nena masthead
SENA Home Staff & Editors For Readers For Authors

An Initial Inventory of Bacteria Found within the Soils and Waters of Great Smoky Mountains National Park
Seán P. O’Connell, Emily A. York, Melissa B. Collins, Derren T. Rosbach, Kristina Reid Black, and Weaver B. Haney

Southeastern Naturalist, Volume 6, Special Issue 1 (2007): 35–72

Full-text pdf (Accessible only to subscribers.To subscribe click here.)


Access Journal Content

Open access browsing of table of contents and abstract pages. Full text pdfs available for download for subscribers.

Current Issue: Vol. 22 (3)
SENA 22(3)

Check out SENA's latest Special Issue:

Special Issue 12
SENA 22(special issue 12)

All Regular Issues


Special Issues






JSTOR logoClarivate logoWeb of science logoBioOne logo EbscoHOST logoProQuest logo

1Department of Biology, 132 Natural Science Building, Western Carolina University, Cullowhee, NC 28723. 2Current address - University of Alberta, Department of Medical Microbiology and Immunology, 1-41 Medical Sciences Building, Edmonton, AB T6G 2H7. 3Current address - Virginia Tech University, Department of Environmental Design and Planning, Blacksburg, VA 24061. 4Current address - Dynamac Corporation, Mail Code DYN-3, Kennedy Space Center, FL 32899. *Corresponding author - An Initial Inventory of Bacteria Found within the Soils and Waters of Great Smoky Mountains National Park Seán P. O’Connell1,*, Emily A. York1, Melissa B. Collins1,2, Derren T. Rosbach1,3, Kristina Reid Black1,4, and Weaver B. Haney1 Abstract - Soils and waters were collected from Great Smoky Mountains National Park (GSMNP), and bacteria were cultured or DNA cloned from the samples. Oconaluftee Visitors Center and Kephart Prong Trail were sampled to examine distributions of heterotrophs inhabiting streams and riparian soil. Soil from All Taxa Biodiversity Inventory plots at Albright Grove, Cataloochee, and Purchase Knob were also sampled. A total of eleven phyla were detected, of which six were only found via culture-independent techniques. Overall, 69 genera were documented, with differences in their detection in soil and water and by methodology. Firmicutes dominated cultures from soil, while Acidobacteria dominated clone libraries; Bacteroidetes was the dominant phylum in water. Three classes of the phylum Proteobacteria were commonly seen as isolates or clones. Prokaryotic diversity is extraordinary; this is the first inventory examining non-photosynthetic bacteria inhabiting GSMNP and it lays the groundwork for investigations exploring the true breadth of diversity in the park and what this diversity means to the broader ecosystem. Introduction The extent of bacterial diversity on Earth has been of great interest to taxonomists, microbial ecologists, and theoretical biologists over the last two decades (Curtis et al. 2002, DeLong and Pace 2001, Gans et al. 2005, Hong et al. 2006, Hughes et al. 2001, Torsvik et al. 1990, Whitman et al. 1998). With estimates ranging as high as ten million species in ten grams of pristine soil (Gans et al. 2005), only a few thousand species globally have been well classified (Janssen 2006). It is clear that much work remains to catalog and understand the role of this diversity in natural ecosystems (Handelsman 2004, Madsen 1998, Zak et al. 2006). Additionally, there is a lack of consensus as to how to define a bacterial species (Rosselló-Mora and Amann 2001) as well as evidence indicating that bacteria evolve quickly (Cohan 2001); both concerns complicate the understanding and assessment of diversity. 57 The Great Smoky Mountains National Park All Taxa Biodiversity Inventory: A Search for Species in Our Own Backyard 2007 Southeastern Naturalist Special Issue 1:57–72 58 Southeastern Naturalist Special Issue 1 There are two chief ways to assess bacterial diversity in natural samples: either by cultivating species using a growth medium followed by isolation of single species (Janssen et al. 2002, Joseph et al. 2003, Sait et al. 2002) or by using cultivation-independent techniques that detect species by characteristic DNA sequences, most often via PCR amplification of ribosomal genes (Dunbar et al. 2002, Hugenholtz et al. 1998, Janssen 2006). The benefit of culturing organisms is that one can measure various metabolic, biochemical, and physiological parameters for an organism once it is isolated from others in its community. This assessment of parameters can lead to better understanding of ecosystem functions performed by microorganisms (Madsen 1998). However, cultivation often selects for organisms that are rare or perhaps ecologically unimportant in the environments from which they were sampled (Hugenholtz et al. 1998, Janssen 2006). Methods which detect DNA from numerically abundant microorganisms should, theoretically, give a less-biased picture of diversity in a given environment (Janssen 2006). A drawback to these molecular techniques, however, is that they do not usually yield information about ecological functions carried out by these species. Recent advances have led to theoretical predictions of microbial roles in their environment via sequencing of unknown genes extracted from samples (Handelsman 2004, Schleper et al. 2005) and have provided the information necessary to cultivate unusual species (Könneke et al. 2005). Creative techniques to narrow the gap between the diversity observed in culture-independent and culturedependent approaches are leading to many discoveries of novel species heretofore known only by their DNA sequences from the environment (Ferrari et al. 2005, Joseph et al. 2003). More complete culture collections will allow microbial ecologists to elucidate the tasks performed by the majority of species in various aquatic and terrestrial environments. Due to the overwhelming number of individual cells and species of bacteria in most natural environments, it is necessary to take a step back to assess microbial diversity in macrobiotic ecosystems. Qualitative data can be gathered by examining bacterial taxa at a level above that of species. There are at least 52 established or proposed phyla in the domain Bacteria (Schloss and Handelsman 2004), but only a small number of these phyla are routinely encountered in culture-based studies (Jansen 2006). Instead, most of the bacterial phyla are detected using only direct DNA-sequence retrieval techniques (Dunbar et al. 2002, Jansen 2006). By first examining the microbial community at the phylum or immediately lower levels, predictive hypotheses may be generated about the relationships between terrestrial or aquatic environments and the microorganisms that drive their biogeochemistry. Such studies have been reported for soil (Barns et al. 1999, Fierer and Jackson 2006, Zhou et al. 2002), streams (Hullar et al. 2006), and marine ecosystems (Rappé and Giovannoni 2003). The purpose of this study was to initiate inventorying bacteria from soils and waters of GSMNP and included comparing cultured and uncultured species in three different forested habitats. 2007 S.P. O’Connell et al. 59 Methods Annual sampling of Oconaluftee and Kephart Prong sites for heterotrophic bacterial isolates Samples from soil and water were taken as part of the Western Carolina University Methods of General Microbiology (BIOL 414/514) class from two sites in GSMNP during the fall semesters from 2002–2005. The sites are near the Oconaluftee Visitor Center (UTM 17S 0291211, 3931770) and Kephart Prong trail (UTM 17S 0286057, 3940540), and samples included aseptically gathered soil and stream water. These locations were selected because of their proximity to the Western Carolina University campus and they serve as long-term study sites for examining bacterial distribution patterns. Individual soil samples were prepared by removing loose leaf litter and then homogenizing the upper 12–15 cm of soil using a propane torch-fl amed spade. A smaller, sterile trowel was then used to subsample the mixed soil to fill 50-mL sterile polypropylene centrifuge tubes. Stream-water samples were collected by immersing a sterile centrifuge tube in the water and filling to 80% volume. Each sample was collected upstream of the previous sample to minimize collecting sediments stirred up from the bottom. All samples were placed into a cooler on ice packs and returned to the lab where they were kept at 4 °C until culture work began within 5–7 days. Cultures were obtained by serially diluting environmental samples in sterile water and spread-plating the dilutions on R2A media plates. Plates were inverted, incubated in the dark at room temperature (≈25 °C), and assessed for growth after one week, at which time each student selected a colony for isolation and further study. Each colony was streaked for purity, and this process was repeated 4–6 times until a pure culture comprised of one species had been obtained. Students then characterized the isolate using growth-based, biochemical, and microscopic techniques before obtaining a portion of its ribosomal DNA sequence to identify it (n = 130 total isolates for the four years). DNA was obtained from the isolates by using a Mo Bio UltraClean Microbial DNA Isolation kit (Mo Bio, Inc., Solana Beach, CA). Polymerase chain reaction (PCR) conditions to amplify ca 550 bp of the 16S rDNA from each isolate consisted of the following: 50 μL total volume in nuclease- free water with final concentrations of 0.05% IgePal (Sigma, Inc., St. Louis, MO), 1.5 mM Mg2+, 1X Taq buffer (Promega, Inc. Madison, WI or Eppendorf, Inc., Westbury, NY), 0.25 μM of bacterial specific 341F and universal 907R primers (Operon, Inc., Huntsville, AL; Casamayor et al. 2000), 2.5 U of Taq polymerase (Promega or Eppendorf), 0.20–0.25 mM dNTPs (Promega or Eppendorf), and genomic DNA from the isolate. PCR was conducted using a “touchdown” approach (Casamayor et al. 2000) on a Mastercycler Personal thermal cycler (Eppendorf) with initial denaturation for 5 min at 94 °C, followed by 30 PCR cycles consisting of: 1 min 60 Southeastern Naturalist Special Issue 1 denaturation at 94 °C; 1 min annealing at decreasing temperature (beginning at 65 °C for two cycles, dropping 1 °C at each cycle for ten cycles, and ending at 55 °C for 18 cycles); 3 min elongation at 72 °C; and a final elongation for 7 min at 72 °C with a sample hold at 4 °C. Genomic DNA extracts and PCR products were screened in 1% agarose gels, and PCR products were cleaned using Montage spin filters (Millipore, Inc., Bedford, MA) prior to DNA sequencing (see below). Comparison of bacteria from Albright Grove, Cataloochee, and Purchase Knob sites Culturable bacteria. Soil samples were collected as above from three established All Taxa Biodiversity Inventory (ATBI) study plots: Albright Grove (old-growth forest in Tennessee), and Cataloochee and Purchase Knob (second-growth forests in North Carolina). These sites are on the eastern side of the GSMNP and were chosen because they have been classified as different forest types (Sharkey 2001) and it was hypothesized that microbial communities would refl ect the differences in vegetation. Samples were collected in 2002 (February), 2004 (September), and 2005 (February). Isolates were obtained on R2A medium for the 2002 samples, 10% tryptic soy agar (TSA) for the 2004 samples, and 0.1% nutrient broth (DDNB) for the 2005 samples. DDNB cultures were either plated from samples that had been frozen at -70° C or samples that had been diluted when fresh to 10-8 strength and incubated at room temperature (≈25 °C) for 3 months. All other isolates were obtained via spread-plating of diluted soil samples within a few days of sampling; DNA extractions and PCR conditions were implemented as described in the previous section. All data from isolates obtained from these four cultivation techniques were pooled (n = 81 total isolates) for comparison to clone libraries generated, as described next. Molecular clones. Paired soil samples were collected with the 2005 samples from the three ATBI plots described above, frozen in the field on dry ice, and stored in the lab at -70 °C. These samples were taken to obtain culture-independent 16S rDNA sequences from soil to compare to the sequences obtained by isolating species on the heterotrophic growth media. A Mo Bio PowerSoil DNA Isolation kit was used with the alternate lysis method and 1 min of bead beating at 2500 RPM (Mini BeadBeater, BioSpec, Inc., Bartlesville, OK) to obtain genomic DNA from the soil. Approximately 1500 base-pair fragments of the 16S rDNA from the mixed bacterial species were amplified using bacterial primers 27F and 1492R (Corinaldesi et al. 2005, Ferrari et al. 2005). Genomic DNA extracts were all diluted to 10% and added to a total volume of 50 μL containing the same PCR reagents and concentrations as for the isolates above. Thermal cycler conditions were as follows: 3 min of initial denaturation at 94 °C, 30 cycles of PCR consisting of 1 min of denaturation at 94 °C, 1 min of annealing at 55 °C, 2 min of elongation at 72 °C, a final elongation of 10 min at 72 °C, and a sample hold 2007 S.P. O’Connell et al. 61 at 4 °C. The resulting 1500 bp PCR products were cloned into Escherichia coli (JM109 strain, Promega) using the pGEM-T-Easy Vector System (Promega), and PCR inserts were reamplified by performing whole-cell PCR (O’Connell et al. 2003) using primers M13F and M13R. Clones that had the appropriate-sized inserts were then screened using restriction-fragment length polymorphism (RFLP) analysis with the restriction enzymes RsaI and MspI (Promega; RM Lehman, USDA, Brookings, SD, pers. comm.). A total of 91, 66, and 22 unique clones were obtained from Albright Grove, Cataloochee, and Purchase Knob, respectively, and sequenced (as described below). One hundred and fifty clones from each site were targeted, but due to cloning inefficiencies, we were only able to recover the above number of unique sequences; however, in only a handful of cases did we find the same RFLP pattern twice. DNA sequencing and DNA sequence analysis PCR using primers 341F/907R was run on the clone inserts from unique RFLP banding patterns to amplify ca 550 bp of the product (the same size and region of the 16S rDNA gene used for all of the isolates). All shortfragment 16S rDNA PCR products (isolates and clones) were then amplified in a sequencing PCR reaction using primer 907R with a BigDye Terminator Version 3.0 or 3.1 Cycle Sequencing Kit (Applied Biosystems, Foster City, CA). The PCR products were cleaned using AutoSeq Sephadex G-50 spin columns (Amersham Biosciences, Piscataway, NJ) and analyzed by either an ABI 377 gel sequencer or a 3130/3130xl automated capillary DNA sequencer (Applied Biosystems). Sequences were compared to sequences of clones and isolates contained in the Ribosomal Database Project II (RDP II) using both the “Classifier” and “Sequence Match” programs (Maidak et al. 2001). Clone sequences were additionally analyzed using the Greengenes suite of software including the Classifier program (DeSantis et al. 2006) to obtain additional phylogenetic inferences. Sequences were classified to the level of phylum, class (for the Proteobacteria), and genus. All sequences were checked for chimeras by using the Bellerophon program (Huber et al. 2004) and also by first aligning them with ClustalW using the Vector NTI program (Invitrogen, Inc., Carlsbad, CA) followed by Mallard software (Ashelford et al. 2006). Sequences called into question by these programs were screened more thoroughly using the Pintail computer program (Ashelford et al. 2005). No chimeric sequences were discovered. Results A total of eleven phyla were identified from riparian soil and water samples from the Kephart Prong and Oconaluftee sites and from the soils at the Albright Grove, Cataloochee, and Purchase Knob ATBI plots (Table 1). These included Actinobacteria, Firmicutes, Bacteroidetes, 62 Southeastern Naturalist Special Issue 1 Table 1. Bacterial genera grouped by phylum, candidate phylum, and class (for the Proteobacteria) and categorized by detection in culture collections or clone libraries and presence in soil or water for all sites and sampling methodologies (number after phylum or class shows how many sequences were analyzed; X = genus found in sample type; -- = genus not detected). Classifications are based on the best available sequence matches to the RDP II for cultured isolates and RDP II Classifier and Greengenes Classifier for the cloned samples. Genus Sample type Phylum Cultured Cloned Soil Water Actinobacteria (31) Acidimicrobineae -- X X -- Arthrobacter X -- X -- Curtobacterium X -- X -- Dermacoccus X -- X -- Kitasatospora X -- X -- Leifsonia X -- X -- Micrococcus X -- -- X Nocardia X -- X X Rhodococcus X -- X X Streptomyces X -- X -- Terrabacter X -- -- X Firmicutes (65) Anaeroglobus -- X X -- Bacillus X -- X X Paenibacillus X -- X -- Staphylococcus X -- X X Acidobacteria (132) Acidobacterium -- X X -- Verrucomicrobia (5) Verrucomicrobium -- X X -- Planctomyces (9) Isosphaera -- X X -- Planctomyces -- X X -- Gemmatimonadetes (1) Unclassified genus -- X X -- Bacteroidetes (40) Chitinophaga X X X -- Chryseobacterium X -- X -- Cytophaga X -- X -- Flavobacterium X -- -- X Flexibacter X -- X -- Pedobacter X -- X X Sphingobacterium X -- X X Taxeobacter X -- -- X Thermonema X -- -- X Proteobacteria (107) Alpha (24) Acidisphaera -- X X -- Agrobacterium X -- X -- Blastochloris -- X X -- Bradyrhizobium -- X X -- Inquilinus X -- X -- Magnetospirillum -- X X -- Methylosinus -- X X -- 2007 S.P. O’Connell et al. 63 Table 1, continued. Genus Sample type Phylum Cultured Cloned Soil Water Odyssella -- X X -- Phenylobacterium -- X X -- Rhizobium X -- X -- Rhodoplanes -- X X -- Roseomonas -- X X -- Sphingomonas X -- -- X Tistrella -- X X -- Beta (41) Acidovorax X -- -- X Aquaspirillum X -- X -- Burkholderia X X X -- Collimonas X -- X -- Caenibacterium -- X X -- Herbaspirillum X -- X -- Janthinobacterium X -- -- X Tepidiphilus -- X X -- Variovorax X -- -- X Zoogloea X -- X X Delta (1) Desulfomonile -- X X -- Gamma (41) Alkalispirillum -- X X -- Buttiauxella X -- -- X Enterobacter X -- -- X Frateuria X -- X -- Isochromatium -- X X -- Klebsiella X -- X -- Pantoea X -- -- X Pseudomonas X -- X X Rhodanobacter X -- X -- Rickettsiella -- X X -- Serratia X -- X X Thiorhodospira -- X X -- Xanthomonas X -- X -- Yersinia X -- -- X Deinococcus (1) Deinococcus X -- -- X Termite Group 1 (1) -- X X -- OP10 (2) -- X X -- Proteobacteria, Deinococcus-Thermus, Acidobacteria, Verrucomicrobia, Planctomyces and Gemmatimonadetes. The first two phyla are the grampositive bacteria, and the others are representatives of gram-negative bacteria. Candidate phyla OP10 and Termite Group 1 were also detected in the clone libraries from the ATBI sites. Because Proteobacteria is such a large phylum and contains a large number of metabolically and genetically diverse species, this phylum is broken down into four classes for further comparison, including Alpha-, Beta-, Delta-, and Gammaproteobacteria. While we did not recover enough clones and isolates from the ATBI plots 64 Southeastern Naturalist Special Issue 1 to make discrete comparisons between our study sites, some general trends are reported below. For the purpose of discussion, we have also combined data from waters at Oconaluftee and Kephart Prong and soils from these sites and will treat the results as a master list of genera detected in GSMNP to date (Table 1). The eleven phyla recovered included DNA sequence data from 390 organisms (211 bacterial isolates and 179 clones), which can be placed into 69 genera (Table 1). Of the 390 sequences, 324 originated from soil and 66 from water samples. At the level of phylum, molecular cloning detected representatives from ten phyla and the four classes of Proteobacteria, while culturing techniques yielded representatives from five phyla and three of the proteobacterial classes. The latter also produced the only record of a Deinococcus-Thermus representative. Cultures and clones accounted for 45 and 26 genera respectively, with only two genera, Chitinophaga (Bacteroidetes) and Burkholderia (Betaproteobacteria) found using both methods. Forty-six genera were found exclusively in soil samples, while waters produced 14 unique genera; nine genera overlapped between the two sample types. With the relatively small number of sequences obtained from water samples and the lack of clones from these samples, we cannot make strong conclusions from these data. However, it is worth noting again the number of unique genera from these samples and the occurrence of Deinococcus, the sole representative of its phylum we found. Additionally, waters were dominated by species from Bacteroidetes (34.8% of sequences) and Betaproteobacteria (22.7% of sequences), while soils had more cultured representatives from Firmicutes and Gammaproteobacteria, with 37.5% and 21.9% of sequences identified to these phyla, respectively, from the Kephart Prong and Oconaluftee sites. Six phyla and Deltaproteobacteria were represented only in the clone libraries. Of these, only Acidobacteria (132 clones), Planctomyces (9 clones), and Verrucomicrobia (5 clones) produced more than two clones (Table 1). For the other groups, only Alphaproteobacteria were more commonly cloned than cultured (14 clones versus 10 isolates), and the highest number of clones for any other group was six for Gammaproteobacteria. The number of genera per phylum reveals some other interesting trends including that the most sequence-rich group detected, Acidobacteria, was represented by only a single genus. Alphaproteobacteria, with 24 sequences, accounted for 14 genera and included the most cloned genera (10). Gammaproteobacteria records show 14 genera as well, but with 41 sequences, and most of these originating from cultures. Ten betaproteobacterial genera were detected from 41 sequences and most were from isolated bacteria (eight versus three genera from clones). Actinobacteria and Bacteroidetes were similar in being represented largely by cultures (ten and nine genera, respectively), with only one genus detected each in clone libraries. Lastly, Firmicutes was relatively genera-poor (four genera) but sequence-rich with 65 records, with all but one sample obtained from culturing techniques. 2007 S.P. O’Connell et al. 65 Table 2 shows the differences in sequence detection of cultures and clones from the three ATBI plots. Interestingly, there is almost no consensus between phyla and proteobacterial classes recovered using the two means of assessing diversity. Only in the case of Proteobacteria does there appear to be some similarity at this higher level of taxonomy, but as can be seen in Table 1, little overlap actually exists toward the level of species. Isolates from the soils of Albright Grove, Cataloochee, and Purchase Knob largely fall into one of three groups: Firmicutes, Betaproteobacteria, or Actinobacteria. Firmicutes accounted for 44.4% of the sequences, Betaproteobacteria 21%, and Actinobacteria 12.3%. Acidobacteria overshadowed all other sequence data and represented 73.7% of the soil clones, with Alphaproteobacteria (8.4%) and Planctomyces (5%) of secondary abundance. Discussion Little to no overlap was seen between the bacteria inhabiting streams and soils (Table 1) or between methods assessing diversity from three forests of different histories (Table 2). The data in this study have been reported at the level of genus rather than species; however, in only a few cases (less than 3% of all samples) have we observed the same DNA sequence on more than one occasion. A species definition for bacteria has not been unanimously established, but ideally a combination of DNA sequence and ecological similarity are criteria for taxonomic placement (Rosselló-Mora and Amann 2001). In the instances where we have retrieved identical 16S rDNA sequences (from Table 2. Percent of diversity attributed to each bacterial phylum, candidate phyla (Termite Group 1 and OP10), and class (for the Proteobacteria) broken down by the approach taken to obtain 16S rDNA sequences from soils from the Albright Grove, Cataloochee, and Purchase Knob ATBI study plots (data are combined for the three sites). Classifications are based on the best available sequence matches to the RDP II database for cultured isolates (n = 81 isolates) and RDP II Classifier and Greengenes Classifier for the cloned samples (n = 179 clones). Phylum Cultured Cloned Actinobacteria 12.3% 0.6% Firmicutes 44.4% 0.6% Acidobacteria - 73.7% Verrucomicrobia - 2.8% Planctomyces - 5.0% Gemmatimonadetes - 0.6% Bacteroidetes 9.9% 1.1% Proteobacteria 33.3% 14.0% Alphaproteobacteria 3.7% 8.4% Betaproteobacteria 21.0% 1.7% Deltaproteobacteria - 0.6% Gammaproteobacteria 8.6% 3.4% Termite Group 1 - 0.6% OP10 - 1.1% 66 Southeastern Naturalist Special Issue 1 isolated bacteria), we have been able to measure some other characteristic that could differentiate the two organisms (e.g., metabolic trait or other growth parameter). What our data suggest is that even from a few small samples from GSMNP, we are only beginning to be able to document the bacterial diversity therein. Further complicating matters is that many of the species we have detected overlap little in 16S rDNA sequence to their closest relatives in the sequence databases that are available for comparison. This diversity of bacteria in natural environments is not surprising as others have shown similar results, especially from soils (Curtis et al. 2002, Gans et al. 2005, Hong et al. 2006). Understanding the meaning of this diversity is the challenge. Soils are incredibly complex at multiple scales (Fierer and Jackson 2006, Zhou et al. 2002) and infl uence and are infl uenced by the plant and microbial communities that they are associated with (Dunbar et al. 2002, Grayston and Prescott 2005, Hackl et al. 2004, Smalla et al. 2001). Floyd et al. (2005) reported on the most easily cultured bacteria from soils as contained in the American Type Culture Collection (ATCC; Manassas, VA), and their summary shows that the gram-positive bacteria are dominant in soils, with Actinobacteria accounting for over 35% of all isolated type strains and Firmicutes making up another 12.4%. In the former phylum, the genus Streptomyces is an important source of antibiotics and therefore may be intentionally over-sampled from soil, while the latter group contains spore-forming bacteria that may also be over-represented in culture. This possibility has been suggested by other work (Janssen 2006) and also likely holds true in our study. Firmicutes were dominant in the cultures from soil at Kephart Prong and Oconaluftee as well as at Albright Grove, Cataloochee, and Purchase Knob (Tables 1 and 2). Actinobacteria were present in the same samples, but not as commonly cultivated since we did not specifically target this phylum. Proteobacteria comprised nearly 10% of the records in the ATCC, while the Bacteroidetes accounted for 1.2% (only genera with greater than 1% representation in the collection were reported). Proteobacteria and Bacteroidetes were likewise commonly cultured in GSMNP, but at greater levels (Tables 1 and 2). Hugenholtz et al. (1998) analyzed over 8000 16S rDNA isolate and clone sequences in public databases and estimated that 65% of all Proteobacteria records were of cultured species, 80% each of Actinobacteria and Firmicutes were represented by cultures, and 40% of Bacteroidetes entries were from cultures. This contrasted with only 1% of the Acidobacteria and 10% of Verrucomicrobia records from culture. The other records came from clone libraries. In our study, the majority of soil bacteria have been assigned to the phylum Acidobacteria (Table 2). This is not unusual, since Acidobacteria have been shown to be dominant or co-dominant in many environments and this group also shows great taxonomic diversity, although they are represented by few genera (Barns et al. 1999, Dunbar et al. 2002, Hugenholtz et 2007 S.P. O’Connell et al. 67 al. 1998, Janssen 2006). What is surprising is how large a proportion of our clone library this group comprises (73.7% of all clones; Table 2). Others have shown Acidobacteria accounting for 40–50% of the clones from arid soils (Dunbar et al. 2002) and 20% of the diversity of soils globally (Janssen 2006). It remains to be seen what roles the species in this phylum play, since so few have been cultured; however, their sheer number and phylogenetic breadth suggest their importance in soil ecosystems (Janssen 2006). Other phyla commonly encountered in clone libraries from soil include Proteobacteria, Actinobacteria, Verrucomicrobia, and Bacteroidetes, with other groups such as Planctomyces and Firmicutes less common (Dunbar et al. 2002 and Janssen 2006). Every soil sample reported by Janssen (2006) contained representatives of the Alpha-, Beta-, and Gammaproteobacteria, and the proteobacterial classes made up a mean of 40% of all soil clones worldwide. Proteobacteria were commonly cloned in GSMNP sites as well, comprising 14% of the clones from Albright Grove, Cataloochee, and Purchase Knob (Table 2). Alphaproteobacteria were more abundant in the clone libraries than in cultures and represent the group with the most genera detected (Table 1), none of which are refl ected in our culture collection and bear more investigation. Unlike in the culture collection where Firmicutes accounted for a large proportion of all DNA sequences obtained, this group was represented by only a single clone. This is interesting since at Albright Grove, this phylum accounted for nearly 80% of all isolates. The species within Firmicutes are likely to be readily cultured from numerous spores. However, owing to their relatively massive cell walls and spores, these species (and Actinobacteria) may resist DNA extraction efforts and therefore, may not be detected via molecular cloning (Janssen 2006). Although we tried to account for as much diversity as possible in our molecular cloning efforts, e.g., using bead beating as part of the DNA extraction step (Miller et al. 1999), we cannot completely rule out inefficiencies in our DNA recovery methods. Other groups that were readily cultured but not well-represented in the clone libraries included Actinobacteria and Bacteroidetes (Table 2), perhaps suggesting their relative scarcity or unimportance in soil ecosystems, since molecular cloning should refl ect numerically abundant organisms (Actinobacteria as discussed above) and gram-negative species such as those in Bacteroidetes (Janssen 2006). Lastly, Planctomyces and Verrucomicrobia represent phyla found globally most often in clone libraries. This was the case in our work as well, with the former producing nine clones, and the latter group producing five unique DNA sequences from soil. The large gap between organisms which are readily cultivated and those dominating in clone libraries has been narrowing somewhat thanks to creative culturing efforts. These either mimic in situ conditions in the lab or field or use dilution and long incubation times to select for oligotrophs, which are the norm in nature (Ferrari et al. 2005, Kaeberlein 68 Southeastern Naturalist Special Issue 1 et al. 2002, Zengler et al. 2002). Recent literature has shown improvements for culturing Acidobacteria (Davis et al. 2005, Sait et al. 2006, Stevenson et al. 2004), Verrucomicrobia (Davis et al. 2005, Janssen et al. 2002, Joseph et al. 2003, Stevenson et al. 2004), Planctomyces (Wang et al. 2002), and Acidobacteria, Actinobacteria, Alpha-, Beta-, and Gammaproteobacteria simultaneously (Janssen et al. 2002, Joseph et al. 2003, Sait et al. 2002). Our goal is to employ similar methods in an attempt to culture the many species represented by the clones in our libraries, which are numerous in the native soil and likely ecologically important. Once we have these species in culture, their roles in the environment may be more readily elucidated (e.g., specific nutrient turnover processes, symbioses, pathogenic interactions, etc.). Potential ecological functions these species participate in, other than in heterotrophic processes (all of the isolates and many of the clones likely act as decomposers), include a huge range of physiologies in the Proteobacteria. These include the chemolithotrophs (methane, sulfur, iron, manganese, hydrogen, ammonia, and nitrate oxidizers), nitrogen-fixers, sulfur-reducers, and bacterial predators (e.g., Bdellovibrio; Perry et al. 2002, Madigan and Martinko 2006). Bacteroidetes include animal gut symbionts and complex polymer degraders and likely play a role in soil biogeochemistry. Actinobacteria are producers of antibiotics, which may regulate population sizes of bacteria and fungi in soils. This phylum also contains nitrogen-fixers. Some Firmicutes are sulfur reducers, and cultured representatives of Verrucomicrobia act as protist symbionts, protecting their host from protist predators. Acidobacteria and Planctomyces certainly have roles as heterotrophs, but the great taxonomic diversity within Acidobacteria, which rivals the betterstudied Proteobacteria, suggests many niches yet to be discovered by microbial ecologists (Janssen 2006). This preliminary inventory of bacteria inhabiting the soils and waters of GSMNP sets the stage for future investigations. The main finding of this work is that different methodologies produced significantly different views of bacterial diversity, even when examining the same samples. Other than for the cyanobacteria (Johansen 2007), no bacterial inventories have been undertaken in the GSMNP. We plan to continue to examine the three forested ecosystems represented in this study by cultures and clones. The long-term study plots established for soils and waters at Kephart Prong and Oconaluftee will be sampled each fall for many years to come to document species distribution patterns over time. Additionally, clone libraries should be generated for these soil and water samples to further elucidate the patterns of and participants in bacterial diversity in the GSMNP. Estimating the total number of bacterial species in GSMNP is improbable using conventional models, especially considering recent estimates that place the number of species in a few grams of soil in the millions (Curtis et al. 2002, Gans et al. 2005). However, with repeated sampling of the same sites, sampling 2007 S.P. O’Connell et al. 69 multiple sites, using standardized methods, and combining data from culture collections and clone libraries, a greater proportion of the total species pool can be detected. Some have called for more intensive cloning of environmental samples to fill out the bacterial phylogenetic tree (Curtis 2006); this would serve to especially detect the numerous and culture-resistant groups and would be a good use of resources. Based on the clone libraries we have generated from GSMNP, we believe the Acidobacteria to be quite important in the forest soil. Understanding this group, in which only a few members have been cultivated, would go a long way to aid in discovering important processes occurring in the GSMNP as well as globally. Acknowledgments The authors would like to acknowledge Western Carolina University and Discover Life in America for funding this work, and Keith Langdon, Paul Super (National Park Service), and Jeanie Hilten (DLIA) for logistical support and advice. Most of the DNA sequences were generated using support from a grant from the National Science Foundation (DBI-0521334). We would also like to acknowledge the 132 students from eight sections of the BIOL 414/514 classes who isolated and characterized bacteria from Kephart Prong and Oconaluftee, and Philip Drummond for invaluable lab assistance. Lastly, the authors thank three reviewers whose comments greatly improved the quality and scope of this manuscript. Literature Cited Ashelford, K.E., N.A. Chuzhanova, J.C. Fry, A.J. Jones, and A.J. Weightman. 2005. At least 1 in 20 16S rRNA sequence records currently held in public repositories is estimated to contain substantial anomalies. Applied and Environmental Microbiology 71:7724–7734. Ashelford, K.E., N.A. Chuzhanova, J.C. Fry, A.J. Jones, and A.J. Weightman. 2006. New screening software shows that most recent large 16S rRNA gene clone libraries contain chimeras. Applied and Environmental Microbiology 72:5734–5741. Barns, S.M., S.L. Takala, and C.R. Kuske. 1999. Wide distribution and diversity of members of the kingdom Acidobacterium in the environment. Applied and Environmental Microbiology 65:1731–1737. Casamayor, E.O., H. Schäfer, L. Bãneras, C. Pedrós-Alió, and G. Muyzer. 2000. Identification of and spatio-temporal differences between microbial assemblages from two neighboring sulfurous lakes: comparison by microscopy and denaturing gradient gel electrophoresis. Applied and Environmental Microbiology 66:499–508. Cohan, F.M. 2001. Bacterial species and speciation. Systems Biology 50:513–524. Corinaldesi, C., R. Danovaro, and A. Dell’Anno. 2005. Simultaneous recovery of extracellular and intracellular DNA suitable for molecular studies or studies from marine sediments. Applied and Environmental Microbiology 71:46–50. Curtis, T.P., W.T. Sloan, and J.W. Scannell. 2002. Estimating prokaryotic diversity and its limits. Proceedings of the National Academy of Sciences 99:10494–10499. Curtis, T. 2006. Microbial ecologists: It’s time to “go large.” Nature Reviews Mi70 Southeastern Naturalist Special Issue 1 crobiology 4:488. Davis, K.E.R., S.J. Joseph, and P.H. Janssen. 2005. Effects of growth medium, inoculum size, and incubation time on culturability and isolation of soil bacteria. Applied and Environmental Microbiology 71:826–834. DeLong, E.F., and N.R. Pace. 2001. Environmental diversity of bacteria and archaea. Systems Biology 50:470–478. DeSantis, T.Z., P. Hugenholtz, N. Larsen, M. Rojas, E.L. Brodie, K. Keller, T. Huber, D. Dalevi, P. Hu, and G.L. Andersen. 2006. Greengenes, a chimera-checked 16S rRNA gene database and workbench compatible with ARB. Applied and Environmental Microbiology 72:5069–5072. Dunbar, J., S.M. Barns, L.O. Ticknor, and C.R. Kuske. 2002. Empirical and theoretical bacterial diversity in four Arizona soils. Applied and Environmental Microbiology 68:3035–3045. Ferrari, B.C., S.J. Binnerup, and M. Gillings. 2005. Microcolony cultivation on a soil substrate membrane system selects for previously uncultured soil bacteria. Applied and Environmental Microbiology 71:8714–8720. Fierer, N., and R.B. Jackson. 2006. The diversity and biogeography of soil bacterial communities. Proceedings of the National Academy of Sciences 103:626–631. Floyd, M.M., J. Tang, M. Kane, and D. Emerson. 2005. Captured diversity in a culture collection: Case study of the geographic and habitat distributions of environmental isolates held at the American Type Culture Collection. Applied and Environmental Microbiology 71:2813–2823. Gans, J., M. Wolinsky, and J. Dunbar. 2005. Computational improvements reveal great bacterial diversity and high metal toxicity in soil. Science 309:1387– 1390. Grayston, S.J., and C.E. Prescott. 2005. Microbial communities in forest fl oors under four tree species in coastal British Columbia. Soil Biology and Biochemistry 37:1157–1167. Hackl, E., S. Zechmeister-Boltensern, L. Bodrossy, and A. Sessitsch. 2004. Comparison of diversities and compositions of bacterial populations inhabiting natural forest soils. Applied and Environmental Microbiology 70:5057–5065. Handelsman, J. 2004. Metagenomics: Application of genomes to uncultured microorganisms. Microbiology and Molecular Biology Reviews 68:669–685. Hong, S-H., J. Bunge, S-O. Jeon, and S.S. Epstein. 2006. Predicting microbial species richness. Proceedings of the National Academy of Sciences 103:117–122. Huber, T., G. Faulkner, and P. Hugenholtz. 2004. Bellerophon: A program to detect chimeric sequences in multiple sequence alignments. Bioinformatics 20:2317– 2319. Hugenholtz, P., B.M. Goebel, and N.R. Pace. 1998. Impact of culture-independent studies on the emerging phylogenetic view of bacterial diversity. Journal of Bacteriology 180:4765–4774. Hughes, J.B., J.J. Hellmann, T.H. Ricketts, and B.J.M. Bohannan. 2001. Counting the uncountable: Statistical approaches to estimating microbial diversity. Applied and Environmental Microbiology 67:4399–4406. Hullar, M.A.J., L.A. Kaplan, and D.A. Stahl. 2006. Recurring seasonal dynamics of microbial communities in stream habitats. Applied and Environmental Microbiology 72:713–722. Janssen, P.H. 2006. Identifying the dominant soil bacterial taxa in libraries of 2007 S.P. O’Connell et al. 71 16S rDNA and 16S rRNA genes. Applied and Environmental Microbiology 72:1719–1728. Janssen, P.H., P.S. Yates, B.E. Grinton, P.M. Taylor, and M. Sait. 2002. Improved culturability of soil bacteria and isolation in pure culture of novel members of the divisions Acidobacteria, Actinobacteria, Proteobacteria, and Verrucomicrobia. Applied and Environmental Microbiology 68:2391–2396. Joseph, S.J., P. Hugenholtz, P. Sangwan, C.A. Osborne, and P.H. Janssen. 2003. Laboratory cultivation of widespread and previously uncultured soil bacteria. Applied and Environmental Microbiology 69:7210–7215. Kaeberlein, T., K. Lewis, and S.S. Epstein. 2002. Isolating “uncultivable” microorganisms in pure culture in a simulated natural environment. Science 296:1127– 1129. Könneke, M., A.E. Bernhard, J.R. de la Torre, C.B. Walker, J.B. Waterbury, and D.A. Stahl. 2005. Isolation of an autotrophic ammonia-oxidizing marine archaeon. Nature 437:543–546. Madigan, M.T., and J.M. Martinko. 2006. Brock Biology of Microorganisms, 11th Edition. Prentice-Hall, Inc., Upper Saddle River, NJ. Madsen, E.L. 1998. Epistemology of environmental microbiology. Environmental Science and Technology 32:429–439. Maidak, B.L., J.R. Cole, T.G. Lilburn, C.T. Parker, Jr., P.R. Saxman, R.J. Farris, G.M. Garrity, G.J. Olsen, T.M. Schmidt, and J.M. Tiedje. 2001. The RDP-II (Ribosomal Database Project). Nucleic Acids Research 29:173–174. Miller, D.N, J.E. Bryant, E.L. Madsen, and W.C. Ghiorse. 1999. Evaluation and optimization of DNA extraction and purification procedures for soil and sediment samples. Applied and Environmental Microbiology 65:4715–4724. O’Connell, S.P., R.M. Lehman, O. Snoeyenbos-West, V.D. Winston, D.E. Cummings, M.E. Watwood, and F.S. Colwell. 2003. Detection of Euryarchaeota and Crenarchaeota in an oxic basalt aquifer. FEMS Microbiology Ecology 44:165–173. Perry, J.J., J.T. Staley, and S. Lory. 2002. Microbial Life. Sinauer Associates, Sunderland, MA. Rappé, M.S., and S.J. Giovannoni. 2003. The uncultured microbial majority. Annual Review of Microbiology 57:369–394. Rosselló-Mora, R., and R. Amann. 2001. The species concept for prokaryotes. FEMS Microbiology Reviews 25:39–67. Sait, M., P. Hugenholtz, and P.H. Janssen. 2002. Cultivation of globally distributed soil bacteria from phylogenetic lineages previously only detected in cultivationindependent surveys. Environmental Microbiology 4:654–666. Sait, M., K.E.R. Davis, and P.H. Janssen. 2006. Effect of pH on the isolation and distribution of members of Subdivision 1 of the phylum Acidobacteria occurring in soil. Applied and Environmental Microbiology 72:1852–1857. Schleper, C., G. Jurgens, and M. Jonuscheit. 2005. Genomic studies of uncultivated archaea. Nature Reviews Microbiology 3:479–488. Schloss, P.D., and J. Handelsman. 2004. Status of the microbial census. Microbiology and Molecular Biology Reviews 68:686–691. Sharkey, M.J. 2001. The all taxa biological inventory of the Great Smoky Mountains National Park. Florida Entomologist 84:556–564. Smalla, K., G. Wieland, A. Buchner, A. Zock, J. Parzy, S. Kaiser, N. Roskot, H. Heuer, and G. Berg. 2001. Bulk and rhizosphere soil bacterial communities studied 72 Southeastern Naturalist Special Issue 1 by denaturing gradient gel electrophoresis: Plant-dependent enrichment and seasonal shifts revealed. Applied and Environmental Microbiology 67:4742–4751. Stevenson, B.S., S.A. Eichorst, J.T. Wertz, T.M. Schmidt, and J.A. Breznak. 2004. New strategies for cultivation and detection of previously uncultured microbes. Applied and Environmental Microbiology 70:4748–4755. Torsvik, V., J. Goksoyr, and F.L. Daae. 1990. High diversity in DNA of soil bacteria. Applied and Environmental Microbiology 56:782–787. Wang, J., C. Jenkins, R.I. Webb, and J.A. Fuerst. 2002. Isolation of Gemmata-like and Isophaera-like plancomycete bacteria from soil and freshwater. Applied and Environmental Microbiology 68:417–422. Whitman, W.B., D.C. Coleman, and W.J. Wiebe. 1998. Prokaryotes: The unseen majority. Proceedings of the National Academy of Sciences 95:6578–6583. Zak, R.Z., C.B. Blackwood, and M.P. Waldrop. 2006. A molecular dawn for biogeochemistry. TRENDS in Ecology and Evolution 21:288–295. Zengler, K., G. Toledo, M. Rappé, J. Elkins, E.J. Mathur, J.M. Short, and M. Keller. 2002. Cultivating the uncultured. Proceedings of the National Academy of Sciences 99:15681–15686. Zhou, J., B. Xia, D.S. Treves, L.-Y. Wu, T.L. Marsh, R.V. O’Neill, A.V. Palumbo, and J.M. Tiedje. 2002. Spatial and resource factors infl uencing high microbial diversity in soil. Applied and Environmental Microbiology 68:326–334.