Introduction
Biomediated calcium carbonate (CaCO3) production is the process by which
organisms induce the precipitation of
calcium carbonate. With reference to bacterial CaCO3 precipitation – also
known as “microbially induced carbonate
precipitation”, “microbially induced calcite precipitation” (MICP) and
“microbially induced calcium carbonate
precipitation” (MICCP) – the phenomenon is well documented (Stocks-Fischer et
al., 1999; DeJong et al., 2006; Whiffin et al., 2007; van Paassen et al., 2010). For example, cyanobacteria
precipitate CaCO3 in microbial processes
related to the shedding of the S layer, forming the stalagmites and
stalactites in limestone caves and adding to the
rocky sediments of coral reefs (Southam, 2000). Crystal aggregation of
CaCO3 in the kidney, urinary tract or
gallbladder have been shown to be induced by microorganisms such as
Proteus mirabilis, a urease-positive organism,
due to secondary infection (Worcester and Coe, 2008). Ureolytic soil organisms
of the genera Sporosarcina or
Bacillus can also induce CaCO3, for example, in their cycling
of nitrogen with a urease enzyme (Hammes et al.,
2003; Gower, 2008; Worcester and Coe, 2008). This last group of MICP producers
has piqued recent engineering
interests to apply them in a bioengineering and repair
context.
MICP biotechnology utilizing ureolytic soil organisms, most notably
Sporosarcina pasteurii, has been shown to
directly reinforce or restore engineered or natural structures – such as the
repair of historical monuments (Le Métayer-Levrela et al., 1999; Webster
and May, 2006), marble slabs (Li and Qu, 2011) and stone heritage sites
(Rodriquez-Navaro et al., 2012) – and to reduce weathering of soil embankments
(Chu et al., 2012). The enzyme urease (urea amidohydrolase, EC 3.5.1.5)
initiates the process, catalyzing the breakdown of urea to raise local pH and
produce CaCO3 in a solution of calcium ions often supplied as calcium
chloride (CaCl2), as summarized in Eqs. (1) and (2). The
produced CaCO3 fills structural gaps or bridges materials (e.g. soil
grains) to form a cemented product with unconfined strengths of up to
20 MPa (Whiffin et al., 2007).
CO(NH2)2+2H2O↔2NH4++CO32-(ureahydrolysis)2NH4++CO32-+CaCl2↔CaCO3+2NH4Cl(CaCO3formation)
Bacterial species such as Bacillus sphaericus (van Tittelboom et
al., 2010) and Bacillus megaterium (Krishnapriya et
al., 2015) have also been applied in material or volume strengthening. The
aforementioned ureolytic soil organisms
are attractive for MICP as they are “generally regarded as safe” (GRAS)
bacteria with accessible substrates (i.e.
urea) and an aerobic metabolism applicable to most engineering and
terrestrial environments (DeJong et al., 2006).
These gram-positive organisms offer other attractive features such as
spore-forming capability, allowing for long-term capsule storage in cements (Jonkers, 2011) and exopolysaccharide (EPS)
secretion for improved material
bonding (Bergdale, 2012).
The application of MICP in industry as a biotechnology is proposed to help
reduce the need for current structure
repair practices such as chemical grouting, which have been found to be
environmentally detrimental in their permanence (DeJong et al., 2010), in some cases posing serious human
health risks (Karol, 2003). That said, ureolytic MICP does produce excess ammonia, which can be harmful (van Paassen
et al., 2010). The use of nitrifying and denitrifying bacteria could help solve this issue by oxidizing
ammonia to nitrate and later nitrogen gas without affecting MICP. In fact,
the work of Gat et al. (2014) has shown co-cultures of ureolytic and
non-ureolytic bacteria can actually be beneficial to MICP. Alternatively,
denitrifying bacteria can be used to directly induce MICP to avoid ammonia
toxicity, though the level of CaCO3 is comparatively less than that of ureolytic
MICP, and harmful nitrites can build up in solution (van Paassen et al., 2010).
Other pathways to achieve MICP have also been explored with
B. subtilis, B. megaterium and B. sphaericus (Kang et al., 2015; Li et al., 2015).
Problems in large-scale application of the MICP technology have occurred too
and remain unsolved. Research by van Paassen et al. (2009) found poor sample
homogeneity of MICP, as well as decreasing biomass and urease-inducing
CaCO3 activity over time and increasing soil depth in a pilot 100 m3
sand study using Sporosarcina pasteurii, attributing these
heterogeneities mostly to the application process. Alternative metabolisms
and bacteria for large-scale applications in biomineralization of CaCO3
have also been investigated by the group (van Paassen et al., 2010). Indeed,
it has been commented that the type of bacteria utilized is one of the major
considerations and potential limitations in large-scale geotechnical
operations (Mitchell and Santamarina, 2005).
Therefore, the search for new bacteria by which to achieve viable levels of
MICP is important for optimizing the protocol best suited (in terms of
performance, economics and environmental impact) for marketing in green
industries (van Paassen et al., 2010; Cheng and Cord-Ruwisch, 2012; Patel, 2015).
Following a literature review of the nine documented species of
Sporosarcina (Claus and Fahmy, 1986), seven species were found to be
urease positive and distinct from Sporosarcina pasteurii as
alternative ureolytic MICP sources. While no candidate improves on some of
the shortcomings of ureolytic MICP (i.e. ammonia toxicity), each candidate
was found to be poorly investigated in the current MICP technology, despite
fitting the ureolytic model for MICP. One candidate, Sporosarcina ureae, was selected at random for investigation as it was deemed appropriate
to explore the feasibility of a single candidate species in thorough
comparison to other already-published species applied in ureolytic MICP.
Thus, the primary goal of this study was to investigate the suitability of
S. ureae as a MICP organism in material improvement by testing it
experimentally against the previously investigated species of
Sporosarcina pasteurii, Bacillus megaterium and Bacillus sphaericus.
In its assessment, a parallel investigation was also
performed to assess how the MICP technology, utilizing S. ureae as
the candidate MICP organism, can perform under various environmental
conditions including acid rain, flooding and freeze–thaw cycling concurrent
with colder North American climates.
Results
NH3–NH4+ production
Among the different bacterial strains considered, S. pasteurii
and S. ureae were capable of producing the first- and second-highest
levels of NH3–NH4+, respectively, per unit of time, in both UB-1
(32.50; 29.00 U mL-1) and UB-2 (32.76; 30.28 U mL-1) media (Fig. 2a, b). Isolates of B. subtilis (2.91 U mL-1), B. megaterium
(4.87 U mL-1) and L. sphaericus (5.89 U mL-1) displayed a lower peak of
NH3–NH4+ production in both media. When urea in medium moved
from the sole source (i.e. UB-2) to one of a number of sources (i.e. UB-1)
for nitrogen, NH3–NH4+ production dropped to near-zero values
(Fig. 2a, b) for B. subtilis (0.44 U mL-1), B. megaterium
(0.56 U mL-1) and L. sphaericus (1.20 U mL-1) that were statistically
significantly different (p < 0.05, n= 6) from the final UB-1
values for each species. However, isolates of S. ureae and S. pasteurii
observed no statistically significant difference (p > 0.05, n= 6) between final values recorded in UB-1 and UB-2
media. Instead, a rise in production (t= 0–5 h) followed by a
levelling-off in value (t= 6–12 h) was the general trend observed in
UB-1 and UB-2 media (Fig. 2a, b).
(a, b) NH3–NH4+ production (U mL-1: µmol of
NH3–NH4+ min-1 mL OD600 of culture); (c, d) pH; and
(e, f) growth of selected bacteria types in (a, c, e) UB-1
(no yeast extract, YE) and (b, d, f) UB-2 (10 g L-1 YE) nutrient conditions (SD, n= 6). YE was a nitrogen
source in the growth medium.
Examination of bacterial abundance in culture
All strains showed a decline in growth progression when medium was restricted
(i.e. UB-2) to urea as nitrogen and
glucose as carbon sources (Fig. 2e, f). Growth repression was
greatest in the cases of B. subtilis (-33.9 %), L. sphaericus (-26.8 %)
and B. megaterium (-23.6 %) compared to S. pasteurii (-17.8 %) and S. ureae
(-16.6 %).
Additionally, the final OD600 (t= 12 h) achieved for all strains in
UB-2 medium was decreased compared to UB-1 medium values (t= 12 h), and the
difference in value for each strain was found to be statistically
significantly different (p < 0.05, n= 6). Growth cessation (i.e.
stationary phase) occurred for S. ureae and S. pasteurii in
both conditions but later in UB-1 (t= 11 h) compared to UB-2 (t= 9–10 h) medium (Fig. 2e, f); they grew logistically in both medium conditions.
In general, growth of L. sphaericus, B. subtilis and
B. megaterium in UB-2 medium followed a logistic growth curve too.
However, in UB-1 medium their growth fit an exponential model, whereby an
exponential growth phase was observed from t= 4 h to t= 12 h following a lag
phase of growth between t= 0 h and t= 3 h.
Changes in pH
The alkalinity increased with the increase in time for the strains of
S. ureae and S. pasteurii studied, in both UB-1
(8.99; 9.2) and UB-2 (8.74; 8.8) media. The lowest final pH values were
observed in L. sphaericus (7.88; 8.16),
B. megaterium (7.85; 7.93) and B. subtilis (7.70; 7.81)
in UB-1 and UB-2 media at the end of 12 h (Fig. 2c, d).
While pH continued to rise for S. pasteurii and S. ureae in
either UB-1 or UB-2 medium, it was constant
for L. sphaericus, B. megaterium and B. subtilis
after time in UB-1 medium as early as 6 h (L. sphaericus and
B. megaterium) in UB-2 medium. While final pH values for L. sphaericus, B. megaterium and B. subtilis
reached higher final (t= 12 h) values in UB-2 medium compared to UB-1,
which were found to be statistically significantly different (p < 0.05, n= 6), the opposite was true for S. pasteurii and
S. ureae; values in UB-2 were lower compared to UB-1, and the
difference was found to be statistically significantly different for each
species (p < 0.05, n= 6). In general, acidity increased with the
increase in time for L. sphaericus, B. megaterium and
B. subtilis in UB-1 medium. This was also true in UB-2 medium except
for L. sphaericus, which showed an increase in pH over time.
Direct shear strengths (τ, kPa) of treated sands
(SE, n= 3).
Mechanical and biological behaviour in MICP reinforced sands
Experiments of sand consolidation with triplicate holding vessels (Fig. 1)
mixed with S. ureae (135.77 kPa) or S. pasteurii (135.5 kPa) and fed MICP medium (i.e. CM-1) had
improvements in their direct shear strength compared to control vessels
(15.77 kPa) fed with MICP medium only. In fact, the difference in direct
shear strength values for S. ureae and S. pasteurii
compared to control vessels was found to be statistically significantly
different (p < 0.05, n= 3). However, the difference in strength
between S. ureae and S. pasteurii was not statistically
significantly different (p > 0.05, n= 3). Mixtures of
non-ureolytic B. subtilis (28.1 kPa) showed no statistically
significant difference (p > 0.05, n= 3) in value when compared
to the control (Fig. 3). While pre-injection (21.9 × 107 CFU mL-1) and
post-incubation (3.2 × 107 CFU mL-1) cell abundance was highest in the
case of B. subtilis (Fig. 4), all bacterial isolates showed a
decrease in cell abundance when comparing pre-injection to post-incubation
cell abundance with statistically significant differences (p < 0.05,
n= 9). Also, the percentage loss of cell abundance, taken as the
difference between post-incubation and pre-injection cell abundances divided
by the initial pre-injection cell abundance (-77.7 % (S. ureae),
-75.4 % (S. pasteurii), -77.7 % (B. subtilis)), was
not statistically significantly different (p > 0.05, n= 9) when
comparing values between species. Of note, the medium-only control had no
cell growth (CFU mL-1) observed before and after incubation.
Microbial viability of treated sands before injection (black
bars) and after incubation (gray bars) (SD, n= 9).
Microstructure investigation
The precipitation of calcium as CaCO3 via MICP was visualized. Sand
granules from approximately the first 1 cm of
sands treated with MICP solution (i.e. CM-1) combined with S. ureae
are shown (Fig. 5a, b), where crystals arranged in rosette peaks (20–40 µm) can be seen across the surface of a sand grain (Fig. 5a, b).
Rod-shaped structures (40–80 µm) can also be visualized, though less
commonly, across grain surfaces (Fig. 5a, b). Calcium, carbon and oxygen
peaks captured by EDS analysis for crystals organized in “rosette” patterns
as well as in rod-shaped structures suggest CaCO3 precipitation (Fig. 5c, d).
SEM image of the (a) whole surface (bar, 100 µm) and
(b) magnified (bar, 10 µm) silica granule with crystalline
(yellow arrow) and amorphous (white arrow) calcium
structures following bacterial treatment. EDS analysis shows the chemical
composition of (c) crystalline and (d) amorphous precipitates.
Environmental durability of MICP
A reduction in the reinforcement of sands by CaCO3 mineralization with
S. ureae inoculations was observed following exposure to acid rain
as direct shear strengths reduced to 39.7 kPa (Fig. 6) or 29.2 % compared
to those with no such treatment (Fig. 3). Treated sands under conditions of flooding (111.7 kPa)
or freeze–thaw (93.5 kPa) rounds had better durability (i.e. strength
retention) compared to acidified states, with differences in strength being
statistically significantly different (p < 0.05, n= 3). In fact,
no severe mechanical damage was incurred by samples treated under conditions of simulated
flooding or freeze–thaw cycles (Fig. 6); when comparing the difference in
their direct shear strengths to sands tested under ideal (i.e.
non-environmental) conditions, these differences were found to be not
statistically significantly different (p > 0.05, n= 3) (Fig. 3).
Direct shear strengths (τ, kPa) of treated sands with
Sporosarcina ureae in flood (water), freeze–thaw
(ice) and acid rain (acid) simulations (SE, n= 3).
Discussion
In characterizing S. ureae as a ureolytic organism in MICP, the
goals of the study were to understand (1) its ability to degrade urea over
time relative to other commonly applied MICP bacterial isolates and (2) its
preference for urea as a
nitrogen source. The strain (BGSC 70A1) was consistent in its total nitrogen
(NH3–NH4+) production regardless of whether the nutrient medium
included (i.e. UB-1) or did not include (i.e. UB-2) yeast extract. This can
be attributed to mostly urea catabolism in UB-1 medium and entirely so in
UB-2 medium as urea was the sole source of nitrogen. It is important to note
that minor mineralization of the yeast extract components in UB-1 medium
would likely have contributed ammonium
(Gat et al., 2014) in this medium condition. This is supported by data
recorded for the negative control (medium-only) in UB-1 medium with production as high as 0.12 U mL-1 (Fig. 2a, b).
Also, degradation of amino acids from
bacterial metabolism, such as ornithine, particularly supplied in UB-1 medium
via yeast extract, could also
contribute to total nitrogen in solution for this condition (Cruz-Ramos et
al., 1997). For both media (UB-1 and
UB-2) dissolution of ammonium as ammonia into the atmosphere would have
reduced available nitrogen for
measurement over time. Thus, a quantitative urea hydrolysis rate cannot be
determined from the data collected, as
nitrogen production over extended periods of time is a complex collection of
some or all of these processes. This limits the conclusions able to be drawn
as only the broad bacterial activity in medium, as regards preferences for
urea as a nitrogen source, over time can be considered. For a quantitative
method determining urease rates a robust protocol is presented by Lauchnor et al. (2015).
Also, urea-hydrolysis-induced CaCO3 precipitation rates can
be determined by measuring the decrease in dissolved Ca2+ ions over time
(Harbottle et al., 2016). However, overall, the total nitrogen production
over time draws support for S. ureae as a promising MICP candidate
in biocement as over the time period measured it was able to produce a
consistent amount of nitrogen as ammonia–ammonium in UB-1 or UB-2 medium, and
ammonia production has been found to be directly proportional to CaCO3
production (Achal et al., 2009) and soil stabilization (Park et al., 2012).
As mentioned, the production of nitrogen by S. ureae in medium is
due mostly, or completely, to urea catabolism, and this process is likely
driven chiefly by its urease enzyme (Gruninger and Goldman, 1988; Mobley and
Hausinger, 1989). Alternatively, an unknown urea-degrading enzyme other than
urease could produce or contribute to the result. Notably, all
Bacillus strains observed a decrease in total ammonia production
when yeast extract was available (i.e. UB-1). This was not observed for
S. ureae, much like S. pasteurii. Urea is a nitrogen source
for bacterial growth, often catabolized by urease (Lin et al., 2012), which
has been found to be controlled by nitrogen levels and pH as well as other
factors which can differ between bacterial species (Mobley et al., 1995, 2001). Our observations indicate that S. ureae
selects for urea in a metabolic pattern potentially similar to S. pasteurii and quite differently from the Bacillus strains
investigated here, which appear to have medium-dependent metabolism of urea.
The observation that the investigated Bacillus strains have
medium-dependent metabolism of urea is particularly interesting for
B. subtilis as it has been applied as a non-ureolytic control
organism in previous literature (Stocks-Fischer et al., 1999; Gat et al.,
2014). In UB-2 medium, a non-zero total ammonia activity was measured for
this strain (Fig 2a, b). This is consistent with previously published
literature linking total ammonia production to urea breakdown from urease
when urea is the sole source of nitrogen and urease is the assumed main
catabolic enzyme – the enzyme expressed constitutively in species of
Sporosarcina (Mobley et al., 1995) but in a repressible manner
(i.e. activated in the absence of NH4+ and other forms of nitrogen
(i.e. NO3-), with urea being the sole nitrogen source) in strains such
as B. megaterium (Mobley and Hausinger, 1989) and B. subtilis (Atkinson and Fisher, 1991; Cruz-Ramos et al.,1997). This is indeed
suggested by our data as it was observed for B. subtilis (and also
for B. megaterium and L. sphaericus) that increased total
ammonia production reached higher values in UB-2 medium compared to near-zero
values in UB-1 medium with yeast extract as an alternative nitrogen source.
In fact, in UB-2 medium peaks were reached within 3–6 h from near-zero
values (t= 0–1 h) for all Bacillus species, further suggesting
an increase in processes related to urea hydrolysis, such as urease
expression, over time following a reduction in genetic repression (Fig 2a, b).
This also corrolates well with growth patterns. A comparatively slow
growth rate occurred (t= 8–12 h) after a comparatively fast (t= 3–7 h)
rate of growth following a lag period (t= 0–2 h) for these
strains, in general (Fig. 2a, b). An increase in urease, or other urea
hydrolysis processes, may account for an ability to grow quickly (t= 3–7 h) despite nitrogen limitation in UB-2, as ureolysis would provide nitrogen
for growth-related processes. However, growth could have been restricted,
over time, due to other nutrient limitations such as glucose depletion. This
would explain a continued but reduced growth rate (t= 8–12 h) (Fig. 2a, b).
Alternatively, or in addition, the decreased growth could be due to
decreased dissolved oxygen content in medium over time, which is required for
aerobic respiration, such that each Bacillus species switched to a
slower anaerobic growth pattern. An increase in harmful metabolites such as
organic acids in solution over time could also have hindered growth. There is evidence
that this occurred for these species in UB-1 medium as a decrease in
pH over time was observed which correlates with organic acid production (Fig. 2c).
Taken together, this has significance as, while B. megaterium
and L. sphaericus have been investigated as candidates in ureolytic
MICP, this has not been extensively the case for B. subtilis, which
in this study shows ureolytic capability under specific conditions. This may
guide future research on ureolytic MICP with B. subtilis,
particularly where cementation media do not contain nutrient-rich additives
such as yeast extract. This has been the case in some literature solutions
for inducing ureolytic MICP (van Paassen et al., 2010; Cheng et al., 2013).
In this study B. subtilis was included in sand solidification as a
non-ureolytic strain control as the cementation medium contained yeast
extract, intended for maximum biomass support and CaCO3 production rates
(van Paassen et al., 2010).
It is clear that S. ureae prefers an alkaline environment, like
S. pasteurii and quite different from the other isolates in trials,
as in both growth conditions samples grew not only exponentially but towards
an increased pH (Fig. 2c, d). Urea hydrolysis, driven potentially by urease,
in this species may maintain ureolytic activity for production of the highly
alkaline environment to which it is suited for growth as an alkaliphile and
for its role as a nitrogen cycler (Gruninger and Goldman, 1988). These
conditions are also important for CaCO3 production (Whiffin et al.,
2007). It can also use the charge gradient generated from ammonium production
for energy (Jahns, 1996) to support growth. A diagram of this adenosine triphosphate (ATP)-generating
system coupled to ureolysis is available in the work of Jahns (1996) and
Whiffin (2004). Additionally, the ammonium is an accessible nutrient (i.e.
nitrogen) source (Gruninger and Goldman, 1988). This may partly account for
S. ureae and S. pasteurii having the smallest change in
growth between UB-1 and UB-2 medium by having the material but also energetic
means to multiply. This is extremely promising as van Paassen et al. (2010)
determined the CaCO3 precipitation rate is positively correlated with
the number of viable microorganisms in solution. Thus, taken together, the
ureolytic, pH and growth data of this study support S. ureae as
superior in ureolytic action to every Bacillus strain considered
except S. pasteurii. Indeed, the work of Harbottle et al. (2016)
likewise found S. ureae and S. pasteurii to be about as
efficient in terms of ureolytic activity. Given the current data,
S. ureae and S. pasteurii are comparable as candidates for
ureolytic MICP. This should prompt interest for further investigations
differentiating between the two strains on such parameters as protease
activity, exopolysaccharide production and biofilm levels, which are also connected to
MICP capability (Achal et al., 2010), so as to identify the superior
candidate. Some differential work has already been done (Sarmast et al.,
2014).
To understand the macroscopic engineering aspects of S. ureae in
MICP application, efforts of this study were focused on measuring and
assessing its ability to strengthen model sands via urea hydrolysis to form
CaCO3. In experiments with a model silica sand featuring poor
geotechnical characteristics (i.e. uniform sand profile) for high
susceptibility to settling and static strength decreases (Conforth, 2005), it
was clearly shown that the S. ureae treatment led to consolidation
of the medium in 48 h with an improvement in strength to 135.77 kPa. This was
8 times that of the control treatment (15.76 kPa) (Fig. 3). In addition,
while average consolidation strengths had no statistically significant
difference (p < 0.05, n= 3) between S. ureae and
S. pasteurii, the peak sample strength recorded for a S. ureae mould (175.8 kPa) exceeded the maximum sample strength recorded for
S. pasteurii (165.7 kPa), the typical model ureolytic organism in
MICP soil strengthening. It was also well above peak average strength
recorded for B. subtilis (28.1 kPa) (Fig. 3). This is as expected;
B. subtilis is a non-ureolytic organism in the “good nitrogen”
(Atkinson and Fisher, 1991) nutrient conditions supplied by the yeast extract
of CM-1 medium. Other Bacillus species were not tested under the
assumption that they too would experience repressive urea hydrolysis
expression in CM-1 medium and would produce similar observations as a result.
This is supported by data provided by the groups of Al Qabany et al. (2012)
and van Paassen et al. (2010), which found that CaCO3 precipitation, and by
inference soil strength, improved with more suitable microorganisms in MICP.
Taken together, this study provides evidence that S. ureae is
capable of soil improvement by ureolytic MICP similarly to S. pasteurii.
The presence of crystals as organized “rosettes” and amorphous “rods” was
observed (Fig. 5a, b) along sand granules treated with S. ureae,
which is evidence that it is capable of inducing prevalent formation of secondary
minerals. The structures were analyzed by EDS, and the results provide support
for CaCO3 formation (Fig. 5c, d). Assuming that the solution was
saturated with respect to CaCO3 and that the nucleation and
crystallization of the calcite polymorph was thermodynamically favoured
over time, the organized deposits should represent calcite (De Yoreo and
Vekilov, 2003). However, fast nucleation and crystallization can result in
amorphous CaCO3 structures and could explain the rod deposits that
appear amorphous in morphology under SEM (Fig. 5a, b) (Addadi et al., 2003).
This observation is limited, though, as SEM cannot discriminate among
CaCO3 polymorphs which can have varying morphology based on the
crystallization conditions (Ni and Ratner, 2008). The exact polymorph of
CaCO3 for each structure could be distinguished with techniques such as
X-ray diffraction (XRD) and/or Fourier transform infrared spectroscopy
(FTIR) (Anthony et al., 2003; Ni and Ratner, 2008). Assuming the rod
structures are amorphous precipitates, this indicates that the treatment
conditions were potentially suboptimal for the maximum precipitation of
crystalline CaCO3 such as calcite over time. This could be due to high
local chemical concentrations (e.g. calcium) which have been found to hinder
CaCO3 crystal formation as calcite (Al Qabany et al., 2012).
Investigators may be prompted to test alternative calcium concentrations from
those used in this study for injections so as to increase the efficiency of
crystalline CaCO3 precipitation in MICP. Finally, medium and B. subtilis treated sands gave no discernible crystal CaCO3 formation
(data not shown). This provides evidence of superficial strengthening in
shear tests for these treatments based on natural biofilm excretion
(B. subtilis) or sporadic mineral crystallization. Thus, overall,
the microscopy evidence does support that S. ureae can precipitate
CaCO3 for strength improvements in soil, which was part of the goal in
studying S. ureae in MICP.
When analyzing the cell viability of injections before and after incubation in
treated sands, it was found that S. ureae maintained higher
post-incubation (2.56 × 107 CFU) cell abundance compared to S. pasteurii (1.21 × 107 CFU) and that these differences were
statistically significant (p < 0.05, n= 9) (Fig. 5). Also, both
species' cell abundance was lower and found to be statistically significantly
different (p < 0.05, n= 9) compared to the cell abundance for
B. subtilis (3.2 × 107 CFU). This difference could be due to
the solution (i.e. TBS) utilized for serial dilution of the growth medium.
The TBS did not include ammonium and was not buffered at a high pH, which are
two necessary conditions for the survival of alkaliphilic genera such as
Sporosarcina (Morsdörf and Kaltwasser, 1989). Thus, a deflated
value for S. pasteurii and S. ureae would result. Also,
moulds become mostly anaerobic over time below the subsurface and within the
microenvironments of sand grains as oxygen is depleted by bacterial
respiration (van Paassen et al., 2010). B. subtilis cells may have
survived anaerobically (Clements et al., 2002) as opposed to the obligate
aerobes S. ureae and textitS. pasteurii (Claus and Fahmy, 1986),
leading to higher post-incubation cell abundance for B. subtilis.
However, considering the percentage loss of cell abundance calculated as
described (Sect. 3.4) is comparable between all three species, this indicates that
neither species outperforms the other in cell survival while in the high-salt,
high-urea CM-1 medium with incubation in treated sands. That being
written, the total cell abundance in S. ureae is higher compared to
S. pasteurii. This is important as cells provide nucleation points
for CaCO3 formation. Indeed, the literature reports designate that
strength enhancement by ureolytic MICP is driven not only by urea hydrolysis activity
but also by the presence of bacteria acting as nucleation sites
(Stocks-Fischer et al., 1999; Gat et al., 2014). While sand surfaces can also
act as nucleation points, the negatively charged bacteria cell wall attracts
positively charged cations (e.g. calcium) preferentially for the controlled
nucleation of CaCO3 over time. In fact, it has been shown that cell
abundance in MICP treatments positively correlate with the precipitation of
CaCO3 in both the rate of production and crystal size (Mitchel and
Ferris, 2006). The group of Hommel et al. (2015) have even developed a model
showing that calcite precipitation is proportional to cell abundance (i.e.
biomass) and potentially improved soil strengths. This model assumes that the
features of the cells such as biofilm production around their cell walls
favour and facilitate the precipitation of CaCO3. It follows that any
intact cell wall part of the biofilm can facilitate the precipitation process
whether the cell itself is alive or dead. Thus, in general, more cells
equates to more CaCO3 precipitation. However, in this study, S. ureae gave rise to strengths in sands that were not statistically
significantly different (p > 0.05, n= 3) versus S. pasteurii treatments. This is unexpected since S. ureae had
comparable ureolytic activity to S. pasteurii but higher cell
abundance over time in precipitation medium. Therefore, more CaCO3
precipitation should have occurred and led to a greater strength increase in
sands in S. ureae treatments. This non-linear increase in strength
compared to cell abundance can be a result of a number of factors. For
example, the ability of cells to precipitate CaCO3 can be hindered when
an abundance of cells injected into porous material (i.e. sands) leads to
pore plugging from the organic matter (i.e. cells). This has been seen to
lead to a varied amount of CaCO3 precipitation throughout the volume of
a mould (van Paassen et al., 2009). Where cells are distributed more evenly,
they can facilitate the precipitation of CaCO3 as nucleation points
(Hommel et al., 2015). This may explain why S. ureae, having a
comparable NH3–NH4+ activity to S. pasteurii, did not
outperform it on average in undrained, direct shear strength tests despite
having a higher cell abundance on average. It may also explain the broader
range of strengths achieved in S. ureae (Fig. 3). For example, a
suboptimal spreading mechanism could have hindered strength achievement in
some moulds of S. ureae treatment where pore plugging by organic
matter (i.e. cells) occurred. With this in mind, optimization of treatment
protocols would help to determine whether or not S. ureae is the
superior candidate compared to S. pasteurii given that it has
consistently increased total cell abundance (Fig. 3) to support more
nucleation of CaCO3 over time, in tandem with a
NH3–NH4+ production comparable to that of S. pasteurii.
However, it is important to note that S. ureae cells are
significantly smaller than cells of S. pasteurii (Claus and Fahmy,
1986). Therefore, the total cellular surface area available for nucleation of
CaCO3 would be similar for the two species. This provides a possible
explanation for why no statistically significant differences in strength were
observed because if total cellular surface area was most important for
precipitating CaCO3 this means there would be no difference in strengths
expected for the same total cellular surface area, whether it was spread over
a relatively high number of smaller cells (i.e. S. ureae) or lower
number of larger cells (i.e. S. pasteurii).
It was the current authors' focus to also apply tests in conditions
reflective of a Canadian environment with a relatively novel bacterial
isolate (S. ureae). Sands treated with S. ureae and which
underwent short-term flooding (111.67 kPa) or freeze–thaw cycling (93.47 kPa)
showed no statistically significant (p > 0.05, n= 3) strength
difference compared to in-lab (135.77 kPa) conditions (Fig. 6). It has been
shown that MICP-treated sands maintain some porosity in materials (Cheng and
Cord-Ruwisch, 2012; Chu et al., 2012) and that good strength maintenance in
seasonal water saturation and freeze–thaw cycling is possible with porous materials
(Litvan, 1980; Cornforth, 2005). Further studies may wish to investigate the permeability of
hardened sands via S. ureae at various levels of CaCO3
precipitation to strike a balance between porosity, peak strength and
endurance over time in weather simulations.
Predictably, it was seen that the acid rain model, reflective of a northern
Ontario rain pH (Sect. 4.4), eroded the shear
strength of sands (Fig. 6) to 35.5 % of originally observed values (Fig. 3). This is a result of the reaction of acid
with CaCO3 producing units of H2O, CO2 and salt, known as
weathering. A study by Cheng et al. (2013) reported similar
results with a Bacillus sphaericus model.
This prompts the idea that a MICP strength model,
regardless of the bacteria treatment selected (S. ureae, S. pasteurii etc.) for strength enhancement, would require a
time-based repair of treated volumes. This realistically limits its
geotechnical and economical practicality in the
industry. However, it does prompt interest to test the ability of natural
buffers, such as limes and sodas, to increase
the life span of MICP-induced strength enhancement by reducing acid rain
degradation.