Kinetics of calcite precipitation by ureolytic bacteria under aerobic and anaerobic conditions

. The kinetics of urea hydrolysis (ureolysis) and induced calcium carbonate (CaCO 3 ) precipitation for engineering use in the subsurface was investigated under aerobic conditions using Sporosarcina pasteurii (ATCC strain 11859) as well as Bacillus sphaericus strains 21776 and 21787. All bacterial strains showed ureolytic activity inducing CaCO 3 precipitation aerobically. Rate constants not normalized to biomass demonstrated slightly higher-rate coefﬁcients for both ureolysis ( k urea ) and CaCO 3 precipitation ( k precip ) for B. sphaericus 21776 ( k urea = 0 . 10 ± 0 . 03 h − 1 , k precip = 0 . 60 ± 0 . 34 h − 1 ) compared to S. pasteurii ( k urea =

Abstract. The kinetics of urea hydrolysis (ureolysis) and induced calcium carbonate (CaCO 3 ) precipitation for engineering use in the subsurface was investigated under aerobic conditions using Sporosarcina pasteurii (ATCC strain 11859) as well as Bacillus sphaericus strains 21776 and 21787. All bacterial strains showed ureolytic activity inducing CaCO 3 precipitation aerobically. Rate constants not normalized to biomass demonstrated slightly higher-rate coefficients for both ureolysis (k urea ) and CaCO 3 precipitation (k precip ) for B. sphaericus 21776 (k urea = 0.10 ± 0.03 h −1 , k precip = 0.60 ± 0.34 h −1 ) compared to S. pasteurii (k urea = 0.07 ± 0.02 h −1 , k precip = 0.25 ± 0.02 h −1 ), though these differences were not statistically significantly different. B. sphaericus 21787 showed little ureolytic activity but was still capable of inducing some CaCO 3 precipitation. Cell growth appeared to be inhibited during the period of CaCO 3 precipitation. Transmission electron microscopy (TEM) images suggest this is due to the encasement of cells and was reflected in lower k urea values observed in the presence of dissolved Ca. However, biomass regrowth could be observed after CaCO 3 precipitation ceased, which suggests that ureolysis-induced CaCO 3 precipitation is not necessarily lethal for the entire population. The kinetics of ureolysis and CaCO 3 precipitation with S. pasteurii was further analyzed under anaerobic conditions. Rate coefficients obtained in anaerobic environments were comparable to those under aerobic conditions; however, no cell growth was observed under anaerobic conditions with NO − 3 , SO 2− 4 or Fe 3+ as potential terminal electron accep-tors. These data suggest that the initial rates of ureolysis and ureolysis-induced CaCO 3 precipitation are not significantly affected by the absence of oxygen but that long-term ureolytic activity might require the addition of suitable electron acceptors. Variations in the ureolytic capabilities and associated rates of CaCO 3 precipitation between strains must be fully considered in subsurface engineering strategies that utilize microbial amendments.

Introduction
Carbonate precipitation is a natural phenomenon that may also be utilized in many subsurface engineering applications (Phillips et al., 2013a) including soil stabilization (van Paassen et al., 2010), immobilization of radionuclides Ferris, 2005, 2006a, b;Tobler et al., 2012;Warren et al., 2001), and mineral plugging for enhanced oil recovery and carbon sequestration (Dupraz et al., 2009;Ferris et al., 1996;Mitchell et al., 2010;Phillips et al., 2013b). Mineral precipitation can be induced by bacteria as a by-product of common microbial processes, such as urea hydrolysis (ureolysis). In this process, bacteria hydrolyze urea (CO(NH 2 ) 2 ), an important nitrogen compound found in natural environments, to ammonia (NH 3 ) and carbonic acid (H 2 CO 3 ) (Eqs. 1-3). The NH 3 and H 2 CO 3 equilibrate in circumneutral aqueous environments to form bicarbonate (HCO − 3 ), two ammonium ions (NH + 4 ) and one hydroxide ion Published by Copernicus Publications on behalf of the European Geosciences Union.
(OH − ) (Eqs. 4-5), or at higher pH values, to one carbonate ion (CO 2− 3 ) and two NH + 4 (Eqs. 4-6). In the presence of dissolved calcium (Ca 2+ ), this increase in carbonate alkalinity shifts the saturation state of the system, allowing for solid calcium carbonate (CaCO 3 ) to form (Eq. 7). The overall reaction from the hydrolysis of urea in the presence of Ca 2+ is summarized by Eq. (8).
CO(NH 2 ) 2 + H 2 O → NH 2 COOH + NH 3 (1) CO(NH 2 ) 2 + 2H 2 O → 2NH 3 + H 2 CO 3 (Eqs. 1 + 2 overall) (3) CO(NH 2 ) 2 + 2H 2 O + Ca 2+ ↔ 2NH + 4 + CaCO 3 (overall process) (8) The use of ureolytic bacteria in biotechnological applications is appealing for many reasons. Ureolytically active microorganisms are common in a wide variety of soil and aquatic environments; thus, indigenous microorganisms capable of ureolysis can be either stimulated in situ or, alternatively, they can be used to augment environments lacking ureolytic microorganisms (Fujita et al., 2000;Warren et al., 2001). Urea is a fairly inexpensive substrate and it is often contained in wastewater (Hammes et al., 2003b), so this waste product may be used to stimulate ureolysis in engineering applications (Mitchell et al., 2010). Moreover, the controlled increase in pH and alkalinity in the subsurface by ureolytic bacteria is preferable to the injection of a basic solution (abiotic process), which could lead to instantaneous CaCO 3 supersaturation and precipitation at the point of injection limiting the radius of influence of the technology. The injection of urea into the subsurface followed by microbially induced ureolysis would allow for controlled, gradual ureolysis further away from the injection point, promoting a wider spatial distribution of CaCO 3 in the subsurface and avoiding uncontrolled plugging at the point of injection (Cuthbert et al., 2013;Ebigbo et al., 2012;Mitchell and Ferris, 2005;Schultz et al., 2011;Tobler et al., 2012Tobler et al., , 2014. Among different ureolytic bacteria, Sporosarcina pasteurii (formerly known as Bacillus pasteurii) has been extensively used as the model urease-producing organism in ureolysis-driven CaCO 3 precipitation studies, due to its high ureolytic activity and constitutive production of urease (Phillips et al., 2013a). The use of S. pasteurii for CaCO 3 precipitation is feasible under aerobic conditions and the kinetics of ureolysis under different conditions have been studied. Most studies have reported first-order ureolysis rates with respect to urea concentration, with the rate constant (k urea ) ranging between 0.002 and 0.090 h −1 under aerobic conditions in artificial groundwater without nutrients added (Dupraz et al., 2009;Ferris et al., 2004;Hammes et al., 2003a;Mitchell and Ferris, 2005;Tobler et al., 2012) and 0.35 h −1 with the addition of nutrients (Lauchnor et al., 2015). Ureolysis rates have been suggested to be temperature-dependent (Ferris et al., 2004), and it seems to also be affected by cell concentration (inoculum size) (Lauchnor et al., 2015;Tobler et al., 2011).
Although the ureolytic activity of S. pasteurii under anoxic conditions has been observed (Martin et al., 2012;Mortensen et al., 2011;Tobler et al., 2012), there is controversy regarding the extent and duration of ureolytic activity that can be achieved in the absence of oxygen. Mortensen et al. (2011) and Tobler et al. (2012) observed extensive ureolytic activity under anoxic conditions, suggesting that the anoxic environment does not inhibit urease activity. Conversely, Martin et al. (2012) observed limited cell growth and poor ureolysis under anoxic conditions and suggested that the ureolytic activity observed was due to the urease already present in the cells.
In this study, the ability of S. pasteurii to grow in the absence of oxygen (with or without nitrate (NO − 3 ), sulfate (SO 2− 4 ) or ferric ion (Fe 3+ ) as possible electron acceptors) was investigated along with the kinetics of ureolysis and CaCO 3 precipitation. Moreover, this study investigates and compares the ureolytic activity of S. pasteurii with different strains of Bacillus sphaericus under aerobic conditions, which have also been suggested to be capable of ureolysisinduced CaCO 3 precipitation (Dick et al., 2006;Hammes et al., 2003a).

Solutions
Kinetic experiments were carried out using the CaCO 3 mineralizing medium (CMM) described by Ferris et al. (1996) (see Table SI1.1 in the Supplement). Both Ca 2+ -inclusive (CMM+) and Ca 2+ -exclusive (CMM−) versions of this medium were used. Aerobic CMM− was prepared as follows. A double-strength solution of nutrient broth was prepared and autoclaved. Nutrient broth was chosen in this experiment to enable cell growth. A separate solution of double-strength urea, ammonium chloride and sodium bicarbonate was prepared and stirred until completely dissolved. These two solutions were combined and adjusted to a pH of 6.0 using concentrated HCl. CMM+ was prepared similarly, but calcium chloride was added after the pH adjustment. Media were filter-sterilized into sterile Pyrex bottles using 0.2 µm pore size filters (Nalgene, Rochester, NY). Anaerobic CMM was produced in the same manner. However, all stock solutions were made in an anaerobic chamber using water that had been degassed by stirring overnight in the oxygenfree atmosphere of the chamber (90 % N 2 , 5 % CO 2 , 5 % H 2 ). Solutions were filter-sterilized into serum bottles and were then capped and sealed inside the chamber. Prior to the

Bacterial strains and culturing conditions
Three strains of ureolytic bacteria were used: S. pasteurii (ATCC 11859) and two isolates from a garden soil and landfill soil -B. sphaericus 21776 and B. sphaericus 21787 (Belgian Coordinated Collections of Microorganisms, Laboratory of Microbiology, Ghent University) (Hammes et al., 2003a). Bacillus subtilis strain 186 (ATCC 23 857), a nonureolytic organism, was used as a control species. Abiotic (i.e., non-inoculated) controls were also set up and run in parallel. Pilot cultures were grown in flasks by adding 100 µL of thawed stock to 100 mL of autoclaved brain heart infusion (BHI) +2 % urea. S. pasteurii and B. sphaericus were grown on an incubator shaker (New Brunswick Scientific, Edison, NJ) at 30 • C and 150 rpm, while B. subtilis was grown on an incubator shaker at 37 • C and 150 rpm. A total of 100 µL of pilot cultures was transferred at 24 and 48 h to new flasks containing 100 mL of BHI +2 % urea.

Aerobic experiments
Once the pilot cultures were ready for inoculation, 40 mL of culture was added to a 50 mL centrifuge tube. This tube was centrifuged at 4303 × g using a Sorvall Instruments (Asheville, NC) RC-5C centrifuge for 10 min at 4-6 • C. The supernatant was poured off and the cell pellet was resuspended using about 40 mL of CMM− and again centrifuged for 10 min. This process was repeated once more. After the third run in the centrifuge, the supernatant was poured off and enough CMM− was added to achieve a final optical density reading at 600 nm (OD 600 ) of 0.4 (CMM-blank corrected, measured in a 96-well plate using a BioTek Synergy HT plate reader). A total of 1 mL of prepared S. pasteurii, B. sphaericus strain 21776 or strain 21787 culture was added to 250 mL Pyrex bottles with 150 mL of media (either CMM+ or CMM−; initial concentration of biomass OD 600 =∼ 0.015). After inoculation, the systems were statically incubated at 30 • C for kinetic experiments.

Anaerobic experiments
Pilot cultures for anaerobic experiments were limited to the use of S. pasteurii and were grown in the same manner as those used in aerobic experiments. However, cells were transferred into an anaerobic chamber and resuspended in anaerobically prepared CMM−, then transferred to a serum bottle, sealed and crimped inside the anaerobic chamber. Optical density measurements for time zero were taken after the final suspension, with the same initial density as the aerobic experiments (OD 600 =∼ 0.015). The first set of experiments investigated cell growth and ureolysis under oxygenfree conditions with a range of potential terminal electron acceptors (TEAs). Experiments were run using a batch system, consisting of 100 mL of CMM− media including 10 mM NO − 3 , SO 2− 4 or Fe 3+ as potential TEAs and inoculated with 1 mL of S. pasteurii in 150 mL serum bottles. Concentrated stock solutions of each TEA were made in the anaerobic chamber and filter-sterilized: (i) a 1 M solution of NaNO 3 ; (ii) a concentrated SO 2− 4 solution, made by combining 1 M Na 2 SO 4 and 1 M Na 2 S, where Na 2 S was added to quench any residual oxygen and make SO 2− 4 reduction possible; and (iii) a stock solution of Fe(III) citrate, using 50 mM Fe(III) citrate as previously described . Appropriate amounts of each stock solution were added to the separate serum bottles containing CMM− and S. pasteurii. The growth survey was also conducted in CMM− without the addition of a TEA. After inoculation, the systems were statically incubated at 30 • C. Abiotic control experiments, without the inclusion of S. pasteurii, were also performed. Aliquots were extracted from the systems in the anaerobic chamber and monitored for pH and OD 600 during the duration of the experiments. Comparative aerobic control experiments were also performed with CMM− media including 10 mM NO − 3 , SO 2− 4 or Fe 3+ and inoculated with 1 mL of S. pasteurii in 150 mL serum bottles.
The second set of experiments investigated the detailed kinetics of ureolysis and CaCO 3 precipitation with S. pasteurii and CMM+ as described above, with NO − 3 as the potential TEA, by monitoring pH, dissolved Ca 2+ and NH + 4 concentrations. Control experiments were also performed with CMM+ without the addition of a TEA and CMM− with NO − 3 as a potential TEA. Here, a stock solution of 10 M NaNO 3 was made by mixing and filter-sterilizing in the anaerobic chamber, and an appropriate amount was added to the CMM+ or CMM− to reach a final concentration of 1 M NO − 3 .

Experimental sampling and analysis
At different time points, 3 mL of sample was aseptically extracted from the systems and measurements were made of pH, biomass, NH + 4 and Ca 2+ concentration. NH + 4 concentrations were determined using the Nessler Assay (Mitchell and Ferris, 2005). Urea concentrations were determined from NH + 4 concentrations, according to Eqs. (3) and (4). Ca 2+ concentrations were measured after appropriate dilution in trace-metal grade 5 % HNO 3 (Fisher Scientific) using an Agilent 7500ce inductively coupled plasma mass spectrometer (ICP-MS). Bacterial biomass was determined using three methods: plate counts, OD 600 and protein assays. OD 600 was used as a growth indicator in experiments carried out in the absence of Ca 2+ (see Sect. 1.2 in the Supplement). Transmission electron microscopy (TEM) images were taken using a LEO 912AB TEM and photographed with a Proscan 2048 × 2048 charge-coupled device (CCD) camera from a batch culture in CMM+ inoculated with S. pasteurii. At the point of crystal formation (after approximately 2.5 h), a mixture of CaCO 3 crystals and cells were extracted from the syswww.biogeosciences.net/16/2147/2019/ Biogeosciences, 16, 2147-2161, 2019 tem. Separate samples of S. pasteurii grown in the absence of Ca 2+ were also collected and imaged. Further details are given in the Supplement Sect. 1.3. The PHREEQC (version 2) speciation-solubility geochemical model (Parkhust and Appelo, 1999) was used to calculate solution speciation and carbonate mineral saturation. The simulation was performed using calcite as the only precipitate as it was identified by X-ray diffraction (XRD) as the calcium carbonate polymorph present in the systems. The MINTEQ database was used for all calculations, and the thermodynamic constants for urea (Stokes, 1967) were added (more information is provided in SI1.4 in the Supplement).

Kinetics of ureolysis and CaCO 3 precipitation
The rate coefficient for ureolysis was determined by integrating the following first-order differential equation assuming constant biomass concentrations during the period of urea hydrolysis (Ferris et al., 2004;Mitchell and Ferris, 2005): to get: where k urea is the first-order rate coefficient for ureolysis, t is the time and X is the concentration of biomass (Sect. SI2.1 in the Supplement). First-order relationships have been successfully used to describe microbial ureolysis (Lauchnor et al., 2015;Connolly et al., 2015). Indeed, while the Michaelis-Menten model has been used when evaluating ureolysis, studies of ureolysis-based carbonate precipitation have demonstrated that the first-order ureolysis rate model fits well for urea concentrations of 330 mM or below (Lauchnor et al., 2015;Connolly et al., 2015), which is the concentration range used in this study. The change in urea concentration was determined according to Eqs. (3) and (4) (Eq. 11).
Biomass-normalized ureolysis rates were calculated. Firstly, it was assumed that biomass (X) is constant in Eqs. (10) and (11), as performed in other studies of ureolysis kinetics (Cuthbert et al., 2012;Ferris et al., 2004;Ferris, 2005, 2006;Schultz et al., 2011;Tobler et al., 2011). Secondly, the obtained first-order rate coefficients with respect to urea concentration (k urea ) were normalized to the biomass concentration by dividing the ureolysis rate coefficient by the biomass at the onset of precipitation (i.e., urea hydrolysis rates were normalized to the initial biomass concentration, determined as either OD 600 or CFU mL −1 ; Supplement Sect. 2.2), which was equivalent to the initial biomass in each system (X = X 0 ). This is an appropriate choice of model, since the biomass analysis indicated that the cell density was constant for the duration of CaCO 3 precipitation and was equivalent to the initial biomass in the systems, as presented in the results section. The biomass-normalized ureolysis rates were compared to other parameters previously published (Ferris et al., 2004;Fujita et al., 2000;Stocks-Fischer et al., 1999;Tobler et al., 2011). The media used in the different studies were similar to those used in this study, all with 25 mM of Ca 2+ , 333 mM urea and including nutrient-broth-based growth media, apart from (i) Ferris et al. (2004), who used a dilute artificial groundwater (nongrowth medium) with Ca 2+ and urea concentrations of 1.75 and 6 mM, respectively, and (ii) Tobler et al. (2011), who used different Ca 2+ concentrations varying from 50 to 500 mM and urea concentrations between 250 and 500 mM. The precipitation of CaCO 3 from the system is dependent on the saturation state of the system, as well as the growth mechanism of CaCO 3 (Ferris et al., 2004;Mitchell and Ferris, 2005;Teng et al., 2000). The literature is ambiguous on defining a set rate expression for CaCO 3 precipitation, so a non-affinity-based first-order rate law was applied to these studies for both its simplicity and the fact that it seems to describe the data well, assuming that for every mole of Ca 2+ removed from the solution 1 mol of CaCO 3 formed (Teng et al., 2000): Integration of the above equation yields where k precip is the first-order rate coefficient for CaCO 3 precipitation. Rate constants were found using a nonlinear regression method utilizing the Solver function in Microsoft Excel. Due to the significant lag phase, the data used for analysis excluded onset time before ureolysis and CaCO 3 precipitation occurred, as previously documented (Tobler et al., 2011).

Results and discussion
3.1 Aerobic experiments

Solution chemistry
Aerobic experiments with a CMM+ medium inoculated with S. pasteurii, B. sphaericus 21776 and B. sphaericus 21787 showed an increase in pH (Table 1) and NH + 4 (displayed as a stoichiometrically equivalent decrease in urea concentrations; see Fig. 1) over time. These results support observations from previous studies confirming the ureolytic capabilities of S. pasteurii (Ferris et al., 2004;Fujita et al., 2000;Warren et al., 2001) and B. sphaericus species (Dick et al., 2006;Hammes et al., 2003a). Differences in the rate of pH change and the amount of urea hydrolyzed between the different bacterial species suggest differences in their ureolytic  (Mitchell and Ferris, 2005). Control experiments, inoculated with the non-ureolytic organism B. subtilis and sterile controls, did not exhibit significant changes in pH or urea concentrations (Table 1, Fig. 1). Geochemical modeling suggested that no CaCO 3 precipitation should occur in the absence of ureolysis and that approximately 0.45 mM of urea would have had to be hydrolyzed to achieve supersaturation and for CaCO 3 precipitation to commence (see Supplement Sect. SI1.4).
In all the experiments containing ureolytic bacteria, Ca 2+ concentration decreased to ∼ 5 % of the initial Ca 2+ concentration in the liquid medium after approximately 30 h (Fig. 1). The decrease in Ca 2+ concentrations suggests the precipitation of CaCO 3 , which was identified by XRD as the polymorph calcite. The onset of CaCO 3 precipitation occurred shortly after the start of the experiment (∼ 3-4 h) in the S. pasteurii and B. sphaericus 21776 experiments, whereas the onset of precipitation was delayed (∼ 9 h) for B. sphaericus 21787. This supports differences in the rate of urea hydrolysis and thus the time at which CaCO 3 saturation was exceeded (Eqs. 2, 4, 5 and 7). Differences in ureolysis and CaCO 3 precipitation rates can be attributed to differences in the specific ureolytic activities of the organisms or the number of ureolytically active cells (Anbu et al., 2016;Hammes et al., 2003a). B. sphaericus 21776 and S. pasteurii exhibit urease activities approximately twice that of B. sphaericus 21787 when Ca 2+ is present (Hammes et al., 2003a), supporting our observations of limited ureolysis and delayed CaCO 3 precipitation by B. sphaericus 21787. Nevertheless, the decrease in Ca 2+ concentrations in the absence of significant urea hydrolysis for B. sphaericus 21787 suggests there was a sufficient carbonate ion concentration in the CMM+ and from ureolysis to sustain CaCO 3 precipitation. Hammes et al. (2003a) observed B. sphaericus 21787 was also able to precipitate CaCO 3 despite lower urease activity in the presence of Ca 2+ , suggesting this strain may enhance precipitation via other mechanisms such as enhanced nucleation on cell surfaces or via organic exudates (Mitchell et al., 2006b).

Aerobic bacterial growth and ureolytic activity
Changes in biomass, measured as protein and colonyforming units (CFUs), were observed during ureolysis in both aerobic CMM+ and CMM− experiments (Fig. 2). CFUs and protein concentrations exhibited similar trends for S. pasteurii and B. sphaericus 21776, where in CMM− experiments CFUs and protein increased over time asymptot- ically (Fig. 2). In CMM+ experiments, CFUs and protein concentrations seemed to slightly decrease or remain quasiconstant while CaCO 3 precipitation occurred (< 10 h), followed by an increase in CFUs and protein once Ca 2+ had been depleted (Figs. 1 and 2). Decrease in biomass growth during CaCO 3 precipitation has been suggested to occur due to the encasement of bacteria within the CaCO 3 precipitates (Tobler et al., 2011). Encasement of S. pasteurii cells in CaCO 3 minerals has been reported (Cuthbert et al., 2012;Ebigbo et al., 2012;Schultz et al., 2011;Stocks-Fischer et al., 1999), and cell indentations in CaCO 3 precipitates have been observed (Mitchell and Ferris, 2005). The recovery (i.e., regrowth) of biomass after CaCO 3 precipitation suggests that  Fig. 2). TEM images and electron energy-loss spectroscopy (EELS) of material collected on 0.2 µm pore size filters from the CMM+ S. pasteurii systems (Fig. 3a-c) confirm that some cells are surrounded by a layer of calcium-containing precipitates. Figure 3d shows S. pasteurii grown in CMM− for comparison. The data suggest that cells are removed from suspension and potentially inactivated by CaCO 3 encasement, either in large crystals Ferris, 2005, 2006a) or by a thin coating (Fig. 3a-c).
Quasi-constant biomass concentrations during CaCO 3 precipitation (Fig. 2) suggest that cell growth might not have to be considered in kinetic descriptions of bacterially induced CaCO 3 precipitation. However, it is unclear whether (i) CaCO 3 -encased cells are ureolytically active or (ii) CaCO 3 precipitates surrounding the cells effectively act as a barrier to urea reaching the cell or to NH 3 , OH − or NH + 4 formed by the hydrolysis of urea from diffusing through the CaCO 3 to the bulk solution. Therefore, a theoretical analysis of urea diffusion in CaCO 3 was performed. The diffusion of oxygen in CaCO 3 at high temperatures has been documented (Farver, 1994), but, to the best of our knowledge, information on urea diffusion in CaCO 3 at 30 • C has not been reported. A number of assumptions were made for the estimates in this study: (1) since urea has a lower diffusion coefficient than oxygen in aqueous solutions at 25 • C (Stewart, 2003), it was assumed that this will hold true at other temperatures and through other substances, like CaCO 3 ; (2) since the diffusion coefficient of oxygen through CaCO 3 at 400 • C and 100 MPa is 2.66 × 10 −22 m 2 s −1 (Farver, 1994), and diffusion coefficients generally increase with increasing temperature and pressure, it can be assumed that the diffusion coefficient of urea in CaCO 3 at atmospheric pressure and 30 • C is smaller than 2.66 × 10 −22 m 2 s −1 ; (3) assuming that the geometry of the CaCO 3 is a uniformly thick slab with a thickness of approximately 200 nm, as determined from the TEM images (Fig. 3b), the time it will take to reach 5 % of the bulk urea concentration can be calculated using the relation presented by Carslaw and Jaeger (1959): where L is the slab thickness, D e is the (estimated) effective diffusion coefficient in CaCO 3 and t 5 is the amount of time it will take to reach 5 % of the bulk concentration. Using the above assumptions, it would take at least 175 d for 5 % of the urea to diffuse through the CaCO 3 surrounding the cells. Because CaCO 3 precipitation takes place over the course of approximately 1 d, it can safely be assumed that even if the encased cells are still alive, urea is not able to diffuse through the CaCO 3 fast enough for them to hydrolyze significant amounts and contribute to the increase in solution alkalinity. Therefore, it is argued that, at least in the systems described here, cell growth does not have to be considered in kinetic expressions describing ureolysis during CaCO 3 precipitation. Instead, the biomass at the onset of precipitation, which is equivalent to the initial biomass in the system (Fig. 2), can be used to normalize the observed ureolysis rates to biomass concentration. While a physical association between cells and CaCO 3 precipitates is evident, precipitation is likely to occur from a combination of (i) homogeneous nucleation in the bulk solution in alkaline microenvironments around bacterial cells (Schultze-Lam et al., 1996;Stocks-Fischer et al., 1999), and (ii) heterogeneous nucleation on nascent crystals, bottle walls

Kinetics of ureolysis and CaCO 3 precipitation
Kinetic analyses were performed on the individual CMM+ and CMM− experimental data for all bacterial strains (Fig. SI2.1 Table 2 (for detailed results  on individual experiments see Table SI2.1 in the Supplement). k urea values for the bacterial species varied according to the presence (CMM+) or absence (CMM−) of Ca 2+ in the medium. S. pasteurii and B. sphaericus 21776 exhibited statistically insignificant differences in k urea values (t test p value = 0.27) from each other in both CMM− and CMM+ systems with k urea values being between 1.6 and 2.5 times higher in the absence of Ca 2+ (S. pasteurii: k urea,CMM+ = 0.07±0.02 h −1 , k urea,CMM− = 0.19±0.10 h −1 ; B. sphaericus 21776: k urea,CMM+ = 0.10 ± 0.03 h −1 , k urea,CMM− = 0.16 ± 0.05 h −1 ). This is likely due to the encasement of cells by CaCO 3 and their inactivation in CMM+ experiments. Some data points were excluded for S. pasteurii CMM− k urea calculations because of an estimated increase in urea concen-tration (based on a decrease in NH + 4 concentration) likely due to significant volatilization of NH + 4 ↔ NH 3 + H + that can occur at pH > 9 (at 34 h; open marker, Fig. SI2.1b). B. sphaericus 21787 exhibited low k urea values in both CMM+ (k urea = 0.02 h −1 ) and CMM− (k urea = 0.05 h −1 ). From triplicate experiments, only one experiment showed values that could be used for kinetic analysis (Fig. S2.1), and some outlying data points were not used for the kinetic calculations; hence, no standard deviations can be provided (Table 2)
The kinetic parameters obtained in this study were compared to other parameters previously published (Tobler et al., 2011;Ferris et al., 2004;Fujita et al., 2000;Stocks-Fischer et al., 1999). k urea values of S. pasteurii obtained in this study as well as in previous publications were standardized to the initial cell concentrations (Table 3). Values of k urea were higher in the present study for both S. pasteurii and B. sphaericus 21776 (k urea = 0.07 and 0.11 h −1 , respectively) than those from S. pasteurii in other studies (k urea = 0.005 to 0.095 h −1 ). This was also apparent once normalized to biomass (this study, B. sphaericus 21776 k urea = 8.02 OD −1 600 h −1 and S. pasteurii 5.251 OD −1 600 h −1 , compared to k urea = 0.11 to 2.80 OD −1 600 h −1 in previous studies). The generally higher k urea values in this study appear to reflect the higher temperature (30 • C) used compared to the previous studies, which ranged from 20 to 25 • C. Higher temperatures generally increase reaction rates where chemical re-Biogeosciences, 16, 2147-2161, 2019 www.biogeosciences.net/16/2147/2019/ actions are advanced through a transient activated complex (Stumm and Morgan, 1996). In urease, the transitional state involves coordination of urea and water at the active catalytic site of the enzyme (Jabri et al., 1995). Formation of such an activated complex tends to impart a greater temperature dependency on the absolute reaction rate than would be encountered if the reactions were mediated solely by collisions arising from molecular diffusion (Ferris et al., 2004;Mitchell and Ferris, 2005). The biomass concentration-normalized k urea values from Fujita et al. (2000) are much lower. This could be due to the highest biomass concentrations (OD 600 = 0.072) used in these studies; very high biomass concentrations could shift the primary kinetic dependency from being catalysts (i.e., enzyme-limited) to substrate-limited. However, this appears to be opposite to what Tobler et al. (2011) reported indicating k urea increased with increasing inoculum density. The rate constants obtained with the three organisms used in the present study are similar to the range of values measured in deeper vadose zone mineral subsoils which were between 0.00375 to 0.07 h −1 (Swensen and Bakken, 1998), suggesting natural levels of ureolytic bacterial activity were reasonably approximated in the aerobic experiments.
On average, B. sphaericus 21776 had the highest k precip (0.60 ± 0.34 h −1 ), although considering its high standard deviation, k precip for S. pasteurii is not significantly different (k precip = 0.25 ± 0.02 h −1 ; t test p value 0.21). R 2 values of the fit to Eq. (12) were relatively high (0.84-0.93) for B. sphaericus 21776 and S. pasteurii ( Table 2). The k precip for B. sphaericus 21787 was lower than the other strains (0.21 h −1 ); although, as noted, rate constants for this strain are not statistically significant. The lag time for CaCO 3 precipitation was 3.3 h for B. sphaericus 21776 and S. pasteurii, which reflects the similar k urea values, and thus the similar time it took to reach CaCO 3 saturation and induce precipitation, whereas the longer lag time for B. sphaericus 21776 reflects the significantly lower k urea value. Tobler et al. (2011) observed similar lag times until CaCO 3 precipitation (2-3 h) in aerobic experiments (artificial groundwater with no nutrients added, 250-500 mM urea and 50-500 mM Ca) with S. pasteurii, as observed here for B. sphaericus 21776 and S. pasteurii. First-order rate constants for CaCO 3 precipitation observed here for B. sphaericus 21776 and S. pasteurii were also higher (0.21 h −1 < k precip < 0.60 h −1 ) compared to other studies (0.01 h −1 < k precip < 0.11 h −1 ) ( Table 3). This is likely associated with the greater k urea values observed in this study than in previous studies. Temperature is unlikely to account for this variation given the modest decrease in calcite solubility (∼ 27 %) that occurs between 20 and 30 • C (Miller, 1952;Stumm and Morgan, 1996). Overall, k urea values are lower than k precip values in this and previous studies, with the exception of Ferris et al. (2004), indicating urea hydrolysis is the rate limiting step during ureolysis-induced CaCO 3 precipitation and that CaCO 3 precipitation rates are rapid and controlled by the rate of urea hydrolysis, once the critical supersaturation is exceeded (Mitchell and Ferris, 2005).

Anaerobic ureolysis and bacterial growth
Given the potential for anaerobic conditions in subsurface environments, screening experiments were performed to assess the capability of S. pasteurii to grow and/or increase pH in CMM− in the presence of NO − 3 , SO 2− 4 and Fe 3+ as potential TEAs. There are contradicting reports in the literature regarding S. pasteurii's ability to grow and hydrolyze urea in the absence of oxygen; some studies suggest that the anoxic environment does not hinder urease activity (Mortensen et al., 2011;Tobler et al., 2011), whereas other studies report limited microbial growth and poor ureolysis (Martin et al., 2012).
In this study, pH increased (pH > 9.0) in all experiments with and without potential TEAs added (Fig. 4), suggesting ureolysis by S. pasteurii took place in the absence of oxygen. The pH of the abiotic controls did not exceed pH 7, except for the medium containing SO 2− 4 as the added TEA, which had an initial pH of 7.7 and remained constant throughout the experiments (Fig. 4c). Growth in the CMM− anaerobic experiments quantified by OD 600 absorbance was lower than that observed in aerobic experiments regardless of the presence or absence of TEAs (Fig. 4). Although some growth might have occurred during the initial period of the experiments (increased absorbance by ∼ 20 h), the lack of sustained growth over time suggests the inability of S. pasteurii to grow under anaerobic conditions. These findings are in agreement with those by Martin et al. (2012), who observed limited cell growth in the absence of oxygen. In the present study, once oxygen was allowed to diffuse into the systems (at 120 h), optical density in all inoculated systems increased, indicating that even though no significant growth was observed in the absence of oxygen, the bacteria were still viable after 120 h of oxygen depletion and can be resuscitated.
Sustained growth of S. pasteurii in the absence of oxygen does not appear to be feasible which might limit the potential use of S. pasteurii for inducing CaCO 3 precipitation in the subsurface to only short-term purposes. However, the potential regrowth of microbes, even after prolonged periods of exposure to oxygen-free conditions, suggests that S. pasteurii could be resuscitated and restimulated through the injection of oxygenated fluids, which could enable bacterial growth and thus ureolytic activity over longer periods of time.

Kinetics of anaerobic ureolysis and CaCO 3 precipitation
After the screening experiments with different TEAs, studies were performed to determine the kinetics of ureolysis and CaCO 3 precipitation in the absence of oxygen. Since there  3.00 × 10 −8 3.73 × 10 −10 1.81 × 10 −9 9.86 × 10 −10 7.13 × 10 −10 7.13 × 10 −10 3.56 × 10 −9 4.56 × 10 −9 2.11 × 10 −9  were similarly low levels of growth in the initial anaerobic screening experiments with different TEAs and no TEA (Fig. 4), kinetic experiments in the absence of oxygen were conducted with no TEA, as well as with NO − 3 as a potential TEA (Fig. 5). Urea was hydrolyzed under all experimental conditions and CaCO 3 precipitation was observed in the presence of Ca 2+ (Fig. 5). However, ureolysis and CaCO 3 precipitation did not occur in all replicates for each experiment, which accounts for the high standard deviations.
Rate coefficients were estimated as previously described for aerobic experiments (Fig. SI2.2) and a summary of the results is presented in Table 2 (for detailed results for the  individual experiments see Table SI2.2). Data points which preceded the onset and completion of ureolysis and calcite precipitation were excluded (Fig. SI2.2) k urea seems to be lowest in CMM+ with NO − 3 (k urea = 0.04 ± 0.01 h −1 ), followed by CMM− with NO − 3 (k urea = 0.07 ± 0.01 h −1 ) and CMM+ without TEA (k urea = 0.08 h −1 ). The same relative rates were apparent when k urea values were normalized to biomass ( Table 2). The presence of NO − 3 appears to have slightly decreased ureolysis rates. While the reasons for this possible decrease in the ureolytic activity due to the presence of NO − 3 are unclear, it could be perceivable that the ureolytic activity is downregulated in the presence of the alternative nitrogen source NO − experiments relative to the aerobic ones. Moreover, comparing the experiments containing NO − 3 , the presence of Ca 2+ in the medium results in a lower k urea value (Table 2). This could be due to the encasement of cells by CaCO 3 .
The onset of CaCO 3 precipitation occurred after approximately 6 h, which was twice the lag time observed for precipitation under aerobic conditions (Table 2); this was expected, as slower ureolysis was observed in the anaerobic experiments (Table 2). Rate constants (k precip ) for anaerobic CaCO 3 precipitation were not statistically different for CMM+ with NO − 3 (0.36 h −1 ± 0.22) and CMM+ without TEA (0.19 h −1 ± 0.05). Tobler et al. (2011) reported a similar k urea value (0.09 h −1 ) for experiments with S. pasteurii under anoxic conditions in natural groundwater containing comparable concentrations of urea and Ca 2+ (250 and 50 mM, respectively); k precip was not reported due to a poor fit of the data with first-order kinetics. Experiments containing only indigenous bacteria exhibited far lower rates of ureolysis (k urea = 0.0016 h −1 ) and CaCO 3 precipitation (k precip = 0.009 h −1 ) than observed in the present study. Differences in ureolysis and CaCO 3 precipitation rates between this study and Tobler et al.'s (2011) study are likely due to the low initial biomass of the indigenous ureolytic population in the natural groundwater.
The comparison of kinetics from aerobic and anaerobic experiments in the present study demonstrates that rates are on the same order of magnitude (Table 2). In CMM+, urea hydrolysis rates for S. pasteurii under aerobic and anaerobic conditions (with or without TEA) were not significantly different (P value = 0.274), even when normalized to initial biomass concentrations (OD 600 or CFU mL −1 ). pH increases in these screening experiments suggest anaerobic ureolysis occurred at the same rate as under aerobic conditions (Fig. 4). Similarly, k precip in aerobic and anaerobic experiments are comparable (0.19 and 0.25 h −1 , respectively). This suggests that oxygen-free environments do not significantly impact the rate of ureolysis or CaCO 3 precipitation initially but that anaerobic growth cannot be conclusively demonstrated under the conditions of this present study. This supports observations by Tobler et al. (2011), who reported similar rates of ureolysis under both oxic and anoxic conditions when amending natural groundwater with S. pasteurii (k urea 0.10 h −1 in oxic conditions, k urea 0.097 h −1 in anoxic conditions with 50 mM Ca 2+ and 250 mM urea; Table 3). Martin et al. (2012) also observed ureolytic activity by S. pasteurii under anoxic conditions but to a lesser extent compared to the extensive activity reported by Tobler et al. (2011); however, rate constants were not reported. The current study therefore suggests ureolytic activity observed under anoxic conditions corresponds to the urease already present in the cells, as suggested by Martin et al. (2012). S. pasteurii could therefore be used for CaCO 3 -induced precipitation in the subsurface in the short term, and bacterial growth could be stimulated through multiple injections of bacterial cells or oxygenated medium to re-enable ureolytic activity and thus CaCO 3 precipitation. This is supported by our calcite precipitation rate constants under anaerobic conditions, the first to be reported for a ureolytic strain, which are comparable to aerobic rate constants, suggesting anaero-