Article pubs.acs.org/est
Soil Bacteria Population Dynamics Following Stimulation for Ureolytic Microbial-Induced CaCO3 Precipitation Daniella Gat,*,† Zeev Ronen,‡ and Michael Tsesarsky†,§ †
The Department of Geological and Environmental Sciences, Ben-Gurion University of the Negev, Beer-Sheva 8410501, Israel The Department of Environmental Hydrology and Microbiology, The Zuckerberg Institute for Water Research, The Jacob Blaustein Institutes for Desert Research, Ben-Gurion University of the Negev, Sede-Boqer Campus 8499000, Israel § The Department of Structural Engineering, Ben-Gurion University of the Negev, Beer-Sheva 8410501, Israel ‡
S Supporting Information *
ABSTRACT: Microbial-induced CaCO3 precipitation (MICP) via ureahydrolysis (ureolysis) is an emerging soil improvement technique for various civil engineering and environmental applications. In-situ application of MICP in soils is performed either by augmenting the site with ureolytic bacteria or by stimulating indigenous ureolytic bacteria. Both of these approaches may lead to changes in the indigenous bacterial population composition and to the accumulation of large quantities of ammonium. In this batch study, effective ureolysis was stimulated in coastal sand from a semiarid environment, with low initial ureolytic bacteria abundance. Two different carbon sources were used: yeast-extract and molasses. No ureolysis was observed in their absence. Ureolysis was achieved using both carbon sources, with a higher rate in the yeast-extract enrichment resulting from increased bacterial growth. The changes to the indigenous bacterial population following biostimulation of ureolysis were significant: Bacilli class abundancy increased from 5% in the native sand up to 99% in the yeast-extract treatment. The sand was also enriched with ammonium-chloride, where ammonia−oxidation was observed after 27 days, but was not reflected in the bacterial population composition. These results suggest that biostimulation of ureolytic bacteria can be applied even in a semiarid and nutrient-poor environment using a simple carbon source, that is, molasses. The significant changes to bacterial population composition following ureolysis stimulation could result in a decrease in trophic activity and diversity in the treated site, thus they require further attention.
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INTRODUCTION The ubiquity of bacteria and the diverse roles they play in natural environments have led to the growing interest in harnessing bacterial activities for various anthropogenic purposes. Microbial-induced CaCO3 precipitation (MICP) is an emerging technique aimed at addressing different environmental and engineering concerns, including sequestration of atmospheric CO2, remediation of contaminated soils, soil stabilization and restoration of stone monuments, among others.1−6 Most soil bacteria are capable of inducing CaCO 3 precipitation through a variety of metabolic pathways, both autotrophic and heterotrophic.7 Hydrolysis of urea, catalyzed by the microbial enzyme urease (urea amidohydrolase, EC 3.5.1.5), is considered the most efficient microbial pathway for MICP.8,9 The hydrolysis of urea produces ammonium and carbonate (eq 1), thus increasing the saturation with respect to CaCO3 which could lead to its precipitation, usually as calcite (eq 2).
Two alternative approaches to promote in situ MICP in soils are biostimulation and bioaugmentation. Biostimulation encourages indigenous urea-hydrolyzing bacteria by providing appropriate enrichment and precipitation media; it relies on the natural ubiquity of ureolytic soil bacteria9−11 and on the bacterial spatial distribution. Bioaugmentation introduces large volumes of bacterial cultures into the treated soil along with growth and precipitation medium, and therefore requires very large volumes of pure-cultured ureolytic bacteria, for excample, Sporosarcina pasteurii. Producing and transporting large volumes of these cultures is an expensive and delicate procedure; their injection and homogeneous distribution throughout the treated site are difficult to achieve and might encounter regulatory hindrances.12 Moreover, the introduced bacteria are likely to decline in numbers due to low compatibility to the environment as well as competition and predation by indigenous bacteria.13 As the two methods involve the introduction of large quantities of urea along with a source of organic carbon, both biostimulation and bioaugmentation
urease
CO(NH 2)2 + 2H 2O ⎯⎯⎯⎯⎯→ 2NH+4 + CO32 −
(1)
CO32 − + Ca 2 + ↔ CaCO3↓
(2)
© 2015 American Chemical Society
Received: Revised: Accepted: Published: 616
August 19, 2015 December 15, 2015 December 21, 2015 December 21, 2015 DOI: 10.1021/acs.est.5b04033 Environ. Sci. Technol. 2016, 50, 616−624
Article
Environmental Science & Technology
v) on site, and were afterward kept at 4◦C for 2−24 h until the experiments began. The sand is classified (ASTM D422-63) as poorly sorted (SP) fine sand, with average D50 of 0.26 mm. Mineral composition is up to 92% (w/w) quartz grain and up to 6% (w/w) CaCO3 (mainly shells fragments); the fines content is below 1.6%. Organic carbon comprises 0.5−1% (w/w) of the sand26,27 and a pH of 7.5 ± 0.1 measured after suspension of the sand in a 0.01 M CaCl2 solution (1:1 w/v) and shaking for 30 min. Natural moisture content of the sand is up to 4% (w/ w) in the sampled depth, as measured gravimetrically on January 2014. Enrichment Media and Culture Conditions. The collected sand was suspended in media based on artificial groundwater (AGW) representing the composition of Israel’s coastal aquifer,26 containing: MgCl2 (1 mM), MgSO4 (1 mM), NaHCO3 (2.56 mM), NaCl (14.35 mM), CaCl2 (2.43 mM) and KCl (0.32 mM); total ionic strength: 31.5 mM. Ten g of sand were suspended in 100 mL of each treatment medium, in duplicates. All treatments were incubated at ambient temperature (approximately 22 °C) with shaking (100 rpm), in 250 mL Erlenmeyer flasks, corked with cellulose plugs. All enrichment media were filter-sterilized using 0.22 μm sterile filter-systems prior to the suspending of the sand. Five different AGW-based enrichment media were prepared: (1) molasses medium, MLS12 and MLS14 for enrichment experiments conducted in 2012 and 2014, respectively, supplemented with 20 g L−1 (333 mM) of urea, 0.01 mM of NiCl2 and 1 g L−1 of molasses ; (2) yeast extract medium, YE12 and YE14 for enrichment experiments conducted in 2012 and 2014, respectively, supplemented with 20 g L−1 (333 mM) of urea, 0.01 mM of NiCl2 and 3 g L−1 of yeast extract; (3) ammonia oxidizing enrichment medium, NH4, supplemented with 100 mM of NH4Cl, in order to encourage ammoniaoxidizing microorganisms; (4) urea medium, Urea12, supplemented with 20 g L−1 (333 mM) of urea and 0.01 mM of NiCl2; and (5) control media, Cont14, comprised only of AGW and 0.01 mM of NiCl2. NH4 and Urea12 treatments were prepared only in the 2012 enrichment experiment; Cont14 was prepared only in the 2014 enrichment experiment. Incubation was terminated at different times for each treatment, ranging from 7 to 81 days, according to the microbial activity as inferred from parameters such as pH, urea hydrolysis and ammonium− oxidation. Molasses represents a simple growth medium, devoid of amino-acids and other required cell building blocks. Yeastextract represents a rich growth medium as it provides the cell with peptides as well as organic carbon. The concentrations of both substances were decided upon in an attempt to compare this work with previous studies of Tobler et al. (2011),23 Burbank et al. (2011),21 Hammes et al. (2003),28 and Gomez et al. (2014).29 Chemical Analysis. After measuring pH, medium samples from enrichment experiments were filtered through 0.22 μm sterile syringe filters and refrigerated at 4°c until urea or ammonium measurement. Urea hydrolysis in the 2014 enrichment experiment was measured spectrophotometrically, according to the method described by Knorst et al. (1997),30 detection limit −1.0 mM urea. The color reagent was prepared for each measurement session and was discarded at the end of each day. A calibration curve of five different urea concentrations and a blank solution was produced anew for each freshly prepared color reagent. Calibration curve R2 values
are likely to affect and change the indigenous microbial population to an unknown extent. Following the hydrolysis of urea, ammonium accumulates in the soil, possibly leading to the stimulation of ammonia− oxidation and nitrification.14 Ammonia-oxidizing and nitrifying bacteria are generally chemo-autotrophs, though heterotrophs are also found, and some are also known to be able to hydrolyze urea.15 Ammonia−oxidation induces a decrease in pH and carbonate concentration (eq 3),16 thus, the oxidation of the accumulated ammonium could result in CaCO3 dissolution. Moreover, the end product of nitrification, nitrate, is a highly soluble contaminant that is likely to infiltrate into the groundwater of the treated site. A study by Reed et al. (2010)17 showed that following molasses and urea amendments to a basaltic aquifer, an increase in bacterial and archaeal amoA gene copies, markers for ammonia-oxidizing microorganisms, was observed, along with a slight increase in nitrate concentrations. 2NH+4 + 3O2 → 2NO−2 + 2H 2O + 4H+
(3)
Ureolytic MICP has been extensively investigated under laboratory conditions, using bacterial isolates from different environments.18−20 A limited number of studies on MICP with enrichment cultures have been performed.21−23 These studies focused mainly on urea hydrolysis and CaCO3 precipitation rates, while little attention has been paid to changes in indigenous bacterial populations as a result of soil MICP treatment, and to the long-term sustainability of precipitated CaCO3 due to microbial succession and ammonia−oxidation. Moreover, only few biostimulation studies were performed on loose sand, despite the high relevance of MICP application to this environment, for example, mitigation of liquefaction, mitigation of dust emissions or mitigation of desertification. In this research we studied the urea-hydrolysis potential of indigenous bacteria from coastal sand in a semiarid environment. The main goal of the research was to determine whether significant ureolysis could be stimulated in a nutrient-poor sand, and to characterize the changes in the soil bacterial population following different enrichment media using 16S rRNA gene sequencing. Our secondary goal was to evaluate the local ureolysis potential via biostimulation using rich and poor carbon sources; yeast-extract and molasses, respectively. The ammonia−oxidation potential of the indigenous soil bacteria following enrichment with NH4+ was also studied.
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EXPERIMENTAL SECTION Sand Source and Sampling. The sand used for the enrichment experiments was collected at the Ziqim site (31.611N 34.503E), a semiarid zone (BSh in Köppen climate classification), located in the southern part of the Israeli coastal plain. The collection site is located about 60 m from the shoreline, in the supratidal zone of the beach. The site was selected for its soil characteristics, a mineral soil low in organic matter, subject to relatively dry conditions. The initial general bacteria and ureolytic bacteria abundancy was considerably low, 8.4 × 104 copies of 16S gene and 101 ureC copies per gram of sand.24 Thus, it differs from sites tested by other research groups, for example, oxic and anoxic aquifers and saturated soils.21,23,25 The sand was sampled twice: in October 2012 and in January 2014. Samples were taken from a depth of 0.4 m using a manual auger, which was washed with ethanol (70% v/ 617
DOI: 10.1021/acs.est.5b04033 Environ. Sci. Technol. 2016, 50, 616−624
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Article
RESULTS Urea Hydrolysis by Indigenous Bacterial Populations. Urea and ammonium concentrations for 2012 and 2014 enrichment cultures are presented as urea hydrolysis percentage (Figure 1a). Both enrichment experiments yielded similar
were above 0.99 for all measurements. In the 2012 enrichment experiment urea hydrolysis and ammonia oxidation were inferred from measurements of ammonium concentration using Spectroquant kit no. 1.00683.0001 (Merck KGaA, Darmstadt, Germany). All spectrophotometric measurements were conducted using Spectroquant Pharo100 spectrophotometer (Merck KGaA, Darmstadt, Germany). Samples were diluted with double-distilled water as necessary. Microbial Analysis. Optical Density. Bacterial growth in the 2012 enrichment experiment was assessed by measuring optical absorbance at two wavelengths: 600 and 540 nm. The results of these two wavelengths were not significantly different, therefore only the 540 nm measurements are presented. The first measurement took place immediately following the suspension of the sand and was therefore influenced by some suspended fine particles. For the following measurements these particles settled and the shaking was insufficient in raising them, thus the turbidity was attributed solely to the presence of bacteria in the solution. DNA Extraction. DNA for 16S rRNA gene sequencing was extracted from each of the enrichment cultures of 2014 upon termination of the enrichment experiment, from the 2012 enrichment cultures that were kept in glycerol (25% v/v) at −80°c, and from the native sand collected in 2014 and kept at −20°c. DNA from enrichment cultures (both 2012 and 2014) was extracted using PowerSoil kit (MoBio, CA), with slight modifications to the manufacturer’s instructions, as follows: varying volumes of each enrichment culture were centrifuged (3600g, 15 min), the supernatant was discarded and the pellet was added to the PowerBead tubes. At the elution step, 50 μL (instead of 100 μL) of solution C6 were used in two elution steps, to further increase concentration and yield. Molecular analysis results of the 2014 experiment are presented for each replicate separately and are marked 14.1 and 14.2. DNA extraction from frozen (−80°) 2012 enrichment cultures took place approximately two years after the experiment was terminated and, due to low yields, duplicates were combined prior to the extraction. These DNA extracts were used for 16S rRNA gene sequencing, as described hereafter. DNA extraction from the native sand was extracted using PowerMax kit (MoBio, CA), with modifications recommended by the manufacturer for low biomass soils with low humics (see Supporting Information). 16S rRNA Gene Sequencing and Bacterial Population Analysis. DNA extracts of concentrations ranging from 2.4 ng/ μL to 50.0 ng/μL were sent to Research and Testing Laboratories (Lubbock, TX), where samples were sequenced using Illumina MiSeq platform, with the primer set Gray28FGray519R.31 The sequencing procedure and primers are described in the Supporting Information. The obtained sequences were submitted to the National Center of Biotechnology Information (NCBI) GenBank and were given the following accession numbers: SAMN03998131SAMN03998137 and SAMN03997516. The sequences were processed using mothur,32 according to the analysis pipeline described in the Web site: http://www.mothur.org/wiki/ MiSeq_SOP,33 accessed on May 20, 2015. Due to the size of the data set (about 200 K reads), subsampling of 25% of the trimmed sequences of each treatment was conducted prior to the calculation of a phylogenetic distance matrix for beta diversity analysis; alpha diversity analysis (diversity and species richness indices inverse-Simpson and Chao-1) and taxonomic analysis were conducted on the entire data set.
Figure 1. Percentage of urea hydrolyzed (a), pH evolution (b) and optical density changes (c) over time in enrichment media from 2012 (marked 12) and 2014 (marked 14) amended with yeast-extract (YE) or molasses (MLS) and urea, urea only (urea12) and control without urea or organic carbon (Cont14). All results are average of duplicates, error bars represent the standard deviation of the measurement.
trends for the respective treatments YE and MLS. Yeast extract (YE) amendments induced the fastest urea hydrolysis, where over 95% of the urea was hydrolyzed within 5 days. Molasses (MLS) supplements induced slower urea hydrolysis, where 60% and 85% of the urea was hydrolyzed within 13 and 14 days for MLS12 and MLS14, respectively. No urea hydrolysis was observed in the absence of an organic carbon source, treatment Urea12, during 81 days (shown up to 336 h, that is, 14 days). Control treatment, Cont14, contained no urea and therefore is not presented in Figure 1a. An increase in pH in both YE12 and YE14 treatments corresponded to the rapid urea hydrolysis observed, with final maximal pH values of 9.3 and 10.4 respectively (Figure 1b). The final pH in treatments MLS12 and MLS14 resembled that in YE12 and YE14 respectively, despite the lower amount of urea hydrolyzed in the MLS treatments. In treatment Urea12, pH did not change throughout the experiment (81 days). In control treatment Cont14, pH fluctuated between values of 8.1 and 9.1, without exhibiting a distinct trend. Optical density (OD) at a wavelength of 540 nm, indicative of bacterial growth, is presented in Figure 1c. A significant difference between MLS12 and YE12 treatments was observed. The increase in OD in treatment YE12 correlated in time with the rapid urea hydrolysis observed in this treatment, whereas in MLS12 and Urea12 treatments, the increase in OD was similar and much lower; final OD values in treatment YE12 were higher than in treatment MLS12 by an order of magnitude, 1.6 and 0.2 respectively. It is important to note that the measured OD is influenced by the presence of suspended fine particles in 618
DOI: 10.1021/acs.est.5b04033 Environ. Sci. Technol. 2016, 50, 616−624
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Environmental Science & Technology the medium, thus, the initial OD in both treatments was relatively high, 0.4−0.5, as samples were taken immediately after the suspension of the sand in the medium, before the finer particles were allowed to settle. Nonetheless, the significant increase in OD in treatment YE12, compared with MLS12, could only be attributed to an increase in bacterial density. Ammonia−Oxidation Biostimulation Through NH4 Amendment. Changes in ammonium concentrations and pH values over time in treatment NH4+ are presented in Figure 2. A decrease in ammonium concentrations commenced after
Table 1. Population Diversity and Richness Following Enrichment in Different Media, Compared to the Native Sand (ZS)a sample
inverseSimpson
Lci, Hci
Chao-1
Lci, Hci 2853, 3435 3810, 4604 1639, 1945 33 253, 42 538 21 323, 27 021 21 951, 26 712 30 860, 39 002 28 391, 34 011 9839, 11 530 5256, 6039 9133, 10 714
Cont14.1 Cont14.2 Urea12 MLS14.1
3.37 (0.06) 5.95 (0.09) 10.1 (0.1) 137 (3)
3.25, 3.51 5.69, 6.24 9.6, 10.6 127.1, 148.5
3118 (156) 4174 (172) 1774 (63) 37571 (1717)
MLS14.2
65 (1.3)
60.3, 69.7
23971 (1057)
MLS12
74 (2.2)
68.5, 81.1
24188 (1055)
YE14.1
57 (1.1)
53.5, 61.6
34658 (1497)
YE14.2
108 (4.6)
96.6, 123.4
31047 (1256)
YE12 NH4 ZS
85 (1.9) 109 (3.2) 458.53 (NA)
79.0, 92.4 101.5, 117.4 423.21, 500.29
10634 (330) 5621 (187) 9875 (NA)
a
In parentheses is the standard deviation due to subsampling; Lci, Hci − Low and high confidence intervals, respectively.
Figure 2. Ammonium concentration (left vertical axis, circles) and pH (right vertical axis, triangles) over time, following enrichment with NH4Cl.
27 days, and by day 65 a 35% decrease was observed. pH decreased only slightly from 7.38 to 7.20, excluding volatilization as a removal mechanism for ammonia. Upon the final sampling of this treatment, sulfanilamide and N-(1naphtyl)-ethylenediamine dihydrochloride were added to the samples; a dark reddish-purple color appeared, indicating the presence of nitrite in the samples. Microbial Analysis. Bacterial Population Alpha Diversity. The effect of different enrichment treatments on the composition of microbial population was examined using 16S rRNA gene sequencing, based on OTU assignment according to 97% similarity between sequences. Table 1 presents the calculated inverse Simpson indices (2D)34 and Chao-1 indices for species richness35 for each treatment and for the native sand. The calculation was made on subsamples of 8464 sequences, the lowest number of sequences obtained by one sample. As can be seen in Table 1, the microbial diversity (2D), decreased significantly in all of the treatments compared to the native sand (ZS). The lowest diversity was observed in the control treatments Cont14 and Urea12. Collector’s curves of observed OTUs for each sample exhibited an ongoing increase with sampling, indicating that the sampling depth was still insufficient in producing a representative estimation of the diversity in each treatment (Supporting Information Figure S1). Species richness according to Chao-1 values indicated that control treatments Cont14 and Urea12 exhibited the lowest species-richness of all treatments. Bacterial Population Composition. In the native sand, the most abundant bacterial class was γ-proteobacteria, comprising about 18% of total population. α-proteobacteria and actinobacteria comprised another 14% of the total population, each. The Bacilli class comprised only 5% of the total population, Gemmatimonadetes were 3% and Nitrospirae about 1% of the population (Figure 3).
Figure 3. Bacterial population composition based on 16S rRNA gene sequencing, presented as fraction of each bacterial class from the total population.
Following enrichment with yeast-extract, the bacterial population was overtaken by Bacilli, increasing from 5% in the native sand, to 85% and 99% of the population in the two duplicates (Figure 3). The remaining population was comprised mainly of γ-proteobacteria. Similar results were observed in the MLS treatments, where Bacilli class bacteria comprised 67− 96% of the bacterial population and γ-proteobacteria comprised 1−19%. The microbial population composition following ammonium-chloride enrichment, NH4, displayed the mildest decrease in taxonomic diversity, compared with the native sand. The relative abundance of Bacilli in this treatment increased significantly, from 5% in the native sand to about 24%. Other abundant phyla found in this treatment were Gemmatimonadetes (19%), Actinobacteria (15%) α- and β-proteobacteria (13% and 10%, respectively). Interestingly, the nitrifying bacterial class Nitrospirae, identified in the native sand, was 619
DOI: 10.1021/acs.est.5b04033 Environ. Sci. Technol. 2016, 50, 616−624
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of the different treatments (see Supporting Information for input data). The lowest I values were of control treatments Cont14 and Urea12 (31−35 mM), intermediate values were found for NH4 treatment (136 mM), and highest values were found for molasses and for yeast-extract enrichment treatments (308−523 mM and 394−544 mM, respectively). Ionic strength of the native sand was not measured due to the relatively low water content, that is, 4% w/w. Table 2 presents weighted Unifrac distances subtracted from 1, thus higher values represent stronger similarity between samples. All pairwise P-values were under 0.001. According to the pairwise distances, the molasses and yeast-extract treatments, in which ureolytic bacteria were most successfully stimulated, exhibited the least similarity to the native sand, indicating that ureolytic biostimulation resulted in a significant alteration to the microbial population.
not detected in the NH4 enrichment, despite chemical evidence of the accumulation of nitrite in this treatment. Bacterial Population Beta Diversity. To compare the microbial composition differences between the different enrichments, Principal Coordinates Analysis (PCoA), based on weighted Unifrac distances,36 was performed (Figure 4).
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DISCUSSION The main goal of this research was to study the effect of urea and nutrient enrichment on an indigenous bacterial population in nutrient-poor sand, as a part of a biostimulated MICP. The Zikim site (southern Israel) was chosen both for its composition and location: a low organic carbon sand in an arid environment. As such this site represents sites that are potential sites for MICP application, that is, loose sand susceptible to liquefaction and migrating sand. The enrichment media were designed to achieve the following main purposes: to establish whether ureolytic bacteria could be stimulated in this soil; to determine whether a simple carbon source, that is, molasses, would suffice to significantly enhance ureolytic activity and to define the changes in bacterial population composition following stimulation of ureolysis. The nitrification potential of this soil was also evaluated. The environment chosen for this study differs from previously studied sites in its location, mineralogy, geochemistry, and moisture content. Previous studies were conducted on oxic25 and anoxic23 groundwater and soils from the saturated zone.21 In this work the unsaturated zone was examined (4% w/w water content), which provides a relatively harsh environment for microorganisms due to high salinity and low availability of resources.38 Nonetheless, effective ureolysis was stimulated, further supporting the feasibility of biostimulation even under harsh initial conditions. Bacterial Growth and Activity. Two different sources of organic carbon were used in this study: a rich organic−carbon source, yeast-extract, and a simple source, molasses. Yeast
Figure 4. PCoA analysis based on weighted Unifrac distances for treatments amended with molasses (MLS), yeast-extract (YE), ammonium-chloride (NH4), urea alone (Urea12), no urea and organic-carbon (Cont14), compared with the native sand (ZS).
Distinct clustering was observed according to the enrichment medium composition, that is, molasses and yeast extract, despite the taxonomic similarity observed between the two types of enrichment (Figure 3). Interestingly, Urea12 enrichment treatment clustered with the AGW-only Cont14 treatments, despite the difference in media composition. The native sand (ZS), control and ammonium enrichment treatments appear to cluster closely together, however, according to weighted Unifrac analysis these groups were significantly distinct one from another (P < 0.001). Axis 1 points at a correlation with final ionic strength (I) values, calculated by PHREEQC37 according to final chemical composition and pH
Table 2. Phylogenetic Similaritya between the Different Enrichment Treatments and the Native Sand (ZS) Cont14.2 Urea12 MLS14.1 MLS14.2 MLS12 YE14.1 YE14.2 YE12 NH4 ZS
Cont14.1
Cont14.2
Urea12
MLS14.1
MLS14.2
MLS12
YE14.1
YE14.2
YE12
NH4
0.75 0.63 0.21 0.26 0.12 0.22 0.23 0.21 0.36 0.45
0.66 0.22 0.26 0.12 0.22 0.23 0.20 0.38 0.49
0.21 0.26 0.12 0.22 0.23 0.21 0.39 0.50
0.56 0.41 0.39 0.35 0.29 0.22 0.22
0.32 0.40 0.35 0.28 0.27 0.26
0.19 0.20 0.19 0.14 0.13
0.58 0.58 0.22 0.23
0.60 0.23 0.23
0.21 0.21
0.45
a
Phylogenetic similarity calculated as weighted Unifrac distances subtracted from 1. All pairwise distances presented were found significant (p-value less than 0.001). 620
DOI: 10.1021/acs.est.5b04033 Environ. Sci. Technol. 2016, 50, 616−624
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Environmental Science & Technology extract contains growth factors such as amino-acids and peptides that accelerate cell growth,39 whereas molasses is comprised mostly of carbohydrates, with trace amount of protein and amino-acids. Molasses is a byproduct of sugar refining and it is thus a low-cost and readily available alternative to carbon sources such as yeast-extract, peptone, etc. Moreover, as molasses does not provide biologically available forms of nitrogen, it is less likely to affect urease expression in ureolytic bacteria exhibiting nitrogen-concentration dependent regulation.21 The different enrichment cultures exhibited significant differences in bacterial growth and activity. Yeast-extract amended cultures, YE12 and YE14, exhibited the highest rate of urea hydrolysis, at least 6.4 mM of urea hydrolyzed per hour. In the molasses enrichment cultures (MLS) urea hydrolysis reached a maximal rate only after 4 days of incubation (Figure 1a), from then on urea was hydrolyzed at a rate of 1 and 2 mM an hour in treatments MLS14 and MLS12, respectively. To better capture the differences between YE12 and MLS12 treatments we normalized urea hydrolysis rate to OD as an indication for biomass. The linearity between biomass and OD is not kept over absorbance values of 0.4−0.6, thus normalizing ureolysis rates of YE12 treatment to the measured OD of this treatment should be referred to as a qualitative indication for the efficiency of ureolysis in this treatment. Nonetheless, it provides a good basis for comparison between these two treatments. The normalized ureolysis rate was 4.2 mM urea per hour per unit OD (calculated by interpolation of OD and ammonium data for the 48th hour of the experiment) for the YE12 treatment, and for MLS12 the rate was 3.9 mM per hour per unit OD (on the 290th hour of the experiment, at the peak rate of hydrolysis and peak OD). The similarity in ODnormalized ureolysis rates between treatments MLS12 and YE12 suggests that the faster ureolysis in the complex carbon source treatment, that is, yeast-extract, resulted from a higher bacterial growth-rate due to the availability of growth factors and/or electron donors in the yeast-extract. This supposition also explains the four day lag in ureolysis observed in the molasses enrichment treatment, proposing that this lag was due to de novo synthesis of proteins and amino-acids that slowed down cell growth in this treatment. Nevertheless, an important point is that despite the longer period of time required to achieve complete ureolysis, using molasses as a cost-effective alternative to yeast-extract for in situ applications of MICP might prevent eutrophication and a complete consumption of oxygen. In the absence of an organic carbon source, represented by treatments Urea12 and Cont14, the bacterial population diversity and richness decreased significantly (Table 2). It can be assumed that the organic matter present in the sand, 0.5−1% w/w, became more accessible due to suspension in AGW, thus enabling this change in population composition. However, no urea hydrolysis was observed in treatment Urea12. The process of urea hydrolysis in model bacteria such as Sporosarcina pasteurii does not require additional substances other than urea and water, provided that a sufficient cell density is available. Thus, it was considered likely to induce little growth of nonureolytic microorganisms.8,10 However, it appears that the initial concentration of ureolytic bacteria was too low, demanding enrichment with organic carbon to enlarge the ureolytic population, as was also suggested by Tobler et al. (2011)23 and by Fujita et al. (2008).25
In natural soils, extracellular urease can be found adsorbed onto clays, minerals and organic matter; urease is thus stabilized and can maintain activity for years.11,40,41 Therefore, urea hydrolysis in soils is attributed partly to extracellular urease and requires no organic carbon.9,10 However, we did not detect any active urease in Ziqim sand since urea was hardly hydrolyzed in treatment Urea12. Similar observations were reported for alkaline soils from arid zones in SE Spain42 and in coastal soil in India, where enzyme activities were reported to decrease with decreasing humidity and increasing salinity.43 Bacterial Population Composition and Diversity. The different enrichment treatments resulted in different microbial populations compared with the native sand. As DNA from the native sand was extracted in a different method than the DNA from enrichment cultures, some bias due to efficiency of extraction methods should be considered.44 The method used to extract DNA from all enrichment treatments (PowerSoil, MoBio) is intended for soil DNA extractions. The samples from the enrichment cultures were relatively rich in suspended particles and sand grains. Therefore, this method was preferred over more subtle methods designed to extract DNA from liquid cultures (e.g., UltraClean, MoBio or GenElute, Sigma-Aldrich, MO). When applying the same method to the native sand, DNA yield was too low, required applying to a different extraction method (PowerMax, MoBio). Differences in bacterial population composition between treatments, however, were significant (P < 0.001 according to weighted Unifrac results) and could only result from the treatment composition and not the extraction method. The similarity between identical treatments from two occasions, 2012 and 2014, was also significant, and suggested that little change occurred during the two-year storage of samples from 2012. As the collection of the sand samples was conducted in the end of the dry season (October 2012) and during the wet season (January 2014), some variation in the indigenous bacterial population should be expected. Nonetheless, identical treatments performed in 2012 and 2014, that is, YE12−YE14 and MLS12−MLS14, exhibited similar ureolysis rates suggesting that seasonal changes had little or no impact on the potential of biostimulation in this environment. The prolonged preservation of samples obtained in 2012 prior to the extraction of DNA may have masked any seasonal variations in the bacterial population. Microbial communities in coastal sand have received little attention compared with agricultural soil, sediments and aquatic environments.45 The most abundant bacterial class in beach sand according to Kostka et al. (2011)46 was γ-proteobacteria (28.4%), similar to the results presented here (17.6%). Similarly, a work reported by Whitman et al. (2014)45 found that the most abundant sand bacteria in three marine and freshwater sites were Proteobacteria, Bacteroidetes, Firmicutes, and Actinobacteria. These four phyla were also found in Ziqim sand, along with Nitrospirae and Gemmatimonadetes that were not reported by Whitman and co-workers. The apparent similarity in bacterial population composition could suggest that the changes to bacterial population following ureolysis biostimulation, as presented here, may be repeated in other coastal environments, which are also susceptible to liquefaction. The greatest difference in bacterial population composition from the native sand was exhibited by the MLS and YE treatments (Table 2), indicating a significant change in bacterial population following the stimulation of ureolysis. These treatments also exhibited an increase in relative abundance of 621
DOI: 10.1021/acs.est.5b04033 Environ. Sci. Technol. 2016, 50, 616−624
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bacteria but not in AO archaea,58 thus the presence of AO archaea following MICP biostimulation should be studied in future research. The results of this study indicate that urea hydrolysis can be stimulated in coastal sand from a semiarid environment by providing urea and a simple carbon source, for example, molasses. The importance of examining this environment lies in its susceptibility to liquefaction, and its relevance to MICP application, along with the relatively harsh conditions it represents that might interfere with biostimulation. Successful biostimulation of ureolytic bacteria would render bioaugmentation unnecessary and might simplify in situ MICP. Following biostimulation, a significant decrease in bacterial diversity is to be expected, as of yet, little attention was paid to the changes in general bacterial population following MICP in soils. This study suggests that these changes might be fundamental and could harm the delicate ecological system. A major concern for the sustainability of MICP is the microbial ammonia−oxidation potential, as this process might reduce ambient pH and carbonate concentration. In this study, the ammonia−oxidation potential was demonstrated and should thus be considered in in situ application of MICP, either via biostimulation or bioaugmentation.
Bacilli, suggesting that urea hydrolyzing bacteria in these treatments belong to this bacterial class. Similar results were observed in the work of Burbank et al. (2012),18 where 7 out of 10 isolates of indigenous ureolytic bacteria were classified as Bacilli. Bacilli are members of the Firmicutes phyla, which are also common in alkaline environments,47 suggesting that the significant increase in Bacilli following ureolysis is also a result of the increase in pH. While stable alkaline environments exhibit a diversity in trophic groups,48 a rapid increase in pH might not enable such diversity to develop, resulting in the loss of various trophic activities, as was also observed following nitrogen fertilization.49,50 These findings suggest that the increase in pH during MICP should be mitigated to prevent significant changes to the indigenous bacterial population. This could be achieved by decreasing the ratio of added urea to added calcium, as CaCO3 precipitation results in a decrease in pH. Despite the apparent taxonomic resemblance between yeastextract enrichment cultures and molasses enrichment cultures, population analysis based on weighted Unifrac distances, indicated that these are distinct populations (Table 2, Figure 4). As pH values in these treatments were similar, we suggest that the difference in population composition resulted from the different types of media.50 The duration of the observed change in bacterial population and activity is yet undetermined; while Ramirez et al. (2012)50 and Fierer et al. (2012)49 suggest that the change in bacterial population and activity following inorganic nitrogen fertilization could persist over a long-term, Stark et al. (2007)51 assert that the population change is transient and will not persist after urea amendment is concluded. The various conjectures presented here, on the outcome of in situ enrichment, should therefore be carefully assessed and observed over large time-scales before MICP is applied to soils. Nitrification Potential. In this experiment, the nitrification potential of the sampled site was studied by adding 100 mM of ammonium-chloride to the AGW medium. Since MICP in soils involves introduction of organic carbon to the treated site, its lingering presence along with the accumulation of ammonium might affect the ammonia-oxidizing (AO) population composition, but probably not to prevent nitrification entirely.52 Thus, the conditions in the AO enrichment treatment in this experiment probably differ from those that would follow ureolysis biostimulation, mainly in the concentration of organic carbon. The ammonium-enriched culture exhibited a decrease in ammonium concentration along with a slight decrease in pH. In addition, nitrite accumulation in this enrichment culture was detected. The decrease in ammonium concentrations commenced about 27 days into the experiment, which is within the reported time scale for growth of ammonia-oxidizing and nitrifying bacteria enrichment cultures (e.g., 3−7 weeks).53−55 It should be noted that the enrichment medium was not amended with microelements such as Cu2+ or Fe2+ that are generally included in AO growth media (e.g., ATCC medium 2265 or DSMZ medium 756c). The microbial population in the AO enrichment treatment consisted mainly of Bacilli (24%) (Figure 3). Surprisingly, no evidence of the common ammoniaoxidizing bacterial genera Nitrosomonas and Nitrosospira (βproteobacteria), Nitrosococcus (γ-proteobacteria),56 was found in this treatment. Ammonia-oxidizing archaea are significant to ammonia−oxidation in soils;57 it was suggested that in a coastal environment the decrease in salinity results in a decrease in AO
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ASSOCIATED CONTENT
S Supporting Information *
The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.est.5b04033. DNA Extraction procedure for the collected sand and MiSeq sequencing method and primers (PDF)
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AUTHOR INFORMATION
Corresponding Author
*Phone: +972-8-6596989; fax: +972-8-6596831; e-mail:
[email protected]. Notes
The authors declare no competing financial interest.
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ACKNOWLEDGMENTS We thank Dr. Tali Bruner of the department of Environmental Hydrology and Microbiology, The Zuckerberg Institute for Water Research, The Jacob Blaustein Institutes for Desert Research, for her assistance and her helpful observations. We thank Dr. Itzhak Katra of the department of Geography and Environmental Development, Ben-Gurion University of the Negev, Amir Wahnon and Aviran Feldheim of the Department of Geological and Environmental Sciences, Ben-Gurion University of the Negev, for their assistance in obtaining sand samples, to Noa Cohen and Osnat Barnea for assistance in analysis.
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REFERENCES
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