Article pubs.acs.org/est
Inhibition of Sporosarcina pasteurii under Anoxic Conditions: Implications for Subsurface Carbonate Precipitation and Remediation via Ureolysis Derek Martin,* Kevin Dodds, Bryne T. Ngwenya, Ian B. Butler, and Stephen C. Elphick School of Geosciences, Grant Institute, University of Edinburgh, The Kings Buildings, Edinburgh EH9 3JW, United Kingdom ABSTRACT: The use of Sporosarcina pasteurii to precipitate calcium carbonate in the anoxic subsurface via ureolysis has been proposed for reducing porosity and sealing fractures in rocks. Here we show that S. pasteurii is unable to grow anaerobically and that the ureolytic activity previously shown under anoxic conditions is a consequence of the urease enzyme already present in the cells of the aerobically grown inoculum. The implications are discussed, suggesting that de novo synthesis of urease under anoxic conditions is not possible and that ureolysis may decline over time without repeated injection of S. pasteurii as the urease enzyme degrades and/or becomes inhibited. Augmentation with a different ureolytic species that is able to grow anaerobically or stimulation of natural communities may be preferable for carbonate precipitation over the long term.
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INTRODUCTION Sporosarcina pasteurii has been widely studied for its role in ureolysis-driven CaCO3 precipitation and its potential role in a number of diverse applications such as soil stabilization,1 radionuclide stabilization,2 and carbon sequestration.3,4 One thing these and other studies have in common is the proposed use of S. pasteurii in the subsurface. This approach is feasible where oxygen is present; however, the growth of S. pasteurii under anaerobic conditions in this context is equivocal, with some authors suggesting it is a facultative anaerobe,5−7 whereas others state that it is an (obligate) aerobe8,9 or that biomass production is inhibited under low oxygen conditions.10 The ambiguity in the metabolism of S. pasteurii may result from the experimental methodology used, since very few studies have been performed under truly anaerobic conditions. Previous investigations into microbially induced carbonate precipitation (MICP) using S. pasteurii can be split into two basic types: (1) Those where the experiments were started in air and were exposed to air throughout, or were closed systems that may have eventually been driven anoxic during the course of the experiment (for example, refs 4,11, and 12). The key point here is that oxygen was present at the start of the experiment either dissolved in the medium or in both the headspace and the medium, and this is true for the majority of studies. (2) Those performed completely under anaerobic conditions from start to finish (refs 7 and 13). Here oxygen was removed from the growth medium and all preparation and the experiment were performed under completely anoxic conditions. In both these cases, ureolysis rates were found to be unaffected by anoxia over time scales of up to ca. 30 h; however, no data were presented to © 2012 American Chemical Society
confirm actual cell division or growth under these conditions. It has been proposed that S. pasteurii could be used as a foreign inoculum for MICP in anoxic environments if indigenous bacteria lack the required ureolytic activity.7 The question remains, however, as to whether the ureolytic activity observed under anoxic conditions is a function of cell growth and/or metabolism or whether ureolysis is proceeding due to urease already present in the initial cell inoculum, which was prepared aerobically but may be inactive under anaerobic conditions. If the latter is the case, the feasibility of using S. pasteurii under anoxic conditions becomes problematic. Although there may be some ureolytic activity when the cells are initially injected into the subsurface, the cells will be dormant or eventually be killed and urease will not be replaced, the existing enzyme will eventually degrade and/or be inhibited,14 and ureolytic activity will cease in the long term. The purpose of this study was to ascertain whether S. pasteurii is able to grow and/or metabolize anaerobically and, therefore, determine whether the ureolytic activity previously observed under anoxic conditions is just a consequence of the urease already present in the cells. The methodology used was to chemically inhibit the growth of S. pasteurii by use of the antibiotic chloramphenicol to be sure that cell division had ceased; comparisons could then be made with cells grown aerobically and anaerobically. Received: Revised: Accepted: Published: 8351
April 20, 2012 June 28, 2012 July 2, 2012 July 9, 2012 dx.doi.org/10.1021/es3015875 | Environ. Sci. Technol. 2012, 46, 8351−8355
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MATERIALS AND METHODS
The base growth medium used for all experiments contained nutrient broth (no. 3, Fluka, 13 g·L−1) supplemented with 250 mM urea at a pH of 6.7 ± 0.1. No calcium was added to the medium to prevent interference from precipitating calcium, since only growth and ureolytic activity were under investigation. S. pasteurii (NCIMB 8841) was grown aerobically at 30 °C to exponential phase in filter-sterilized (0.22 μm) medium. The cells were harvested via centrifugation (22000g, 10 min) and then washed by resuspending the pellet in sterile urea-free nutrient broth (pH 6.7) and centrifuging again. The cells were then resuspended in nutrient broth to an optical density at 600 nm of 0.14 (Camspec, M501 UV/vis spectrophotometer, 1 cm path length). Experiments were performed in 100 mL of medium; therefore, 50-mL aliquots of this suspension were centrifuged and the pellet subsequently used for the inoculum gave a final optical density of 0.07 when resuspended in 100 mL of medium. For oxic experiments, the pellet was resuspended in sterile medium and added to conical flasks at a final volume of 100 mL stoppered with foam plugs. Anoxic medium was prepared by sparging sterile medium with oxygen-free nitrogen via a sterile 0.22 μm filter for ≥40 min. The medium was transferred into an anaerobic chamber (Saffron alpha, 100% nitrogen atmosphere) and 100-mL aliquots were measured into sterile vials that were crimp-sealed with sterile butyl septa. The cell pellets, contained in 50-mL centrifuge tubes, were transferred into the anaerobic chamber and resuspended in a small volume of medium taken from the crimp vial via a sterile syringe. This whole inoculum was then injected back into the crimp vial to obtain the final inoculum. Where required, 0.5 mL of chloramphenicol was provided from a stock solution (made up in 100% ethanol) to give a final concentration of 30 μg·mL−1, and 0.5 mL of 100% ethanol was added to untreated experiments accordingly. Both oxic and anoxic experiments were incubated at 30 °C on a rotary shaker (60 rpm), and all experiments were performed in triplicate. At each sampling 4 mL of solution was removed aseptically, and anoxically where required, and the pH and optical density were measured immediately. One milliliter of sample was centrifuged for 10 min and the supernatant was used to measure NH4+ via the Nessler assay,1 whereby 50 μL of Nessler reagent was added to 1 mL of sample, which was diluted accordingly, and absorbance was measured at 425 nm. Repeated analysis of a standard showed the precision to be ±2%. Anoxic experiments were also performed with the addition of nitrate as a terminal electron acceptor and glucose as a possible substrate for fermentation to explore growth more fully under different conditions. The medium consisted of nutrient broth (13 g·L−1) and urea (300 mM) with the addition of either KNO3 (10 mM) or glucose (1 g·L−1). The initial optical density used in these experiments was lower at 0.003.
Figure 1. Changes in (A) optical density, (B) pH, and (C) NH4+ for S. pasteurii grown aerobically and anaerobically with and without the addition of chloramphenicol. Data points represent the mean of triplicate experiments with error bars to show the standard deviation.
higher concentration of NH4+ than the anaerobic treatments after 24 h, although the difference is small and the overall concentration is still much lower than the increase shown in the aerobic treatment without chloramphenicol (Figure 1C). Figure 2A shows that no significant growth was observed in medium containing either nitrate or glucose, although a slight increase in pH was observed in both treatments (Figure 2B). Our results demonstrate that ureolytic activity is still present in cultures of S. pasteurii even where cells appear not to be actively growing, either under anaerobic conditions or where the growth inhibitor chloramphenicol is present. However, under anoxic conditions the rate of NH4+ production is much lower than that observed by Tobler et al.,7 who measured NH4+ production an order of magnitude higher than this study over the same time period of ca. 30 h, or by Mortensen et al.,13 who show 5−10 times the concentration of NH4+ being produced in 1 h, depending on the inoculum size used. In the case of the chloramphenicol-free anoxic treatment, even where an incubation period of up to 96 h was used, the NH 4 +
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RESULTS AND DISCUSSION In all treatments, pH and NH4+ increased when incubated both oxically and anoxically, although the only significant cell growth, measured as optical density at 600 nm, is in the aerobically grown cells without the addition of chloramphenicol (Figure 1). The increase in NH4+ is lower in the anoxic treatments than the oxic treatments, and this is reflected in the slightly lower pH values, which also reflect the individual NH4+ concentrations (compare Figure 1 panels B and C). The aerobic treatment with addition of chloramphenicol shows a 8352
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pKa value of the NH4+/NH3 equilibrium is about 9.25, as the pH increases, outgassing of ammonia increases, thus reducing the measured concentration of NH4+. NH4+ is used as a surrogate measurement to estimate the amount of urea that has been hydrolyzed over time (for example, refs 4 and 7). Urease activity may be affected by the concentration of NH4+ and is pH-dependent, with activity peaking at a pH of about 8,16,17 although Mortensen et al.13 found no inhibitory effect of NH4+ concentration on urease activity over the range tested. Consequently, the relationship between the pH of the growth medium and ureolysis rates, as measured by NH4+ production, can complicate comparisons. Overall, however, it is the actual production of NH4+ and pH increase, rather than absolute rates, which is important. None of these effects has a negative impact on the comparisons and conclusions established in this study. The purpose of chemically inhibiting the cells was to show that urease was still active within the cells even where no cell division was possible. The antibiotic chloramphenicol was chosen since it is known to have a bacteriostatic effect by inhibiting protein synthesis. Other antibiotics with bacteriolytic effects, such as penicillins or cephalosporins,18 were not used so as to reduce the chances of cell lysis and subsequent release of urease into the medium, since the initial inoculum would have been actively dividing (though that is not to say that cell permeability was not affected at all in this study). Other physical techniques to kill cells, such as heating or freezing, were also likely to physically break open the cells. Application of chloramphenicol at 30 μg·mL−1 was shown to inhibit cell division (Figure 1A), but whether the cells were actually killed, rather than rendered unable to divide, was not determined. The possibility exists that cell division and urease production could have been decoupled, meaning urease production continued while cell division ceased. However, this seems unlikely given the fact that chloramphenicol is known to inhibit protein synthesis and the low concentration of NH4+ observed in the chloramphenicol treatments (much larger concentrations of NH4+ would be expected if urease production continued). Additionally, chloramphenicol may have some effect on the activity of urease itself,19 but the fact that the anoxic chloramphenicol-free treatment showed similar NH4+ concentrations as that of the chloramphenicol-treated cells suggests any effect was minimal at this concentration of chloramphenicol. Although this study does not categorically show that chloramphenicol inhibits the production of urease, the fact that chloramphenicol is a known protein production inhibitor and little, if any, new cell growth is observed suggests that de novo synthesis of urease is minimal, at best, and a comparison with the anaerobic treatment can be made. If the ureolytic activity observed is purely due to that already present in the cells, the use of S. pasteurii in completely anoxic environments would appear to be redundant as a means of producing urease in situ. Enzyme supply would be proportional to the amount of cells added to the system; indeed, Tobler et al.7 and Mortensen et al.13 showed greater ureolytic activity the larger the inoculum they used. Similarly, increasing the concentration of free urease (from jack beans) has been shown to increase ureolysis and subsequent CaCO3 production under oxic conditions at least.20 Moreover, where a much smaller inoculum was used for the nitrate and glucose treatments in the current study, a smaller increase in pH was observed than in those experiments where a much larger inoculum was used (compare Figures 1B and 2B),
Figure 2. Changes in (A) optical density and (B) pH for S. pasteurii grown anaerobically with 10 mM NO3 and 1 g·L−1 glucose. Data points represent the mean of triplicate experiments with error bars to show the standard deviation. Note that initial cell density of the inoculum is lower than in the other experiments.
concentration in this study is still 9 times smaller than that observed by Tobler et al.7 after 30 h incubation (see Figure 1C). Sneath et al.15 reported that the ureolytic activity of S. pasteurii may be lost when it is maintained on synthetic media; however, the substantial increase in growth and NH 4 + production observed under standard aerobic conditions relative to the other treatments shows that this is probably not relevant in the current study (Figure 1C). Why the rates of ureolysis under anoxic conditions should differ so widely between this study and those of Mortensen et al.13 and Tobler et al.7 is unclear. The inoculum size and concentration of urea used in this study were identical to that used by Tobler et al.,7 who did show that inoculum size changed the rate constant for urea hydrolysis. The growth medium and inoculum sizes used in the study of Mortensen et al.13 were different than those used in the current study; however, even where Mortensen et al.13 used a smaller inoculum, ureolysis rates were much faster than in the current study. Although the same strain of S. pasteurii has been used in most MICP studies, including the current one, it is conceivable that differences in activity may be observed between different laboratory cultures depending on storage conditions and use. One thing to note is that the medium used to grow the initial inoculum and the time of harvesting is different in all three studies, which may have had an effect on the urease per cell in the initial inoculum. For instance, high nitrogen concentrations in the medium may suppress urease expression; consequently, under anaerobic conditions, where no cell growth is observed, ureolysis rates could potentially differ widely. Whiffin10 did show that even with the same concentration of biomass, urease activity could vary by a factor of more than 10. Also, since the 8353
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suggesting a smaller initial concentration of urease (NH4+ was not measured in the nitrate and glucose experiments shown in Figure 2). The fact that ureolysis is observed with S. pasteurii under anoxic conditions is not in question. Both Mortensen et al.13 and Tobler et al.7 have shown extensive activity under anoxic conditions, and in the current study some ureolysis does occur, though to a lesser extent. The question remains as to the possible uses of S. pasteurii in the subsurface. For example, Ferris et al.21 suggested that, for remediation via MICP, the addition of ureolytic bacteria is preferable to a direct application of an alkaline solution because the gradual hydrolysis of urea is likely to promote a wider spatial distribution of precipitation than the precipitation brought about by the direct addition of an alkaline chemical. Tobler et al.7 proposed that if nutrient stimulation of indigenous subsurface communities fails, S. pasteurii could be grown in large numbers aerobically and then injected into the subsurface to increase urea hydrolysis. They note that S. pasteurii may have to be injected repeatedly to ensure continued hydrolysis because the cells are continually being killed by being embedded in precipitating calcite; however, the results presented here suggest that S. pasteurii does not grow anaerobically (at least under the conditions presented in this study) and that any urease activity is likely to be a function of the urease already present in the cells at the time of injection. As well as being embedded in the precipitating calcite, any decrease in ureolytic activity is just as likely to be due to metabolic inhibition, enzyme degradation, and cell death under anaerobic conditions. Ureolysis may still be a feasible option in the anoxic subsurface when the cells are initially applied; however, whether ureolytic activity could be sustained long-term in any significant amount without the application of oxygen is open to question. Since in situ growth in the anoxic subsurface is unlikely, the same effect could be obtained by injecting pure enzyme. Urease has been applied as free enzyme to induce carbonate precipitation under oxic conditions,20,22 and there is no reason to presume that this would not be feasible under anoxic conditions, although the use of pure enzyme in solution is likely to produce a diffuse treatment area, whereas cell retention onto the surrounding matrix may help constrain and allow more control over MICP, for instance, into engineered mineralized biofilms.4,11 However, that is not to say that S. pasteurii could not be used as a source of “unrefined” urease itself, which may be cheaper and easier to produce directly in the field rather than using purified enzyme from other sources. Potentially, it may also be possible to exploit the fact that S. pasteurii is unable to grow under anoxic conditions yet still produce short-term ureolytic activity. Less biomass growth would reduce the possibility of organic cell material being included in the mineral precipitate, which may affect its long-term stability. Cells could be injected into the subsurface for MICP without the excessive biomass production that could clog the treatment area and prevent the flowthrough of treatment solutions. Repeated injection of oxygenated treatment fluids may allow S. pasteurii to survive over longer periods, although this may be dependent on the distance from injection points and would increase treatment expense. Stimulation of natural ureolytic bacteria in the subsurface has been shown by the addition of nutrients and urea to natural communities in the field23,24 and experimentally in the laboratory using natural communities under completely anaerobic conditions,7 so biological treatment may still be feasible without the use of S. pasteurii. In reality, circumstances
may dictate the treatment type and further research is required to investigate ureolysis and MICP in the field. Obviously, the type of growth medium is critical and the one used in the present study may not supply the correct constituents for anaerobic growth. Preliminary experiments where the anoxic medium was supplemented with 10 mM NO3 and 1 g·L−1 glucose showed no growth after 6 days of incubation (Figure 2A). One cannot state categorically that anaerobic growth is not possible, but at the very least, growth is severely restricted under anoxic conditions. The confusion in the status of S. pasteurii growth (formerly Bacillus pasteurii25) under anaerobic conditions probably arises from use of the standard descriptions of B. pasteurii taken from Bergey’s Manual of Determinative Bacteriology (various years) that have propagated throughout the literature. Edition 6 of Bergey26 states that B. pasteurii is an aerobe, whereas the seventh edition states that it is a facultative anaerobe and growth is observed on urea glucose broth under anaerobic conditions,27 although Norris28 clearly states that B. pasteurii is unable to utilize glucose. It appears that part of the description updated in the seventh edition is based on work by Smith et al.29 during their revision of the Bacillus group, where they examined four cultures of putative B. pasteurii. The contradiction between the present study and that of Smith et al.29 is likely to be due to the different methodologies used. In the present study, the medium was sparged with oxygen-free nitrogen and all preparation was performed in an anaerobic chamber to eliminate all but traces of oxygen. Smith et al.29 steamed the medium to drive off free oxygen, cooled it, and then inoculated it before sealing with a mixture of paraffin and Vaseline. Butler et al.30 have subsequently shown that boiling water to remove oxygen is one of the least effective methods of the four they testedto remove dissolved oxygen from solution, with oxygen concentrations up to 5 times higher than just sparging water with nitrogen, as was the case in the present study. This, coupled with the fact that Smith et al.29 presumably performed their manipulations in the open air (the use of an anaerobic chamber or oxygen-free gassing jets is not mentioned), suggests that the original tests for anaerobic growth may be questionable due to the presence of some oxygen in the growth medium. Indeed, Smith et al.29 did state that gas production from anaerobic growth on nitrate was variable. Additionally, Knight and Proom31 noted that there was only a trace reaction to anaerobic growth, although they used various different strains of B. pasteurii and the exact methodology was not stated. S. pasteurii (NCIMB 8841) is unable to grow anaerobically under the conditions used in this study. The ureolytic activity observed under anoxic conditions is due to the urease enzyme already present in the cells as a consequence of their initial aerobic production; de novo synthesis of urease is unlikely to take place once cells are exposed to anoxia. S. pasteurii could still be used as a source of enzyme in the anoxic subsurface, but ureolysis may decline over time as the urease degrades or becomes inhibited. Repeated injections of cells and/or oxygenated medium may have to be made, especially if the intention is to induce ureolysis in areas distant from the injection point. Augmenting with a different ureolytic species that is able to grow anaerobically or stimulation of natural communities may be preferable for carbonate precipitation over the long term, although the precise treatment type is likely to depend on individual circumstances. 8354
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(16) Fidaleo, M.; Lavecchia, R. Kinetic study of enzymatic urea hydrolysis in the pH range 4−9. Chem. Biochem. Eng. Q. 2003, 17, 311−318. (17) Stocks-Fischer, S.; Galinat, J. K.; Bang, S. Microbiological precipitation of CaCO3. Soil Biol. Biochem. 1999, 31, 1563−1571. (18) Madigan, M. T. Biology of Microorganisms, 10th ed.; Prentice Hall: Upper Saddle River, NJ, 2003. (19) Jahns, T.; Zobel, A.; Kleiner, D.; Kaltwasser, H. Evidence for carrier-mediated, energy-dependent uptake of urea in some bacteria. Arch. Microbiol. 1988, 149, 377−383. (20) Nemati, M.; Voordouw, G. Modification of porous media permeability, using calcium carbonate produced enzymatically in situ. Enzyme Microb. Technol. 2003, 33, 635−642. (21) Ferris, F. G.; Phoenix, V.; Fujita, Y.; Smith, R. Kinetics of calcite precipitation induced by ureolytic bacteria at 10 to 20°C in artificial groundwater. Geochim. Cosmochim. Acta 2003, 68, 1701−1710. (22) Bachmeier, K. L.; Williams, A. E.; Warmington, J. R.; Bang, S. S. Urease activity in microbiologically-induced calcite precipitation. J. Biotechnol. 2002, 93, 171−181. (23) Fujita, Y.; Taylor, J.; Gresham, T.; Delwiche, M.; Colwell, F.; McLing, T.; Petzke, L.; Smith, R. Stimulation of microbial urea hydrolysis in groundwater to enhance calcite precipitation. Environ. Sci. Technol. 2008, 42, 3025−3032. (24) Burbank, M. B.; Weaver, T. J.; Green, T. L.; Williams, B. C.; Crawford, R. L. Precipitation of calcite by indigenous microorganisms to strengthen liquefiable soils. Geomicrobiol. J. 2011, 28, 301. (25) Yoon, J. H.; Lee, K. C.; Weiss, N.; Kho, Y. H.; Kang, K. H.; Park, Y. H. Sporosarcina aquimarina sp. nov., a bacterium isolated from seawater in Korea, and transfer of Bacillus globisporus (Larkin and Stokes 1967), Bacillus psychrophilus (Nakamura 1984) and Bacillus pasteurii (Chester 1898) to the genus Sporosarcina as Sporosarcina globispora comb. nov., Sporosarcina psychrophila comb. nov. and Sporosarcina pasteurii comb. nov., and emended description of the genus Sporosarcina. Int. J. Syst. Evol. Micr. 2001, 51, 1079−1086. (26) Breed, R. S., Murray, E. G. D., Smith, N. R., Eds.; Bergey’s Manual of Determinative Bacteriology, 6th ed.; Bailliere, Tindall & Cox: London, 1948. (27) Breed, R. S., Murray, E. G. D., Smith, N. R., Eds.; Bergey’s Manual of Determinative Bacteriology, 7th ed.; Bailliere, Tindall & Cox: London, 1957. (28) Norris, J. R. Sporosarcina and Sporolactobacillus. In The Aerobic, Endosporeforming Bacteria; Berkeley, R. C. W., Goodfellow, M., Eds.; Academic Press: London, 1981; pp 337−357. (29) Smith, N. R.; Gordon, R. E.; Clark, F. E. Aerobic Sporeforming Bacteria; Agricultural Monographs; U.S. Dept. of Agriculture: WA, 1952. (30) Butler, I. B.; Schoonen, M. A. A.; Rickard, D. T. Removal of dissolved oxygen from water: A comparison of four common techniques. Talanta 1994, 41, 211−215. (31) Knight, B. C. J. G.; Proom, H. A comparative survey of the nutrition and physiology of mesophilic species in the genus Bacillus. J. Gen. Microbiol. 1950, 4, 508−538.
AUTHOR INFORMATION
Corresponding Author
*Telephone: +44 (0)131 6508608; e-mail:
[email protected]. Notes
The authors declare no competing financial interest.
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ACKNOWLEDGMENTS The research leading to these results has received funding from the European Union’s Seventh Framework Programme (FP7/ 2007-2013) under Grant Agreement 226306. We acknowledge the comments of three anonymous reviewers who helped to improve the manuscript.
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REFERENCES
(1) Whiffin, V. S.; van Paassen, L. A.; Harkes, M. P. Microbial carbonate precipitation as a soil improvement technique. Geomicrobiol. J. 2007, 24, 417. (2) Mitchell, A. C.; Ferris, F. G. The coprecipitation of Sr into calcite precipitates induced by bacterial ureolysis in artificial groundwater: Temperature and kinetic dependence. Geochim. Cosmochim. Acta 2005, 69, 4199−4210. (3) Dupraz, S.; Parmentier, M.; Ménez, B.; Guyot, F. Experimental and numerical modeling of bacterially induced pH increase and calcite precipitation in saline aquifers. Chem. Geol. 2009, 265, 44−53. (4) Mitchell, A. C.; Dideriksen, K.; Spangler, L. H.; Cunningham, A. B.; Gerlach, R. Microbially enhanced carbon capture and storage by mineral-trapping and solubility-trapping. Environ. Sci. Technol. 2010, 44, 5270−5276. (5) Achal, V.; Mukherjee, A.; Basu, P. C.; Reddy, M. S. Lactose mother liquor as an alternative nutrient source for microbial concrete production by Sporosarcina pasteurii. J. Ind. Microbiol. Biotechnol. 2008, 36, 433−438. (6) Achal, V.; Mukherjee, A.; Basu, P.; Reddy, M. Strain improvement of Sporosarcina pasteurii for enhanced urease and calcite production. J. Ind. Microbiol. Biotechnol. 2009, 36, 981−988. (7) Tobler, D. J.; Cuthbert, M. O.; Greswell, R. B.; Riley, M. S.; Renshaw, J. C.; Handley-Sidhu, S.; Phoenix, V. R. Comparison of rates of ureolysis between Sporosarcina pasteurii and an indigenous groundwater community under conditions required to precipitate large volumes of calcite. Geochim. Cosmochim. Acta 2011, 75, 3290− 3301. (8) Benini, S.; Borsari, M.; Ciurli, S.; Dikiy, A.; Lamborghini, M. Modulation of Bacillus pasteurii cytochrome c 553 reduction potential by structural and solution parameters. J. Biol. Inorg. Chem. 1998, 3, 371−382. (9) DeJong, J. T.; Fritzges, M. B.; Nusslein, K. Microbially induced cementation to control sand response to undrained shear. J. Geotech. Geoenviron. 2006, 132, 1381−1392. (10) Whiffin, V. S. Microbial CaCO3 precipitation for the production of biocement. Ph.D. Dissertation, Murdoch University, Perth, Australia, 2004. (11) Cunningham, A. B.; Gerlach, R.; Spangler, L.; Mitchell, A. C. Microbially enhanced geologic containment of sequestered supercritical CO2. Energy Proc. 2009, 1, 3245−3252. (12) Dupraz, S.; Ménez, B.; Gouze, P.; Leprovost, R.; Bénézeth, P.; Pokrovsky, O. S.; Guyot, F. Experimental approach of CO 2 biomineralization in deep saline aquifers. Chem. Geol. 2009, 265, 54−62. (13) Mortensen, B. M.; Haber, M. J.; DeJong, J. T.; Caslake, L. F.; Nelson, D. C. Effects of environmental factors on microbial induced calcium carbonate precipitation. J. Appl. Microbiol. 2011, 111, 338− 349. (14) Vaughan, D.; Ord, B. G. Influence of natural and synthetic humic substances on the activity of urease. J. Soil Sci. 1991, 42, 17−23. (15) Sneath, P. H. A., Mair, N. S., Sharpe, M. E., Holt, J. G., Eds.; Bergey’s Manual of Systematic Bacteriology, Vol. 2; Williams and Wilkins: Baltimore, MD, 1986. 8355
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