Email updates

Keep up to date with the latest news and content from Microbial Cell Factories and BioMed Central.

Open Access Research

Role of bicarbonate as a pH buffer and electron sink in microbial dechlorination of chloroethenes

Anca G Delgado12, Prathap Parameswaran1, Devyn Fajardo-Williams1, Rolf U Halden13 and Rosa Krajmalnik-Brown13*

Author Affiliations

1 Swette Center for Environmental Biotechnology, Biodesign Institute, Arizona State University, PO Box 875001, Tempe, AZ 85287-5701, USA

2 School of Life Sciences, Arizona State University, Tempe, USA

3 Ira A Fulton Schools of Engineering, Arizona State University, Tempe, USA

For all author emails, please log on.

Microbial Cell Factories 2012, 11:128  doi:10.1186/1475-2859-11-128

The electronic version of this article is the complete one and can be found online at: http://www.microbialcellfactories.com/content/11/1/128


Received:19 July 2012
Accepted:4 September 2012
Published:13 September 2012

© 2012 Delgado et al.; licensee BioMed Central Ltd.

This is an Open Access article distributed under the terms of the Creative Commons Attribution License ( http://creativecommons.org/licenses/by/2.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

Abstract

Background

Buffering to achieve pH control is crucial for successful trichloroethene (TCE) anaerobic bioremediation. Bicarbonate (HCO3) is the natural buffer in groundwater and the buffer of choice in the laboratory and at contaminated sites undergoing biological treatment with organohalide respiring microorganisms. However, HCO3 also serves as the electron acceptor for hydrogenotrophic methanogens and hydrogenotrophic homoacetogens, two microbial groups competing with organohalide respirers for hydrogen (H2). We studied the effect of HCO3 as a buffering agent and the effect of HCO3-consuming reactions in a range of concentrations (2.5-30 mM) with an initial pH of 7.5 in H2-fed TCE reductively dechlorinating communities containing Dehalococcoides, hydrogenotrophic methanogens, and hydrogenotrophic homoacetogens.

Results

Rate differences in TCE dechlorination were observed as a result of added varying HCO3 concentrations due to H2-fed electrons channeled towards methanogenesis and homoacetogenesis and pH increases (up to 8.7) from biological HCO3 consumption. Significantly faster dechlorination rates were noted at all HCO3 concentrations tested when the pH buffering was improved by providing 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) as an additional buffer. Electron balances and quantitative PCR revealed that methanogenesis was the main electron sink when the initial HCO3 concentrations were 2.5 and 5 mM, while homoacetogenesis was the dominant process and sink when 10 and 30 mM HCO3 were provided initially.

Conclusions

Our study reveals that HCO3 is an important variable for bioremediation of chloroethenes as it has a prominent role as an electron acceptor for methanogenesis and homoacetogenesis. It also illustrates the changes in rates and extent of reductive dechlorination resulting from the combined effect of electron donor competition stimulated by HCO3 and the changes in pH exerted by methanogens and homoacetogens.

Keywords:
Acetogen; Alkalinity; Bicarbonate competition; Dehalococcoides; pH range; Trichloroethylene

Background

Organohalide respiring microorganisms represent a unique, efficient, and sustainable approach to detoxifying chloroethenes contamination from soil, water, and groundwater [1-3]. These microbes are important because they can use priority pollutants such as trichloroethene (TCE), dichloroethene (DCE), and vinyl chloride (VC) as electron acceptors for energy metabolism [4]. Dehalococcoides bacteria hold a prominent role among the organohalide respirers isolated to date, as these are the only ones having the proven ability to detoxify chloroethenes to the innocuous end product, ethene [1,5]. Dehalococcoides have a constrained metabolism; they strictly utilize hydrogen (H2) as the electron donor and acetate as the carbon source [6]. The most common method for delivery of H2 and acetate at bioremediation sites is by addition of fermentable substrates as precursors [2,7,8]. H2 gas has also been supplied for groundwater field applications [9] and in engineered ex situ treatment technologies for chloroethenes remediation [10-12]. In systems fed with H2, the pH tends to rise as a result of competing biological reactions, whereas dechlorination and/or fermentation of H2-releasing compounds decrease the pH. As a consequence, buffering and pH management are important parameters for assessing in situ and ex situ remediation approaches, and are crucial for sustained dechlorination [12-14].

In groundwater, dissolution of carbonate (CO32−)-containing minerals serves as the natural pH buffer. Among the CO32− species, bicarbonatea (HCO3) is the most abundant at neutral pH, and it contributes substantially to the alkalinity of groundwater. Typical HCO3 concentrations in groundwater are in the range of 0.7-10 mM [15,16]. Additionally, HCO3 is supplemented to groundwater as a common strategy when biostimulation or bioaugmentation are employed in order to buffer the protons produced by the biological reactions [2,8].

In laboratory settings, pH management is also commonly achieved through the addition of HCO3 buffer in the form of NaHCO3 or KHCO3. HCO3 has been used for growth of Dehalococcoides strains [17] and for mixed dechlorinating communities [18-20] to maintain a favorable pH. Dehalococcoides optimum pH has been reported to range from 6.9-7.5 [6]; yet, to date, there is a lack of systematic studies defining both the pH boundaries for these important organisms, and the effect pH exerts on each step in the TCE reduction pathway. Beyond its function as a buffer, HCO3 also serves as an electron acceptor for other microorganisms commonly encountered with organohalide respirers in the environment and in laboratory-cultured consortia. For example, at neutral pH, hydrogenotrophic methanogens consume HCO3 and H2 to generate methane [21]:

Hydrogenotrophic methanogenesis:

<a onClick="popup('http://www.microbialcellfactories.com/content/11/1/128/mathml/M1','MathML',630,470);return false;" target="_blank" href="http://www.microbialcellfactories.com/content/11/1/128/mathml/M1">View MathML</a>

(1)

The competition for H2 among organohalide respirers and methanogens has been well documented [22-28]. However, none of these studies have addressed how consumption of H2, whether added as gas or as a result of fermentation, is affected by varying HCO3 concentrations.

Homoacetogens are other important microorganisms commonly encountered with organohalide respirers. Homoacetogens produce H2 from fermentation of complex substrates and/or consume available H2[29,30]. Hydrogenotrophic homoacetogens catalyze the formation of acetate from H2 and HCO3 in their energy metabolism [29]:

Hydrogenotrophic homoacetogenesis:

<a onClick="popup('http://www.microbialcellfactories.com/content/11/1/128/mathml/M2','MathML',630,470);return false;" target="_blank" href="http://www.microbialcellfactories.com/content/11/1/128/mathml/M2">View MathML</a>

(2)

They, too, compete for H2 with organohalide respirers. To date, the limited number of studies that have investigated hydrogenotrophic homoacetogenesis in TCE dechlorinating consortia [27,31] has not included the HCO3 concentration as a variable driving the extent and the rates of reductive dechlorination.

Hydrogenotrophic methanogens and homoacetogens can also affect pH in dechlorinating communities. Methanogens produce methane as the end product (Equation 1) by expending one proton and one HCO3, while hydrogenotrophic homoacetogens generate acetate (Equation 2) from one proton and two HCO3. Both reactions increase the pH while consuming HCO3, which often is the only buffer in the system. The effect of HCO3 concentration in TCE dechlorinating microbial communities has not been studied. Few studies focusing on other dechlorinating systems have recognized its importance and examined the effect of HCO3 concentrations on the formation of chlorinated daughter products, thus motivating this work. For example, removal of chlorophenols from simulated wastewater in upflow anaerobic sludge blanket (UASB) reactors revealed significant inhibition on dechlorination at high HCO3 (3500 mg L−1 as CaCO3) and high pH [32]. In microcosms showing microbial dechlorination of polychlorinated biphenyls with H2 gas as electron donor, 100 mg L−1 HCO3 (1.64 mM) yielded the fastest rates of dechlorination, whereas addition of 1000 mg L−1 HCO3 (16.4 mM) resulted in the slowest polychlorinated biphenyls dechlorination rates and triggered the most acetate to form [33].

In this study, we evaluate the role of HCO3 as a buffering agent and as an electron acceptor in TCE reductively dechlorinating mixed communities using a previously described culture, DehaloR^2, as a model consortium [20]. H2, and not fermentable substrates, was used as the sole electron donor to directly and accurately measure hydrogenotrophic production of methane and acetate from HCO3. The concentrations of HCO3 tested reflect typical groundwater concentrations (2.5-10 mM), as well as commonly reported laboratory concentrations (30 mM).

Results and discussion

Chloroethenes reductive dechlorination at different HCO3 concentrations

The time course dechlorination measurements presented in Figure 1 show a short lag time for the onset of dechlorination of 0.55 mmol L−1 TCE. TCE to cis-DCE conversion was the fastest dechlorination step in all cultures, with only VC and ethene detected after day 5, regardless of the concentration of HCO3 added. A close monitoring of VC to ethene reduction rates between each GC measurement revealed that after day 5, dechlorination rates had slowed down at all HCO3 concentrations, especially in the cultures containing 30 mM (Figure 1G-H), suggesting an electron donor limitation. The measured H2 levels on day 12 were 1.5 mmol L–1 at 2.5 mM HCO3 and 0.5 mmol L–1 at 5 mM HCO3. At 10 and 30 mM HCO3, no H2 peak was detected on the GC-TCD on day 12. Immediately after injecting an additional 8.2 mmol L–1 H2 on day 12, we observed an increase in the rates of VC consumption and ethene formation (Figure 1A-H).

thumbnailFigure 1. Chloroethenes dechlorination at different HCO3concentrations. Time course of chloroethenes dechlorination to ethene in cultures amended with H2 as the sole electron donor and with HCO3 buffer (graphs A, C, E, and G) and a combination of HCO3 and HEPES buffers (graphs B, D, F, and H). The arrows represent the 2nd and 3rd addition of 8.2 mmol L−1 H2. The error bars are standard deviations of triplicate cultures.

Following the second addition of H2, all cultures reached ≥70% conversion of TCE to ethene. Complete TCE dechlorination (Figure 1D) was first observed between days 17 and 18 in cultures containing 5 mM HCO3 and 5 mM 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES), which was provided as an additional buffer. Complete conversion to ethene was further observed in the cultures with 2.5 mM HCO3 & HEPES on day 26. A threefold increase in the 16S rRNA Dehalococcoides genes (data not shown) from 1.13 × 1011 (±4.98 × 109) copies L−1 (time 0) to 3.71 × 1011 (±2.78 × 1010) and 3.67 × 1011 (±8.04 × 109) copies L−1 was detected after complete dechlorination at 5 mM HCO3 & HEPES and 2.5 mM HCO3 & HEPES, respectively. Chloroethenes conversion rates in the cultures containing 10 and 30 mM HCO3 were the slowest, as seen in Figure 1. The Dehalococcoides 16S rRNA gene copies per L in the cultures with HCO3 and HEPES after complete conversion to ethene were 2.07 × 1011 (± 5.79 × 109) at 10 mM and 2.03 × 1011 (± 5.59 × 109) at 30 mM (data not shown). The lower resulting cell density coupled to decreased dechlorination rates indicates that Dehalococcoides growth was diminished at the higher HCO3 concentrations (Student’s t test; ≥70% confidence level).

We observed a second H2 limitation at 10 and 30 mM HCO3, with the complete cessation of VC reduction at 30 mM between days 18 and 26 (Figure 1G-H). Consequently, an additional dose of H2 (8.2 mmol L–1) was injected into all cultures still undergoing dechlorination. With the 3rd addition of electron donor, the 10 and 30 mM HCO3 cultures supplemented with HEPES dechlorinated all TCE to ethene by day 28 and 32 (Figure 1F and H), respectively. The parallels without HEPES showed incomplete conversion to ethene even by day 40 (Figure 1E and G) and VC dechlorination had stalled once again on day 35, or it was proceeding at very reduced rates.

Methane and acetate production during TCE reductive dechlorination

In Figure 1, we show how H2 was limiting dechlorination rates before the 2nd and 3rd H2 addition at the different concentrations of HCO3 tested. The theoretical H2 demand for 0.55 mmol L−1 TCE is 1.65 mmol L−1H2. Considering that the H2 at time 0 was 8.2 mmol L−1, five times in excess of the theoretical demand for dechlorination, the slower dechlorination rates observed, together with H2 depletion, indicated that competing microorganisms were consuming H2 faster than the dechlorinators. An increase in methane of only 0.01 mmol L−1 was detected at all HCO3 concentrations before day 4 (Figure 2A), which coincides with the disappearance of TCE and formation of less chlorinated daughter products (Figure 1). The lack of methane production was also confirmed by the qPCR data which show no relative increase in the numbers of Archaea gene copies L−1 at this time point compared to the 10% inoculum (Figure 2B, day 7). The lag time for methane production could have been due to the previously reported longer lag time of the methanogenic microorganisms [34] and the toxic effect of TCE on methanogens [31]. Additionally, besides Dehalococcoides, other dechlorinators can use TCE as electron acceptor and H2 as electron donor to produce cis-DCE. A competitive advantage of Geobacter spp., the other identified TCE to cis-DCE respirers in the inoculum culture [20], over methanogens could have also contributed to a delayed onset of methanogenesis.

thumbnailFigure 2. Methanogenesis and homoacetogenesis during active evolvement of reductive dechlorination. Methane (A) and acetate (C) production during reductive dechlorination in medium buffered with 2.5 (circles), 5 (triangles), 10 (squares), and 30 (diamonds) mM HCO3. The error bars are standard deviations of triplicate cultures and the arrows represent the 2nd and 3rd addition of 8.2 mmol L−1 H2. Quantification of methanogens, Archaea (B), and homoacetogens, FTHFS (D) using qPCR. The error bars are standard deviations of triplicate analytical runs.

Methanogenesis was mostly stimulated at 2.5 mM HCO3 and 5 mM HCO3, and it was less active with increasing concentrations of HCO3 (Figure 2A). The methane production trends observed are supported by a higher increase in Archaea numbers at the lower HCO3 concentrations (2.5 and 5 mM in Figure 2B) compared to 10 and 30 mM (Figure 2B). At 30 mM HCO3, we detected no net increase in methane between day 10 and 12, suggesting that methanogens, like dechlorinators, were also experiencing H2 limitation. Once H2 became available after the second addition, methane production rates quickly increased in all cultures (Figure 2A).

Upon the third addition of H2 (day 26), methane no longer increased at 2.5 mM HCO3 even though H2 was provided (Figure 2A, day 26–32), indicating a HCO3, and not a H2 limitation. Even though HCO3 was not measured due to analytical limitations, we were able to track HCO3 consumption via production of methane and acetate, as illustrated in Additional file 1. The HCO3 utilization balance presented in Additional file 1 shows that production of methane (and to a lesser degree acetate) exhausted all the HCO3 in the systems initially supplemented with 2.5 mM.

Additional file 1. Calculated HCO3consumption for methane and acetate production.

Format: PDF Size: 211KB Download file

This file can be viewed with: Adobe Acrobat ReaderOpen Data

Homoacetogenesis exhibited the opposite trend to methanogenesis. According to the time course concentrations recorded and shown in Figure 2C, more acetate was produced when more HCO3 buffer was present. Additionally, among all conditions tested, the greatest increase in copies L−1 culture by day 7 of the formyltetrahydrofolate synthase (FTHFS) gene, a functional marker for acetogens, was detected at 30 mM HCO3 (Figure 2D), and the relative numbers of gene copies were lower with decreasing concentrations of HCO3. Before the second addition of H2, all cultures showed an increase of 0.3-1.3 mM acetate (Figure 2C). However, after injecting the second dose of H2, only a small rise in acetate was observed at 2.5 and 5 mM HCO3. In contrast, at 10 and 30 mM HCO3, we detected a boost in homoacetogenesis (Figure 2C) and corresponding higher increases in the FTHFS gene (Figure 2D).

The qPCR data for both methanogens and homoacetogens correlate well with our analytical data. The resulting increased levels of homoacetogens coupled to the lowest levels of methanogens at 30 mM HCO3 indicate benefits for the first group at the higher HCO3 concentrations. Unlike homoacetogens, the resulting methanogenic microorganisms were present at similar levels in cultures initially containing 2.5 and 5 mM HCO3 and less plentiful in cultures initially containing 10 and 30 mM HCO3 (Figure 2B). Overall, our findings are consistent with the lower HCO3 requirement for methane production: one mol HCO3 consumed for one mol methane (Equation 1) vs. two mol HCO3 consumed for one mol acetate (Equation 2). Additionally, these data are in agreement with the findings of Florencio et al., 1995 [35] on substrate competition between methylotrophic methanogens and methanol-utilizing acetogens in UASB reactors, where acetogenesis was significant and outcompeted methanogenesis only in the presence of exogenously supplemented HCO3.

Distribution of electrons for H2-consuming processes

The fate of electrons fed as H2 is depicted in Figure 3. By day 12 (after one addition of H2; Figure 3A), 70% or greater of the total added electrons can be accounted for towards the three main energy-deriving reactions, dechlorination, homoacetogenesis and methanogenesis, under all conditions tested. Biomass was not included in these balances, however, a 10-20% fraction of the total electrons can be assumed for cell synthesis [36]. 1.65 mmol H2, the theoretical H2 requirement for dechlorination of 0.55 mmol TCE, equals to 3.3 me- equivalents H2, and each 8.2 mmol L–1 H2 addition represents 16.4 me- equivalents. Out of the three main processes occurring in our test systems, TCE dechlorination utilized a small fraction of 9.3% out of the total me- equivalents for the cultures that completed dechlorination with two H2 additions (Figure 1B and D), and 6.7% of the total me- equivalents for those that received three H2 additions (Figure 1A, C, E, F, G, and H).

thumbnailFigure 3. Distribution of electrons fed as H2 towards dechlorination, methanogenesis, and homoacetogenesis at various HCO3concentrations. (A) Average data from triplicate cultures on day 12 after addition of 16.4 me- equiv. H2 (8.2 mmol L−1). (B) Average final data from triplicate cultures after addition of 33 me- equivalents (16.4 mmol L−1 H2 in 2.5 mM HCO3 & HEPES and 5 mM HCO3 & HEPES) and 49 me- equivalents (32.8 mmol L−1 in all other sets). The presence of the additional buffer, HEPES, is denoted as H on the X-axis.

From the H2 me- equivalents provided at time 0, only 18.3% would have been required to completely reduce TCE to ethene. As seen in Figures 1 and 3A, none of the cultures, regardless of their H2 demand, completed dechlorination with the initial H2. Additionally, the 10 and 30 mM HCO3 amendments with or without HEPES received H2 fifteen times in excess of the theoretical demand for dechlorination, yet only the sets supplemented with HEPES completed dechlorination, implicating an important pH factor, which is discussed in the next section.

Overall, the fate of most H2 me- equivalents was to HCO3-driven reactions towards the production of methane and acetate. Acetate from hydrogenotrophic homoacetogenesis was also found to be the main sink of electrons in a field study that used H2 gas for remediation of chlorinated ethenes in groundwater [9]. Moreover, Duhamel and Edwards 2007 [18] investigated the growth and yields of hydrogenotrophic methanogens, acetogens and dechlorinators during the process of dechlorination. The authors found that most of the electrons fed as methanol in 30 mM HCO3- buffered medium went towards acetogenesis and, that methanogens were outcompeted by acetogens. Our data from 10 and 30 mM HCO3 corroborate their findings; however, one important additional finding from our experiments, as seen in Figure 2 and 3, is that methanogens can outcompete homoacetogens at low HCO3 concentrations (2.5 and 5 mM).

The results on TCE dechlorination, methanogenesis and homoacetogenesis from this work at different HCO3 concentrations offer some insights into which competing microbial groups will prevail and how HCO3 consumption affects rates of dechlorination. Furthermore, our study also alludes to how HCO3 drives the H2 competition between organohalide respirers, methanogens, and homoacetogens. This important aspect has not been determined previously in reductive dechlorination, to our knowledge. In addition, for application purposes, it is important to consider how temperature could affect these findings, as these predictions might be somewhat different at lower temperatures, such as those in groundwater. Our experiments were performed at 30°C, however, temperature studies on organohalide respirers (i.e. Dehalococcoides) have documented slower rates of dechlorination at 10-15°C compared to their maximum rates at 30-35°C [37]. Homoacetogens are even greater H2 and HCO3 consumers than methanogens at lower temperatures [34,38], hence, the predominance of homoacetogens would be greater in groundwater systems. Furthermore, because many homoacetogens can consume fermentables and/or H2 to produce acetate [29], it is important to consider homoacetogenesis as an electron sink and alkalinity-consuming process in dechlorination at the laboratory and field scale. Although comprehensive models on in situ reductive dechlorination have been developed [13,14,22,39], the introduction of hydrogenotrophic homoacetogenesis in these models has not been considered.

Effect of pH on dechlorination in HCO3 -amended cultures

We supplemented HEPES to all HCO3 concentrations tested to separate between the effect of HCO3 as an electron acceptor/sink and the effect of pH changes resulting from microbial processes that use HCO3 as an electron acceptor, i.e. methanogenesis and homoacetogenesis. The time course pH measurements presented in Additional file 2 and the final measurements in Figure 4 uncovered a trend when HCO3 was the sole buffer: a higher pH increase with increasing HCO3 concentrations due to methanogenesis and homoacetogenesis HCO3-consuming reactions. This was not the case at 30 mM HCO3, where we recorded a lower final pH than at 10 mM HCO3 (Figure 4) due to the buffering capacity from the remaining 20 mM unconsumed HCO3 (Additional file 1). However, in a separate experiment where we increased the total concentration of H2 to 41.2 mmol L−1 in cultures containing 30 mM HCO3, we recorded a final pH of 9.6 under these conditions (data not shown). These cultures also exhibited slower rates of dechlorination compared to the data from Figure 1 and no ethene formed by day 40 of the experiments (data not shown).

Additional file 2. Time course pH measurements.

Format: PDF Size: 222KB Download file

This file can be viewed with: Adobe Acrobat ReaderOpen Data

thumbnailFigure 4. pH changes resulting from biological HCO3consumption. Average initial (t = 0) and final pH measurements in all HCO3 amendments from this study in the absence (closed symbols) or presence (open symbols) of HEPES. The error bars are standard deviations of triplicate cultures. The buffer HEPES is denoted as H on the X-axis.

An increase in pH at all HCO3 concentrations tested was also observed when HEPES was present as an additional buffer but the pH increase was within a much narrower range (Figure 4). We ran statistical analyses and determined that, because of better pH buffering, the rates of dechlorination were significantly faster (Student t-test, P <0.05) in the presence of HEPES, compared to when HCO3 was the sole buffer (Figure 1). In this study, we show that high pH can also occur in dechlorinating systems, especially in engineered systems fed with H2, and this pH change can negatively impact chloroethenes reduction. A detrimental effect on TCE dechlorination that resulted in accumulation of mainly cis-DCE at pH 8.3 was previously observed in an anaerobic biotrickling filter [40]. Our results show that high pH is stressful to TCE dechlorinating microorganisms, hence, research on bioremediation of chloroethenes will greatly benefit from comprehensive pH studies.

Conclusions

Despite the fact that HCO3 is a common natural buffer and addition of more HCO3 can counteract pH deviations from the optimum range for dechlorination, the results of our study point out that 1) high HCO3 concentrations increase the H2 demand, and that 2) consumption of HCO3 contributes to pH increases that could adversely affect TCE dechlorination rates or result in accumulation of toxic intermediate by-products (i.e., DCE and VC). Our findings regarding the effect of pH increases from HCO3-consuming reactions are relevant for ex situ chloroethenes remediation technologies that provide H2 and for laboratory amendments. When fermentable substrates are used to stimulate reductive dechlorination, or, in the case of groundwater where HCO3 is replenished from minerals dissolution or organics oxidation, this increase in pH will likely be offset by the protons produced from fermentation or by the constant supply of buffer.

However, the lessons learned from this study on dechlorination, methanogenesis, and homoacetogenesis highlight that HCO3, especially when abundant, could be an important variable for biologically-driven TCE dechlorination, as it has a prominent role as an electron acceptor by stimulating competing H2-consuming processes. Our findings also point out that a shift in the main H2 competitors occurs depending on the HCO3 concentration available in the environment, with homoacetogens as the greater electron sink at high HCO3, and methanogens as the main H2 competitors at low HCO3.

Methods

Microbial inoculum and preparation of batch cultures

The sediment-free microbial consortium, DehaloR^2, described by Ziv-El et al., 2011 [20] was used as the inoculum. For the experiments in this study, we preconditioned the inoculum culture by growing it in 10 mM HCO3 medium, with excess H2 as electron donor, and two consecutive feedings of 10 μL neat TCE in 120 mL medium.

Reduced anaerobic mineral medium was prepared containing the following reagents per liter: 1 g NaCl, 0.06 g MgCl2 × 6H2O, 0.2 g KH2PO4, 0.3 g NH4Cl, 0.3 g KCl, 0.005 g CaCl2 × 2H2O, and 1 mL of Trace A and Trace B solutions described elsewhere [17]. During medium preparation, nitrogen was the sole gas for boiling and bottling and the reducing agents were 0.2 mM L-cysteine and 0.2 mM Na2S × 9 H2O. No buffer was added to the medium before autoclaving. For bottling, we used 160-mL glass serum bottles containing 90 mL liquid and 70 mL headspace sealed with black butyl rubber stoppers and aluminum crimps.

The concentrations of NaHCO3 tested were 2.5, 5, 10, and 30 mM. In the cultures where both NaHCO3 and HEPES (pKa = 7.55) were used as buffers, we supplemented 5 mM HEPES in combination with 2.5, 5 and 10 mM HCO3, and 10 mM HEPES in the 30 mM HCO3 cultures. NaHCO3 and HEPES were delivered to each bottle from 1 M sterile anaerobic stock solutions. The initial pH was adjusted with sterile 2.25 N HCl or NaOH to 7.5 (± 0.1). At time 0, we added to each culture bottle 0.55 mmol L–1 TCE (5 μL neat or 71.3 mg L−1), 1 mL ATCC vitamin mix, 50 μL of 1 g L−1 vitamin B12 solution, 8.2 mmol L–1 H2 (20 mL H2 gas), and 10 mL DehaloR^2 microbial culture, corresponding to a 10% inoculum. The working volume was 100 mL of liquid with 60 mL of headspace. The bottles were incubated in the dark at 30°C without shaking. An additional 8.2 mmol L–1 H2 was added on day 12 (all cultures) and on day 26 (only to cultures still undergoing dechlorination). All experimental conditions were tested in triplicates and the experiments were also performed on two separate occasions.

Chemical and pH measurements for the time course experiments

We measured TCE, cis-DCE, VC, ethene, and methane using a gas chromatograph (GC) (Shimadzu GC-2010; Columbia, MD) equipped with a flame ionization detector (FID). The compounds were carried by helium gas through an Rt-QS-BOND capillary column (Restek; Bellefonte, PA). The oven temperature was maintained at 110°C for 1 min, followed by a temperature increase of 50°C min−1 to 200°C. Then, the temperature ramp was further raised to 240°C with a 15°C min−1 gradient and held for 1.5 mins. The temperatures of the FID and the injector were 240°C. Chloroethenes, ethene and methane calibrations in 160-mL bottles with 100 mL liquid volume were performed in a range of 0.05-2.45 mmol L–1. The detection limit for all compounds measured on the GC-FID is ≤0.018 mmol L–1.

A GC instrument equipped with a thermal conductivity detector (TCD) was employed to measure H2 before reinjecting additional H2 to the cultures on day 12. The instrument settings used were those previously outlined [41]. The H2 detection limit for the GC-TCD is 0.8% vol/vol.

We quantified acetate, propionate, and formate from 0.75-mL liquid samples filtered through a 0.2 μm polyvinylidene fluoride membrane syringe filter (Pall Corporation; Ann Arbor, MI) into 2-mL glass vials (VWR; Radnor, PA) via high performance liquid chromatography (HPLC) using a previously published method [41]. Five point calibration curves (0.5-10 mM) were generated for acetate, propionate, and formate during every HPLC run. The detection limit for the compounds measured on the HPLC was ≤0.1 mM.

0.29 ± 0.06 mM propionate was carried over from the inoculum culture and the final measured concentration was 0.33 ± 0.04 mM, indicative that propionate did not serve as a significant source of electrons. Formate was sometimes also detected at low concentrations (0.1-0.3 mM), however, we did not identify a clear trend on the formation/consumption of this product. Therefore, propionate and formate were omitted from the electron balances in Figure 3.

The pH was measured using an Orion 2-Star pH bench top meter (Thermo Scientific, USA) that was calibrated regularly with 4.01, 7.00, and 10.01 standard solutions from the manufacturer.

All cultures were sampled for gas and liquid analyses until dechlorination of TCE to ethene was complete or until the end of experiments on day 40.

DNA extraction and molecular microbial characterization

Pellets were formed by centrifugation from 2-mL liquid from each culture replicate and they were stored at −20°C until the DNA extraction. Genomic DNA was extracted for two time points for all sets of HCO3 & HEPES, and two time points for the set with 30 mM HCO3 only. Before DNA extraction, the replicate pellets were thawed, resuspended in the supernatant, and combined, so that only extraction per set per time point was performed. This was done to increase total biomass and DNA yield. The DNA extraction was performed as previously described [20].

We employed quantitative real-time PCR to target the 16S rRNA gene of Dehalococcoides and Archaea (TaqMan® assays) and the FTHFS gene of homoacetogens (SYBR Green assay). Triplicate reactions were setup for the six point standard curves and the samples in 10 μL total volume using 4 μL of 1/10 diluted DNA as template. We generated standard curves by serially diluting 10 ng μL–1 plasmid DNA. The primers and probes, reagents concentrations, and thermocycler (Realplex 4S thermocycler; Eppendorf, USA) conditions were those described for Dehalococcoides[42], Archaea[43,44], and FTHFS [44,45]. Acetoclastic methanogens (the order Methanosarcinales) were not assayed because they are absent in the culture employed in this study, which this was confirmed by qPCR previously [20].

Time 0 for all qPCR assays was generated by amplifying genomic DNA from the inoculum culture and assigning 10% as the starting concentrations of gene copies per L culture.

Calculations

The distributions of electrons from Figure 3 were calculated in units of me- equivalents for each compound from the equation below:

<a onClick="popup('http://www.microbialcellfactories.com/content/11/1/128/mathml/M3','MathML',630,470);return false;" target="_blank" href="http://www.microbialcellfactories.com/content/11/1/128/mathml/M3">View MathML</a>

(3)

The number of me- equivalents for dechlorination is 2, 4, and 6 for DCE, VC and ethene, respectively, 8 for acetate and methane, and 2 for H2.

End notes

aThroughout this work, HCO3 is used to denote the buffer HCO3/CO2. At the pH ranges observed in this study, HCO3 accounted for 90% or greater of the two species.

Competing interests

The authors declare that they have no competing interests.

Authors’ contributions

AGD and RKB designed the experiments that led to the writing of the manuscript. AGD and DFW carried out the experimental work. PP participated in the performance and analyses for quantitative PCR. AGD drafted the manuscript; RKB, PP, and RUH critically reviewed and contributed to the intellectual merit of the paper. All authors read and approved the final version of the manuscript.

Acknowledgements

This project was supported by NSF CAREER Award Number 1053939 to RKB. The content is solely the responsibility of the authors and does not necessarily represent the official views of NSF.

References

  1. Maymo-Gatell X, Chien YT, Gossett JM, Zinder SH: Isolation of a bacterium that reductively dechlorinates tetrachloroethene to ethene.

    Science 1997, 276:1568-1571. PubMed Abstract | Publisher Full Text OpenURL

  2. Ellis DE, Lutz EJ, Odom JM, Buchanan RJ, Bartlett CL, Lee MD, Harkness MR, Deweerd KA: Bioaugmentation for accelerated in situ anaerobic bioremediation.

    Environ Sci Technol 2000, 34:2254-2260. Publisher Full Text OpenURL

  3. Marzorati M, Balloi A, de Ferra F, Corallo L, Carpani G, Wittebolle L, Verstraete W, Daffonchio D: Bacterial diversity and reductive dehalogenase redundancy in a 1,2-dichloroethane-degrading bacterial consortium enriched from a contaminated aquifer.

    Microb Cell Fact 2010, 9:12. PubMed Abstract | BioMed Central Full Text | PubMed Central Full Text OpenURL

  4. Tas N, van Eekert MHA, de Vos WM, Smidt H: The little bacteria that can - diversity, genomics and ecophysiology of 'Dehalococcoides' spp. in contaminated environments.

    Microb Biotechnol 2010, 3:389-402. PubMed Abstract OpenURL

  5. He JZ, Ritalahti KM, Yang KL, Koenigsberg SS, Loffler FE: Detoxification of vinyl chloride to ethene coupled to growth of an anaerobic bacterium.

    Nature 2003, 424:62-65. PubMed Abstract | Publisher Full Text OpenURL

  6. Loffler FE, Yan J, Ritalahti KM, Adrian L, Edwards EA, Konstantinidis KT, Müller JA, Fullerton H, Zinder SH, Spormann AM: Dehalococcoides mccartyi gen. nov., sp. nov., obligate organohalide-respiring anaerobic bacteria, relevant to halogen cycling and bioremediation, belong to a novel bacterial class, Dehalococcoidetes classis nov., within the phylum Chloroflexi.

    Int J Syst Evol Microbiol

    in press

    OpenURL

  7. Major DW, McMaster ML, Cox EE, Edwards EA, Dworatzek SM, Hendrickson ER, Starr MG, Payne JA, Buonamici LW: Field demonstration of successful bioaugmentation to achieve dechlorination of tetrachloroethene to ethene.

    Environ Sci Technol 2002, 36:5106-5116. PubMed Abstract | Publisher Full Text OpenURL

  8. Schaefer CE, Lippincott DR, Steffan RJ: Field-scale evaluation of bioaugmentation dosage for treating chlorinated ethenes.

    Ground Water Monit Rem 2010, 30:113-124. Publisher Full Text OpenURL

  9. Edstrom JA, Semmens MJ, Hozalski RM, Clapp LW, Novak PJ: Stimulation of dechlorination by membrane-delivered hydrogen: small field demonstration.

    Environ Eng Sci 2005, 22:281-293. Publisher Full Text OpenURL

  10. Ma X, Novak PJ, Clapp LW, Semmens MJ, Hozalski RM: Evaluation of polyethylene hollow-fiber membranes for hydrogen delivery to support reductive dechlorination in a soil column.

    Water Res 2003, 37:2905-2918. PubMed Abstract | Publisher Full Text OpenURL

  11. Villano M, De Bonis L, Rossetti S, Aulenta F, Majone M: Bioelectrochemical hydrogen production with hydrogenophilic dechlorinating bacteria as electrocatalytic agents.

    Bioresource Technol 2011, 102:3193-3199. Publisher Full Text OpenURL

  12. Ziv-El M, Popat SC, Cai K, Halden RU, Krajmalnik-Brown R, Rittmann BE: Managing methanogens and homoacetogens to promote reductive dechlorination of trichloroethene with direct delivery of H2 in a membrane biofilm reactor.

    Biotechnol Bioeng 2012, 109:2200-2210. PubMed Abstract | Publisher Full Text OpenURL

  13. Robinson C, Barry DA: Design tool for estimation of buffer requirement for enhanced reductive dechlorination of chlorinated solvents in groundwater.

    Environ Model Softw 2009, 24:1332-1338. Publisher Full Text OpenURL

  14. Robinson C, Barry DA, McCarty PL, Gerhard JI, Kouznetsova I: pH control for enhanced reductive bioremediation of chlorinated solvent source zones.

    Sci Total Environ 2009, 407:4560-4573. PubMed Abstract | Publisher Full Text OpenURL

  15. Abdelouas A, Lu YM, Lutze W, Nuttall HE: Reduction of U(VI) to U(IV) by indigenous bacteria in contaminated ground water.

    J Contam Hydrol 1998, 35:217-233. Publisher Full Text OpenURL

  16. Wilkin RT, Digiulio DC: Geochemical impacts to groundwater from geologic carbon sequestration: controls on pH and inorganic carbon concentrations from reaction path and kinetic modeling.

    Environ Sci Technol 2010, 44:4821-4827. PubMed Abstract | Publisher Full Text OpenURL

  17. Loffler FE, Sanford RA, Ritalahti KM: Enrichment, cultivation, and detection of reductively dechlorinating bacteria.

    Environ Microbiol 2005, 397:77-111. OpenURL

  18. Duhamel M, Edwards EA: Growth and yields of dechlorinators, acetogens, and methanogens during reductive dechlorination of chlorinated ethenes and dihaloelimination of 1,2-dichloroethane.

    Environ Sci Technol 2007, 41:2303-2310. PubMed Abstract | Publisher Full Text OpenURL

  19. Vainberg S, Condee CW, Steffan RJ: Large-scale production of bacterial consortia for remediation of chlorinated solvent-contaminated groundwater.

    J Ind Microbiol Biotechnol 2009, 36:1189-1197. PubMed Abstract | Publisher Full Text OpenURL

  20. Ziv-El M, Delgado AG, Yao Y, Kang DW, Nelson KG, Halden RU, Krajmalnik-Brown R: Development and characterization of DehaloR^2, a novel anaerobic microbial consortium performing rapid dechlorination of TCE to ethene.

    Appl Microbiol Biotech 2011, 92:1063-1071. Publisher Full Text OpenURL

  21. Cordruwisch R, Seitz HJ, Conrad R: The capacity of hydrogenotrophic anaerobic bacteria to compete for traces of hydrogen depends on the redox potential of the terminal electron acceptor.

    Arch Microbiol 1988, 149:350-357. Publisher Full Text OpenURL

  22. Fennell DE, Gossett JM: Modeling the production of and competition for hydrogen in a dechlorinating culture.

    Environ Sci Technol 1998, 32:2450-2460. Publisher Full Text OpenURL

  23. Fennell DE, Gossett JM, Zinder SH: Comparison of butyric kid, ethanol, lactic acid, and propionic acid as hydrogen donors for the reductive dechlorination of tetrachloroethene.

    Environ Sci Technol 1997, 31:918-926. Publisher Full Text OpenURL

  24. Carr CS, Hughes JB: Enrichment of high rate PCE dechlorination and comparative study of lactate, methanol, and hydrogen as electron donors to sustain activity.

    Environ Sci Technol 1998, 32:1817-1824. Publisher Full Text OpenURL

  25. Smatlak CR, Gossett JM, Zinder SH: Comparative kinetics of hydrogen utilization for reductive dechlorination of tetrachloroethene and methanogenesis in an anaerobic enrichment culture.

    Environ Sci Technol 1996, 30:32850-2858. OpenURL

  26. Aulenta F, Gossett JM, Papini MP, Rossetti S, Majone M: Comparative study of methanol, butyrate, and hydrogen as electron donors for long-term dechlorination of tetrachloroethene in mixed anerobic cultures.

    Biotechnol Bioeng 2005, 91:743-753. PubMed Abstract | Publisher Full Text OpenURL

  27. Yang YR, McCarty PL: Competition for hydrogen within a chlorinated solvent dehalogenating anaerobic mixed culture.

    Environ Sci Technol 1998, 32:3591-3597. Publisher Full Text OpenURL

  28. Ballapragada BS, Stensel HD, Puhakka JA, Ferguson JF: Effect of hydrogen on reductive dechlorination of chlorinated ethenes.

    Environ Sci Technol 1997, 31:1728-1734. Publisher Full Text OpenURL

  29. Drake HL: Acetogenesis. New York, NY: Chapman & Hall; 1994. OpenURL

  30. Rittmann S, Herwig C: A comprehensive and quantitative review of dark fermentative biohydrogen production.

    Microb Cell Fact 2012, 11:115. PubMed Abstract | BioMed Central Full Text | PubMed Central Full Text OpenURL

  31. Yang YR, McCarty PL: Biologically enhanced dissolution of tetrachloroethene DNAPL.

    Environ Sci Technol 2000, 34:2979-2984. Publisher Full Text OpenURL

  32. Majumder PS, Gupta SK: Effect of influent pH and alkalinity on the removal of chlorophenols in sequential anaerobic-aerobic reactors.

    Bioresource Technol 2009, 100:1881-1883. Publisher Full Text OpenURL

  33. Yan T, LaPara TM, Novak PJ: The effect of varying levels of sodium bicarbonate on polychlorinated biphenyl dechlorination in Hudson River sediment cultures.

    Environ Microbiol 2006, 8:1288-1298. PubMed Abstract | Publisher Full Text | PubMed Central Full Text OpenURL

  34. Fey A, Conrad R: Effect of temperature on carbon and electron flow and on the archaeal community in methanogenic rice field soil.

    Appl Environ Microbiol 2000, 66:4790-4797. PubMed Abstract | Publisher Full Text | PubMed Central Full Text OpenURL

  35. Florencio L, Field JA, Lettinga G: Substrate competition between methanogens and acetogens during the degradation of methanol in UASB reactors.

    Water Res 1995, 29:915-922. Publisher Full Text OpenURL

  36. Rittmann BE: McCarty PL: Environmental Biotechnology: Principles and Applications. Boston: McGraw-Hill Book Co; 2001. OpenURL

  37. Friis AK, Heimann AC, Jakobsen R, Albrechtsen HJ, Cox E, Bjerg PL: Temperature dependence of anaerobic TCE-dechlorination in a highly enriched Dehalococcoides-containing culture.

    Water Res 2007, 41:355-364. PubMed Abstract | Publisher Full Text OpenURL

  38. Kotsyurbenko OR, Glagolev MV, Nozhevnikova AN, Conrad R: Competition between homoacetogenic bacteria and methanogenic archaea for hydrogen at low temperature.

    FEMS Microbiol Ecol 2001, 38:153-159. Publisher Full Text OpenURL

  39. Clapp LW, Semmens MJ, Novak PJ, Hozalski RM: Model for in situ perchloroethene dechlorination via membrane-delivered hydrogen.

    J Environ Eng-ASCE 2004, 130:1367-1381. Publisher Full Text OpenURL

  40. Popat SC, Deshusses MA: Reductive dehalogenation of trichloroethene vapors in an anaerobic biotrickling filter.

    Environ Sci Technol 2009, 43:7856-7861. PubMed Abstract | Publisher Full Text OpenURL

  41. Parameswaran P, Torres CI, Lee HS, Rittmann BE, Krajmalnik-Brown R: Hydrogen consumption in microbial electrochemical systems (MXCs): the role of homo-acetogenic bacteria.

    Bioresource Technol 2011, 102:263-271. Publisher Full Text OpenURL

  42. Holmes VF, He JZ, Lee PKH, Alvarez-Cohen L: Discrimination of multiple Dehalococcoides strains in a trichloroethene enrichment by quantification of their reductive dehalogenase genes.

    App Environ Microb 2006, 72:5877-5883. Publisher Full Text OpenURL

  43. Yu Y, Lee C, Kim J, Hwang S: Group-specific primer and probe sets to detect methanogenic communities using quantitative real-time polymerase chain reaction.

    Biotechnol Bioeng 2005, 89:670-679. PubMed Abstract | Publisher Full Text OpenURL

  44. Parameswaran P, Zhang HS, Torres CI, Rittmann BE, Krajmalnik-Brown R: Microbial community structure in a biofilm anode fed with a fermentable substrate: the significance of hydrogen scavengers.

    Biotechnol Bioeng 2010, 105:69-78. PubMed Abstract | Publisher Full Text OpenURL

  45. Xu KW, Liu H, Du GC, Chen J: Real-time PCR assays targeting formyltetrahydrofolate synthetase gene to enumerate acetogens in natural and engineered environments.

    Anaerobe 2009, 15:204-213. PubMed Abstract | Publisher Full Text OpenURL