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6 Organohalide Respiration Rates in OHRB Communities and Modeling of Community Food Webs
14 Organohalide-Respiring Bacteria as Members of Microbial …
respiring chlorinated ethenes. For example, rates of approximately 100 µmole
Cl−/L/h have been reported in cultures respiring chlorinated ethenes and containing Dhc populations near 1E9 cells/mL (Rowe et al. 2008; Delgado et al. 2014;
Ziv-El et al. 2011; Duhamel et al. 2002; Yu et al. 2005; Maymó-Gatell et al. 1997;
Mansfeldt et al. 2014). The fastest reported culture to fully dechlorinate TCE to
ethene was grown in a chemostat with a three-day retention time and achieved 130
µmole Cl−/L/h (Delgado et al. 2014). The original report on the isolation of Dhc
strain 195 presented data suggesting that pure cultures can reach high density (1E9
cells/mL) and rates (120 µmoles Cl−/L/h) approaching those of mixed cultures
(Maymó-Gatell et al. 1997). Sulfurospirillum is a pure culture with high rates of
dechlorination. For example, Sulfurospirillum multivorans was shown to dechlorinate PCE to TCE at a rate of 45 µmoles Cl−/L/h (Scholz-Muramatsu et al. 1995).
However, in most studies, pure culture rates are much lower than rates reported for
mixed cultures or defined cocultures.
Organohalide respiration rates for other organohalide classes are generally
lower than for chlorinated ethenes. Chlorinated ethane-respiring cultures show
rates on the order of 1 µmole Cl−/L/h (Grostern and Edwards 2006a; Manchester
et al. 2012). Chlorobenzene-enriched cultures have shown rates up to 58 µmoles
Cl−/L/h (Nelson et al. 2011). In communities respiring high molecular weight
organohalides with low aqueous solubility (e.g., polychlorinated dioxins, polychlorinated furans, PCBs), organohalide respiration rates are more likely to be
limited by mass transfer rates than for higher solubility organohalide compounds.
Cultures respiring trichlorodibenzo-p-dioxin were reported to have rates of 0.22
µmole Cl−/L/h (Ballerstedt et al. 2004) and culture GY respired PBDE with rates
near 0.006 µmole Cl−/L/h (Lee et al. 2011b). Overall, reported rates range by five
orders of magnitude across the different organohalide classes.
Effective models for community behavior require accounting for important
physiological groups that impact OHRB activity. The production and consumption of H2 and, to a lesser extent, acetate are central processes in these OHRB
communities. Several models based on Monod kinetics exist for chloroethenerespiring OHRB populations growing in mixed cultures (Heavner 2013; Yu and
Semprini 2004; Yu et al. 2005; Haest et al. 2010a, b; Baelum et al. 2013; Becker
2006; Becker and Seagren 2009; Lai and Becker 2013; Fennell and Gossett 1998;
Chen et al. 2013; Schneidewind et al. 2014; Maphosa et al. 2010). Chambon et al.
(2013) summarizes models of organohalide respiration in a recent review. Many of
these existing modeling efforts consider the activity of only the OHRB. The concentrations of H2 and acetate are usually assumed to be constant (Schneidewind
et al. 2014). A few models have also biokinetically modeled the activities of nonOHRB community members including the effects of fermenters and methanogens
upon H2 and acetate availability (Fennell and Gossett 1998; Heavner et al. 2013).
The use of DNA, RNA, and protein biomarkers to quantify different populations
and their key enzymes is allowing development and validation of community models in ways not previously possible and should improve predictive models in the
14.7 Community Structure in Contaminated Aquifer/
As discussed in Chap. 22, successful bioremediation at contaminated sites hinges
upon OHRB activities being realized in situ. In a remediation context, enhanced
bioremediation efforts historically involved adding electron donor in excess of
what would be needed to completely reduce site organohalide concentrations—to
ensure complete dechlorination. Electron donor additions stimulate many microbes
in addition to OHRB—including fermenters, methanogens, and others physiological groups supported by site geochemistry (e.g., sulfate reducers, iron reducers).
At natural attenuation sites, electron donor may be limiting and may be derived
from hydrocarbon co-contaminants at the site (e.g., petroleum products) or decay
of endogenous biomass. Compared to their relatively high abundance in laboratory
cultures, OHRB are a much smaller percentage of the biomass at contaminated
field sites. This is true even in dense nonaqueous phase liquid (DNAPL) source
zones where organohalide levels are high and/or when OHRB cultures are bioaugmented into the aquifer (commonly at 100–10,000 fold dilutions of inoculum into
aquifer porewater) (Major et al. 2002; Baelum et al. 2013; Tas et al. 2009; Ellis
et al. 2000; Lendvay et al. 2003).
Several surveys of in situ biomass have been conducted from materials
retrieved from field sites undergoing enhanced bioremediation (e.g., biostimulation alone or combined with bioaugmentation) or natural attenuation. Many studies have assayed biomass specifically for known OHRB via methods targeting
their 16S rRNA genes and/or their RDase genes. The assays include qualitative/
semiquantitative methods (e.g., PCR amplification and clone library analysis)
as well as quantitative assays by qPCR and/or competitive PCR (Matturro et al.
2013; Hatt and Loffler 2012; Kjellerup et al. 2008; Hendrickson et al. 2002; Pérezde-Mora et al. 2014; Lu et al. 2006; Rahm et al. 2006; Lee et al. 2008). In general,
these studies show that native OHRB are often present in situ, can be biostimulated by electron donor additions, and can even persist following bioaugmentation
with other OHRB cultures (Pérez-de-Mora et al. 2014; Baelum et al. 2014). Other
researchers have utilized the benefits of degenerate primers for either ribosomal
RNA genes of the Dhc-like group of the Chloroflexi (Lowe et al. 2002) or RDase
genes (Tas et al. 2009, 2011; Dugat-Bony et al. 2012; Lee et al. 2008). DugatBony et al. (2012) have introduced a DechlorArray microarray with probes to
92 distinct RDase genes covering a range of available RDase diversity. Using this
array on groundwater biomass from four chlorinated ethene-impacted sites undergoing biostimulation, the authors found that there was great temporal and spatial
variability of diverse OHRB genera. The VC RDases, bvcA, and vcrA were below
detection at a location stalled at cis-DCE but present where dechlorination past
cis-DCE occurred. Tas et al. (2009) examined biomass from a PCE-contaminated
aquifer using functional gene arrays including probes for 153 RDase genes as well
as probes for many other functional genes. They found diverse RDase genes of the
14 Organohalide-Respiring Bacteria as Members of Microbial …
Dhc type in the DNA pool and they also witnessed high variability across time and
General community structures at contaminated sites have been examined using
methods such as high-throughput sequencing of 16S rRNA gene amplicons (Kotik
et al. 2013; Lee et al. 2011a), phylogenetic microarrays (Conrad et al. 2010; Lee
et al. 2012c; Nemir et al. 2010), and 16S rRNA gene clone libraries (Rahm et al.
2006; Dojka et al. 1998; Bowman et al. 2006; Macbeth et al. 2004; Lee et al.
2012c). Macbeth et al. (2004) surveyed biomass from a TCE-contaminated deepfractured basalt aquifer that had been biostimulated. The Firmicute Acetobacterium
was the most abundant sequence in the small clone library (32/93 clones in bacterial library). Other phylogenetic groups present included Sphingobacteria,
Bacteroides, Spirochaetes, Mollicutes, Proteobacteria, and candidate divisions
OP11 and OP3. Bowman et al. (2006) studied acidic groundwater near a DNAPL
source zone (mostly chlorinated ethanes) using both general and Dhc-specific 16S
rRNA gene-targeted primers to establish clone libraries. In the library established
with general 16S rRNA gene primers, Firmicutes sequences were dominant (62 %
of the library) and no Dhc sequences were detected. However, Dhc-specific PCR
did confirm presence of Dhc populations. Rahm et al. (2006) had similar observations at the Seal Beach site—that Dhc were not observed in clone libraries
created with degenerate primers, but direct PCR with Dhc primers was positive.
Direct comparisons between results from clone libraries and phylogenetic microarrays found that clone libraries would need to be very large to enable detection
of Dhc at their low in situ populations (Lee et al. 2012c). It is notable that recommended densities for in situ bioremediation are above 1E4/mL (1E7/L) (Lu et al.
2006). This may be a very small fraction of the community overall and detection
limits may be high in community survey methods (e.g., clone libraries, microarrays, and high-throughput sequencing of 16S rRNA gene amplicons). However,
some studies have found OHRB in clone libraries created using bacterial 16S
rRNA gene-targeted primers or general 16S rRNA gene-targeted primers (Dojka
et al. 1998; Lowe et al. 2002). Lowe et al. (2002) found obligate OHRB 16S rRNA
gene sequences in clone libraries established with DNA obtained from locations
impacted with chlorinated ethanes but not from pristine locations—specifically
Dehalobacter, not Dehalococcoidales.
Remarkable advances in sequencing are enabling much more thorough investigations of microbial community changes at field sites using PCR amplicon
sequencing and even metagenome sequencing. Such studies are improving resolution of community structure. With increased numbers of samples, and appropriate metadata, microbial responses to contamination and remediation practices
can now be traced. Studies utilizing phylogenetic microarrays or 454 pyrosequencing on chlorinated ethene plume biomass agreed in observing a wide range
of physiological types in the plume communities—with obligately anaerobic
and aerobic organisms both present in samples (Conrad et al. 2010; Kotik et al.
2013). Conrad et al. (2010) performed a transect along a two-kilometer-long
plume and found that methane produced in the active biostimulation area (likely
by Methanosarcinaceae populations) was stimulating methanotrophs (particularly
Methylosinus) further down the plume. These methanotrophs can degrade both
methane and chlorinated ethenes (Group 4 in Fig. 14.1), the latter via cometabolic
reactions catalyzed by their methane monooxygenase. In work by Kotik et al.
(2013), bacterial rRNA gene amplicons were analyzed by 454 pyrosequencing.
Though Albidiferax (beta Proteobacteria) were predominant in all locations, the
two most highly contaminated sites did have several putative facultative OHRB
genera (Anaeromyxobacter, Desulfuromonas, Desulfovibrio, and Geobacter),
though it is not known if the strains contained genes for organohalide respiration.
These populations may be respiring with sulfate or iron and some can grow fermentatively. Even though the deep sequencing method had a detection level of
0.25 % of total amplicons, no obligate OHRB sequences were detected. Instead,
the community structure suggested that cis-DCE and VC may be acted upon
by the aerobic chlorinated ethene degraders detected by their analyses. These
included Polaromonas, the only known aerobe that grows with cis-DCE as its
sole carbon and energy source. They also found possible aerobic cometabolizers
of chlorinated ethenes (Burkholderia and Methylobacter). Lee et al. (2012c) used
phylogenetic microarrays to track groundwater communities during biostimulation and bioaugmentation for TCE bioremediation. Archaea grew over time—consistent with a two order of magnitude increase in methane at the site. Dhc were
detected by their microarray method.
It is clear that biostimulation efforts at contaminated groundwater sites stimulate a wide range of microbial populations directly and indirectly—including
various anaerobes (e.g., OHRB, methanogens, fermenters) and aerobes (methanotrophs, aerobic organohalide-mineralizing populations, cometabolic organohalide
14.8 Roles of OHRB in Natural Systems
Though releases of pure organohalide compounds into the environment creates an
unnatural situation, organohalides in general are not solely anthropogenic—they
are produced naturally and can support native populations of OHRB (see Chap. 2).
Figure 14.1 includes several processes that are proposed to play important roles in
natural organohalide cycles.
The naturally occurring organohalides are diverse (Leri et al. 2007). Gribble
(2003) reported that more than 3800 individual organohalides are present in natural systems and most of these are chlorinated and brominated organohalides,
though fluorinated and iodinated organohalides do exist. A variety of studies have
clearly demonstrated the existence of natural organohalide pools in weathered
plant material, soil, and sediment (freshwater and marine) (Asplund and Grimvall
1991; Gribble 1994, 2003; Leri et al. 2007). Organohalides are also found associated with living animals (e.g., marine sponges produce brominated organics intentionally) (Ahn et al. 2003).
14 Organohalide-Respiring Bacteria as Members of Microbial …
With respect to the abundance of organohalides, in several studies of soils the
organic chloride pool was actually larger than the inorganic chloride pool (Oberg
et al. 2005; Rohlenova et al. 2009; Redon et al. 2011). Chlorinated organics were
shown to correlate strongly with the prevalence of organic matter content of
soils (Gustavsson et al. 2012). In humus layers, nearly 100 % of the soil chlorine
atoms were tied up as chlorinated organics (Redon et al. 2011). Absolute levels of
chlorinated organics across 51 soils in France ranged from 34 to 689 mg/kg soil
(Redon et al. 2011).
Biogeochemical cycling of natural organohalides involves both formation
and destruction. The OHRB likely play a major role in organohalide destruction.
Formation of organohalides occurs by both abiotic and biotic mechanisms. Several
recent studies have concluded that in organic soils, the biotic route predominates
(Aeppli et al. 2013; Bastviken et al. 2007, 2009; Clarke et al. 2009; Rohlenova
et al. 2009). Biological halogenation reactions can occur by different enzymatic classes including haloperoxidases and flavin dehydrogenases (Reaction 6
in Fig. 14.1) (Aeppli et al. 2013; Krzmarzick et al. 2012; Bengtson et al. 2009).
Leri et al. (2007) used microscopic techniques to visualize hotspots of chlorinated
organics and colocalization of fungi suggests that they play a substantial role in
chlorinated organics formation. The reasons for organohalide production by biota
(fungi and bacteria and animals) include antagonism, defense (against predation),
and signaling (Bengtson et al. 2009; Clarke et al. 2009).
The production of less halogenated and nonhalogenated organic matter by
OHRB detoxifies antagonistic compounds and also provides cross-feeding opportunities. The cross-feeding takes two forms in native soils/sediments. As discussed
earlier (and in Chap. 13), successive reductive dehalogenation can set up an OHRB
food web. Additionally, these partially dehalogenated organohalides can be oxidized by organisms utilizing alternative electron acceptors such as O2, iron, sulfate,
and nitrate (Groups labeled “4” and “5” in Fig. 14.1). The aerobes include those
that cometabolically degrade organohalides by methane-, toluene-, and alkeneoxygenase enzyme systems (Arp et al. 2001; Mattes et al. 2010) as well as those
that can use partially dehalogenated organics as sole carbon and energy sources
such as specific Polaromonas and Nocardiodes strains (Coleman et al. 2002a, b;
Jennings et al. 2009). Less chlorinated ethenes and aromatics can drive aerobic
catabolism even at extremely low O2 levels (Gossett 2010). The chloride released
by OHRB can be recycled back to be used by halogenating enzymes (Reaction 6 in
Fig. 14.1) (Aeppli et al. 2013; Bengtson et al. 2009; Krzmarzick et al. 2012).
The increasing speed and power of phylogenetic and metagenomic profiling methods is enabling studies that shed light on the ecology of OHRB communities from
simple two-member cocultures to highly diverse communities. We see trends in
the reactions mediated by non-OHRB populations including the provision of
cofactors, carbon, and reducing power (especially H2) to OHRB. Many OHRB
(especially the obligate OHRB) utilize H2 and acetate in the presence of suitable
organohalide electron acceptors. In OHRB communities, the OHRB populations
often compete with methanogens for H2 and acetate. In fact, many OHRB communities and their food webs resemble anaerobic digester communities in their
non-OHRB populations, although clustering of metagenomic sequencing raw
reads did suggest distinct composition among the OHRB communities versus
anaerobic methanogenic digester communities (Hug et al. 2012). Euryarcheaota,
Proteobacteria, Bacteroidetes, Firmicutes, and Spirocheata are the most commonly
found non-OHRB groups in organohalide-respiring communities. Another common observation is that multiple OHRB usually coexist in enrichment cultures
even after decades of enrichment—presumably due to cross-feeding or niche specialization. In pristine, carbon-rich settings, a natural organohalide cycle exists
where organohalides are produced biotically for signaling, defense, and antagonism purposes. The various organohalide molecules are dehalogenated by OHRB
and the organic products of organohalide respiration support respirers of other terminal electron acceptors (e.g., oxygen, iron and sulfate). Organisms that use these
alternate electron acceptors can also directly compete with OHRB for H2 and acetate. Significant mysteries remain, such as the full suite of cofactors, nutrients, and
signaling molecules transferred among different community members as well as
the ecological reason for the general lack of full dehalogenation capabilities on
individual OHRB genomes. Further meta-omic studies will help find answers to
these remaining questions.
Acknowledgments The author acknowledges the collective work of many researchers
worldwide and the public and private funding agencies that have made this area of research
possible. Thanks to Elizabeth Edwards and Cresten Mansfeldt for input on the chapter content.
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