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6 Organohalide Respiration Rates in OHRB Communities and Modeling of Community Food Webs

6 Organohalide Respiration Rates in OHRB Communities and Modeling of Community Food Webs

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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

near future.



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14.7 Community Structure in Contaminated Aquifer/

Sediment Environments

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



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Dhc type in the DNA pool and they also witnessed high variability across time and

location.

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



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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

degraders).



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).



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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).



14.9 Summary

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



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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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