W.J. Davis, P.M. Letcher, and M.J. Powell 2013 Southeastern Naturalist Vol. 12, No. 4 666 2013 SOUTHEASTERN NATURALIST 12(4):666–683 Chytrid Diversity of Tuscaloosa County, Alabama William J. Davis1,*, Peter M. Letcher1, and Martha J. Powell1 Abstract - Alabama is a biodiversity hotspot. The diversity of chytrid fungi, however, is underexplored. For this reason, we used standard bait-culturing techniques to sample habitats in Tuscaloosa County for chytrids. We cultured 100 isolates; the majority was assigned to 23 taxa belonging to 6 of the 7 recognized orders. Some could not be assigned to a currently described taxon. The majority of isolates belonged to one of three taxa: Chytriomyces hyalinus, Rhizoclosmatium globosum, and Boothiomyces macroporosum. This result demonstrates that chytrid communities in Tuscaloosa County, as elsewhere, are composed of a few common and many uncommon to rare taxa. The presence of unidentified chytrid isolates demonstrates the need for further sampling in Alabama, and the potential for this sampling to broaden our understanding of chytrid diversity. Introduction Alabama is a biodiversity hotspot for several taxa (Boschung and Mayden 2004) and is home to many endemic, endangered, and threatened animals and plants (ANHP 2011). Alabama also has a diverse fungal biota (Atkinson 1897, Gray and Morgan-Jones 1979, Hollis 1954, Morgan-Jones 1974). Our understanding of the scope of this diversity is limited by a lack of recent activity and attempts to update the taxonomy of former records. As a result, Alabama’s mycobiota is poorly documented (Morgan-Jones 1974). Due to the importance of agriculture in the state, most fungal surveys have focused on pathogens and potential pathogens (Morgan-Jones 1974), a trend that continues to the present (e.g., Diamond et al. 2006, Palmateer et al. 2004, Rong et al. 2001, Vargas-Ayala et al. 2000). With a better understanding of fungal taxonomy and new techniques, there have been recent inventories of other groups, such as salt marsh saprophytes (Walker et al. 2010) and trichomycetes (Nelder 2003, Nelder et al. 2010). With the exception of Lefèvre et al. (2012), Nelder (2003), and Nelder et al. (2010), past and contemporary inventories have excluded the early-diverging fungal lineages, such as the chytrid fungi. Chytridiomycota (sensu Hibbett et al. 2007, = chytrid fungi) is an early-diverging lineage of fungi characterized by a motile zoospore with a single, posterior flagellum (Sparrow 1960). Chytrids are primarily aquatic, although numerous species survive in the capillary network of water found around soil particles and can be considered terrestrial (Sparrow 1960). As a group, chytrids appear to be ubiquitous and cosmopolitan across soil and aquatic ecosystems (Barr 1990, Czeczuga et al. 2005, Sparrow 1960). Within habitats, some chytrid taxa occur frequently and regularly while others occur more rarely (Willoughby 1961, 1962); thus, chytrid communities are structured with a few common taxa and many uncommon to rare taxa (Letcher and Powell 2001, Marano et al. 2008). 1Department of Biological Sciences, University of Alabama, Tuscaloosa, AL 35487. *Corresponding author - wjdavis1@crimson.ua.edu. 667 W.J. Davis, P.M. Letcher, and M.J. Powell 2013 Southeastern Naturalist Vol. 12, No. 4 Chytrids are often overlooked due to their inconspicuous, microscopic nature and difficulties with identification (Powell 1993). Nevertheless, early researchers recognized that they are potentially key components of aquatic and terrestrial ecosystems (Christensen 1951, Sparrow 1960). Indeed, chytrids act as important parasites and saprobes (Powell 1993, Sparrow 1960, Wakefield et al. 2010). Recent molecular inventories have indicated that chytrids may dominate the fungal communities in the pelagic zones of lakes (Monchy et al. 2011) and in alpine soils (Freeman et al. 2009, Schmidt et al. 2012); consequently, they likely impact the flow of nutrients and energy through the food webs in these habitats. Although research is beginning to unravel how parasitic chytrids influence aquatic food webs (e.g., Grami et al. 2011, Ibelings et al. 2004, Kagami et al. 2007, Miki et al. 2011, Niquil et al. 2011, van Donk 1989), many aspects of chytrid ecology—for example, the abiotic and biotic factors determining the spatial structure of chytrid communities—remain to be explored. Prior to molecular inventories, most studies of chytrid diversity were observation- or culture-based local inventories. For this reason, chytrid diversity has been well documented in only a few locations, such as the English Lake District (Willoughby 1961, 1962) and the Douglas Lake region of Michigan (Dogma 1969, Paterson 1967, Sparrow 1952), and poorly documented in others. Recent taxonomic revisions have sampled more broadly and have included 12 isolates from Alabama (e.g., Letcher et al 2006, 2012; Simmons 2011; Vélez et al. 2011; Wakefield et al. 2010). Some of these isolates are phylogenetically unique, such as unidentified species WB235A in Fig. 1 of Vélez et al. (2011), and may represent new species. A recent report from AL has expanded the known range of Blyttiomyces spinulosus (Blytt) Bartsch (Blackwell et al. 2011). Lefèvre et al. (2012) inventoried two lakes in Tuscaloosa County, AL using a combination of culture-based and molecular techniques. Their results indicated that chytrids dominated the fungal communities in these lakes. Moreover, approximately half of the chytrid sequences found did not cluster with currently described taxa (Lefèvre et al. 2012). This finding suggests that an exploration of chytrid diversity in AL would benefit a broader understanding of chytrid diversity as well as add to the knowledge of Alabama’s mycobiota. Thus, the purpose of this study was to investigate the chytrid diversity within Tuscaloosa County, with an emphasis on Lake Lurleen. We predicted that chytrids found in Tuscaloosa County consist of an assemblage of a few common and many uncommon to rare taxa, with some unique taxa, and that the common taxa are the same as those found globally. Study Sites Soil and water samples were collected from a variety of locations and habitats around Tuscaloosa County. Tuscaloosa County has a total area of approximately 3500 km2 and is located in west-central AL. Soils in the northeast portion of the county are derived from the Appalachian Highlands, while soils to the west and south are part of the Gulf Coast Plain (Johnson et al. 1981). Four main aquatic sites were sampled: Lake Lurleen, Lake Nicol, the Black Warrior River, and Marr’s Spring. Lake Lurleen (33.291014, -87.511253) is a 101-ha W.J. Davis, P.M. Letcher, and M.J. Powell 2013 Southeastern Naturalist Vol. 12, No. 4 668 reservoir located in a 658-ha acre state park. Lake Nicol (33.307315, -87.479989) is a 156-ha reservoir. According to Johnson et al. (1981), the soils surrounding both reservoirs are mainly acidic, well-drained loams with moderate to high water capacity. Lefèvre et al. (2012) designated Lake Lurleen as meso-eutrophic and found it to have a metalimnetic oxygen maximum during the stratified period (May to November). To date, basic limnological data for Lake Nicol have not been measured. The Black Warrior River is a 268-km tributary of the Tombigbee River and part of the Mobile River drainage basin (Ward et al. 2005). It begins in the Appalachian Plateau region of AL (northeast corner) and crosses the Fall Line, the transition from the Appalachian Plateau to the Coastal Plain, near the city of Tuscaloosa (Ward et al. 2005). The Black Warrior River has a mean flow of 277 m3s-1 with maximum flow occurring in March (Ward et al. 2005). Unfortunately, the Black Warrior River has been heavily impacted by activities associated with urban centers, such as Tuscaloosa and Birmingham, mining activity, and agriculture (Mette et al. 1989). The Black Warrior River has been impounded by a system of locks and dams and is kept at a width of 61 m and a depth of 3 m for the transportation of coal (Mette et al. 1989). Marr’s Spring is a modified spring located on the University of Alabama campus. It has a concrete bottom and is surrounded by flower beds. In addition, a few samples were taken opportunistically from roadside ditches, ponds, yards, and pastures across Tuscaloosa County, AL. Methods Sampling and isolation Soil and aquatic samples were opportunistically collected from the banks, shallows, and pelagic regions of Lake Lurleen, Lake Nicol, the Black Warrior River, Marr’s Spring, and other locations across Tuscaloosa County. Collections of approximately 200 mL of water or 200 g of soil were made by hand (i.e., dipping a container in the water or scooping soil into a bag). Samples were stored in whirl-top plastic bags, kept cool with ice, and transferred to the laboratory. In the laboratory, portions of each sample were placed in Petri dishes, and sterile water was added to the soil samples. Samples were baited with sterile pollen from Pinus spp. (pine) and Liquidambar styraciflua L. (Sweetgum), cellulose (onion epidermal cells), keratin (snake skin), and chitin (shrimp exoskeleton; Couch 1939, Sparrow 1960). Baits (added substrates) and natural substrates already present (e.g., senescing or dead algae, insect exuviae, etc.) were examined microscopically and periodically for chytrid thalli. Standard techniques (Couch 1939, Sparrow 1960) were used to bring observed chytrids into pure culture. Due to the limitations of the methods employed, only saprophytic chytrids were brought into culture and used in subsequent analyses. Any chytrid that was observed but not brought into culture was not included in the analyses. Chytrids brought into pure culture, hereafter isolates, were maintained on PmTG (1 g peptonized milk, 1 g tryptone, 5 g glucose, and 8 g agar per liter of water), mPmTG (0.4 g peptonized milk, 0.4 g tryptone, 2 g glucose, and 8 g agar per liter of water), or Archimycete Media (2 g peptonized milk, 3 g malt extract, 5 g glucose, and 8 g agar per liter of water) nutrient agar plates (CBS 2013) and transferred at 2 month 669 W.J. Davis, P.M. Letcher, and M.J. Powell 2013 Southeastern Naturalist Vol. 12, No. 4 intervals to maintain viability. Plates were sealed and maintained in the dark at room temperature. Isolates were tentatively identified using morphological features with the aid of Sparrow (1960), Karling (1977), and relevant literature. DNA extraction, PCR, and sequencing For DNA extractions, isolates were grown in 50 mL of nutrient broth, which is the same as nutrient agar without the agar. Broth cultures were then centrifuged in 50-mL Falcon tubes for 20 minutes at 3000 rpm in a ThermoIEC I-703a centrifuge (ThermoIEC, Needham Heights, MA) to pellet chytrid thalli. DNA was extracted from the pellet using the NucleoSpin Plant II DNA extraction kit (Macherey-Nagel, Inc., Bethlehem, PA) and the NucleoSpin protocol for fungal cultures (Macherey-Nagel, Inc. 2008). DNA concentration was determined with spectrophotometry using a Nanodrop (Nanodrop, Wilmington, DE). The DNA was diluted to 10 ng/μL for PCR. The D1/D2 region of the 28s large ribosomal subunit (800–900bp from the 5’ end) has been used as a molecular marker to delineate taxa within Chytridiomycota (e.g., Letcher et al. 2006, Longcore and Simmons 2012, Simmons 2011, Simmons et al. 2009, Vélez et al. 2011, Wakefield et al. 2010), and phylogenies inferred with this region are congruent with those inferred from zoospore ultrastructure (Letcher et al. 2005). As a result, there is a database of taxonomically reliable sequences available, and we chose this region for use in our study. The region was amplified using the LROR/LR5 primer pair (Rehner and Samuels 1994, Vilgalys and Hester 1990) for 30 cycles of 1 min at 94 °C, 1 min at 50 °C, 1 min at 72 °C with an initial denaturing at 94 °C for 2 mins, and final elongation at 72 °C for 5 mins. Four replicate amplifications were pooled and cleaned following the protocols of the Nucleospin Extraction II kit (Macherey-Nagel, Inc. 2009). Amplicons were sequenced (Macrogen USA, Rockville, MD) and assembled into contiguous sequences using Sequencher 4.5 (Genecodes) as described by Letcher et al. (2004b). Sequences were searched against the GenBank database using the blastn algorithm (Altschul et al. 1990) to corroborate the tentative morphology-based iden tification. Alignment and phylogenetic analysis Taxonomically reliable reference sequences used in revisions of the Chytridiomycota were downloaded from GenBank. In two cases (MP53 and EL102), the reference sequences were also obtained from the study area and so were included in subsequent calculations. All sequences were aligned with ClustalX (Thompson et al. 1997) and manually adjusted with BioEdit (Hall 1998). Maximum parsimony (MP) trees were inferred using PAUPRat (Sikes and Lewis 2002), and maximum likelihood (ML) trees were inferred using RAxML 7.0.3 (Stamatakis 2006) under the GTR + G model of nucleotide substitution as determined by ModelTest 3.7 (Posada and Crandall 1998). A 50% majority rule consensus-tree was generated from the MP trees and bootstrapped in PAUP* (Swofford 2002). The best ML-tree was bootstrapped with 1000 replicates using the rapid bootstrap option (Stamatakis et al. 2008). The inferred trees were rooted with Monoblepharella mexicana (UCB 78-1 from James et al. 2006), a member of the Monoblepharidiomycota (Doweld 2001), the sister group of Chytridiomycota. W.J. Davis, P.M. Letcher, and M.J. Powell 2013 Southeastern Naturalist Vol. 12, No. 4 670 Results Phylogenetic analysis The alignment contained 100 isolates and 992 characters, of which 521 were parsimony-informative. The inferred MP (2615 steps, CI = 0.408, RI = 0.891) and ML (-lnL = 14427.072300) trees were incongruent concerning the placement of the Spizellomycetales and Polychytriales. Neither could resolve the relationships between the orders. However, the trees were congruent concerning the placement of taxa into families, genera, and species. Thus, only the ML tree is illustrated (Fig. 1). Phylogenetic distribution The 100 isolates were delineated into taxa at the order, family, genus, and species level using the monophyletic phylogenetic species concept (Mayden 1999). Specifically, an isolate was considered a member of a taxon if it formed a monophyletic clade with a reference sequence of that taxon. Chytridiomycota contains seven orders: Chytridiales, Rhizophydiales, Spizellomycetales, Rhizophlyctiales, Polychytriales, Cladochytriales, and Lobulomycetales. Isolates obtained in this study grouped with reference sequences in the Chytridiales, Rhizophydiales, Spizellomycetales, Rhizophlyctidales, Polychytriales, and Cladochytridales. The isolates were unevenly distributed among these orders (Fig. 1). Lobulomyces poculatus (Willoughby) Simmons, a representative of the Lobulomycetales, was observed but attempts to culture it failed. The majority of the isolates (52%) was placed in Chytridiales and represented one of two families within the order (Vélez et al. 2011). Approximately one-third (34%) of the isolates represented eight of ten families within the Rhizophydiales (Letcher et al. 2006, 2008b). All of the families within Spizellomycetales were represented by 9% of the isolates. Two isolates (2%) represented one of four families in the Rhizophlyctidales (Letcher et al. 2008a). Two isolates (2%) also represented the Polychytriales, and one isolate (1%) represented the Cladochytriales. The vast majority (96%) of the isolates was assigned to 23 described taxa at the genus or species level (Appendix 1). The isolates were also unevenly distributed among these taxa, with 24% of the isolates grouping into the clade corresponding to the morphospecies Chytriomyces hyalinus Karling, 15% grouping with the morphospecies Rhizoclosmatium globosum Petersen, and 17% grouping with Boothiomyces macroporosum (Karling) Letcher (Fig. 1). Most taxa were represented by few isolates. Geographic distribution Thirty-nine of the isolates came from Lake Lurleen (Appendix 1). These isolates represented nine taxa at the genus and species level: Rhizoclosmatium globosum, Rhizidium sp., Siphonaria petersenii Karling, Chytriomyces hyalinus, Geranomyces variabilis (Powell) Simmons, Boothiomyces macroporosum, Kappamyces laurelensis Letcher, Polychytrium aggregatum Ajello, and Angulomyces argentinensis Letcher. Two of the Lake Lurleen isolates—WB228 and WB235A— did not form monophyletic clades with currently described taxa. 671 W.J. Davis, P.M. Letcher, and M.J. Powell 2013 Southeastern Naturalist Vol. 12, No. 4 Figure 1. Maximum likelihood tree (-lnL = 14427.072300) of Tuscaloosa County chytrid isolates. Only clades with >50% bootstrap support are shown, with the exception of the Rhizophlyctidales. Branches cut with a │are half their original length W.J. Davis, P.M. Letcher, and M.J. Powell 2013 Southeastern Naturalist Vol. 12, No. 4 672 Polychytrium aggregatum, Kappamyces laurelensis, and Siphonaria petersenii were only isolated from Lake Lurleen. Thirteen isolates came from Lake Nicol (Appendix 1) and were grouped into six taxa: Chytriomyces hyalinus, Rhizoclosmatium globosum, Globomyces pollinis-pini (Braun) Letcher, Protrudomyces lateralis (Braun) Letcher, Rhizophydium sp., and Gorgonomyces haynaldii (Schaaraschm) Letcher. Protrudomyces lateralis and G. haynaldii were only isolated from Lake Nicol. The Black Warrior River yielded eleven isolates that were identified as Alphamyces chaetifer (Sparrow) Letcher, Rhizophydium sp., G. pollinis- pini, C. hyalinus, Rhizidium sp., Fimicolochytrium alabamae Simmons, and Fimicolochytrium jonesii Simmons (Appendix 1). The Black Warrior River was the only source of F. jonesii. Nine isolates and eight taxa (Appendix 1) were from Marr’s Spring: Rhizidium sp., Rhizophydium globosum (Braun) Rabenhorst, A. chaetifer, Cladochytrium replicatum Karling, Boothiomyces macroporosum, Gorgonomyces sp., Angulomyces argentinensis, and C. hyalinus. Cladochytrium replicatum was isolated only from Marr’s Spring. Discussion The purpose of our study was to explore chytrid diversity in Tuscaloosa County, AL. We were motivated by the need to document Alabama’s diverse mycobiota, specifically the understudied chytrid fungi. Approximately 95% of the isolates form monophyletic clades with previously identified taxa, which have global distributions. For example, Boothiomyces macroporosum has been found in Australia, Argentina, and Canada (Letcher et al. 2006, 2008b). Gaertneriomyces semiglobifer (Uebelmesser) Barr has been observed or isolated from Germany, Israel, and Australia (Wakefield et al. 2010). Globomyces pollinis-pini has been observed or isolated from Russia, China, and Cuba (Sparrow 1960); Douglas Lake, MI (Sparrow 1952); and Argentina (Letcher et al. 2008b). Thus, the chytrid taxa found in AL are the same as those found globally, which is in agreement with our initial prediction. This is the first time that many of these taxa have been reported from AL. Thus, our sampling in AL has expanded the known distribution of these taxa and has corroborated the view that chytrids are cosmopolitan (Barr 1990, Czeczuga et al. 2005, Sparrow 1960). Approximately half of the isolates belong to the clades of Chytriomyces hyalinus, Rhizoclosmatium globosum, and Boothiomyces macroporosum. The rest of the taxa were represented by few isolates. Although frequency and abundance were not calculated, it can be inferred from the phylogenetic distribution of isolates that chytrid communities in AL are dominated by a few common species; thus, there are a few common species and many uncommon to rare species in Alabama. This is in agreement with general ecological theory, the results of surveys conducted in Virginia (Letcher and Powell 2001), Australia (Letcher et al. 2004a, b), Brazil (Nascimento et al. 2011a, b), Argentina (Marano et al. 2008), Canada (Lee 2000), and Great Britain (Willoughby 1961, 1962), and our initial prediction. We were able to assign some isolates to a genus but not to a species, e.g., isolates in the Triparticalcar, Gorgonomyces, Rhizidium, and Rhizophydium clades. This is 673 W.J. Davis, P.M. Letcher, and M.J. Powell 2013 Southeastern Naturalist Vol. 12, No. 4 a common occurrence and has been reported for other taxa such as Triparticalcar and Powellomyces (Wakefield et al. 2010) and Cladochytrium (Mozley-Standridge et al. 2009). The lack of resolution at the species level could be due to exclusive use of the D1/D2 region of the 28s rDNA gene, which is good at resolving chytrid families and genera but may have limited utility at the species level (Letcher et al. 2004b). It could also be the result of an insufficient number of available sequences from those genera. Alternatively, these isolates could represent unknown phylogenetic diversity, which could correspond to new species. Such phylogenetic diversity was previously reported for the genus Powellomyces by Wakefield et al. (2010), which was delineated into new species by Simmons (2011) and Simmons and Longcore (2012). Four isolates could not be assigned to a genus or a species: WB228, WB235A, MP041, and WJD150. Isolate WB235A was previously reported by Vélez et al. (2011) as sister to Chytriomyces hyalinus. In our study, isolate MP041 is sister to it, and their position within Chytridiales is unresolved. Isolate WB228 is an early diverging lineage within the Chytridiales. Isolate WJD150 is sister to isolate PL157, an undescribed isolate from Argentina (Letcher et al. 2008b). These isolates could represent undescribed taxa or described but not sequenced taxa. Thus, it can also be concluded that AL samples may reveal novel taxa and unknown phylogenetic diversity within described taxa. As a result, our AL samples can aid in the current exploration and refinement of chytrid species concepts (Longcore 2004, Simons and Longcore 2012). A total of 23 described taxa were isolated in our study. This is comparable to the diversity reported by other inventories. Marano et al. (2008, 2011) reported 16 species from the Las Cañas stream near Buenos Aires, Argentina. Thirteen species have been recorded from the Reserva Natural Selva Marginal Punta Lara, Argentina (Arellano et al. 2009, Marano et al. 2008). Nascimento et al. (2011a, b) have reported 20 species from the Reserva Biológica de Mogi Guaçu, Brazil. However, given that 34 species were recorded from the English Lake District by Willoughby (1961, 1962) and approximately 60 species have been recorded from the Douglas Lake Region, MI (Dogma 1969, Paterson 1967, Sparrow 1952), it is likely that Tuscaloosa County is under-sampled for chytrid diversity. Of the 23 taxa isolated from Tuscaloosa County, nine were reported from Lake Lurleen, the most heavily sampled site. This number is comparable to the findings of Kiziewicz and Nalepa (2008), who reported five species from a site on Lake Michigan near Muskegon, MI. It is also comparable to the numbers reported for the Reserva Natural Selva Marginal Punta Lara and Las Cañas stream. However, a comparison of our results for Lake Lurleen to Lefèvre et al.’s (2012) results suggests much of the chytrid diversity in Lake Lurleen was missed in our study. Lefèvre et al. (2012) reported 18 unique chytrid phylotypes and 3 cultured chytrids from Lake Lurleen for a total of 21 taxa. Of those 21 taxa, only three—Chytriomyces hyalinus, Rhizoclosmatium globosum, and Kappamyces laurelensis—were the same as ones isolated in our study. Also, the majority of the phylotypes did not form clades with previously described species or genera (Lefèvre et al. 2012), which further supports the contention that Lake Lurleen has been under-sampled. W.J. Davis, P.M. Letcher, and M.J. Powell 2013 Southeastern Naturalist Vol. 12, No. 4 674 Part of this under-sampling is due to the intensity of sampling. In order to fully sample chytrid diversity, repeated temporal (Willoughby 1961, 1962) and spatial (Letcher and Powell 2001) sampling is necessary. Most sites included in this study were not sampled as intensely as those in Willoughby (1961, 1962) and Letcher and Powell (2001). Thus, the chytrid diversity at each site was undersampled, resulting in an overall under-sampling of the chytrid diversity across Tuscaloosa County. This under-sampling can also be explained by the limitations of a culture-based inventory, which have been reviewed by Lozupone and Klein (2002). Our results include only those chytrids capable of saprophytic growth on nutrient media. All of the parasitic lineages have been excluded, which reduces the total number of species reported and the phylogenetic diversity sampled (Letcher et al. 2004b). The inclusion of these taxa will require the development of new techniques in chytrid isolation and culturing. Alternatively, a molecular inventory would reveal more diversity because it would include difficult-to-culture taxa, such as those in the Lobulomycetales (Simmons et al. 2009). However, molecular inventories are only useful when there is reliable and abundant sequence information available for a lineage (Lozupone and Klein 2002). This fact is demonstrated by the findings of Lefèvre et al. (2012). It remains to be seen whether Lefèvre et al.’s (2012) phylotypes are novel taxa or merely unsequenced, described taxa, and this ambiguity highlights the need for more taxonomic work in Chytridiomycota and an increase in the number of taxonomically reliable sequences. Although recent molecular revisions of Chytridiomycota have greatly increased the number of chytrid sequences available, these studies were all bait-culture based. Since the majority of chytrid species are described based on morphological characters (Longcore 1996, Sparrow 1960), bait-culture studies will be crucial to building a molecular database necessary for molecular inventories to be useful. The sequences we have generated represent the beginning of such a database for future molecular inventories that might take place in Alabama. Our results suggest that the same phylogenetic depth and diversity (Faith 1992) seen with a global sampling is mirrored on a local scale. Further analysis is required to determine the amount of biodiversity not detected by local sampling. Increased sampling of the state will document a previously excluded portion of Alabama’s mycobiota biodiversity and broaden our understanding of chytrid diversity. Acknowledgments Thanks are extended to Dr. Carol Duffy, Rebecca Holland, Sharmeka Lewis, Scotty De- Priest, Samantha Perkins, Trey Milton, Adam Fuller, Nichole Mattheus, Sarah Duncan, and Alissa Vincent for help with collecting the samples. Appreciation is expressed to Dr. Will Blackwell, Antijuan Spivy, Leeanne Bertram, Ben Swann, Keith Atkinson, and Michael Brooks for help in bringing isolates into culture and to Dr. Satoshi Sekimoto and Dr. Emilie Lefèvre for help with extractions and sequencing. Special thanks to Jonathan Antonetti who was involved in multiple areas of the project and to Richard Baird and two anonymous reviewers for careful review and helpful suggestions. Funding was kindly provided by the National Science Foundation (NSF REVSYS 0949305), the McNair Graduate Fellowship, and the Department of Biological Sciences of the University of Alabama. 675 W.J. Davis, P.M. Letcher, and M.J. Powell 2013 Southeastern Naturalist Vol. 12, No. 4 Literature Cited Alabama Natural Heritage Program (ANHP). 2011. Home page. Auburn University Environmental Institute, Auburn University, Auburn, AL. Available online at http://www. alnhp.org. Accessed 25 February 2013. Altschul, S.F., W. Gish, W. Miller, and E.W. Myers. 1990. 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GenBank 28s Culture number/taxonomic affiliation accession # Location Isolates Chytridiales MP053 Chytriomyces hyalinus JN049526 Lake Lurleen, AL AQ Chytriomyces hyalinus KC691310 Lake Lurleen, AL WJD105 Chytriomyces hyalinus KC691369 Lake Lurleen, AL WJD107 Chytriomyces hyalinus KC691370 Lake Lurleen, AL WJD108 Chytriomyces hyalinus KC691371 Lake Lurleen, AL WJD112 Chytriomyces hyalinus KC691375 Lake Lurleen, AL WJD136 Chytriomyces hyalinus KC691386 Lake Lurleen, AL WJD139 Chytriomyces hyalinus KC691389 Lake Lurleen, AL BD Chytriomyces hyalinus KC691311 Lake Nicol, AL JA003 Chytriomyces hyalinus KC691314 Lake Nicol, AL MP083 Chytriomyces hyalinus KC691351 Black Warrior River, AL MP089 Chytriomyces hyalinus KC691354 Black Warrior River, AL WJD138 Chytriomyces hyalinus KC691388 Black Warrior River, AL WJD140 Chytriomyces hyalinus KC691390 Black Warrior River, AL JA001 Chytriomyces hyalinus KC691312 Marr’s Spring Pond, AL PL181 Chytriomyces hyalinus KC691357 Lake Tuscaloosa, AL MP004 Chytriomyces hyalinus DQ273836 University of Alabama, AL MP070 Chytriomyces hyalinus KC691345 Cottondale, AL MP080 Chytriomyces hyalinus KC691348 Cottondale, AL WB216 Chytriomyces hyalinus KC691358 Cottondale, AL MP066 Chytriomyces hyalinus KC691342 Northport, AL MP068 Chytriomyces hyalinus JX905526 Northport, AL MP069 Chytriomyces hyalinus KC691344 Northport, AL WB241 Chytriomyces hyalinus KC691366 Northport, AL MP081 Chytriomyces hyalinus KC691349 Tuscaloosa, AL MB001 Rhizoclosmatium globosum KC691318 Lake Lurleen, AL MB007 Rhizoclosmatium globosum KC691322 Lake Lurleen, AL MB037 Rhizoclosmatium globosum KC691329 Lake Lurleen, AL MB038 Rhizoclosmatium globosum KC691330 Lake Lurleen, AL MB048 Rhizoclosmatium globosum KC6911331 Lake Lurleen, AL WB219 Rhizoclosmatium globosum KC691360 Lake Lurleen, AL WJD111 Rhizoclosmatium globosum KC691374 Lake Lurleen, AL WJD143 Rhizoclosmatium globosum KC691391 Lake Lurleen, AL WB235C Rhizoclosmatium globosum KC691364 Lake Lurleen, AL JA002 Rhizoclosmatium globosum KC691313 Lake Nicol, AL JA004 Rhizoclosmatium globosum KC691315 Lake Nicol, AL JA005 Rhizoclosmatium globosum KC691316 Lake Nicol, AL WB236B Rhizoclosmatium globosum KC691365 Lake Nicol, AL 681 W.J. Davis, P.M. Letcher, and M.J. Powell 2013 Southeastern Naturalist Vol. 12, No. 4 GenBank 28s Culture number/taxonomic affiliation accession # Location WB218 Rhizoclosmatium globosum KC691359 Cottondale, AL WB224 Rhizoclosmatium globosum KC691361 Cottondale, AL EL102 Rhizoclosmatium aurantiacum JN049529 Lake Lurleen, AL MP067 Rhizoclosmatium aurantiacum KC691343 Lake Nicol, AL MP046 Rhizoclosmatium sp. KC691335 Lake Lurleen, AL MP056 Rhizoclosmatium sp. KC691339 Marr’s Spring, AL MB013 Rhizidium sp. KC691324 Lake Lurleen, AL MP051 Rhizidium sp. KC691338 Lake Lurleen, AL MP040 Rhizidium sp. KC691332 Marr’s Spring, AL MP087 Rhizidium sp. KC691352 Black Warrior River, AL MP088 Rhizidium sp. KC691353 Black Warrior River, AL WB235A Chytriomyces sp. FJ822968 Lake Lurleen, AL MP041 Chytriomyces sp. JX905522 Tuscaloosa, AL WB235B Siphonaria petersenii KC691363 Lake Lurleen, AL WB228 Unidentified Chytridiales sp. KC691362 Lake Lurleen, AL Rhizophydiales MB006 Boothiomyces macroporosum KC691321 Lake Lurleen, AL MB012 Boothiomyces macroporosum KC691323 Lake Lurleen, AL MB016 Boothiomyces macroporosum KC691325 Lake Lurleen, AL MB017 Boothiomyces macroporosum KC691326 Lake Lurleen, AL MB019 Boothiomyces macroporosum KC691327 Lake Lurleen, AL MB020 Boothiomyces macroporosum KC691328 Lake Lurleen, AL MP063 Boothiomyces macroporosum KC691340 Lake Lurleen, AL MP075 Boothiomyces macroporosum KC691347 Lake Lurleen, AL WJD102 Boothiomyces macroporosum KC691367 Lake Lurleen, AL WJD109 Boothiomyces macroporosum KC691372 Lake Lurleen, AL WJD110 Boothiomyces macroporosum KC691373 Lake Lurleen, AL WJD117 Boothiomyces macroporosum KC691376 Lake Lurleen, AL WJD128 Boothiomyces macroporosum KC691381 Marr’s Spring, AL P065 Boothiomyces macroporosum KC691341 Northport, AL WJD118 Boothiomyces macroporosum KC691377 Northport, AL WJD127 Boothiomyces macroporosum KC691380 Northport, AL PL133 Terramyces subangulosum DQ485584 Northport, AL JA006 Gorgonomyces haynaldii KC691317 Lake Nicol, AL WJD130 Gorgonomyces sp. KC691383 Marr’s Spring, AL MP045 Alphamyces chaetifer JF809855 Black Warrior River, AL MP048 Alphamyces chaetifer JF809857 Marr’s Spring, AL WJD154 Kappamyces laurelensis KC691397 Lake Lurleen, AL MP050 Rhizophydium globosum KC691337 Tuscaloosa, AL WJD145 Rhizophydium sp. KC691393 Lake Nicol, AL MP043 Rhizophydium sp. KC691334 Black Warrior River, AL MP042 Rhizophydium sp. KC691333 Tuscaloosa, AL MP049 Rhizophydium sp. KC691336 Tuscaloosa, AL WJD132 Globomyces pollinis-pini KC691384 Lake Nicol, AL WJD133 Globomyces pollinis-pini KC691385 Lake Nicol, AL W.J. Davis, P.M. Letcher, and M.J. Powell 2013 Southeastern Naturalist Vol. 12, No. 4 682 GenBank 28s Culture number/taxonomic affiliation accession # Location MP082 Globomyces pollinis-pini KC691350 Black Warrior River, AL WJD137 Angulomyces argentinensis KC691387 Lake Lurleen, AL WJD129 Angulomyces argentinensis KC691382 Marr’s Spring, AL WJD144 Protrudomyces lateralis KC691392 Lake Nicol, AL WJD150 Unidentified Rhizophydiales sp. KC691395 University of Alabama, AL Spizellomycetales MB004 Geranomyces variabilis KC691319 Lake Lurleen, AL MB005 Geranomyces variabilis KC691320 Lake Lurleen, AL PL166 Geranomyces variabilis HQ901699 Tuscaloosa, AL WJD125 Fimicolochytrium alabamae KC691379 Black Warrior River, AL WJD152 Fimicolochytrium jonesii KC691396 Black Warrior River, AL WJD148 Fimicolochytrium jonesii KC691394 Northport, AL MP074 Gaertneriomyces semiglobifer KC691346 Tuscaloosa Co., AL WJD101 Triparticalcar sp. KC788571 Duncanville, AL WJD156 Triparticalcar sp. KC691398 Tuscaloosa Co., AL Rhizophlyctidales JM001 Rhizophlyctis rosea EU379183 Tuscaloosa, AL RT003 Rhizophlyctis rosea EU379197 Tuscaloosa, AL Polychytriales PL071 Polychytrium aggregatum KC691355 Lake Lurleen, AL WJD104 Polychytrium aggregatum KC691368 Lake Lurleen, AL Cladochytriales WJD123 Cladochytrium replicatum KC691378 Marr’s Spring, AL Reference sequences MP053 Chytriomyces hyalinus JN049526 JEL006 Rhizoclosmatium globosum AY439061 EL102 Rhizoclosmatium aurantiacum JN049529 JEL378 Rhizidium sp. DQ273832 KP013 Rhizophydium phycophilum FJ214802 JEL102 Siphonaria petersenii AY439072 PLAUS021 Boothimyces macroporosum AY439040 ARG026 Gorgonomyces haynaldii EF585607 ARG025 Alphamyces chaetifer EF585606 PL098 Kappamyces laurelensis DQ485581 JEL222 Rhizophydium globosum DQ485551 ARG068 Globomyces pollinis-pini EF585625 ARG008 Angulomyces argentinensis EF585595 ARG071 Protrudomyces lateralis EF585628 PL157 Unidentified Rhizophydiales sp. DQ485594 MP003 Geranomyces variabilis HQ901689 JEL538 Fimicolochytrium alabamae HQ901669 JEL569 Fimicolochytrium jonesii HQ901681 683 W.J. Davis, P.M. Letcher, and M.J. Powell 2013 Southeastern Naturalist Vol. 12, No. 4 GenBank 28s Culture number/taxonomic affiliation accession # Location BR043 Gaertneriomyces semiglobifer FJ827702 BR059 Triparticalcar arcticum DQ273826 BR186 Rhizophlyctis rosea AY349079 JEL109 Polychytrium aggregatum AY546686 JEL180 Cladochytrium replicatum NG_027614 Outgroup UCB781 Monoblepharella mexicana DQ273777 Appendix 2. List of described species, with authorities, included in Appendix 1. State records are designated with (**). Taxon name Chytridiales Chytriomyces hyalinus Karling Rhizoclosmatium globosum Petersen ** Rhizoclosmatium aurantiacum (Petersen) Sparrow Siphonaria petersenii Karling ** Rhizophydiales Boothiomyces macroporosum (Karling) Letcher ** Terramyces subangulosum (Braun) Letcher Gorgonomyces haynaldii (Schaarschmidt) Letcher ** Alphamyces chaetifer (Sparrow) Letcher Kappamyces laurelensis Letcher Rhizophydium globosum (Braun) Rabenhorst ** Globomyces pollinis-pini (Braun) Letcher ** Angulomyces argentinensis Letcher ** Protrudomyces lateralis (Braun) Letcher ** Spizellomycetales Geranomyces varibilis (Powell) Simmons Fimicolochytrium alabamae Simmons Fimicolochytrium jonesii Simmons Gaertneriomyces semiglobifer Barr ** Triparticalcar arctiacum Barr Rhizophlyctidales Rhizophlyctis rosea (de Bary and Woronin) Fischer Polychytriales Polychytrium aggregatum Ajello ** Cladochytriales Cladochytrium replicatum Karling ** Outgroup Monoblepharella mexicana Shanor