Microbially generated H2S above 90 °C in the deep biosphere

doi:10.21203/rs.3.rs-3117180/v1

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Microbially generated H₂S above 90 °C in the deep biosphere

https://doi.org/10.21203/rs.3.rs-3117180/v1

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Thermophilic microorganisms play critical roles in sulfur cycling within the deep biosphere, but knowledge of these biogeochemical reactions at temperatures >80 °C is generally restricted to marine hydrothermal systems^1,2. Hydrocarbon extraction activities routinely interact with the deep subsurface, providing access to thermophilic environments where the thermal limits of life within sedimentary basins can be explored. Paramount to this, is understanding of how microbial communities in deep, hot biospheres are activated by and respond to disturbances such as the introduction of fluids from the surface. Here we show through chemical, stable isotope and microbiological analyses that microbial populations can actively catalyze sulfur cycling in subsurface hotter than 90 °C generating H₂S via microbial sulfate reduction. These temperatures exceed previously reported thermal limits of sulfate reducing microorganisms³ thus narrowing the gap between temperatures permissive for biogenic and thermochemical sulfate reduction⁴. Our analyses demonstrate that the introduction of surface-derived fluids into nutrient-limited deep environments stimulates microbial sulfate reduction at temperatures >90 °C previously considered too high for this process with implications for underground storage activities such as CO₂ and hydrogen storage.

Earth and environmental sciences/Biogeochemistry/Element cycles

Biological sciences/Microbiology/Biogeochemistry/Element cycles

Studies of microbial sulfate reduction (MSR) within the hot biosphere are dominated by investigations of deep-sea hydrothermal vents and other modern marine ecosystems^1,2,5. This is despite abundant drilling into deep, hot environments in continental settings related to industrial activities including hydrocarbon production and increasingly geological CO₂ sequestration, hydrogen storage, and disposal of waste fluids. Understanding the potential geochemical and biological reactions related to the injection of surface-derived fluids into extreme environments (e.g., > 80°C⁶) assumed to be biologically inactive is critical.

Within modern marine environments the estimates for the upper temperature limit for MSR has increased from 110°C as measured at deep-sea hydrothermal vents¹, potentially up to 120°C based on laboratory incubation experiments^2,7,8. In these hot marine environments, production of readily degradable organic substrates (e.g., acetate) permits the development of highly active populations of sulfate reducing microorganisms with rapid doubling times of hours as opposed to weeks for other sulfate reducing microorganisms². To date, reports of biological activity at such temperatures are almost exclusively associated with marine environments^1,2,5. In comparison, biological activity in the deep biosphere, hosted in sedimentary basins, is thought to cease at temperatures in excess of 80°C based on observations of hydrocarbon biodegradation in petroleum systems⁶.

In the deep biosphere, drilling into continental environments where temperatures exceed 80°C provide exceptional opportunities to study biogeochemical processes in non-hydrothermal extreme environments. Advances in directional drilling and enhanced hydrocarbon recovery technology has led to exploitation of hydrocarbons from low-permeability rocks such as shales and siltstones via hydraulic fracturing (HF)⁹. This process enables sampling of the otherwise inaccessible terrestrial deep biosphere via continuously produced fluids permitting time-resolved sampling of biogeochemical reactions in the deep biosphere.

HF of a hydrocarbon reservoir involves the high-pressure injection of water and sand to create fracture networks thereby forming fluid exchange pathways while generating space for active microbial communities. Along with water and sand, different chemicals (Table SI-1) are co-injected during HF to promote efficient recovery of hydrocarbons (e.g., oxidizing breakers such as persulfates¹⁰), and to stabilize the fracture networks (e.g., sand, guar gum). In general, great efforts are made to limit biological reactions as previous research has shown that microbial metabolisms and growth negatively impacts production of hydrocarbons due to production of H₂S¹¹ which is toxic and corrosive even at low concentrations. Injected fluids are treated with biocides, yet some microbial groups including resilient endospore-forming lineages¹² are consistently reported in produced fluids¹³. Despite originating from entirely different geological formations and generally at cooler temperatures, microbial communities in HF produced fluids are remarkably consistent^11,13,14, leading to the interpretation that microorganisms are introduced via surface-derived populations from nearby water sources followed by environmental selection in the extreme subsurface environment during and subsequent to HF¹⁴.

Montney shale formation

Here, we use aqueous and isotopic geochemistry combined with microbial DNA sequencing of HF fluids to evaluate biogeochemical responses to HF within a hot (> 90°C) and deep (> 3 km) environment. We analyzed produced fluids from a hydraulically fractured horizontal well at 3,411 meters depth in Canada’s Montney Formation, an Early Triassic siltstone formation¹⁵(Fig. 1). Injected water was sourced from local rivers characterized by low total dissolved solids (TDS < 1,000 mg/L) that mixes upon injection with high salinity (TDS > 170,000 mg/L) formation water in the target reservoir. The measured temperature within the reservoir is 91.6°C and is likely a conservatively low value given that cooling is often associated with the drilling process. Other local Montney reservoirs temperature is typically ≥ 95°C (Fig. SI-1). Injection of HF fluids initially reduces the temperature of fracture-filling fluids as cooler surface water (12°C) interacts with the hot reservoir (91.6°C). Following injection, the well was “shut-in” for 30 days during which time fluid temperature increases, reaching equilibrium on the order of weeks¹⁶ with simulations indicating < 2°C difference between geothermal and fluid temperatures¹⁷ during the production phase.

Water samples were collected at high density sampling over the first week of production (samples 20 to 900 m³), and then monthly to capture the initial conditions immediately following the onset of production, and monthly changes over the course of one year of production. This enabled the investigation of initial evolution of produced water geochemistry and microbiology related to reactions of highly reactive oxygen-bearing compounds within a reducing environment, the degradation of HF chemicals, and the introduction of microbial populations within the injected fluids. Monthly sampling revealed biological activation of the subsurface sulfur cycle as redox conditions shifted back to a highly reducing environment over time. Previous work and industry experience indicates that the geochemistry of produced water reflects the composition of the original formation water following a year of production^18,19.

Oxygenating an anoxic environment

Introducing surface-derived fluids into the subsurface exposes a hypersaline anoxic environment to low salinity, oxygen-rich fluids. Analysis of major anions and cations revealed that many species behave conservatively (e.g., Na, Cl, Li etc.; Fig. SI-2) displaying binary mixing of HF fluids and formation waters as noted by others¹⁹. However, non-conservative species including iron (Fe), sulfate, and sulfide reveal water-rock interactions and biological metabolism (i.e., MSR) that are consistent with the initiation of a biologically active subsurface sulfur cycle. Sulfate concentrations in produced water quickly increased to ~ 500 mg/L (Fig. 2a) and remained relatively constant throughout the sampling period (Fig. 3a). Core analysis of reservoir rocks from this region shows evidence of abundant anhydrite (CaSO₄) within pore-space (Fig. SI-3) which provides the sulfate source that is buffering dissolved sulfate concentrations in produced fluids.

Injection of oxygen-bearing compounds like persulfates and dissolved O₂ results in the oxidation of iron sulfides (i.e., pyrite occurs at up to 3.7 weight %) in the fractured host rock releasing Fe²⁺ ions. This is shown by the sharp increase in dissolved Fe in initial samples (Fig. 2a). Rapid iron sulfide oxidation through either microbial (Fig. 2) or abiotic reactions produces sulfate causing a corresponding shift in δ¹⁸O_SO4 from + 12.8 to + 15.3‰, reflecting dissolved oxygen being close to the atmospheric oxygen isotope ratio (δ¹⁸O_atm = + 24.2‰) ²⁰. Rapid consumption of reactive oxygen corresponded with δ¹⁸O_SO4 decreasing to baseline values (Fig. 2a). Previous studies suggest that removal of oxygen via sulfide oxidation occurs rapidly at elevated temperatures consistent with our observations²¹. The trajectory of the slope of δ¹⁸O_SO4 plotted vs. δ³⁴S_SO4 of initial samples (i.e., 20–100 m³ Fig. 2a) (Fig. 4) is analogous to those in other iron-rich environments where high rates of sulfate recycling²² result in considerable changes in δ¹⁸O_SO4 coupled with minimal δ³⁴S_SO4 variability. Any H₂S produced during this period is rapidly re-oxidized or sequestered via reactions with Fe²⁺ which is consistent with the presence of sulfate reducing and sulfide oxidizing microbial populations in the initial water samples (Fig. 2b).

Microbial life in a mixed redox environment

Initial microbial populations in produced water (samples 20 to 900 m³, Fig. 2b) reflect mixed redox conditions where contemporaneous sulfate reduction and oxidation occur. Relatives of the sulfate reducing, oxygen tolerant bacterium Desulfosporosinus fructosivorans (Fig. 2b) were observed in these initial samples, followed by elevated populations of putative sulfur oxidizing populations in subsequent samples (Fig. 2b). This suggests that biological activation of the subsurface sulfur cycle commences rapidly following injection of surface-derived fluids during HF. Desulfosporosinus spp. harbour multiple oxygen detoxification systems allowing these bacteria to persist in aerobic environments²³ together with an ability to form endospores likely conferring resistance to biocides in what could be a surface-derived dormant “seed bank” population^24,25 transported into the subsurface via injection fluids where the cells are activated by new environmental conditions.

Halotolerant species are enriched following injection especially Halanaerobium spp. (Fig. SI-4), which are routinely detected in shale reservoir produced waters ^14,26,27 where they are suspected to metabolise guar gum and/or produce sulfides via thiosulfate reduction²⁶. Thiosulfate was not detected in produced waters, suggesting organotrophic metabolism by Halanaerobium led to enrichment of these populations. Relatives of the halotolerant Pseudomonas songnenensis (Fig. 2b)²⁸ that is capable of aerobic hydrocarbon degradation²⁹ and Methylophaga spp. (Fig. SI-4) observed at hydrothermal vents and oil spill sites³⁰ capable of degrading higher molecular weight hydrocarbons³¹ were also observed.

This mixed anoxic-hypoxic phase may be required for initiating subsurface biological sulfur cycling, given that many sulfate reducing microorganisms do not directly metabolize n-alkanes and instead require metabolic residues such as organic acids and alcohols generated as by-products of organotrophic metabolism including the aerobic degradation of hydrocarbons³². Acetate and propionate in produced fluids (Fig. SI-5) may result from the fermentation/degradation of organic compounds used in HF additives³³ and/or in situ hydrocarbons or via biodegradation or thermal cracking of kerogen at temperatures between 80 and 200°C³⁴. Increasing acetate concentrations over the sampling period suggest a fermentative source. Fermentation of carbohydrates by Halanaerobium spp. may produce acetate and propionate that subsequently supports the growth of sulfur utilizing microbial communities³⁵. This hints at the importance of an aerobic phase to provide the necessary reductants for future anaerobic populations to take hold.

Re-establishment of anoxic conditions

Over time as HF chemicals are degraded and specific microbial populations become enriched conditions evolve from a mixed redox environment to an anoxic one. Over a 12-month period, we observed large changes to microbial populations and their impacts on fluids as revealed by geochemical and isotopic analyses. After the first three months iron concentrations decreased to below the detection limit coinciding with the onset of measurable H₂S in the produced water (Fig. 3a). Dissolved iron is removed via sulfide precipitation reactions, signaling a change in redox state in the subsurface.

The isotopic composition of sulfate during the initial sampling period (20–900 m³ Fig. 2a) and those at the beginning of monthly sampling (Fig. 3a) resemble δ³⁴S_SO4 and δ¹⁸O_SO4 values of in situ anhydrite minerals within the Montney Formation³⁶. This coupled with relatively constant sulfate concentrations in produced waters through time indicates that anhydrite dissolution is the principal source of sulfate. Over the course of the monthly sampling δ³⁴S_SO4 and δ¹⁸O_SO4 values of dissolved sulfate increased by 7.8‰ and 2.1‰, respectively (Fig. 3a) coincident with the onset of H₂S production. This shift in δ³⁴S_SO4 and δ¹⁸O_SO4 values is indicative of kinetic S and O isotope fractionation via semi-closed Rayleigh distillation during MSR. δ³⁴S_SO4 and δ¹⁸O_SO4 values increase during MSR as enzymatic processes within the cell preferentially metabolise bonds with lighter sulfur (³²S) and oxygen (¹⁶O) isotopes resulting in a residual pool of sulfate becoming progressively enriched in ³⁴S and ¹⁸O^37,38. It has been postulated that the magnitude of this isotope fractionation may be a function of microbial community and metabolites³⁹ and the availability of sulfate ^40,41. Additionally, laboratory and field studies indicate that higher MSR rates are associated with decreased sulfur^37,42,43 and oxygen⁴⁴ isotope fractionation. Moreover, lower oxygen isotope fractionations compared to sulfur isotope fractionations have been associated with higher sulfate reduction rates ^44,45. The ratio of sulfur to oxygen isotope fractionation in remaining sulfate of produced waters is remarkably low (0.358) corresponding to a rapid sulfate reduction rate of approximately 3.1·10^− 5 mol·cm^− 3·year^{− 1 44} (Fig. 4) implying minimal sulfate recycling via disproportionation. A similar sulfur-oxygen isotope fractionation relationship and sulfate reduction rate was observed at an oil seep in the Gulf of Mexico⁴² suggesting that oil seeps may represent good analogues for processes occurring in the subsurface during HF operations. Indeed, many of the microbes (e.g., Pseudomonas songnenensis, Methylophaga thalassica) observed in this study are similar to those recovered from oil seeps and oil degradation sites³¹.

Sulfide δ³⁴S values ranged from 17.1‰ to 19.7‰ equating to an average sulfur isotope enrichment factor of − 16.5‰ (δ³⁴S_SO4 – δ³⁴S_H2S). Sulfur isotope enrichment factors associated with MSR in natural populations typically range from − 16 to − 43‰⁴⁶, but can be as large as − 72‰⁴⁵. Smaller S isotope fractionation is associated with higher sulfate reduction rates coupled with ample sulfate supply⁴⁶. The relatively small S isotope fractionation between sulfate and sulfide in this system implies high sulfate reduction rate, supporting the δ³⁴S_SO4–δ¹⁸O_SO4 relationship described above. Desulfosporosinus spp. are the dominant sulfate reducing microorganism within most of the monthly samples (Fig. 3b), however species diversity increases toward later samples as the system evolves towards more anoxic conditions. Despite relatively low abundances compared to other bacteria, Desulfosporosinus is capable of high sulfate reduction rate⁴⁷ making it a keystone species having a disproportionally large effect compared with its abundance⁴⁸, here estimated to be 2% or less.

In concert with increasing δ³⁴S_SO4 we observed an increase in the diversity and relative abundance of sulfate reducing microorganism and higher H₂S concentrations in produced gas (Fig. 3a,b). The abundance of sulfate reducing microorganisms increases until 7 months post-injection, coincident with peak H₂S concentrations in produced gas. Following this peak, the abundance of sulfate reducing microorganisms and H₂S concentrations decreased potentially signalling the end of nutrient availability or the system poisons itself. Acetate, propionate, and sulfate concentrations remain constant (Fig. SI-5) potentially favouring the system poisoning hypothesis. Together, this data suggests that rapid sulfate reduction via MSR persists in these environments despite ‘hot’ (> 90°C) ambient temperatures in excess of previously assumed MSR thresholds.

Chemical data and stable isotope compositions of sulfate and sulfide coupled with DNA sequencing of microbial populations from fluids produced during and after hydraulic stimulation of a hydrocarbon reservoir demonstrated MSR at ≥ 90°C, establishing a new upper temperature limit for biogenic H₂S in deep environments. Our comprehensive dataset indicates that hydraulic fracturing of the deep, hot biosphere results in the injection of oxygen and nutrient-rich fluids (e.g., fermentable metabolites) along with microbial populations into an environment previously assumed inhospitable to biological activity, resulting in the activation of subsurface biogeochemical cycles. The activation of the subsurface sulfur cycle and the production of H₂S at temperatures in excess of 90°C requires a reassessment of the limits of biological activity in deep environments. This not only narrows the temperature gap between MSR and thermochemical sulfate reduction but is also analogous to systems experiencing freshwater injections into thermophilic environments including industrial activities such as CO₂ sequestration or waste storage and natural processes such as meteoric fluid migration along deep-seated faults, subglacial groundwater recharge⁴⁹, and hydrothermal fluid flow within sedimentary basins.

Water Time series Sampling

Water samples were collected from a separator on site in 1L liter Nalgene bottles. Two 1L samples were collected for geochemical analysis, and one bottle was amended with cadmium acetate to trap H₂S immediately upon sampling. Both samples were cooled and immediately transferred to the laboratory for analysis. Two 1L samples were also collected for microbiological analysis. One bottle was frozen and the other cooled to 3°C. Initial samples were collected every 20 m³ until 100 m³ of produced water, then samples were collected every 100 m³ of water until 900m³. After this samples were collected every month until one year from the onset of production.

Stable Isotope Analysis

Dissolved sulfide was extracted from water samples immediately following sampling via a reaction with added cadmium acetate. Resulting cadmium sulfide is converted to silver sulfide with silver nitrate prior to stable isotope analysis. Dissolved sulfate is extracted by first acidifying the sample with ~ 5 mL of 3N HCl, then ~ 5 mL of BaCl₂ solution is added precipitating BaSO₄ which is then filtered out and dried before stable isotope analysis.

Sulfur isotope analysis was performed via continuous flow isotope ratio mass spectrometry (CF-IRMS) on a Carlo Erba NA 1500 elemental analyzer coupled to a Thermo Delta + XL isotope ratio mass spectrometer at the Isotope Science Laboratory in the University of Calgary. Sulfide and sulfate samples were weighed in tin cups and combusted at 1050°C creating SO₂ which is separated from CO₂ and N₂ prior to analysis in the mass spectrometer. Samples are corrected for instrument effects using a suite of internal standards and normalized to the VCDT scale using the international standards IAEA S1, S2, S3 for sulfides and IAEA SO5, SO6, and NBS-127 for sulfates. Stable isotope results are presented in standard delta (δ) notation in ‰ VCDT. δ³⁴S precision is generally better than ± 0.3‰. Oxygen isotope analysis of sulfates was performed via CF-IRMS on a Heka High Temperature Oxygen analyzer coupled to a Thermo Delta + XL isotope ratio mass spectrometer at the Isotope Science Laboratory in the University of Calgary. Sulfate samples are weighed into silver cups and then converted to carbon monoxide at 1450°C and separated from other evolved gases by GC before analysis in the mass spectrometer. Samples are corrected for instrumental effects (linearity, drift, etc.) using a suite of internal and international reference materials (IAEA SO5, SO6, NBS-127) and normalized to the VSMOW scale via international standards.

Aqueous Geochemistry

Aqueous geochemistry was performed at the Applied Geochemistry Group at the University of Calgary. Water for alkalinity analysis (expressed as bicarbonate, HCO₃) was filtered through a 0.45 µm nitrocellulose filter (Millipore Corporation) prior to analysis. Total alkalinity was determined using an Orion 960 Titrator (Thermo Electron Corporation). A 10 mL aliquot of each sample was titrated with 0.01 M sulfuric acid (Fisher Scientific) until a pH of 2.5 was reached. End-point determination is accomplished via double differentiation of titration results.

Water for cation analysis was filtered through a 0.45 µm nitrocellulose syringe filter (Millipore Corporation), diluted (1:100, 1:1000) with deionized water, and placed in 15 mL Falcon tubes. Major/minor cations (Al, B, Ba, Ca, Fe, K, Li, Mg, Mn, Na, Si and Sr) were analyzed using a Varian 725-ES Inductively Coupled Plasma Optical Emission Spectrophotometer (ICP-OES). The spectrophotometer was calibrated (r² > 0.99) via serial dilution of certified standards (BDH Ltd.) of each individual element. Analytical precision and accuracy for cation analysis is typically ± 5.0%.

Water for anion analysis was filtered through a 0.45 µm nitrocellulose syringe filter (Millipore Corporation), diluted (1:100, 1:1000) with deionized, water and placed in 2.0 mL glass vials prior to analysis. Major/minor anions (Br, Cl, and SO₄) were analyzed using a Thermo-Dionex ICS 2000 ion chromatograph (Thermo Fisher Scientific Inc.). A 25 µL aliquot of each sample was injected onto an Ion Pac AS18 anion column (Thermo Fisher Scientific Inc.) and then separated by isocratic elution using 35.0 mM potassium hydroxide, a flow rate of 1.1 mL/min, and column temperature of 30°C. Analytical precision and accuracy for anion analysis is typically ± 5.0%.

Microbiology Analysis

Sample collection and processing for biological analyses

Produced water collected at the site were shipped to the laboratory for microbiological analyses. Upon arrival, 1 liter of produced water was frozen for DNA preservation until filtration could be done. Samples were thawed overnight at 4°C, and 400 mL of produced water was transferred to a sterilized filtration unit containing a 0.1µm, 47 mm diameter MilliporeSigma isopore polycarbonate membrane filter (Fisher Scientific, MA, USA). Filters were preserved at -20°C until DNA extractions were performed.

Molecular biology work

DNA was extracted using the PowerSoil DNA extraction kit from Qiagen (Hilden, Germany) by following the manufacturer’s guidelines. A volume of 32 µL was eluted in PCR grade water and used for quantification of DNA level and PCR.

The V6-V8 hypervariable region of the 16S rRNA gene was amplified using the 515F (GTGYCAGCMGCCGCGGTAA) and 806R (GGACTACNVGGGTWTCTAAT) primer pair. PCR amplification was done in a final volume of 25 µL using the 2x KAPA HiFi Hot Start Ready Mix (KAPA Biosystems), a final concentration of 0.1 mM of each primer and 5 µL of DNA sample. PCR reactions were done in triplicate and pooled together. PCR reaction went as follows: an initial denaturation at 95°C for 5 minutes, then followed by 30 cycles of denaturation at 95°C for 30 seconds, annealing at 55°C for 45 seconds and extension at 72°C for 1 min. This cycle ended with a final extension for 5 minutes at 72°C and PCR products followed a post-PCR cleanup and indexing. Indexed amplicon samples were sequenced using Illumina’s v3 600-cycle (paired-end) reagent kit on an Illumina MiSeq benchtop sequencer after all DNA extraction blanks and PCR reagent blanks were confirmed for negative amplification.

Biological data analyses

Primers were removed from the amplicon sequence variants (ASV) using Cutadapt, and further analyzed using DADA2. Taxonomy was assigned using the SILVA database version 138. Further information was obtained using BlastN to obtain taxonomy at the species level. ASV abundances were converted to relative abundances and GraphPad was used for visualization. To determine if microorganisms were most likely aerobic, anaerobic or facultative, a modified procedure from⁵⁰ was conducted. ASVs that were classified to the genus level and over 2% relative sequence abundance in one of the produced water samples analyzed in this study were used for further analysis. A total of 1346 species belonging to 36 different genera listed within the List of Prokaryotic Names with Standing in Nomenclature were classified as putative facultative, aerobe or anaerobe. When they were unassigned through inspection through the literature, they were removed from the calculations. The same ratio was followed as An et al.⁵⁰ to determine classification of microorganisms between the three groups.

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Microbially generated H₂S above 90 °C in the deep biosphere

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Version 1

Abstract

Figures

Introduction

Montney shale formation

Oxygenating an anoxic environment

Microbial life in a mixed redox environment

Re-establishment of anoxic conditions

Conclusions

Methods

Water Time series Sampling

Stable Isotope Analysis

Aqueous Geochemistry

Microbiology Analysis

Sample collection and processing for biological analyses

Molecular biology work

Biological data analyses

References

Additional Declarations

Supplementary Files

Status:

Version 1