Microbial Ecology Soil Fungal Communities Underneath Willow Canopies on a Primary Successional Glacier Forefront: rDNA Sequence Results Can Be Affected by Primer Selection and Chimeric Data Ari Jumpponen Division of Biology, Kansas State University, 125 Ackert Hall, Manhattan, KS 66506, USA Received: 7 January 2004 / Accepted: 9 March 2004 / Online publication: 3 November 2006 Abstract Soil fungal communities underneath willow canopies that had established on the forefront of a receding glacier were analyzed by cloning the polymerase chain reaction (PCR)-amplified partial small subunit (18S) of the ribosomal (rRNA) genes. Congruence between two sets of fungus-specific primers targeting the same gene region was analyzed by comparisons of inferred neighbor-joining topologies. The importance of chimeric sequences was evaluated by Chimera Check (Ribosomal Database Project) and by data reanalyses after omission of potentially chimeric regions at the 50- and 30-ends of the cloned amplicons. Diverse communities of fungi representing Ascomycota, Basidiomycota, Chytridiomycota, and Zygomycota were detected. Ectomycorrhizal fungi comprised a major component in the early plant communities in primary successional ecosystems, as both primer sets frequently detected basidiomycetes (Russulaceae and Thelephoraceae) forming mycorrhizal symbioses. Various ascomycetes (Ophiostomatales, Pezizales, and Sordariales) of uncertain function dominated the clone libraries amplified from the willow canopy soil with one set of primers, whereas the clone libraries of the amplicons generated with the second primer set were dominated by basidiomycetes. Accordingly, primer bias is an important factor in fungal community analyses using DNA extracted from environmental samples. A large proportion (930%) of the cloned sequences were concluded to be chimeric based on their changing positions in inferred phylogenies after omission of possibly chimeric data. Many chimeric sequences were positioned basal to existing classes of fungi, suggesting that PCR artifacts may cause frequent discovery of new, higher level taxa (order, class) in direct PCR analyses. Longer extension times during the PCR Correspondence to: Ari Jumpponen; E-mail: ari@ksu.edu DOI: 10.1007/s00248-004-0006-x & Volume 53, 233–246 (2007) & * amplification and a smaller number of PCR cycles are necessary precautions to allow collection of reliable environmental sequence data. Introduction Fungi perform important ecosystem functions by participating in the decomposition of dead tissues as well as plant uptake of water and nutrients [6, 34]. Assessment of fungal community composition is difficult because of unreliable and ephemeral production of identifiable macroscopic fruiting bodies [11, 27, 35]. Many fungi also produce microscopic, sexual or asexual fruiting structures or fruit below ground escaping detection in assessments relying exclusively on the collection of epigeous fruiting bodies. Pure culture techniques allow fungal community assays of soil and tissue samples in the absence of identifiable macroscopic fruiting bodies. However, similar to bacteria [38], it is likely that large numbers of fungi would be missed in such pure culture assays (see [31, 41]). To overcome these problems in fungal community analysis, molecular means specifically targeting fungi in environmental samples have been developed [3, 9, 14, 25, 28, 32, 33, 40]. Direct molecular assessment of the fungal communities allows analyses without relying on whether or not the fungi can be grown in pure culture or produce fruiting bodies. However, polymerase chain reaction (PCR) artifacts, such as chimeric sequences resulting from amplification of more than one template, can cause problems in environmental samples with unknown sources of diverse initial template DNA [13, 19, 24, 42, 43]. Various coextracted substances and low concentrations of the target template in the presence of highly similar competing target and nontarget templates may further influence the fidelity of PCR reactions [42]. Springer Science + Business Media, Inc. 2006 233 Chimera at RDP Yes (G20) Yes (G40) Yes (G80) Yes (G40) Yes (G20) Yes (G20) Yes (G20) Yes (G20) Yes (G20) Yes (G40) No Yes (G100) Yes (G40) No Yes (G80) Yes (G40) Yes (G40) Yes (G40) Yes (G40) Yes (G40) Yes (G40) Yes (G40) Yes (G40) Yes (G40) No Yes (G20) Yes (G40) Yes (G40) Yes (G80) Yes (G40) Yes (G40) Yes (G40) Yes (G80) Yes (G20) Yes (G20) No Environmental clone B_Canopy_300_01_08 [AY382401] B_Canopy_300_01_14 [AY382402] B_Canopy_300_01_16 [AY382403] B_Canopy_300_01_18b [AY382404] B_Canopy_300_02_04b [AY382405] B_Canopy_300_02_05 [AY382406] B_Canopy_300_02_06 [AY382407] B_Canopy_300_02_10 [AY382408] B_Canopy_300_02_12 [AY382419] B_Canopy_300_02_14b [AY382410] B_Canopy_300_03_06 [AY382411] B_Canopy_300_03_12b [AY382412] B_Canopy_300_03_17 [AY382413] B_Canopy_300_03_19b [AY382414] B_Canopy_450_01_02 [AY382415] B_Canopy_450_01_06 [AY382416] B_Canopy_450_01_13 [AY382417] B_Canopy_450_01_14 [AY382418] B_Canopy_450_01_18 [AY382419] B_Canopy_450_02_02 [AY382420] B_Canopy_450_02_13 [AY382421] B_Canopy_450_03_02 [AY382422] B_Canopy_450_03_05 [AY382423] B_Canopy_450_03_07 [AY382424] B_Canopy_450_03_14 [AY382425] B_Canopy_750_01_01b [AY382426] B_Canopy_750_01_07b [AY382427] B_Canopy_750_01_10b [AY382428] B_Canopy_750_01_15b [AY382429] B_Canopy_750_02_13 [AY382430] B_Canopy_750_02_15b [AY382431] B_Canopy_750_02_19b [AY382432] B_Canopy_750_03_03b [AY382433] B_Canopy_750_03_04 [AY382434] B_Canopy_750_03_08b [AY382435] B_Canopy_750_03_11 [AY382436] Spilocaea oleaginea [AF338393] (Chaethothyriales/Dothidiales) Spilocaea oleaginea [AF338393] (Chaethothyriales/Dothidiales) Hymenoscyphus ericea [AY228753] (Helotiales) Inocybe geophylla [AF287835] (Agaricales) Dark septate endophyte DS16b [AF168167] (Unknown) Peziza griseorosea [AF133150] (Pezizales) Peziza griseorosea [AF133150] (Pezizales) Tetracladium marchalianum [AY204613] (Incertae sedis) Spilocaea oleaginea [AF338393] (Chaethothyriales/Dothidiales) Oidiodendron tenuissimum [AB015787] (Onygenales) Prismatolaimus intermedius [AF036603] (Enoplida; Prismatolaimidae) Cladonia sulphurina [AF241544] (Lecanorales) Hypoxylon submonticulosum [AF346544] (Xylariales) Neobulgaria premnophila [U45445] (Helotiales) Pulvinula archeri [U62012] (Pezizales) Hypomyces chrysospermus [AB027339] (Hypocreales) Oidiodendron tenuissimum [AB015787] (Onygenales) Oidiodendron tenuissimum [AB015787] (Onygenales) Oidiodendron tenuissimum [AB015787] (Onygenales) Rhizoctonia solani [D85643] (Ceratobasidiales) Hypomyces chrysospermus [AB027339] (Hypocreales) Connersia rilstonii [AF096174] (Eurotiales) Raciborskiomyces longisetosum [AY016351] (Chaetothyriales) Herpotrichia juniperi [U42483] (Pleosporales) Mycosphaerella mycopappi [U43463] (Chaetothyriales) Inocybe geophylla [AF287835] (Cortinariaceae) Peziza griseorosea [AF133150] (Pezizales) Anamylopsora pulcherrima [AF119501] (Agyriales) Pulvinula archeri [U62012] (Pezizales) Hypomyces chrysospermus [M89993] (Hypocreales) Ophiostoma piliferum [AJ243294] (Ophiostomatales) Hypomyces chrysospermus [M89993] (Hypocreales) Sarcinomyces petricola [Y18702] (Chaetothyriales) Laccaria pumila [AF287838] (Agaricales) Laccaria pumila [AF287838] (Agaricales) Peziza griseorosea [AF133150] (Pezizales) BLAST match [accession number] (Order) Ascomycota Ascomycota Ascomycota Basidiomycota Ascomycota Ascomycota Ascomycota Ascomycota Ascomycota Ascomycota Contaminant Ascomycota Ascomycota Ascomycota Ascomycota Ascomycota Ascomycota Ascomycota Ascomycota Basidiomycota Ascomycota Ascomycota Ascomycota Ascomycota Ascomycota Basidiomycota Ascomycota Ascomycota Ascomycota Ascomycota Ascomycota Ascomycota Ascomycota Basidiomycota Basidiomycota Ascomycota Phylum 98 96 95a 97 98 99 98 99a 98 97 97 93 96 98 94 96 98 97 98 95 96 99/99a,c 99 97 98 97 99 97 97 96 97/97c 95 95 98 96 99 Similarity 0.60 0.20 0.10 0.10 0.22 0.11 0.11 0.11 0.33 0.11 0.08 0.15 0.31 0.46 0.27 0.20 0.07 0.33 0.13 0.06 0.94 0.14 0.14 0.43 0.29 0.20 0.40 0.20 0.20 0.33 0.44 0.22 0.14 0.29 0.14 0.43 Frequency Table 1. BLAST and RDP analyses of the environmental sequences obtained from underneath the willow canopies established on the forefront of a receding glacier 234 A. JUMPPONEN: FUNGI IN THE WILLOW CANOPY SOIL ON A GLACIER FOREFRONT Yes Yes No No No Yes No Yes Yes Yes Yes Yes No Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes (G100) (G40) (G80) (G160) (G100) (G40) (G20) (G100) (G80) (G60) (G60) (G20) (G20) (G20) (G40) (G160) (G100) (G20) (G40) (G40) (G20) (G20) (G40) (G40) (G80) (G40) (G80) (G40) (G60) (G80) (G40) (G40) Russula compacta [AF026582] (Agaricales) Hypomyces chrysospermus [AB027339] (Hypocreales) Russula compacta [AF026582] (Agaricales) Oidiodendron tenuissimum [AB015787] (Onygenales) Chaetomium elatum [M83257] (Sordariales) Thelephora sp. [AF026627] (Thelephorales) Dark septate endophyte DS16b [AF168167] (Unknown Pulvinula archeri [U62012] (Pezizales) Hypomyces chrysospermus [AB027339] (Hypocreales) Polyporoletus sublividus [AF287840] (Cantharellales) Thelephora sp. [AF026627] (Thelephorales) Bulgaria inquinans [AJ224362] (Helotiales) Inocybe geophylla [AF287835] (Agaricales) Bulgaria inquinans [AJ224362] (Helotiales) Mortierella chlamydospora [AF157143] (Mucorales) Bulgaria inquinans [AJ224362] (Helotiales) Limnoperdon incarnatum [AF426952] (Aphyllophorales) Panellus serotinus [AF026590] (Agaricales) Spizellomyces acuminatus [M59759] (Spizellomycetales) Laccaria pumila [AF287838] (Agaricales) Byssoascus striatosporus [AB015776] (Onygenales) Entoloma strictius [AF287832] (Agaricales) Bulgaria inquinans [AJ224362] (Helotiales) Ophiostoma stenoceras [M85054] (Ophiostomatales) Thelephora sp. [AF026627] (Thelephorales) Inocybe geophylla [AF287835] (Agaricales) Inocybe geophylla [AF287835] (Agaricales) Sporothrix schenkii [M85053] (Ophiostomatales) Dendrocorticium roseocarneum [AF334910] (Aphyllophorales) Bulgaria inquinans [AJ224362] (Helotiales) Termitomyces sp. [AB051891] (Agaricales) Inocybe geophylla [AF287835] (Agaricales) Laccaria pumila [AF287838] (Agaricales) Cyathrus striatus [AF026617] (Nidulariales) Inocybe geophylla [AF287835] (Agaricales) Russula compacta [U59093] (Agaricales) Thelephora sp. [AF026627] (Thelephorales) Basidiomycota Ascomycota Basidiomycota Ascomycota Ascomycota Basidiomycota Ascomycota Ascomycota Ascomycota Basidiomycota Basidiomycota Ascomycota Basidiomycota Ascomycota Zygomycota Ascomycota Basidiomycota Basidiomycota Chytridiomycota Basidiomycota Ascomycota Basidiomycota Ascomycota Ascomycota Basidiomycota Basidiomycota Basidiomycota Ascomycota Basidiomycota Ascomycota Basidiomycota Basidiomycota Basidiomycota Basidiomycota Basidiomycota Basidiomycota Basidiomycota 96 97 98 99 98 99 98 97 94 94 98 98 98 95 97 96 94 94 97 97 94 93 98 95 97/94c 96 98 93 92 95 94 97 95 97 98 97 97 0.25 0.13 0.63 0.09 0.18 0.55 0.09 0.09 0.75 0.13 0.13 0.11 0.89 0.14 0.29 0.14 0.14 0.29 0.67 0.33 0.25 0.25 0.50 1.00 0.40 0.40 0.20 0.92 0.08 0.14 0.43 0.14 0.14 0.14 0.77 0.23 1.00 IN THE WILLOW CANOPY SOIL ON A Chimera Check scores in parentheses. Frequency refers to the occurrence of a clone in the library obtained from one sample. RDP = Ribosomal Database Project; BLAST = basic local alignment search tool. a Sequence omitted from the neighbor-joining analyses because of a large insert. b Sequence determined chimeric in analyses after omission of data beyond chimera points. c BLAST matches were partial and did not span over the entire cloned sequence. B_Canopy_900_01_03b [AY382437] B_Canopy_900_01_16 [AY382438] B_Canopy_900_01_19 [AY382439] B_Canopy_900_02_02 [AY382440] B_Canopy_900_02_04 [AY382441] B_Canopy_900_02_06 [AY382442] B_Canopy_900_02_10 [AY382443] B_Canopy_900_02_12b [AY382444] B_Canopy_900_03_09 [AY382445] B_Canopy_900_03_11b [AY382446] B_Canopy_900_03_17 [AY382447] S_Canopy_300_01_01 [AY382448] S_Canopy_300_01_07 [AY382449] S_Canopy_300_02_01b [AY382450] S_Canopy_300_02_11 [AY382451] S_Canopy_300_02_13b [AY382452] S_Canopy_300_02_14b [AY382453] S_Canopy_300_02_19b [AY382454] S_Canopy_300_03_04 [AY382455] S_Canopy_300_03_18 [AY382456] S_Canopy_450_01_02b [AY382457] S_Canopy_450_01_07 [AY382458] S_Canopy_450_01_19 [AY382459] S_Canopy_450_02_05 [AY382460] S_Canopy_750_01_10 [AY382461] S_Canopy_750_01_11 [AY382462] S_Canopy_750_01_18 [AY382463] S_Canopy_750_02_09 [AY382464] S_Canopy_750_02_17b [AY382465] S_Canopy_750_03_01b [AY382466] S_Canopy_750_03_13 [AY382467] S_Canopy_750_03_15 [AY382468] S_Canopy_750_03_18b [AY382469] S_Canopy_750_03_19 [AY382470] S_Canopy_900_01_06 [AY382471] S_Canopy_900_01_11 [AY382472] S_Canopy_900_03_02 [AY382473] A. JUMPPONEN: FUNGI GLACIER FOREFRONT 235 236 A. JUMPPONEN: FUNGI Furthermore, primer sets designed to obtain broad specificity to a target group (e.g., fungi) may have biases and preferentially amplify one target group but not another [2, 36]. The overall goal of the presented studies was to characterize fungal community composition within established willow (Salix spp.) canopies on the forefront of a receding glacier. The nuclear small subunit (18S) of the ribosomal RNA gene (rDNA) was amplified with two different sets of fungus-specific primers to estimate the influence of primer selection on the observed community structure. To evaluate the influence of chimeric amplicons on the obtained 18S phylogenies, data sets were reanalyzed after omission of the chimeric regions identified using Chimera Check software of the Ribosomal Database Project (RDP, version 2.7 [26]). The results indicate that diverse fungal communities exist within the willow canopies, that primer selection strongly influences the observed fungal community structure, and that chimeras are a serious concern in direct PCR applications targeting fungi in environmental samples. Methods Lyman Glacier (48-1005200N, 120-5308700W) is located in the Glacier Peak Wilderness Area in the North Cascade Mountains (Washington, USA). The site has been utilized in several studies on early plant community assembly in recently deglaciated substrate (e.g., [20, 23]). Similarly, it has been a focus of studies aiming to examine fungal community assembly in such an environment [19, 21, 22]. The elevation of the present glacier terminus is about 1800 m. The deglaciated forefront is approximately 1000 m long over an elevation drop of only 60 m with no distinctive recessional moraines [4, 20]. The glacier has receded since the 1890s, opening the forefront to colonization by plants and fungi. Periodic photographs and snow survey data have allowed the reconstruction of the glacier retreat over the last century [20]. Study Site. Sampling and DNA Extraction. Shrub willows (Salix commutata and S. planifolia) comprise the early perennial plant communities and are the largest plant individuals during early vegetation development [22]. Twelve shrub canopies–three of approximately equal size at distances of 300, 450, 750, and 900 m from the glacier terminus–were selected, and 200-mL soil samples were IN THE WILLOW CANOPY SOIL ON A GLACIER FOREFRONT collected in August 2001. Samples were stored on ice until processed. In the laboratory, roots were handpicked from soil, and soil was homogenized manually in plastic bags. Approximately 0.25 g of soil was transferred to the extraction buffer, and DNA was extracted using UltraClean Soil kit (Molecular Biology Laboratories Inc., Carlsbad, CA) following manufacturer’s protocol. Extracted DNA was stored frozen (_20-C) until further processing. PCR Amplification of the Fungal DNA. A partial sequence of the 18S of the fungal rDNA was amplified with two different primer sets in 50-2L PCR reaction mixtures. First, the reaction to collect data set B contained final concentrations or absolute amounts of reagents as follows: 400 nM of each of the forward and reverse primers (nu-SSU-0817-50 and nu-SSU-1536-30 [3]), 2 2L of the extracted template DNA, 200 2M of each deoxynucleotide triphosphate, 2.5 mM MgCl2, 1 U of Taq DNA polymerase (Promega, Madison, WI), and 5 2L of manufacturer’s PCR buffer. The PCR cycle parameters consisted of an initial denaturation at 94-C for 3 min, then 40 cycles of denaturation at 94-C for 1 min, annealing at 56-C for 1 min and extension at 72-C for 1 min, followed by a final extension step at 72-C for 10 min. Second, the reaction to collect data set S contained final concentrations or absolute amounts of reagents as follows: 300 nM of each of the forward and reverse primers (EF4 and EF3 [32]), 2 2L of the extracted template DNA, 200 2M of each deoxynucleotide triphosphate, 1.7 mM MgCl2, 2 U of Taq DNA polymerase (Promega), and 5 2L of manufacturer’s PCR buffer. The PCR cycle parameters consisted of an initial denaturation at 94-C for 3 min, then 40 cycles of denaturation at 94-C for 1 min, annealing at 48-C for 1 min and extension at 72-C for 1 min, followed by a final extension step at 72-C for 10 min. All PCR reactions were performed in a Hybaid OmniCycler (Hybaid Ltd., Middlesex, UK). Possible PCR amplification of airborne and reagent contaminants was determined using a blank sample ran through the extraction protocol simultaneously with the actual samples and a negative PCR control in which the template DNA was replaced with ddH2O. These remained free of PCR amplicons in all trials. Small-Subunit rDNA Clone Library Construction and Primers specific to fungi and stringent PCR Analysis. conditions resulted in amplicons of expected size (about Figure 1. Neighbor-joining analysis of environmental partial 18S sequences (see Table 1 for accession numbers; AY382401–AY382473) 0 0 obtained with primer set B (nu-SSU-0817-5 and nu-SSU-1536-3 [3]) from willow canopy soil on the forefront of a receding glacier. Accession numbers of the GenBank-obtained sequences are shown in parentheses. Sequence data were aligned in Sequencher and neighbor-joining analyses performed in PAUP* [37]. Numbers above the nodes refer to the occurrence of that node in 1000 bootstrap replicates. Values 9 50% are shown. A. JUMPPONEN: FUNGI IN THE WILLOW CANOPY SOIL ON A GLACIER FOREFRONT 237 238 780 bp in set B and about 1400 bp in set S) when the PCR products were visualized on 1.5% agarose gels. The mixed populations of PCR products were ligated into a linearized pGEM-T vector (Promega). The circularized plasmids were transformed into competent JM109 cells (Promega) by heat shock, and the putative positive transformants were identified by !-complementation [30]. Twenty putatively positive transformants from each clone library were randomly sampled, and the presence of the target insert was confirmed by PCR amplification in 15-2L reaction volumes under the same reaction conditions as described above. To select different plasmids for sequencing, these PCR products were digested with endonucleases (HinfI, AluI; New England BioLabs, Beverly, MA) and were resolved on 3% agarose gels [15]. The PCR screening of clone libraries combined with restriction fragment length polymorphisms (RFLP) enabled the selection of different RFLP phenotypes for sequencing. Sequences from each different RFLP phenotype in all clone libraries were obtained by use of fluorescent dideoxy-terminators (ABI Prism\ BigDyei Applied Biosystems, Foster City, CA) and an automated ABI Prism\ 3700 DNA Analyzer (Applied Biosystems) at the DNA Sequencing and Genotyping Facility at Kansas State University (GenBank accession numbers AY382401–AY382473). Vector contamination was removed with the automated vector trimming function in Sequencher (Version 4.1, GeneCodes, Ann Arbor, MI). The similarities to existing rDNA sequences in the GenBank database were determined at the National Center for Biotechnology Information (http://www. ncbi.nlm.nih.gov/BLAST/ [1]) by standard nucleotide basic local alignment search tool (BLAST, version 2.2.1) without limiting queries and Sequence Match (version 2.7) at the RDP (http://rdp.cme.msu.edu/html/ [26]). The environmental sequences and sequences from GenBank were aligned in 830 positions (data set B) and in 1623 positions (data set S) using Sequencher and were manually adjusted to maximize conservation. Regions adjacent to the priming sites were omitted because of high frequency of ambiguous sites. Data set B contained one nontarget contaminant (B_Canopy_300_03_06 most similar to Prismatolaimus intermedius, Enoplida, in BLAST searches; Table 1) and three clones that contained large insertions and were unalignable with other fungal sequences (B_Canopy_300_01_16, B_Canopy_300_ 02_10, and B_Canopy_450_03_02; Table 1). Although A. JUMPPONEN: FUNGI IN THE WILLOW CANOPY SOIL ON A GLACIER FOREFRONT large insertions have been observed in the rDNA of Helotiales, Lecanorales, and Onygenales (see [3, 16, 17, 29]), the unalignable sequences were omitted because true insertions and chimeric PCR products could not be identified reliably. The taxonomic relationships among the fungal sequences were inferred by neighbor-joining (NJ) analyses in phylogenetic analysis using parsimony (PAUP*) [37]. A chytridiomycetous fungus (Monoblepharis hypogyna) was selected for the outgroup. Data matrices were left uncorrected, rates for variable sites were assumed equal, and no sites were assumed invariable. Sites with missing data, ambiguous nucleotides, or gaps, were randomly distributed among taxa. The robustness of the inferred NJ topologies was tested by 1000 bootstrap replicates. The most parsimonious trees were obtained using random addition sequence and a branch-swapping algorithm with tree bisection reconnection. The number of equiparsimonious trees was expected to be high attributable to several closely related sequences in the clone libraries. As a result, the maximum number of retained trees was restricted to 1000. The consensus (50% majority rule) and NJ topologies placed the environmental sequences similarly (data not shown). Detection and Analysis of Chimeric Sequences. Chimeric sequences may be frequent in environmental samples with diverse, mixed populations of competing templates [19, 24, 42]. To identify the most likely chimera breakpoints, all sequenced clones were analyzed by the Chimera Check program of the RDP (version 2.7 [26]). To test the effects of the chimeric sequences on the placement of the environmental clones in the obtained NJ topologies, the data were reanalyzed after exclusion of data upstream and downstream of the most commonly encountered chimera breakpoints (positions 1–391 and 502–830 in data set B alignment and positions 1–730 and 902–1623 in data set S). The obtained topologies were compared to detect clones that clearly changed positions in different analyses. Results Fungal Community Analyses. A total of 480 rDNA clones in 24 libraries were screened, and unique RFLP phenotypes were identified and sequenced to assay fungal community composition within established Salix spp. canopies in a primary successional ecosystem. After Figure 2. Neighbor-joining analysis of environmental partial 18S sequences (see Table 1 for accession numbers; AY382401–AY382473) obtained with primer set S (EF4 and EF3 [32]) from willow canopy soil on the forefront of a receding glacier. Accession numbers of the GenBank-obtained sequences are shown in parentheses. Sequence data were aligned in Sequencher and neighbor-joining analyses performed in PAUP* [37]. Numbers above the nodes refer to the occurrence of that node in 1000 bootstrap replicates. Values 9 50% are shown. A. JUMPPONEN: FUNGI IN THE WILLOW CANOPY SOIL ON A GLACIER FOREFRONT 239 240 exclusion of likely chimeric sequences, data set B contained 24 and data set S 18 unique clones. BLAST (Table 1) and NJ analyses (Figs. 1 and 2) placed the cloned environmental sequences into the kingdom Fungi. The target sequences broadly represented fungi including Ascomycota, Basidiomycota, Chytridiomycota, and Zygomycota. Overall, the cloned sequences indicated the presence of various groups of fungi in the soil underneath the willow canopies at the receding glacier forefront. Nontarget contaminants were rare; one clone was determined to be a nematode (P. intermedius). Three additional sequences were omitted because they contained large unalignable inserts whose origin could not be confirmed to be fungal. The majority of clones obtained with both primer pairs were placed among hymenomycetes and filamentous ascomycetes. The clones included taxa with likely affinities within the ascomycetous Sordariomycetes and basidiomycetous Russulales and Thelephorales (Figs. 1 and 2; Table 1). Two general points are noteworthy. First, various basidiomycete clones likely represent ectomycorrhizal fungi. Clones in data sets B and S had well-supported affinities within Russulaceae (B_Canopy_900_01_19 in data set B and S_Canopy_900_01_11 in data set S) and Thelephoraceae (B_Canopy_900_02_06 and B_Canopy_900_03_17 in data set B and S_Canopy_750_01_10 and S_Canopy_900_03_02 in data set S). Second, some ascomycete clones, similarly, are likely to form associations with willow roots. Both data sets contained clones with well-supported affinities to Sordariales (B_Canopy_900_02_04 in data set B and S_Canopy_450_02_05 and S_Canopy_750_02_09 in data set S). These sordarialean fungi are likely similar to those forming ectomycorrhizas with willows as reported earlier by Trowbridge and Jumpponen [39]. Most clone libraries were dominated by a single sequence type (Table 1). In two cases (samples S_Canopy_ 450_2 and S_Canopy_900_03), the libraries contained only one sequence type. These libraries were unlikely to be representative because data set B contained more than one sequence type in those samples. The dominant, nonchimeric sequence types in data set B were not identical with those in data set S suggesting primer bias (see below). Congruence in Fungal Community Composition Analysis of the 18S Among the Two Data Sets. rDNA with two different primer sets designed to be specific to fungi congruently identified several groups. A. JUMPPONEN: FUNGI IN THE WILLOW CANOPY SOIL ON A GLACIER FOREFRONT These included well-supported groups with affinities within Sordariales, Russulaceae, and Thelephoraceae. However, after exclusion of all suspected chimeric data, several incongruences were also evident (Figs. 3 and 4). Data set B (20 ascomycete clones of the 24 total clones) contained a larger number of ascomycete sequences than did data set S (4 ascomycete clones of the 18 total clones). Many of the groupings were not supported in bootstrap analyses, but three ascomycete groups exemplify the more abundant detection of ascomycetes in data set B. First, two clones (B_Canopy_450_03_14 and B_Canopy_450_03_17) were placed among Dothideomycetes with reasonably high bootstrap support in NJ analyses (Fig. 1). Second, three clones (B_Canopy_300_ 02_05, B_Canopy_300_02_06, and B_Canopy_750_ 03_11) were grouped with Peziza griseorosea with 100% bootstrap support, strongly indicating an affinity within Pezizaceae. Third, five clones (B_Canopy_300_03_17, B_Canopy_450_01_06, B_Canopy_450_02_13, B_Canopy_750_02_13, and B_Canopy_900_03_09) from five different samples were placed on a sister clade to Ophiostomatales. None of these well-supported groups occurred in data set S. Data set S contained well-supported groups within Chytridiomycota (S_Canopy_300_03_04; Fig. 2) and Zygomycota (S_Canopy_300_02_11; Fig. 2). In contrast, data set B contained no clones representing lower fungi. This result was not attributable to mere exclusion of chimeric data, as no lower fungi were detected in data set B in BLAST analyses. Data set S also included a large group of basidiomycetes with likely affinities within Cortinariaceae representing at least two distinct taxa (Cortinarius sp. and Inocybe sp.). No clones had wellsupported affinities to Cortinariaceae in data set B, although at least three sequences were determined most similar to Inocybe geophylla in BLAST analyses. Detection and Importance of Chimeric Sequences. A majority of the environmental sequences were determined to be likely chimeric by Chimera Check of the RDP. Further testing by reanalyses identified 17 chimeras in data set B and 8 in data set S (Figs. 3 and 4). Exceptionally high scores (980) in Chimera Check were always confirmed chimeric in the reanalyses. Lower scores did not indicate nonchimeric origin of a sequence, but many sequences could be confirmed chimeric in the NJ analyses (Table 1). Many of the chimeric sequences were likely a result of combined PCR products of templates representing fungi from different Figure 3. Reanalyses of data set B. Phylogram obtained by neighbor-joining analysis after the omission of potentially chimeric upstream data (positions 502–830) as identified by Chimera Check. Arrows on the right show the new placement of environmental sequences after the omission of potentially chimeric downstream data (positions 1–391). The environmental sequences with unstable placements in these reanalyses were concluded to be chimeric and were excluded from analyses shown in Fig. 1. A. JUMPPONEN: FUNGI IN THE WILLOW CANOPY SOIL ON A GLACIER FOREFRONT 241 242 A. JUMPPONEN: FUNGI IN THE WILLOW CANOPY SOIL ON A GLACIER FOREFRONT A. JUMPPONEN: FUNGI IN THE WILLOW CANOPY SOIL ON A GLACIER FOREFRONT divisions as indicated by placement among Ascomycota in analyses utilizing only 50-end of the sequences and among Basidiomycota in analyses utilizing only 30-end of the sequences (e.g., B_Canopy_900_02_12 and B_Canopy_750_03_03 in data set B–Fig. 3; and S_Canopy_750_02_17 in data set S–Fig. 4). Data set S was expected to have a greater proportion of chimeras, as their likelihood was anticipated to increase with increasing amplicon length. Surprisingly, data set B contained 40% (17/43) chimeric sequences, whereas data set S contained only 31% (8/26) chimeras. Discussion Fungal Communities Within Willow Canopies in the Fungal PCR amplicons were Glacier Forefront Soil. successfully obtained from environmental soil samples collected at the forefront of a receding glacier. A large proportion of the sequences was determined to be chimeric by the Chimera Check software of the RDP. Analyses conducted after exclusion of the sequence data potentially obtained from another target organism confirmed many chimeras, but the placement of most cloned sequences was insensitive to the exclusion of the potentially chimeric data. In other words, the placement of a majority of the cloned sequences was similar whether or not the data identified as possibly chimeric by Chimera Check were included in the analyses. After exclusion of chimeric data, 24 and 18 environmental sequences were analyzed in the two data sets. Most of the basidiomycetes detected in these analyses likely represented ectomycorrhizal associates of the willow plants. Earlier studies on sporocarp occurrence have indicated that Cortinariaceae (Inocybe spp. and Cortinarius spp.) and Tricholomataceae (Laccaria spp.) are common throughout the primary successional glacier forefront [21, 22]. Neither primer set produced cloned sequences that would find strongly supported affinity to Laccaria spp. in the NJ analyses, although both data sets contained nonchimeric sequences that were deemed similar to Laccaria pumila in BLAST analyses. The absence of support in NJ analyses is likely because of the poor resolution within the Agaricales that the 18S rDNA data provide. Several sequences similar to Cortinariaceae were detected in both data sets, although only data set S had well-supported affinities to Cortinarius iodes and I. geophylla. Additional infrequently fruiting ectomycorrhizal fungi exclusive to areas adjacent to the terminal moraine (Russulales representing genera Lactar- 243 ius and Russula [21, 22]) were detected in the soil samples collected 900 m from the glacier terminus by both primer sets. Finally, ectomycorrhizal fungi with inconspicuous fruiting bodies (Thelephoraceae) were detected within the willow canopies furthest from the glacier terminus by both primers. Although functional roles of the ectomycorrhizal basidiomycetes are often simple to decipher from their affinities to taxa available in sequence databases, the function of a majority of ascomycetes detected in these analyses remain unclear. Data set B contained clones with affinities to Pezizales (P. griseorosea), and both data sets contained clones with well-supported affinities to Sordariales. Several taxa within Pezizales have various associations ranging from pathogenicity to mycorrhizal symbiosis with ectomycorrhizal hosts [7, 8, 10]. Recent studies at the Lyman glacier site have suggested that taxa with affinities to Sordariales may, unexpectedly, be common mycorrhizal associates of the shrub willows [39]. Although it is very likely that many cloned ascomycetes represent these (facultative) biotrophic associations, various groups of the detected ascomycetes (e.g., taxa with affinities to Dothideales) are soil-inhabiting saprobes. Congruence in Fungal Community Composition Differential PCR ampliAmong the Two Data Sets. fication may be a result of various factors including template concentration, numbers of template molecules, GC content of the template molecules, efficiency of primer-template hybridization, polymerase extension efficiency for different templates, relative substrate exhaustion for different templates, and primer specificity [5, 12, 36, 42, 44]. The presented results of rDNA analyses using two sets of primers confirmed predicted EF4–EF3 primer bias toward basidiomycetes and lower fungi [2, 32]. Only 4 of the 18 nonchimeric clones in data set S were ascomycetous, whereas ascomycetes comprised a majority of nonchimeric clones in data set B (20 ascomycetes of the total of 24 nonchimeric sequences). Although not observed in the present study, primers for data set B do amplify chytridiomycetes and zygomycetes from environmental samples [3, 19]. The observed incongruences are therefore likely to have resulted either from true primer bias or from stochastic variation within an environmental DNA extract. However, the two different fungus-specific primers congruently identified several groups. These included well-supported groups with affinities within Sordariales, Russulaceae, and Thelephoraceae. The congruence among Figure 4. Reanalyses of data set S. Phylogram obtained by neighbor-joining analysis after the omission of potentially chimeric upstream data (positions 902–1623) as identified by Chimera Check. Arrows on the right show the new placement of environmental sequences after the omission of potentially chimeric downstream data (positions 1–730). The environmental sequences with unstable placements in these reanalyses were concluded to be chimeric and were excluded from analyses shown in Fig. 2. 244 A. JUMPPONEN: FUNGI the data sets could possibly have been improved by increasing the number of clones sampled from each library. However, there is often a compromise between the number of clones sampled from each library and the number of samples to be processed. Clearly, choice of primers and the number of sampled transformants within the clone libraries have a pivotal importance on the observed community structure. Comparisons among multiple extracts of the same sample, two or more primer sets, as well as multiple replicate samples may be necessary to obtain a more comprehensive view of the fungal communities. IN THE WILLOW CANOPY SOIL ON A GLACIER FOREFRONT after the omission of potentially chimeric regions included such groups. Both B and S data sets included cloned sequences that were basal to ascomycetous Saccharomycetales (e.g., B_Canopy_750_02_19 in Fig. 3 and S_Canopy_300_02_01 in Fig. 4) and basidiomycetous hymenomycetes (e.g., B_Canopy_300_01_18 and S_Canopy_750_03_18). Data set S included a sequence (S_Canopy_300_02_19) that was positioned basal to higher fungi (i.e., Ascomycota and Basidiomycota). None of the sequences placed in these basal positions were consistent in the reanalyses of the partial data sets and were therefore concluded to be PCR artifacts. Detection and Importance of Chimeric Sequences. Chimera Check overestimated the number of chimeric sequences as determined in confirmatory NJ analyses. However, sequences with high scores (980) in the Chimera Check were always confirmed chimeric. Lower scores included sequences that were determined chimeric and many that appeared stable in their position in confirmatory NJ analyses. The NJ analyses presented here aimed to identify and detect sequences whose positions in the obtained topologies were inconsistent when only 50-ends or only 30-ends of the sequences were utilized. Reanalyses of partial data sets identified 17 chimeras in data set B and 8 in data set S, more than 30% of all analyzed sequences. Similar chimera frequencies have been observed in bacterial community analyses [43] and analyses of somatic mutations [13]. Chimeric sequences are particularly frequent if sequence similarity among the competing templates and the number of PCR cycles are high [13, 43]. Accordingly, simple precautionary measures, such as longer extension times and fewer PCR cycles [42, 43], to minimize the generation of chimeras seem necessary. It was hypothesized that longer target amplicons would be more susceptible for chimera formation. Unexpectedly, the data set with shorter target amplicon had greater number of identified chimeras. This observation may be a result of the larger number of competing templates with fairly high similarity when primers with lesser bias were used (data set B; see [13, 43]). Overall, more data (longer amplicons) are usually beneficial, as they often allow better resolution in inferred topologies [18]. This is especially important when using conserved gene regions such as the 18S of the rDNA. It appears that the generation of chimeras is stochastic, and that targeting shorter amplicons may be unnecessary in fear of poor-quality environmental sequence data if steps to minimize chimera formation have been taken. Recent studies that utilize direct PCR from environmental samples have suggested frequent occurrences of novel fungal phyla, which find positions basal to filamentous ascomycetes or hymenomycetes [31, 41]. The preliminary analyses conducted prior to exclusion of chimeras as well as the analyses using partial sequences Conclusions The results indicate that ascomycetous and basidiomycetous ectomycorrhizal fungi comprise a substantial component in the fungal communities associated with the established willow canopies in primary successional ecosystems on the forefront of a receding glacier. Use of different primers yielded different results and supported different conclusions. It seems therefore necessary to view the results of direct molecular assessments with some caution. Finally, chimeras seem to comprise a large proportion of the environmental sequence data as determined by the Chimera Check of RDP and data reanalyses. Many of the chimeric reads appeared to comprise novel taxa at least on the level of an order. However, because it is possible that these sequences may be but PCR artifacts, the discovery of novel taxa without microscopic or culture-based confirmation may be premature. Acknowledgments This work was supported by Kansas State University BRIEF program, National Science Foundation EPSCoR Grant No. 9874732 with matching support from the State of Kansas, and National Science Foundation Grant No. OPP-0221489. I am grateful to Dr. Francesco T. Gentili, Nicolo Gentili, Anna Jumpponen, and Dr. James M. Trappe for their assistance during sample collection, transport, and preparation in August 2001 and to Emily L. King and Justin Trowbridge for their assistance in clone library screening and plasmid preparation. Dr. Charles L. Kramer, Nicholas B. Simpson, and Dr. James M. Trappe provided helpful comments on early drafts of this manuscript. Nicholas B. Simpson edited the manuscript. References 1. Altschul, SF, Madden, TL, Schäffer, AA, Zhang, J, Zhang, Z, Miller, DJ, Lipman, DJ (1997) Gapped BLAST and PSI-BLAST: a new generation of protein database search programs. Nucleic Acids Res 25: 3389–3402 A. JUMPPONEN: FUNGI IN THE WILLOW CANOPY SOIL ON A 245 GLACIER FOREFRONT 2. Anderson, IC, Campbell, CD, Prosser, JI (2003) Potential bias of fungal 18S rDNA and internal transcribed polymerase chain reaction primers for estimating fungal biodiversity in soil. Environ Microbiol 5: 36–47 3. Borneman, J, Hartin, RJ (2000) PCR primers that amplify fungal rRNA genes from environmental samples. Appl Environ Microbiol 66: 4356–4360 4. Cázares, E (1992) Mycorrhizal fungi and their relationship to plant succession in subalpine habitats. PhD Thesis, Oregon State University 5. Chandler, DP, Fredrickson, JK, Brockman, FJ (1997) Effect of PCR template concentration on the composition and distribution of total community rDNA clone libraries. Mol Ecol 6: 475–482 6. Cornelissen, JHC, Aerts, R, Cerabolini, MJA, Werger, MJA, van derHeijden, MGA (2001) Carbon cycling traits of plant species are linked with mycorrhizal strategy. Oecologia 129: 611–619 7. Dahlstrom, JL, Smith, JE, Weber, NS (2000) Mycorrhiza-like interaction by Morchella with species of the Pinaceae in pure culture synthesis. Mycorrhiza 9: 279–285 8. Danielson, RM (1984) Ectomycorrhiza formation by the operculate discomycete Sphaerosporella brunnea (Pezizales). Mycologia 76: 454–461 9. Dickie, IA, Xu, B, Koide, RT (2002) Vertical niche differentiation of ectomycorrhizal hyphae in soil as shown by T-RFLP analysis. New Phytol 156: 527–535 10. Egger, KN, Paden, JW (1986) Biotrophic associations between lodgepole pine seedlings and post-fire ascomycetes (Pezizales) in monoxenic culture. Can J Bot 64: 2719–2725 11. Egli, S, Ayer, F, Chatelain, F (1997) Die Beschreibung der Diversität von Macromyceten. Erfahrungen aus pilzölologischen Langenzeitstudien im Pilzreservat La Chanéaz, FR. Mycol Helv 9: 19–32 12. Farrelly, V, Rainey, FA, Stackebrandt, E (1995) Effect of genome size and rrn gene copy number on PCR amplification of 16S rRNA genes from a mixture of bacterial species. Appl Environ Microbiol 91: 2798–2801 13. Ford, JE, McHeyzer-Williams, MG, Lieber, MR (1994) Chimeric molecules created by gene amplification interfere with the analyses of somatic hypermutation of murine immunoglobulin genes. Gene 142: 279–283 14. Gardes, M, Bruns, TD (1993) ITS primers with enhanced specificity for higher fungi and basidiomycetes: application to identification of mycorrhizae and rusts. Mol Ecol 2: 113–118 15. Gardes, M, Bruns, TD (1996) ITS-RFLP matching for the identification of fungi. In: Clapp, JP (Ed.) Methods in Molecular Biology, Vol. 50: Species Diagnostics Protocols: PCR and Other Nucleic Acid Methods. Humana Press Inc., Totowa, NJ, pp 177– 186 16. Gargas, A, DePriest, PT, Taylor, JW (1995) Positions of multiple insertion in SSU rDNA of lichen-forming fungi. Mol Biol Evol 12: 208–218 17. Holst-Jensen, A, Vaage, M, Schumacher, T, Johansen, S (1999) Structural characteristics and possible horizontal transfer of group I introns between closely related plant pathogenic fungi. Mol Biol Evol 16: 114–126 18. Hugenholtz, P, Goebel, BM, Pace, NR (1998) Impact of cultureindependent studies on the emerging phylogenetic view of bacterial diversity. J Bacteriol 180: 4765–4774 19. Jumpponen, A (2003) Soil fungal community assembly in a primary successional glacier forefront ecosystem as inferred from rDNA sequence analyses. New Phytol 158: 569–578 20. Jumpponen, A, Mattson, K, Trappe, JM, Ohtonen, R (1998) Effects of established willows on primary succession on Lyman Glacier forefront: evidence for simultaneous canopy inhibition and soil facilitation. Arct Alp Res 30: 31–39 21. Jumpponen, A, Trappe, JM, Cázares, E (1999) Ectomycorrhizal 22. 23. 24. 25. 26. 27. 28. 29. 30. 31. 32. 33. 34. 35. 36. 37. 38. 39. fungi in Lyman Lake Basin: a comparison between primary and secondary successional sites. Mycologia 91: 575–582 Jumpponen, A, Trappe, JM, Cázares, E (2002) Occurrence of ectomycorrhizal fungi on a receding glacier forefront. Mycorrhiza 12: 43–49 Jumpponen, A, Väre, H, Mattson, KG, Ohtonen, R, Trappe, JM (1999) Characterization of Fsafe sites_ for pioneers in primary succession on recently deglaciated terrain. J Ecol 87: 98–105 Kopczynski, ED, Bateson, MM, Ward, DM (1994) Recognition of chimeric small-subunit ribosomal DNAs composed from genes from uncultivated microorganisms. Appl Environ Microbiol 63: 3614–3621 Kowalchuk, GA, Gerards, S, Woldendorp, JW (1997) Detection and characterization of fungal infections of Ammophila arenaria (marram grass) roots by denaturing gradient gel electrophoresis. Appl Environ Microbiol 63: 3858–3865 Maidak, BL, Cole, JR, Jr, Parker, CT, Garrity, GM, Larsen, N, Li, B, Lilburn, TG, McCaughey, MJ, Olsen, GJ, Overbeek, R, Pramanik, TM, Schmidt, TM, Tiedje, JM, Woese, CR (1999) A new version of the RDP (Ribosomal Database Project). Nucleic Acids Res 27: 171– 173 O’Dell, TE, Smith, JE, Castellano, M, Luoma, D (1996) Diversity and conservation of forest fungi. In: Pilz, D, Molina, R (Eds.) Managing Forest Ecosystems to Conserve Fungus Diversity and Sustain Wild Mushroom Harvests. U.S. Forest Service General Technical Reports, PNW-GTR-317. US Department of Agriculture, Forest Service, Pacific Northwest Research Station. Portland, OR, pp 5–18 Pennanen, T, Paavolainen, L, Hantula, J (2001) Rapid PCR-based method for the direct analysis of fungal communities in complex environmental samples. Soil Biol Biochem 33: 697–699 Perotto, S, Nepote-Fus, P, Saletta, L, Bandi, C, Young, JPW (2000) A diverse population of introns in the nuclear ribosomal genes of ericoid mycorrhizal fungi includes elements with sequence similarity to endonuclease-coding genes. Mol Biol Evol 17: 44–59 Sambrook, J (1989) Molecular Cloning—A Laboratory Manual, 2nd ed. In: Fritsch, EF, Maniatis, T (Eds.) Cold Spring Laboratory Press, New York Schadt, CW, Martin, AW, Lipson, DA, Schmidt, SK (2003) Seasonal dynamics of previously unknown fungal lineages in tundra soils. Science 301: 1359–1361 Smit, E, Leeflang, P, Glandorf, B, vanElsas, JD, Wernars, K (1999) Analysis of fungal diversity in the wheat rhizosphere by sequencing of cloned PCR-amplified genes encoding 18S rRNA and temperature gradient gel electrophoresis. Appl Environ Microbiol 65: 2614–2621 Smit, E, Veenman, C, Baar, J (2003) Molecular analysis of ectomycorrhizal basidiomycete communities in Pinus sylvestris L. stand reveals long-term increased diversity after removal of litter and humus layers. FEMS Microbiol Ecol 45: 49–57 Smith, SE, Read, DJ (1997) Mycorrhizal Symbiosis. Academic Press. London Straatsma, G, Krisai-Greilhuber, I (2003) Assemblage structure, species richness, abundance, and distribution of fungal fruitbodies in a seven year plot-based survey near Vienna. Mycol Res 107: 632–640 Suzuki, MT, Giovannoni, SJ (1996) Bias caused by template annealing in the amplification mixtures of 16S rRNA genes by PCR. Appl Environ Microbiol 62: 625–630 Swofford, DL (2001) PAUP, Phylogenetic Analysis Using Parsimony (and Other Methods), Version 4. Sinauer Associates. Sunderland, MA Torsvik, V, Goksøyr, J, Daae, FL (1990) High diversity on DNA of soil bacteria. Appl Environ Microbiol 56: 782–787 Trowbridge, J, Jumpponen, A (2004) Fungal colonization of shrub willow roots at the forefront of a receding glacier. Mycorrhiza 14: 283–293 246 40. Vainio, EJ, Hantula, J (2000) Direct analysis of wood-inhabiting fungi using denaturing gradient gel electrophoresis of amplified ribosomal DNA. Mycol Res 104: 927–936 41. Vandenkoornhuyse, P, Baldauf, SL, Leyval, C, Straczek, J, Young, JPW (2002) Extensive fungal diversity in plant roots. Science 295: 2051 42. vonWintzingerode, F, Göbel, UB, Stackebrandt, E (1997) Determination of microbial diversity in environmental samples: pitfalls of PCR-based rRNA analysis. FEMS Microbiol Rev 21: 213–229 A. JUMPPONEN: FUNGI IN THE WILLOW CANOPY SOIL ON A GLACIER FOREFRONT 43. Wang, GCY, Wang, Y (1996) The frequency of chimeric molecules as a consequence of PCR co-amplification of 16S rRNA genes from different bacterial species. Microbiology 142: 1107–1114 44. Zheng, D, Alm, EW, Stahl, DA, Raskin, L (1996) Characterization of universal small-subunit rRNA hybridization probes for quantitative molecular microbial ecology studies. Appl Environ Microbiol 62: 4504–4513