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Microbiological and molecular determination of mycobiota in fresh and ensiled maize silage

     Department of Plant Pathology, 321 Buckhout Laboratory, Pennsylvania State University, University Park, Pennsylvania 16802

	ABSTRACT

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The mycobiota of fresh and ensiled maize was studied with culturing techniques and a DNA sequence-based approach. Freshly chopped and ensiled maize were collected for 2 y from 12 farms in Pennsylvania. Samples were plated on selective media and isolates identified by morphology and sequences of the internal transcribed spacer regions of rDNA, 800–900 bp of the 5' end of the translation elongation factor 1-alpha gene and a portion of the rodA gene (Aspergillus fumigatus only). ITS regions were amplified from total silage DNA, cloned, sequenced and compared to fungal ITS sequences in GenBank with the BLAST-N algorithm. For samples analyzed by both methods, the molecular technique detected a greater number of species than selective plating. Plating recovered several Penicillium and Fusarium species and Aspergillus fumigatus, while molecular analysis detected Alternaria, Penicillium and Fusarium species. Data from both methods found that Fusarium and Penicillium were the dominant mycotoxigenic fungi in silage, while yeast made up the majority all fungi recovered or detected. Known mycotoxigenic species often accounted for 50% or more of the total number of species isolated or detected at each site. Viable Fusaria were not isolated from or detected in ensiled maize, suggesting that Fusarium species do not survive the ensiling process. Results from this study suggest that given the numerous species of fungi present in silage with mycotoxin producing ability, there is a strong possibility that silage may be contaminated with multiple toxins simultaneously.

Key words: Alternaria, Aspergillus, corn silage, Fusarium, Gibberella, Penicillium

	INTRODUCTION

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Fungal spoilage of maize silage reduces the nutritional value and palatability of the feed, increases its allergenic potential and may result in mycotoxin contamination (Scudamore and Livesey 1998). Mycotoxigenic fungi in silage are associated with animal health problems such as acute toxicoses, decreased productivity, fertility and increased disease susceptibility (Aghina 1978, D’Mello et al 1999, Scudamore and Livesey 1998, Smith and Lynch 1973, Still et al 1972). Human health also be may affected because dust from molded silages has been implicated as a causal agent in organic dust toxic syndrome, a neurological and respiratory illness (Perry et al 1998).

There are reports on the co-occurrence of mycotoxigenic fungi (Aleksandrov 1986, Baath et al 1990, Khristov 1981, Le Bars and Escoula 1973, Muller 1991) and mycotoxins (Muller 1997, Yu et al 1999) in silage. It also is well documented that many mycotoxigenic fungi are capable of producing more than one toxin simultaneously (Boysen et al 2000, Cole et al 1977, Lepom et al 1990, Melo dos Santos et al 2002). Multitoxin contamination is of particular concern because of potential additive or synergistic effects on animals and humans exposed to molded silages (Riley 1998). Another issue is that some fungal species present in silage have been linked to mycotic infections in cattle, particularly Aspergillus fumigatus (Jensen et al 1992). To date known toxigenic genera reported in silage include Aspergillus, Alternaria, Fusarium and Penicillium. Other commonly isolated fungi include Mucor, Epicoccum, Cladosporium and a variety of yeasts (Aleksandrov 1986, Le Bars et al 1973, Middelhoven and van Baalen 1988, Morrison et al 2001, Scurti 1971).

Mycotoxigenic fungi often are associated with maize either as field or storage organisms. For example many Fusarium spp. are maize pathogens and/or endophytes, while Alternaria spp. are epiphytic saprophytes or weak pathogens (Bilgrami et al 1998). As such toxins produced by these genera usually are considered to be fieldborne problems. Conversely, because Aspergillus and Penicillium often are isolated as postharvest spoilage organisms, their toxins are regarded as a storage concern (Pitt 2002, Scudamore and Livesey 1998). However, under favorable conditions, including hot, dry weather and insect damage, members of both genera and their toxins may be found on maize in the field (Ciegler et al 1970, Jones et al 1980, Marsh and Payne 1984). In ensiled maize the majority of fungal growth is limited by low oxygen content and the production of organic acids by lactic acid bacteria (McDonald et al 1991). However some species, such as Penicillium roqueforti and Aspergillus fumigatus, can survive the microaerophilic low pH environment of silage likely explaining the high reported frequency of toxigenic fungi isolated from ensiled maize (Aleksandrov 1986, Boysen et al 2000, Cole et al 1977, Le Bars et al 1973, Melo dos Santos et al 2002, Pelhate 1977, Vesely et al 1981 ). Fusarium and Alternaria species also may grow and contaminate silage if the low oxygen tension in the feed is not maintained (Aleksandrov 1986, Baath et al 1990, Pelhate 1977).

The development of strategies to reduce toxin contamination of silage will be aided by improved surveys of fungi present. Methods to determine the fungi present in silage usually rely on selective media (Richard-Molard 1986, Skaar and Steinwig 1996). However this approach only isolates viable fungi and may have difficulty recovering those with fastidious growth requirements (Smit et al 1999). Another concern is that selective media may mask the presence of slower growing fungi. Likewise antagonistic interactions among fungi in culture make it difficult to recover sensitive species. An alternative approach is to identify fungi present in silage by characterizing their ribosomal DNA sequences. This approach successfully has detected fungi in environmental samples including soil from the wheat rhizosphere (Smit et al 1999), grasslands (Hunt et al 2004) and arctic tundra soils (Schadt et al 2003). Another benefit of this approach, as illustrated by the work of Schadt (2003), is that it can detect previously unidentified species.

Many studies have been conducted to characterize fungi in maize silage (Aleksandrov 1986, Baath et al 1990, Khristov 1981, Le Bars et al 1973, May et al 2001, Muller 1991, Scurti 1971) but the majority have been conducted in Europe and Asia, making it difficult to directly relate their findings to what might be true in North America. In addition most of these studies relied on culturing techniques, with the exception of May et al (2001). These researchers used denaturing gradient gel electrophoresis on fungal SSU rDNA amplified from total silage to identify fungi in maize silage. However this approach had some difficulty resolving closely related fungal sequences. To provide a more comprehensive assessment of mycoflora in northeastern North American silage, we analyzed fresh and ensiled whole plant maize with two methods, a microbiological evaluation with selective media and morphological identification and a molecular assessment using DNA sequences. This design let us compare our results to earlier work, assess the efficacy of the DNA sequence-based technique and potentially reveal species not previously reported in silage. By examining fresh and ensiled maize we can investigate what fungi are more prevalent in the field vs. those predominant in storage and better understand how the process of ensiling affects the fungal community in maize silage.

	MATERIALS AND METHODS

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Sample collection.— – Maize silage was collected from four diverse geographic regions in 12 counties in Pennsylvania: northeast (NE) counties (Tioga, Susquehanna and Wayne); northwest (NW) counties (Erie, Crawford and Clarion); southeast (SE) counties (York, Lancaster and Berks); and south central (SC) counties (Centre, Blair and Bedford). Samples were collected in 2001 and 2002 from farms with a history of known or suspected mycotoxin contamination. Samples were collected at harvest and then again at the same site 3–6 mo after ensiling. Fresh samples consisted of maize that was collected from a mechanical harvester as it made several passes in a field intended for ensiling. Ensiled maize either was removed from several locations at the face where bunker or trench silos were used or as the silage was being emptied from the silo during "feed out" where upright silos were used. All samples were mixed by hand and 1–3 kg was removed for further study. Samples for molecular analysis were stored at –80 C while those for microbiological assessment were stored at 4 C.

Fungal isolation.— – Microbiological assessment was conducted on freshly harvested and ensiled maize collected in 2001 and 2002 from three farms in each of the four regions. Fungi were isolated with a protocol developed by Skaar and Steinwig (1996) with some modifications. Fifty grams of fresh and ensiled samples were ground with an M-2 model Stein mill (Fred Stein Lab Inc., Atchinson, Kansas) and a 10 g portion of the sample soaked in 50 mL of sterile 0.9% aqueous NaCl solution (wt/vol) for 30 min at room temperature. After incubation the mixture was manually shaken 1 min and filtered through four layers of sterile cheesecloth. The resulting suspension was diluted to 10^–3 with sterile 0.9% NaCl solution, and 100 µL of the diluted solution was spread onto a plate of malt-yeast sucrose agar (MYSA). The media consisted of 15 g malt extract, 2 g tryptone, 2 g oxgall, 30 g sucrose, 5 g yeast extract 20 g of agar (all materials purchased from Difco, Sparks, Maryland), and 0.5 g of sodium nitrate (Sigma-Aldrich, St Louis, Missouri) to which 1 L of water was added. A total of 15 MYSA plates were inoculated for each silage sample. MYSA medium was prepared according to the original protocol (Skaar and Steinwig 1996). Plates were incubated 7 d at room temperature. During this period plates were examined daily and fungal colonies were transferred at first observation to potato-dextrose agar (PDA) (Becton-Dickinson Microbiology Systems, Sparks, Maryland). Plating was done in duplicate for each silage sample where the second plating took place approximately 14 d after the initial plating for a total of 30 MYSA plates. Samples were stored at 4 C between platings.

Because the MYSA method yielded low numbers of Fusarium species, a separate protocol was developed for isolation of these fungi. After grinding (described above) 10 g of silage, the sample was soaked in 50 mL of 50% aqueous Clorox solution for 5 min. The sample was rinsed three times with 100 mL of sterile distilled water for 2 min per wash. Approximately 0.5 g of surface-sterilized silage was placed directly onto a plate containing Nash medium (Nash 1962). Nash medium was prepared with materials purchased from and consisted of 15 g peptone (Difco), 1 g potassium dihydrogen phosphate (Sigma-Aldrich), 0.5 magnesium sulfate *7H₂O, 20 g agar (Sigma-Aldrich), 1 g of pentachloronitrobenezene (Uniroyal Chemical Co. Inc, Middlebury, Connecticut) to which 1 L of water was added. A total of five plates were used for each sample. This procedure was performed twice for a total of 10 plates and a maximum of 20 Fusarium isolates per silage sample. For each sample, with the MYSA and Nash isolation protocols, a total of 100 isolates was collected.

Ensiled samples (n = 12) from 2002 only were analyzed with a method to detect Aspergillus fumigatus that was modified from Melo dos Santos et al (2002). Silage was ground (described above) and a 10 g portion was removed for analysis. One hundred mL of sterile distilled water with 0.01% Tween 20^® (Merck, Darmstadt, Germany) was added to the sample and shaken by hand 1 min. The extract was diluted to 10^–1 and 200 µL spread onto plates of Dichloran Rose Bengal Medium (DRBM) (Horn and Dorner 1998) and grown 5 d at 45 C. The medium consisted of 10 g glucose (Difco), 2.5 g peptone (Difco), 0.5 g yeast extract (Difco), 1 g potassium dihydrogen phosphate (Sigma-Aldrich), 0.5 g magnesium sulfate*7H₂O (Sigma-Aldrich), 20 g agar (Difco), and 25 mg rose bengal to which 1 L of water was added and the pH adjusted to 5.0. Five plates of DRBM were used per silage sample. After incubation 5–20 isolates per sample were collected.

Isolate identification.— – Isolates from the MYSA, Nash, and DRBM plates were inoculated onto PDA, grown 7 d and transferred to appropriate media for identification. Aspergillus and Penicillium species were transferred to Czapek yeast autolysate agar and malt-extract agar (Pitt 2000), while Fusarium species were placed on fresh PDA and carnation leaf agar (Nelson et al 1983). All other fungi were transferred to fresh PDA before identification. Isolates were identified morphologically with reference cultures (listed in TABLE I) and identification guides (Barnett and Hunter 1998, Klich and Pitt 1988, Nelson et al 1983, Pitt 2000).

Fungal isolates were grown in 10 mL of potato-dextrose broth (Becton Dickinson Microbiology Systems), shaking at 150 rpm at room temperature for 7 d. Mycelia were harvested by filtering the culture and scraping the material into a 1.7 mL centrifuge tube. Yeast cells were collected by centrifugation (Sorvall RT7 centrifuge with RTH-750 rotor; Thermo Electron Corp., Boston, Massachusetts) of the cultures at 6000 x g for 5 min, decanting the media and resuspending the cells in 1 mL of sterile distilled water. This suspension was transferred to a 1.7 mL centrifuge tube, spun at 13 000 x g for 1 min (Heraeus Biofuge pico centrifuge, Thermo Electron Corp.) and the water decanted. After harvest fungal isolates were lyophilized in an ADVantage Freeze Dryer (VirTis, Gardiner, New York). Cultures of known mycotoxigenic species are available on request.

DNA was extracted with the DNeasy Plant MiniKit (QIAGEN, Valencia, California). Isolates for DNA sequence analysis then were subjected to the polymerase chain reaction (PCR) in 50 µL reactions with 1 µL of sample DNA, and a final concentration of 0.2 µM of each primer, 0.2 mM dNTP mixture, 0.1 x PCR buffer (0.5 M KCl, 0.1 M TrisHCl pH 8.3 and 0.025 M MgCl₂) and 0.01 U/µL of Taq polymerase (Promega, Madison, Wisconsin). All reactions were carried out in a PTC-200 thermocycler (MJ Research, Waltham, Massachusetts). The PCR and sequencing primers used were ITS5 and ITS4 (White et al 1990) for the ITS region, EF1 and EF2 for the translation elongation factor gene (O’Donnell et al 1998) and rodA1 and rodA2 for the RodA gene (Geiser et al 1998). The PCR amplification conditions were: for ITS (i) 96 C for 1 min, (ii) 55 C for 30 s, (iii) 72 C for 1 min, 30 cycles of (iv) 94 C for 30 s, (v) 55 C for 20 s, (vi) 72 C for 1 min and a final polymerization step of (viii) 72 C for 2 min. The amplification conditions for TEF analysis were: (i) 95 C for 1 min, followed by 35 cycles of (ii) 94 C for 1 min, (iii) 53 C for 1 min, (iv) 72 C for 1 min, followed by a final polymerization step of (v) 72 C for 5 min. The amplification conditions for the RodA gene were: (i) 94 C for 2 min, followed by 35 cycles of (ii) 94 C for 1 min, (iii) 56 C for 1 min, (iv) 72 C for 1 min, followed by a final polymerization step of (v) 72 C for 2 min.

PCR products were cleaned up for DNA sequencing with the QIAquick^® PCR Purification Kit (QIAGEN). Sequencing reactions were performed with the BigDye^® Terminator v3.1 (Applied Biosystems, Foster City, California) kit according to the manufacturer’s instructions, except that reaction volumes were scaled down to 8 µL. Sequence reactions were purified with Performa^® DTR 384-Well Plates (Edge BioSystems, Gaithersburg, Maryland). Sequence analysis was performed on a 3730 DNA Analyzer (Applied Biosystems).

Genomic DNA extraction and purification from maize silage.— – For molecular analysis, one sampling site was selected from each of the four regions from which one fresh and one ensiled sample from 2001 and 2002 was analyzed, for a total of 16 samples. Fifty grams of maize silage was ground as described above and 10 g of sample removed for DNA extraction. Total genomic DNA was extracted with the UltraClean^TM Mega Prep Soil DNA Kit (MoBio Laboratories Inc., Solona Beach, California). DNA was purified further with Genomic-tip 20/G DNA clean-up columns (QIAGEN).

PCR amplification and purification.— – PCR was performed with purified total genomic DNA from silage, with primers ITS5 and ITS4 (White et al 1990) and the high fidelity PfuTurbo^® DNA Polymerase (Stratagene, La Jolla, California). The remaining reaction reagents and conditions are described above. A total of 10 amplification reactions were performed for each sample. After amplification the reactions were pooled and subjected to electrophoresis in a 1.5% agarose gel in 1 x TAE buffer (Ausubel 1999). The gel was viewed with ethidium bromide and the ITS fragments (a smear 400–900 bp) were excised manually from the gel with a scalpel and purified using QIAquick^® Gel Extraction Kits (QIAGEN).

Generation of ITS clone library.— – Purified ITS fragments were ligated into the pGEM^®-T Easy vector (Promega) according to the manufacturer’s instructions and recombinant plasmids were transformed into competent JM106 Escherichia coli cells (Promega). One hundred µL of undiluted transformed cells were plated onto Luria-Bertani (LB) medium containing 100 µg/mL ampicillin (Shelton Scientific, Shelton, Connecticut), 40 µg/mL X-gal (Inalco, Milano, Italy) and 0.5 mM IPTG (Inalco) for blue-white screening. Clones included in the libraries were selected by picking white colonies with a sterile toothpick and inoculating wells in a 96-well microtiter dish (BD Biosciences, Bedford, Massachusetts) containing 200 µL of LB broth amended with 100 µg/mL ampicillin. Cultures were incubated overnight at 37 C with shaking at 100 rpm. Libraries were preserved for long-term storage by adding 60 µL of 50% glycerol (Shelton Scientific) before storage at –80 C.

Clone library analysis.— – Preliminary sequencing analysis revealed that the majority of the population of each library consisted of several yeast species. To avoid repetitively sequencing clones of a few species, a hybridization method was used to identify clones of interest. To determine predominant yeast species in each library, plasmid DNA was extracted from 24 randomly selected clones with the QIAprep^® Miniprep kit (QIAGEN) and sequenced using the SP6 or T7 primer according to the manufacturer’s instructions. Based on the resulting data, yeast sequences, found to make up 25% or more of the library, were chosen for selective screening to avoid repetitious sequencing. Template DNA for the hybridization probes was generated by growing clones of the selected yeast species and extracting their plasmid DNA as noted above. EcoR1 restricted plasmid DNA was run on a 1.0% agarose gel and the 400–900 bp ITS insert band manually excised and purified as described above. After template DNA purification, the NEBlot^® Phototype Kit (New England Biolabs, Beverly, Massachusetts) was used to generate biotinylated ITS probes for colony hybridization with the manufacturer’s protocol.

Southern blots were prepared by stamping each library with a 96-well replicating tool onto a Hybond-N+ nylon membrane (Amersham Pharmacia Biotech Inc., Piscataway, New Jersey) set on a plate of ampicillin-amended LB media (100 µg/µL). Membranes were incubated overnight at 37 C. After incubation, membranes were soaked 1 min in 10% aqueous sodium dodecyl sulfate solution (SDS) and washed. Membranes then were prepared according to the NEBlot^® Phototope^® Kit instructions for colony hybridizations.

Membranes were prehybridized at 68 C for 1 h in a prehybridization solution and hybridization was performed immediately afterward under the same conditions with approximately 20 ng/mL of probe added to the prehybridization solution according to the NEBlot^® Phototope^® Kit instructions. The membrane was hybridized overnight, washed and incubated in CDP-Star reagent (New England Biolabs) per the Phototope^®-Star Detection Kit manual. The treated membranes were exposed 5 min to BioMax Light Audioradiography film (Eastman Kodak, Rochester, New York) and the film developed according to company instructions. Membranes were stripped and stored according to the Phototope^®-Star Detection Kit instructions.

After screening the libraries, 96 clones that did not hybridize with any of the targeted yeast species were selected for sequencing analysis. Selected clones were grown overnight and subjected to plasmid extraction, sequencing and data analysis as described above.

	RESULTS

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Microbiological assessment.— – Using a combination of three plating methods, we isolated six species of Penicillium (P. roqueforti, P. paneum, P. expansum, P. crustosum, P. commune and P. citrinum) seven species of Fusarium (F. avenaceum, F. culmorum, F. graminearum, F. pseudograminearum, F. proliferatum, F. sporotrichioides and F. verticillioides; FIG. 1) and one species of Aspergillus, A. fumigatus. In terms of species abundance, P. roqueforti and F. graminearum were the most commonly isolated species known to be mycotoxigenic. P. roqueforti was isolated from 50% of the fresh (n = 24) and 75% of the ensiled (n = 24) samples, while F. graminearum was found in 58% of the fresh samples and in none of the ensiled samples. P. paneum, which is closely related to P. roqueforti and only recently defined as a separate species (Boysen et al 2000), was isolated from two fresh (n = 24) and four ensiled samples (n = 24). The third member of the P. roqueforti group, P. carneum, was not isolated from any samples.

View larger version (11K): [in this window] [in a new window]   FIG. 1. Mycotoxigenic species isolated by plating from maize at harvest and after ensiling. Isolations were made from fresh and ensiled silage plated on to MYSA and Nash media.

In addition to the mycotoxigenic species isolated from silage, several other filamentous and yeast species were recovered as well. Filamentous fungal genera isolated included Acremonium, Cladosporium, Cordyceps, Epicoccum, Mortierella and Mucor (TABLE II). In general yeast made up the majority of the fungi isolated from silage. Geotrichum candidum was the most frequently encountered species identified by sequence and was isolated in 75% of the fresh samples (n = 24) and 21% of the ensiled samples (n = 24). Other yeasts that occurred at relatively high frequency included Candida intermedia, Candida sake, Debaryomyces hansenii, Issatchenkia orientalis, Pichia anomala, Pichia fermentans and Pichia membranifaciens.

For silage samples analyzed by plating and DNA sequences, molecular analysis detected at least twice the number of species identified by plating. Data from plating and molecular analysis revealed that mycotoxigenic fungi often made up 50% or more of the species isolated or molecularly detected from fresh and ensiled samples at each site (FIG. 2a, b). However we found that in general species diversity decreased as a result of ensiling (FIG. 3). Ten out of 12 sites examined by plating had fewer fungal species in their ensiled samples compared to those from fresh (FIG. 3a), whereas all four sites analyzed with the molecular technique had fewer species in their ensiled samples (FIG. 3b). Of note, we observed that while Fusarium and Alternaria species could be isolated or detected from fresh samples none were found in ensiled material (FIG. 1). In contrast several Penicillium species, including P. roqueforti, P. paneum, P. crustosum and P. commune, were found in both ensiled and fresh samples.

View larger version (28K): [in this window] [in a new window]   FIG. 2. Number of toxigenic and nontoxigenic species isolated from each site. Comparison of the number of toxigenic and nontoxigenic fungal species isolated by A. plating and B. molecular detection from maize at harvest and after ensiling. The asterisks under the samples in A correspond to samples analyzed with the molecular technique in B.

View larger version (25K): [in this window] [in a new window]   FIG. 3. Number of fungal species identified from study sites in fresh and ensiled maize. A. Number of fungal species isolated by plating and B. detected by the molecular method from fresh and ensiled maize. The asterisks under the samples in 2A correspond to samples analyzed with the molecular technique in 2B.

	DISCUSSION

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Because the majority of the Fusarium species encountered in this study are known pathogens or endophytic inhabitants of maize (Payne 1999), their presence in the fresh samples was not unexpected. In particular F. graminearum, the most frequently isolated Fusarium species from fresh samples, is one of the most commonly encountered maize pathogens in North America (Payne 1999) and it is reported to be a late season colonizer of maize stalk tissue (Windels and Kommendahl 1984), which might explain its high frequency in the fresh samples. We did find it surprising that F. verticillioides was not isolated with greater frequency (isolated from only four of 24 samples), although it was detected in six of the eight fresh samples analyzed with the molecular technique. F. verticillioides, like F. graminearum, is a pathogen of maize (Payne 1999) but also can be isolated with relatively high frequency from symptom-less kernels, stalks and leaves (Bacon et al 1996).

Consistent with Aleksandrov (1986), Auerbach et al (1998), Le Bars et al (1973), Pelhate (1977) and Vesely et al (1981), we found P. roqueforti to be the most commonly isolated mycotoxigenic fungus in ensiled samples and the second most frequently isolated from fresh samples. Although A. fumigatus also was recovered we did not isolate it as frequently as P. roqueforti. This might be due in part to unfavorable culturing conditions before the addition of the DRBM method, although other methods have successfully isolated A. fumigatus onto nonselective media at 25 C (Cole et al 1977). Therefore it might be that A. fumigatus was simply not present as abundantly as P. roqueforti in silage from this study. Similar to observations of Boysen et al (2000), we did recover P. paneum from maize silage but at a lower frequency than P. roqueforti. Likewise we did not isolate or detect P. carneum, the third member of the P. roqueforti complex, in any of the silage samples. The isolation of P. roqueforti and P. paneum is expected because their ability to survive ensiling is well documented (Auerbach et al 1998, Boysen et al 2000). Penicillium expansum also was isolated in this study, and like P. roqueforti and P. paneum it is reported to tolerate a low oxygen concentration and the presence of organic acids (Pitt and Hocking 1997), which likely explains its presence in silage (Boysen et al 2000).

Of interest, no viable Fusarium species were isolated from ensiled samples. The inability of Fusarium to persist in ensiled maize was observed by Golosov et al (1967) and suggests that they are not capable of surviving the low oxygen, low pH environment of silage. The optimal pH for growth of two commonly isolated Fusarium species from maize, F. proliferatum and F. verticillioides, are respectively 5.5 and 7.0 (Marin et al 1995), while the mean pH of the silage samples in our study was 3.7 (Mansfield and Kuldau unpublished). Low pH in combination with low oxygen tension likely contributes to the overall inability of Fusarium species to persist in silage. As such the majority of Fusarium mycotoxins in silage probably are produced in the field and strategies to reduce Fusarium mycotoxins in silage should focus on the crop before ensiling. This is particularly important because although ensiling may successfully eliminate fusaria from maize many of their toxins, such as the trichothecenes and zearalenone, are stable in storage (Lepom et al 1990, 1991). Therefore once formation has occurred toxin levels in the silo may not drop appreciably before consumption. However based on our observations the concentration of one trichothecene, deoxynivalenol does decrease from harvest to approximately 6 mo after ensiling (Mansfield et al 2005). One possible explanation for this phenomenon is that microbial agents within silage are capable of degrading or binding this toxin (El-Nezami et al 2002, He et al 1992). Although Fusarium spp. are not tolerant of conditions during ensiling, most Penicillium isolated in this study were found both in fresh and ensiled maize, so it is possible that toxin formation by these fungi might occur in the field and during storage. Toxin production by Penicillium in the field is perhaps less common than it is for Fusarium because most Penicillium spp. are cosmopolitan saprophytes rather then pathogens. However toxins may be produced in the field if damaged kernels are colonized (Ciegler et al 1970) in a disease known as "blue-eye" on maize (Payne 1999). Most Penicillium species grow well in low water activity, which is why they usually are considered postharvest spoilage organisms and their toxins are generally produced in maize after ensiling (Pitt 2002, Scudamore and Livesey 1998). Consequently, because Penicillium species require some oxygen to proliferate and produce toxins, prevention of toxin production during storage should focus on maintaining the anaerobic conditions of the feed.

Although mycotoxigenic fungi accounted for 50% or more of the fungal species isolated from each of the study sites, the majority of the fungal population consisted of yeast. This also was also true for the clone libraries, which in many cases were made up almost entirely by three or more species of yeast. Middlehoven and van Baalen (1988) isolated several predominant species in ensiled maize including Issatchenkia orientalis, Geotrichum candidum, Pichia anaomala and Pichia fermentans. They found that while some species, such as the basidiomycete yeast Cryptococcus laurentii (detected in this study by molecular analysis in fresh samples only), did not persist after ensiling the species listed above were present for the duration of the 122 d study. May et al (2001) also noted that yeasts, including P. anomala, were the dominant fungi present 2–3 mo after ensiling. Of interest, Middlehoven and van Baalen also noted that when the silage was exposed to oxygen P. fermentans, P. anomala and G. candidium all were able to assimilate lactic and acetic acids from the feed (1988). By degrading organic acids, yeast have been suggested to precondition silage for colonization by other fungi when anaerobic conditions are not maintained (Melo dos Santos et al 2002). Contamination of silage with high populations of yeast also may be problematic because species such as G. candidum can produce unpleasant odors that cause the cattle to reject the feed (Scudamore and Livesey 1998).

The results from this work confirm that multiple mycotoxigenic species can occur in a single whole-plant maize sample both at harvest and after ensiling and suggest that a mixture of toxins can be present simultaneously (Baath et al 1990, Le Bars et al 1973, Muller 1997, Scurti et al 1971). Many of these toxins, including several trichothecenes, zearalenone, AAL-toxin TA, and those produced by Penicillium roqueforti and P. paneum, have been detected in maize silage (Boysen 2000, Mansfield and Kuldau unpublished, Muller 1997, Yu 1999). It is likely that the other toxins also are present but they have yet to be investigated. For example, although several studies have cultured A. fumigatus from silage and tested the ability of isolates to produce mycotoxins (Cole et al 1977, Melo dos Santos et al 2002, Smith et al 1973), as far as we are aware no studies have been conducted to directly detect these toxins in silage. Data from this study provide additional guidance as to what mycotoxins may be expected in northeastern North American silages and invites further research toward their detection. In addition by expanding our understanding of what fungi occur in North American silages more targeted management strategies can be developed to prevent fungal and mycotoxin contamination in the field and during storage.

	ACKNOWLEDGMENTS

 
This research was financed in part by agricultural research funds administered by the Pennsylvania Department of Agriculture. We acknowledge participating dairies as well as Pennsylvania State University Extension Educators and representatives from Agway Inc., Pennfield Inc. and F.M. Brown & Sons Inc. Thanks to Brian Dombroski for assistance with sample collection. Thanks to Drs Kelly Ivors and Seogchan Kang for technical advice; and thanks to Dr David Geiser for the RodA primers. Special thanks to Dr Steve Peterson, Dr David Geiser and Jean Juba for providing reference cultures and Dr K. O’Donnell for sample analysis. Thanks to Drs Geiser and Kang for critical review of this manuscript.

	FOOTNOTES

 
Accepted for publication December 21, 2006.

¹ Corresponding author. E-mail: kuldau{at}psu.edu

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