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Walla Walla College, College Place, Washington 99324
David S. Hibbett
Department of Biology, Clark University, 950 Main St., Worcester, Massachusetts 01610-1477
Joseph F. Ammirati, Jr.
Department of Botany, University of Washington, Seattle, Washington 98195
Russell J. Rodriguez 1
U.S. Geological Survey, WFRC, 6505 N.E. 65th St., Seattle, Washington 98115, and Department of Botany, University of Washington, Seattle, Washington 98195
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ABSTRACT |
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The genetic diversity and phylogeny of Bridgeoporus nobilissimus have been analyzed. DNA was extracted from spores collected from individual fruiting bodies representing six geographically distinct populations in Oregon and Washington. Spore samples collected contained low levels of bacteria, yeast and a filamentous fungal species. Using taxon-specific PCR primers, it was possible to discriminate among rDNA from bacteria, yeast, a filamentous associate and B. nobilissimus. Nuclear rDNA internal transcribed spacer (ITS) region sequences of B. nobilissimus were compared among individuals representing six populations and were found to have less than 2% variation. These sequences also were used to design dual and nested PCR primers for B. nobilissimus-specific amplification. Mitochondrial small-subunit rDNA sequences were used in a phylogenetic analysis that placed B. nobilissimus in the hymenochaetoid clade, where it was associated with Oxyporus and Schizopora.
Key words: Abies procera, Basidiomycete, conk, fungi, Fuzzy Sandoze, hymenochaetoid clade, molecular systematics, Noble fir, Noble polypore, old-growth forest, Oxyporus, Polyporaceae
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONLITERATURE CITED |
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, Castellano et al 1999).
The fruiting bodies of B. nobilissimus demonstrate three general shapes, depending largely on their location on the host. Hoof-shaped and shelf-like conks are found on the sides of hosts. Short, oblong-topped conks with tapering pore surfaces are found growing on the main roots of the host and centrally substipitate conks are located on the tops of stumps (Hibler and O'Dell 1997).
A description of macro- and microscopic morphological characteristics by Burdsall et al (1996) includes: The basidioma is characterized by a fibrous pileal surface with cracks and crevices from which vascular plants (e.g., Oxalis sp. and pteridophytes) and bryophytes often grow epiphytically. The pileal surface is covered with a dense mat of white mycelial fibers in young basidiomata, often somewhat agglutinated at the tips, which become darker with age, reaching several mm in length. These fibers often are green, due to epimycotic associations with several species of algae, including Coccomyxa sp. and Charicium species. B. nobilissimus lacks clamp connections at the septa and has a monomitic hyphal system with pseudocystidia arising from the trama. The spores are 5.5–6.5 x 3.5–4.5 µm, broadly ovoid, hyaline, smooth, thin-walled and inamyloid. The round pores are stratified with a layer of sterile tissue 2–3 mm thick between successive annual pore layers.
This polypore initially was named Oxyporus nobilissimus by William Bridge Cooke in 1949. A study by Burdsall et al (1996) identified characteristics of O. nobilissimus that were incompatible with the genus Oxyporus and could find no other described genus appropriate for this species. Although the stratification of tube layers and context is a unique characteristic shared by O. populinus and O. nobilissimus (Cooke 1949, Gilbertson and Ryvarden 1987 p 499), all species of Oxyporus have true cystidia arising from the subhymenium, whereas O. nobilissimus has pseudocystidia of tramal origin. Burdsall et al (1996) considered the genus Rigidoporus the "closest placement" for O. nobilissimus because of these shared characteristics: pseudocystidia of tramal origin, thin- and thick-walled hyphae and similar basidium and basidiospore shape and size. Both Rigidoporus and Oxyporus cause a white rot in which lignin as well as other wood components are decomposed, whereas O. nobilissimus consistently has been associated with brown rot (Burdsall et al 1996), in which lignin is not appreciably degraded (Boddy and Watkinson 1995). However, the true rot characteristics of this species remain unresolved because efforts to culture O. nobilissimus have been unsuccessful to date (Cooke 1949; Burdsall et al 1996; Mueller G., pers comm, 1997). Because of these incompatibilities, the genus Bridgeoporus was created to accomodate O. nobilissimus (Burdsall et al 1996).
Bridgeoporus nobilissimus was placed on the Oregon Natural Heritage program's endangered species list in 1995 (as Oxyporus nobilissimus), making it one of the first fungi to be so listed by any private or public agency in the United States (Lizon 1995). As of 2000, 81 sporocarps of B. nobilissimus, many of them in various states of decline, had been located at 12 sites (Hibler and O'Dell 1997; Castellano et al 1999; Cowden, pers comm, 2002). Genetic studies of B. nobilissimus have not been reported nor have the spores been germinated successfully under laboratory conditions to date.
The purpose of this study was to conduct a genetic analysis of spores from 22 individual B. nobilissimus sporocarps in six populations in the Pacific Northwest (Table I) and to develop a diagnostic system for detecting B. nobilissimus in wood or soil substrates. Nuclear rDNA was sequenced for genetic analysis to develop Bridgeoporus-specific polymerase chain reaction (PCR) primers and to examine relationships between B. nobilissimus spore DNA and the DNA of a putative filamentous fungus cultured from B. nobilissimus tissue and spore drops.
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METHODS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONLITERATURE CITED |
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) agar plates that were incubated at 25 C. The isolated colonies (referred to as spore-drop cultures) were cultured further in Luria Broth, M3S and SOC broth. Control samples of Fomitopsis pinicola, Ganoderma applanatum and G. oregonense were collected from conks in the South Fork of the Hoh River watershed in Olympic National Park. Tissue samples (approximately 0.5 cm3) from the growing hymenium of each species were placed in 1.5 mL microcentrifuge tubes containing 0.5 mL of a DNA preservative (150 mM EDTA, 50 mM Tris pH 8.0, 2.0% n-lauroylsarcosine), and the DNA was extracted as below. Paul Stamets (Fungi Perfecti, P.O. Box 7634, Olympia, WA) provided three mycelial cultures derived from spores and tissue of B. nobilissimus for sequencing (Table I).
DNA extraction –
DNA was extracted from the mycelial cultures using Rodriguez's (1993) method. DNA was extracted from the spore samples using Bio 101 DNA FastPrep kit and a modification of Rodriguez's (1993) method. Approximately 100 µL of an aqueous spore suspension were placed in each Bio 101 homogenization tube containing 250 mg of 0.8 mm zircon beads with 2.0 mm spheres. Eight hundred µL of CLS-Y buffer were added and the tubes placed in the FastPrep cell disrupter for three cycles at speed 5.0 for 40 s/cycle. Tissue samples from the hymenium of control fungi were pulverized with a small pestle and incubated at 65 C for 30 min, then processed along with spore samples. Tubes were centrifuged for 5 min at 14 000x g and the supernatant transferred into 1.5 mL microcentrifuge tubes. 0.7 volumes of PEG/NaCl (20% polyethylene glycol mw 8000, and 2.5 N sodium chloride) solution were added, mixed thoroughly and iced 5–20 min to precipitate nucleic acids, proteins and polyphosphates. Tubes were centrifuged for 5 min at 14 000x g, decanted, and pellets resuspended in 0.5 mL TE buffer (10 mM Tris pH 8.0, 1.0 mM EDTA). Any scum present on the surface of the supernatant was retained with the pellet. After the pellets dissolved, 0.5 volumes 7.5 N ammonium acetate were added and tubes were incubated in ice 5–20 min. Protein, RNA and polyphosphates, etc. were pelleted at 14 000x g for 5 min. The supernatant was transferred, 0.6 volumes of isopropyl alcohol added, and samples gently mixed and incubated in ice for 5–30 min to precipitate DNA. DNA was pelleted for 5 min at 14 000x g and resuspended in 0.5 mL TE Buffer. DNA was reprecipitated by the addition of NaCl to 0.1 M and two volumes of chilled 95% ethanol. Samples were incubated in ice for 5–30 min, DNA pelleted at 14 000x g for 5 min and resuspended in 0.5 mL TE Buffer. The size of the DNA was assessed by electrophoresis in a 0.7% agarose gel. The gels were stained with ethidium bromide and DNA visualized with 305 nm ultraviolet light (Sambrook et al 1989).
Bacterial and fungal DNA from sporedrop cultures were extracted in a manner similar to that of the spores. This exception applied: Instead of using the cell disrupter, the cultures were centrifuged and the pellets treated with 300 µL lysis buffer (EDTA 150 mM, Tris pH 8.0 at 50 mM, Sarcosyl 2.0%, Protease, and water to volume) vortexed for 1–5 s, and heated at 65 C. After 15–30 min the cultures were centrifuged at 14 000x g for 5 min and the supernatant transferred to a new 1.5 mL tube and the DNA purified as described above.
PCR amplification –
The ITS regions of nuclear rDNA of all samples were PCR amplified with primer sets ITS1/ITS4 (White et al 1990), ITS1-F/ITS4-B (Gardes and Bruns 1993) and Eubac27F/Eubac492R (DeLong 1992) to distinguish between ascomycete, basidiomycete and eubacterial DNA. PCR reactions were performed in 20 µL reactions containing 10 mM Tris-HCl (pH 9.0), 50 mM KCl, 2.5 mM MgCl2, 0.2% Triton X-100, 200 µM each of dATP, dCTP, dGTP, dTTP (Pharmacia), 0.2 units Taq DNA polymerase, 250 ng of each oligonucleotide primer, 20–200 ng of fungal genomic DNA.
Amplifications involved 35 cycles of this temperature regime: Denaturing at 93 C for 15 s, annealing at 64 C for 1.5 min and synthesis at 72 C for 1.5 min. No ramp times were imposed from denaturing to annealing temperatures or from annealing to synthesis temperatures. The cycles were preceded by 2 min of denaturation at 93 C.
PCR primers MS1 and MS2 (White et al 1990) were used to amplify the mitochondrial small-subunit rDNA from B. nobilissimus spore samples and from mycelia of st-2 samples. The protocol was the same as described above, except that the annealing temperature was 60 C.
Electrophoresis of PCR-amplified products was performed in 2.0% agarose gels for 1.5 h at 7.0 V/cm2 (Sambrook et al 1989). PCR products were stained with ethidium bromide and visualized with 305 nm ultraviolet light.
Sequence analysis and design of diagnostic primers –
The QIAEX II Agarose Gel Extraction Protocol (Qiagen Inc., 28159 Avenue Stanford, Valencia, CA 91355) was used to purify the ITS1-F/ITS4-B and MS1/MS2 PCR products that then were quantified by fluorescence and sequenced by the chain termination method (Sanger et al 1977). ITS sequences were aligned using SequencherTM to generate consensus sequences. The consensus sequences for B. nobilissimus and the mycelial cultures (st-2, tc-a and tc-b) were aligned and primers were designed from regions of the sequences showing the greatest differences between these organisms. The B. nobilissimus primers Bn215-118 (ATGTGCTCGTTGCCCTTGAC) and Bn217-190 (AGTTGTCCGAAGACGATTAG) were located 118 and 190 bp from primers ITS1-F and ITS4-B, respectively. The primers for the mycelial cultures, St215-184 (GGTCTTAATCGACTCTGCTC) and St217-35 (GGACGATTGGAAGCCGAATA) were located 184 and 35 bp from ITS1-F and ITS4-B, respectively. Sequences of B. nobilissimus can be obtained from GenBank with the access numbers shown in Table I.
Nested primer PCR – To perform nested primer PCR (npPCR), dual primer reaction products amplified with ITS1-F/ITS4-B primers were diluted from 100x to 10 000x with 10 mM Tris pH 8.5 buffer and 2 µL transferred to a 0.5 mL microcentrifuge tube containing 18 µL fresh PCR reaction buffer and primers Bn215-118/Bn217-190 or St215-184/St217-35. The thermocyclers were programmed similarly to the dpPCR reactions, except the annealing temperatures were 68 C.
Phylogenetic analysis –
To evaluate the phylogenetic relationships of B. nobilissimus, mt-ssu rDNA sequences obtained from three spore preparations (gm-g, rn-b, sp-h) and one putative mycelial isolate (st-2) were added to a reference dataset containing 85 species of homobasidiomycetes, emphasizing wood-decaying taxa. The heterobasidiomycetes Auricularia auricula-judae and Dacrymyces chrysospermus were included for rooting purposes. All species other than B. nobilissimus were represented by sequences of nuclear small-subunit rDNA (ca 1.8 kb) as well as the mt-ssu rDNA. Sequences in the reference dataset were used in a study on the evolution of wood-rotting homobasidiomycetes (Hibbett and Donoghue 2001), which should be consulted for strain information and accession numbers.
Sequences were manually aligned to the reference dataset, and divergent regions were excluded from analysis. Phylogenetic analyses in PAUP*4.0 (Swofford 1999) used equally weighted parsimony. An initial search with 1000 heuristic searches was performed, with random taxon addition sequences and TBR branch swapping, keeping up to 10 trees per replicate. Trees found in the initial search were used as input trees for TBR branch swapping with MAXTREES set to autoincrease. Bootstrap analysis used 1000 replicates, with one heuristic search per replicate. Each bootstrap replicate used a single random taxon addition sequence with TBR swapping, keeping up to 10 trees.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONLITERATURE CITED |
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, were noted frequently. Culture analysis indicated that each of the spore samples contained low levels of bacteria and yeast (data not shown). Although these cultures were not defined taxonomically, they were used to extract genomic DNA for PCR analysis of rDNA (see below). In addition, only one filamentous fungus (st-2, tc-a, tc-b) was cultured from the spores and tissue of B. nobilissimus (Paul Stamets unpubl) and it was characterized by rDNA sequence analysis (described below).
Nuclear rDNA sequence analysis –
Microscopic analysis indicated that either the physical separation of spores from contaminating organisms or the germination of B. nobilissimus spores would be required for the extraction of pure B. nobilissimus DNA. However, efforts to accomplish either were unsuccessful. Therefore, DNA from spore samples and from the spore-drop cultures was PCR amplified with primers specific to the rDNA of ascomycetes (ITS1/ITS4, White et al 1990), basidiomycetes (ITS1-F/ITS4-B, Gardes and Bruns 1993) and eubacteria (Eubac27F/Eubac492R, DeLong 1992) for comparative analysis.
An 850 bp product was amplified from all 22 B. nobilissimus samples from six populations with the primers ITS1-F/ITS4-B (Fig. 1). The mycelial cultures (st-2, tc-b) also amplified with ITS1-F/ITS4-B producing an 800 bp product (Fig. 2). When amplified with the primer set ITS1/ITS4 (Fig. 3a, b) and with the primer set Eubac27F/492R (Fig. 4), B. nobilissimus spore DNA produced products that were similar in size to those produced by the yeast and bacterial spore-drop cultures. However, with one exception (gm-fy), the yeast and bacterial cultures did not amplify with the Basidiomycete primers ITS1-F/ITS4-B. Sequence analysis of this anomalous 850 bp yeast product (GenBank accession # AF509231) showed no similarity to the B. nobilissimus sequences.
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The ITS2 region of 19 isolates from all geographic areas displayed 100% identity. Sequences from wm, sp-m and gm-g were not included in the ITS2 analysis due to poor sequence quality. A BLAST analysis with the B. nobilissimus ITS sequences revealed no similarities with other taxa in GenBank.
The ITS2 region of the mycelial cultures (st-2 and tc-b) had 98.5% identity with Fomitopsis pinicola. The ITS1 region of F. pinicola was not found in the database, and no comparison could be made. As a result, the mycelia of st-2 and tc-b cultures were relabeled as a Fomitopsis sp.
Fidelity of taxon-specific primers – Nested PCR reactions, with the primer sets ITS1-F/ITS4-B and Bn215-118/Bn 217-190, specifically amplified B. nobilissimus and did not amplify DNA from other polypores or the Fomitopsis sp. cultures (Fig. 5). In addition, npPCR allowed B. nobilissimus DNA to be detected at concentrations of 100 pg/µL (Fig. 6). The Fomitopsis sp.-specific primer set St215-184 and St217-35 detected Fomitopsis sp. at concentrations of 1 pg/µL (data not shown) and did not amplify B. nobilissimus or other polypore DNA, including F. pinicola (Fig. 7). Thus, although this culture (st-2) likely represents a species of Fomitopsis, it probably is not Fomitopsis pinicola.
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DISCUSSION |
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). Participants in the 1996 study also noted several variations on cell morphologies during microscopic analysis of B. nobilissimus spore samples (Volk, T., pers comm, 1997). Therefore, it is not surprising that the spore samples of the current study contained other fungi, yeast and bacteria. While there is significant evidence of bacteria initiating basidiome formation (Rainey 1991) and stimulating germination of spores (Mayo et al 1986, Ali and Jackson 1989) in other fungi it is not yet known what role these microorganisms play in the biology of B. nobilissimus.
Although the intermingling of organisms has made the study of this polypore challenging, the use of taxon-specific primers to differentially amplify DNA mixtures is effective (Camacho et al 1997) and enabled us to analyze genetically B. nobilissimus. The regions of the nuclear genome coding for the ribosomal subunits have evolved at different rates. While the areas coding for the large and small subunits have been highly conserved among eukaryotes, the spacer regions between the subunits are highly variable among morphologically distinct fungal species. And yet, in these same spacer regions, intraspecific variation is low (Hillis and Dixon 1991, Gardes and Bruns 1993, Redecker et al 1997). Distinguishing among taxa was made possible by using primers embedded in the highly conserved regions. These same primers amplified the highly variable spacer regions that we were able to sequence and thus detect genetic differences among individual sporocarps.
Sequence analysis of both the ITS regions and the mitochondrial small rDNA region indicated that all of the fruiting bodies from which spore samples were obtained are of the same species. Because B. nobilissimus is an endangered species, we did not harvest any tissue samples and thus no comparison was made between spore and tissue DNA. However, the likelihood of the same contaminant being present on all 22 widely separated conks is remote. Morphological variation of the fruiting bodies might be due to differences in the location of the sporocarp on the substrate, nutritional inconsistencies or variation in types of colonizing organisms. It also is clear from the ITS sequencing that the three mycelial cultures (st-2, tc-a, tc-b) derived from tissues and spore-drops of B. nobilissimus are probably a species of Fomitopsis (Fig. 8).
The Forest Ecosystem Management Assessment Team (FEMAT) Report 1993 (p. IV 86–89) stated the need for information on old-growth associated fungi of the Pacific Northwest. It specifically noted that a long-term study of Oxyporus nobilissimus should be initiated and that it should include distance and effectiveness of spore dispersal. Such a study would require a molecular diagnostic system to detect B. nobilissimus in its vegetative stage.
The vegetative stage of a polypore may grow for a considerable span of time as it accumulates resources to form its large fruiting bodies (Gilbertson and Ryvarden 1986, p 5). In the case of B. nobilissimus, which is found only on ancient trees, this stage could span centuries, perhaps the entire life of the tree. During the vegetative stage it is not identifiable without the use of molecular techniques. A useful diagnostic system to determine the location of B. nobilissimus mycelium before it develops the sporocarp is the B. nobilissimus-specific PCR primer set. This primer set will enable future studies to be conducted on spore dispersal patterns and colonization patterns of the substrate by amplifying DNA extracted from soil and wood samples near existing conks.
Some questions this system could address regarding B. nobilissimus are: i. What are the specific habitat requirements (age and diameter range of host trees)? ii. How long does B. nobilissimus inhabit a substrate before fruiting? iii. What is its relationship with the associated yeast, bacteria and the Fomitopsis sp.? iv. Is it saprobic, pathogenic or possibly a commensal or mutualisitic endophyte to Abies procera?
The taxonomic placement of B. nobilissimus has been problematical (Burdsall et al 1996). Phylogenetic analysis of four mt-ssu rDNA sequences in the context of a broad dataset of lignicolous homobasidiomycetes placed three of the sequences in the hymenochaetoid clade (Hibbett and Thorn 2001) and the misidentified sequence in the polyporoid clade (Fig. 8). Taxa in the hymenochaetoid clade include members of the Hymenochaetaceae and other white-rotting, lignicolous species that traditionally have been placed in the Corticiaceae s. lat. and Polyporaceae s. lat. (Hibbett and Thorn 2001). The latter includes Oxyporus, which is represented in the reference dataset by one sequence identified as Oxyporus sp. (probably O. populinus, D. Hibbett pers obs). Burdsall et al (1996) segregated Bridgeoporus from Oxyporus, based on anatomical features and rot type. Nevertheless, results of the molecular phylogenetic analysis place Bridgeoporus in the white-rotting hymenochaetoid clade (Hibbett and Donoghue 1995) and in a weakly supported clade containing Bridgeoporus, Oxyporus and Schizopora (Fig. 8). If this is correct, then Bridgeoporus might represent another independent origin of the brown-rot mode of wood decay in the homobasidiomycetes (Hibbett and Donoghue 2001). On the other hand, B. nobilissimus actually might cause a white rot and the brown rot observed in the association is due to another species, possibly Fomitopsis. Gilbertson and Ryvarden (1986) noted that O. populinus resembles B. nobilissimus, which they classified in Oxyporus, in having a "perennial sessile basidiocarp with the tube layers separated by thin layers of context." In addition, B. nobilissimus and O. populinus both occur on living trees (hardwoods, in the case of O. populinus) and the pilei typically support mosses and other epimycota. Although the tramal origin of the cystidia makes this unlikely, B. nobilissimus indeed might be an Oxyporus, as described by Cooke (1949). However, the weak support for many nodes in the hymenochaetoid clade and limited sampling of Oxyporus spp. limit the conclusions that can be made at this time. The inclusion of Bridgeoporus and other nonxanthochroic taxa (e.g., Trichaptum, Schizopora) in the hymenochaetoid clade is at odds with the traditional delimitation of the Hymenochaetales. Nevertheless, members of the hymenochaetoid clade that have been studied at the ultrastructural level are united by possession of a nonperforated parenthesome (Hibbett and Thorn 2001). It would be valuable to determine whether Bridgeoporus has a nonperforated parenthesome, which would support its placement in the hymenochaetoid clade. Future analyses also should include additional isolates of Oxyporus spp., as well as Rigidoporus spp., and to seek genes that resolve relationships within the hymenochaetoid clade.
While rotting characteristics are considered important indicators at the genus level, they are not necessarily so at the family level. The characteristic of producing brown rot appears to have evolved repeatedly. Past phylogenetic studies of Polyporaceae have implicated a history of extensive convergence and parallelism in macro-morphological features, as well as considerable homoplasy in such physiological features as rotting characteristics (Hibbett and Donoghue 1995, 2001). Therefore it is not uncommon for fungi within the same family to have different rotting characteristics. Because B. nobilissimus has not been cultured successfully and its life history is poorly understood, its association with brown rot can be classified only as correlation rather than causal at this time.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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Accepted for publication February 10, 2003.
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LITERATURE CITED |
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