Mycologia, 103(1), 2011, pp. 10–21. DOI: 10.3852/09-297 # 2011 by The Mycological Society of America, Lawrence, KS 66044-8897
Diversity and distribution of soil fungal communities in a semiarid grassland Andrea Porras-Alfaro1
INTRODUCTION
Department of Biological Sciences, Waggoner Hall 372, 1 University Circle, Western Illinois University, Macomb, Illinois 61455
In aridland ecosystems biological activity is constrained by episodic water availability, high soil pH, extreme soil surface temperature and intense solar radiation. These abiotic characteristics create patchy distributions of grass tussocks and biological soil crust (BSC) and select microbial communities that are compositionally and functionally distinct from those of mesic ecosystems (Collins et al. 2008). BSCs are considered fundamental regulators of the biogeochemistry and geomorphology of semiarid grasslands, accounting for up to 70% of the biotic cover (Belnap 2002). BSCs are dominated by photoautotrophic communities (e.g. algae, mosses, cyanobacteria and/ or lichens) associated with microfauna, bacteria and fungi (Belnap 2002, States et al. 2001). Studies have shown that the top 1 cm of crusted soil has the greatest bacterial diversity (Maestre et al. 2005, Garcia-Pichel et al. 2003). The presence of these microbial communities increases soil fertility and reduces erosion by fixing carbon and nitrogen and aggregating the soil surface (Eldridge et al. 2000). In many desert ecosystems nitrogen fixation in BSC is the primary source of bio-available N (Belnap 2002, Eldridge et al. 2000). The extreme conditions of desert ecosystems constrain primary producers (plants and the BSC phototrophic components) to form symbiotic relationships that aid their establishment and survival (Porras-Alfaro et al. 2008, Khidir et al. 2010). Root exudates are known to attract and maintain symbiotic fungal communities that form mutualistic associations with plant roots and colonize surrounding soil (Morgan et al. 2005). Our limited knowledge about the composition and diversity of BSC and rhizosphere fungal communities precludes a broader understanding of the ecological function of fungi in desert ecosystems. Work at the Sevilleta LTER site and elsewhere indicates that fungal metabolism can dominate N transformation reactions in the soil (McLain and Martens 2005, 2006; Crenshaw et al. 2008) and that this fungal-driven N cycle is supported by biotrophic C from grasses and cyanobacterial crusts (Green et al. 2008) and by the rapid turnover of soil proteins (McLain and Martens 2006, Stursova et al. 2006) rather than by saprotrophic decomposition of an accumulated store of plant litter and SOM (Collins et al. 2008). However little is known about the effect of N enrichment on soil fungal communities in semiarid ecosystems.
Jose Herrera Department of Biology, 100 E. Normal, Truman State University, Kirksville, Missouri 63501
Donald O. Natvig Kendra Lipinski Robert L. Sinsabaugh Department of Biology, MSC03 2020, University of New Mexico at Albuquerque, New Mexico 87131-0001
Abstract: The fungal loop model of semiarid ecosystems integrates microtopographic structures and pulse dynamics with key microbial processes. However limited data exist about the composition and structure of fungal communities in these ecosystems. The goal of this study was to characterize diversity and structure of soil fungal communities in a semiarid grassland. The effect of long-term nitrogen fertilization on fungi also was evaluated. Samples of rhizosphere (soil surrounding plant roots) and biological soil crust (BSC) were collected in central New Mexico, USA. DNA was amplified from the samples with fungal specific primers. Twelve clone libraries were generated with a total of 307 (78 operational taxonomic units, OTUs) and 324 sequences (67 OTUs) for BSC and rhizosphere respectively. Approximately 40% of soil OTUs were considered novel (less than 97% identity when compared to other sequences in NCBI using BLAST). The dominant organisms were dark-septate (melanized fungi) ascomycetes belonging to Pleosporales. Effects of N enrichment on fungi were not evident at the community level; however the abundance of unique sequences, sampling intensity and temporal variations may be uncovering the effect of N in composition and diversity of fungal communities. The fungal communities of rhizosphere soil and BSC overlapped substantially in composition, with a Jaccard abundance similarity index of 0.75. Further analyses are required to explore possible functions of the dominant species colonizing zones of semiarid grassland soils. Key words: biological soil crust, Bouteloua gracilis, dark septate fungi, rhizosphere soil, semiarid grassland Submitted 24 Nov 2009; accepted for publication 19 May 2010. 1 Corresponding author. E-mail:
[email protected]
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PORRAS-ALFARO ET AL.: SOIL FUNGI IN GRASSLAND Descriptions of the fungal components of arid ecosystems based on culture-based and molecular techniques showed that dark-septate fungi (DSF) dominate (States et al. 2001, Zak 2005, Bates and Garcia-Pichel 2008). These phylogenetically diverse fungi have pigmented hyphae due to the presence of fungal melanin that provides resistance to UV radiation, drought and high temperatures (Zak 2005). However the taxonomic determinations and diversity estimations for DSF have been limited by the lack of sexual stages for many of these fungi (States et al. 2001, Grishkan et al. 2006) or by the use of genetic markers or methodologies with limited ability to determine fungal diversity at species rank (e.g. Bates and Garcia-Pichel 2008). The incorporation of fungal components in aridland ecosystems models has been limited by a paucity of information concerning their composition within the soil, BSC and plant roots and their response to environmental changes (Maestre et al. 2005). In this study we described and compared the fungal community composition in the rhizosphere and BSC of a semiarid grassland on the basis of the ITS rDNA region. The study was conducted in the Sevilleta National Wildlife Refuge (SNWR), a Long-Term Ecological Research (LTER) site in New Mexico, USA. Our goals were (i) to describe taxonomic composition of soil fungal communities in a semiarid ecosystem, (ii) to contrast the soil fungi with previously described root-associated fungal communities at the same site (Porras-Alfaro et al. 2008), and (iii) to determine whether the microtopographic distribution of fungal communities within rhizosphere, BSC and roots is consistent with the proposed ‘‘fungal loop’’ model in semiarid ecosystems (Collins et al. 2008). The fungal loop model proposed by Collins et al. (2008) incorporates the role of microbial communities in nitrogen and carbon cycling within a threshold-delay nutrient dynamics model. In arid systems plants respond to pulsed precipitation, soils have limited organic matter, N and C, and often have high temperatures. These conditions favor fungaldominated processes vs. bacteria and belowground fungal networks or loops that integrate BSC with tussocks (FIG. 1). We hypothesized that rhizosphere soils and BSC will have overlapping fungal communities even though they differ in their primary producer composition (plants vs. cyanobacteria respectively). MATERIALS AND METHODS
Site description.—Soil samples were collected from experimental plots maintained by the Sevilleta Long-Term Ecological Research (LTER) program and located within Sevilleta National Wildlife Refuge (SNWR) in central New
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Mexico. SNWR includes an extensive grassland dominated by C4 perennial grasses (Bouteloua gracilis, B. eriopoda, Sporobolus spp., Pleuraphis jamesii and Muhlenbergia spp.). In 1995 an N-addition experiment was established within the semiarid grassland biome (McKenzie Flats, 34u249N, 106u419, 1630 m). The experiment includes 20 5 3 10 m plots, all located in the same area to limit variability and maintain similar plant composition. Ten plots were untreated controls and 10 were fertilized twice per year with 50 kg N ha21 NH4NO3. The treatments were assigned randomly and plots were spaced 2 m apart (Johnson et al. 2003). Ambient N deposition was approximately 2 kg ha21y21 (Ba´ez et al. 2007). Vegetation cover averaged 60%, with the exposed soil surface between plants colonized by light cyanobacterial crusts dominated by Microcoleus spp. (Lipinski and Porras-Alfaro unpubl). Mean annual precipitation is 250 mm, although annual precipitation and its distribution throughout the year vary widely (Pennington and Collins 2007). The soils, formed by alluvial and aeolian deposition, are part of the Turney loamy sand series. Additional information about this site, including studies by Johnson et al. (2003), Khidir et al. (2010), Crenshaw et al. (2008), Porras-Alfaro et al. (2007, 2008) and Stursova et al. (2006) is available through the Sevilleta LTER Website (http://sev.lternet.edu). Sample collection.—Rhizosphere soil (soil surrounding plant roots) and BSC samples were collected from six randomly selected plots (three soil samples from control plots and three soil samples from N-addition of each soil type) in May 2005. Each sample was a mixture of three subsamples per plot collected along transects spaced 2 m apart; samples were pooled to create a composite sample (25 mL soil in 50 mL tubes). BSC samples correspond to the first 0.5 cm of soil where the majority of the microbial biomass (cyanobacteria and fungi) are found. Roots of B. gracilis also were collected; detailed molecular and microscopic analyses of the fungi associated with these grama roots were presented by Porras-Alfaro et al. (2007, 2008). The BSCs at SNWR are easily identified by texture and pigmentation. We collected only the upper 1 cm of crust, which forms an adhesive layer that can be lifted intact from the soil (approx. 8 mL soil per sample). Rhizosphere soil was gathered by dislodging a plant with a trowel and collecting soil associated with the root mass (5–10 mL deep, approximately 8 mL soil per sample). Soil and root samples were transported on ice and stored at 220 C for 5 d or fewer before DNA extraction. Molecular methods.—Soil was mixed in the tubes and small roots were separated with a stereomicroscope and fine forceps. DNA was extracted with the PowerSoilTM DNA Isolation Kit (MO BIO Laboratories Inc), following manufacturer’s instructions. Subsamples of soil and crust for DNA extraction were examined under a stereomicroscope to verify that they did not contain fine roots or fragments of plant tissue. DNA amplification, cloning and sequencing were conducted as described in Porras-Alfaro et al. (2008). DNA was amplified by with fungal-specific primers ITS1-F and ITS4 (Gardes and Bruns 1993) and Glomeromycotaspecific 18S primers NS31 and AM1 (Helgason et al. 1998,
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FIG. 1.
Microtopographic distribution of primary producers and fungal networks in semiarid ecosystems.
Simon et al. 1992). PCRs were performed with this protocol: initial denaturation at 95 C for 5 min, followed by 30 cycles of 95 C for 30 s, 53 C for 30 s and 72 C for 45 s with a final extension of 72 C for 7 min. DNA was amplified in 25 mL reactions with 12.5 mL Premix Taq (Takara Bio), 1.0 mL each primer (5 mM), 3 mL BSA 1%, 6.5 mL milliQ water and 1 mL template DNA. PCR products were cloned with the TOPOTA cloning kit (Invitrogen, Carlsbad, California). Cloned sequences were amplified with rolling circle amplification (TempliPhi, Amersham, Buckinghamshire, England) and sequenced with the BigDyeH Terminator 1.1 Sequencing Kit (Applied Biosystems, Foster City, California). Negative controls (reactions without DNA template) were included in all amplification and cloning reactions. Sequencing was conducted at the Molecular Biology Facility at the University of New Mexico. Sequences were assembled and edited with Sequencher 4.0 (Gene Codes Corp., Ann Arbor, Michigan). Chimeric sequences were determined following O’Brien et al. (2005): ITS1 and ITS2 regions of the same sequence were queried independently in GenBank. If the ITS1 and ITS2 sequences matched unrelated sequences in the NCBI database (i.e. taxa from different genera) they were considered chimeric, particularly if the sequences were recovered only once or in only one sample. Taxonomic classification followed Lumbsch and Huhndorf (2007), James et al. (2006a, b), Hansen and Pfister (2006), Matheny et al. (2006), Parker et al. (2006), Schoch et al. (2006), Scorzetti et al. (2002), Spatafora et al. (2006) and Zhang et al. (2006). Fungal sequences
were deposited in GenBank under accession numbers EU479716–EU480346 for ITS nrDNA sequences and EU555331–EU555335 for 18S rDNA Glomeromycotaspecific primers. EU144389–EU144746, deposited by Porras-Alfaro et al. (2008), of the root-associated fungal communities also were included in subsequent analyses for comparison (see DISCUSSION). Sequences were aligned with Clustal W, and DNA distance matrices were constructed using the F84 evolutionary model with DNADIST program from PHYLIP (Felsenstein 2005). DNA distance matrices include only the 5.8S rDNA and ITS partial sequences (193 nucleotides, sequence alignment number, ALIGN_001232 deposited in Webin-Align: ftp://ftp.ebi.ac.uk/pub/databases/embl/ alig). Only the 5.8s rDNA gene and small partial regions of ITS1 and ITS2 were included in the alignments because ITS regions are highly variable. Alignments of the 5.8S and ITS flanking regions were edited by eye in Jalview. Operational taxonomic units (OTUs) and rarefaction curves were determined based on 99% similarity with the aid of the DOTUR program to determine abundance of unique taxa (Schloss and Handelsman 2005). We used 99% similarity in DOTUR instead of 97% (i.e. identity value that was used to identify unique species in BLAST) because only partial sequences were used to create the DNA matrices. Consistent results were obtained for groups created with DOTUR at 99% similarity and BLAST results at 97% identity (data not shown). DOTUR uses distance-based methods with random sampling without replacement to define
PORRAS-ALFARO ET AL.: SOIL FUNGI IN GRASSLAND OTUs. DNA distance matrices were created with PHYLIP and used as input files. Differences based on community structure were calculated with the program UniFrac (Lozupone and Knight 2005). UniFrac uses a phylogenetic tree with sequences from different environments as input file. A text file describing the sample type and abundance of each unique OTU also was created. The phylogenetic tree included only unique OTUs obtained with DOTUR at 99% similarity (137 sequences were included in the tree). PAUP 4.0b10 was used to construct the most parsimonious tree with Toxoplasma gondii (X75429) and Drosophila melanogaster (M21017) as outgroups (James et al. 2006a, b). Principal coordinate analysis (PCA) was calculated in UniFrac using non-normalized weighted parameters (similar results were obtained with normalized weighted PCA) with fungal communities from roots included in this analysis for comparison. We also conducted a Jackknife cluster analysis to determine whether the composition and abundance of fungal communities were affected by nitrogen fertilization or soil type (rhizosphere vs. BSC). Default parameters were used, and the results were identical for normalized and nonnormalized abundance weighted cluster analysis. The program SONS (shared OTUs and similarity) was used to compare similarities between fungal communities by sample (Schloss and Handelsman 2006). SONS used non-parametric tests to determine similarities between microbial communities based on species abundance and distribution. The OTU designation file (99% similarity) from DOTUR was used as the input file for SONS (Schloss and Handelsman 2005). Fungal communities also were compared with the SLibshuff program. S-Libshuff uses the Cramer-von Mises test statistic to compare the microbial populations of two clone libraries. A distance matrix also was used as the input file. P values were calculated based on 10 000 permutations (Schloss et al. 2004). Phylogenetic analyses of different phyla (e.g. Chytridiomycota) and the most common orders (e.g. Pleosporales, Pezizales, Sordariales and Hypocreales) obtained from soil samples were conducted with PAUP (4.0b10, Swofford 2002). Alignments were created with Clustal W, edited in Jalview and sequences deposited in Webin-Align under numbers ALIGN_001239 (Ascomycota, Pleosporales, 714 nucleotides, 218 highly variable characters excluded from analysis), ALIGN_001237 (Ascomycota, Hypocreales and Sordariales, 409 nucleotides, 51 characters excluded), ALIGN_001235 (Ascomycota, Pezizales, 159 nucleotides), ALIGN_001236 (Basidiomycota, Filobasidiales, 712 nucleotides, 223 characters excluded), ALIGN_001238 (Chytridiomycota and Blastocladiomycota, 173 nucleotides). Trees were constructed with a full heuristic search for parsimony analysis with all characters equally weighted and unordered. One hundred trees were retained during the analysis. Bootstrap values were obtained based on 1000 replicates with 50% majority rule. Outgroups were selected based on James et al. (2006a, b), Hansen and Pfister (2006), Matheny et al. (2006), Parker et al. (2006), Schoch et al. (2006), Scorzetti et al. (2002), Spatafora et al. (2006), Zhang et al. (2006).
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FIG. 2. Average fungal diversity in rhizosphere and BSC in a semiarid grassland at SNWR. Rarefaction curves were calculated with DOTUR at 99% similarity. Error bars represent standard deviations (n 5 6). RESULTS
Twenty sequences from the dataset were eliminated after chimeric analysis. Twelve clone libraries, six from BSCs and six from rhizosphere soils, yielded a total of 307 and 324 sequences respectively with a minimum of 48 sequences per library. A total of 78 OTUs were found in BSC and 67 OTUs were found in rhizosphere soils. Rhizosphere soils and BSCs did not show significant differences in diversity and rarefaction curves did not show saturation (FIG. 2). Diversity was not significantly different between rhizosphere and BSC soil as determined by Chao diversity estimators of 107 (CI 83–161) and 149 (CI 108–241) respectively. The fungal community of semiarid grassland at SNWR has a large number of undescribed taxa, ascertained from the more than 130 000 fungal ITS rDNA sequences available in GenBank in Jun 2009. A total of 46.2% of 307 sequences from BSC and 36.5% of 324 sequences of rhizosphere soils had , 97% sequence identity with NCBI-deposited sequences. Based on queries of the NCBI core nucleotide database, at least 23 fungal orders inhabit BSC and rhizosphere soils. Five percent of the sequences in our database could not be classified with certainty even at order rank due to low similarity to sequences in the NCBI database; most of these unidentified sequences were within Chytridiomycota. The Ascomycota represented 86% of all sequences (TABLE I) while Pleosporales was the most common order in BSC and rhizosphere soils (, 50% of sequences, FIG. 3). Orders Hypocreales, Pezizales and Dothideales (Aureobasidium) also were common
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T ABLE I. Composition of biological soil crust and rhizosphere soil fungal communities sampled from a semiarid grassland in central New Mexico (numbers correspond to number of sequences) Sample type Crust
Rhizosphere
Total
Phylum Ascomycota 268 (82.7%) 274 (89.3%) 542 (85.9%) Basidiomycota 28 (8.6%) 25 (8.1%) 53 (8.4%) Chytridiomycota 20 (6.2%) 8 (2.6%) 28 (4.4%) Glomeromycota 0 0 0 Zygomycota 7 (2.2%) 0 7 (1.1%) Unknown 1 (0.1%) 0 1 (0.2%) Total 324 307 631 Common orders Pleosporales Sordariales Agaricales Pezizales Hypocreales Dothideales Filobasidiales
178 3 0 51 6 9 14
(54.9%) 170 (0.9%) 6 3 (15.7%) 27 (1.9%) 37 (2.8%) 22 (4.3%) 18
(55.4%) 348 (2%) 9 (1%) 4 (8.8%) 78 (12.1%) 43 (7.2%) 31 (5.9%) 32
(63.9%) (1.7%) (0.7%) (14.3%) (7.9%) (5.7%) (5.9)
in the soil samples (FIG. 4a–c, TABLE I). Most of the Pleosporales OTUs belong to Pleosporaceae and Phaeosphaeraceae (FIG. 3). The most common fungi are related to species in Alternaria, Cochliobolus/ Bipolaris, Phaeosphaeria, Leptosphaeria, Preussia and Phoma complexes. In addition phylogenetic analyses disclosed several novel clades and putative new families. Fungi closely related to Fusarium (Hypocreales, FIG. 4a) and Ascobolus (Pezizales, FIG. 4b) also were common (with 39 and 59 sequences respectively). Basidiomycota represented only 8% (53 sequences) of soil samples. Cryptococcus spp. (Filobasidiales) were the most common Basidiomycota found in the crust and rhizosphere with a total of 32 sequences (FIG. 4c). Other Basidiomycota genera found in the soil included close relatives of Rhizoctonia sp., Pterula sp., Tulostoma sp. and Panaeolus sp. Zoosporic fungi were found mainly in the BSC (70% of 29 Chytridiomycota-Blastocladiomycota sequences). At least five of the 11 major chytrid clades described by James et al. (2006a) were found colonizing the rhizosphere and BSC (FIG. 4d). Zygomycota were rare; only seven sequences were obtained from BSC (TABLE I). More than 70% of the BSC and rhizosphere soil fungal sequences are closely related to other soil fungal sequences deposited in GenBank. We did not recover any AMF sequences from soil samples with the fungal-specific nrITS primers. With AMF specific primers we recovered only five Glomeromycota
sequences related to Glomus from 74 amplicons (data not shown), reflecting the difficulty of specific amplification from a taxonomically rich and compositionally more complex soil community. The fungal communities of the rhizosphere soil and BSC exhibited similar composition. Analysis of Chao diversity estimators showed that rhizosphere and BSC fungal communities share more than 50% of OTUs (FIG. 5) with a Jaccard abundance index of 0.75 (SE 5 0.09). The effects of long-term N fertilization on species richness and fungal community composition were not significant. A total of 22 (SD 5 4) and 20 (SD 5 4) OTUs were found respectively in control and nitrogen plots in BSC. In rhizosphere soils a total of 21 (SD 5 3) and 17 (SD 5 3) OTUs were found per sample. Chao estimators and rarefaction curves showed the same tendency. Comparisons of community similarity (jackknife cluster analysis, FIG. 5) did not clearly segregate fungal communities based on N treatment or microhabitat. Although differences between nitrogen and control treatments were not observed at the community level, our data suggested that nitrogen enrichment might have affected the abundance of specific taxa. For example a total of 75 sequences (12.5 sequences per sample, SD 5 3.6) were found in nitrogen plots versus 25 sequences in control plots (average 3.8 sequences per samples, SD 5 1.9) for the most common OTU in the soil, a fungus within the Alternaria alternata complex. DISCUSSION
Species richness and novel fungal clades.—This study was the first comprehensive sequencing survey of soil fungal communities at the Sevilleta National Wildlife Refuge. Sequence-based techniques are useful in the identification of novel groups and estimation of fungal diversity, especially in environments with large numbers of sterile mycelial fungi, as is the case for arid ecosystems (Herrera et al. 1997, Zak 2005). Fungal richness in SNWR semiarid grassland soil is similar to that reported for Alaskan soils (Allison et al. 2007) but less than that reported for Duke Forest (a Pinus taeda plantation) in North Carolina, USA (O’Brien et al. 2005). Unlike boreal and temperate forests, which are dominated by Basidiomycota (O’Brien et al. 2005, Allison et al. 2007), the soils and plant roots of semiarid grasslands are colonized predominantly by Ascomycota, including a large number of DSF. The most common genera of DSF at SNWR (e.g. Alternaria, Phoma, Bipolaris, Aureobasidium) also have been reported from soils in other desert ecosystems in Utah and Wyoming, USA (States
PORRAS-ALFARO ET AL.: SOIL FUNGI IN GRASSLAND
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FIG. 3. Phylogenetic relationships within Pleosporales, the most common order found in a semiarid grassland rhizosphere and BSC soil at Sevilleta National Wildlife Refuge. Bootstrap values (1000 replicates) greater than 70% are indicated above branches. Sequences obtained from this study are shown in boldface.
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FIG. 4. Common fungal orders among different phyla in a semiarid grassland. Bootstrap values (1000 replicates) greater than 70% are indicated above branches. Sequences obtained from this study are shown in boldface. a. AscomycotaSordariomycetes, Pleosporales and Hypocreales. b. Ascomycota, Pezizomycetes, Pezizales. c. Basidiomycota, Filobasidiales. d. several orders of Chytridiomycota and Blastocladiomycota.
PORRAS-ALFARO ET AL.: SOIL FUNGI IN GRASSLAND
FIG. 5. Cluster environmental analysis. Jackknife values were calculated with UniFrac. Samples collected from nitrogen (N) and control (C) plots followed by plot number.
and Christensen 2001), Nevada and Death Valley, USA (Durrell and Shields 1960), and the Negev Desert, Israel (Grishkan et al. 2006). Although Johnson et al. (2003) reported presence of AMF fungal spores at the same site, we did not recover a large number of sequences for this group, despite using AMF fungal-specific primers. Differences in abundance of the different fungal groups, temporal variation, bias in DNA extractions and unspecific amplifications are plausible explanations for the low recovery of AMF sequences in these complex soil microbial communities. Phylogenetic analyses of the most common orders in the soil showed that soils at SNWR are colonized by a large group of unrepresented or undescribed fungal clades (FIGS. 3, 4). Although several clades did not have strong bootstrap support, all phylogenetic trees presented here agreed with published phylogenies (e.g. James et al. 2006a, b; Hansen and Pfister 2006; Matheny et al. 2006; Parker et al. 2006; Schoch et al. 2006; Scorzetti et al. 2002; Spatafora et al. 2006; Zhang et al. 2006). Because of the high variability of the ITS rDNA region more extensive analyses with a more conserved region such as 28S rRNA are required to corroborate the phylogenetic placement of some of these novel clades. Pleosporales is the most abundant and diverse order represented in SNWR semiarid grassland soils, and members of this group accounted for the largest number of novel sequences (FIG. 3, TABLE I). This study and those of rootassociated fungi revealed several unique clades within this order, suggesting that arid lands could be
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hotspots of Pleosporalean diversity (Porras-Alfaro et al. 2007, 2008; Khidir et al. 2010). Although it is the largest order within Dothidiomycetes, Pleosporales has been poorly surveyed because most investigations have focused primarily on plant pathogens of economically important crops (Schoch et al. 2006). The great abundance of Pleosporales and other DSF in arid grasslands may be explained by the presence of melanin pigments in their hyphae. Fungal melanins have been associated with mitigating environmental stresses (including high temperatures, high UV radiation and extended drought) (Bell and Wheeler 1986). States and Christensen (2001) showed with the culture-based techniques that fungal communities of BSCs from Utah and Wyoming, USA, and the Negev Desert in Israel are both diverse and dominated by DSF. Our study also found that BSCs are dominated by DSF and identified similar genera to those reported by States and Christensen (2001). The availability of a molecular database will provide a fundamental framework that will aid future cross-site comparisons because of the large number of anamorphic fungi present in these ecosystems. Effect of N enrichment on fungal communities.—The effects of N availability on the fungal communities of aridlands has received little study relative to mesic ecosystems. The few studies of responses to N manipulation variously reported increases, decreases or no net changes in the diversity and structure of fungal communities (Johnson 1993, Jumpponen and Johnson 2005, Frey et al. 2004). Most studies have focused on the effects of N availability on arbuscular mycorrhizal fungal development or on colonization rates and generally showed declines with increased N availability (e.g. Porras-Alfaro et al. 2007, Johnson et al. 2003). Other studies suggested that losses of oxidative enzyme activity and lignin degradation capacity in N saturated soils might be a result of losses of saprophytic fungi, particularly filamentous basidiomycetes (Frey et al. 2004, Sinsabaugh et al. 2002). Filamentous basidiomycetes are rare at SNWR. More than a decade of experimental N amendment has not significantly altered the richness and composition of fungal communities associated with rhizosphere and BSC (FIGS. 5, 6), even though plant productivity is N limited when soil moisture is adequate and significant changes in soil enzyme activity and N transformation have been reported (Stursova et al. 2006, Crenshaw et al. 2008). Microbial communities are spatially and temporally heterogeneous on both small and large scales (e.g. Bayman et al. 1998), and because a large portion of the diversity is associated with primary producers it is difficult to
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MYCOLOGIA al. 2003, Schlesinger et al. 1996). Rhizosphere soils and BSC are areas where carbon and nutrients accumulate through the activity of phototrophic organisms. The associations of fungi with phototrophs (e.g. cyanobacteria or plants) are likely to enhance their survival through extreme conditions. Development of specific symbioses between B. gracilis and endophytic and mycorrhizal fungi might be fundamental in protecting roots against pathogenic fungi through a variety of mechanisms that may include improvement of plant growth, competition for nutrients and the production of extracellular enzymes that limit pathogen growth (FIG. 6, PorrasAlfaro et al. 2007, 2008).
FIG. 6. Principal coordinate analysis (PCA) of root and soil fungal communities. Roots, rhizosphere soil and BSC are represented respectively by triangles, circles and squares. Samples collected from N-amended plots are represented in white, and samples from control plots are in black. Fungal communities were significantly different in soil vs. root samples. S-Libshuff (root vs. soil samples): P , 0.0001. UniFrac significance: P 5 0.01.
establish on an ecosystem scale the extent to which fungal communities are affected by N enrichment or other environmental disturbances (e.g. Jumpponen et al. 2005, Jumpponen and Johnson 2005). Fungal distribution patterns.—Limited information is available about the distribution of fungi in relation to the characteristic microtopography of semiarid grasslands (FIG. 1). Jumpponen and Johnson (2005) demonstrated with molecular data that the structure of fungal communities of soils and roots in a tallgrass prairie are distinct, and our study also documented the same pattern for semiarid grasslands (FIG. 6). Taxonomic richness in rhizosphere and crust soil samples was nearly three times greater than the richness described for fungal communities associated with roots of dominant grasses at the same site. Sordariales and Agaricales, the second and third most common orders associated with plant roots (PorrasAlfaro et al. 2008), were rarely found in soil (TABLE I). For example only five Agaricales sequences were found colonizing rhizosphere soils and none were found associated with BSC. Fungal assemblages of rhizosphere soils and BSC substantially overlap in composition (FIGS. 5, 6), and the similarities in fungal richness and community composition of BSC and rhizosphere soils may reflect similarities in the biogeochemistry of plant-centered ‘‘islands’’ and crust-covered ‘‘mantles of fertility’’ characteristic of semiarid ecosystems (Garcia-Pichel et
Fungal networks in semiarid grasslands.—The threshold-delay nutrient dynamics model for arid ecosystems suggested that fungal communities in semiarid grasslands might have a fundamental role in N and C transformation, decomposition and nutrient translocation processes through the establishment of a hyphal network that links BSCs and plants (Collins et al. 2008). Studies at SNWR using isotopically labeled substrates have shown that grasses and biotic crusts exchange N (and C) through a fungal network. 15 N derived from both nitrate and glutamate was translocated from point applications on crusts and grasses at up to 100 cm/d (Green et al. 2008). The high similarity of BSC and rhizosphere soil fungal communities detected in this study support this fungal network hypothesis (FIGS. 1, 5, 6). Hyphal networks between roots and BSC with similar community composition are more likely to aid nutrient absorption by plants because they can extend beyond the area of nutrient depletion surrounding roots, grow into protected soil pores and produce extracellular enzymes that release nutrients from soil organic matter (Morgan et al. 2005). DSF-dominated communities of semiarid grassland may extend these advantages at lower water activities because these fungi are able to metabolize at low water potentials and to confer drought and UV protection to plants and cyanobacteria (States et al. 2001, Garcia-Pichel et al. 2003). In most ecosystems AMF provide fundamental support to plant communities through the translocation of nutrients and water (Smith and Read 1997). Because most AMF hyphae are not highly tolerant to drought their abundance and contributions in support of primary production diminish with low water availability. Few AMF were found in this and previous studies at SNWR grassland 2004–2007 (Porras-Alfaro et al. 2007, 2008; Khidir et al. 2010). There is evidence that AMF were more abundant during wetter periods in the 1980s and 1990s (Allen
PORRAS-ALFARO ET AL.: SOIL FUNGI IN GRASSLAND et al. 1981, Johnson et al. 2003). Allen (2006) showed that in desert ecosystems with low moisture conditions and infrequent rain AMF are important in preventing water loss mainly during transition to drought. The high colonization rates by DSF during a dry period 2004–2007 (Porras-Alfaro et al. 2007, 2008) suggested that DSF might contribute to plant survival. DSF may be a key component of arid environments specifically because they are adapted to high temperatures and high salt concentrations (States et al. 2001). ACKNOWLEDGMENTS
This paper is dedicated to the memory of Kendra Lipinski (1983–2009). This research was financially supported by the National Science Foundation through the Long Term Ecological Research Program at Sevilleta National Wildlife Refuge (DEB-027774 and DEB-0217774) and a grant from the National Science Foundation Ecosystems Sciences Program (DEB 0516113).
LITERATURE CITED
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