Analog Sinkhole

Microbial communities and organic biomarkers in a Proterozoic-analog sinkhole

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As Published http://dx.doi.org/10.1111/GBI.12252

Publisher Wiley Blackwell

Version Final published version

Citable link http://hdl.handle.net/1721.1/118593

Detailed Terms http://creativecommons.org/licenses/by/4.0/ Received: 26 September 2016 | Accepted: 07 July 2017 DOI: 10.1111/gbi.12252

ORIGINAL ARTICLE

Microbial communities and organic biomarkers in a Proterozoic-analog sinkhole

1Department of Biological Sciences, University of Cincinnati, Cincinnati, OH, USA Abstract 2Department of Plant and Microbial Biology, Little Salt Spring (Sarasota County, FL, USA) is a sinkhole with groundwater vents at University of Minnesota, St. Paul, MN, USA ~77 m depth. The entire water column experiences sulfidic (~50 μM) conditions sea- 3Department of Earth System Science, sonally, resulting in a system poised between oxic and sulfidic conditions. Red pinnacle Stanford University, Stanford, CA, USA 4Department of Geosciences and the mats occupy the sediment–water interface in the sunlit upper basin of the sinkhole, Penn State Astrobiology Research Center and yielded 16S rRNA gene clones affiliated with Cyanobacteria, Chlorobi, and sulfate- (PSARC), The Pennsylvania State University, University Park, PA, USA reducing clades of Deltaproteobacteria. Nine bacteriochlorophyll e homologues and 5Department of Geosciences, Baylor isorenieratene indicate contributions from Chlorobi, and abundant chlorophyll a and University, Waco, TX, USA pheophytin a are consistent with the presence of Cyanobacteria. The red pinnacle mat 6 Department of Earth, Atmospheric and contains hopanoids, including 2-methyl structures that have been interpreted as bio- Planetary Sciences, Massachusetts Institute of Technology, Cambridge, MA, USA markers for Cyanobacteria. A single sequence of hpnP, the gene required for methylation of hopanoids at the C-2 position, was recovered in both DNA and cDNA libraries Correspondence T. L. Hamilton, Department of Plant and from the red pinnacle mat. The hpnP sequence was most closely related to cyanobac- Microbial Biology, University of Minnesota, terial hpnP sequences, implying that Cyanobacteria are a source of 2-methyl hopa- St. Paul, MN, USA Email: [email protected] noids present in the mat. The mats are capable of light-dependent primary productivity and as evidenced by 13C-bicarbonate photoassimilation. We also observed 13C-bicarbonate J. L. Macalady, Department of Geosciences and the Penn State Astrobiology Research photoassimilation in the presence of DCMU, an inhibitor of electron transfer to Center (PSARC), The Pennsylvania State Photosystem II. Our results indicate that the mats carry out light-driven primary pro- University, University Park, PA, USA. Email: [email protected] duction in the absence of oxygen production—a mechanism that may have delayed the oxygenation of the Earth’s oceans and atmosphere during the Proterozoic Eon. Funding information National Science Foundation, Grant/ Furthermore, our observations of the production of 2-methyl hopanoids by Award Number: NSF EAR-0525503; NASA Cyanobacteria under conditions of low oxygen and low light are consistent with the Astrobiology Institute, Grant/Award Number: NNA04CC06A and NNA13AA90A recovery of these structures from ancient black shales as well as their paucity in modern marine environments.

1 | INTRODUCTION present-day levels. The prevailing view suggests that during the Proterozoic, ~2–0.5 billion years ago, atmospheric oxygen levels were A growing body of evidence indicates a substantial increase in within 1–10% of present-day levels (Kump, 2008). Although recent atmospheric oxygen ~2.5 billion years ago. However, following the ini- research has revealed some complexity in the evolution of the global tial rapid rise of oxygen, a prolonged delay in the further rise of oxygen oceans during the Proterozoic, substantial evidence from marine sed- lasted until ~500 million years ago, when atmospheric oxygen reached imentary deposits indicate that oxygen-poor and sulfidic conditions

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784 | wileyonlinelibrary.com/journal/gbi Geobiology. 2017;15:784–797. HAMILTON et al. | 785 have been an intermittent feature of the global oceans since the ini- in Earth’s evolution. The signal of 2-MeBHPs in the rock record may tial rise of oxygen (Canfield, 2005). Euxinic conditions have accompa- provide additional insights. nied remarkable events in Earth history including several Phanerozoic Little Salt Spring is a cover-collapse, karst-solution sinkhole lo- crises (Meyer & Kump, 2008). Sulfidic, anoxic environments are still cated 14 miles from the Gulf of Mexico in Sarasota, FL. This sinkhole widespread today although they tend to be confined to basins with has proven to be a valuable archaeological site because its anoxic bot- highly restricted circulation. They feature isotopic, mineralogical, or- tom waters preserve artifacts from some of North America’s earliest ganic, and morphological biosignatures that have the potential to be human populations (Alvarez Zarikian, Swart, Gifford, & Blackwelder, preserved in the geologic record, thereby enhancing our ability to in- 2005; Clausen, Brooks, & Weslowsky, 1975; Clausen, Cohen, Emiliani, terpret the presence or absence of the same biosignatures in earth’s Holman, & Stipp, 1979; Haynes, 2009; Peres & Simons, 2006). Data geologic record or on other planets. presented here and elsewhere (Alvarez Zarikian et al., 2005) indicate Hydrocarbon biomarkers derived from biologic lipid membranes the geochemical attributes of the sinkhole water are similar to those have historically played an important role in efforts to reconstruct thought to be common in the Proterozoic oceans, poised between events and environmental conditions in the deep past, including the oxic and sulfidic conditions. The microbial diversity of Little Salt Spring evolution of Cyanobacteria and oxygenic photosynthesis (Brocks, has not been investigated previously. Here, we report the discovery Buick, Summons, & Logan, 2003; Summons, Jahnke, Hope, & Logan, of conspicuous populations of microbial photoautotrophs that form a 1999). Hopanes, the diagenetic product of hopanoids (membrane lip- red pinnacle mat covering the sediment–water interface in the upper ids produced by numerous varieties of bacteria), are preserved in both basin. The primary populations in the mat are Cyanobacteria (oxygenic modern sediments and ancient sedimentary rocks, and are the most phototrophs) and green sulfur bacteria, Chlorobium spp. (anoxygenic abundant molecular fossils in the rock record (Brocks & Pearson, 2005; phototrophs). The mat contains multiple 2-methyl hopanoids and the Ourisson & Albrecht, 1992). Methylation of hopanoid A-rings is re- only hpnP gene recovered from the mat is affiliated with Cyanobacteria. tained upon diagenesis (Summons & Jahnke, 1992) and can have tax- Thus, Little Salt Spring provides an extraordinary opportunity to in- onomic relevance. For instance, C-3 methylated hopanes are thought vestigate the factors affecting competition among oxygenic and an- to be diagnostic of microaerobic methanotrophs and acetic acid bac- oxygenic phototrophs and how these interactions may have affected teria (Zundel & Rohmer, 1985). Hopanoids modified by methylation at the oxygen budget throughout the Proterozoic Eon. Furthermore, the the C-2 position, or 2-methyl hopanoids, have been recovered from recovery of 2-methyl hopanoids derived from Cyanobacteria in a low modern environments as well as from ancient sedimentary rocks and oxygen, sulfidic environment is consistent with both a cyanobacterial were long considered to be diagnostic of Cyanobacteria, and thus origin of these structures in ancient black shales and the paucity of a proxy for oxygenic photosynthesis in ancient sedimentary rocks 2-methyl hopanoids in modern marine sediments. (Summons et al., 1999). However, the discovery of their precursors, the 2-methylbacteriohopanepolyols (2-MeBHPs) in members of the 2 | MATERIALS AND METHODS Alphaproteobacteria (Bisseret, Zundel, & Rohmer, 1985; Bravo, Perzl, Hartner, Kannenberg, & Rohmer, 2001; Rashby, Sessions, Summons, & 2.1 | Site description, sample collection, and Newman, 2007; Renoux & Rohmer, 1985; Sahm, Rohmer, Bringermeyer, geochemistry Sprenger, & Welle, 1993) highlights the challenge associated with eval- uating the significance of 2-methylhopanes in sedimentary rocks. Little Salt Spring (Fig. S1) is a 78 meter diameter sinkhole lake located The function of 2-MeBHPs remains unknown, as well as the con- in Sarasota County, FL (lat. 27°04′30″N, long. 82°14′00″W). The ditions favoring the production of these lipids. These knowledge gaps geology and hydrology of the spring have been described previously further hinder our ability to interpret hopane signatures in the rock by Alvarez Zarikian et al. (2005). Groundwater vents enter the lower record. The geologic distribution of hopanes indicates a higher abun- basin of the sinkhole near the maximum explored depth of 77 m. dance of the C-2 methylated form in Proterozoic rocks compared to Flocculent brown sediments blanket the floor of the lower basin to Phanerozoic rocks and in carbonates compared to non-carbonate sed- a depth of several meters or more. The sunlit upper basin hosts rich imentary rocks (Knoll, Summons, Waldbauer, & Zumberge, 2007), as archaeological remains (Clausen et al., 1979; Dietrich & Gifford, 1997) well as increased hopane abundance during oceanic anoxic events in and an extensive seasonal red microbial mat at the sediment–water the Phanerozoic (Cao et al., 2009; Kasprak et al., 2015; Talbot et al., interface. The temperature of the water column averages ~27°C and 2008). The increased abundance of hpnP genes in environments where has a salinity of ~3 ppt. Based on diver observations, parts or all of the anoxic conditions prevail and fixed nitrogen is limiting could represent water column are often weakly sulfidic to within 6 m of the surface, conditions that favored their production and subsequent deposition and sulfidic conditions extend to the surface water and stream out- on early Earth (Ricci et al., 2013). As such, geomicrobiological and flow during seasonal events. biogeochemical investigations of modern environmental analogues to Dissolved sulfide concentration was measured at the field site in early Earth are necessary to shed light on the relationship between January 2009, June 2012, and October 2013 with a portable spectro- biosignatures, specifically 2-MeBHPs, and photic zone anoxia or eu- photometer (Hach Co., Loveland, CO), using methylene blue for total xinia. A better understanding of conditions that favor the production sulfides (Hach method 690, detection limit ~0.2 μM). Water column of 2-MeBHPs could greatly aid in the unraveling of current mysteries samples were collected using a 3.8 L Van Dorn bottle (Wildlife Supply 786 | HAMILTON et al.

Company, Yule, FL) deployed from a platform at the approximate center plasmid-specific primers. An average of 46 clones was sequenced of the sinkhole. Water samples were analyzed immediately for sulfide for the January 2009, January 2012, and October 2013 samples. as they were brought to the surface. Sulfide analyses were conducted in Sequences were assembled and manually checked using Bio-Edit duplicate and were within 5% of each other. Water depth, temperature, (v7.2.5) and checked for chimeras using CHIMERA_CHECK (Cole conductivity, oxidation–reduction potential (ORP), and dissolved oxy- et al., 2003). Putative chimeras were excluded from subsequent gen were measured in situ using a calibrated mulitparameter YSI 6600 analyses. Sequences were aligned using the Silva aligner avail- sonde (Yellow Springs, OH, USA). The mulitparameter YSI 6600 sonde able at http://www.arb-silva.de, imported into a database of more measures ORP using an Ag/AgCl reference electrode and a luminescent- than 200,000 reference sequences in the software program ARB based dissolved oxygen sensor (ROX®). The sonde was calibrated daily (Ludwig et al., 2004) and manually refined. Operational taxonomic according to the manufacturer’s protocol and a detection limit of units (OTUs) were identified using a sequence identity threshold of 0.3 μM was determined for the ROX sensor. Photosynthetically active 99% (0.01) using mothur (v.1.34.4) (Schloss et al., 2009). Rarefaction radiation (PAR) was measured at the surface and underwater using a analyses indicate this sequencing depth was sufficient to capture the scalar quantum PAR sensor (LiCor LI-193S) attached to a LiCor LI-1400 majority of diversity of 16S rRNA sequences in the red pinnacle mat data logger (Lincoln, NE). The sonde and PAR instruments were also (Fig. S2). Maximum parsimony and bootstrap consensus phyloge- deployed from the platform at the approximate center of the sinkhole. netic trees were computed for unique Chlorobi and Cyanobacteria PAR measurements were made between 12:00 and 13:00 when the operational taxonomic units (OTUs) with 1000 bootstrap replicates sky was cloudless. Diver-collected pinnacle mat samples for lipid and using PAUP* 4.0b10 (Swofford, 2003). Phylogenetic analyses were nucleic acid analyses were photographed in situ (Figs S1 and S2) before populated with closely related environmental sequences and cultured sampling. Red mat was collected in January 2009, January of 2012, and representatives identified using BLAST. October of 2013 at the sediment–water interface in the upper basin Total RNA was extracted in triplicate from preserved mat samples at a depth of 8–10 m. Mat and sediment samples were preserved in (collected in 2009, 2012, and 2013) as previously described (Hamilton, RNA later (two parts per sample volume; Ambion/Applied Biosystems, Peters, Skidmore, & Boyd, 2013b). RNA extracts were screened Foster City, CA, USA) and stored on ice until frozen (within 6 hr). for the presence of contaminating genomic DNA by performing a PCR using ~1 ng of RNA as template and bacterial 16S rRNA gene primers primers 1100F (5′-YAACGAGCGCAACCC-3′) and 1492R 2.2 | Microscopy and nucleic acid analyses (5′-GGTTACCTTGTTACGACTT-3′) and an annealing temperature of Pinnacle mat samples were examined using light and epifluorescence 55°C as described in Boyd et al. (2007). cDNA was synthesized from microscopy before and after staining with the nucleic acid-binding ~20 ng of purified RNA using the iScript cDNA Synthesis Kit (Bio-Rad fluorophor 4,6-diamidino-2-phenylindole (DAPI). Genomic DNA was Laboratories, Hercules, CA, USA) and the following reaction cycling extracted in triplicate from ~250 mg (wet weight) preserved pinnacle conditions: 5 min at 25°C, 30 min at 42°C, and 5 min at 85°C. cDNA mat samples collected in January 2009, January of 2012, and October was purified by ethanol precipitation and re-suspended in nuclease- of 2013 using a FastDNA Spin for Soil kit (MP Biomedicals, Solon, free water for use in PCRs. OH). Quality of extracted DNA was assessed on an agarose gel (1%) A ~570 bp fragment of hpnP was amplified from purified gDNA using the HiLo DNA Marker (Bionexus, Oakland, CA) and visualized and cDNA using the primers 1F (5′-GSBTTYATGCCDCCBCARGG-3′) by ethidium bromide staining and using a NanoDrop ND-1000 spec- and 1R (5′ TCNARKCCVAKRATRATNCC-3′) followed by 2F (5′-GYB trophotometer (NanoDrop Technologies, Wilmington, Delaware). VTBGGHGGNCCNTCNGT-3′) and 2R (5′-TCNGGNGTYTCDATN Equal volumes of each extraction were pooled and quantified using CC-3′) in a nested PCR as previously described (Ricci et al., 2013). PCR a Qubit dsDNA HS Assay kit (Molecular Probes, Eugene, OR, USA) products (n = 24 gDNA; n = 24 cDNA) were purified using a QIAquick and a Qubit 2.0 Fluorometer (Invitrogen, Carlsbad, CA, USA), respec- PCR Purification Kit (Qiagen, Valencia, CA) and sequenced using prim- tively. Bacterial 16S rRNA primers 27F and 1492R (Lane, 1991) were ers 2F and 2R the at Genomics Core Facility of the Huck Institutes of used to amplify nearly full-length small subunit rDNA genes from the Life Sciences at Penn State University. Sequences were assembled the pooled environmental genomic DNA. Thermal cycling conditions and manually checked using Bio-Edit (v7.2.5). were as follows: 5 min at 94°C for initial denaturation followed by 30 cycles of PCR consisting of 60 s at 94°C, 30 s at 52°C and 90 s at 2.3 | Pigment analysis 72°C, followed by 20 min at 72°C for final elongation. Reactions were performed in triplicate, purified using a QIAquick PCR Purification Kit Pigment extractions were performed on frozen samples of the pinnacle (Qiagen, Valencia, CA), and pooled. The purified and pooled PCR- mat. Samples were thawed, sonicated twice in acetone, and the com- amplified DNA was cloned using a TOPO TA cloning kit according bined extracts were filtered through a dichloromethane pre-extracted to the manufacturer’s instructions (Life Technologies, Carlsbad, CA). cotton wool plug and dried at ca. 21°C in a TurboVap LV concentration Colonies containing inserts (n = 138) were isolated by streak-plating workstation (Caliper LifeSciences). Throughout the procedure, care was and amplified by colony PCR using plasmid-specific M13 primers. taken to avoid exposure to light and the extracts were stored dry under

PCR products were sequenced at the Genomics Core Facility of the N2 at −20°C prior to analysis. The total pigment extract was analyzed by Huck Institutes of the Life Sciences at Penn State University using reversed-phase high performance liquid chromatography (HPLC) on an HAMILTON et al. | 787

Agilent 1200 system using a four-solvent gradient (0.01 M ammonium MS settings follow those reported in Talbot, Squier, Keely, & acetate, methanol, acetonitrile, and ethyl acetate) and two consecutive Farrimond, 2003; : positive ion mode, mass scanning from m/z 150– Waters Spherisorb ODS2 columns (3 μm, 150 mm × 4.6 mm I.D.) pro- 1300, and “Smart” fragmentation setting (i.e., 8000 nA corona voltage, tected by a Waters guard column (10 mm × 4.6 mm) and a Phenomenex 60 psi nebulizer pressure, dry gas of 5 l/min, dry temperature of 350

Security Guard C18 pre-column (4 mm × 3 mm) (Airs, Atkinson, & Keely, C, vaporizer temperature 490 C). The run was divided into three seg- 2001). Pigments were characterized by their mass spectra and resonance- ments targeting m/z 285 (0–10 min), m/z 1002 (10–17 min), and m/z induced fragmentation patterns (Airs & Keely, 2002; Airs et al., 2001; 655 (17–50 min), all with normal optimization. Auto MSn settings for Glaeser & Overmann, 2003) using atmospheric pressure chemical ioni- two precursor ions were an absolute threshold of 100,000 and a rela- zation (APCI) on an Agilent 6310 Ion Trap LC/MSn (positive ion mode, tive threshold of 5%. The most abundant ion was excluded after two 4000 nA corona current, 60 psi nebulizer pressure, 5 L/min drying gas measurements and released after 0.5 min. The acquisition parameter flow, 350°C drying temperature, and 400°C vaporizer temperature). had a fragment amplitude 1.0 V, with an average of 5.0 and an isola- Quantitation was based on light absorbance (Agilent 1200 MWD) using tion width of m/z 3.0. Auto MS (n > 2) settings with one precursor ion chromatographic peak areas determined at absorbance maxima (Fulton, were as follows: absolute threshold of 1000, relative threshold 5%, 2010). For the carotenoid isorenieratene, peak area was calculated at and fragment amplitude 1.0V. Compounds were identified by compar- 487 nm. For the tetrapyrrole pigments, absorbance was measured in the ison to published MS2 spectra and identification of the molecular ion

Qy band, at 658 nm for bacteriochlorophyll e (Bchl e), 665 nm for chlo- from Talbot et al., 2003, 2008. These molecular ions are as follows: rophyll a (Chl a) and pheophytin a (Phe a), and 746 nm for bacteriopheo- BHT (m/z 655), 2-Me BHT (m/z 669), Anhydro BHT (m/z 855), and phytin a (Bphe a). Response factors (peak area nmol−1) were calculated 2-Me Anhydro BHT (m/z 869). directly for Chl a and pyropheophytin a (Pphe a) by HPLC analysis of pure Standards consisted of individual BHP compounds from sam- standards. Response factors for Bchl e, isorenieratene, Bphe a, and Phe a ples collected at Bear Meadows Bog, Rothrock State Forest, PA. were calculated by comparison with those of Chl a and Pphe a, correcting Standards were fraction-collected following the initial reverse-phase for differences in absorbance using ratios of published molar extinction separation. Each collected fraction was additionally purified via coefficients in acetone or hexane (Borrego, Arellano, Abella, Gillbro, & normal phase separation on a silica column (150 × 3.0 mm, Waters) Garcia-Gil, 1999; Britton, 1995; Coolen & Overmann, 1998; Pennington, using a isocratic flow of 80:30 hexanes:isopropyl alcohol (IPA) at Strain, Svec, & Katz, 1964; Watanabe et al., 1984). An additional correc- 1 ml/min. Fractions were then transferred, dried, and weighed using tion was made for each response factor using the ratio of absorbance for tin elemental analysis cups on a nanogram scale. Compounds col- each pigment in its corresponding HPLC eluent and the solvent in which lected for use as standards include BHT, and BHTriol cyclitol ether, its extinction coefficient was published, as measured on a wavelength- with the cycltiol ether used as a standard for Anhydro BHT and 2-Me scanning spectrophotometer (Fulton, 2010). Anhydro BHT. Using these, standard response curves were produced by analyzing 1 ng, 10 ng, 20 ng, 50 ng, 100 ng, and 200 ng of each BHP. Peak areas were normalized to the internal standard of preg- 2.4 | Hopanoid analyses nanediol to account for instrument drift. A linear regression line of Pinnacle mat samples were freeze-dried, homogenized, weighed, and the ratio of the known amounts against the peak area was produced lipids were extracted using a modified Bligh–Dyer method after Talbot, and used to quantify the amounts in samples. The percent error for Rohmer, and Farrimond (2007). Specifically, samples were submerged triplicate HPLC-MS quantitations of individual BHPs ranged be- in a monophasic solution of 4:10:5 water:methanol:dicholorometh tween 15% and 27%. ane, disrupted in a sonicator bath for 1 hr at 40°C, shaken at 200 rpm Confirmation of the methyl group positioning on C2 was per- for 1 hr, and centrifuged at 13,000 g for 15 min. The supernatant was formed by GC-MS analysis of isolated BHP compounds fraction removed and the extraction repeated twice. Dichloromethane (10 ml) collected as clean, separated peaks in repeated HPLC runs. After iso- and water (10 ml) were added to the supernatant to induce phase sep- lation of each structure, a microscale version of the Innes, Bishop, aration, and the organic phase was removed. The aqueous phase was Head, and Farrimond (1997) protocol, based on the earlier Rohmer, extracted twice more with dichloromethane (10 ml) and the pooled Bouvier-Nave, and Ourisson (1984) side-chain cleavage, was used to organic phases were dried under N2 and weighed. An aliquot of the convert the BHPs into their hopanol forms. In short, acetyl groups total lipid extract (TLE) was acetylated using 1:1 pyridine:acetic anhy- were reductively removed with lithium aluminum hydride. The result- dride at 60°C for 1 hr. A 500 ng sample of pregananediol was added ing polar BHP was reacted with periodic acid (1 ml of a THF-and- to the TLE as an internal standard before derivatization. water solution 8:1, THF:water ratio) to cleave the carbon–carbon Bacteriohopanpolyols (BHPs), like the pigments, were character- bond in vicinal diols to yield aldehydes. Finally, aldehydes were re- n ized by APCI LC/MS . Reversed-phase HPLC was performed using a duced to form alcohols with sodium borohydride (10 mg of NaBH4

Phenomenex Gemini C18 column (5 μm × 150 mm × 3.0 mm) and a in 1 ml of ethanol, shaken for 1 hr and quenched with 1 ml KH2PO4). 5 μm guard column with the same solid phase. Mobile phase flowed Before GC-MS analysis, hopanol structures were derivatized with at 0.5 ml/min and was composed of (A) water, (B) methanol, and (C) acetic anhydride and pyridine. An Agilent 6890 GC-MS with a 60 m, isopropyl alcohol (0 min A:B:C 10:90:0, with a ramp to 25 min 1:59:40 0.32 mm, 25um DB-5 column was used for analysis. The methyl po- and an isocratic flow to 45 min). sition was determined based on comparison of retention times and 788 | HAMILTON et al. elution order with known hopane and methylhopane structures aluminum foil. The contribution of Photosystem II-independent an- (Summons & Jahnke, 1990, 1992). oxygenic photo assimilation of inorganic carbon was determined in assays amended with the Photosystem II inhibitor 3-(3,4-dichlorphenyl)- 1,1-dimethylurea (DCMU) (Sigma-Aldrich, St Louis, MO) at 10 M final 2.5 | Phylogenetic analyses of hpnP μ concentration. Light and dark controls were amended with unlabeled

All amplicons generated from gDNA and cDNA resulted in a single hpnP NaHCO3 (Sigma-Aldrich, St Louis MO). All assays—biological, control, sequence. The nucleotide sequence was translated into the amino acid dark, DCMU-amended—were performed in triplicate. The vials were sequence and identified as HpnP using BLAST. HpnP homologs were incubated in situ at a depth of 10 m in the center of the sinkhole for detected in the IMG genomic databases by BLASTP (Altschul et al., 3 hr of sunlight. Following incubation, mat samples were treated with 1997). Geneious (Biomatters Limited, Newark, NJ) was utilized to align concentrated HCl (1 M) to remove excess carbonate, washed with

302 HpnP protein sequences from genomes via MUSCLE (Edgar, 2004). de-ionized H2O, and filtered through pre-combusted (12 h, 450°C) Redundancy in the alignments was reduced through the Decrease GF/F filters (0.3 μM pore size) (Sterlitech Corporation, Kent, WA) and Redundancy Program (http://web.expasy.org/decrease_redundancy/) dried (8 h, 60°C). Dried filters were weighed and placed into tin dishes, which resulted in a final alignment with 75 HpnP sequences includ- sealed, and analyzed via an elemental analyzer with an isotope ratio ing the sequence recovered from Little Salt Spring. A maximum likeli- mass spectrometer (EA-IRMS). hood tree was constructed by PhyML (Guindon et al., 2010) using the LG+gamma model, four gamma rate categories, 10 random starting 2.7 | GenBank accession numbers trees, NNI branch swapping, and substitution parameters estimated from the data. The resulting HpnP tree was viewed and edited using 16S rRNA gene sequences representing each operational taxonomic iTOL (http://itol.embl.de/) (Letunic & Bork, 2016). unit (OTU) recovered in this study were submitted to the GenBank, DDBJ, and EMBL databases and assigned the following accession numbers: KP728178-KP728230. The hpnP sequence was submitted 2.6 | Inorganic carbon photoassimilation to the GenBank, DDBJ, and EMBL databases and assigned accession To examine the ability of the red pinnacle mat to photoassimilate number KP728177. inorganic carbon, mat samples (~500 mg wet weight) collected at ~10 m were placed in pre-combusted (12 hr, 450°C) serum vials and 3 | RESULTS overlaid with source water (10 mL) collected from the same location as the mat, capped, and sealed. Assays were initiated by addition of 13 3.1 | Water column geochemistry NaH CO3 (Cambridge Isotope Laboratories, Inc., Andover MA) at 200 μM final concentration. To assess inorganic carbon assimilation Geochemical analyses showed that the Little Salt Spring water col- in the absence of light, a subset of vials were completely wrapped in umn is relatively homogeneous with depth regardless of sampling

(a) Salinity (b) (c) (d) 2.25 2.50 2.75 3.00 3.25 + D.O. (µM) NH (µM) H S (µM) -1 -2 pH 4 2 ORP (mV) PAR (µmols-s m ) 7.00 7.13 7.25 7.38 7.50 0.015.0 30.045.060.0 0.015.0 30.045.0 60.0 0.0 500 10001500 2000 0.0 0.0 0.0 0.0

10.0 10.0 10.0 10.0

20.0 20.0 20.0 20.0 Depth (m) Depth (m)

Depth (m) Depth (m) 2009 30.0 30.0 30.0 30.0 2012 2013

40.0 40.0 40.0 40.0

50.0 50.0 50.0 50.0 27.00 27.25 27.50 27.75 28.00 0.02.5 5.0 7.510.0

o 2– Temperature ( C) SO4 (mM)

FIGURE 1 Water column geochemistry of Little Salt Spring, a cover collapse sinkhole in Sarasota County, FL. (a) Salinity, pH, and temperature + 2− of Little Salt Spring measured during a bloom of the red pinnacle mat (October 2013). (b) Dissolved oxygen, NH4 , H2S, and SO4 in the Little Salt Spring water column measured during a bloom of the red pinnacle mat (October 2013). (c). Variation in the oxidation reduction potential of the sinkhole water column measured in 2009, 2012, and 2013. (d) Photosynthetically active radiation (PAR) measured in October of 2013. Red bar indicates approximate depth of sample collection and depth of incubation of the inorganic carbon photoassimilation assays [Colour figure can be viewed at wileyonlinelibrary.com] HAMILTON et al. | 789 time. In October 2013 during a full bloom of the red pinnacle mat, Red pinnacle mat clones specific conductivity (4.9 mS/cm) and sulfate (~5 mM) were con- (n = 138) stant with depth (Figure 1a). The pH ranged between 7.3 and 7.4 and the temperature ranged from 27.5 to 28.0°C, with the highest values in the uppermost 10 m (Figure 1a). The total dissolved sulfide Cyanobacteria + diverse SRB concentrations ranged from 7 to 55 μM and NH ranged from 6 to 4 (Deltaproteobacteria) 20 μM, with the lowest values observed at the surface (Figure 1b). Dissolved oxygen concentrations in the upper 10 cm of the Little Chlorobi Gammaproteobacteria Salt Spring water column are well below saturation (34 μM, approxi- Other mately 20% saturation) even in full sunlight, and declined with depth Uncultured groups to <3 μM (the detection limit of the sensor) (Figure 1b). Water column chemistry was similar in January 2009, June 2012, and October Bacteriodetes 2013 with two notable exceptions. First, H S was not detected 2 FIGURE 2 Phylogenetic distribution of 16S rRNA clones retrieved (<0.2 μM) in the water column during the 2009 and 2012 sampling from upper basin red pinnacle mat in January 2009, June 2012, and trips. Second, while ORP declined with depth from +90 mV at the October 2013. Nearly full-length sequences were assigned to taxa surface to −280 mV at 55 m (the maximum depth rating of the sen- based on comparison with reference sequences in the software program ARB. “Other” includes taxa representing <5.0% of the total sors) in October 2013, there was very little change in ORP with depth in 2009 and a decrease from ~−110 mV at the surface to −280 mV at 55 m was observed in 2012 (Figure 1c). Dissolved oxygen profiles Cyanobacteria capable of anoxygenic photosynthesis (Figure 3a). throughout the water column were consistent between sampling Cyano5 is most closely related to 16S rRNA gene sequences from times. Photosynthetically active radiation (PAR) was consistent be- Geitlerinema sp. PCC 7407 (CP003591) and Cyanothece sp. PCC 7425 tween sampling times and decreased by over an order of magnitude (KM109978) (94% sequence identity). Cyano5 shares 90% sequence −1 −2 −1 −2 in the first 5 m (~1890 μmols-s m at 0 m to ~270 μmols-s m ) identity with 16S rRNA gene sequences from Geitlerinema sp. 9228 −1 −2 and was at ~50 μmols-s m at the pinnacle mat surface (~8–10 m) (formerly Oscillatoria limnetica “Solar Lake”) and Phormidiaceae cyano- (Figure 1d). The distribution of the red pinnacle mat in the upper basin bacterium SAG 31.92 (formerly Microcoleus chthonoplastes strain 11), was most extensive in October 2013 and more limited at the January both of which are known to carry out anoxygenic photosynthesis. 2009 and June 2012 time points. Six OTUs affiliated with the Chlorobi were recovered from the red pinnacle mats. The majority of the sequences (GSB1, GSB2, and GSB3) were closely related to clones recovered from the shallow 3.2 | Microscopy and 16S rRNA cloning chemocline of Sawmill Sink in the Bahamas (Figure 3b) (accession Pinnacles in the mat range in height up to ~4 cm (Fig. S3A). The mat number FJ716314, FJ716319; FJ716321; Macalady, J. L. unpub- surface is composed of strongly autofluorescent, long thin filaments, lished data). These sequences are also closely related (98% sequence consistent with the morphology of filamentous Cyanobacteria (Fig. identity) to Chlorobium limicola. GSB5 was closely related to clones S3B). Bacterial clones retrieved from the red pinnacle mat in January recovered from meromictic Fayetteville Green Lake in upstate New 2009, June 2012, and October 2013 were dominated by relatives York, US (FJ437791, FJ437915) (Meyer et al., 2011). Clones repre- of filamentous Cyanobacteria in the order Oscillatoriales, the ob- senting the same populations of Cyanobacteria and Chlorobi (100% ligately anoxygenic phototrophic order Chlorobiales, and diverse sequence identity) were obtained from samples of red pinnacle mat Deltaproteobacteria affiliated with sulfate-reducing clades (Figure 2). from January 2009, June 2012, and October 2013, indicating that the We recovered roughly equal numbers of 16S rRNA clones affiliated main mat-forming taxa are constant over seasonal and interannual with Oscillatoriales and Chlorobiales. Eight unique sequences (opera- time scales. tional taxonomic units (OTUs) defined at 99%) of Oscillatoriales spp. were recovered from the pinnacle mats. The most abundant, Cyano1, 3.3 | Pigment analyses shared 99% sequence identity with 16S rRNA sequences recovered from a sulfur-rich spring on the shore of Lake Erie (FJ967916). Based Pigments in the red pinnacle mat included nine homologs of BChl e, on % sequence identity of 16S rRNA gene sequences, Cyano1 is isorenieretane, Chl a, Bphe a, and Phe a (Table 1). Similar diversity also closely related to Leptolyngya sp. CENA538 (KF246502) from in Bchl e structures was also found in the chemocline of a holomic- an alkaline saline lake, Geitlerinema sp. PCC 8501 (FM210758), and tic lake (Glaeser & Overmann, 2003). BChl e, the most abundant pig- Leptolyngbya laminosa (FM210757), isolated from Euganean thermal ment in Little Salt Spring, and the carotenoid isorenieratene are only muds. Cyano1 shares 92% sequence identity with the 16S rRNA gene known to occur in Chlorobi, which also produce a small amount of sequence of Leptolyngbya sp. FYG, a cone-forming Cyanobacteria Bphe a (Hauska, Schoedl, Remigy, & Tsiotis, 2001), as observed in the isolated from a hot spring in Yellowstone National Park FJ933259 Little Salt Spring mat. Chl a is the primary photosynthetic pigment of (Figure 3a) (Bosak, Liang, Sim, & Petroff, 2009). Another cyanobacterial Cyanobacteria and Phe a is its initial decomposition product in senes- OTU, Cyano5, branches within a clade containing other characterized cent cells. The high concentration of Phe a suggests that there was 790 | HAMILTON et al.

(a) 99 Leptolyngbya sp. C1 (KC182752) FIGURE 3 Neighbor-joining phylogenetic tree showing the Leptolyngbya sp. FYG (FJ933259) Biological soil crust clone (KC463200) placement of cyanobacterial (a) and Chlorobi (b) 16S rRNA clones Cf. Leptolyngbya sp. Greenland 9 (DQ431004) from the red mat among closely related sequences. Neighbor- Hawaiian lava cave clone (EF032779) Leptolyngbya sp. JS2 (JX524204) joining bootstrap values >85% based on 1000 samplings are Phormidesmis priestleyi ANT.L52.6 (AY493579) 91 Pseudanabaenaceae cyanobacterium CENA529 (KF246495) shown. Accession numbers are given in parentheses. Stars indicate Arthronema africanum SAG 12.89 (KM019974) sequences recovered in clone libraries from all 3 years (2009, 2012, Cf. Leptolyngbya sp. Greenland 10 (DQ431005) 100 LSS Cyano OTU1 and 2013). Orange dots indicate Cyanobacteria that are capable Lake Eric sulfur-rich spring clone (FJ967916) of performing anoxygenic photosynthesis (Cohen et al., 1975a,b; 99 Leptolyngbya sp. CENA538 (KF246502) Geitlerinema sp. PCC 8501 (FM210758) De Wit & van Germerden, 1987) [Colour figure can be viewed at Leptolyngbya laminosa (FM210757) Lake Taihu clone (HM151383) wileyonlinelibrary.com] 90 East Antarctic lake benthic clone (DQ181685) 92 Leptolyngbya frigida ANT.REIDJ.1 (AY493611) Pseudanabaena tremula UTCC 471 (AF218371) LSS Cyano OTU2 TABLE 1 Pigment composition for Little Salt Spring red pinnacle 100 Synechococcus elongatus PCC 6301 (KM019981) Synechococcus elongatus PCC 7942 (CP000100) mat Gloeothece sp. KO68DGA (AB067580) LSS Cyano OTU3 −1 LSS Cyano OTU4 Pigment nmol gdw Esterifying alcohol 93 Limnothrix redekei 2LT25S01 (FM177493) Stanieria cyanosphaera PCC 7437 (CP003653) Bchl ea 243.8 ± 23.2 Halomicronema sp. Cyano39 (DQ058860) Highborne Cay stromatolite clone (EU249119) b Bchl e1-3 161.3 ± 13.4 Farnesol Benthic clone (DQ786167) Calothrix sp. 96/26 LPP3 (KM019977) c Bchl e4-6 52.5 ± 8.6 Hexadecenol 89 Marine cyanobacteria clone (KF793929) Nodosilinea sp. CENA144 (KC695838) Bchl e d 30.0 ± 2.6 Hexadecanol Cyanothece sp. PCC 7425 (CP001344) 7-9 94 Acaryochloris marina strain MBIC11017 (NR_074407) Isorenieratane 39.3 ± 19.5 Aphanocapsa muscicola VP3-03 (FR798916) Thermosynechococcus elongatus BP-1 (NR_074328) Phormidiaceae cyanobacterium SAG 31.92(EF654088) Bphe a 0.8 ± 0.1 Phytol Microcoleus chthonoplastes PCC 7420 (AM709630) 86 Chl a 82.2 ± 8.8 Phytol 99 Endoevaportie clone (EF106448) 91 Gypsum-precipitating environment clone (EU687425) Geitlerinema sp. 9228 (U96443) Phe a 133.8 ± 49.7 Phytol 90 99 Geitlerinema sp. PCC 7407 (CP003591) Oscillatoria acuminata PCC 6304 (KM109978) Bchl, bacteriochlorophyll; Bphe, bacteriopheophytin; Chl, chlorophyll; Phe, Oscillatoriales cyanobacterium UVFP2 (AJ630648) East Antarctic lake benthic clone (DQ181675) pheophytin. 94 LSS Cyano OTU5 aCombined concentrations of nine homologs. Banana plantation clone (JX133481) b Gloeocapsa sp. PCC 7428 (NR_102460) 22% [Et,Et]; 35% [n-Pr,Et]; 43% [i-Bu,Et]. Microcoleus chthonoplastes PCC 7420 (AM709630) c26% [Et,Et]; 42% [n-Pr,Et]; 32% [i-Bu,Et]. 99 Kamptonema formosum PCC 6407 (AM398782) d 89 Oscillatoria sp. PCC 6506 (AY768397) 35% [Et,Et]; 37% [n-Pr,Et]; 28% [i-Bu,Et]. Oscillatoria nigro-viridis st. PCC 7112 (NR_102469) 99 Phormidium autumnale Ant-Ph68 (DQ493874) Lake Huron Genome Bin (FJ866618) 94 Uncultured clone (JF733394) abundant organic matter accumulated from dead Cyanobacteria, from Biological soil crust clone (KC463193) populations either in the water column or mat. The pigment results 85 Thermal spring clone (DQ471447) Biological soil crust clone (JQ769661) are consistent with the clone libraries we obtained from the red mat, 94 Terrestrial sulfidic spring clone (JX521366) 100 Leptolyngbya antarctica ANT.BFI.1 (AY493590) in which 16S rRNA sequences related to Chlorobium spp. (green sul- LSS Cyano OTU6 100 LSS Cyano OTU7 fur bacteria) and Cyanobacteria were the dominant taxa recovered LSS Cyano OTU8 100 Clostridium clariflavum DSM 19732 (NC_016627) (Figure 2). Desulfotomaculum carboxydivorans CO-1-SRB (NC_015565) 0.05

88 Sawmill Sink clone (FJ716314) (b) Sawmill Sink clone (FJ716319) 3.4 | Hopanoid analyses 97 Sawmill Sink clone (FJ716321) 95 LSS GSB OTU1 LSS GSB OTU2 Multiple bacteriohopanepolyols (BHPs) were recovered from the red 96 Lake Kinneret sediment clone (AM086114) LSS GSB OTU3 pinnacle mat (Table S1). The pinnacle mat samples contain bacterioho- Wastewater bioreactor clone (HQ602909) 91 Chlorobium limicola DSM 245 (CP001097) panetetrol (BHT) (m/z 655), 2Me-BHT (m/z 669), anhydro-BHT (m/z 100 92 Chlorobium limicola strain E2P2 (AM050126) Chlorobium phaeobacteroides strain Glu (AM050128) 855), and 2-Me-anhydro-BHT (Table S1, Fig. S4). The red pinnacle mat LSS GSB OTU4 contains a small proportion of BHT and 2 Me-BHT and large propor- 100 Fayetteville Green Lake clone (FJ437781) Fayetteville Green Lake clone (FJ437915) tions of anhydro-BHT and 2Me-anhydro-BHT (Table S1; Fig. S4). BHT 93 Chlorobium phaeovibrioides strain DSM 269 (Y10654) Sawmill Sink clone (FJ716300) 100 and Me-BHT were detected at 0.10 μg/mg total lipid extract in the 100 Sawmill Sink clone (FJ716285) Biological sulfide-removal reactor clone (DQ383305) red mat. Anhydro-BHT and 2-Me-anhydro-BHT were present in much Pelodictyon luteolum (AM050131) Prosthecochloris vibrioformis (Y10648) larger amounts, 117 μg/mg TLE and 176 μg/mg TLE, respectively. The LSS GSB OTU5 99 100 LSS GSB OTU6 percent error for triplicate HPLC-MS quantitations of individual BHPs 92 Pond sediment clone (JF428832) 100 Environmental clone (HM243993) ranged between 15% and 27%. Shule River clone (JQ978595) Environmental clone (GQ342332) Lower Kane Cave clone(AM490708) 93 Soil clone (EU043618) 100 Chloroflexus aurantiacus strain J-10-fl (NR_074263) 3.5 | Phylogenetic analyses of HpnP Roseiflexus sp. RS-1 (NR_074197) 0.05 PCR amplification of hpnP, a gene encoding an enzyme respon- sible for adding the C-2 methyl to the hopanoid ring structure HAMILTON et al. | 791

8 Methylocystis parvus OBBP

Tardiphaga sp. OK246 Methylocystis sp. Rockw ) 1 ) RHD008 U Hyphomicrobium facile DSM 1565

Afipia sp. P52-10 Rhodovulum sp.

Methylocapsa aureae KYG Methylocapsa acidiphilall B2 Bradyrhizobium sp. (1 ) opseudomonas sp. (5 PH10 Afipia sp. (4 Methylocapsa palsarum NE

Nitrobacter winogradskyi Nb-255 Rhod Methylocella silvestris BL2 Rhodospirillales bacterium

T ) Methyloferula stellata 2 Methylobacterium sp. (6 Beijerinckia mobilis UQM 1969 AR4

Beijerinckia indica ATCC 9039 Streptomyces viridochromogenes

Rhodoplanes sp. Z2-YC6860 n345

Acidobacteria bacterium Elli

imnion genome Trout Bog Hypol

Cyanothece sp. PCC 7425 ThermosySynechococcus sp. PCC 6312 Synechoco nechococcu s sp ccus sp. CAUP . NK55a ng HpnP A 6 F 1101 Little Salt Spri ischerella sp. PCC 9605 CC 7310 Fischerella muscicola PCC 7414 P To Mastigocladopsis repens MORA Has Chlorogl 2 1 Tolypothrix bouteillNost 06 lypothrix campylonemoides VB51 Nostoc piscinales CENA21Chlorogloeopsis sp. PCC 77 Microcoleus sp. PCC 71 allia by Gloeocapsa sp. e 4 d o Gloeobacter kilaueensis JS1 c spoeop. Calothrix desertica PCCssoidea 7102 VB51217 4 sp Gloeobacterlandica violaceus PCC PCC90 7421 8 sis l 2 fritschi 0 Oscillatoriales sp. JSC-1 2 i PCC 69 e 0 ochlorothrix ho i Iicb Pr Cyanobacterium sp. ESFC- ma tolypothrichoi

1 12

Leptolyngbya sp. NIES-210 13 Cyanothece sp. PCC 7822 Leptolyngbya sp. PCC 6306 cytone

Cyanothece sp. PCC 742

288

Geminocystis herdmanii PCC 630

Cyanobacterium sp. PCC 10605

FIGURE 4 Unrooted phylogenetic tree of HpnP. Cyanobacterial HpnP branching is shown in green with the translated hpnP transcript recovered from Little Salt Spring indicated by a black rectangle. The scale bar represents 0.1 substitutions per nucleotide site [Colour figure can be viewed at wileyonlinelibrary.com]

(Welander, Coleman, Sessions, Summons, & Newman, 2010), was Salt Spring HpnP groups with other cyanobacterial HpnP sequences successful for both gDNA and cDNA templates derived from the (Figure 4). red mat. All amplicons were identical. The translated partial HpnP sequence from Little Salt Spring was 78% identical to the Nostoc 3.6 | Inorganic carbon photoassimilation punctiforme PCC 73102 HpnP and 62% identical to the HpnP from Rhodopseudomonas palustris TIE-1, the organism in which the radical Aliquots of the red pinnacle mat assimilated more inorganic carbon in −1 −1 SAM methylase enzyme HpnP was first described (Welander et al., the presence of light (0.70 ± 0.03 μmol C mg dry weight hr ) than −1 −1 2010). Phylogenetic analysis of the HpnP sequence from Little Salt in the dark (0.48 ± 0.0.05 μmol C mg dry weight hr ) (Figure 5). In Spring with characterized HpnP sequences indicates that the Little the presence of the Photosystem II inhibitor DCMU, the mats still 792 | HAMILTON et al.

0.80 data (Planavsky et al., 2014b) and a lack of terrestrial Mn oxidation in mid-Proterozoic paleosols (Zbinden, Holland, Feakes, & Dobos, 1988) are consistent with low atmospheric oxygen concentrations ) throughout this period. –1 0.60 In the water column of Little Salt Spring, the sulfate concentration (5 mM) is low compared to current marine levels (28 mM), and near the expected concentration of sulfate in mid-Proterozoic

sample hr 0.40 oceans (Canfield, 1998; Planavsky, Bekker, Hofmann, Owens, & –1 Lyons, 2012). Dissolved oxygen concentrations in the upper 10 cm of the Little Salt Spring water column are below saturation (34 μM, C assim/g sample

13 approximately 20% saturation) even in full sunlight, and decline 0.20

( µ mol C mg with depth to <10 μM at 50 m depth. The entire water column is sulfidic (~50 μM) (Figure 1; Fig. S1). Little Salt Spring thus provides a Proterozoic Ocean analog environment, poised between oxic and 0.00 sulfidic conditions. Data collected over a 4-year period indicate that while conditions fluctuate within limits, fully oxic, or fully sulfidic Dark Light conditions do not develop in the spring. The cause of this apparent DCMU redox stability is not known, but likely involves feedbacks between Dark Control Light Control the hydrologic system of the spring and the biologic system. Similar FIGURE 5 Inorganic carbon assimilation by the red pinnacle mats. feedback may have acted to stabilize low-oxygen conditions in the Controls were amended with unlabeled substrate. Error bars are from Proterozoic oceans. triplicate incubations PAR levels at Little Salt Spring decrease with depth by an order of magnitude in the first ~5 m (Figure 1c). Although these conditions incorporated more inorganic carbon (0.11 ± 0.0.01 μmol C mg dry w would challenge many phototrophs, the spring hosts a pinnacle mat eight−1 hr−1) than those incubated in the dark. that is comprised largely of photoautotrophs related to Oscillatoriales- group Cyanobacteria and Chlorobi. Many Oscillatoriales spp. synthesize phycoerythrin (red pigment) and thrive at deep chlorophyll 4 | DISCUSSION maxima in stratified pelagic systems. This chromatic niche is gener- ally located below the thermocline, where light intensities are low The protracted delay between the initial rise of oxygen and the sec- but nutrient concentrations are high. Cyanobacteria are metabolically ond rise of oxygen ~0.5 Gyr ago, when oxygen rose to present-day diverse and include species that perform anoxygenic photosynthesis levels, is the subject of ongoing debate. It is clear from geologic evi- in the presence of sulfide (Cohen, Padan, & Shilo, 1975a; Jørgensen, dence that the evolution of oxygenic photosynthesis predates the Cohen, & Revsbech, 1986). The Little Salt Spring Cyanobacteria are GOE. For instance, the enrichment of Mo and Re in organic-matter- distantly related to other Cyanobacteria that are capable of perform- rich shales dated to 2.5 Gyr led to the idea that “whiffs” of O2 accu- ing anoxygenic photosynthesis (Figure 3a), including Geitlerinema sp. mulation occurred before the GOE (Anbar et al., 2007); Mo isotopes PCC 9228 (formerly Oscillatoria limnetica) (Cohen, Jørgensen, Padan, & as proxies for manganese oxides date oxygen production to ~3 Gyr Shilo, 1975b), mat-forming Phormidiaceae cyanobacterium SAG 31.92 (Planavsky et al., 2014a); and Cr isotopes and redox-sensitive metals (formerly Microcoleus chthonoplastes strain 11) from Solar Lake, Sinai in rocks from the Pongola Supergroup in South Africa indicate the (Jørgensen et al., 1986), and Phormidium autumnale-like strains domi- presence of atmospheric oxygen ~3 Gyr (Crowe et al., 2013). The nating purple mats in sunlit sinkholes in Lake Huron (Nold et al., 2010; disappearance of mass-independent sulfur isotope fractionation in Voorhies et al., 2012). The Phormidiaceae cyanobacterium SAG 31.92 mineral sulfides and sulfates provides the most convincing evidence dominated Solar Lake mats and cyanobacterial mats at the Frasassi of the rise in oxygen ~2.5 Gyr ago (Bekker et al., 2004; Farquhar, sulfidic springs (Hausler, 2010) carry out anoxygenic and oxygenic Bao, & Thiemens, 2000) and there is strong evidence that diverse photosynthesis concurrently. microbial metabolisms existed at this time (David & Alm, 2011). Recently, Johnston, Wolfe-Simon, Pearson, and Knoll (2009) pro- Therefore, it is reasonable to hypothesize that the delayed second posed that contributions to primary production by anoxygenic pho- rise of oxygen involves feedbacks between early Earth microbiota totrophs effectively limited oxygen production throughout Earth’s and elemental cycles. Multiple feedbacks that would act to destabi- middle age. This model requires a flux of sulfide to the photic zone lize Proterozoic-like (sulfidic, low-oxygen) conditions in the world’s sufficient to support anoxygenic photosynthesis. In Little Salt Spring, oceans have been identified (Lyons and Gill, 2010). These nega- we observed sulfide conditions throughout the water column suggest- tive feedback must not have been strong since sulfidic, low-oxygen ing the flux of upwelling of sulfide-rich water could exceed the down- conditions in the oceans appear to have been common for 2 bil- ward mixing of oxygen, similar to Proterozoic ocean models (Meyer & lion years after the GOE (Catling & Buick, 2006). Chromium isotope Kump, 2008). This observation is supported by low levels of dissolved HAMILTON et al. | 793 oxygen throughout the upper water column—7–15 μM in the present in Cyanobacteria is a dynamic trait that can be gained or lost, in- study and hypoxic below 3 m in previous investigations (Clausen et al., cluding the variable tolerance of photosystem II to sulfide (Miller & 1979). Furthermore, in the absence of ample wind-mixing, sulfide Bebout, 2004). Clearly, more information regarding the physiology could reach the surface due to an upward sulfide flux that exceeds of the Little Salt Spring Cyanobacteria and Chlorobi are necessary to wind-mixed oxygen flux (Kump, Pavlov, & Arthur, 2005). The small di- predict which factors control the proportion of oxygenic and anoxy- ameter (~78 m) of Little Salt Spring and the surrounding landscape of genic photosynthesis in this system. Additional information regarding mature trees and dense shrubs suggest little wind mixing. Regardless, the daily and seasonal dynamics of oxygen and sulfide in the water the combination of mixing with the atmosphere and net oxygen pro- column and the source of sulfide to the pinnacle mats will further duction from both the red pinnacle mats and the dense layer of chara inform the redox balance within the mat ecosystem and surrounding (eukaryotic algae) observed down to ~5 m is not sufficient to maintain water column. a fully oxygenated water column in the upper basin. In our study, red pinnacle mat samples (collected from ~8.5 m) incubated in the pres- 4.1 | Hopanoids as biomarkers ence of DCMU assimilated more inorganic carbon in the light than those incubated in the dark (Figure 5). Since DCMU blocks electron Hopanoids, which originate almost exclusively from bacteria, have transfer to Photosystem II in Cyanobacteria, our experiment shows been found throughout the geologic record where thermal history al- that the mat photoassimilates a large percentage of inorganic carbon lows (French et al., 2015). Increasing numbers of studies that detect via anoxygenic photosynthesis. Although Chlorobi likely account for hopanoids in extant organisms and modern environments raise the some of the anoxygenic photosynthetic activity, the dominance of fil- expectation that they may aid in interpreting the rock record with re- amentous Cyanobacteria in the mats at Little Salt Spring where the spect to geochemical conditions or taxonomic composition of micro- water column is sulfidic and anoxic strongly suggests that they also bial communities present at the time of deposition. The majority of can carry out anoxygenic photosynthesis. Two additional lines of evi- sedimentary 2-methylhopanes were thought to derive from ancient dence support a role for cyanobacterial anoxygenic photosynthesis in shallow marine environments (Eigenbrode, Freeman, & Summons, situ: (i) characterization of photosynthesis in situ indicates oxygenic 2008; Waldbauer, Sherman, Sumner, & Summons, 2009), where photosynthesis occurs for only a few hours a day, whereas anoxygenic phototrophs would have been dominant populations; however, con- photosynthesis occurs throughout the photoperiod (de Beer et al., tamination is the likely source of 2-methylhopanes in these studies 2017); and (ii) the dominant Cyanobacteria from the red pinnacle mat (French et al., 2015). Regardless, hopanoid production is not observed is capable of performing anoxygenic photosynthesis (Hamilton, T. L., in pure cultures of the most abundant modern marine Cyanobacteria Klatt, J. M., de Beer, D., Macalady, J. L., unpublished data). In either (Sáenz, Eglinton, Waterbury, & Summons, 2012; Talbot et al., 2008). case, the relative proportion of oxygenic (Cyanobacteria) and anoxy- Furthermore, most of the sequenced Cyanobacteria that contain the genic (Chlorobi ± Cyanobacteria) photosynthesis being carried out in hpnP methylase gene were not isolated from marine environments the mat has strong implications for the redox balance within the mat (Welander et al., 2010; Figure 4). C-2 methylated hopanoids are, ecosystem and surrounding water column. however, common in environmental samples from certain modern In sunlit environments where low oxygen concentrations and sul- marine environments (Talbot et al., 2008) including Shark Bay stro- fide are present, some Cyanobacteria can use sulfide as the electron matolites, where the majority of partial hpnP sequences recovered donor to photosystem I in the absence of oxygen generation (Cohen were most closely related to Cyanobacteria (Garby, Walter, Larkum, et al., 1975a,b; Garcia-Pichel & Castenholz, 1990; Padan, 1979). In & Neilan, 2013). In the moderately saline waters of Little Salt Spring these cases, photosystem II-independent photoassimilation of CO2 (~3 ppt), the red mat is a source of BHPs and 2-MeBHPs (anhydro- (anoxygenic photosynthesis) occurs with the same efficiency as ox- BHT and 2-methylanhydro-BHT) (Table S1; Fig. S4). The amount of ygenic photosynthesis (Oren, Padan, & Avron, 1977; Oren & Paden, 2-MeBHP observed in the biofilm is exceptional (176 μg/mg TLE 1978). In a few rare cases, Cyanobacteria capable of performing both 2-Me-anhydro-BHT from the red pinnacle mat), and nearly 30 times oxygenic and anoxygenic photosynthesis simultaneously have been that reported for other environments. For instance, only low amounts documented (De Wit & van Gemerden, 1987; Klatt et al., 2015). of 2-MeBHPs (0.5–6 μg/mg TLE for 2-methyl BHPs) were recovered Sulfide is the preferred substrate of Chlorobi and these organisms from microbial mats from Yellowstone National Park (Jahnke et al., typically have a high affinity for sulfide (Brune, 1995; Frigaard & 2004; Summons et al., 1999; Talbot et al., 2008), hypersaline lakes Bryant, 2008). In contrast, a low affinity for sulfide has been observed (Blumberg et al., 2013), Antarctica (Talbot et al., 2008), and stroma- in anoyxgenic photosynthetic Cyanobacteria compared to Chlorobi. tolites and mats in the Tswaing Crater of South Africa (Talbot et al., These observations suggest that facultative anoxygenic photosyn- 2008). Both anhydro-BHT and 2-methylanhydro-BHT have been pre- thesis among Cyanobacteria may not be a competitive strategy where viously thought to be diagenetic products of BHT, BHT-cyclitol, or these species co-exist and sulfide is limiting (De Wit & van Germerden, BHT-glucosamine (with 2-Me-anhydro-BHT being formed from the 1987; Garcia-Pichel & Castenholz, 1990). 2-Me equivalents) or an analytical artifact (Bednarczyk et al., 2005; In Cyanobacteria, Photosystem II can also be inhibited by sul- Talbot et al., 2005; Handley, Talbot, Cooke, Anderson, & Wagner, fide. Taxonomically diverse Cyanobacteria display a range of sulfide 2010; Eickhoff, Birgel, Talbot, Peckmann, & Kappler, 2014). A bio- tolerances and several lines of evidence suggest sulfide tolerance logic source of anhydro-BHT and 2-methylanhydro-BHT has yet to be 794 | HAMILTON et al. demonstrated and our data cannot discern between early diagenesis the ecologic factors that promote the co-existence of oxygenic and in the mat or analytical artifact; however, we have recovered BHT and anoxygenic phototrophs in the mat; and (iii) describing the timescale 2-MeAnhydroBHT from pure cultures of the abundant red pinnacle and chemical changes associated with production and diagenesis of mat cyanobacterium (Bird, 2016; Hamilton et al., 2013a). According organic biomarkers. to the processes forming 2-MeAnhydroBHT from BHT outlined by Schaeffer, Schmitt, Adam, and Rohmer (2008, 2010), 2-MeBHT (the potential source of 2-MeAnhydroBHT) should also be present; how- ACKNOWLEDGMENTS ever, we have not detected AnhydroBHT or 2-MeBHT in the pure Sampling at Little Salt Spring was carried out in cooperation with culture. Regardless, elevated levels of hopanoids, particularly those J. Gifford (U. Miami/RSMAS) and divers S. Koski (U. Miami/RSMAS), methylated at the C-2 position, and the recovery of a cyanobacterial R. Riera-Gomez (U. Miami/RSMAS), and C. Coy (Florida Aquarium). hpnP transcript from the red pinnacle mat support a cyanobacterial We thank S. Koski for help with field operations, and K. Dawson, source of these lipids in situ. D. Jones, and D. Tobler for geochemical analyses. We are grateful to The recovery of a single hpnP transcript most closely related to D. Walizer for invaluable technical assistance and S. Lincoln for in- cyanobacterial hpnP genes (Figure 4) strongly indicates the source of sightful discussions and detailed assistance with the hopanoid data. 2-MeBHPs in the Little Salt Spring pinnacle mat is cyanobacterial. The We are grateful to S. Kopf and an anonymous reviewer for their con- synthesis of 2-Me-BHPs by Cyanobacteria under low oxygen condi- structive suggestions. This project was funded by the National Science tions has important implications for the interpretation of these struc- Foundation (NSF EAR-0525503 to J.L.M.), the NASA Astrobiology tures when they are found in the rock record, especially in Precambrian Institute (PSARC, NNA04CC06A to J.L.M. and K.H.F.), and the PSU rocks. The purple cyanobacterial mats in sunlit sinkholes of Lake Huron, Science Diving Program. Work at MIT was supported by the NASA which are dominated by relatives of Phormidium autumnale recovered Astrobiology Institute (NNA13AA90A) Foundations of Complex Life, from Arctic and Antarctic environments (Nold et al., 2010), do not ap- Evolution, Preservation, and Detection on Earth and Beyond to R.E.S. pear to synthesize 2Me-BHPs (Voorhies et al., 2012). In contrast, cya- T.L.H. graciously acknowledges support from the NASA Astrobiology nobacterial mats growing under low light beneath permanent ice cover Institute Postdoctoral Program and the University of Cincinnati. in Antarctic Lake Vanda appear to produce abundant 2-MeBHP (Matys et al., 2015). These results, combined with the Little Salt Spring data presented here, demonstrate the need for further studies of model or- REFERENCES ganisms representative of early-evolving Cyanobacteria to understand the physiologic role of 2-MeBHPs in these organisms and shed light on Airs, R. L., Atkinson, J. E., & Keely, B. J. (2001). 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