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Phylogenetic Diversity of Chytridiomycetes in a Temporary Forest Pond Surveyed using Culture-Based Methods
William J. Davis, Jonathan Antonetti, Peter M. Letcher, and Martha J. Powell

Southeastern Naturalist, Volume 15, Issue 3 (2016): 534–548

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Southeastern Naturalist W.J. Davis, J. Antonetti, P.M. Letcher, and M.J. Powell 2016 Vol. 15, No. 3 534 2016 SOUTHEASTERN NATURALIST 15(3):534–548 Phylogenetic Diversity of Chytridiomycetes in a Temporary Forest Pond Surveyed using Culture-Based Methods William J. Davis1,*, Jonathan Antonetti1, Peter M. Letcher1, and Martha J. Powell1 Abstract - Temporary forest ponds are depressions that annually cycle between non-inundated and inundated stages and are recognized as biodiversity hotspots for amphibians and other taxa. However, little is known about the microbial communities of temporary forest ponds and less is known about chytrids, an inconspicuous, early diverging lineage of fungi, present in such ponds. We sampled soil and water from a temporary forest pond in the Oakmulgee District of the Talladega National Forest. Chytrids were isolated from the samples using baits and standard isolating techniques. Isolated strains were identified using the monophyletic-species concept based on a molecular phylogeny inferred from an alignment of the nuclear large subunit rDNA. A total of 30 strains were isolated, which represented 7 species. Due to the limitations of a culture-based method in being able to culture all chytrids that are present, we have described only a portion of the chytrid community. This study, however, is foundational to future inventories that apply a multiphasic approach to further reveal chytrid diversity in this understudied and variable habitat. Introduction Chytridiomycota (= chytrid fungi; Hibbett et al. 2007) inhabit aquatic and terrestrial habitats on all continents (Freeman et al. 2009, Hassett and Gradinger 2016, Richards et al. 2015, Sparrow 1960). Early studies of chytrid fungi relied on the practice of baiting samples and using light microscopy to observe and report chytrid species; they focused primarily on inventorying species and alpha taxonomy. Some authors sampled globally (i.e., on multiple continents; e.g., Karling 1945, 1946), whereas others focused on a region (i.e., a country/state; e.g., Canter 1946, 1947, 1949; Canter and Jaworski 1982, 1986; Canter and Lund 1948; Willoughby 1961, 1962a) or a single habitat (e.g., Czeczuga 1994, Dogma 1969, Paterson 1967). Researchers often encountered the same morphological species in multiple locations and in both soil and aquatic habitats, which lead to the view that chytrids are ubiquitous and cosmopolitan (Sparrow 1960). More recently, chytrid systematists, working regionally (Chen and Chien 1995, 1998; Chen et al. 2000; Jerônimo et al. 2015) or globally (Letcher et al. 2008a, b; Simmons et al. 2009), have isolated chytrids into axenic culture before studying and reporting them. Zoospore ultrastructural characters and rDNA sequences from the strains were used to infer new phylogenies and revise the phylum (Barr 1980; Letcher et al. 2008a, b; Longcore and Simmons 2012; Mozley-Standridge et al. 2009). Researchers found cryptic phylogenetic diversity within common morphological species. For example, Barr and Hartmann (1977) and Barr and Désaulniers 1Department of Biological Sciences, University of Alabama, Tuscaloosa, AL 35487. *Corresponding author - Manuscript Editor: Richard Baird Southeastern Naturalist 535 W.J. Davis, J. Antonetti, P.M. Letcher, and M.J. Powell 2016 Vol. 15, No. 3 (1986) observed variation in zoospore ultrastructure of Rhizophlyctis rosea (de Bary and Woronin) A. Fisch. Letcher et al. (2008a), using molecular and zoospore ultrastructural characters, described new genera based on Rhizophlyctis rosea-like organisms. Despite the progress in chytrid systematics, chytrids are still considered to be among the “dark-matter fungi”, i.e., those fungi for which little knowledge exists (Grossart et al. 2016), and most habitats remain un- or undersampled. One habitat that is undersampled for chytrid taxa is temporary forest ponds. Temporary forest ponds are depressions that annually cycle between non-inundated and inundated stages (Colburn 2004, Williams 2006). This cycling creates temporal environmental variability. For example, temperature and oxygen content can fluctuate rapidly over the course of a day. As a second example, pH and nutrient content increase or decrease from the start of the inundated stage to the start of non-inundated stage (Colburn 2004, Modlin 1980, Williams 2006). Due to their environmental variability, temporary ponds are centers of genetic diversity in microcrustaceans (Herbert 1998) and salamanders (Zamudio and Wieczorek 2007) and hypothesized epicenters of evolution (Williams 1988), which have allowed them to become biodiversity hotspots for a number of aquatic taxa, such as amphibians, microcrustaceans, and aquatic insects (Colburn 2004, Williams 2006). Food webs within temporary forest ponds are based on the degradation of terrestrial inputs, such as leaf litter and pollen (Colburn 2004, Rubbo et al. 2006, Williams 2006). However, little is known about the structure and dynamics of microbial communities responsible for degrading these terrestrial inputs (Bärlocher et al. 1978, Carrino-Kyker and Swanson 2008, Carrino-Kyker et al. 2011, Colburn 2004, Williams 2006). Bacteria are often numerous and likely make significant contributions to nitrogen-cycling, the degradation of cellulose, and the diets of filter feeders (Carrino-Kyker and Swanson 2008, Felton et al. 1967). Fungi, specifically terrestrial, aero-aquatic, and aquatic hyphomycetes, play a key role in degrading the complex compounds found in the annual terrestrial inputs (Bärlocher et al. 1978). As such, both control the flow of energy and nutrients through the food web, the production of humus, and the fertility of the basin soil (Williams 2006). Members of Chytridiomycota are present in temporary ponds and may play an important role in the ecosystem functioning of temporary forest ponds. Ingold (1941) described Phlyctochytrium proliferum, a parasite of the green alga Chlamydomonas, from a temporary pond. Laird (1988) lists Micromyces sp., a chytrid parasite of green algae, in a survey of a temporary pond. Longcore et al. (1995) included a strain of Geranomyces variabilis (Longcore, D.J.S. Barr, & Désauln.) D.R. Simmons (Barr 350) isolated from a temporary pond. In a molecular inventory of temporary-pond microbial communities, Carrino-Kyker and Swanson (2008) and Carrino-Kyker et al. (2011) detected chytrid fungi phylotypes identified as Rhizophydium elyensis Sparrow and Blyttiomyces helicus Sparrow & M.E. Barr. In streams reliant on terrestrial inputs, it has been shown that chytrids are a substantial and potentially important component of the microbial community degrading the leaf litter (Bärlocher et al. 2012, Marano et al. 2011, Nikolcheva and Bärlocher 2004, Seena et al. 2008, Sridhar et al 2011). In lakes, it has been shown that chytrid Southeastern Naturalist W.J. Davis, J. Antonetti, P.M. Letcher, and M.J. Powell 2016 Vol. 15, No. 3 536 zoospores provide an important link in aquatic food webs, called the mycoloop, that transfers energy and nutrients from lower trophic levels to higher trophic levels, and that this link potentially has a stabilizing effect on the overall food web (Kagami et al. 2014). Since temporary ponds are rich in the organic substrates that chytrids utilize (e.g., cellulose, chitin exo-skeletons, and pollen) and filter-feeding crustaceans make up one of the largest trophic levels of temporary ponds (Colburn 2004, Williams 2006), it is likely that chytrids are important components of temporary- pond food webs. The purpose of this study is to use a culture-based method and phylogenetic analysis to add to the knowledge of microbial communities by inventorying the chytrids present in a temporary pond. Materials and Methods Site description We collected soil, sediment, and water samples randomly from the basin edge and middle of a temporary forest pond located in the Oakmulgee District of the Talladega National Forest, Hale County, AL (32.97207°N, 87.47812°W; Fig. 1). The pond had standing water (inundated) from December to June (Fig. 2B). From June to November, standing water was not present, and the sediments were exposed to the atmosphere (non-inundated; Fig. 2A). The 50 m × 16 m pond inundates in late December, reaches peak inundation stage in March, and is noninundated by June. At peak inundation stage, pond depth ranges from 0.15 m at the edge to 2 m in the middle. Collection of samples for culturing We sampled monthly from January 2011 to August 2012, with an average of 10 samples taken at each time point. We collected soil in sterile whirl-top bags when the basin was not inundated. When the basin was inundated, we collected water in 50-mL Falcon tubes. Samples were transported on ice and stored at 4 °C until processed. In the laboratory, we flooded samples with sterile water in a Petri dish and placed sterile pollen (Pinus spp. and Liquidambar styraciflua L.), cellulose (onion epidermal cells), keratin (snake skin), and chitin (shrimp exoskeleton) in each Petri dish (Barr 1987, Couch 1939, Emerson 1950, Sparrow 1960). We sterilized the pollen and cellulose at 100 °C for 1 hr, treated the keratin (human hair) with chloroform to remove lipids, and decalcified and purified the chitin according to Karling (1945). We examined baits and naturally occurring substrates (e.g., insect exuviae, microcrustacean exuviae, and algae) periodically as wet mounts with a Nikon Eclipse E200 microscope (Davis et al. 2013). When chytrid thalli released zoospores, we used a glass pipette to remove the water, thalli, and zoospores from the microscope slide and streak them on a nutrient agar plate (Couch 1939, Emerson 1950). We used PmTG (1 g peptonized milk, 1 g tryptone, 5 g glucose, and 8 g agar per liter of water; Barr 1986) or mPmTG (0.4 g peptonized milk, 0.4 g tryptone, 2 g glucose, and 8 g agar per liter of water; Longcore 1992b) nutrient agar plates with antibiotics (0.25 g penicillin G and 0.25 g streptomycin sulfate per liter of water). Over 2 Southeastern Naturalist 537 W.J. Davis, J. Antonetti, P.M. Letcher, and M.J. Powell 2016 Vol. 15, No. 3 Figure 1. Sampling locations. (A) Map of Alabama showing the location of the Oakmulgee District of the Talladega National Forest in green. (B) Enlarged view of the Oakmulgee District with location of 2 temporary forest ponds indicated by the filled circles. (C) Detailed map showing the position of the ponds (filled circles) in the Oakmulgee National Forest; the pond focused on in this study is located on FS-705. Map created in ArcGIS using data from TIGER and the United States Forest Service. Southeastern Naturalist W.J. Davis, J. Antonetti, P.M. Letcher, and M.J. Powell 2016 Vol. 15, No. 3 538 days, we removed chytrid thalli from the streaked plate and placed them on fresh plates using sterile insect needles under a stereomicroscope (Barr 1987, Emerson 1950). Strains were observed and transferred until they were in axenic culture. We labeled axenic strains with the initials of the isolator and a number and kept them maintained on PmTG or mPmTG nutrient agar without antibiotics in Parafilmed Petri dishes at room temperature (20–25 °C) in the dark. We tentatively identified strains based on morphological features (Karling 1977, Sparrow 1960). DNA extraction, amplification, and sequencing of strains We extracted, purified, amplified, and sequenced genomic DNA from the cultured strains as described in Davis et al. (2013). We incubated strains in 50 mL of Figure 2. Photographs of the temporary forest pond. (A) Photograph of the non-inundated pond basin; note the large amounts of woody debris and leaf litter. (B) Photograph of the inundated pond basin. Southeastern Naturalist 539 W.J. Davis, J. Antonetti, P.M. Letcher, and M.J. Powell 2016 Vol. 15, No. 3 PmTG or mPmTG nutrient broth at room temperature for ~3 days on an Innova 2100 platform shaker (New Brunswick Scientific) at 120 rpm and centrifuged them for 30 min at 3000 rpm in a ThermoIEC-703a centrifuge (ThermoIEC) to pellet chytrid thalli. We extracted DNA with the NucleoSpin Plant II DNA extraction kit (Macherey-Nagel, Inc.) following the NucleoSpin protocol for fungal cultures. We determined the DNA concentration with a ND-1000 Nanodrop (Nanodrop) spectrophotometer. For PCR amplification, we diluted DNA to 10 ng/μL, and amplified the D1–D3 domains of the nuclear large subunit rDNA (28S) using the LROR/LR5 primer pair (Rehner and Samuels 1994, Vilgalys and Hester 1990). We chose this region because it is used to delineate taxa within Chytridiomycota (e.g., Letcher et al. 2008a, 2008b, 2015; Longcore and Simmons 2012). DNA was initially denatured at 94 °C and amplified through 30 cycles of 1 min at 94°C, 1 min at 50°C, and 1 min at 72 °C with a final elongation step at 72 °C for 5 min in a DNA Engine PTC- 200 Thermal Cycler (MJ Research). We pooled and purified Amplicons from 4 replicates following the protocols in NucleoSpin Extraction II kit. Amplicons were sequenced by Macrogen USA (Rockville, MD). We assembled contiguous sequences with Sequencher 4.5 (Genecodes) and searched them with the discontiguous megablast algorithm (Ma et al. 2002) against the GenBank database to corroborate or refute the morphological identification (Davis et al. 2013). Phylogenetic analysis of strains We downloaded reference sequences from GenBank and aligned them with the pond strain sequences with ClustalX (Thompson et al. 1997) followed by manual adjustment in BioEdit (Hall 1999). The alignment was deposited in TreeBase ( We used MrModeltest 2.3 to determine the best-fit model of base substitution (Letcher et al. 2015), inferred maximum likelihood (ML) trees with the GTR + Gamma model of sequence evolution in RAxML 7.0.3 (Stamatakis 2006), and bootstrapped with 1000 replicates using the rapid bootstrap function (Stamatakis et al. 2008). We rooted the inferred trees with Monoblepharella mexicana Shanor (UCB 78-1; James et al. 2006), a member of the Monoblepharidomycetes, the sister group of the Chytridiomycetes (Dee et al. 2015). We used the monophyletic-species concept (Mayden 1999) to identify strains; specifically, a strain was considered a member of a taxon if it formed a monophyletic clade with a reference sequence of that taxon (Davis et al. 2013). Results We isolated 30 strains representing 7 species: Rhizoclosmatium aurantiacum Sparrow (1 strain; Fig. 3B), Rhizoclosmatium sp. (1 strain), Rhizoclosmatium globosum H. E. Petersen (11 strains; Fig. 3A), Fayochytriomyces spinosus (Fay) W. J. Davis, Letcher, Longcore, and M.J. Powell (1 strain; Fig. 3C), Chytriomyces hyalinus Karling (12 strains; Fig. 3D), Borealophlyctis nickersoniae W.J. Davis, Letcher, and M.J. Powell (1 strain; Fig. 3E), Geranomyces variabilis (1 strain; 3F), Southeastern Naturalist W.J. Davis, J. Antonetti, P.M. Letcher, and M.J. Powell 2016 Vol. 15, No. 3 540 and Boothiomyces macroporosum (Karling) Letcher (2 strains; Fig. 3G). The 7 species are from 4 orders: Chytridiales (4 species), Rhizophlyctidales (1 species), Spizellomycetales (1 species), and Rhizophydiales (1 species). Two strains, WJD187 and WJD185, were morphologically identified as Rhizoclosmatium aurantiacum. Strain WJD187 formed a monophyletic clade with the representative sequence of Figure 3. Representative light micrographs of the observed chytrid taxa. (A) Rhizoclosmatium globosum on fairy shrimp (a microcrustacean commonly observed in temporary forest ponds) exoskeleton. (B) Rhizoclosmatium aurantiacum on chitin. (C) Fayochytriomyces spinosus on cellulose. (D) Chytriomyces hyalinus on agar. (E) Borealophlyctis nickersoniae on agar. (F) Geranomyces variabilis on agar. (G) Boothiomyces macrosporosum on pollen. Scale bars in A and G = 25 μm; in B, C, D, and E = 10μm; and in F = 50 μm. Abbreviations: Ap = apophysis, Ce = cellulose, Ch = chitin, P = pollen, R = rhizoid, Sp = sporangium, and ZC = zoospore cyst. Southeastern Naturalist 541 W.J. Davis, J. Antonetti, P.M. Letcher, and M.J. Powell 2016 Vol. 15, No. 3 R. aurantiacum in GenBank, strain EL002, with 100% bootstrap support (Fig. 4) and was 99% similar. Strain WJD185, which was 10% divergent from strain WJD187, was sister to R. aurantiacum with weak support (57% bootstrap support). Most strains were isolated from the baits, especially cellulose and pollen. However, strains WJD183 (Chytriomyces hyalinus) and WJD185 (Rhizoclosmatium sp.) were isolated from a dragonfly wing in the collected sample, and strain WJD187 (Rhizoclosmatium aurantiacum) was isolated from a Daphnia exoskeleton. Figure 4. Molecular phylogeny of strains isolated from the temporary forest pond inferred with an alignment of partial 28S sequences. Numbers at the nodes are ML bootstrap values. Southeastern Naturalist W.J. Davis, J. Antonetti, P.M. Letcher, and M.J. Powell 2016 Vol. 15, No. 3 542 Discussion The purpose of this preliminary study was to determine what chytrids were present in a temporary forest pond. Food webs within temporary forest ponds are dependent upon the decomposition of terrestrial inputs, such as leaf litter and pollen, yet little is known about their microbial communities (Bärlocher et al. 1978, Carrino-Kyker and Swanson 2008, Carrino-Kyker et al. 2011, Colburn 2004, Williams 2006). Chytrids are a substantial and potentially important component of the microbial community that degrade leaf litter in streams (Bärlocher et al. 2012, Marano et al. 2011, Nikolcheva and Bärlocher 2004, Seena et al. 2008, Sridhar et al 2011), and it has been demonstrated that chytrids provide a vital link between higher and lower trophic levels in lakes (Kagami et al. 2014). Our study supplements earlier observations of chytrids in temporary ponds, demonstrates the phylogenetic diversity of the chytrids present, and lays the foundation for determining their role in food webs of temporary forest ponds. Our results suggest that a mycoloop (Kagami et al. 2014) exists in temporary forest ponds, and that through it the energy and nutrients contained in recalcitrant terrestrial inputs and autochthonous biomass (e.g., microcrustacean exuviae) are made available to higher trophic levels. Leaf litter and pollen are important terrestrial inputs to temporary forest ponds (Colburn 2004, Williams 2006). We observed chytrids that degraded cellulose (e.g., Fayochytriomyces spinosus [Davis et al. 2015]) and pollen (e.g., Borealophlyctis nickersoniae [Davis et al. 2016] and Boothiomyces macroporosum; Fig. 3G). We also observed chytrids degrading fairy shrimp exoskeletons (Fig. 3A), dragonfly wings (strains WJD183 and WJD185), and Daphnia exoskeletons (strain WJD187). As these chytrids complete their life cycle, they produce zoospores that can be a high-quality food source for various filter feeders (Kagami et al. 2014). The filter feeders then provide food for predators such as Lithobates sylvaticus (LeConte) (Wood Frog) tadpoles, salamanders, and aquatic insects (Colburn 2004). Thus, chytrids are potentially a stabilizing link in the food web that supports amphibian populations and biodiversity, and our study is one step in elucidating that link. As well, the strains cultured in this study have and may contribute to chytrid systematics and taxonomy. Strain WJD186 was used as the ex-type culture for the new combination Fayochytriomyces spinosus (Davis et al. 2015). Strain WJD170 was used to inform the description of Borealophlyctis nickersoniae (Davis et al. 2016). Strains WJD185 and WJD187 both represent the morphological species Rhizoclosmatium aurantiacum, but their divergence suggests that the broad scope of the morphological concept of the species may encompass additional new species. However, due to the limitations of culture-based methods, our study likely underestimates chytrid diversity in the temporary forest pond. Finding and culturing chytrids from baited collections are time consuming and are limited by the ability of organisms to live on artificial media. Culture-based methods are biased toward the detection of chytrids capable of axenic, saprobic growth (Lozupone and Klein 2002). This excludes parasitic lineages and taxa that require co-culture with other organisms, such as Rhizidium phycophilum K.T. Picard (Picard et al. 2009, Southeastern Naturalist 543 W.J. Davis, J. Antonetti, P.M. Letcher, and M.J. Powell 2016 Vol. 15, No. 3 2013). Since we cultured on general media, and chytrids have varied nutritional needs (Barr 1970, Digby et al. 2010, Midgley et al. 2006, Willoughby 1962b), the method probably selected for generalist species. We incubated all strains at room temperature; however, some chytrids require cooler temperatures for growth (Longcore 1992a, b; Longcore and Simmons 2012; Shin et al. 2001; Van Donk and Ringelberg 1983). One way to compensate for the limitations of a culture-based study is to supplement it with other methods. Early researchers used light microscopy and camera lucida drawings to report chytrid diversity (e.g., Dogma 1969, Paterson 1967). Molecular inventories of aquatic habitats revealed many chytrid phylotypes that do not correspond to any taxa for which we have sequence data (e.g., Lefèvre et al. 2007, 2008, 2012; Monchy et al. 2011). Before molecular inventories are meaningful, however, databases need to be populated with DNA sequences from classical or newly described strains in axenic culture; such strains only come from the time-consuming practice of baiting followed by observation and isolation. Thus, to inventory the chytrids present in any habitat, a multiphasic approach is needed. Acknowledgments We thank Eric Van Abel, Kimberly Peden, Andrew DeAtkine, Jessica Fults, Rebecca Holland, and Trey Melton for helping with the collection and processing of samples. We thank Walter Smith for telling us where we could find the study pond. We thank Joe Koloski and Cynthia Ragland for aid in applying for a research permit and the National Forest Service for granting permit 2660. 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