*Article* **Patulin Detoxification by Recombinant Manganese Peroxidase from** *Moniliophthora roreri* **Expressed by** *Pichia pastoris*

**Shuai Wang 1, Xiaolu Wang 2, Leena Penttinen 3, Huiying Luo 2, Yuhong Zhang 1, Bo Liu 1, Bin Yao 2, Nina Hakulinen 3, Wei Zhang 1,\* and Xiaoyun Su 2,\***


**Abstract:** The fungal secondary metabolite patulin is a mycotoxin widespread in foods and beverages which poses a serious threat to human health. However, no enzyme was known to be able to degrade this mycotoxin. For the first time, we discovered that a manganese peroxidase (*Mr*MnP) from *Moniliophthora roreri* can efficiently degrade patulin. The *Mr*MnP gene was cloned into pPICZα(A) and then the recombinant plasmid was transformed into *Pichia pastoris* X-33. The recombinant strain produced extracellular manganese peroxidase with an activity of up to 3659.5 U/L. The manganese peroxidase *Mr*MnP was able to rapidly degrade patulin, with hydroascladiol appearing as a main degradation product. Five mg/L of pure patulin were completely degraded within 5 h. Moreover, up to 95% of the toxin was eliminated in a simulated patulin-contaminated apple juice after 24 h. Using *Escherichia coli* as a model, it was demonstrated that the deconstruction of patulin led to detoxification. Collectively, these traits make *Mr*MnP an intriguing candidate useful in enzymatic detoxification of patulin in foods and beverages.

**Keywords:** patulin; mycotoxin; manganese peroxidase; apple juice; detoxification

**Key Contribution:** *Mr*MnP efficiently degraded patulin, particularly in the malonate/Mn2+ system, lead to detoxification. It removed 95% of patulin in a simulated patulin-contaminated apple juice after 24 h treatment.

### **1. Introduction**

Patulin (PAT) is a secondary metabolite and a food-born mycotoxin produced by at least 60 different filamentous fungi including *Penicillium expansum*, *Penicillium shell*, *Penicillium clavum*, and *Aspergillus clavatus* [1]. This mycotoxin has been detected in many kinds of fruits (such as apples, pears, grapes, kiwifruit, blueberries, and peaches) and their products (such as juice, jam, and cider) [2–4]. Based on cellular and animal toxicological studies, it has been found that patulin can cause genotoxicity, embryonic toxicity, cytotoxicity, neurotoxicity, immunotoxicity, carcinogenicity, and teratogenicity [5]. At the molecular level, patulin induces DNA damage, leading to cell-cycle arrest, which inhibits the activity of cell survival proteins and induces apoptosis until cell death [5]. Because of its toxicity and high frequency of contamination, the World Health Organization (WHO), some European countries, the Food and Drug Administration of the United States (FDA), and the Ministry of Health of China have all set up their recommended maximum concentrations of patulin in foods and beverages. For example, 50 μg/kg, 25 μg/kg, and 10 μg/kg of patulin are allowed in apple juice, solid apple products, and fruit baby foods, respectively, in the European Union [6]. However, it was noted that, although legal provisions enforced

**Citation:** Wang, S.; Wang, X.; Penttinen, L.; Luo, H.; Zhang, Y.; Liu, B.; Yao, B.; Hakulinen, N.; Zhang, W.; Su, X. Patulin Detoxification by Recombinant Manganese Peroxidase from *Moniliophthora roreri* Expressed by *Pichia pastoris*. *Toxins* **2022**, *14*, 440. https://doi.org/10.3390/ toxins14070440

Received: 8 June 2022 Accepted: 27 June 2022 Published: 29 June 2022

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**Copyright:** © 2022 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/).

in Serbia are in line with the EU regulations, from the 142 kinds of fruit juices (apple or multiple fruits) collected from the market in three consecutive years (2013–2015), patulin was detected in 51.4% of the fruit juices, and 0.7% of the samples exceeded the legal limit of 50 μg/kg [7]. In Spain, when analyzing PAT in 161 apple juice, 77 solid apple food, and 146 apple baby foods, PAT was discovered in 42% of the apple sauce samples, 32% in multiple fruit plates, and 25% in apple juice [8]. In Italy, 65% of 120 fruit plates and fruit paste samples tested positive for patulin [9]. The wide existence of patulin, in conjunction with its detrimental effects to the human health, points to the necessity to eliminate this mycotoxin in foods.

To remove the patulin contamination in foods, many trials have been carried out previously. These can be classified as physical, chemical, and biological treatments. Ultraviolet radiation is a physical means approved by both Canada and the United States for degradation of patulin in food [10], but the turbidity of apple juice and cider, plus the ascorbic acid present in large quantities, can significantly diminish the effect of this treatment. Treatment with chemicals such as ozone was reported to rapidly remove up to 98% of patulin within 1 min [11]. However, this manipulation can also generate new, undefined, chemicals and cause loss of important nutrients, thereby limiting its wide application in the food industry. These drawbacks prompt the scientific community to pursue other economically viable, efficient, safe, and environmentally friendly ways to remove the patulin and ensure food safety. Thus, biological decomposition of patulin in food by microorganisms or enzymes is emerging as an attractive alternative.

Up to now, the microorganisms that have been reported to be able to effectively degrade patulin include *Pichia caribbica* [12], the marine yeast *Kodameae ohmeri* [13], the biocontrol yeast *Rhodosporidium kratochvilovae* [14], and *Rhodotorula mucilaginosa* [15]. The rate of decomposition can reach as high as 97.66% [13]. Although the use of microbial cells has been proven to be an effective strategy, the presence of abundant residual cells and high concentrations of metabolites after treatment may change the final quality of the product. In essence, microbes transform patulin by their encoding enzymes. Unlike the microorganisms, the enzyme biocatalysts retain the ability to degrade patulin but do not introduce unwanted cells or their metabolites. Therefore, the use of detoxifying enzymes, which has the potential to ensure food safety and quality, should be a promising strategy in controlling patulin contamination in foods. However, even in the patulin-degrading microorganisms, the enzymes responsible for eliminating patulin have not been identified, thus impeding the use of an enzyme to detoxify patulin. Moreover, some microbes employ a strategy to import the mycotoxin and degrade it within the cell. This commonly involves the participation of expensive co-factors such as NAD+/NADH or NADP+/NADPH, which may not be economically viable in practical applications [16,17].

Manganese peroxidases (MnP) are a kind of heme-containing peroxidases, which are generally produced by the lignocellulose-degrading basidiomycete filamentous fungi and symbolized by their ability to oxidize Mn2+ to Mn3+ [18]. In nature, the oxidized magnesium is chelated by dicarboxylic acids (such as oxalate [19]) excreted by the fungi, which can stabilize Mn3+ and penetrate into lignocellulose to attack the recalcitrant parts. Previously, we have demonstrated that manganese peroxidases can detoxify four major feed mycotoxins including aflatoxin, zearalenone, deoxynivalenol, and fumonisin [20], highlighting their potential to be used in feed. Moreover, this enzyme can act by either directly interacting with the substrate or indirectly through an oxidized Mn3+. Neither of the two ways involves the necessity of an expensive co-factor, saving the cost in mycotoxin degradation. However, the ability of MnP to degrade patulin remains unknown. Therefore, to understand whether a manganese peroxidase can degrade patulin and whether the degradation leads to detoxification, in this study, *Mr*MnP, a manganese peroxidase originating from *Moniliophthora roreri* [21] and recombinantly prepared in *Pichia pastoris*, was explored for its ability to degrade patulin (Figure 1). The nature of the degradation products was analyzed by mass spectrometry and the toxicity of the degradation products was determined.

**Figure 1.** Schematic diagram showing the use of a recombinant *Mr*MnP expressed in *P. pastoris* to degrade and detoxify patulin, a mycotoxin commonly discovered in fruits.

#### **2. Results and Discussion**

#### *2.1. Recombinant Production of the Manganese Peroxidase MrMnP*

The most convenient way of characterizing the ability of a manganese peroxidase to degrade patulin would be to use a recombinant enzyme. However, the manganese peroxidases are notoriously difficult to be recombinantly produced, which might be, at least in part, ascribed to the presence of the heme prosthetic group in this kind of enzyme. Accordingly, many MnPs are expressed as insoluble, non-functional inclusion bodies in *Escherichia coli* and require tedious denaturation and refolding processes to obtain active enzymes [22–24]. However, in the commonly used eukaryotic microbial expression systems *P. pastoris* and *Aspergillus* spp., there were occasional reports indicating that a few MnPs could be functionally expressed, despite commonly being found in minor amounts [25–30]. The *M. roreri* MnP (*Mr*MnP) is such an enzyme that can be successfully produced in *P. pastoris* [21]. The gene encoding *Mr*MnP was thus artificially synthesized, cloned into the *Eco*RI and *Not*I restriction sites of pPICZα(A) to generate the recombinant plasmid pPICZα(A)-*Mr*MnP, and transformed into the *P. pastoris* X33 strain (Figure 2A). The transformant bearing the *Mr*MnP gene was first cultured in BMGY medium until the optical density of the culture at 600 nm (OD600) reached 6.0. Then, the medium was changed to BMMY to induce *Mr*MnP and the culture was continued for 5 d. The recombinant strain produced 271.4 and 3659.5 U/L of MnP (using ABTS as the substrate) in flask and fedbatch fermentations (Figure 2B), respectively. This is comparable to that described by Agathe et al. [21], thus providing a sound basis for subsequent enzymatic characterization and degradation of patulin.

**Figure 2.** Expression of *Mr*MnP in *P. pastoris*. (**A**) The plasmid map of pPICZα(A)-*Mr*MnP. (**B**) SDS-PAGE analysis of the *Mr*MnP enzyme recombinantly produced in *P. pastoris*. The arrow indicates the recombinant *Mr*MnP protein. Lane M: protein molecular mass marker; 1: *Mr*MnP protein recombinantly produced in *P. pastoris*.

#### *2.2. The Di-Carboxylic Acids Play a Critical Role in Degradation of Patulin by MrMnP*

For an enzyme, the buffering system normally exerts a profound effect on its activity. Specifically, for manganese peroxidase, the buffer components may be directly involved in the reaction. For example, eight MnPs from different microbial sources have been demonstrated to be able to degrade four major feed mycotixins in presence of the di-carboxylic acid malonate [20], indicative of the potency of MnPs in detoxifying mycotoxins. However, in that study, the tested buffers were restricted to malonate, lactate, and acetate, while the effects of the buffers were only tested on aflatoxin B1 and zearalenone. Herein, the effects of the buffer components on degradation of patulin were systematically examined by including the tri-carboxylic acid (citrate), di-carboxylic acids (malonic acid and oxalic acid), the α-hydroxyl carboxylic acid (lactate), the mono-carboxylic acid (acetate), the inorganic acid (phosphate), and the zwitterionic buffers (MES, standing for 2-morpholinoethanesulphonic acid; and HEPES, standing for 2-[4-(2-hydroxyethyl)-1-piperazinyl] ethanesulfonic acid). In the previous study, it was noted that zearalenone, but not aflatoxin B1, could be transformed at a similarly high rate in the acetate buffer to that in the malonate buffer. Similarly, *Mr*MnP was highly efficient in degrading patulin, eliminating all the mycotoxin after 5 h of incubation in the malonate buffer (Figure 3). Contrary to the situation for zearalenone, patulin was only slightly degraded in the acetate buffer (10.6 ± 0.1%). The degradation in oxalate was not as high as that in malonate, but still obvious (54.3 ± 0.3%). The degradation was reduced to 28.5 ± 0.1% in citrate buffer and decreased to be marginal in the lactic acid (15.2 ± 3.4%) and phosphate (11.5 ± 2.7%). The reason for ineffective degradation may best be explained by the knowledge that Mn2+ cannot form stable chelates with either of the two acids. Degradation of mycotoxins in oxalate and citrate is of physiological relevance since, in nature, the fungi expressing MnPs can also produce these two compounds in their cellular metabolism [22]. Additionally, it has been reported that gluconic acid, cellobionic acid, and other organic acids as well, can aid manganese peroxidases to exert their degradation on lignin molecules [31]. This allows the fungi to use manganese peroxidases, in assistance with these naturally occurring molecules, to break the lignin barrier apart and capture energy from the plant cell wall polysaccharides [32]. Unexpectedly, there were 42.6 ± 0.3% and 33.1 ± 0.5% degradation in the MES and HEPES buffers (Figure 3), higher than those in acetate, citrate, lactate, and phosphate buffers. In the univariate analysis, there was an extremely significant difference (*p* < 0.001), indicating that the degradation of patulin was affected by the buffer.

**Figure 3.** Effects of the buffer components on patulin transformation. The reactions were carried out by incubating 0.5 U/mL of *Mr*MnP with 5 mg/L of patulin in one of the buffers containing malonate, acetate, oxalate, citrate, lactate, phosphate, MES, and HEPES at 30 ◦C for 24 h. The data in the picture is the average ± standard deviation, \*\*\* *p* < 0.001, One Way anova test.

#### *2.3. Mn2+ Is Another Key Determinant in Degradation of Patulin by MrMnP*

Next, we sought to determine if the presence of Mn2+ would affect the activity of *Mn*MnP in patulin degradation. This is because the manganese peroxidases commonly have two substrate channels. One is the δ-heme edge, responsible for oxidation of hydrophobically-bound substrates including ABTS and phenolic compounds. The other one is the γ-heme edge, which is involved in catalysis of Mn2+ [33]. Patulin is a small chemical with ring structures, which might best fit for the first substrate channel, but could not enter the γ-heme edge. Incubation of *Mr*MnP with patulin in the acetate buffer, with or without Mn2+, did not lead to obvious degradation of patulin. The degradation rate is 1.1 ± 0.2% in acetic acid without Mn2+ and 10.6 ± 0.1% in the system containing Mn2+. The degradation was only observed in presence of both malonate and Mn2+ (Figure 4). The degradation rate reached 52 ± 0.3% at 2 h, and patulin was completely degraded at 5 h. These results collectively indicate that *Mr*MnP degraded patulin via a Mn3+-mediated indirect way, and patulin could not enter the δ-heme edge for degradation. The enzyme first catalyzed oxidation of Mn2+ to Mn3+, which then formed a chelate with malonate and subsequently oxidized patulin. Therefore, the enzyme's activity against patulin appeared to be related to the stability of Mn3+ in complex with the buffer components in the reaction systems. The stability and reactivity of the Mn3+ ion are strongly dependent on the nature and concentrations of the Mn3+-complexing agents [18]. Both oxalate and malonate are dicarboxylic acids, but the degradation rate of patulin in malonate was higher, indicating that the chelate of Mn3+ and malonate is more stable and hence has higher activity for patulin. Based on this assumption, it was also suggested that, in the reactions with MES and HEPES serving the buffer systems, the newly generated Mn3+ was stabilized by these

two buffers to a higher extent than that in acetate, citrate, lactate, and phosphate buffers. Although the reason for putatively increased Mn3+ stability remains unknown, it was noted that both chemicals have the sulfonate group, which could be involved in this stabilization.

**Figure 4.** Mn2+ played an important role in degrading patulin. The reactions were carried out by incubating 0.5 U/mL of *Mr*MnP with 5 mg/L of patulin in the acetate or malonate buffer in absence or presence of Mn2+ at 30 ◦C for 24 h. Then the products were analyzed by HPLC.

#### *2.4. MrMnP-Catalyzed Degradation of Patulin Led to Detoxification*

The pre-requisite to apply *Mr*MnP in patulin detoxification is that the degradation products should have much less toxicity. *Escherichia coli* has been used as a microbial sensor system for successful monitoring of the toxicity of patulin [34]. In this study, increasing concentrations of untreated patulin were first incubated with *E. coli*. At 1 mg/L and 10 mg/L of patulin. There was no significant negative impact on the bacterial growth in a culturing period of 10 h. However, when 50 mg/L and 100 mg/L of patulin were added, the growth of *E. coli* was significantly retarded, as manifested by the drop of OD600 from 1.03 to 0.89 (for 50 mg/L) and 0.79 (for 100 mg/L) at 10 h after incubation (Figure 5A). These results indicated that *E. coli* could indeed be used to monitor the toxicity of patulin.

Patulin samples at final concentrations of 1, 10, 50, and 100 mg/L, respectively, were individually treated with 0.5 U/mL of *Mr*MnP. The HPLC analysis indicated that under all concentrations tested, the patulin was completely degraded (data not shown). It was observed that, at all these concentrations, the treated patulin no longer retarded the growth of *E. coli*, which suggested that the degradation of patulin by *Mr*MnP led to detoxification (Figure 5B).

**Figure 5.** *Mr*MnP-catalyzed degradation of patulin led to detoxification. (**A**) Patulin was toxic to *E. coli* as demonstrated by retarded growth of the bacterium. A series of concentrations (1, 10, 50, and 100 mg/L) of patulin were added to equal amounts of *E. coli* and the culture was continued at 37 ◦C for 12 h. (**B**) *Mr*MnP-catalyzed degradation of patulin alleviated the retarding effect of patulin on *E. coli*. Patulin (1, 10, 50, and 100 mg/L) was first treated with 0.5 U/mL of *Mr*MnP at 30 ◦C for 24 h and then added to *E coli*.

#### *2.5. Structural Analysis of the Degradation Products*

As *Mr*MnP-catalyzed degradation of patulin led to detoxification, it would be interesting to know the chemical nature of the degradation products. Degradation and detoxification of mycotoxins are a process that, in essence, involves the transformation of mycotoxins into less-toxic or even non-toxic compounds [35]. Therefore, the degradation products of *Mr*MnP on PAT were further identified by UPLC-MS/MS. It was found that hydroascladiol (5-(2-hydroxyethyl)-4-(hydroxymethyl)furan-2(5*H*)-one) was one of

the main intermediate degradation products of PAT. The parent ion appeared at *m*/*z* 157.1 [M−H]−, producing daughter ions of 129.0 [M−H−CO]<sup>−</sup> and 113.1 [M−H−CO2] − (Figure 6). Daughter ions were produced by continuous loss of carbon dioxide [36]. In a previous study using *Lactobacillus plantarum* to degrade PAT, hydroascladiol was also obtained as the degradation product [37]. However, other intermediates including (E)-ascladiol and (Z)-ascladiol identified in their study were not discovered in our study. The toxicity of patulin is related to the hemiacetal and lactone rings in its structure. The generation of hydroascladiol, and hence concurrent destruction of the hemiacetal ring, can significantly reduce the toxicity of patulin [38]. With the prolonged reaction, hydroascladiol was further diminished (data are not shown), leading to further destruction of the lactone acid in hydroascladiol and lower toxicity. This is consistent with the observed decreased toxicity of the degradation products on *E. coli*.

**Figure 6.** Identification of hydroascladiol as one major transformation product of patulin. PAT was incubated with 0.5 U/mL of *Mr*MnP in 50 mM malonate buffer (pH 5.0) supplemented with 1 mM MnSO4 and 0.1 mM H2O2 and the reaction was carried out at 30 ◦C for 8 h. The degradation products were analyzed by HPLC–MS/MS.

#### *2.6. Degradation in a Simulated Patulin-Contaminated Apple Juice*

The ability of an enzyme to degrade a mycotoxin does not necessarily mean that the enzyme can efficiently degrade the mycotoxin in real foods/feeds. This is because the foods or feeds contain numerous components that could either adsorb the mycotoxins or act as competitors or inhibitors of the enzyme. For example, lignin phenolic compounds are naturally the substrate of MnPs [39–42]. In addition, many mycotoxins including aflatoxin B1, ochratoxin A, and zearalenone all have high affinity for lignocellulose, which also hinders the degradation process [43]. Therefore, to investigate whether *Mr*MnP can degrade patulin in the real environment, i.e., in foods with possibly interfering components, patulin was added to apple juice which then acted as a simulated patulin-contaminated beverage. In the control group, when no apple juice was added, rapid degradation of patulin was observed: the degradation rate was 76.8 ± 1.2% at 2 h of incubation and reached 100% after 5 h (Figure 7). When apple juice was present, the degradation rate of patulin was decreased to 19.8 ± 2.5% after incubation for 2 h. However, the degradation rate increased to 72.1 ± 4.3% after 5 h of incubation and then steadily increased to 95 ± 2.1% after 24 h. In the univariate analysis of 24 h data, there was a significant difference (*p* < 0.05). Therefore, it was evident that some components negatively affected degradation of patulin. However, the degradation rate of patulin was still highly comparable to that in absence of apple juice at 12 h and 24 h of incubation. Therefore, *Mr*MnP-catalyzed degradation of patulin can serve as an effective means to control the pollution of patulin in fruit juice.

**Figure 7.** *Mr*MnP efficiently catalyzed degradation of patulin in a simulated patulin-contaminated apple juice. The reactions were carried out by incubating 0.5 U/mL of *Mr*MnP with 5 mg/L in absence (or presence) of apple juice. The samples were periodically taken out for HPLC analysis. The data in the picture is the average ± standard deviation, \*\*\* *p* < 0.001, \*\* *p* < 0.01, \* *p* < 0.05, One Way anova test.

Manganese peroxidases are present in many different microorganisms, such as *Irpex lacteus* [44], *Rhizoctonia* sp. [45], *Stereum Ostrea* [46], and *Phanerochaete chrysosporium* [47]. Regardless of the source of manganese peroxidases, they can all catalyze the oxidation of Mn2+ to Mn3+, which can be stabilized by forming complexes with a specific chelator in the reaction system. Therefore, it is expected that manganese peroxidase from other sources can also degrade patulin, and these enzymes will be a rich resource of candidate MnPs with physiochemical properties satisfying the demands of practical applications.

#### **3. Conclusions**

*Mr*MnP can degrade patulin most rapidly in the malonate/Mn2+ system, with 0.5 U/mL enzyme completely removing 5 mg/L of pure patulin within 5 h. One major degradation intermediate was identified by mass spectrometry to be hydroascladiol. In a simulated patulin-contaminated apple juice, 95% of patulin was eliminated after a 24 h treatment. Use of the enzyme to eliminate patulin contamination in juice appears to be advantageous to other detoxifying strategies such as microbial cells. Importantly, as the degradation led to detoxification, *Mr*MnP, and perhaps other manganese peroxidases, may serve as candidates for enzymatic detoxification of patulin in foods and beverages. As MnPs are commonly fragile, and the expression level of *Mr*MnP is still not comparable to those of other proteins such as glycoside hydrolases, in future efforts should be made to improve the stability and expression level of MnPs to reduce the cost in juice detoxification.

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

#### *4.1. Strains and Plasmids*

The *Escherichia coli* Trans1-T1 (TransGen, Beijing, China) was used for gene cloning and plasmid propagation. The *E. coli* DH5α (Vazyme, Nanjing, China) was used to detect the residual toxicity of patulin after *Mr*MnP treatment. The yeast used for recombinant *Mr*MnP expression was the *Pichia pastoris* X-33 strain (Invitrogen, Carlsbad, CA, USA). The plasmid used for construction of the expression plasmid was pPICZα(A) (Invitrogen, Carlsbad, CA, USA).

#### *4.2. Cloning and Expression of MrMnP*

The coding sequence of the *Mr*MnP manganese peroxidase gene (GenBank accession number: ESK95360.1) from *M. roreri* was codon-optimized according to the codon bias of *Pichia pastoris* and synthesized by the GenScript Biotech Corp. (Nanjing, China). Then, the *Mr*MnP gene was amplified from the synthesized gene by gene specific primers MrMnP-F and MrMnP-R (MrMnP-F: 5 -GCGGAATTCGCTGTTCCACAAAGAGTTGCTT-3 , where the underlined sequence indicates the *Eco*RI restriction site; MrMnP-R: 5 - GCGGCGGCCGCAGATGGTGGAACAGCTGGAAC-3 , where the underlined sequence indicates the *Not*I restriction site). The PCR product was treated with *Eco*RI and *Not*I and ligated into the expression vector pPICZα(A) pre-digested with the same two restriction enzymes to generate the recombinant plasmid pPICZα(A)-MrMnP, which was transformed into *E. coli* Trans1-T1 for cloning and sequencing. The integrity of the recombinant plasmid was confirmed by DNA sequencing. Then, the *Dra*I-linearized plasmid was transformed into the *P. pastoris* X33 competent cells by electroporation. Clones were selected on YPDS agar-plates including 100 μg/mL of zeocin.

A single colony of the transformant bearing the *Mr*MnP gene was inoculated into 10 mL of the YPD (yeast potato dextrose) medium, cultured at 30 ◦C overnight with an agitation of 200 rpm, and then transferred to 50 mL BMGY medium (containing 1% yeast extract, 2% peptone, 1% glycerol, 1.34% YNB, 100 mM sodium phosphate buffer pH 6, 0.4 mg/L biotin). The culture was continued at 30 ◦C and 200 rpm until the optical density of the culture at 600 nm (OD600) reached 6.0. The cells were then collected by centrifugation and suspended in 50 mL BMMY medium (containing 1% yeast extract, 2% peptone, 1% methanol, 1.34% YNB, 100 mM sodium phosphate buffer pH 6.0, 0.4 mg/L biotin). Methanol was added every 24 h to a final concentration of 1% (*v*/*w*) and the induction of enzyme expression was continued at 30 ◦C for 5 d. The secreted crude *Mr*MnP enzyme was collected from the culture supernatant.

#### *4.3. Fed-Batch Fermentation of MrMnP in a Bioreactor*

The X33 transformant integrated with the *Mr*MnP gene was inoculated in 50 mL YPD and incubated at 30 ◦C for 48 h with shaking at 220 rpm. This "primary seed" culture was transferred into another three 200 mL fresh YPD medium (with a ratio of one tenth, *v*/*v*) and the culture was continued overnight, which served as the "secondary seed". This secondary seed culture was added into a bioreactor filled with 6 L of minimal salt medium, and the culture parameters were set with the pH to 4.0, temperature to 30 ◦C, and the rotation speed to 300 rpm. After the carbon source in the culture medium was exhausted, a mixture of glucose/methanol (glucose: 40%; glucose:methanol = 6:1) and 75 μM heme solution were added at a flow rate of 36 mL/(L h). The fermentation lasted for 120 h, during which the heme solution was replenished appropriately.

#### *4.4. Degradation of Patulin by MrMnP*

To determine the ability of *Mr*MnP to degrade patulin, the manganese peroxidase activity was first calibrated using 2,2 -azino-bis (3-ethylbenzothiazoline-6-sulfonic acid) (ABTS), which was carried out by monitoring the oxidation of ABTS (ε420 = 36,000 M−<sup>1</sup> cm<sup>−</sup>1) at 420 nm in a buffer containing 50 mM malonate, 1 mM ABTS, 1 mM MnSO4, and 0.1 mM H2O2 (pH 5.0 and 25 ◦C) as described in Qin et al. [48]. Then, *Mr*MnP with an activity of 0.5 U/mL against ABTS was incubated with 5 mg/L of patulin in 50 mM malonate buffer, 1 mM MnSO4, and 0.1 mM H2O2. The reaction was carried out at 30 ◦C. At the end of the reaction, 3 volumes of methanol were added to the mixture for termination and the reaction products were analyzed by HPLC.

#### *4.5. Effect of the Buffer Componesnts on Degradation of Patulin by MrMnP*

To determine the effect of the buffer components on the degradation of patulin, the malonate (50 mM) was replaced with one of the other buffers, which included acetate, lactate, citrate, oxalate, phosphate, MES, and HEPES. The pH of the buffer was adjusted to 5.0 in all tested reaction systems. The reaction was carried out at 30 ◦C for 24 h. The reaction products were analyzed by HPLC and the degradation rate of patulin was then determined.

#### *4.6. Effect of Mn2+ on Degradation of Patulin*

In order to study the effect of Mn2+ on the degradation of patulin, the degradation rate of patulin in the malonate and acetate systems with or without Mn2+ was tested. All the reactions were carried out at 30 ◦C, and the samples were periodically (2 h, 5 h, 8 h, and 24 h) taken out for analysis of the degradation rate of patulin.

#### *4.7. Toxicity Assay*

The *E. coli* DH5α was used as a microbial sensor system to determine if the *Mr*MnPcatalyzed degradation products of patulin would still have toxicity [34]. *E. coli* was cultured in a Luria–Bertani broth (LB: 1.0% tryptone, 0.5% yeast extract, and 1.0% sodium chloride) to an OD600 of 0.3. Then, 0.5 U/mL of *Mr*MnP was added to varying concentrations (0, 1, 10, 50, and 100 mg/L) of patulin and the reaction was incubated at 30 ◦C for 24 h. Then, 5 mL of *E. coli* cells were mixed with 1 mL of patulin treated with or without *Mr*MnP. The growth of *E. coli* was carried out at 37 ◦C in a 96-well microplate. At 0, 2, 4, 6, 8, and 10 h, the OD600 was measured as an indicator of the growth of *E. coli* and toxicity of patulin to the bacterium.

#### *4.8. HPLC and LC-MS/MS Analyses*

Degradation of patulin was analyzed by High-Performance Liquid Chromatography (HPLC), which was carried out using a SHIMADZU 20A series instrument (Kyoto, Japan) with an Agilent ZORBAX SB-C18 column (5 μm, 4.6 mm × 250 mm) (Santa Clara, CA, USA). The elution condition included the use of 10% acetonitrile as the mobile phase, and the flow rate was set to 0.75 mL/min. Patulin was monitored under ultraviolet light of wavelength 276 nm.

The degradation products of patulin were further analyzed by using LC-MS/MS, which was carried out by coupling a SHIMADZU Nexera UHPLC system (Kyoto, Japan) to an AB-SCIEX 5600+ Triple TOF mass spectrometer (Waltham, MA, USA). The chromatographic column was XBrige BHE C18 (2.5 μm, 2.1 mm × 150 mm) and the column temperature was 40 ◦C. The mobile phase A was acetonitrile and the mobile phase B was 0.1% formic acid. One μL of the sample was injected and the flow rate was 0.3 mL/min. The elution procedure was as follows: initial 10% phase A; 0.5 min, 10% phase A; 1.5 min, 50% phase A; 5.0 min, 90% phase A; 6.0 min, 90% phase A; 6.2 min, 10% phase A; 8.0 min, 10% phase A. The detection conditions of mass spectrometry were as follows: negative ion; TOF-mass (Da) 50–500; ion source: Duo Spray Ion Source; ion source gas 1:50; ion source gas 2:50; curtain gas: 25; temperature: 450, IonSpray voltage floating (ISVF): −4500; declustering potential: −57.0; collision energy: −12.0; accumulation time: 0.1 s. ion scanning

conditions: declustering potential: −80.0; collision energy: −30.0; collision energy spread: 0; ion release delay: 67; ion release width: 25.

#### *4.9. MrMnP-Catalyzed Degradation of Patulin in a Simulated Patulin-Contaminated Apple Juice*

The commercially available fresh apple juice (Huiyuan, Beijing, China) was obtained from a local retail market. No patulin was detected in this juice by HPLC analysis. Patulin was added to this product to a final concentration of 5 mg/L to simulate mycotoxincontaminated juice. Then, the reaction was carried out by adding a final concentration of 0.5 U/mL of *Mr*MnP to the juice in the presence of 50 mM of malonic acid solution and 1 mM of MnSO4. The reaction was initiated by addition of 0.1 mM of H2O2. At 0, 2, 5, 8, and 24 h, samples were taken out for HPLC analysis.

#### *4.10. Statistical Analysis*

The results of three repetitions were expressed as mean ± standard deviation (SD). The data were analyzed by one-way analysis of variance (ANOVA).

**Author Contributions:** S.W. conducted experiments and wrote the manuscript. X.W., L.P., H.L., Y.Z., B.L., B.Y. and N.H. conducted experiments. W.Z. and X.S. conceived and designed the research. All authors contributed to data analyses, results interpretation. All authors have read and agreed to the published version of the manuscript.

**Funding:** This study was supported by the National Key R&D Program of China (No. 2021YFA0910602, 2021YFC2103002 and 2021YFC2102400) and the National Chicken Industry Technology System of China (CARS-41).

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

**Informed Consent Statement:** Not applicable.

**Data Availability Statement:** The data presented in this study are available on request from the corresponding author.

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

#### **Abbreviations**


#### **References**


### *Article* **Sustainable Strategies to Counteract Mycotoxins Contamination and Cowpea Weevil in Chickpea Seeds during Post-Harvest**

**Claudia Pisuttu 1, Samuele Risoli 1,2, Lorenzo Moncini 3, Cristina Nali 1,4, Elisa Pellegrini 1,4,\* and Sabrina Sarrocco 1,4**


**Abstract:** Mycotoxins contamination and pest infestation of foods and feeds represent a pivotal threat for food safety and security worldwide, with crucial implications for human and animal health. Controlled atmosphere could be a sustainable strategy to reduce mycotoxins content and counteract the vitality of deleterious organisms in foodstuff. Ozone treatment (O3, 500 ppb for 30, 60 or 90 min) and high nitrogen concentration (N2, 99% for 21 consecutive days) were tested in the post-harvest management of four batches of *Cicer arietinum* grains to control the presence of mycotoxigenic fungi and their secondary metabolites, as well as pest (i.e., *Callosobruchus maculatus*) infestation. At the end of the treatment, O3 significantly decreased the incidence of *Penicillium* spp. (by an average of −50%, independently to the time of exposure) and reduced the patulin and aflatoxins content after 30 min (−85 and −100%, respectively). High N2 concentrations remarkably reduced mycotoxins contamination (by an average of −94%) and induced pest mortality (at 100% after 5 days of exposure). These results confirm the promising potential of O3 and N2 in post-harvest conservation strategies, leading to further investigations to evaluate the effects on the qualitative characteristics of grains.

**Keywords:** *Cicer arietinum* L.; mycotoxigenic fungi; mycotoxin occurrence; pest attack; nitrogencontrolled atmosphere; ozone

**Key Contribution:** The findings of the present study are relevant in enhancing the shelf-life of chickpea seeds by controlling fungal growth, mycotoxin contamination and pest infestation with eco-friendly and low-cost storage practices such as ozone treatment and high nitrogen concentrations.

### **1. Introduction**

The increasing occurrence of food/feed contaminants worldwide poses a huge threat to human and animal health. One of the major contaminants are mycotoxins, which annually cause enormous economic losses in the food industry and animal husbandry [1,2]. These low molecular weight metabolites produced by filamentous fungi (belonging to the phylum Ascomycota) contaminate various categories of foods and feeds [3]. Two groups of mycotoxigenic fungi exist: field fungi (such as *Fusarium* and *Aspergillus* spp.) that infect crops before harvest, and storage fungi (such as *Penicillium* spp.), which only occur after harvest [4,5]. According to a recent world survey based on around 97,000 analyses performed between January and December 2020 on more than 21,000 finished feed and raw commodity sources collected from 79 countries, the most prevalent mycotoxins were those produced by *Fusarium*, affecting more than the 60% of tested samples [Biomin, Inzersdorf-Getzersdorf, Austria, https://www.biomin.net/science-hub/world-mycotoxin-survey-impact-2021/, accessed on 4 January 2023], In 2020, Mesterhazy et al. highlighted how toxins are responsible for a loss of almost 700 mt during the harvest and storage of grains [6]. At any stage

**Citation:** Pisuttu, C.; Risoli, S.; Moncini, L.; Nali, C.; Pellegrini, E.; Sarrocco, S. Sustainable Strategies to Counteract Mycotoxins Contamination and Cowpea Weevil in Chickpea Seeds during Post-Harvest. *Toxins* **2023**, *15*, 61. https:// doi.org/10.3390/toxins15010061

Received: 21 December 2022 Revised: 5 January 2023 Accepted: 7 January 2023 Published: 11 January 2023

**Copyright:** © 2023 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/).

of the food production process (in the field, during harvest, during drying and transport, as well as during storage), the fungal production of mycotoxins can occur by exposing consumers to the risk of contamination, either directly through food consumption or indirectly through feed [7]. The most important mycotoxins are aflatoxins [mainly represented by aflatoxin B1 (AFB1), B2 (AFB2), G1 (AFG1), G2 (AFG2) and M1 (AFM1)], ochratoxins, fumonisins, trichothecenes, zearalenone, the emerging *Fusarium* mycotoxins, ergot alkaloids, *Alternaria* toxins, and patulin [8]. Of the approximately 400 compounds identified as mycotoxins, 30 have received significant consideration due to their harmful effects on both human and animal health (including genotoxicity and endocrine disruption [9]). Despite a huge number of published papers reporting the occurrence of mycotoxins on cereals and cereal-derived food products, in 2017, an analysis of 104 papers—from 2006 to 2016—was carried out, summarizing that mycotoxins are ubiquitously present in cereals and cereal-derived food products throughout the world [10]. If Africa and Asia showed the highest incidence (%) of cereals contaminated by aflatoxins and ochratoxins, respectively, South and North America registered the highest level of fumonisins and Europe the highest percentage of deoxynivalenol (trichothecenes)/zearalenone contamination [10].

Different physical, chemical and biological factors affect fungal colonization and mycotoxins production. Physical factors include environmental conditions such as temperature, relative humidity, pH, water activity, nutrients, insect infestation and other associated factors, which at specific rates are known to favor the growth of many types of fungi and the production of mycotoxins [2]. Biological factors are mainly related to the interactions between the colonizing toxic fungal species and the host, thus including features such as fungal species, strain specificity, levels of inoculation, the nature of the substrate, strain variation, the instability of fungal toxic properties, and insect damage [11].

Regulatory agencies have established strict legislative thresholds in order to keep the levels of mycotoxins in food/feed commodities under control. These limits range from below one to thousands of μg kg−1, depending on the (i) mycotoxin, (ii) type of product and (iii) country considered [12,13]. Consequently, there is an urgent need to develop a feasible and highly sensitive analytical method for mycotoxins detection [14] and reduce the contamination of mycotoxins in food/feed, in order to protect/preserve their quality and safety. Overall, the control of mycotoxin contamination follows two strategies: the prevention of their production (i.e., microbial inactivation) and detoxification (e.g., mycotoxin degradation [3]). In pre-harvest, the control of mycotoxins is based on control of the contamination levels in crops to be used as food/feed components. Generally, these systems are based on preventive strategies (such as the use of resistant varieties, crop rotation, tillage and the management of irrigation) which aim to avoid the development of contamination, operating on the predisposing factors that facilitate the production of mycotoxins. Although pre-harvest approaches should be preferred, in the perspective of preventing mycotoxin contamination, the development of toxic fungi is inevitable under certain environmental conditions [15]. Therefore, appropriate storage practices and other post-harvest control systems (at the microbiological, physical and chemical level) are necessary to minimize the final mycotoxin content of foods/feeds [16]. These traditional methods for the elimination/inactivation of mycotoxins have some limitations regarding (potential) safety issues, loss in the nutritional value and the palatability of feeds, cost implication and limited efficacy [17]. In recent decades, various detoxification approaches have demonstrated to be (i) highly effective in degrading mycotoxins into less toxic products, (ii) economically favorable and (iii) not environmentally harmful [18]. Among these, cold plasma—containing reactive oxygen and nitrogen species and free radicals—has received attention in recent years for use on cereals during storage, due to its lethal effect on microorganisms and its potential to decontaminate surfaces and improve shelf-life [19]. Nevertheless, their practical application in food/feed matrices is limited, since the degradation process under conditions of large-scale production is much more complex, and the experiments at lab-scale might not always reflect practices in industrial processing [1]. Possible reasons for this are that the degradation process can be easily

affected by multiple factors such as temperature, relative humidity, pH, water activity, nutrients, insect infestation and types of contamination [2]. The relevance of studying naturally contaminated samples is consistent with the actual distribution conditions of mycotoxins in the field and/or in grains.

Since gas composition is considered one of the most important abiotic conditions that impacts fungal and pest growth, ozonation (i.e., the application of gaseous ozone, O3) is a simple technology for controlling insects and reducing mycotoxins in stored products, which does not leave harmful residues after application. Being unstable, O3 quickly degrades into oxygen (and related cytotoxic radicals) in a short period, oxidizing the vital cellular components (such as unsaturated lipids and proteins) of pathogenic microbes and storage pests by causing lysis and rapid cell death [20]. Consequently, O3 can inhibit fungal growth, sporulation and germination by offering a negligible loss of nutrients or sensory qualities in food/feed [21], making it a suitable candidate as a residue-free fumigant. For this reason, the application of O3 in food chains has been considered safe and effective by the WHO and is now recognized as a "green technology" for the fumigation of grains, fruits and vegetables [22]. In fact, O3-treated products are safe for consumption and their microbiological shelf life can be greatly enhanced. However, the efficacy of O3 in fungi count reductions, mycotoxin degradation and insect control depend on the (i) method, concentration and timing of the O3 application; (ii) microorganisms/contaminants to inactivate; (iii) the type and mass of food/feed processed; and (iv) other co-factors such as temperature, relative humidity and water activity [16,20]. Similarly, the use of a controlled or modified atmosphere by using a very high nitrogen (N2) concentration is a valid alternative to chemical fumigation to control mycotoxigenic fungi contamination and pest challenge post-harvest [23]. Its effects on different stored products (such as wheat, maize, corn and rye) are well documented [2,24]. In particular, a N2-controlled atmosphere can control fungal growth and proliferation by improving the quality of stored products [25]. The action of N2 at high concentration is mainly due to the significant reduction in O2 (1% or less [26]) and offers several advantages at the economic and environmental level [27]. A major advantage of N2 is that all gas is free of pollutants, leaving no residue in food/feed. Consequently, N2-treated products are safe for consumption and their microbiological shelf-life can be greatly enhanced [28]. Consequently, a N2-controlled atmosphere might represent an eco-friendly tool that could be transferred to a large-scale system for grain storage as an alternative strategy to the use of conventional residue-producing chemical fumigants [29]. However, the efficacy of a N2-controlled atmosphere in fungi count reductions, mycotoxin degradation and insect control depends on the (i) concentration of gas, (ii) the timing of the application, (iii) microorganisms/contaminants to inactive, (iv) the type and mass of food/feed processed, and (v) other co-factors such as temperature and water activity [30].

The chickpea (*Cicer arietinum* L.) is a legume of the family Fabaceae, subfamily Faboideae. It is one of the most cultivated pulses in terms of world production due to its low content of fat and sodium, absence of cholesterol and being an excellent source of both soluble and insoluble fiber, complex carbohydrates, vitamins, folate and minerals (such as calcium, phosphorus, iron and magnesium [31]). With a worldwide production of more than 12 million tons per year [FAOSTAT, https://www.atlasbig.com/en-in/countriesby-chickpea-production, accessed on 4 January 2023], chickpea represents one of the five leading pulses based on sales value. In 2021, India was the largest chickpea producer in the world, with around 11 million metric tons of production, followed by Turkey, with around 600,000 metric tons (https://www.statista.com/statistics/722203/chickpeas-productionvolume-by-country-worldwide/, accessed on 4 January 2023). Chickpea is often attacked by fungi pre- and post-harvest, significantly affecting its productivity. Many fungal genera/species commonly isolated from chickpea seeds and chickpea by-products are potential mycotoxin-producers, especially of aflatoxins, ochratoxin A and patulin, so there would be a potential risk of contamination [32]. Another issue threatening chickpea quality is *Callosobruchus maculatus* (Fab.) (Coleoptera: Chrysomelidae: Bruchinae [33]), which is also known

as the "cowpea weevil". The granivorous larvae of cowpea weevil are the considerable causative agent of severe losses in the grain germination, weight and nutritional values of chickpea (in some cases reaching 60% of the grain [34]). In addition, *C. maculatus* can favor the occurrence of infections due to mycotoxigenic fungi (*Aspergillus* and *Penicillium* [35]).

The aim of this work was to investigate the possibility of using a single pulse of O3 or high N2 concentrations as storage technologies (at lab- and large-scale) for the purpose of (i) containing the fungal population present on the chickpea seeds surface; (ii) reducing the mycotoxins content (such as aflatoxins and patulin; Figure 1); and (iii) limiting the *C. maculatus* infestations (only in the case of N2 treatment). We postulated that O3 and N2 can be an alternative to traditional chemical-based fumigants for controlling spoilage pathogens and insects in stored chickpea seeds.

**Figure 1.** Molecular structures of measured mycotoxins (patulin, aflatoxin B1, aflatoxin B2, aflatoxin G1 and aflatoxin G2; modified by [3]), showing numbering of carbon atoms (3a and 7a represent small fragments).

#### **2. Results**

#### *2.1. Effect of O3 and N2 Treatments on Fungal Infection*

To evaluate the effect of O3 treatment as well as of the conservation under a N2 controlled atmosphere on a fungal population naturally occurring on chickpea seeds, with particular attention to mycotoxigenic fungi, a grain health test was performed on control stocks stored at 4 ◦C and on seeds after O3 or N2 treatments. After the morphological and molecular identification of single colonies isolated from seeds, all the batches were contaminated with *Penicillium* spp. isolates, but at a different extent depending on the quality of the batches. Molecular identification performed on the ITS region sequence confirmed the membership of all the isolates to this important fungal genus, with the species *P. pinophilum* and *P. polonicum* as the most represented.

Of particular interest was batch n. 2, where almost 80% of the analyzed seeds were infected by *Penicillium* isolates, thus, confirming the non-marketability of this batch for commercial purposes. The other three batches showed a percentage of natural occurring infection varying around 10–20% (Figure 2).

**Figure 2.** Contamination levels (% of infected seeds) by *Penicillium* spp. in four batches (n. 1 (**a**), n. 2 (**b**), n. 3 (**c**) and n. 4 (**d**)) of chickpea seeds (CTRL, white fill) exposed to ozone [500 ppb O3 for 30 (O3 30', light grey fill), 60 (O3 60', grey fill) and 90 (O3 90',dark grey fill) minutes] or nitrogen treatment (99% N2 for 21 consecutive days, dark fill). Data are shown as mean + standard deviation (n=3). Results of two-way ANOVA are reported; asterisks show the significance of factors/interaction for: \*\*\* *p* ≤ 0.001. According to Tukey's HSD post hoc test, different letters indicate significant differences (*p* ≤ 0.05).

After O3-exposure, as well as after incubation under the N2-controlled atmosphere, the grain health test, in most cases, resulted in a reduction in the *Penicillium* contamination of the seeds. A significant effect of single factors and their interaction was observed. For batch n. 1, O3 treatment did not result in any significant reduction if compared to the control, as well as among treatments. Conversely, a significant reduction was registered after N2 incubation (−16% of infected grains compared with no seed developing *Penicillium* spp. colonies; Figure 2a). Much more evident was the effect of the O3 treatment on batch n. 2, where, independently from the time of exposure, a significant reduction in the naturally occurring *Penicillium* population was observed (−55% as average), with the exposure lasting 90 min being the most efficient (−75%). On the other hand, any significant difference was observed after the incubation in the N2-controlled atmosphere (Figure 2b). For batches n. 3 and 4, a similar behavior was observed, where any significant difference in the percentages of infected seeds was observed if compared with the controls, as well as among treatments (Figure 2c,d). However, in these last two cases, the initial infection was lower if compared with the other two batches, particularly with batch n. 2.

#### *2.2. Effects of O3 and N2 Treatments on Mycotoxin Levels*

In the four batches of chickpea grains maintained in filtered air or incubated in control silos, the mycotoxins patulin, AFG2, AFB2, AFG1 and AFB1 were found. Figure 3 shows an overview of the concentrations of these mycotoxins and their variability among the batches.

**Figure 3.** Box and whiskers representation of the average content of measured mycotoxins in four batches of chickpea seeds ((**a**) patulin and aflatoxin G2 (AFG2); (**b**) aflatoxin B2 (AFB2), aflatoxin G1 (AFG1) and aflatoxin B1 (AFB1)). For each mycotoxin, the top line represents the 90th percentile; the bottom line represents the 10th percentile; and the box represents the 75th percentile (upper side), the 25th percentile (lower side) and the median (50th percentile, central line), respectively.

The comparison between the content of selected mycotoxins and the batches of chickpea grains maintained in filtered air or incubated in control silos revealed that the concentrations of patulin were significantly higher in batch n. 4 than those in batch n. 1 (+55%; Table S1). No significant differences were observed among the remaining batches of chickpea grains. The levels of AFB1, AFB2 and AFG1 were significantly higher in batch n. 3 than those in the remaining batches of chickpea grains (about six-fold higher on average). Conversely, the levels of AFG2 were significantly higher in batch n. 2 than those in the remaining batches of chickpea grains (about 100-fold higher than batch n. 1, and five-fold higher than batches n. 2 and 3; Table S1).

The effects of the O3 and N2 treatments on the patulin levels in the four batches' grains are reported in Figure 4. The two-way ANOVA test revealed that the interaction "batch of chickpea grains × treatment" and the effects of each factor were significant. Ozone treatment induced a significant decrease in patulin in batch n. 2, independently of its duration (an average of −50% compared to chickpea grains maintained in filtered air; Figure 4b), and even more in the remaining batches (about 150-, 230- and 240-fold lower than CTRL in batches n. 1, 3 and 4, respectively). Similarly, a reduction in patulin was observed in batches n. 3 and 4 incubated in silos under high N2 concentrations (−70 and −82% than CTRL, respectively; Figure 4c,d), and even more in the remaining batches (about 160-fold lower than those in batches n. 1 and 2 incubated in control silos, respectively).

**Figure 4.** Patulin content in four batches (n. 1 (**a**), n. 2 (**b**), n. 3 (**c**) and n. 4 (**d**)) of chickpea seeds (CTRL, white fill) exposed to ozone (500 ppb O3 for 30 [O3 30', light grey fill), 60 (O3 60', grey fill) and 90 (O3 90', dark grey fill) minutes] or nitrogen treatment (99% N2 for 21 consecutive days, dark fill). Data are shown as mean + standard deviation (n = 3). Results of two-way ANOVA are reported; asterisks show the significance of factors/interaction for: \*\*\* *p* ≤ 0.001. According to Tukey's HSD post hoc test, different letters indicate significant differences (*p* ≤ 0.05).

The effects of the O3 and N2 treatments on the total aflatoxin levels in four batches of chickpea grains are reported in Figure S1. The two-way ANOVA test revealed that the interaction "batch of chickpea grains × treatment" and the effects of each factor were significant. Ozone treatment induced a complete reduction in total aflatoxins in batches n. 2, 3 and 4, independently of its duration (about 80-fold lower compared to chickpea grains maintained in filtered air). Similarly, a reduction in total aflatoxins was observed in batches n. 2, 3 and 4 incubated in silos under high N2 concentrations (about 150-fold on average). No other significant differences were found in batch n. 1, independently of the kind of treatment.

#### *2.3. Effect of N2 Treatment on Pest Survival*

The effect of 5-day exposure in the N2-controlled atmosphere (in the 60 L lab-scale silos) on the number of emerged adults and on Abbott's index is reported in Figure 5.

A statistical analysis highlighted how not only the treatment and the time had a significant effect on the adult emergence, but also the interaction between these two sources of variability resulted as highly significant (*p* ≤ 0.001). The exposure to the N2-controlled atmosphere resulted in a reduction in the number of emerged adults from the first day of exposure. No significant differences were observed after 24 h of exposure. From the second until the fifth day, a continuous and significant reduction was observed, with the adult emergence close to zero after five days of exposure of the eggs within the silos under the N2-controlled atmosphere.

With respect to Abbott's index, which is conventionally used to evaluate the effect of treatments on pest mortality compared to naturally occurring mortality, the incubation of eggs under high N2 concentration resulted in an increasing mortality over time of exposure, with 100% reached at the end of the experiment, i.e., after 5 days of exposure (Figure 5, dashed line).

**Figure 5.** Number of emerged adults of *Callosobruchus maculatus* and mortality (measured using Abbott's index, dotted line and dark triangle) on batch n. 1 (CTRL, open circle) and exposed to nitrogen treatment (60 L lab-scale; dark circle). Data are shown as mean ± standard deviation (n = 3). Results of two-way ANOVA are reported; asterisks show the significance of factors/interaction for: \*\*\* *p* ≤ 0.001. According to Tukey's HSD post hoc test, different letters indicate significant differences (*p* ≤ 0.05).

When the effect of the high N2 atmosphere was evaluated on *C. maculatus* adult number in real-scale silos after 5 days of exposure (Figure 6), a highly significant reduction in emergence was registered, compared to the control. This result was also confirmed by Abbott's index, which resulted in approximately 80% of mortality due to the N2-controlled atmosphere.

**Figure 6.** Number of emerged adults of *Callosobruchus maculatus* (Fab.) and mortality (measured using Abbott's index, dotted line) on batch n. 1 (CTRL, white fill) and exposed to nitrogen (N2) treatment (real-scale; gray fill). Statistical differences were examined by paired Student's *t*-test; asterisks indicate statistical significance for: \*\*\* = *p* ≤ 0.001.

#### **3. Discussion**

In the present study, the fungicidal efficacy of O3 was only observed in batch n. 2 (the most infected seeds), as confirmed by the large reduction in *Penicillium* contamination. According to Mendez et al. [36], at the beginning of O3 treatment, this gas reacts with a mass of grains and quickly decomposes. In its second phase, O3 moves freely through the grains with little degradation. Moreover, O3 reacts faster with the whole mass of grains when higher dosages are used. This was also confirmed by our results: a higher reduction in the naturally occurring fungal population was observed by using O3 at 500 ppb for 90 min. To date, few studies have investigated the direct effect of O3 on fungal growth and proliferation, preferring to focus on the reduction in mycotoxins in different products (such as wheat, corn, corn flour, peanuts and pistachio [37]). Most studies have been carried out on aflatoxins. Using O3 in patulin, degradation has only been studied in apple juice, apples and pear with brown rot, flour, and malt feed [38]. In the present study, O3 treatment induced a significant decrease in patulin in batch n. 2 (independently of its duration), and even more in the remaining batches. The mechanism of patulin degradation might be associated with the oxidation of a polyketide lactone on its structure, which made it highly susceptible to O3 attack [39]. The experimental conditions used in this study (e.g., O3 and/or the timing of treatment) were sufficient, by attacking two conjugated ethylenic double bonds on the chemical structure of patulin and inducing its partial or full degradation [40]. Similarly, all aflatoxins were easily attacked and degraded by O3 (independently of its duration), confirming its efficacy as a detoxifying agent. The mechanism of AFB1 and AFG1 degradation might be associated with the oxidation of the C8-C9 double bond at the terminal furan, resulting in the production of primary ozonide [26]. This product may rapidly rearrange to a molozonide derivative, yielding a variety of carbonyl compounds or organic acids. Since AFB2 and AFG2 lack a susceptible double bond for oxidation, their degradation requires higher levels of O3 and/or longer exposure until the lactone ring is opened [41]. The experimental conditions used in this study (e.g., O3 concentration and/or the timing of treatment) were sufficient, by rapidly and effectively detoxifying aflatoxins without any difference in degradation rate between AFB1 and AFG1 with AFB2 and AFG2 (the most abundant aflatoxin in batches n. 2, 3 and 4). It is worth noting that the production of aflatoxins was not associated with the presence of the fungal itself, confirming that the absence of *Aspergillus* spp. from chickpea seeds does not guarantee the absence of aflatoxins because of their resistant chemical nature [42].

In the present study, high N2 concentrations only induced a reduction in *Penicillium* contamination in batch n. 1 confirming that certain fungal species might continue to grow, albeit at a greatly reduced rate under low O2 concentrations [43]. However, a significant decrease in patulin was observed in batches n. 3 and 4, and even more in the remaining ones, indicating that low O2 concentrations may (partially or totally) depress patulin production by *Penicillium* spp. on chickpea seeds. In addition, the experimental conditions used in this study (e.g., N2 concentration and/or the timing of treatment, temperature, relative humidity and water activity) were sufficient, by totally removing aflatoxins. Consequently, it is possible to speculate that high N2 concentrations are more effective in inhibiting selected mycotoxins (aflatoxins > patulin) than in preventing the development of mycotoxigenic fungi [44]. Although a controlled atmosphere is used to control both mycotoxigenic fungi and insects in stored products, it has been documented that the experimental conditions sufficient for controlling fungal proliferation are not always effective against insect pests that can survive, due to the dependence on other environmental factors (e.g., temperature and humidity [45]). This is in line with the results of the present work, where the same experimental conditions (e.g., N2 concentration and/or the timing of treatment, temperature, relative humidity and water activity) that were partially effective for reducing the growth of *Penicillium* spp. were effective for detoxifying mycotoxins (as previously reported) and limiting *C. maculatus* infestation (as confirmed by the reduction in adult emergence already observed starting from the first day of the experiment, and the concomitant pest mortality). This is of relevance, since it is generally

recognized that pest attack can damage grains and favor moisture accumulation by creating suitable conditions for fungal development and mycotoxin production (as observed in batch n. 2 [46]). Consequently, it is possible to speculate that the experimental conditions used in this study were sufficient, by limiting the pest infestation and guaranteeing seed quality, as already reported in wheat [23,25]. In addition, the possibility of moving to a realscale dimension, as here preliminary reported, with results comparable with those obtained in the 60 L lab-scale silos, marks a further step made in the direction of the scaling-up of the method. In fact, the set-up chosen for the present study is not only a suitable way to provide a proof-of-concept of its efficacy before scale-up, but also a valid choice for small and medium farms [23,25].

The findings of the present study are relevant in enhancing the shelf-life of chickpea seeds by controlling fungal growth, mycotoxin contamination and pest infestation with eco-friendly and low-cost storage practices (the mechanisms of action of O3 and N2 are summarized in Figure 7).

**Figure 7.** Gaseous ozone (O3) and nitrogen (N2) mechanisms of action during post-harvest of grains, and their effects on (i) mycotoxigenic fungi and storage pests (only N2), (ii) patulin and (iii) aflatoxins.

In particular, O3 can contain the fungal population present on the chickpea seeds surface and reduce the content of patulin and aflatoxins. This is a fundamental goal in the development of emerging new techniques, since these metabolites are recognized as a Group I carcinogen by the International Agency of Research on Cancer (Lyon, France), and their allowable levels in human foods and animal feedstuff are strictly regulated by governmental jurisdictions in about 100 countries [47]. It is worth noting that the antimicrobial activity of O3 is highly dependent on vegetable/fungus species, growth stage, concentration and timing of exposure. Improvements and innovations in O3 generation and application systems will be evaluated more effectively in the future by facilitating the enhanced control of both the quality and safety parameters of ozonized foods/feeds. Similarly, a N2-controlled atmosphere represents a valid sustainable method to limit mycotoxin accumulation and *C. maculatus* infestation. This system requires low energetic costs, offers a negligible loss of nutrients or sensory qualities in food/feed, and demonstrates a reduced hazard to employees with no need for registration and no contamination of the environment [23,25]. Consequently, it can be considered as a promising alternative method that could be transferred to a large-scale grain storage system. For effective and safe use in processing, optimum O3 and N2 concentrations, contact time and other treatment conditions should be defined for foods and feeds. Here, a pilot test was conducted by offering scientific evidence to support the commercial application of these innovative strategies.

#### **4. Conclusions**

In conclusion, our pioneering study demonstrated that the experimental conditions used (e.g., O3 and N2 concentrations and/or the timing of treatment) were enough to rapidly and effectively (i) reduce the growth and proliferation of *Penicillium* spp. population present on the chickpea seeds' surface (in the case of O3 treatment); (ii) detoxify patulin and aflatoxins; and (iii) limit *C. maculatus* infestation (only in the case of N2 high concentrations). These are fundamental goals in the development of emerging new techniques and novel methods to control mycotoxin infections, intoxications and diseases, by offering a negligible loss of nutrients or sensory qualities in stored products and without leaving toxic chemical residues in the food and feed chain.

Therefore, our results suggest that a single pulse of O3 and high N2 concentrations are a promising alternative to traditional chemical-based fumigants for controlling spoilage pathogens and insects in stored chickpea seeds. An industrial facility of O3 technology remains to be developed for the large-scale treatment of food/feed products, requiring input from different disciplines. For effective and safe use in processing, optimum O3 and N2 concentrations, contact time and other treatment conditions (e.g., vegetable/fungus species) should be defined for foods and feeds.

Additional research is obviously required to evaluate the responses of stored products to these effective and straightforward solutions, in order to control mycotoxin contamination and pest infestation at post-harvest. This would provide a clearer picture of the practical advantages of O3 and N2 treatment and allow further endorsement of our present results.

#### **5. Materials and Methods**

#### *5.1. Reagents and Standards*

Sodium hypochlorite, ethanol and streptomycin sulphate were supplied by Sigma-Aldrich (Milan, Italy). Potato Dextrose Agar was purchased from Biolife (Milan, Italy). Acetonitrile, methanol and water were HPLC-grade (Carlo Erba, Milan, Italy). Standards of patulin and aflatoxins were chromatographically pure and purchased from Sigma-Aldrich (Milan, Italy) and Romer Lab (Getzersdorf, Austria).

#### *5.2. Raw Materials*

Four *C. arietinum* batches, produced by a local farm located in Tuscany (Italy), were used in the present work. Before commercialization, all the batches (except in the case of n. 3) were submitted to a quality check by the producer, thus resulting in batches n. 1 and n. 4 being validated for sale, and batch n. 2 rendered non-compliant. All seeds were stored at 4 ◦C until submitted to a grain health test, mycotoxin determination and treatments under a controlled atmosphere.

#### *5.3. Isolation and Identification of Fungal Contaminants Naturally Associated with Chickpea Seeds*

To assess the presence of potential mycotoxigenic fungi naturally associated with chickpea seeds, all four batches were submitted to a grain health test. In detail, seeds from each batch were surface sterilized on a rotary shaker for 1 min in a solution containing NaClO (1% active chlorine) in 50% ethanol, then washed three times in sterile distilled water for 1 min each. After drying on filter paper, the seeds were transferred to 100 mm diameter Petri dishes containing Potato Dextrose Agar (PDA, 42 g L−1) with the addition of 300 mg L−<sup>1</sup> of streptomycin sulphate. Since batch n. 2 showed a profuse development of *Mucor* spp., the seeds from this sample were plated on PDA that contained the antibiotic, as previously described, and with hymexazol fungicide (at the final concentration of 300 mg L<sup>−</sup>1; [48]). For each batch, four replicates (each consisting of twenty-five seeds) were made. The plates were incubated at room temperature (24 ± 2 ◦C). Then, from the

second to the tenth day of incubation, colonies morphologically attributable to *Aspergillus* and *Penicillium* spp., developing from seeds, were transferred to new PDA + streptomycin sulphate plates and incubated under the same conditions previously reported. Then, when sporulated, they were used for single-spore cultivation. The single-spore *Penicillium* spp. colonies were used for molecular identification. Genomic DNA was extracted from each single-spore culture according to the Chelex 100 method [49]. For molecular identification, the complete internal transcribed spacers (ITS) 1 and 2 sequences—including the 5.8S gene—of the nuclear ribosomal DNA were amplified and sequenced as described in Sarrocco et al. [30]. All the sequences were then submitted to GeneBank (NCBI) to assign, where possible, the species for a preliminary evaluation of the risk of mycotoxin contamination that could occur on the seeds.

#### *5.4. O3 and N2 Treatment Systems for Chickpea Seeds*

#### 5.4.1. O3 Treatment System for Chickpea Seeds

Chickpea seeds (300 g from each batch) were placed in two Perspex chambers (60 × 60 × 110 cm) in a controlled environment fumigation facility. The system was adapted by including commercial colanders (34 × 23 cm, stainless steel), collocated in the middle of the chambers, in which seeds were placed to allow their complete exposure to O3, maintained in the dark throughout the whole period of the experiment (temperature 25 ± 1 ◦C, relative humidity (RH) 50 ± 5%). The fumigation system was continuously ventilated (two complete air changes per min) with charcoal-filtered air. Fumigation was performed by generating O3 from pure oxygen by electrical discharge, using a Fisher 500 air-cooled apparatus (Fisher America Inc., Houston, TX, USA). The O3 concentration was monitored with a Serinus 10 analyzer (Ecotech Acoem Group, Milan, Italy) set at <sup>500</sup> ± 50 ppb of O3 (for O3, 1 ppb = 1.96 <sup>μ</sup>g m−3, at 20 ◦C and 101.325 kPa) for 30, 60 and 90 min, in which the chickpea seeds were mixed every 15 min. At the end of each treatment, the samples were collected and immediately used for the subsequent analyses (fungal infection and mycotoxin contamination). The entire methodology was performed according to Marchica et al. [50].

#### 5.4.2. Nitrogen Treatment System for Chickpea Seeds

Nitrogen treatments were performed using a NitrosepAgri system (Eurosider sas, Grosseto, Italy) based on selective membrane (MNS, Membrane Nitrogen Separator) to separate N2 from atmospheric air [25]. The N2-enriched atmosphere was driven into silos where it was maintained under a slight overpressure. All the environmental parameters, such as temperature and RH, were constantly monitored, and the N2 percentage could be set, automatically maintained and quickly reintegrated if needed. To perform the experiments here reported, two lab-scale (20 and 60 L silos, respectively) prototypes, already described in Moncini et al. [25], and one field-scale apparatus were used. For each chickpea batch, 300 g of seeds were transferred into a 1.5 L glass jar (9 cm diameter) that was closed with a micro-perforated nylon layer (350 μm pore size) to facilitate gas exchange and incubated under a 99% N2 atmosphere for 21 consecutive days. For each treatment and for each batch, three replicates were made. The field-scale apparatus consisted of four 15 m3 volume fiberglass silos connected to the MNS system. Each silos had a stainless steel (786 mm diameter) top hatch for grains charging, equipped with an overpressure valve and, on the bottom, with a stainless-steel ball (90 mm diameter) valve for discharging the product. The field-scale apparatus is located at the Azienda Agraria Macchiascandona (Castiglione della Pescaia, Grosseto, Italy).

#### *5.5. Effect of O3 and N2 Treatments on Fungal Infection, Mycotoxin Contamination and Pest Survival*

At the end of the O3 and N2 treatments, the seeds were collected and used to evaluate fungal contamination according to the grain health protocol. Mycotoxins determination was performed by using the clean-up aflatoxins and patulin (AFP) columns (OR SELL,

S.p.a., Modena, Italy), according to the manufacturing protocols, with a few modifications. The samples were extracted by adding 50 mL of acetonitrile:water (ACN:H2O, 84:16 *v*/*v*) solution to 25 g of finely ground chickpea seeds, and vigorously vortexed for 6 min. The samples were centrifuged for 10 min at 12,000× *g* at room temperature, and the supernatants were filtered through Whatman® paper (Cytiva, Marlborough, MA, USA) and subsequently by using the clean-up AFP columns, which enable the contemporary purification of aflatoxins and patulin. The obtained solutions were equally separated, dried at 40 ◦C and finally resuspended in 400 μL of 45% methanol (*v*/*v*) or 75% ACN (*v*/*v* in HPLC-demineralized water) for aflatoxins and patulin separation, respectively. The separation was performed in a UHPLC Dionex UltiMate 3000 system (Thermo Scientific, Waltham, MA, USA) equipped with a ZORBAX Eclipse Plus C18 column (150 × 4.6 mm, 5 μm particle size, Agilent technologies, Santa Clara, CA, USA). Aflatoxins determination was carried out by using an UltiMate™ 3000 Fluorescence Detector (Thermo Scientific, Waltham, MA, USA) with excitation and emission at 362 and 420 nm, after post-column derivatization through a UVE™ Photochemical Reactor for Aflatoxin Analysis (254 nm lamp; 240 VAC, 50/60 Hz, LCTech, Obertaufkirchen, Germany). The run conditions were set at a flow rate of 0.8 mL min−<sup>1</sup> of a mobile phase 45% methanol (*v*/*v* in HPLC-demineralized water), for 30 min at 30 ◦C. Patulin quantification was performed using a Dionex UV-Vis Detector (Dionex UVD 170 U UV-Vis detector, Thermo scientific, Waltham, MA, USA) at 276 nm, at a flow rate of 0.6 mL min−1, for 30 min at 30 ◦C, and the same mobile phase reported above. Known amounts of pure (Patulin HPLC standard) or mixed (Aflatoxin Mix 4 solution) standards were injected into the UHPLC system (range 0.1−100 ng mL<sup>−</sup>1), and the quantity of each mycotoxin was obtained by correlating the peak area to the related standard concentration by using the Chromeleon Chromatography Management System software, version 7.2.10-2019 (Thermo Scientific, Waltham, MA, USA). The sum of AFB1, AFB2, AFG1 and AFG2 was considered as a measure of the total aflatoxins content.

Stock cultures of *C. maculatus*, kindly provided by Graham J. Holloway (University of Reading, Reading Berkshire, UK), were maintained in a climatic chamber (25 ± 1 ◦C, 70 ± 5% RH) on chickpea seeds in a 150 mL PP jar (5.5 cm diameter) that was closed with a micro-perforated nylon layer (350 μm pore size) to facilitate air exchange. To obtain chickpea infested with eggs, pest sub-cultures were set up by placing 100 unsexed adults into glass Petri dishes (14 cm diameter) containing 100 g of chickpea seeds. Adults were allowed to oviposit for 24 h in a climatic chamber under the same conditions as described before. At the end of the oviposition period, the adults were removed. Seeds with the addition of age-synchronized eggs were immediately used for the N2-controlled atmosphere test [51]. The lab-scale experiment was set up in six 60 L silos (three under a 99% N2 atmosphere and three under an unmodified atmosphere used as control). Two biotests, each consisting of 15 g of chickpea seeds with the addition of 1-3 eggs (up to a total of 30 eggs) placed into a 30 mL PP jar (4.3 cm diameter) and covered with a micro-perforated nylon layer (350 μm pore size), were transferred into each of the 60 L silos, which were already full of chickpea seeds; one bio-test was placed at the center and the other 10 cm under the grains' surface. The infected seeds were incubated for 24, 48, 72, 96 and 120 days. To avoid the opening of the silos at each sampling, the experiment was repeated each time, with three replicates for each experiment.

With respect to the field-scale experiment, two silos filled with 9 tons of chickpea seeds (the same batch as the laboratory) were used for the tests: one of these was automatically kept at a 99% N2-controlled atmosphere, and the other one, with the lid partially opened and disconnected from the N2 supply, was used as a control. In order to simulate a storage condition inside the full grain mass, three bio-tests previously placed into a jute bag filled with 8 kg of chickpea seeds were transferred into the silos. The test was carried out for 5 days and replicated three times. At the end of each experiment (lab-scale and large-scale test), the bio-tests were removed from the silos and their contents transferred into larger PP jars (like those used for pest stock cultures), closed with a micro-perforated nylon layer and placed in a climate chamber at 25 ± 2 ◦C, 70 ± 5% RH. The adults' emergence was periodically recorded from 35 to 70 days after oviposition. The number of adults was registered, and mortality was corrected using Abbott's formula [52].

$$\text{Mortality (\%)} = [(\text{N}\_{\text{c}} - \text{N}\_{\text{t}})/(\text{N}\_{\text{c}})] \times 100$$

where Nc = No. of emerged adults in control and Nt = No. of emerged adults in treatment.

#### *5.6. Statistical Analysis*

The robustness of data among the replicates was verified according to the results of the Shapiro–Wilk test for normality and Levene's tests for homogeneity of variance. Data were submitted to an analysis of variance (one-way or two-way ANOVA), and comparisons among the means were determined by Tukey's HSD post hoc test or Student *t*-test by using JMP Pro 14 software (SAS Institute Inc., Cary, NC, USA), in order to evaluate the effect of the treatments (control silos; O3 exposure for 30, 60 and 90 min; and N2 exposure for 21 consecutive days), batch (B1-4), and their interaction. For all the analyses, *p* ≤ 0.05 was assumed as a significant level.

**Supplementary Materials:** The following supporting information can be downloaded at: https://www.mdpi.com/article/10.3390/toxins15010061/s1, Table S1: Analysis of variance of mycotoxins content in the four chickpea batches between the mean values determined on three different replicates; Figure S1: Total content of aflatoxins in four batches [n. 1 (a), n. 2 (b), n. 3 (c) and n. 4 (d)] of chickpea grains (CTRL, white fill), exposed to ozone [500 ppb O3 for 30 (light grey fill), 60 (grey fill) and 90 (dark grey fill) minutes] and nitrogen treatment [99% N2 for 21 consecutive days, dark fill]. Results of two-way ANOVA are reported, asterisks show the significance of factors/interaction for: \*\*\* *p* ≤ 0.001. According to Tukey's HSD post hoc test, different letters indicate significant differences (*p* ≤ 0.05).

**Author Contributions:** Conceptualization, L.M., C.N. and S.S.; methodology, software, validation, investigation and formal analysis, C.P., S.R., L.M. and S.S.; resources, C.N., S.S. and E.P.; data curation, C.P., S.R., L.M. and S.S.; writing—original draft preparation, C.P., S.R., S.S. and E.P.; writing—review and editing, L.M. and C.N. All authors have read and agreed to the published version of the manuscript.

**Funding:** The experiments were carried out at CRISBA–ISIS Leopoldo II di Lorena (GR) and the construction of the field scale-N2 supplier device apparatus was funded by Fondazione CR Firenze.

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

**Informed Consent Statement:** Not applicable.

**Data Availability Statement:** Not applicable.

**Acknowledgments:** We gratefully acknowledge Andrea Parrini and Grazia Puntoni for their technical support for the fumigation facilities and nitrogen treatment, respectively; Enrico Giussani for furnishing the raw material; and Luca Mussi for the experimental activities. We also acknowledge Gabriele Simone for the technical support in setting up the tests on pests.

**Conflicts of Interest:** Authors declare no conflict of interest.

#### **References**


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