Next Article in Journal
Response to Leopoldo Palma. Comments on Ekino et al. Cloning and Characterization of a Unique Cytotoxic Protein Parasporin-5 Produced by Bacillus thuringiensis A1100 Strain. Toxins 2014, 6, 1882–1895
Previous Article in Journal
Use of Monoclonal Antibodies in the Sensitive Detection and Neutralization of Botulinum Neurotoxin Serotype B
 
 
Search for Articles:
Title / Keyword
Author / Affiliation / Email
Journal
Article Type
 
 
Advanced Search
 
Section
Special Issue
Volume
Issue
Number
Page
 
Journals
Toxins
Volume 7
Issue 12
10.3390/toxins7124864
toxins-logo
Submit to this Journal Review for this Journal Propose a Special Issue
► Article Menu

Article Menu

share Share announcement Help format_quote Cite question_answer Discuss in SciProfiles thumb_up
...
Endorse textsms
...
Comment
first_page
settings
Order Article Reprints
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:
Background:
Article

Biodegradation of Ochratoxin A by Bacterial Strains Isolated from Vineyard Soils

by
Palmira De Bellis
1,†,
Mariana Tristezza
2,†,
Miriam Haidukowski
1,
Francesca Fanelli
1,
Angelo Sisto
1,
Giuseppina Mulè
1 and
Francesco Grieco
2,*
1
Institute of Sciences of Food Production, National Research Council, Unit of Bari, via Amendola 122/O, Bari 70126, Italy
2
Institute of Sciences of Food Production, National Research Council, Unit of Lecce, via Provinciale Lecce-Monteroni, Lecce 73100, Italy
*
Author to whom correspondence should be addressed.
†
These authors contributed equally to this work.
Toxins 2015, 7(12), 5079-5093; https://doi.org/10.3390/toxins7124864
Submission received: 7 October 2015 / Revised: 9 November 2015 / Accepted: 13 November 2015 / Published: 27 November 2015
(This article belongs to the Section Mycotoxins)
Browse Figures
Versions Notes

Abstract

:
Ochratoxin A (OTA) is a mycotoxin with a main nephrotoxic activity contaminating several foodstuffs. In the present report, five soil samples collected from OTA-contaminated vineyards were screened to isolate microorganisms able to biodegrade OTA. When cultivated in OTA-supplemented medium, OTA was converted in OTα by 225 bacterial isolates. To reveal clonal relationships between isolates, molecular typing by using an automated rep-PCR system was carried out, thus showing the presence of 27 different strains (rep-PCR profiles). The 16S-rRNA gene sequence analysis of an isolate representative of each rep-PCR profiles indicated that they belonged to five bacterial genera, namely Pseudomonas, Leclercia, Pantoea, Enterobacter, and Acinetobacter. However, further evaluation of OTA-degrading activity by the 27 strains revealed that only Acinetobacter calcoaceticus strain 396.1 and Acinetobacter sp. strain neg1, consistently conserved the above property; their further characterization showed that they were able to convert 82% and 91% OTA into OTα in six days at 24 °C, respectively. The presence of OTα, as the unique OTA-degradation product was confirmed by LC-HRMS. This is the first report on OTA biodegradation by bacterial strains isolated from agricultural soils and carried out under aerobic conditions and moderate temperatures. These microorganisms might be used to detoxify OTA-contaminated feed and could be a new source of gene(s) for the development of a novel enzymatic detoxification system.

1. Introduction

Mycotoxins are natural contaminants in foodstuffs that, according to the Food and Agricultural Organization (FAO), contaminate more than 25% of the world’s commodities [1]. Among more than 300 already known mycotoxins [2,3], ochratoxin A (OTA) is one of the most important not only for its toxicological potential but also because of its extensive distribution. OTA is nephrotoxic, teratogenic, embryotoxic, immunotoxic, genotoxic, and neurotoxic in animals and humans [4,5]. Moreover, OTA is considered a possible human carcinogen and is classified in Group 2B by the International Agency of Research on Cancer [6]. OTA is produced by species belonging to Aspergillus and Penicillium genera, mainly Aspergillus niger, A. ochraceus, A. carbonarius, and Penicillium verrucosum. Ochratoxin A, an isocoumarin derivative linked by its carboxyl group to l-β-phenylalanine, is one of the most relevant mycotoxins because of both its toxicological potential and widespread distribution [7]. Its occurrence has been reported in a wide variety of food and beverages including cereals, cocoa, olives, coffee, beer, and spices [4,8,9,10]. Grapes and their derived products, such as grape juice, dried vine fruits, and wine, are frequently contaminated with this toxin [11,12] and are among the major sources of OTA dietary exposure [13]. OTA contamination severity in vineyards depends on several factors, including environmental conditions, vintage, vineyard location, cropping system [14,15,16,17], soil type, characteristics of the grapes [18], and competitive growth of microorganisms [19]. For example, OTA occurrence in Europe is higher in wines from southern regions because of the warmer climates [11] and red wines are contaminated more frequently than white ones probably due to the different winemaking methods [20,21].
Due to the impact of this mycotoxin on health, preventive measures for the reduction of fungal growth and mycotoxin levels have received much attention in recent years. The usual methods to reduce the levels of OTA comprised several approaches, such as physical treatment, addition with absorbent materials, or solvent extraction. Nevertheless, these systems are often expensive and they cause organoleptic changes in treated foodstuffs. Decontamination or detoxification procedures have been considered and, in particular, several physical, chemical, or biological approaches have been proposed for OTA reduction in musts and wine [21]. Biodegradation is considered one of the most promising approaches to control mycotoxins. Several enzymes may be involved in the microbiological degradation of OTA [22]. However, little information is available and very few of them have been purified and characterized. The first reported protease able to hydrolyze OTA was carboxypeptidase A (CPA) (EC 3.4.17.1) from bovine pancreas, thus releasing l-β-phenylalanine and ochratoxin α (OTα) [23,24,25]. OTα is considered a non-toxic compound and has an elimination half-life 10-times shorter than OTA [26]. Several studies have reported degradation of ochratoxin A by different microorganisms, such as Phenylobacterium immobile [27], Acinetobacter calcoaceticus [28], Trichosporon mycotoxinivorans [29], Brevibacterium spp. [30], Pediococcus parvulus [31], and Aspergillus spp. [32,33,34]. Several reports also showed that agricultural soils could represent an interesting source of microorganisms with ability to degrade mycotoxins [35,36]. In fact, soil bacteria, such as Pseudomonas spp. and Acinetobacter spp. are capable of converting a large number of aromatic compounds, thus playing an important role in the biodegradation of toxic molecules in contaminated soils [37]. For example, Shima and coworkers [38] identified from a soil sample a Gram-negative bacteria able to aerobically metabolize the mycotoxin deoxynivalenol (DON) into 3-keto-4-deoxynivalenol. Additionally, bacterial strains of soil origin belonging to Nocardioides and Devosia genera were characterized for their capacity to degrade DON to 3-epi-deoxynivalenol [36], whereas Islam et al. [35] isolated from agricultural soils bacterial populations able to biodegrade DON under aerobic conditions and moderate temperatures via a de-epoxydation mechanism. With this regard, in our knowledge, this is the first study aimed to evaluate degradation of ochratoxin A in soil samples from vineyards contaminated by high levels of OTA in order to isolate new microorganisms with OTA-degrading ability.

2. Results

2.1. Degradation Activity of Soil and Branch Samples

Soil and branch samples were first analyzed in three different culture media to promote the growth of fungi, yeast, and bacteria in the presence of OTA. The results shown in Figure 1 indicate that microbial populations from three soil samples, when grown in Minimal Medium Peptone (MMP) substrate, achieved the total degradation of OTA to OTα in six days, while cultures growing in Potatto Dextrose Broth (PDB) and Minimal Medium Sucrose MMS substrates did not attain a complete biodegradation of OTA. The cultures also failed the biodegradation of OTA in MM (data not shown). The microbial populations associated to branches did not show any ability to biodegrade the OTA (data not shown). An aliquot of each one of the three soil suspensions with OTA-degrading ability was then used to inoculate MMP and PDB supplemented with 10 µg/mL of OTA and a given antibiotic or an antifungal agent. The results (Figure 2) revealed the bacterial nature of microorganisms with degrading properties present within each microbial soil population, since this degrading activity was not modified by the presence of nystatine, but was inhibited by the addition of chloramphenicol.
Toxins 07 04864 g001 1024
Figure 1. Degradation of OTA (grey) in OTα (white) by microbial populations associated to five soil samples in three liquid substrates.
Figure 1. Degradation of OTA (grey) in OTα (white) by microbial populations associated to five soil samples in three liquid substrates.
Toxins 07 04864 g001
Toxins 07 04864 g002 1024
Figure 2. Ochratoxin A degradation by microbial populations associated with the three soil samples respectively cultured in presence of nystatine or chloramphenicol. The OTA degradation in absence of antibiotic addition is also shown.
Figure 2. Ochratoxin A degradation by microbial populations associated with the three soil samples respectively cultured in presence of nystatine or chloramphenicol. The OTA degradation in absence of antibiotic addition is also shown.
Toxins 07 04864 g002
Toxins 07 04864 g003 1024
Figure 3. Gel-like image obtained using the Agilent Expert software of the 27 different rep-PCR profiles identified after the analysis of the 225 OTA-degrading isolates. The profiles were obtained using primers REP-1R-Dt/REP-2R-Dt. L: DNA 7500 ladder.
Figure 3. Gel-like image obtained using the Agilent Expert software of the 27 different rep-PCR profiles identified after the analysis of the 225 OTA-degrading isolates. The profiles were obtained using primers REP-1R-Dt/REP-2R-Dt. L: DNA 7500 ladder.
Toxins 07 04864 g003

2.2. Genotypic Characterization and Identification of Bacteria from OTA-Degrading Soils

The results obtained by chemical analysis showed that bacterial populations from soil samples 3PR, 4NG, and 5NG included isolates able to degrade OTA to OTα, while this property was not detected in populations associated with soil samples 1NG and 2PR. On the basis of this evidence, a total of 225 bacterial isolates with degrading activity were obtained from those three soil samples and then investigated for possible clonal relationships, using the rep-PCR technique. Twenty-seven different rep-PCR patterns, each associated with a number of isolates ranging from 1 to 64, were obtained using the REP primers (Figure 3). In particular, 13 profiles differentiated the isolates from soil sample 3PR, while six and eight patterns distinguished those from soil samples 4NG and 5NG, respectively.
An isolate representative of each rep-PCR profile was identified by sequencing about 1400 bp of the 16S rRNA gene. Primer pair P0/P6 was suitable for the amplification of the 16S rRNA gene from isolates belonging to the genus Acinetobacter, while the primer pair 27F-YM/1492r provided better results for the other isolates. Database searches with these sequences showed identities >99% with type strain sequences available in database, leading to the assignment of the following bacterial species: Acinetobacter calcoaceticus, Pseudomonas taiwanensis, Pseudomonas reinekei, Pseudomonas koreensis, Leclercia adecarboxylata, Pantoea agglomenans, Enterobacter aerogenes, and Enterobacter xiangfangensis (Table 1). The genus Pseudomonas was the one more frequently isolated from each soil and Pseudomonas koreensis was the only detected species from two soils (3PR and 5NG). Acinetobacter sp. strain neg1 was an exception because its 16S rRNA gene showed the best alignment with Acinetobacter baumanni but at a percentage of identity <99% (i.e., 98.29% and four gaps); in fact, this strain was assigned to a new species on the basis of genomic data (Genome GenBank accession number JSZD00000000; [39]). The 16S rRNA gene sequence of strain 396.1, due to its relevance as an OTA-degrading strain, was deposited under GenBank accession number KR856228.
Table
Table 1. Bacterial strains isolated from OTA-degrading soils.
Table 1. Bacterial strains isolated from OTA-degrading soils.
SpeciesStrain (REP-PCR Profile)Isolation Soil
Pseudomonas taiwanensis106 (1), 115 (2), 154 (3)4NG
Leclercia adecarboxylata158 (4)4NG
Acinetobacter sp.neg1 (5)4NG
Pantoea agglomerans195 (6)4NG
Pseudomonas reinekei207 (7), 217 (8), 261 (13), 291 (14)5NG
Pseudomonas koreensis221 (9), 244 (10), 254 (11), 256 (12)5NG
303 (15), 308 (16), 310 (17), 321.2 (18), 334.1 (19), 372 (23)3PR
Enterobacter aerogenes339 (20), 348 (21), 358 (22), 376 (24), 390 (26)3PR
Acinetobacter calcoaceticus396.1 (25)3PR
Enterobacter xiangfangensis399.2 (27)3PR

2.3. OTA Degradation by Acinetobacter spp.

The twenty-seven identified strains were preliminarily characterized for their ability in OTA degradation. A marked degradation of OTA level was observed after six days (Figure 4). However, when these strains were further characterized for their ability in OTA conversion, the only strains that maintained the OTA-degrading ability were the neg1 and 396.1, both belonging to the Acinetobacter genus. These two strains were from two out of three soil samples showing degrading activity, in particular, Acinetobacter sp. strain neg1 from soil 4NG and A. calcoaceticus strain 396.1 from 3PR. A significant degradation of OTA level was observed over six days. The results of the analyses revealed that these strains (viable cells) degraded OTA to OTα. The average degradation levels of OTA with respect to the control (with 100% OTA) were 20% (RSD, 20%) after three days and 82% (RSD, 12%) after six days for Acinetobacter sp. strain neg1, while for A. calcoaceticus strain 396.1 were 54% (RSD, 12%) after three days and 91% (RSD, 1%) after six days at 24 °C (Figure 5). Based on our evidence, when tested in a temperature range spanning from 22 to 28 °C, the bacterial degrading activity was slightly more efficient at 24 °C. Degrading activity was even functional after one day of incubation, when, for example at 24 °C, 6% and 14% of OTA resulted in being degraded by strain neg1 and strain 396.1, respectively. Results confirmed the ability of enzymes produced by strains neg1 and 396.1 of degrading OTA to OTα. According to the molecular structure of OTA, the microbial degradation through the hydrolysis of the amide bond results in l-β-phenylalanine molecule and OTα [23,24,25]. The production of OTα metabolite was ascertained by HPLC-HRMS (Figure 6). Our analyses did not reveal the presence of l-β-phenylalanine, possibly indicating that l-β-phenylalanine was immediately used or further degraded by bacterial cells.
Toxins 07 04864 g004 1024
Figure 4. Percentage of OTA degradation by the twenty-seven strains identified by the REP-PCR analysis. Cultures were incubated at 24 °C and mycotoxin degradation was evaluated by HPLC-FLD assay after six days. Bars represent standard errors. (A) Pseudomonas taiwanensis; (B) Leclercia adecarboxylata; (C) Acinetobacter sp. neg1; (D) Pantoea agglomerans; (E) Pseudomonas reinekei; (F) Pseudomonas koreensis; (G) Enterobacter aerogenes; (H) Acinetobacter calcoaceticus; and (I) Enterobacter xiangfangensis.
Figure 4. Percentage of OTA degradation by the twenty-seven strains identified by the REP-PCR analysis. Cultures were incubated at 24 °C and mycotoxin degradation was evaluated by HPLC-FLD assay after six days. Bars represent standard errors. (A) Pseudomonas taiwanensis; (B) Leclercia adecarboxylata; (C) Acinetobacter sp. neg1; (D) Pantoea agglomerans; (E) Pseudomonas reinekei; (F) Pseudomonas koreensis; (G) Enterobacter aerogenes; (H) Acinetobacter calcoaceticus; and (I) Enterobacter xiangfangensis.
Toxins 07 04864 g004
Toxins 07 04864 g005 1024
Figure 5. OTA degradation by Acinetobacter sp. neg1 (A) and A. calcoaceticus 396.1; and (B) in MMP medium added with 1 µg/mL of OTA. Cultures were incubated at 22, 24 and 28 °C and mycotoxin degradation was evaluated by HPLC-FLD assay at 13 and six days post inoculation.
Figure 5. OTA degradation by Acinetobacter sp. neg1 (A) and A. calcoaceticus 396.1; and (B) in MMP medium added with 1 µg/mL of OTA. Cultures were incubated at 22, 24 and 28 °C and mycotoxin degradation was evaluated by HPLC-FLD assay at 13 and six days post inoculation.
Toxins 07 04864 g005
Toxins 07 04864 g006 1024
Figure 6. LC-HR-MS Total Ion Current (TIC) and extracted ion chromatograms (XIC) filtered on the accurate mass of ochratoxin A and ochratoxin alpha from the supernatant of liquid culture sample added with ochratoxin A and inoculated Acinetobacter sp. neg1. In the upper panels are reported (A) the full mass chromatogram; (B) the extracted ion chromatogram filtered on the accurate mass of ochratoxin alpha; and (C) the extracted ion chromatogram filtered on the accurate mass of ochratoxin A.
Figure 6. LC-HR-MS Total Ion Current (TIC) and extracted ion chromatograms (XIC) filtered on the accurate mass of ochratoxin A and ochratoxin alpha from the supernatant of liquid culture sample added with ochratoxin A and inoculated Acinetobacter sp. neg1. In the upper panels are reported (A) the full mass chromatogram; (B) the extracted ion chromatogram filtered on the accurate mass of ochratoxin alpha; and (C) the extracted ion chromatogram filtered on the accurate mass of ochratoxin A.
Toxins 07 04864 g006

3. Discussion

To date, no biological treatment is currently employed to reduce the content of OTA in foods, beverages, and feed. To fulfill this urgent demand and because of previous evidence showing [30,40] that soil bacteria are capable of converting a wide assortment of aromatic compounds, we decided to analyze the fate of OTA in the presence of bacterial populations associated with soil samples. Since it has been shown that OTA can absorb relatively strongly to soil organic matter [41], the soil samples used in the present study were collected from vineyards infected by ochratoxigenic fungi and contaminated by high levels of OTA [42]. A rapid degradation of OTA in soil was demonstrated by Mortensen and coworkers [43], who showed that the degradation was faster in planted soil compared to unplanted soil, possibly due to a higher microbial activity in the rhizosphere of the planted soil [44]. In fact, our analyses confirmed an OTA-degrading activity also in our collected soil samples. In our initial screening steps, upwards of 144 h of incubation were required to achieve >70% OTA degradation by bacterial populations associated to the analyzed soil samples. The extent of the time required for toxin conversion can be explained by the presence of quickly growing microbes in artificial media that provoke a suppressive action on the growth of the OTA degrading bacterial strains [35,45]. The use of antibiotics and antifungal compounds with different modes of actions [46] allowed us to determine the bacterial nature of the OTA-degrading microbial populations associated to the soil samples. In this regard, the present investigation also describes the selective isolation from media supplemented with OTA of 225 soil bacterial isolates that, in a first screening test, showed OTA-degrading activity. A rep-PCR analysis enabled us to reveal the clonal relationships among the isolates leading to the detection of 27 different rep-PCR profiles, each associated to a variable number of isolates. The identification at the species level of 27 representative isolates, each characterized by a different rep-PCR profile, indicated that the 225 isolates belonged to nine different bacterial species. Three of these were of the Pseudomonas genus, while the other belonged to the genera Acinetobacter, Enterobacter, Pantoea, and Leclercia. Therefore, these results could suggest the presence in the analyzed soils of a relevant biodiversity, at the species and strain levels, of bacteria able to grow in the presence of OTA and to degrade this toxic compound. However, when the 27 representative isolates were further analyzed to confirm their OTA-degrading activity only two of them, namely Acinetobacter calcoaceticus (strain 396.1) and Acinetobacter sp. (strain neg1; [39]), consistently conserved this capability which, on the contrary, was definitely lost by the other strains.
The two abovementioned bacterial strains isolated from agricultural soil have confirmed their own capacity to convert OTA to OTα in aerobic conditions and at moderate temperatures. The loss of OTA-degrading activity by the other strains could be possibly due to the loss of genes responsible for this capability occurring during subculturing under not selective conditions in media without OTA. A similar behavior has been already described for OTA degradation by Lactobacillus vitulinus [29], chlorinated biphenyls conversion by Acinetobacter sp. strain B6 [47], and methyl parathion removal by Pseudomonas sp. [48]. Therefore, the two Acinetobacter strains, i.e., A. calcoaceticus (strain 396.1) and Acinetobacter sp. (strain neg1), were selected for further investigations.
OTA-degrading activity of A. calcoaceticus was also described in a previous investigation in which strain NRRL B-551 was shown to be able to degrade OTA in ethanol minimal salts medium with an initial concentration of 10 μg/mL at both 25 and 30 °C [28]; nonetheless, it is important to note that the OTA-degrading strain neg1 isolated in this study, based on genomic data [39], belongs to a new species.
A relevant interest should be addressed to the abovementioned Acinetobacter strains, isolated in this study, because of the high percentage of OTA removal. In addition, it should be considered the ecological importance of the genus Acinetobacter recently used as a model microorganism for studies in environmental microbiology and industrial production of chemicals [49]. Furthermore, the OTA degradation mechanism of both Acinetobacter strains was also denoted by a technologically relevant feature, that is, the ability to efficiently convert OTA in OTα at a moderate temperature (<25 °C). Indeed, with the exception of that concerning Phenylobacterium immobile [27], the available reports about OTA degradation by bacterial strains indicated that Bacillus licheniformis [50], lactic acid bacteria [51,52]; Brevibacterium spp. [30], Bacillus amyloliquefaciens [53], and Pediococcus parvulus [31] were able to convert OTA at a temperature equal or higher than 30 °C. The advantage of the present finding is that OTA conversion activity of the culture was optimal in a temperature range that matches the temperature range of agronomical crops. Therefore, it is reasonable to hypothesize that the gene(s)/enzyme(s) responsible for the activity have potential application for enzymatic detoxification of OTA in food and feed of vegetal origin [22].
Both Acinetobacter strains described here may utilize the enzymatic pathway to biodegrade OTA that has been recognized in other microorganisms [22], degrading OTA by means of the hydrolysis of the OTA amide group and the consequent release of OTα, that is considered non-toxic [54,55]. Different peptidases are known to catalyze the above hydrolysis, such as the carboxypeptidase A enzyme isolated from the bovine pancreas [56], a commercially available lipase-like enzyme [57], an OTA-degrading protein enzyme from Aspergillus niger [58], the carboxypeptidase Y isolated from Saccharomyces cerevisiae [22], and a carboxypeptidase-like from Trichosporon mycotoxinivorans [29].
On the above basis, the biological degradation of OTA is a very feasible approach for the decontamination of foodstuffs, since the use of chemical or physical tools is likely to also eliminate, along with the mycotoxin, nutrients and organoleptic-relevant compounds. The possibility to reduce OTA levels by the here described Acinetobacter strains isolated from agricultural soils is of great interest for the control of OTA contaminations. In fact, these strains will be a source of efficient enzymes for OTA biodegradation in other food or feeds. Accordingly, further studies are in progress to determine which enzymes take part in the detoxification process by Acinetobacter sp. strain neg1, and to establish the absence of byproducts with any residual toxicity. Moreover, experiments are now underway to characterize and to clone gene(s) responsible for the detoxification reaction, in order to develop a novel enzymatic detoxification system.

4. Experimental Section

4.1. Soil and Branch Sampling from Vineyards

Soil and branch samples were collected from five vineyards of Negroamaro and Primitivo grape cultivars, in the most significant production areas for these varieties in Salento (Apulia, Southern Italy). In particular, three vines planted with the variety Negroamaro were trained in a bower system in a head system, or in spur-pruned cordon, and two vineyards planted with the variety Primitivo were trained in a bower system or in an espalier system (Table 2). All of the sampled vineyards were conducted by the methods of organic agriculture and reported to be contaminated by A. carbonarius and by high levels of OTA in previous years [59]. In each vineyard, during the “dormant” period, a random sampling of soil and branches was performed from each source. The samples were placed in sterile plastic bags, cooled, and analyzed within 4 h after sampling.
Table
Table 2. Sampling from different ecological niches of the “system vineyard”.
Table 2. Sampling from different ecological niches of the “system vineyard”.
Sample IDOriginVineyard n.CultivarSiteTraining System
1 NGsoil1NegroamaroNardò (LE)Bower system
1 NGTbranches1NegroamaroNardò (LE)Bower system
2 PRsoil2PrimitivoNardò (LE)Bower system
2 PRTbranches2PrimitivoNardò (LE)Bower system
3 PRsoil3PrimitivoManduria (TA)Espalier
3 PRTbranches3PrimitivoManduria (TA)Espalier
4 NGsoil4NegroamaroCellino S. Marco (BR)Head system
4 NGTbranches4NegroamaroCellino S. Marco (BR)Head system
5 NGsoil5NegroamaroBrindisiSpur-pruned cordon
5 NGTbranches5NegroamaroBrindisiSpur-pruned cordon

4.2. Evaluation of the OTA-Degrading Ability of Soil and Branch Samples and Isolation of Microorganisms

Five grams of each sample (Table 2) were suspended in 50 mL of sterile distilled water. The suspension was incubated at room temperature with shaking (180 rpm) for 20 min. An aliquot (0.5 mL) of the suspension was added to 4.5 mL of three liquid substrates.
The three different liquid substrates used to support the growth of the microorganisms were: minimal medium (MM = K2HPO4 (2.5 g/L), KH2PO4 (2.5 g/L), (NH4)2HPO4 (1.0 g/L), MgSO4·7H2O (0.2 g/L), FeSO4 (0.01 g/L), MnSO4·7H2O (0.007 g/L)) with Bacto Peptone (0.5%, w/v) (MMP), minimal medium with sucrose (1.0%, w/v) (MMS), and potato dextrose broth (PDB) (Oxoid, Basingstoke UK) [35]. OTA (Sigma-Aldrich, St Louis, Missouri, USA) was added to MMP, MMS, and PDB media at a final concentration of 10 µg/mL. When the mycotoxin was used as the sole carbon source, it was added to MM medium at a final concentration of 100 µg/mL. Cultures were incubated at 24 °C with continuous shaking (180 rpm) and aliquots (0.5 mL) from each culture were collected after 0, 72, and 144 h. Cells were harvested by centrifugation (10,000× g for 10 min at 4 °C), the supernatants were filter sterilized (0.2 µm, Millipore, Billerica, MA, USA) and used for chemical analyses [60]. Each sample was tested in triplicate.
In order to assess the type of the OTA-degrading microbial populations, soil samples with OTA-degrading ability were treated as follows: an aliquot (0.5 mL) of the soil suspension, prepared as above described, was added to 4.5 mL of PDB and MMP medium, respectively, added with 50 µg/mL of chloramphenicol (for bacteria) or nystatine (for fungi), in the presence of 10 µg/mL of OTA. The amount of OTA biodegradation was determined after five days of incubation at 24 °C with continuous shaking (180 rpm). Moreover, to isolate OTA-degrading microorganisms, after three days of incubation, the cultures showing OTA-degrading ability were serially diluted and plated in triplicate on MMP agar. Plates were incubated for 24 h at 24 °C. From each samples, approximately 100 colonies were singly picked up from the agar plates and checked for purity prior to storage at −80 °C.

4.3. rep-PCR

In order to perform a molecular characterization of isolates, bacterial DNA was extracted from overnight cultures grown in Brain Heart Infusion (BHI) broth (Becton, Dickinson and Company, Franklin Lakes, NJ, USA) at 26 °C, using the Wizard Genomic DNA Purification Kit (Promega, Madison, WI, USA) and analyzed by repetitive element sequence-based PCR (rep-PCR) [61,62]. In the first phase of the study, in order to identify primers able to provide the best results in discriminating the strains, different primers were used to amplify the DNA of 12 isolates.
In particular, the primer pair REP-1R-Dt/REP-2R-Dt and the primer (GTG)5 were evaluated (Table 3). The PCR reactions and the PCR product analyses were performed as previously described [63]. The primer pair REP-1R-Dt/REP-2R-Dt discriminated better than the primer (GTG)5 the different strains, so it was chosen for the study of all the isolates.
Table
Table 3. Nucleotide sequences of the different primers used in this study for rep-PCR and for amplification and sequencing of 16S rRNA gene.
Table 3. Nucleotide sequences of the different primers used in this study for rep-PCR and for amplification and sequencing of 16S rRNA gene.
PrimerPrimer SequenceReference
REP-1R-Dt5′-IIINCGNCGNCATCNGGC-3′[63]
REP-2R-Dt5′- NCGNCTTATCNGGCCTAC-3′[63]
(GTG)55′-GTGGTGGTGGTGGTG-3′[64]
P05′-GAGAGTTTGATCCTGGCTCAG-3′[65]
P65′-CTACGGCTACCTTGTTACGA-3′[65]
27f-YM5′-AGAGTTTGATYMTGGCTCAG-3′[66]
1492r5′-TACCTTGTTACGACTT-3′[66]
ENT-16S-for5′-CAGCCACACTGGAACTGAGA-3′This study
ENT-16S-rev5′-GACAGCCATGCAGCACCT-3′This study
ENT-16S-revII5′-TTATGAGGTCCGCTTGCTCT-3′This study
PSE-16S-for5′-GGTCTTCGGATTGTAAAGCAC-3′This study
PSE-16S-rev5′-GACGACAGCCATGCAGC-3′This study

4.4. Identification of Bacterial Strains

Isolates representative of each rep-PCR profile were identified by 16S rRNA gene sequencing [67] The universal primer pair P0/P6 or when appropriate the primer pair 27f-YM/1492r (Table 3) were used for 16S rRNA gene amplification and their locations at the 5′ and 3′ ends of the 16S rRNA gene allow to amplify almost all the gene. Each 50 μL reaction mixture contained 5 μL of 10× AccuPrime™ Pfx Reaction Mix, 1.25 U of AccuPrime™ Pfx DNA polymerase (Thermo Fisher Scientific, Waltham, MA, USA), 0.3 μM of each primer and 1 μL of genomic DNA. PCR amplifications were performed in a GeneAmp PCR system 9700. The reaction mixtures were first incubated for 2 min at 95 °C, and then cycled for 30 cycles according to the following temperature profiles: 30 s at 95 °C, 30 s at an annealing temperature of 60 °C for the first five cycles, 55 °C for the next five cycles and 50 °C for P0/P6 or 48 °C for 27F-YM/1492r for the last 25 cycles, then 3 min at 68 °C, followed by a final extension for 10 min at 68 °C. Sequencing of the 16S rRNA gene PCR products were performed in both forward and reverse directions by using the universal primers P0, P6, 27f-YM, 1492r and internal primers (Table 3). PCR products were sequenced by using the BigDye™ Terminator cycle sequencing kit on an ABI Prism 3730xl DNA Analyzer (Thermo Fisher Scientific, Waltham, MA, USA). The Blast N program, available through the National Center for Biotechnology Information [68] was used to compare the sequenced PCR products with sequences of type strains in database (“Sequences from type material” option). Bacterial strains were assigned to the species on the basis of the highest scores of alignment and percentage of identity (>99%) between their 16S rRNA gene sequences and those of type strains in database.

4.5. Biodegradation of OTA by Bacterial Isolates

Bacterial isolates were analyzed to evaluate their OTA-degrading ability. Isolates were cultured (2% v/v) in BHI broth and incubated at 24 °C with shaking (120 rpm). After overnight incubation, 5 mL of MMP containing 1 µg/mL of OTA were inoculated with subcultures (OD600nm: 1.0) at 2% (v/v) in triplicate. Controls were prepared by dissolving 1 µg/mL of OTA in MMP liquid in triplicate (100% OTA). Both inoculation and control tubes were incubated at 22, 24, and 28 °C with shaking. After one, three, and six days, 0.5 mL aliquots were collected, cells were harvested by centrifugation (10,000× g for 10 min at 4 °C), the supernatants were filter sterilized (0.2 µm, Millipore) and used for chemical analyses.

4.6. Preparation of Standards

OTA stock solution was prepared by dissolving the solid commercial toxin (Sigma-Aldrich, St. Louis, MO, USA) in methanol (1 mg/mL). Appropriate aliquots of the stock solution were brought to dryness and reconstituted with acetonitrile-water-acetic acid (99:99:2 v/v/v) to obtain standard solutions of OTA over the range 0.05–0.10 µg/mL. The standard of OTα was purchased from Biopure (Romer Labs Diagnostic GmbH, Austria) at a concentration of 10 µg/mL in acetonitrile. Aliquots of the stock solution were brought to dryness and reconstituted with acetonitrile-water-acetic acid (99:99:2, v/v/v) to obtain standard solutions of OTα from 0.01 to 0.10 µg/mL.

4.7. Chemical Analyses

HPLC-FLD. Decimal dilutions of the supernatants were made with acetonitrile-water-glacial acetic acid (99:99:2, v/v/v; Baker, Italy). Samples were filtered using RC 0.2 µm micro spin filter tubes (Phenomenex, Torrance, CA, USA). Fifty µL were injected into the HPLC apparatus (Technology series 1100, Agilent, Santa Clara, CA, USA) with a full loop injection system. Direct injection of liquid culture was possible since no interfering peaks eluted at retention times of OTα and OTA.
Measurement of toxins was carried out as described by Gallo et al. [69], with minor modifications. OTα and OTA were measured by comparing peak areas with calibration curves; the retention times were 7.75 and 10.36 min, respectively.
The detection limit was 0.1 ng/mL for OTA and OTα (based on a signal-noise ratio 3:1). HPLC-HRMS. Structural confirmation of OTα and identification of others possible degradation metabolites were carried out by LC-HRMS. Twenty microliters of supernatants were also analyzed by LC-HRMS according to the procedure reported by Gallo et al. [69].

Acknowledgments

We acknowledge the Italian Ministry of Education, University and Research for supporting this research by the Project S.I.Mi.S.A—PON02_00186_3417512/1. We thank Stella Lisa Lonigro, Roberto Schena, Giuseppe Cozzi and Gianni Colella for their valuable technical assistance.

Author Contributions

All authors significantly contributed to this paper: Francesco Grieco, Angelo Sisto, and Giuseppina Mulè conceived and designed the experiments; Mariana Tristezza, Palmira De Bellis, Francesca Fanelli and Miriam Haidukowski performed the experiments; Francesco Grieco, Angelo Sisto, Giuseppina Mulè, Mariana Tristezza, Palmira De Bellis, Francesca Fanelli and Miriam Haidukowski analyzed the data; Francesco Grieco was responsible for manuscript preparation and submission; Francesco Grieco, Angelo Sisto, Giuseppina Mulè, Mariana Tristezza and Palmira De Bellis, reviewed the paper.

Conflicts of Interest

The authors declare no conflict of interest.

References

  1. Reddy, K.R.N.; Salleh, B.; Saad, B.; Abbas, H.K.; Abel, C.A.; Shier, W.T. An overview of mycotoxin contamination in foods and its implications for human health. Toxin Rev. 2010, 29, 3–26. [Google Scholar] [CrossRef]
  2. Bennett, J.W.; Klich, M. Mycotoxins. Clin. Microbiol. Rev. 2003, 16, 497–516. [Google Scholar] [CrossRef] [PubMed]
  3. Stoev, S.D. Food safety and increasing hazard of mycotoxin occurrence in foods and feeds. Crit. Rev. Food Sci. Nutr. 2013, 53, 887–901. [Google Scholar] [CrossRef] [PubMed]
  4. Benford, D.; Boyle, C.; Dekant, W.; Fuchs, R.; Gaylor, D.W.; Hard, G.; McGregor, D.B.; Pitt, J.I.; Plestina, R.; Shephard, G.; et al. Ochratoxin A. WHO Food Additives Series: 47. Safety evaluation of certain mycotoxins in food. In Proceedings of the 56th Joint FAO/WHO Expert Committee on Food Additives (JECFA), Geneva, Switzerland, 6–15 February 2001; Available online: http://www.inchem.org/documents /jecfa/jecmono/v47je04.htm (accessed on 10 June 2015).
  5. Wangikar, P.B.; Dwivedi, P.; Sinha, N.; Sharma, A.K.; Telang, A.G. Teratogenic effects in rabbits of simultaneous exposure to ochratoxin A and aflatoxin B1 with special reference to microscopic effects. Toxicology 2005, 215, 37–47. [Google Scholar] [CrossRef] [PubMed]
  6. IARC. Monograph on the Evaluation of Carcinogenic Risks to Human. In Some Traditional Herbal Medicines, Mycotoxins, Naphthalene and Styrene, Volume 82; World Health Organization, International Agency for Research on Cancer: Lyon, France, 2002. [Google Scholar]
  7. Kuiper-Goodman, T.; Scott, P.M. Risk assessment of the mycotoxin ochratoxin A. Biomed. Environ. Sci. 1989, 2, 179–248. [Google Scholar] [PubMed]
  8. Birzele, B.; Prange, A.; Krämer, J. Deoxynivalenol and ochratoxin A in German wheat and changes of level in relation to storage parameters. Food Addit. Contam. 2000, 17, 1027–1035. [Google Scholar] [CrossRef] [PubMed]
  9. Fazekas, B.; Tar, A.K.; Zomborszky-Kovács, M. Ochratoxin A contamination of cereal grains and coffee in Hungary in the year 2001. Acta Vet. Hung. 2002, 50, 177–188. [Google Scholar] [CrossRef] [PubMed]
  10. Roussos, S.; Zaouia, N.; Salih, G.; Tantaoui-Elaraki, A.; Lamrani, K.; Cheheb, M.; Hassouni, H.; Verhe, F.; Gaime, I.; Augur, C.; et al. Characterization of filamentous fungi isolated from Moroccan olive and olive cake: Toxinogenic potential of Aspergillus strains. Mol. Nutr. Food Res. 2006, 50, 500–506. [Google Scholar] [CrossRef] [PubMed]
  11. Battilani, P.; Magan, N.; Logrieco, A. European research on ochratoxin A in grapes and wine. Int. J. Food Microbiol. 2006, 111, S2–S4. [Google Scholar] [CrossRef] [PubMed]
  12. Rosa, C.A.R.; Magnoli, C.E.; Fraga, M.E.; Dalcero, A.M.; Santana, D.M.N. Occurrence of ochratoxin A in wine and grape juice marketed in Rio de Janeiro, Brazil. Food Addit. Contam. 2004, 21, 358–364. [Google Scholar] [CrossRef] [PubMed]
  13. El Khoury, A.; Atoui, A. Ochratoxin A: General overview and actual molecular status. Toxins 2010, 2, 461–493. [Google Scholar] [CrossRef] [PubMed]
  14. Bellí, N.; Mitchell, D.; Marín, S.; Alegre, I.; Ramos, A.J.; Magan, N.; Sanchis, V. Ochratoxin A-producing fungi in Spanish wine grapes and their relationship with meteorological conditions. Eur. J. Plant Pathol. 2005, 113, 233–239. [Google Scholar] [CrossRef]
  15. Battilani, P.; Giorni, P.; Bertuzzi, T.; Formenti, S.; Pietri, A. Black aspergilli and ochratoxin A in grapes in Italy. Int. J. Food Microbiol. 2006, 111, S53–S60. [Google Scholar] [CrossRef] [PubMed]
  16. Lucchetta, G.; Bazzo, I.; Cortivo, G.D.; Stringher, L.; Bellotto, D.; Borgo, M.; Angelini, E. Occurrence of black aspergilli and ochratoxin A on grapes in Italy. Toxins 2010, 2, 840–855. [Google Scholar] [CrossRef] [PubMed]
  17. Chiotta, M.L.; Ponsone, M.L.; Sosa, D.M.; Combina, M.; Chulze, S.N. Biodiversity of Aspergillus section Nigri populations in Argentinian vineyards and ochratoxin A contamination. Food Microbiol. 2013, 36, 182–190. [Google Scholar] [CrossRef] [PubMed]
  18. Jiang, C.; Shi, J.; Zhu, C. Fruit spoilage and ochratoxin A production by Aspergillus carbonarius in the berries of different grape cultivars. Food Control. 2013, 30, 93–100. [Google Scholar] [CrossRef]
  19. Belkacem-Hanfi, N.; Fhoula, I.; Semmar, N.; Guesmi, A.; Perraud-Gaime, I.; Ouzari, H.I.; Boudabous, A.; Roussos, S. Lactic acid bacteria against post-harvest moulds and ochratoxin A isolated from stored wheat. Biol. Control 2014, 76, 52–59. [Google Scholar] [CrossRef]
  20. Visconti, A.; Perrone, G.; Cozzi, G.; Solfrizzo, M. Managing ochratoxin A risk in the grape-wine food chain. Food Addit. Contam. Part A 2008, 25, 193–202. [Google Scholar] [CrossRef] [PubMed]
  21. Quintela, S.; Villarán, M.C.; López de Armentia, I.; Elejalde, E. Ochratoxin A removal in wine: A review. Food Control 2013, 30, 439–445. [Google Scholar] [CrossRef]
  22. Abrunhosa, L.; Paterson, R.R.M.; Venâncio, A. Biodegradation of ochratoxin A for food and feed decontamination. Toxins 2010, 2, 1078–1099. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  23. Pitout, M.J. The hydrolysis of ochratoxin A by some proteolytic enzymes. Biochem. Pharmacol. 1969, 18, 485–491. [Google Scholar] [CrossRef]
  24. Wu, Q.; Dohnal, V.; Huang, L.; Kuča, K.; Wang, X.; Chen, G.; Yuan, Z. Metabolic pathways of ochratoxin A. Curr. Drug Metab. 2011, 12, 1–10. [Google Scholar] [CrossRef] [PubMed]
  25. Ferenczi, S.; Cserháti, M.; Krifaton, C.; Szoboszlay, S.; Kukolya, J.; Szőke, Z.; Kőszegi, B.; Albert, M.; Barna, T.; Mezes, M.; et al. A New ochratoxin a biodegradation strategy using Cupriavidus basilensis Őr16 Strain. PLoS ONE 2014, 9. [Google Scholar] [CrossRef]
  26. Li, S.; Marquardt, R.R.; Frohlich, A.A.; Vitti, T.G.; Crow, G. Pharmacokinetics of ochratoxin A and its metabolites in rats. Toxicol. Appl. Pharmacol. 1997, 145, 82–90. [Google Scholar] [CrossRef] [PubMed]
  27. Wegst, W.; Lingens, F. Bacterial degradation of ochratoxin A. FEMS Microbiol. Lett. 1983, 17, 341–344. [Google Scholar] [CrossRef]
  28. Hwang, C.A.; Draughon, F.A. Degradation of ochratoxin A by Acinetobacter calcoaceticus. J. Food Protec. 1994, 57, 410–414. [Google Scholar]
  29. Schatzmayr, G.; Zehner, F.; Täubel, M.; Schatzmayr, D.; Klimitsch, A.; Loibner, A.P.; Binder, E.M. Microbiologicals for deactivating mycotoxins. Mol. Nutr. Food Res. 2006, 50, 543–551. [Google Scholar] [CrossRef] [PubMed]
  30. Rodriguez, H.; Reveron, I.; Doria, F.; Costantini, A.; de Las Rivas, B.; Muñoz, R.; Garcia-Moruno, E. Degradation of ochratoxin A by Brevibacterium species. J. Agric. Food Chem. 2011, 59, 10755–10760. [Google Scholar] [CrossRef] [PubMed]
  31. Abrunhosa, L.; Inês, A.; Rodrigues, A.I.; Guimarães, A.; Pereira, V.L.; Parpot, P.; Mendes-Faia, A.; Venâncio, A. Biodegradation of ochratoxin A by Pediococcus parvulus isolated from Douro wines. Int. J. Food Microbiol. 2014, 188, 45–52. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  32. Varga, J.; Rigó, K.; Téren, J. Degradation of ochratoxin A by Aspergillus species. Int. J. Food Microbiol. 2000, 59, 1–7. [Google Scholar] [CrossRef]
  33. Abrunhosa, L.; Venâncio, A. Isolation and purification of an enzyme hydrolyzing ochratoxin A from Aspergillus niger. Biotechnol. Lett. 2007, 29, 1909–1914. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  34. Amézqueta, S.; González-Peñas, E.; Murillo-Arbizu, M.; López de Cerain, A. Ochratoxin A decontamination: A review. Food Control 2009, 20, 326–333. [Google Scholar] [CrossRef]
  35. Islam, R.; Zhou, T.; Young, J.C.; Goodwin, P.H.; Pauls, K.P. Aerobic and anaerobic de-epoxydation of mycotoxin deoxynivalenol by bacteria originating from agricultural soil. World J. Microbiol. Biotechnol. 2012, 28, 7–13. [Google Scholar] [CrossRef] [PubMed]
  36. Sato, I.; Ito, M.; Ishizaka, M.; Ikunaga, Y.; Sato, Y.; Yoshida, S.; Koitabashi, M.; Tsushima, S. Thirteen novel deoxynivalenol-degrading bacteria are classified within two genera with distinct degradation mechanisms. FEMS Microbiol. Lett. 2012, 327, 110–117. [Google Scholar] [CrossRef] [PubMed]
  37. Cao, B.; Nagarajan, K.; Loh, K.C. Biodegradation of aromatic compounds: Current status and opportunities for biomolecular approaches. Appl. Microbiol. Biotechnol. 2009, 85, 207–228. [Google Scholar] [CrossRef] [PubMed]
  38. Shima, J.; Takase, S.; Takahashi, Y.; Iwai, Y.; Fujimoto, H.; Yamazaki, M.; Ochi, K. Novel detoxification of the trichothecene mycotoxin deoxynivalenol by a soil bacterium isolated by enrichment culture. Appl. Environ. Microbiol. 1997, 63, 3825–3830. [Google Scholar] [PubMed]
  39. Fanelli, F.; Chiara, M.; Liuzzi, V.C.; Haidukowski, M.; Tristezza, M.; Caterina, M.; D’Erchia, A.M.; Pesole, G.; Horner, D.S.; Mulè, G. Draft genome sequence of Acinetobacter sp. neg1 capable of degrading ochratoxin A. FEMS Microbiol. Lett. 2015, 362. [Google Scholar] [CrossRef]
  40. Karigar, C.S.; Rao, S.S. Role of microbial enzymes in the bioremediation of pollutants: A review. Enzyme Res. 2011, 805187. [Google Scholar] [CrossRef] [PubMed]
  41. Mortensen, G.K.; Strobel, B.W.; Hansen, H.C.B. Determination of zearalenone and ochratoxin A in soil. Anal. Bioanal. Chem. 2003, 376, 98–101. [Google Scholar] [CrossRef] [PubMed]
  42. Somma, S.; Perrone, G.; Logrieco, A.F. Diversity of black Aspergilli and mycotoxin risks in grape, wine and dried vine fruits. Phytopathol. Mediterr. 2012, 51, 131–147. [Google Scholar] [CrossRef]
  43. Mortensen, G.K.; Strobel, B.W.; Hansen, H.C.B. Degradation of zearalenone and ochratoxin A in three Danish agricultural soils. Chemosphere 2006, 62, 1673–1680. [Google Scholar] [CrossRef] [PubMed]
  44. Chekol, T.; Vough, L.R.; Chaney, R.L. Phytoremediation of polychlorinated biphenyl-contaminated soils: The rhizosphere effect. Environ. Int. 2004, 30, 799–804. [Google Scholar] [CrossRef] [PubMed]
  45. Davis, K.E.R.; Joseph, S.J.; Janssen, P.H. Effects of growth medium, inoculum size, and incubation time on culturability and isolation of soil bacteria. Appl. Environ. Microbiol. 2005, 71, 826–834. [Google Scholar] [CrossRef] [PubMed]
  46. Guan, S.; He, J.; Young, J.C.; Zhu, H.; Li, X.Z.; Ji, C.; Zhou, T. Transformation of trichothecene mycotoxins by microorganisms from fish digesta. Aquaculture 2009, 290, 290–295. [Google Scholar] [CrossRef]
  47. Furukawa, K.; Chakrabarty, A.M. Involvement of plasmids in total degradation of chlorinated biphenyls. Appl. Environ. Microbiol. 1982, 44, 619–626. [Google Scholar] [PubMed]
  48. Chaudhry, G.R.; Ali, A.N.; Wheeler, W.B. Isolation of a methyl parathion-degrading Pseudomonas sp. that possesses DNA homologous to the opd gene from a Flavobacterium sp. Appl. Environ. Microbiol. 1988, 54, 288–293. [Google Scholar] [PubMed]
  49. Jung, J.; Park, W. Acinetobacter species as model microorganisms in environmental microbiology: Current state and perspectives. Appl. Microbiol. Biotechnol. 2015, 99, 2533–2548. [Google Scholar] [CrossRef] [PubMed]
  50. Petchkongkaew, A.; Taillandier, P.; Gasaluck, P.; Lebrihi, A. Isolation of Bacillus spp. from Thai fermented soybean (Thua-nao): Screening for aflatoxin B1 and ochratoxin A detoxification. J. Appl. Microbiol. 2008, 104, 1495–1502. [Google Scholar] [CrossRef] [PubMed]
  51. Skrinjar, M.; Rasić, J.L.; Stojicić, V. Lowering of ochratoxin A level in milk by yoghurt bacteria and bifidobacteria. Folia Microbiol. (Praha) 1996, 41, 26–28. [Google Scholar] [CrossRef] [PubMed]
  52. Böhm, J.; Grajewski, J.; Asperger, H.; Rabus, B.; Razzazi, E. Study on biodegradation of some A- and B-trichothecenes and ochratoxin A by use of probiotic microorganisms. Mycotoxin. Res. 2000, 16, 70–74. [Google Scholar] [CrossRef] [PubMed]
  53. Chang, X.; Wu, Z.; Wu, S.; Dai, Y.; Sun, C. Degradation of ochratoxin A by Bacillus amyloliquefaciens ASAG1. Food Addit. Contam. Part A Chem. Anal. Control Expo. Risk Assess. 2014, 32, 564–571. [Google Scholar] [CrossRef] [PubMed]
  54. Xiao, H.; Marquard, R.R.; Frohlich, A.A.; Ling, Y.Z. Synthesis and structure elucidation of analogues of ochratoxin A. J. Agric. Food Chem. 1995, 43, 524–530. [Google Scholar] [CrossRef]
  55. Bittner, A.; Cramer, B.; Harrer, H.; Humpf, H.U. Structure elucidation and in vitro cytotoxicity of ochratoxin α amide, a new degradation product of ochratoxin A. Mycotoxin Res. 2015. [Google Scholar] [CrossRef] [PubMed]
  56. Doster, R.C.; Sinnhuber, R.O. Comparative rates of hydrolysis of ochratoxins A and B in vitro. Food Cosmet. Toxicol. 1972, 10, 389–394. [Google Scholar] [CrossRef]
  57. Stander, M.A.; Steyn, P.S.; van der Westhuizen, F.H.; Payne, B.E. A kinetic study into the hydrolysis of the ochratoxins and analogues by carboxypeptidase A. Chem. Res. Toxicol. 2001, 14, 302–304. [Google Scholar] [CrossRef] [PubMed]
  58. Abrunhosa, L.; Santos, L.; Venâncio, A. Degradation of ochratoxin A by proteases and by a crude enzyme of Aspergillus niger. Food Biotechnol. 2006, 20, 231–242. [Google Scholar] [CrossRef] [Green Version]
  59. Battilani, P.; Logrieco, A.; Giorni, P.; Cozzi, G.; Bertuzzi, T.; Pietri, A. Ochratoxin A production by Aspergillus carbonarius on some grape varieties grown in Italy. J. Sci. Food. Agric. 2004, 84, 1736–1740. [Google Scholar] [CrossRef]
  60. Bleve, G.; Lezzi, C.; Chiriatti, M.A.; D’Ostuni, I.; Tristezza, M.; Di Venere, D.; Sergio, L.; Mita, G.; Grieco, F. Selection of non-conventional yeasts and their use in immobilized form for the bioremediation of olive oil mill wastewaters. Bioresour. Technol. 2011, 102, 982–989. [Google Scholar] [CrossRef] [PubMed]
  61. Versalovic, J.; Koeuth, T.; Lupski, J.R. Distribution of repetitive DNA sequences in eubacteria and application to fingerprinting of bacterial genomes. Nucl. Acids Res. 1991, 19, 6823–6831. [Google Scholar] [CrossRef] [PubMed]
  62. Tobes, R.; Ramos, J.L. REP code: Defining bacterial identity in extragenic space. Environ. Microbiol. 2005, 7, 225–228. [Google Scholar] [CrossRef] [PubMed]
  63. Hyytiä-Trees, E.; Lyhs, U.; Korkeala, H.; Björkroth, J. Characterisation of ropy slime-producing Lactobacillus sakei using repetitive element sequence-based PCR. Int. J. Food Microbiol. 1999, 50, 215–219. [Google Scholar] [CrossRef]
  64. Coudeyras, S.; Marchandin, H.; Fajon, C.; Forestier, C. Taxonomic and strain-specific identification of the probiotic strain Lactobacillus rhamnosus 35 within the Lactobacillus casei group. Appl. Environ. Microbiol. 2008, 74, 2679–2689. [Google Scholar] [CrossRef] [PubMed]
  65. Di Cello, F.; Bevivino, A.; Chiarini, L.; Fani, R.; Paffetti, D.; Tabacchioni, S.; Dalmastri, C. Biodiversity of a Burkholderia cepacia population isolated from maize rhizosphere at different plant growth stages. Appl. Environ. Microbiol. 1997, 63, 4485–4493. [Google Scholar] [PubMed]
  66. Frank, J.A.; Reich, C.I.; Sharma, S.; Weisbaum, J.S.; Wilson, B.A.; Olsen, G.J. Critical evaluation of two primers commonly used for amplification of bacterial 16S rRNA genes. Appl. Environ. Microbiol. 2008, 74, 2461–2470. [Google Scholar] [CrossRef] [PubMed]
  67. Capozzi, V.; Ladero, V.; Beneduce, L.; Fernández, M.; Alvarez, M.A.; Benoit, B.; Laurent, B.; Grieco, F.; Spano, G. Isolation and characterization of tyramine-producing Enterococcus faecium strains from red wine. Food Microbiol. 2011, 28, 434–439. [Google Scholar] [CrossRef] [PubMed]
  68. National Center for Biotechnology Information. Available online: http://www.ncbi.nlm.nih.gov (accessed on 10 July 2015).
  69. Gallo, A.; Bruno, K.S.; Solfrizzo, M.; Perrone, G.; Mulè, G.; Visconti, A.; Baker, S.E. New insight into the ochratoxin A biosynthetic pathway through deletion of a nonribosomal peptide synthetase gene in Aspergillus carbonarius. Appl. Environ. Microbiol. 2012, 78, 8208–8218. [Google Scholar] [CrossRef] [PubMed]

Share and Cite

MDPI and ACS Style

De Bellis, P.; Tristezza, M.; Haidukowski, M.; Fanelli, F.; Sisto, A.; Mulè, G.; Grieco, F. Biodegradation of Ochratoxin A by Bacterial Strains Isolated from Vineyard Soils. Toxins 2015, 7, 5079-5093. https://doi.org/10.3390/toxins7124864

AMA Style

De Bellis P, Tristezza M, Haidukowski M, Fanelli F, Sisto A, Mulè G, Grieco F. Biodegradation of Ochratoxin A by Bacterial Strains Isolated from Vineyard Soils. Toxins. 2015; 7(12):5079-5093. https://doi.org/10.3390/toxins7124864

Chicago/Turabian Style

De Bellis, Palmira, Mariana Tristezza, Miriam Haidukowski, Francesca Fanelli, Angelo Sisto, Giuseppina Mulè, and Francesco Grieco. 2015. "Biodegradation of Ochratoxin A by Bacterial Strains Isolated from Vineyard Soils" Toxins 7, no. 12: 5079-5093. https://doi.org/10.3390/toxins7124864

APA Style

De Bellis, P., Tristezza, M., Haidukowski, M., Fanelli, F., Sisto, A., Mulè, G., & Grieco, F. (2015). Biodegradation of Ochratoxin A by Bacterial Strains Isolated from Vineyard Soils. Toxins, 7(12), 5079-5093. https://doi.org/10.3390/toxins7124864

Article Metrics

Back to TopTop