lower level than that of the NIL-R, which showed the highest constitutive expression of the *PxMAP4K4* gene [27]. In particular, the high Cry1Ac resistance levels in the NIL-R strain does not involve the Cry1Ac protoxin activation mechanism [30]. However, the relationship between the protoxin activation mechanisms in other resistant strains of *P. xylostella* remains unclear.

Here, we compared data from midgut transcriptome and RNA-Seq analyses that were previously done, showing a significant decrease expression of a novel trypsin-like protease in a *P. xylostella* strain SZ-R that shows resistance to Cry1Ac toxin [31,32]. Then, we cloned and characterized the midgut

trypsin protease gene of *P. xylostella* (*PxTryp\_SPc1*). Finally, we demonstrated that down-regulation of the *PxTryp\_SPc1* gene in the midgut tissue of SZ-R strain is related to Cry1Ac resistance by using different genomic, molecular, biochemical, and genetic tools. The conclusions of this study provide a new insight into the Bt resistance mechanism that could give hints for the control of insect pests.

#### **2. Results**

#### *2.1. Comparison of Midgut Protease Activities and Cry1Ac Protoxin Activation between Susceptible DBM1Ac-S and Resistant SZ-R Strains*

Previously, differential expression of midgut trypsin-like serine protease (*Tryp\_SP*) genes was identified in the Cry1Ac-resistant strain SZ-R in contrast to the susceptible *P. xylostella* strain DBM1Ac-S [31,32]. To determine whether the potential altered *PxTryp\_SP* gene expression can change the midgut protease activities and affect Cry1Ac protoxin activation in SZ-R strain, we first compared the midgut protease activities and Cry1Ac protoxin activation in both susceptible DBM1Ac-S and resistant SZ-R strains. The resistant SZ-R strain showed significantly lower caseinolytic protease activity in the midgut extracts than the susceptible DBM1Ac-S larvae (*p* < 0.05; Duncan's tests; n = 3), likewise, the trypsin activity of resistant SZ-R was also significantly lower than the susceptible DBM1Ac-S larvae, but the chymotrypsin activity was similar between these two *P. xylostella* strains (*p* < 0.05; Duncan's tests; n = 3) (Figure 1A). Subsequently, the incubation of Cry1Ac protoxin with midgut protease extracts from susceptible DBM1Ac-S or resistant SZ-R larvae were compared (Figure 1B). After 1-h incubation, a strong single band of about 65 kDa was produced in DBM1Ac-S gut extracts, corresponding to the processed Cry1Ac protein, having a similar band size as produced by the control treatment with bovine trypsin. In contrast, two bands were observed in the activation produced by SZ-R gut extracts: one similar to that produced by the control bovine trypsin or DBM1Ac-S and the other of higher molecular weight. These results confirmed that the potential altered *PxTryp\_SP* gene expression might be involved in Bt Cry1Ac resistance in SZ-R strain via decreasing midgut protease activities and protoxin activation.

**Figure 1.** Midgut protease activities (caseinolytic proteases, trypsin, and chymotrypsin) and the activation differences between the *P. xylostella* DBM1Ac-S and SZ-R strains. (**A**). Protease activities were calculated relative to the activities shown by the susceptible DBM1Ac-S strain (100%). Different letters stand for statistically significant differences within the three replicates and four technical repeats (*p* < 0.05; Duncan's test; n = 3). (**B**) Activation of Cry1Ac protoxin with protease midgut extracts from control (DBM1Ac-S) or resistant (SZ-R) strains. Lane 1: protein maker; Lane 2: Cry1Ac protoxin; Lane 3: Cry1Ac incubated with bovine trypsin (positive control); Lane 4: Cry1Ac incubated with protease midgut extracts from DBM1Ac-S strain; Lane 5: Cry1Ac incubated with midgut extracts from SZ-R strain.

#### *2.2. Bioassay Analyses of Cry1Ac Protoxin and Activated Toxin*

To further validate the influence of altered *PxTryp\_SP* gene expression on the resistance to Cry1Ac toxin in SZ-R strain, bioassays with Cry1Ac (protoxin and activated toxin) were further conducted. Bioassays revealed that the resistance levels of Cry1Ac protoxin or activated toxin by trypsin were different in the SZ-R strain (Table 1). The resistance ratios (RR) were 662 for Cry1Ac protoxin and 422 for the activated Cry1Ac toxin in the SZ-R strain. The LC50 values of activated toxin and protoxin showed slightly but significant differences since the LC50 value of activated toxin was less than two fold lower to that of Cry1Ac protoxin (Table 1). These data correlated with the partial activation of Cry1Ac protoxin as shown above (Figure 1B). Furthermore, to verify whether Cry1Ac protoxin is about half as potent as the activated toxin due to their different molecular weights (130 vs. 65 kDa), we estimated the potency of Cry1Ac protoxin relative to activated toxin as reported before [33]. The potency ratios (PR) were 0.83 for the DBM1Ac-S strain and 0.53 for the SZ-R strain, which did not differ significantly from the predicted value of 0.50 (one sample *t*-test, df = 1, t = 3.25, *p* = 0.19), implying that the Cry1Ac protoxin was no more effective than the activated toxin in both strains analyzed (Table 1).

**Table 1.** Bioassays of Cry1Ac protoxin and activated toxin in DBM1Ac-S and resistant SZ-R larvae.


<sup>a</sup> Slope of the dose response-mortality. SEM stands for standard error of the mean. <sup>b</sup> LC50 (95% FL): Toxin concentration (mg/L) killing 50% of larvae and its 95% fiducial limits (lower-upper). <sup>c</sup> RR: Resistance ratio calculated by the ratio between the LC50 value of SZ-R by the LC50 of DBM1Ac-S. <sup>d</sup> PR: Potency was calculated as the ratio of LC50 value of activated toxin by the LC50 of protoxin as reported [33,34]. Potency values < 1 indicate the activated Cry1Ac toxin is more potent than protoxin, while potency values > 1 indicate that Cry1Ac protoxin is more potent than activated toxin. \* Asterisks represent significantly different LC50 value by the conservative criterion of non-overlapped 95% CL value.

#### *2.3. Cloning, Characterization, and Phylogenetic Analyses of the PxTryp\_SPc1 Gene*

During the previous characterization of differentially altered genes in SZ-R, we identified that a *PxTryp\_SPc1* gene was possibly down-regulated [31,32]. Thus, we further explored this gene in *P. xylostella*. Based on the unigene sequences from the midgut transcriptome database of *P. xylostella* [32], the full-length cDNA sequence of *PxTryp\_SPc1* gene (GenBank accession no. MN422356) was cloned from fourth-instar *P. xylostella* larval midgut tissue using specific primers (Supplementary Materials Table S1). The cDNA sequence of the *PxTryp\_SPc1* gene (799 bp) contains an ORF of 768 nucleotides encoding 222 amino acid residues. The genomic DNA (gDNA) sequence of this gene can be found in the *P. xylostella* genome (DBM-DB: http://iae.fafu.edu.cn/DBM, Gene ID: Px016056). The genomic analysis revealed that it contains four exons (Figure 2A). The amino acid sequence of the *PxTryp\_SPc1* showed structural features characteristic of members of the trypsin family, as three catalytic residues His (70H), Ser (116S), and Asp (211D) (Figure 2B).

The PxTryp\_SPc1 protein shares sequence identity from 17% to 52% with other insect trypsin orthologs, as revealed by the BLASTp homology search of the GenBank database (Supplementary Materials Figure S1). Moreover, phylogenetic analysis of different insect trypsin orthologs showed that trypsin proteins from different insect orders are clustered in independent branches and are evolutionarily conserved (Figure 2C). Additionally, the phylogenetic tree revealed close relationship among trypsin proteins from Lepidoptera and PxTryp\_SPc1, which indicated that these trypsin proteins are homologous. Moreover, those trypsin proteins that were reported to be related to Bt resistance were not identified as PxTryp\_SPc1 orthologs and were not included in this phylogenetic tree.

**Figure 2.** Structural and phylogenetic relationship analyses of *PxTryp\_SPc1* gene. (**A**) Genomic structure of *PxTryp\_SPc1* gene. Orange boxes represent exons, and the spaces between two boxes represent introns. The figure is drawn to scale. (**B**) Conserved domain annotation obtained from NCBI annotation of the PxTryp\_SPc1 protein sequence. The protein sequence was considered as a characteristic member of the trypsin family. The location of the signal peptide, cleavage site, active sites, and substrate binding sites are indicated by orange triangles. (**C**) Phylogenetic analysis of the PxTryp\_SPc1 protein and its orthologs in diverse insects by the neighbor-joining (NJ) method. The unrooted phylogenetic tree was constructed by ClustalW alignment of amino acid sequences in MEGA-X. The bootstrap values with 1000 replications are shown on branches. The amino acid sequences of these trypsins were retrieved from the GenBank database (GenBank accession numbers are listed below). The scale bar shows the evolutionary distances. Abbreviations: 1. Lepidoptera (Pm (*Papilio machaon*, KPJ14943); Pr (*Pieris rapae*, XP\_022118678); Pxy (*Plutella xylostella*, MN422356); Hv (*Heliothis virescens*, AFO68329); Sl (*Spodoptera litura*, XP\_022815738); Dp (*Danaus plexippus*, OWR45697); Ms (*Manduca sexta*, CAM84320); Ha (*Helicoverpa armigera*, ABU98624); Bm (*Bombyx mori*, XP\_004923288); Pxu (*Papilio xuthus*, KPJ03461)); 2. Hemiptera (Dn (*Diuraphis noxia*, XP\_015367971); Hh (*Halyomorpha halys*, XP\_024219146); Ls (*Laodelphax striatellus*, RZF38227); Bt (*Bemisia tabaci*, XP\_018896298); Nl (*Nilaparvata lugens*, XP\_022184709)); Ago (*Aphis gossypii*, XP\_027850262); Ap (*Acyrthosiphon pisum*, XP\_001943273)); 3. Coleoptera (Obo (*Oryctes borbonicus*, KRT83696); Tm (*Tenebrio molitor*, AFB81537); Nv (*Nicrophorus vespilloides*, XP\_017773892); Ot (*Onthophagus taurus*, XP\_022900611)); 4. Blattodea (Bg, (*Blattella germanica*, AAZ78212); Pa (*Periplaneta americana*, AIA09342); Zn (*Zootermopsis nevadensis*, XP\_021914447)); 5. Diptera (As (*Anopheles sinensis*, KFB42846); Aga (*Anopheles gambiae*, XP\_317171.2); Aa (*Aedes aegypti*, XP\_001657786); Bd (*Bactrocera dorsalis*, XP\_011214086); Cq (*Culex quinquefasciatus*, XP\_001847028); Dm (*Drosophila melanogaster*, NP\_001285772)); 6. Orthoptera (Sg, (*Schistocerca gregaria*, CAA70820)); 7. Hymenoptera (Ar (*Athalia rosae*, XP\_020711972); Am (*Apis mellifera*, XP\_623564); Ac (*Apis cerana*, XP\_016922703); Af (*Apis florea*, XP\_012344846); Si (*Solenopsis invicta*, XP\_011166798); Hs (*Harpegnathos saltator*, EFN81462); Cf (*Camponotus floridanus*, XP\_011266670); Ob (*Ooceraea biroi*, XP\_011336015)); 8. Anoplura (Phc (*Pediculus humanus corporis*, AAV48634)).

#### *2.4. Tissue Expression Profiles of the PxTryp\_SPc1 Gene*

Expression analysis of the *PxTryp\_SPc1* gene by qPCR in the different tissues of the fourth-instar larvae indicated that it was specifically expressed in the midgut (MG) tissue, in contrast to its expression in the head, integument, testis, and Malpighian tubules (*p* < 0.05; Duncan's test; n = 3) (Figure 3A). Moreover, expression analysis of *PxTryp\_SPc1* gene in various developmental stages showed that its expression levels gradually raised from egg (EG) into the larval stages and reached the highest peak in the fourth-instar larvae (L4), while it showed low expression in pre-pupae, pupae, female, and male adults (Figure 3B). The expression of *PxTryp\_SPc1* gene was high in midgut and larval stages of *P. xylostella*.

**Figure 3.** qPCR expression profile of the *P. xylostella PxTryp\_SPc1* gene in different tissues and developmental stages. (**A**) Relative expression levels of *PxTryp\_SPc1* in different tissues including head (HD), integument (IN), midgut (MG), testis (TS), and Malpighian tubules (MT) of fourth-instar larvae. (**B**) Expression profile of *PxTryp\_SPc1* in different developmental stages: eggs (EG), first-instar larvae (L1), second-instar larvae (L2), third-instar larvae (L3), fourth-instar larvae (L4), prepupae (PP), pupae (P), male adults (MA), and female adults (FA). *RPL32* gene expression was used as the internal reference gene to normalize and calculate the gene expression levels. Expression level was calculated according to the value of the lowest expression identified (Tissues: HD; developmental stages: P), which was given an arbitrary value of 1. The means and the corresponding standard errors are shown. Different letters stand for statistically significant differences within the three replicates and four technical repeats (*p* < 0.05; Duncan's test; n = 3).

#### *2.5. The Expression of PxTryp\_SPc1 Gene in Susceptible DBM1Ac-S and Resistant SZ-R Strains*

Expression difference of the *PxTryp\_SPc1* gene by qPCR was compared in the resistant SZ-R and susceptible DBM1Ac-S strains (Figure 4). In general, the transcript levels of *PxTryp\_SPc1* showed a significantly reduced expression (about 2.8-fold down) in the SZ-R strain compared to the DBM1Ac-S strain (*p* < 0.05; Duncan's test; n = 3). Furthermore, treatment of third-instar SZ-R larvae with a high concentration of Cry1Ac protoxin (2000 mg/L), showed that the transcript level of *PxTryp\_SPc1* gene was further down-regulated (*p* < 0.05; Duncan's test; n = 3), showing a ratio of ~5.1-fold down compared to the DBM1Ac-S strain (Figure 4).

**Figure 4.** The expression differences of *PxTryp\_SPc1* gene between susceptible and resistant strains. Expression levels of *PxTryp\_SPc1* by qPCR in fourth-instar larval midgut tissue from susceptible and resistant strains. Lane 1: DBM1Ac-S; Lane 2: SZ-R; Lane 3: SZ-R intoxicated with 2000 mg/L Cry1Ac protoxin. *RPL32* gene was considered as a reference gene to normalize and calculate the level of gene expression. The expression level was calculated based on the value of the highest expression (DBM1Ac-S, arbitrary value of 1). The means and standard errors are shown. Different letters represent statistically significant differences with three independent repeats and four technical replications (*p* < 0.05; Duncan's test, n = 3).

#### *2.6. Linkage between Decreased PxTryp\_SPc1 Gene Expression and Cry1Ac Resistance in SZ-R Strain*

To determine the genetic linkage of decreased *PxTryp\_SPc1* expression with Cry1Ac resistance in the SZ-R strain, a single-pair cross between a male SZ-R larva and a female DBM1Ac-S larva was performed to obtain F1 progeny. Subsequently, backcross family a or b generated from reciprocal crosses between SZ-R moths and F1 progeny were selected and fed on cabbage leaves without or with a diagnostic dose of Cry1Ac protoxin (20 mg/L), and the midgut samples from fourth-instar *P. xylostella* larvae were subjected to qPCR analysis. The qPCR results indicated that *PxTryp\_SPc1* gene expression levels in individual fourth-instar larval midguts from F1 generation resemble those in their susceptible DBM1Ac-S strain (Figure 5), implying that the resistance trait in SZ-R is recessive. Nevertheless, the expression levels of *PxTryp\_SPc1* in midgut tissue from two backcross families (backcross a and b) showed two different groups; one displayed notable decreased expression levels of *PxTryp\_SPc1* (< ~2.8-fold), but another group demonstrated similar expression levels to those of larvae midgut tissue from the original susceptible DBM1Ac-S strain or the F1 generation from the DBM1Ac-S and SZ-R strains cross (Figure 5). The ratios between the two families of individuals were found to be 10:8 (backcross a) and 9:9 (backcross b), following the calculated random assortment ratio 1:1 basically (*p* > 0.1 or *p* = 1.0; χ<sup>2</sup> test). On the contrary, all of the survivals from Cry1Ac exposure in the two backcross families showed decreased expression levels of *PxTryp\_SPc1* (<~2.8-fold) compared to larvae of the DBM1Ac-S strain or the F1 progenies, testifying a co-segregation (linkage) with resistance to Cry1Ac in SZ-R (*p* < 0.05, χ<sup>2</sup> test) (Figure 5). Thus, the decreased expression level of the *PxTryp\_SPc1* gene was tightly linked to Cry1Ac resistance in the *P. xylostella* SZ-R strain.

(*rs*) untreated (*rs:rr*) treated (*rr*) untreated (*rs:rr*) treated (*rr*)

**Figure 5.** Genetic linkage analysis of the decreased *PxTryp\_SPc1* expression level in the SZ-R strain of *P. xylostella* and resistance to Cry1Ac. The expression levels of *PxTryp\_SPc1* in F1 larvae, Cry1Ac-treated backcross families (family a and b), and non-selected (untreated) are shown in relation to the levels in the DBM1Ac-S strain. Corresponding intensity of PCR bands for the *PxTryp\_SPc1* and the reference *RPL32* gene are exhibited (Upper).

#### *2.7. RNAi-Mediated Functional Assay of the PxTryp\_SPc1 Gene*

The *PxTryp\_SPc1* gene expression was silenced by microinjection of *P. xylostella* susceptible larvae with *PxTryp\_SPc1* dsRNA to determine the potential role of the *PxTryp\_SPc1* gene in Cry1Ac resistance. The expression levels were statistically reduced after 24 h post dsRNA injection, and the reduction effect lasted almost 96 h. In contrast, controls treated with buffer or dsEGFP, did not show any silencing effect on *PxTryp\_SPc1* expression (Figure 6A). The subsequent bioassays revealed that silencing of *PxTryp\_SPc1* gene reduced larval susceptibility to Cry1Ac protoxin at 1 mg/L (the LC50 value) or at 2 mg/L (the LC90 value) after 48 h post-injection compared to control larvae injected with buffer or dsEGFP (Figure 6B), suggesting that the reduced expression of *PxTryp\_SPc1* gene correlated with higher tolerance of *P. xylostella* to Cry1Ac. We also determined the effect of *PxTryp\_SPc1* gene silencing on Cry1Ac protoxin activation using larval midgut extracts from different RNAi treatments. After 1 h of Cry1Ac incubation from larvae injected with nonspecific dsEGFP and buffer-only, a single band of ~65 kDa was observed (Figure 6C, lanes 4 and 6). In contrast, larvae treated with dsPxTryp\_SPc1 showed the two bands of Cry1Ac as previously observed with midgut extract from the SZ-R strain (Figure 6C, lane 5 compared to Figure 6C lane 5).

**Figure 6.** Influences on Cry1Ac toxicity of *PxTryp\_SPc1* expression in larval midgut after RNAi silencing. (**A**) Impacts on injection of larvae with buffer, dsEGFP, or dsPxTryp\_SPc1 on *PxTryp\_SPc1* expression after 120 h RNAi silencing from *P. xylostella*. Different letters represent statistically significant differences within three repeats and four technical replications (*p* < 0.05; Duncan's test, n = 3). (**B**) Mortality of *P. xylostella* larvae after treatment with two concentrations of Cry1Ac protoxin; larvae were injected with buffer, dsEGFP, or dsPxTryp\_SPc1. Within each group, different letters denote statistically significant differences between treatments (*p* < 0.05; Duncan's test, n = 3). (**C**) Activation analysis of Cry1Ac protoxin by *P. xylostella* midgut extracts from larvae injected with: dsEGFP (lane 4), dsPxTryp\_SPc1 (lane 5), and buffer only (lane 6). Lane1: protein maker; Lane 2: Cry1Ac protoxin; Lane 3: Cry1Ac incubated with bovine trypsin (positive control).

#### **3. Discussion**

The role and function of the insect proteases present in midgut tissue in mediating Bt resistance has been analyzed in different *P. xylostella* resistant strains. It was shown before that decreased activation of protoxin to toxin could be a major Bt resistance mechanism in the Cry1Ac-resistant Cry1Ac-SEL strain of *P. xylostella* [35] and a Bt resistance strain of *P. xylostella* [36], but the specific midgut protease gene involved was not identified. Here, we show that the reduced expression of the *PxTryp\_SPc1* trypsin gene contributes to Cry1Ac resistance in the Cry1Ac-resistant SZ-R strain. The activities of caseinolytic and trypsin proteases in midgut extracts significantly decreased in contrast to the susceptible strain, suggesting that reduced trypsin protease activity is associated with a resistant phenotype of the SZ-R strain. Based on previously published transcriptome, RNA-Seq, and proteomics-based studies [31,32,37], we identified a new trypsin gene, *PxTryp\_SPc1* (GenBank accession no. MN422356, DBM-DB gene ID: Px016056), which is mainly expressed in the midgut and that is down-regulated in the SZ-R resistant strain. Previous identified trypsin proteins involved in Bt resistance in different lepidopteran insects including OnT23 (~50%, GenBank accession no. AAR98919), SfT6 (~43%, GenBank accession no. ACR25157), HaSP2 (~33%, GenBank accession no. ABP96915), and HaTryR (~28%, GenBank accession no. AHL46496) have been reported to be associated with Bt resistance in *O. nubilalis* [14], *S. frugiperda* [15], and *H. armigera* [16,17] (the percentage in parentheses means the identity between the amino acid sequences of these trypsins and PxTryp\_SPc1). Although the protein identity between PxTryp\_SPc1 and OnT23 or SfT6 is as high as 50% and 43%, respectively, another *P. xylostella* trypsin that shared 57% protein similarity with PxTryp\_SPc1 was identified in the *P. xylostella* genome database (DBM-DB gene ID: Px015403). This *P. xylostella* trypsin has higher protein identity (55% and 63%, respectively) with OnT23 or SfT6, indicating that PxTryp\_SPc1 and OnT23 or SfT6 are actually not orthologs. Some of the orthologs of PxTryp\_SPc1 in other lepidopteran insects are shown in the phylogenetic tree constructed in this study (Figure 2C). The role of other orthologs from different lepidopteran insects in Cry toxin activation still remains to be identified. These results indicated that PxTryp\_SPc1 is a novel trypsin member related to Bt resistance, which enriches the Bt-responsive midgut trypsin gene repertoire in insects.

The most common mechanism of high resistance levels to Bt Cry toxins in lepidopteran insects is related to reduced toxin binding to midgut receptors [7,38], while an altered protease expression mechanism has been associated with low or moderate resistance in lepidopteran insects [39]. Indeed, altered processing of Cry1Ac protoxin by midgut proteases is not related to high-level field-evolved Cry1Ac resistance in the *P. xylostella* NIL-R strain [30]. The laboratory-selected strain SZ-R shows moderate Cry1Ac resistance to Cry1Ac activated toxin or protoxin (Table 1). We reported that the *PxCAD* and *PxABCH1* genes are not associated with Cry1Ac resistance in the SZ-R strain [24], but differential expression of the *PxmALP*, *PxABCB1*, *PxABCC1*, *PxABCC2*, *PxABCC3*, and *PxABCG1* genes was shown to be associated with Cry1Ac resistance in the SZ-R strain [27–29], suggesting that reduction in toxin binding is associated with Cry1Ac resistance in the SZ-R strain. Thus, the resistance mechanism related to reduced expression of the trypsin gene *PxTryp\_SPc1* identified in this study is an additional mechanism in this *P. xylostella* strain. The reduced expression of the *PxTryp\_SPc1* gene correlated well with altered Cry1Ac protoxin activation, suggesting that incomplete activation of protoxin is conducive to developing the resistant phenotype (Figure 1B). However, our data showed that protoxin activation was not completely blocked in the SZ-R strain since treatment of Cry1Ac protoxin with midgut juice from the resistant population resulted in two protein bands; one correlated with the 65 kDa Cry1Ac activated toxin, implying that other midgut proteases may still participate in the activation of Cry1Ac protoxin in the resistant SZ-R strain. These data correlated with the toxicity bioassays revealed that SZ-R was only two-fold more susceptible to the Cry1Ac activated toxin compared with the protoxin (Table 1). Nevertheless, RNAi analysis showed that silencing *PxTryp\_SPc1* gene did reduce the larval susceptibility to Cry1Ac toxin supporting that this protein contributes to the Cry1Ac resistance phenotype of SZ-R strain (Figure 6). Moreover, reduced expression of *PxTryp\_SPc1* was linked to Cry1Ac resistance in SZ-R strain (Figure 5).

The regulation mechanism involved in the reduced expression of *PxTryp\_SPc1* in the SZ-R strain still remains to be determined. Interestingly, down-regulation of the trypsin gene *HaTryR* in *H. armigera* is caused by a promoter sequence mutation mediated *cis*-regulatory mechanism [17]. Our previous studies demonstrated that different expression of the *PxmALP*, *PxABCB1*, *PxABCC1–3*, and *PxABCG1* genes can be modulated by the MAPK signaling pathway [27,29]. Thus, whether reduced expression of *PxTryp\_SPc1* in SZ-R strain is conferred by the promoter mutation-induced *cis*- or MAPK-induced *trans*-regulatory mechanism warrants further study. In addition, considering that the *PxABCC2* and *PxABCC3* genes were successfully knocked out by a novel CRISPR/Cas9 genome editing tool confirming their involvement in Bt Cry1Ac resistance [40], we will further utilize the CRISPR/Cas9 system to edit the *PxTryp\_SPc1* gene to offer in vivo reverse genetic evidence of its involvement in Cry1Ac resistance, which thus could help us in the future to determine what is the function of *PxTryp\_SPc1* gene during Bt Cry1Ac toxin activation processing in *P. xylostella*. Moreover, the *P. xylostella* genome contains many different trypsin genes [41]; some other trypsin genes displaying marked differential expression levels between Cry1Ac-susceptible and -resistant *P. xylostella* strains have also been found previously by transcriptome, RNA-Seq, and proteomics analyses, and further investigations of their potential functions in Bt Cry1Ac resistance in *P. xylostella* are also needed.

Overall, our data confirmed that down-regulation of a novel trypsin gene (*PxTryp\_SPc1*) is associated with Cry1Ac resistance in the SZ-R strain of *P. xylostella*. This study is helpful to elucidate the complex causes of Bt Cry1Ac resistance in *P. xylostella*. The deeper understanding that we have of these mechanisms, the stronger and better strategies we will be able to propose to cope with the evolution of insect resistance to Bt toxins.

#### **4. Materials and Methods**

#### *4.1. Insect Strains*

The *P. xylostella* susceptible DBM1Ac-S and resistant SZ-R strains that were used in this study were previously described [24,28]. The SZ-R strain was originated from field collected moths at Shenzhen in China (2003), and it was constructed by constant selection with a concentration of Cry1Ac protoxin that generally kills 50%–70% of larvae in the laboratory for more than 200 generations. Both *P. xylostella* strains were reared on Chinese cabbage, JingFeng No. 1 (*Brassica oleracea* var. *capitata*), at 65% RH, 25 ◦C, with a photoperiod of 16 h light:8 h dark, and adults were fed with a 10% sucrose solution.

#### *4.2. Midgut Protease Activity Assays*

The caseinolytic protease was measured at 28 ◦C using the substrate azocasein (Sigma, St. Louis, MO, USA), as previously reported [42]. In brief, midgut extracts (20 μL) were mixed with 1% azocasein in 50 mM NaHCO3-Na2CO3 buffer (150 μL) and incubated for 2 h at 28 ◦C. Then 10% trichloroacetic acid (TCA) (170 μL) was used to stop the reaction. The solution was incubated at 25 ◦C for 1 h and centrifuged for 15 min at 16,000× *g* at room temperature to remove the debris. Then, 1 M NaOH (340 μL) was mixed and the optical density (OD) of collected supernatant was measured at 450 nm in a SpectraMaxM2*<sup>e</sup>* microplate reader (Molecular Devices, Sunnyvale, CA, USA).

Chymotrypsin and trypsin activities were detected using 1 mM Nα-benzoyl-L-arginine-*p* -nitroanilide (BApNA, Sigma) and 1 mM *N*-succinyl-Ala-Ala-Pro-phenylalanine *p*-nitroanilide (SAApFpNA, Sigma) as respective specific substrates. For chymotrypsin activity determination, 5 μL midgut extract was mixed with 3 mL of 1 mM SAApFpNA in 50 mM NaHCO3-Na2CO3 buffer. For trypsin activity examination, 10 μL midgut extract was mixed with 3 mL of 1 mM BApNA in 50 mM NaHCO3-Na2CO3 buffer. The peptidolytic reaction was tested immediately by recording continuously the optic density (OD) value at 405 nm every 15 s at 28 ◦C for 30 min. The enzyme activities are exhibited as relative activities of the DBM1Ac-S midgut extract protease activities, which were considered as 100%. Biological assay was performed in triplicate and four technical repetitions

each were used to confirm the protease activities. For analysis of statistical differences among samples, one-way ANOVA with Duncan's tests (*p* < 0.05) was used.

#### *4.3. Bioassays*

The Cry1Ac protoxin and trypsin-activated toxin were obtained as previously described [30]. The Cry1Ac toxin was finally dissolved in 50 mM Na2CO3 (pH 9.6) and stored at −20 ◦C. The respective toxicity of the Cry1Ac protoxin and trypsin-activated toxin was determined by 72-h leaf-dip bioassays using a total of 280 third-instar *P. xylostella* larvae per bioassay as described before [27,28]. In short, ten larvae that were exposed to seven different concentrations of Cry1Ac toxin in each group, and four repeats were performed for all bioassays. The control mortality did not exceed 5%. We used the POLO Plus 2.0 software (LeOra Software, Berkeley, CA, USA) to calculate the LC50 values (median lethal concentrations killing 50% of the tested larvae) and 95% CL (95% confidence limits of the LC50) values by Probit analysis.

#### *4.4. RNA Extraction and cDNA Synthesis*

The methods of RNA extraction and cDNA synthesis from *P. xylostella* were previously described [27]. The midgut samples were extracted in TRIzol Reagent (Invitrogen, Carlsbad, CA, USA); then the concentration of RNA was quantified by a NanoDrop 2000c spectrophotometer (Thermo Fisher Scientific Inc., Waltham, MA, USA). PrimeScript II 1st strand cDNA Synthesis Kit (TaKaRa, Dalian, China) was used to synthetize first-strand DNA. For qPCR analysis, 1 μg total RNA was used to perform the first-strand cDNA with the PrimeScript RT kit (containing gDNA Eraser, Perfect Real Time) (TaKaRa, Dalian, China) following the manufacturer's instructions. The synthesized cDNA was immediately used or stored at −20 ◦C until used.

#### *4.5. Gene Identification and Cloning*

The candidate cDNA sequence of *PxTryp\_SPc1* gene was identified in our previous midgut transcriptome database of *P. xylostella* [32] and was further in silico corrected by the *P. xylostella* genome database (DBM-DB: http://iae.fafu.edu.cn/DBM, Gene ID: Px016056); then the specific primers were designed (Supplementary Materials Table S1) and were used in subsequent PCR amplification assays. The full-length cDNA sequence *of PxTryp\_SPc1* gene was finally obtained and deposited in the GenBank database (accession no. MN422356).

As described previously [27], the PCR reaction (25 μL total volume) contained 0.2 μL LA Taq HS polymerase (TaKaRa, Dalian, China) in an C1000 Thermal Cycler PCR system (BioRad, Philadelphia, PA, USA) for 35 cycles using LA Taq polymerase (TaKaRa, Dalian, China). A gel extraction kit (CWBIO, Beijing, China) was used for purification of the PCR products of *PxTryp\_SPc1*, which were further cloned into the pEASY-T1 vector (TransGen, Beijing, China). For gene sequencing, *Escherichia coli* TOP10 competent cells (TransGen, Beijing, China) were transformed with candidate plasmids.

#### *4.6. Gene Sequence Analysis*

DANMAN 8.0 (Lynnon BioSoft, San Ramon, CA, USA) software was used for gene sequence assembly, exon-intron analysis, and multiple sequence alignment. The open reading frame (ORF) was identified by the ORF finder tool at the NCBI (https://www.ncbi.nlm.nih.gov/orffinder/), and predicted amino acid sequences were achieved by ExPASy online tool to translate (https://web.expasy. org/translate/). The BLAST tool at the GenBank database (https://blast.ncbi.nlm.nih.gov) was used for the sequence-similarity analyses. The protein-specific motifs and active sites were found and annotated at the GenBank database (https://www.ncbi.nlm.nih.gov/). The signal peptide was predicted by SignalP-5.0 Server online (http://www.cbs.dtu.dk/services/SignalP/).

#### *4.7. Phylogenetic Tree Construction*

To verify the classification of the *PxTryp\_SPc1* gene, phylogenetic analysis of the PxTryp\_SPc1 protein was done by using the full-length amino acid sequences of its orthologs from other insects. MEGA-X software (https://www.megasoftware.net/) with ClustalW algorithm was used to construct the phylogenetic tree. An unrooted neighbor-joining (NJ) phylogenetic tree was done choosing the "p-distance" as the amino acid substitution model; the bootstrap value was determined from 1000 replicates.

#### *4.8. Sample Preparation*

Samples from different developmental stages were collected, and different tissues were also dissected from the fourth-instar DBM1Ac-S larvae to characterize the spatio-temporal expression patterns of the *PxTryp\_SPc1* gene. Moreover, in order to resolve whether the expression level of *PxTryp\_SPc1* was related to Cry1Ac resistance, third-instar SZ-R larvae were treated with a high concentration of Cry1Ac protoxin (2000 mg/L). After the midguts from the survivors were dissected, the extraction of total RNA and cDNA was synthesized as mentioned above. Data were obtained from three biological replications performed in all samples.

#### *4.9. Gene Expression Analysis*

Gene expression differences were determined by real-time quantitative PCR (qPCR) as described before with slight modification [27,28]. Briefly, Primer Premier 5.0 (PREMIER Biosoft international, Palo Alto, CA, USA) was used for defining specific *PxTryp\_SPc1* gene primers (Supplementary Materials Table S1). PCR reactions (20 μL) contained 7.4 μL RNase-Free ddH2O, 10 μL of 2 × FastFire qPCR PreMix Plus (TIANGEN, Beijing, China), 5 μM of each specific primer, 1 μL of first-strand cDNA template, and 0.4 μL 50 × ROX Reference Dye (TIANGEN, Beijing, China). The running program consisted of a denaturation at 95 ◦C for 10 min followed by 40 denaturalized cycles at 95 ◦C for 15 s, annealing at 57 ◦C for 30 s, and extension at 72 ◦C for 30 s. All reactions were performed in an Applied Biosystems QuantStudio 3 Real-Time PCR System (Applied Biosystems, Forster City, CA, USA). As an internal control for relative quantification, the *ribosomal protein L32* (*RPL32*) gene (GenBank accession no. AB180441) was used in qPCR data analysis. Three biological repetitions and four technical repetitions were conducted for each sample. To define the statistically differences, one-way ANOVAs with Duncan's test (overall significance level *p* < 0.05) were used.

#### *4.10. Linkage Analysis*

Genetic linkage analysis was performed as previously described [27,28]. F1 progeny was generated by a single-pair mating between a SZ-R male and a DBM1Ac-S female. A diagnostic Cry1Ac protoxin diagnostic dose (20 mg/L) killed all the F1 (heterozygous) larvae was determined in a toxicity bioassay. Reciprocal crosses between SZ-R moths and F1 progeny were made to generate backcross family a and b. Forty larvae from each backcross families of progeny were fed on cabbage (non-Cry1Ac-selected) or cabbage with 20 mg/L of Cry1Ac protoxin (Cry1Ac-selected), and midguts tissues from the survived fourth-instar *P. xylostella* larvae were dissected for qPCR analysis as mentioned above.

#### *4.11. RNA Interference (RNAi)*

To determine the impact of *PxTryp\_SPc1* gene expression in *P. xylostella* resistance to Cry1Ac, RNAi-mediated down expression of *PxTryp\_SPc1* gene was performed. Early third-instar *P. xylostella* larvae were microinjected with specific dsRNA targeting *PxTryp\_SPc1* gene (dsPxTryp\_SPc1), as described previously [24]. Briefly, Primer Premier 5.0 (PREMIER Biosoft international, Palo Alto, CA, USA) was used to design the dsRNA primers containing the T7 promoter on the 5 end targeting gene-specific region of *PxTryp\_SPc1* (GenBank accession no. MN422356) or *EGFP* gene (GenBank accession no. KC896843) (Supplementary Materials Table S1). To further validate the specificity of

these dsRNAs, BLASTn searches of the GenBank database (https://www.ncbi.nlm.nih.gov/) and the *P. xylostella* genome database (DBM-DB: http://iae.fafu.edu.cn/DBM) were performed showing no unspecific hit diminishing potential off-target effects. The amplicons (389 bp for dsPxTryp\_SPc1 and 469 bp for dsEGFP) were used as templates for in vitro transcription reactions to produce dsRNAs using the T7 RiboMAX Express RNAi System (Promega, Madison, WI, USA). The synthesized dsRNA was dissolved in 10 mM Tris–HCl (pH 7.0), and 1 mM EDTA was used as injection buffer and mixed with Metafectene PRO transfection reagent (Biontex, Planegg, Germany). A total of 300 ng (70 nL) dsEGFP or dsPxTryp\_SPc1 were injected into the hemocoel of DBM1Ac-S larvae, resulting in less than 20% larval mortality determined after 5 days. Finally, to determine the silencing efficiency at 48 h post-injection, midgut tissue was dissected from injected larvae. The control group was injected with equal volumes of buffer alone. At least 30 larvae were analyzed for each treatment, and three replicate experiments were conducted. The bioassay data were processed as mentioned above. Statistically significant differences between qPCR and bioassay analyses were determined by one-way ANOVAs with Duncan's tests (overall significance level *p* < 0.05).

**Supplementary Materials:** The following are available online at http://www.mdpi.com/2072-6651/12/2/76/s1, Figure S1. Pairwise comparisons of protein sequence identities among nine trypsin orthologs of PxTryp\_SPc1 from different insect species. Table S1. List of primers used in this study.

**Author Contributions:** L.G. (Lijun Gong), Z.G., L.W., and Y.Z. conceived and designed the experiments. L.G. (Lijun Gong), Z.G., S.K., J.Z., D.S., L.G. (Le Guo), J.Q., L.Z., Y.B., F.Y., M.A., Q.W., S.W., B.X., and Z.Y. performed the experiments. L.G. and Z.G. analyzed the data. L.G. (Lijun Gong), A.B., M.S., Z.G., L.W., and Y.Z. wrote the paper. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research was funded by the National Natural Science Foundation of China (31630059; 31701813), the Beijing Key Laboratory for Pest Control and Sustainable Cultivation of Vegetables, and the Science and Technology Innovation Program of the Chinese Academy of Agricultural Sciences (CAAS-ASTIP-IVFCAAS).

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**


© 2020 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).

*Communication*

## **Study of the** *Bacillus thuringiensis* **Cry1Ia Protein Oligomerization Promoted by Midgut Brush Border Membrane Vesicles of Lepidopteran and Coleopteran Insects, or Cultured Insect Cells**

**Ayda Khorramnejad 1,2, Mikel Domínguez-Arrizabalaga 3, Primitivo Caballero 3, Baltasar Escriche <sup>1</sup> and Yolanda Bel 1,\***


Received: 16 December 2019; Accepted: 19 February 2020; Published: 21 February 2020

**Abstract:** *Bacillus thuringiensis* (Bt) produces insecticidal proteins that are either secreted during the vegetative growth phase or accumulated in the crystal inclusions (Cry proteins) in the stationary phase. Cry1I proteins share the three domain (3D) structure typical of crystal proteins but are secreted to the media early in the stationary growth phase. In the generally accepted mode of action of 3D Cry proteins (sequential binding model), the formation of an oligomer (tetramer) has been described as a major step, necessary for pore formation and subsequent toxicity. To know if this could be extended to Cry1I proteins, the formation of Cry1Ia oligomers was studied by Western blot, after the incubation of trypsin activated Cry1Ia with insect brush border membrane vesicles (BBMV) or insect cultured cells, using Cry1Ab as control. Our results showed that Cry1Ia oligomers were observed only after incubation with susceptible coleopteran BBMV, but not following incubation with susceptible lepidopteran BBMV or non-susceptible Sf21 insect cells, while Cry1Ab oligomers were persistently detected after incubation with all insect tissues tested, regardless of its host susceptibility. The data suggested oligomerization may not necessarily be a requirement for the toxicity of Cry1I proteins.

**Keywords:** Cry1Ab; oligomer formation; Sf21 cell line; *Ostrinia nubilalis*; *Lobesia botrana*; *Leptinotarsa decemlineata*; bioassay

**Key Contribution:** The paper studies the oligomer formation of trypsin activated Cry1I in vitro, after incubation with insect BBMV or insect cultured cells. The results show that Cry1Ia oligomers are only visualized after incubation with coleopteran susceptible BBMV. This could suggest that oligomerization may not be a limiting step in the mode of action of Cry1I protein.

#### **1. Introduction**

The entomopathogenic Gram-positive bacterium, *Bacillus thuringiensis* (Bt), is the most successful bioinsecticide commercialized to date. It generates a wide variety of insecticidal proteins that can be produced at different growth stages. Cry, Cyt and parasporin proteins are synthetized during the stationary growth phase, Cry1I proteins are secreted in the initial phase of sporulation and Vip and Sip proteins are secreted during the vegetative phase of bacterial growth [1–5]. The success of Bt-based insecticides is due to its narrow spectrum of activity, environmental safety and because it is

harmless to animals and plants [6]. To date, the most studied Bt entomopathogenic proteins are the three domain (3D) Cry proteins such as Cry1 and Cry2 toxins. Their mode of action is not completely known, but it is commonly accepted that specific binding to insect midgut receptors is essential to exert their toxicity [7,8]. The current models of Cry toxin action include the "signal transduction model" that claims that the toxicity is mediated by intracellular pathways [9], and the "sequential mode of action"—the most accepted model so far, that is based on the sequential binding of Bt toxin to several midgut receptors, the promotion of a pre-pore oligomer, the insertion of pre-pore oligomer into the midgut membrane, pore formation, osmotic imbalance, midgut epithelium disruption, septicemia and insect death [10]. Regarding this model, some authors have stated that the formation of the oligomer prior to toxin insertion into membrane is a major step in the toxicity process, and that the oligomeric (tetrameric) structure, is necessary for the final pore formation; also, it has been claimed that the oligomer formation is a conserved mechanism in the mode of action of the Cry proteins [11–16].

Oligomerization has been studied in several wild type and mutant toxins such as Cry1Aa, Cry1Ab, Cry1Ac, Cry1Ca, Cry1Da, Cry1Ea, Cry1Fa, Cry2Ab, Cry3Aa, Cry3Ba, Cry3Ca, Cry4Ba, Cry11Aa and Cry46Aa1 [15,17–25]. Oligomerization has also been detected in other Bt toxins such as Cyt [26] and Vip [27]. However, regarding the Cry1I protein family, only one study has described the possible oligomerization of this protein in solution, in the absence of insect midgut proteins [28]. So far, the promotion of Cry1I oligomer formation after incubation with insect tissues or insect cell-derived models has never been experimentally addressed.

Cry1I proteins are included within the Cry1 family (3D Cry proteins) because of their sequence similarities and structural characteristics. However, Cry1I proteins display several unique, specific and remarkable characteristics, that include their secretion early in the stationary growth phase of Bt (instead of forming part of the Bt crystals), their unusual protoxin molecular weight (80 kDa), and the dual insecticidal activity against lepidopteran and coleopteran pests [29,30]. This, together with the lack of cross-resistance with other Cry1A and Cry1F insecticidal proteins [31–33], make Cry1I proteins interesting for developing new insecticidal products and indeed, they have been recently introduced in Bt crops [34,35].

Understanding the mode of action of Bt toxins is critical for enhancing and sustaining their efficacy against pests. In this context, in the present work, for the first time, the oligomer formation of Cry1Ia toxins has been examined after incubation with midgut insect tissues, trying to clarify its relevance in the Cry1I mode of action. Cry1Ab, for which oligomerization has been reported in several studies [36–39], has been used as control. In this study, midgut brush border membrane vesicles (BBMV) of different insect species from Lepidoptera (*Ostrinia nubilalis* and *Lobesia botrana*) and Coleoptera (*Leptinotarsa decemlineata*), as well as a cell line derived from *Spodoptera frugiperda* (Sf21) were used to promote the Cry oligomer formation. This selection covered both susceptible and tolerant insect species for Cry1Ia and Cry1Ab. Based on this, the oligomer formation of Cry1Ia toxin compared to Cry1Ab, and its possible correlation with insecticidal activity, have been examined.

#### **2. Results**

#### *2.1. Toxicity of Cry1Ia and Cry1Ab against Lepidopteran and Coleopteran Species*

The toxicity of Cry1Ia and Cry1Ab protoxins towards the insects' species used in the present study were assessed by performing bioassays. The results are summarized in Table 1. The Cry1Ia protein was found to be toxic for the two selected lepidopteran species, *L. botrana*, and *O. nubilalis*, with LC50 values of 80 and 273 ng/cm<sup>2</sup> respectively, as well as for the coleopteran *L. decemlineata* (LC50 = 22 μg/mL). On the other hand, Cry1Ab was only toxic for the lepidopteran insect species.


**Table 1.** Toxicity parameters of Cry1Ab and Cry1Ia protoxins.

LC50 values are expressed as ng/cm<sup>2</sup> for *L. botrana* and *O. nubilalis*, and in μg/mL for *L. decemlineata*. NT: non-toxic at 100 μg/mL.

#### *2.2. BBMV of Susceptible Lepidopteran Insects Promoted Oligomerization of Cry1Ab But Not of Cry1Ia*

The oligomer formation of Cry1Ab and Cry1Ia was studied by incubating the proteins with lepidopteran BBMV from *O. nubilalis* and *L. botrana*, susceptible hosts (Table 1). In order to favor oligomer detection, the milder SDS-PAGE denaturing conditions used in the bibliography to observe Cry1 oligomers were employed (see Section 4.8 in Materials and Methods). The results showed that Cry1Ab toxin was able to form oligomers after incubation with BBMV from *O. nubilalis*, but this incubation did not promote Cry1Ia oligomer formation (Figure 1). The oligomers observed (band of about 250 kDa) were associated to the *O. nubilalis* BBMV fraction (Figure 1a, lane P); these oligomeric structures could be inserted into the membranes or just bound to the surface of the BBMV due to interaction with specific membrane proteins. For Cry1Ab, the Western blot revealed another band with a molecular weight of approximately 60 kDa, corresponding to the monomeric form of Cry1Ab (Figure 1a, lane P). Nevertheless, the Cry1Ia protein associated with *O. nubilalis* BBMV was detected as a single band of about 50 kDa, corresponding to Cry1Ia monomers, and no band corresponding to Cry1Ia oligomers was detected (Figure 1b, lane P). The Cry1Ab and Cry1Ia monomeric proteins were also recovered in the supernatants, as bands of about 60 and 50 kDa, respectively (Figure 1, lanes S). Controls of Cry1Ab or Cry1Ia proteins without BBMV, but subjected to the same process as the rest of the samples, were conducted to assess the possible spontaneous formation of oligomers. The results (Figure 1, lanes C) showed the presence of Cry1Ab and Cry1Ia monomers mostly while the oligomers (tetramers) were not detected in these lanes, pointing to the fact that Cry1Ab and Cry1Ia tetramers did not form spontaneously in the solution in the absence of insect BBMV. In the case of Cry1Ia some minor bands of MW about 65 and 90 kDa were detected, which were most probably traces of Cry1Ia protoxin and partially trypsinized products; a third minor band of about 130 kDa (MW that does not match with the size of Cry1Ia dimers or trimers) was also observed.

The oligomer formation was also tested using BBMV from *L. botrana*. The results obtained were similar to the ones found with *O. nubilalis* BBMV. The incubation of Cry1Ia with BBMV from *L. botrana*, did not render bands with molecular weight consistent with Cry1Ia oligomeric structures (Figure 2b, lane P), whilst, after incubation of Cry1Ab with *L. botrana* BBMV, a clear band corresponding to a Cry1Ab oligomer (tetramer) was observed (Figure 2, lane P). On the other hand, monomers of both proteins were detected in the corresponding supernatants recovered after the BBMV-protein incubation assays (Figure 2, lanes S). In the Cry1Ia experiments, minor bands of MW of about 65 and 90 kDa were observed in the supernatant (S) and control (C) lanes that probably represent a minor fraction of partially trypsinized Cry1Ia.

**Figure 1.** Cry1Ab and Cry1Ia oligomer formation promoted by *O. nubilalis* BBMV: (**a**) Cry1Ab; (**b**) Cry1Ia. Lanes B: *O. nubilalis* BBMV incubated without Cry1Ab or Cry1Ia proteins. Lanes C: Control of Cry1Ab or Cry1Ia proteins, incubated without BBMV. Lanes S: Supernatant obtained after incubation of Cry1Ab or Cry1Ia proteins, with the BBMV. Lanes P: Pellet obtained after incubation of Cry1Ab or Cry1Ia with the BBMV. Lanes M: Molecular weight marker. The arrowhead points to the Cry1Ab oligomer (about 250 kDa).

**Figure 2.** Cry1Ab and Cry1Ia oligomer formation promoted by *L. botrana* BBMV: (**a**) Cry1Ab; (**b**) Cry1Ia. Lanes B: *L. botrana* BBMV incubated without Cry1Ab or Cry1Ia proteins. Lanes C: Controls of Cry1Ab or Cry1Ia proteins incubated without BBMV. Lanes P: Pellet obtained after incubation of the respective protein with the *L. botrana* BBMV. Lanes S: Supernatant obtained after incubation of the respective protein with the *L. botrana* BBMV. Lanes M: molecular weight marker. The arrowhead points to the Cry1Ab oligomer (about 250 kDa).

#### *2.3. Oligomerization of Cry1Ia Was Promoted by BBMV from L. decemlineata*

Cry1Ia exhibits a dual toxic activity towards lepidopteran and coleopteran hosts, whereas Cry1Ab is only toxic to lepidopteran species. In this work, *L. decemlineata* BBMV was employed as a coleopteran Cry1Ia susceptible host tissue (Table 1) to study Cry1Ia oligomer promotion as well to study if these BBMV promoted the oligomerization of the Cry1Ab protein, non-toxic for this insect species (Table 1).

The results, summarized in Figure 3, showed that the incubation of either Cry1Ab or Cry1Ia toxins with the coleopteran BBMV provided bands of a molecular weight of about 250 kDa for both Cry proteins, corresponding to the respective oligomers. Interestingly, the oligomers were detected in both, pelleted (BBMV) and supernatant fractions (Figure 3, lanes P and S respectively). Bands corresponding

to the monomers of both proteins were also observed in both fractions (pelleted BBMV and supernatant) for both Cry proteins. Incubation with Cry1Ab rendered also a band of about 150 kDa that could correspond to a dimer. It is worth highlighting that the band corresponding to the Cry1Ia oligomer had a molecular weight of about 250 kDa, which would indicate that Cry1Ia oligomer could be composed by more than four subunits, since the Cry1Ia monomer has a size of about 50 kDa. Moreover, a strong band of molecular weight higher than 250 kDa was also observed as being associated to the BBMV (Figure 3, lane P) that could correspond to Cry1Ia aggregates with high number of units.

**Figure 3.** Cry1Ab and Cry1Ia oligomer formation promoted by *L. decemlineata* BBMV: (**a**) Cry1Ab; (**b**) Cry1Ia. Lanes B: *L. decemlineata* BBMV incubated without Cry1Ab or Cry1Ia proteins. Lanes C: Controls of Cry1Ab or Cry1Ia proteins incubated without BBMV. Lanes P: Pellet obtained after incubation of the respective protein with the *L. decemlineata* BBMV. Lanes S: Supernatant obtained after incubation of the respective protein with the *L. decemlineata* BBMV. Lanes M: molecular weight marker. The arrowheads in panels (**a**,**b**) point to the Cry1Ab (about 250 kDa) and the Cry1Ia (about 250 kDa) oligomer bands respectively.

#### *2.4. Oligomerization Promoted by Sf21 Insect Cells*

It has been described that Sf21 cells (insect cultured cells derived from *S. frugiperda* ovaries) are tolerant to both Cry1Ab and Cry1Ia proteins [40,41]. These insect cells were selected in order to check if oligomer formation could be promoted by tolerant insect tissues. Figure 4 shows the results of the incubation of Sf21 cells with each one of these proteins. In the case of Cry1Ab, the incubation of the protein with the insect cells resulted in the detection of two main bands of approximately 60 and 250 kDa associated with the cell fraction, corresponding to Cry1Ab monomers and oligomers, respectively (Figure 4a, lane P). Other minor bands detected were also present in the control lanes of Sf21 cells that had been incubated without Cry proteins, showing that these bands probably correspond to natural biotinylated proteins present in the cells (Figure 4, lanes B). In the case of Sf21 cells incubated with Cry1Ia, only a band with the molecular weight of the monomer (50 kDa) was found to be associated with the cells (Figure 4b, lane P). In summary, results pointed out the absence of Cry1Ia oligomer formation after incubation with Sf21 cells, in contrast to what is observed after incubation with Cry1Ab. It is worth noting that in the supernatants, only a main band with the molecular weight of the Cry1Ab or Cry1Ia monomers, was detected (Figure 4, lanes S).

**Figure 4.** Cry1Ab and Cry1Ia oligomer formation promoted by Sf21 cells: (**a**) Cry1Ab; (**b**) Cry1Ia. Lanes B: Sf21 cells incubated without Cry1Ab or Cry1Ia proteins. Lanes C: Controls of Cry1Ab or Cry1Ia proteins incubated without Sf21 cells. Lanes P: Pellet obtained after incubation of the respective protein with the Sf21 cells. Lanes S: Supernatant obtained after incubation of the respective protein with the Sf21cells. Lanes M: molecular weight marker. The arrowhead in Cry1Ab panel points to the Cry1Ab oligomer.

#### **3. Discussion**

Cry1I proteins share sequence and structural similarities to the most-known three domain Cry proteins present in the parasporal crystal of Bt [10,29,42]. Therefore, so far, it has been assumed that their mode of action is similar to its crystal protein counterparts, despite their special features such as that the Cry1I proteins do not form crystals [5,43], their protoxin MW is smaller (about 81 kDa [29]), and they show dual toxic activity against lepidopteran and coleopteran insect pests [29,44].

The mode of action of the Cry proteins accumulated in the crystals is not completely understood [10,12]. It is commonly accepted that includes the solubilization of the crystals in the insect midgut to yield the protoxin form, and a proteolytic processing to produce the activated forms. Then, the "sequential binding model", suggests that the activated proteins bind to several midgut membrane receptors to finally form an oligomeric structure that inserts in the midgut membrane and forms pores, leading to cellular osmotic imbalance, cell lysis, septicaemia and eventually insect death [13,14]. In support of this model, several studies have reported that oligomerization plays a crucial role in the insecticidal activity of *B. thuringiensis* Cry toxins [18,38,45,46]. Moreover, Cry protein mutants that did not form oligomers, showed severely decreased toxicity [17,24,45]. Likewise, it has been shown that some Cry1A mutants that had lost their toxicity, were unable to oligomerize and to form pores [38,47]. However, despite these studies, the sequential binding to several insect membrane proteins as well as the oligomer formation and insertion events prior to pore formation are not clearly defined yet [12]. On the other hand, as an alternative to the "sequential binding model", the "signaling pathway model" has been proposed. This model claims that the Cry toxicity can be due to the activation of intracellular cell death pathways [9]. Indeed, some authors claim that both mechanisms could coexist [48].

In this context, the occurrence of an oligomerization step in the mode of action of Cry1Ia protein, and the study of Cry1Ia oligomerization promotion by membranes of susceptible and tolerant insect BBMV or insect cells, have been the main goals of this research.

In 2009, the occurrence of a spontaneous oligomerization of trypsinized Cry1Ie in solution was reported [28]. According to the mentioned research, the oligomer fraction contained a small amount of dimer and a large amount of aggregates larger than tetramers. The toxicity of the oligomers against *Plutella xylostella* was about 70 times lower than the toxicity of the monomer or the toxicity of the non-trypsinized Cry1Ie protein, which led the authors to claim that the Cry1Ie spontaneous aggregation most likely differ from the oligomers occur by the insect midgut membranes. In the present work, the oligomerization of Cry1Ia protein after incubation with susceptible insect BBMV or with non-susceptible cultured insect cells, has been determined for the first time.

To properly select the insect species for this study, the insecticidal activity of Cry1Ia and Cry1Ab protoxins towards two lepidopteran species (*O. nubilalis* and *L. botrana*) and a coleopteran insect (*L. decemlineta*) has been assessed. The toxicity data obtained were expected based on the published data [29,44,49,50]. Moreover, Sf21 cells were also used in this study, on the bases that Cry1Ia and Cry1Ab are not toxic for them [40,41].

So far, Cry oligomerization studies have focused on the incubation of the Cry proteins with BBMV of susceptible and resistant populations of the same insect, and, as a result, an association has been found between resistance and reduced oligomerization [51,52]. In the present work, we have used Cry1Ab as an oligomeric control protein [36–39], and have been able to clearly detect Cry1Ab oligomers in the expected tetramer form (about 250 kDa molecular weight size) after incubation with all tested insect tissues, regardless to their susceptibility. Thus, oligomers have been observed after incubation of the trypsinized Cry1Ab protein with *O. nubilalis* or *L. botrana* BBMV (susceptible insects), but also with *L. decemlineata* BBMV (non-susceptible insect) and Sf21 cells (non-susceptible cultured insect cells). The oligomeric structures were found associated to the insect BBMV or to the insect cells, indicating that they were either inserted or bound to the membrane proteins. The finding of Cry1Ab oligomers after incubation with tolerant insect BBMV or associated with tolerant insect cells could be explained by an improper insertion of oligomers into the membranes (and therefore inefficiency to produce damages), or by the inability to induce the post-pore subsequent events in the cells (e.g., no triggering of cell death mechanisms).

In this study, the Cry1Ab oligomer was mainly found to be associated to the membrane fractions in accordance with previous reports, but oligomers were also observed in the supernatants after incubation with BBMV from the non-susceptible insect *L. decemlineata*. This suggested that the Cry1Ab oligomers (tetramers and dimers), promoted by the coleopteran BBMV, are not only associated to the BBMV (whether inserted or not) but also free in the supernatant (Figure 3a, lane S). The results obtained resemble the ones shown by Rodríguez-Almazán et al. [47], with the Cry1Ab helix α-4 mutants which had a mutation in domain I, involved in membrane insertion and pore formation, and had lost drastically their insecticidal activity towards *M. sexta* larvae. Similarly, in our results, the incubation of Cry1Ab with BBMV from *L. decemlineata* rendered a relatively high amount of oligomeric structures (dimers and tetramers) in the supernatant, apparently indicating that these oligomers were not able to insert into the membranes, resulting in the absence of toxicity. Nevertheless, in this study, after the incubation of the Cry1Ia protein with *L. decemlineata* BBMV, a high proportion of Cry1Ia oligomers were also observed in the supernatants (Figure 3b, laneS), and in this case, the protein showed a high toxicity against this coleopteran pest (Table 1).

The Cry1Ia oligomers were observed after incubation of the protein with BBMV from the susceptible coleopteran *L. decemlineata*. The molecular weights of the bands that correspond to the Cry1Ia oligomers (about 250 kDa) point out that the Cry1Ia oligomer could be composed of more than four units.

After incubation of Cry1Ia with BBMV of susceptible lepidopterans (*O. nubilalis* and *L. botrana*) or with insect Sf21 cells, no oligomers were detected, suggesting that the Cry1Ia proteins associated to the lepidopteran membranes could be mainly in monomeric forms. In this case, oligomers would not be a limiting step in the toxicity of Cry1Ia proteins, and the toxicity could be mediated by intracellular signaling pathways [9]. Moreover, other hypotheses could be mentioned to explain the reason for the absence of Cry1Ia oligomers. Firstly, it can be suggested that the observed monomers come from disassembled oligomers produced due to the SDS-PAGE technique conditions, similarly to what was proposed by Ocelotl et al. [51] working with Cry1Ab oligomers. However, this reasoning would be in conflict with the observation of oligomers after the incubation of Cry1I with coleopteran BBMV, which were detected using the same SDS-PAGE conditions. Secondly, it could be considered that the biotinlylation of Cry1I could interfere with oligomerization. This hypothesis was examined following the incubation of the biotin labelled Cry1Ab with *O. nubilalis* BBMV. The results showed no influence of biotin in oligomerization (Figure S1), similarly to what had been already claimed by other authors showing that biotinylation of Cry proteins does not prevent their oligomerization, binding and insecticidal activity [12,53,54]. Thirdly, it has to be noticed that, in the present study, the oligomer formation experiments were performed with trypsinized Cry1Ab and Cry1Ia proteins to mimic the in vivo conditions. It has been described that Cry1Ie protoxin and trypsinized protein have the same toxicity [28]. However, other Cry1I proteins such as Cry1Ia1 have shown some differences in toxicity amongst protoxins (more active) and trypsinized proteins [49]. We can speculate that in vitro trypsinization of Cry1Ia could alter the toxicity by impairing oligomer formation, maybe provoking a flawed ability to form oligomers when they are promoted by lepidopteran BBMV.

The spontaneous formation of Cry1Ab and Cry1Ia oligomers in solution has been questioned in this study. Cry1Ab submitted to the same experimental situation of treatments, but without being in contact with insect BBMV or cells, did not oligomerize (Figures 1, 2, 3 and 4a, lanes C). Regarding Cry1Ia, some bands of smaller sizes than the expected tetramer, and with MW sizes that were not multiples of 50 kDa (monomer size) were observed (Figures 1, 2, 3 and 4b, lanes C). Most probably, these bands are residual incomplete trypsinized Cry1Ia forms, dragged through the toxin purification process. In conclusion, although some studies have reported the spontaneous formation of Cry protein oligomers in solution without being in contact with the membrane-like environment (i.e., Cry4Ba [55] or Cry2Ab [24]), in our study neither Cry1Ab nor Cry1Ia formed oligomers without being exposed to insect BBMV or cultured insect cells.

In summary, our findings indicated that the oligomers of a classical 3D crystal forming Cry protein of Bt such as Cry1Ab were promoted and could be detected after incubation of activated Cry1Ab with susceptible or non-susceptible insect midgut BBMV and with non- susceptible Sf21 cells. In contrast, in the same assay conditions, Cry1Ia oligomers were detected only after incubation with BBMV of *L. decemlineata* (coleopteran susceptible host), but no Cry1Ia oligomeric structures were found following incubation of Cry1Ia with lepidopteran BBMV or with the Sf21 cells, regardless of its host susceptibility. Hence, our results, using trypsin processed Cry1Ab and Cry1Ia as an in vitro model of what might occur in vivo, suggest that: (1) The promotion of oligomers can occur by incubation of the Cry toxin with susceptible insect BBMV but also with non-susceptible insect tissues, and (2) The oligomerization may not be a determining step in the toxicity of Cry1I proteins.

#### **4. Materials and Methods**

#### *4.1. Production and Purification of Cry Proteins*

The Cry1Ab protein used for oligomer formation was obtained from a recombinant *Escherichia coli* strain GG094-208 (kindly supplied by Dr. R.A. de Maagd, Wageningen University, The Netherlands). Protein expression, inclusion bodies purification, solubilization and protoxin activation by trypsin, were performed as described previously [56]. The activated Cry1Ab was purified by anion-exchange chromatography using Äkta 100 explorer system (GE Healthcare, Amersham, UK) following Crava et al. [57]. The eluted fractions from the column were individually analysed by sodium dodecyl sulphate-12% polyacrylamide gel electrophoresis (SDS-PAGE).

The *cry1Ia7* gene was cloned and expressed in *E. coli*, BL21(DE3) cells [40]. The purification of the protein by affinity chromatography using a HisTrapTM FF crude column (GE Healthcare Bio-Sciences, Upsala, Sweden), protein dialysis and protoxin activation by trypsin, were performed as reported by Khorramnejad et al. [40]. The activated Cry1Ab and Cry1Ia were visualized after SDS-PAGE, and their concentration was estimated by densitometry using TotalLab Quant program version 12.3 (Newcastle, UK), employing bovine serum albumin as standard.

The Cry1Ab protein used in bioassays (protoxin) was obtained from a recombinant Bt strain that produced a crystal composed solely of the Cry1Ab protein, kindly supplied by Dr. Colin Berry, Cardiff University, Cardiff, UK. This Bt strain was grown in CCY medium [58] supplemented with erythromycin and the crystal formation was observed daily. When at least 95% of the cells were lysed, the fermentation process was stopped and then spores and crystals were collected by centrifugation (8600× *g*, 4 ◦C), washed with a saline solution (NaCl 1M, EDTA 10mM) and resuspended in KCl (10 mM). For Cry1Ia7 protoxin production, the recombinant *E. coli* BL21 (DE3) was grown and purified as described above. Both Cry1Ab and Cry1Ia7 protoxins were quantified by the method of Bradford [59] using BSA as standard, and kept at 4 ◦C until used.

#### *4.2. Insect Rearing*

Two different lepidopteran species, *L. botrana* (Lep: Tortricide) and *O. nubilalis* Hübner (Lep.: Crambidae), and one coleopteran species, *L. decemlineata* (Col.: Chrysomelidae) (the Colorado potato beetle, CPB) were used in this study. Both lepidopteran pests were maintained in the insectary of the Universidad Pública de Navarra (UPNA, Pamplona, Spain) at 25 ◦C ± 1 ◦C with 70% ± 5% RH and a photoperiod of 16/8 h (light/dark) on the artificial diet described by MacIntosh et al. [60]. The population of beetles was raised on potato plants (Desiré variety) grown throughout the year in a phytotron to provide a continuous supply of food substrate. This population was refreshed 1–2 times every year by the introduction of wild adults collected in the field during the spring-summer.

#### *4.3. Insect Cell Line*

The lepidopteran cell line Sf21, from ovaries of fall armyworm *S. frugiperda* (Lep.: Noctuidea), was obtained from Wageningen University (Wageningen, The Netherlands). The Sf21 cells were maintained at 25 ◦C on 1X Grace's medium (Gibco® Life technologiesTM, Carslbad, CA, USA) supplemented with 10% heat-inactivated fetal bovine serum (FBS) in 25 cm<sup>2</sup> cell culture flasks. Cells were passaged weekly. Cell concentrations were measured by using an automatic cell counter (Countess Automated Cell Counter from Invitrogen, Carlsbad, CA, USA).

#### *4.4. Insect Bioassays*

The bioassays for the three insect species were performed with first instar larvae. Five different protoxin concentrations, ranging from 0.39 to 100 μg/mL, were prepared to determine the concentration-mortality responses in order to calculate the mean lethal concentration (LC50). For the lepidopteran insect species, *O. nubilalis* and *L. botrana*, the diet surface contamination assay was used [61], and for *L. decemlineata*, the leaf dip bioassay described by Iriarte et al. [62], was performed. For each bioassay, several protein concentrations were tested, using 28 larvae per concentration. Each bioassay was repeated at least three times. Total insect mortality was recorded after 7 days for lepidopteran insects, and after 4 days for *L. decemlineata*. The concentration-mortality data obtained for each insect species were analyzed after transformation of the concentration-response curve to fit a linear model using POLO-PC program (LeOra Software, Berkeley, CA, USA, 1987), based on the Probit analysis [63].

#### *4.5. Midgut Isolation and BBMV Preparation*

Midguts were dissected from fifth-instar larvae of *O. nubilalis* and forth-instar larvae of *L. decemlineata*. The dissected midguts were rinsed in ice-cold MET buffer (0.3 M mannitol, 5 mM EGTA, 17 mM Trsi-HCl, pH 7.5), snap frozen in liquid nitrogen and kept at −80 ◦C until use.

Brush border membrane vesicles (BBMV) were obtained from the dissected midguts of *O. nubilalis* and *L. decemlineata* and from the whole last instar larvae of *L. botrana*, following the differential magnesium precipitation method [64,65]. Proteins in the purified BBMV were quantified following Bradford protein assay [59] and stored at −80 ◦C.

#### *4.6. Biotin Labelling*

Trypsin activated Cry1Ia protein was biotinylated by using the protein biotinylation kit from GE Healthcare (GE Healthcare, Little Chalfont, UK), as described elsewhere [66]. The eluted fractions were quantified by NanoDrop 2000 spectrophotometer (ThermoFisher Scientific, Waltham, MA, USA), analyzed by 12% SDS-PAGE and verified by Western blot. The protein fractions were concentrated by using an Amicon Ultra-4 10K centrifugal filter device (Merck Millipore, Tullagreen, Ireland) and stored at 4 ◦C.

The interference of biotin in oligomerization was tested following incubation of the biotin labelled Cry1Ab with *O. nubilalis* BBMV. The detection of biotin labelled Cry1Ab oligomerization was performed following the same protocol than has been described for Cry1Ia. The results showed no influence of biotin in oligomerization (Figure S1), as had been already claimed by other authors that have used biotin labelled proteins to detect oligomers [12,20,53,54].

#### *4.7. Oligomerization Assays with Sf21 Cells*

The confluent monolayer growing Sf21 cells were suspended in fresh Grace's medium without FBS. The cell concentration was measured (by using the Countess Automated Cell Counter from Invitrogen, Carlsbad, California, USA), and 100 <sup>μ</sup>L of cell suspension at a concentration of <sup>2</sup> <sup>×</sup> <sup>10</sup><sup>6</sup> cells/mL were seeded into 96-well plates. The oligomerization assays were performed as described by Portugal et al. [38] with slight modifications. In short, cells were incubated at 25 ◦C for at least 30 min. Later, 4 μg of activated toxins (biotinylated Cry1Ia or unlabeled Cry1Ab) were added to the cells (final concentration of 0.03 μg protein/μL) except in controls, which received 50 mM carbonate buffer pH 10.5. The plates were incubated for 3 h at 25 ◦C. After the incubation, the treated cells were collected and pelleted by centrifugation at 16,200× *g*, 4 ◦C for 15 min. The supernatants containing unbound proteins were kept for further analysis. The controls (proteins alone, without cells) were submitted to the same experimental conditions as treatments. After centrifugation of the controls, as there was no pellet due to the absence of cells, a dilution of supernatant (containing 200 ng of selected protein) was analyzed in the gel. The Sf21 cells in the pellet were washed once with 200 μL of 50 mM carbonate buffer pH 10.5, and recovered by centrifugation (45 min, 18,800× *g*). The final pellet was resuspended in 10 μL of buffer and heated at 50 ◦C for 3 min. The proteins present in the sample were separated by SDS-PAGE 10% and electrotransferred onto polyvinylidene difluoride (PVDF) Western Blotting membrane (Roche Diagnostics GmbH, Mannheim, Germany). The membrane was incubated overnight in blocking buffer (PBST; 0.1% Tween 20 in phosphate-buffered saline, supplemented with 5% skimmed milk) with gentle shaking, and washed three times with PBST, before incubation with the corresponding antibodies. Cry1Ab protein was detected with polyclonal rabbit anti-Bt Cry1Ab/1Ac (1:10,000; 60 min) from Abraxis (Warminster, PA, USA) followed by secondary antibody (1:20,000; 60 min) coupled with horseradish peroxidase (HRP) (Sigma-Aldrich, Saint Louis, MO, USA), whereas biotinylated Cry1Ia protein was detected by streptavidin-conjugated horseradish peroxidase (1:2000; 60 min) (GE Healthcare, Amersham, UK). Both Cry1Ab and Cry1Ia proteins were visualized by chemiluminescence using ECLTM prime western blotting detection reagent (GE Healthcare, Little Chalfont, UK) using an ImageQuant LAS400 image analyzer (GE Healthcare Bio-Sciences, Upsala, Sweden). The molecular weight marker used was Precision Plus Protein™ Dual Color Standard (Bio-Rad, Carlsbad, CA, USA). Each oligomerization assay was repeated at least three times.

#### *4.8. Oligomerization Assays with BBMV*

The BBMV were centrifuged for 10 min at 16,000× *g* and suspended in 50 mM carbonate buffer pH 10.5. The oligomerization protocol was set up after reviewing the procedures described in the literature for Cry1 proteins [17,18,37–39,45,47,51,52,54]. Finally, the oligomerization assays with BBMV were performed following Ocelotl et al. [51] who employed the milder SDS-PAGE denaturing conditions (heating the samples 3 min at 50 ◦C), with some modifications. Briefly, 2 μg of biotin labelled activated Cry1Ia and activated Cry1Ab toxins were incubated for one hour with 5 μg of *L. botrana* or *L. decemlineata* BBMV, or with 20 μg of *O. nubilalis* BBMV, at 37 ◦C, in a final volume of 50 μL. Activated proteins incubated in the absence of BBMV and samples containing only BBMV were used as controls. Then, phenylmethylsulfoyl fluoride (PMSF) was added (final concentration 1 mM) and BBMV were recovered by centrifugation at 18,400× *g* for 45 min at 4 ◦C. The supernatant containing unbound protein was recovered and stored. The controls (samples with proteins and without BBMV) went through the same experimental conditions as treatments. After centrifugation of the controls, a dilution of the supernatant (containing 200 ng of selected protein) was analyzed in the gel to avoid the observation of the over saturated signal in the membrane. In the samples containing BBMV, the pellet was washed once with 100 μL ice-cold buffer. The final BBMV pellets were resuspended in 10 μl of the buffer. After incubating the samples for 3 min at 50 ◦C, the proteins were separated by 10% SDS-PAGE and blotted onto PVDF Western Blot membranes (Roche Diagnostics GmbH, Mannheim, Germany). After Western blot, Cry1Ab was detected with polyclonal rabbit anti-Bt Cry1Ab/1Ac, and Cry1Ia was detected by streptavidin-conjugated horseradish peroxidase as has been described in the previous section. The experiments were repeated, at least, three times.

**Supplementary Materials:** The following is available online at http://www.mdpi.com/2072-6651/12/2/133/s1, Figure S1: Biotin labelled Cry1Ab oligomer formation promoted by *O. nubilalis* BBMV.

**Author Contributions:** B.E. and Y.B. contributed to the design of the study. A.K. and M.D.-A. performed the experiments. A.K., Y.B., B.E., P.C. and M.D.-A. analyzed the data. A.K., Y.B. and M.D.-A. wrote the manuscript. All authors have read and agreed to the published version of the manuscript.

**Funding:** This work was supported by grants from the Spanish Ministry of Science, Innovation and Universities, the State Research Agency of Spain and the European FEDER founds (Refs. AGL2015-70584-C2 and RTI2018-095204-B-C21), and by the Generalitat Valenciana (GVPROMETEOII-2015-001). M. Domínguez received a predoctoral fellowship from the Universidad Pública de Navarra, Spain.

**Acknowledgments:** We are deeply grateful to Patricia Hernández-Martínez for her valuable comments. We thank Rosa Maria González-Martínez and Óscar Marin Vázquez for their support in laboratory assistance.

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**


© 2020 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).

#### *Article*

## **A** *Bacillus thuringiensis* **Chitin-Binding Protein is Involved in Insect Peritrophic Matrix Adhesion and Takes Part in the Infection Process**

**Jiaxin Qin 1,**†**, Zongxing Tong 1,**†**, Yiling Zhan 1,2, Christophe Buisson 3, Fuping Song 2, Kanglai He 2, Christina Nielsen-LeRoux 3,\* and Shuyuan Guo 1,\***


Received: 10 March 2020; Accepted: 10 April 2020; Published: 13 April 2020

**Abstract:** *Bacillus thuringiensis* (Bt) is used for insect pest control, and its larvicidal activity is primarily attributed to Cry toxins. Other factors participate in infection, and limited information is available regarding factors acting on the peritrophic matrix (PM). This study aimed to investigate the role of a Bt chitin-binding protein (CBPA) that had been previously shown to be expressed at pH 9 *in vitro* and could therefore be expressed in the alkaline gut of lepidopteron larvae. A ΔcbpA mutant was generated that was 10-fold less virulent than wild-type Bt HD73 towards *Ostrinia furnacalis* neonate larvae, indicating its important role in infection. Purified recombinant *Escherichia coli* CBPA was shown to have a chitin affinity, thus indicating a possible interaction with the chitin-rich PM. A translational GFP–CBPA fusion elucidated the localization of CBPA on the bacterial surface, and the transcriptional activity of the promoter P*cbpA* was immediately induced and confirmed at pH 9. Next, in order to connect surface expression and possible *in vivo* gut activity*,* last instar *Galleria mellonella* (Gm) larvae (not susceptible to Bt HD-73) were used as a model to follow CBPA in gut expression, bacterial transit, and PM adhesion. CBPA-GFP was quickly expressed in the Gm gut lumen, and more Bt HD73 strain bacteria adhered to the PM than those of the ΔcbpA mutant strain. Therefore, CBPA may help to retain the bacteria, via the PM binding, close to the gut surface and thus takes part in the early steps of Bt gut interactions.

**Keywords:** *Bacillus thuringiensis*; chitin-binding protein; adhesion; peritrophic matrix

**Key Contribution:** Our findings add a new step to the understanding of *Bacillus thuringiensis* (Bt) infection and suggest that the chitin binding protein A (CBPA) plays an important role in the first steps of the infection for insects where the Bt bacteria itself plays a role, which is for instance the case for *Ostrinia*. Indeed, due to a better bacterial binding to the larval peritrophic matrix (PM), it may increase the efficacy of the gut cell and PM damaging virulence factors produced from the outgrown bacteria.

#### **1. Introduction**

*Bacillus thuringiensis* (Bt) is a prominent insect pathogen of the *Bacillus cereus* group. Several strains are used worldwide as microbial control agents against major agricultural and forest insect pests [1]. The primary insecticidal factors of Bt are the Cry toxins, which comprise parasporal protein crystals

produced by Bt, and numerous studies have focused on the structural resolution of the crystals and on the mode of action of Cry toxins [2]. As an insect pathogen [3], the roles of Bt itself in pathogenesis has been much less investigated and may depend on the insect species, larval stage, and Bt strain.

For pathogenic bacteria, the successful establishment of infection generally requires adhesion, colonization, and host cell degradation or active invasion. The capacity for host cell and tissue adherence is a key feature of pathogenic bacteria [4,5]. Orally acting entomopathogenic bacteria including Bt face the peritrophic matrix (PM) just after ingestion. The PM is an important component of the insect digestive tract: It serves both as a physical barrier to separate food particles, digestive enzymes, and pathogens, and it serves as a biochemical barrier, sequestering or even inactivating ingested toxins [6]. Therefore, Bt must bypass the PM barrier to establish persistent infections [7] in order to develop and complete the process of infection and life cycle, ending with sporulation in the insect cadaver [8,9].

The PM is a chitin- and glycoprotein-rich matrix, separating intestinal cells from the gut content [10]. Chitin is a linear polymer of N-acetylglucosamine (GlcNAc) linked via β-1,4 linkage [11]. Chitinases can hydrolyze chitin, thus fragmenting the PM and suggesting that chitinases may be part of the enzymes involved in the degradation of the PM [12]. Chitinases can enhance the insecticidal activity of Bt, irrespective of chitinase activity derived from a chromosomal gene, the co-expression of chitinase with a Cry toxin gene, or even from the addition of commercial chitinases [13–16]. The most probable role of the endogenous chitinases of Bt is to weaken the integrity of the insect PM, facilitating the better access of the bacterial toxins and the bacteria to the gut epithelia [17].

Chitin-binding proteins (CBP) are present in numerous microorganisms. They belong to the 14, 18, or 33 families of the carbohydrate-binding domain proteins [18]. Various microorganisms simultaneously synthesize chitinases and CBPs [19]. The subcellular localization of CBP differs in accordance with the organism, most of them being secreted proteins [19,20]. Structural analyses have revealed the presence of the aromatic amino acid residues exposed on most CBPs, which play an important role in substrate binding [21–25]. From viruses to invertebrate organisms, CBP participates in various biological processes in different species, such as antifungal activity [26], synergistic effects with chitinase [19], and the detection of hydrophobic surfaces [27].

The alkaline pH of the midgut—in lepidopteron larvae, in particular—is needed for Bt to exert insecticidal activity, since an alkaline pH permits the solubilization of several Cry toxin crystals [1]. Hence, it is important for bacteria to adhere to host tissue and survive in this alkaline environment in order to pursue infection. Transcriptome gene microarray data previously indicated that *cbp3189* is up-regulated more than eight-fold after alkaline induction [28]. The protein encoded by *cbp3189* is referred to as chitin binding protein A (CBPA) in this study. CBPA is a conserved protein in Bt strains, as its amino acid sequence homology in 34 different Bt strains is greater than 97%, and, among them, nine strains have a 100% sequence identity. This gene encodes a protein containing a signal peptide and a transmembrane structure, and localization prediction has revealed that CBPA may be localized on the bacterial cell wall [29].

In this study, we addressed several questions related to the function, expression, and localization of CBPA. First, to determine whether CBPA plays a role during infection, a ΔcbpA mutant was constructed and assessed for virulence in *Ostrinia furnacalis* and *Galleria mellonella* larvae. Further, its subcellular localization in Bt and the activity of its promoter were assessed during in vitro growth. CBPA was expressed and purified from *Escherichia coli* through binding to chitin beads, and it was further analyzed for chitinase activity. Finally, the interaction of CBPA with the gut and the PM in vivo was assessed in *G*. *mellonella*, with a focus on the early stages of infection. Our results may provide functional insights into the role of CBPA in adhesion to the PM, thus improving the current understanding of the mode of action of Bt insecticidal strains, particularly for insects where the action of Cry toxins, spores, and out-grown bacteria are important for full virulence.

#### **2. Results**

#### *2.1.* Δ*cbpA Mutant,* Δ*cbpA::cbpA-Complemented Strain Construction*

To determine the role of CBPA in insect infection, an interruption mutant strain ΔcbpA mutant was constructed via homologous recombination. Cry1Ac protein expression levels were not changed in the ΔcbpA mutant in comparison with wild-type HD73 upon protein quantification (Figure 1A). Spore count results showed that the wild-type strain and ΔcbpA mutant contained equal CFU values (Figure 1), indicating that the absence of *cbpA* did not influence spore formation.

**Figure 1.** Comparison of Cry protein expression and spore formation in *Bacillus thuringiensis* (Bt) HD73 wild-type strain and ΔcbpA strain. (**A**) Cry protein production analyzed via SDS-PAGE. (**B**) Spore counts. 1. Bovine serum albumin (BSA) (1 μg); 2. BSA (5 μg); 3. BSA (10 μg); 4. Cry1Ac protein in wide-type (10-μL spore crystal suspension); 5. Cry1Ac protein in deletion mutant (10-μL spore crystal suspension). "a" indicates there was no significant difference (*p* > 0.05).

#### *2.2. Role of CBPA in Ostrinia Furnacalis and Galleria Mellonella Mortality*

As the Bt HD73 produces Cry1Ac that is toxic against the Asian corn borer (*Ostrina*), we selected this insect to evaluate the difference in mortality of larvae fed a diet supplemented with spore-crystal suspensions of wild-type or ΔcbpA mutant HD73 at various concentrations. Mortality induced by the ΔcbpA mutant was lower than that of the wild-type strain (Figure 2 and Table 1). Figure 2 shows the comparison of mortality rates between the wild-type HD73 strain and the ΔcbpA mutant strain for seven days post-feeding. For both strains, almost no change in mortality was observed after the fourth day of feeding. Mortality rates on the seventh day after feeding with seven different concentrations are listed in Table 1. The mortality between the wild-type and mutant strains was significantly different from the third concentration (Cry1Ac protein: 0.05 <sup>μ</sup>g/g; spore: 5.4 <sup>×</sup> 107/g).

**Figure 2.** Comparison of mortality rates between the wild-type HD73 strain and the ΔcbpA mutant strain against the Asian corn borer (Cry protein: 2.5μg/g diet; spore: 2.7 <sup>×</sup> <sup>10</sup>9/g diet).


**Table 1.** Mortality of crystal and spore mixture against Asian corn borer larvae.

\* Means within a line were significantly different (*p* ≤ 0.01) via the t-test. \*\* significance: *p* = 0.000130. \*\*\* significance: *p* = 0.000478.

Further, the difference in LC50 was evaluated by administering larvae with a diet supplemented with spores of the HD73 wild-type strain, the ΔcbpA mutant, or Δ*cbpA*::*cbpA*-complemented strains at different doses (104–108 CFU/per gram diet) and with the concentration of Cry1Ac at 0.01 μg/g diet. Table 2 shows the LC50 values of the three strains. Deionized water was used as the negative control, and the mortality was 2% in this control. The inferred larval mortality of the ΔcbpA mutant was significantly lower than that of the wild-type strain, while that of the complemented strain reverted to levels of the wild-type strain. These data indicate that CBPA significantly contributes to Bt virulence in *Ostrinia*.


**Table 2.** LC50 values of different strains against Asian corn borer.

˅

Each concentration of spore was mixed with Cry1Ac at a final concentration of 0.01 ug/g.

To further investigate the role of CBPA, we tested the mutant for virulence towards *G. mellonella*; this insect needs bacteria associated with Cry1Ca for complete virulence in the synergy model [30,31]. Therefore, *G. mellonella* is a suitable model to elucidate the role of Bt and *B*. *cereus* chromosomal carried factors, and large last instars are easy to manipulate for accurate feeding and for dissection. Infections were induced through controlled force-feeding at a dose of 5 <sup>×</sup> <sup>10</sup><sup>6</sup> mid log-phase vegetative bacteria (OD600 = 1) or with spores associated with 3 μg of activated Cry1Ca toxin for each larva, as described previously [32]. The results (Figure S1) showed no differences in mortality between infection with wild-type HD73 and the ΔcbpA mutant strains under all tested conditions. Indeed, no mortality was observed with spores or log-phase bacteria alone, and 90–100% mortality was observed when associated with 3 μg of Cry1Ca for both strains. Therefore, under these infection conditions, no clear role of CBPA was elucidated in Bt HD-73 mortality towards *G. mellonella* last instars.

#### *2.3. Localization of CBPA in Bt HD-73*

GFP-conjugated CBPA was used to investigate the subcellular localization of CBPA in HD73 bacteria cells. Samples were harvested at different growth stages (T4, T7, T8, and T10 after the onset of the stationary phase). GFP expression was visualized via laser-scanning confocal microscopy. The cell membrane was stained with FM 4-64 dye solution. Red fluorescence indicated the cell membrane, while green fluorescence indicated the expression of the GFP–CBPA fusion protein. No green fluorescence was detected during early growth; however, it stabilized from T8 onwards (Figure 3A). Figure 3B shows the green fluorescence at T12. Fluorescence observed on the bacteria cell surfaces was indicated by a yellow arrow and on the prespore surface with a red arrow. Consistent with the in silico predictions, GFP fusion experiments revealed that CBPA was located on the cell surface.

**Figure 3.** *Cont*.

**Figure 3.** Green fluorescence detection of the chitin binding protein A (CBPA)–GFP fusion at different stages of culturing, observed via laser-scanning confocal microscopy. (**A**) Stages T4, T7, T8, and T10. (**B**) Localization of CBPA (T12). Yellow arrow denotes green fluorescence on the bacterial cell surface. Red arrow denotes green fluorescence on the spore surface. GFP (green fluorescent protein) signal in the bacterial cytosol. FM 4-64, (red fluorescent signal of bacterial membrane stain). The overlay shows green and red fluorescent signals. PC: phase-contrast microscopy.

#### *2.4. Analysis of cbpA Promoter Activity under Alkaline Induction*

To investigate the effect of alkali on CBPA expression, we selected an early culture stage wherein the CBPA protein was not expressed. Both the *cbpA-gfp* transduction fusion and the transcriptional activation of the *cbpA* promoter-*lacZ*fusion were analyzed under similar growth conditions. The bacteria were cultured to the late exponential growth stage up to an OD600 nm between 1.5 and 2.0, and an NaOH solution was added to yield a final concentration of 24 mM (pH 9). Samples were maintained under this alkaline environment for 15 and 30 min. GFP expression was then visualized via laser-scanning confocal scanning microscopy (Figure 4A). Cells not treated with the NaOH solution were considered as the negative control. GFP was expressed after alkaline induction (pH 9) but not in the negative control, indicating that CBPA protein expression was rapidly induced under alkaline conditions. In the transcriptional *cbpA* promoter-*lacZ* fusion strain, β-galactosidase activity significantly increased after the addition of NaOH (15 or 30 min) in comparison with non-induced conditions (Figure 4B). Consequently, the cytological observation of the GFP fusion strain and the enzymatic activity analysis in P*cbpA*-*lacZ* promoter fusion strain yielded consistent results, indicating that CBPA expression was induced at an alkaline pH.

**Figure 4.** Analysis of transcriptional activity. (**A**) Observation of alkaline induction via laser-scanning confocal fluorescence microscopy. Row 1: non-induced for 15 min. Row 2: induced under alkaline conditions for 15 min. Row 3: non-induced for 30 min. Row 4: induced under alkaline conditions for 30 min. (**B**) Analysis of β-galactosidase activity of the P*cbpA*-*lacZ* fusion +/- alkaline induction.

#### *2.5. Chitin Binding Ability and Chitinase Activity of CBPA*

To follow-up on the in silico information indicating CBPA as a chitin binding protein, the next step was to analyze if CBPA really has chitin binding capacity. Therefore, we expressed CBPA as a heterologous recombinant protein. *cbpA* (gene 3189) from Bt HD73 was cloned and expressed in an *E. coli* BL21/DE3 strain. The expected size of the HD73-CBPA protein was 49.78 kDa. Protein expression was induced through an isopropyl-β-D-thiogalactopyranoside (IPTG) gradient, and expression was assessed via SDS-PAGE as a ~50-kDa protein at 0.4–1.0 mM IPTG (Figure 5A). Thereafter, CBPA was purified via chitin affinity chromatography and eluted at a gradient of 0.4 M NaCl, thus showing the chitin-binding capacity of CBPA (Figure 5B). The purified protein band was excised and analyzed via matrix-assisted laser desorption/ionization and time-of-flight peptide mass spectrometry analysis after in-gel digestion, confirming that the heterologous *E. coli*-cloned and -expressed CBPA protein contained the expected peptide composition. Having confirmed its chitin-binding capacity, we assessed for the

eventual chitinase activity of the protein. CBPA displayed no chitobiosidase and endochitinase activity (Table 3). Indeed, the fluorescence intensity of chitinase degradation products from various substrates obtained with CBPA approached the same values as those of the negative control. Therefore, CBPA can probably not degrade chitin-rich structures, at least those analyzed herein.

**Figure 5.** Expression and purification of CBPA proteins harvested for SDS-PAGE analysis. (A) Lane 1: the non-induced expression of CBPA in *E. coli* BL21/DE3. Lanes 2-4: induced expression of HD73-*cbpA* by the isopropyl-β-D-thiogalactopyranoside (IPTG) gradient of 0.4, 0.7 mM, and 1.0 mM. (B) purified CBPA eluted by a gradient of 0.4 M NaCl.


**Table 3.** Fluorescence value of chitinase activity.

#### *2.6. Expression of CBPA-GFP Fusion in Vivo in G. mellonella*

Despite the lack of an evident role of CBPA in virulence in the final instar of *Galleria* larvae, we aimed at determining the possibility of CPBA to bind the PM since chitin is a structural element of the PM in all insects. First, we investigated whether CBPA was expressed in the *Galleria* gut, since the aforementioned *in vitro* studies (Figures 3 and 4) indicated that CBPA was expressed on the surface of HD73 cells and under alkaline pH, which may occur in the *Galleria* larval gut. The pH of the *Galleria* midgut was measured via the injection of a liquid pH indicator into three sites of the gut; the pH was between 8.5 and 9 from the anterior to posterior midgut, as directly observed under binoculars with four times magnification. Thereafter, we assessed, via epi-fluorescence microscopy, the presence of fluorescent bacteria from the anterior and posterior midgut of *Galleria* larvae infected with mid log-phase Luria–Bertani (LB) grown vegetative HD73 bacteria carrying the CBPA–GFP plasmid fusion protein. Observations were recorded at 1 and 4 h post-ingestion. The CFU values and the fluorescence intensities were scored upon arbitrary visual observation (Figure 6). Greater CFU values were recovered at 1 than at 4 h post-ingestion (Figure 6A), indicating a relatively rapid intestinal transit and that fluorescent bacteria (Figure 6B) were more abundant at the early time point.

**Figure 6.** Arbitrary scores of bacteria (**A**) and the expression of the CBPA–GFP fusion protein (**B**) recovered in the *Galleria mellonella* intestine. At one hour and 4 h post-force-feeding with a wild-type HD73 (pHT*cbpA*-*gfp*) strain, samples from the anterior midgut and posterior midgut were analyzed via fluorescence microscopy at 1000× magnification from five chilled, dissected larvae. Scores are as follows: 0 = no bacteria and no fluorescence, 1 = few bacteria <10 per field, 2 = between 10 and 50 bacteria and 3 = more than 50 bacteria per observation field.

#### *2.7. HD73 and HD73* Δ*cbpA Intestinal Transit and Localization Assays*

Since purified CBPA can bind to chitin, is expressed on the bacterial surface (Figures 3 and 5), and is activated in the gut of *Galleria*, we performed a tight analysis of the persistence of the HD73 wild-type and ΔcbpA mutant strains with vegetative bacteria, presuming that the PM binding capacity of CBPA *in vivo* would lead to a difference between the wild-type and the mutant strains during intestinal transit.

First, the presence of bacteria was estimated in whole larvae and dissected whole intestines (gut with the PM) (Figure 7A,B) at three time points. Immediately after ingestion (T0), no difference (≈5000 CFU) was observed between wild-type HD73 and ΔcbpA mutant strains, while at T3 h post-ingestion, a significant difference was observed between wild-type HD73 (≈100 CFU) and the ΔcbpA mutant (approximately 5000 CFU were still observed). At 24 h, no bacteria were observed in larvae fed with wild-type HD73, and approximately 100 CFU were recorded for larvae infected with the ΔcbpA mutant. A similar analysis was performed with the dissected whole intestines (gut and the PM) (Figure 7B). No difference was observed at T0; however, at T3 h post-ingestion, almost no bacteria were observed with the wild-type HD73 strain, while approximately 3000 CFU were still observed with the ΔcbpA mutant strain. Thus, bacteria not expressing CBPA persist longer in the gut than the wild-type bacteria, suggesting that the wild-type HD73 bacteria are more easily excreted with the PM during natural food bolus transit, resulting in feces production. The feces are surrounded by the PM [6].

Therefore, we further analyzed the speed of transit after force-feeding with spores in order to uncover the time where we would still find all bacteria in the insect before they would be excreted with the feces. Feces were collected and assessed for the presence of bacteria, and the mean numbers of feces per larvae were recorded at four time points. Feces were observed at 2 h (0.3 feces/larvae) and displayed an increase in the mean number of feces at 3 and 4 h to 1.5 feces per larva. The presence of the bacteria was found in feces from the 2 to early 3 h post ingestion, showing that the mean transit time under these conditions was approximately 2 h. Based on these observations, we thereafter tested for the presence of the bacteria adhering to the PM. Hence, we selected the time point of 1 h post-ingestion, since feces were excreted at 2 h per the aforementioned results and since at 3 h post-ingestion, only a few residual bacteria were observed in the wild-type HD73-treated larvae (Figure 7A,B). The CFU value recovered from the dissected PM alone (Figure 7C) was approximately 5000 for wild-type HD73 and three-fold lesser for the ΔcbpA mutant, which was significantly different from the wild-type. In addition, the *cbpA*-complemented ΔcbpA mutant recovered a better adhesion to the PM. One way ANOVA analysis and the Bonferroni's multiple comparison test showed significant differences

between the PM from the ΔcbpA mutant and wild-type HD73, while the complemented strain PM had no significant differences with the others. The observed variations in CFU that were associated with the dissected intestine, separated intestine, and the PM may have been due to the difficulty of the dissection approach (see Materials and Methods). The results indicated that CBPA *in vivo* has an affinity for the PM (Figure 7C) and therefore may help retain the bacteria to this tissue, which could then increase the infection efficacy of Bt.

**Figure 7.** Presence of bacteria in *Galleria mellonella* whole larvae (**A**) dissected complete intestine (**B**), and dissected dissociated intestine and the PM (**C**) from chilled, fifth-instar larvae after force-feeding at stages T0, T3, and T24 h post-feeding for (**A**,**B**) and after 1 h for (**C**). Whole larvae, the larval intestine, and the PM were homogenized to determine the CFU counts for each sample. Assays were repeated at least three times with two replicates per sample time and sample type. CFU counts were analyzed with the PRISM software and one way ANOVA associated with the Bonferroni's multiple comparison test. \*\* <0.01 and \* <0.05 level, ns (non-significant) different.

#### **3. Discussion**

During infection, a pathogen interacts with the host and circumvents the host's defense mechanisms. For many pathogens, the first step of colonization depends on the capacity to adhere to the host tissue via multiple factors [5]. Therefore, pathogenic bacteria produce surface molecules and appendages, such as flagella and pili. They sense host surfaces, thus facilitating their adhesion with host cells and thereby bringing secreted molecules close to the host cell targets. Pore-forming Cry endotoxins are the major *B. thuringiensis* insecticidal effectors. They bind to specific membrane receptors on the larval midgut epithelium [1,33]. Meanwhile, the spores and their outgrown vegetative form participate in infection [3]. Several proteases, lipases, chitinases, toxins, or adaptation factors are involved in pathogenesis [8,31] and in fulfilling the insect phase of the Bt life cycle. The germination and growth of *B. thuringiensis* in the gut of insect larvae have been previously photographically studied. It was reported that the spores of *B. thuringiensis* germinated at the surface of the epithelium 40–120 min after inoculation [34]. Additionally, histological studies have shown that the vegetative bacteria of *B. thuringiensis* are found in the gut lumen

of *Chrysomela* [35]. However, thus far, limited information is available regarding the role of factors associated with Bt spores and newly outgrown vegetative bacteria in the very early stages of gut infection, particularly with respect to their interaction with the PM.

This study investigated the expression and functions of a yet unknown chitin-binding protein (CBPA) from the Bt HD73 strain and explored its role in insect infection. The gene encoding CBPA was previously identified among genes activated under alkaline conditions via an *in vitro* transcriptomic screening [28], and our previous *in silico* analysis indicated its putative chitin-binding function and its presence in several *B. cereus* genomes. Herein, we investigated the spatiotemporal aspects of its expression. The confocal microscopic imaging of Bt HD73 harboring a CBPA–GFP fusion protein revealed that CBPA was expressed in vitro at the late stationary growth stage and localized on both the bacterial and prespore surfaces (Figure 3). Furthermore, when the bacteria were exposed to an alkaline pH, the protein was expressed as soon as 15 min post-induction, as revealed through both CBPA–GFP fusion and a lacZ transcriptional promoter fusion (Figure 4). These observations indicate that expression is inducible at an alkaline pH, which is known for the Lepidopteran midgut environment. This has also been reported in the case of the Bt CBP-21 chitin-binding protein [36] and is concurrent with our former transcriptome findings [28] and with the present study, wherein the CBPA–GFP protein was observed in the *Galleria* midgut at 1 h post-ingestion with vegetative bacteria.

The widespread presence of CBP proteins in bacteria and other organisms implies their importance, with chitin binding being the most common function. CBP21 from the Bt HD1 strain binds chitin in insects [36], and CBP50 from the Bt *konkukian* serotype and CBP33A from *Lactococcus lactis* bind insoluble chitin (α and β), colloidal chitin, and cellulose [37]. Furthermore, *Streptomyces* can secrete small proteins that specifically bind α-chitin [38]. ChbB produced by *Bacillus amyloliquefaciens* preferentially binds β-chitin [39]. The present study showed the capacity of recombinant purified CBPA to bind chitin, which is concurrent with previous reports with similar CBP proteins.

Some bacteria simultaneously produce CBP and chitinase, thus improving the hydrolysis efficiency of chitinases. For example, CBP24 and CBP50 produced by Bt *serovar konkukian* act synergistically with bacterial chitinases for chitin degradation [40,41], CBP21 from *Serratia marcescens* exerts a synergistic effect with chitinase on its hydrolysis efficiency [19], and CBP33A from *Lactococcus lactis* can increase the hydrolysis efficiency of chitinase Chi18A [20]. As expected, chitinase activity was not observed for CBPA itself, but, based on our findings, we may suggest that CBPA on the bacterial surface can help target the bacteria to the chitin-rich PM, where the chitinases are of particular relevance. Indeed, the present results showed that the ΔcbpA mutant adhered less well to the *Galleria mellonella* PM, thus indicating that CBPA is involved in the adhesion of Bt HD73 out-grown vegetative bacteria to the PM (Figure 7C).

CBPA localized on the cell surfaces in a manner similar to CBP21 from the Bt HD1 strain, which was also reported to be present in the spore crystal preparation [36]. CBP21 and CBPA have low global sequence homology, and an *in silico* analysis indicated that CBPA comprises the chitin-binding-3 domain, two FN3 domains, and a CBM-5-12 domain, while CBP21 comprises a peptidase M73 domain. It was speculated that Bt-CBP21 interacts with Cry1Ac to potentiate its insecticidal activity [36], which may be concurrent with an earlier observation with strain Bt HD73, wherein Cry1Ac localized at the spore surface [42]. In the present study, Cry1Ac and spores from ΔcbpA mutant displayed higher LC50 (approximately 10-fold) values against *Ostrinia furnacalis* (Asian Corn borer) neonate larvae in comparison with the wild-type Bt HD73 strain, and the *cbpA*-complemented ΔcbpA mutant strain displayed a similar LC50 value to the wild-type strain, indicating the role of CBPA in virulence in the HD73 strain. As the first barrier in the digestive tube in most insects is the PM, orally acting pathogens require factors that can interfere with the PM. In the present study, CBPA increased the adherence to the PM, thus playing a role in the early stage of infection. In the susceptible insect, the Asian corn borer, the absence of CBPA strongly reduced mortality, which was not the case for *Galleria,* where no mortality was recorded with the wild-type HD73 strain or ΔcbpA mutant strain alone. Therefore, under the present conditions, no obvious function of CBPA in virulence was discerned in *Galleria*, probably because the strong synergism with Cry1Ca [31] concealed a rather subtle bacterial effect or

because Cry1Ca is acting directly on the PM, consequently reducing the role of CBPA in that synergy model. However, the *Galleria* model was optimal to assess bacteria–PM interactions *in vivo*. Indeed, the transit studies in *Galleria* clearly indicated that CBPA plays a role *in vivo*, as its presence increases the capacity of vegetative bacteria to adhere to the PM.

Concurrent with previous reports, the present study proposes an additional step in the mode of action of *B. thuringiensis* (Figure 8). The present results indicated that CBPA can be induced in vegetative Bt cells in the alkaline midgut environment, thus facilitating the adhesion of Bt bacteria to the PM and thereby increasing the performance of various virulence factors. Accordingly, chitinases or enhancin-like proteins (Bel and MpbE) [43,44] may be produced and destabilize the chitin structure of the PM. This, along with the role of the active pore-forming Cry toxins known to damage the midgut cells resulting in reduced PM renewal and reduced intestinal transit time, further facilitates bacterial adhesion with intestinal cells and increases colonization. This might be followed up by bacterial translocation from the midgut to the insect hemocoel, through the action of non-specific adaptation and virulence factors, notably those from the PlcR regulon, which was earlier shown to being important for virulence toward *Galleria* [31]. Therefore, the present results further the current understanding of the complex pathogenesis and ecology of Bt, owing to studies on two insects: *Ostrina*, which is naturally sensitive to Bt HD73 and its Cry1Ac toxin, and the non-sensitive model insect *Galleria*, which allows for the easy manipulation of the PM. Further studies are required to analyze the importance of CBPA in other Bt strains and insects in order to understand the mode of action of CBPA and to validate our findings as a general feature in the early stages of Bt insect larva infection.

**Figure 8.** Proposed working model for the site of action of the chitin binding protein CBPA. The figure indicates where CBPA, during the oral infection with *B. thuringiensis* in a Cry toxin susceptible lepidopteron larva, plays a role. The green blocks of the steps refer to the spore/bacteria actions, and the red blocks refer to the role of the Cry toxins. The numbers, in the time scale arrow (in blue), indicates the order of events of which some occurs simultaneously. Our results showed that CBPA is expressed on the surface of vegetative bacteria (step 1) and is induced at alkaline pH. The proposed major role of CBPA is its adhesion to the peritrophic matrix (PM) (steps 1 and 3), which permits outgrown bacteria to bind to the PM and to be closer to the intestinal surface (4 and 5) in order to facilitate the tissue damaging action of secreted enzymes and toxins, e.g., from the PlcR regulon [31].

#### **4. Materials and Methods**

#### *4.1. Bacterial Strains*

The plasmids, primers, and sequences used herein are enlisted in Tables 4 and 5. Bt strains were cultured at 30 ◦C, and *Escherichia coli* was cultured at 37 ◦C with agitation at 220 rpm in an LB medium (1% NaCl, 1% tryptone, and 0.5% yeast extract) [45]. *B. thuringiensis* HD73 (the wild-type strain, producing crystals exclusively comprising the Cry1Ac toxin) was used to clone the target gene and monitor promoter activity as the recipient strain [46], as well as for bioassays and mutant construction. DNA sequences were obtained from the NCBI database (https://www.ncbi.nlm.nih.gov/) and compared using the Basic Local Alignment Search Tool (BLAST).


#### **Table 5.** Primers and sequences used in this study.


Underline indicates the restriction enzyme site.

#### *4.2. Insects*

For *O. furnacalis* (Asian corn borer), larvae for bioassays and mortality tests (see below), were provided by the rearing at the Chinese Academy of Agriculture Sciences (Beijing, China). For mortality tests and other in vivo analyses, 5th instar larvae of the greater wax moth *G. mellonella* were used. Insects were reared at the INRAE-Micalis, Jouy en Josas, France, facilities at 27 ◦C and fed with pollen

and bee wax (La ruche Roannaise, France). Prior to assays, the larvae weighting approximately 250 mg were selected and stored under starvation conditions for 24 h.

#### *4.3. DNA Manipulation and Transformation*

PCR amplifications were performed using Taq DNA polymerase and Pfu DNA polymerase (TIANGEN Biotechnologies Corporation, Beijing, China). PCR products were separated on agarose gels and recovered using the HiPure Gel Pure DNA Mini Kit (Magen Biotechnology Corporation, Guangzhou, China), and plasmid DNA was extracted from *E. coli* using the Plasmid Miniprep Kit (Axygen Biotechnology Corporation, Hangzhou, China) in accordance with the manufacturers' instructions. Restriction enzymes and T4 DNA ligase (Thermo Fisher Scientific, Beijing, China) were used in accordance with the manufacturer's instructions. Oligo-nucleotide primers were synthesized by Sangon Biotech (Shanghai, China). All constructs were confirmed by DNA sequencing (GENEWIZ, Beijing, China). *E. coli* cells were transformed via standard procedures [53], and Bt cells were transformed via electroporation, as described previously [54].

#### *4.4. Cloning of the HD73-cbpA Gene*

The *HD73-cbpA* gene (ID:14557228) was cloned from the wild-type HD73 genome via PCR, using specific primers *cbpA*-a and *cbpA*-b (Table 5) under the following cycling conditions: denaturation at 94 ◦C for 30 s, annealing at 50 ◦C for 1 min, and extension at 72 ◦C for 1 min for 34 cycles. The size of the PCR products was 1368 bp, which was digested with *Bam*H I and *Sal* I. Thereafter, the fragment of the *cbpA* gene was inserted into the expression vector pET21b (Novagen, Bloemfontein, South Africa) and digested with the aforementioned restriction enzymes. The recombinant plasmid was transformed into *E. coli* JM110 for amplification and preservation. The recombinant plasmid was selected using Ampr on the vector to select transformants and to obtain potentially positive clones via PCR, followed by NCBI BLAST to verify the correct sequence. Finally, the recombinant plasmid was transformed into *E. coli* BL21/DE3 for expression.

#### *4.5. Expression and Purification of CBPA*

*E. coli* BL21-harboring pET*cbpA* were cultured in an LB medium up to the logarithmic phase (A600 = 0.8 to 1.0), and the culture was cooled to 20 ◦C and induced with IPTG at a step-down gradient of the final concentration from 0.4 to 1.0 mM at 150 rpm for 20 h. The cells were harvested via centrifugation (6000 rpm/min, 10 min, and 4 ◦C) and resuspended in a 20 mM Tris-HCl buffer (pH = 8.0). Thereafter, the supernatant (cytosol) and pellet of the crude protein extract were obtained via centrifugation (12,000 rpm/min, 20 min, and 4 ◦C), followed by bacterial cell lysis using an ultrasonic cell disruption system. Protein expression was analyzed via SDS-PAGE (10% resolving gel). The protein was incubated with chitin beads, and the bound protein was purified after elution with an NaCl solution containing a step-up gradient from 0 to 1.0 M.

#### *4.6. Determination of Chitinase Activity*

The chitinase activity of the CBPA protein was detected using the Sigma-Aldrich Chitinase Assay Fluorimetric kit (Sigma-Aldrich) in accordance with the manufacturer's instructions. First, the 4-MU standard solution was prepared, and fluorescence was measured. Thereafter, the CBPA protein (1.21 mg/mL) was added to three different substrates. Green Trichoderma chitinase was used as a positive control. The substrate reaction solution and the standard solution were equilibrated in a 37 ◦C water bath for 5–10 min. The standard sample and the reaction sample were prepared (10 μL) in accordance with the manufacturer's instructions before being subjected to agitation in parallel. The sample was incubated in a 37 ◦C warm bath for 30–60 min. Finally, 200 μL of a stop solution was added, and fluorescence was measured at an excitation wavelength of 360 nm and an emission wavelength of 450 nm. The concentration of the target solution was determined from a standard curve with chitinase as the positive control.

#### *4.7. Construction and Expression of Recombinant gfp-conjugated cbpA*

Both the 1379-bp fragment of the *cbpA* ORF and the 541-bp upstream sequence comprising the promoter were amplified via PCR with the specific primers *gfp*-1 and *gfp*-2 (Table 5), using the HD73 genome as the template. A 48-bp linker fragment (TCAGGTGGAGGCGGTTCAGGCGG AGGTGGCTCTGGCGGTGGCGGATCG) and a 717-bp GFP ORF were amplified via PCR with specific primers *gfp*-3 and *gfp*-4 (Table 5), using the Cry1Ac-GFP plasmid as the template [55]. The fusion fragment was amplified via overlapping PCR and inserted into the shuttle vector pHT315, as described previously [49], using the *Sph*I and *Bam*HI restriction sites. Thereafter, the recombinant plasmid was transformed into HD73 via electroporation.

#### *4.8. Laser-Scanning Confocal Microscopy of CBPA-GFP Fusions*

A single colony was inoculated in an LB medium, cultured overnight at 30 ◦C with agitation at 220 rpm, and 1% of the inoculum was seeded in 100 mL of the LB medium and incubated until the OD600 value approached 2.0–2.2, which is the end point of the exponential phage (T0) according to the previously established growth curve. One-milliliter bacterial aliquots were taken every 1 h for centrifugation to obtain the precipitation (30 ◦C, 12,000 rpm for 1 min) and washed twice with deionized water (200 μL). Thereafter, bacterial cells were resuspended in a specific amount of deionized water. Different samples were analyzed at time points T1, T2, etc., (Tn means n hours after T0 entrance into stationary phase). A red fluorescent membrane stain FM4-64 (Molecular Probes, Inc., Eugene, OR, USA) was suspended in dimethyl sulfoxide to a final concentration of 100 μM and incubated on ice for 1 min. Five-hundred nanoliters of the bacterial sample and an equal volume of FM4-64 were mixed and placed on a glass slide, covered with a coverslip, and sealed with a transparent nail polish. FM4-64-stained bacteria were observed using a 63× oil-immersion lens and scanned using a laser-scanning confocal microscope (Leica TCS SL; Leica Microsystems, Wetzlar, Germany). The FM4-64 was detected at an excitation wavelength of 514–543 nm; GFP was detected at 633 nm.

#### *4.9. Construction of the Transcriptional Promoter P*cbpA-lacZ *Fusion Gene*

The sequence upstream from the *cbpA* gene, where the promoter fragment of P*<sup>3189</sup>* is located, was cloned using the specific primers P*cbpA*-F and P*cbpA*-R (Table 5) from the wild-type HD73 genome, under the following cycling conditions: denaturation at 95 ◦C for 20 s, annealing at 50 ◦C for 20 s and extension at 72 ◦C for 1 min for 30 cycles. The fragment of the P*cbpA* promoter was inserted into vector pHT304-18Z using the *Pst*I and *Bam*HI restriction sites. The vector pHT304-18Z harbored a promoter-less *lacZ* gene [50]. Thereafter, the recombinant plasmid pHTP*cbpA* was transformed into HD73 via electroporation, and positive strains were selected on the basis of the Erm<sup>r</sup> phenotype and via PCR identification.

#### *4.10.* β*-Galactosidase Assays*

Bt strains containing *lacZ* fusion transcripts were cultured in LB at 30 ◦C and at 220 rpm with appropriate antibiotics up to an OD600 value of 1.5–2.0. Thereafter, NaOH was added to a final concentration of 24 mM (pH 9). No NaOH was added to the control culture. Two-milliliter aliquots were harvested from the experimental and control cultures at 15 and 30 min, and β-galactosidase activity (Miller units per milligram of protein) of the cell pellets was measured as described previously [56]. Final values were determined using the data processing software Original 8.0 and SPSS.

#### *4.11. Construction of the HD-73* Δ*cbpA Mutant*

The interruption mutant was obtained via homologous recombination and insertion-replacement with a kanamycin resistance-encoding DNA cassette. The upstream gene fragment *cbpA*-u (317 bp) was obtained with the use of specific primers *cbpA*-A and *cbpA*-B and the downstream gene fragment 3189-d (504 bp) using primers *cbpA*-C and *cbpA*-D, with the genomic DNA of wild-type strain HD73 as the template. The kanamycin resistance gene cassette was a 1495-bp fragment. Primers *cbpA*-A and *cbpA*-D were used to ligate the aforementioned three fragments via overlapping PCR. Thereafter, the cassette fragment (2243 bp) was inserted into the temperature-sensitive suicide mutant erythromycin-resistant plasmid pRN5101 [51] at the *Bam*HI and *Sal*I restriction sites, finally yielding the pRN5101Ω*cbpA* plasmid. Positive transformant mutants were selected as reported previously [48].

#### *4.12. Complementation of the HD-73* Δ*cbpA Mutant*

Oligonucleotide primers *cbpA*-c (with a pHT 304 upstream homologous arm) and *cbpA*-d (with a pHT 304 downstream homologous arm) (Table 5) were used to amplify the *cbpA* gene and its own promoter by using HD73 genomic DNA as the template. The amplified fragment (1946 bp) was ligated into the shuttle vector pHT304 to generate pHTC*cbpA* using a recombinant enzyme. The resulting plasmid (pHTC*cbpA*) was amplified in *E. coli* and introduced into the HD-73 ΔcbpA mutant strain. Genetically-complemented mutant HD73 strains (Δ*cbpA*::*cbpA*) were obtained by transforming pHTC*cbpA* into HD73Δ*cbpA* cells.

#### *4.13. Crystal Spore Mixture Preparation for Asian Corn Borer Bioassays*

A single colony of wild-type and mutant strains of Bt HD73 was inoculated in 20 mL of an LB liquid medium and cultured overnight at 30 ◦C with agitation. The activated bacteria were then transferred to 300 mL of an LB broth at a ratio of 1% and cultured at 180 rpm for 4–5 h to the logarithmic growth phase. Thereafter, the bacterial culture supernatant was transferred to 300 mL of an LP beef extract peptone medium at a ratio of 1% for approximately 40 h at 30◦C, and the crystal cleavage rate was observed to be greater than 50%. The crystals, spores, and debris were harvested via centrifugation at 6000 rpm for 20 min (20 ◦C). The pellet was washed with water and centrifuged at 6000 rpm for 20 min (20 ◦C), and the supernatant was discarded. The pellet was resuspended in deionized water to obtain the final crystal and spore suspension.

#### *4.14. Spore Preparation for Asian Corn Borer Bioassays*

A single colony of the HD73, ΔcbpA mutant, and the complemented strains was inoculated in 20 mL of the LB broth and cultured overnight at 30 ◦C with agitation. Activated bacteria were transferred to 300 mL of a CCY liquid medium (MgCl2·6H2O: 0.5 mmol/L, MnCl2·4H2O: 0.01 mmol/L, FeCl3·6H2O: 0.05 mmol/L, ZnCl2: 0.05 mmol/L, CaCl2.6H2O: 0.2 mmol/L, KH2PO4: 13 mmol/L, K2HPO4: 26 mmol/L, L-Glutamine: 20 mg/L, Casamino acids hydrolysate: 1 g/L, Yeast extract: 0.4 g/L, glycerol: 0.6g/L) based on a ratio of 1% and cultured at 220 rpm for 72 h up to a spore percentage of >99%, as reported previously [57]. The spores were harvested via centrifugation at 6000 rpm for 10 min (4 ◦C). The pellet was washed with water and centrifuged at 6000 rpm for 10 min, and the supernatant was discarded. The pellet was resuspended in deionized water to obtain a spore suspension.

#### *4.15. Spore Counts*

Spore suspensions of the wild-type HD73, ΔcbpA mutant, and the complemented strains were administered a heat treatment (65 ◦C for 30 min) to eliminate all vegetative cells. Thereafter, 100 μL of the 10−<sup>6</sup> dilutions of each sample were plated on the LB agar medium and incubated at 30 ◦C for 12 h. The colony characteristics of the Bt culture were assessed as described previously [58].

#### *4.16. Dose-Mortality Response Bioassays Against Asian Corn Borer*

SDS-PAGE was performed to analyze protein profiles and concentrations, using bovine serum albumin (BSA) as the standard. Bioassays were performed as described previously [59]. Cry1Ac was prepared as described previously [60]. Insecticidal activity against the Asian corn borer, *O. furnacalis* (Guenée) was assayed by administering neonates with an artificial diet supplemented with a 0.01 μg Cry1Ac/g diet and different concentrations of spores (104–108 CFU/g diet) prepared from the HD73 wild-type, ΔcbpA mutant, and the complemented strains. The spores were heat-treated (65 ◦C for 30 min) to eliminate all vegetative cells and crystals. The feed was uniformly distributed into 48-well trays, each well containing approximately 200 mg of feed and infested with one neonate larva. Assays were carried out at 27 ± 1 ◦C with a L16/D 8-h photoperiod and 70–80% relative humidity. Survivors were recorded after 7 d. Deionized water was used as the negative control. Each assay was performed in triplicate.

#### *4.17. In Vivo G. mellonella Virulence Assays with HD73 and* Δ*cbpA Mutant Strains*

Oral force-feeding assays were performed as described previously [32,61]. A dose of 3–5 <sup>×</sup> 106 spores or mid-log LB grown bacteria from HD-73 wild-type and ΔcbpA mutants was suspended in 10 μL of 0.3 mg/mL Cry1C activated toxin or in 1% NaCl water (negative control) using a needle and syringe for the accurate distribution to each larvae. At least 20 larvae per condition were incubated at 37 ◦C under starvation conditions, and mortality was scored at 24 h post-infection. Experiments were performed in triplicate. The inocula were evaluated via plating after serial dilution.

#### *4.18. Expression of the CBPA-GFP Fusion Protein in Vivo in G. mellonella*

CBPA expression during intestinal transit was scored via the fluorescence microscopic examination of intestinal samples from the anterior and posterior midgut of larvae infected with HD73 (pHT-*cbpA-gfp*). At least 5 larvae were dissected, and gut samples were observed using the Fluorescence Zeiss-Observer microscope with a 100× oil-immersion objective lens with a GFP filter at 1 and 3 h post-ingestion. CFU counts and fluorescence levels were scored through arbitrary visual observation, since only few bacteria were present, and the fluorescence intensity was too low for imaging or fluorescence-activated cell sorting analysis.

#### *4.19. Intestinal Particle Transit Time for Final-Instar G. mellonella*

Final-instar larvae starved for 24 h, similar to those in the virulence assays, were used. The transit time was recorded by observing the first appearance of feces and the presence of bacteria in the feces. Thirty larvae were force-fed with the same dose of spores as in the virulence assay. Larvae were placed individually in a 24-well micro-titer plate. Feces were harvested and enumerated at 1, 3, 4, and 24 h after force-feeding. The CFU value was evaluated in the feces at 1 and 2 h post-ingestion via the plating of 200 μL suspension in 1% NaCl water.

#### *4.20. HD73 and HD-73* Δ*cbpA Intestinal Transit and Localization Assays*

To analyze the effect of *cbpA* deletion on bacterial transit following oral infection, a condition devoid of the Cry1C toxin was used. In total, 3–5 <sup>×</sup> 106 mid-log-phase bacteria were force-fed to 5th instar *G. mellonella* larvae. CFU values were recorded after plating at different time points and with four different larval samples: whole larvae, dissected whole intestine, intestine without the PM, and the PM alone. First, alcohol (70%, 1 min)-cleaned larvae were homogenized in a PBS (phosphate buffed saline pH 7.4) buffer using an Ultraturax mixer at the rate of 2 larvae per 4 mL. Two completely dissected intestines were homogenized with Ultraturax in 1.5 mL 1% NaCl water. CFU values per larva were recorded at 0, 3, and 24 h post-feeding in at least 2 larvae and repeated 3–4 times. To record bacteria adhering to the PM, larvae were incubated on ice 1 h post-ingestion and gently dissected with the help of chirurgical scissors and tweezers to obtain the PM and intestine alone. The cooled larvae were placed on the dorsal under the binoculars and gently opened with the scissors from the rectum to the head, and the skin is maintained with needles. Fat body, silk glands, and other tissue were gently moved to only expose the digestive tube (DT) (the whole intestine from foregut to hindgut). The DT was cut at two sites, one just above the foregut and one above the rectum, and moved to a clean glass slide. Then, a small cut with the scissors was performed just above the hindgut, and the PM was gently pooled out and separated from the intestine with the fine tweezers; the PM and the intestine were placed directly in the respective tubes prior to homogenization. Two PMs were crushed

with a small pestle and 5–6 glass beads in 200 μL 1% NaCl water, and an additional 200 μL of NaCl were added before serial dilution and plating. This experiment was performed in triplicate for each strain. The intestines alone were processed as for the above whole intestines method.

**Supplementary Materials:** The following are available online at http://www.mdpi.com/2072-6651/12/4/252/s1, Figure S1: Analysis of mortality of *Galleria mellonella*.

**Author Contributions:** Conceptualization, S.G., F.S. and C.N.-L.; validation, J.Q., Z.T., Y.Z. and C.B.; formal analysis, S.G., F.S. and C.N.-L.; investigation, J.Q., Z.T., C.B.; data Curation, K.H.; writing—original draft preparation, J.Q. and Z.T.; writing—review and editing, F.S., S.G. and C.N.-L.; visualization, Z.T.; funding acquisition, F.S. and C.N.-L. All authors have read and agreed to the published version of the manuscript.

**Funding:** This study was supported by the grants from The National Key Research and Development Program of China (2017YFD0200400) and from the National Natural Science Foundation (31530095). The project was also financially supported from France, INRAE, MICA department and the French National Research Program RISKOGM (11-MERES-RISKOGM-3-CVS-59).

**Conflicts of Interest:** We have no conflicts of interests.

#### **References**


© 2020 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).

## **Potential of Cry10Aa and Cyt2Ba, Two Minority** δ**-endotoxins Produced by** *Bacillus thuringiensis* **ser.** *israelensis***, for the Control of** *Aedes aegypti* **Larvae**

**Daniel Valtierra-de-Luis 1, Maite Villanueva 1,2, Liliana Lai 1, Trevor Williams <sup>3</sup> and Primitivo Caballero 1,2,4,\***


Received: 17 April 2020; Accepted: 26 May 2020; Published: 29 May 2020

**Abstract:** *Bacillus thuringiensis* ser. *israelensis* (Bti) has been widely used as microbial larvicide for the control of many species of mosquitoes and blackflies. The larvicidal activity of Bti resides in Cry and Cyt δ-endotoxins present in the parasporal crystal of this pathogen. The insecticidal activity of the crystal is higher than the activities of the individual toxins, which is likely due to synergistic interactions among the crystal component proteins, particularly those involving Cyt1Aa. In the present study, Cry10Aa and Cyt2Ba were cloned from the commercial larvicide VectoBac-12AS® and expressed in the acrystalliferous Bt strain BMB171 under the *cyt1Aa* strong promoter of the pSTAB vector. The LC50 values for *Aedes aegypti* second instar larvae estimated at 24 hpi for these two recombinant proteins (Cry10Aa and Cyt2Ba) were 299.62 and 279.37 ng/mL, respectively. Remarkable synergistic mosquitocidal activity was observed between Cry10Aa and Cyt2Ba (synergistic potentiation of 68.6-fold) when spore + crystal preparations, comprising a mixture of both recombinant strains in equal relative concentrations, were ingested by *A. aegypti* larvae. This synergistic activity is among the most powerful described so far with Bt toxins and is comparable to that reported for Cyt1A when interacting with Cry4Aa, Cry4Ba or Cry11Aa. Synergistic mosquitocidal activity was also observed between the recombinant proteins Cyt2Ba and Cry4Aa, but in this case, the synergistic potentiation was 4.6-fold. In conclusion, although Cry10Aa and Cyt2Ba are rarely detectable or appear as minor components in the crystals of Bti strains, they represent toxicity factors with a high potential for the control of mosquito populations.

**Keywords:** *Bacillus thuringiensis*; *Aedes aegypti*; minor proteins; synergy; mosquito control; Bti

**Key Contribution:** Cry10Aa and Cyt2Ba are found as minor components in the crystals of some strains of *Bacillus thuringienis* ser. *israelensis*. Both proteins have a high insecticidal activity against insects (e.g., *Aedes aegypti* larvae) and when ingested together they exhibit one of the strongest synergistic activities that have been described so far.

#### **1. Introduction**

*Bacillus thuringiensis* ser. *israelensis* (Bti) was the first Bt serotype found to be toxic for dipteran species [1]. Bti forms parasporal inclusion bodies composed of insecticidal proteins (δ-endotoxins) that are widely used as the basis for microbial larvicides against dipteran species of medical importance, including mosquitoes, blackflies and chironomids [2,3]. Bti based products are considered to be powerful and highly selective larvicides for the control of disease vectors [4–6]. Indeed, Bti has been used to control mosquitoes for more than 35 years with almost no resistance report in vector populations [7,8]. The absence of resistance is likely due to the different modes of action and the synergistic effects of the multiple crystal proteins present in Bti-based products [9–11].

The parasporal crystal of Bti contains large amounts of four toxins: Cry4A, Cry4B, Cry11A and Cyt1A [12]. In addition, Cry10Aa and Cyt2Ba have also been described in some Bti strains, although these are expressed and accumulate in the crystal in much smaller quantities that the four main components [13,14]. All six of these proteins are encoded in the fully sequenced Bti plasmid pBtoxis [15]. The Cry10Aa protein was cloned and named CryIVC, according to the existing classification at that time, but was described as a protein with a low larvicidal potency against *A. aegypti* (Diptera; Culicidae) [16]. Later, the *cry10Aa* gene was identified as part of an operon that comprises two open reading frames (*orf1* and *orf2*) separated by a 66 bp gap [15]. Cloning of the complete operon, linked to the strong promoter of the *cyt1A* gene, revealed that Cry10Aa was expressed at high levels and exhibited high larvicidal activity, both alone and in combination with Cyt1A [17]. In contrast, although present at relatively low abundance in the Bti crystal [18], Cyt2Ba exhibited activity against *A. aegypti* larvae, but lower than the better-studied Cyt1Aa protein [19].

The interactions among the Cry and Cyt proteins of Bti have received more attention than any of the other Bt serovars [20–24]. Interactions involving the Cyt1A protein have attracted particular attention given the capacity of this protein to enhance the insecticidal activity of Cry proteins in strains of Bti [9,17,20,24], and those of Bt strains belonging to other subspecies [25]. Conversely, studies on the interactions of Cyt2Ba with other components of the Bti crystal are restricted to a single report of low synergistic activity of Cyt2Ba with the Cry4Aa protein [21].

The objective of this study was to quantify the larvicidal activity of the δ-endotoxins Cry10A and Cyt2Ba, which are minor components of the parasporal crystal of some Bti strains, against *A. aegypti*. To produce high amounts of these minority proteins, two recombinant Bt strains were constructed. One of these recombinants produced a crystal whose only component were the two Cry10Aa proteins, while the other only produced Cyt2Ba. We provide evidence that these proteins interacted synergistically to a remarkable degree when simultaneously ingested by *A. aegypti* larvae.

#### **2. Results**

#### *2.1. Insecticidal Cry and Cyt Genes Identified in Bti Strain from VectoBac-12AS*®

A bioinformatic analysis of the genome of the Bti strain isolated from the commercial product VectoBac-12AS®, revealed that this strain contains a complex of insecticidal genes, including *cry* genes (*cry4Aa*, *cry4Ba*, *cry10Aa*, *cry11Aa*, and *cry60Aa*/*cry60Ba*) and *cyt* genes (*cyt1Aa*, *cyt2Ba*, and *cyt1Ca*). Unfortunately, it was not possible to obtain the complete sequence of the *cry4Aa* and *cry4Ba* genes, because they appeared distributed in various contigs. The rest of the *cry* genes shared 100% identity and similarity with some of the gene variants that have been previously described. Thus, *cry10Aa* was completely identical to *cry10Aa3* [15], *cry11Aa* was identical to *cry11Aa1* [26], and *cry60Aa*/*cry60Ba* were identical to *cry60Aa2*/*cry60Ba2* [27]. The three *cyt* genes (*cyt1Aa*, *cyt2Ba*, and *cyt1Ca*) identified in the VectoBac-12AS® strain were also fully identical to *cyt1Aa1* [28], *cyt2Ba1* [18], and *cyt1Ca1* [15], respectively.

#### *2.2. Cloning of Cyt2Ba, Cry10Aa and Cry11Aa*

The pairs of primers designed for *cyt2Ba, cry10Aa* and *cry11Aa* amplified fragments of 1536, 3813 and 2634 bp, respectively. The *cry10A* cloned fragment contained two open reading frames (*orf1* and *orf2*) in the nucleotide sequence, codifying for proteins of 680 and 489 amino acids, respectively. The *cry11Aa* amplicon encoded a protein of 646 amino acids but it also contained the *p19* gene located

before *cry11Aa,* in line with the usual order of genes in this operon. The amplicon was cloned in a pSTAB plasmid containing the *p20* chaperon, which improves Cry11Aa synthesis and crystal formation [29]. The amplified fragment of *cyt2Ba* contained a single ORF which codified for a protein of 263 residues. The *cry4A* and *cry4Ba* genes, previously described and cloned by Delecluse et al. [30], were used in this study.

#### *2.3. Characterization of Bt Recombinant Strains Expressing cyt2Ba, cry10Aa, cry4Aa, cry4Ba and cry11Aa*

Daily microscopical observation of the growth of the recombinant Bt strains in CCY medium confirmed that all of them produced spores and crystals between 36 and 48 h after the medium was inoculated. As expected, vegetative cells of BMB171 strain transformed with an empty plasmid produced endospores but no crystals.

SDS-PAGE showed that the recombinant BMB171-Cry10Aa expressed two proteins with molecular masses of approximately 68 and 56 kDa, which corresponded to the predicted sizes of the proteins encoded by *orf1* and *orf2*, respectively, of the *cry10Aa* operon (Figure 1, lane 3). Samples of spores and crystals from the rest of the recombinant strains (BMB171-Cyt2Ba, 4Q2-81-Cry4Aa, 4Q2-81-Cry4Ba and BMB171-Cry11Aa) generated characteristic major bands of approximately 29, 134, 128, and 73 kDa, respectively (Figure 1). The electrophoretic mobility of all these bands correlated well with the molecular mass of the proteins Cyt2Ba (lane 2), Cry4Aa (lane 4), Cry4Ba (lane 5) and Cry11Aa (lane 6).

**Figure 1.** SDS-PAGE gel showing the protein profiles of the recombinant Bt strains and the strain present in VectoBac-12AS®. Lane M, molecular mass marker; lane 1, BMB171 acrystalliferous strain with an empty plasmid; lane 2, BMB171-Cyt2Ba; lane 3, BMB171-Cry10Aa; lane 4, 4Q2-81-Cry4Aa; lane5, 4Q2-81-Cry4Ba; lane 6, BMB171-Cry11Aa; lane 7, wild-type Bti strain from VectoBac-12AS®. Arrows indicate major protein bands.

#### *2.4. Mosquitocidal Activity of the* δ*-Endotoxins Produced by Bti*

Single-concentration bioassays involving an estimated LC30 concentration of inoculum in all cases were performed on mixtures of Cyt2Ba with each of the Cry4Aa, Cry4Ba, Cry10Aa and Cry11Aa proteins. The results of these assays indicated that *A. aegypti* second instar larvae treated with combinations of Cry10Aa+Cyt2Ba and Cry4Aa+Cyt2Ba experienced high mortality compared to the mortality values observed in insects treated with each of the toxins separately (Table 1). In contrast, no evidence of potentiation of larval mortality was observed for mixtures of Cry4Ba+Cyt2Ba or Cry11Aa+Cyt2Ba. For this reason, the 1:1 mixtures of Cry10Aa+Cyt2Ba and of Cry4Aa+Cyt2Ba were selected for subsequent concentration-mortality studies.

**Table 1.** Mortality of *A. aegypti* second instar larvae at 24 h after inoculation with individual Bti δ-endotoxins and the binary combinations Cyt2Ba/Cry10Aa, Cyt2Ba/Cry4Aa, Cyt2Ba/Cry4Ba and Cyt2Ba/Cry11Aa.


<sup>1</sup> Control insects experienced no mortality in all cases.

Table 2 shows the raw mortality data of a series of concentrations for Cry10Aa, Cyt2Ba and the combination of both. Analogously, Table 3 shows the raw mortality data of a series of concentrations for Cry4Aa, Cyt2Ba and the combination of both.

**Table 2.** Mortality of *A. aegypti* second instar larvae at 24 h after inoculation with Cry10Aa, Cyt2Ba and combination of both.


Control insects experienced no mortality in all cases.

**Table 3.** Mortality of *A. aegypti* second instar larvae at 24 h after inoculation with Cry4Aa, Cyt2Ba and combination of both.


Control insects experienced no mortality in all cases.

Regression lines were performed for the individual toxins and the mixture of toxins (Figure 2) which were then used to estimate median lethal concentrations (LC50) (Table 4).


*Toxins* **2020**

, *12*, 355

comprisedspore+crystalexperiencedmortality (95%). ExpectedLC50 by(1992). (d) Synergism factor defined as the ratio of the expected LC50and the observed LC50. (e) Toxins were present in equal amounts in the experimental inocula.

**Figure 2.** Graphical representation of the logit regression lines for the individual toxins and the toxin combinations. (**a**) Regression lines for Cyt2Ba, Cry10Aa and Cry10Aa+Cyt2Ba. (**b**) Regression lines for Cyt2Ba, Cry4Aa and Cry4Aa+Cyt2Ba.

Recombinant Cry10Aa and Cyt2Ba proteins exhibited a high insecticidal activity against *A. aegypti* second instar larvae when inoculated individually. The LC50 values estimated for Cry10Aa and Cyt2Ba were 299.62 ng/mL and 279.37 ng/mL, respectively. The VectoBac-12AS® wild-type strain, incorporated into the bioassays as a positive control, had an LC50 value of 1.02 <sup>×</sup> 10−<sup>1</sup> ng/mL and the BMB171 strain with the empty plasmid resulted in no mortality (Table 4). The slopes of the regression lines corresponding to Cry10Aa and Cyt2Ba did not differ significantly (F1,8 = 0.620, *p* = 0.454), whereas the slope of the mixture of Cry10Aa+Cyt2Ba was significantly lower than that of the individual toxins (F2,12 = 7.359, *p* = 0.008). The observed LC50 value for Cry10Aa+Cyt2Ba was 4.22 ng/mL whereas the expected LC50 value was 289.27 ng/mL, assuming additive action of each of the toxins [31]. The estimated potentiation of Cyt2Ba and Cry10Aa proteins when ingested together and in the same relative proportions, was 68.6-fold (Table 4).

In contrast, the slopes of the regressions of the individual Cyt2Ba and Cry4Aa toxins differed significantly (F1,8 = 11.405, *p* = 0.010). The LC50 value for Cry4Aa was estimated at 34.63 ng/mL. The observed LC50 value for the binary combination of Cry4Aa+Cyt2Ba was 13.41 ng/mL, whereas the expected LC50 value was 61.62 ng/mL, assuming additive action of each of the toxins [31]. These results indicate potentiation in the Cry4Aa+Cyt2Ba protein mixture by a factor of 4.6 (Table 4).

#### **3. Discussion**

The δ-endotoxins that constitute the major parasporal crystal components of Bti strains (Cry4Aa, Cry4Ba, Cry11Aa and Cyt1Aa) are the best studied of the Bt crystal proteins, both in terms of the insecticidal properties of individual proteins and the interactions among them in the digestive tract of susceptible mosquito species. The present study provides evidence that additional proteins, such as Cry10Aa and Cyt2Ba, which are usually present as minor components in Bti strains, are also important toxicity factors that act in a highly synergistic manner when these proteins are inoculated simultaneously in *A. aegypti* larvae. Cyt2Ba was previously described as a synergy factor for Cry4Aa and in this study we quantified the effects of the interaction on larval mosquito mortality [21].

The insecticidal activity of Cry10Aa in fourth instar larvae of *A. aegypti* was previously estimated at LC50 = 2061 ng/mL [17], which is about 7-fold higher than the value that we estimated in second instar larvae of the same species. A decrease in the susceptibility of larvae to infection by pathogens with increasing growth stage is common in insects [32], including their susceptibility to Bt toxins [22,33,34].

Several previous studies have described the larvicidal activity of Cyt2Ba protein in mosquito species belonging to the genera *Culex*, *Aedes* and *Anopheles* [19,35,36], although for a given toxin concentration the mortality that was recorded in Cyt2Ba-treated *A. aegypti* larvae was lower than that produced by the Cyt1Aa protein [19]. The estimated 24 h LC50 value of Cyt2Ba in second instar larvae of *A. aegypti* obtained in this study was approximately 27-fold lower than the value estimated by others [36]. This may be due to differences in the origin of the mosquito population and history of exposure to Bt toxins, and the fact that Wirth et al. [36] used lyophilized powdered inoculum rather than the freshly-prepared spore + crystal preparations that we employed.

The high potential of Bti proteins against mosquito larvae is mainly attributed to the interactions that occur among the component toxins. Although present at low abundance, Cry10Aa and Cyt2Ba contribute to the insecticidal activity of Bti by potentiation of toxin interactions [5]. Cry10Aa shows synergistic activity with Cyt1Aa [17] and Cry4Aa [37], whereas Cyt2Ba shows synergistic interaction with Cry4Aa [21] and *L. sphaericus* [36] in *A. aegypti*. When large amounts of these proteins were produced in an acrystalliferous Bt strain in the present study, very high levels of toxicity against *A. aegypti* larvae were obtained.

The LC50 value obtained here for the Cry10Aa/Cyt2Ba mixture was about 19 times lower than the value described for the Cry10Aa/Cyt1Aa mixture. This degree of potentiation is one of the highest observed so far for Bti crystal proteins, only comparable to that described for Cyt1A with Cry4Aa and Cry11Aa [38], or Cry4Ba in mixtures with Cyt2Aa2 from Bt *darmstadiensis* against *A. aegypti* larvae [39]. It seems, therefore, that Cyt2 proteins have a greater involvement in toxin synergy than has been attributed to date.

The molecular mechanisms underlying synergistic or other combinatorial effects between Bt insecticidal proteins have been the subject of several studies, although none have focused specifically on Cyt2Ba. The synergistic interaction of Cyt1Aa and Cyt2Aa with Cry4Ba appears to involve binding to the Cry protein through the domain II loops [9,40]. For the interaction between Cyt1Aa and Cry11Aa, specific charged residues have been identified on the Cyt1Aa protein that are involved in binding to Cry11Aa prior to insertion in the midgut epithelial cell membrane [41], although others have proposed that Cyt1Aa is a membrane-bound receptor that uses the exposed charged residues to bind Cry11Aa, thereby facilitating the interaction of the Cry protein with the target cell membrane [42,43]. Oligomerization of the Cyt1Aa toxin is essential for its toxicity in *A. aegypti* [44].

Bti has high larvicidal activity against mosquitoes, although its repeated use can lead to the appearance of resistance. The major components of the crystal, such as Cyt1A, Cry11A, or Cry4 are likely to be the main targets of such resistance. In this study, we demonstrated that Cry10Aa and Cyt2Ba, minor components of the parasporal crystal, have a high mosquitocidal potency and marked synergistic activity when present in a mixture. The optimization of culture conditions that result in improved production of Cry10A and Cyt2Ba may offer a rapid means to produce more effective Bti-based mosquitocidal products.

#### **4. Conclusions**

The toxicities of Cry10Aa and Cyt2Ba against *A. aegypti* are comparable to the major toxins of Bti and show one of the strongest potentiation effects observed for Bti crystal components to date. This potentiation was much stronger than occurred between Cyt2Ba and Cry4Aa. Further study of the minor crystal components of Bti is likely to provide additional opportunities for the development of safe and effective tools for the biological control of mosquito vectors of medical importance.

#### **5. Materials and Methods**

#### *5.1. Bacterial Strains and Plasmids*

*B. thuringiensis* ser. *israelensis* (Bti) was isolated from the commercial insecticide VectoBac-12AS® (Kenogard, Barcelona, Spain). *Escherichia coli* XL1 blue was used for transformation. The recombinant vector pSTAB [45] was used as the protein expression vector, engineered with the gene of interest. The acrystalliferous Bt strain BMB171 was used as the host strain for protein expression [46]. Bt recombinant strains 4Q2-81 pHT606:*cry4Aa* and 4Q2-81 pHT611:*cry4Ba* were kindly provided by Dr. Colin Berry (Cardiff University, Cardiff, UK) [30]. The Bt strains were grown in CCY medium containing 13 mM KH2PO4, 26 mM K2HPO4, 10 mL/L Nutrient stock solution (comprising L-glutamine, casein hydrolysate, casitone, yeast extract and glycerol), 1 ml/L metal salts solution [47] at 28 ◦C with continuous shaking at 200 rpm. All *E. coli* strains were cultured at 37 ◦C with continuous shaking (200 rpm) in Luria-Bertani (LB) broth (1% tryptone, 0.5% yeast extract, and 1% NaCl, pH 7.0). When required for selective growth, LB medium was supplemented with 20 μg/mL erythromycin (Em) and 100 μg/mL ampicillin (Amp).

#### *5.2. Insect Culture*

A laboratory colony of *A. aegypti* was started using eggs obtained from Dr. Susana Vilchez, (Universidad de Granada, Granada, Spain). The colony was maintained, under controlled environmental conditions (25 ± 1 ◦C and 85% RH, and a 16 h:8 h light: dark photoperiod), in the insectary facilities of the Instituto Multidisciplinario de Biología Aplicada (IMBA), Universidad Pública de Navarra, Spain. Adults of both sexes were maintained in BugDorm-1 insect rearing cages (MegaView Science, Taichung, Taiwan) and had continuous access to 20% sucrose solution and intermittent access (3 h/day) to defibrinated horse blood (Thermo Scientific, Waltham, MA, USA) to complete their gonotrophic cycle. Larvae were reared in 250 mL glass beakers (40–50 larvae/beaker) with 100 mL distilled water and brewer's yeast (1 mg/mL) as food.

#### *5.3. Total DNA Extraction and Genomic Sequencing*

Genome sequencing was performed to ensure that our Bti clone contained all the expected plasmids and genes, some of which may be lost during laboratory culture. Total genomic DNA (chromosomal + plasmid) was extracted from VectoBac-12AS® strain, following the protocol for DNA isolation from Gram-positive bacteria using the Wizard® Genomic DNA Purification Kit (Promega, Madison, WI, USA). A DNA library was prepared from total DNA and was subsequently sequenced in an Illumina NextSeq500 Sequencer (Genomics Research Hub Laboratory, School of Biosciences, Cardiff University, Cardiff, UK).

#### *5.4. Identification of Cry and Cyt Insecticidal Genes in VectoBac-12AS*®

Genomic raw sequence data were processed and assembled using CLC Genomics Workbench 10.1.1. Reads were trimmed, filtered by low quality and reads of less than 50 bp were eliminated. Processed reads were assembled de novo using stringent criteria of at least 95 bp overlap and 95% identity. Reads were then mapped back to the contigs for assembly. Genes were predicted using GeneMark [48].

To assist in the identification of potential insecticidal proteins, local BLASTP [49] was deployed against a database built in our laboratory comprising the amino acid sequences of known Bt toxins available at http://www.lifesci.sussex.ac.uk/home/Neil\_Crickmore/Bt [50], as well as other protein toxins of interest.

#### *5.5. Amplification, Cloning and Sequencing of Cyt2Ba, Cry11Aa and Cry10Aa*

Primers were designed to amplify the full-length coding sequence of *cyt2Ba, p19-cry11Aa* (including *p19* and *cry11Aa* genes) and the *cry10Aa* operon including *orf1* and *orf2* (Table 5). Primer sequences included XbaI and PstI restriction sites for *cyt2Ba*, as well as SalI and PstI restriction sites for *p19*-*cry11Aa* and SalI and PaeI restriction sites for *cry10Aa*. PCR reactions were performed, from total genomic DNAs, using Phusion DNA polymerase (NEB, Ipswich, UK) and amplicons were gel-purified using NucleoSpin Extract II kit (Macherey-Nagel, Düren, Germany). Purified products were then ligated into pJET1.2/blunt plasmid (CloneJET PCR Cloning Kit, Waltham, MA, USA) following the manufacturer's instructions. Ligation mixtures were transformed into *E. coli* XL1-Blue using standard procedures [51]. Colony-PCR was applied in order to check positive clones from which plasmid DNA was purified, using the NucleoSpin<sup>R</sup> plasmid kit (Macherey-Nagel Inc., Bethlehem, PA, USA) following the manufacturer's instructions. Subsequently, pJET plasmids were verified by sequencing (STABVida, Caparica, Portugal), digested with the appropriate combination of restriction enzymes, electrophoresed in 1% agarose gel and ligated into pre-digested pSTAB vector using the Rapid DNA ligation kit (Thermo Scientific) to obtain the recombinant plasmids pSTAB-*cyt2Ba* and pSTAB-*cry10Aa*. To clone *cry11Aa* the amplicon was ligated in a pSTAB in which *p20* gene was previously introduced, to obtain the recombinant plasmid pSTAB-*p19-cry11Aa-p20*. Ligation products were then electroporated into *E. coli* XL1 blue cells following standard protocols [51]. Positive clones were verified by colony-PCR and plasmids were purified and verified by restriction endonuclease digestion and electrophoresis. Once pSTAB-*cyt2Ba*, pSTAB-*cry10Aa* and pSTAB-*p19-cry11Aa-p20* were obtained, they were introduced into the acrystalliferous Bt strain BMB171.



Restriction enzyme sites are underlined.

*Bacillus* electrocompetent cells were generated as described previously [52]. Briefly, bacteria were grown in 300 mL of Brain heart infusion broth (Pronadisa) at 28 ◦C under shaking conditions (200 rpm) until the culture reached an OD600 nm of 0.4. Glycine was then added to the culture at 2% and bacterial cells were incubated for another hour, at 28 ◦C under shaking conditions (200 rpm). Bacterial cells were kept on ice for 5 min, centrifuged at 9000× *g* (4 ◦C) for 10 min and the pellet was washed three times with F buffer (272 mM sucrose, 0.5 mM MgCl2, 0.5 mM K2HPO4, 0.5 mM KH2PO4, pH 7.2). Cells were then resuspended in 600 μL of ice-cold F buffer and stored in aliquots of 50 μL at −80 ◦C. Plasmids were transformed into the BMB171 strain by electroporation, as described previously [53]. Positive clones were selected by colony-PCR. BMB171 was also transformed with an empty plasmid as a negative control.

#### *5.6. Expression of Cyt2Ba, Cry10Aa, Cry4Aa, Cry4Ba and Cry11Aa Recombinant Proteins and SDS-PAGE Analysis*

Wild-type Bti and recombinant Bt strains were grown at 28 ◦C, under shaking conditions (200 rpm), in CCY medium supplemented with 20 μg/mL erythromycin, if required. Crystal formation was observed daily under the optical microscope. After 2–3 days, when ~95% of the cells had lysed, the mixture of spores and crystals was collected by centrifugation at 10,000× *g*, for 10 min at 4 ◦C. The pellet was washed once with saline solution (1 M NaCl, 10 mM EDTA) and three times with 10 mM KCl. The spore + crystal mixture was finally resuspended in 10 mM KCl and kept at 4 ◦C until used. Samples of spores and crystals were mixed with 2x sample buffer (Bio-Rad, Hercules, CA, USA), boiled at 100 ◦C for 5 min, and then subjected to electrophoresis as previously described [54], using Criterion TGX™ 4–20% Precast Gel (Bio-Rad). Gels were stained with Coomassie brilliant blue R-250 (Bio-Rad) and then distained in 30% ethanol and 10% acetic acid. For protein quantification, a 10 μL volume of spore and crystal suspension was solubilized in vitro in 1 mL of alkaline solution (50 mM Na2CO3, 10 mM DTT, pH 11.3) for 2 h at 37 ◦C. The protein concentration of each preparation was measured by the Bradford assay (Bio-Rad), using bovine serum albumin (BSA) as a standard.

#### *5.7. Mosquitocidal Activity of the* δ*-Endotoxins Produced by Bti*

The toxicity of Cry10Aa and Cyt2Ba was determined by bioassay against *A. aegypti* second instar larvae. Concentration-mortality bioassays were performed following a modified method described previously [33]. Groups of 10–15 second instar larvae were placed in one well of a 6-well cell culture plate (Costar) and they were exposed to one concentration of Bt (spores+crystals). Each well contained 5 mL of Bt suspension with the corresponding toxin concentration and 0.5 mg of brewer's yeast as food. Toxin concentrations were 2000, 666, 222, 74, 24.7 and 8.2 ng/mL for Cry10Aa; 4000, 1333, 444, 148, 49.4 and 16.4 ng/mL for Cyt2Ba and 4 <sup>×</sup> <sup>10</sup><sup>−</sup>1, 2 <sup>×</sup> <sup>10</sup><sup>−</sup>1, 1 <sup>×</sup> <sup>10</sup><sup>−</sup>1, 5 <sup>×</sup> <sup>10</sup><sup>−</sup>2, 2.5 <sup>×</sup> <sup>10</sup><sup>−</sup>2, 1.2 <sup>×</sup> <sup>10</sup>−<sup>2</sup> ng/mL for Bti (VectoBac-12AS®) as the positive control. Each bioassay was performed at least three times, depending on the toxin. Control insects were mock-infected. Insects were incubated at 25 ◦C and 16 h:8 h L:D photoperiod. Mortality was recorded at 24 h post-treatment. The concentration-mortality raw data are represented in Tables 2 and 3. Graphical representation of logit regressions for the individual toxins are summarized in Figure 2. These regressions were used to estimate the median lethal concentration (LC50) for the toxins.

To study the synergistic larvicidal activity of Cyt2Ba in binary mixtures with other components of the Bti crystal, a series of preliminary bioassays were made, using a single protein concentration (below 30% mortality). The binary combinations studied, as well as the concentration of proteins used in each case, are shown in Table 1. For those binary combinations that resulted in the highest mortality of inoculated insects, quantitative bioassays were performed in order to determine the potentiation between Cyt2Ba and other toxins. Concentration-mortality bioassays were performed for Cry4Aa at concentrations of 486, 162, 54, 18, 6, 2 ng/mL. Mixtures of Cyt2Ba with either Cry10Aa or Cry4Aa in equal proportions were tested at concentrations of 300, 60, 12, 2.4, 4.8 <sup>×</sup> 10−<sup>1</sup> and 9.6 <sup>×</sup> <sup>10</sup>−<sup>2</sup> ng/mL, and 54, 27, 13.5, 6.74, 3.36 and 1.68 ng/mL, respectively. Each bioassay was performed between five and ten times. In all other aspects, the bioassay procedure and data curation was as described above. Graphical representation of logit regressions for all toxin mixtures are summarized in Figure 2. These regressions were used to estimate the median lethal concentration (LC50) for the mixture of the toxins.

#### *5.8. Statistical Analysis*

Concentration-mortality data were subjected to logit regression to estimate the median lethal concentration (LC50) for individual toxins and the mixture of toxins. The significance of treatment and interaction terms was determined by sequential removal of terms from the complete regression model. The observed and expected LC50 values for the individual toxins and the toxin mixture in *A. aegypti* were used to evaluate the interaction of Cyt2Ba with Cry10Aa and Cry4Aa. To calculate the expected LC50 values for the toxin mixture under the null hypothesis of no interaction the "simple similar action" model was used [31]. This model assumes that concentration-response regression lines for different components of a mixture are parallel and is suitable for testing synergism in chemically similar compounds such as Bt toxins. Because Cyt2Ba and Cry4Aa regression lines are not parallel the synergism factor calculated is only correct for the LC50 single point.

The expected LC50 was calculated as follows:

$$\rm{LC}\_{50(m)} = \left[ \frac{r\_A}{\rm{LC}\_{50(A)}} + \frac{r\_B}{\rm{LC}\_{50(B)}} \right]^{-1}$$

where LC50(m) is the expected LC50 of the mixture of toxin A and toxin B, LC50(A) is the observed LC50 for toxin A alone, LC50(B) is the observed LC50 for toxin B alone and rA and rB represent the relative proportions of toxin A and toxin B in the mixture, respectively. All statistical procedures were performed using R software (v.3.5.1).

**Author Contributions:** Conceptualization, D.V.-d.-L., M.V. and P.C.; Data curation, D.V.-d.-L.; Formal analysis, D.V.-d.-L. and T.W.; Funding acquisition, P.C.; Investigation, D.V.-d.-L., M.V. and L.L.; Methodology, D.V.-d.-L. and M.V.; Project administration, P.C.; Resources, P.C.; Software, D.V.-d.-L.; Supervision, M.V., T.W. and P.C; Validation, D.V.-d.-L. and M.V.; Visualization, D.V.-d.-L. and M.V.; Writing—original draft, D.V.-d.-L. and M.V.; Writing—review & editing, D.V.-d.-L., M.V., T.W. and P.C. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research was funded by Spanish Ministry of Science and Innovation (RTI2018-095204-B-C22). D.V.-d.-L. received a doctoral grant from Universidad Pública de Navarra, Pamplona, Spain. L.L. received a doctoral grant from the European Union's H2020 research and innovation programme under Marie Sklodowska-Curie grant agreement N◦ 801586.

**Acknowledgments:** The authors thank Colin Berry (Cardiff University, Cardiff, UK) for providing Bt recombinant strains 4Q2-81 pHT606:*cry4Aa* and 4Q2-81 pHT611:*cry4Ba* used in this study.

**Conflicts of Interest:** The authors declare no conflict of interest. The funders had no role in the design of the study; in the collection, analyses, or interpretation of data; in the writing of the manuscript, or in the decision to publish the results.

#### **References**


© 2020 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).

### *Article* **Protein-Lipid Interaction of Cytolytic Toxin Cyt2Aa2 on Model Lipid Bilayers of Erythrocyte Cell Membrane**

#### **Sudarat Tharad 1,\*, Boonhiang Promdonkoy <sup>2</sup> and José L. Toca-Herrera 1,\***


Received: 3 March 2020; Accepted: 1 April 2020; Published: 3 April 2020

**Abstract:** Cytolytic toxin (Cyt) is a toxin among *Bacillus thuringiensis* insecticidal proteins. Cyt toxin directly interacts with membrane lipids for cytolytic action. However, low hemolytic activity is desired to avoid non-specific effects in mammals. In this work, the interaction between Cyt2Aa2 toxin and model lipid bilayers mimicking the erythrocyte membrane was investigated for Cyt2Aa2 wild type (WT) and the T144A mutant, a variant with lower hemolytic activity. Quartz crystal microbalance with dissipation (QCM-D) results revealed a smaller lipid binding capacity for the T144A mutant than for the WT. In particular, the T144A mutant was unable to bind to the phosphatidylcholine lipid (POPC) bilayer. However, the addition of cholesterol (Chol) or sphingomyelin (SM) to the POPC bilayer promoted binding of the T144 mutant. Moreover, atomic force microscopy (AFM) images unveiled small aggregates of the T144A mutant on the 1:1 sphingomyelin/POPC bilayers. In contrast, the lipid binding trend for WT and T144A mutant was comparable for the 1:0.4 POPC/cholesterol and the 1:1:1 sphingomyelin/POPC/cholesterol bilayers. Furthermore, the binding of WT and T144A mutant onto erythrocyte cells was investigated. The experiments showed that the T144A mutant and the WT bind onto different areas of the erythrocyte membrane. Overall the results suggest that the T144 residue plays an important role for lipid binding.

**Keywords:** Cyt2Aa2 toxin; protein-lipid binding; erythrocyte membrane; AFM; QCM-D

**Key Contribution:** The alanine replacement of the threonine 144 residue reduces the lipid binding ability of the Cyt2Aa2 toxin onto model lipid bilayers (and erythrocyte cells). In particular, it was found that the Cyt2Aa2 T144A mutant did not bind onto POPC bilayers.

#### **1. Introduction**

The most widely known bacteria as a bioinsecticidal agent is *Bacillus thuringiensis* (Bt). It is a Gram-positive rod shape bacterium originally hosted in soil. In the last few decades Bt has been used to control insect larvae especially for pest insects in the form of a bioactive agent or a transgenic plant. The active proteins, Crystal (Cry) and Cytolytic (Cyt) toxins are produced as crystalline proteins during the sporulation phase of the Bt growth cycle [1]. After toxin ingestion by insect larvae, the protein crystals are solubilized and concomitantly activated by proteases in alkaline condition of the mid gut [2–4]. Consequently, the toxins interrupt a cell membrane permeability of the gut cells leading to cell burst because of the osmotic pressure imbalance [5,6]. However, both toxins disrupt the cell membrane with different mechanisms. Cry toxin requires a protein receptor for the cell membrane binding whereas Cyt toxin interacts directly with the membrane lipids [7–9], in particular with the

unsaturated phospholipids [10]. Cry toxin has been used more in crop fields to control insect larvae than Cyt toxin because of its efficiency and specificity [11]. Nevertheless, the long-term application of Cry toxin has led to insect larvae resistance [12]. Accordingly, Cyt toxin has been taken into the strategy to overcome such resistance. Thus, the Cyt toxin is able to be a receptor for the Cry toxin and these toxins can synergize their activities together [13].

Cytolytic toxin Cyt2Aa2 is produced from *Bacillus thuringiensis* subsp. *darmstadiensis* [14]. The Cyt toxin shows the cytolytic activity against a broad range of cell types, e.g., insect cells, mammalian cells [7], and bacterial cells [15]. Previous experiments suggest that the protein-lipid binding mechanism of the Cyt toxin is driven by (i) pore formation [16,17], (ii) detergent-like action [18,19], and (iii) carpet action (protein aggregate) [20]. However, the precise mechanism is still unclear and devotes further investigation. In particular, hemolytic activity has been tested to determine the cytolytic activity of Cyt2Aa2 toxin against erythrocyte cells (in relation to mammalian cells). Therefore, we have tried to obtain a variant with lower hemolytic activity by performing amino acid mutation (in order to reduce the non-specific target to mammalian cells). The effect of the amino acid point mutation of the Cyt2Aa2 molecule on the toxin activity has been reported. Previous studies have shown that the amino acids located in the helix A, helix C [21], and helixD-beta4 loop [22,23] alter the activity of the Cyt2Aa2 toxin. Particularly, we have investigated the amino acid mutation of T144A (alanine replacement of T144 residue) placed in the helixD-beta4 loop (Figure S1) because it keeps its larvicidal activity, although its hemolytic activity is reduced [24].

To elucidate the influence of the point mutation T144A on the interaction of Cyt2Aa2 protein with model lipid bilayers (which mimic the erythrocyte membrane), we have carried out binding studies with the Cyt2Aa2 wild type (WT) and the mutant Cyt2Aa2-T144A. The prepared lipid bilayers containing phospholipid (POPC), sphingomyelin (SM), and cholesterol (Chol) were mixed in various molar ratios to build the different membranes, which can be found in erythrocyte cells [25,26]. The combination of quartz crystal microbalance with dissipation (QCM-D) and atomic force microscopy (AFM) indicated that the T144A mutant had a lower binding capability than the WT, especially for the POPC bilayer. Moreover, the T144A mutant formed small aggregates on the 1:1 SM/POPC bilayer (showing a different binding trend from the Cyt2Aa2 WT). Finally, both toxins were also exposed to lysed erythrocytes cells. It was found that WT and T144A bound to different parts of the erythrocyte membrane.

#### **2. Results**

#### *2.1. Determination of the Cyt2Aa2-Lipid Interaction with Di*ff*erent Model Lipid Bilayers that Mimic the Erythrocyte Membrane by QCM-D*

The lipid components of the cell membrane, phospholipid (POPC), sphingomyelin (SM) and cholesterol (Chol), were mixed in different molar ratios in order to form different lipid bilayers that could mimic the erythrocyte cell membrane. The interaction of both Cyt2Aa2 wild type (WT) and T144A mutant with pure POPC (Figure 1A), 1:0.4 POPC/Chol (Figure 1B), 1:1 SM/POPC (Figure 1C), and 1:1:1 SM/POPC/Chol bilayers was investigated with QCM-D (Figure 1D). The results showed that Cyt2Aa2 WT (black plot) interacted with all kind of lipid bilayers, whereas no interaction between the T144A mutant and the POPC bilayer could be detected. In this case, the frequency (ΔF) and the dissipation (ΔD) signals did not change with time (blue plot). The measurements indicated that the lipid binding of both the Cyt2Aa2 WT and T144A mutant was saturated for ΔF values between −25 to <sup>−</sup>40 Hz. The difference in dissipation (ΔD) achieved values between 2 <sup>×</sup> 10−<sup>6</sup> to 5 <sup>×</sup> 10−<sup>6</sup> (Table 1). Here it is worth remembering that ΔF relates to changes in the mass adsorption (on the sensor surface), and that ΔD refers to the viscoelastic properties of the formed hybrid protein-lipid layer.

**Figure 1.** Protein-lipid binding of Cyt2Aa2 wild type and the T144A mutant on different lipid bilayers. The lipid bilayers were formed on the surface of silica sensors. Once the bilayer was built, the value of the frequency was set to zero. Thus, the reported difference in frequency relates to the adsorption of the protein toxin on the lipid bilayers. The protein solution (25 μg/mL) was filled into the quartz crystal microbalance with dissipation (QCM-D) chamber, and then the flow was paused in order to evaluate the Cyt2Aa2-lipid binding for 2 h. The black arrow and red arrow indicate protein exposure and buffer rinsing, respectively. (**A**) phospholipid (POPC), (**B**) 1:0.4 POPC/cholesterol (Chol), (**C**) 1:1 sphingomyelin (SM)/POPC and (**D**) 1:1:1 SM/POPC/Chol.


**Table 1.** ΔF, ΔD, and lipid binding rate values for wild type (WT) and T144A on different lipid bilayers.

In addition, ΔD-ΔF plots can be used to compare the binding behavior between the WT and the mutant T144A. For binding onto POPC bilayers (Figure 2A), the ΔD-ΔF signal for the T144A remained mostly constant with increasing time, while the WT showed a proportional increasing of ΔD and ΔF. However, the WT and the T144A seemed to bind in a similar way onto 1:0.4 POPC/Chol and 1:1:1 SM/POPC/Chol bilayers (Figure 2B,D), suggesting similar viscoelastic properties. On the contrary, a different trend occurred for the binding onto 1:1 SM/POPC bilayers (Figure 2C). It can be observed that for the same frequency change (ΔF) the mutant induced a final less rigid protein-lipid layer.

**Figure 2.** ΔD-ΔF plots of the binding of Cyt2Aa2 WT (black) and the T144A mutant (blue) on different model lipid bilayers. The dissipation value (ΔD) was plotted against the frequency value (ΔF) to elucidate the interplay between the protein binding and the viscoelasticity of the hybrid protein-lipid layer. The similarity of the slopes indicates an analogous qualitative behavior. (**A**) POPC, (**B**) 1:0.4 POPC/Chol, (**C**) 1:1 SM/POPC, and (**D**) 1:1:1 SM/POPC/Chol bilayers.

Furthermore, the binding kinetics were determined by fitting the experimental data to a single exponential decay equation:

$$F\_t = F\_0 + \mathcal{A}e^{-t/\Gamma} \tag{1}$$

(see Figure S2). In Table 1, the constant decay (Γ) indicates the lipid binding rate. Thus, a lower value means a faster binding rate, and vice versa. It can be observed that the WT showed lower Γ values than the T144A mutant for all types of model lipid bilayers. Hence, the binding rate of the WT was faster than the T144A mutant. Remarkably, the Γ values of the WT corresponding to the 1:0.4 POPC/Chol, 1:1 SM/POPC and 1:1:1 SM/POPC/Chol bilayers (ca. 2.0 min) were approximately five times smaller than the rate for the POPC bilayers (ca. 10.0 min). It seemed that the lipid bilayers containing either cholesterol or sphingomyelin favored the binding of Cyt2Aa2 WT. Similarly, the binding of the T144A mutant could be detected when cholesterol or sphingomyelin were present in the lipid bilayers. However, the Γ values indicate that the lipid binding ability of T144A mutant onto 1:1 SM/POPC bilayers (Γ = 52.1 min) was lower than 1:0.4 POPC/Chol (Γ = 11.2 min) and 1:1:1 SM/POPC/Chol (Γ = 10.9 min) bilayers, respectively. Unlike the WT case, sphingomyelin promoted a lower binding capability of the T144A mutant than cholesterol. These findings indicate that the replacement of threonine 144 with alanine results in a reduction of the lipid binding ability of Cyt2Aa2 toxin, especially for POPC bilayers.

#### *2.2. AFM Imaging of the Cyt2Aa2 (WT and Mutant) Interaction with Di*ff*erent Model Lipid Bilayers*

AFM experiments were carried out in order to investigate the topographic structure of the different Cyt2Aa2-lipid layers. The model lipid bilayers were successfully formed on the silica surface via lipid vesicle fusion and revealed a smooth surface (Figure S3). Subsequently, the protein solutions with the Cyt2Aa2 WT and the T144A mutant were incubated with the lipid bilayers. The surface topography of the hybrid protein-lipid layers was visualized after 30 min of incubation. Figure 3 shows no binding of the T144A mutant on POPC bilayers, which agrees with the QCM-D results. In contrast, the WT toxin almost covered the whole lipid bilayer surface (black areas refer to protein-free lipid bilayer). A longer incubation time of 120 min did not promote the binding of the T144A mutant on the POPC bilayer (Figure S4A). Subsequently, cholesterol and sphingomyelin were included into the lipid mixtures. Cyt2Aa2 WT bound onto the lipid surfaces reaching saturation. Thus, the lipid surfaces were fully covered with Cyt2Aa2 WT (note that the black areas disappeared). In addition, the binding between the T144A mutant and the lipid bilayers could be observed. The binding behavior of the T144A mutant onto 1:0.4 POPC/Chol and 1:1:1 SM/POPC/Chol bilayers showed a similar trend than the trend depicted by the WT; the protein-free membrane (black area) was observed prior to reaching a saturation after 120 min (see Figure S4). Remarkably, the T144A mutant formed small protein aggregates onto 1:1 SM/POPC bilayers. These aggregates seemed to be different from the ones observed for the WT and the T144A mutant on other lipid membranes (Figure 3).

**Figure 3.** Atomic force microscopy (AFM) height images showing the interaction of the Cyt2Aa2 WT and the T144A mutant with the different lipid bilayers. First, the lipid bilayers were formed on silica surfaces. After, both protein solutions (WT and mutant) were exposed to the lipid bilayers for 30 min. The AFM images were collected in tapping mode with a scan rate of 1–2 Hz. Note that the scan size of every image is 5 μm × 5 μm. The vertical scale (until 10 nm) is indicated on the right. Image processing was carried out with the Nanoscope program. (**A**) POPC, (**B**) 1:0.4 POPC/Chol, (**C**) 1:1 SM/POPC, and (**D**) 1:1:1 SM/POPC/Chol bilayers.

Furthermore, the influence of sphingomyelin (SM) on the binding capability of the Cyt2Aa2 toxins was determined. For this purpose, 1:1 SM/DOPC (1,2-dioleoyl-*sn*-glycero-3-phosphocholine) bilayers were exposed to the toxins. For the SM/DOPC bilayers, a phase separation was observed where the sphingomyelin domains appeared as a liquid disordered-solid phase (ld-So). In particular, the So domains of SM were distributed over the lipid bilayer surface being about 1 nm thicker than the DOPC-enriched domains (Figure S5). The 1:1 SM/DOPC bilayers were firstly exposed to the T144A mutant. The observed protein aggregates looked similar to the aggregates found on the SM/POPC bilayers. Subsequently, the WT protein was introduced into the system. The AFM micrographs indicate that the WT fully occupied the remaining areas (DOPC-enriched domains). Furthermore, no protein could be observed on SM domains (Figure 4). This suggests unfavorable binding of the Cyt2Aa2 toxins

onto SM bilayers. Concordantly, AFM and QCM-D results support each other (i.e., no binding of the T144A mutant on the POPC bilayer). Moreover, AFM topography studies provided additional information about the T144A-lipid complex formation onto SM/POPC bilayers and its binding inability onto sphingomyelin bilayers.

**Figure 4.** AFM height images of the Cyt2Aa2 wild type and the T144A mutant on 1:1 SM/DOPC (1,2-dioleoyl-*sn*-glycero-3-phosphocholine) bilayers. The SM domains are indicated as white asterisks. The lipid bilayers were initially exposed to the T144A mutant (25 μg/mL) for 2 h (**A**). After buffer rinsing, the Cyt2Aa2 wild type solution (25 μg/mL) was exposed to the lipid bilayers for 1 h (**B**). The topographic images were collected in tapping mode at a scan rate of 1–2 Hz. Note that the scan size of both images is 5 μm × 5 μm. The vertical scale (until 10 nm) is indicated on the right. Image analysis was performed with the Nanoscope program.

#### *2.3. Cyt2Aa2 (WT and Mutant) Toxin Interaction with Erythrocyte Cell Membranes*

Sheep erythrocytes presented a round and concave shape with diameter of ~3.0 μm under the light microscope. To prepare the erythrocyte membrane layers, a low salt solution (1/3 dilution PBS) was used to break the cell attached on the supporter surface. The ghost erythrocytes appeared as flat cells because of the releasing of cytoplasmic fluid (Figure S6). AFM images revealed a size of ca. 3.0–4.0 μm for the erythrocytes, which agreed with light microscopy observations.

After the erythrocytes were lysed, it was assumed that two types of erythrocyte membrane could be observed: (i) a single layer (inner cytoplasmic membrane) formed by cell opening (inside membrane facing up), and (ii) a double layer presenting the outer surface of the membrane (inner and outer cytoplasmic membranes) (Figure 5). Unlike the model lipid bilayers, the erythrocyte cytoplasmic membrane had a rougher surface. The membranes were firstly exposed to the T144A mutant. The binding of the T144A revealed a change in the topography of the height surface (area surrounding the asterisks). However, some areas of the membrane remained free of the T144A protein (no binding). In a second step, Cyt2Aa2 WT was introduced into the system. Cyt2Aa2 WT bound to the remaining areas leading to a smoother surface (compared to the erythrocyte-T144A ones) (Figure 5). The experiments with model lipid bilayers showed that the T144A mutant could not bind to the POPC bilayer bilayers. Therefore, it was not expected that the mutant would bind on the POPC areas of the erythrocyte membrane. On the contrary, the results of Cyt2Aa2 WT could indicate that WT bind POPC-enriched domains of the erythrocyte membrane.

**Figure 5.** AFM images of Cyt2Aa2 toxin binding on the erythrocyte membrane. The erythrocyte membrane was prepared on a lysine-coated glass (**A**). The cell membranes were initially exposed to the T144A mutant (25 μg/mL) for 2 h (**B**). After buffer rinsing, the Cyt2Aa2 wild type solution (25 μg/mL) was exposed to the same membrane for 1 h (**C**). The topographic images were collected in tapping mode at a scan rate of 1–2 Hz. The images were analyzed with the Nanoscope program. The images have a scan size of 3 μm × 3 μm (upper panel) and 1.8 μm × 1.8 μm (lower panel). Note that the vertical scale differs: from 0 to 30 nm (upper panel), and from 0 to 15 nm (lower panel). The white asterisks mark the same areas on the erythrocyte membrane.

#### **3. Discussion**

In this work, we have studied the interaction of two Cyt2Aa2 proteins, the WT and the less hemolytic T144A mutant [24], with model lipid bilayers and erythrocyte membranes. QCM-D results showed that the T144A mutant could not bind to the POPC bilayer. In turn, the mutant protein retained its binding capability when exposed to 1:0.4 POPC/Chol, 1:1 SM/POPC, and 1:1:1 SM/POPC/Chol bilayers (Figure 1). Moreover, the ΔD–ΔF plots suggest that the binding behavior of the WT and T144A mutant followed similar adsorption trends except for 1:1 SM/POPC bilayers (Figure 2). The final values of ΔF and ΔD were not significantly different for the Cyt2Aa2 WT and the T144A mutant (Table 1). Our values are similar to the reported values for the binding of perforin on lipid membranes [27]. However, the mutant presented a lipid binding rate of at least five times lower than the WT, suggesting a smaller binding ability of the T144A mutant. The presence of either cholesterol or sphingomyelin in the lipid bilayer increased the binding rate of the WT and promoted the binding of the T144A mutant on the model lipid bilayers. At room temperature (25 ◦C), the POPC bilayer exists in a liquid disordered phase (ld). Addition of cholesterol or sphingomyelin into lipid membranes reduces the fluidity of the lipid bilayer (less lateral diffusion) [28]. Less dynamic membranes seem to increase the possibility for the Cyt2Aa2-lipid interaction. Hence, Cyt2Aa2 WT bound faster on the 1:0.4 POPC/Chol, 1:1 SM/POPC and 1:1:1 SM/POPC/Chol bilayers than on the single POPC bilayer. In addition, it was observed that the T144A mutant could also bind onto these heterogeneous membranes, but not onto POPC bilayers. It seems that although the membranes of insect cells and mammalian cells are different in composition, both support the larvicidal activity and the low hemolytic activity of the T144A mutant. In comparison to mammalian cells, insect cell membranes present a lower amount of cholesterol and sphingomyelin [29]. This might suggest that the binding of the Cyt2Aa2 toxin on insect cells is possibly promoted by other components of the cell membrane (e.g., membrane proteins) besides cholesterol and sphingomyelin. This different binding mechanism might play a role in the in vivo larvicial activity of the Cyt2Aa2 toxin.

Atomic force microscopy (AFM) was carried out to investigate the surface structure of the hybrid Cyt2Aa2-lipid bilayers. The AFM measurements confirmed that the T144A mutant does not bind onto POPC bilayers, while the WT toxin adsorbs on such lipid bilayers (Figure 3). With the addition of cholesterol into this lipid bilayer, the 1:0.4 POPC/Chol bilayer led to the binding of the T144A mutant. On the contrary, sphingomyelin (1:1 SM/POPC bilayer) also promoted the T144A mutant binding but a dissimilar binding was observed as well as small protein aggregates. Correspondingly, the ΔD–ΔF plots of QCM-D results suggest dissimilar binding behavior between the WT and the T144A mutant on the 1:1 SM/POPC bilayers (Figure 2). Furthermore, the binding behavior of Cyt2Aa2 WT and the T144A mutant were very much alike once cholesterol was included into the lipid membranes, as found for the 1:0.4 POPC/Chol and 1:1:1 SM/POPC/Chol bilayers (Figure 3). Moreover, both Cyt2Aa2 WT and the T144A mutant did not bind onto sphingomyelin (So) domains (Figure 4). These results suggest that although the lipid head group plays a role in the interaction between Cyt2Aa2 toxin and lipid bilayers, the lipid phase might also be taken into account [30,31]. This could be feasible since sphingomyelin contains the same choline head group as POPC. Besides, the small aggregates of the T144A mutant might imply a coexistence of a different fluid membrane in the 1:1 SM/POPC bilayer. This particular result indicates that cholesterol is more important for the binding behavior of the T144A mutant than for the binding of Cyt2Aa2 WT.

Furthermore, the interaction of Cyt2Aa2-lipid (WT and mutant) with sheep erythrocyte membranes was investigated. Unlike model lipid bilayers, a rougher surface was detected for the sheep erythrocyte membrane, which is comparable to the chicken erythrocyte membrane [32], and the human erythrocyte membrane [33]. The T144A mutant revealed a limited binding capability on the erythrocyte membrane, leaving part of the membrane uncovered. Further experiments indicated that the Cyt2Aa2 WT could bind on the remaining free areas (Figure 5). This binding study provides insight about the different properties of the erythrocyte membranes, e.g., lipid composition and lipid fluidity, when compared with model lipid membranes. Nowadays, a detection of the lipid phase coexistence in biological cell membranes is still a challenge for a biologist. Although lipid phase separation could not be observed in this experiment, the two Cyt2Aa2 proteins enabled us to distinguish the different components of the lipid membrane of sheep erythrocytes.

In conclusion, the combination of QCM-D and AFM permitted us to monitor the interaction between CytAa2 toxin with model lipid bilayers and supported erythrocyte membranes. The lower protein-lipid binding capability of the T144A mutant (in comparison with the WT) could lead to its small hemolytic activity. In particular, the alanine replacement of threonine 144 residue disables the binding properties of the Cyt2Aa2 toxin onto the POPC bilayer. Although certain hemolytic activity still remains for the T144A mutant, it can be said that the T144 residue located in the αD-β4 loop plays an important role in the Cyt2Aa2-lipid binding. Furthermore, the modification of amino acid residues in the αD-β4 loop of the Cyt2Aa2 toxin will be investigated for specific cell targeting. In future work, the effect of different amino acid properties (e.g., polar charge and positive charge) on the Cyt2Aa2-lipid interaction will be investigated. In addition, protein concentration, lipid phase and lipid charge will be taken into account for further investigations.

#### **4. Materials and Methods**

#### *4.1. Reagents and Bu*ff*er*

1-palmitoyl,2-oleoyl-sn-glycero-3-phosphocholine (POPC), 1,2-dioleoyl-*sn*-glycero-3-phospho choline (DOPC), chicken egg yolk sphingomyelin (SM), and cholesterol (Chol) were purchased from Sigma-Aldrich (Darmstadt, Germany). The lipids were dissolved in chloroform and divided into 1 mg aliquots. Then, the organic solvent was evaporated under nitrogen stream and kept at −20 ◦C.

Phosphate buffered saline (PBS) pH 7.4 (137 mM NaCl, 2.7 mM KCl and 10 mM phosphate) was prepared from PBS tablet (Sigma-Aldrich, Darmstadt, Germany). The buffer tablet was dissolved in ultrapure water (Milli-Q, Merck, Darmstadt, Germany) and filtrated through a 0.22 μm filter (Whatman, GE Health care life science, Chicago, IL, USA).

#### *4.2. Protein Preparation*

The Cyt2Aa2 toxin from *Bacillus thuringiensis* subs. *darmstadiensis* was expressed in *Escherichia coli* as previously described by B. Promdonkoy [14]. The amino acid replacement at the threonine 144 residue with alanine was carried out by means of site-directed mutagenesis as described in a previous publication [24]. To obtain activated Cyt2Aa2 toxin (25 kDa), the Cyt2Aa2 inclusion was solubilized in 50 mM carbonate buffer, pH 10.0 at 30 ◦C for 1 h. The soluble Cyt2Aa2 toxin (29 kDa-protoxin) was collected by centrifugation at 10,000× *g* for 10 min. Then, the Cyt2Aa2 toxin was activated by 2% (*w*/*w*) chymotrypsin (Sigma-Aldrich, Darmstadt, Germany) at 30 ◦C for 2 h. The purity of the protein was determined by SDS-PAGE (Invitrogen, Waltham, MA, USA). Protein concentration was determined by UV adsorption (Hitachi, Tokyo, Japan). Stock protein solution was prepared to 2.0 mg/mL (80 μM) and kept at −20 ◦C.

#### *4.3. Lipid Vesicle Preparation*

The lipids were mixed in chloroform with the desired lipid ratios. After that, the organic solvent was evaporated under a gentle nitrogen stream to form lipid films. The residual solvent was removed by further keeping the lipid films under nitrogen stream for 1 h. Furthermore, the lipid films were hydrated with PBS solution to a concentration of 1 mg/mL and incubated above the melting transition temperature (*Tm*) for 2 h. The hydrated films were intermittently vortexed during incubation until complete suspension. The vesicles were homogenized by extrusion method for low *Tm* lipid mixtures (POPC/Chol system). The vesicles were pressed through a 50 nm Øpolycarbonate membrane for 21 times at room temperature by using a mini-extruder (Avanti, Alabaster, AL, USA). For the lipid mixtures with higher *Tm* (SM system) tip sonication with a 50% duty cycle of 10 min was used (Branson sonifier, Emerson, Ferguson, MO, USA). Then, the residual material was removed by centrifugation at 10,000× *g* for 10 min. After that, the vesicles size, in a range of 100–130 nm, was determined by Zetasizer Nano ZS (Malvern Instrument, Worcestershire, UK). The vesicle solutions were stored at a temperature higher than *Tm* and were used within a week.

#### *4.4. Supported Erythrocyte Cell Membrane Preparation*

Sheep blood (Oxoid, Thermo scientific, Waltham, MA, USA) was removed by washing with PBS pH 7.4 three times. The sheep blood was gentle mixed with PBS in a ratio of 1:7. Then, the erythrocytes were collected by centrifugation at 3000× *g* and 4 ◦C for 5 min. The erythrocyte pellet was kept and resuspended in PBS pH 7.4, 2% (*V*/*V*), as a working solution.

The erythrocyte membrane was prepared on a poly-lysine coated glass. The round-glass cover slips were cleaned as follows: soaking in 1.0 M hydrochloric acid for 2 h, rinsing thoroughly with ultrapure water (MilliQ, Merck, Darmstadt, Germany), sonication in 70% (*v*/*v*) ethanol for 10 min, and final treatment with plasma cleaner (Diener electronic, Ebhausen, Germany). Prior cell attachment, the glass cover slips were coated with 30–70 kDa poly L-lysine (Sigma, Darmstadt, Germany). The glass slips were immersed in a 0.1 mg/mL lysine solution (in PBS) for 30 min at room temperature. The excess of lysine was removed by buffer rinsing. After that, the erythrocytes were attached on the glass surface by incubation over the surface for 30 min at room temperature. The unbound erythrocytes were removed and the attached cells were opened under shear flow by using a low content salt solution (1/3 dilution PBS; 45.7 mM NaCl, 0.9 mM KCl and 3.3 mM phosphate). Finally, the cell membrane was rinsed with PBS pH 7.4.

#### *4.5. Quartz Crystal Microbalance with Dissipation (QCM-D) Measurement*

The protein-lipid bilayer interaction was evaluated with quartz crystal microbalance with dissipation from Q-Sense E4 (Biolin Scientific, Gothenburg, Sweden) using silica-coated sensors (QSX 303, Biolin Scientific, Sweden). Before use, the sensors were subsequently cleaned as follows: sonication in 2% (*w*/*w*) SDS solution for 15 min, rinsing with ultra-pure water, drying under nitrogen stream, and organic residues-eliminating with UV/Ozone cleaner (Bioforce Nanosciences, Salt Lake City, UT, USA) for 30 min. The frequencies of the sensors were evaluated prior to running the experiments. The outcome of the experiments delivers changes in frequency and dissipation. The change in frequency (Δ*F*) is proportional to changes in the adsorbed mass (Δm) on the crystal surface through the Sauerbrey equation:

$$
\Delta m = -\frac{\mathcal{C}}{n} \Delta F \tag{2}
$$

where (*C*) is the sensitivity constant (−17.7 ng cm−<sup>2</sup> Hz−1) and (*n*) is the overtone number. Simultaneously, the change in dissipation (Δ*D*) indicates the viscoelastic properties of the new forming layer on the crystal surface (in our case, of the hybrid protein-bilayer system). Low dissipation values are typical for a rigid (elastic) layer whereas high values relate to softer (viscoelastic) layers. The changes in frequency (Δ*F*) and dissipation (Δ*D*) values are presented for the 5th overtone unless otherwise stated.

The lipid bilayers were formed by the lipid vesicle fusion method. After a stable baseline with PBS solution was achieved, 0.1 mg/mL lipid vesicle solutions were slowly flowed in the QCM-D chamber with a flow rate of 50 μl/min. Once the characteristic patterns (for the frequency and dissipation) of lipid bilayer formation were observed, the excess of vesicles was removed by buffer rinsing. Some of the lipid bilayers were completely formed by additional water rinsing (through osmotic stress). Finally, all lipid bilayers were incubated under PBS flow until reaching a stable baseline.

To study the interaction of the toxin with the model lipid bilayers, the different Cyt2Aa2 toxin solutions were introduced into the system at a flow rate of 50 μL/min. After that, the flow was stopped in order to evaluate when the protein-lipid binding could reach a saturated state. Furthermore, the unbound protein was flushed from the chamber with PBS solution at a flow rate of 50 μL/min for 30 min. The experiments were carried out with at least three replications at 25 ◦C (298 K). The protein-lipid binding kinetics was determined by curve fitting. The frequency (ΔF) vs. time plots were fitted with a single exponential decay equation (Equation (1)). All the plots and data fitting were carried out with Origin 8.0 (OriginLab Corporation, Northampton, MA, USA).

#### *4.6. Atomic Force Microscopy (AFM) Imaging*

The AFM cantilevers with a nominal spring constant of 0.24 N/m (DNP-S10, Bruker, Billerica, MA, USA) and the silica substrate were mounted inside a closed fluid cell with an O-ring. The 1 cm × 1 cm silica wafers (IMEC, Leuven, Belgium) were cleaned before using with the following procedure: sonication in 2% (*w*/*w*) SDS solution for 15 min, rinsing with ultrapure water, and drying under nitrogen stream. Finally, the substrates were treated by plasma cleaner (Diener electronic, Ebhausen, Germany). The lipid bilayers were formed by means of lipid vesicle fusion. 0.1 mg/mL of lipid vesicle solutions were incubated over the silica surface for at least 10 min and then the vesicle excess was rinsed from the chamber. Afterwards, the two Cyt2Aa2 proteins, wild type (WT) and the T144A mutant (25 μg/mL or 1.0 μM), were incubated with the lipid bilayers or with supported erythrocyte membrane for the desired experimental time. The surface topography was imaged in tapping mode with a JV-scanner controlled by a NanoScope V controller (Bruker, Billerica, MA, USA) at a scan rate of 1–2 Hz. The images were processed and analyzed with the Nanoscope program. The experiments were carried out at room temperature (298 K).

*Toxins* **2020**, *12*, 226

**Supplementary Materials:** The following are available online at http://www.mdpi.com/2072-6651/12/4/226/s1, Figure S1: Three dimensional structure of the Cyt2A toxin (PDB 1CBY). Figure S2: Curve fitting of the frequency vs. time plots of the binding between the Cyt2Aa2 toxins and the model lipid bilayers. Figure S3: AFM images of the lipid bilayers. Figure S4: Time sequence AFM imaging of the binding of the Cyt2Aa2 toxins on lipid bilayers. Figure S5: AFM height images and profile analysis of hybrid Cyt2Aa2-1:1 SM/DOPC bilayers. Figure S6: Cell shape of sheep erythrocytes under light microscopy.

**Author Contributions:** Conceptualization, S.T. and J.L.T.-H.; methodology, S.T.; validation, J.L.T.-H.; formal analysis, S.T.; investigation, S.T. and J.L.T.-H.; resources, J.L.T.-H. and B.P.; data curation, S.T. and J.L.T.-H.; writing—original draft preparation, S.T.; writing—review & editing, J.L.T.-H. and B.P.; visualization, S.T.; supervision, J.L.T.-H.; project administration, J.L.T.-H.; funding acquisition, J.L.T.-H. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research was funded by The Austrian Science Fund (FWF), grant number P29562-N28 and the APC was funded by BOKU Vienna Open Access Publishing Fund.

**Acknowledgments:** The authors thank Jacqueline Friedmann for technical support.

**Conflicts of Interest:** There are no conflicts to declare.

#### **References**


© 2020 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).

## **Structural and Functional Insights into the C-terminal Fragment of Insecticidal Vip3A Toxin of** *Bacillus thuringiensis*

#### **Kun Jiang 1,**†**, Yan Zhang 1,**†**, Zhe Chen 1,**†**, Dalei Wu 1,2, Jun Cai <sup>3</sup> and Xiang Gao 1,\***


Received: 10 June 2020; Accepted: 3 July 2020; Published: 5 July 2020

**Abstract:** The vegetative insecticidal proteins (Vips) secreted by *Bacillus thuringiensis* are regarded as the new generation of insecticidal toxins because they have different insecticidal properties compared with commonly applied insecticidal crystal proteins (Cry toxins). Vip3A toxin, representing the vast majority of Vips, has been used commercially in transgenic crops and bio-insecticides. However, the lack of both structural information on Vip3A and a clear understanding of its insecticidal mechanism at the molecular level limits its further development and broader application. Here we present the first crystal structure of the C-terminal fragment of Vip3A toxin (Vip3Aa11200–789). Since all members of this insecticidal protein family are highly conserved, the structure of Vip3A provides unique insight into the general domain architecture and protein fold of the Vip3A family of insecticidal toxins. Our structural analysis reveals a four-domain organization, featuring a potential membrane insertion region, a receptor binding domain, and two potential glycan binding domains of Vip3A. In addition, cytotoxicity assays and insect bioassays show that the purified C-terminal fragment of Vip3Aa toxin alone have no insecticidal activity. Taken together, these findings provide insights into the mode of action of the Vip3A family of insecticidal toxins and will boost the development of Vip3A into more efficient bio-insecticides.

**Keywords:** *Bacillus thuringiensis*; Vip3A; 3D-structure; mode of action; biological control

**Key Contribution:** Here we showed the first atomic structure of the C-terminal fragment of Vip3A toxin. Our study revealed the general domain organization and the potential function of each domain of C-terminal Vip3A family toxin. It also showed the interesting convergent evolution between Vip3A toxin and Cry toxin.

#### **1. Introduction**

The entomopathogenic bacteria *Bacillus thuringiensis* (Bt), is the most widely used microbial insecticide in the world [1,2]. It is renowned for its ability to produce insecticidal crystal proteins (Cry toxins) during its sporulation phase, which have been widely used in the prevention and control of agricultural pests through the development of transgenic plants or Bt-based biopesticides [3–5]. However, many pests are not sensitive to Cry toxins, and the development of insect resistance has also been reported [1,6,7]. The successful application of Cry proteins, coupled with their limitations,

has spurred on intensive research seeking to identify and characterize novel classes of insecticidal toxins that can be developed for agricultural purposes.

Vegetative insecticidal proteins (Vips), which are produced by Bt during its vegetative stages, have a wide spectrum of insecticidal activity, especially against lepidopteran pests [8]. To date, ~150 distinct Vip toxins have been identified, which have been classified into four families (Vip1, Vip2, Vip3 and Vip4) based on their sequence similarity [9]. Among the Vip toxin family, Vip3A toxins are the most abundant and most studied [8]. Compared with known Cry toxins, Vip3A toxins share no sequence homology, bind to different receptors [10–13], and lack cross-resistance [14–17], therefore they are considered as ideal options to complement and expand the use of Bt in crop protection and resistance management. At present, the Vip3Aa toxin is the only family member that has been used in commercial transgenic crops together with Cry toxins, and no field-evolved resistance has yet been reported [1,8,18]. However, the lack of structural information and incomplete understanding of its mechanisms of action have severely limited the further development of Vip3A as a tool in pest control.

Vip3A toxins are large proteins (~789 amino acids) consisting of a conserved N-terminus and a variable C-terminal region. The ~88kDa Vip3A protoxin could be digested by insect midgut juices into two fragments: a ~20 kDa fragment corresponding to the N-terminal 198 amino acids, and a ~65 kDa fragment corresponding to the C-terminal fragment of Vip3A protein, which is regarded as an essential step for its activation and toxicity [12,19–23]. Since their discovery in 1996 [24], Vip3A proteins have been the subject of intensive research. It has been reported that Vip3A stimulates membrane pore formation and apoptosis upon binding to target cells, which is proposed to be responsible for its cytotoxic effects [12,25–27]. The scavenger receptor class C like protein (Sf-SR-C) and the fibroblast growth factor receptor (Fgfr) have been reported as potential receptors for Vip3A [10,11]. Vip3Aa16 and Vip3Af1 have been subjected to in silico modelling, and three domains and five domains were proposed respectively [28,29]. Quan et al. propose a map of Vip3Af protein with five domains based on the altered protease digestion patterns through the Vip3Af alanine mutants [23]. In addition, Vip3Ag protoxin and the trypsin-activated toxin were found to be a potential tetrameric complex according to the surface topology obtained by transmission electron microscopy [20]. Recently, Zheng et al. reported the crystal structure of a Vip3B protoxin like protein: Vip3B2160 [30], which shares around 60% sequence identity to Vip3A. The overall structure of Vip3B2160 shows a five-domain organization and forms a novel tetramer structure assembly. However, the atomic structure of Vip3A is still not available, which makes it difficult to reveal the relationship between its structure and function accurately.

Here, we report the crystal structure of the C-terminal fragment of Vip3A toxin (Vip3Aa11200–789). The structure shows a four-domain organization which is likely to be conserved for all insecticidal Vip3A family toxins. We identify conserved hydrophobic α-helices in domain II, which we predict to be involved in the membrane insertion process. Structure-guided cell binding assays reveal that domain III may have a central role in host cell targeting and binding of Vip3A toxins. Structural analysis indicates that Vip3A toxins have potential for glycan binding through domains IV and V. Together, our structural and functional studies provide new insights into the molecular mechanisms underlying the mode of action of insecticidal Vip3A toxins.

#### **2. Results**

#### *2.1. Overall Structure of Vip3Aa11200–end*

We used a Vip3A toxin from Bt strain C9, which has been named Vip3Aa11 (GenBank accession No. AY489126.1) in this study. Full-length Vip3Aa11 consists of 789 amino acids, which have been demonstrated to be digested between residues K198 and D199 by insect midgut juice [19–21]. We initiated our crystallization trial with both Vip3Aa11 protoxin and Vip3Aa11199–end. Using spare matrix crystallization screening, we only identified one condition that yielded needle-shaped crystals of Vip3Aa11 protoxin. However, the crystals diffracted to only ~15 Å and could not be improved despite extensive effort. No crystals were observed for the Vip3Aa199–end construct despite screening more than

1000 crystallization conditions. However, when we deleted the N-terminal amino acid (Asp199) from the Vip3Aa11199–end, we obtained the crystal of Vip3Aa11200–end, which diffracted to ~6 Å. Through the addition of an N-terminal MBP (Maltose Bind Protein) tag, we were able to isolate crystals with improved diffraction. The structure was solved using a combination of anomalous phasing with a selenomethionine derivative crystal of Vip3Aa200–end and molecular replacement using MBP as a model in native crystals. The final structure of Vip3Aa11200–end was refined to a 3.2 Å resolution with R and Rfree values of 0.1980 and 0.2389, respectively (Table S1).

The structure shows that the Vip3Aa11 C-terminal fragment is comprised of four domains (Figure 1A,B). Vip3Aa11 could be digested between residues K198 and D199, and residues 1–198 are lacking in our structure. For a better description based on the full-length Vip3A, we assume that Vip3Aa111–198 is a separate domain. Then the protoxin can be divided into five domains, starting from N-terminus: domain I, 1–198; domain II, 199–327; domain III, 328–518; domain IV, 537–667; and domain V, 679–789 in Vip3Aa11 (Figure 1A and Figure S1). The overall structure of the Vip3Aa11200–end resembles a lobster, wherein domains II and III form the body, and domains IV and V are the claws (Figure 1B,C). The connection between domain II and domain III is compact. However, domains III/IV and IV/V are connected by long and flexible loops, which indicates that the relative locations and orientations of these two domains could change under different biological circumstances. There are over 100 known proteins of the Vip3A family. Based on their high degree of sequence conservation and previous studies [8], they are very likely to share similar overall structures and domain compositions.

**Figure 1.** Overall structure of vegetative insecticidal protein Vip3Aa11200–end. (**A**) Domain organization of Vip3A. (**B**) Two views of the overall structure of Vip3Aa11200–end monomer colored as in (**A**). (**C**) Two views of the surface model of Vip3Aa11200–end monomer colored as in (**A**). The black arrow indicates the angle of rotation around the central axis.

The crystal belongs to the P21 space group and four MBP-Vip3Aa11 molecules were found in one asymmetric unit (Figure S2). These four molecules form two dimers in different orientations. PISA [31] determined that there was limited interaction between the two dimers, indicating that their association was caused by crystal packing. Notably, the two Vip3Aa11 molecules in the "dimer" showed moderate conformational variations, with a core root mean square deviation (r.m.s.d) of 1.234 Å among 468 Cα atoms (Figure S3). Superimposition of separate domains between the two molecules revealed better alignment for domains III, IV and V, but not for domain II (Figure S3), suggesting that domain II might potentially be involved in the conformational changes during the activation of Vip3A toxins. Due to their high similarity, we used the monomeric structure of Vip3Aa11200–end for subsequent analysis.

#### *2.2. Domain II Contains a Conserved Hydrophobic Architecture*

Domain II of Vip3Aa11 (residues199–327) consists of five helices, which form two layers (Figure 2A). The outer layer facing the solution contains two short helices, α2 and α3, while the inner layer that contacts with domain III consists of three anti-parallel helices α1, α4 and α5. The outer layer contacts with the upper portion of the inner layer and is almost perpendicular to the inner layer. Among these five helices, helix α4 is the longest. It spans around 45 Å and contains 30 amino acid residues, starting from E267 at the N-terminus to L296 at the C-terminus. Electrostatic surface potentials analysis shows that the majority of charged and polar amino acid residues locate at the N-terminal and C-terminal ends of helix α4 (Figure 2B). For the middle portion of helix α4, from F274 to L289, 75% amino acid residues are hydrophobic residues. Sequence alignment through Vip3 family shows that the hydrophobic region of helix α4 is very much conserved and it is also the most agminated hydrophobic region of Vip3 family proteins (Figure S1). Close to helix α4, helix α1 also shows several conserved hydrophobic amino acid residues facing helix α4 (Figures S1 and S4).

**Figure 2.** Domain II of Vip3Aa11 shows a conserved hydrophobic surface. (**A**) Two views of structure of Vip3Aa11 domain II shown as a ribbon cartoon. (**B**) Two views of the surface model of helix α4 from domain II show its surface charge distribution. (**C**) The highly conserved amino acid residues from Vip3 family sequence alignment (Figure S1) are highlighted in the Vip3Aa structure with red color. (**D**) Two views of the surface model of Vip3Aa11 domain II show its surface charge distribution. The conserved hydrophobic surface is highlighted by black square. (**B**,**D**) The surface is colored as the basis of electrostatic potential with positive charged surface in blue and negatively charged area in red. The black arrow indicates the angle of rotation around the central axis.

Based on the sequence alignment (Figure S1), all the conserved amino acid residues were highlighted on the Vip3Aa11200–end structure (Figure 2C). This shows that the sequence of domain II is highly conserved (about 62%), only slightly lower than that of domain I (about 68%). Electrostatic surface potential analysis shows that there is an obvious hydrophobic surface, which is mainly contributed to by the conserved helix α1 and α4 (Figure 2D).

#### *2.3. Domain III Is Involved in Cell Binding of Vip3A Toxin*

Domain III of Vip3Aa11 (residues 328–518) consists of twelve β strands and one short α-helix at the C-terminal end (Figure 3A). Twelve β strands comprise three antiparallel β sheets sharing a similar "Greek-key" topology (Figure 3A) with a hydrophobic center featuring highly conserved residues V349, F360, I362 and L370 from β sheet I, I425 and F427 from β sheet II, and I481, F492 and L505 from β sheet III (Figure 3B). The results from the DALI server [32] showed that the fold of domain III is similar to that of domain II of the three domain Cry (3d-Cry) family of insecticidal toxins, which has been shown to be involved in host cell receptor recognition and binding [4]. We therefore sought to explore whether domain III serves as a receptor binding domain for Vip3A toxins.

To explore this hypothesis, we used *Spodoptera frugiperda* cells (Sf9 cells), which have previously been shown to be specifically targeted by Vip3 toxins [11,33]. To determine which domain(s) of Vip3Aa11 interact with Sf9 cells, we carried out fluorescence-based cell binding assays using different C-terminal RFP-tagged Vip3Aa truncation derivatives (shown schematically in Figure 3C). As shown in Figure 3D,E and Figure S5, while domain IV-V does not show detectable binding to Sf9 cells, the binding of a construct featuring only domain I-III or II-III to Sf9 cells is indistinguishable from that of full-length Vip3Aa. The interaction of domain III alone with Sf9 cells is significantly stronger than that of the domain I-II construct, indicating that domain III may have a central role in Vip3A receptor binding to Sf9 cells. In addition, Domain II-III shows higher binding than Domain III alone to Sf9 cells, and structural analysis shows that domain II and domain III have close interaction, suggesting that the presence of domain II is also important for cell binding.

#### *2.4. Domains IV and V Are Glycan Binding Motifs*

Both domains IV and V are all β-sheets folds (Figure 4A,B). Unlike domains II and III, which have compact organization, domains III/IV and IV/V are connected by long and flexible loops (Figure 4A). In addition to these loops, there are several polar interactions between domains IV/V and domain III, that reduce the flexibility and fix domains IV and V at the observed positions and orientations (Figure 4A).

Domains IV and V are both built from two anti-parallel sheets of β sandwich, forming the "jelly-roll" topology. Despite showing only 17% sequence identity (Figure 4C), domains IV and V align very well structurally, with a root-mean-squared deviation (r.m.s.d) of 1.299 Å over 61 Cα atoms (Figure 4D). To examine the potential function of these two domains, we searched for their structural homologues using the DALI server [32]. The results for both domains show a very high similarity (Z score > 10) to family 16 carbohydrate binding module (CBM16) of S-Layer associated multidomain endoglucanase (RCSB ID 2ZEY). Superimposition of domains IV, V and CBM16 demonstrates that these three motifs share a similar fold (Figure 4E), suggesting that they are likely to share a related function as well. CBM16 is a carbohydrate-binding domain of the highly active mannanase from the thermophile *Thermoanaerobacterium polysaccharolyticum* with high specificity toward β-1,4-glucose or β-1,4-mannose polymers [34]. Analysis of the electrostatic surface potential shows that both domains IV and V have a surface pocket at a similar position to a sugar-binding pocket of the CBM16 domain (RCSB ID 2ZEY), although all three pockets have different shapes and charge distributions (Figure 4F). Taken together, our structural analysis indicates that domains IV and V of Vip3A both contain a conserved glycan binding motif and that these motifs may target different sugars.

**Figure 3.** Domain III is a potential receptor binding domain. (**A**) Overall structure of Vip3Aa11 domain III shown as a ribbon cartoon. Two views of three antiparallel β sheets from domain III are shown in three different colors, the black arrow indicates the angle of rotation around the central axis. (**B**) Two views of the surface model of domain III of Vip3Aa11. Inside the domain III, there is a conserved hydrophobic core, and the conserved hydrophobic amino acid residues from three antiparallel β sheets are shown as sticks, the black arrow indicates the angle of rotation around the central axis. (**C**) The schematics of C-terminal RFP (red fluorescent protein)-tagged Vip3Aa and its truncation derivatives. (**D**) Fluorescence microscope images of Sf9 cells treated with Vip3Aa-RFP or its truncations, which were labeled with C-terminal RFP tag, for 6 h. Nuclei are stained with DAPI (blue). (**E**) Quantification of the number of Sf9 cells that can be bound by RFP-tagged Vip3Aa and its truncations of Figure 3D in a blind fashion (*n* = 100 cells per sample). Data are expressed as the mean ± SD from three independent experiments. ns, nonsignificant; \*\*, *p* < 0.01; \*\*\*, *p* < 0.001. Scale bar: 10 μm.

**Figure 4.** Domains IV and V of Vip3Aa11 have glycan binding motifs. (**A**) Domain architectures of domains III, IV and V of Vip3Aa11. The polar interactions between domain IV, V and domain III are shown as sticks. (**B**) Overall structure of Vip3Aa11 domain IV and V shown as a ribbon cartoon. (**C**) Amino acid sequence alignment between domain IV and domain V of Vip3Aa11. The identical residues are denoted in white characters and red background, and the similar residues are denoted in red. ClustalX2 was used to perform the sequence alignment [35]. ESPript-3.0 was used to generate the figure [36]. (**D**) Two views of structure superimposition between domain IV and domain V of Vip3Aa11 shown as a ribbon cartoon. Color of each domain is consistent with Figure 4B. (**E**) Structure superimposition between domains IV, V of Vip3Aa11 and glycan bound CBM16 (RCSB ID 2ZEY) shown as a ribbon cartoon. Domains IV and V are colored as Figure 4B, and CBM16 is shown in magenta color. The glycan in CBM16 is shown as stick in light brown color. (**F**) Surface charge distribution of the sugar-binding pocket of CBM16 (RCSB ID 2ZEY) and potential sugar-binding pocket of domains IV, V of Vip3Aa11, highlighted with the orange circle.

#### *2.5. Purified Vip3Aa11200–end Has no Insecticidal Activity*

The C-terminal fragment of Vip3A has been considered to be the toxic core [8]. To verify whether the purified Vip3Aa200–end still have insecticidal activity, cytotoxicity assays and insect bioassays were carried out. As shown in Figure 5A, the purified full length Vip3Aa toxin has significant toxicity to Sf9 cells, while Vip3Aa199–end, Vip3Aa200–end and MBP-Vip3Aa200–end have no toxicity to Sf9 cells. In addition, bioassay results showed that wild-type Vip3Aa was highly toxic against *S. exigua* at the concentration of 200 ng/cm2. However, the purified Vip3Aa199–end, Vip3Aa200–end and MBP-Vip3Aa200–end have no obvious insecticidal activity to *S. exigua* larvae even at the concentration of 2000 ng/cm<sup>2</sup> (Figure 5B). These results indicate that the purified C-terminal fragment of Vip3A alone has no insecticidal activity.

**Figure 5.** Cytotoxicity assays and insect bioassays of different Vip3A constructs. (**A**) Cell viability of Sf9 treated with different Vip3A constructs (50 and 100 μg/mL). (**B**) Mortality analysis of *S. exigua* caused by different Vip3A constructs (200 and 2000 ng/cm2). Data are expressed as the mean <sup>±</sup> SD from three independent experiments; ns, nonsignificant; \*\*\* *p* < 0.001; one-way ANOVA with Dunnett's method.

#### **3. Discussion**

Vip3A toxins show a wide spectrum of specific insecticidal activities and are functionally distinct compared to the Cry toxins. These features make them good candidates for combined application with Cry toxins in transgenic crops to broaden the insecticidal spectrum and to prevent or delay resistance [1,8,11]. The structural features and insecticidal mechanisms of Cry toxins have been studied in detail, which has been crucial to their widespread application [2–4,6]. However, despite the fact that Vip3A toxins were identified almost 25 years ago [24], their mode of action remains poorly understood. One of the main reasons for this is the lack of a high-resolution three-dimensional structure, which significantly impedes detailed molecular-level functional and mechanistic studies, and thus limits the development of their insecticidal potential. In this study, we report the first crystal structure of the C-terminal fragment of Vip3A toxin, which provides a badly-needed framework to explore the molecular-level functional details of Vip3A-family toxins.

Although the amino acid sequence similarity between the Vip3A family toxin and the 3d-Cry toxin is very low, our three-dimensional structural analysis showed interesting convergent evolution between these two families. Domain II of Vip3A has an all α-helix fold, including two conserved hydrophobic α-helices. Similarly, domain I of 3d-Cry also has an all α-helix fold and two hydrophobic α-helices, although it has additional α helices surrounding the conserved hydrophobic helices [4,37]. Several studies have reported that domain I of 3d-Cry toxin is involved in its membrane insertion and pore formation processes through its conserved hydrophobic α-helices [4]. This therefore suggests that domain II of Vip3A may also take part in these processes through its conserved hydrophobic α-helices.

Both domain III of Vip3A and domain II of 3d-Cry are comprised of three β sheets with a conserved hydrophobic core. Extensive studies of domain II of 3d-Cry toxins showed that it plays a key role in the recognition of midgut receptors [4]. The results of our cell binding assay indicate that Vip3A domain III is also central to cell binding. Furthermore, the binding ability of domains II-III to Sf9 cells is similar to that of full-length Vip3Aa and stronger than domain III alone, and the Vip3Aa11200–end structure shows that domain II and domain III have very compact interaction, which revealed that domain II is also involved in the binding of Vip3A to sensitive cells.

Domain III of 3d-Cry toxins was predicted to bind glycans with a classic glycan binding motif [38–40]. Based on amino acid sequence analysis, previous studies also predicted that all Vip3A proteins contain a carbohydrate-binding motif (CBM\_4\_9 superfamily; pfam02018) in the C-terminus (amino acids 536 to 652 in Vip3Aa) [8]. In the present structure, we found that, instead of the single CBM found in Cry toxins, there were two potential different CBM domains in the C-terminus of the Vip3A toxin, forming domains IV and V, respectively. Our structural analysis indicates that the putative glycan-binding pockets of these two domains differ significantly, suggesting that they are likely to have different glycan binding specificities. This multiplicity of CBMs in Vip3A toxins may increase the diversity of their target polysaccharides. However, in our cell binding assay, domains IV-V did not show binding ability to the Sf9 cells (Figure 3D,E), which may be due to the lack of the specific glycans recognized by domain IV-V on the Sf9 cells' surface. The effect of domains IV and V on the toxicity of Vip3A toxins in insect midgut needs further study.

Taken together, we find here that although the overall structure and domain organization are very different between Vip3 toxin and 3d-Cry toxin, these two families are comprised of functionally and structurally related modules that are assembled in different ways, which may expand the insecticidal spectrum of Bt and make Bt more powerful and efficient to target and kill its hosts.

In addition, the ~65 kDa C-terminal fragment of Vip3A used to be considered as the toxic core [8]. However, recent studies indicated that the ~20 kDa N-terminal fragment and the ~65 kDa C-terminal fragment of Vip3A still bind together after digestion, and the N-terminus is required for the stability and toxicity of Vip3A [20,21,41]. Moreover, several studies further demonstrated that Vip3A remains tetrameric after being activated by trypsin or midgut fluid [20,22]. In our work, the C-terminal fragment of Vip3Aa alone has shown no toxicity through cytotoxicity assays and insect bioassays, and it forms a dimer in the crystal structure, which is consistent with the fact that the C-terminal fragment of Vip3B2160 will form a dimer instead of a tetramer without the N-terminal 21-kDa segment [30]. It is possible that, without N-terminal assistance, the C-terminal fragment cannot correctly assemble into an active tetramer; or, maybe without the protection of N-terminal, the C-terminal fragment loses stability and is degraded by protease.

Vip3A and Vip3B share about 65% sequence similarity and have different insecticidal specificity [42], and recently the C-terminal fragment was found to be related to insecticidal specificity of Vip3 [42,43]. Our structure provides a good opportunity to further study the mechanism of insecticidal specificity between Vip3A and Vip3B. The recently reported structure of Vip3B2160 showed a five-domain organization [30] (Figure S6). When these two Vip3 protein structures are superposed, the C-terminal fragment of Vip3B2160 shows similar folds and organization to Vip3Aa11 (Figure 6A). According to our division, the domain I of Vip3B2160, which is lacked in Vip3Aa11200–end structure, formed a unique fold containing five α-helices wrapping around domain II. Domains III, IV and V of Vip3B2160 have similar folds as their counterparts from Vip3Aa11, respectively (Figure 6C–E). Although domain V in the two structures share similar folds, their positions in their respective structures are obviously different, which suggests that the location of domain V is flexible, and this flexibility of domain V may be related to the insecticidal specificity of Vip3 toxins. However, there are dramatic conformational differences in their domain II (Figure 6B); in the Vip3B2160 structure, the highly conserved hydrophobic α-helix (corresponding to the helix α4 in Vip3Aa11200–end domain II) is surrounded by other helices from domains I and II (Figure 6A, Figure S6). In Vip3Aa11200–end structure, the helices α1 and α2 of domain II have significant conformational changes and expose the hydrophobic region in domain II

(Figure 6B). Hence, we hypothesize that the structural difference in domain II between the full-length and cleaved Vip3 proteins may represent the conformational change after the proteolysis of Vip3A toxins inside the insect midgut. In this scenario, once the cleavage site between domain I and II is processed by insect midgut juice, the α-helices of domain II may undergo a dramatic structural shift that enables helix α1 to rotate and form a hairpin-like structure with helix α4. However, a complex structure of ~20 kDa N-terminal fragment and the ~65 kDa C-terminal fragment of Vip3 after protease digestion will be needed to further prove this hypothesis and to further understand the function of the N-terminal fragment for Vip3 insecticidal activity.

**Figure 6.** Structural comparation between corresponding domains of Vip3Aa11200–end and Vip3B2160. (**A**) Structural comparation of Vip3Aa11200–end and Vip3B2160. The domain I of Vip3B2160 which is lacking in Vip3Aa11200–end structure is colored in light purple. (**B**) Structural overlay of domain II between Vip3A (green) and Vip3B2160 (blue). The cylindrical cartoon shows a detailed view of the conformational changes. (**C**–**E**) Structure superimposition for domain III, domain IV and domain V between Vip3Aa11200–end and Vip3B2160. Each domain is color-coded as the indication.

Collectively, these data provide important structural and functional insights into Vip3A family toxins as well as a valuable resource to guide future studies and to re-evaluate the previous genetic and functional studies that are crucial for the development of Vip3A as a new generation of bio-insecticides.

#### **4. Materials and Methods**

#### *4.1. Bacterial Strains, Cell Lines and Plasmids*

*E. coli* BL21(DE3) for plasmid constructions and protein purification were cultured at 37 ◦C in lysogeny broth (LB) or agar. Methionine auxotrophic *E. coli* strain B834 (DE3) (Novagen, Madison, WI, USA) were used for selenomethionine-substituted (SeMet) Vip3Aa200–end expressing. The S. frugiperda Sf9 cells (Thermo fisher Scientific, Grand Island, NY, USA) were maintained and propagated in Sf-900 II SFM (Gibco, Grand Island, NY, USA) culture medium at 27 ◦C.

The DNA of Vip3Aa200–end was amplified from the Vip3Aa11 gene (GenBank accession No. AY489126.1) using oligonucleotide primer Vip200-F and Vip200-R and cloned into the pET28a vector with an N-terminal 6×His-MBP tag. Plasmids used for RFP (red fluorescent protein) and C-terminal RFP tagged Vip3Aa (Vip3Aa-RFP) expression were constructed as described by Jiang et al. [11]. The different Vip3Aa DNA truncations were amplified from the Vip3Aa11 gene using oligonucleotide primer pairs, DmI-III-F and DmI-III-R, DmIV-V-F and DmIV-V-R, DmI-II-F and DmI-II-R, DmII-III-F and DmII-III-R, and DmIII-F and DmIII-R, and cloned into the pET28a vector with a C-terminal RFP-6×His tag, respectively. All plasmids were generated by the Gibson assembly strategy [44]. The nucleotide sequences of recombinant plasmid were verified by DNA sequencing. All the primers used in this study are shown in Table S2.

#### *4.2. Protein Expression and Purification*

Native His-MBP-Vip3Aa200–end (Vip3Aa200–end) protein was expressed in *E. coli* B21(DE3) at 25 ◦C for 48 h in autoinduction Terrific broth (TB) medium. The cells were harvested by centrifugation at 5000× *g* at 4 ◦C for 15 min and the pellet was resuspended in lysis buffer (20 mM Tris–HCl pH 8 and 150 mM NaCl). After the cells were lysed by high pressure cell crusher (Union-Biotech co., LTD, shanghai, China), the supernatant was collected after centrifuged at 12000× *g* at 4 ◦C for 60 min. The proteins were purified using Ni-NTA agarose resin, washed with 20 mM Tris-HCl, 150 mM NaCl, 20 mM imidazole, pH 8.0, and then eluted with 300 mM imidazole. The Vip3Aa200–end proteins were further purified by HiTrap Q HP ion-exchange chromatography and Superdex 200 gel filtration chromatography (GE Healthcare Life Sciences, Marlborough, MA, USA). Fractions containing the Vip3Aa200–end protein were concentrated to ~7 mg/mL for crystallization. The expression and purification steps of other Vip3Aa truncations were the same as those of Vip3Aa200–end.

SeMet-substituted Vip3Aa200–end was expressed in *E. coli* B834(DE3) strain. Briefly, the cells were cultured in the LB medium at 37 ◦C along with shaking until the OD600 of the bacterial culture reached 1.0. The cells were harvested by centrifugation at 4000× *g* at 4 ◦C for 15 min and the pellet was washed once with PBS. The pellet was resuspended in 1 L Medium A (M9 medium plus 0.4% glucose, 1 mM MgCl2, 1 mM CaCl2, 1 mg Biotin, 1 mg thiamin, 50 mg EDTA, 8.3 mg FeCl3, 0.84 mg ZnCl2, 0.13 mg CuCl2, 0.1 mg CoCl2, 0.1 mg H3BO3 and 0.016 mg MgCl2) and incubated for 3 h at 37 ◦C. We added 50 mg seleno-methionine in the medium and incubated for a further 30 min. The protein was incubated to express for a further 10 h by adding 200 mM IPTG (isopropyl-β-D-thiogalactopyranoside). The SeMet-Vip3Aa200–end was purified by the same procedure as for the native Vip3Aa200–end protein.

#### *4.3. Crystallization, Data Collection and Structural Determination*

The purification of His6-tagged MBP-Vip3Aa200–end used for crystallization is described above. MBP-Vip3Aa200–end (5 mg/mL) was used to perform initial spare matrix crystal screening with a crystallization robot. After crystal optimization trials, MBP-Vip3Aa200–end (7 mg/mL) crystals grew in 3 days at 18 ◦C using the hanging-drop vapor-diffusion method in a mix of 1 μL of protein with 1 μL of reservoir solution consisting of 0.1 M sodium acetate pH 4.2, 0.5 M potassium formate, 0.1 M ammonium sulfate and 11% PEG4000. SeMet MBP-Vip3Aa200–end crystals grew in a similar condition.

A native data set with the space group of P212121 was collected at 3.62 Å (native I). A weak selenomethionine (SeMet) derivative data set was collected at 3.9 Å with the same symmetry as the native I crystal for the amino-acid assignment using the difference Fourier map of the SeMet derivative. After further crystallization optimization, another native crystal (native II) was obtained with the space group of P21 that could diffract to around 3.2 Å. Diffraction data were collected on BL17U1 and BL18U beamlines at Shanghai Synchrotron Radiation Facility (Shanghai, China) and processed by HKL2000 [45].

Molecular replacement was carried out to identity the MBP positions in the native crystals [46] by PHASER [47]. The initial phases were further improved with multi-crystal averaging [48]. Model building was performed manually in COOT [49], and the sequence assignment was helped with the SeMet anomalous difference map. Figures were prepared using PyMol (v.2.3.2, https://pymol.org/, Schrödinger, New York, NY, USA). Structure refinement was done by PHENIX [50]. The data collection and refinement statistics are summarized in Table S1.

#### *4.4. Immunofluorescence*

Sf9 cells with a density of 5 <sup>×</sup> 10<sup>4</sup> cells per ml were seeded into 6-well culture plates separately. After overnight culture, the cells were respectively treated with RFP tagged Vip3Aa or its truncations (0.15 μM) for 6 h. After treatment, the cells were washed three times with PBS to remove unbound proteins, and fixed with 4% paraformaldehyde at 37 ◦C for 15 min. The cell nuclei were labeled with DAPI (0.2 μg/mL) for 30 min. Cell images were captured using a Nikon TI-E inverted fluorescence Microscope (Nikon, NIKON TI-E, Tokyo Metropolis, Japan).

#### *4.5. Cytotoxicity Assays*

Cell viability assays were performed as described by Jiang et al. [25]. Briefly, cells with a density of 5 <sup>×</sup> 104 cells per ml were seeded into 96-well culture plates separately. After overnight incubation, the cells were treated with different Vip3Aa toxins for 72 h. WST-8 reagent was then added to each well. After incubating at 27 ◦C for 2 h, the absorbance was measured in microplate reader at 450 nm. Treatment with protein buffer was used as a control. All tests were performed in triplicate and were repeated at least three times. Cell viability (%) = average absorbance of treated group/average absorbance of control group × 100%.

#### *4.6. Bioassay*

Bioassays were assessed using surface contamination method with *S. exigua* first instar larvae and maintained in a rearing chamber at 27 ◦C, with 50% relative humidity, and 16:8 h light:dark photoperiod. The artificial diet was poured in a 1.8-cm<sup>2</sup> 24 well plate (about 5 mm thick per hole). 200 and 2000 ng/cm<sup>2</sup> concentrations of Vip3Aa proteins (full length Vip3Aa, Vip3Aa199–end Vip3Aa200–end and MBP-Vip3Aa200–end) were spread on the diet. A tris buffer (20 mM Tris-HCl, 300mM NaCl, pH 8.0) was used as a blank control. Three independent replicates and 16 first instar larvae of *S. exigua* were used for each concentration. Mortality was recorded after 5 days.

#### *4.7. Statistical Analysis*

All functional assays were performed at least three times independently. Data were shown as means ± SD. All statistical data were calculated using GraphPad Prism version 8.0 (GraphPad Software, San Diego, CA, USA). One-way ANOVA followed by Dunnett's test were used to identify statistically significant differences between treatments. Significance of mean comparison is annotated as follow: ns, nonsignificant; \*\*, *p* < 0.01; \*\*\*, *p* < 0.001. A *p* value of < 0.05 was considered to be statistically significant.

#### *4.8. Data Availability*

Coordinate for the atomic structure has been deposited in the RCSB Protein Data Bank under RCSB ID: 6VLS. The data that support the findings of this study are available from the corresponding author upon reasonable request.

**Supplementary Materials:** The following are available online at http://www.mdpi.com/2072-6651/12/7/438/s1, Figure S1. Sequence alignment of selected Vip3 family members. Each domain is indicated by the lines bellow sequences, colored as in Figure 1A. Secondary structural elements of Vip3Aa11 are shown above the sequences. The conserved hydrophobic amino acid residues discussed in domain II and domain III are marked with green and magenta triangles, respectively. The potential cleavage site between domain III and domain IV is highlighted with blue triangle. ClustalX2 was used for the sequence alignment. ESPript-3.0 was used to generate the figure. Figure S2. Structure of MBP-Vip3Aa11200–end in the *P*21 space group. Two views of MBP-Vip3Aa11200–end structure in one asymmetric unit. There are four molecules of MBP-Vip3Aa11200–end in one asymmetric unit and they are arranged into two copies of dimer in the different orientations. The molecule A, B, C and D are shown in green, cyan, magenta and yellow, respectively. The MBP (Maltose Bind Protein) tags are shown in silver color in all four molecules. The interaction area between molecule B and C is less than 500 Å2, as calculated by PISA server. Figure S3. Structural alignment between Molecule A and B from the Vip3Aa11200–end dimer. Structure superimposition for the Vip3Aa11200–end and each domain between molecule A and B from the Vip3Aa11200–end dimer structure. Molecule A is colored as Figure 1A, and Molecule B is shown in cyan color. The root means

*Toxins* **2020**, *12*, 438

square deviation (r.m.s.d) of each alignment is listed. Figure S4. Two hydrophobic helices from domain II. The hydrophobic amino acid residues are shown as sticks and labelled with residue numbers. The amino acid residues involved in the hydrophobic (red) and polar (yellow) interactions between α1 and α4 helices are shown as sticks, and the polar interactions are shown in black dashes. Figure S5. Images of Sf9 cells treated with RFP. Fluorescence microscope images of Sf9 cells treated with RFP protein only for 6 h as control. The images are representative of three independent experiments. Nuclei are stained with DAPI (blue), Scale bar, 10 μm. Figure S6. Overall structure of Vip3B2160. Domains I, II, III, IV and V are colored in light purple, blue, light brown, magenta and green, respectively. Table S1. X-ray and refinement statistics. Table S2: Primers used in this study

**Author Contributions:** Conceptualization, K.J. and X.G.; methodology, K.J., Y.Z., Z.C., D.W., J.C. and X.G.; software, Z.C. and X.G.; validation, K.J., Y.Z., Z.C. and X.G.; formal analysis, K.J., D.W., J.C. and X.G.; investigation, K.J., Y.Z., Z.C. and X.G.; resources, J.C. and X.G.; data curation, X.G.; writing—original draft preparation, K.J., D.W. and X.G.; writing—review and editing, K.J. and X.G.; visualization, K.J., Y.Z., Z.C., D.W. and X.G.; supervision, X.G.; project administration, K.J. and X.G.; funding acquisition, K.J. and X.G. All authors have read and agreed to the published version of the manuscript.

**Funding:** This work was supported by the National Natural Science Foundation of China (31901943 and 31770143), the Major Basic Program of Natural Science Foundation of Shandong Province (ZR2019ZD21), China Postdoctoral Science Foundation funded project (2019T120585 and 2019M652370), Youth Interdiscipline Innovative Research Group of Shandong University (2020QNQT009) and Taishan Young Scholars (tsqn20161005).

**Acknowledgments:** We thank Jiawei Wang for providing the suggestion for structure determination, Casey Flower and Jorge Galan for constructive proofreading of this manuscript, the staffs from BL17U1/BL18/BL19U1 beamlines of National Facility for Protein Science Shanghai (NFPS) at Shanghai Synchrotron Radiation Facility (SSRF) for assistance during data collection, and Xiaoju Li from Shandong University Core facilities for life and environmental sciences for her help with the XRD.

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**


© 2020 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).

## **Domain Shu**ffl**ing between Vip3Aa and Vip3Ca: Chimera Stability and Insecticidal Activity against European, American, African, and Asian Pests**

**Joaquín Gomis-Cebolla 1, Rafael Ferreira dos Santos 2, Yueqin Wang 3, Javier Caballero 4, Primitivo Caballero 4, Kanglai He 3, Juan Luis Jurat-Fuentes <sup>2</sup> and Juan Ferré 1,\***


Received: 20 December 2019; Accepted: 29 January 2020; Published: 4 February 2020

**Abstract:** The bacterium *Bacillus thuringiensis* produces insecticidal Vip3 proteins during the vegetative growth phase with activity against several lepidopteran pests. To date, three different Vip3 protein families have been identified based on sequence identity: Vip3A, Vip3B, and Vip3C. In this study, we report the construction of chimeras by exchanging domains between Vip3Aa and Vip3Ca, two proteins with marked specificity differences against lepidopteran pests. We found that some domain combinations made proteins insoluble or prone to degradation by trypsin as most abundant insect gut protease. The soluble and trypsin-stable chimeras, along with the parental proteins Vip3Aa and Vip3Ca, were tested against lepidopteran pests from different continents: *Spodoptera exigua*, *Spodoptera littoralis*, *Spodoptera frugiperda, Helicoverpa armigera*, *Mamestra brassicae*, *Anticarsia gemmatalis*, and *Ostrinia furnacalis*. The exchange of the Nt domain (188 N-terminal amino acids) had little effect on the stability and toxicity (equal or slightly lower) of the resulting chimeric protein against all insects except for *S. frugiperda*, for which the chimera with the Nt domain from Vip3Aa and the rest of the protein from Vip3Ca showed a significant increase in toxicity compared to the parental Vip3Ca. Chimeras with the C-terminal domain from Vip3Aa (from amino acid 510 of Vip3Aa to the Ct) with the central domain of Vip3Ca (amino acids 189–509 based on the Vip3Aa sequence) made proteins that could not be solubilized. Finally, the chimera including the Ct domain of Vip3Ca and the Nt and central domain from Vip3Aa was unstable. Importantly, an insect species tolerant to Vip3Aa but susceptible to Vip3Ca, such as *Ostrinia furnacalis*, was also susceptible to chimeras maintaining the Ct domain from Vip3Ca, in agreement with the hypothesis that the Ct region of the protein is the one conferring specificity to Vip3 proteins.

**Keywords:** Bacillus thuringiensis; Spodoptera spp., Helicoverpa armigera; Mamestra brassicae; Anticarsia gemmatalis; Ostrinia furnacalis

**Key Contribution:** Chimeric proteins between Vip3Aa and Vip3Ca were generated combining fragments of the Nt, the central part, and the Ct of the proteins. The exchange of the Nt domain had little effect on the stability and toxicity (equal or slightly lower), except for *S. frugiperda*, for which a gain of function was observed. Specificity to *O. furnacalis* followed the Ct domain from Vip3Ca.

#### **1. Introduction**

*Bacillus thuringiensis* (Bt) is an aerobic, spore-forming, Gram-positive, and entomopathogenic bacterium belonging to the *Bacillus cereus* group. The Bt bacterium produces a wide variety of insecticidal proteins [1] along with other virulence factors contributing to its pathogenicity [2]. Two major categories of insecticidal proteins produced by Bt are δ-endotoxins (Cry and Cyt toxins) that form crystals within the sporangium in the sporulation phase, and vegetative insecticidal proteins (Vip), which are secreted into the growth medium during vegetative growth [1,3,4]. The Vip proteins are classified into four groups (Vip1, Vip2, Vip3, and Vip4) based on their protein sequence similarity [4,5]. The Vip1 and Vip2 proteins act as binary toxins against coleopteran pests [1,4], while for the Vip4 protein no insecticidal activity has been reported yet.

The Vip3 proteins, mainly those of the Vip3A family, are active against a wide range of lepidopteran pests [1,4]. These proteins do not share structural homology with the Cry proteins, but the toxic action follows the same sequence of events: ingestion, activation by midgut proteases, binding to specific receptors in the midgut epithelium, and pore formation [1,4]. Recent studies indicate that Vip3 proteins (either as protoxins or in the activated form of toxin) spontaneously form tetramers in solution [6–10]. In addition, when the Vip3 proteins are activated by proteases, the oligomer remains stable and the cleaved Nt fragment (19 kDa) remains associated to the main fragment (65 kDa) of the protein [6–10]. In agreement with their diverse structure, Vip3 proteins do not share receptors with Cry proteins [11–15], but share receptors with other Vip3 proteins, either from the same (Vip3Aa, Vip3Af, Vip3Ae, and Vip3Ad) or different (Vip3Ca) protein families [14,16].

Five domains have been proposed for the structure of Vip3A proteins from in silico modelling [17,18]. Based on structural features and stability to trypsin, Quan and Ferré [19] identified five domains from Vip3Af: Domain I encompassing amino acids (aa) 12-198, domain II aa 199-313, domain III aa 314–526, domain IV aa 527–668, and domain V aa 669–788. As far as the structural role of the proposed Vip3Af domains, Quan and Ferré [19] found that domains I–III were required to form the tetrameric structure, the role for domain IV was unclear, and domain V was not necessary for oligomerization. Wang et al. [20] generated a disabled Vip3A protein with two site-engineered mutations (S175C and L177C) in domain I, which was not toxic but retained the ability to compete for the wild type binding sites. Taken together, these results suggest that domain I may be involved in post-binding events, such as membrane insertion, and domain V in binding recognition and specificity.

In this work, we capitalized on the high sequence similarity among Vip3 proteins to test, by domain shuffling, the compatibility of the proposed Vip3Af domains in protein stability and toxicity using representatives from two different Vip3 protein families (Vip3Aa45 and Vip3Ca2). Six chimeric Vip3 proteins (Vip3\_ch1, Vip3\_ch2, Vip3\_ch3, Vip3\_ch4, Vip3\_ch5, and Vip3\_ch6) were designed, where the amino acids (aa) phenylalanine and serine at positions 188 and 509 were chosen as the sites to generate the chimeric Vip3 proteins (Figure 1).

Sequence exchange at these sites coincided approximately with domains I, II+III, and IV+V in the proposed Vip3Af domain model. For the sake of simplicity, we named these domains as the Nt domain (domain I), the central domain (domains II+III), and the Ct domain (domains IV+V), respectively (Figure 1). The objectives of the current research were to determine which main regions of the Vip3 proteins are exchangeable while maintaining the stability and toxicity of the proteins, with the aim to evaluate if any of the new chimeric proteins might confer an increase in the toxicity compared to the parental proteins.

**Figure 1.** Protein sequence alignment of Vip3Aa45 and Vip3Ca2. Black background shading is used to highlight the conserved amino acid between proteins. The proposed structural domains (based on the Vip3Af proteolysis mutants) are indicated with colored lines above the sequences [19]. The purple box indicates the position of the cleavage site (PPS1), while the red vertical lines show the sites chosen to generate the chimeric proteins.

#### **2. Results**

#### *2.1. Sequence Analysis of the Vip3Aa and Vip3Ca Proteins and Determination of the Vip3 Protein Fragment Combinations that Generate Stable Chimeric Proteins*

The amino acid sequence alignment of the two Vip3 proteins indicate that most of the differences are located in their Ct domain (Figure 1). Chimeric proteins were constructed by exchanging the Nt domain (aa 1–188), the central domain (aa 189–509), and the Ct domain (aa 510–788), using as a reference the Vip3A sequence (Figure 1). The Nt domain is highly conserved, with only eight residue differences between the two proteins. The main difference in the protein sequence of the central domain between Vip3Ca and Vip3Aa was two insertions (in Vip3Ca), one located immediately after the main proteolytic processing site (PPS1) (188GIFNE), and the other at aa position 464 (464TF) [21]. To determine the combinations of the different Vip3 protein domains that generated soluble chimeric Vip3 proteins, all the possible combinations were expressed in *Escherichia coli* (Figure 2). The results indicated that the six chimeric proteins could be expressed, but only the Vip3\_ch1, Vip3\_ch2, Vip3\_ch4, and Vip3\_ch5 proteins could be solubilized from the respective inclusion bodies (Figure 2). The exchange of Nt domain did not affect the solubility of the generated chimeric proteins (Vip3\_ch1 and Vip3\_ch2) (Figure 2). However, the exchange of the Ct domain had, in most cases, a negative effect on the solubility of the chimeric protein. The Ct domain from Vip3Aa combined with the central domain from Vip3Ca produced insoluble proteins (Vip3\_ch3 and Vip3\_ch6), whereas the reciprocal combination produced a little soluble protein, with tendency to precipitate (Vip3\_ch5), and a soluble one (Vip3\_ch4) (Figure 2).

**Figure 2.** Summary of the combinations of the different Vip3 protein domains expressed in the heterologous *Escherichia coli* expression system. The "single" chimeric Vip3 proteins (Vip3\_ch1, Vip3\_ch2, Vip3\_ch5, and Vip3\_ch6) were obtained from the Vip3Aa and Vip3Ca as a template, while the "double" chimeric Vip3 proteins (Vip3\_ch3 and Vip3\_ch4) were amplified from the Vip3\_ch5 and Vip3\_ch6. The percentage of similarity of the different proteins vs. the parental proteins, Vip3Aa and Vip3Ca, was calculated with the NCBI Blast align tool.

#### *2.2. Proteolytic and Thermal Stability of The Parental and Chimeric Proteins*

To determine whether the chimeric proteins were stable to the activation by proteases, the proteins were digested with 1% trypsin (w:w). The results showed that Vip3Aa, Vip3Ca, and the chimeric proteins Vip3\_ch1, Vip3\_ch2, and Vip3\_ch4 were processed into the two expected protein fragments of 65-67 kDa and 19-22 kDa (Figure 3). However, the proteolytic pattern of the Vip3\_ch5 chimera differed from the rest of the Vip3 proteins; this phenomenon could be due to (i) instability to proteases, (ii) instability of the Vip3\_ch5 protein in solution, or (iii) problems in the production and purification of the protein. (Figure 3B). Thermal stability of the more soluble and highly purified Vip3 proteins (Vip3Aa, Vip3Ca, Vip3\_ch1, Vip3\_ch2, and Vip3\_ch4) resistant to the trypsin treatment was tested by the thermofluor method (Figure S1). The parental protein Vip3Aa showed two thermal transitions (melting temperature, Tm, of Vip3A-Peak (1): 59.4 ± 0.4 and Tm of Vip3A-Peak (2): 75.5 ± 0.0), while Vip3Ca only showed one thermal transition (Tm of Vip3C-Peak (2): 73.7 ± 0.0) (Figure S1). The chimeric proteins also showed two thermal transitions, but with the first negative peak less pronounced than in the parental Vip3Aa, indicating that the first denaturation involved a lesser part of the protein (Figure S1).

**Figure 3.** Time course of trypsin activation of Vip3 parental and chimeric protoxins. The reaction was carried out using 1% trypsin (w:w) at 37 ◦C for increasing incubation periods. (**A**) Vip3Aa protein and Vip3\_ch1, (**B**) Vip3\_ch5. (**C**) Vip3Ca and Vip3\_ch2; (**D**) Vip3\_ch4. The arrowheads indicate the protein bands corresponding to the 62–67 kDa fragment, while the asterisks indicate the protein bands corresponding to the 19–22 kDa fragment. M1: Molecular Mass Marker.

#### *2.3. Insecticidal Activity of the Parental and Chimeric Vip3 Proteins*

The insecticidal activity of the soluble chimeric proteins (Vip3\_ch1, Vip3\_ch2, Vip3\_ch4, and Vip3\_ch5) was compared to that of the parental proteins by testing eight insect species with different susceptibilities to Vip3Aa and Vip3Ca (Table 1). The Vip3Aa protein was toxic for all the insect species tested except for *Ostrinia furnacalis* (for this insect species the Vip3Aa protein is only toxic at very high concentration). The Vip3Ca protein showed high toxicity to *O. furnacalis* and moderate toxicity to *A. gemmatalis*; for the other insect species tested, this protein was slightly active at very high concentrations (Table 1).

