*Review* **Experimental Infection of Ticks: An Essential Tool for the Analysis of** *Babesia* **Species Biology and Transmission**

**Sarah I. Bonnet 1,2,\* and Clémence Nadal 3,4**


**Abstract:** Babesiosis is one of the most important tick-borne diseases in veterinary health, impacting mainly cattle, equidae, and canidae, and limiting the development of livestock industries worldwide. In humans, babesiosis is considered to be an emerging disease mostly due to *Babesia divergens* in Europe and *Babesia microti* in America. Despite this importance, our knowledge of *Babesia* sp. transmission by ticks is incomplete. The complexity of vectorial systems involving the vector, vertebrate host, and pathogen, as well as the complex feeding biology of ticks, may be part of the reason for the existing gaps in our knowledge. Indeed, this complexity renders the implementation of experimental systems that are as close as possible to natural conditions and allowing the study of tick-host-parasite interactions, quite difficult. However, it is unlikely that the development of more effective and sustainable control measures against babesiosis will emerge unless significant progress can be made in understanding this tripartite relationship. The various methods used to date to achieve tick transmission of *Babesia* spp. of medical and veterinary importance under experimental conditions are reviewed and discussed here.

**Keywords:** ticks; *Babesia* sp.; biological cycle; experimental transmission; experimental models

## **1. Introduction**

Babesiosis remains prevalent worldwide and represents an important threat for both humans and animals [1,2]. The disease, impacting mainly cattle, sheep, goat, equidae, canidaecanidae, and accidentally humans, is caused by apicomplexan parasites belonging to the *Babesia* genus that exclusively infect erythrocytes of their vertebrate hosts [3]. To date, more than 100 *Babesia* species have been identified [1]. *Babesia* spp. are transmitted by hard ticks—occasionally by blood transfusion—and require both a competent vertebrate and invertebrate host to maintain the transmission cycle [3].

Human babesiosis is caused by *Babesia microti,* a *Babesia crassa*-like pathogen, *Babesia divergens, Babesia duncani,* and *Babesia venatorum*, as well as other parasites closely genetically related to these pathogens, such as *B. divergens*-like, *B. duncani*-like, and *B. microti*-like [4]. Infections in otherwise healthy individuals is usually mild to moderate and most cases of severe disease occur in immunocompromised individuals. *B. microti* is endemic in the northeastern and upper midwestern regions of the United States, while *B. duncani* is present on the west coast of the country [4]. In Europe, most of the human cases are due to *B. divergens,* whereas in Asia they are due to *B. venatorum, B. crassa*-like, and *B. microti* [4]. In cattle, *Babesia* spp. have a significant worldwide economic, social, and epidemiological impact and include, among the most important species, *B. bovis, B. bigemina, Babesia major,* and *B. divergens* [5]. *B. bovis* and *B. bigemina* are present in many countries in Africa, Asia,

**Citation:** Bonnet, S.I.; Nadal, C. Experimental Infection of Ticks: An Essential Tool for the Analysis of *Babesia* Species Biology and Transmission. *Pathogens* **2021**, *10*, 1403. https://doi.org/10.3390/ pathogens10111403

Academic Editors: Estrella Montero, Jeremy Gray, Cheryl Ann Lobo and Luis Miguel González

Received: 9 August 2021 Accepted: 26 October 2021 Published: 29 October 2021

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**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/).

Australia, Central and South America, and Southern Europe between 40◦ N and 32◦ S. *B. major* is present in Europe, Northwest Africa and Asia, and *B. divergens* is present in northern Europe [5]. Ovine babesiosis due to *Babesia ovis* and *Babesia motasi* is considered as the most critical blood-borne parasitic disease of small ruminants in tropical and nontropical regions (occurring in South-eastern Europe, North Africa, and Asia) [6]. In equids, *Babesia caballi* is (with *Theileria equi* and *Theileria haneyi*) the agent of equine piroplasmosis known to be endemic in several countries of Africa, Asia, the Americas, and mainly in the Mediterranean basin for Europe [7–9]. The disease represents a significant animal health issue and causes notable economic losses for the equine industry. Finally, babesiosis is one of the most important globally extended and quickly spreading tick-borne diseases in dogs worldwide. *Babesia canis* is the main cause of canine babesiosis in Europe and is only sporadically found around the world, whereas *Babesia gibsoni,* the most prevalent species, and *Babesia vogeli* have a global distribution. *Babesia rossi*, one of the most pathogenic species, is endemic in southern Africa [10].

The current approaches available for babesiosis control have many important limitations, including increased resistance to acaricides by ticks, as well as the numerous drawbacks of these acaricides and of the current vaccines and babesicidal drugs (e.g., efficacy, toxicity, environmental effects) [11]. The development of improved control measures against babesiosis is limited by the numerous and significant gaps in our understanding of the biology of *Babesia* spp., especially regarding molecular interaction between parasites, vectors, and vertebrate hosts, as well as the factors that may influence both the development and the transmission of the parasite [12]. To fill these gaps, it is essential to be able to reproduce the life cycle of *Babesia* species in controlled experimental conditions, including transmission by ticks [13,14]. In addition, the validation of new methods of interruption of the cycle requires that it can be first carried out entirely under such conditions. Finally, it is important to mention that despite the promise of in vitro culture systems [15], maintenance by in vitro culture [16] or needle-passage in the vertebrate host [17] in the absence of tick passage may generate significant changes in the parasite population, potentially creating a bias in research results.

Several laboratory studies that aimed to understand babesiosis pathogenesis have focused their interest on infecting laboratory animals through artificial parasite injection [18,19]. Regarding tick-parasite interaction, the great majority of studies carried out concern mainly epidemiological studies focusing on the detection of parasites in ticks collected into the field [20]. Some in vitro studies have also been performed in order to understand the interactions between parasites and the cells of their vertebrate [21–23] or invertebrate hosts [24–27]. However, and certainly because of the difficulties inherent to the studied model—including *Babesia* spp. culture, tick colony maintenance, and animal models—relatively few studies have been based on the establishment, under experimental conditions, of complete parasite transmission cycles from one vertebrate host to another via the tick bite. Thanks to the experimental models developed for that purpose, these few studies have nevertheless made significant advances in (1) the definition/confirmation of the vector competence of various tick species; (2) the understanding of the modalities of parasite acquisition and transmission by ticks; (3) the discovery of the molecular interactions between the parasite and its invertebrate hosts; (4) the evaluation of some control methods. The aim of this review is to summarize studies that include both tick infection on *Babesia*-infected animals and *Babesia* infection of ticks through artificial systems, and to comment on the major results they achieved.

#### **2. General Description of the** *Babesia* **Life Cycle**

*Babesia* spp. are transmitted by hard tick (ixodid) vectors. The tick vectors and reservoir hosts differ depending on *Babesia* sp. and geographical location considered [1]. Several tick species have been mentioned in the literature as vectors of *Babesia* sp., but, as shown in Table 1, vector competence through experimental transmission has not been validated for all of them. For those mentioned here as "suspected vectors" without realization of the complete transmission cycle under experimental conditions, their involvement is mostly based on epidemiological evidence (e.g., correlation of tick species presence with disease occurrence). Each of the three active stages of hard ticks (larva, nymph, and adult) takes a single blood meal from a vertebrate host in order to mature to the next stage or lay eggs for the female. Most of the tick species listed as confirmed or suspected vectors of some *Babesia* species are three-hosts ticks—meaning that they take their three blood meals on three different hosts. Some of them, however, have a two-host cycle such as some *Hyalomma* spp., *Rhipicephalus evertsi,* and *Rhipicephalus bursa,* whereas *Dermacentor nitens, Rhipicephalus* annulatus, *Rhipicephalus microplus,* and *Rhipicephalus decoloratus* are one-host ticks that use the same individual host animal for all active tick stages.

**Table 1.** Major *Babesia* species infecting humans, dogs, cattle, sheep, goat, and equids; their suspected or confirmed main vector and vertebrate hosts in the field; and the realization of the whole transmission cycle under experimental conditions.


ND: no identified data.

The *Babesia* life cycle includes both asexual multiplication in the erythrocytes of the vertebrate host and sexual reproduction in the tick vector [3,61]. The general life cycle of the *Babesia* species is summarized in Figure 1 for *Babesia* sensu stricto (s.s.) species.

Indeed, it is necessary to specify here that, quite recently, molecular phylogeny studies using the 18S rRNA gene have led to the division of *Babesia* species into two large groups: *Babesia* s.s. and *Babesia* sensu lato (s.l.). Species belonging to the latter group, such as *B. microti*, are not capable of transovarial transmission within the tick but only a transstadial mode of transmission [62,63]. Vertebrate hosts are infected by the injection of sporozoites present in tick saliva during the tick bite. Each sporozoite penetrates the cell membrane of an erythrocyte with the aid of a specialized apical complex. Once inside, the parasite produces two merozoites by a process of merogony. Merozoites are then intermittently released following erythrocyte lysis to infect new erythrocytes. The parasite may then persist asymptomatically within its host for several years or lead to acute disease. When they are ingested by the tick during the blood meal, some parasites present in infected erythrocytes (pre-gametocytes) undergo further development in the passage from host blood to the midgut of the tick vector to evolve into gametocytes. The sexual reproduction between gametocytes takes place in the tick gut and leads to a zygote that penetrates the gut epithelium, where further multiplication occurs, with development to motile and haploid kinetes that escape into the tick hemolymph. The kinetes then infect a variety of tick cell types and tissues, including the ovary in the female tick—for *Babesia* s.s. species—and the tick salivary glands, where successive cycles of asexual multiplication take place. In this last organ, sporozoite development usually only begins when the infected tick attaches to the vertebrate host. The ticks thus transmit the sporozoites to a new host during a new blood meal of the next life-stage for the ticks with several hosts or of the next generation after transovarian transmission for the one-host ticks. it is necessary to specify here that, quite recently, molecular phylogeny studies using the 18S rRNA gene have led to the division of *Babesia* species into two large groups: *Babesia* s.s. and *Babesia* sensu lato (s.l.)*.* Species belonging to the latter group, such as *B. microti*, are not capable of transovarial transmission within the tick but only a transstadial mode of transmission [62,63]. Vertebrate hosts are infected by the injection of sporozoites present in tick saliva during the tick bite. Each sporozoite penetrates the cell membrane of an erythrocyte with the aid of a specialized apical complex. Once inside, the parasite produces two merozoites by a process of merogony. Merozoites are then intermittently released following erythrocyte lysis to infect new erythrocytes. The parasite may then persist asymptomatically within its host for several years or lead to acute disease. When they are ingested by the tick during the blood meal, some parasites present in infected erythrocytes (pre-gametocytes) undergo further development in the passage from host blood to the midgut of the tick vector to evolve into gametocytes. The sexual reproduction between gametocytes takes place in the tick gut and leads to a zygote that penetrates the gut epithelium, where further multiplication occurs, with development to motile and haploid kinetes that escape into the tick hemolymph. The kinetes then infect a variety of tick cell types and tissues, including the ovary in the female tick—for *Babesia* s.s. species—and the tick salivary glands, where successive cycles of asexual multiplication take place. In this last organ, sporozoite development usually only begins when the infected tick attaches to the vertebrate host. The ticks thus transmit the sporozoites to a new host during a new blood meal of the next life-stage for the ticks with several hosts or of the next generation after transovarian transmission for the one-host ticks.

The *Babesia* life cycle includes both asexual multiplication in the erythrocytes of the vertebrate host and sexual reproduction in the tick vector [3,61]. The general life cycle of the *Babesia* species is summarized in Figure 1 for *Babesia* sensu stricto (s.s.) species. Indeed,

*Pathogens* **2021**, *10*, x FOR PEER REVIEW 4 of 14

**Figure 1***.* Life cycle of *Babesia* spp. sensu stricto. Vertebrate hosts are infected following the bite of an infected hard tick, through the invasion of the host erythrocytes by sporozoites excreted in the tick saliva. Inside the erythrocyte, sporozoites develop into trophozoites that undergo an asexual multiplication called merogony, ending in the formation of either merozoites that can infect other erythrocytes, or of gametocytes, which eventually develop into gametes. Infected erythrocytes **Figure 1.** Life cycle of *Babesia* spp. sensu stricto. Vertebrate hosts are infected following the bite of an infected hard tick, through the invasion of the host erythrocytes by sporozoites excreted in the tick saliva. Inside the erythrocyte, sporozoites develop into trophozoites that undergo an asexual multiplication called merogony, ending in the formation of either merozoites that can infect other erythrocytes, or of gametocytes, which eventually develop into gametes. Infected erythrocytes are taken up by the tick during its blood meal, but only the gametocytes survive, and they then undergo further development, changing into gametes in the tick midgut. Then, sexual multiplication—gamogony—takes place with the fusion

of two gametes to form a motile zygote that enters the midgut epithelial cells to develop into motile kinetes through meiotic division. Kinetes disseminate to tick tissues, including ovarian and salivary gland cells. The invasion of tick ovaries results in transovarial transmission while those of salivary glands leads to transmission to the vertebrate host through injection of sporozoites with the saliva. *Babesia* microti-like species, which belong to *Babesia* spp. s.l. species, only invade the salivary glands, not the ovary. The schematic representation was made using the software biorender.com.

#### **3. Experimental Models of** *Babesia* **Species—Transmission by Ticks**

#### *3.1. Tick Infestations on Babesia spp. Infected Animals*

The first experiments to transmit *Babesia* spp. naturally were carried out by applying ticks suspected of being vectors on infected animals. Due to the huge economic importance of bovine babesiosis, these studies on vector competence for *Babesia* sp. were first conducted on species that infect cattle. Indeed, in 1893, two American researchers, Smith and Kilborne—the first authors to demonstrate the transmission of a disease organism from an arthropod to a mammalian host—showed the vector competence of *R.* annulatus for *B. bigemina* by placing ticks infected on animal onto naïve cattle that developed the associated disease [42]. Thereafter, different species of animals were used depending on the species of *Babesia* studied. Over time, the animal models and methodology used were refined to optimize the infection of both animals and ticks, and to comply with health and safety rules, and animal accommodation and tick containment methods during the tick feeding process have been the subject of several tests and evolutions (see examples in Figure 2A–F). In most cases, for cattle, animals were housed in individual, tick-proof pens surrounded by moats with or without detergent or insecticide [43,45]. Concerning the tick containment methods, in 1961, Callow and Hoyte used a hessian rug to protect the larvae until adult repletion to demonstrate the transmission of *B. bigemina* to cattle by *R. microplus* [43]. Ticks were either allowed to spread at will over the animal, or were confined to one site by releasing them under a fabric patch (nylon or organdie), which was glued along its edges to the flank of the bovine. A few years later, for the demonstration of the vector competence of *R. decoloratus* for *B. bigemina* in Kenya, ticks were fed on cattle until adult repletion by sprinkling larvae on the backs of the animals [45]. For the first experimental transmission of *B. divergens* by *Ixodes ricinus* achieved by Joyner et al in 1963, the ticks were contained in ear bags [36], a method also used later for sheep [64]. Regarding rodents, several laboratory studies involving complete transmission cycles of *B. microti* to the vertebrate host through the tick bite were performed [28–33]. In most instances, rodents were maintained over trays of water from which detached engorged ticks—applied by brush to animals—were harvested, whereas in some cases, ticks were contained in plastic capsules attached with different adhesives. For horses, the first studies aimed to validate the vector competence of *D. nitens* for *B. caballi* used larvae applied by brush on the animal [51], whereas in subsequent studies, tick feedings were accomplished by placing the larvae under a cloth patch glued to the back of the host [52].

The identification of a pathogen or pathogen DNA alone—which is even less convinc ing—in an arthropod cannot be sufficient to prove its ability to transmit this pathogen. Indeed, demonstrations of parasite presence in unfed field ticks, in tick salivary glands, eggs, or unfed larvae, while more convincing than detection in ticks collected from animals, also require confirmation only provided by the validation of vector competence in a controlled experimental model. The best illustration of this corresponds to the following studies performed in Iran in order to identify the vector of *B. ovis* to sheep. The kinetes of *B. ovis* were observed in hemolymph and egg smears of *Rhipicephalus sanguineus* and *Hyalomma marginatum* field ticks collected from sheep infected with *B. ovis* [69], so the vector competence of both tick species was further evaluated by placing pairs of adult ticks on sheep inoculated with *B. ovis,* but no transmission by any of the succeeding tick stages could be demonstrated, thus showing that these tick species are not vectors [64].

(**E**) (**F**)

**Figure 2.** Experimental feeding of *Ixodes ricinus* ticks on (**A**) Siberian chipmunks (*Tamias sibiricus barberi)* [65], (**B**) mouse [66], (**C**,**D**) sheep [67], (**E**,**F**) rabbit [68], and (**F**,**H**) a membrane artificial feeding system [35]. **Figure 2.** Experimental feeding of *Ixodes ricinus* ticks on (**A**) Siberian chipmunks (*Tamias sibiricus barberi)* [65], (**B**) mouse [66], (**C**,**D**) sheep [67], (**E**,**F**) rabbit [68], and (**G**,**H**) a membrane artificial feeding system [35].

The identification of a pathogen or pathogen DNA alone—which is even less convincing—in an arthropod cannot be sufficient to prove its ability to transmit this pathogen. Indeed, demonstrations of parasite presence in unfed field ticks, in tick salivary glands, eggs, or unfed larvae, while more convincing than detection in ticks collected from animals, also require confirmation only provided by the validation of vector competence in a controlled experimental model. The best illustration of this corresponds to the following studies performed in Iran in order to identify the vector of *B. ovis* to sheep. The kinetes of *B. ovis* were observed in hemolymph and egg smears of *Rhipicephalus sanguineus* and *Hyalomma marginatum* field ticks collected from sheep infected with *B. ovis* [69], so the vector competence of both tick species was further evaluated by placing pairs of adult ticks on sheep inoculated with *B. ovis,* but no transmission by any of the succeeding tick stages could be demonstrated, thus showing that these tick species are not vectors [64]. As recently reviewed by Gray and co-workers, the establishment of experimental models of pathogens transmission by ticks using live animals has led to significant advances in the understanding of transmission modalities and tick-parasite interactions [70]. For example, it allowed the demonstration that transovarial transmission does not occur in *B. microti,* leading to no longer classifying this parasite in *Babesia* spp. s.s. [31–33], and that, in *I. ricinus,* the infection only survives one molt [31]*.* Laboratory models have also demonstrated that *B. microti* may promote its transmission in rodents by enhancing the feeding success and survival of its tick vector, *Ixodes trianguliceps* [29]. Likewise, the estab-As recently reviewed by Gray and co-workers, the establishment of experimental models of pathogens transmission by ticks using live animals has led to significant advances in the understanding of transmission modalities and tick-parasite interactions [70]. For example, it allowed the demonstration that transovarial transmission does not occur in *B. microti,* leading to no longer classifying this parasite in *Babesia* spp. s.s. [31–33], and that, in *I. ricinus,* the infection only survives one molt [31]. Laboratory models have also demonstrated that *B. microti* may promote its transmission in rodents by enhancing the feeding success and survival of its tick vector, *Ixodes trianguliceps* [29]. Likewise, the establishment of the transmission of *B. divergens* by *I. ricinus* in a gerbil experimental model [35] provided proof of sexual development of *Babesia* through DNA measurements on the developmental stages of *B. divergens* in the blood of the vertebrate host and in the gut, hemolymph, and salivary glands of the tick vector [38,39]. In addition, several studies have been performed in order to establish which tick life stages are able to acquire and/or to transmit the parasite. Most of them have concluded that only adult stages were able to acquire *Babesia* sp. s.s. from infected animals, while all succeeding stages (larvae, nymphs, and adults) were able to re-transmit the parasites to susceptible animals [36,37,43,44,48]. However, Schwint and co-workers demonstrated that only the first of three subsequent generations from *D. nitens* females was able to transmit *B. caballi* to naïve horses, showing that the parasite is unable to persist in ticks without continuing alimentary infection of adult females [52], whereas other studies have also shown the acquisition of this parasite by nymphs of *R. evertsi* [53].

lishment of the transmission of *B. divergens* by *I. ricinus* in a gerbil experimental model [35] provided proof of sexual development of *Babesia* through DNA measurements on the developmental stages of *B. divergens* in the blood of the vertebrate host and in the gut, hemolymph, and salivary glands of the tick vector [38,39]. In addition, several studies have been performed in order to establish which tick life stages are able to acquire and/or to transmit the parasite. Most of them have concluded that only adult stages were able to acquire *Babesia* sp. s.s. from infected animals, while all succeeding stages (larvae, nymphs, and adults) were able to re-transmit the parasites to susceptible animals [36,37,43,44,48]. However, Schwint and co-workers demonstrated that only the first of three subsequent generations from *D. nitens* females was able to transmit *B. caballi* to naïve horses, showing that the parasite is unable to persist in ticks without continuing alimentary infection of adult females [52], whereas other studies have also shown the acquisition of this parasite by Models of transmission of *B. microti* to rodents via ticks in the laboratory have also made it possible to carry out studies on the phenomena of co-infections. In fact, a lower transmission efficiency of *B. microti* than *Borrelia burgdorferi* to *Ixodes dammini* from both hamsters [71] and white-footed mice [72] has been demonstrated. In the meantime, it was shown that ticks that fed on mice with these concurrent pathogen infections exhibited twice the incidence of *B. burgdorferi* infection compared with *B. microti* [72]. Twentyfive years later, however, this laboratory model also showed an increase of the frequency of *B. microti*-infected *Ixodes scapularis* (formerly *I. dammini*) nymphs when they fed as larvae on white-footed mice coinfected with *B. burgdorferi,* as well as an increase of *B. microti* parasitemia in co-infected mice [28]. This enhancement of *B. microti* establishment by *B. burgdorferi* has been attributed to an immunological conflict in the adaptive immune response of the vertebrate host against the two tick-borne pathogens [73].

nymphs of *R. evertsi* [53]. Models of transmission of *B. microti* to rodents via ticks in the laboratory have also made it possible to carry out studies on the phenomena of co-infections. In fact, a lower transmission efficiency of *B. microti* than *Borrelia burgdorferi* to *Ixodes dammini* from both The development of experimental models using animals also provided knowledge on the infection acquisition by the vertebrate host following a *Babesia*-infected tick bite. For example, by developing a laboratory model of *B. bovis* infection of calves through the bite of *R. microplus*-infected ticks, Smith and co-workers demonstrated, in 1978, that tick-induced

hamsters [71] and white-footed mice [72] has been demonstrated. In the meantime, it was

infection was more severe than in calves infected with carrier blood, even when very low numbers of infected larvae were applied [41]. They attributed this difference in virulence to the large number of infective doses injected by each infected tick but Salivary-Assisted Transmission of tick-borne pathogens (see review in [74]) probably also contributed to this observation.

Although, compared to other pathogens, few molecular studies have involved *Babesia* parasites [70,75], experimental tick-transmission models of *Babesia* spp. using animals have also made it possible to identify molecules potentially involved in this transmission. Such studies are helping us to better understand the interactions involved and to identify potential targets for blocking parasite transmission. Thus, infection of *H. longicornis* on dogs infected with *B. gibsoni* has allowed to implicate a tick protein, longipain, in the transmission of the parasite by its vector [76], and to demonstrate that the vitellogenin receptor on the surface of tick oocytes is essential for its transovarial transmission [77]. Experimental models of *B. bovis* infection of cattle by *R. microplus* has also been used to study the function of the tick protein Bm86 during *B. bovis* infection [78]. In the same way, experimental tick infection models of both *B. bigemina* and *B. bovis*-infected cattle were used to perform functional genomics studies on *R. annulatus* and *R. microplus* genes that are differentially expressed in response to parasite infection [79,80].

Both the discovery and validation of methods to control the transmission of tickborne pathogens require experimental designs that include complete transmission cycles. Regarding the *Babesia* spp. that infect cattle, experimental tick infection models for *B. bigemina*-infected cattle were used to evaluate the efficiency of some vaccine candidates and drugs against parasite transmission by *R. microplus* [81,82]. *Rhipicephalus microplus* experimentally infected with *B. bovis* were also used to demonstrate the inefficiency of the injectable and pour-on forms of both ivermectin and moxidectin to prevent parasite transmission by ticks [82]. In Argentina, Mangold and co-workers developed a laboratory model of *B. bovis* transmission to cattle by *R. microplus* in order to demonstrate the nontransmissibility to ticks of an attenuated vaccine strain of the parasite [83]. Several studies also involved experimental transmission of *B. canis* to dogs through the bite of infected *D. reticulatus* ticks in order to evaluate the usefulness of different acaricides to prevent parasite transmission [55–57,84], whereas similar experiments were performed regarding *B. canis* transmission by *R. sanguineus* [85,86], and *B. rossi* transmission by *Haemaphysalis elliptica* [60].

#### *3.2. Tick Infection through Artificial Feeding Systems*

The use of natural hosts for direct infection of ticks on infectious animals remains the best method to obtain conditions that are closest to the physiological reality of tick-borne pathogen transmission. However, firstly, ethical considerations lead us to limit the use of animals as much as possible. Secondly, in addition to the constraints associated with licensing the experiments and host specificity, obtaining the animals, keeping them in the laboratory, and handling them can be expensive and difficult or even impossible in the case of most species of wildlife. Finally, the difficulty of controlling parasitaemias of infected animals for the whole feeding period of the ticks is an important consideration. All these reasons have led to the development of artificial methods of tick infection in order to complete the lifecycle of tick-borne pathogens under laboratory conditions. Artificial feeding of ticks, mimicking the natural process, has been used for different purposes including tick rearing, the study of tick physiology and the effects of antibodies or drugs on tick physiology, functional genomic studies, and vaccine candidate discovery, as well as the analysis of tick-borne pathogen transmission (see review by Bonnet and Liu [13]). Nevertheless, relatively few studies have involved *Babesia* parasites. One of the factors limiting these studies is undoubtedly the need for successful in vitro cultivation of the *Babesia* species of interest [15].

The use of blood-filled capillary tubes placed over the mouthparts of ticks was first reported in 1938 by Gregson, who used this technique to collect saliva from *D. andersoni* [87].

Since then, this technique has been used to infect ticks with several tick-borne pathogens, mainly bacteria (see review in [13]). Although this infection process has the advantage of using the natural route of infection, i.e., the digestive tract of the tick, and allows control of the quantity of pathogen ingested, it is quite far from natural conditions because the tick absorbs a large amount of pathogen at once and this, regardless of the "true" full blood meal. The other drawback is that it requires the use of animals before or after infection to feed ticks to repletion. However, its use has led to some significant advances in the study of *Babesia* spp. transmission. In 1998, Inokuma and Kemp used the capillary feeding technique in order to infect *R. microplus* with *B. bigemina* [88]. Adult ticks were pre-fed on cattle and then *B. bigemina*-infected red blood cells were offered to the ticks using glass capillaries on a warm plate at 35 ◦C for 18 h in the dark. The authors subsequently demonstrated that ticks were able to acquire the parasite and to transmit it to their progeny. The capillary feeding technique has also been used by Antunes and co-workers in order to evaluate the impact of purified rabbit polyclonal antibodies on some *R. microplus* proteins associated with both tick physiology and tick infection by *B. bigemina* [89]. While an effect on tick weight and oviposition was observed, no effect was observed on pathogen DNA levels. However, this work has made it possible to set up an alternative system to the use of animals both to test vaccine candidates and to obtain essential data on tick-pathogen interaction, as this could also be done later with another protein, the calreticulin, identified as being involved in *B. bigemina* infection in *R. annulatus* ticks [90].

The membrane feeding technique—consisting of feeding ticks on blood or culture media through a membrane—was first developed by Pierce and Pierce in 1956 in order to feed *R. microplus* using embryonated hen eggs [91]. Since then, several membranes of animal or artificial origin have been used both to feed and to infect ticks with different pathogens (see review in [13]). In this case, as the pathogen is mixed in blood and absorbed throughout the blood meal via the digestive tract, the method mimics the natural conditions of tick infection more closely than other methods. Its main disadvantages, however, are the need to regularly change the blood used, and the need to use antibiotics and antifungals to avoid contamination. It is also necessary to test pathogen viability under these feeding conditions at regular intervals. In the case of artificial membranes such as silicone ones, feeding systems also need olfactory stimuli for attachment and feeding. Regarding their use to infect ticks with *Babesia* spp., and due to red blood cell sedimentation, it is important to note that this requires the placing of blood above the membrane to produce a continuous gravitational pressure, ensuring tick absorption of the intraerythrocytic parasites. In 2007, Bonnet and co-workers developed a skin-feeding technique using the skins of both gerbils (for larvae and nymphs) and rabbits (for nymphs and adults) to infect *I. ricinus* with *B. divergens* without the need for additional stimuli (Figure 2G,H) [35]. To our knowledge, this is the only membrane-feeding technique that has been used to date to infect ticks with *Babesia* spp. All tick instars were allowed to acquire the parasite from infected red blood cells maintained at 37 ◦C in a glass feeder through the animal skin until repletion. Contrary to what was previously observed [36,37], this system showed that in addition to adult females, *I. ricinus* larvae and nymphs can also acquire *B. divergens* infections, which persists transtadially in the subsequent nymphal and adult stages (as determined by the detection of DNA in their salivary glands). *Babesia divergens* DNA was also detected in eggs and larvae produced by females that had fed on parasitized blood, demonstrating the transovarial transmission of the parasite. Later, the use of this artificial tick infection system also allowed the discovery of molecular markers for *B. divergens* sexual stages [92,93]. Lastly, the same membrane feeding technique allowed the validation of the vector competence of *I. ricinus* for *Babesia* (EU1) *venatorum* [40].

Finally, in order to understand tick-*Babesia* interactions and to follow parasite development in the vector, Maeda and co-workers used a "semi-artificial" mouse skin membrane feeding technique to infect *H. longicornis* with *B. ovata* [94]. In this case, female adult ticks were first allowed to feed on the shaved back of mice, and after 4 to 5 days, a section of the mouse skin with the ticks attached was removed immediately after euthanasia,

and set up in artificial feeding units. The ticks were then fed on a mix of media and *B. ovata*-infected red blood cells through the piece of mouse skin. This technique was then used to demonstrate the transovarial persistence of *B. ovata* DNA in *H. longicornis* [50]. Thus, the mixing of animal use and membrane feeding makes it possible to control the parasitemia of the meal offered to ticks, but does not prevent the use of live animals.

## *3.3. Tick Infection through Injection*

Although this method is more distant from physiological reality than the ones previously detailed, some studies have also performed tick infections by injecting the pathogen through the cuticle of the tick. In addition to requiring live animals to feed ticks after the infection, this invasive method also has the disadvantage of a low survival rate of ticks after injection [95]. Nevertheless, its use by some authors has led to important results concerning *Babesia* spp. In 2018, Antunes and co-workers used a *R. bursa-B. ovis*-sheep infection model to characterize tick salivary gland genes that were differentially expressed in response to blood feeding and *B. ovis* infection [96]. In that experiment, female ticks were inoculated with *B. ovis* in the trochanter-coxae articulation and allowed to feed on rabbits. Vector competence was then confirmed by feeding *B. ovis*-infected ticks on a naïve lamb. This study allowed both increased understanding of the role of tick salivary gland genes in *Babesia* infections and identification of potential candidate vaccine antigens for innovative control strategies.

#### **4. Conclusions**

It is likely that vector competence for *Babesia* spp. has yet to be determined in some tick species. Furthermore, our understanding of the life cycles of *Babesia* spp. is still incomplete, especially regarding the intimate mechanisms of molecular dialogue between the parasite and its vertebrate and invertebrate hosts. Field experiments are less easy to control than those under laboratory conditions, and the gaps in our knowledge can probably only be filled by experimentally reproducing the transmission cycle as closely as possible to reality and by involving all three actors of the vectorial system. Indeed, this review has shown that the use of experimental systems for tick-borne pathogen infections that permit the complete transmission cycles of *Babesia* parasites has led to major scientific advances in the study of these pathogens. Despite these successes, efforts are still needed to standardize and simplify laboratory protocols to improve our ability to exploit tick artificial infection systems. It is hoped that, in the future, such models of artificial infection will be further developed in order to acquire new knowledge and develop new control strategies while avoiding the use of animals.

**Author Contributions:** Conceptualization, S.I.B.; formal analysis, S.I.B. and C.N.; writing—original draft preparation, S.I.B. and C.N.; writing—review and editing, S.I.B. and C.N.; supervision, S.I.B. All authors have read and agreed to the published version of the manuscript.

**Funding:** The PhD of CN is funded by the Institut Français du Cheval et de l'Equitation (IFCE) and by the Fonds Eperon.

**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**


## *Review* **Preventing Transfusion-Transmitted Babesiosis**

**Evan M. Bloch 1,\* , Peter J. Krause <sup>2</sup> and Laura Tonnetti <sup>3</sup>**


**Abstract:** *Babesia* are tick-borne intra-erythrocytic parasites and the causative agents of babesiosis. *Babesia*, which are readily transfusion transmissible, gained recognition as a major risk to the blood supply, particularly in the United States (US), where *Babesia microti* is endemic. Many of those infected with *Babesia* remain asymptomatic and parasitemia may persist for months or even years following infection, such that seemingly healthy blood donors are unaware of their infection. By contrast, transfusion recipients are at high risk of severe babesiosis, accounting for the high morbidity and mortality (~19%) observed in transfusion-transmitted babesiosis (TTB). An increase in cases of tick-borne babesiosis and TTB prompted over a decade-long investment in blood donor surveillance, research, and assay development to quantify and contend with TTB. This culminated in the adoption of regional blood donor testing in the US. We describe the evolution of the response to TTB in the US and offer some insight into the risk of TTB in other countries. Not only has this response advanced blood safety, it has accelerated the development of novel serological and molecular assays that may be applied broadly, affording insight into the global epidemiology and immunopathogenesis of human babesiosis.

**Keywords:** *Babesia*; blood transfusion; prevention; screening; babesiosis

## **1. Introduction**

*Babesia* are tick-borne apicomplexan parasites and the causative pathogens of the clinical illness, babesiosis. Over 100 species of *Babesia* infect a wide array of vertebrates, yet only six species have been implicated in human infections, of which *Babesia microti* is overwhelmingly predominant [1]. While *B. microti* has been reported frequently from the northeastern and northern midwestern United States (US), cases of babesiosis have been described globally [2]. Findings from *Babesia* surveillance and clinical case reporting suggest a significant increase in *B. microti* incidence in the United States (US) over the past two decades [3]. Factors that have been postulated for the emergence of *Babesia* include an increase in the deer population that amplifies the number of ticks, an increase in the human population, and building homes in tick infested areas [3–5]. *Babesia* was historically under-investigated, whereby greater attention (i.e., awareness) following its becoming a notifiable disease in many US states in 2011 likely contributed to the observed increase in cases.

*Babesia* are transmissible through blood transfusion [6]. The increase in reported cases of naturally acquired and transfusion-transmitted babesiosis (TTB) in the US drew the attention of the blood banking community, thus prompting over a decade of donor surveillance studies, along with the development of laboratory-based diagnostic and donor screening strategies to contend with TTB [7,8]. This culminated in 2019 with the publication of nonbinding recommendations from the US Food and Drug Administration (FDA) in favor of regional blood donor screening for *Babesia* in the US using an approved molecular

**Citation:** Bloch, E.M.; Krause, P.J.; Tonnetti, L. Preventing Transfusion-Transmitted Babesiosis. *Pathogens* **2021**, *10*, 1176. https:// doi.org/10.3390/pathogens10091176

Academic Editors: Estrella Montero, Jeremy Gray, Cheryl Ann Lobo and Luis Miguel González

Received: 31 July 2021 Accepted: 8 September 2021 Published: 13 September 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/).

assay [9]. Prior to the adoption of laboratory-based screening, *B. microti* was a leading infectious risk to the US blood supply. The risk of TTB in the US is now low as a result of routine testing for *Babesia* [10]. We describe the evolution of the response to TTB in the US as a means to contextualize the risk of *Babesia* in general with a view to guide future research efforts.

#### **2. Epidemiology: Geographic Distribution, Seasonality, and Transmissibility**

*Babesia* species have different geographic distributions. Cases of *B. microti* have been reported widely, notably in the northeastern and upper midwestern US, but also in other countries [11–15]. *Babesia duncani* occurs in the far western US [16]. *Babesia venatorum* and *Babesia crassa*-like agent have been reported in Europe and northeastern China [17–19]. *Babesia divergens*/*Babesia divergens*-like agents has been reported in Europe [17] and the United States [20]. *Babesia motasi*-like agent has been implicated in human cases in Korea [21].

#### *2.1. Blood Donor Surveillance in the US*

Beginning in the late 1990's, a series of surveillance studies were conducted to determine the seroprevalence of *Babesia* (specifically *B. microti*), as well as rates of parasitemia (using molecular positivity as a surrogate of active infection) specific to the blood donor population (Table 1). In one of the earliest studies, blood donors (*n* = 3490) in endemic and nonendemic areas of Connecticut were evaluated for *B. microti* [22]. In this study, 30 (0.9%) donors were confirmed positive for antibodies against *B. microti*; over half (10/19) of seropositive donors who were subsequently tested by PCR were shown to be positive [22]. In another study, about a fifth (21%) of 84 seropositive blood donors (IFA titers ≥ 64), who were followed for up to three years in Connecticut and Massachusetts, were found to be parasitemic [23]. Over the course of follow-up, protracted low-level parasitemia was variably and intermittently detectable.


**Table 1.** Transfusion-transmitted babesiosis: blood donor surveillance and follow-up studies in the United States.


**Table 1.** *Cont.*


**Table 1.** *Cont.*

AFIA—arrayed fluorescent immunoassay (AFIA); IFA—indirect fluorescent antibody; EIA—enzyme immunoassay; PCR—polymerase chain reaction; TTB—transfusion-transmitted babesiosis; NA—not applicable; CT—Connecticut; MA—Massachusetts; MN—Minnesota; WI—Wisconsin; VT—Vermont; AZ—Arizona; NY—New York; NM—New Mexico; OK—Oklahoma; FL—Florida; RI—Rhode Island.

#### *2.2. TTB in the US*

*Babesia* are intraerythrocytic parasites and are readily transmissible through transfusion of any product containing red blood cells. TTB has been reported following transfusion of whole blood, packed red blood cells (RBCs), and even frozen RBCs [6]. Confirmed cases of TTB have not been ascribed to transfusion of apheresis platelets and acellular blood products such as plasma and cryoprecipitate [6]. Rare cases of TTB have been reported after transfusion of whole blood-derived platelets [6]. This may have been due to contamination of red cells and/or the presence of extraerythrocytic parasites [34]. The minimum infectious dose of *B. microti* that can cause TTB is low (10–100 parasites), based on murine models [35] (Table 2). TTB following transfusion of pediatric red cell aliquots and whole blood-derived platelets suggests that infectivity is high.




**Table 2.** *Cont.*

**Abbreviations:** IFA—indirect fluorescent antibody; EIA—enzyme immunoassay; PCR—polymerase chain reaction; TTB—transfusiontransmitted babesiosis; IDT—individual donor testing; MP-NAT—minipool nucleic acid testing.

> To date, over 250 cases of TTB have been reported in the US, almost all (98%) of which were caused by *B. microti*. There are three reports of TTB due to *B. duncani* and one due to *B. divergens*-like parasites [20,41]. The risk of TTB is more widespread in the US than that associated with tick-borne transmission of parasites. Blood is often transfused far from where it is collected. It is not uncommon for blood products to cross state lines, where distribution is driven by clinical need, disproportionately being drawn toward major urban centers. In addition, residents from nonendemic areas may become asymptomatically infected during travel to endemic areas, return home, and donate blood [42]. *B. microti* can persist for long periods of time, even after standard antimicrobial therapy, whereby asymptomatic individuals may donate long after becoming infected [42,43]. These factors have accounted for cases of TTB in nonendemic states [44,45].

> Natural acquisition of *Babesia* is predominantly seasonal, with peak incidence spanning late spring to early fall, following the life cycle of the tick vector. By contrast, cases of TTB are not strictly confined to peak periods of vector-borne transmission, although they still have a similar time distribution pattern as tick-borne disease, having been reported throughout the year [6,46]. Prolonged storage of blood components enables transfusion of parasitemic blood long after donor acquisition of infection and expands transmission time to include the entire calendar year [8,43]. In addition, the incubation period for development of symptoms after transfusion is as long as six months [6]. Furthermore, donor

surveillance studies and prospective screening have also identified parasitemic donations (i.e., positive nucleic acid test) throughout the year, although positive donations still tend to occur from June to October [33].

#### **3. The Risk of Transfusion-Transmitted Babesiosis outside of the US**

To date, cases of TTB have been almost exclusively described in the US, with rare exceptions of reports in Japan and Canada (Table 3) [47]. Although it is well established that *Babesia* is globally ubiquitous, few studies have been undertaken to quantify risk of TTB outside of the US.

**Table 3.** Transfusion-transmitted babesiosis: blood donor surveillance and quantification of transfusion-associated risk outside of the United States.



**Table 3.** *Cont.*

A seroprevalence study was undertaken of Tyrolean blood donors (*n* = 988): 2.1% were IgG-positive against the *B. divergens* complex and 0.6% were seropositive for *B. microti* [48]. While both species are causes of human infections, *B. divergens* has not been found to be transmitted through blood transfusion.

Canada has plausible risk given its proximity to endemic US states, as well as previously described autochthonous cases. In one study, passive surveillance was utilized to guide follow-up active surveillance and intervention [49]. Specifically, ~12,000 ticks that had been submitted by the public were tested for evidence of *Babesia* infection. Fourteen were found to be *B. microti*-positive, 10 of which originated in Manitoba. This guided selection of regions for active surveillance (2009–2014) using tick drag sampling. The ticks were tested by PCR: 6/361 (1.7%) were positive in Manitoba and 3/641 (0.5%) were positive in Quebec. None were positive from other sites. Blood donations (July and December, 2013) at selected sites near endemic US regions were tested for antibodies to *B. microti*. A donor questionnaire was used to enquire about travel-related risk and possible tick exposure. A total of 13,993/ 26,260 (53%) donors were tested, none of whom were found to have antibodies to *B. microti.* Further, almost half (47%) reported having visited forested areas in Canada and 41% had traveled to the US. During a more extensive study performed in 2018, over 50,000 donations that had been collected near the US border were tested for *Babesia* nucleic acid by transcription-mediated amplification (TMA). In addition, a subset of 14,758 TMA-nonreactive samples was also screened for *B. microti* antibodies. The study identified one TMA-reactive donation that had been collected in Winnipeg, Manitoba, the only region in Canada where autochthonous infections have been reported, and four antibody-positive donations in the TMA-negative group [51]. Collectively, these findings suggest that the risk of TTB is low in Canada and that a risk-based deferral for *Babesia* is not needed at the moment.

A study was conducted in blood donors in China [50]. Again, there is a plausible regional risk given prior reports of human babesiosis in China, as well as in Mongolia, Korea, and Japan [14,54–58]. A total of 1000 donor samples representing 888 whole blood and 112 platelet donations that had been collected in Heilongjiang province were evaluated by IFA against *B. microti:* 13/1000 (1.3%) were seroreactive.

In Australia, a fatal case of autochthonous babesiosis due to *B. microti* raised concern pertaining to the national blood supply [11,59]. A total of 7000 donations were tested for anti-*B. microti* IgG by IFA [52]. Initial reactive samples were subjected to *B. microti* IgG and IgM (immunoblot), as well as PCR. Five donors were initially reactive by IFA, none of whom were confirmed during repeat testing. All were PCR-negative. In addition, clinically suspected cases of babesiosis (*n* = 29) were also evaluated; none were *B. microti* IgG, IgM, or DNA positive.

#### **4. Clinical Presentation**

Clinically, about a fifth of *Babesia* infections in adult immunocompetent hosts are subclinical or manifest as mild flu-like illnesses that are not diagnosed and often clear without treatment [60]. Most patients experience a mild to moderate febrile illness that typically consists of fatigue, headache, chills, and sweats. However, selected patient subsets are at high risk of severe disease with complications. The latter include hemolytic anemia; cardiorespiratory, renal, and/or liver failure; disseminated intravascular coagulopathy; and death [2]. Transfusion recipients harbor many of the risk factors for severe or even fatal babesiosis, such as advanced age, comorbid cardiac or pulmonary disease, immunodeficiency due to asplenia, cancer, HIV/AIDS, or sickle cell disease [2]. This helps to explain the severity of illness and high fatality rate (~19%) associated with transfusion-transmitted babesiosis (TTB) [1,6,60]. Indeed, variability in reported fatality rates from babesiosis, in general, largely reflects a difference in clinical penetrance that is governed by the immune status of the host [61–63]. Importantly, transfusion of red blood cells and whole blood is indicated for the treatment of severe, decompensated anemia. Therefore, parasite-induced hemolysis that might otherwise be tolerated in the immunocompetent individual can have dire consequences in the transfusion recipient.

#### **5. Prevention Strategies**

## *5.1. Risk-Based Deferral*

Historically, prevention of TTB has relied on donor selection (Table 4). Individuals who reported a history of babesiosis were permanently deferred from blood donation. This proved suboptimal, as evidenced by the number of cases of TTB that escaped detection using this approach. There are a number of reasons why this approach was problematic. For one, *Babesia* are able to persist chronically in donors without apparent adverse effects [42]. Even when clinically overt, the symptoms of babesiosis in immunocompetent adults are nonspecific. Risk factors for tick exposure are also nonspecific (e.g., outdoor activities, residence in highly endemic states), offering little diagnostic utility [32]. Vector (i.e., tick) borne transmission is seasonal, largely aligning with the tick life cycle, whereby most infections occur late spring to early fall [2]. By contrast, cases of TTB are less prone to seasonality, given that blood can be stored for prolonged periods. Further, persistent, asymptomatic infection is well described, in some cases being detectable for more than two years following infection [42,43].


**Table 4.** Approaches to address the risk of TTB.


**Table 4.** *Cont.*

A history of tick bites is also a poor predictor of infection. Recall of tick bite is unreliable. One study observed no significant difference in *Babesia* seroprevalence between those who reported tick bites as compared to those who did not [64]. The investigators postulated that those who report tick exposure are the same group who take precautions against tick bites. Importantly, a high proportion of infections are ascribed to the bites from nymphs rather than adult ticks. Nymphal ticks are the size of poppy seeds, rendering them highly inconspicuous.

#### *5.2. Laboratory-Based Methods for Donor Screening*

Laboratory testing is necessary for any meaningful donor screening intervention. Laboratory approaches in routine use for clinical diagnosis of babesiosis (e.g., microscopy of peripheral blood smears and manual indirect fluorescent antibody [IFA] testing) are not suitable for donor screening. Microscopy is neither scalable nor sufficiently sensitive or specific to detect the low level of parasitemia that is often encountered in blood donors. Manual IFA testing is not amenable to high-throughput screening. Molecular testing for *Babesia* is a more suitable approach for blood donor screening but poses novel challenges. *Babesia*—unlike the major transfusion-transmitted viruses— is primarily red-cell-based, thereby requiring additional processing steps for optimal sensitivity of detection. Given the large numbers of donors, automation is critical. Therefore, a process needed to be devised to better access the target parasites in the infected red blood cells.

#### *5.3. Serological Testing*

The initial approach for evaluating *Babesia* in the blood donor population was focused on serology (i.e., antibody capture)—specifically of anti-*B. microti* antibodies— in endemic areas. Experimental research assays were developed for the detection of *B. microti*. One approach used an enzyme immunoassay (EIA) (i.e., targeting the recombinant protein BMN-17 and MN-10) [36]; the other employed a semi-automated IFA test [22]. Although less labor-intensive, the EIA assay showed poor specificity as compared to the semiautomated IFA test. By contrast, IFA testing is sensitive and specific and is still used today to supplement positive nucleic acid test results. The semi-automated version of the IFA test, the arrayed fluorescent immunoassay (AFIA), was applied successfully in a series of donor surveillance studies [8,10]. The combination of AFIA and real-time PCR were the first tests to receive FDA licensure for screening of blood donations, but have since been discontinued for blood screening by the manufacturer [65].

Another antibody test, an enzyme-linked immunoassay (ELISA), was developed to detect antibodies against *B. microti*. The assay employed four immunodominant peptides from the BMN1 family that had been shown to be immunodominant and highly specific to *B. microti* [36]. The assay was capable of detecting both IgM and IgG against *B. microti* [29]. In a pilot study, 15,000 blood donor samples from high-risk, low-risk, and nonendemic areas of New York State (5,000 each) were tested. Rates of reactivity following application of a revised cutoff were 0.92%, (46/5000), 0.54% (27/5000), and 0.16% (8/5000), respectively [29]. ELISA repeat-reactive samples were also tested by IFA with a concordance rate of 99.34%. Although the ELISA was evaluated in a formal IND (investigational new drug) trial, which was a preliminary step along the regulatory pathway to licensure, the assay was never licensed and is no longer in use.

#### *5.4. Molecular Testing*

Molecular testing better detects active infection/parasitemia than antibody testing. This is important because active infection rather than *Babesia* exposure alone (i.e., antibodies), is required for transmission by blood transfusion. Nevertheless, mitigation strategies for blood donors focused initially on serological methods. Molecular assays (i.e., nucleic acid testing or NAT) have been used since ~1999 to detect the major transfusiontransmissible viruses (e.g., HIV, hepatitis B, and hepatitis C viruses) [66]. Those agents are detectable in plasma. By contrast, *Babesia* are primarily intraerythrocytic, requiring additional processing of whole blood to ensure adequate target capture.

A variety of PCR research assays, from nested to real-time, have been developed using the 18S ribosomal RNA gene of *B. microti* as a target and used to determine parasitemia in antibody-positive blood donations during surveillance studies [27,67]. In most cases, these assays have been shown to be sensitive and specific; however, the methodologies used to access the red cell compartment represented a limiting factor for the sensitivity of these assays for blood donor screening. In addition, hemoglobin is also a known inhibitor of PCR [68]. The first real-time PCR assays for donor screening utilized an automated membrane-based isolation system (Taigen Bioscience) and had a limit of detection of 66 piroplasms per mL [8]. Later, larger manufacturers such as Grifols Diagnostics and Roche developed assays, and ultimately obtained FDA licensure [69,70]. These assays are exquisitely sensitive and specific, attaining limits of detection for *Babesia* as low as 2–3 parasites/mL [66]. Both assays can detect ribosomal DNA or RNA of four major species of *Babesia* that infect humans (*B. microti*, *B. divergens*, *B. duncani*, and *B. venatorum*) [38]. The assays can be performed on an automated platform and in pools of 6 (Cobas Babesia, Roche Diagnostics) to 16 samples (Procleix Babesia assay, Grifols diagnostic solutions), allowing for the screening of large numbers of donations. One of the two assays in current use (Cobas Babesia, Roche Diagnostics) employs proprietary whole blood collection tubes containing lysing agents [38].

#### **6. Economic Impact**

The cost implications of donor screening have been assessed in three studies undertaken by different groups. The first study examined four different testing strategies as applied to endemic areas: universal antibody screening, universal molecular screening, universal combined testing (antibody/molecular), and recipient-risk-targeted combined (antibody/molecular) testing [71]. The strategies were compared to the then-current standard practice of using a questionnaire. The authors concluded that use of a questionnaire was most wasteful, followed by a risk-targeted combined approach. Universal molecular screening would incur an incremental cost-effectiveness ratio (ICER) of \$26,000 to \$44,000/quality adjusted life year (QALY) and would serve to prevent 24 to 31 TTB cases/100,000 units transfused, incurring no wastage. The combined approach would be more effective, albeit at a higher cost. By contrast, antibody-based screening was lower in cost, yet was less effective and incurred higher wastage than the molecular options.

The second analysis evaluated the cost utility of a similar repertoire of screening approaches in endemic areas [72]. The results were substantially different. For one, the ICER for combined testing as compared to antibody screening was in excess of \$8.7 million, preventing 3.6 cases of TTB per 100,000 units transfused. Universal endemic antibody screening was projected to prevent 3.39 cases of TTB at an ICER of \$760,000/QALY when compared to the recipient-risk-targeted strategy. The authors concluded that antibody was the most cost-effective strategy when applying the threshold of cost effectiveness specific to transfusion safety initiatives in the US, i.e., \$1 million/QALY.

The third study examined the cost-utility of different screening strategies, both by mode of testing (IFA, ELISA, PCR), as well as extent of geographic inclusion [73]. The authors concluded that even a strategy that was to be confined to highly endemic states would likely exceed the implicit threshold for cost-effectiveness of \$1 million per QALY.

#### **7. US Policy**

Babesiosis has long been recognized as posing a risk to the US blood supply [9,74]. However, availability of validated tests that were of sufficient level of performance for donor screening, impeded rapid adoption of preventive strategies [75]. In 2019, the US FDA published their recommendations, thus supporting regional molecular screening of blood donors in 14 states and Washington DC using any of the approved assays [9]. Over 95% of all cases of TTB and 99% of clinical cases of babesiosis have occurred in the selected locations. The recommendations also allowed for pathogen reduction (PR) as an alternative to laboratory testing. At the time of this writing, at least one PR technology had been FDA approved for use in plasma and platelets. Of note, neither plasma nor apheresis platelets pose significant—if any—risk of transfusion transmission. A history or babesiosis or a positive test for *Babesia* previously led to permanent deferral from blood donation. Under the new guidance, donor re-entry is allowable after 2 years in the event that the donor has not had a positive test result for *Babesia* during the interval, remains negative by requalification using one of the licensed *Babesia* NAT assays, and meets all other eligibility criteria for blood donation [9].

#### **8. Discussion**

Successful strategies to reduce the risk of the major transfusion-transmitted viruses (e.g., HIV, hepatitis B, and C viruses) have rendered blood transfusion remarkably safe, at least in the US and other high-income countries [66,76]. These successful strategies have contributed to the investigation of risk posed by other pathogens (e.g., *Babesia*) and classes of pathogens (e.g., bacteria) to the blood supply. Implementation of donor screening for *Babesia* in the US has been a success, having —arguably— removed one of the last major transfusion-transmissible infections, thus serving to advance blood transfusion safety nationally.

Nonetheless, the donor screening policy was long overdue. A potential contributing factor for the delayed development of *Babesia* blood screening assays was the evolution of *T. cruzi* screening in the US. *T. cruzi*, the causative parasite for Chagas disease, is transfusiontransmissible. The agent is endemic to Central and South America, where longstanding public health efforts coupled with serological testing of blood donors have contributed to a decline in cases [77]. Universal donor screening for Chagas disease began in the US in 2006. Following implementation, studies determined the risk in the US to be low. This prompted a revision of the policy at that time to restrict screening to first-time donor testing only. While rational in outlook, that shift in policy impaired commercial investment in testing. The downstream effect may have been the later, tepid support from the major test manufacturers—at least initially—for *Babesia* testing. Instead, the larger blood collection agencies, such as American Red Cross, Vitalant (then Blood Systems), and New York Blood Center, partnered with small businesses to develop assays.

The path to regulatory approval and development of a screening policy for *Babesia* took almost a decade. By way of comparison, implementation of routine testing for West Nile Virus in 2003 (lauded as a major success) took less than a year from recognition of transfusion-transmitted disease [78], a timeline bettered by the later adoption of screening for Zika in 2016 within weeks [79–81]. Of note, Zika has yet to show any evidence of clinical effect following the rare accounts of possible transfusion transmission. Collectively, this underscores the myriad of factors and competing priorities that guide blood transfusion policy, not all of which are scientific in nature [82].

While there may be an element of closure on TTB in the US, *Babesia* remain global pathogens. *Babesia* species have been described in both ticks and animal populations over a wide geographic distribution spanning the Americas, Europe, Asia, Africa, and Australia [2,11–13,17,55,58,83]. Outside of the US, perception of risk is low and the US remains the only country to have implemented blood donor screenings [77]. Over the last two decades, only six studies (and two case reports) pertaining to TTB have originated outside of the US (Table 3). Those studies did not find comparable risk to that encountered in the US [49,52,53]. Nonetheless, surveillance is lacking, with a grossly skewed geographic sampling that remains focused on the US. One of the challenges that previously impeded surveillance was the lack of diagnostic tools that could be applied to high-throughput testing. The advent of licensed, high-performance commercial *Babesia* PCR and TMA assays should enable testing across a more diverse geography, with the caveat that implementation of molecular testing, even for research use, is challenging for low- and middle-income countries [84]. While robust molecular assays may be available, the lack of local expertise and infrastructure may still necessitate the transfer of samples to settings where equipment is available.

#### **9. Conclusions**

*Babesia* are major transfusion-transmissible parasites. A concerted effort by the blood banking community has yielded effective policy and testing strategies that have been integrated into routine donation practices in the US. Nonetheless, these efforts have not been matched elsewhere and deserve greater attention from the international blood banking community. Further, the lessons learned from *Babesia* (e.g., related to sample preparation, thus enabling automated testing of an intraerythrocytic pathogen) can be applied to P*lasmodium* (malaria), a related parasite that remains a leading cause of transfusionassociated morbidity in much of the World.

**Author Contributions:** Conceptualization, E.M.B.; data curation, E.M.B.; writing—original draft preparation, E.M.B.; writing—review and editing, E.M.B.; P.J.K. and L.T. All authors have read and agreed to the published version of the manuscript.

**Funding:** EMB is supported by National Heart Lung and Blood Institute 1K23HL151826 (EMB). PJK is supported in part by the Gordon and Llura Gund Foundation.

**Conflicts of Interest:** EMB reports personal fees and nonfinancial support from Terumo BCT, Grifols Diagnostics Solutions, and Abbott Laboratories outside of the submitted work; EMB is a member of the United States Food and Drug Administration (FDA) Blood Products Advisory Committee. Any views or opinions that are expressed in this manuscript are that of the author's, based on his own scientific expertise and professional judgment; they do not necessarily represent the views of either the Blood Products Advisory Committee or the formal position of FDA, and also do not bind or otherwise obligate or commit either the Advisory Committee or the Agency to the views expressed. PJK reports research collaboration with Gold Standard Diagnostics. LT has no relevant conflicts of interest to disclose.

#### **References**


## *Review* **Sickle Cell Anemia and** *Babesia* **Infection**

#### **Divya Beri <sup>1</sup> , Manpreet Singh <sup>1</sup> , Marilis Rodriguez <sup>1</sup> , Karina Yazdanbakhsh <sup>2</sup> and Cheryl Ann Lobo 1,\***

	- New York, NY 100065, USA; kyazdanbakhsh@nybc.org
	- **\*** Correspondence: clobo@nybc.org

**Abstract:** *Babesia* is an intraerythrocytic, obligate Apicomplexan parasite that has, in the last century, been implicated in human infections via zoonosis and is now widespread, especially in parts of the USA and Europe. It is naturally transmitted by the bite of a tick, but transfused blood from infected donors has also proven to be a major source of transmission. When infected, most humans are clinically asymptomatic, but the parasite can prove to be lethal when it infects immunocompromised individuals. Hemolysis and anemia are two common symptoms that accompany many infectious diseases, and this is particularly true of parasitic diseases that target red cells. Clinically, this becomes an acute problem for subjects who are prone to hemolysis and depend on frequent transfusions, like patients with sickle cell anemia or thalassemia. Little is known about *Babesia*'s pathogenesis in these hemoglobinopathies, and most parallels are drawn from its evolutionarily related *Plasmodium* parasite which shares the same environmental niche, the RBCs, in the human host. In vitro as well as in vivo *Babesia*-infected mouse sickle cell disease (SCD) models support the inhibition of intra-erythrocytic parasite proliferation, but mechanisms driving the protection of such hemoglobinopathies against infection are not fully studied. This review provides an overview of our current knowledge of *Babesia* infection and hemoglobinopathies, focusing on possible mechanisms behind this parasite resistance and the clinical repercussions faced by *Babesia*-infected human hosts harboring mutations in their globin gene.

**Keywords:** *Babesia*; sickle-cell anemia; hemolysis; haemoglobinopathies

## **1. Introduction**

Human babesiosis is a zoonotic disease in which the natural acquisition of human cases is most often the result of an interaction with established zoonotic cycles [1,2]. A number of factors have contributed to the emergence of human babesiosis, including increased awareness among physicians, changing ecology, and an increased population of immuno-compromised individuals who exhibit severe disease. *Babesia* belongs to the Phylum Alveolata, Class Apicomplexa Family Piroplasmida and Genus *Babesia* which comprises more than 100 classified species. The four identified *Babesia* species that can infect humans are: *B. microti*, *B. divergens*, *B. duncani* and *B. venatorum*. As molecular techniques are becoming more available and accessible, new species described as "*microti*-like" or "*divergens*-like" are being described [3].

*Babesia* is an intra-erythrocytic parasite that causes malaria-like symptoms in infected people. *Plasmodium*, the causative agent of malaria, is the most studied Apicomplexan parasite and, like *Babesia*, resides within red blood cells. *Plasmodium* has a long association with its human host dating back to the first report in 1857 [4]. As the erythrocyte provides the parasite with the infra-structure to grow and multiply, it is expected that any perturbation to the cell should impact parasite homeostasis and viability. Clinical, epidemiological, and genome-wide association studies have identified multiple polymorphisms in the globin protein of hemoglobin within the red blood cell (RBC), commonly referred to as hemoglobinopathies, that attenuate or completely abrogate malaria pathogenesis.

**Citation:** Beri, D.; Singh, M.; Rodriguez, M.; Yazdanbakhsh, K.; Lobo, C.A. Sickle Cell Anemia and *Babesia* Infection. *Pathogens* **2021**, *10*, 1435. https://doi.org/10.3390/ pathogens10111435

Academic Editors: Estrella Montero, Jeremy Gray, Luis Miguel González and Lawrence S. Young

Received: 5 October 2021 Accepted: 1 November 2021 Published: 4 November 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/).

Malaria has thus imposed extreme selective pressure on the human genome, far more than any other infectious disease, and the RBC has been the prime target for evolutionary adaptation. The evolutionary proximity of *Plasmodium* and *Babesia* [5], and the fact that they both infect RBCs, raises important clinical questions of *Babesia* infections in patients harboring hemoglobinopathies.

In this paper, we review the literature documenting the effects of hemoglobinopathies on the life cycle of the *Babesia* parasite, using both in vitro and in vivo models of *Babesia* infection. We provide an overview of available clinical cases of the severity of *Babesia* infection in patients harboring these mutations and emphasize why it is essential to focus research in this area. We also describe plausible mechanisms that could exert this protective effect and discuss ways we can use this double-edged sword to develop better therapeutics against blood-borne parasites.

#### **2. Pathogenesis and Anemia in Babesiosis**

Hemolytic anemia is the central feature of sickle cell anemia (SCA) that contributes to its severe clinical outcomes. Epidemiological studies and basic research point to the pathogenic role of intravascular hemolysis as the primary cause of clinical complications in SCA. Interestingly, the primary pathological event in babesiosis is also hemolysis, resulting in hemolytic anemia and jaundice. In the absence of aggressive intervention, the anoxia and toxic effects that follow often lead to organ failure and death. Parasitemias do not always relate directly to the degree of anemia, suggesting that erythrocyte destruction is not only due to lysis of infected cells or their removal by splenic and liver macrophages, but also due to lysis of bystander cells which might be a significant contributing factor to the process. Some symptoms, such as fever, myalgia, renal insufficiency, coagulopathy, and hypotension, that occur in *B. microti* infections with parasitemias of less than 1%, may be caused by excessive production of pro-inflammatory cytokines, as also seen in malaria [6,7].

Clinical features in heavy infections, particularly those caused by another major human species, *B. divergens*-like parasites occurring in immuno-compromised patients, exhibit acute illness that appears suddenly with hemoglobinuria as a presenting symptom [8,9]. The clinical presentation also includes persistent non-periodic high fever (40–41 ◦C), shaking chills, intense sweats, headaches, myalgia, and lumbar and abdominal pain. Jaundice may develop as a result of the high level of hemolysis; vomiting and diarrhea may be present, and the toxins and anoxia, resulting from the hemolysis and the host immunological response, may cause respiratory, cardiac, renal, or hepatic failure [10–12]. The few known infections with *B. venatorum* have shown similar though generally milder manifestations [12].

In our previous report of *B. microti* infections in mice, we reported the increase in hemolysis in *Babesia*-infected mice, which was highly accentuated in mice harboring the SCA genotype, as observed by significantly reduced hematocrit and enhanced hemoglobinuria in these mice [13]. Therefore, these studies indicate that hemolysis is a central mechanism of clinical manifestations of both babesiosis and SCA individually, which is further accentuated and becomes life-threatening in *Babesia* infections in SCA mice/humans.

We found in our infected SCA mouse model that they mount an equally robust adaptive immune response despite exhibiting low parasitemia. This underscores the importance of examining both the fate of *B microti* and the immunological consequences of parasite infection in individuals with SCA to establish whether a similar hyperimmune response against the parasite occurs in humans too. Patients with SCA require transfusions, with some undergoing chronic transfusion therapy, placing them at greater risk of acquiring transfusion-transmitted infections like babesiosis. Thus, these individuals, if transfused from an infected donor, would be exposed to a larger infectious dose compared with a tick bite. The outcome of these infections, whether one of immune protection mediated by the first infection or a more deleterious pathological sequel, is required to be studied to establish effective treatments for these patients [13].

Laboratory findings that are consistent with mild-to-moderate hemolytic anemia include a low hematocrit, low hemoglobin level, low haptoglobin level, elevated reticulocyte

count, and elevated lactate dehydrogenase level [14]. Thrombocytopenia is commonly observed. The illness usually lasts for 1 or 2 weeks, but fatigue may persist for months. Asymptomatic parasitemia may persist for several months after standard therapy is initiated or for more than a year if the patient does not receive treatment. Illness may relapse in severely immuno-compromised patients despite 7 to 10 days of antimicrobial therapy and may persist for more than a year if not adequately treated [15].

Patients infected by *B. microti* show a wider range of signs and symptoms. A study on Block Island, Rhode Island, USA, concluded that about 25% of adults and 50% of children are asymptomatic or only show very mild 'flu-like' symptoms in cases that may not result in medical consultation and are therefore rarely diagnosed [16]. At the other end of the spectrum, very severe manifestations, similar to those seen in *B. divergens* infections, may occur in patients who have been splenectomized, are receiving immune-suppressive therapy, or are elderly. These cases typically show high fever, chills, night sweats, myalgia, hemolytic anemia, and hemoglobinuria [17]. Life-threatening complications include acute respiratory failure, disseminated intra-vascular coagulation, congestive heart failure, coma, and renal failure [18]. Immuno-compromised individuals are also likely to develop persistent relapsing disease despite treatment [15]. The symptoms caused by *B. duncani* and related parasites (CA1–4) closely resemble those of *B. microti* infections [19].

#### **3.** *Babesia* **and the Red Blood Cell**

When *Babesia* sporozoites are first injected into the human host, they target the host RBCs immediately, unlike *Plasmodium* spp. which are required to undergo an exoerythrocytic phase in hepatic cells. Furthermore, *Babesia*-infected RBCs remain circulating in the peripheral blood stream, including regularly passing through the hosts' spleen, and do not sequester to the fine capillaries of the bone marrow or organs. It is the parasite's ability to first recognize and then invade host RBCs that is central to human babesiosis, and the parasites invade RBCs using multiple complex interactions between parasite proteins and the host cell surface, which are not fully elucidated yet [20]. Once inside the RBC, the parasite begins a cycle of maturation and growth exhibited by intense intra-cellular proliferation leading to populations described as 1N, 2N, 4N and >4N [21]. The parasite population can expand inside the RBCs or egress at multiple points in the life cycle [21]. Previous work from our lab has led to the development of synchronized parasite populations and showed the sequential progression of the seven morphological forms of *B. divergens*in culture along with the dynamics of parasite proliferation and differentiation. These processes are maintained through controls that secure the constituent infected-RBC populations in strict ratios to enable rapid movement between new invasion events or further intra-RBC development and replication cycles, as dictated by the environment of the parasite. The early stages of the cycle are morphologically indistinguishable from *Plasmodium* spp., with both appearing as ring-like parasites. However, unlike *Plasmodium*, *Babesia* exhibits plasticity in its life cycle and is thus able to swiftly respond to environmental conditions like host RBCs and nutritional availabilities [22].

#### **4. Hemoglobinopathies**

For intracellular parasites, the environment of host red cells plays a key role in the development and success of the pathogen; therefore, perturbations in the RBCs are most likely to modify parasite survival and viability. For blood-borne parasites like *Plasmodium* and *Babesia*, the environments of the RBC, membrane proteins, and hemoglobin, the primary oxygen carrier, are important determinants of parasite success. Human hemoglobin is comprised of alpha and beta globin chains encoded from multiple globin genes. The α-globin gene cluster is at the end of chromosome 16 and contains three genes. The human β-globin gene cluster consists of five genes arranged in chromosome 11 in the same order in which they are expressed during human development: *e*, Gγ-, Aγ-, δ-, and β-globin gene. The Hb switching event which occurs after birth in the β-globin cluster leads to the suppression of the γ-globin gene accompanied by the complementary increase of the previously silent β-globin gene. Understanding the regulation of Hb switching can have direct

therapeutic applications for sickle cell disease in which the γ-globin gene can functionally substitute for mutations in the β-globin gene of these diseases [23]. Hemoglobinopathies are genetic disorders of the globin protein and are classified as structural hemoglobin variants including HbS, HbC and HbE, as described ahead, or thalassemia syndromes [24]. The term 'thalassaemias' collectively refers to several different genetic mutations that result in either reduced or absent expression of one or more of these globin alleles. Specifically, individuals described as having 'α-thalassaemia' have a loss of one or more α-globin allele(s). Additionally, there is also HbH disease (loss of 3 α-globin alleles) and, finally, hydrops fetalis (loss of all 4 α-globin alleles), which leads to the death of the fetus in the uterus. Individuals with mutations in HBB can also have a range of genetic defects referred to as 'β-thalassaemia', including β-thalassaemia minor (reduced expression of one β-globin allele), and β-thalassaemia major (reduced expression of both β-globin alleles) [25].

Sickle cell disease (SCD) is the most common monogenic blood disorder of hemoglobin synthesis, encompassing the single replacement mutation of glutamic acid at position 6 of the β-globin chain by valine (HbSS genotype) [26–28]. The hallmark of SCD is "the sickleshaped" red blood cells due to the polymerization of mutated sickle hemoglobin (HbS) under low oxygen tension. Chronic blood transfusion is one of the most effective treatments in SCD and results in the reduction of the frequency of acute pain episodes and acute chest syndrome but causes a dramatic increase in the risk of transfusion-transmitted infection [29]. The HbSS and HbAS (heterozygous) genotypes are commonly found in populations from sub-Saharan Africa. The Hemoglobin C (HbC) mutation (HbAC–heterozygotes; HbCC– homozygotes) also involves a point mutation at the 6th codon in the HBB gene, resulting in a glutamic acid to lysine substitution and is most common in West Africa, with prevalence reported as high as 15% in parts of Burkina Faso [30]. The Hemoglobin E (HbE) mutation is a point mutation that results in a glutamic acid to lysine switch at position 26 of the HBB gene and is most commonly found in parts of Southeast Asia and India and reaches a prevalence of up to 60% in some areas [31,32]. HbS, HbC, and HbE are characterized as structural hemoglobin variants. The major human hemoglobinopathies and related genetic mutations are summarized in Table 1. According to CDC Reports in 2010, the total incidence estimate for sickle cell trait was 15.5 cases per 1000 births in USA, ranging from 0.8 cases per 1000 births in Montana to 34.1 cases per 1000 births in Mississippi. The U.S. incidence estimate for sickle cell trait (based on information provided by 13 states) was 73.1 cases per 1000 black newborns, 3.0 cases per 1000 white newborns, and 2.2 cases per 1000 Asian or Pacific Islander newborns. The incidence estimate for Hispanic ethnicity was 6.9 cases per 1000 Hispanic newborns. The total number of babies born with sickle cell trait in 2010 was estimated to be greater than 60,000. The study showed that as many as 1.5% of babies born in the United States have the sickle cell trait [33]. With approximately 7% of the worldwide population being carriers, hemoglobinopathies are the most common monogenic diseases and one of the world's major health problems. This makes it very essential to understand the pathogenesis of blood-borne parasites in human hosts harboring these mutations in their RBCs.


**Table 1.** Major hemoglobinopathies and related genetic mutations.

Denotes: (-) loss of α-globin gene. (β ◦ ) loss of β-globin [34,35].

#### **5. Natural Resistance against Blood-Borne Parasites**

The long association and co-evolution of the malaria parasite with humans is reflected in the fact that almost all examples of molecular evolution in humans, like sickle cell anemia, G6PD-deficiency, and thalassemia, are attributed to a selection of mutations that attenuate malaria pathogenesis. Though these mutations lead to unpleasant consequences, as of 2015 it was estimated that about 4.4 million people have sickle cell disease, while an additional 43 million have sickle cell traits [36]. Zones that are endemic for malaria have a high proportion of humans carrying these mutations either in the homozygous form (the subject suffers from the disease caused due to the mutation) or heterozygous form (one copy of normal gene and one copy of mutated gene). These genes have all arisen in areas in which falciparum malaria is endemic, and their rise to high levels of prevalence is thought to result from their conferring significant degrees of protection against this dreaded pathogen. It is well-established that the homozygotes suffer from sickling of RBCs but do not support the rapid growth of the parasite; however, in rare cases, subjects with SCD and malaria can suffer from hyper-hemolytic crisis [37]. Heterozygotes do not suffer from sickling and have lesser severity of malaria. AS subjects can get malaria, but the number of parasitized cells is low, and they rarely suffer from cerebral malaria or severe anemia [37]. The enhanced resistance of persons with sickle traits to falciparum malaria is substantial. Infected AS children have lower parasite densities than AA children and are 50–90% less likely to progress to a severe form of malaria or to die from the disease [38]. *Babesia*'s association with hemoglobinopathies is not completely understood but is an important field of research as hemolytic anemia is common in hemoglobinopathies and can be life-threatening when coupled with *Babesia* infection.

There have been several explanations as to why β-globin might confer resistance to malaria. Researchers have reported that parasitized AS cells sickle more readily and show enhanced HbS polymerization under hypoxic conditions and are therefore removed from circulation. Further, it has been shown that parasites are fragile and killed by these HbS polymers. A compelling cause of reduced parasite load in AS and SS RBCs is the extent of oxidative damage which is inherent in these host cells added to the oxidative stress due to parasite growth; the cumulative oxidant damage can cause considerable damage to the host RBC and impairment of parasite development [39]. Interestingly, accumulated reactive oxygen species (ROS)-mediated damage is a common mechanism shared by AS, SS, G6PD-deficient, β- and α-thalassemia RBCs in mediating resistance to malaria [40,41]. However, the mechanism for ROS-mediated protection in malaria remains elusive. It was also observed that AS RBCs parasitized with *P. falciparum* late stages bound to human microvascular endothelial cells and blood monocytes half as effectively as did comparably infected AA RBCs. Moreover, infected AS RBCs displayed slightly reduced and highly uneven distribution of expression of PfEMP-1 on their surface. There is also evidence based on host microRNAs playing a role in protection in AS and SS RBCs [38,42–50]. There have been several plausible mechanisms proposed for resistance of hemoglobinopathic RBCs to malaria, but little is known about *Babesia* in this regard. Given the parallels between the two parasites, it is tempting to speculate that they might share mechanisms of resistance to growth in hemoglobinopathic RBCs.

#### **6.** *Babesia* **and the Sickle Red Cell**

The RBC serves as the home for this intra-erythrocytic parasite for its entire life cycle in the human host. The interactions between the parasite and the RBC can be classified into three broad areas: invasion, growth and maturation within the RBC, and egress. Previous studies from our group have examined these phases of the intra-erythrocytic lifecycle of *Babesia divergens* in homozygous SS and heterozygous AS human blood [51]. While the invasion was similar across all RBCs, there was atypical population progression, a potential loss of merozoite infectivity, and defective egress of the parasite in SS cells (as explained in Figure 1). Unlike previous reports in *Plasmodium*, AS cells supported invasion, growth, and egress of *Babesia* much like AA cells. While parasites grew from their characteristic 1N to

2N, 4N and >4N populations in AA and AS cells, in SS host cells beyond 24 h, the majority of the parasites were stuck in the 1N phase as demonstrated in Figure 1. Interestingly, even when parasites growing in SS RBCs were supplemented with fresh AA RBCs, they did not grow [51]. This indicates that the initial invasion and growth in SS host cells programs the parasite irreversibly to poor growth and/or defects in egress. Our work on the mouse model using *B. microti* also showed poor growth of the parasite in mice harboring the SS gene (HbSS-Townes mice), and normal growth in AS mice when compared to the wildtype AA mice [13]. For AA and AS mice, parasitemias peaked on day 7 of infection, while the SS mice exhibited a sluggish increase in parasitemias. In all three genotypes, parasites were cleared by day 21 and all mice survived. Interestingly, while the parasitemia was 4–5-fold lower in SS mice, the extent of immune response mounted was the same in AA, AS and SS mice. The adaptive immune response was measured by a robust GC reaction and significant expansion of TFH cells. Currently, it is not known how these SS mice respond to subsequent *Babesia* infection. This becomes especially critical to understand as babesiosis is primarily a transfusion-transmitted infection and since several sickle cell patients undergo repeated blood transfusions, they may be exposed to a parasite load that could be more than the parasites in one tick bite. yond 24 h, the majority of the parasites were stuck in the 1N phase as demonstrated in Figure 1. Interestingly, even when parasites growing in SS RBCs were supplemented with fresh AA RBCs, they did not grow [51]. This indicates that the initial invasion and growth in SS host cells programs the parasite irreversibly to poor growth and/or defects in egress. Our work on the mouse model using *B. microti* also showed poor growth of the parasite in mice harboring the SS gene (HbSS-Townes mice), and normal growth in AS mice when compared to the wildtype AA mice [13]. For AA and AS mice, parasitemias peaked on day 7 of infection, while the SS mice exhibited a sluggish increase in parasitemias. In all three genotypes, parasites were cleared by day 21 and all mice survived. Interestingly, while the parasitemia was 4–5-fold lower in SS mice, the extent of immune response mounted was the same in AA, AS and SS mice. The adaptive immune response was measured by a robust GC reaction and significant expansion of TFH cells. Currently, it is not known how these SS mice respond to subsequent *Babesia* infection. This becomes especially critical to understand as babesiosis is primarily a transfusion-transmitted infection and since several sickle cell patients undergo repeated blood transfusions, they may be exposed to a parasite load that could be more than the parasites in one tick bite.

into three broad areas: invasion, growth and maturation within the RBC, and egress. Previous studies from our group have examined these phases of the intra-erythrocytic lifecycle of *Babesia divergens* in homozygous SS and heterozygous AS human blood [51]. While the invasion was similar across all RBCs, there was atypical population progression, a potential loss of merozoite infectivity, and defective egress of the parasite in SS cells (as explained in Figure 1). Unlike previous reports in *Plasmodium*, AS cells supported invasion, growth, and egress of *Babesia* much like AA cells. While parasites grew from their characteristic 1N to 2N, 4N and >4N populations in AA and AS cells, in SS host cells be-

*Pathogens* **2021**, *10*, x FOR PEER REVIEW 6 of 10

**Figure 1.** *Babesia* infection progression in wild type (AA) RBCs, heterozygous for sickle cell anemia (AS) RBCs, and homozygous for sickle cell anemia (SS) RBCs. (**a**) In AA and AS host cells, the merozoite invades and the parasite develops inside the RBCs to 1N, 2N, 4N and >4N populations. Egress can take place at 2N, 4N or >4N stage. (**b**) The distribution percentage of 1N, 2N and 4N parasites is similar. (**c**) In SS host RBCs, parasites mostly retain their "ring form" and very few "Maltese cross" forms are seen. (**d**) As shown in the pie chart, a high population of parasites get stuck in the 1N form. (**e**) List of probable reasons for compromised growth of *Babesia* parasites in SS cells.

CDC reported that from 1979 to 2009, 159 transfusion-related *Babesia microti* cases were identified, most (77%) of which were from 2000 to 2009 [52]. A recent review in 2016 has summarized the state-wise seropositivity of *B. microti* in the blood used for transfusion in the USA [53]. However, no data are available on the number of sickle patients transfused with *Babesia*-contaminated blood.

### **7. Clinical SCA and Babesiosis**

There have been scattered case reports of babesiosis in sickle cell patients transmitted via blood transfusions. Transfusion-acquired babesiosis can result in severe hemolytic anemia in patients with sickle cell disease. The infection can be difficult to treat and may require a prolonged treatment duration [54]. A recent study presented a case of two sickle cell patients who had delayed diagnosis post transfusion due to confusing symptoms: as patients who receive chronic transfusions are also at risk for the development of allo- and autoantibodies, the hemolytic anemia caused by the former can often obscure a different pathophysiology, such as babesiosis, which occurred with these patients. In another case, diagnosis took 4 months for a patient with HbSS and babesiosis, after repeated visits to the hospital [54,55]. Another study reported a young female with HbSC who presented in the emergency department multiple times with pain and shortness of breath, eventually developing unresponsiveness and a brief episode of pulseless electrical activity. She was admitted to the intensive care unit with multisystem organ failure and found to have diffuse ischemic strokes. Infectious workup revealed disseminated anaplasmosis and babesiosis, which had likely caused sickle cell crisis. The patient continued to show a significant neurologic burden, despite months of treatment [56]. Evidently, with increased awareness about babesiosis among physicians and more sensitive diagnostic tests available, the number of case reports on babesiosis have increased. Given that there is a significant incidence of SCA carriers/other hemoglobinopathies in *Babesia*-endemic zones (like northeastern USA) who require frequent transfusions, it becomes increasingly necessary to understand *Babesia* pathogenesis in such subjects.

#### **8. Plausible Mechanisms for Resistance of** *Babesia* **in Sickle Cells**

From previous work of our group, the outstanding question is why the parasite exhibits developmental/egress abnormalities when growing in a sickle cell as opposed to a wild type AA RBC host. There are multiple possible causes for this: It is well known that sickle cells have a high burden of oxidative stress due to repeated polymerization and depolymerization of hemoglobin. It is also well known that a wide variety of intracellular pathogens like *Plasmodium*, *Mycobacterium,* and several viruses impose redox stress on their host cells. Therefore, unfavorable host cell conditions, like increased hemoglobin autoxidation, accumulation of iron in membranes, increased membrane damage, and a shorter red cell life span, could justify the reason why SS cells do not promote growth of the *Babesia* species. Further, the bystander effect, whereby uninfected RBCs are also affected leading to increased hemolysis, has been widely described in malaria. It is possible that in SS subjects there is an accentuated bystander effect leading to massive hemolysis and therefore unfavorable conditions for the parasite to grow and proliferate. It is also possible that invasion/growth of the parasite modifies the shape of the sickling RBCs, making them more prone to splenic removal. Our study in the mouse model has clearly shown that even though parasitemias are much lower in SS mice as compared to AA mice, the adaptive immune response is almost as severe; therefore, the heightened immune response against the parasite might be another SS-specific strategy to abrogate the growth of *Babesia*. In future studies, it would be interesting to monitor the growth of *Babesia* in other hemoglobinopathic disorders like thalassemia and RBC enzymopathy like G6PD-deficiency, both of which are known to afford resistance to malaria. However, unlike *Plasmodium*, AS mice showed no protection against babesiosis and followed the same parasite growth curve both in vitro and in vivo. Thus, from the current experiments, heterozygotes for the mutated beta globin gene do not seem to be protected against babesiosis.

#### **9. Concluding Remarks and Future Directions**

Hemoglobin-associated genetic disorders affect millions throughout the world and are concentrated in humans living in malaria-endemic countries. However, as borders of countries are becoming more porous, these genetic traits are now seen throughout the world. For several years, researchers have observed that hemoglobinopathies afford protection from malaria and the studies from our group in *Babesia* also point in that direction. Given the evolutionary proximity between these two parasites, it is possible that resistance to their growth in SS cells has a common mechanism. Further studies are needed to understand if the growth of *Babesia* parasites in thalassemic RBCs and those with an inherent deficiency in the G6PD enzyme is similarly impaired and to determine how these mutations hinder intraerythrocytic parasite growth. These results will provide researchers with an opportunity to discover the Achilles' heel of two deadly parasites and learn how nature has evolved a way to protect against these diseases. Uncovering the mechanism behind this protection will lead us to a better understanding of their pathogenesis as well as in designing better drugs against these parasites. As described above, multiple mechanisms of resistance against parasite proliferation in sickle cells may operate to confer protection. A detailed study of these pathways is needed to identify the main pathways in *Babesia*-infected red cells and this, in turn, will shed light on the intricate interplay between polymorphisms of the human host red cells and intruding parasites.

**Funding:** This research was funded by National Institutes of Health P01 HL149626 (K.Y. and C.A.L.) and R01HL140625 (C.A.L.) and a grant from BNY Mellon (K.Y. and C.A.L.).

**Institutional Review Board Statement:** Not applicable.

**Informed Consent Statement:** Not applicable.

**Data Availability Statement:** Not applicable.

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

#### **References**

