The Oman Drilling Project established an “Active Alteration” multi-borehole observatory in peridotites undergoing low-temperature serpentinization in the Samail Ophiolite. The highly serpentinized rocks are in contact with strongly reducing fluids. Distinct hydrological regimes, governed by differences in rock porosity and fracture density, give rise to steep redox (Eh +200 to −750 mV) and pH (pH range 8.5–11.2) gradients within the 300–400 m deep boreholes. The serpentinites and fluids host an active subsurface ecosystem. Microbial cell abundances in serpentinite vary at least six orders of magnitude, from ≤3.5 × 10¹ to 2.9 × 10⁷ cells/g. Low levels of biological sulfate reduction (2–1,000 fmol/cm³/day) can be detected in rock cores, particularly in rocks in contact with reduced groundwaters with pH < 10.5. Thermodesulfovibrio is the predominant sulfate reducer identified via metagenomic sequencing of adjacent groundwater communities. We infer that transport and reaction of microbially generated sulfide with the serpentine and brucite assemblages gives rise to optical darkening and sulfide overprinting, including the formation of tochilinite-vallerite group minerals, potentially serving as an indicator that this system is inhabited by microbial life. Olivine mesh-cores replaced with ferroan brucite and minor awaruite, abundant veins containing hydroandradite garnet and polyhedral serpentine, and late-stage carbonate veins are suggested as targets for future spatially resolved life-detection investigations. The high-quality whole-round core samples that have been preserved can be further probed to define how life distributes itself and functions within a system where chemical disequilibria are sustained by low-temperature water/rock interaction, and how biosignatures of in situ microbial activity are generated.

Plain Language Summary

Ultramafic rocks undergoing water/rock interaction, and storing fluids that are far from chemical equilibrium, may be one of the most common habitats in our solar system. Through the Oman Drilling Project we collected >1 km of intact serpentinite in contact with groundwaters. These cores capture parts of the rock-hosted biosphere and show how cells are distributed within serpentinites that vary in their mineralogical, physical and chemical properties. The cores are also biologically active, enabling us to detect specific metabolisms, such as when microorganisms combine hydrogen as reductant and sulfate as an oxidant to fuel their metabolism. Although the distribution of microbial cells in the rock cores is very heterogeneous, there are many intervals where the abundance of cells constitutes robust biomass. In the deeper cores, slow, albeit detectable, microbial sulfate reduction proceeds. We suggest that this pervasive biological activity releases byproducts such as sulfide that can react with the serpentinite and change the optical and chemical properties of the rocks. The feedbacks between the rock alteration and microbial activity produce markers that enable us to focus our search for rock-hosted life and any specific biosignatures it may produce on Earth and perhaps on other planetary bodies.

Key Points

Highly serpentinized subsurface rocks exhibit steep redox gradients and host microbial cell abundances that vary >6 orders of magnitude
Low rates of microbial sulfate reduction in rock cores are inferred to result in optical darkening and sulfide overprinting of the mineralogy
Widespread andradite garnet, abundant ferroan brucite, and rare carbonate are targets for future spatially resolved life-detection efforts

1 Introduction

1.1 Interest in the Biology of Low-Temperature Serpentinizing Systems

Serpentinization, a geochemical process that occurs during the hydration of ultramafic rocks, is of biological interest for its potential to support microbial communities through available chemical energy (i.e., chemosynthesis), independent of the input of sunlight energy and photosynthetic by-products. The oxidation of Fe(II) derived from olivine and pyroxene in ultramafic rocks releases enormous amounts of reducing power, as well as a diversity of secondary minerals dominated by serpentine (Andreani et al., 2013; McCollom & Bach, 2009). The generation of aqueous reductants (e.g., H₂ from reduction of water, or organic acids from reduction of CO₂) could power life serpentinite rocks, if sufficient oxidants (e.g., O₂, nitrate, sulfate, CO₂) are delivered through time. Serpentinizing systems may have sustained some of the earliest life on Earth, and may have promoted life's emergence, through the evolution of H₂-driven metabolisms such as methanogenesis (Boyd et al., 2020; Colman et al., 2017; Nealson et al., 2005; Russell et al., 2014; Schulte et al., 2006; Sleep et al., 2011; Spear et al., 2005). Moreover, serpentinizing systems may be one of the most common potential habitats beyond Earth, such as on Mars, Europa, Enceladus and other rocky bodies (Ehlmann et al., 2010; McKay et al., 2014; Schulte et al., 2006; Vance & Melwani Daswani, 2020). However, we need to better assess how life distributes itself within an actively serpentinizing system, and identify the best evidence of biological activity that can be preserved from fractured rock ecosystems (Hays et al., 2017). Such information would inform life-detection strategies in Earth's serpentinite-hosted environments and would provide a roadmap for analysis of materials that may be sampled during future landed missions to Mars, Europa or Enceladus.

Local conditions in any serpentinizing system span highly variable water/rock ratios and fluid mixing regimes, giving rise to a spectrum of chemical environments and disequilibria that could sustain biological activity. Recent information about serpentinite-adapted life has been derived from intensive geochemical and microbiological analysis of surface fluids discharged from the subsurface serpentinites along fractures and faults in the Cedars, Del Puerto, Chimaera, Tablelands, Cabeço de Vide, Liguria, and Zambales ophiolites (Blank et al., 2009; Brazelton et al., 2012, 2017; Morrill et al., 2013, 2014; Suzuki et al., 2013; Szponar et al., 2013; Tiago & Veríssimo, 2013; Woycheese et al., 2015). Substantive insights into the potential microbial community structure and function of the serpentinite-hosted biosphere has also been recently derived from analysis of deeper fluids and rocks recovered from the Atlantis Massif, California Coast Range Microbial Observatory, Leka ophiolite, and Oman (Crespo-Medina et al., 2014; Daae et al., 2013; Fones et al., 2019, 2020; Früh-Green et al., 2018; Kraus et al., 2020; Miller et al., 2016; Motamedi et al., 2020; Quéméneur et al., 2019; Rempfert et al., 2017; Sabuda et al., 2020; Seyler et al., 2020). However, subsurface serpentinizing rocks and mineralized fractures have not yet been comprehensively examined to determine how the distribution and activity of the rock-hosted life is structured and functions. An observatory located within a massive section of uplifted upper mantle rocks that currently store water can function as an excellent model environment for better understanding the role of low-temperature water-rock reactions in shaping the serpentinite-hosted biosphere on and beyond Earth. Therefore, microbiological investigations became an integral part of the International Continental Scientific Drilling Program “Oman Drilling Project,” where multiple boreholes were drilled beneath the water table within dunite and harzburgite lithologies, and >1 km of high-quality core was successfully recovered from rocks undergoing active serpentinization in the Samail Ophiolite, Sultanate of Oman.

1.2 Oman Drilling Project Phase II at the “Active Alteration” Multi-Borehole Observatory

The Oman Drilling Project (OmanDP) established a “Active Alteration” Multi-Borehole Observatory (MBO) in the Wadi Tayin massif of the Samail Ophiolite in 2018 (Kelemen et al., 2013, 2020) (Figure 1). The dunite and harzburgite lithologies are actively undergoing hydration and carbonation at temperatures <50°C, and therefore provide an excellent opportunity for unraveling successive stages of water/rock reaction at low-temperatures, and deciphering how the rock alteration and aqueous geochemistry is coupled to microbial activity. Specifically, whole-round core samples from numerous subsurface geochemical regimes were sought in order to detect and characterize the endolithic microbial communities and to identify the geological and geochemical controls on cell distributions and activity. We focus here on three 4″ diameter boreholes labeled “BA1B,” “BA3A,” and “BA4A” that were wireline core drilled between January and March 2018. We place the initial analysis of biological sub-cores within the context of the physical and mineralogical properties of the cores described by the Oman Drilling Project Science Party (Kelemen et al., 2020). We provide information on how rock core samples were collected for biological investigations, how the mineralogy of the core subsamples varies, and how cell abundances vary from hole to hole. The cores preserved for biological experiments are now being used in myriad ways, in particular to determine how Fe, C, N, and S cycling are intimately connected under distinct geochemical and mineralogical regimes. Here, we share the first activity measurements testing whether or not biological sulfate-reduction was occurring in OmanDP core samples, following on from recent studies that have highlighted the importance of biological S dynamics in terrestrial serpentinites such as the California Coast Range Microbial Observatory (Sabuda et al., 2020) and the Oman ophiolite (Glombitza et al., 2021). Altogether, the study presented herein provides a first snapshot of the structure of this inhabited serpentinite-biosphere, and establishes an essential framework for investigations that will characterize the active versus fossil subsurface microbial community structure, function and biosignature preservation potential.

Details are in the caption following the image — **Figure 1**
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Field context of the Oman Drilling Project “Active Alteration BA” multi-borehole observatory in Wadi Lawayani, Northern Sharqiyah, in the Samail Ophiolite, Oman. GPS coordinates: BA4A, site 10A (22°53.101′ 58°41.703′), BA1B, site 5A (22°52.874′ 58°42.035′), BA3A, site 9B (22°51.970′ 58°42.634′). A site map placed in regional context can be found at https://omandrilling.ac.uk/drilling program. A map showing the relationships of BA holes to nearby wells can be found in Figure S5.

2 Materials and Methods

2.1 Core Procedures at the “Active Alteration” Multi-Borehole Observatory

In order to obtain samples suitable for biological analysis and experimentation from the Oman Drilling Project (OmanDP) Phase II drilling at the “Active Alteration” MBO sites in Wadi Lawayni, we developed several drilling and core-handling procedures that could be integrated into the core-flow. The lubricant for the wireline diamond core drilling was EZ-Mud® (Haliburton), a polyacrylamide/polyacrylamate borehole stabilizer, which was mixed with drinking-water that was delivered to the site in water-tanker trucks each day. We did not have the ability to detect perfluorocarbon tracers on site (French et al., 2015; Lever et al., 2006; Orcutt et al., 2017; Smith et al., 2000), so we chose to instead mix a non-toxic fluorescent “Invisible Blue®” tracer (DayGlo Color Corp, pigment SPL-594N) into the drilling fluids that would not be sensitive to pH or silicate adsorption. The “Invisible Blue®” tracer is a DayGlo® pigment with blue fluorescence under UV excitation and particle sizes ranging from 0.25 to 0.45 µm (Friese et al., 2017). The initial particle concentration of ∼1 × 10¹⁵ particles/L was diluted into the drilling mud at an estimated ratio of 1:10,000 to target an average concentration of 10⁸ particles/mL. When drilling fluids were lost into the formation and needed to be replenished by mixing more EZmud® and drinking water, additional “Invisible Blue®” pigment was added to restore the mud to the target tracer concentration.

During the drilling of holes BA3A, BA1B, and BA4A, excellent core recovery (100%) was achieved, with numerous sections recovered as intact or cleanly fractured 3 m lengths. A 50 cm section of whole-rock core was identified for microbiological (“BIO”) studies at approximately 10 m intervals. Upon recovery of a full 3 m section of core that would be subsampled for BIO, the core was scrubbed and washed along the outside with ultrapure deionized water. The 3 m core was then laid onto combusted carbon-free foil inside the core liner for the initial photography and description, and kept within the initial core flow established by the Oman Drilling Project (Kelemen et al., 2020). The core was kept on combusted foil during all stages of preparation except for when it was placed on a core scanner, whose surface was pre-cleaned with bleach and ethanol. After rubbing the exterior surface of the core again with an ethanol wipe, a specific 50 cm section was separated out for BIO investigations using a diamond blade saw lubricated with deionized water to remove it from the 3 m core length. This 50 cm section was held in combusted foil and moved into a BIO trailer on site.

The BIO trailer was a small enclosure on the drill site that contained an open workbench, a deionized water system producing 18.2 MΩ cm, 0.2 μm filter-sterilized water, a HEPA-filtered and UV sterilized laminar flow hood, and a COY® anaerobic glovebox purged with nitrogen gas. Once inside the BIO trailer, the 50 cm BIO subcore was subsampled using an ethanol-washed hammer, chisel and tweezers, while resting on combusted foil. Discrete 5–15 cm sections were separated for DNA, lipid, biological activity and mineralogy analyses. Sub-cores designated for DNA or lipids studies were transferred to the laminar flow hood, washed twice more with deionized water, and wrapped in a combusted carbon-free foil and then placed in a Teflon® sleeve. These cores were then transferred into the N₂ glovebox, sealed into mylar bags flushed with nitrogen, and frozen at −20°C for shipment. Sub-cores designated for activity assays or mineralogy were transferred directly into the N₂ glovebox, washed twice with N₂-purged deionized water, wrapped in foil, sealed into mylar bags flushed with N₂ together with an O₂-scrubbing sachet (BD GasPak 260001) and stored at 4°C.

Samples for cell and tracer counts were obtained by capturing mixed pieces of core fractures, veins, surfaces and interiors during subsampling of the 50 cm core. These chips were loaded into two sterile 2 ml Eppendorf tubes and fixed with 4% paraformaldehyde (for fluorescence microscopy) or 2.5% glutaraldehyde (for electron microscopy) in phosphate-buffered saline (PBS) overnight, and stored at 4°C in PBS. Additional material was also loaded into pre-sterilized glass serum bottles, filled with fluids pumped from 50 m depth in nearby hole BA1A, and sealed with butyl stoppers under nitrogen gas for future cultivation experiments.

2.2 Mineralogical and Spectroscopic Analysis of BA Cores Preserved for Microbiological Investigations

All BIO samples for mineralogical analysis were stored in anaerobic pouches at 4°C until analysis. The dominant mineral components were determined using quantitative powder X-ray diffraction (XRD) (Table S1). Four hundred mg of crushed and sieved sample was mixed with 100 mg corundum and ground in a McCrone micronizing mill with 4 mL ethanol for 5 min, generating particle sizes on the order of 10–30 μm (a modified method based on Eberl, 2003). After drying at 60°C, the mixture was transferred to a plastic scintillation vial with three acrylic balls (∼1 cm in diameter) along with 200–800 μL Vertrel® solution and shaken for 10 min. The powder was passed through a 250 μm sieve to break up larger aggregates and loaded onto an XRD sample holder. Samples were analyzed using a Siemens D500 X-ray diffractometer from 5° to 65° 2θ using Cu Kα X-ray radiation, with a step size of 0.02° 2θ and a dwell time of 2 s per step. Quantitative mineralogy was calculated using the USGS software, RockJock (Eberl, 2003), which fits XRD intensities of individual mineral standards to the measured diffraction pattern.

Thin sections were also made by cutting a 1 × 2 × 4 cm billet from pieces of whole-round core using a water-cooled diamond sectioning saw. This section was then was cut into two duplicate, ∼5 mm thick billets to be prepared as standard petrographic thin section by Spectrum Petrographics (Vancouver, WA) and an “anoxic” thin section made by hand under O₂-free conditions (∼3% H₂ in N₂, <1 ppm O₂) in a Coy chamber. These thin sections were stored in an anoxic atmosphere until analysis by Raman and synchrotron X-ray methods.

Raman spectroscopy and hyperspectral mapping were completed using a Horiba LabRam Evolution Raman spectrometer equipped with a 532 nm laser and 600 gratings/mm diffraction grating. The laser was focused onto the thin section surface with a 50X or 100X microscope objective. Individual acquisitions were averaged to remove noise and filter out cosmic ray spikes, and spectra were baseline-subtracted using a polynomial baseline fit in LabSpec 6 (Horiba Scientific). Hyperspectral maps were modeled by a non-negative, least squares fit using endmember spectra composed of averages from relatively pure areas within the map.

Synchrotron-based X-ray fluorescence mapping was used to image the distribution of trace metals and to derive microscale Fe(II)/Fe(III) redox maps using the pre-edge multiple energy mapping method of Ellison et al. (2020). Maps were collected at the Stanford Synchrotron Radiation Lightsource Beamline 2-3, with a Si(111) Φ = 0° double-crystal monochromator. The beam was focused to a ∼4 µm spot, and the energy of the incoming beam was calibrated by defining the first inflection point of Fe foil as 7,112 eV, and by monitoring the position of a crystal glitch in every collected spectrum. X-ray fluorescence was detected with a silicon drift detector and multichannel analyzer. For trace metal mapping, samples were mapped with an incoming X-ray energy of 11 keV. For quantitative Fe redox mapping, a set of mineral Fe redox standards (described in Ellison et al., 2020) were measured to calibrate the Fe redox measurement. Maps were collected using a 4 μm step size and 90 ms dwell time at a series of 9 energies across the pre-edge and one above the main edge. The centroid position and integrated intensity of the pre-edge peak feature were determined in every pixel of the map and the Fe(III)/∑Fe ratio was determined by comparison to the calibrated variogram of Wilke et al. (2001).

Electron probe microanalysis (EPMA) was used to identify garnet and estimate the hydrous content of the garnet. EPMA was performed using a JEOL JXA-8230 instrument. Garnet was analyzed using 15 kV accelerating voltage, 20 nA beam current, and a 1 µm spot size. Acquisition and analysis were performed with Probe for EPMA software (Probe Software, Inc.) with ZAF background correction. Water content was calculated from the excess of charge on Ca, Fe, Al, and Cr versus Si, and garnet analyses were normalized to 24 cation charges (including H).

Focused ion beam (FIB) sections (approximately 20 μm long, 10 μm tall, and ∼100–150 nm thick) were cut from polished petrographic thin sections after applying a conductive Au coating using a FEI Helios NanoLab 600i with a Ga⁺ ion beam. The FIB sections were imaged by transmission electron microscopy (FIB-TEM) on a FEI Talos F200X field emission microscope.

2.3 Sulfate Reduction Activity Assays Using ³⁵S-Sulfate

Twenty-three depth intervals in the BA1B, BA3A, and BA4A cores were selected to measure potential biological sulfate reduction rates (pSRR), using methods adapted from Glombitza et al. (2016) and Glombitza et al. (2021). For each depth, a 10 cm whole-round core sample was prepared by removing the outer parts of the core using an ethanol cleaned chisel inside an anoxic chamber under N₂ atmosphere, where oxygen free conditions were maintained by circulating the gas atmosphere through a FeCl₂ solution. The inner parts of each core subsection were powdered by hand with a clean tungsten carbide mortar and pestle, and then 5–8 g of homogenized powder was dispensed into each of ten 20 ml serum vials. All equipment was cleaned between samples by washing with sterile water and additional wiping with 70% ethanol. Five milliliter of a synthetic anoxic medium was added to the mineral powders and each vial was sealed under a N₂ atmosphere. The medium was designed to represent a local groundwater with a pH of 9.5, using the following salt concentrations per liter: CaCl₂: 114 mg, Ca(OH)₂: 50 mg, MgSO₄·7H₂O: 90 mg, Na₂SO₄: 88.5 mg, KNO₃: 5.3 mg, KCl: 7.5 mg, K₂SO₄: 2.5 mg, H₄SiO₄: 0.5 mg, NaBr: 1.1 mg, NaCl: 275 mg, NaF: 0.02 mg. The effective sulfate concentration was 1 mM, which is similar to some of the local groundwaters (Nothaft, Templeton, Boyd, et al., 2021). Prior to adding the medium to the vials containing homogenized rock powder, the medium was subjected to autoclave sterilization. Following autoclave sterilization, a few drops of a filter-sterilized Resazurin solution (stock concentration 1 mg/L) was added as redox indicator and the redox potential was lowered by adding a few drops of a concentrated, filter-sterilized Na₂S₂O₄ solution until the indicator became colorless. Three different types of electron donor amendments (none, H₂ and CH₄) were tested in triplicate. H₂ and CH₄ amendments were achieved by replacing the N₂ headspace with 100% H₂ gas in three vials and by 100% CH₄ in another 3 vials. One additional vial with a N₂ headspace was used as an “autoclaved control” by autoclaving 3 times for 50 min at 134°C.

The 10 incubation vials for each depth interval were pre-incubated at 35°C for 3 days to allow the microbial communities to acclimate to the incubation conditions. Three MBq (in 15 μL of Milli-Q® water) of ³⁵S-labeled sulfate were injected into each incubation vial and incubated for 14 days at 35°C. The carrier-free ³⁵S-labeled sulfate (Na₂³⁵SO₄, American Radiolabeled Chemicals) was diluted with filter-sterilized (0.22 μm) Milli-Q® water to an activity of 200 kBq/μL prior to use. Six medium “blanks,” which comprised 5 mL samples of anoxic medium with a N₂ headspace but lacking rock core material, were incubated with 3 MBq (15 μL) sulfate tracer for 14 days and subjected to the standard work-up procedure in order to determine the assay detection limit (see below). Following the incubation, the vials were opened and the content was transferred into sterile 50 mL Falcon tubes containing 10 mL 20% zinc acetate to precipitate sulfide (as ZnS) and to terminate biological activity. Samples were then frozen at −20°C until further treatment.

To calculate the sulfate turnover rates, we determined the radioactivity of the reduced sulfur species and the total radioactivity in each sample. Frozen and fixed slurries were thawed and a 100 μL aliquot was taken and mixed with 5 mL zinc acetate (5%) and 15 mL scintillation cocktail (Ecoscint™ XR, National Diagnostics) to determine the activity of the unreacted ³⁵S-sulfate in the sample by scintillation counting. Subsequently, the remainder of the samples were subjected to the cold-chromium distillation procedure, which liberates dissolved and solid-phase reduced inorganic sulfur as H₂S (Kallmeyer et al., 2004). Subsamples were placed in the distillation apparatus (Røy et al., 2014) together with 10 mL of dimethylsulfoxide and 0.2 mL of a 0.5 M NaS₂ solution serving as sulfide carrier. Subsequently, 8 mL of 6 N HCl and 12 mL of a 1 M CrCl₂ solution in 2 N HCl were slowly added via an injection port into the air-tight apparatus. The generated H₂S gas was directed through an acetic acid wash and trapped as zinc sulfide using a 5% zinc acetate solution. Dinitrogen was used as carrier gas for the entire procedure. The content of the zinc sulfide trap was collected and mixed with 15 mL scintillation cocktail, and the radioactivity was measured by scintillation counting. The measured radioactivity of this fraction (a_TRIS: total reduced inorganic sulfur) is used together with the measured total activity (a_TOT: activity of the unreacted sulfate plus a_TRIS), to calculate the pSRR in the rock material according to Kallmeyer et al. (2004) and Glombitza et al. (2016).

$urn:x-wiley:21698953:media:jgrg22043:jgrg22043-math-0001$ (1)

In Equation 1, (SO₄²⁻) is the sulfate concentration (1 mM), ф is the average porosity of the rocks (2% for harzburgite and 3% for dunite; Kelemen et al., 2020), a_TRIS is the radioactivity measures in the TRIS fraction, a_TOT the radioactivity measured in the total samples (both in counts per minutes, CPM, on the scintillation counter), t is the incubation period and 1.06 represents an empirically determined fractionation factor between ³⁵SO₄²⁻ and ³²SO₄²⁻ that corrects for the slightly slower turnover of ³⁵SO₄²⁻ relative to ³²SO₄²⁻ (Jørgensen, 1978). The volume of rock material used for each subsample was calculated from subsample weights assuming an average density of 2.65 g/cm³ (Kelemen et al., 2020), in order to report an estimated sulfate reduction rate per cm³, since any biological activity occurs in available pore space, and is measured by measuring the turnover of sulfate dissolved in the fluid. Blank samples (n = 20) of 5 mL of 5% zinc acetate and 15 mL scintillation cocktail without tracer addition were measured on the scintillation counter and the mean counts per minute (CPM) of these “counter blanks,” representing a background of the scintillation counting was used to correct the sample measurements. A mean detection limit (MDL) was determined from the analysis of “medium blanks” (n = 6), where 3-MBq aliquots of the ³⁵S-tracer were added to 5 mL of medium and incubated and processed with the rock samples. The additional background measured from the medium blanks can result from turnover of sulfate tracer in the medium (i.e., any potential biological contamination and any radiotracer impurities). This turnover represents a minimum detection level below which we cannot attribute the measured turnover to an intrinsic sample activity. The MDL was then estimated from mean CPM of the TRIS fraction + 1σ (22 CPM), the lowest used sample amount (5.6 g) and the highest applied porosity value (0.03). MDL was estimated at 0.5 fmol/cm³/day.

2.4 Identification of Putative Sulfate Reducing Organisms.

Previous studies of the composition of 16S rRNA genes in filtered biomass collected from a variety of existing wells in the Samail Ophiolite, Oman, identified sequences closely related to Thermodesulfovibrionaceae, a bacterial family comprising sulfate reducers (Umezawa et al., 2021), as the most abundant organism likely catalyzing sulfate reduction (Nothaft, Templeton, Rhim, et al., 2021; Rempfert et al., 2017). In lieu of genomic data from the rock cores themselves, we examined available metagenomic sequence from biomass filtered from nearby subsurface waters as generated as a part of a previous study (Kraus et al., 2020) to characterize the potential for Thermodesulfovibrionaceae to function as sulfate reducers, and to identify potential electron donors capable of fueling this activity. Briefly, metagenomic sequences were generated from biomass collected from wells that intersect both gabbro and peridotite bedrock and include WAB188 (pH 7.6), WAB105 (pH 8.3), WAB104 (pH 8.5), WAB55 (pH 9.2), NSHQ4 (pH 10.5), WAB71 (pH 11.1), and NSHQ14. Two depth intervals were sampled at NSHQ14: 50 m (pH 11.1) and 85 m (pH 11.3).

Metagenomic sequences from WAB188, WAB105, WAB104, WAB55, NSHQ4, WAB71, and NSHQ14 have been previously assembled and binned into metagenome assembled genomes (MAGs) (Fones et al., 2019; Kraus et al., 2020). The completeness and contamination of MAGs was determined using CheckM (Parks et al., 2015). Metagenomic sequences are available in the MG-RAST database under accession numbers mgm4795805.3 to mgm4795809.3 and mgm4795811.3. MAGs that were most closely with family Thermodesulfovibrionaceae, specifically with the genus Thermodesulfovibrio, were identified and compiled for further phylogenetic characterization using the Genome Taxonomy Database Toolkit (GTDB-Tk), GTDB-Tk assigns taxonomy by placing MAGs in a phylogenetic tree using the GTDB database and then verifying its phylogenetic position with sequence alignments, average nucleotide identity (ANI), and average amino acid identity (AAI) calculations (Chaumeil et al., 2019). The average nucleotide identities (ANI) of MAGs affiliated with Thermodesulfovibrio were calculated using the program MUMmer v.4.0 (Kurtz et al., 2004). Pairwise ANI calculations were conducted with MAGs from Oman and several closely related genomes and MAGs reported previously, including Thermodesulfovibrio sp. N1 (SAMN05362001) (Frank et al., 2016). A dendrogram visualizing pairwise ANI calculations was generated using the default R package.

PROKKA was used to identify and annotate protein coding genes (Seemann, 2014). Manual assessment of protein annotations was performed by compiling protein sequences for metabolic pathways of interest from the facultatively alkaliphilic Thermodesulfovibrio sp. N1 assembled genome since this organism has been cultivated and subjected to physiological characterization (Frank et al., 2016). Additional protein sequences demonstrated in vitro to facilitate biofilm formation were compiled from Pseudomonas aeruginosa, Vibrio cholerae, Staphylococcus aureus, and Escherichia coli, since they are models for understanding biofilm formation (Wolska et al., 2016).This included searches for diguanylate cyclase (DosC) (Tuckerman et al., 2009) and biofilm PGA synthesis protein (PgaD) (Itoh et al., 2008) that facilitate EPS formation, the sensor histidine kinase CpxA (Dorel et al., 1999), which is required for transcription of biofilm relevant genes, and rugosity and biofilm modulator A (RbmA) (Smith et al., 2015) and surface protein G (SasG) (Belyi et al., 2018; Roche et al., 2003) that are required for adhesion. Local databases were generated for each Thermodesulfovibrio affiliated MAG from Oman samples. Protein queries were used in a BLASTp analysis against each local database specifying a query coverage cut-off of 50% and minimum e-value of 10⁻³⁰. Retrieved sequences were then subjected to reciprocal BLASTp analysis against the NCBI non-redundant database to add further support to protein annotations.

2.5 Enumeration of Microbial Cells and Tracer Beads in Rock and Fluid Samples

Paraformaldehyde-fixed samples from each sub-core were crushed by ceramic pestle and mortar in Super Clean Room in Kochi Institute for Core Sample Research, JAMSTEC, and mixed with a solution consisting of 3% (w/v) sodium chloride and 10% (v/v) neutralized formalin (containing 3.8% formaldehyde) at 1:1 volume ratio. To detach microbial cells and tracer particles from the rock matrix, a 1 mL aliquot of the fixed sample was amended with 2.2 mL of 2.5% NaCl, 400 μL of 100% methanol, and 400 μL of detergent mix containing 100 mM ethylenediamine tetraacetic acid (EDTA), 100 mM sodium pyrophosphate, and 1% (v/v) Tween-80 (Kallmeyer et al., 2008). The mixture was thoroughly shaken for 60 min (Shake Master, Bio Medical Science, Japan), and subsequently sonicated at 160 W for 30 s for 10 cycles (Bioruptor UCD-250HSA; Cosmo Bio, Japan). The detached cells were recovered by centrifugation based on the density difference of microbial cells and rock matrix, which allows collection of microbial cells in a lower-density layer. To achieve density separation, the sample was transferred into a set of four density layers composed of 30% Nycodenz (1.15 g/cm³), 50% Nycodenz (1.25 g/cm³), 80% Nycodenz (1.42 g/cm³), and 67% sodium poly-tungstate (2.08 g/cm³). Cells were separated from rock powders by centrifugation at 10,000 × g for 1 h at 25°C. All supernatant was collected using a 20G needle syringe. With the remaining rock powder pellet, the density separation was repeated. The rock powder was resuspended using 2.2 mL of 2.5% NaCl, 400 μL of methanol, and 400 μL of detergent mix (see above) and shaken at 500 rpm for 10 min at 25°C, before the slurry sample was transferred into a fresh centrifugation tube where it was layered onto another density gradient and separated by centrifugation just as before. All the supernatant was collected using a 20G needle syringe, and combined with the previously collected supernatant to form a single suspension for cell counting. Separately, the concentration of tracer beads in fluid samples was examined by trapping onto polycarbonate membrane without density separation.

The number of SYBR Green I-stained cells and UV-fluorescent tracer particles were enumerated separately by direct microscopic observation (Morono et al., 2009). For cell enumeration, all of the collected cell suspension was passed through a 0.22-μm polycarbonate membrane filter. Cells on the membrane filter were treated with SYBR Green I nucleic acid staining solution (1/40 of the stock concentration of SYBR Green I diluted in Tris-EDTA [TE] buffer). Unless at least 20 cells were found, we analyzed at least 900 fields of view for each whole membrane filter. The minimum quantification limit (MQL) for this cell enumeration procedure was determined by the counts, plus three-times standard deviation, of 8 procedural blanks, in which rock powder suspension was replaced by filter-sterilized ultrapure deionized water and processed in parallel with the samples. MQL in this study was 35 cells/g.

3 Results: Mineralogy, Cell Distribution and Cellular Activity Data From “BIO” Sub-Cores

Phase II of the Oman Drilling Project drilled 3 cored holes from 300 to 400 m in length, at the “Active Alteration” MBO site in Wadi Lawayni, located within the mantle section of the Wadi Tayin massif in the Samail Ophiolite (Figure 1). The excellent core recovery (100%) yielded 1 km of intact core. The three BA holes that were sampled for this study intersected dunite, harzburgite and interspersed gabbroic dikes located beneath a water table at ∼8–15 m depth. In Hole BA1B, the upper 160 m are dominated by dunite, and the lower 240 m are within harzburgite. In contrast, Hole BA4A is dominated by dunite throughout, with interspersed harzburgite and gabbro lenses, whereas Hole BA3A is dominated by harzburgite across the full 300 m depth (Figure 2). Most of the initial core characterization was conducted by the Oman Drilling Project Phase II Science Party aboard the D/V Chikyu in July and August 2018. Some of the most important Phase II results relevant to the microbiology investigations are summarized in the following paragraphs. Within this context, we also present data obtained by our analysis of BIO sub-cores.

3.1 Geochemistry and Mineralogy at BA1B, BA4A, and BA3A

The drilling into harzburgite and dunite at the “Active Alteration” MBO sites intersected distinct geochemical regimes within 300–400 m of the surface. When profiling the aqueous geochemistry as a function of depth in BA1B, BA4A, and BA3A in March 2018, there are notable changes in pH (pH 8.5–11.2), Eh (+200 to −750 mV) and conductivity (700–3700 μS/cm²). The most gradual changes in redox potential as a function of depth are observed in BA1B, whereas both BA4A and BA3A steeply transition to highly reducing potentials within the upper 30–50 m, approaching the lower stability limit of water (where H₂O is reduced to H₂) (Figure 3). The fluid temperatures range from 33°C at the surface to 43°C at 400 m depth, following the local geothermal gradient. In fluids pumped from adjacent boreholes BA1A and BA1D, the most abundant oxidants measured were nitrate (<1–240 μM) and sulfate (270–946 μM) (Nothaft, Templeton, Boyd, et al., 2021).

Reducing conditions occurred in the subsurface along almost the entire length of each BA core. The only samples where visible oxidation of the rock had occurred were located within the first ∼10 m of the surface, where the rocks were bright red and orange in contrast to the dark green to black of almost all the remaining core (Figure S1). For all boreholes, the peridotite rocks are ∼80–100% serpentinized, as confirmed by optical petrology, XRD, and X-ray computed tomography analysis (Kelemen et al., 2020). In quantitative XRD analysis, serpentine dominates all BIO subsamples, mixed with minor amounts of relict orthopyroxene, clinopyroxene and olivine (Figure 2). Brucite is notably abundant in the BA rocks (Figure 2). This is particularly evident in dunite rocks, such as the upper 160 m dunite section of BA1B (up to 8 wt% of the mineral assemblage). Brucite is also present at several weight percent in harzburgite protoliths as well. For example, in BA3A, which is almost entirely harzburgite throughout, Ellison et al. (2021) show that Fe-rich brucite is common in the mesh textured serpentine, often intimately intergrown with serpentine. Minor phases that we detected at times in bulk XRD of the BIO cores include garnet, chlorite, biotite, amphibole and dolomite (Figure 2; Table S1). Fortunately, we were able to preserve reduced mineral assemblages, including Fe(II)-bearing sulfides and hydroxides that can rapidly react and transform in air, by using an anaerobic chamber for sample processing and storing core samples in N₂-flushed mylar bags.

Numerous generations of veins, mineralized fractures, open joints and faults are abundant in the BA cores. Extensive networks of millimeter to centimeter scale serpentine veins can be observed in all core segments (Figure S1). The initial core characterization onboard D/V Chikyu has shown that there is an abundance of waxy green serpentine veins that cross cut almost all other veins types, suggesting that they have formed most recently (Kelemen et al., 2020). Calcite bearing veins are restricted to the top 10 s of meters of each hole, with the greatest amount of carbonate vein formation in the shallow portions of BA1B. In the BIO samples, we also commonly observe late-stage white serpentine veins that contain hydroandradite. Altogether, there are more veins in the upper part of BA1B than in BA3A and BA4A. Vein frequency decreases by almost an order of magnitude at depth in BA1B (Kelemen et al., 2020). The vein area in BA1B spans from 1% to 40%, with the highest vein proportions arising from the high frequency of calcite and waxy green serpentine veins in the upper 150 m, whereas BA4A and BA3A exhibit vein areas varying from ∼0.1% to 10% (Kelemen et al., 2020).

Open, unmineralized fractures and joints are common in all the holes, with decreasing abundance with depth. Their proportions will be quantified in the future by detailed analysis of optical and acoustic televiewer logs (see Kelemen et al., 2020 for more detail). BA1B has a notable fault zone near 30 m depth that released an almost liquefied, black serpentine mud. The adjacent intact core was also black and bubbled with gas (Figure S1). Tiny black particles were released from several upper BA1B core sections upon sawing with a diamond blade (Figure S1). The largest, meter scale fault in BA1B is located near 160 m, at the contact between the upper dunite and lower harzburgite. Numerous faults and cataclastic zones (average thickness ∼9 cm) also occur in BA4A and BA3A (Kelemen et al., 2000).

The porosity in most samples varies from 2.0% to 5.5% in the dunite and between 1.3% to 2.4% in the harzburgite, with an average rock bulk density of ∼2.65 g/cm³ (Kelemen et al., 2020). However, porosity is notably more variable in the upper 100–150 m of the BA holes, reaching 32% porosity and bulk densities as low as 2.2 g/cm³ in areas of high deformation (Kelemen et al., 2020).

For many BIO core subsamples, we used hyperspectral Raman imaging, in addition to optical petrology, to spatially resolve the numerous forms of serpentine, hydroxides, sulfides, carbonates, garnets, and relict minerals in BA samples. As part of this effort, we conducted an in-depth mineralogical study of BA3A, mapping out several stages of serpentinization and showing that ferroan brucite formed during primary serpentinization is present at depth, but reacts out to form ferric phases (Fe(III)-rich serpentine, hydroandradite and minor magnetite) in the upper 100 m (Ellison et al., 2021). Awaruite (Fe-Ni alloy) is also present throughout the BA3A core and becomes more Ni-rich closer to the surface (ranging from Ni₃Fe at 230 m depth to Ni₇Fe at 100 m depth) (Ellison et al., 2021). Fine opaque awaruite filaments similar to those detected in BA3A by Ellison et al. (2021) were also observed optically in BA1B and BA4A thin sections. In our petrological characterization of other BIO sub-cores we note that, unlike BA3A, many depths in BA1B and BA4A contain opaque replacement of the mesh in the cores (Figure 4). The dark mesh cores after olivine were also noted for many BA4A and BA1B intervals during the shipboard characterization on D/V Chikyu (Kelemen et al., 2020). Small amounts of sulfur are detected in some BA1B and BA4A samples by elemental analysis onboard D/V Chikyu, although the total sulfur concentrations are often less than 0.2 wt%, with a maximum of 0.6 wt% in BA1B at 120 m and BA4A at 40 m depth.

The mesh within the dark serpentinites cannot be characterized optically, and there is no new phase that shows up as quantitatively significant in the bulk XRD results. However, in the viscous serpentine mud material collected from the major fault zone at 30 m in BA1B, a small peak at ∼16° two-theta is consistent with the dominant peak of tochilinite (e.g., 6Fe_0.9S.5(Mg,Fe(OH)₂)), in addition to the lizardite, chlinochlore and quintinite that dominate the fault gouge material (Figure S2). A similar peak that varies slightly in position (from 16° to 16.5° two-theta) is also observed in opaque samples from BA1B 30 m (adjacent to the fault zone) and BA1B 70 m, as well as at 150 and 200 m in BA4A. The tochilinite-vallerite group (TVG) minerals comprise alternating sulfide (e.g., (Fe,Ni,Cu)S) and hydroxide components (e.g., (Fe, Mg, Al, Ca) (OH, CO₃)_1-2) (Beard & Hopkinson, 2000), and tochilinite specifically contains alternating mackinawite-like and brucite-like layers (e.g., Fe_1-xS and (Fe, Mg) (OH)₂), with a needle-like habit and repeated layer spacing of 10.4 Å (McKinnon & Zolensky, 2003). Although the single peak cannot be used for definitive tochilinite detection, and any such phase in the TVG minerals might be quite chemically variable, tochilinite was definitively detected by XRD in one sample during shipboard analyses on D/V Chikyu, and was observed to exist as a discrete phase within a BA4A vein (Tutolo & Evans, 2018). Thus TVG group minerals may be widespread through the opaque samples but near the limit of detection.

We used microscale hyperspectral Raman imaging to characterize several of the opaque samples in greater detail, such as BA1B 70 m (Figure 4). The blackened cores that have replaced the olivine mesh are visible in the optical image of BA1B 70 m, and are dominated by the sulfide peak shown in the corresponding Raman map. Two types of serpentine veins that are spectrally distinct are also present in the Raman maps, as well as hydroandradite garnet bounding the larger late-stage veins in BA1B 70 m. The position of the Raman sulfide peak maxima in the opaque cores is most often located at ∼275–290 cm⁻¹, which is most consistent with tochilinite, chalcopyrite, troilite or niningerite, but not pyrite, mackinawite, pentlandite, millerite, bornite, heazlewoodite or pyrrhotite (see Figure S3).

Hydroandradite garnet was detected in many BA samples by bulk XRD. Hydroandradite garnet was also identified throughout the mesh and late-stage serpentine veins using optical imaging and hyperspectral Raman mapping of altered dunites and harzburgites. An example from BA1B at 140 m is shown in Figure 5. Electron microprobe analyses were used to determine the chemistry and water content, providing an average formula of $urn:x-wiley:21698953:media:jgrg22043:jgrg22043-math-0002$ from samples from BA3A (60 and 140 m) and BA1B (140 m) (Table S2). Thus hydroandradite is a notable reservoir of ferric iron. We then used FIB-TEM to further characterize the hydroandradite present in the late-stage veins, revealing polyhedral serpentine surrounding hydroandradite (Figure 5).

Synchrotron-based x-ray fluorescence (XRF) mapping at variable energies around the Fe K-edge was used on numerous thin sections, including BA1B 140 m, to quantitatively map the Fe redox state of veins and the alteration assemblages, by correlating XRF maps to specific mineral phases identified by microscale Raman mapping (e.g., Ellison et al., 2020, 2021). One notable feature in the BA core samples is that the late stage veins that cross-cut most alteration are notably dominated by reduced Fe (Fe(III)/∑Fe ∼ 10%) compared to the wide range of Fe oxidation stages in the background alteration which includes contributions from serpentine as well as hydroandradite (Fe(III)/∑Fe ∼30%–90%) (Figure 5).

3.2 BA Cores: Quantification of Cells and Tracer

BA core subsamples fixed in 4% paraformaldehyde were used for cell separation and direct counting following procedures developed by Morono et al. (2013). In samples in which material from veins and fractures was preferentially preserved, cell densities varied from 10³ to 10⁷ cells/gram (Figure 6). Although the separated cells were typically dispersed, and were easily resolved from the tracer particles on the basis of their shape and fluorescence characteristics, large aggregates containing >60 cells were observed in several samples (Figure 7). Cells were also counted for interior portions of anoxically preserved cores that were separated and pulverized for sulfate reduction rate assays ∼8 months after drilling and core recovery. Before incubation, cell numbers varied from values near the mean quantification limit (35 cells/g) to 10² cells/g in samples used for measurements of potential sulfate reduction rates (Figure 6; Table S3).

Fluorescent “Invisible Blue®” micro-particles were measured at an average concentration of 1.3 × 10⁷ particles/mL (±6%) for the drilling mud through time. Tracer particles were also counted in each fixed rock sample to identify the most contaminated samples before downstream biological analyses, since the tracer particles are spectrally distinct from SYBR-green labeled cells. The “Invisible Blue®” tracer counts in the core samples exhibited an enormous dynamic range, varying from a minimum of 5 × 10² particles/g rock to 1.6 × 10⁶ particles/g rock (Table S3), comparable to the four order of magnitude range observed for cells. However, cell counts are not correlated to tracer counts, and the samples of greatest interest for downstream analysis have 1–3 orders of magnitude more cells than tracer particles. The viscous fault gouge serpentine mud sample from 30 m in BA1B—a sample expected to be contaminated by drill fluid—and exhibited a high load of 3 × 10⁷ tracer particles/g rock, whereas the adjacent intact core only contained 5 × 10⁴ particles/g rock.

3.3 Cell Activity: Measuring Potential Biological Sulfate Reduction Rates

Potential rates of microbial sulfate reduction (pSRR) in the BA cores were first measured in assays without electron donor amendment. Here, we were measuring the extent that sulfate reduction could be fueled by electron donors that may be endogenous to the rock (e.g., dissolved H_2, or organic acids, which are present at micromolar concentrations in fracture fluids, or mineral electron donors, such as ferroan brucite and metal sulfides). Efforts were also made to stimulate sulfate reduction by providing amendments of H₂ and CH₄. Observed potential sulfate reduction rates lie within the range of 2–100 fmol/cm³/day for almost all BA cores, with notably active samples exhibiting maximum rates as high as almost 1 pmol/cm^3/day (Figure 8).

The rates are highly variable in incubated samples from all three holes, and the slowest rates (∼2 fmol/cm³/day) are near the mean detection limit of our assay (0.5 fmol/cm³/day). In BA3A, the native and stimulated rates reach a maximum ∼200 fmol/cm³/day. In holes BA1B and BA4A, higher rates were observed at depth, with a maximum of almost 1,000 fmol/cm³/day in some samples below 150 m. However, there is great variability amongst the replicates for all depths and experimental conditions. Amendments with H₂ or CH₄ as potential electron donors resulted in modest stimulation of pSRR, up to a factor of 2–3 relative to unamended samples (Figure 8).

We also show data for the “autoclaved controls” (Figure 8), which were derived from rock powders that were triply autoclaved at 134°C before addition of the medium and ³⁵SO₄²⁻. Many of the “autoclaved control” samples exhibited sulfate reduction activity up to 50 fmol/cm³/day, which is up to 100 times higher than the MDL estimated from the incubation of the media used in sulfate reduction rate determination. We do not subtract the activity in the “autoclaved control” experiments from the rates measured in the “live” samples; instead, they are all shown for comparative purposes and addressed in the Discussion (Section 4.3) (Figure 8).

3.4 Identification of an Abundant, Putative Sulfate Reducing Bacterium

Previously generated metagenomic data from planktonic biomass filtered from wells near the “Active Alteration” MBO were used to identify MAGs from existing metagenomic libraries (Fones et al., 2020; Kraus et al., 2020) that are affiliated with the bacterial family Thermodesulfovibrionaceae (Umezawa et al., 2021), the most abundant group of putative sulfate reducing organisms identified in previous studies of Oman fracture fluids (Nothaft, Templeton, Boyd, et al., 2021; Rempfert et al., 2017). MAGs affiliated with Thermodesulfovibrionaceae, more specifically the bacterial genus Thermodesulfovibrio, were identified in wells WAB188, WAB55, WAB71, WAB105, NSHQ04, and two depth intervals in NSHQ14 (50 and 85 m) (Table S4; Figures S4 and S5). MAGs affiliated with Thermodesulfovibrio represented 0.5% of the total reads in the NSHQ14 50 m depth interval metagenome, and 6.2% of the NSHQ14 85 m depth interval metagenome. Among cultivated organisms with available genomes, the Thermodesulfovibrio MAGs from Oman were most closely related to Thermodesulfovibrio sp. N1 (0.858–0.917 pairwise ANI), which was isolated from a 2 km deep aquifer in western Siberia (Frank et al., 2016). ANI values between Oman Thermodesulfovibrio MAGs ranged from 0.810 to 0.997.

The Oman Thermodesulfovibrio MAGs do not show genetic evidence for nitrate, nitrite or O₂ utilization but do predict Thermodesulfovibrio will respire sulfate through the combined activities of sulfate adenylyl transferase (SAT), adenosine 5′-phosphosulphate reductase (AprAB), and dissimilatory sulfite reductase (DsrABCD) (Wolfe et al., 1994). Homologs of DsrJKMOP genes, which encode accessory proteins that stabilize core Dsr proteins and channel sulfur (Dahl et al., 1993), were also detected. However, homologs of DsrFH genes were not detected, consistent with Dsr likely operating reductively (Weissgerber et al., 2014). The Thermodesulfovibrio MAGs also encode homologs of heterodisulfide reductase (HdrABC) and methyl-viologen reducing and cytoplasmic [NiFe]-hydrogenase (MvhDGA), which function together to couple the oxidation of H₂ to the simultaneous reduction of FAD⁺ and heterodisulfide through electron bifurcation (Kaster et al., 2011). The MAGs also encode a reversible and NAD⁺ reducing [NiFe]-hydrogenase and a [NiFe]-hydrogenase (HyfA through HyfG and HoxSL, respectively), both of which are predicted to be biased toward H₂ oxidation (Peters et al., 2015). Proteins allowing for use of oxidants such as O₂ (cytochrome c oxidases) or nitrate (NarGHI) were not identified. Homologs of formate dehydrogenase (FdhAB) and carbon monoxide dehydrogenase/acetyl-CoA synthase subunit B (CODH/AcsB) are encoded in the genome, both of which are involved in the Wood Ljungdahl pathway of CO₂ fixation (Ragsdale & Pierce, 2008). The apparent absence of homologs of other proteins involved in this CO₂ fixation pathway, such as proteins involved in the entire methyl branch, indicates that formate dehydrogenase is running in the oxidative direction allowing for utilization of formate as an electron donor (Hattori et al., 2005). This also indicates that the Oman Thermodesulfovibrio are incomplete organic carbon oxidizers and incapable of CO₂ fixation. Likewise, Oman Thermodesulfovibrio MAGs do not encode proteins known to be necessary for other CO₂ fixation pathways.

Pyruvate ferrodoxin reductase (PorABC) may also facilitate the use of pyruvate as an additional reductant/carbon source (Blamey & Adams, 1993). Nutrient uptake is likely facilitated by ATP-binding Cassette (ABC) transporters including those specific for cysteine, thiosulfate, and sulfate import. Additional putative substrates potentially transported include phosphate and tungstate. Oman Thermodesulfovibrio MAGs encode genes for all the requisite subunits of Mo-dependent nitrogenase (NifHDKENB) (Boyd & Peters, 2013), indicating an ability to fix N_2. The MAGs encode a complete Embden-Meyerhoff pathway and a near complete oxidative TCA cycle, lacking only succinyl CoA synthase. A homolog of acetate kinase, which facilitates generation of acetyl CoA from acetate, was detected.

Homologs of several proteins associated with biofilm formation were not identified in Thermodesulfovibrio affiliated MAGs. This included searches for DosC (Dorel et al., 1999), PgaD (Smith et al., 2015), CpxA (Roche et al., 2003), RbmA (Smith et al., 2015) and SasG (Roche et al., 2003). This finding is consistent with characterizations of other strains closely related to the MAGs, including Thermodesulfovibrio sp. N1 (Frank et al., 2016). However, Thermodesulfovibrio sp. N1 has been documented as a member of a biofilm community (Frank et al., 2016), but it is not known if this biofilm was produced by Thermodesulfovibrio sp. N1 or a commensal community member.

Given the hyperalkaline pH of Oman subsurface fracture fluids, the Thermodesulfovibrio MAGs from Oman were examined for adaptations that may confer an advantage to these cells under such conditions. Like the closely related facultative alkaliphile Thermodesulfovibrio sp. N1 (Frank et al., 2016), the Oman MAGs encode multiple multi-subunit (Mnh type) Na⁺/H⁺ antiporters that function to maintain cytoplasmic osmotic balance (Krulwich et al., 2011). The MAGs also encode Na⁺/H⁺ antiporters from other families including the multiple resistance and pH family (MrpAD) (Ito et al., 2017). Similarly, the three homologs of previously mentioned [NiFe]-hydrogenases, which are inferred to be cytoplasmic, may play a role in producing protons intracellularly thereby allowing for cells to better modulate intracellular pH. The Oman MAGs also encode DnaK, a chaperone protein that is upregulated in response to osmotic shock (Bianchi & Baneyx, 1999). It is also important to note that many responses to alkalinity likely occur at the transcriptional level or in the form of single amino acid substitutions for key proteins (Kaya et al., 2018), which cannot be readily assessed with the data at hand. These putative adaptive traits may help to explain the abundance of Thermodesulfovibrio in Oman waters with hyperalkaline pH.

4 Discussion

The OmanDP “Active Alteration” Multi-Borehole Observatory encompasses ultramafic rocks storing water at habitable temperatures in a long-lived reactive system where redox disequilibria persist. The serpentinites have experienced several prior episodes of alteration and deformation, and are almost fully serpentinized. The rocks are continuing to undergo hydration and oxidation within a fracture-dominated hydrologic system, with greater density of transmissive fractures within the upper 50–100 m (Dewandel et al., 2005; Lods et al., 2020). Despite the lack of much relict olivine or pyroxene, the primary alteration assemblages contain abundant ferroan brucite, and widespread awaruite and sulfide phases, which serve as pervasive reductants.

In the March 2018 profiles of the aqueous geochemistry as a function of depth (Figure 3), we repeatedly observe regimes where there are large changes in pH (>2 pH units) and Eh (changes of >600 mV). The strong variations in pH persist through time, as shown by additional downhole measurements of pH for BA1B and BA3A conducted 2 yr later in March 2020; BA4A has not yet been relogged (Figure S6). Similar fluid disequilibria exist in adjacent BA1A and BA4A rotary boreholes (see Nothaft, Templeton, Boyd, et al., 2021). We infer that the fluids at depth are longer residence time, highly reducing fluids that contain dissolved hydrogen, methane and ammonium derived from extensive water/rock interaction, as characterized in groundwater well NSHQ14 that is immediately adjacent to BA3A (Miller et al., 2016; Nothaft, Templeton, Rhim, et al., 2021; Paukert Vankeuren et al., 2019), whereas the most shallow fluids are anoxic Mg−HCO₃^- type waters observed in nearby wells WAB104 and WAB105 that do contain oxidants such as nitrate and CO₂ derived from the atmosphere (Barnes & O’Neil, 1969; Nothaft, Templeton, Rhim, et al., 2021; Rempfert et al., 2017). Chemical disequilibrium occurs over the 10 s of meters scale, likely sustained by an upward flux of rock-buffered fluids that mix with shallow groundwater. We are likely observing evidence of the upward flux of a reduced fluid within all the large, late-stage veins. When mapped by synchrotron-based x-ray fluorescence mapping at the Fe K-edge, these serpentine veins are notably dominated by Fe(II) and are more reduced than the surrounding serpentine (FeIII/Fe_T is 10%–30% in late stage veins vs. 35%–90% in other BA rocks).

As discussed by Leong et al. (2021) and Nothaft, Templeton, Boyd, et al. (2021), mixing between a moderately alkaline, oxidizing fluid and a hyperalkaline, highly reduced end-member fluid close to equilibrium with serpentine, brucite and calcite (or diopside or hydroandradite instead of calcite) is likely occurring in the BA holes. The large chemical gradients, and the localization of the mixing zones, should strongly shape the microbial distribution. The greatest amount of chemical energy and dissolved inorganic carbon available to sustain chemolithoautotrophic metabolisms in the subsurface fracture system are predicted to be in zones of mixing between these endmember fluids (Canovas et al., 2017; Leong & Shock, 2020; Rempfert et al., 2017). Indeed, mixed fluids have been shown to harbor more cells than endmember fluids (Fones et al., 2019).

We also note that these reaction zones could also be important for abiotic organic synthesis, although the non-hydrothermal temperature and pressure conditions would likely render many reactions to be extremely sluggish (McCollom & Seewald, 2007). Although this topic is beyond the scope of the study presented here, it is important to recognize that highly reactive and metastable minerals such as iron hydroxides (including green rust like phases) and transition metal sulfides, including iron-nickel sulfides, are present across the steep redox and pH gradients we observe downhole; these minerals may be capable of catalyzing abiotic reduction of CO₂ and/or condensations, transformations and reductive aminations of small molecules under the ambient conditions (Barge et al., 2019; Cody et al., 2004; Hudson et al., 2020; Russell, 2018; Russell & Hall, 1997), although this will be hard to discern given the fact that these rocks are biologically alive. Several phases also associated with potential abiotic synthesis of carbonaceous matter during rock alteration, such as hydrogarnets, spinels and ferric saponites (Sforna et al., 2018) are also variably abundant.

4.1 Microbial Distributions and Activity in Highly Serpentinized BA Cores

One of the overarching goals of obtaining samples of subsurface peridotites undergoing active serpentinization is to identify whether these rocks are inhabited by living microorganisms, and what geochemical and hydrological processes may control the distribution and activity of microbial life. Yet it is challenging to accurately count cells and detect activity in fractured bedrock, given the highly heterogeneous nature of the cell distributions. The rocks recovered from the “Active Alteration” MBO sites exhibit a wide range of cell abundances, from robust values as high as 10⁷ cells/g at the top of BA1B to the lower detection limit of 10¹ cells/g in the interior of some BA cores (Figure 6). The highest cell concentrations are higher than those reported to date for other subsurface fractured igneous mafic and ultramafic rocks, such as the Atlantis Massif (10¹–10³ cells/g for International Ocean Discovery Program IODP) Exp. 357 (Früh-Green et al., 2018; Orcutt et al., 2017), the Atlantis Bank at the Southwest Indian Ridge (10²–10³ for IODP Exp. 360; Li et al., 2020), upper oceanic crust basalts buried under seafloor sediments adjacent to the Mid-Atlantic ridge (10⁴ cells/g for IODP Exp. 336; Jørgensen & Zhao, 2016), and deep continental basalts in the Decca traps (10⁵ cells/g; Dutta et al., 2018).The cell abundances may also be an underestimate, since the cell separation and enumeration protocol captures materials larger than 0.22 μm, and there is increasing evidence that ultra-small cells that would pass through such a filter are present in groundwater communities, including in serpentinitizing systems (Luef et al., 2015; Suzuki et al., 2017).

There is a difference in mean cell abundance between BA cores, which decrease in the order BA1B > BA4A > BA3A (1.1 × 10⁶, 4.3 × 10⁵, and 1.9 × 10⁴ cells/g, respectively; Table S3). As can be seen in Figure 2, the geochemical conditions do vary distinctly between BA1B, BA3A, and BA4A. The conditions are the most challenging for biology in BA3A, with pH > 11 and highly reducing, oxidant- and carbon-limited conditions below the top 30 m. In comparison, the pH is ∼1 unit lower in BA4A than BA3A, and the interface between the alkaline versus hyperalkaline fluids occurs slightly deeper (Figure 2). In BA1B, pH (and Eh) is much more variable, likely due to greater fluid mixing that is controlled by strong contrasts in the lithology, extent of fracturing, and porosity.

The high cell densities in the rocks are not necessarily surprising in light of those measured previously in nearby fracture fluids. Prior work in deep groundwater wells in the Samail Ophiolite has shown that the cell abundance was on average ∼10⁵ cells/mL for alkaline and hyperalkaline fluids circulating within these serpentinites (Fones et al., 2019). This fluid cell density is at the higher end of the range observed in ophiolite fluids, which can vary from 10¹ to 10⁶ cells/ml at the Voltri Massif, the Cedars, Cabeço de Vide, and the Santa Elena, Leka, Tablelands and California Coast Range ophiolites (Brazelton et al., 2012, 2017; Crespo-Medina et al., 2014; Daae et al., 2013; Morrill et al., 2013; Schrenk et al., 2004, 2013; Tiago & Veríssimo, 2013; Twing et al., 2017), and the majority of biomass can often be attached to the rock or fractures (Flemming & Wuertz, 2019). Some of the highest cell abundances in the BA cores may be localized along the numerous fractures present. Many of these are open fractures that are currently transmissive for fluids, as determined from analysis of optical and acoustic televiewer images, X-ray computed tomography (XCT) data, and pumping and monitoring hydraulic tests (e.g., Kelemen et al., 2020; Lods et al., 2020). It is possible that these fractures could host dense biofilms (e.g., Bochet et al., 2020; Jägevall et al., 2011; Wanger et al., 2006), and we may have sampled some of this material when fractures were present within the 50 cm BIO sub-cores.

The high cell abundances we detected should be considered overestimates for the bulk rock. There is a several order of magnitude difference between the cell counts for samples containing abundant veins and fracture surfaces than for samples that were derived only from the interior of cores (Figure 6). Moreover, during the cell counting, clusters with >60 cells were sometimes observed (Figure 7), and so cell counts could be skewed by the probability of capturing these clusters. Forming such microcolonies within open fractures may be common: high cell densities (e.g., 10⁶–10⁹ cells/g) can be found at fracture surfaces in subsurface rock ecosystems, particularly those significantly mineralized with secondary Fe-silicates and oxides (e.g., Suzuki et al., 2020; Trias et al., 2017; Wanger et al., 2006). In addition, recent studies have shown that large aggregations of cells can be preserved. For example, in 120 Ma low-temperature (<50°C) veins filled with calcite-brucite mineral assemblages in deep serpentinites of the Iberian Margin, significant accumulations of fossil microbial cells and complex biogenic organic matter has been detected via Raman micro-spectroscopy and further characterized by analysis of the preserved lipid biomarkers (Klein et al., 2015).

The protocols used to preserve samples for cell counting were not fully optimal, although we did ensure to wash the outer surface of core multiple times after sampling, first with deionized water and then with ethanol wipes, which we expected to reduce contamination by at least 1–2 orders of magnitude (e.g., Jørgensen & Zhao, 2016; Lever et al., 2006). However, we were not able to pare away the exterior core while in field, and only subsample interior materials, due to the competence of the intact cores and the lack of a clean-room to cut to the rock interior, as can sometimes be done shipboard. Instead, the materials we collected for paraformaldehyde fixation preferentially included veins and fracture surfaces that cross-cut samples, since the rock would most easily split along these interfaces with a sterile hammer and chisel. However, we do note that cores to be used for downstream biological analysis were subjected to an additional double-washing procedure before they were preserved frozen or at 4°C under anoxic conditions. Thus we do anticipate lower loads of contamination in samples that are currently being used for DNA and intact polar lipid extraction and analysis.

4.2 Microbial Activity: Potential Sulfate Reduction Rates

Sulfate reduction is often one of the most important anaerobic metabolisms fueling the subsurface biosphere (Chivian et al., 2008; Magnabosco et al., 2016; Robador et al., 2015; Teske, 2005). Sulfate is also expected to be an important oxidant in extraterrestrial systems where peridotite/water interactions and H₂ generation may occur, such as in subsurface aquifers on Mars (Boston et al., 1992; Clifford et al., 2010; Marlow et al., 2014; Michalski et al., 2013; Tarnas et al., 2018), and within the ocean of Europa (Hand et al., 2007; Kargel et al., 2000; McKinnon & Zolensky, 2003; Zolotov, 2003). Understanding the range of physiological conditions under which sulfate reduction can proceed will be valuable for future life-detection efforts in such systems. Here, we report the first cellular activity measurements in Oman serpentinite rocks, using assays focused on microbial sulfate reduction. This specific metabolism was chosen because sulfate is present up to 1 millimolar concentration in the Oman subsurface fluids, and therefore is one of the most abundant oxidants (Nothaft, Templeton, Rhim, et al., 2021; Rempfert et al., 2017).

Multiple lines of evidence have shown that oxidant limitation is pervasive in subsurface serpentinite systems, and is a major factor structuring the taxonomic and functional composition of subsurface communities in the Samail Ophiolite (Fones et al., 2019, 2020; Kraus et al., 2020; Rempfert et al., 2017). Prior work characterizing the 16S rRNA genes of microbial biomass present in fracture fluids has demonstrated that sulfate reducing organisms, particularly those within the family Thermodesulfovibrionia, are important members of the microbial communities characterized in the Samail Ophiolite. For example, sequences affiliated with Thermodesulfovibrionia comprise up to 40% of the total community in biomass collected from fluids pumped from nearby pre-existing groundwater wells such as WAB71 (Nothaft, Templeton, Rhim, et al., 2021). In addition, Nothaft, Templeton, Rhim, et al. (2021) found that sequences affiliated with Thermodesulfovibrionia comprised 20%–92% of the total community in the fluids sampled from holes BA1A and BA1D, which are within 120 m of BA1B. Likewise, our analysis of available metagenomic data from NSHQ14, which is within 10 m of BA3A, reveal that the MAGs affiliated with Thermodesulfovibrio sp. are inferred to constitute 6.2% of the community at 85 m. Pairwise ANI comparisons between Thermodesulfovibrio MAGs from NSHQ14 and Thermodesulfovibrio MAGs from other wells in Oman indicate that the MAGs from NSHQ14 are largely representative of the diversity of the MAGs detected in the sampled well waters (Figure S4). Examination of the MAGs for key metabolic genes involved in electron donor and acceptor usage indicate that these organisms encode the capacity to reduce sulfate, with electrons to fuel sulfate reduction likely derived from substrates such as H₂, formate, and carbon monoxide. However, we must acknowledge that the relative importance of Thermodesulfovibrionia within the rock matrix is unknown, although other recent studies have shown that Thermodesulfovibrionia can be strongly mineral-associated in the deep subsurface (Casar et al., 2020). The rock-hosted community likely differs strongly from the planktonic community, and any particle-attached sulfate reducers may be different phyla. Yet even if they are dominantly planktonic, Thermodesulfovibrionacea are likely present within some of the fracture volume and connected porosity sampled within these cores.

The use of radio-isotopically labeled ³⁵SO₄²⁻ provides a robust and sensitive method for quantifying any sulfate reduction activity. To our knowledge, the pSRR measured in the BA rocks (2–1000 fmol/cm³/day) are the lowest rates thus far reported in subsurface systems. Comparable studies have only been conducted in marine sediments, yielding higher rates (Beulig et al., 2018; Glombitza et al., 2016; Jørgensen et al., 2019), and as yet have not been conducted in deep crustal rocks. Recent work has provided the first quantification of microbial sulfate reduction in serpentinite fluids, specifically showing that this metabolism can proceed slowly across a wide-range of fluid compositions (Glombitza et al., 2021). In those assays, which used alkaline and hyperalkaline fluids ranging in pH from 8.5 to 11.2, the measured sulfate reduction rates varied from ∼0.5 fmol/ml/day to 2.1 pmol/ml/day (Glombitza et al., 2021).

We do not calculate cell specific sulfate reduction rates for the rock core assays, since we do not know what proportion of cells in any given sample might be active and functionally capable of sulfate reduction. Furthermore, sulfate reduction rates in the rock samples were measured in slurries using crushed rock powder and a synthetic incubation medium. This treatment renders the sample significantly different from in situ conditions and rates can thus only be considered as potential rates that do not justify cell specific turnover calculation. Clearly, cell numbers are lower for the materials used in the rock experiments (10¹–10² cells/g), whereas the fracture fluids tested by Glombitza et al. (2021) contained on average ∼10⁵ cells/ml (Fones et al., 2019).

The highest potential sulfate reduction rates between 100 and 1,000 fmol/cm³/day were found in the deeper cores of BA1B and BA4A (Figure 8). The maxima do not correlate with cell abundance, but instead occur at depth intervals where the redox potential in borehole fluids during logging is ≤250 mV, and the pH is ≤10.5 (Figure 2). In contrast, the lowest sulfate reduction rates are observed in BA3A, where the downhole fluid pH is ≥10.5 and the redox potential is held near the lower stability limit for water (i.e., reduction of H₂O to H₂). Glombitza et al. (2021) also noted a significant decline in sulfate reduction rates in fluids with pH > 10.5. These fluids also exhibit the lowest rates of CO and formate oxidation (Fones et al., 2019), potential reductants used by Thermodesulfovibrio. A possible inhibition of sulfate reduction at the highly alkaline pH values typically obtained during extensive serpentinization, despite the availability of sulfate and suitable electron donors, may define an important boundary condition for this metabolism that should be more fully explored. For example, isolates of alkalaphilic sulfate reducing bacteria from soda lakes also do not grow above pH 10.7 (Pérez-Bernal et al., 2020; Pikuta et al., 2003).

We only observe a small stimulation of microbial sulfate reduction activity with the addition of H₂ or CH₄ to the assays, despite the presence of several [NiFe]-hydrogenases that are putatively biased toward H₂ oxidation in the most predominant sulfate reducing Thermodesulfovibrio MAG. This observation may be consistent with the hypothesis that electron donors are not limiting, as proposed when Glombitza et al. (2021) did not find a significant difference between the native versus stimulated sulfate reduction rates in serpentinite fracture fluids. A similar phenomenon was recently demonstrated for hydrogenotrophic methanogens in serpentinite fracture fluids in Oman, where cells were shown to be more limited by carbon source than electron donor (Fones et al., 2020). To better test the potential for stimulation, it might have been preferable to (a) amend with small molecular weight organic acids, and (b) to increase the incubation time with ³⁵SO₄²⁻ in order to enable the rock-hosted organisms, which likely persist under low energy and oxidant fluxes, to respond to the amendments. However, the ³⁵SO₄²⁻ tracer, although well designed for sensitive measurements, only has a half-life of 87.4 days, and therefore is better suited to short experimental durations prior to extraction and counting. Future work probing hard rock samples for sulfate reduction activity (or other S transforming activities) could include a pre-stimulation of endogenous core populations with non-radioactive sulfate over months-long pre-incubation time scales, followed by the addition of ³⁵SO₄²⁻, in order to better test whether or not H₂, CH₄, formate or other electron donors might significantly stimulate microbial sulfate reduction activity.

It is important to note that the “autoclaved control” samples also showed sulfate turnover, albeit to a low extent relative to several of the core intervals examined (Figure 8). Although the sulfate turnover in the “autoclaved controls” was lower than in most of the “live” samples, this result is unexpected. As such, rates of sulfate reduction in biotic assays that do not exceed 50 fmol/cm³/day cannot be definitively attributed to biology. The only possible biological process we can invoke would be the germination of robust microbial endospores that survived the autoclave treatment and then germinated during the incubations (e.g., as observed with Desulfotomaculum spores by O’Sullivan et al., 2015). Although numerous spore-forming Firmicutes, including Clostridia members of the family Peptococcacea such as Desulfotomaculum sp., Desulfitobacterium were detected in subsurface fluids (Kraus et al., 2020; Rempfert et al., 2017), it is unlikely they can survive triple autoclaving and respond so quickly to the addition of sulfate. However, we cannot readily attribute these background rates of sulfate reduction to abiotic processes either. The rates of non-biological sulfate reduction are not predicted to be important <200°C (Ohmoto & Lasaga, 1982). Specifically, the incubation temperature of 35°C is well below the 120°C lower threshold temperature for where thermochemical sulfate reduction is observed (Machel, 2001), and thus falls within a regime where biological processes are known to drive sulfate reduction. To resolve this conundrum, we suggest that potential abiotic mechanisms to facilitate sulfate reduction within serpentinizing systems should be further investigated.

The biological sulfate reduction data raise several additional questions, including the source of the sulfate, and the potential impact of in situ microbial sulfate reduction on the serpentinite mineralogy and geochemistry. In fluids sampled from other BA holes at the “Active Alteration” MBO sites (e.g., BA1A, BA1D), as well as nearby wells, there is detectable sulfate (270–946 μM) (Nothaft, Templeton, Rhim, et al., 2021) and sulfide (0.4–24 μM) in the fluids. This sulfate may be derived from biotic or abiotic oxidation of the sulfide assemblages in the primary alteration, such as pentlandite, heazlewoodite, chalcopyrite and bornite, that are present at depth in the BA cores (Kelemen et al., 2020). There may also be some amount of sulfate salts stored in the serpentinite from early seafloor water/rock alteration prior to obduction, as well as subaerial weathering in the late Cretaceous (de Obeso & Kelemen, 2018; Oeser et al., 2012). Altogether, at the scale of the Wadi Tayin massif, there may be significant cycling of sulfur within a relatively closed system, where sulfur is mobile and oxidized in regions of fluid recharge, and where sulfur is reduced and mineralized in conditions with low redox potentials and active in situ biological sulfate reduction. This hypothesis could be tested in the future by establishing bulk and spatially resolved sulfide mineralogy and δ³⁴S isotopic analysis of secondary sulfides within the BA cores, as has been done in other serpentinites (e.g., Alt & Shanks, 1998; Delacour et al., 2008; Schwarzenbach et al., 2012).

At the OmanDP “Active Alteration” MBO, we suggest that the pervasive formation of secondary sulfides that are difficult to characterize in the BA cores is a result of biological sulfate reduction activity during ongoing low-temperature alteration and fluid circulation. It is also possible that other oxidized sulfur species such as thiosulfate, polysulfides or elemental sulfur, if present, are reduced by microbial cells, leading to formation of sulfide. Importantly, although the measurements of potential rock-hosted biological sulfate reduction activity are low, the prolonged production of a flux of sulfide may exert a direct geochemical imprint on the serpentinite alteration. In BA1B and BA4A there is extensive optical darkening of the rocks. The shipboard petrology identified “black cored” olivine in BA4A and BA1B, and the geochemical analyses showed enrichments in sulfur concentrations in bulk powders up to 0.6 wt% in BA1B and BA4A, whereas sulfur was not detectable in BA3A (Kelemen et al., 2020). In our petrological and Raman spectroscopic characterization of BA1B and BA4A rocks, we commonly detected the presence of a sulfide phase in the cores and rims of replaced olivine (Figure 4; Figure S3). The Raman spectra extracted from such maps show some variability, likely due to variations in the amount or type of sulfide mixed with surrounding serpentine. The dominant peak often lies between 275-300 cm⁻¹ and can be consistent with tochilinite (e.g., Vacher et al., 2019) as well as phases such as chalcopyrite (CuFeS₂), niningerite (i.e., MgS variably substituted with minor Fe and Mn) and troilite (FeS) (Avril et al., 2013). To further constrain the variability in secondary sulfide phases that are replacing olivine mesh cores, a comprehensive synchrotron-based X-ray absorption study of the opaque cores could be conducted, where the S X-ray absorption near-edge structure (XANES) spectra should be able to distinguish between the relevant sulfide species (Anzures et al., 2020; Fleet, 2005).

In multiple BA1B and BA4A samples, we also observed a small peak in the bulk X-ray diffraction data that is consistent with tochilinite (Figure S2), and tochilinite was definitely detected in a BA4A sample by Tutolo and Evans (2018). In Iberian Abyssal Plain rocks from ODP site 1068, where microbial reduction of seawater sulfate was inferred, Beard and Hopkinson (2000) and Whitmarsh et al. (1998) note that tochilinite is a major sulfur sink and rock-forming mineral that can be detected by XRD in the upper serpentinites. Given that the amount of sulfate present in the groundwater fluids at the BA sites is much lower than the average seawater sulfate concentration, we expect much less sulfur addition to these rocks even if microbial sulfate reduction is pervasive.

The potential links between in situ biological activity and the optical and chemical overprinting of the BA rocks is important to decipher. We specifically suggest that there may be a continuous flux of biologically generated sulfide that could be transported through the “Active Alteration” system, ultimately reacting with Fe(II), as well as Mg and trace metals such as Ni and Cu, derived from serpentine and brucite. Such sulfurization by sulfide could result in a diversity of opaque FeS-MgS-NiS-CuS phases, as well as mixed hydroxide-sulfide phases such as tochilinite. Any sulfide is likely quite mobile in the fluids, and thus optical darkening of the rocks might not (only) occur directly surrounding sulfate reducing cells. However, optical darkening might serve as an important chemical and spectroscopic signature that can be used at a variety of spatial scales to recognize a system strongly impacted by microbial sulfate reduction, as a first order way to identify rocks of interest for further biological analysis. In addition, if there is organic matter present in such rocks, sulfurization can often lead to preferential preservation of organics (Eigenbrode et al., 2018; Werne et al., 2004), potentially enabling the detection of biosignatures (Picard et al., 2019).

4.3 Future Outlook: Minerals and Interfaces of Specific Interest for Life Detection

Ultramafic rocks that have experienced sustained low-temperature water/rock interaction may have provided habitable environments on several rocky bodies in our Solar System. There exists a need to increase our capabilities to recognize active and fossil rock-hosted biospheres in fractured-rock systems, such as serpentinites, at a variety of scales (Hays et al., 2017). As suggested by Onstott et al. (2019), the search for biosignatures requires identifying the best type of interfaces at several spatial scales, including where there are notable redox disequilibria and contrasts in the alteration zone between high and low permeability regions of aquifers. From our preliminary investigation of the Oman “Active Alteration” MBO sites, we suggest that the rocks recovered from drilling in this system match these criteria and will provide excellent materials for developing targeted life-detection strategies. There are large contrasts in rock porosity and permeability, and fluid redox potential and pH, between the upper 100 m and greater depths in both dunite and harzburgite that may control the abundance of cells. In addition, there is significant secondary mineralization, dominated by sulfide or Fe(III)-bearing phases, that can play a critical role in preserving cellular structures that can serve as organic biosignatures and microbial fossils (Parenteau et al., 2014). Cells mineralized in veins could be excellent repositories for microbial membrane lipids (e.g., Klein et al., 2015; Newman et al., 2020; Zwicker et al., 2018). Moreover, since this is still an active system, it provides an excellent opportunity to design studies that can distinguish between lipid biomarkers associated with the active versus fossil microbial communities, by analysis of intact polar lipids versus core lipids (e.g., Li et al., 2020 vs. Newman et al., 2020), to correlate the signature of active microbial communities to distinct geochemical regimes, and to examine the relative preservation potential of different membrane lipids.

Our next stage of biological work with the BA cores focuses on characterizing the active biosphere in whole-rock samples by probing the Fe and C cycling dynamics, as well as extracting DNA and intact polar lipids and characterizing them in comparison to a deep collection of contamination controls. There are so many open questions regarding the microbial diversity, metabolic potential and functional activities of the serpentinite-hosted biosphere that has been accessed through drilling at the BA sites. In particular, we are focused on distinguishing regimes where Fe(II)-oxidation processes (e.g., biotic and abiotic oxidation of Fe(II) in brucite, sulfides and even serpentine) underpin the electron flux, where Fe(III)-reduction processes (e.g., reduction of ferric iron in serpentine, hydroandradite) provide critical electron acceptors, and where CO₂, formate or carbonate minerals serves as essential C sources that may control the turnover rates of the microbial communities. These cores should also be used for future spatially resolved investigations of fossil microbial activity that would have been sustained through progressive stages of water/rock interaction and fluid mixing. Recent microscale oxygen isotope studies of late-stage serpentine in BA1B veins suggests that crystallization temperatures varied from ∼38°C to 58°C (Scicchitano et al., 2020), which is well within the habitable range of temperatures to support microbial activity. In addition, Kelemen et al. (2020) show that there is a remarkably high abundance of veins, particularly in the upper 100 m, that have formed in response to recent water/rock interaction.

4.3.1 Hydroandradite

Hydroandradite would be one mineralogical target for searching for preserved cellular biomass. Hydroandradite garnet and polyhedral serpentine veins can form from serpentinization under low SiO₂ activity (Andreani et al., 2008; Beard & Hopkinson, 2000). The potentially large H₂ fluxes associated with Fe-oxidation and hydrandradite formation may be conducive to abiotic CO₂ reduction and/or in situ H₂-dependent microbial activity (Frost, 1985; Ménez et al., 2012; Plümper et al., 2014). Hydroandradite has previously been interpreted to be a low-temperature alteration phase when formed in association with polyhedral serpentine (Beard & Hopkinson, 2000; Ménez et al., 2012; Plümper et al., 2014). Ménez et al. (2018) suggest that organic carbon specifically plays a crucial role in the formation of polyhedral serpentine. In turn, hydroandradite can be intimately associated with organic matter, such as the condensed carbonaceous matter surrounding bastite-associated hydroandradite in the Ligurian ophiolite (Sforna et al., 2018), as well as the disordered carbonaceous matter, and a biopolymeric carbon, associated with hydroandradite and polyhedral serpentine from the Mid Atlantic ridge (Ménez et al., 2012). Hydroandradite is also a concentrated reservoir of Fe(III) that may serve as a oxidant for biological metabolism under highly reducing conditions. Thus, the veins containing hydroandradite and polyhedral serpentine in the Oman BA cores (e.g., Figure 5) are good targets in the search for any accumulations of abiotic and biological organic matter that might be preserved.

4.3.2 Carbonate Minerals

Carbonate veins (e.g., calcite, dolomite and magnesite) are another focal point for future efforts to determine where cells may be localized in the serpentinite subsurface. Carbonate minerals may be essential sources of carbon for autotrophic organisms, due to the low dissolved inorganic carbon concentrations in the hyperalkaline fluids, which are predicted to be ∼8–20 micromolar in water/rock reaction path modeling for low-temperature harzburgite alteration reaching chrysotile-brucite-calcite equilibrium (Leong & Shock, 2020). For example, hydrogenotropic methanogenesis, one of the most widespread and putative primordial metabolisms, may be prevalent in this system, as inferred from the detection of high abundances of 16S rRNA gene sequences affiliated with Methanobacterium sp. in fracture fluids, as well as metatranscriptomic and isotopic evidence for active methane production (Kraus et al., 2020; Miller et al., 2016; Nothaft, Templeton, Rhim, et al., 2021; Rempfert et al., 2017). However, CH₄ production by Methanobacterium sp. requires a source of dissolved inorganic carbon or alternative carbon sources such as formate (Fones et al., 2020). To date, most hyperalkaline fluid samples with abundant Methanobacterium only contain low micromolar levels of formate and other small molecular weight organics (Miller et al., 2016; Rempfert et al., 2017), and dissolved inorganic carbon is often below the ∼12µM detection limit (Nothaft, Templeton, Rhim, et al., 2021). Yet methanogens, including Methanobacterium sp., can grow at alkaline and hyperalkaline pH if CaCO₃ minerals provide a carbon source (Kral et al., 2014; Miller et al., 2018), Methanobacterium sp. have also been found to be abundant in hyperalkaline pH > 11 fluids in the Voltri massif and within carbonate deposits (Brazelton et al., 2017; Quéméneur et al., 2014). Particularly since the rock-hosted community may significantly differ from the fluid communities, carbonate minerals may play an important role in sustaining localized biological methane production in hyperalkaline systems and serpentinites, which should be directly investigated. Similarly, carbonate minerals may be a critical carbon source for other alkaliphilic H₂-based metabolisms, as shown in the Cedars (Suzuki et al., 2013).

4.3.3 Ferroan Brucite

Brucite is notably abundant in these low-temperature serpentinites (Figure 2). The brucite (Mg_xFe_1-x(OH)₂) phases are highly ferroan, and thus are particularly important reservoirs of labile Fe(II) (Bach et al., 2006; Miller et al., 2016; Templeton & Ellison, 2020). The Fe(II) in brucite could be utilized as an abundant electron-donor in anaerobic and aerobic Fe-oxidation; in addition, brucite in the BA cores should react with modern fluids to generate H₂ (Ellison et al., 2021), which may then serve as an electron donor to sustain biological activity. Tracing where brucite is actively reacting with fluids could provide a strategy for searching where H₂-consuming biological activity and disequilibrium fluid mixing is currently localized, and assessing whether this represents a hot-spot of biological activity. With such a study, it would also be possible to investigate whether brucite is preferentially colonized and serves as a direct electron donor for microbial growth when oxidants are present (Templeton & Ellison, 2020). The loci where brucite is reacting out could also be investigated to determine whether cells are preserved within newly formed ferric phases, such as Fe(III)-bearing serpentine and magnetite.

5 Conclusions

The Samail Ophiolite in Oman hosts a non-hydrothermal subsurface fractured rock ecosystem. Although the ultramafic rocks are almost fully serpentinized, chemical energy is available for chemosynthetic microbial communities where fluids exhibit steep redox gradients. In this system, it is possible to trace the intersection of moderately alkaline and more oxidizing fluids transmitted in the fractured and more permeable upper aquifers with the extremely reducing and hyperalkaline fluids stored in the deeper, lower permeability serpentinites. Across the system, cell densities vary from 10¹ to 10⁷ cells/g. The variability in cell abundances may reflect high biomass in fractures and late-stage veins, which are variably mineralized with serpentine, hydroandradite, and carbonate, and that lower cells densities are associated with the primary mesh serpentine and brucite alteration of the harzburgites and dunites. There are likely several additional physical controls on cell abundance, including changes in rock porosity that vary over orders of magnitude. In future work, spatially resolved approaches should be used to define exactly where within these rocks the majority of active and fossil biomass resides.

In the first efforts to measure microbial activity within the BA serpentinites, low levels of microbial sulfate reduction were measured using rock incubations with alkaline fluids amended with sulfate, including a ³⁵SO₄²⁻ label. Sulfate reduction coupled to oxidation of H₂, CH₄ or small molecular weight organic acids could sustain long-term microbial activity decoupled from surface inputs of oxidants such as O₂ or nitrate. Similar processes may be possible within the Martian subsurface, or the oceans on Europa and Enceladus, where fluids or brines rich in sulfate might be stored in contact with ultramafic rocks. The optical darkening and sulfide replacement observed in these BA serpentinites may be due to reaction with sulfide generated by microbial reduction of sulfate circulating in the subsurface fluids, as hypothesized here and in analogous systems such as the Iberian Abyssal Plain (Beard & Hopkinson, 2000). If so, this sulfide overprinting should be sought on other planetary bodies to broadly define a reaction zone that may have been biologically inhabited. However, we note that the abiotic formation of secondary sulfides and phases such as tochilinite is also common on meteorites, particularly on highly aqueously altered carbonaceous chondrites (Zolensky, 1984), and so such alteration can only be used to identify reaction zones of interest, but cannot be used directly as a biosignature. Instead, biosignatures, which could be assessed by studies of the sulfur isotopic systematics, successful extraction and sequencing of DNA, the preservation and characterization of microbial membrane lipid biomarkers, and the detection and/or imaging of other preserved cellular constituents, should be sought and tested once the most promising BA samples have been identified for detailed analysis.

Acknowledgments

The authors would like to thank Alexander Sousa and Kalen Rasmussen for volunteering effort to the OmanDP BIO team efforts in 2018, and Tyler Kane at the USGS for assistance with quantitative X-ray diffraction analysis of BIO samples. The authors also thank J. Coggon, D. Teagle and M. Al Sulaimani for all their logistical support, and M. Goddard and K. Michibayashi for their leadership and in the BA core characterization onboard D/V Chikyu. In addition, the authors thank J. Kallmeyer for guidance in the selection of particle tracers, and M. Schrenk, W. Bach and E. Shock for their efforts to help establish a biogeochemical component to the Oman Drilling Project. The authors thank Aaron Bell for assistance with Electron microprobe analyses, which were performed in the Electron Microprobe Laboratory, Department of Geological Sciences, University of Colorado-Boulder. The authors also thank David Dierks for assistance with FIB and TEM at the Electron Microscopy Laboratory, Colorado School of Mines. Raman spectroscopy and mapping were completed at the Raman Microspectroscopy Laboratory, Department of Geological Sciences, University of Colorado-Boulder. Use of the Stanford Synchrotron Radiation Lightsource, SLAC National Accelerator Laboratory, is supported by the U.S. Department of Energy, Office of Science, Office of Basic Energy Sciences under Contract No. DE-AC02-76SF00515. This work was supported by funding from the NASA Astrobiology Institute (NNA15BB02A), as well as the NASA Postdoctoral Program and the NASA Habitable Worlds Program (80NSSC19K0705). This research used logistical support, samples and data provided by the Oman Drilling Project. The Oman Drilling Project has been possible through co-mingled funds from the International Continental Scientific Drilling Project (ICDP, lead PI's Kelemen, Matter & Teagle), the Sloan Foundation-Deep Carbon Observatory (Grant 2014-3-01, Kelemen PI), the National Science Foundation (NSF-EAR-1516300, Kelemen PI), the NASA Astrobiology Institute (NNA15BB02A, Templeton PI), the German Research Foundation (DFG, Koepke PI), the Japanese Society for the Promotion of Science (JSPS, 16H06347, Michibayashi PI, and 19H00730, Morono PI), the European Research Council (Jamtveit PI), the Swiss National Science Foundation (Früh-Green PI), the Japanese Marine Science and Technology Center (JAMSTEC), the TAMU-JR Science operator, and in-kind contributions from the Sultanate of Oman Ministry of Regional Municipalities and Water Resources, the Oman Public Authority of Mining, Sultan Qaboos University, CRNS- Univ. Montpellier II, Columbia University, and the University of Southampton.

Open Research

Data Availability Statement

Data is provided in supplementary tables and figures associated with this manuscript. Metagenomic data can be accessed through the MG-RAST database under accession numbers mgm4795805.3 to mgm4795809.3 and mgm4795811.3.

Supporting Information

References

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Citing Literature

Volume126, Issue10

October 2021

e2021JG006315

This article also appears in:

Ophiolites and Oceanic Lithosphere, With a Focus on the Samail Ophiolite in Oman

Accessing the Subsurface Biosphere Within Rocks Undergoing Active Low-Temperature Serpentinization in the Samail Ophiolite (Oman Drilling Project)

Abstract

Plain Language Summary

Key Points

1 Introduction

1.1 Interest in the Biology of Low-Temperature Serpentinizing Systems

1.2 Oman Drilling Project Phase II at the “Active Alteration” Multi-Borehole Observatory

2 Materials and Methods

2.1 Core Procedures at the “Active Alteration” Multi-Borehole Observatory

2.2 Mineralogical and Spectroscopic Analysis of BA Cores Preserved for Microbiological Investigations

2.3 Sulfate Reduction Activity Assays Using 35S-Sulfate

2.4 Identification of Putative Sulfate Reducing Organisms.

2.5 Enumeration of Microbial Cells and Tracer Beads in Rock and Fluid Samples

3 Results: Mineralogy, Cell Distribution and Cellular Activity Data From “BIO” Sub-Cores

3.1 Geochemistry and Mineralogy at BA1B, BA4A, and BA3A

3.2 BA Cores: Quantification of Cells and Tracer

3.3 Cell Activity: Measuring Potential Biological Sulfate Reduction Rates

3.4 Identification of an Abundant, Putative Sulfate Reducing Bacterium

4 Discussion

4.1 Microbial Distributions and Activity in Highly Serpentinized BA Cores

4.2 Microbial Activity: Potential Sulfate Reduction Rates

4.3 Future Outlook: Minerals and Interfaces of Specific Interest for Life Detection

4.3.1 Hydroandradite

4.3.2 Carbonate Minerals

4.3.3 Ferroan Brucite

5 Conclusions

Acknowledgments

Open Research

Data Availability Statement

Supporting Information

References

Citing Literature

Figures

References

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2.3 Sulfate Reduction Activity Assays Using ³⁵S-Sulfate