Robust Archaeal and Bacterial Communities Inhabit Shallow Subsurface Sediments of The

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1 Robust archaeal and bacterial communities inhabit shallow subsurface sediments of the

2 Bonneville Salt Flats

3 Julia M. McGonigle1, Jeremiah A. Bernau2, Brenda B. Bowen2, William J. Brazelton1

4 1School of Biological Sciences, University of Utah 2Department of Geology & Geophysics, University of Utah

25 ABSTRACT

26 We report the first census of natural microbial communities of the Bonneville Salt Flats

27 (BSF), a perennial salt pan at the Utah–Nevada border. Environmental DNA sequencing of

28 archaeal and bacterial 16S rRNA genes was conducted on samples from multiple evaporite

29 sediment layers of the surface salt crust. Our results show that at the time of sampling

30 (September 2016), BSF hosted a robust microbial community dominated by diverse

31 Halobacteriaceae and Salinibacter species. Desulfuromonadales from GR-WP33-58 are also

32 abundant in all samples. We identified taxonomic groups enriched in each layer of the salt crust

33 sediment and revealed that the upper gypsum sediment layer found immediately under the

34 uppermost surface halite contains a robust microbial community. We found an increased

35 presence of Thermoplasmatales, Nanohaloarchaeota, Woesearchaeota, Acetothermia,

36 Halanaerobium, Parcubacteria, Planctomycetes, Clostridia, Gemmatimonadetes, Marinilabiaceae

37 and other Bacteroidetes in this upper gypsum layer. This study provides insight into the diversity,

38 spatial heterogeneity, and geologic context of a surprisingly complex microbial ecosystem within

39 this macroscopically-sterile landscape.

41 IMPORTANCE

42 Over the last ~13,000 years the Pleistocene Lake Bonneville, which covered a large

43 portion of Utah, drained and desiccated leaving behind the Bonneville Salt Flats (BSF). Today

44 BSF is famous for its use as a speedway, which has hosted many land-speed records and a

45 community that greatly values this salty landscape. Additionally, the salts that saturate BSF basin

46 are extracted and sold as an additive for agricultural fertilizers. The salt crust is a well-known

47 recreational and economic commodity, but the roles of microbes in the formation and

48 maintenance of the salt crust are generally unknown. This study is the first geospatial analysis of

49 microbial diversity at this site using cultivation-independent environmental DNA sequencing

50 methods. Identification of the microbes present within this unique, dynamic, and valued

51 sedimentary evaporite environment is an important step toward understanding the potential

52 consequences of perturbations to the microbial ecology on the surrounding landscape and

53 ecosystem.

55 INTRODUCTION

56 The Great Salt Lake and Bonneville Salt Flats (BSF) are both remnants of Pleistocene

57 Lake Bonneville which drained at the end of the last glacial maximum and desiccated over the

58 last ~13,000 years (Crittenden, 1963). BSF is famous for its use as a speedway through the last

59 century, which has hosted many land-speed records and a community that greatly values this

60 salty landscape (Noeth, 2002). Over this same time period, the salts that saturate BSF basin have

61 been extracted as an economic commodity, particularly as potash (e.g. KCl) that is sold as an

62 additive for agricultural fertilizers. The hydrology and sedimentology of BSF have been studied

63 periodically throughout the 20th century in relation to potash mining and concerns about the

64 impacts of land use on the environment (Associates, 1979; Brooks, 1991; Eardley, 1962; Kaliser,

65 1967; L. J. Turk, S.N. Davis, 1973; Lines, 1979; Turk, 1970; White III and Terrazas, n.d.).

66 Human land use has altered many aspects of the hydrology and morphology of the environment

67 that facilitated deposition of the ~2m thick evaporite salt crust that caps BSF, and the amount

68 and extent of salt present at the site have been observed to change through time (Bowen, B.B.,

69 Bernau, J., Kipnis, E.L., Lerback, J., Wetterlin, L. and Kleba, 2018; Bowen et al., 2018).

70 Examination of modern and ancient salt pan deposits demonstrates that these

71 environments undergo repeated cycles of desiccation (dry saline pan), flooding (brackish lake),

72 and evaporative concentration (drying to dry saline pan) (Bowen and Benison, 2009;

73 Lowenstein, T.K. and Hardie, 1985). While the desiccation stage is most common, it is

74 repeatedly interrupted by flooding events related to individual storms, wet seasons, and spring

75 thaws driving runoff from surrounding mountains. The environmental parameters that influence

76 the hydrology and timing of flooding, evaporation, and desiccation cycles are dynamic (Bowen

77 et al., 2017). For this study, multiple individual strata from several sites spanning the salt crust

78 were sampled at BSF during an extensive sampling campaign to measure the overall stratigraphy

79 and volume of the salt crust (Bowen et al., 2018). The samples were collected in early

80 September 2016 near the end of a three-month long desiccation period.

81 Microbial communities provide critical ecosystem services by cycling nutrients and

82 providing a biological foundation upon which other organisms can establish themselves. In

83 environments considered “extreme” by human standards such as BSF, microbes are often the

84 only life forms that can survive. The study of microbial life in these ecosystems can elucidate

85 how extremophilic organisms have evolved unique attributes, strategies, and metabolic

86 capabilities that enable them to thrive where most other organisms cannot (Ma et al., 2010;

87 Ventosa et al., 2015). Extreme hypersaline ecosystems select for organisms capable of not only

88 tolerating, but actually requiring high salt for growth (Litchfield and Gillevet, 2002). Microbes in

89 these ecosystems employ either a “salt-in” or “organic-solutes-in” strategy to overcome the

90 problems associated with osmoregulation in a high salt environment (Oren, 1999). In addition,

91 halophiles found in these environments produce enzymes with potential biotechnological,

92 bioremediation, and medical applications (Ventosa and Nieto, 1995).

93 The role of microbes in BSF salt crust processes are generally unknown. Identification of

94 the microbes present within this unique, dynamic, and valued sedimentary evaporite environment

95 is an important step toward understanding the potential consequences of perturbations to the

96 microbial ecology on the surrounding landscape and ecosystem. This study is the first geospatial

97 analysis of microbial diversity at BSF using cultivation-independent environmental DNA

98 sequencing methods. This study provides insight into the diversity, spatial heterogeneity, and

99 geologic context of a surprisingly complex microbial ecosystem within this macroscopically-

100 sterile landscape.

101

102 METHODS

103 Sampling Location and Collection

104 Samples were collected from eight pits dug with sterile tools in September 2016 (Table

105 S1, Fig. S1). Distinct sediment layers visually identified by color and textural changes were

106 sampled at each site. Samples were immediately placed on ice and transported within 12 hours to

107 the University of Utah where they were stored at -80 °C until microbiological analyses were

108 performed in the following 6 months.

109

110 Elemental and Sedimentological Analysis

111 A portion of each homogenized frozen sample was separated and analyzed for major and

112 trace elements by ActLabs. Major and trace elements were analyzed via Aqua Regia ICP

113 (Inductively Coupled Plasma; method 1E3). Anions were analyzed via Ion Chromatography

114 (method 6B). Using this elemental data along with visual characteristics, each sample was

115 grouped into one of five sediment categories. Elemental concentrations were used to reconstruct

116 relative weight mineralogy based upon stoichiometry of gypsum, halite, and clay. Elemental

117 concentrations beyond detection limits were derived using this method.

118 In addition to these samples, additional blocks of representational material were collected

119 at the same time of sampling from sites 56, 35, and 12B. These blocks were processed by

120 Wagner Petrographic to create petrographic thin sections for visual microscopy analysis. Blocks

121 were desiccated to remove any pore waters, impregnated with clear or colored epoxy. Several

122 thin sections were made from each block by cutting 50 by 75 mm blocks and mounting them on

123 glass slides. Samples were cut to thicknesses ranging from 0.5 to 2 mm. The samples were then

124 analyzed at the University of Utah using an Olympus BX53M microscope with 1000x

125 magnification and a Zeiss M2 Petroscopic Microscope with an Axiocam Camera and Zen 2 Pro

126 software.

127

128 DNA Extraction and Sequencing

129 DNA was extracted from salt crust samples using a protocol from Brazelton et al. (2010)

130 modified for extremely high-salt material (Brazelton et al., 2010). The modified protocol is

131 available on our lab's website (https://baas-becking.biology.utah.edu/data/category/18-protocols)

132 and summarized here. Sediment samples were crushed and homogenized with a sterile mortar

133 and pestle, and 0.25 g subsamples were placed in a DNA extraction buffer containing 0.1 M Tris,

134 0.1 M EDTA, 0.1 M KH2PO4, 0.8 M guanidium HCl, and 0.5 % Triton-X 100. For lysis, samples

135 were subjected to one freeze-thaw cycle, incubation at 65 ºC for 30 minutes, and to beating with

136 0.1 mm glass beads in a Mini-Beadbeater-16 (Biospec Products). Purification was performed via

137 extraction with phenol-chloroform-isoamyl alcohol, precipitation in ethanol, washing in Amicon

138 30K Ultra Centrifugal filters, and final clean-up with 2x SPRI beads (Rohland and Reich, 2012).

139 DNA quantification was performed with a Qubit fluorometer (ThermoFisher).

140 Archaeal and bacterial 16S rRNA gene amplicon sequencing was conducted by the

141 Michigan State University genomics core facility. The V4 region of the bacterial 16S rRNA gene

142 was amplified with dual-indexed Illumina fusion primers with the 515F/806R primers as

143 described by Kozich et al. (Kozich et al., 2013). The V4 region of the archaeal 16S rRNA gene

144 was amplified with the A519F/Arch958R primers (Klindworth et al., 2013). Amplicon

145 concentrations were normalized and pooled using an Invitrogen SequalPrep DNA Normalization

146 Plate. After library QC and quantitation, the pool was loaded on an Illumina MiSeq v2 flow cell

147 and sequenced using a standard 500 cycle reagent kit. Base calling was performed by Illumina

148 Real Time Analysis (RTA) software v1.18.54. Output of RTA was demultiplexed and converted

149 to fastq files using Illumina Bcl2fastq v1.8.4.

150

151 Analysis of 16S rRNA Amplicon Data

152 Quality screening and processing of all 16S rRNA amplicon sequences was conducted

153 with the DADA2 R package (Callahan et al., 2016). Using this package primer contaminants and

154 chimeras were removed, reads were trimmed and filtered based on quality, and sequence variants

155 likely to be derived by sequencing error were identified. The final amplicon sequence variants

156 (ASVs) are considered to be true variants and are analyzed in the same method as traditional

157 operational taxonomic units (OTUs). Taxonomic classification of all ASVs was performed with

158 the DADA2 package and the SILVA reference database (NRv123).

159 Sequence counts for each ASV were transformed with a biomass correction by

160 multiplying the proportion of each ASV's contribution to each sample's total sequence count by

161 the estimated number of total cells in that sample. Total cells were estimated by the total ng of

162 extracted DNA (as measured by the Qubit fluorometer) and an assumption of 2x10-6 ng of DNA

163 per cell. All downstream analyses were performed with these biomass-weighted relative

164 abundances. Differential abundance of ASVs in each layer was performed using edgeR

165 (McMurdie and Holmes, 2013a) and phyloseq (McMurdie and Holmes, 2013b). An ASV was

166 considered to be significantly enriched in a layer if its differential abundance passed a false

167 discovery rate threshold of 0.05. Multivariate analysis was done using phyloseq (McMurdie and

168 Holmes, 2013b) and Bray-Curtis and Simpson indices were calculated using vegan (Jari

169 Oksanen, F. Guillaume Blanchet, Michael Friendly, Roeland Kindt, Pierre Legendre, Dan

170 McGlinn, Peter R. Minchin, R. B. O’Hara, Gavin L. Simpson, Peter Solymos, M. Henry H.

171 Stevens, 2018).

172

173 Accession numbers

174 All sequence data are publicly available at the NCBI Sequence Read Archive under BioProject

175 PRJNA522308. All SRA metadata, supplementary material, and protocols are archived with the

176 following DOI: http://doi.org/10.5281/zenodo.2564239. All custom software and scripts are

177 available at https://github.com/Brazelton-Lab.

178

179 RESULTS

180 Elemental and Sedimentological Analysis

181 Thin sections and reconstructed mineralogy from elemental data were used to group each

182 sample into one of five classifications: Group 1 (surface halite), Group 2 (upper gypsum), Group

183 3 (lower halite), Group 4 (lower gypsum), and Group 5 (halite mixed with gypsum).

184 Thin section and elemental analysis and microscopy reveal different layers of distinct

185 sedimentological morphology (Fig. 1, Fig. S2). Primary minerals were identified as halite

186 (NaCl), gypsum (CaSO4*2H2O), and clay minerals (Fe, Mg, and Al). The majority of samples

187 were above reporting limits for sodium. All minerals are reported as weight % of composition

188 (Table S1, Figs. S3-S5). The relative enrichment of select major and trace elements was

189 determined for each sediment classification (Table 1). Groups with mean values above the cutoff

190 limit were reported as enriched with an element.

191 Thin sections revealed Group 1 (surface halite) consists of puffy fine efflorescent halite

192 crystals followed by larger halite crystals (up to 0.5 cm wide) with fluid inclusions. Group 1

193 samples are composed of predominantly halite with 2.5-27% gypsum and trace amounts (<0.5%)

194 of clay. Group 1 is relatively enriched in chloride, potassium, zinc, and lead (from most to least

195 abundant).

196 Group 2 (upper gypsum) consists of layered fine to medium sized gypsum grains. Group

197 2 samples contain 29-39% gypsum and 6-22% halite. Samples 56-2 and 35-2 are mineralogical

198 outliers in this group as they have higher amounts of halite. Thin sections from sites 35 and 56

199 show relatively thin (~5-20 mm thick) layers of fine-grained gypsum adjoined by halite (Fig.

200 S2). Group 2 samples contain the highest amount of clay minerals of any strata at 0.3-1.7%.

201 Reconstructed mineralogy only accounted for 41-61% of Group 2’s mass. Group 2 is enriched in

202 sulfur, calcium, magnesium, aluminum, potassium, iron, strontium, zinc, lead, and manganese.

203 Group 3 (lower halite) samples are cemented halite with porous vertical dissolution pipes

204 running through it. These pipes are typically 10mm - 30mm wide, but some are > 1 cm.

205 Elemental data from group 3 is similar to Group 1 (surface halite). Group 3 is enriched in

206 chloride, and zinc.

207 Group 4 (lower gypsum) samples contain coarse black organic or clay coated gypsum

208 grains (medium to coarse, some very coarse) that smell of sulfur. Samples in this category have

209 similar amounts of gypsum (32.5-38%) but differing amounts of halite (6.5-54%). Group 4

210 samples are enriched in sulfur, calcium, magnesium, aluminum, potassium, iron, strontium, and

211 barium.

212 Group 5 (halite mixed with gypsum) consists of a more chaotic halite (53-61%)

213 supported framework made from vestigial dissolution pipes filled by pore space and 28-33%

214 gypsum. Group 5 is enriched in chloride, calcium, sulfur, strontium, and barium.

215

216 Bacterial and Archaeal Community Composition

217 All sediment samples produced 1,845,510 merged paired reads with the bacterial primer

218 set and 552,906 merged paired reads with the archaeal primer set. From these, DADA2 identified

219 2,740 amplicon sequence variants (ASVs) in the bacterial dataset and 1,646 ASVs in the archaeal

220 dataset. DNA extraction yields and ASV counts per sample are shown in Table S2. These ASV

221 counts were transformed with the DNA extraction yields with the procedure described in the

222 methods to obtain biomass-weighted environmental abundances for downstream analyses. At

223 most sites, samples in Group 1 and Group 2 have higher DNA yields and ASV counts than

224 samples from Group 3, 4, and 5. Samples from Group 1 and 2 sequenced more successfully than

225 those in other groups, particularly with archaeal primers.

226 There were 1,552 ASVs (56% of the total ASVs) in the bacterial dataset that were

227 assigned archaeal taxonomy. The 515F/806R bacterial primers are known to amplify both

228 bacteria and archaea, so we chose to include the archaeal ASVs in the bacterial dataset rather

229 than remove such a large proportion of the total sequence diversity (Apprill et al., 2015; Parada

230 et al., 2016; Walters et al., 2015). Most archaeal ASVs in the bacterial dataset overlap

231 classifications found in the archaeal dataset, but the archaeal dataset generally provided more

232 specific classifications, particularly for the class Thermoplasmata. The exceptions are

233 Woesearcheota and Nanohaloarchaeota, which are only found in the bacterial dataset.

234 All samples in the bacterial dataset are dominated by diverse Halobacteriaceae and

235 Salinibacter species (Fig. S6). All samples in the archaeal dataset are dominated by

236 Halobacteriaceae from 27 different classified genera (Fig. S7). Desulfuromonadales from GR-

237 WP33-58 are also abundant in all samples. Less dominant taxa generally increase in abundance

238 with depth at most sites. The most dominant cyanobacteria are classified as Lyngbya and are

239 more abundant in upper surface halites. Nanohaloarchaeota and ASVs classified to the Soil

240 Crenarchaeotic Group also appear more abundant in these upper sediments. Woesearchaeota,

241 Archaeoglobaceae, Aenigmarchaeota (DSEG), and Methanomicrobia (ST-12K10A) appear more

242 abundant in lower layers. In the bacterial dataset, the relative abundances of Acetothermia,

243 Rhodovibrio, Marinilabiaceae, Desulfovermiculus, Gemmatimonadetes, Phycisphaeraceae,

244 Thiohalorhabdus, and Ectothiorhodospiraceae are greater in lower layers. In both datasets, the

245 relative abundances of ASVs classified to Thermoplasmata are greatest in the middle layer.

246 Multivariate analysis of beta diversity for the bacterial and archaeal datasets did not

247 reveal any significant correlations between community compositions and location or sediment

248 depth. (Fig. S8, Fig. S9). Similarly, alpha diversity, as measured by the Simpson index, did not

249 exhibit any consistent patterns with location or sediment depth (Table S3). These results

250 highlight the general lack of significant variation among the whole-community compositions

251 prior to differential abundance analyses.

252

253 Differential Abundance

254 Despite the lack of variation at a whole community level, we investigated whether

255 specific ASVs are significantly enriched in one or more sediment layers. Differential abundances

256 of ASVs were calculated edgeR (McMurdie and Holmes, 2013a) by comparing each of the

257 sediment groups to each other (Fig. 1). For the bacterial dataset, comparisons were done

258 between: Group 1-2, Group 2-3, and Group 2-4. For the archaeal dataset, comparisons were done

259 between: Group 1-2 and Group 2-5. All other comparisons were not possible due to lack of data

260 from particular samples.

261 In the bacterial dataset, comparisons between Group 1 and 2 identified 429 ASVs with

262 significantly greater abundance (determined by a false discovery rate of <0.05) in Group 1

263 (surface halite) and 660 ASVs with significantly greater abundance in Group 2 (upper gypsum)

264 (Fig. 2, Fig. S10). Diverse Halobacteriaceae make up 37% of the surface halite-enriched ASVs.

265 Salinibacter species comprise another 43% of these ASVs. The remaining ASVs enriched in

266 Group 1 are related to the cyanobacteria Euhalothece, Desulfuromonadales, Nanohaloarchaeota,

267 and unclassified members of Chitinophagaceae. The enriched taxa become more diverse as the

268 sediment transitions from surface halite to the upper gypsum layer. Taxa with a greater

269 abundance in these Group 2 sediments include Thermoplasmatales, Nanohaloarchaeota,

270 Woesearchaeota, Acetothermia, Halanaerobium, Parcubacteria, Planctomycetes, Clostridia,

271 Gemmatimonadetes, Marinilabiaceae and other Bacteroidetes. Unclassified bacterial and

272 archaeal ASVs are also more abundant in Group 2.

273 In the archaeal dataset, comparisons between Group 1 and 2 identified 231 ASVs with

274 significantly higher abundance in Group 1 (surface halite) and 579 ASVs with significantly

275 higher abundance in Group 2 (upper gypsum). Diverse Halobacteriaceae make up 99% of the

276 surface halite-enriched ASVs, with the majority of these belonging to unclassified species. The

277 remaining 1% belong to members of DSEG and Thaumarchaeota. The ASVs enriched in Group

278 2 are also dominated by diverse Halobacteriaceae as well as Thermoplasmatales. A few ASVs

279 enriched in Group 2 belong to DSEG and unclassified Archaea.

280 The number of enriched ASVs identified by differential abundance comparisons between

281 upper gypsum and deeper sediment layers are much lower. In the bacterial dataset, comparing

282 Group 2 (upper gypsum) and 3 (lower halite) revealed 8 enriched ASVs in Group 2, mostly from

283 Aenigmarchaeota, Rhodovibrio, and Desulfobacteraceae, and 25 Group 3-enriched ASVs heavily

284 represented by Halobacteriaceae.

285 In the bacterial dataset, comparison of Group 2 (upper gypsum) and 4 (lower gypsum)

286 identified only one differentially abundant ASV in Group 2 samples, which was classified as

287 Halonotius, a species of Halobacteriaceae. 33 ASVs were identified as enriched in Group 4,

288 including diverse Halobacteriaceae, DPANN (Nanohaloarchaeota and Woesearchaeota),

289 candidate division SR1, chloroplasts, Halobacteroidaceae, and Clostridia such as Halanaerobium.

290 In the archaeal dataset, comparisons between Group 2 (upper gypsum) and 5 (halite

291 mixed with gypsum) found only 4 ASVs that were more abundant in Group 2 samples, all

292 belonging to Halobacteriaceae. There were 96 ASVs enriched in Group 5 samples. These ASVs

293 were classified as Aenigmarchaeota, diverse Halobacteriaceae, Methanomicrobia,

294 Thermoplasmata, and Thaumarchaeota.

295

296 DISSCUSION

297 The Bonneville Salt Flats hosts a robust microbial ecosystem

298 Despite its barren appearance on the surface, the Bonneville Salt Flats (BSF) appears to

299 harbor a robust microbial ecosystem featuring a range of metabolic capabilities potentially

300 including phototrophy, sulfur and nitrogen cycling metabolisms, and heterotrophy. Although

301 hypersaline systems can be harsh environments for life, they have been previously reported to

302 contain high biomass (Çınar and Mutlu, 2016; Litchfield and Gillevet, 2002; Maturrano et al.,

303 2006). Our results indicate that BSF is no exception. If we assume 2x10-6 ng of DNA per cell, we

304 estimate 107 - 109 cells per gram of sample from our quantifications of environmental DNA. This

305 is likely to be an over-estimate, since it is known that salt can preserve DNA, and our

306 environmental DNA could include a significant extracellular fraction (Borin et al., 2008; Rhodes

307 et al., 2011). Nevertheless, the low range of our biomass estimate is consistent with other

308 reported values around 106- 107 cells mL-1 (Çınar and Mutlu, 2016; Litchfield and Gillevet,

309 2002; Maturrano et al., 2006).

310 The surface halite and upper gypsum sediment layers (Groups 1 and 2) have higher DNA

311 yields than deeper samples (Table S2). These upper sediments have availability to sunlight and

312 oxygen while still retaining protection from UV radiation within the evaporite crusts.

313 Additionally, sulfur minerals in the upper gypsum layer likely provide an energy source for

314 chemotrophic microbes living in these upper layers. Therefore, the upper gypsum layer is likely

315 to have more abundant and more active phototrophic and chemotrophic microbial communities

316 compared to deeper layers.

317

318 A microbial community shift occurs along a mineralogical shift in the sediment

319 Although the depth of each sediment layer varied per site, microbial communities seemed

320 homogenous among samples within each sediment category. For example, at site 12B, the

321 surface halite was only 3 mm, while at site 56 the surface halite was 25mm (Fig. S11, Table S1).

322 In spite of these differences in depth, these surface halite samples hosted remarkably similar

323 archaeal and bacterial communities (Fig. S6, Fig. S7). This trend is still evident even between

324 the two samples with the greatest spread between sampling depths, site 12B where the upper

325 gypsum layer was sampled at a depth of 3mm-7mm and site 41 where the gypsum layer was

326 sampled at a depth of 25mm - 90mm. Both of these samples appear similar in mineral

327 classification and relative abundance of microbial taxa. These results suggest microbial

328 communities shift along mineralogical rather than depth gradients at BSF.

329 Upper halite crusts (Group 1) are dominated by heterotrophs from Salinibacter and

330 Halobacteriaceae. Halobacteriaceae gain most of their energy through heterotrophy but are also

331 capable of harnessing sunlight for supplemental ATP production with the specialized pigment

332 bacteriorhodopsin (Hartmann et al., 1980; Sharma et al., 2007; Spudich, 1998). Key primary

333 producers in the top halite layer are likely cyanobacteria, particularly Euhalothece, and the salt

334 tolerant algae Dunaliella salina. Although we did not perform 18S rRNA sequencing to

335 characterize the eukaryotic community, one ASV has 100% identity to the 16S rRNA gene of the

336 Dunaliella chloroplast. This ASV was only absent at 3 sites: 12B, 67B, and 41. Sites 12B and

337 67B have the highest proportion of gypsum for samples in Group 1. These sites are also located

338 in a region with the thinnest surface halite (Fig. S11).

339 Thin sections show surface halites contain large pore spaces and fluid inclusions within

340 the halite crystals (Fig. 1). This upper phototrophic community at BSF likely resides in tandem

341 with heterotrophs in these pore spaces of the halite structure or as a filamentous biofilms

342 attached to the surface of halite crystals (Spear et al., 2003). Microbes in this surface layer may

343 also be encapsulated within fluid inclusions. Members of this community are likely active during

344 the flooding stage of BSF, when upper halite dissolves and brine exists on the surface. They may

345 become encapsulated once again as the brine precipitates into halite crystals during the next

346 desiccation cycle.

347 Our results indicate that more genera are enriched in the upper gypsum sediments (Group

348 2) found beneath the surface halite layer than in other types of sediment (Fig. 2, Table 2). These

349 genera may be concentrated in this layer because gypsum, halite, and other evaporite minerals

350 provide some protection from harsh environmental conditions (such as UV radiation and extreme

351 temperature fluctuations) while still transmitting enough sunlight to support photosynthesis

352 (Friedmann, 1982). The higher surface area of clay minerals present in these samples likely

353 provides a more suitable substrate for growth than the surface halite.

354 In general, bacterial taxa associated with sulfur metabolism (e.g. Thiohalorhabdus,

355 Desulfovermiculus, Desulfobacteraceae, and Desulfobulbaceae) are enriched in the upper

356 gypsum (Group 2) layer. Phototrophs enriched in upper gypsum sediments include members of

357 Ectothiorhodospiraceae, the purple sulfur bacteria. These anaerobic organisms produce sulfur

358 globules outside of their cells. Thermophilic organisms (Thermoplasmatales, Archaeoglobaceae,

359 and Thermosulfurimonas) commonly found in environments containing sulfur compounds are

360 also enriched in this upper gypsum layer. These results are strongly suggestive that the abundant

361 sulfur minerals found in this layer (Table S1) are metabolized by the resident microbes.

362 Interestingly, methanogens and acetogens who often compete with sulfate-reducing bacteria, are

363 also enriched in the upper gypsum layer (Kato et al., 2014). Methanogens in salt flats often

364 utilize methylated compounds, rather than carbon dioxide, allowing them to coexist with

365 acetogens and sulfate-reducing bacteria (García-Maldonado et al., 2015).

366 The lowest layers sampled (Groups 4 and 5) contain chemoheterotrophic

367 (Thermoplasmata and Planctomycetes) and fermentative taxa perhaps dependent on primary

368 production of the phototrophic community in upper layers. Strictly anaerobic fermenters from

369 the class Clostridia (particularly the genus Orenia found within the order Halanaerobiales) were

370 enriched in deeper samples where oxygen is likely more depleted. Aerobic heterotrophs from the

371 Proteobacteria, such as Acinetobacter and Sandaracinus, are also enriched here when compared

372 to higher sediments suggesting oxygen may not be completely depleted at this depth. The

373 dissolution pipes and larger grain sizes found in the lower gypsum, lower halite, and lower halite

374 mixed with gypsum layers may enable more air circulation at these depths (Fig. S2).

375

376 The microbial community of BSF is similar to previously described hypersaline ecosystems

377 Only one culture-independent study of microorganisms at the Bonneville Salt Flats (BSF)

378 has been conducted previously (Lynch, 2015). This study focused on the adjacent basin Pilot

379 Valley but also included one sampling location at the BSF. The genus Halomina was the most

380 dominant member of the Halobacteriaceae in this BSF sample. Halomina sequences were also

381 identified in all of our samples, but the most abundant Halobacteriaceae genera in our dataset

382 were Haloarcula and Halapricum (Fig. S6, Fig. S7). Culture-dependent studies at BSF have

383 only been successful at isolating Haloarcula, Halorubrum, Halobacterium, and Salicola species

384 (Boogaerts, 2015; Gary M. King, 2015).

385 The microbial diversity at BSF is comparable to that of other dry saline environments.

386 Archaeal communities are often dominated by members of Halobacteriaceae (Di Meglio et al.,

387 2016; Kambourova et al., 2017; Stivaletta et al., 2009). Bacterial communities are often

388 dominated by Bacteroidetes (Salinibacter) and Proteobacteria (Rasuk et al., 2014; Stivaletta et

389 al., 2011; Vogt et al., 2017). Members from Crenarchaeota, Gemmatimonadetes,

390 Verrucomicrobia, SRB from Deltaproteobacteria, and Clostridia are also commonly reported

391 (Caton and Schneegurt, 2012; Kim et al., 2012; McKay et al., 2016). We found ASVs at BSF

392 that are closely related to all of these commonly reported species (Fig. S6, Fig. S7). Euhalothece

393 is the most commonly reported dominant Cyanobacteria in hypersaline environments(Caton and

394 Schneegurt, 2012; Lindsay et al., 2017; McKay et al., 2016; Schneider et al., 2013; Spear et al.,

395 2003; Stivaletta et al., 2011; Vogt et al., 2017). Some of our ASVs classified to Euhalothece, but

396 we found Lyngbya to be the most dominant Cyanobacteria at BSF.

397 Interestingly, microbial species commonly reported in hypersaline lake environments,

398 such the Great Salt Lake, Lake Chaka, and the Salton Sea are also found in our BSF samples

399 (Almeida-Dalmet et al., 2015; Bowman et al., 2000; Dillon et al., 2013; Jiang et al., 2007;

400 Lindsay et al., 2017; Schneider et al., 2013; Swan et al., 2010), suggesting physiological and/or

401 ecological connections between hypersaline aquatic and sediment systems.

402

403 CONCLUSION

404 The Bonneville Salt Flats hosts thriving microbial communities dominated by diverse

405 Halobacteriaceae and Salinibacter species that are apparently adapted to the challenges of this

406 extreme ecosystem. Many of the archaeal and bacterial taxa are most abundant beneath the halite

407 crust within the upper gypsum sediments, where microbes are likely metabolizing sulfur-

408 containing minerals and can still rely on light penetration for phototrophy.

409 This study represents the first comprehensive culture-independent characterization of the

410 microbial communities inhabiting the Bonneville Salt Flats, and future work in this understudied

411 ecosystem could address the following questions. Which microbes in this ecosystem are crucial

412 for the cycling of nutrients, such as sulfur and nitrogen? Are the haloarchaea dependent on light

413 filtered through the surface halite crust, and are they involved in surface halite formation? Is

414 there evidence for dispersal and exchange with nearby hypersaline environments such as Pilot

415 Valley and the Great Salt Lake? How does the microbial community shift with seasonal

416 desiccation and flooding stages? What sediment interfaces are microbes inhabiting? Do the

417 compositions of endolithic microbes and microbes preserved in fluid inclusions differ from

418 microbes within sediment interfaces? Has anthropogenic use of this environment for racing and

419 brine mining changed the microbial composition of this area?

420 Answering these questions would extend our understanding of the presence and

421 preservation of life not only in extreme environments, but in any environment where salt

422 deposits are formed. Halite crystal growth bands can preserve microbial life present in the

423 original air-water and water-sediment interface where the salt formed thousands to hundreds of

424 millions of years ago (Lowenstein, T.K. and Hardie, 1985; Sankaranarayanan et al., 2014).

425 Evaporite ecosystems may even reflect life on Earth prior to the Cambrian explosion (Ventosa et

426 al., 2015). Furthermore, because evaporite deposits on other planetary bodies, like Mars, are

427 evidence of past or present water (Barbieri, R. and Stivaletta, 2012; Barbieri and Stivaletta, 2011;

428 Douglas, 2005; Rothschild, 1990), further advances in the characterization of live and preserved

429 microbes in salt deposits will have implications for the development of extraterrestrial life

430 detection strategies.

431

432 ACKNOWLEDGEMENTS

433 This research was supported by the National Science Foundation Award 1617473, CNH-L:

434 Adaptation, Mitigation, and Biophysical Feedbacks in the Changing Bonneville Salt Flats. We

435 would like to thank Emily Dart and Betsy Kleba for assistance with sample collection, DNA

436 extraction, and general insight on the BSF system. We would also like to thank Alex Hyer and

437 Christopher Thornton for computational assistance.

438

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642 FIGURES/TABLES

Surface Halite Upper Gypsum Lower Halite Lower Gypsum Halite/Gypsum Cutoff Value (G1) (G2) (G3) (G4) (G5)

Mn 15 mg/L 10.25 38 5.2 19.5 10.5

Zn 10 mg/L 47.5 80.29 41.2 5 19

Al 0.10% 0.01 0.14 0.02 0.1 0.05

Ba 20 mg/L 1.75 29.29 4.6 22.5 22

Ca 5% 1.99 7.97 3.19 7.92 6.93

Fe 0.05% 0.02 0.13 0.02 0.09 0.04

K 0.15% 0.2 0.21 0.09 0.17 0.15

Mg 0.20% 0.13 0.4 0.07 0.25 0.16

S 2% 1.54 5.67 2.6 6.75 5.86

Sr 200 mg/L 106.38 505.29 146.2 465 472

Cl 450,000 mg/L 541125 139700 520800 201750 384500 643 Table 1: Sample means for sediment categories. Bold values indicate element is enriched in

644 samples.

645

Grp 1 Grp 2 Grp 3 Grp 4 Grp 5 # Taxonomic Groups 34 66 19 24 - in Bacterial Dataset # Taxonomic Groups 10 16 - - 6 in Archaeal Dataset 646

647 Table 2: Number of different Orders (or lowest assigned taxonomic group if not classified to

648 Order) represented in the enriched ASVs from differential abundance comparisons.

649

650

651 Figure 1: Sedimentology and differential abundance results (EdgeR) for archaeal and bacterial

652 datasets. Red dots indicate an ASV with a significantly higher abundance (false discovery rate

653 <0.05) in the indicated category. Representative stratigraphic column presented here. Dashed

654 lines represent stratigraphic shifts that only occurred at some sites. General Stratigraphy consists

655 of groups 1-5. Group 1, surface halite, contains efflorescent halite and halite crystals with

656 abundant fluid inclusions. Group 2, upper gypsum, consists of gypsum grains with varying

657 proportions of halite, this group has the highest proportion of clays. Group 3, lower halite, has

658 the highest proportion of pore space and is mineralogically similar to group 1. Group 4, lower

659 gypsum, contains coarser gypsum grains than group 2, this group also smelled of sulfur. Group

660 5, lower halite and gypsum, forms when pores in lower halite fill with gypsum.

661

662

663 Figure 2: Abundance of ASVs enriched in each group as determined by differential abundance

664 (EdgeR) comparisons shown in Figure 1. Number of ASVs making up the total plot are indicated

665 below the comparison ID. A full legend can be found in Fig. S11.