microbial dechlorination of dioxins in estuarine ... · bated with pcbs, dechlorination ensued...

MARINE

ENVIRONMENTAL

www.elsevier.com/locate/marenvrev

Marine Environmental Research 59 (2005) 177–195RESEARCH

Microbial dechlorination of dioxins in estuarineenrichment cultures: effects of respiratory

conditions and priming compound on communitystructure and dechlorination patterns

Q. Shiang Fu a,*, Andrei L. Barkovskii b, Peter Adriaens c

a Environmental Engineering and Science, Department of Civil and Environmental Engineering,

Stanford University, Stanford, CA 94305-4020, USAb Department of Biological and Environmental Sciences, Georgia College and State University,

Milledgeville, GA 31061-0490, USAc Environmental and Water Resources Engineering, Department of Civil and Environmental Engineering,

The University of Michigan, Ann Arbor, MI 48109-2125, USA

Received in revised form 22 January 2004; accepted 6 April 2004

Abstract

The effect of respiratory conditions and priming compound on dechlorination patterns of

heptachlorodibenzo-p-dioxins (HpCDD) was investigated using estuarine sediment-eluted cul-

tures in the presence and absence of 20 mM sulfate, and 0.2 lM 2-bromodibenzo-p-dioxin (2-

BrDD) as a priming compound. Electron balance calculations based on fatty acid turnover,

hydrogen production, and electron acceptor depletion/methane formation indicated that

whereas fermentative processes dominated in sulfate-free incubations, sulfate-reduction was

predominant in the sulfate-amended incubations. The dechlorination of 1,2,3,4,6,7,8-HpCDD

exhibited the following trends: (i) the relative yields of product formation did not exceed 30%and

the presence of 2-BrDD increased the yield by up to 10%; (ii) sulfidogenic conditions resulted in a

lower 2,3,7,8-tetrachlorodibenzo-p-dioxin (2,3,7,8-TCDD) formation, and the presence of 2-

BrDD decreased the formation of 2,3,7,8-TCDD by additional 4–5-fold; (iii) the presence of

2-BrDD effected a predominance in lateral (2,3,7,8 positions) over peri (1,4,6,9 positions)-

dechlorination. Denaturing gradient gel electrophoresis (DGGE) banding patterns indicated

significant shifts of microbial community structure in response to terminal electron accepting

*Corresponding author. Tel.: +1-650-724-5310; fax: +1-650-723-7058.

E-mail address: [email protected] (Q. Shiang Fu).

0141-1136/$ - see front matter � 2004 Elsevier Ltd. All rights reserved.

doi:10.1016/j.marenvres.2004.04.003

mail to: [email protected]

178 Q.Shiang Fu et al. / Marine Environmental Research 59 (2005) 177–195

processes as well as to the presence of the priming compound. The latter resulted in a similar

community structure independent of dioxin spike, indicating that subsets of populations in the

sediment are capable of exploiting the new niche provided by the priming compound.

� 2004 Elsevier Ltd. All rights reserved.

Keywords: Chlorinated dioxins; Reductive dechlorination; Community structure; Degradation; Sediment

pollution; Microorganisms

1. Introduction

Fate and transformation of polychlorinated dibenzo-p-dioxins (PCDD) in the

environment are of particular importance due to their high toxicity, bioaccumula-

tion, and suspected carcinogenicity. Freshwater, estuarine, and marine sediments

serve as the primary sinks for this group of compounds. Chlorinated dioxins, es-

pecially 2,3,7,8-substituted polychlorinated dibenzo-p-dioxins, have been shown to

undergo reductive dechlorination reactions under reducing conditions (Adriaens,Fu, & Grbic’-Grlic’, 1995; Adriaens, Chang, & Barkovskii, 1996; Albrecht, Bar-

kovskii, & Adriaens, 1999; Barkovskii & Adriaens, 1996, 1998; Beurskens et al.,

1995; Bunge, Ballerstedt, & Lechner, 1999; Fu, Barkovskii, & Adriaens, 1999, 2001).

Using cultures eluted from Passaic River sediments, microbial dechlorination has

been demonstrated under freshwater and estuarine conditions (Barkovskii &

Adriaens, 1996; Fu et al., 2001). Lateral dechlorination patterns (chlorine removal at

2,3,7,8 positions) were found to be dominant under freshwater conditions.

Little is known about the microbiology of PCDD dechlorination, through severallines of evidence indicate that non-methanogenic subpopulations may be responsible

for reductive dechlorination under freshwater conditions (Barkovskii & Adriaens,

1996). Microbial PCDD dechlorination under estuarine conditions is less clear.

Recently, we documented microbial PCDD dechlorination under high salinity and

sulfate conditions by estuarine microbial consortia from Passaic River sediments (Fu

et al., 2001). However, nothing is known about the impact of PCDD on microbial

community structure and the relationships between microbial community compo-

sition and dechlorination patterns in estuarine and marine environments. Numerousinvestigations have demonstrated that the presence of organic compounds or con-

taminants affects the diversity and structure of microbial communities (Abed et al.,

2002; Colores, Macur, Ward, & Inskeep, 2000). Recent insights in the microbial

ecology of polychlorinated biphenyl (PCB) dechlorination have indicated that ortho-

or meta/para-dechlorination activity can be enriched (Cutter, Sowers, & May, 1998;

Pulliam-Holoman, Elberson, Cutter, May, & Sowers, 1998). Furthermore, com-

munity analysis using genetic methods shows that the community structure associ-

ated with ortho-dechlorination patterns differs markedly from that of cultures notspiked with PCBs (Prieur et al., 1987).

Polychlorinated biphenyl studies also revealed that various compounds structurally

related to PCBs exhibit a ‘priming’ effect towards PCB dechlorination. The actual role

of the priming compounds is not trivial. In particular, it is unknown whether such

Q.Shiang Fu et al. / Marine Environmental Research 59 (2005) 177–195 179

primers cause increased production of proteins involved in reductive dehalogenation

(e.g., components of electron transport chains) or the priming effect is the result of the

emergence of new populations in response to selective pressure by the priming agents.

The natural production of halogenated compounds in marine environments

(Gribble, 1994) has prompted investigations on the potential of these compounds to

induce or stimulate microbial degradation activities of (structurally analogous) an-thropogenic compounds (Allard, Hynning, Remherger, & Nielson, 1992; King,

1988). Dechlorination of historical PCB contamination in freshwater sediments has

been stimulated using both chloro- and bromobiphenyls (Bedard & van Dort, 1997;

Bedard, van Dort, & DeWeerd, 1998; DeWeerd & Bedard, 1993), and bro-

mobenzoates (DeWeerd & Bedard, 1993). More recently, it was demonstrated that

dioxin-contaminated sediments amended with 2-bromodibenzo-p-dioxin (2-BrDD)

exhibited extensive dechlorination activity, resulting in the production of 2-chloro-

dibenzo-p-dioxin (2-monoCDD) (Albrecht et al., 1999).In estuarine environments, microbial diversity and metabolic activity are mainly

affected by salinity gradients, high sulfate concentrations, and competition for en-

ergy sources. Sulfate reduction has been reported to be the dominant microbial re-

spiratory process in estuarine and marine environments (Capone & Kiene, 1988).

Whereas earlier studies suggested that methanogenesis and sulfate reduction were

spatially separated or that methanogenesis only became significant after sulfate was

depleted, the coexistence of sulfate reduction and methanogenesis has been shown in

marine sediments that exhibit sulfate reduction as the dominant respiration (Kiene &Capone, 1994).

Previously, freshwater sediments were shown to dechlorinate PCBs under meth-

anogenic, but not under sulfate-reducing conditions (Alder, H€aggblom, Openheimer,

& Young, 1993). However, when estuarine sulfate-reducing sediments were incu-

bated with PCBs, dechlorination ensued (Øfjord, Puhakka, & Ferguson, 1994). This

suggests that freshwater and estuarine environments select for different populations,

which exert differences in PCB dechlorination activity (Bedard & Quensen, 1995).

One of the most challenging aspects of microbial ecology remains the demon-stration of cause–effect relationships between the effect of external amendments on

community structure and activity endpoints. The goals of this study are to deter-

mine: (i) whether there are structural and physiological changes related to the

presence of PCDDs and their structural analogues (brominated dioxins), (ii) whether

the latter can prime PCDD dechlorination under estuarine and marine-mimicking

conditions, and (iii) whether the diversity in structural and physiological composi-

tion affects PCDD dechlorination.

2. Materials and methods

2.1. Inoculum

The sediment core was collected from the lower Passaic River (NJ) near the

outfall of 24 Lister Ave., one of the presumed sources of the dioxin contamination in


this estuary (Albrecht et al., 1999; Bopp et al., 1991). A 30-cm core section derived

from approximately 1 m below the riverbed was used to obtain the microbial frac-

tion. Estuarine bottom water was collected at the same time from the same location.

The salinity of the water column 1 m above the sediment ranged from 16‰ to 22 ‰.

2.2. Experimental design

Estuarine microbial consortia were prepared under strictly anaerobic conditionsin a glove box (N2–CO2–H2, 85:10:5) (Coy Laboratory Products, Inc., Ann Arbor,

MI). Two estuarine media for the experiment included modified basal medium

(Sowers, Johnson, & Ferry, 1984) (in g/L): Na2CO3, 3.0; Na2HPO4 � 7H2O, 1.12;

NH4Cl, 0.5; cysteine-HCl �H2O, 0.25; and resazurine, 0.001. Trace element and vi-

tamin solutions (1% each) were supplemented to the media (Wolin, Wolin, & Wolfe,

1963). Estuarine medium with sulfate (E-Sulfate medium) (Berkaw, Sowers, & May,

1996) contained (g/L): NaCl, 8.4; MgSO4 � 7H2O, 4.78; KCl, 0.27; and CaCl2 � 2H2O,

0.05. The final sulfate concentration in the E medium was 19.4 mM. Estuarinemedium without sulfate (E medium) was similar to E-Sulfate medium with the ex-

ception that MgSO4 was replaced by MgCl2 (3.9 g/L). All of the components except

for Na2CO3 and the vitamins were subjected to a stream of N2–CO2 (75:25). The

vitamin solution was filter-sterilized. The pH of the media was about 6.8. The salinity

in the two estuarine media was 17‰.

Wet sediment (1.2 kg) was transferred from the sediment core into glass jars in the

glove box and amended with 2.4 L of the Passaic River bottom water and 100 mg/L

(final concentration) of a primary growth substrate cocktail (acetic acid, 75 mg/L;butyric acid, 15 mg/L; and benzoic acid 10 mg/L). The jar was covered with alu-

minum foil and the sediment suspension was incubated in the anaerobic chamber for

6 months at room temperature in the dark and was shaken daily for 1 min. After 6

months, a Tween 80 solution (0.1% v/v) was added to the sediment suspension,

which was then transferred into 800-ml centrifuge tubes, capped with a butyl rubber

stopper, and sonicated for 5 min. The suspension was then centrifuged (500 rpm; 5

min) to separate solid particles, and the supernatant containing cells and colloidal

material was harvested. The supernatant was centrifuged (3700 rpm; 15 min) topellet the cells, which were resuspended separately in E and E-Sulfate medium (600

ml), and amended with 100 mg/L (final) of substrate cocktail. These cell suspensions

were incubated for another 6 months to enrich communities in the presence and

absence of sulfate and to increase biomass densities, centrifuged (3700 rpm; 20 min),

and left overnight for sedimentation. The pelleted cells were resuspended separately

with E and E-Sulfate media (700 ml each) and spiked with substrate cocktail (100

mg/L). These enriched microbial consortia were used as the inoculum for subsequent

incubations. A 5-ml aliquot of the enrichment cultures was transferred into 20-mlsterilized serum bottles sealed with Teflon lined butyl rubber stoppers (The West Co.,

Lionville, PA).

The cell suspensions were set up in five replicates in addition to background (no

spike of dioxin congeners) and sterile controls (autoclaved for 2 h on two consecutive

days prior to the addition of dioxin congeners). One set of active cultures was spiked


separately with 0.47� 0.01 lM of either 1,2,3,4,6,7,8- or 1,2,3,4,6,7,9-HpCDD

(AccuStandard Inc., New Haven CT) from a 90 lM stock solution in decane. An-

other set of cultures was spiked with the HpCDD congeners and 2-bromodibenzo-p-

dioxin (2-BrDD) (0.19 lM) (Dow Chemical, Midland, MI). Duplicate incubations

were used for analysis of methane, hydrogen, sulfate, protein, and molecular (16S

rRNA) characterizations. All cultures (a total of 240) were incubated with dailymanual shaking (twice a day and 2 min each time) at 30 �C in the dark for a year.

Substrate cocktail (100 mg/L, final concentration) was added every two months.

2.3. Sample preparation and PCDD analysis

Triplicate microcosms were sacrificed for congener-specific dioxin (mono-

through heptaCDD) analysis. Octachlorodibenzofuran (100 ll of a 5 ppm stock) was

used as the internal standard to monitor extraction efficiency. A NaOH solution

(2 ml, 5 M) was added to the cell suspensions to break up bacterial cell walls. Manual

triplicate extractions with toluene (4 ml) were performed and the extracts werepooled and dried over a gentle stream of nitrogen gas to approximately 2 ml. Ele-

mental sulfur was removed from E-Sulfate medium samples based on a modified

procedure with tetrabutylammonium sulfate and sodium sulfite (Jensen, Renberg, &

Reutergardh, 1977). The treated samples were dried again with nitrogen to reduce

the amount of solvent to about 2 ml, and approximately 0.5 g of anhydrous sodium

sulfate was added to remove residual water from the organic phase. Sample cleanup

was performed in a micro Florisil column (�0.2 g in a disposable Pasteur pipette

with glass-wool at the bottom) eluted with 10 ml of hexane. After evaporation ofhexane, 100 ll of toluene was added to the sample to concentrate the dioxins prior to

analysis.

Mono- to heptaCDD congeners were analyzed on a 5890-II Hewlett–Packard GC

equipped with a 5972A Mass Selective Detector (MSD) in selective ion monitoring

mode (SIM) as described previously (Albrecht et al., 1999). Recovery efficiencies for

PCDDs were 85–100% based on the internal standard used. Quantitation was based

on a six-level calibration curve established for a custom-made mixture of dioxin

congeners. The detection limit for native dioxins was 0.1 ng on-column. Identifica-tion of congeners was based on the comparison of the retention times of known

congeners and the ratios of relative abundance of key ion clusters in the mass

spectrum corresponding to the values calculated with the natural isotopic abundance

of chlorines (EPA, 1986).

2.4. Hydrogen analysis

A 0.4-ml aliquot of the gas sub-sample was taken from microcosms with aHamilton (Reno, NV) gas-tight sample-lock syringe, and the Teflon valve was closed

until analysis. Hydrogen gas was analyzed using a Trace Analytical (Menlo Park,

CA) reduction gas analyzer equipped with Carbosorb and molecular sieve 5A col-

umns in series and quantified against a 10-point calibration curve. The detection

limit of this analysis was 20 ppb (v/v). The gas concentration was then converted to


an equilibrium liquid phase concentration using the Oswald phase partitioning

constant for hydrogen of 0.08211 L atm./Kmole (Lovley, Chapelle, & Woodward,

1994).

2.5. Sulfate analysis

Sulfate in filtered culture medium was analyzed using a Dionex 4500i ion chro-

matograph (IC) (Sunnyvale, CA) equipped with an IonPac AS4A column (4� 250mm) (Dionex, Sunnyvale, CA), and a conductivity detector with an eluent consisting

of both sodium bicarbonate (1.7 mM) and sodium carbonate (1.8 mM) pumped at 2

ml/min. Ions were identified and quantified by comparing retention times and peak

areas, respectively, with those of external standards. External calibration was based

on a 10-point standard curve (0.1–10.0 mM, R2 ¼ 0:99). Time zero microcosms were

diluted 2-fold prior to analysis.

2.6. Methane analysis

Headspace gases were collected from GC vials via purge with helium carrier gas.

The subsample was injected onto a 30 m DB-624 column through a split/splitless

port operating at 140 �C in the splitless mode. Methane was detected within 4 min by

Flame Ionization Detector (FID). A 4-point calibration was used for quantitation.

Standards were prepared by taking 1 ml methane from a sealed gas-tight container

and diluted into a 160-ml sealed vial: 0.25, 0.5, 0.75, and 1 ml subsamples were in-

jected into four 10-ml headspace GC vials.

2.7. Organic acid analysis

HPLC analysis was performed on a Hewlett–Packard Series II 1090 Liquid

Chromatograph equipped with an HP Chemstation and diode array detector. Pri-

mary substrate cocktail mixtures, benzoic, butyric, and acetic acids were analyzed on

a Spherisorb C8 column (250 mm� 4.6 mm� 5 lm) (Alltech, Deerfield, IL). The

mobile phase consisted of 90% 50 mM KH2PO4 (pH was adjusted to 2.5 withH3PO4) and 10% CH3CN. The 50 mM KH2PO4 solution was prepared with Milli-Q

water and filtered with 0.2-lm filter using a Masterflex pump. Identification and

quantification was performed by retention time and peak area of external standards.

External calibration was based on a 9-point standard curve for each compound

(0.10–10.0 mM, R2 ¼ 0:99).

2.8. Protein analysis

Bacterial growth was determined by measuring the amount of protein produced in

the replicate experimental cultures, based on 1-ml aliquots. A 10-ml culture tube

containing the cells and 0.4 ml of 5 N NaOH (to break up cell membranes) was

heated for 10 min at 98 �C with a dry bath incubator (Fisher Scientific). The test

tubes were then centrifuged at 13,000 rpm for 15 min with a Tomy MC-140 high-


speed micro centrifuge (Tomy Tech USA, Inc., Palo Alto, CA). The protein con-

centration was measured by using the Bio-Rad Protein Assay Kit II (Bio-Rad

Laboratories, Hercules, CA) with bovine serum albumin as standard.

2.9. DNA extraction procedure

DNA was extracted from each active culture for 16S ribosomal DNA (rDNA)

analysis based on a reported procedure (Oelm€uller, Kr€ugger, Steinb€uchel, &Friedrich, 1990). The concentration and purity of the extracted DNA were deter-

mined by absorption spectrophotometry.

2.10. PCR amplification of rDNA fragments

The 16S rRNA was amplified by PCR with a Mastercycler personal� thermo-

cycler (Eppendorf Scientific, Inc. Westbury, NY). The universal primers were

f984GC (forward primer with a GC clamp) (50CGCCCGGGGCGCGCCCCGGG-CGGGGCGGGGGCACGGGGGGAACGCGAAGAACCTTAC30) and r1378

(reverse primer) (50CGGTGTGTACAAGGCCCGGGAACG30). These two primers

amplify the bacterial 16S rDNA fragment at positions 968–1401 (fragment 968–

1401; Escherichia coli numbering; Brosius, Dull, Sleeter, & Noller, 1981). The GC

clamp was attached to the forward primer 984 to prevent melting of the PCR

products during separation of DNA bands in the denaturing gradient gel electro-

phoresis (DGGE) (Myers, Maniatis, & Lerman, 1987). The PCR mixtures were

added to a 1.5-ml autoclaved Eppendorf tube as follows: 5 ll 10� PCR buffer (100mM Tris–HCl, 15 mM MgCl2, and 500 mM KCl, pH 8.3), 200 lmol of each de-

oxyribonucleoside triphosphate, 100 nmol of each primer, 2 ll of the extracted DNA

sample, and 40.5 ll of autoclaved ultra-filtered water (pH 7.0). The mixture was

centrifuged with a microcentrifuge (Micro12, Fisher Scientific) to pellet all compo-

nents. After initial denaturation at 94 �C for 5 min, Taq DNA polymerase of 0.5 ll(1.25 U) was added to each reaction tube without removing the tubes from the

thermocycler. This hot start was effective to prevent non-specific annealing of the

primers to non-target DNA molecules (Chou, Russel, Birch, Raymond, & Bloch,1992). The reaction tubes were centrifuged again before reaction cycles were

initiated.

Amplification was performed with 31 cycles of reaction: 1 min denaturing at

94 �C, 1 min annealing at 54 �C for primer annealing, 1 min at 72 �C for primer

extension, followed by final extension at 72 �C for 5 min, and cooling to 4 �C.Products of PCRs were first examined by running electrophoresis in 1% (w/v) aga-

rose gels and ethidium bromide staining (Sambrook, Fritsch, & Maniatis, 1989).

2.11. DGGE analysis

A D-Code 16/16-cm gel system with a 1.5-mm gel width (Bio-Rad, Hercules, CA)

was used to perform DGGE analysis. The DNA separation was maintained at a

constant temperature of 60 �C in 6 L of 0:5� TAE buffer (20 mM Tris acetate and


0.5 m MEDTA [pH 8.0]). Gradients were formed with 6% (w/v) acrylamide stock

solutions (acrylamide–N,N 0-dimethylene-bis-acrylamide, 37:1) which contained 15%

and 55% denaturant (with 100% denaturant defined as 7 M urea dissolved in 40% [v/

v] formamide). Gels were run at 35 V for 16 h. Gels were stained in Milli-Q water

containing ethidium bromide (0.5 mg/L) and destained twice in 0:5� TAE buffer for

15 min each. Images were obtained with the Alpha-Imager software (Alpha Inno-tech, San Leandro, CA). A tif-formatted image of the gel was loaded into Gel-

Compar� (Applied Maths, Kortrijk, Belgium) for comparative correlation analyses

of the lanes. The Pearson correlation algorithm in association with either Un-

weighted Pair Group Method with Arithmatic Mean (UPGMA) or Ward’s methods

was used to establish clustering patterns. Analysis was limited to the region of the gel

containing bands (top region near wells were excluded). The dendrogram is from

clustering analysis with UPGMA using the ‘‘Fine Optimization’’ option of Gel-

Compar�. The observed pattern was detected with both UPGMA and Ward’sclustering methods using either the global optimization or fine optimization options

in GelCompar.

2.12. Extraction of DNA from DGGE gels and sequence analysis

The central 1-mm2 portions of strong DGGE bands were excised with a razor

blade and soaked in 50 ll of sterile Milli-Q water overnight. A 15 ll portion of the

solution was reamplified by PCR as described in the PCR procedure. The reamplifiedPCR products were purified by electrophoresis though a 1.2% agarose gel in 0:5�TAE running buffer followed by glass-milk extraction (Gene-Clean Kit; Bio 101).

Purified DNA was sequenced with an ABI-Prism model 373 automatic sequencer.

Sequence identification was performed by use of BLASTN facility of the National

Center for Biotechnology Information and the Sequence Match facility of the

Ribosomal Database Project (BLASTN, 1997; Maidak et al., 1999; Sequence Match,

1999). Not all bands produced high quality sequence and no further sequence

analysis was done on those sequences due to lack of PCR products.

3. Results

3.1. Impact of environmental conditions and dioxin isomers on microbial respiratory

activity

Respiratory activity was evaluated by means of sulfate, methane, and hydrogen

analysis. The data for sulfate consumption and methane production over the period

of incubation are presented in Table 1. It is clear that salinity and sulfate affect the

distribution of predominant respiratory activities. Methane production in the sul-

fate-free incubations accounted for the consumption of over one-third of theavailable reducing equivalents (primary substrates). The methane concentrations

were comparable in all incubations regardless of dioxin congener spiked and the

presence of 2-BrDD. Methane concentration in the sulfate-amended systems indi-


cated a low level of methanogenic activity. Less than 1% of the reducing equivalents

was converted via methanogenesis, most likely due to the presence of high sulfate

concentrations. Under these conditions, methane formation was not likely hydro-

genotrophic as sulfate reducers compete more efficiently for substrates such as hy-

drogen (Yang & McCarty, 1998). The sulfate consumption was nearly stoichiometric

based on the loss of the reducing equivalency, indicating that the predominantrespiration in the sulfate-amended incubations was sulfate reduction. Even

though sulfate consumption was not monitored during the course of the incubation,

residual sulfate (0.94–2.29 mM) was found in all active incubations spiked with di-

oxins in sulfate-amended microcosms at the end of the incubation. The presence of

brominated dioxin appeared to have no effect on either methane production or

sulfate consumption. Electron balance calculations based on turnover of primary

substrates, dissolved hydrogen concentration, and sulfate reduction/methane for-

mation indicated that whereas sulfate-free incubations were dominated by fermen-tative processes, sulfate-reduction was predominant in the sulfate-amended

incubations. Dissolved hydrogen concentrations were measured in order to deter-

mine their correlation with the dominant terminal electron accepting processes

(TEAP) and dechlorination patterns. Dissolved hydrogen concentrations were

measured for both sulfate-amended (in the range of 2.11–3.76 nM) and sulfate-free

incubations (in the range of 1.96–3.53 nM).

Table 1

Stoichiometry of sulfate reduction under sulfate-rich conditions; methane production under sulfate-rich

and sulfate-depleted conditions in estuarine medium (all values except ratios are in the unit of mM and

presented as the average of high and low values of duplicate samples; high and low values deviate in less

than 5% from the average)

Incubations Sulfate reduction Methane production

Sulfate amended Sulfate-amended Sulfate-free

Actual

loss

Predicted

lossa (% of

theoreticalb)

Productionc Predictedd

(% of

theoreticalb)

Production Predicted

(% of

theoreticalb)

Unspiked control 9.96 16.19 (61.5) 0.14 16.19 (0.87) 3.13 16.75 (19.5)

1234678-HpCDD 15.67 16.22 (96.6) 0.14 16.20 (0.86) 5.16 16.71 (32.0)

1234678-HpCDD+

2-BrDD

17.38 16.15 (107.6) 0.15 16.15 (0.93) 5.01 16.68 (31.1)

1234679-HpCDD 16.29 16.2 (100.5) 0.14 16.21 (0.85) 5.49 16.77 (34.1)

1234679-HpCDD+

2-BrDD

15.96 16.17 (98.7) 0.18 16.25 (1.10) 5.98 16.72 (37.2)

aCalculated based on the amount of primary substrates added and the equation (use acetate as an

example): CH3COO� +SO2�4 ¼ 2HCO�

3 +HS�.bRatio of actual loss over predicted loss expressed in percent.cMethane production was estimated by the subtracting of the time zero from the endpoint.dObtained by multiplying the number of moles of total loss of the reducing equivalent added by the

theoretical stoichiometric ratio of methane production without cell biomass production from the equation

(use acetate as example): CH3COO� +H2O¼CH4 + HCO�3 .


3.2. Impact of environmental conditions and dioxin compounds on microbial community

structure

The bacterial community structure of each incubation type was characterized and

compared by PCR-DGGE analysis (Fig. 1). Several features were observed from the

DGGE banding patterns: (i) the addition of the priming compound caused majorshifts in microbial composition in both sulfate-amended and sulfate-free incubations;

(ii) the presence of the priming compound resulted in a similar community structure

independent of the dioxin spiked, but dependent on the geochemical conditions.

The sulfate-free incubation without dioxins (unspiked control) showed two highly

visible bands (A-1 and A-2), indicating two specific populations. The derived se-

quences of the first band (A-1) indicated its close relation (98% match) to Pseudo-

monas stutzeri within the c subgroup of the proteobacteria. This band remained in

incubations with either heptaCDD isomers alone (lanes B and D), but almost en-tirely disappeared in the presence of 2-BrDD (lanes C and E). The second band (A-2)

was present in incubations spiked with either heptaCDD isomer and in the presence

of 2-BrDD. The sequence analysis revealed its close match (98%) in sequence to

Marinobacter sp., also within the c subgroup of the proteobacteria. Sulfate-free in-

cubations with 2-BrDD (lanes C and E) generated a novel unidentified band (C-1).

The unspiked control (lane F) in the sulfate-amended incubations showed a high

diversity in microbial community structure with several distinct populations. One

band (H-1) showed close resemblance (97% match) in sequence to Thiorhodovibrio

winogradskyi, a sulfur oxidizer within the c subgroup of the proteobacteria.

Cluster analysis of DGGE gel banding patterns demonstrated significant diversity

between microbial communities originating from sulfate-amended and sulfate-free

incubations as shown by two separate clusters in Fig. 2, with exception for the first

lane (represents lane G in Fig. 1). While the impact of PCDDs on community

structure under sulfate-amended conditions was not quite clear due to lane G as an

outlier in Fig. 1, under sulfate-free conditions both heptaCDD-spiked communities

clustered with the dioxin-free control, demonstrating the absence of selective pres-sure due to the presence of PCDDs. This is counter to observations with PCBs,

where the addition of 2,3,5,6-tetrachlorobiphenyl effected the selection of species

(e.g., Thermotogales, Dehalococcoides ethenogenes-similar organisms) that were ei-

ther absent or less predominant in the absence of the spiked congener (Pulliam-

Holoman et al., 1998).

3.3. Microbial dechlorination activity

The dechlorination patterns are similar between the two heptachlorodioxins

investigated. The dechlorination pattern interpretation was based on the dechlori-

nation of 1,2,3,4,6,7,8-HpCDD as it can potentially produce 2,3,7,8-tetrachlo-

rodibenzo-p-dioxin, which is arguably the most toxic chlorinated compound. Theeffect of 2-BrDD on dechlorination of 1,2,3,4,6,7,8-heptaCDD in sulfate-amended

and sulfate-free incubations was assessed in terms of both dechlorination yield and

homologue distribution (Figs. 3 and 4). Autoclaved controls did not yield lesser

Fig. 1. DGGE banding patterns of bacterial communities based on 16S rRNA sequences: (A) Unspiked

control /sulfate-free; (B) 1,2,3,4,6,7,9-HpCDD/sulfate-free; (C) 1,2,3,4,6,7,9-HpCDD+2-BrDD/sulfate-

free; (D) 1,2,3,4,6,7,8-HpCDD/sulfate-free; (E) 1,2,3,4,6,7,8-HpCDD+2-BrDD/sulfate-free; (F) Unspiked

control/sulfate-amended; (G) 1,2,3,4,6,7,9-HpCDD/sulfate-amended; (H) 1,2,3,4,6,7,9-HpCDD+2-

BrDD/sulfate-amended; (I) 1,2,3,4,6,7,8-HpCDD/sulfate-amended; and (J) 1,2,3,4,6,7,8-HpCDD+2-

BrDD/sulfate-amended.

Fig. 2. A dendrogram representation of a hierarchical cluster analysis (with percent of similarity) of the

DGGE banding patterns.


chlorinated products (data not shown). Not all the loss of the starting 1,2,3,4,6,7,8-

heptaCDD can be accounted for as only dechlorination products were monitored

and analyzed. In all active incubations, PCDD dechlorination has occurred. The


main effect of the presence of 2-BrDD is that the percentage of lateral reactions in

PCDD dechlorination increased under all the conditions studied, even though lat-

erally (2,3,7,8)-substituted PCDDs were formed, including 2,3,7,8-TCDD. The yields

(amount of products formed as a percent of total parent compound removed) of

2,3,7,8-substituted hexaCDDs among dechlorination products in 2-BrDD incuba-

tions represented 10% in the presence and 30% in the absence of sulfate (Figs. 3 and4), whereas in the absence of 2-BrDD these values were 21% and 40% for sulfate-

amended and sulfate-free incubations, respectively (data not shown). 1,2,3,7,8-Pen-

taCDD was also detected, but its level was below the quantitation limit. In the

presence of 2-BrDD, the yield of 2,3,7,8-TCDD resulting from dechlorination of

1,2,3,4,6,7,8-heptaCDD under both tested conditions was up to one order of mag-

nitude lower than in the absence of 2-BrDD (1% and 2% in the presence of 2-BrDD;

10% and 20% in the absence of 2-BrDD).

Under sulfate-free conditions, the presence of 2-BrDD also caused the greatestextent of the dechlorination resulting in the formation of monoCDDs (Figs. 3 and

4). A similar trend was reported earlier for PCDD dechlorination in Passaic River

sediments (Albrecht et al., 1999). Moreover, the presence of 2-BrDD caused a sig-

nificant increase in dechlorination yield. Dechlorination of 1,2,3,4,6,7,8-heptaCDD

yielded a total of 55 and 72 nM of lesser-chlorinated dioxin congeners in sulfate-

amended and sulfate-free incubations corresponding to a stoichiometric conversion

of, respectively, 26% and 32% of the 1,2,3,4,6,7,8-heptaCDD (data derived from the

difference in 1,2,3,4,6,7,8-heptaCDD concentration before and after the incubations,211 and 226 nM in sulfate-amended and sulfate-free conditions respectively). The

absolute yield of products increased nearly 2-fold due to the presence of 2-BrDD

(Fig. 5).

Hexa-through diCDD isomers were produced under both conditions, while

monoCDDs were the end products under sulfate-free conditions only, and only in

the presence of 2-BrDD (Fig. 4). Here, 1- and 2-monoCDD were formed in nearly

equimolar concentrations. Tri- and dichlorodibenzo-p-dioxin congeners were the

major congeners accounting for 28–41% of all dechlorination products except forsulfate-free incubations without 2-BrDD (Fig. 6).

4. Discussion

The presence of 2-BrDD caused the most distinctive shifts in microbial commu-

nity structure under both sulfate-amended and sulfate-free conditions, relative to the

control community. Moreover, DGGE profiles originating from 2-BrDD incuba-

tions exhibited highest structural similarity among themselves both in the presence or

the absence of sulfate (78% and 69%, respectively, Fig. 2). The distinctive patterns of

community structure in the presence of 2-BrDD indicated that subsets of popula-

tions in the estuarine enrichment cultures are capable of exploiting the new nicheprovided by this compound, even at the low concentrations provided (0.2 lM).

Bedard et al. (1998) observed that the addition of specific brominated PCBs primed

specific dechlorinating processes. It was shown that brominated congeners with a

Fig. 3. Dechlorination pathway of 1,2,3,4,6,7,8-heptaCDD in sulfate-amended incubations amended with

2-BrDD under estuarine conditions. The microcosms were incubated at room temperature for 12 months.


Fig. 4. Dechlorination pathway of 1,2,3,4,6,7,8-heptaCDD in sulfate-free incubations amended with 2-

BrDD under estuarine conditions. The microcosms were incubated at room temperature for 12 months.


meta bromine primed Dechlorination Process N (flanked meta dechlorination) and

congeners with an unflanked para bromine primed Dechlorination Process P

(flanked para dechlorination). The authors hypothesized that brominated analoguesprimed specific PCB-dechlorinating populations (Bedard et al., 1998). However, no

Fig. 5. Total concentration of lesser chlorinated dioxin congeners from dechlorination of 1,2,3,4,6,7,8-

heptaCDD (initial measured conc. 410 nM) under estuarine conditions in the presence (+) and absence ())of 2-BrDD. The microcosms were incubated at room temperature for 12 months. Autoclaved controls did

not yield lesser chlorinated products.


observations on microbial community shifts have been demonstrated following

amendments of PCB-dechlorinating systems with brominated analogues.

The result from our experiment clearly indicates that the changes are directly

attributable to 2-BrDD as evidenced by the new bands (e.g., band #1 in lane C forsulfate-free microcosms and band #2 in lane J for sulfate-amended microcosms). It is

unclear, however, whether those species associated with the new bands possess

specific dechlorinating activities towards either highly or lesser chlorinated dioxins.

In order to understand their actual role in PCDD dechlorination, those microor-

ganisms should be isolated and further studied, and the temporal changes in mi-

crobial community structure as related to the degree and extent of PCDD

dechlorination in the presence of 2-BrDD should also be followed.

Recently, Yang and McCarty (1998) reported on the use of different organic acids(which ferment to produce hydrogen) to stimulate reductive dechlorination in a

benzoate-acclimated dehalogenating mixed culture. Dissolved hydrogen concentra-

tions between 2 and 11 nM were found to be optimal for dechlorination reactions

and did not result in competition by methanogenic bacteria. Dissolved hydrogen

concentrations in the presence of 2-BrDD correlated with sulfate-reduction as the

dominant TEAP (Vroblesky & Chapelle, 1994). In the absence of sulfate, and a

predominance of methanogenic activity, H2 concentrations are expected to be in the

5–25 nM range, unless other coupling (e.g., halorespiring) processes are active. Sincemixed cultures were used in this study and no dechlorinating organisms were iden-

tified, the measured dissolved hydrogen concentration itself does not necessarily

indicate halorespiration as the respiratory process for the observed dechlorination

activity.

Fig. 6. Molar percent of lesser chlorinated dioxin congeners from dechlorination of 1,2,3,4,6,7,8-hepta-

CDD (initial measured conc. 410 nM) in the presence (+) and absence ()) of 2-BrDD under estuarine

conditions. The microcosms were incubated at room temperature for 12 months. Autoclaved controls did

not yield lesser chlorinated products.


No correlation between respiratory and dechlorination activity could be inferred

from the metabolic endpoints. The onset of dechlorination activity was unknown

since no kinetic analysis was performed. It has been reported that dechlorination of

PCBs started at a later stage in the presence of sulfate in estuarine media (Berkawet al., 1996) and that inhibition of sulfate-reduction negatively impacted PCB de-

chlorination (Cutter et al., 1998). Several observations can be made regarding PCDD

dechlorination as a function of the presence of 2-BrDD and incubation conditions:

(i) PCDD dechlorination occurred under sulfate-reducing conditions; (ii) the pres-

ence of 2-BrDD caused more extensive dechlorination; (iii) the homologue distri-

bution among dechlorination products was significantly affected by the presence of

2-BrDD; (iv) concentrations of lesser chlorinated dioxin congeners were higher

under sulfate-free conditions; and (v) monoCDDs were only produced in the pres-ence of 2-BrDD under sulfate-free conditions.

We conclude that in the presence of 2-BrDD, shifts among the patterns and the

extent of PCDD dechlorination correlate to changes in microbial community


structure as revealed by 16S rRNA analysis. Therefore, the presence of 2-BrDD

likely resulted in the priming of microbial populations with novel dechlorinating

capabilities and may indicate that PCDD dechlorination is not as fortuitous as

postulated earlier. Whether this indicates that populations with predominantly peri-

and lateral-dechlorinating capabilities can be stimulated in sediments remains to be

investigated. The indications of cause-and-effect relationships between PCDD de-chlorination activity and community composition may lead to further development

of predictive tools for natural and enhanced dechlorination of PCDD in estuarine

and marine environments.

Acknowledgements

Funding for this research was provided by the Office of Naval Research (ONR).

Since this paper has not been reviewed by this agency, no endorsement should be

inferred. We thank Upassri Sorachart for technical assistance. We are indebted to

Dr. Terence L. Marsh of the Center for Microbial Ecology and Department of

Microbiology at Michigan State University for performing cluster analysis of the

DGGE gel banding patterns.

References

Abed, R. M. M., Safi, N. M. D., K€oster, J., de Beer, D., El-Nahhal, Y., Rullk€otter, J., & Garcia-Pichel, F.

(2002). Microbial diversity of a heavily polluted microbial mat and its community changes following

degradation of petroleum compounds. Applied and Environmental Microbiology, 68, 1674–1683.

Adriaens, P., Fu, Q., & Grbic’-Grlic’, D. (1995). Bioavailability and transformation of highly chlorinated

dibenzo-p-dioxins and dibenzofurans in anaerobic soils and sediments. Environmental Science &

Technology, 29, 2252–2260.

Adriaens, P., Chang, P. R., & Barkovskii, A. L. (1996). Dechlorination of chlorinated PCDD/F by organic

and inorganic electron transfer molecules in reduced environments. Chemosphere, 32, 433–441.

Albrecht, I. D., Barkovskii, A. L., & Adriaens, P. (1999). Production and dechlorination of 2,3,7,8-

tetrachlorodibenzo-p-dioxin in historically-contaminated estuarine sediments. Environmental Science &

Technology, 33, 737–744.

Alder, A. C., H€aggblom, M. M., Openheimer, S. R., & Young, L. Y. (1993). Reductive dechlorination of

polychlorinated biphenyls in anaerobic sediments. Environmental Science & Technology, 27, 530–538.

Allard, A.-S., Hynning, P.-�A, Remherger, M., & Nielson, A. H. (1992). Role of sulfate concentration in

dechlorination of 3,4,5-trichlorocatechol by stable enrichment cultures grown with cumarin and

flavanone glycones and aglycones. Appl. Environ. Microbiol., 58, 961–968.

Barkovskii, A. L., & Adriaens, P. (1996). Microbial dechlorination of historically present and freshly

spiked chlorinated dioxins and diversity of dioxin-dechlorinating populations. Applied and Environ-

mental Microbiology, 62, 4556–4562.

Barkovskii, A. L., & Adriaens, P. (1998). Impact of model humic constituents on microbial reductive

dechlorination of polychlorinated dioxins. Environmental Toxicology and Chemistry, 17, 1013–1021.

Bedard, D. L., & Quensen, J. F., III (1995). Microbial reductive dechlorination of polychlorinated

biphenyls. In L. Y. Young & C. E. Cerniglia (Eds.), Microbial transformation and degradation of toxic

organic chemicals. New York, NY: Wiley-Liss.

Bedard, D. L., & van Dort, H. M. (1997). The role of microbial PCB dechlorination in natural restoration

and bioremediation. In G. S. Sayler, J. Sanseverino, & K. Davis (Eds.), Biotechnology in the sustainable

environment (pp. 65–71). New York, NY: Plenum.


Bedard, D. L., van Dort, H. M., & DeWeerd, K. A. (1998). Brominated biphenyls prime extensive

microbial reductive dehalogenation of Aroclor 1260 in Houstonic River Sediment. Applied and

Environmental Microbiology, 64, 1786–1795.

Berkaw, M., Sowers, K. R., & May, H. D. (1996). Anaerobic ortho dechlorination of polychlorinated

biphenyls by estuarine sediments from Baltimore Harbor. Applied and Environmental Microbiology, 62,

2534–2539.

Beurskens, J. E. M., Toussaint, M., de Wolfe, J., van der Steen, J. M. D., Slot, P. C., Commandeur, L. C.

M., & Parson, J. R. (1995). Dehalogenation of chlorinated dioxins by an anaerobic microbial

consortium from sediment. Environmental Toxicology and Chemistry, 14, 939–943.

BLASTN. (1997). Posting date. [Online] National Center for Biotechnology Information, Bethesda, MD.

Available: http://www.ncbi.nlm.nih.gov/BLAST.

Bopp, R. F., Gross, M. L., Tong, H., Simpson, H. J., Monson, S. J., Deck, B. L., & Moser, F. C. (1991). A

major incident of dioxin contamination: Sediments of New Jersey estuaries. Environmental Science &

Technology, 25, 951–956.

Brosius, J., Dull, T. J., Sleeter, D. D., & Noller, H. F. (1981). Gene organization and primary structure of

a ribosomal RNA operon from Escherichia coli. Journal of Molecular Biology, 148, 107–127.

Bunge, M., Ballerstedt, H., & Lechner, U. (1999). Different regioselectivities of reductive dioxin

dechlorination by anaerobic bacteria from river sediments of the Bitterfeld district (Germany).

Organohalogen Compounds, 41, 387–390.

Capone, D. G., & Kiene, R. P. (1988). Comparison of microbial dynamics in marine and freshwater

sediments: Contrasts in anaerobic carbon metabolism. Limnology and Oceanography, 33, 725–749.

Chou, Q., Russel, M., Birch, D., Raymond, J., & Bloch,W. (1992). Prevention of pre-PCRmis-priming and

primer dimerization improves low-copy-number amplifications. Nucleic Acids Research, 20, 1717–1723.

Colores, G. M., Macur, R. E., Ward, D. M., & Inskeep, W. P. (2000). Molecular analysis of surfactant-

driven microbial population shifts in hydrocarbon-contaminated soil. Applied and Environmental

Microbiology, 66, 2959–2964.

Cutter, L., Sowers, K. R., & May, H. R. (1998). Microbial dechlorination of 2,3,5,6-tetrachlorobiphenyl

under anaerobic conditions in the absence of soil or sediment. Applied and Environmental Microbiology,

64, 2966–2969.

DeWeerd, K. A., & Bedard, D. L. (1993). Q-1447 stimulation of anaerobic dechlorination of PCBs with

halogenated benzoic acids in Woods Pond sediment slurries. Abstr. Annu. Meet Am. Soc. Microbiol.,

Atlanta, GA.

Environmental Protection Agency (1986). Test methods for evaluating solid wastes, Vol. IB. Laboratory

manual physical/chemical methods (pp. 8280/1–8280/33). Washington, DC: Environmental Protection

Agency.

Fu, Q. S., Barkovskii, A. L., & Adriaens, P. (1999). Reductive transformation of dioxins: An assessment of

the contribution of dissolved organic matter to dechlorination reactions. Environmental Science &

Technology, 33, 3837–3842.

Fu, Q. S., Barkovskii, A. L., & Adriaens, P. (2001). Dioxin cycling in aquatic sediments: The Passaic River

estuary. Chemosphere, 43, 643–648.

Gribble, G. M. (1994). The natural production of chlorinated compounds. Environmental Science &

Technology, 28, 310A–319.

Jensen, S., Renberg, L., & Reutergardh, L. (1977). Residue analysis of sediment and sewage sludge for

organochlorines in the presence of elemental sulfur. Analytical Chemistry, 49, 316–318.

Kiene, R. P., & Capone, D. G. (1994). Effects of organic pollutants on methanogenesis, sulfate reduction

and carbon-dioxide evolution in salt-marsh sediments. Marine Environmental Research, 13, 141–160.

King, G. M. (1988). Dehalogenation in marine sediments containing natural sources of halophenols.

Applied and Environmental Microbiology, 54, 3079–3085.

Lovley, D. R., Chapelle, F. H., & Woodward, J. C. (1994). Use of dissolved H2 concentrations to

determine distribution of microbially catalyzed redox reactions in anoxic groundwater. Environmental

Science & Technology, 28, 1205–1210.

Maidak, B. L., Cole, J. R., Parker, C. T., Jr., Garrity, G. M., Larsen, N., Li, B., Lilburn, T. G.,

McCraughey, M. J., Olsen, G. J., Overbeek, R., Pramanik, S., Schmidt, T. M., Tiedje, J. M., & Woese,

http://www.ncbi.nlm.nih.gov/BLAST


C. R. (1999). A new version of the RDP (Ribosomal Database Project). Nucleic Acids Research, 27,

171–173.

Myers, R. M., Maniatis, T., & Lerman, L. S. (1987). Detection and localization of single base changes by

denaturing gradient gel elecrophoresis. Methods Enzymology, 155, 501–527.

Oelm€uller, U., Kr€ugger, N., Steinb€uchel, A., & Friedrich, C. G. (1990). Isolation of prokaryotic RNA and

detection of specific mRNA with biotinylated probes. Journal of Microbiological Methods, 11, 73–84.

Øfjord, G. D., Puhakka, J. A., & Ferguson, J. F. (1994). Reductive dechlorination of Aroclor 1254 by

marine sediment cultures. Environmental Science & Technology, 28, 2286–2294.

Prieur, D., Troussellier, M., Romana, A., Chamroux, S., Mevel, G., & Baleux, B. (1987). Evolution of

bacterial communities in the Gironde Estuary (France) according to a salinity gradient. Estuarine

Coastal and Shelf Science, 24, 95–108.

Pulliam-Holoman, T. R., Elberson, M. A., Cutter, L. A., May, H. D., & Sowers, K. R. (1998).

Characterization of a defined 2,3,5,6-tetrachlorobiphenyl-ortho-dechlorinating microbial community

by comparative sequence analysis of genes coding for 16S rRNA. Applied and Environmental

Microbiology, 64, 3359–3367.

Sambrook, J., Fritsch, E. F., & Maniatis, T. (1989). Molecular cloning: A laboratory manual (second ed.).

Cold Spring Harbor, New York: Cold Spring Harbor Laboratory Press.

Sequence Match (1999). posting date. [Online] Ribosomal Database Project, Michigan State University,

East Lansing, MI. Available: http://www.cme.msu.edu/RDP/analyses.html.

Sowers, K. R., Johnson, J. L., & Ferry, J. G. (1984). Phylogenetic-relationships among the methylotrophic

methane-producing bacteria and emendation of the family methanosarcinaceae. International Journal

of Systematic Bacteriology, 34, 444–450.

Vroblesky, D. A., & Chapelle, F. H. (1994). Temporal and spatial changes of terminal electron accepting

processes in a petroleum hydrocarbon-contaminated aquifer and the significance for contaminant

biodegradation. Water Resources Research, 30, 1561–1570.

Wolin, E. A., Wolin, M. J., & Wolfe, R. S. (1963). Formation of methane by bacterial extracts. Journal of

Biological Chemistry, 238, 2882–2886.

Yang, Y., & McCarty, P. L. (1998). Competition for hydrogen within a chlorinated solvent dehalogenating

anaerobic mixed culture. Environmental Science & Technology, 32, 3591–3597.

http://www.cme.msu.edu/RDP/analyses.html

microbial dechlorination of dioxins in estuarine ... · bated with pcbs, dechlorination ensued...

Documents