Microbial Growth under Supercritical CO₂

Abstract

Growth of microorganisms in environments containing CO₂ above its critical point is unexpected due to a combination of deleterious effects, including cytoplasmic acidification and membrane destabilization. Thus, supercritical CO₂ (scCO₂) is generally regarded as a sterilizing agent. We report isolation of bacteria from three sites targeted for geologic carbon dioxide sequestration (GCS) that are capable of growth in pressurized bioreactors containing scCO₂. Analysis of 16S rRNA genes from scCO₂ enrichment cultures revealed microbial assemblages of varied complexity, including representatives of the genus Bacillus. Propagation of enrichment cultures under scCO₂ headspace led to isolation of six strains corresponding to Bacillus cereus, Bacillus subterraneus, Bacillus amyloliquefaciens, Bacillus safensis, and Bacillus megaterium. Isolates are spore-forming, facultative anaerobes and capable of germination and growth under an scCO₂ headspace. In addition to these isolates, several Bacillus type strains grew under scCO₂, suggesting that this may be a shared feature of spore-forming Bacillus spp. Our results provide direct evidence of microbial activity at the interface between scCO₂ and an aqueous phase. Since microbial activity can influence the key mechanisms for permanent storage of sequestered CO₂ (i.e., structural, residual, solubility, and mineral trapping), our work suggests that during GCS microorganisms may grow and catalyze biological reactions that influence the fate and transport of CO₂ in the deep subsurface.

INTRODUCTION

Geologic carbon dioxide sequestration (GCS) is an emerging strategy to abate CO₂ emissions associated with the burning of fossil fuels by capture, compression, and subsurface injection of generated CO₂ (1, 2). Although many subsurface geologic formations targeted for storage of compressed CO₂ are known to be biologically active environments (3–6), the extent to which biological processes may play a role in the fate and transport of CO₂ remains unknown (7). CO₂ exists as a supercritical fluid (scCO₂) at the temperature and pressures of the vast majority of reservoirs targeted for sequestration (i.e., >31°C and 72.9 atm). scCO₂ has generally been regarded as a microbial sterilizing agent due to a combination of factors, including cytoplasm acidification, increased CO₂ anion concentration, osmotic stress, membrane permeabilization and leakage via CO₂ extraction, and physical cell rupture (8–14).

While there has been no direct evidence that microorganisms can sustain metabolic activity and grow in environments containing scCO₂, previous work indicates this possibility. Survival of spores and biofilms after short-term scCO₂ exposure (i.e., minutes to hours) (8, 9, 15, 16) is well documented, and recent studies show that mineral matrices may enhance microbial survival of scCO₂ exposure by providing substrates for biofilm formation and/or by creating buffered microenvironments (10, 17). Biogeochemical models also suggest that diverse forms of microbial metabolism are thermodynamically favorable under reservoir conditions post-CO₂ injection (3, 18), where an aqueous phase in direct contact with scCO₂ may have dissolved CO₂ concentrations exceeding 2.5 M (3). Furthermore, high-pressure incubations to simulate reservoir conditions with elevated (but not supercritical) CO₂ have demonstrated activity of acetoclastic methanogens (under 49.3-atm pressure with 86.4 mM CO₂) (19) and homoacetogens (under 395-atm total pressure with 126 mM CO₂) (20). Recently, field studies at the Ketzin CO₂ sequestration site in Germany and the Otway Basin site in Australia provided evidence that changes in microbial community composition occur following CO₂ injection, suggesting that a combination of processes, including differential survival and possibly growth, occurs in the subsurface after exposure to near- and supercritical levels of CO₂ (21, 22).

Whether microorganisms survive and remain active post-CO₂ injection is relevant for predicting the fate and stability of the injected CO₂. Microbial activity can influence the various trapping mechanisms that are crucial to permanent storage of sequestered CO₂. Prior work has documented that microbial biofilms can be employed to plug pore spaces and impede the flow of scCO₂ through sandstone cores, providing a means of “structural trapping” for a mobile CO₂ phase (15, 23). Trapping of CO₂ residuals in pore spaces by capillary forces (residual trapping) may be affected by biosurfactant effects on wetting (24). Bacterial surfaces may provide sites for carbonate mineral nucleation, while bacterial activity can increase the rate of mineral weathering and therefore liberate the metal cations necessary for incorporation of CO₂ into carbonate minerals (mineral trapping) (25–27). Finally, increased dissolution of CO₂ into an aqueous phase (solubility trapping) has been demonstrated by pH increases induced by bacterial ureolysis under high partial CO₂ pressure (pCO₂) (28).

In this study, we tested whether environmental microbes could be isolated with the ability to survive and exhibit microbial activity (growth) during exposure to scCO₂. We performed a series of experimental enrichment cultures inoculated with subsurface fluid filtrate or well core samples from three subsurface environments targeted for CO₂ sequestration: the Frio-2 site near Liberty, TX (29); the Otway Basin site in southeastern Australia (22, 30, 31); and the King Island site near Stockton, CA (32). These three sites are geologically attractive as prospective CO₂ injection zones because they consist of high-porosity/permeability sandstone formations overlaid by low-permeability sealing layers capable of structurally trapping buoyant scCO₂ in the underlying zone. Enrichment cultivation was followed by isolation and characterization of strains able to grow under scCO₂. Microbial growth under scCO₂ is surprising given its inhibitory properties and indicates the possibility that microbial activities will influence CO₂ trapping during geologic carbon dioxide sequestration.

MATERIALS AND METHODS

Subsurface sample collection and storage.

Samples from GCS sites were utilized as inocula for microbial enrichment cultures using scCO₂ as the selective agent. Samples from the Frio-2 site were collected as part of the Frio-2 project and shared courtesy of Tommy Phelps (Oak Ridge National Laboratory). For sample collection, 10 to 20 liters of formation fluids was collected by U-tube from the Frio-2 CO₂ sequestration site near Liberty, TX, before, during, and after CO₂ injection (29, 33). Frio-2 formation fluids from 1,528- to 1,534-m depth were filtered through borosilicate glass filters (nominal pore size, 0.8 μm) and frozen on site. Three samples screened for this study correspond to samples collected prior to CO₂ injection, 7.5 h after injection, and 372 days postinjection. Otway Basin samples consisted of rock cores from 929- to 1,530-m depth from the Pemble, Paaratte, and Skull Creek formations (30). Otway cores were collected as part of the CO2CRC project in Southeast Australia (www.co2crc.com.au). The King Island core sample was obtained from the Mokelumne River formation at ∼1,447-m depth during drilling of the Citizen Green #1 deep-characterization well by the West Coast Regional Sequestration Partnership (WESTCARB), San Joaquin County, CA. Rock cores were kept refrigerated at 4°C prior to analysis.

Enrichment cultivation.

Inocula for enrichment cultures were prepared in an anaerobic glove bag with an O₂ and H₂ monitor (95% CO₂-5% H₂) and added to 4- or 10-ml pressure vessels containing a 50% volume of growth medium (below). For Frio-2 samples, inocula consisted of 10 μl of hydrocarbon and particulate residue from the surface of glass fiber filters, collected with a sterile scalpel. For Otway Basin cores, drilling fluid tracer penetration data were used to guide collection from the core interior where contamination was least likely. The selected regions of Otway cores were pulverized with a stainless steel mortar and pestle, and 1 g of crushed rock was used as inoculum. For the King Island formation core, no tracer-free interior could be identified, as the sediment was highly permeable and unconsolidated. Thus, a representative sample from the center of the King Island core was used as inoculum and processed in the same manner as Otway cores.

Medium for enrichment cultivation of Frio-2 samples was modified GYP sodium acetate mineral salts broth (GYP) consisting of (in g/liter) 2.0 glucose, 1.0 yeast extract, 1.0 tryptic peptone, 1.0 sodium acetate, 0.2 MgSO₄·7H₂O, 0.01 NaCl, 0.01 MnSO₄·4H₂O, and 0.01 FeSO₄·7H₂O. Both GYP and MS media were used for enrichment cultivation from the Otway Basin and King Island cores with supplements targeting different microbial functional groups added to MS medium (34). MS medium consisted of (in g/liter) 0.5 yeast extract, 0.5 tryptic peptone, 10.0 NaCl, 1.0 NH₄Cl, 1.0 MgCl₂·6H₂O, 0.4 K₂HPO₄, 0.4 CaCl₂, 0.0025 EDTA, 0.00025 CoCl₂·6H₂O, 0.0005 MnCl₂·4H₂O, 0.0005 FeSO₄·7H₂O, 0.0005 ZnCl₂, 0.0002 AlCl₃·6H₂O, 0.00015 Na₂WoO₄·2H₂O, 0.0001 NiSO₄·6H₂O, 0.00005 H₂SeO₃, 0.00005 H₃BO₃, and 0.00005 NaMoO₄·2H₂O. MS medium supplements consisted of 0.5 g/liter glucose for fermenters; 1.3 g/liter MnO₂, 2.14 g/liter Fe(OH)₃, and 1.64 g/liter sodium acetate for metal reducers; 0.87 g/liter K₂SO₄, 0.83 g/liter FeSO₄, and 0.82 g/liter sodium acetate for sulfate reducers; or 1.3 ml trimethylamine and 0.82 g/liter sodium acetate to target methanogens (34). Culture media were added to serum bottles and degassed with a stream of 100% CO₂ or 100% N₂ gas for 30 min prior to pressurization. Na₂S (at 0.25 g/liter), a reducing agent to maintain anaerobicity, and resazurin (at 0.001 g/liter), a visual redox indicator, were added to culture media. Following inoculation, vessels were pressurized and incubated for 2 to 4 weeks at 37°C for Frio-2 samples and 37 and 60°C for Otway Basin and King Island samples, respectively. In addition to the above media, Luria-Bertani broth (LB) (Difco) and LB agar were used for strain cultivation at 37°C under ambient aerobic conditions.

The pH of ambient and CO₂-saturated medium was measured at 1 atm and 21°C using an Orion model 520A pH meter. The pH of medium under an scCO₂ headspace was measured by visualization of a pH indicator strip (EMD Chemicals) through the sapphire window of a 25-ml view cell (Thar Technologies; 05422-2). In addition, PhreeqC version 2 was used to predict the equilibrium pH and potential precipitation of chemical species in the growth medium (Table 1) under a CO₂ or N₂ atmosphere and as a function of temperature and pressure. Thermodynamic data were obtained from the Lawrence Livermore National Library (LLNL) database.

TABLE 1.

Observed and predicted pH and predicted CO₂ concentration as a function of GYP culture medium composition,^a headspace gas, and pressure at 37°C

Headspace	Pressure (atm)	Predicted pHb	Observed pHc	Predicted [CO2] (M)b
CO2	1	5.3	5.1	0.026
	100	3.6	3.9–4.5	2.7
N2	1	7.0	7.0	0
	100	7.0	NDd	0

^aGrowth medium components (g/kg): 0.72 acetate, 0.431 Na⁺, 0.607 Cl⁻, 0.002 Fe²⁺, 0.0025 Mn²⁺, 0.0196 Mg²⁺, 0.08575 SO₄²⁻, 0.1028 S²⁻. Buffering of pH by the acetate system in GYP maintained the final pH above the value predicted for deionized water alone.

^bGeochemical modeling performed in PhreeqC calling the LLNL database.

^cpH measured by Orion model 520A pH meter (P = 1 atm) or visualized (P = 120 atm) via indicator strip (EMD Chemicals).

^dND, not determined.

High-pressure incubation.

Vessels for high-pressure growth (see Fig. S10 in the supplemental material) were 316 stainless steel high-performance liquid chromatography (HPLC) column bodies or 316 stainless steel tubing (4- and 10-ml capacity). Vessels were fitted with ball valves (Supelco) or quarter-turn plug valves (Swagelok or Hylok). Vessels were filled to one-half capacity (2 or 5 ml) with cultivation medium, and following inoculation, the headspace of the stainless steel culture vessels, representing 50% of the total vessel volume, was pressurized at a rate of 2 to 3 atm min⁻¹ to 100 to 136 atm with industrial-grade N₂ gas (Airgas) or with extraction-grade CO₂ gas (Airgas) with a helium (He) head pressure such that the final gas mixture was 97 to 99% CO₂. Pressurized vessels were incubated in a 37°C warm room, with shaking at 100 rpm to increase mixing of media and subsurface inocula. Following incubation, the vessels were connected with 316 stainless steel tubing and fittings to Swagelok pressure gauges to measure the final vessel pressure before samples were depressurized at a rate of 3 to 5 atm min⁻¹ over approximately 30 min. Generally, culture vessels with initial headspace pressures of >100 atm lost between 5 and 25 atm of pressure, due to slow leakage through fittings, over the course of a multiple-week incubation, with greater losses associated with longer incubations. Unless specifically noted, all vessel incubation data reported here maintained scCO₂ headspace pressures of >72.9 atm, the critical pressure for CO₂ mixed with ≤3% inert helium at 37°C (35), or for incubations at lower pressures, final pressure (P_final) was ≥70% of P_initial. All pressures were measured at room temperature (21°C). Based on the ideal gas law, we can estimate a maximum pressure increase of 6% associated with incubation of reactors at 37°C, although since scCO₂ is a nonideal gas, this may be an overestimate. Following depressurization, cultures were transferred to an anaerobic chamber (Coy Lab Products) containing a 95% CO₂-5% H₂ atmosphere for subsampling and passaging. All pressure vessels and valves were cleaned and autoclaved between uses, and high-pressure tubing was flushed before use with 10% bleach for 30 min, rinsed with MilliQ H₂O, rinsed with 100% ethanol, and dried with CO₂ gas.

Enrichment cultures from the Frio-2, Otway, and King Island sites were serially passaged by diluting 10% (vol/vol) of the previous culture in fresh growth medium under a 95% CO₂-5% H₂ atmosphere, with pressurization to 120 atm with CO₂, followed by incubation at 37°C. The contents of enrichment vessels were analyzed for cell abundance at the end of passages and in the inocula prior to incubations. Frio-2 passages 1 to 3 were incubated for 15 days each, while passage 4 was incubated for 60 days; subsequent passages were incubated for 9 to 15 days (Table 2). Otway and King Island passages were incubated for longer time periods (i.e., 1 to 2 months) to increase the likelihood of cellular growth based on earlier observations. Samples from each passage were subjected to microscopic enumeration and archived as glycerol stocks at −80°C.

TABLE 2.

ScCO₂ enrichment cultivation summary

Sample origin (incubation time [days] for initial enrichment and passages 1–3, respectively)	Growth mediuma	Biomass (cells/ml) observed for enrichment/passage at temp (°C)c								Isolateb
		Initial		1		2		3
		37	60	37	60	37	60	37	60
Frio-2; formation water particles; 1,500 m (15, 14, 15, 15)	GYP	+++	NDd	+++		+++		+++		MIT0214
Otway core 2; 1,164.12–1,164.33 m; silty claystone (21, 33, 61, 41)	MS + fermenter	−	+		+
	MS + metal reducer	−	+		−
	MS + sulfate reducer	−	+		+
	MS + methanogen	−	+		+		+
	GYP	−	−
Otway core 3; 1,193.59–1,193.69 m; sandstone (14, 28, 81, 40)	MS + fermenter	+	+	+	+	−	−
	MS + metal reducer	+	−	+		+++		+++		MITOT1
	MS + sulfate reducer	+	−	−
	MS + methanogen	−	−
	GYP	−	−
Otway core 4; 1,239.31–1,239.48 m; silty claystone (28, 81, 41, ND)	MS + fermenter	+	+	+	+	−	−
	MS + metal reducer	+	+	−	+		−
	MS + sulfate reducer	+	+	−	−
	MS + methanogen	−	+		−
	GYP	−	−
Otway core 5; 1,273.86–1,273.95 m; sandstone (29, 42, 49, ND)	MS + fermenter	+	−	+		−
	MS + metal reducer	−	−
	MS + sulfate reducer	−	−
	MS + methanogen	−	−
	GYP	−	−
King Island core; 1,447 m; unconsolidated sandstone (29, 38, 49, 37)	MS + fermenter	+	+	+	−	++		++
	MS + metal reducer	+	−	+		+++		+++		WMR1
	MS + sulfate reducer	−	−
	MS + methanogen	+	−	+		+++		+++		WMe1, WMe2
	GYP	+	−	+		+++		+++		WG1

^aGrowth medium consisted of GYP or basal MS medium with supplementation targeting various microbial metabolic groups (see Materials and Methods).

^bStrain identification by 16S rRNA gene BLASTN: MIT0214, B. cereus; MITOT1, B. subterraneus; WMR1, B. safensis; WMe1, B. safensis; WMe2, B. megaterium; WG1, B. amyloliquefaciens.

^c+, ++, and +++, biomass observed at <10⁴, 10⁴ to 10⁵, and >10⁵ cells/ml, respectively. −, biomass not observed.

^dND, not determined.

Enumeration of cell density.

To quantify biomass, we used a combination of methods, including direct cell counts, viable cell counts, and optical density. Cell density at the beginning and end of incubations was determined by microscopic epifluorescent cell staining using 4′,6-diamidino-2-phenylindole (DAPI; Sigma) or SYTO9 (Invitrogen) with shaking for 10 min in the dark. Five hundred microliters to 1 ml of sample was then filtered onto 25-mm, 0.2-μm-pore-size, black polycarbonate filters (Nuclepore), followed by 2 washes with 1 ml of phosphate-buffered saline (PBS). PBS was incubated on the filtered sample for 1 min to help wash off excess dye. Filters were laid on slides under microscope immersion oil with a coverslip (Thermo Scientific) and were stored at 4°C in the dark until counting. The cell density (in cells/ml) was calculated by multiplying the mean cell counts (in one 10-by-10 microscope grid) by the dilution factor and then by 3.46 × 10⁴ (as one 10-by-10 grid at ×1,000 magnification corresponds to 1/3.46 × 10⁴ of a 25-mm filter). Samples were visualized on a Zeiss Axioplan fluorescence microscope. Images were captured on a Nikon D100 camera using the NKRemote live imaging software. Viability counts, i.e., CFU plating, were performed using Luria broth agar. Viable spore counts were carried out by heating aliquots to 80°C for 10 min to kill vegetative cells (36) prior to plating on Luria broth agar. Culture optical density (600 nm [OD₆₀₀]) was measured on a Bausch and Lomb spectrophotometer (1-cm path length) or via a 96-well microplate reader (BioTek Synergy 2) (200 μl per well). Optical density was not measured for incubations using metal reducer medium due to confounded readings from solid particulate content.

For MIT0214, growth was defined by increased cell density and evidence of vegetative cell morphologies by microscopy. Growth-positive cultures had at least 45-fold-increased direct cell counts relative to initial inocula of less than 1 × 10⁶ spores/ml, or at least a 5-fold increase in direct cell counts for cultures with inocula greater than 1 × 10⁶ spores/ml, as MIT0214 final cell densities generally varied between 1 × 10⁷ and 1 × 10⁸ cells/ml. For MITOT1, growth was defined by observation of culture turbidity accompanied by at least a 4-fold increase in viable cell counts (CFU/ml) above the initial spore density (on average, increases were >50-fold) and at least 25-fold-higher viable counts compared to replicates without observed turbidity since samples without evident growth all showed a decline in viable counts relative to the initial viable counts of the inoculated cultures, presumably due to a loss of spore viability during the incubation period.

Extraction of DNA and analysis of 16S rRNA gene diversity in enrichment cultures.

DNA extraction from Frio-2 sample enrichment passages was performed using a protocol modified for Gram-positive bacteria (37). DNA extraction from Otway Basin project passages was performed using two methods, the Qiagen Blood and Tissue DNA extraction kit protocol for Gram-positive cells (Qiagen) or the MoBio soil DNA extraction kit (MoBio). Amplification of 16S rRNA genes from extracted DNA was performed using universal bacterial primers 515F 5′-GTG CCA GCM GCC GCG GTA A-3′ and 1406R 5′-ACG GGC GGT GWG TRC AA-3′ (Frio-2 passages 1, 2, and 7) and 27F 5′-AGA GTT TGA TCM TGG CTC AG-3′ and 1492R 5′-TAC GGY TAC CTT GTT ACG ACT T-3′ (Frio-2 passage 9, Otway passage 3, and colony-purified isolates). PCR mixtures (20 μl per reaction mixture) contained 25 to 75 ng of genomic DNA, 1× Phusion polymerase buffer, 0.4 μM (each) primer (IDT), 0.4 μM deoxynucleotide mixture, and 1 U Phusion polymerase (New England BioLabs). Thermal cycling conditions consisted of an initial 3 min at 95°C followed by 35 cycles of 95°C for 30 s, 52°C for 30 s, and 72°C for 90 s, followed by a final extension time of 5 min. Every PCR included negative and positive controls.

Amplified 16S rRNA gene fragments from Frio-2 samples were gel purified (Qiagen gel extraction kit) and ligated into the pJET1.2 vector (Fermentas) according to the manufacturer's protocol. Ligation products were transformed into Escherichia coli DH5α or E. coli Top10 cells, and clones were selected for sequencing (using LacZ/isopropyl-β-d-thiogalactopyranoside [IPTG] plating). For the Frio-2 enrichment, sequencing reaction mixtures were prepared using BigDye Terminator 3.1 according to the manufacturer's instructions and sequencing was performed on an ABI 3130 platform. Otway project 16S rRNA gene fragments were gel purified (Qiagen gel extraction kit) and sequenced commercially (Genewiz, Cambridge, MA). Removal of vector and primer sequences and manual editing and clustering of operational taxonomic units (OTUs) at 99% nucleotide identity were performed using Sequencher 4.5 (Gene Codes Corp.). Chimeric sequences were identified by Chimera Check 2.7 (RDP II Database) software and removed from analysis. 16S rRNA gene sequences obtained from passages of the scCO₂ enrichments from the Frio-2 and Otway Basin sites and from all colony-purified isolates were uploaded to the Ribosomal Database Project (RDP) server (38) for multiple-sequence alignment using their weighted neighbor-joining tree-building algorithm. The stability of the groupings was estimated by bootstrapping on 100 trees, and phylogenetic tree files were downloaded and visualized with MABL (39).

Isolation of strains and preparation of spores.

Samples from passages 2, 5, and 3 of the enrichment cultures from Frio-2 (sample 9-26-1039-20L), Otway Basin (core 3), and the King Island core, respectively, were plated on LB agar and incubated aerobically at ambient pressure at 37°C to obtain colonies, followed by colony purification by restreaking on LB agar and identification using DNA extraction and 16S rRNA gene sequencing. Since isolates had 16S rRNA sequence types matching cloned sequences corresponding to endospore-forming bacteria, spores were prepared as described in reference 40 to serve as inocula for subsequent characterization. For spore preparation, overnight stationary-phase cultures grown under aerobic ambient conditions in LB medium were diluted 1:50 into modified G medium which consists of the following (in g/liter): 2.0 yeast extract, 0.025 CaCl₂·2H₂O, 0.5 K₂HPO₄, 0.2 MgSO₄·7H₂O, 0.05 MnSO₄·4H₂O, 0.005 ZnSO₄·7H₂O, 0.005 CuSO₄·5H₂O, 0.0005 FeSO₄·7H₂O, 2.0 (NH₄)₂SO₄, adjusted to pH 7.1 after autoclaving. Cells were incubated with shaking for 72 h to induce sporulation and then centrifuged for 15 min at 4,000 × g. Samples were resuspended and centrifuged 5 times in autoclaved wash buffer containing 0.058 g/liter NaH₂PO₄·H₂O and 0.155 g/liter Na₂HPO₄·7H₂O with 0.01% (vol/vol) Tween 20 to prevent aggregation of spores. Spore preparations were heat treated at 80°C for 10 min to kill residual vegetative cells. Spores for each isolated strain were stored in wash buffer at 4°C until further use.

Physiological characterization.

Physiological tests for isolates displayed in Table 3 were conducted in triplicate. LB medium was used for assaying temperature, pH, and salinity ranges, with positive growth scored by an optical density at 600 nm (OD₆₀₀) of greater than 0.05. Medium pH was adjusted with NaOH or HCl followed by incubation at 37°C. Salinity was adjusted with NaCl in LB medium followed by incubation at 37°C. LB agar was used for colony morphology determination after 20 h of incubation at 37°C. Spore formation was determined by confirming viability after heat-killing cultures grown in modified G sporulation medium and by microscopic evaluation after staining for spores (41).

TABLE 3.

ScCO₂ enrichment isolate physiology

Bacillus sp. strain	16S rRNA BLAST identification (% identity)	Isolation environment	O2 requirement	Spore formation	Temp range (°C)a	pH rangeb	Salinity range (g/liter)c	Colony morphologyd
MITOT1	B. subterraneus (98.6)	Sandstone core, 1,193.6 m, 39°C; Otway Basin, Australia	Facultative anaerobe	+	23–37	4–8	0.5–50	0.2 mm, circular, entire, raised, smooth, grayish
MIT0214	B. cereus (99.8)	Formation water, 1,528–1,534 m, 55°C; Frio-2, Texas	Facultative anaerobe	+	23–45	4–10	0–40	5 mm, circular, undulate, raised, smooth, off-white
WG1	B. amyloliquefaciens (98.6)	Unconsolidated sandstone core, 1,447 m; King Island, CA	Facultative anaerobe	+	23–55	2–10	0–60	4 mm, circular, undulate, umbonate, wrinkly, off-white
WMR1	B. safensis (99.2)	Unconsolidated sandstone core, 1,447 m; King Island, CA	Facultative anaerobe	+	23–55	4–12	0–60	2 mm, circular, entire, raised, smooth, yellow-white
WMe1	B. safensis (99.1)	Unconsolidated sandstone core, 1,447 m; King Island, CA	Facultative anaerobe	+	23–45	4–12	0–60	2 mm, circular, entire, raised, smooth, yellow-white
WMe2	B. megaterium (99.4)	Unconsolidated sandstone core, 1,447 m; King Island, CA	Facultative anaerobe	+	23–45	4–10	0–60	2 mm, circular, entire, convex, smooth, yellow-white

^aTemperatures tested were 9, 23, 30, 37, 45, 55, and 65°C over 72 h.

^bpHs tested were 2, 4, 6, 7, 8, 10, and 12 over 123 h.

^cSalinities tested were 0, 0.5, 1, 10, 20, 30, 40, 50, and 60 g/liter over 72 h.

^dColony morphologies were determined after 20 h of growth on LB agar.

Measuring growth and survival in anaerobic CO₂ and N₂ atmospheres at variable pressure.

The dynamics of growth and survival of strains isolated from CO₂ sequestration sites and the type strains Bacillus subtilis PY79, Bacillus mojavensis JF-2 (ATCC 39307), and Bacillus cereus (ATCC 14579) were investigated under variable headspace gas composition and pressures. Inocula consisted of either cells or spores; vegetative cell cultures were grown overnight to stationary phase under aerobic conditions and diluted approximately 100-fold to 10⁷ cells/ml in fresh growth medium while spores maintained in wash buffer were diluted to approximately 10⁵ spores/ml unless otherwise specified. Media for MIT0214 and MITOT1 were GYP and MS media with the metal reducer supplement, respectively. Incubations at pressures from 3 to 136 atm were conducted in 316 stainless steel pressure vessels at 37°C with shaking at 100 rpm. Pressurization was achieved by regulating backpressure from the CO₂-He tank. Determination of growth dynamics under 1 atm N₂ or 1 atm 95% CO₂-5% H₂ was conducted in 100-ml serum bottles at 37°C with shaking at 100 rpm.

To test whether Bacillus spores could tolerate indirect or direct exposure to scCO₂, spores of B. cereus MIT0214 (Frio-2 isolate), B. subtilis PY79, and B. mojavensis JF-2 were aliquoted into Durham vials in equal volumes of spore storage buffer corresponding to 5 × 10⁷, 7 × 10⁵, and 1 × 10⁸ spores, respectively, and dried for 2 to 3 days at 70°C to achieve a desiccated state. Dried spores were either covered in 2 ml of fresh spore storage buffer or left desiccated and then incubated under scCO₂ (100 atm, 37°C) in a 1-liter High Pressure Equipment Company (HIP) vessel for 2 weeks. After depressurization, samples were plated for viable counts on LB medium. Desiccated samples were resuspended in 2 ml of buffer before plating.

Antibiotic-amended control experiments were used to confirm growth under scCO₂. We amended reactors containing spores with both kanamycin and chloramphenicol antibiotics at 100 and 10 μg/ml, respectively, in parallel with reactors without antibiotics. Additionally, we included no-cell controls for all experiments. Presence or absence of growth was assessed with a combination of metrics, including direct cell counts, changes in cell morphology from spores to vegetative cells, viable cell counts, and optical density.

Statistical methods.

Logistic regression analyses were performed using JMP Pro v. 10 where growth outcome (growth/no growth) was the dependent variable and incubation time and inoculating spore density were the independent variables. Student's t tests were performed in Microsoft Excel.

Nucleotide sequence accession numbers.

16S rRNA gene sequences from isolates and clones characterized in this study have been deposited in the NCBI database with accession numbers KP873174 to KP873199.

RESULTS

Enrichment cultivation under scCO₂.

Enrichment cultivation of subsurface samples in bioreactors containing nutrients, water, and scCO₂ yielded isolates from all three GCS sites examined in this study (Fig. 1; Table 2). Enrichment cultures inoculated with filtered formation fluid (Frio-2) or crushed rock cores (Otway or King Island) were evaluated for growth under an scCO₂ headspace after 14 to 30 days as evidenced in some cases by an increase in the turbidity of culture medium. All samples were evaluated by epifluorescence microscopy to confirm biomass in turbid cultures and to screen for microbial cells in nonturbid samples (Fig. 1). Microscopy and extraction of total nucleic acids confirmed biomass generation after successive rounds of dilution and growth (Table 2).

FIG 1.

Epifluorescent microscopy of enrichment cultures grown under scCO₂, filter concentrated onto 25-mm 0.2-μm-pore-size membranes, and stained with DAPI (A) or SYTO9 (B to H). Bars, 10 μm. (A) Cells from Frio-2 initial enrichment. (B) Cells from Frio-2, passage 1, counterstained with propidium iodide (red) to identify membrane-compromised cells. (C) Cells from Otway core 3, passage 3, showing larger vegetative cells and smaller cells that may be spores observed within the same sample. (D) King Island initial enrichment, metal reducer medium. (E to H) Images correspond to cultures of MIT0214, MITOT1, WMe2, and WG1, respectively. Cultures in panels E to H were inoculated with spores, with a mixture of mostly vegetative cells and some spores visible after 30 days of growth under scCO₂. Black particles (denoted by white arrowheads in panel F) correspond to metal solids from the growth medium for metal reducers.

The composition of microbial assemblages enriched under an scCO₂ atmosphere was determined by analysis of cloned 16S rRNA gene sequences for enrichments from the Frio-2 and Otway samples achieving cell biomass greater than approximately 10⁵ cells/ml. The libraries from the initial enrichment and first passage of the Frio-2 sample revealed a mixed community of microbes dominated by members of the Bacillus genera (see Fig. S1 in the supplemental material). DNA isolated from Frio-2 passages 7 and 9 revealed a single dominant Bacillus sequence type sharing 99.8% 16S rRNA sequence identity (1,463/1,465 nucleotides) with B. cereus ATCC 14579 (see Fig. S1 in the supplemental material).

Biomass was detected in enrichment cultures inoculated with four different core samples from the Otway Basin under a variety of media and environmental conditions (Table 2). However, sparse cell densities (≤10⁴ cells/ml) did not yield PCR-amplifiable DNA. Subsequent rounds of dilution and incubation in scCO₂ bioreactors were performed, leading to increased biomass consisting of a mixture of vegetative and spore-like cells for core 3 in MS medium with an oxidized metal supplement (Fig. 1C; Table 2). The 16S rRNA gene was amplified and cloned from DNA extracted from passage 3 of Otway core 3, yielding 26 clones of a single ribotype matching B. subterraneus with 98.5% identity (1,502/1,521 nucleotides) (Table 3; see also Fig. S2 in the supplemental material). scCO₂ enrichment cultures inoculated with material from the King Island core revealed positive growth in 5 out of 8 incubations (Table 2). 16S rRNA clone libraries of King Island enrichments were not pursued as the core was unconsolidated and thus fully permeated by drilling tracer fluid, which likely contaminated the microbial diversity. The 16S rRNA gene of isolates from the scCO₂ enrichment of King Island core material was sequenced following colony purification (Table 3).

Colony purification and strain characterization.

Since the Bacillus spp. identified in clone libraries are known to be predominantly aerobic and heterotrophic/copiotrophic, we attempted to isolate Bacillus strains as colonies on LB agar plates from scCO₂ enrichment passages of all samples with significant growth (>10⁵ cells/ml). When growth under scCO₂ was observed, depressurized cultures were used as inocula and subsequent colony formation under ambient aerobic conditions (37°C) was observed after 1 to 3 days. Individual colonies with distinct morphologies were colony purified and identified by 16S rRNA gene sequencing and corresponded to B. cereus (Frio-2 sample, passages 2 and 7, isolate MIT0214); Bacillus subterraneus (Otway core 3, passage 5, isolate MITOT1); and Bacillus amyloliquefaciens, Bacillus megaterium, and Bacillus safensis (2 strains) (King Island core, passage 3, isolates WG1, WMe2, WMe1, and WMR1, respectively) (Table 3; also see Fig. S2 in the supplemental material). Isolates were subjected to physiological characterization and were facultative anaerobes with variable ranges of pH, temperature, and salinity similar to previously characterized Bacillus spp. (Table 3). Isolates were subjected to regrowth experiments to confirm bacterial growth under scCO₂.

Growth of Bacillus strains MIT0214 and MITOT1 and Bacillus type strains under scCO₂.

Based on our observations of biomass increases in enrichment cultures, spore formation in isolated strains, and the lethal effects of scCO₂ on vegetative cells of isolates and type strains grown under ambient conditions (see Fig. S3 and S4 in the supplemental material), we hypothesized that spores but not vegetative cells would germinate and grow under scCO₂. To test our hypothesis, we inoculated growth medium with a range of spore concentrations (2.4 × 10⁴ to 9.9 × 10⁶ spores/ml for isolate MIT0214 and 6.9 × 10³ to 5.6 × 10⁶ spores/ml for isolate MITOT1), followed by incubation under scCO₂ from 1 week to 49 days (Fig. 2), accompanied by antibiotic (see Fig. S5 in the supplemental material) and no-cell controls. As we observed highly variable outcomes (i.e., growth/no growth) in replicate cultures of MIT0214 and MITOT1 incubated under an scCO₂ headspace, we used logistic regression (Fig. 2A and B) to determine the relationship between frequency of growth under scCO₂, incubation time, and initial spore density. For >20-day incubations of strain MIT0214, 26/78 cultures exhibited increased cell density (average final cell density of 5.6 × 10⁷ cells/ml), while 0/15 no-cell controls and 0/30 antibiotic controls displayed growth (Fig. 2A; see also Fig. S5 in the supplemental material). For ≥20-day incubations of strain MITOT1, 32/58 cultures exhibited increased cell density (average final cell density of 1.2 × 10⁷ cells/ml), while no increase in cell density was seen for no-cell controls (0/15) or antibiotic controls (0/4) (Fig. 2B; see also Fig. S5 in the supplemental material). For logistic regression analysis, no-cell controls were input at half the detection level (i.e., at 870 cells/ml), while antibiotic controls were excluded from the model. As expected, growth under scCO₂ is significantly affected by both initial inoculum concentration and incubation time (Fig. 2A: P = 0.045 and P = 0.0005, respectively, for MIT0214, n = 78; Fig. 2B: P = 0.0014 and P = 0.023, respectively, for MITOT1, n = 73). We noted that growth of MITOT1 consistently accompanied more reduced conditions in growth medium that contained the resazurin redox indicator dye and oxidized metal supplements (see Fig. S6 in the supplemental material). The incubation times associated with a 50% likelihood of growth in cultures inoculated with at least 1 × 10⁴ spores/ml are 33 days for MIT0214 and 36 days for MITOT1.

FIG 2.

Logistic regressions of MIT0214 (A) and MITOT1 (B) growth outcomes under scCO₂ show a significant increase in the frequency of observed growth with increasing incubation time (MIT0214, P = 0.0005; MITOT1, P = 0.023) and increasing density of spore inocula (MIT0214, P = 0.045; MITOT1, P = 0.0014). Model results predicting frequency of growth are represented by contour lines. Experimental data (MIT0214, n = 78; MITOT1, n = 73) are overlaid on top of the contours with symbol shade indicating frequency of growth and symbol size proportional to number of replicates represented by each point. Circles are cell-containing samples, and squares are no-cell controls that were entered into the model at half the detection limit (870 cells/ml).

Growth of Bacillus type strains and King Island isolates under scCO₂.

After observing growth under scCO₂ of isolates MIT0214 and MITOT1, we investigated whether other Bacillus strains, including subsurface isolates from the King Island site and additional Bacillus type strains, could germinate and grow under scCO₂. Spores of three isolates from the King Island core (WG1, WMe1, and WMe2) and three Bacillus type strains, B. mojavensis JF-2, B. subtilis PY79, and B. cereus ATCC 14579, were inoculated into growth medium and incubated under scCO₂ for 30 days. King Island isolates and Bacillus type strains yielded increased cell densities after incubation (Fig. 3A and B, respectively). However, growth patterns were variable between replicates as previously observed during experiments with strains MIT0214 and MITOT1 (Fig. 2). This unexpected result appears to indicate that the ability to grow in environments containing scCO₂ may be widespread among Bacillus spp.

FIG 3.

Microscopic enumeration of cell abundance in cultures grown under scCO₂ for King Island isolates (A) and three Bacillus type strains (B. subtilis PY79, B. mojavensis JF-2, and B. cereus ATCC 14579) (B). Data shown are total direct cell counts after 30 days of incubation, with horizontal lines representing initial cell counts. Initial spore densities ranged from 3.2 × 10⁵ to 9.5 × 10⁵ spores/ml.

Variable survival of cells and spores after exposure to scCO₂.

In contrast to the observed germination and growth of spores in bioreactors containing scCO₂, vegetative cells grown under aerobic conditions and ambient pressure were not able to acclimate to scCO₂ exposure and grow in our experimental system. Vegetative cells from overnight stationary-phase cultures of isolate B. cereus MIT0214 and type strains B. cereus ATCC 14579, B. subtilis PY79, and B. mojavensis JF-2 were exposed to scCO₂ for 6 h at 37°C. Reductions in viable cell counts of 4 to 8 orders of magnitude were observed for all strains exposed to scCO₂ relative to controls incubated under ambient conditions (see Fig. S3 and S4 in the supplemental material). Cells surviving exposure to scCO₂ were most likely spores based on similar proportions of cells surviving heat-kill treatments (see Fig. S4 in the supplemental material).

Spores from B. cereus MIT0214, B. subtilis PY79, and B. mojavensis JF-2 were robust to scCO₂ exposure under both aqueous and desiccated conditions. Spores that were dried and then resuspended in aqueous buffer and exposed to scCO₂ showed no significant change in viability after 2 weeks relative to the initial viable count or controls incubated under ambient conditions (P > 0.05) (Fig. 4A). While dried spores directly exposed to scCO₂ were more susceptible to the killing effect, 14 to 24% of spores remained viable after 2 weeks relative to ambient-incubated controls for B. mojavensis and B. cereus (P < 0.05) and for B. subtilis (P = 0.08) (Fig. 4B). Strain B. cereus MIT0214, isolated early in the project, was used as a model scCO₂-tolerant isolate for most analyses. Anecdotally, spores of strain B. subterraneus MITOT1 appeared to be less robust than spores of other Bacillus spp. tested. In the spore tolerance experiment, the pretreatment protocol to desiccate the spores resulted in total loss in viability for MITOT1, and thus, no data are available for this strain. Overall, this experiment indicated that spores from diverse Bacillus strains can withstand both direct and indirect exposure to scCO₂, with enhanced survival of spores under aqueous conditions (Fig. 4; also see Fig. S7 in the supplemental material).

FIG 4.

Change in viable cell count (CFU/ml, final/initial) in spores from three Bacillus species exposed to scCO₂ for 2 weeks. (A) Spores resuspended in spore preparation buffer (dark triangles) revealed no decrease in viability upon exposure to scCO₂ relative to starting spore counts and ambient-atmosphere-incubated controls (open circles). B. subtilis spores may have germinated and grown during the incubation periods under ambient conditions. (B) Spores exposed to dry scCO₂ (dark triangles) decreased in viability by 66 to 86%.

Growth of MIT0214 under variable CO₂ and N₂ pressure.

We incubated strain MIT0214 under subcritical pressures (<72.9 atm) of N₂ and CO₂ to observe whether more consistent growth outcomes (growth/no growth) would accompany more permissive growth conditions. At 1-atm pressure, anaerobic growth under CO₂ or N₂ was reproducible among triplicate incubations with increases in turbidity and viable cell counts revealing lag, log, stationary, and decline phases over a 1-week time frame (see Fig. S8A and B in the supplemental material). Considerably lower germination frequencies were observed under 1-atm CO₂ (<19%) than 1-atm N₂ (>99%) (see Fig. S8A in the supplemental material). In contrast to reproducible growth observed at ambient pressure, growth at all pressures above 1 atm under either N₂ or CO₂ revealed positive growth but variability in outcome (i.e., growth/no-growth) between replicates over 1- and 2-week time frames, respectively, similar to the variability observed for cultures incubated under scCO₂ (see Fig. S9 in the supplemental material).

DISCUSSION

We demonstrated that bacterial growth under an scCO₂ atmosphere is possible using strains isolated through enrichment cultivation of samples from three sites targeted for geologic sequestration of CO₂: filtrate samples from the Frio-2 project in Texas; consolidated rock cores from the Paaratte formation in the Otway Basin, Australia; and an unconsolidated core from the Mokelumne formation at the King Island site in California. In addition, we observed germination and growth under scCO₂ of three Bacillus type strains isolated from subsurface (B. mojavensis JF-2) and surface (B. subtilis PY79 and B. cereus ATCC 14579) environments. These observations of microbial growth under scCO₂ have several implications for geological carbon dioxide sequestration (GCS). (i) Microbial survival and activity are possible, and perhaps likely, at the CO₂ plume-water interface, even in areas previously exposed to pure-phase scCO₂. (ii) Engineering biofilm barriers or stimulating biomineralization in situ at the CO₂ plume-water interface may be feasible, despite previously documented lethal properties of scCO₂ for nonacclimated microorganisms. (iii) Organisms likely to survive and proliferate after CO₂ injection include spore-forming microbes that can withstand the initial CO₂ stresses.

Phylogenetic and physiological analyses reveal that strains isolated from the three GCS sites are similar to previously described Bacillus isolates that are readily cultured copiotrophs. Anaerobic enrichment medium formulations, although supplemented to target diverse metabolic groups, contained a base of organic carbon (yeast extract and peptone), potentially explaining the dominance of Bacillus spp. in our enrichments. While the catabolic pathways for different strains recovered were not determined in this study, previous studies suggest that closely related Bacillus spp. may grow anaerobically by fermentation and/or anaerobic respiration. Strain MIT0214 is most closely related to B. cereus—a widely distributed bacterium that has been isolated from a diverse range of environments, including sites characterized by heavy metals, deep oil reservoirs, and/or hypersaline conditions (42–45). Strain MITOT1 is closely related to strains B. subterraneus and Bacillus infernus, both of which were isolated from deep subsurface environments (46, 47). King Island isolates are closely related to strains of B. megaterium, B. safensis, and B. amyloliquefaciens, which have all been isolated from diverse terrestrial environments (48–53). In light of the diversity of Bacillus strains able to grow under scCO₂, we hypothesize that the ability to grow under high pCO₂ is a widespread feature of Bacillus spp. and possibly other organisms.

All Bacillus spp. isolated in this study were sporeformers capable of facultative anaerobic growth under both N₂ and CO₂ atmospheres. Notably, we have observed that all strains exhibit the fastest growth under aerobic conditions and atmospheric pressure but are also able to grow anaerobically at elevated pressures beyond the critical point for CO₂ (greater than 37°C and 73 atm). Growth dynamics of strain MIT0214 at 1 atm are consistent with anaerobic growth via fermentation where end product inhibition leads to a reduction of cell and spore viability in stationary phase (see Fig. S8A and B in the supplemental material). In strain MITOT1, the observed change in culture oxidation state that accompanied cell growth (as demonstrated by the resazurin indicator) suggests that MITOT1 may be capable of Fe(III) or Mn(IV) reduction (see Fig. S6 in the supplemental material), although whether oxidation state changes occur via fermentation or respiration has not been established in this study. We note that several close phylogenetic relatives of strain MITOT1 are anaerobic metal reducers capable of Fe(III), Mn(IV), Se(VI), and As(V) respiration (46, 47, 54).

Spores of three Bacillus strains, including isolate B. cereus MIT0214, are tolerant of both direct and indirect exposure to scCO₂. Direct exposure of dried spores to scCO₂ proved to be a more severe stress, resulting in a 66 to 86% loss in spore viability over a 2-week interval, while no significant loss was observed in spores indirectly exposed via aqueous phase (Fig. 4; also see Fig. S7 in the supplemental material). The decrease in viability of spores directly exposed to scCO₂ may be due to its solvating and desiccating properties, as spores in the aqueous phase do not experience the same degree of contact with scCO₂ and are protected from dehydration. While it remains to be determined whether the viability decrease is time dependent, our observation of a decrease of <1 order of magnitude in survival of desiccated spores exposed to scCO₂ suggests that spores in subsurface environments will likely be able to tolerate at least brief periods of direct exposure to pure-phase scCO₂. This finding is especially relevant in the GCS context because as buoyant scCO₂ migrates toward the cap-rock interface, spores may experience fluctuations in scCO₂ contact in areas where formation fluids have been displaced, creating local areas of more extreme and desiccating conditions. Persistence of a subpopulation of spores after direct scCO₂ exposure suggests that scCO₂ injection during GCS may not effectively sterilize the subsurface, enabling the resumption of microbial activities upon return of an aqueous phase. Our results, and previous demonstrations of spore resistance to scCO₂ sterilization (9), suggest that spore-forming microbial taxa may be able to survive and acclimate to in situ conditions post-CO₂ injection during geologic carbon dioxide sequestration.

Previous work suggests that several features of Bacillus physiology are likely responsible for the variable, but predictable, time- and density-dependent growth observed under scCO₂. First, germination frequencies under CO₂ headspaces are known to be significantly lower than those under aerobic or anaerobic nitrogen headspaces, with only 10 to 30% of B. cereus spores germinating under 1 atm CO₂ (55). Growth dynamics for isolate B. cereus MIT0214 (see Fig. S8A in the supplemental material) were consistent with these findings, displaying a germination frequency at 1 atm CO₂ no greater than 19%, in contrast to a 99% germination frequency under 1 atm N₂ (see Fig. S8A in the supplemental material). Second, it has been shown that certain stresses, such as pH and oxidative stresses, significantly extend the lag time for germination of Bacillus spores (55, 56). We have observed extended lag phases for growth of MIT0214 under 1 atm CO₂ compared to 1 atm N₂ (see Fig. S8A and B in the supplemental material) and have also observed extended lag times for growth of all isolates at low pH (pH <6) (data not shown) during physiological tests (Table 3). The third mechanism that may explain the observed variability in growth has been termed the “microbial scout hypothesis,” where activation of cells from dormant bacterial populations (e.g., spores) is stochastic but occurs at a constant low frequency (i.e., ∼0.01% per day) independent of environmental cues. This behavior would allow a clonal population to sense and respond to permissive growth conditions without risking activation of the entire dormant population (57). Fourth, it has been shown that modulation of the sporulation conditions has a significant effect on spore physiology and resistance to stress (58, 59), and we suggest that a population of spores may contain a range of this physiological plasticity where some, but not all, spores are able to grow following germination under stressful conditions. Finally, microenvironments within cultures (e.g., associated with biofilms or mineral surfaces) may influence the degree to which cells are exposed to stresses associated with scCO₂ (17, 60). Thus, variability in exposure and stress tolerance of spores in combination with extended lag times under conditions of high pCO₂ and low pH, and inherently stochastic and suppressed germination frequencies, may account for observed variability of growth under scCO₂.

Microbial biomass obtained from ongoing GCS projects supports the notion that spore-forming Firmicutes populations, such as Bacillus spp., persist and potentially grow in the subsurface after scCO₂ injection. Recovery of phospholipid fatty acid (PLFA) from Frio-2 groundwater samples indicated the presence of viable microorganisms after CO₂ injection, especially Firmicutes, as indicated by the observation of characteristic terminally branched saturated fatty acids. In particular, i15:0, i17:0, and a17:0, which are prominent fatty acids in B. cereus (61, 62), were observed in the sample from the Frio-2 project from which B. cereus strain MIT0214 was isolated and in samples following CO₂ breakthrough (S. Pfiffner, personal communication). Microbial communities recovered in formation waters of the Otway Basin (Australia) and Ketzin (Germany) GCS sites (10 to 12 days and 2 days to 10 months post-CO₂ injection, respectively) include taxa of the Firmicutes (e.g., Bacilli and Clostridia) among the detectable diversity. Finally, in experimental incubation of formation fluids under scCO₂, enrichment of Firmicutes (Clostridiales) was observed after removal of scCO₂, demonstrating the resilience of native Firmicutes populations to scCO₂ exposure (63).

While some populations of Bacillus are well established as members of subsurface biospheres (46, 47), others may be introduced to the subsurface through drilling activities. Although sampling methods for Frio-2 formation water and Otway cores were designed to reduce the possibility of drilling fluid contamination through use of a U-tube sampling device (33) or fluorescein drilling fluid penetration data, respectively, drilling activities remain a potential source for some of the strains recovered in this study. Indeed, the King Island core, which yielded the highest number of scCO₂-tolerant isolates, was unconsolidated sandstone and fully penetrated by drilling fluid based on tracer data. Notably, drilling fluid characterized by Mu et al. (22) was dominated by sequences of Bacilli.

High concentrations of local dissolved organic carbon at the edges of injected CO₂ plumes, as observed during the Frio-2 project (29, 64), may stimulate growth of native or introduced bacteria. Enrichment of dissolved organic carbon near the plume may be due to extraction of the subsurface organic matrix by the nonpolar solvent-like properties of scCO₂. We suggest that these enrichments of dissolved organic carbon at the leading edge of the CO₂ plume may serve as potential hot spots for microbial activity fueled by anaerobic metabolism. Previous observations that DNA and lipid biomass from Firmicutes populations persist in scCO₂-exposed environments (21, 22), coupled with the presented evidence that a diverse set of Bacillus strains and isolates may grow in the presence of scCO₂, water, and nutrients, suggest that microbially mediated processes may continue to occur up to the scCO₂ plume-water interface during geologic carbon dioxide sequestration. Understanding how CO₂ injection shifts the balance of microbial community structure, metabolic potential, and associated biogeochemical processes is therefore necessary to predict the long-term fate of injected CO₂.