Mycotoxin-producing fungi occurring in sorghum grains from Saudi Arabia

Fungal Diversity (2010) 44:45 52 DOI 10.1007/s13225-010-0058-9 Mycotoxin-producing fungi occurring in sorghum grains from Saudi Arabia Mohamed A. Yassin & Abdel-Rheem El-Samawaty & Ali Bahkali & Mohamed Moslem & Kamel A. Abd-Elsalam & Kevin D. Hyde Received: 26 May 2010 / Accepted: 16 August 2010 / Published online: 11 September 2010 # Kevin D. Hyde 2010 Abstract Thirty one species belonging to 17 fungal genera were found to be associated with sorghum (Sorghum bicolor) grain samples imported to the Kingdom of Saudi Arabia. An agar plate method was used to screen 24 samples collected from different markets located in Riyadh region. Statistical comparisons of fungal isolation frequencies revealed that Aspergillus niger, Penicillium funiculesum and Rhizopus stolonifer were most frequently isolated from untreated grains, while A. niger, P. funiculesum and F. semitectum, were most dominant in surface-sterilized grains. Significant correlation coefficients were observed among frequencies of some fungal species. Mycotoxin investigations using HPLC revealed that mycotoxinproducing isolates varied in the type and concentration of toxins produced. A. niger isolates were the highest producers of aflatoxins, followed by A. flavus var. columnaris and A. terreus, P. funiculosum was the highest producer of the Penicillium toxin Patulin (41 ppb), while P. oxalicum was the highest producer of citreoviridin (10 ppb). F. verticillioides was the highest producer of the M. A. Yassin : A.-R. El-Samawaty : A. Bahkali : M. Moslem : K. A. Abd-Elsalam (*) : K. D. Hyde College of Science, Botany and Microbiology Department, King Saud University, Riyadh, Saudi Arabia e-mail: abdelsalamka@gmail.com M. A. Yassin : A.-R. El-Samawaty : K. A. Abd-Elsalam Agricultural Research Center, Plant Pathology Research Institute, Giza, Egypt K. D. Hyde School of Science, Mae Fah Laung University, Chiang Rai 57100, Thailand Fusarium toxins fumonisin (19.1 ppb) and Zearalenone (21.4 ppb). F. nygamai was the highest producer of vomitoxin (31.3 ppb). Two of the three Alternaria isolates were altenuene producers. Keywords Seed-borne. Fungi. Sorghum. Toxigenic Introduction Grain sorghum (Sorghum bicolor L.) is one of the main cereal crops that are extensively used for human food as well as poultry, cattle and horse feed. Total world production of sorghum reached about 54 million tons in 2003 (FAO 2004). Grain sorghum is the principal source of energy, protein, vitamins and minerals for more than 500 million humans in more than 30 countries of the semi-arid tropics (National Academic Science 1996). Industrially, wax, starch, syrup, alcohol, dextrose agar, edible oils and gluten feed are manufactured from sorghum grains (Komlaga et al. 2001; Murty and Renard 2001). Sorghum is attacked by several diseases that affect crop growth and productivity (Prom 2004; Shinde et al.2004; Navi et al. 2005). Of these, seed-borne diseases are the most widespread and devastating diseases that affect crop yield quantitatively and qualitatively (Dawson and Bateman 2001; Singh and Navi 2001; Erpelding and Prom 2006). International spread of grain sorghum plant pathogens occurs on imported infected or contaminated seeds (Agarwal and Sinclair 1996). Sorghum is an important cereal crop imported to Kingdom of Saudi Arabia for food and animal feed purposes. External or internal seed-borne fungi associated with sorghum grains may cause seed deterioration, reduce

46 Fungal Diversity (2010) 44:45 52 seed germination capacity, seedling diseases and may result in systemic plant diseases (Khanzada et al. 2002; Rodriguez et al. 2006). Several seed-borne fungal taxa such as A. alternata, A. flavus, A. fumigatus, A. niger, Cladosporium sp., Colletotrichum sublineolum, Curvularia spp., Fusarium moniliforme, F. oxysporum, F. pallidoroseum, Drechslera tetramera, Nigrospora sp., Penicillium spp., Phoma sp., and Rhizopus sp. have been frequently recovered from sorghum grains (Fakhrunnisa and Ghaffar 2006; Gwary et al. 2006). In addition to the effects of seed-borne fungi on seed health, such fungi may also produce harmful mycotoxins that can affect both human and animal health (Palanee et al. 2001; David et al. 2005). Occurrence of mycotoxin producing fungal species in grains and grain based food has been reported in many countries with a high frequency of Aspergillus and Fusarium taxa (Da Silva et al. 2000; Kumar et al. 2008). Mycotoxigenic seed-borne Aspergillus and Fusarium strains have been isolated from both freshly harvested and stored Brazilian sorghum (Da Silva et al. 2004). Recovery of several other mycotoxigenic fungi from sorghum grains have been frequently documented (Thakur et al. 2006; Makun et al. 2009). The objective of the present study was (1) to investigate the contaminant mycoflora of imported sorghum grains in Saudi Arabia markets and, (2) to evaluate mycotoxin productivity of the toxigenic isolated fungi. Materials and methods Isolation of seed-borne fungi Twenty four sorghum grain samples collected from different localities of Riyadh, in the Kingdom of Saudi Arabia, were examined for seed-borne fungi. Fungi were isolated and cultured according to the method described by Hussaini et al. (2009). Two sets of 10 g/each of sorghum grains were used either after they were surface sterilized or without sterilization. Surface sterilization was accomplished by immersing the seeds in 5% sodium hypochlorite solution for 10 20 min and followed by washing in three changes of sterile distilled water. Ten grains, in each case, were placed randomly onto potato dextrose agar (PDA) in three, 9 cm diam. Petri dishes. Petri dishes were incubated at 25 C and examined daily for 5 days, after which the colonies were counted. Isolates were purified either by single spore or hyphal tip methods and then transferred to PDA slants. The isolation frequencies of fungal species were calculated according to the method of Gonzalez et al. (1995). Identification of fungal isolates was carried out based on morphological and microscopic characteristics in the Mycological Center, Assiut University, Egypt. Mycotoxins assays Aflatoxin Isolates were grown on sterilized SMKY liquid medium [sucrose 200 g, magnesium sulphate 0.5 g, potassium nitrate 3 g, yeast extract 7 g and distilled water 1,000 ml (Diener and Davis 1966) in 100 ml flasks for 10 days at 27±2 C with three replicates per isolate. Cultures were then blended for 2 min using a high speed homogenizer and filtered though Whatman s filter paper. Aflatoxins were extracted from the homogenized filtrates using methanol solution (80:20 methanol/isolate filtrate). Solvents were evaporated under vacuum at 35 C; dried residues containing aflatoxin were dissolved in 1 ml of the same liquid mobile phase solution which contained methanol: acetic acid: water and stored in dark vials. The method of Christian (1990) was used to detect and determine aflatoxin production. The extract was passed through a 0.45 μm micro-filter. Analysis of compounds was performed on HPLC model PerkinElmer Brownlee validated C18, 100 mm 4.6 mm, 3 micron. The HPLC was equipped with an UV detector and the wave length in the UV detector was 365 nm. The mobile phase consisting of methanol: acetic acid: water (20.0/20.0/60.0 v/v/v). The total run time for the separation was approximately 25 min at a flow rate of 1 ml/min. Penicillium toxins Tested isolates were grown on sterilized malt extract prepared in 100 ml flasks for 7 10 days at 27±2 C with three replicates per isolate. Cultures were blended for 2 min using a high speed homogenizer and filtered using glass filter paper. Patulin was extracted from homogenized filtrate using acetonitrile: water (5:95 v:v) (liquid mobile phase) solution. The solvent was then evaporated at 35 C under vacuum. The dried residues containing Patulin were dissolved in 1 ml of the same liquid mobile phase solution. The method described by Christian (1990) was used to determine patulin. The extract was passed through a 0.45 μm microfilter. Analysis of compounds was performed on an HPLC model PerkinElmer Brownlee validated C18, 250 mm. The HPLC was equipped with UV detector and the wave length in the UV detector was 280 nm. The total run time for the separation was approximately 25 min at a flow rate of 1 ml/min. A reliable analytical quantitative method described by Stubblefield et al. (1988) was used for citreoviridin determination. The toxin was extracted with dichloromethane, and the extract was partially purified on silica and amino solidphase extraction (SPE) columns. The extract was analyzed for citreoviridin by normal-phase liquid chromatography,

Fungal Diversity (2010) 44:45 52 47 using a mobile phase of ethyl acetate: hexane (75:25) at 1.5 ml/min and a fluorescence detector to measure the yellow fluorescence (388 nm excitation, 480 nm emission). Fusarium toxins Fumonisin toxins were determined according to the method described by Mazzani et al. (2001). Isolates were grown on sterilized SMKY liquid medium prepared in 50 ml flasks for 10 days at 27±2 C, with three replicates per isolate. Fungal culture of each flask was blended with 5 g sodium chloride and 100 ml of methanol: water (80:20) solution at a high speed for one min, then filtered through glass microfiber filter paper. Ten ml of the filtrate was diluted with 40 ml of wash buffer and filtered again through 1 μm micro-fiber filter. Ten ml of the diluted extract were passed through a fumontest column (VICAM Company) and the column was washed using 10 ml of the same dilute solution. The fumonisin was eluted by passing one ml of HPLC grade methanol through the column and then elutes were re-collected. One ml of each of developer A (VICAM product No. G5005) and developer B (VICAM product No. G5004) were added to the elute and placed in a calibrated fluorometer (Series-4/VICAM). The zearalenone and vomitoxin concentration was determined as described above for fumonisin, but the dilution was made with 49 ml distilled water, which were passed through a Zearatest and/or vomitoxin column (VICAM Company) and then measured in a calibrated fluorometer model (Series-4/VICAM). Alternaria toxins The method described by Li et al. (2001) was used to examine the ability of Alternaria isolates to produce toxins. Flasks containing 12.5 g of autoclaved modified polished rice at 40% moisture were inoculated with agar plugs of one-week-old Alternaria spp. cultures. Flasks were incubated in the dark at 25 C for 21 days after which culture materials were homogenized with 30 ml of methanol and filtered through a Whatman s filter paper (no. 1). The filtrates were clarified with 60 ml of 20% ammonium sulphate and then about 40 ml of clear filtrate was extracted three times with 10 ml of chloroform. The organic phases were combined, evaporated to dryness, and dissolved in 4 ml of methanol for analysis by HPLC. The HPLC system consisted of a Shimadzu liquid chromatography equipped with a Rheodyne sample valve fitted with a 20 μl loop and a Shimadzu SPD-M10Avp UV photodiode array detector. The analytical column was Jupiter 4.6 250 mm 5 μ C18. The liquid mobile phase was methanol/water (80:20) containing 300 mg ZnSO4 H2O. A flow rate of 0.4 ml/min was used. The wavelength for recording chromatograms was 258 nm. A calibration curve was constructed for quantification purposes using the toxin standards and correlating peak-area versus concentration. Statistical analysis The isolation frequency (Fq) of genera was calculated according to the method described by Marassas et al. (1988). A randomized complete block design was used in the present study. Analysis of variance (ANOVA) of the fungal isolation frequency was performed with the MSTAT-C statistical package, Michigan State Univ., USA). Least significant difference (LSD) was used to compare fungi means. Cluster analysis by the unweighted pair-group method based on arithmetic mean (UPGMA) was performed using SPSS6.0 software package. Results Isolation frequencies of fungi recovered from non-sterilized and sterilized sorghum grains Thirty one species belonging to 17 fungal genera were obtained from the test samples (Table 1). Of these, R. stolonifer, A. niger and P. funiculosum were the dominant fungi isolated from untreated grains, with isolation frequencies of 40.39, 28.47 and 9.55%, respectively. Sterilized sorghum seeds yielded the dominant species A. niger, P. funiculosum and F. semitectum with isolation frequencies of 13.11 13.04 and 11.22%, respectively. These results indicate that R. stolonifer, A. niger and P. funiculosum may colonize the surface of sorghum grains. The other fungi isolated occurred in frequencies ranging from 0.00 to 2.44%. Analysis of variance of the fungal isolation frequency (Table 2) revealed that fungus and fungus treatment interaction were all highly significant sources of variation in frequencies of fungi isolated from sorghum grains (Table 2 & Fig. 1). The fungus was the most important as a source of variation in isolation frequency, while fungus treatment interaction was of second importance (Fig. 1). Due to the significant effect of fungus treatment interaction, the isolation frequencies of some fungi differed according to whether the seeds were surface sterilized or not. For example, the isolation frequencies of R. stolonifer and A. niger decreased from 40.39% and 28.47% to 00.00% and 13.11% respectively, when seeds were surface sterilized (Table 1). Significant positive and negative associations were observed between the incidence of some fungal species when compared with the frequency of others. A. alternata exhibited significant positive correlation with A. flavus, P. oxalicum and Setosphaeria rostrata. Significant positive correlation was also found among F.

48 Fungal Diversity (2010) 44:45 52 Table 1 Isolation frequencies of fungi recovered from nonsterilized and sterilized sorghum grains a The difference was significant at *P 0.05, **P 0.01, or not significant (NS) LSD for the difference=6.03 (P 0.05) or 7.93 (P 0.01) Fungi Treatments Difference a Non-sterilized seeds (%) Sterilized seeds (%) 1 Absidia cylindrospora 2.10 5.38 3.28 NS 2 Alternaria alternata 0.71 4.71 4.00 NS 3 Alternaria chlamydospora 1.33 0.00 1.33 NS 4 Alternaria sp. 1.16 2.49 1.33 NS 5 Aspergillus flavus 1.69 1.22 0.47 NS 7 Aspergillus flavus var. columnaris 0.01 0.01 0.00 NS 6 Aspergillus niger 28.47 13.11 15.36 ** 8 Aspergillus terreus 0.00 2.44 2.44 NS 9 Botryodiplodia theobromae 0.00 0.52 0.52 NS 10 Chrysosporium indicum 0.00 5.56 5.56 NS 11 Cochliobolus lunatus 2.29 0.98 1.31 NS 12 Curvularia pallescens 0.56 5.11 4.55 NS 13 Curvularia tritici 1.57 6.40 4.83 NS 14 Drechslera pluriseptata 0.00 0.60 0.60 NS 15 Epicoccum nigrum 2.44 1.76 0.68 NS 16 Eurotium amstelodami 0.00 6.00 6.00 NS 17 Fusarium nygamai 0.27 4.76 4.49 NS 18 Fusarium semitectum 0.78 11.22 10.44 ** 19 Fusarium sp. 0.67 4.44 3.77 NS 20 Fusarium thapsinum 0.00 2.39 2.39 NS 21 Fusarium verticillioides 0.78 3.00 2.22 NS 26 Papulaspora irregularis 0.00 0.97 0.97 NS 22 Penicillium chrysogenum 0.67 0.00 0.67 NS 23 Penicillium funiculosum 9.55 13.04 3.49 NS 24 Penicillium griseofulvum 0.60 0.00 0.60 NS 25 Penicillium oxalicum 0.93 0.00 0.93 NS 27 Phoma sp. 0.00 0.49 0.49 NS 28 Rhizopus stolonifer 40.39 0.00 40.39 ** 29 Setosphaeria rostrata 0.44 0.75 0.51 NS 30 Stenocarpella sp. 0.00 1.60 1.60 NS 31 Sterile mycelia 0.53 1.04 0.51 NS verticillioides and F. nygamai with P. grisofulvum, and Cochliobolus lunatus and F. semitectum. On the other hand isolate frequencies of Epicoccum nigrum and R. stolonifer exhibited negative correlation (Data not shown). Table 2 Analysis of variance of effect of sterilization on frequency of fungi isolated from sorghum seeds Source of variation D.F M.S F. value P<F Replication a 24 1.513 0.0128 Treatment (T) 1 1.634 0.0138 Fungus (F) 30 1,249.694 10.5708 0.000 T X F 30 853.249 7.2174 0.000 Error 1,560 118.221 a Samples were used as replicates Fungi isolated from non-sterilized sorghum grains formed several distinct groups based on their distribution patterns over the samples (Fig. 2). Within each group, fungi were associated strongly and positively in their distribution patterns over samples, whereas between groups, fungi were associated weakly or negatively (Aly et al., 2004). This phenogram implies the potential existence of sample (environment) related groups of fungi. Mycotoxin production Aflatoxins Most isolates were capable of producing detectable levels of both B and G aflatoxins, although four of the seven A. niger isolates failed to produce any detectable amount. The

Fungal Diversity (2010) 44:45 52 49 Table 3 Production of aflatoxin by Aspergillus spp. isolated from sorghum grains Isolates Aflatoxins (ppb) B1 B2 G1 G2 Fig. 1 Relative contribution of treatment (T), fungus (F) and their interaction (T x F) to variation in frequencies of fungi isolated from sorghum grains 1 A. flavus var. columnaris 2 1 2 3 2 A. niger 0 0 0 0 3 A. niger 0 0 0 0 4 A. niger 0 0 0 0 5 A. niger 0 0 0 0 6 A. niger 9 8 5 3 7 A. niger 6 6 3 2 8 A. niger 3 2 0 1 9 A. terreus 4 2 1 2 other three A. niger isolates were the best producers of aflatoxins over all, followed by A. flavus var. columnaris and A. terreus (Table 3). Penicillium toxins Isolates varied in the type and concentration of toxins produced. While one of the P. oxalicum isolates produced 25 and 37 ppb of patulin and citreoviridin, respectively, P. funiculosum produced only patulin and P. griseofulvum produced only citreoviridin (Table 4). The highest level of patulin was obtained from P. funiculosum, while the highest level of citreoviridin was obtained from P. oxalicum. Fusarium and Alternaria toxins All isolates except F. thapsinum were toxin producers. Toxin-producing isolates varied in the kind and concentrations of toxin produced. F. verticillioides was the highest producer of fumonisin and zearalenone (19.1 and 21.4 ppb, respectively), while F. nygamai was the highest producer (31.3 ppb) of vomatoxin (Table 5). Two of the three Alternaria isolates were altenuene producers with highest productivity by Alternaria sp. (Table 6). Discussion Analysis of fungal contamination of sorghum grains in the present study yielded 31 fungal species belonging to 17 genera. Table 4 Production of citreoviridin and patulin by Penicillium spp. isolated from sorghum grains Isolates Mycotoxins (ppb) Patulin Citreoviridin Fig. 2 Phenogram based on average linkage cluster analysis of frequency of 17 fungi isolated from 24 nonsterilized samples of sorghum grains. 1 P. funiculosum 41 0 2 P. funiculosum 12 0 3 P. griseofulvum 0 10 4 P. oxalicum 25 37 5 P. oxalicum 14 0 6 P. oxalicum 38 0

50 Fungal Diversity (2010) 44:45 52 Table 5 Production of fumonisin, zeralenone and vomatoxin by Fusarium spp. isolated from sorghum grains Isolates Mycotoxins (ppb) Fumonisin Zearalenone Vomatoxin 1 Fusarium nygamai 11.7 0.0 31.3 2 F. thapsinum 0.0 0.0 0.0 3 F. semitectum 4.6 17.9 23.4 4 Fusarium sp. 2.1 12.3 14.1 5 F. verticillioides 19.1 21.4 12.1 6 F. verticillioides 3.5 0.0 0.0 Most of these taxa have been previously reported to be responsible for grain sorghum damage in several parts of the world (Girish et al. 2004; Shinde et al. 2004; Pawinde et al. 2008). The predominance of A. niger, P. funiculesum, F. semitectum and R. stolonifer in the grains differed from the results of Fakhrunnisa and Ghaffar (2006). They found that A. candidus, A. flavus, A. niger, Penicillium spp., Piptocephalis sp. and Rhizopus sp. were predominant fungi recovered from sorghum grains in Pakistan. F. semitectum was most frequently recovered from sorghum grains in Puerto Rico (Erpelding and Prom 2006). These results indicate that these fungi may colonize the grain surface, hence, their sensitivity to surface sterilization. On the other hand, the isolation frequency of F. semitectum increased from 0.78% to 11.22% following surface sterilization. Other species isolates were not significantly affected by surface sterilization because they colonize the internal parts of the grains. Our results also differ from those of Hemanth et al. (2007) and Broggi et al. (2007) who found that A. alternata and F. moniliforme were the predominant pathogens in sorghum grain samples in Argentina. Recovery of diverse fungal groups from sorghum grains in the present study could be attributed to long term storage and/or transport of grains under damp environmental conditions, which would promote fungal growth. In addition, probable differences in sample genotypes and origins could play a critical role in mycoflora colonizing ability (Funnell and Pedersen 2006; Thakur et al. 2006). Differences in colonization could also be attributing to sample locations and storage structures (Hemanth et al. 2007; Islam et al. 2009). A significant positive correlation between the incidences of some fungi with others in this study may indicate the possibility of synergism in colonizing the sorghum grains by these fungi. A negative correlation may reflect an antagonism or competitive exclusion between such fungi. Erpelding and Prom (2006) found significant negative associations between the incidence of F. semitectum with the frequency of C. lanata and F. thapsinum. Most grain molding fungi are known to produce diverse toxic secondary metabolites generally called mycotoxins. These are mainly produced by Aspergillus, Penicillium, Fusarium species and other toxigenic taxa (Kang et al. 2001; Calvo et al. 2002). Amadi and Adeniyi (2009) reported that many of the Aspergillus and Penicillium isolates recovered from stored grains produced more than one toxin. Aflatoxins, the most widespread of toxins, are mainly produced by Aspergillus spp. In Morocco A. flavus strains isolated from sorghum produced AFB1 and AFB2 mycotoxins (Kichou and Walser 1993). Several aflatoxigenic Aspergillus strains produce both B and G aflatoxins, while other produces only B aflatoxins (Creppy 2002). Fumonisins, vomitoxin and zearalenone are well established fusarial mycotoxins (Yazar and Omurtag 2008) and were produced by isolates recovered from sorghum grains (Ayalew et al. 2006). Fumonisin-producing species have been recovered from sorghum grains by several investigators (Da Silva et al. 2000, 2004; Isakeit et al. 2008). Involvement of sorghum grain molding fusaria in producing zearalenone has also been documented (Makun 2007; Hussaini et al. 2009). Vomitoxin is another dominant fusarial toxin found to be produced by several cereal infecting species (Argyris et al. 2003; Hopeetal. 2005). Patulin and citreoviridin are toxic secondary metabolites produce by Penicillium spp. (Wicklow and Cole 1984; Kosemura 2003; Kurtzman and Blackburn 2005). Plant pathogenic penicillia may produce these toxins in culture media, and in agricultural commodities as well as in food waste (Wicklow et al. 1984; Rundberget et al. 2004; Abramson et al. 2009). Production of such toxins by Penicillium species has been frequently reported (Rundberget and Wilkins 2002; Kosemura 2003). Alternaria species are well known to contaminate a wide variety of crops in the field and to cause post-harvest decay of various grains. In addition, several toxic compounds belonging to different structural groups have been documented to produce by Alternaria spp. in cereals (Feng-Gin and Yoshizawa 2000; Li et al. 2001). Dibenzo-a-pyrones or altenuene is one of the most important toxic secondary metabolites produced by A. alternata on sorghum (Sauer et Table 6 Production of altenuene by Alternaria spp. isolated from sorghum seeds Isolates 1 Alternaria alternata 14 2 Alternaria chlamydospora 0 3 Alternaria sp. 49 Altenuene (ppb)

Fungal Diversity (2010) 44:45 52 51 al. 1978), wheat (Magan et al. 1984), linseed and peas (Kralova et al. 2006). Variation in mycotoxin productivity, as well as in the kinds of toxic compound produced, might be due to the probable genetic diversity among and within tested species. In addition, the genetic base of mycotoxin production, genetic signaling pathways and the functions of involved genes have been elucidated in several toxigenic species (Geiser et al. 2001; Kale et al. 2003; Bhatnagar et al. 2006; Brodhagen and Keller 2006). Isolation of these toxicogenic seed-borne fungi from sorghum grain samples should alert plant pathologists to make more effort to minimize the risks of post harvest fungi in storage grains. Rigorous quarantine and healthy storage conditions should be undertaken to minimize fungal contamination and prevent further hazard to human and animal health. 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