RESEARCH ARTICLE
Culturable fungal assemblages growing within Cenococcum sclerotia in forest soils Keisuke Obase1, Greg W. Douhan2, Yosuke Matsuda3 & Matthew E. Smith1 1
Department of Plant Pathology, University of Florida, Gainesville, FL, USA; 2Department of Plant Pathology and Microbiology, University of California, Riverside, CA, USA; and 3Laboratory of Forest Pathology and Mycology, Graduate School of Bioresources, Mie University, Tsu, Mie, Japan
Correspondence: Keisuke Obase, Department of Plant Pathology, University of Florida, 2523 Fifield Hall, Gainesville, FL 32611-0680, USA. Tel.: +13522262181; fax: 13523926532; e-mails: [email protected], [email protected] Received 8 August 2014; revised 11 September 2014; accepted 12 September 2014. Final version published online 30 September 2014.
MICROBIOLOGY ECOLOGY
DOI: 10.1111/1574-6941.12428 Editor: Ian C. Anderson Keywords Cenococcum geophilum; decomposition; mycoparasite; root endophyte; pathogen.
Abstract The ectomycorrhizal fungus Cenococcum geophilum (Ascomycota, Dothideomycetes) forms black, round to irregular sclerotia in forest soils. Fungi that colonize the sclerotia appear to affect sclerotia viability and may play an important role in the life history of Cenococcum. Some of the fungi could also affect nutrient cycling by decomposing Cenococcum sclerotia, which are melanized and recalcitrant to decay. We used a culture-based method to document the fungal communities growing inside surface-sterilized sclerotia that were collected from forest soils. Cenococcum was successfully isolated from 297 of 971 sclerotia whereas 427 sclerotia hosted fungi other than Cenococcum. DNA barcoding of the internal transcribed spacer rDNA followed by grouping at 97% sequence similarity yielded 85 operational taxonomic units (OTUs) that consisted primarily of Ascomycota (e.g. Chaetothyriales, Eurotiales, Helotiales, Pleosporales) and a few Basidiomycota and Mucoromycotina. Although most fungal OTUs were infrequently cultured, several OTUs such as members of Asterostroma, Cladophialophora, Oidiodendron, and Pleosporales were common and found across many sites. Our results suggest that Cenococcum sclerotia act as a substrate for diverse fungi. The occurrence of several OTUs in sclerotia across many sites suggests that these fungi may be active parasites of Cenococcum sclerotia or may preferentially use sclerotia as a nutrient source.
Introduction Sclerotia are hard, compact masses of fungal mycelium that usually form in soil or plant tissue. They are thought to serve as resting structures that can survive and remain quiescent in adverse environmental conditions until circumstances become favorable for fungal growth (ColeySmith & Cooke, 1971; Willetts, 1971). Sclerotia of many fungal species, including Cenococcum geophilum (Massicotte et al., 1992), Rhizoctonia solani (Naiki & Ui, 1975), and Sclerotinia minor (Bullock et al., 1980) have thick, pigmented fungal cell walls that can reduce colonization by other microorganisms and aid in their survival in the environment (Hurst & Wagner, 1969; Malik & Haider, 1982). Other fungi, such as Rhizoctonia spp. (Burton & Coley-Smith, 1985) and Aspergillus spp. (e.g. Wicklow & Cole, 1982; Frisvad et al., 2014), produce antibiotic compounds to resist attack by microorganisms. However,
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sclerotia are often infected by a number of soil fungi that may enter via small cracks on the surfaces of the sclerotia or through active attack by fungal pathogens (Makkonen & Pohjakallio, 1960; Naiki & Ui, 1975; Huang & Hoes, 1976; Gladders & Coley-Smith, 1980; Willetts & Wong, 1980; Alexander & Stewart, 1994). Although the interactions between sclerotia and sclerotia-associated fungi have not been fully explored, several studies have shown that sclerotia germination is suppressed by some sclerotia-associated fungi (Gladders & Coley-Smith, 1980; Zazzerini & Tosi, 1985). Moreover, some mycoparasitic fungi, such as Trichoderma harzianum, actually degrade sclerotia cell walls and exploit the cytoplasmic contents of the attacked cells (Elad et al., 1984). Consequently, infection by sclerotia-associated fungi could be an important biological factor that may reduce the viability of sclerotia and therefore negatively impact the fitness of sclerotia-forming fungi such as
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C. geophilum. Sclerotia-associated fungi may also affect nutrient cycling by enhancing the decomposition rate of sclerotia, which otherwise remain recalcitrant to decay (Fernandez et al., 2013). The fungal biota that colonize sclerotia and the rate of infection have been investigated for several plant pathogenic fungi (e.g. Makkonen & Pohjakallio, 1960; Huang, 1985) with the idea that these sclerotia-associated fungi may be useful for biocontrol (Tu, 1984; Whipps & Gerlagh, 1992). However, there have been no studies of the fungal communities associated with the sclerotia of nonpathogenic fungi. The sclerotia of C. geophilum (Fig. 1) are 0.5–4.0 mm in diameter, spherical with a shiny smooth surface, and frequently found in soils of pine and oak forests (e.g. Trappe, 1962; Jany et al., 2002; Douhan & Rizzo, 2005; Obase et al., 2010). Cenococcum geophilum is one of the most common ectomycorrhizal fungi in the world and this fungus forms symbiotic associations with a wide variety of woody host plants (Trappe, 1969; Massicotte et al., 1992; LoBuglio, 1999). Previous studies indicate that C. geophilum is a monophyletic species complex, but the cryptic species within this group are not distinguishable without molecular methods (LoBuglio et al., 1991, 1996; Douhan & Rizzo, 2005; Douhan et al., 2007). Cenococcum sclerotia consist of a well-differentiated rind and a medulla with pseudoparenchymatous organization composed of melanized fungal mycelium (Massicotte et al., 1992). The longevity of Cenococcum sclerotia in forest soils has not been explicitly tested, but several lines of evidence suggest that sclerotia are long-lived and important in the biology and life history of Cenococcum (Trappe, 1962; Shaw & Roy, 1983; Miller et al., 1994). First, Cenococcum is ubiquitous both as sclerotia and on ectomycorrhizal roots in a wide variety of forest habitats (Trappe, 1962; LoBuglio, 1999). Second, Cenococcum has no known sexual or asexual spore-producing structures and is believed to be an ancient asexual fungal lineage (Douhan & Rizzo, 2005; but see Fernandez-Toiran & Agueda, 2007). Third, sclerotia can be surface-sterilized and grown as weak saprotrophs in axenic culture and are thought to act similarly when they contact nutrient resources in nature (Trappe, 1962). The high abundance and resilience of Cenococcum sclerotia in forest soils also means that these structures may sequester carbon and other compounds which could therefore impact nutrient cycling (Watanabe et al., 2001, 2007). Trappe (1962) showed that dead Cenococcum sclerotia persist intact for several years in soil, suggesting that they likely remain intact for long periods of time and sequester nutrients, so that they are unavailable to other organisms. Several studies have measured the size and abundance of Cenococcum sclerotia and found that these structures represent a massive amount of fungal tissue in FEMS Microbiol Ecol 90 (2014) 708–717
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many forests: 440 kg ha1 in an old-growth Norway spruce forest (Dahlberg et al., 1997), 2785 kg ha1 in a second-growth Douglas-fir stand (Fogel & Hunt, 1979), and 2300–3000 kg ha1 in stands of Abies amabilis (Vogt et al., 1981). This evidence suggests that understanding the diversity and community structure of microbial parasites or decomposers of Cenococcum sclerotia may provide insights into the ecology of Cenococcum and also nutrient cycling in forest soils. To assess the fungal diversity and community structure of sclerotia-associated fungi, we collected sclerotia of C. geophilum from ten sites in mixed oak–pine forests in Florida and Georgia. We isolated all culturable fungi from surface-sterilized Cenococcum sclerotia on nutrient agar media and then used DNA sequencing of the internal transcribed spacer (ITS) rDNA region to identify the sclerotia-associated fungi. In some cases, ITS was insufficient to accurately identify the sclerotia-associated fungi, so we also sequenced a portion of the 28S rDNA to refine our phylogenetic placement of these fungi.
Materials and methods Site descriptions and sampling procedures
Between August and October 2013, soil samples (c. 7 9 7 9 10 cm) were collected from ten different forest sites in Florida and Georgia, USA (Fig. 2). All study sites were natural or managed forest ecosystems dominated by species of Quercus and Pinus except for a single sampling site (PO) in an orchard of cultivated Carya illinoinensis (Wangenh.) K. Koch. The closest forest sites were roughly 2 km apart whereas the most distant sites were 170 km apart. At each individual forest site, we selected 2–10 locations, which were 2–500 meters apart each other, and collected one soil sample beneath 1–4 trees spread over an area of 1–25 m2 at each location. Climatic data from the Gainesville weather station during 2012 (Gainesville Regional Airport; 29°410 24″N, 82°160 18″ W; 46.0 m asl) indicated annual precipitation of 56.25 inches (c. 1428.7 mm) and mean temperature of 21.6 °C, ranging between 14.1 °C in January and 28.6 °C in July. All samples were stored in plastic bags at 8 °C for less than a week before isolations were made. Sampling sclerotia and isolation of fungi
Forest soils were washed in running tap water over a 2.0-mm sieve to remove excess plant debris and soil particles. Filtered soil slurry (including both small sclerotia (0.5–2.0 mm in diameter) and larger sclerotia > 2.0 mm in diameter) was taken from the sieve, stirred vigorously in a glass beaker filled with water, and then left standing ª 2014 Federation of European Microbiological Societies. Published by John Wiley & Sons Ltd. All rights reserved
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(a)
(b)
(c)
Fig. 1. (a) Ectomycorrhizas of Cenococcum geophilum with sclerotia (arrows) attached via hyphal connections. (b) Hyphal growth of C. geophilum on MMN agar media. (c) Closeup view of C. geophilum sclerotium.
Fig. 2. Ten study sites where Cenococcum sclerotia were collected in Florida and Georgia, USA.
for 10 s. Floating plant debris was removed by decanting. The remaining slurry was spread on a glass dish (9 cm in diameter) and observed under a dissecting microscope. Sclerotia that were black, c. 0.5–3.0 mm in diameter, round, and sank in water were considered likely to be viable C. geophilum sclerotia based on the findings of Trappe (1969) and Douhan & Rizzo (2005). Forceps were used to transfer these putatively viable sclerotia to a moist kimwipe paper towel and the kimwipe was then used to vigorously scrub the sclerotia to remove adhering soil particles. Sclerotia that broke apart during the scrubbing stage were discarded and excluded from further analysis. Putatively viable sclerotia were then transferred to a ª 2014 Federation of European Microbiological Societies. Published by John Wiley & Sons Ltd. All rights reserved
50-mL tube with 20–30 mL water and shaken vigorously by hand for 30 s. All sclerotia were transferred to a glass dish and observed for a second time under the dissecting microscope. Soil particles still adhering to the sclerotia were then removed by hand with clean foreceps. Thirtynine sclerotia that floated in water were also collected from two study sites (LW and SW) and subjected to the same treatment as above to test whether floating sclerotia were more likely to be parasitized than sinking sclerotia. Prior to culturing, we surface-sterilized sclerotia by immersing them for 20 min in commercially available bleach (containing 2% chlorine) followed by two rinses in c. 20 mL of sterilized distilled water. Following sterilization FEMS Microbiol Ecol 90 (2014) 708–717
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and rinsing, we used flame-sterilized forceps to transfer sclerotia to a new, sterile kimwipe to remove excess water. Dry, surface-sterilized sclerotia were then axenically transferred to 90-mm petri dishes containing modified Melin– Norkran’s (MMN) media (Marx, 1969) with streptomycin (100 mg mL1) and chloramphenicol (50 mg mL1). Petri dishes were incubated at 23 °C. Fungi that grew from sclerotia in the first 2 days after incubation were considered likely soil contaminants that occurred due to incomplete surface sterilization and therefore discarded. Fungal isolates derived from one fungal species and emerging after 3 days were subcultured and retained in the analysis. Culture dishes inhabited by two or more species were discarded due to the difficulties in separating intermingled fungal colonies.
Fungal isolates were classified into operational taxonomic units (OTUs) based on 97% similarity of the ITS sequences. All unique ITS, ITS + LSU, or gpd sequences were subjected to BLAST searches (Altschul et al., 1997) against GenBank (http://blast.ncbi.nlm.nih.gov). Sequences obtained from this study were aligned with identified fungal sequences using MAFFT (Katoh & Standley, 2013). Maximum Likelihood (ML) analyses were performed with bootstrap analysis with 100 replicates with MEGA5 (Tamura et al., 2011), to verify our phylogenetic identification of the isolates based on BLAST searches. The general time-reversible model incorporating invariant sites and a gamma distribution (GTR + I + G) was selected as the appropriate model based on the results of the corrected Akaike information criterion value.
DNA extraction and identification of fungi
Statistical analysis
DNA was extracted from fresh mycelium of each fungal isolate with the Extract-N-Amp kit (Sigma-Aldrich) according to the manufacturer’s instructions. We amplified the ITS rDNA region (ITS1-5.8s-ITS2) using Taq DNA polymerase (New England BioLabs) with a fungispecific primer ITS1F (Gardes & Bruns, 1993) and a universal primer ITS4 (White et al., 1990). For isolates that morphologically matched C. geophilum, we verified their identity by amplifying and sequencing the glyceraldehyde 3-phosphate dehydrogenase (gpd) gene region with primers gpd1 and gpd2 (Berbee et al., 1999). We have determined that sequencing the gpd locus is the most effective way to verify the identity of Cenococcum isolates because gpd was the most phylogenetically informative DNA region examined by Douhan & Rizzo (2005). Unfortunately, many non-Cenococcum isolates could not be readily identified to class or order level based on ITS sequences alone. To refine the identification of these isolates, we also amplified partial LSU sequences from one representative culture of each OTU using primers ITS3 (White et al., 1990) and LR5 (Vilgalys & Hester, 1990). PCR followed the amplification conditions of Obase & Matsuda (in press) for ITS, Duong et al. (2012) for LSU, and Douhan et al. (2007) for gpd except that we used 44 instead of 25 cycles and an annealing temperature of 58 °C instead of 55 °C. Positive PCR amplicons were cleaned with EXO and SAP enzymes (Glenn & Schable, 2005), and then sequenced bidirectionally with the same primers as above with the BigDye Terminator v. 3.1 Cycle Sequencing Kit on an ABI3700 DNA sequencer (Applied Biosystems). The fungal ITS, LSU and gpd sequence data were submitted to Genebank under accession numbers AB986283–AB986537. All of the fungal DNA sequences obtained in this study are detailed in Supporting Information, Tables S1 and S2.
Kruskal–Wallis analyses were conducted to compare the isolation ratio of non-Cenococcum fungi among study sites. Study sites that included more than three soil samples and had more than 10 sclerotia were included in the analysis (Table S3). Differences among study sites were determined using the Mann–Whitney pairwise post hoc tests (P < 0.05). An analysis of similarity (ANOSIM) was performed to compare the biota of non-Cenococcum fungi among study sites. The ANOSIM were computed with 9999 permutations and based on presence/absence data using the Raup–Crick similarity index (Raup & Crick, 1979) or abundance data using the Bray– Curtis similarity index (Bray & Curtis, 1957). Twelve soil samples, from four study sites that included more than seven non-Cenococcum isolates were subjected to the analysis. The analyses were performed using PAST (Hammer et al., 2001). OTU accumulation curves were drawn by plotting the mean of the accumulated number of expected OTUs in pooled soil samples after 1000 randomizations without replacement, using ESTIMATES version 9.1.0 (Colwell, 2013) to estimate whether the sampling effort was sufficient to fully describe the community of sclerotia-associated fungi. Estimated OTU richness was calculated using the Chao 2 estimator.
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Results Isolation success for Cenococcum and nonCenococcum sclerotia-associated fungi
No sclerotia were present in 22 soil samples but the 33 remaining soil samples each contained 1–66 Cenococcum sclerotia (Table S3). In total, 971 sclerotia were collected from 1 to 9 soil samples from each study site and subjected to our isolation procedures. Thirty sclerotia (3.1%) were rapidly contaminated in the first 2 days with fastgrowing fungi and removed from further analysis. ª 2014 Federation of European Microbiological Societies. Published by John Wiley & Sons Ltd. All rights reserved
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Cenococcum geophilum was successfully obtained from 297 sclerotia (31.6%) whereas other fungi were recovered in 427 (45.4%) of 941 total sclerotia. There was no fungal occurrence from the remaining 217 sclerotia (23.1%). When soil samples that included < 10 sclerotia were removed from the analysis, the isolation success rate for Cenococcum ranged from 0 to 80.4% (Ave. SD; 30.0 19.8%) and the isolation success rate for nonCenococcum fungi ranged from 0 to 95.2% (Ave. SD; 47.8 26.2%). The isolation success rate for non-Cenococcum fungi was significantly different between study sites (Kruskal–Wallis analysis; H = 17.2, P = 0.016). However, there were no significant differences in the isolation success rate of C. geophilum (H = 12.3, P = 0.091) between sites. There was also no difference in the number of sclerotia from which no fungi were isolated between sites (H = 7.83, P = 0.34) (Fig. 3). Identification of sclerotia-associated fungi
Six hundred and sixty-seven of 724 isolates (92.1%) were successfully identified based on ITS and/or LSU sequences (for non-Cenococcum fungi) or based on a combination of ITS and/or gpd sequences (for C. geophilum). Of these, 297 isolates were identified as C. geophilum whereas the remaining 370 non-Cenococcum isolates were classified into 85 different OTUs.
Fig. 3. Average success rate for isolation of Cenococcum geophilum (filled black) and non-Cenococcum fungi (filled gray) from sclerotia in each study site. Proportions of sclerotia where no fungal isolates were obtained are indicated in white. Two study sites (FW and PO) were removed from the analysis due to low numbers of soil samples that contained Cenococcum sclerotia. Different letters indicate significant differences at P < 0.05 based on the Mann–Whitney pairwise post hoc tests.
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BLAST searches revealed that 313 isolates (81 OTUs) were Ascomycota, 56 isolates (3 OTUs) were Basidiomycota, and one isolate was a member of Mucoromycotina (Fig. 4). Isolates that were identified as belonging to the Ascomycota consisted of Eurotiomycetes (91 isolates in 25 OTUs representing Eurotiales and Chaetothyriales), Dothideomycetes (93 isolates in 22 OTUs representing Pleosporales and Venturiales), Leotiomycetes (68 isolates in 20 OTUs representing Helotiales sensu lato), Sordariomycetes (54 isolates in eight OTUs representing Hypocreales, Sordariales, and Xylariales), and other Pezizomycotina (seven isolates in six OTUs). All Basidiomycota isolates belonged to the Agaricomycetes (56 isolates in 3 OTUs representing Hymenochaetales and Russulales). One Umbelopsis OTU was the sole member of the Mucoromycotina.
Community structure of sclerotia-associated fungi
The accumulation curve for OTUs did not reach an asymptote (Fig. S1) and estimated a total community of 436 OTUs of sclerotia-associated fungi by the Chao2 estimator, indicating that many rare sclerotia-associated fungi would likely be identified with further sampling effort. The Asterostroma (54 isolates) and Sordariales A (45 isolates) OTUs were the most frequently isolated sclerotia-associated fungi followed by Pleosporales A (n = 36), Oidiodendron A (n = 19), Pleosporales B (n = 18), Hyaloscyphaceae A (n = 16), Cladophialophora A (n = 13), and Hyaloscyphaceae B (n = 11) (Fig. 5). Of these, Asterostroma (14 soil samples from eight sites), Oidiodendron A (nine soil samples from six sites), Pleosporales A (nine soil samples from five sites), and Cladophialophora A (six soil samples from five sites) were the most widely distributed fungi across the 10 sites sampled. Although Sordariales A was only isolated from two study sites, this species was locally abundant and was detected in 10 of 13 soil samples at these two locations. We also noted that some wellresolved genera or families of fungi were particularly diverse and abundant from sclerotia, including Cladophialophora (17 OTUs, 58 isolates), Hyaloscyphaceae (11 OTUs, 39 isolates), and Oidiodendron (5 OTUs, 25 isolates). Sixty-three OTUs were cultured from only one study site and 50 of these were only isolated one time. The ANOSIM analysis detected significant dissimilarity in the occurrence of taxa (R = 0.55, P = 0.005) and the community structure (R = 0.38, P = 0.016) among the different study sites. Fungal isolation from floated sclerotia in water
Isolation success from Cenococcum sclerotia that floated in water was low. Only one isolate of C. geophilum was FEMS Microbiol Ecol 90 (2014) 708–717
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Fig. 4. Ratio of OTUs and isolates of sclerotiaassociated fungi in different phylogenetic groups. OTUs that could not be identified to the order level are left unlabeled.
Fig. 5. Total number of isolates in each OTU and the number of sites where each OTU was found. Fifty OTUs that were only isolated one time are not shown.
obtained from the 39 sclerotia that floated in water. Of the 30 isolates obtained from these 39 floating sclerotia, 21 represented Asterostroma, five represented species of Cladophialophora (4 OTUs), five represented species of Eurotiales (2 OTUs), one was Oidiodendron C, and the remaining isolate was Pezizomycotina B.
Discussion This study documented both a high frequency and high diversity of non-Cenococcum fungi that were isolated from within Cenococcum sclerotia. This high frequency of colonization by other fungi was surprising because a previous study documented a relatively high success rate for isolating Cenococcum from sclerotia (Trappe, 1969). Trappe (1969) successfully isolated Cenococcum c. 90% of the time (54 of 60) when he used viable sclerotia (which were defined as sclerotia with a more-or-less shiny appearance that sank in water). Although we used methods that are similar to those of Trappe (1969), we had a comparatively lower success rate
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for Cenococcum isolations (average 30.0 19.8%, range 0– 80.4%) that was apparently due to the higher colonization rate of sclerotia by non-Cenococcum sclerotia-associated fungi. Although the Cenococcum isolation success rate was not statistically different between sites, the rate of isolation for non-Cenococcum fungi was lower at some sites (e.g. SW, BA and LW) than for others (e.g. SR, SL OS and GA). One possible reason for the high rate of colonization by nonCenococcum fungi may be differences in the average freshness of sclerotia from various collection sites. While Trappe (1969) used only shiny sclerotia that sank in water, we found it was challenging to differentiate between shiny and dull sclerotia, so we used all sclerotia that sank in water for our experiment. However, we observed that the study sites with the highest isolation success rates and the lowest percentages of non-Cenococcum isolates (SW, BA, LW) tended to have more shiny sclerotia that were directly attached to black hyphae (K. Obase, personal observ.). We hypothesize that these may have been the freshest sclerotia and that this may account for the lower proportion of non-Cenococcum ª 2014 Federation of European Microbiological Societies. Published by John Wiley & Sons Ltd. All rights reserved
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fungi and the higher proportion of Cenococcum at these sites. Trappe (1962) documented that sclerotia surfaces roughen, and the sheen disappears when the sclerotia rest before germinating, possibly due to loss of oil content and cell death in the peripheral sclerotia layers. He also mentioned that dead peripheral cell layers peeled off as a husk. We also observed that some sclerotia had a fragile exterior surface that peeled away when the sclerotia were scrubbed. These findings suggest the presence of structural vulnerability in old sclerotia, for example tiny cracks on the surface layers. This structural vulnerability may allow older sclerotia to be invaded by diverse soil fungi that may not otherwise have the physiological capacity to penetrate through the highly melanized cell walls of fresh sclerotia. Thus, it seems likely that as sclerotia age, they will be vulnerable to an increasingly diverse group of fungi. Another important consideration that may account for the success rate of Cenococcum isolations and rate of colonization of sclerotia by other fungi is that antagonistic sclerotia-associated fungi may be more common in some geographic locations than in others. Our experiments were conducted in temperate to subtropical forests with relatively high temperatures, high precipitation, and rare freezing and drought. Thus, the environmental conditions in Florida’s forests may be conducive to decay and/or parasitism of sclerotia as compared to other sites. Comparison in the success rate of both Cenococcum and nonCenococcum fungi among diverse geographic locations (e.g. comparing several boreal, temperate, tropical sites) and edaphic conditions would help to clarify the mechanisms for fungal infection within Cenococcum sclerotia. The majority of fungal OTUs were cultured from only a few sclerotia and were only encountered at one or two sites. However, several sclerotia-associated fungi were isolated frequently (e.g. Sordariales A), occurred across many of the 10 forest sites (e.g. Oidiodendron A, Cladophialophora A, Eurotiales A), or were both frequent and abundant (e.g. Asterostroma, Pleosporales A) (Fig. 5). In fact, these common sclerotia-associated fungi that were isolated more than 10 times each dominated our dataset and accounted for roughly 57% of all sclerotia-associated fungi isolates obtained in this study. The high frequency and/or abundance of these common species was unexpected given the potential difficulties of using sclerotia as a nutrition source. It suggests that these common sclerotia-associated fungi may be specialized mycoparasites or saprobes that preferentially decay fungal tissues or act as endophytes, which colonize sclerotia without any aggressive interactions. Consistent with this hypothesis, we noted that most of the common sclerotia-associated fungi (e.g. species of Cladophialophora, Oidiodendron, and ª 2014 Federation of European Microbiological Societies. Published by John Wiley & Sons Ltd. All rights reserved
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Pleosporales but not Asterostroma) grow slowly on MMN media compared to common soil saprobes. This is particularly surprising because the MMN media we used is a rich source of nutrients and the majority of fungi, including putatively obligate ectomycorrhizal symbionts such as Cenococcum, have the capacity to grow on this medium. Despite the fact that we sequenced the ITS rDNA for all isolates and the more conserved LSU region for most OTUs, it was challenging to determine the phylogenetic placement of many fungi. As exemplified by the OTU names in Fig. 5, many common fungal OTUs appear to be distantly related to any described fungal species and could only be resolved to the fungal order (e.g. Sordariales A, Pleosporales A). This low resolution makes it challenging to infer trophic interactions. For example, Sordariales A is most closely related to a fungal root endophyte of Woollsia pungens (Ericaceae), but the ecological role of this fungus remains unknown (Midgley et al., 2004). Nevertheless, some fungi were readily identified to genus, and their phylogenetic positions can be used to infer something about their biology. For example, the relatively rapid growth rate of Asterostroma on media and the frequent isolation of this OTU (particularly on the dead sclerotia that floated in water) is not surprising given that members of this genus are known for their strong saprobic abilities, including the ability to grow on rigid insulating materials and wood (Schroeder & French, 1964) and to cause damage to buildings (Singh, 1999). On the other hand, the other common sclerotia-associated fungi may have abilities to endophitically colonize the inside of sclerotia as well as to utilize sclerotia as a nutrient source. Oidiodendron A is closely related to O. maius, and this species is known as both a saprobe on Sphagnum and peat (Tsuneda et al., 2001; Rice & Currah, 2002) as well as a root endophyte and ericoid mycorrhizal fungus (Sigler & Gibas, 2005; Kernaghan & Glenn, 2011). Several species of Cladophialophora act as animal pathogens (including on humans), as endophytes of plants, and as specialized mycoparasites (Park & Shin, 2011), although there are scant ecological data for the species. Recent studies suggest that members of this genus are particularly adept at degradation of aromatic hydrocarbons, which may aid these fungi in decomposing sclerotia (Prenafeta-Boldu et al., 2002). Interactions with microorganisms influence successful colonization and survival of soil fungi. For example, Shaw et al. (1995) documented combative hyphal interactions between C. geophilum and saprotrophic fungi on MMN media whereas several other studies have documented competition for roots between C. geophilum and other ectomycorrhizal fungi (Koide et al., 2005; Pickles et al., 2012). However, no studies have yet documented interactions FEMS Microbiol Ecol 90 (2014) 708–717
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between Cenococcum sclerotia and other fungi directly in the soil. The colonization of Cenococcum sclerotia by several widely distributed and common soil fungi suggests that these fungi may actively parasitize Cenococcum sclerotia or specifically target sclerotia as an energy source or substrates for unknown purposes. Understanding the effects of sclerotia-associated fungi on the viability of Cenococcum sclerotia will be important to fully understand the biology and lifecycle of Cenococcum in nature. In the future, in vitro studies that combine microscopy, inoculations of specific sclerotiaassociated fungi, and the use of both fresh and old sclerotia for experiments may help to elucidate the ecological interactions between Cenococcum sclerotia and other soil fungi.
Acknowledgements This study was supported by Grant-in-Aid for JSPS Postdoctoral Fellow for Research Abroad (to K.O.) with additional funding from the University of Florida Institute for Food and Agricultural Sciences (IFAS) (to M.E.S.). We thank Interdisciplinary Center for Biotechnology Research (ICBR) at the University of Florida for performing DNA sequencing and collaborators at the Smith Mycology Lab at the University of Florida for their assistance in collecting samples. The authors would like to thank University of Florida’s Ordway-Swisher Biological Station and Austin Cary Forest for access to forest sites.
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Supporting Information Additional Supporting Information may be found in the online version of this article: Fig. S1. Species accumulation curve for OTUs of soil fungi isolated from Cenococcum sclerotia. Table S1. Table of molecular data obtained for all nonCenococcum fungal isolates obtained in this study. Table S2. Table of molecular data obtained for all Cenococcum geophilum fungal isolates obtained in this study. Table S3. Abundance and percentage of sclerotia that occurred Cenococcum (Cg) or other sclerotia-associating fungi (non-Cg) or that did not occurred any fungi in each soil sample.
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Culturable fungal assemblages growing within Cenococcum sclerotia in forest soils Keisuke Obase1, Greg W. Douhan2, Yosuke Matsuda3 & Matthew E. Smith1 1
Department of Plant Pathology, University of Florida, Gainesville, FL, USA; 2Department of Plant Pathology and Microbiology, University of California, Riverside, CA, USA; and 3Laboratory of Forest Pathology and Mycology, Graduate School of Bioresources, Mie University, Tsu, Mie, Japan
Correspondence: Keisuke Obase, Department of Plant Pathology, University of Florida, 2523 Fifield Hall, Gainesville, FL 32611-0680, USA. Tel.: +13522262181; fax: 13523926532; e-mails: [email protected], [email protected] Received 8 August 2014; revised 11 September 2014; accepted 12 September 2014. Final version published online 30 September 2014.
MICROBIOLOGY ECOLOGY
DOI: 10.1111/1574-6941.12428 Editor: Ian C. Anderson Keywords Cenococcum geophilum; decomposition; mycoparasite; root endophyte; pathogen.
Abstract The ectomycorrhizal fungus Cenococcum geophilum (Ascomycota, Dothideomycetes) forms black, round to irregular sclerotia in forest soils. Fungi that colonize the sclerotia appear to affect sclerotia viability and may play an important role in the life history of Cenococcum. Some of the fungi could also affect nutrient cycling by decomposing Cenococcum sclerotia, which are melanized and recalcitrant to decay. We used a culture-based method to document the fungal communities growing inside surface-sterilized sclerotia that were collected from forest soils. Cenococcum was successfully isolated from 297 of 971 sclerotia whereas 427 sclerotia hosted fungi other than Cenococcum. DNA barcoding of the internal transcribed spacer rDNA followed by grouping at 97% sequence similarity yielded 85 operational taxonomic units (OTUs) that consisted primarily of Ascomycota (e.g. Chaetothyriales, Eurotiales, Helotiales, Pleosporales) and a few Basidiomycota and Mucoromycotina. Although most fungal OTUs were infrequently cultured, several OTUs such as members of Asterostroma, Cladophialophora, Oidiodendron, and Pleosporales were common and found across many sites. Our results suggest that Cenococcum sclerotia act as a substrate for diverse fungi. The occurrence of several OTUs in sclerotia across many sites suggests that these fungi may be active parasites of Cenococcum sclerotia or may preferentially use sclerotia as a nutrient source.
Introduction Sclerotia are hard, compact masses of fungal mycelium that usually form in soil or plant tissue. They are thought to serve as resting structures that can survive and remain quiescent in adverse environmental conditions until circumstances become favorable for fungal growth (ColeySmith & Cooke, 1971; Willetts, 1971). Sclerotia of many fungal species, including Cenococcum geophilum (Massicotte et al., 1992), Rhizoctonia solani (Naiki & Ui, 1975), and Sclerotinia minor (Bullock et al., 1980) have thick, pigmented fungal cell walls that can reduce colonization by other microorganisms and aid in their survival in the environment (Hurst & Wagner, 1969; Malik & Haider, 1982). Other fungi, such as Rhizoctonia spp. (Burton & Coley-Smith, 1985) and Aspergillus spp. (e.g. Wicklow & Cole, 1982; Frisvad et al., 2014), produce antibiotic compounds to resist attack by microorganisms. However,
ª 2014 Federation of European Microbiological Societies. Published by John Wiley & Sons Ltd. All rights reserved
sclerotia are often infected by a number of soil fungi that may enter via small cracks on the surfaces of the sclerotia or through active attack by fungal pathogens (Makkonen & Pohjakallio, 1960; Naiki & Ui, 1975; Huang & Hoes, 1976; Gladders & Coley-Smith, 1980; Willetts & Wong, 1980; Alexander & Stewart, 1994). Although the interactions between sclerotia and sclerotia-associated fungi have not been fully explored, several studies have shown that sclerotia germination is suppressed by some sclerotia-associated fungi (Gladders & Coley-Smith, 1980; Zazzerini & Tosi, 1985). Moreover, some mycoparasitic fungi, such as Trichoderma harzianum, actually degrade sclerotia cell walls and exploit the cytoplasmic contents of the attacked cells (Elad et al., 1984). Consequently, infection by sclerotia-associated fungi could be an important biological factor that may reduce the viability of sclerotia and therefore negatively impact the fitness of sclerotia-forming fungi such as
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C. geophilum. Sclerotia-associated fungi may also affect nutrient cycling by enhancing the decomposition rate of sclerotia, which otherwise remain recalcitrant to decay (Fernandez et al., 2013). The fungal biota that colonize sclerotia and the rate of infection have been investigated for several plant pathogenic fungi (e.g. Makkonen & Pohjakallio, 1960; Huang, 1985) with the idea that these sclerotia-associated fungi may be useful for biocontrol (Tu, 1984; Whipps & Gerlagh, 1992). However, there have been no studies of the fungal communities associated with the sclerotia of nonpathogenic fungi. The sclerotia of C. geophilum (Fig. 1) are 0.5–4.0 mm in diameter, spherical with a shiny smooth surface, and frequently found in soils of pine and oak forests (e.g. Trappe, 1962; Jany et al., 2002; Douhan & Rizzo, 2005; Obase et al., 2010). Cenococcum geophilum is one of the most common ectomycorrhizal fungi in the world and this fungus forms symbiotic associations with a wide variety of woody host plants (Trappe, 1969; Massicotte et al., 1992; LoBuglio, 1999). Previous studies indicate that C. geophilum is a monophyletic species complex, but the cryptic species within this group are not distinguishable without molecular methods (LoBuglio et al., 1991, 1996; Douhan & Rizzo, 2005; Douhan et al., 2007). Cenococcum sclerotia consist of a well-differentiated rind and a medulla with pseudoparenchymatous organization composed of melanized fungal mycelium (Massicotte et al., 1992). The longevity of Cenococcum sclerotia in forest soils has not been explicitly tested, but several lines of evidence suggest that sclerotia are long-lived and important in the biology and life history of Cenococcum (Trappe, 1962; Shaw & Roy, 1983; Miller et al., 1994). First, Cenococcum is ubiquitous both as sclerotia and on ectomycorrhizal roots in a wide variety of forest habitats (Trappe, 1962; LoBuglio, 1999). Second, Cenococcum has no known sexual or asexual spore-producing structures and is believed to be an ancient asexual fungal lineage (Douhan & Rizzo, 2005; but see Fernandez-Toiran & Agueda, 2007). Third, sclerotia can be surface-sterilized and grown as weak saprotrophs in axenic culture and are thought to act similarly when they contact nutrient resources in nature (Trappe, 1962). The high abundance and resilience of Cenococcum sclerotia in forest soils also means that these structures may sequester carbon and other compounds which could therefore impact nutrient cycling (Watanabe et al., 2001, 2007). Trappe (1962) showed that dead Cenococcum sclerotia persist intact for several years in soil, suggesting that they likely remain intact for long periods of time and sequester nutrients, so that they are unavailable to other organisms. Several studies have measured the size and abundance of Cenococcum sclerotia and found that these structures represent a massive amount of fungal tissue in FEMS Microbiol Ecol 90 (2014) 708–717
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many forests: 440 kg ha1 in an old-growth Norway spruce forest (Dahlberg et al., 1997), 2785 kg ha1 in a second-growth Douglas-fir stand (Fogel & Hunt, 1979), and 2300–3000 kg ha1 in stands of Abies amabilis (Vogt et al., 1981). This evidence suggests that understanding the diversity and community structure of microbial parasites or decomposers of Cenococcum sclerotia may provide insights into the ecology of Cenococcum and also nutrient cycling in forest soils. To assess the fungal diversity and community structure of sclerotia-associated fungi, we collected sclerotia of C. geophilum from ten sites in mixed oak–pine forests in Florida and Georgia. We isolated all culturable fungi from surface-sterilized Cenococcum sclerotia on nutrient agar media and then used DNA sequencing of the internal transcribed spacer (ITS) rDNA region to identify the sclerotia-associated fungi. In some cases, ITS was insufficient to accurately identify the sclerotia-associated fungi, so we also sequenced a portion of the 28S rDNA to refine our phylogenetic placement of these fungi.
Materials and methods Site descriptions and sampling procedures
Between August and October 2013, soil samples (c. 7 9 7 9 10 cm) were collected from ten different forest sites in Florida and Georgia, USA (Fig. 2). All study sites were natural or managed forest ecosystems dominated by species of Quercus and Pinus except for a single sampling site (PO) in an orchard of cultivated Carya illinoinensis (Wangenh.) K. Koch. The closest forest sites were roughly 2 km apart whereas the most distant sites were 170 km apart. At each individual forest site, we selected 2–10 locations, which were 2–500 meters apart each other, and collected one soil sample beneath 1–4 trees spread over an area of 1–25 m2 at each location. Climatic data from the Gainesville weather station during 2012 (Gainesville Regional Airport; 29°410 24″N, 82°160 18″ W; 46.0 m asl) indicated annual precipitation of 56.25 inches (c. 1428.7 mm) and mean temperature of 21.6 °C, ranging between 14.1 °C in January and 28.6 °C in July. All samples were stored in plastic bags at 8 °C for less than a week before isolations were made. Sampling sclerotia and isolation of fungi
Forest soils were washed in running tap water over a 2.0-mm sieve to remove excess plant debris and soil particles. Filtered soil slurry (including both small sclerotia (0.5–2.0 mm in diameter) and larger sclerotia > 2.0 mm in diameter) was taken from the sieve, stirred vigorously in a glass beaker filled with water, and then left standing ª 2014 Federation of European Microbiological Societies. Published by John Wiley & Sons Ltd. All rights reserved
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(a)
(b)
(c)
Fig. 1. (a) Ectomycorrhizas of Cenococcum geophilum with sclerotia (arrows) attached via hyphal connections. (b) Hyphal growth of C. geophilum on MMN agar media. (c) Closeup view of C. geophilum sclerotium.
Fig. 2. Ten study sites where Cenococcum sclerotia were collected in Florida and Georgia, USA.
for 10 s. Floating plant debris was removed by decanting. The remaining slurry was spread on a glass dish (9 cm in diameter) and observed under a dissecting microscope. Sclerotia that were black, c. 0.5–3.0 mm in diameter, round, and sank in water were considered likely to be viable C. geophilum sclerotia based on the findings of Trappe (1969) and Douhan & Rizzo (2005). Forceps were used to transfer these putatively viable sclerotia to a moist kimwipe paper towel and the kimwipe was then used to vigorously scrub the sclerotia to remove adhering soil particles. Sclerotia that broke apart during the scrubbing stage were discarded and excluded from further analysis. Putatively viable sclerotia were then transferred to a ª 2014 Federation of European Microbiological Societies. Published by John Wiley & Sons Ltd. All rights reserved
50-mL tube with 20–30 mL water and shaken vigorously by hand for 30 s. All sclerotia were transferred to a glass dish and observed for a second time under the dissecting microscope. Soil particles still adhering to the sclerotia were then removed by hand with clean foreceps. Thirtynine sclerotia that floated in water were also collected from two study sites (LW and SW) and subjected to the same treatment as above to test whether floating sclerotia were more likely to be parasitized than sinking sclerotia. Prior to culturing, we surface-sterilized sclerotia by immersing them for 20 min in commercially available bleach (containing 2% chlorine) followed by two rinses in c. 20 mL of sterilized distilled water. Following sterilization FEMS Microbiol Ecol 90 (2014) 708–717
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and rinsing, we used flame-sterilized forceps to transfer sclerotia to a new, sterile kimwipe to remove excess water. Dry, surface-sterilized sclerotia were then axenically transferred to 90-mm petri dishes containing modified Melin– Norkran’s (MMN) media (Marx, 1969) with streptomycin (100 mg mL1) and chloramphenicol (50 mg mL1). Petri dishes were incubated at 23 °C. Fungi that grew from sclerotia in the first 2 days after incubation were considered likely soil contaminants that occurred due to incomplete surface sterilization and therefore discarded. Fungal isolates derived from one fungal species and emerging after 3 days were subcultured and retained in the analysis. Culture dishes inhabited by two or more species were discarded due to the difficulties in separating intermingled fungal colonies.
Fungal isolates were classified into operational taxonomic units (OTUs) based on 97% similarity of the ITS sequences. All unique ITS, ITS + LSU, or gpd sequences were subjected to BLAST searches (Altschul et al., 1997) against GenBank (http://blast.ncbi.nlm.nih.gov). Sequences obtained from this study were aligned with identified fungal sequences using MAFFT (Katoh & Standley, 2013). Maximum Likelihood (ML) analyses were performed with bootstrap analysis with 100 replicates with MEGA5 (Tamura et al., 2011), to verify our phylogenetic identification of the isolates based on BLAST searches. The general time-reversible model incorporating invariant sites and a gamma distribution (GTR + I + G) was selected as the appropriate model based on the results of the corrected Akaike information criterion value.
DNA extraction and identification of fungi
Statistical analysis
DNA was extracted from fresh mycelium of each fungal isolate with the Extract-N-Amp kit (Sigma-Aldrich) according to the manufacturer’s instructions. We amplified the ITS rDNA region (ITS1-5.8s-ITS2) using Taq DNA polymerase (New England BioLabs) with a fungispecific primer ITS1F (Gardes & Bruns, 1993) and a universal primer ITS4 (White et al., 1990). For isolates that morphologically matched C. geophilum, we verified their identity by amplifying and sequencing the glyceraldehyde 3-phosphate dehydrogenase (gpd) gene region with primers gpd1 and gpd2 (Berbee et al., 1999). We have determined that sequencing the gpd locus is the most effective way to verify the identity of Cenococcum isolates because gpd was the most phylogenetically informative DNA region examined by Douhan & Rizzo (2005). Unfortunately, many non-Cenococcum isolates could not be readily identified to class or order level based on ITS sequences alone. To refine the identification of these isolates, we also amplified partial LSU sequences from one representative culture of each OTU using primers ITS3 (White et al., 1990) and LR5 (Vilgalys & Hester, 1990). PCR followed the amplification conditions of Obase & Matsuda (in press) for ITS, Duong et al. (2012) for LSU, and Douhan et al. (2007) for gpd except that we used 44 instead of 25 cycles and an annealing temperature of 58 °C instead of 55 °C. Positive PCR amplicons were cleaned with EXO and SAP enzymes (Glenn & Schable, 2005), and then sequenced bidirectionally with the same primers as above with the BigDye Terminator v. 3.1 Cycle Sequencing Kit on an ABI3700 DNA sequencer (Applied Biosystems). The fungal ITS, LSU and gpd sequence data were submitted to Genebank under accession numbers AB986283–AB986537. All of the fungal DNA sequences obtained in this study are detailed in Supporting Information, Tables S1 and S2.
Kruskal–Wallis analyses were conducted to compare the isolation ratio of non-Cenococcum fungi among study sites. Study sites that included more than three soil samples and had more than 10 sclerotia were included in the analysis (Table S3). Differences among study sites were determined using the Mann–Whitney pairwise post hoc tests (P < 0.05). An analysis of similarity (ANOSIM) was performed to compare the biota of non-Cenococcum fungi among study sites. The ANOSIM were computed with 9999 permutations and based on presence/absence data using the Raup–Crick similarity index (Raup & Crick, 1979) or abundance data using the Bray– Curtis similarity index (Bray & Curtis, 1957). Twelve soil samples, from four study sites that included more than seven non-Cenococcum isolates were subjected to the analysis. The analyses were performed using PAST (Hammer et al., 2001). OTU accumulation curves were drawn by plotting the mean of the accumulated number of expected OTUs in pooled soil samples after 1000 randomizations without replacement, using ESTIMATES version 9.1.0 (Colwell, 2013) to estimate whether the sampling effort was sufficient to fully describe the community of sclerotia-associated fungi. Estimated OTU richness was calculated using the Chao 2 estimator.
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Results Isolation success for Cenococcum and nonCenococcum sclerotia-associated fungi
No sclerotia were present in 22 soil samples but the 33 remaining soil samples each contained 1–66 Cenococcum sclerotia (Table S3). In total, 971 sclerotia were collected from 1 to 9 soil samples from each study site and subjected to our isolation procedures. Thirty sclerotia (3.1%) were rapidly contaminated in the first 2 days with fastgrowing fungi and removed from further analysis. ª 2014 Federation of European Microbiological Societies. Published by John Wiley & Sons Ltd. All rights reserved
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Cenococcum geophilum was successfully obtained from 297 sclerotia (31.6%) whereas other fungi were recovered in 427 (45.4%) of 941 total sclerotia. There was no fungal occurrence from the remaining 217 sclerotia (23.1%). When soil samples that included < 10 sclerotia were removed from the analysis, the isolation success rate for Cenococcum ranged from 0 to 80.4% (Ave. SD; 30.0 19.8%) and the isolation success rate for nonCenococcum fungi ranged from 0 to 95.2% (Ave. SD; 47.8 26.2%). The isolation success rate for non-Cenococcum fungi was significantly different between study sites (Kruskal–Wallis analysis; H = 17.2, P = 0.016). However, there were no significant differences in the isolation success rate of C. geophilum (H = 12.3, P = 0.091) between sites. There was also no difference in the number of sclerotia from which no fungi were isolated between sites (H = 7.83, P = 0.34) (Fig. 3). Identification of sclerotia-associated fungi
Six hundred and sixty-seven of 724 isolates (92.1%) were successfully identified based on ITS and/or LSU sequences (for non-Cenococcum fungi) or based on a combination of ITS and/or gpd sequences (for C. geophilum). Of these, 297 isolates were identified as C. geophilum whereas the remaining 370 non-Cenococcum isolates were classified into 85 different OTUs.
Fig. 3. Average success rate for isolation of Cenococcum geophilum (filled black) and non-Cenococcum fungi (filled gray) from sclerotia in each study site. Proportions of sclerotia where no fungal isolates were obtained are indicated in white. Two study sites (FW and PO) were removed from the analysis due to low numbers of soil samples that contained Cenococcum sclerotia. Different letters indicate significant differences at P < 0.05 based on the Mann–Whitney pairwise post hoc tests.
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BLAST searches revealed that 313 isolates (81 OTUs) were Ascomycota, 56 isolates (3 OTUs) were Basidiomycota, and one isolate was a member of Mucoromycotina (Fig. 4). Isolates that were identified as belonging to the Ascomycota consisted of Eurotiomycetes (91 isolates in 25 OTUs representing Eurotiales and Chaetothyriales), Dothideomycetes (93 isolates in 22 OTUs representing Pleosporales and Venturiales), Leotiomycetes (68 isolates in 20 OTUs representing Helotiales sensu lato), Sordariomycetes (54 isolates in eight OTUs representing Hypocreales, Sordariales, and Xylariales), and other Pezizomycotina (seven isolates in six OTUs). All Basidiomycota isolates belonged to the Agaricomycetes (56 isolates in 3 OTUs representing Hymenochaetales and Russulales). One Umbelopsis OTU was the sole member of the Mucoromycotina.
Community structure of sclerotia-associated fungi
The accumulation curve for OTUs did not reach an asymptote (Fig. S1) and estimated a total community of 436 OTUs of sclerotia-associated fungi by the Chao2 estimator, indicating that many rare sclerotia-associated fungi would likely be identified with further sampling effort. The Asterostroma (54 isolates) and Sordariales A (45 isolates) OTUs were the most frequently isolated sclerotia-associated fungi followed by Pleosporales A (n = 36), Oidiodendron A (n = 19), Pleosporales B (n = 18), Hyaloscyphaceae A (n = 16), Cladophialophora A (n = 13), and Hyaloscyphaceae B (n = 11) (Fig. 5). Of these, Asterostroma (14 soil samples from eight sites), Oidiodendron A (nine soil samples from six sites), Pleosporales A (nine soil samples from five sites), and Cladophialophora A (six soil samples from five sites) were the most widely distributed fungi across the 10 sites sampled. Although Sordariales A was only isolated from two study sites, this species was locally abundant and was detected in 10 of 13 soil samples at these two locations. We also noted that some wellresolved genera or families of fungi were particularly diverse and abundant from sclerotia, including Cladophialophora (17 OTUs, 58 isolates), Hyaloscyphaceae (11 OTUs, 39 isolates), and Oidiodendron (5 OTUs, 25 isolates). Sixty-three OTUs were cultured from only one study site and 50 of these were only isolated one time. The ANOSIM analysis detected significant dissimilarity in the occurrence of taxa (R = 0.55, P = 0.005) and the community structure (R = 0.38, P = 0.016) among the different study sites. Fungal isolation from floated sclerotia in water
Isolation success from Cenococcum sclerotia that floated in water was low. Only one isolate of C. geophilum was FEMS Microbiol Ecol 90 (2014) 708–717
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Fig. 4. Ratio of OTUs and isolates of sclerotiaassociated fungi in different phylogenetic groups. OTUs that could not be identified to the order level are left unlabeled.
Fig. 5. Total number of isolates in each OTU and the number of sites where each OTU was found. Fifty OTUs that were only isolated one time are not shown.
obtained from the 39 sclerotia that floated in water. Of the 30 isolates obtained from these 39 floating sclerotia, 21 represented Asterostroma, five represented species of Cladophialophora (4 OTUs), five represented species of Eurotiales (2 OTUs), one was Oidiodendron C, and the remaining isolate was Pezizomycotina B.
Discussion This study documented both a high frequency and high diversity of non-Cenococcum fungi that were isolated from within Cenococcum sclerotia. This high frequency of colonization by other fungi was surprising because a previous study documented a relatively high success rate for isolating Cenococcum from sclerotia (Trappe, 1969). Trappe (1969) successfully isolated Cenococcum c. 90% of the time (54 of 60) when he used viable sclerotia (which were defined as sclerotia with a more-or-less shiny appearance that sank in water). Although we used methods that are similar to those of Trappe (1969), we had a comparatively lower success rate
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for Cenococcum isolations (average 30.0 19.8%, range 0– 80.4%) that was apparently due to the higher colonization rate of sclerotia by non-Cenococcum sclerotia-associated fungi. Although the Cenococcum isolation success rate was not statistically different between sites, the rate of isolation for non-Cenococcum fungi was lower at some sites (e.g. SW, BA and LW) than for others (e.g. SR, SL OS and GA). One possible reason for the high rate of colonization by nonCenococcum fungi may be differences in the average freshness of sclerotia from various collection sites. While Trappe (1969) used only shiny sclerotia that sank in water, we found it was challenging to differentiate between shiny and dull sclerotia, so we used all sclerotia that sank in water for our experiment. However, we observed that the study sites with the highest isolation success rates and the lowest percentages of non-Cenococcum isolates (SW, BA, LW) tended to have more shiny sclerotia that were directly attached to black hyphae (K. Obase, personal observ.). We hypothesize that these may have been the freshest sclerotia and that this may account for the lower proportion of non-Cenococcum ª 2014 Federation of European Microbiological Societies. Published by John Wiley & Sons Ltd. All rights reserved
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fungi and the higher proportion of Cenococcum at these sites. Trappe (1962) documented that sclerotia surfaces roughen, and the sheen disappears when the sclerotia rest before germinating, possibly due to loss of oil content and cell death in the peripheral sclerotia layers. He also mentioned that dead peripheral cell layers peeled off as a husk. We also observed that some sclerotia had a fragile exterior surface that peeled away when the sclerotia were scrubbed. These findings suggest the presence of structural vulnerability in old sclerotia, for example tiny cracks on the surface layers. This structural vulnerability may allow older sclerotia to be invaded by diverse soil fungi that may not otherwise have the physiological capacity to penetrate through the highly melanized cell walls of fresh sclerotia. Thus, it seems likely that as sclerotia age, they will be vulnerable to an increasingly diverse group of fungi. Another important consideration that may account for the success rate of Cenococcum isolations and rate of colonization of sclerotia by other fungi is that antagonistic sclerotia-associated fungi may be more common in some geographic locations than in others. Our experiments were conducted in temperate to subtropical forests with relatively high temperatures, high precipitation, and rare freezing and drought. Thus, the environmental conditions in Florida’s forests may be conducive to decay and/or parasitism of sclerotia as compared to other sites. Comparison in the success rate of both Cenococcum and nonCenococcum fungi among diverse geographic locations (e.g. comparing several boreal, temperate, tropical sites) and edaphic conditions would help to clarify the mechanisms for fungal infection within Cenococcum sclerotia. The majority of fungal OTUs were cultured from only a few sclerotia and were only encountered at one or two sites. However, several sclerotia-associated fungi were isolated frequently (e.g. Sordariales A), occurred across many of the 10 forest sites (e.g. Oidiodendron A, Cladophialophora A, Eurotiales A), or were both frequent and abundant (e.g. Asterostroma, Pleosporales A) (Fig. 5). In fact, these common sclerotia-associated fungi that were isolated more than 10 times each dominated our dataset and accounted for roughly 57% of all sclerotia-associated fungi isolates obtained in this study. The high frequency and/or abundance of these common species was unexpected given the potential difficulties of using sclerotia as a nutrition source. It suggests that these common sclerotia-associated fungi may be specialized mycoparasites or saprobes that preferentially decay fungal tissues or act as endophytes, which colonize sclerotia without any aggressive interactions. Consistent with this hypothesis, we noted that most of the common sclerotia-associated fungi (e.g. species of Cladophialophora, Oidiodendron, and ª 2014 Federation of European Microbiological Societies. Published by John Wiley & Sons Ltd. All rights reserved
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Pleosporales but not Asterostroma) grow slowly on MMN media compared to common soil saprobes. This is particularly surprising because the MMN media we used is a rich source of nutrients and the majority of fungi, including putatively obligate ectomycorrhizal symbionts such as Cenococcum, have the capacity to grow on this medium. Despite the fact that we sequenced the ITS rDNA for all isolates and the more conserved LSU region for most OTUs, it was challenging to determine the phylogenetic placement of many fungi. As exemplified by the OTU names in Fig. 5, many common fungal OTUs appear to be distantly related to any described fungal species and could only be resolved to the fungal order (e.g. Sordariales A, Pleosporales A). This low resolution makes it challenging to infer trophic interactions. For example, Sordariales A is most closely related to a fungal root endophyte of Woollsia pungens (Ericaceae), but the ecological role of this fungus remains unknown (Midgley et al., 2004). Nevertheless, some fungi were readily identified to genus, and their phylogenetic positions can be used to infer something about their biology. For example, the relatively rapid growth rate of Asterostroma on media and the frequent isolation of this OTU (particularly on the dead sclerotia that floated in water) is not surprising given that members of this genus are known for their strong saprobic abilities, including the ability to grow on rigid insulating materials and wood (Schroeder & French, 1964) and to cause damage to buildings (Singh, 1999). On the other hand, the other common sclerotia-associated fungi may have abilities to endophitically colonize the inside of sclerotia as well as to utilize sclerotia as a nutrient source. Oidiodendron A is closely related to O. maius, and this species is known as both a saprobe on Sphagnum and peat (Tsuneda et al., 2001; Rice & Currah, 2002) as well as a root endophyte and ericoid mycorrhizal fungus (Sigler & Gibas, 2005; Kernaghan & Glenn, 2011). Several species of Cladophialophora act as animal pathogens (including on humans), as endophytes of plants, and as specialized mycoparasites (Park & Shin, 2011), although there are scant ecological data for the species. Recent studies suggest that members of this genus are particularly adept at degradation of aromatic hydrocarbons, which may aid these fungi in decomposing sclerotia (Prenafeta-Boldu et al., 2002). Interactions with microorganisms influence successful colonization and survival of soil fungi. For example, Shaw et al. (1995) documented combative hyphal interactions between C. geophilum and saprotrophic fungi on MMN media whereas several other studies have documented competition for roots between C. geophilum and other ectomycorrhizal fungi (Koide et al., 2005; Pickles et al., 2012). However, no studies have yet documented interactions FEMS Microbiol Ecol 90 (2014) 708–717
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between Cenococcum sclerotia and other fungi directly in the soil. The colonization of Cenococcum sclerotia by several widely distributed and common soil fungi suggests that these fungi may actively parasitize Cenococcum sclerotia or specifically target sclerotia as an energy source or substrates for unknown purposes. Understanding the effects of sclerotia-associated fungi on the viability of Cenococcum sclerotia will be important to fully understand the biology and lifecycle of Cenococcum in nature. In the future, in vitro studies that combine microscopy, inoculations of specific sclerotiaassociated fungi, and the use of both fresh and old sclerotia for experiments may help to elucidate the ecological interactions between Cenococcum sclerotia and other soil fungi.
Acknowledgements This study was supported by Grant-in-Aid for JSPS Postdoctoral Fellow for Research Abroad (to K.O.) with additional funding from the University of Florida Institute for Food and Agricultural Sciences (IFAS) (to M.E.S.). We thank Interdisciplinary Center for Biotechnology Research (ICBR) at the University of Florida for performing DNA sequencing and collaborators at the Smith Mycology Lab at the University of Florida for their assistance in collecting samples. The authors would like to thank University of Florida’s Ordway-Swisher Biological Station and Austin Cary Forest for access to forest sites.
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Supporting Information Additional Supporting Information may be found in the online version of this article: Fig. S1. Species accumulation curve for OTUs of soil fungi isolated from Cenococcum sclerotia. Table S1. Table of molecular data obtained for all nonCenococcum fungal isolates obtained in this study. Table S2. Table of molecular data obtained for all Cenococcum geophilum fungal isolates obtained in this study. Table S3. Abundance and percentage of sclerotia that occurred Cenococcum (Cg) or other sclerotia-associating fungi (non-Cg) or that did not occurred any fungi in each soil sample.
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