Investigating Bacterial Communities in a Warm, Alkaline Pool in the Sanford Underground Research Facility Using Culture-Based and Culture-Independent Methods

Proceedings of the South Dakota Academy of Science, Vol. 97 (2018) 129

INVESTIGATING BACTERIAL COMMUNITIES IN A WARM, ALKALINE POOL IN THE SANFORD UNDERGROUND RESEARCH FACILITY USING CULTURE-BASED AND CULTURE-INDEPENDENT METHODS

Taylor Liu1 and David J. Bergmann2* 1University of California Berkeley, CA 94720 2Black Hills State University Spearfish, SD 57799 *Corresponding author e-mail: [email protected]

ABSTRACT

We examined the bacterial communities in a pool of water under a flowing rock fracture at a depth of 1470 m in the Sanford Underground Research Facility in Lead, SD. High-throughput sequencing of 16S rDNA indicated that Acidobacteria, Chlorobi, Nitrospirae, Planctomycetes, and Gammaproteobacteria (especially Acidiferribacter) were abundant in pool sediments; while Nitrospirae, Plactomycetes, and Proteobacteria (especially Thiobacillus and Methylococcus) were abundant in pool water. Samples of diluted pool water were mixed with agar media, placed in diffusion chambers surrounded by 0.2 μm pore membranes, and incubated in situ within the pool for 19 days. 16S rDNA analysis of media from diffusion chambers indicated Alphaproteobacteria (including Hyphomicrobium), Betaproteobacteria (including Inhella and Methyloversatilis), and Gammaproteobacteria (mainly Pseudomonas) were dominant. Low-nutrient media were used to isolate bacteria from both pool water and media inside diffusion chambers. The majority of these isolates were Methyloversatilis, Inhella, and Bacillus. Although the genera isolated were not very abundant in the pool water, the presence of Methylococcus and Methyloversatilis suggests the importance of methane and methanol oxidation in this community.

Keywords

Bacteria, deep subsurface biome, SURF

INTRODUCTION

Deep underground rocks and aquifers extending over 1500 m below the earth’s surface constitute a massive region of the biosphere and may contain more prokaryotic cells than all other habitats (Madigan and Martinko 2006). For example, dense communities of Bacteria and Archaea have been found in aquifers 1500 m 130 Proceedings of the South Dakota Academy of Science, Vol. 97 (2018) deep in basalt, where microbial metabolism involving methanogenesis, acetogen- esis, and sulfate respiration are important (Stevens et al. 1995; Anderson et al. 1998). The Sanford Underground Research Facility (SURF), located in Lead, South Dakota, is one of few sites where deep subsurface microbial environments can directly accessed. SURF is located in the former Homestake gold mine, compris- ing over 590 km of tunnels and extending to a depth of 2460 m. Between the cessation of gold mining in 2003 and the opening of SURF (then known as the Deep Underground Science and Engineering Laboratory) in 2007, pumping of the water accumulating in the lower tunnels of the site was discontinued, and the lower levels of the site (including the 1470 m level) became flooded. At present, water is pumped out to depth of about 1700 m, and the 1470 level is now the site of a number of major physics experiments (Lesko 2011; Sanfordlab). Sediments from the 1470 m level of SURF are rich in chlorite-chamosite [(Fe5Al)(AlSi3)

O10(OH)8], annite [KFe3AISi3O10(OH)2], and quartz (SiO2) (Rastogi et al. 2010). The tunnels in SURF provide an environment where air in the tunnels comes in contact with anaerobic fracture water from deep aquifers, making chemoau- trophy and methanotrophy possible. Sediments and drainage water in SURF have been found to contain diverse assemblages of prokaryotes, including ammonia-oxidizing Archaea, sulfide-oxidizing bacteria, and various chemo- heterotrophic bacteria (Waddell et al 2010; Rastogi et al. 2009; Rastogi et al. 2010). Furthermore, SURF contains a wide variety of biofilms, often sustained by fracture water seepage of different redox potentials and chemical composition, supporting diverse microbial communities (Osburn et al. 2014). One difficulty in studying microbial communities is that often less than 1% of microbial species can be cultured on standard media. The cultured species may be minor constituents of their communities, but may be easily isolated because of their rapid growth and their lack of specialized requirements for organic cofactors (Epstein 2013). One way to investigate the taxonomic diversity of microbial communities is to use culture-independent techniques involving extraction of microbial DNA, PCR amplification of the 16S rRNA genes, and sequencing PCR amplicons (Madigan et al. 2006). For example, using next-generation 16S rDNA sequencing methods, Osburn et al. (2014) found that samples of water sand biofilms from SURF contained from 0.3 percent to 19% taxa from candi- date phyla such as BRC1, M7, and W53, none of which have been isolated in culture. Despite the difficulties encountered in culturing microbes from environmental samples, it is often desirable to isolate microbes in culture so their phenotypic characteristics can be fully characterized. Recently, the use of diffusion chambers, in which environmental samples are initially incubated in situ within small volumes of media surrounded by permeable membranes (diffusion chambers), has enabled a greater diversity of bacterial groups to be isolated in culture from some habitats (Bollmann et al 2006), especially slow-growing bacteria or bacteria requiring certain organic cofactors. During an exploration of the 17 Ledge region of the 1470 m level of SURF, we found an extensive area of microbial biofilms where deep aquifer water from Proceedings of the South Dakota Academy of Science, Vol. 97 (2018) 131 a fracture in the rock flows into an old mine tunnel, forming extensive pools of water. This site was located about 324 m from the ramp to the 1410 m level, about 105 m beyond a flowing borehole known as Thiothrix“ Falls,” and is near the farthest portion of the tunnel which can be safely visited. The pools of water at the site had a temperature about 32 °C, and a pH about 8.8. Because of the unusual physical conditions at the site, we decided to conduct a detailed inves- tigation of the microbial communities in the pool of water, which might harbor novel, uncharacterized taxa of Bacteria and Archaea. In this study, we investigate the bacterial communities in this warm, alkaline pool of water on the 1470 m level of SURF using a variety of techniques: 1) culture-independent analysis of bacterial 16S rDNA, 2) direct culturing on a standard bacteriological medium, and 3) culturing of bacteria using in situ incubation of samples in low-nutrient media in diffusion chambers. Analysis of 16S rDNA sequences will allow us to directly evaluate the taxonomic diversity of a possibly unique microbial community. By using the two culture-based techniques of direct isolation and isolation from diffusion chambers, we hope to isolate and culture microbes, including some of those groups not readily cultured by standard bacteriological techniques, for phenotypic analysis and future study.

METHODS

Location and Sampling. The pool of water sampled in this study is located on the 17 Ledge of the 1470 m level of SURF about 150 m beyond a flowing borehole known as “Thiothrix Falls” which is presently capped by a manifold as part of a NASA biology experiment. The pool is about 1 m wide, a few m long, and 10 cm deep, and is fed by water flowing from a fracture in the rock. The rock in the area consists of the Poorman formation with metamorphic rocks, such as phyllite, predominating (Caddey et al. 1991). For some chemical analyses, unfiltered samples of water were collected with a glass beaker. For other chemical analyses, filtered samples were collected with sterilized #15 silicon tubing (Core-Parmer Instruments Co., IL) connected to a Masterflex E/S Portable Sampler peristaltic pump (Core-Parmer Instruments) with a Sterivex GS 0.2 μm pore cartridge filter (Millipore). Two filter cartridges were used; the first filtering about 3.9 L and the second 2.8 L. Filtered water was collected in 250 ml polyethylene bottles and 40 ml glass vials. Filtered and unfiltered samples were preserved as directed by Mid-Continent Testing Laboratories Rapid City, SD, which conducted chemical analyses within two days of collection. The filter cartridges, containing microbial cells from the pool water, were frozen in dry ice. Samples of sediment from the pool were collected in 50 mL Falcon tubes and frozen on dry ice. Filters and sediment samples were stored at -80 °C until microbial DNA was extracted (see 16S rDNA Library Preparation).

Diffusion Chambers.Direct isolation of microbes from the pool water was performed by collecting a water sample in a sterile Falcon tube and taking it to 132 Proceedings of the South Dakota Academy of Science, Vol. 97 (2018) the Black Hills State University Underground Campus cleanroom (BHUC) on the 1470 m level of SURF. The sample was diluted from 10-1 to 10-5 in filtered, autoclaved SURF drainage water (pH 8.5) from the 1470 m level of SURF. Diluted samples were spread across petri plates of 1/10X R2B medium (Reasoner and Geldrich 1985) with 1.5% agar, with 1% ATCC vitamin extract (American Type Culture Collection), and fungicides (100 μg/mL cycloheximide, and 50 μg/ mL Nystatin (Sigma-Aldrich). The plates were then returned to the surface and incubated at 30 °C for two weeks at the Life Sciences building at Black Hills State University, Spearfish, SD. Diffusion chambers forin situ culturing were constructed of sterilized 27 mm (inner diameter) steel washers, 2 mm thick, with 47 mm diameter, 0.2 μm pore size polycarbonate IsoporeTM membrane filters (Millipore, Inc.) covering the upper and lower sides. The lower filter was attached with silicone adhesive, cov- ered in foil, and taken to the pool in SURF. A sample of water was taken from the pool and processed in the BHUC cleanroom. Samples were diluted in filtered, autoclaved SURF drainage water with 1/10X R2B medium. The water samples were diluted to a final concentration of 10-3 in 1/50X R2B medium with 1.0% agar (50 °C), with 1% vitamin extract, and fungicides. About 1.8 mL of diluted sample in media was added to each of five diffusion chambers in a laminar flow hood, and the top filter membrane was glued in place. A sixth diffusion chamber, a negative control, contained sterile media only. The diffusion chambers were taken immediately to the pool on the 17 ledge, placed sideways with one end embedded in sediment and membranes exposed to water, and left in place for 19 days. After 19 days, diffusion chambers were retrieved and taken to the surface. The negative control chamber was opened in a laminar flow hood and half of the contents used for DNA extraction with a PowerSoil kit and the other half homogenized in 10 ml of sterile SURF drainage water and plated out on the 1/10 R2B media mentioned previously. Two sample diffusion chambers were homogenized in 10 mL of sterile SURF drainage water, diluted, and plated out on 1/10X R2B media with vitamins and fungicides. The contents of one diffusion chamber were homogenized, and DNA was extracted (see 16S rDNA Library Preparation), while another chamber was opened and examined under a microscope at 50X to confirm the presence of bacterial micro-colonies.

16S rDNA Library Preparation. Genomic DNA was extracted from water filters using the PowerWater kit (MoBio, Inc.), while DNA was extracted from sediment and agar in diffusion chambers using a PowerSoil kit (MoBio, Inc.). DNA concentrations were determined using QuibitTM dsDNA (Life Technologies). Samples with concentrations <5 ng/μL were cleaned and concentrated with Zymogen Research DNA Clean & ConcentratorTM. Samples >5.0 ng/μL were diluted to 5.0 ng/μL; samples <5.0 ng/μL had double the normal amount of DNA for the following step to compensate for the low DNA concentration. Bacterial 16S rDNA metagenomic libraries for NextGen sequencing were pre- pared with an Illumina NexteraTM kit. DNA was diluted to 7.5 ng/μL, thereby normalizing DNA concentrations among samples. PCR for 16S rDNA V3-V4 Proceedings of the South Dakota Academy of Science, Vol. 97 (2018) 133 region library preparation was done according to Illumina “16S Metagenomic Sequencing Library Preparation,” Part # 15044223 Rev. B. The first PCR protocol called for the following reagents: respective environmental DNA templates, 2x KAPA HiFi HotStart ReadyMix (KAPA), primers 16S Forward (5’-TCG- TCG-GCA-GCG-TCA-GAT-GTG-TAT-AAG-AGA-CAG-CCT-ACG-GG(A/ C/G/T)-GGC-(A/T)GC-AG-3’) and 16S Reverse (5’-GTC-TCG-TGG-GCT- CGG-AGA-TGT-GTA-TAA-GAG-ACA-GGA-CTA-C(A/C/T)(A/C/G)-GGG- TAT-CTA-ATC-C-3’). PCR was conducted with the following condtions: 95 °C (3 min), followed by 25 cycles of 95 °C (30 s), 55 °C (30 s), 72 °C (30 s) and a final 5 min extension step of 72 °C. This first PCR amplified 16S rRNA V3 and V4 regions. PCR products were centrifuged at 1,000 × g at 20 °C for 1 minute to collect condensation in the tubes. AMPure XP beads were added to the amplicon plate and mixed. After 5 minutes of incubating, the plate was placed on a magnetic stand for 2 minutes, the supernatant removed, and the beads washed briefly with 80% ethanol twice. After the beads had been air-dried for 10 minutes, the PCR plate was removed from the magnetic stand. Ten mM Tris (pH 8.5) was added, and the beads were allowed to incubate at room temperature for 2 minutes. The PCR plate was placed on the magnetic stand for 2 minutes, and the supernatant, containing the PCR products for Nextera XT indexing, was placed into a new PCR plate. This step purified the 16S V3 and V4 regions of residual primers and primer dimers. To attach dual indices and sequencing adapters, we arranged 5 μL of samples (PCR amplicon) in the PCR plate according to the TruSeq Index Plate Fixture. Nextera XT Index Primers 1 and 2 were N7xx and S5xx, respectively. 2x KAPA HiFi HotStart ReadyMix was added along with PCR grade water, mixed, and centrifuged at 1,000 × g for 1 minute. PCR was performed with the protocol: 95 °C (3 min), followed by 8 cycles of 95 °C (30 s), 55 °C (30 s), 72 °C (30 s) and a final 5-minute extension step of 72 °C. PCR products were centrifuged, and AMPure XP beads (Agencourt) were added to the plate, washed with ethanol, and suspended in Tris buffer. The purified samples were transferred to a new plate as before to prepare the final library for sequencing. Library quantity was determined on a Qubit 2 Fluorometer, and quality was determined with a Caliper GX DNA High Sensitivity Lab Chip. Samples were multiplexed for sequencing on an Illumina MiSeq instrument. Paired 300-bp read sequencing was done using MiSeq v3 reagents. Samples were pooled twice, resulting in superior normalization. Samples were run on the MiSeq, and results were sent to IlluminaTM BaseSpace.

16S rDNA Sequence Analysis. Sequences were downloaded from Illumina’s BaseSpace in .fastq format and imported to the CLC Bio Workbench as “Paired Reads, Paired-end.” “Remove Failed Reads” was selected, and quality scores were 134 Proceedings of the South Dakota Academy of Science, Vol. 97 (2018) set to the NCBI/Sanger or Illumina Pipeline 1.8 and later. Paired sequences were merged with mismatch costs at 1, minimum score of 40, gap cost of 4, and maximum unaligned end mismatches of 5. “Trimming Primer Sequences” was not needed as MiSeq automatically removes primer sequences during the quality control steps in BaseSpace. “Fixed Length Trimming” set all sequences to a uni- form length. Samples were filtered based on the number of reads with 100 as the minimum number of reads and 50 as the minimum percent from the median. “Copy Samples with Sufficient Coverage” was chosen. All samples passed quality control procedures. Operational Taxonomic Unit (OTU) clustering was conducted using 97% SILVA v. 119 database with the following parameters: 97% similarity percentage; 2 minimum occurrences; 3 chimera crossover cost; 6 Kmer size; and with “Find Best Match” chosen. OTUs with combined abundances less than 10 were removed. A maximum likelihood phylogenetic tree was constructed with the matched OTUs of the SURF samples. Sequences were aligned with known OTUs. Muscle alignment of OTU sequences was chosen with the following parameters: 1,000 maximum hours; 1,000 Mb maximum memory; 16 maximum iterations; 10 minimum combined abundances; 0.0% minimum combination percentage; 100 maximum number of sequences. The resulting alignment was used in making a maximum likelihood phylogeny. The phylogeny used the Neighbor-Joining method with the Jukes-Cantor nucleotide substitution method (Jukes and Cantor 1969) and a 100 bootstrap analysis. Alpha Diversity was conducted on the resulting tree and its given OTU Clustering Table. Diversity measures chosen were: The Number of OTUs, Chao1, Simpson’s Index, and Shannon-Weiner Entropy (Chao 2005; Shannon 1948; Simpson 1949). These indexes were chosen for comparison analyses of other studies and samples. Rarefaction Analysis used 1 Minimum Depth to Sample, 100,000 Maximum Depth to Sample, 20 Number of Depths to be Sampled, and 100 Replicates at Each Depth. Because some abundant OTUs were not classified (N/A) by CLC Bio, FASTA sequence files from those unclassified OTUs with at least 100 sequences (combined from all sites) were input into the Ribosomal Database Classifier Program at https://rdp.cme.msu.edu/classifier/classifier.jsp (Wang et al. 2007) to further attempt classification. The fixed-rank classification of each OTU, if greater than 50% reliability, was used to determine the taxonomic placement of the OTU.

Culturing Bacterial Isolates from Water and Diffusion Chambers. Bacterial colonies from pool water or diffusion chambers were streaked onto 1X R2A plates with nystatin and cycloheximide to obtain well-separated pure colonies, and the isolates were grown on slants of 1X R2A. Gram-staining, growth on Simmon’s Citrate agar, and testing for catalase and cytochrome oxidase were performed as described in Leboffe and Pierce (2010).

Random Amplified Polymorphic DNA Analysis of Isolates. Genomic DNA from 66 isolates directly cultured from pool water and 52 isolates from diffusion Proceedings of the South Dakota Academy of Science, Vol. 97 (2018) 135 chambers were extracted using a DNeasy Tissue Kit (Qiagen, Germany), following the supplied protocol for gram-positive bacteria. The amplification of inter- spersed repetitive DNA sequences was performed using the BOX-AIR primer (5’-CTA CGG CAA GGC GAC GCT GAC G-3’) (Versalovic et al. 1994). The PCR reaction mixture (15 μL) for CN based isolates contained 2 ul of template DNA, 10x BSA, 10 mM BOX-AR1 primer, deoxynucleoside triphosphates (each 10 mM), 10X HotMaster Taq buffer with magnesium (5 Prime Inc, USA), and 0.5 U HotMaster Taq DNA polymerase. PCR was carried out under the following conditions: initial denaturation at 94 °C for 2 min; 30 cycles of denaturation at 94 °C for 20 s, annealing at 53 °C for 1 min, and extension at 65 °C for 4 min, and a final extension at 65 °C for 8 min. For DNA fingerprinting of 0.1X R2A agar isolates, we used 5 Prime HotMasterMix (2.5X) for preparing the reaction mixture. The PCR products were separated by electrophoresis on a 2% agarose gel as described by Suel-Silva et al. (2013). The gels were stained with Ethidium Bromide for 30 min and de-stained in distilled water for 30 min before visual- izing the band patterns under a transilluminator. The degree of similarity of PCR amplicon sizes from the different isolates was evaluated by visual observation, and the isolates were separated into RAPD OTU groups. Representatives, at least one for of each RAPD OTU group, were selected for PCR amplification and sequencing of the nearly complete 16S rRNA gene (~ 1,400 base-pair product). The 16S rRNA gene was amplified by PCR with oligonucleotide primers 27F (5’-GAGTTTGATCMTGGCTCAG-3’) and 1492R (5’-GGT TAC CTT GTT ACG ACT T-3’ (Weisburg et al. 1991). The reaction mixture (15 μL) contained 5 Prime HotMasterMix (2.5x), 10x BSA, 27F and 1492R primers (10 μM) and 2μl genomic DNA. The PCR reaction started with pre-denaturation at 94 °C for 2 min, followed by 32 cycles of denaturation at 94 °C for 30 sec, annealing at 57 °C for 15 sec and 4 min of extension at 65 °C with a final extension at 65 °C for 7 min. After cleaning the amplified product with ExoSAP-IT (Affymetrix, USA), we performed sequencing with primers 27F (AGAGTTTGATCMTGGCTCAG), 338F (ACT CCT ACGGGAGGCAGCAG) and 1390R (CGGTGTGTACAAGGCCC) using BigDye Terminator v1.1 Cycle Sequencing Kit in ABI 3130xl Genetic Analyzer Applied Biosystems, USA) at Black Hills State University (BHSU) Western South Dakota DNA Core Facility.

RESULTS

Water Chemistry. At the site of collection, the pool of fracture water was warm (about 32 °C) and alkaline (pH 8.87). The main cation was sodium, while the main anions were carbonate, bicarbonate, and sulfate (Table 1). A more com- prehensive list of cations and anions from this same pool is presented by Barnes and Zehfus (2017). Although sulfide was not detectable by water chemistry mea- surements (less than 0.05 mg/L), a slight odor of sulfide was noted at the site. A significant concentration of methane was present (36.8 μg/mL). 136 Proceedings of the South Dakota Academy of Science, Vol. 97 (2018)

Table 1. pH and concentration of selected dissolved chemicals in the pool of water sampled on the 1770 m level of SURF. Note: dissolved organic carbon, ammonia, nitrite, sulfide, aluminum, arsenic, copper, iron, selenium, and zinc were below the limits of detection.

Chemical Concentration (mg/L) Carbonate 483 Sodium 254 Sulfate 86.7 Bicarbonate 33.4 Chloride 8.33 Silicon 7.74 Potassium 6.55 Calcium 2.82 Magnesium 0.87 Strontium 0.127 Barium 0.109 Nitrate 0.095 Phosphorous (total) 0.014 pH 8.87 Methane 36.8 mg/L

Taxa Represented in 16S rDNA Sequences from Pool Sediments, Water, and Diffusion Chambers.A diverse group of phyla were represented in pool sediments, including Proteobacteria (31% of sequences, including Acidiferribacter, Thauera, and Thiobacillus), Acidobacteria (17%, mainly subgroup 22), Planctomycetia (12%), Nitrospirae (11%), Chlorobi (7%), Actinobacteria (4%), Chlamydiae (3%), Verrucomicrobia (2%), and others (Figure 1). Most OTUs belonged to uncultured genera. About 63% of 16S rDNA sequences from pool water belonged to Proteobacteria (Figure 2). The most abundant genera of Proteobacteria were Methylococcus (16%) and Thiobacillus (9%); Thauera, Methyloversatilis, Acidiferribacter, and Aquatilis were also present. Planctomycetia, Nitrospirae, Bacteroidetes (especially Ohtaekwangia), Acidobacteria, and other phyla were also present. Proceedings of the South Dakota Academy of Science, Vol. 97 (2018) 137

Figure 1. Proportion of microbial OTUs belonging to different bacterial phyla, classes, and genera from pool sediments. 138 Proceedings of the South Dakota Academy of Science, Vol. 97 (2018)

Figure 2. Proportion of microbial OTUs belonging to different bacterial phyla, classes, and genera from pool water.

About 91% of the 16s rDNA sequences from agar in diffusion chambers were from the phylum Proteobacteria (Figure 3). Pseduomonas (32%) and Hyphomicrobium (14%) were the most abundant genera; Aquabacterium, Inhella, Sphingomonas, Sphingobium, Azoarchus, Aquimonas, Methyloversatilis, and Variovorax were also present. Small numbers of Bacteroidetes, Nitrospirae, Verrucomicrobia, and other phyla were also present. Proceedings of the South Dakota Academy of Science, Vol. 97 (2018) 139

Figure 3. Proportion of microbial OTUs belonging to different bacterial phyla, classes, and genera from agar in diffusion chambers.

Alpha-Diverity of OTUs in Pool Samples. Chao1 bias-corrected estimates for OTU richness were 435 for pool sediment, 528 for pool water, and 294 for agar in diffusion chambers. Shannon-Wiener diversity for samples was 6.48 for pool sediment, 6.48 for pool water, and 4.89 for diffusion chambers. Simpson diversity for samples was 0.97 for sediment, 0.96 for water, and 0.90 for diffusion chambers. 140 Proceedings of the South Dakota Academy of Science, Vol. 97 (2018)

Bacterial Isolates from Direct Isolation from Pool Water and from Diffusion Chambers—Ninety-one isolates were characterized from direct culturing of pool water and 89 from diffusion chambers. In both cases, gram-negative bacilli that were catalase positive predominated, although coccus shaped cells, and oxidase positive cultures were more common in the directly cultured isolates (Table 2). A total of 25 different RAPD groups were defined from the isolates, with some groups found only among the direct isolates, others only diffusion chamber isolates, and some in both kinds of isolates (Table 3).

Table 2. Summary of phenotypic characteristics of bacteria isolated from the pool of water sampled on the 1770 m level of SURF.

Direct Isolates Diffusion Chamber Total Growing 91 89 Percent Bacilli 75 95 Percent Cocci 25 5 Percent Gram-Positive 11 5 Percent Oxidase-Positive 89 47 Percent Catalase-Positive 71 66 Percent using Citrate 32 44

Table 3. Number of bacterial isolates from the pool from either direct isolation or isolation from diffusion chambers belonging to different RAPD genotypic groups. The genus represented by sequencing a member of the RAPD group is shown.

RAPD Group Direct Isolation Diffusion Chamber Bacterial Genus A 25 6 Methyloversatilis B 1 0 Rhizobiales C 12 9 D 2 9 Parvibaculum E 1 0 Methyloversatilis F 1 0 G 2 0 Inhella H 1 0 Burkholderiales I 2 1 J 4 1 Inhella K 1 1 Bacillus L 2 0 M 1 1 N 2 0 Proceedings of the South Dakota Academy of Science, Vol. 97 (2018) 141

Table 3. Continued.

RAPD Group Direct Isolation Diffusion Chamber Bacterial Genus O 3 1 P 1 0 Q 2 1 R 1 2 S 1 0 T 0 11 Inhella U 0 2 V 0 1 W 0 3 X 0 3 Y 1 0 Total 66 52

Sequencing of 16S rDNA from Isolates. Nearly full-length 16S rDNA sequences were obtained from 19 isolates obtained by direct culturing and by culturing from diffusion chambers. Sixteen were identified to genus to 100% confidence by RDB Classifier: seven wereMethyloversatilis , four Bacillus, three Inhella, one Hyphomonas, and one Parvibaculum. Two isolates were identified only to order Burkholderiales, and one to order Rhizobiales.

DISCUSSION

The sediments in the pool have a diverse bacterial community with some likely chemoautotrophic members, such as Nitrospira, which oxidizes nitrite, and Acidiferribacter, which oxidizes ferrous iron or sulfide, and probable che- moheterotrophs, such as Acidobacteria. Pool water microbes had a large proportion of chemoautotrophic sulfide oxidizers Thiobacillus( ), methanotrophs (Methylococcus), and facultative methanol/methylamine oxidizers (methylotrophs, including Methyloversatilis and Hyphomicrobium). Osburn et al. (2014) sampled from a flowing borehole (their Site 8) less than 10 m from the pool sampled in this study. They measured sulfide (382 μg/L) and methane (38 nM), and noted the presence of sulfide and methane-oxidizing bacterial taxa, the genera of which were not listed. As in the pool studied here, sulfide oxidizing microbes, especially Thiobacillus, and a likely methanol oxidizer (Methylotenera) were abundant in drainage water in another region of the same tunnel in SURF sampled by Bergmann et al. (2014). The importance of one-carbon metabolism in this community is not surprising given the presence of methane in fracture and pool water. 142 Proceedings of the South Dakota Academy of Science, Vol. 97 (2018)

The bacterial communities of the diffusion chambers were lacking in obligate chemoautotrophs, perhaps due to the addition of organic nutrients from the diluted R2A media. However, facultative methylotrophs, such as Hyphomicrobium and Methyloversatilis, persisted in the diffusion chambers. Various chemohetero- trophic Proteobacteria, especially Pseudomonas, dominated the bacterial community in diffusion chambers. Bacterial isolates from pool water, both directly isolated from water and isolated from the diffusion chambers, included the generaMethyloversatilis, Hyphomicrobium, and Inhella, which were common in 16S rDNA from both pool water and diffusion chambers. However, none of the 19 isolates whose 16S rDNA was sequenced was identified asPseudomonas , the dominant genus observed from diffusion chamber 16S rDNA. Our inability to isolate this abundant member of the microbial community is puzzling. Also, three of the 19 isolates were identified as Bacillus, despite the fact that very few 16S rDNA sequences from either pool water or diffusion chambers were fromBacillus or other Firmicutes. In this case, unlike the study described by Bollmann et al. (2007), the use of diffusion chambers did not result in the isolation of a more representative or diverse group of bacterial taxa from the pool in SURF than direct culturing.

ACKNOWLEDGEMENTS

This project was funded through NSF REU Grant # 1560474. We thank Brianna Mount for administering the REU program, and SURF personnel, including Tom Regan and others. We thank Oxana Gorbatenko and Bethany Reman at BHSU-CCBR for their help with many technical aspects of this project, such as high-throuput 16S rDNA sequencing. We also thank Cynthia Anderson, Naveen Malik, and the students of Biology 371 at BHSU (Elizabeth Anderson, Cassandra Carter, Cathryn Hester, Nicole Davis, Ranni Hopkins, Zoe Langseth, Tayler Lenz, Kathryn Messler, Maddison Miller, Brian Mischel, Hannah Owens, Abby Reilly, Zeb Rozencranz, Tada Vargas-Black Bear, Cody Wellman, and Gabriel Yellowhawk) for their help with Sanger DNA Sequencing.

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