In cooperation with the

U.S. Army Garrison, Aberdeen Proving Ground

Environmental Conservation and Restoration Division

Aberdeen Proving Ground, Maryland

Anaerobic Degradation of 1,1,2,2-Tetrachloroethane and Association

with Microbial Communities in a Freshwater Tidal Wetland,

Aberdeen Proving Ground, Maryland: Laboratory Experiments and

Comparisons to Field Data

Water-Resources Investigations Report 02–4157

U.S. Department of the Interior

U.S. Geological Survey
U.S. Department of the Interior

U.S. Geological Survey

Anaerobic Degradation of 1,1,2,2-Tetrachloroethane and Association

with Microbial Communities in a Freshwater Tidal Wetland,

Aberdeen Proving Ground, Maryland: Laboratory Experiments and

Comparisons to Field Data

By Michelle M. Lorah, Mary A. Voytek, Julie D. Kirshtein, and Elizabeth J. (Phillips) Jones

Water-Resources Investigations Report 02–4157

In cooperation with the

U.S. Army Garrison, Aberdeen Proving Ground

Environmental Conservation and Restoration Division

Aberdeen Proving Ground, Maryland

The contents of this report have been approved for public release and unlimited distribution by the U.S. Army--

clearance number 4269-A-6

Baltimore, Maryland

2003
U.S. Department of the Interior

GALE A. NORTON, Secretary

U.S. Geological Survey

Charles G. Groat, Director

The use of trade, product, or firm names in this report is for 

descriptive purposes only and does not imply endorsement 

by the U.S. Government.

For additional information contact:

District Chief

U.S. Geological Survey

8987 Yellow Brick Road

Baltimore, MD 21237

Copies may be purchased from:

U.S. Geological Survey

Branch of Information Services

Box 25286

Denver, CO 80225-0286
v

CONTENTS

Abstract .................................................................................................................................................................................1

Introduction ...........................................................................................................................................................................2

Purpose and scope ........................................................................................................................................................6

Description of study area ..............................................................................................................................................6

Background on anaerobic degradation pathways .........................................................................................................8

Acknowledgments ......................................................................................................................................................10

Methods and data analysis ...................................................................................................................................................10

Laboratory microcosm experiments ...........................................................................................................................12

Sediment and ground-water collection ..............................................................................................................12

Microcosm preparation and incubation .............................................................................................................12

Geochemical and microbial community analyses ..............................................................................................12

Calculation of degradation rates ........................................................................................................................13

Laboratory enrichment experiments ...........................................................................................................................14

Enrichment preparation and incubation .............................................................................................................14

Geochemical and microbial community analyses ..............................................................................................14

Field data collection and analyses ..............................................................................................................................15

Surficial wetland sediment samples ...................................................................................................................15

Ground-water samples .......................................................................................................................................16

Calculation of degradation rates ........................................................................................................................16

Laboratory experiments on anaerobic degradation of 1,1,2,2-tetrachloroethane and association with

microbial communities .....................................................................................................................................................16

Degradation rate of 1,1,2,2-tetrachloroethane ............................................................................................................17

Daughter compound production and degradation ......................................................................................................22

Spatial variability ...............................................................................................................................................22

Spatial variability of vinyl chloride production ........................................................................................23

Spatial variability of vinyl chloride degradation .......................................................................................28

Seasonal variability ............................................................................................................................................29

Substrate type .....................................................................................................................................................30

Redox conditions ...............................................................................................................................................32

Toxicity ..............................................................................................................................................................33

Pre-exposure to contaminants ............................................................................................................................34

Microbial communities and associations with degradation pathways ........................................................................35

Degradation of 1,1,2,2-tetrachloroethane ..........................................................................................................36

Daughter compound production and degradation ..............................................................................................41

Spatial variability of vinyl chloride production ........................................................................................41

Spatial variability of vinyl chloride degradation .......................................................................................45

Seasonal variability ...................................................................................................................................47

Substrate type ............................................................................................................................................49

Comparisons to field data on anaerobic degradation of 1,1,2,2-tetrachloroethane and association

with microbial communities .............................................................................................................................................50

Degradation rate of 1,1,2,2-tetrachloroethane ............................................................................................................50

Daughter compound production and degradation ......................................................................................................51

Microbial communities ...............................................................................................................................................53

Implications for natural attenuation and remediation .................................................................................................56

Summary and conclusions ...................................................................................................................................................59

References cited ..................................................................................................................................................................61
vi

Figures

 1. Map showing sampling sites and locations of sections A-A′ and C-C′ in the wetland

study area along the West Branch Canal Creek, Aberdeen Proving Ground, Maryland .................................4

 2. Diagram showing anaerobic degradation pathways for 1,1,2,2-tetrachloroethane (TeCA) ................................5

 3. Section showing hydrogeology and 1,1,2,2-tetrachloroethane (TeCA) distribution in 

ground water along section A-A′ , June–October 1995 ..................................................................................7

 44-41. Graphs showing:

 4A,B. Degradation of 1,1,2,2-tetrachloroethane in previous anaerobic microcosm

experiment (30E4) with wetland sediment (site WB30), May–June 1996:

(A) 1,1,2,2-tetrachloroethane (TeCA) and the sum of daughter products 

in the TeCA-amended live microcosms and sterile controls, and 

(B) Daughter products 1,2-dichloroethene (12DCE, total of the cis and 

trans isomers); vinyl chloride (VC); 1,1,2-trichloroethane (112TCA); and

1,2-dichloroethane (12DCA) in theTeCA-amended microcosms ...................................................................9

5A,B. Apparent first-order degradation rates of (A) 1,1,2,2-tetrachloroethane (TeCA) and 

(B) 1,1,2-trichloroethane (112TCA) in anaerobic microcosm experiments, 1998–2000 ..............................19

6A-C. Degradation of 1,1,2,2-tetrachloroethane (TeCA) and production of daughter 

compounds in anaerobic microcosm experiments: (A) 23TeCA.1, (B) 23TeCA.2, 

and (C) 23TeCA.3 conducted in February–April 1998 using sediment from 

site WB23, and amended with methanol and three different initial TeCA 

concentrations (noted on figures) ..................................................................................................................20

7A,B. Degradation of 1,1,2,2-tetrachloroethane (TeCA) and production of daughter

compounds in anaerobic microcosms, March–April 1999, constructed with

wetland sediment from (A) site WB23 and (B) site WB30 ...........................................................................21

8A,B. Degradation of 1,1,2,2-tetrachloroethane (TeCA) and production of daughter

compounds in anaerobic microcosms, July–August 1999, constructed with 

wetland sediment from (A) site WB23 and (B) site WB30 ...........................................................................21

9A,B. Degradation of 1,1,2,2-tetrachloroethane (TeCA) and production of daughter

compounds in anaerobic microcosms, October–November 2000, constructed

 with wetland sediment from (A) site WB23 and (B) site WB30 ..................................................................22

10,10. Mass balance between initial (day 1) concentration of the parent compound 

1,1,2,2-tetrachloroethane (TeCA) and the sum of daughter compounds 

produced in the live anaerobic microcosms, 1998–2000 .............................................................................22

11A,B. Daughter product distribution in anaerobic microcosms amended with 

1,1,2,2-tetrachloroethane (TeCA) and constructed with wetland 

sediment from sites WB23 and WB30 in microcosm experiments in

(A) March–April 1999 and (B) October–November 2000 ...........................................................................23

12,12. Daughter product distribution in anaerobic microcosms amended with 

1,1,2-trichloroethane (112TCA) and constructed with wetland sediment

from site WB23 (March–April 1999) and site WB30 (March–April 1999 and 

October–November 2000) ............................................................................................................................24

13A-D. Methane and ferrous iron concentrations in 1999 and 2000 microcosms

amended with 1,1,2,2-tetrachloroethane (TeCA) (and live controls, LC):

(A) ferrous iron in microcosms constructed with sediment from site WB23;

(B) ferrous iron in microcosms constructed with sediment from site WB30;

(C) methane in microcosms constructed with sediment from site WB23; and

(D) methane in microcosms constructed with sediment from site WB30 ....................................................25
vii

Figures—Continued

44-41. Graphs showing–Continued

14A,B. Degradation of 1,1,2-trichloroethane (112TCA), production of daughter

compounds 1,2-dichloroethane (12DCA) and vinyl chloride (VC), and 

production of methane in anaerobic microcosms, March–April 1999, 

constructed with wetland sediment from (A) site WB23 and (B) site WB30 ..............................................26

 15. Degradation of 1,1,2-trichloroethane (112TCA), production of daughter 

compounds 1,2-dichloroethane (12DCA) and vinyl chloride (VC), and

production of methane in anaerobic microcosms, October–November 2000,

constructed with wetland sediment from site WB30 ....................................................................................26

16A,B. Production of vinyl chloride in enrichment cultures inoculated with WB23 

or WB30 microcosm slurry, October–November 2000 and (A) amended

with 1,1,2-trichloroethane and either acetate (TCA30A and TCA23A) or

hydrogen (TCA30H and TCA23H), or (B) amended with 1,2-cis-dichloroethene

(cDCE) or 1,2-trans-dichloroethene (tDCE) and with either acetate (cDCE30A

and cDCE23A) or hydrogen (cDCE30H and cDCE23H) ............................................................................27

17,17. Percent of vinyl chloride (VC) degraded in anaerobic microcosms amended 

with 1,1,2,2-tetrachloroethane (TeCA) or amended with TeCA and methanol

(MeOH), 1996–2000 ....................................................................................................................................28

18A,B. (A) Degradation of vinyl chloride and (B) production of methane in sediment

enrichments inoculated with WB23 or WB30 microcosm slurry,

October–November 2000, and amended with only vinyl chloride (“no

addition”), with vinyl chloride and 2-bromoethanesulfonic acid (BES), 

or with vinyl chloride and amorphous ferric oxyhydroxide (FeOOH) .........................................................29

19A,B. Daughter product distribution in anaerobic microcosms constructed with 

wetland sediment from site WB23, February–April 1998, that were 

amended with methanol and with (A) 1,1,2,2-tetrachloroethane (TeCA) and 

(B) 1,1,2-trichloroethane (112TCA) .............................................................................................................31

20A,B. Methane concentrations in February–April 1998 anaerobic microcosms 

amended with methanol and (A) 1,1,2,2-tetrachloroethane (TeCA)

and (B) 1,1,2-trichloroethane (112TCA)........................................................................................................32

 21 ,21. Degradation of 1,1,2,2-tetrachloroethane (TeCA) and production of daughter

compounds in anaerobic microcosms amended with 2-bromoethanesulfonic

acid (BES), methanol, and TeCA and constructed with wetland sediment

from site WB23, February–April 1998 ........................................................................................................33

22A,B. Degradation of (A) 1,1,2,2-tetrachloroethane (TeCA) and production of daughter

compounds in anaerobic microcosms amended with sulfate (SO4 ), methanol,

and TeCA and constructed with wetland sediment from site WB23,

February–April 1998, and (B) effect of SO4 and 2-bromoethanesulfonic

acid (BES) addition on methane and SO4 concentrations ...........................................................................34

 23,23. Degradation of 1,1,2,2-tetrachloroethane (TeCA) and production of daughter compounds

in anaerobic microcosms conducted in February–April 1998 using sediment

from background site WB19 and amended with methanol ..........................................................................35

24A,B. (A) Degradation of vinyl chloride and (B) production of methane in sediment

enrichment experiments amended with vinyl chloride and constructed

with fresh, pre-exposed, and preincubated wetland sediment from

sites WB23 and WB30 .................................................................................................................................36
viii

Figures—Continued

44-41. Graphs showing–Continued

25A-D. Bacteria terminal-restriction fragment length polymorphism profiles in selected TeCA-amended

microcosms constructed with sediment from site WB23, February–April 1998:

(A) live control [no 1,1,2,2-tetrachloroethane (TeCA) added, experiment 23LC],

(B) killed control for microcosms with low initial TeCA concentration 

[experiment killed 23TeCA.1], (C) killed control for microcosms with

TeCA and sulfate amendment [experiment killed 23SO4-TeCA.2], and 

(D) live microcosm with low initial TeCA concentration [experiment 23TeCA.1] ....................................37

26A,B. Bacteria terminal-restriction fragment length polymorphism profiles in

1,1,2,2-tetrachloroethane- (TeCA-) amended microcosms 

constructed with sediment from site WB23 compared to site WB30,

July–August 1999: (A) site WB23 (experiment 23TeCA.7/99) and 

(B) site WB30 (experiment 30TeCA.7/99) ..................................................................................................38

27A,B. Bacteria terminal-restriction fragment length polymorphism profiles in

1,1,2,2-tetrachloroethane- (TeCA-) amended microcosms constructed

with sediment from site WB23 compared to site WB30, March–April 1999: 

(A) site WB23 (experiment 23TeCA.3/99), and (B) site WB30

(experiment 30TeCA.3/99) ...........................................................................................................................39

28A,B. Bacteria terminal-restriction fragment length polymorphism profiles in

1,1,2,2-tetrachloroethane- (TeCA-) and 1,1,2-trichloroethane- (112TCA-)

amended microcosms constructed with sediment from site WB30,

October–November 2000: (A) site WB30, amended with TeCA

(experiment 30TeCA.10/00), and (B) site WB30, amended with 112TCA

(experiment 30TCA.10/00) ..........................................................................................................................40

29A,B. Terminal-restriction fragment length polymorphism (TRFLP) profiles for (A) bacteria

and (B) methanogens in live control microcosms (experiment 23LC.3/99)

constructed with sediment from site WB23, March–April 1999 .................................................................42

30A,B. Bacteria terminal-restriction fragment length polymorphism profiles in

1,1,2-trichloroethane- (112TCA-) amended microcosms constructed

with sediment from site WB23 compared to site WB30, March–April 1999:

(A) site WB23 (experiment 23TCA.3/99), and (B) site WB30 (experiment

30TCA.3/99) .................................................................................................................................................43

31A,B. Methanogen terminal-restriction fragment length polymorphism profiles in

1,1,2,2-tetrachloroethane- (TeCA-) amended microcosms constructed

with sediment from site WB23 compared to site WB30, March–April 1999:

(A) site WB23 (experiment 23TeCA.3/99), and (B) site WB30 (experiment

30TeCA.3/99) ...............................................................................................................................................44

 32,32. Proportion of Methanosarcinaceae and the relative abundance of total

methanogens in 1,1,2,2-tetrachloroethane- (TeCA-) amended microcosms

constructed with WB23 and WB30 sediments, March–April 1999

(experiments 23TeCA.3/99 and 30TeCA.3/99) ...........................................................................................45

 33,33. Proportion of Methanosarcinaceae in 1,1,2,2-tetrachloroethane- (TeCA-)

amended microcosms constructed with WB30 sediments, with or 

without addition of methanol (MeOH), October–November 2000 ..............................................................46

 34,34. Ferrous [Fe(II)] and ferric [Fe(III)] iron concentrations (in micromoles

per gram wet weight) in wetland sediment samples that were collected 

by coring at sites WB23 and WB30, August 2000 .......................................................................................46

 35. Bacteria terminal-restriction fragment length polymorphism profiles in

sediment samples from site WB23 prior to microcosm preparation 

and incubation, March and August 1999 ......................................................................................................48
ix

Figures—Continued

44-41. Graphs showing–Continued

36A,B. Methanogen terminal-restriction fragment length polymorphism profiles

in 1,1,2,2-tetrachloroethane- (TeCA-) amended microcosms constructed 

with sediment from sites WB23 and WB30, July–August 1999: 

(A) site WB23 (experiment 23TeCA.7/99), and (B) site WB30 

(experiment 30TeCA.7/99) ...........................................................................................................................49

37A,B. Natural log of 1,1,2,2-tetrachloroethane (TeCA) concentrations [(ln (C)] 

(normalized by chloride concentrations) and traveltime in wetland

porewater (A) from piezometer siteWB30 in July–August 1995 and 

(B) from a multi-level sampler (WBM30) at the same site in June–July 2000 ............................................52

38A,B. Concentrations of (A) ferrous iron and (B) methane in wetland porewater 

collected from multi-level samplers (WBM23 and WBM30) at sites WB23

and WB30, June–July 2000 ..........................................................................................................................53

 39,39. Distributions of 1,1,2,2-tetrachloroethane (TeCA) and daughter compounds

in wetland porewater collected from multi-level samplers (WBM23 and 

WBM30) at sites WB23 and WB30, June–July 2000 ..................................................................................54

40A,B. Bacteria terminal-restriction fragment length polymorphism profiles in the

surficial wetland sediment along (A) transect A-A′ and (B) transect C-C′, 1999 .......................................55

4 1,41. Bacteria terminal-restriction fragment length polymorphism profile in

surficial wetland sediment at background site WB19, March 1999 .............................................................56

Tables

1. Treatments used in anaerobic microcosm experiments to examine 1,1,2,2-tetrachloroethane

or 1,1,2-trichloroethane degradation in wetland sediments from the West Branch

Canal Creek study area, Aberdeen Proving Ground, Maryland, May 1996–November 2000 ......................11

2. Treatments used in enrichment experiments with anaerobic daughter products of

1,1,2,2-tetrachloroethane (TeCA) degradation ..............................................................................................15

3. First-order rate constants and half-lives for degradation of 1,1,2,2-tetrachloroethane 

(TeCA) or 1,1,2-trichloroethane (112TCA) in anaerobic microcosm experiments

conducted using wetland sediments, May 1996–November 2000 ................................................................18

4. Percentage of each daughter product observed in the TeCA-amended microcosms

constructed with sediment from site WB23 and incubated at 5 degrees Celsius 

(experiment 23TeCA5.3/99) compared to those incubated at 19 degrees Celsius in 

March–April 1999 (experiment 23TeCA.3/99) and July–August 1999 (experiment

 23TeCA.7/99) ...............................................................................................................................................30

5. Degradation rates for 1,1,2,2-tetrachloroethane (TeCA) and trichloroethene (TCE) 

estimated from field data in July–August 1995 and June–July 2000 along upward 

flowpaths in the wetland sediments, using TeCA and TCE concentrations normalized 

with chloride and an estimated linear flow velocity of 0.9 meters per year ..................................................51

6. Abundance of Methanosarcinaceae as a percentage of the total methanogen community

 in surface grab samples of wetland sediment, March 1999 ..........................................................................56
x

Conversion Factors, Vertical Datum, and Abbreviations

_____________________________________________________________________________________________________________________________________

Multiply By To obtain

_____________________________________________________________________________________________________________________________________

Length

centimeter (cm) 0.3937 inch

millimeter (mm) 0.03937 inch

meter (m) 3.281 foot

meter (m) 1.094 yard

Volume

liter (L) 33.82 ounce, fluid

liter (L) 2.113 pint

liter (L) 1.057 quart

liter (L) 0.2642 gallon

liter (L) 61.02 cubic inch

Flow rate

meter per year (m/yr) 3.281 foot per year

Mass

gram (g) 0.03527 ounce, avoirdupois

kilogram (kg) 2.205 pound, avoirdupois

____________________________________________________________________________________________________________________________________

Temperatures in degrees Celsius ( °C) may be converted to degrees Fahrenheit ( °F) as follows:

°

F = (1.8 x °C) + 32 

Vertical Datum: In this report, “sea level” refers to the National Geodetic Vertical Datum of 1929—a geodetic datum 

derived from a general adjustment of the first-order level nets of the United States and Canada, formerly called Sea Level Datum 

of 1929.

Concentrations of chemical constituents in water are given either in milligrams per liter (mg/L), micrograms per liter

(μg/L), millimoles per liter (mmol/L), or micromoles per liter (μmol/L).
Abstract 1

Anaerobic Degradation of 1,1,2,2-Tetrachloroethane and Association 

with Microbial Communities in a Freshwater Tidal Wetland, 

Aberdeen Proving Ground, Maryland: Laboratory Experiments and 

Comparisons to Field Data

By Michelle M. Lorah, Mary A. Voytek, Julie D. Kirshtein, and Elizabeth J. (Phillips) Jones

Abstract

Defining biodegradation rates and processes is 

a critical part of assessing the feasibility of monitored natural attenuation as a remediation method 

for ground water containing organic contaminants. 

During 1998–2001, the U.S. Geological Survey 

conducted a microbial study at a freshwater tidal 

wetland along the West Branch Canal Creek, 

Aberdeen Proving Ground, Maryland, as part of an 

investigation of natural attenuation of chlorinated 

volatile organic compounds (VOCs) in the wetland sediments. Geochemical analyses and molecular biology techniques were used to investigate 

factors controlling anaerobic degradation of 

1,1,2,2-tetrachloroethane (TeCA), and to characterize the microbial communities that potentially 

are important in its degradation. Rapid TeCA and 

daughter product degradation observed in laboratory experiments and estimated with field data 

confirm that natural attenuation is a feasible remediation method at this site. The diverse microbial 

community that seems to be involved in TeCA 

degradation in the wetland sediments varies with 

changing spatial and seasonal conditions, allowing 

continued effective natural attenuation throughout 

the year.

Rates of TeCA degradation in anaerobic microcosm experiments conducted with wetland sediment collected from two different sites (WB23 and 

WB30) and during three different seasons 

(March–April 1999, July–August 1999, and 

October–November 2000) showed little spatial 

variability but high seasonal variability. Initial 

first-order degradation rate constants for TeCA 

ranged from 0.10±0.01 to 0.16±0.05 per day (halflives of 4.3 to 6.9 days) for March–April 1999 and 

October–November 2000 microcosms incubated 

at 19 degrees Celsius, whereas lower rate constants of 0±0.03 and 0.06±0.03 per day were 

obtained in July–August 1999 microcosms incubated at 19 degrees Celsius. Microbial community 

profiles showed that low microbial biomass and 

microbial diversity in the summer, possibly due to 

competition for nutrients by the wetland vegetation, could account for these unexpectedly low 

degradation rates. In microcosms incubated at 

5 degrees Celsius, about 50 percent of the initial 

TeCA in solution was converted to daughter products within a 35-day incubation period, indicating 

that biodegradation in the wetland sediments can 

continue during cold winter temperatures.

Initial pathways of TeCA degradation were the 

same in the wetland sediment microcosms regardless of the season or sediment collection site, the 

reduction-oxidation conditions, and the previous 

exposure of the sediment to contamination. 

Immediate and simultaneous dichloroelimination 

and hydrogenolysis, producing 1,2-dichloroethene (12DCE) and 1,1,2-trichloroethane 

(112TCA), respectively, were the initial TeCA 

degradation pathways in all live microcosm experiments. The production and degradation of vinyl 

chloride (VC), which is the most toxic of the 

TeCA daughter compounds, was affected by spatial and seasonal variability, reduction-oxidation 

condition, and pre-exposure of the wetland sediment. TeCA-amended microcosms constructed 

with WB30 sediment showed approximately twice 

as much VC production as those constructed with 

WB23 sediment. Results of 112TCA-amended 

microcosms indicated that the greater production 

of VC in the WB30 sediment resulted from a 

greater predominance of the 112TCA dichloro-
2 TeCA Degradation and Microbial Communities in a Freshwater Tidal Wetland, APG, Md.

elimination pathway in these sediments. VC degradation also was substantially higher in microcosms constructed with WB30 sediment than 

those constructed with WB23 sediment, resulting 

in lower VC concentrations at the end of WB30 

microcosms. Enrichment experiments in which 

microcosm slurry was amended with high initial 

VC concentrations showed that the spatial difference in VC degradation was negligible after prolonged incubation under methanogenic conditions. 

Inhibition of methanogenic activity in microcosms 

by addition of sulfate or of 2-bromoethanesulfonic 

acid inhibited production and degradation of VC. 

Inhibition of methanogenesis by addition of ferric 

iron or of 2-bromoethanesulfonic acid also completely inhibited VC degradation in VC-amended 

enrichment experiments. Pre-exposure to VC substantially increased degradation in VC-amended 

enrichment experiments.

A microbial consortium, rather than one microbial species or group, likely is involved in the degradation of TeCA, as indicated by the occurrence 

of multiple degradation pathways and the variability in VC production and degradation. A bacterial 

peak at 90 base pair (bp) fragment length in terminal-restriction fragment length polymorphism 

(TRFLP) profiles was associated with TeCA 

hydrogenolysis to 112TCA, and bacterial species 

represented by 198 and 170 bp fragment lengths 

were associated with TeCA dichloroelimination to 

12DCE. Dichloroelimination of 112TCA to VC 

was associated with increasing dominance of the 

198 bp bacterial peak in March–April 1999 and 

October–November 2000 microcosms, whereas an 

86 bp or the 170 bp bacterial peak was associated 

with 112TCA dichloroelimination in the summer 

experiment. Hydrogenolysis of 12DCE to VC was 

associated with a carbon dioxide-utilizing methanogen at 307 bp in the March–April 1999 and 

October–November 2000 microcosm experiments, 

whereas production of VC occurred despite low 

methanogen biomass and methane production in 

the July–August 1999 experiments. Production of 

VC in the absence of methane production also 

occurred in 12DCE-amended enrichment cultures. The exponential production of VC in the 

12DCE-amended enrichment cultures after an initial lag indicated growth of a microbial species or 

group, possibly one of the known dehalorespiring 

bacteria. Molecular analyses using specific primers targeting dehalorespiring bacteria of the Dehalococcoides group (Dehalococcoides ethenogenes

and Dehalococcoides sp. strain FL2) and of the 

acetate-oxidizing Desulfuromonas group (Desulfuromonas sp. strain BB1 and Desulfuromonas chloroethenica) showed the presence of these bacteria 

in microcosm slurry from site WB30 but not from 

site WB23. Addition of hydrogen, which is the 

favored substrate of Dehalococcoides, tripled VC 

production in 12DCE-amended enrichment cultures. VC degradation showed a marked association with an increase in the relative proportion of 

Methanosarcinaceae, a family of methanogens that 

includes all those capable of utilizing acetate as a 

substrate, in the total methanogen community.

Half-lives for TeCA and TCE estimated from 

field data were in the range of 60 to 100 days, 

which agrees well with laboratory estimates of 

degradation rates considering the inherent differences in the laboratory and field systems. Both 

laboratory microcosm experiments and field data 

showed that 12DCE and VC are the predominant, 

persistent daughter compounds from TeCA degradation. In addition, porewater chemistry showed 

higher accumulation of VC in the wetland sediment at site WB30 than at site WB23, as was 

observed in the microcosm experiments. Molecular analyses of grab samples of surficial wetland 

sediment showed that all the microbial species or 

groups linked to TeCA degradation in the microcosm experiments were present in all sediment 

samples. Microbial biomass and diversity were 

lowest in an area of the wetland (transect C-C′) 

where porewater VOC concentrations are highest, 

indicating that the higher VOC concentrations 

could result from lower degradation rates. The 

lower microbial biomass and diversity in this area 

could be caused by toxic effects of the contaminants, or possibly from differences in frequency 

and duration of tidal inundation.

Introduction

Chlorinated solvents such as trichloroethene (TCE) and 

1,1,2,2-tetrachloroethane (TeCA) tend to be relatively resistant to biodegradation in aquifers, compared to the biodegradation potential of petroleum hydrocarbons (National 

Research Council, 2000). This has led to the development of 
Introduction 3

long chlorinated solvent plumes at a number of Department 

of Defense (DoD) and private facilities that have reached, or 

may reach, discharge areas in surface-water bodies or wetlands. About 75 percent of Resource Conservation and 

Recovery Act (RCRA) and Superfund sites, including DoD 

sites, are located within a half mile of a surface-water body 

(Tomassoni, 2000), where wetlands often are part of the 

landscape. Conventional engineered remediation of ground 

water contaminated with chlorinated solvents can be 

extremely costly and could be detrimental to ecologically 

sensitive wetland environments. Although chlorinated solvents tend to be relatively resistant to degradation within 

most aquifers, rapid and complete transformations can occur 

within the organic-rich reducing environment typical of wetland sediments, indicating that monitored natural attenuation 

may be an effective remediation option for discharges into 

wetlands (Lorah and others, 1997; Lorah and Olsen, 1999a, 

b; Lorah and others, 2001). The study of TeCA degradation 

and associated microbial communities in wetland sediments 

reported here is part of a larger ongoing study by the U.S. 

Geological Survey (USGS) of natural attenuation of chlorinated volatile organic compounds (VOCs) in a freshwater 

tidal wetland along the West Branch Canal Creek, Aberdeen 

Proving Ground (APG), Maryland (fig. 1) (Lorah and others, 

1997). TeCA and TCE, which can be a daughter product of 

TeCA degradation in addition to being a widely used solvent, are the major contaminants in the shallow aquifer (less 

than about 15 m deep) and wetland sediments at this site 

(Lorah and others, 1997; Lorah and Olsen, 1999a, b; Lorah 

and others, 2001).

Defining biodegradation rates and reactions is a critical 

part of assessing the feasibility of monitored natural attenuation as a remediation method for ground water containing 

organic contaminants because biodegradation is commonly 

the primary destructive process for these contaminants 

(Wiedemeier and others, 1998; National Research Council, 

2000). Laboratory experiments to define biodegradation 

processes and rates may be warranted during investigation of 

natural attenuation at a site, depending on the availability of 

information in the literature and on the expected complexity 

of biodegradation processes for contaminants at the site 

(Wiedemeier and others, 1998). Relatively few studies of 

TeCA degradation in the environment have been conducted 

compared to the numerous field and laboratory studies of 

TCE degradation. In addition, degradation of TeCA is 

expected to be complex because there are three main initial 

pathways of reaction that can occur, producing both chlorinated ethene and chlorinated ethane daughter products (fig. 

2). Each of these intermediate daughter products could then 

be degraded through several different reaction pathways. 

Elucidation of factors controlling the occurrence of these 

diverse degradation pathways for TeCA is critical because 

of differing toxicity and persistence of the intermediate 

daughter products. Of the chlorinated ethenes and ethanes, 

accumulation of vinyl chloride (VC) might be considered of 

the greatest concern for remediation of a contaminated site 

because it is the only known carcinogen and has the lowest 

maximum contaminant level (MCL) for drinking water 

(2 μg/L, or micrograms per liter) (U.S. Environmental Protection Agency, 2001). Higher MCLs of 5, 70, 100, 5, and 

5 μg/L, respectively, have been established for TCE; cis-1,2-

dichloroethene (cDCE); trans-1,2-dichloroethene (tDCE); 

1,1,2-trichloroethane (112TCA); and 1,2-dichloroethane 

(12DCA) (U.S. Environmental Protection Agency, 2001). 

Although TeCA currently (2002) is not a regulated drinkingwater contaminant, the U.S. Environmental Protection 

Agency (1998) has placed it on the Drinking Water Contaminant Candidate List, where it is listed as a priority contaminant for determination of regulatory needs based on potential 

to occur in public water systems with a relatively high frequency and at concentrations that could pose a threat to public health.

Site characteristics, in addition to contaminant properties, determine the potential complexity of biodegradation 

processes. Wetlands can be extremely complex because they 

typically have a high diversity of reduction-oxidation 

(redox) zones and of microbial populations. In addition, 

wetlands are dynamic surficial systems that can have a high 

degree of spatial and temporal variability in hydrology, 

microbiology, and water chemistry because of variable surface-water flow, ground-water recharge, air temperature, 

sediment organic carbon content, vegetation type, density, 

and activity. Wetlands, therefore, can show greater spatial 

and seasonal variability in contaminant distribution and biodegradation rates than observed in most deeper ground-water 

systems. This variability needs to be understood to determine whether natural attenuation is sufficient as a remediation method at all times of the year, and to predict the 

amount of time needed for complete attenuation of contaminants in the wetland sediment. In addition, this information 

can improve long-term monitoring plans by identifying possible factors that could cause changes in natural attenuation 

efficiency. Because hydrogeologic and geochemical data do 

not always indicate obvious reasons for variations in degradation processes and rates, molecular biology techniques to 

characterize microbial communities can be useful (Flynn and 

others, 2000; Fennel and others, 2001).

The purpose of the USGS microbial study at the freshwater tidal wetland at APG (fig. 1) is to use a combination of 

geochemical analyses and molecular biology techniques to 

investigate factors controlling the occurrence of different 

anaerobic TeCA degradation pathways, and to characterize 

the microbial communities that potentially are important in 

these degradation reactions (fig. 2). The objectives of this 

ongoing study are to (1) evaluate spatial and seasonal variations in TeCA degradation pathways and rates in the wetland 

sediment, (2) identify microbial groups and/or species 

involved in biodegradation of TeCA in the wetland sediment, 

(3) determine the spatial and seasonal variations in microbial 

communities in the wetland sediment, and (4) ultimately 

develop gene probes for indicator microbial species that are 

critical for different reaction pathways of TeCA degradation 

to occur. Gene probes could be used for long-term monitoring of the presence of critical microbial species over time in 
4 TeCA Degradation and Microbial Communities in a Freshwater Tidal Wetland, APG, Md.
Introduction 5
6 TeCA Degradation and Microbial Communities in a Freshwater Tidal Wetland, APG, Md.

the wetland sediment and for evaluating the potential for natural attenuation at other contaminated sites. Because anaerobic TeCA degradation encompasses chlorinated ethane and 

ethene pathways (fig. 2), results of this study are transferable 

to other sites at APG and elsewhere that are contaminated 

with TeCA, TCE, or their anaerobic degradation products.

Purpose and Scope

The purpose of this report is to describe the anaerobic 

TeCA degradation reactions occurring in wetland sediments 

in the West Branch Canal Creek study area at APG (fig. 1), 

and the association between these degradation reactions and 

the microbial communities in the wetland sediment. Anaerobic laboratory microcosm experiments and enrichment cultures are used to evaluate spatial and seasonal variability in 

TeCA degradation rates and pathways, and to evaluate other 

potential factors controlling TeCA degradation, including 

redox conditions, substrate type, and pre-exposure of the 

sediment to the contaminants. In addition to microcosms 

incubated with TeCA, microcosms and/or enrichment cultures amended with the daughter products VC, 112TCA, 

cDCE, or tDCE (fig. 2) assist in elucidation of TeCA degradation pathways and controlling factors. Enrichment experiments focus primarily on VC production and degradation 

because of the importance of this toxic TeCA daughter 

product that potentially can be produced through three reaction pathways (fig. 2). Concomitant geochemical and microbial analyses made during anaerobic microcosm experiments 

are used to define TeCA degradation pathways and relate 

these pathways to the microbial communities in the wetland 

sediment. Geochemical analyses include measurements of 

the distributions of TeCA, its daughter products, and redoxsensitive constituents over time in the microcosms. Microbial analyses include terminal-restriction fragment length 

polymorphism (TRFLP) analysis of polymerase chain reaction (PCR)-amplified bacterial 16S rDNA and mcrA (methylcoreductase gene) to obtain profiles of the bacterial and 

methanogen communities, respectively. These genetic analyses commonly are referred to as “DNA fingerprinting.” 

Although the primary objective of this report is to present 

results of laboratory experiments, selected field data are used 

to estimate biodegradation rates, evaluate TeCA daughter 

compound distributions in the wetland porewater, and evaluate distribution of microbial communities in the wetland sediment.

Between 1998 and 2000, four laboratory microcosm 

experiments were conducted with wetland sediment and 

porewater collected at different times of the year and from 

two contaminated sites in the wetland area, sites WB23 and 

WB30 (fig. 1). During the 35- to 43-day incubation period 

for each experiment, water in the microcosms was analyzed 

for TeCA and its potential daughter products and for redoxsensitive constituents, while sediment in the microcosms 

was analyzed for microbial communities. To allow comparisons of TeCA degradation rates among the experiments, all 

microcosms were incubated at a temperature of 19 °C 

(degrees Celsius), which was the average summer temperature in the shallow wetland sediment (Lorah and others, 

1997). One additional set of microcosms was incubated at a 

winter temperature of 5 °C. The effect of redox conditions 

on TeCA degradation is evaluated using microcosm and 

enrichment experiments with added iron or sulfate, or with 

an inhibitor of methanogenesis. Substrate type effects are 

examined through the addition of methanol, hydrogen, or 

acetate in some experiments. The effect of prior exposure, 

or “pre-exposure,” of the wetland sediment to the chlorinated organic contaminants is evaluated using TeCAamended microcosms constructed with wetland sediment 

and porewater collected from a background site to compare 

to those constructed with sediment and porewater from 

contaminated sites. In addition, pre-exposure effects on VC 

degradation are examined in VC-amended sediment enrichments constructed using microcosm slurry that was or was 

not previously incubated with TeCA.

For comparison to laboratory-derived rates, changes in 

porewater concentrations along flowpaths through the wetland sediment are used to estimate TeCA degradation rates 

at several multi-level sampling and piezometer sites. Distributions of daughter products in the wetland porewater at 

sites WB23 and WB30, which were the two main sites used 

for sediment collection for the microcosm experiments, are 

compared to distributions observed in laboratory microcosms. To evaluate how the microbial communities change 

seasonally and spatially in the wetland sediment, surficial 

sediment samples were collected at 10 piezometer sites 

(including sites used to obtain sediment for microcosm 

experiments) in March 1999 and August 1999 (fig. 1). The 

laboratory- and field-derived data are compared, and the 

implications of these results to natural attenuation of the 

contaminants in the wetland are discussed.

Description of Study Area

The study area is located along the West Branch Canal 

Creek at APG, a U.S. Army base in Maryland near the head 

of the Chesapeake Bay (fig. 1). Freshwater tidal wetlands 

surround much of West Branch Canal Creek, which has tidal 

fluctuations in the range of 0.15 to 0.61 m (meters). Release 

of waste products by spills, landfilling, and discharge to sewers during chemical manufacturing operations near the creek 

resulted in ground-water contamination (Lorah and Clark, 

1996; Lorah and others, 1997). As part of the study of natural attenuation of VOCs begun by the USGS in 1992, nested 

drive-point piezometers were installed in the wetland study 

area primarily along two transects on the eastern side of 

West Branch Canal Creek (Lorah and others, 1997) (figs. 1 

and 3). Transects are aligned perpendicular to the creek, 

along the general direction of ground-water flow in the aquifer (Lorah and others, 1997). As part of a study under the 

DoD Environmental Security Technology Certification Program (ESTCP), additional multi-level samplers and piezometers, described in Spencer and others (2002) and Dyer and 

others (2002), were installed in the wetland and shallow 

aquifer sediments at six of the pre-existing drive-point piezometer sites (fig. 1).

The contaminated aquifer, known as the Canal Creek 

aquifer, is about 12 to 14 m thick in the study area and 
Introduction 7

consists mainly of medium- to coarse-grained sand and 

gravel (fig. 3). Wetland sediments that overlie the aquifer 

consist of two distinct layers that have a combined thickness 

of about 1.8 to 3.6 m in the study area--a lower unit of silty 

to sandy clay or clayey sand, and an upper unit of peat mixed 

with variable amounts of clay and silt (Lorah and others, 

1997). The total organic carbon content in the wetland sediment averaged about 1 percent in the lower clayey unit and 

18 percent in the upper peat unit (Lorah and others, 1997; 

Olsen and others, 1997). Ground-water-flow directions 

within the wetland area are predominantly upward. At low 

tide, water from the aquifer discharges through both the 

wetland and channel sediments. At high tide, however, 

reversals in ground-water-flow directions can occur at some 

sites (Lorah and others, 1997). The linear ground-waterflow velocity in the wetland sediments is estimated to be 

about 0.6 to 0.9 m/yr (meters per year) (Lorah and others, 

1997).

The contaminant plume in the aquifer contains both 

TeCA and TCE from disposal sources, and previous field 

and laboratory studies have shown that rapid TeCA and TCE 

degradation occurs as the plume flows upward from the 

aerobic sand aquifer and through the anaerobic wetland sediments (Lorah and others, 1997, 2001; Lorah and Olsen, 

1999a, b). Although carbon tetrachloride and chloroform 

also are present in the aquifer along the A-A′ and C-C′

transects, their concentrations are below detection levels 

(0.5 μg/L, or micrograms per liter) in the wetland sediments 

(Lorah and others, 1997). TeCA and TCE concentrations 

were highest (about 4,700 and 300 μg/L, respectively) 
8 TeCA Degradation and Microbial Communities in a Freshwater Tidal Wetland, APG, Md.

upgradient from the eastern edge of the wetland at site 

CC 27 (figs. 1 and 3), which is near a suspected source from 

a past sewerline discharge point (Lorah and Clark, 1996; 

Lorah and others, 1997). TCE is a major contaminant in the 

aquifer along section A-A′, but its concentrations are an 

order of magnitude lower along section C-C′ (Lorah and 

others, 1997; Lorah and Olsen, 1999b). Concentrations of 

VOCs observed in ground water at site CC 27 have remained 

relatively constant from 1987 through 2001 (Lorah and 

Clark, 1996; Lorah and others, 1997; Spencer and others, 

2002), indicating that there is a continuous source of VOCs 

to the shallow aquifer and that the contaminant plume discharging to the wetland is at steady state. Moving downgradient into the wetland area, the TeCA and TCE plume 

remains shallow in the aquifer, reflecting the upward head 

gradients in the aquifer (fig. 3). Total concentrations of 

VOCs are in the range of 400 to 500 μg/L in a thin zone that 

lies directly beneath the wetland sediments (Lorah and 

others, 1997). Along the upward direction of flow through 

the overlying wetland and creek-bottom sediments, total 

concentrations of VOCs decrease substantially until concentrations generally are below detection levels within 0.15 to 

0.30 m of land surface (Lorah and others, 1997; Lorah and 

Olsen, 1999a,b) (fig. 3).

The decrease in VOC concentrations is associated with 

the natural increase in dissolved organic carbon concentrations and decrease in redox state of the ground water along 

the upward flow direction in the wetland sediments (Lorah 

and others, 1997; Lorah and Olsen, 1999b). The aquifer 

typically is aerobic or hypoxic. Iron-reducing conditions are 

predominant in the lower clayey unit of the wetland sediments, whereas methanogenic or methanogenic/mixed zones 

that contain variable concentrations of ferrous iron and sulfide in addition to methane are predominant in the upper peat 

unit. Production of anaerobic daughter products (fig. 2) 

coincides with a decrease in TeCA and TCE concentrations 

and a decrease in redox state of the ground water along 

upward flowpaths through the wetland sediments at each site 

(Lorah and others, 1997; Lorah and Olsen, 1999b). Laboratory microcosm experiments conducted during 1996–1997 

with TeCA and TCE confirmed the rapid degradation of 

these compounds in the wetland sediments (Lorah and 

others, 1997, 2001; Lorah and Olsen, 1999a).

Background on Anaerobic Degradation Pathways

TeCA can degrade under anaerobic conditions by three 

main reaction pathways—a hydrogenolysis pathway that 

produces 1,1,2-trichloroethane (112TCA) and 1,2-dichloroethane (12DCA) as intermediate daughter compounds; a 

dichloroelimination pathway that produces 1,2-dichloroethene (12DCE, both cis- and trans- isomers) and VC as 

intermediate daughter products; and an abiotic dehydrochlorination reaction that produces TCE (fig. 2) (Vogel and 

others, 1987; Lorah and others, 1997; Chen and others, 

1996). Hydrogenolysis entails the sequential replacement 

of a single chlorine atom by hydrogen, whereas dichloroelimination entails the simultaneous replacement of two 

adjacent chlorine atoms by hydrogen to produce a double 

bond. For both of these types of reductive dechlorination 

reactions, the contaminant serves as an electron acceptor, 

resulting in production of more reduced, less-chlorinated 

daughter compounds. Microorganisms require the presence 

of suitable electron donors (substrates) for reductive dechlorination to occur. Possible electron donors include natural 

compounds such as hydrogen, acetate, and methanol, and 

anthropogenic organic compounds such as benzene and 

toluene. Low natural organic carbon content may limit the 

extent of reductive dechlorination in many aquifers 

(Chapelle, 1993; Natural Research Council, 2000), whereas 

a source of organic substrates is not expected to be a limiting 

factor in organic-rich wetland sediments (Lorah and Olsen, 

1999b).

Microbially catalyzed reductive dechlorination reactions 

often are believed to be cometabolic (a fortuitous reaction 

from which the microorganism does not derive energy), but a 

few bacteria capable of obtaining energy from reductive 

dechlorination reactions also have been isolated in the past 

decade (Löffler and others, 1996). Cometabolic reductive 

dechlorination can occur through reactive transition metal 

cofactors, such as vitamin B12, heme, and coenzyme F430, 

in anaerobic microorganisms that catalyze the replacement 

of chlorines by hydrogen atoms (Fathepure and Boyd, 1988; 

Gantzer and Wackett, 1991; Schanke and Wackett, 1992; 

Yager and others, 1997; Novak and others, 1998). These 

reactive coenzymes are especially abundant in methanogens 

(anaerobic microorganisms that produce methane predominantly through metabolism of hydrogen plus carbon dioxide 

or metabolism of acetate) and acetogens (anaerobic microorganisms that synthesize acetate from carbon dioxide 

through the acetyl-CoA pathway) (Fathepure and Boyd, 

1988; Maymó-Gatell and others, 1997; Yager and others, 

1997). The highly chlorinated solvents such as TeCA and 

TCE have been shown to biodegrade faster and undergo 

more complete dechlorination under methanogenic conditions than under the less reducing conditions of nitrate or 

sulfate reduction (McCarty and Semprini, 1994; Lorah and 

others, 1997). Methanogenic conditions often are predominant in freshwater wetland sediments (Capone and Kiene, 

1988). In addition to methanogens and bacteria capable of 

cometabolic reactions, pure cultures of anaerobic dehalorespiring bacteria that can use chlorinated ethenes, including 

tetrachloroethene (PCE), TCE, and cDCE, as terminal electron acceptors to derive energy and grow have been isolated 

(Holliger and others, 1993; Krumholz and others, 1996; 

Maymó-Gatell and others, 1997, 1999, 2001). This growthcoupled metabolism of halogenated VOCs generally is 

believed to be faster than cometabolic reductive dechlorination, and thus may be more effective for remediation of contaminated ground water (Maymó-Gatell and others, 1997; 

Yager and others, 1997). The only chlorinated ethane 

reported to be degraded by dehalorespiring bacteria is 

12DCA (Maymó-Gatell and others, 1997, 1999).

Few previous studies have examined the in situ degradation pathways of TeCA in ground water or soil. In previous 

USGS field and laboratory investigations at the West Branch 
Introduction 9

Canal Creek wetland study area at APG, 12DCE (both cisand trans- isomers) and VC were the predominant persistent 

daughter compounds (Lorah and others, 1997; Lorah and 

Olsen, 1999a, b). These daughter compounds primarily 

resulted from the combined hydrogenolysis and dichloroelimination pathway 1B and dichloroelimination pathway 2 

(fig. 2), whereas abiotic production of TCE was a minor degradation pathway (accounting for less than 2 percent of the 

initial TeCA) (Lorah and others, 1997; Lorah and Olsen, 

1999a, b). Although 12DCE and VC temporarily accumulated, complete anaerobic degradation to non-chlorinated 

end products was observed within 34 days in microcosms 

constructed with the wetland sediment and amended with an 

initial TeCA concentration of about 2.9 μmol/L (micromoles 

per liter) (Lorah and others, 1997; Lorah and Olsen, 1999a) 

(fig. 4). In laboratory experiments conducted using anaerobic municipal digester sludge, a substantially higher percentage of TCE and lower percentage of 112TCA and VC was 

observed from TeCA degradation (Chen and others, 1996) 

than was observed in the anaerobic wetland sediment (Lorah 

and Olsen, 1999a). In contrast, TeCA degraded solely to 

cDCE by pathway 2 (fig. 2) in anaerobic batch experiments 

conducted with bottom sediment from a slow-moving stream 

in the Netherlands (Peijnenburg and others, 1998). Dichloroelimination pathway 2 (fig. 2) also was reported to be the 

major transformation route for TeCA in anaerobic batch 

experiments conducted with bottom sediment from a pond or 

a slow-moving stream in Georgia (Jafvert and Wolfe, 1987). 

Continued degradation of 12DCE to VC was not observed in 

either the experiments by Jafvert and Wolfe (1987) or by 

Peijnenburg and others (1998), possibly because they were 

conducted for less than 10 and 25 days, respectively. Alternatively, the sediments used in these experiments may have 

been lacking a critical microbial species or group.

Field and laboratory studies of reductive dechlorination 

of TCE, which are more numerous than studies of TeCA 

degradation, have shown that dehalorespiring bacteria that 

can reductively dechlorinate 12DCE to VC and ethene are 

relatively uncommon, whereas biotic conversion of TCE 

to 12DCE occurs at many chlorinated solvent sites 

(Maymó-Gatell and others, 1995; Fennell and others, 2001). 

Complete dechlorination of TCE and PCE is more commonly observed in the field or in microcosms and reactors 

that contain mixed cultures than in experiments with pure 

cultures (Flynn and others, 2000). Pure cultures of dehalorespiring bacteria that can use PCE and TCE as terminal electron acceptors include Dehalospirillum multivorans, 

Dehalobacter restrictus strains PER-K23A and TEA, 

Desulfuromonas sp. strain BB1 and Desulfuromonas chloroethenica, Enterobacter sp. strain MS1, Desulfitobacterium 

sp. strain PCE-S, and Dehalococcoides ethenogenes strain 

195 (Löffler and others, 2000). Dehalococcoides ethenogenes strain 195 is the only known isolate, however, that 

completely dechlorinates PCE or TCE to the non-chlorinated 

end product ethene (Maymó-Gatell and others, 1997). 

Although Dehalococcoides ethenogenes strain 195 achieves 

complete reduction of PCE and TCE, the reduction of VC 

does not support its growth. Complete dechlorination to 

ethene could be sustained only by periodic additions of 

filtered supernatant from the mixed culture of which it was 
10 TeCA Degradation and Microbial Communities in a Freshwater Tidal Wetland, APG, Md.

isolated originally (Maymó-Gatell and others, 1995, 1997, 

2001), implying that mixed cultures (or a consortium) are 

required to achieve complete dechlorination. The accumulation of VC that commonly is observed during anaerobic 

reduction of chlorinated ethenes in field studies and laboratory experiments may indicate that one or more critical 

microorganisms are lacking.

In addition to reduction of VC, anaerobic oxidation or 

mineralization of VC to carbon dioxide (CO2 ) or to CO2 and 

methane (CH4 ) has been reported in laboratory experiments 

under iron-reducing, sulfate-reducing, humic acid-reducing, 

and methanogenic conditions (Vogel and McCarty, 1985; 

Bradley and Chapelle, 1996, 1998, 1999a; Bradley and 

others, 1998). 12DCE also can undergo anaerobic oxidation, 

but at substantially slower rates than those observed for 

VC (Bradley and others, 1998). Recent experiments with 

organic-rich streambed sediments incubated with 14C-radiolabeled VC showed the transient presence of 14C-acetate as 

an intermediate of VC degradation under methanogenic conditions, and the production of approximately equal molar 

concentrations of CO2 and CH4 (Bradley and Chapelle, 

1999b, 2000). Furthermore, although addition of an inhibitor of methanogenesis (BES, or 2-bromoethanesulfonic acid) 

caused a decrease in VC degradation rate, VC degradation 

was not stopped completely. These results led Bradley and 

Chapelle (2000) to propose that VC mineralization to CO2

and CH4 involves two steps: (1) oxidation of VC by acetogens, which produces acetate, (2) and metabolism of acetate 

to CO2 and CH4 by acetotrophic methanogens. This theory 

proposes that methanogens are indirectly, rather than 

directly, involved in the anaerobic mineralization of VC. 

The relative importance of reductive dechlorination reactions (either cometabolic or dehalorespiratory) and of anaerobic oxidation reactions in controlling the fate of VC and 

other chlorinated daughter products in the environment is 

unknown.

Acknowledgments

The authors thank the following people and agencies for 

their contributions to this study. John Wrobel, U.S. Army 

Environmental Conservation and Restoration Division at 

APG, and Allison O’Brien and Amy Burgess, contractors for 

the U.S. Army Environmental Conservation and Restoration 

Division, are thanked for their support throughout this 

project. Although this study was predominantly funded 

through the U.S. Army Environmental Conservation and 

Restoration Division at APG, the installation and sampling 

of multi-level wells in the study area and the 2000 laboratory 

microcosm experiment were part of another study (Natural 

Attenuation of Chlorinated Solvents in Wetlands) that was 

funded through the DoD Environmental Security Technology Certification Program (ESTCP). Enrichment experiments also were funded partially by the ESTCP project and 

the USGS National Research Program.

Alan Rosencrance and Thomas Shedd of the U.S. Army 

Center for Environmental Health Research, Fort Detrick, 

Maryland, have assisted this study for many years by 

maintaining the analytical chemistry equipment and 

on-site laboratory facility. We thank Lisa D. Olsen and 

Tracey S. Spencer (USGS, Baltimore, Maryland) for performing VOC analyses of microcosm and ground-water 

samples in the on-site laboratory facility. Linda Jo Dyer, 

Mastin M. Mount, and Daniel J. Phelan (USGS, Baltimore, 

Maryland) assisted in microcosm preparation, microcosm 

and ground-water sample collection, and chemical analyses 

over the course of this study. In addition, the following personnel from (or formerly from) the U.S. Air Force Research 

Laboratory, Tyndall AFB, Florida assisted in the ESTCPsupported ground-water sampling at APG: David R. Burris 

(currently with Integrated Science and Technology, Panama 

City, Florida), Christopher P. Antworth, Eila Burr, Bob 

Nichols, Lt. David Kampisty, and Marlene Cantrell. Amy 

Deal, a student working with Mary Voytek (USGS, Reston, 

Virginia), conducted some of the October 2000 microcosm 

experiments (iron-amended microcosms and microcosms 

with sediment from site WB23). The authors also thank the 

Publications Section of the USGS, Baltimore, Maryland for 

their support in publishing this report, especially Timothy 

W. Auer for illustrations.

Methods and Data Analysis

Anaerobic laboratory microcosm experiments to examine TeCA degradation were conducted between 1998 and 

2000 (February–April 1998, October 1998, March–April 

1999, July–August 1999, and October–November 2000). A 

previously reported experiment (Lorah and others, 1997; 

Lorah and Olsen, 1999a) that was conducted in May–June 

1996 is included for comparisons (table 1). Microcosm 

preparation, incubation, and geochemical analysis techniques generally were the same in the 1998–2000 experiments as those methods reported previously (Lorah and 

others, 1997; Lorah and Olsen, 1999a). The only differences 

in microcosm construction methods from the May–June 

1996 experiment are that wetland sediment was collected 

at a different site and during a different season, and that 

methanol (instead of deionized organic-free water) was used 

in preparing TeCA and 112TCA stock solutions for the 

February–April 1998 experiment (table 1). The February–

April 1998 experiment was designed to test the effect of different initial concentrations of TeCA, the presence of sulfate 

as an electron acceptor, and the inhibition of methanogenesis 

on TeCA degradation rates and pathways. The 1999 and 

2000 experiments were designed to evaluate the spatial and 

seasonal variability in TeCA degradation rates and pathways. 

In the October 1998 and October–November 2000 experiments, treatments also were prepared with and without addition of methanol (MeOH) to evaluate the effect of the added 

MeOH (table 1). Enrichment experiments discussed in this 

report were conducted in January–June 2001.

Anaerobic microcosm experiments and geochemical 

analyses were performed by USGS personnel in Baltimore, 

Maryland, whereas enrichment experiments and all micro- 
Methods and Data Analysis 11

 

Table 1. Treatments used in anaerobic microcosm experiments to examine 1,1,2,2-tetrachloroethane or 

1,1,2-trichloroethane degradation in wetland sediments from the West Branch Canal Creek 

study area, Aberdeen Proving Ground, Maryland, May 1996–November 2000

 [Microcosms were amended with 1,1,2,2-tetrachloroethane (TeCA) or 1,1,2-trichloroethane (TCA) at day 0. Treatment names beginning 

with “23,” “30,” or “Bck” were constructed with sediment from site WB23, WB30, or background site WB19, respectively. Treatment 

names containing “LC” indicate live unamended controls. Killed controls were prepared for all treatments, except 23BES-TeCA.2, by 

adding 1 percent formaldehyde by volume. Initial concentrations (Ci

) of TeCA or TCA, which are given in micromolar (μM) followed (in 

parentheses) by micrograms per liter (μg/L), are the highest concentration detected in solution during days 1 to 3 (Ti

). Other amendments 

included 4 millimolar (mM) methanol (MeOH), used in preparing stock solutions of TeCA or TCA in some experiments; 11 mM sulfate

(SO4); and 30 mM 2-bromoethanesulfonic acid (BES) added on day 5. Dashes indicate no amendment.]

Treatment

name

Ci TeCA 

[μM (μg/L)]

Ci TCA

[μM (μg/L)]

Ti

(day)

Other

amendments

May–June 1996:

30E4 1.2 (210) -- 1 --

February–April 1998:

23TeCA.1 1.4 (240) -- 2 MeOH

23TeCA.2 3.6 (610) -- 3 MeOH

23TeCA.3 13 (2200) -- 1 MeOH

Bck-TeCA.2 3.3 (550) -- 2 MeOH

23SO4-TeCA.2 3.1 (520) -- 2 MeOH; SO4

23BES-TeCA.2 3.2 (540) -- 1 MeOH; BES

23TCA.1 -- 2.4 (320) 1 MeOH

23TCA.2 -- 6.3 (840) 1 MeOH

23LC -- -- -- --

Water control 3.3 (550) 3.9 (520) 0 MeOH

October 1998:

23TeCA.2 2.8 (480) -- 1 MeOH

23TeCA.2 2.0 (340) -- 1 --

23TCA.1 -- 1.6 (220) 1 MeOH

23TCA.1 -- 1.9 (250) 1 --

March–April 1999:

23TeCA.3/99 2.8 (470) -- 1 --

23TeCA5.3/99 3.0 (500) -- 1 --

23TCA.3/99 -- 3.2 (430) 1 --

23LC.3/99 -- -- 1 --

30TeCA.3/99 2.2 (370) -- 1 --

30TCA.3/99 -- 3.1 (410) 1 --

30LC.3/99 -- -- 1 --

Water Control 3.2 (540) 3.4 (450) 2 --

July–August 1999:

23TeCA.7/99 4.3 (720) -- 3 --

23TCA.7/99 -- 2.0 (270) 1 --

23LC.7/99 -- -- 1 --

30TeCA.7/99 4.0 (670) -- 3 --

Water Control 5.6 (940) 4.6 (620) 0 --

October-November 2000:

23TeCA.10/00 2.0 (340) -- 1 --

30TeCA.10/00 2.5 (420) -- 3 --

30MeOH.10/00 3.1 (520) -- 1 MeOH

30TCA.10/00 -- 2.6 (350) 1 --

Water Control 4.8 (800) 1.6 (330) 0 --
12 TeCA Degradation and Microbial Communities in a Freshwater Tidal Wetland, APG, Md.

bial community analyses were performed by USGS National 

Research Program personnel in Reston, Virginia. Groundwater-sampling methods for selected chemical data that are 

used in this report for comparison to laboratory data were 

published in Spencer and others (2002). The same analytical 

techniques were used to determine geochemical constituents 

in water samples collected from the laboratory microcosms 

and in ground-water samples collected in the field. The 

same microbial community analytical techniques also were 

used for sediment samples collected from the laboratory 

microcosms and from the field. These analytical techniques, 

therefore, are described only in the first section on laboratory 

microcosm experiments.

Laboratory Microcosm Experiments

Sediment and Ground-Water Collection To evaluate 

spatial and seasonal effects on biodegradation pathways and 

rates, microcosms were constructed with shallow wetland 

sediment collected during three seasons (late winter/early 

spring samples in March 1999, summer samples in July 

1999, and fall samples in October 2000) and from two different contaminated sites (near piezometer sites WB23 and 

WB30 (fig. 1). Sites WB23 and WB30 are separated horizontally by a distance of about 9 m (fig. 1). As part of an 

effort to evaluate the possible effect of pre-exposure to the 

contaminants on degradation, sediment also was collected 

from the upper peat unit at background site WB19 (fig. 1) for 

one microcosm experiment in February–April 1998 (table 

1). Measurements of dissolved oxygen, ferrous and ferric 

iron, sulfide, and methane indicate that the upper peat unit of 

the wetland sediment at and near sites WB23, WB30, and 

WB19 is methanogenic or mixed iron-reducing and methanogenic (Lorah and others, 1997; Olsen and others, 1997). 

The total organic carbon content in sediment samples collected in the upper 30 cm (centimeters) at or near these two 

sites was between 6.9 and 10.6 percent (Olsen and others, 

1997). Wetland sediment from the upper peat unit at both 

sites was collected from a depth of 0 to 25 cm below land 

surface. The sediment was placed in mason jars and packed 

to leave minimum airspace.

Wetland porewater for the microcosms constructed 

with WB23 and WB30 sediment was collected from a nearby 

contaminated piezometer, WB24B (figs. 1 and 3). The 

chemistry of water from piezometer WB24B is similar to 

that from shallow piezometers at WB23 and WB30, but piezometer WB24B has a more rapid recovery rate during sampling. For microcosms constructed with sediment from 

background site WB19, wetland porewater was collected 

from piezometer WB19B. Porewater was collected using 

a peristaltic pump with tubing extended to the top of the 

screened interval. No effort was made to prevent loss of 

VOCs from the water during sample collection, and the 

water was purged with nitrogen gas immediately prior to 

microcosm construction to remove oxygen that potentially 

was added during sample collection. Sediment and porewater for the microcosms were collected during the week prior 

to microcosm construction and then were stored at the 

planned incubation temperature.

Microcosm Preparation and Incubation All microcosms 

were constructed under a nitrogen atmosphere in 162-mL 

(milliliter) serum bottles using a 1.5:1 volumetric ratio of 

ground water to wetland sediment (with no headspace) and 

incubated upside down in the dark. All microcosms were 

incubated at the summer temperature of 19 °C (Lorah and 

others, 1997), although an additional set of microcosms in 

the March–April 1999 experiment was incubated at the winter temperature of 5 °C.

Duplicate microcosm bottles were prepared for each 

treatment and sacrificed for chemical and microbiological 

analyses at each time step. Microcosms generally were 

incubated for 35 to 42 days and sampled at 9 or 11 time steps 

during this period. One exception was the October 1998 

microcosms, which were designed solely to observe the 

effect of MeOH addition, and were incubated for only 15 to 

20 days and sampled at 5 or 6 time steps (Lorah-Devereux, 

1999). Experiments included killed controls that were prepared with 1 percent by volume of formaldehyde for each 

treatment. Unamended live controls that did not contain 

added TeCA or 112TCA also were prepared for each sediment (WB23 or WB30) used to construct microcosms. 

Water controls (microcosms containing only deionized water 

and amended with TeCA or 112TCA) were constructed to 

account for any potential losses due to volatilization or to 

sorption to the serum bottle or Teflon-coated rubber stoppers. Stock TeCA and 112TCA solutions were made from 

neat chemicals (Supelco, Bellefonte, Pennsylvania) and 

added to microcosms using gas-tight syringes. Stock solutions that were prepared using only deionized water were 

placed in the dark on magnetic stir plates for 24 hr before 

spiking the microcosms. For stock solutions mixed in a flask 

with 50 percent water and 50 percent MeOH (table 1), the 

stock solutions were stirred for 4 hr before spiking. Aliquots 

of the stock solutions were analyzed before addition to the 

microcosms to estimate the injection volumes needed to 

obtain the planned initial concentrations at day 1. Because 

of expected losses due to sorption during the first 24 hr 

(Lorah and others, 1997; Lorah and Olsen, 1999a), the 

microcosms were spiked on day 0 with a concentration about 

2.4 times greater than the desired initial (day 1) aqueous 

concentration. Most microcosms were expected to reach 

the wetland sediments’ natural methanogenic conditions 

within several days of incubation, but high sulfate concentrations (11 mM, or millimolar) were added in one treatment to 

induce sulfate-reducing conditions (table 1). In addition, 

BES, a known specific inhibitor of methanogenesis (Oremland and Capone, 1988), was added to one set of TeCAamended microcosms (table 1).

Geochemical and Microbial Community Analyses At 

each time step, supernatant from duplicate microcosm bottles was withdrawn for analyses of pH, VOCs, ferrous iron, 

sulfate, and CH4 by the same methods as previously reported 

(Lorah and others, 1997; Lorah and Olsen, 1999a, b). The 

initial pH in all microcosms ranged from 6.2 to 6.6, which is 

in the range of pH measured in the wetland porewater in the 

field (Lorah and others, 1997; Lorah and Olsen, 1999b), and 
Methods and Data Analysis 13

increased to as high as 7.2 at the end of the experiments. 

VOCs were analyzed at the on-site laboratory facility at 

APG using a purge and trap capillary gas chromatography/

mass spectrometry (GC/MS) method that is equivalent to 

U.S. Environmental Protection Agency Method 524.2 (Rose 

and Schroeder, 1995). Average relative percent differences 

for analyses of replicate VOC samples ranged from 5 to 

13 percent for TeCA and its potential daughter compounds 

(Spencer and others, 2002). Samples for ferrous iron were 

filtered through 0.2-μm (micrometer) filters and analyzed 

on-site by the colorimetric bipyridine technique. Sulfate was 

determined on 0.45-μm filtered samples by ion chromatography. Methane was determined by collecting unfiltered water 

in sealed serum bottles and analyzing the headspace by gas 

chromatography with a flame ionization detector (GC/FID) 

at the on-site laboratory. Methane concentration in the water 

then was calculated using the ideal gas law and Henry’s Law 

constant (Baedecker and Cozzarelli, 1992).

After supernatant was removed for chemical analyses at 

each time interval, the remaining sediment slurry from the 

microcosms was transferred to plastic bags and immediately 

frozen for later microbial analyses. For most experiments, 

sediment samples from the sites used in microcosm construction also were collected in the field and immediately 

frozen for microbial analyses. The bacterial and methanogen communities from TeCA- or 112TCA-amended microcosm slurries were characterized using a polymerase chain 

reaction (PCR) procedure of amplifying specific targeted 

DNA and performing terminal restriction fragment length 

polymorphism (TRFLP) analysis on the PCR products 

(Clement and others, 1998). TRFLP provides a molecularly 

derived microbial "fingerprint" that provides a relative estimate of the total microbial abundance and of the diversity of 

bacterial and methanogen communities. In order to evaluate 

bacterial communities, 16S rDNA was targeted for analysis 

because it codes for a molecule present in all bacteria. For 

microcosm samples from the March–April 1999 experiment, 

a newly available nested PCR methodology also was used 

that targeted specific dehalorespiratory bacteria (Löffler and 

others, 2000). The two primer pairs are specific to hypervariable regions of the 16S rRNA genes of the Dehalococcoides group (Dehalococcoides ethenogenes and Dehalococcoides sp. strain FL2) and of the acetate-oxidizing Desulfuromonas group (Desulfuromonas sp. strain BB1 and Desulfuromonas chloroethenica) (Löffler and others, 2000).

To characterize the methanogenic community, a key 

functional gene involved in methanogenesis, mcrA, was 

amplified and analyzed. The gene mcrA codes for the 

enzyme methylcoreductase that is involved in the final reaction in methanogenesis, which is release of CH4 after reduction of a methyl group bound to a coenzyme (Klein and 

others, 1988). The size of the PCR mcrA products (464 and 

481 base pairs, respectively) differs between the Methanosarcinaceae, the only group that contains acetate-utilizing 

methanogens (acetoclasts), and the other main methanogen 

groups Methanococcales, Methanobacteriales, and Methanomicrobiales. Because of this fragment length heterogeneity, the proportion of Methanosarcinaceae in the total 

methanogen community can be determined to evaluate the 

potential importance of acetoclasts in the microcosm experiments.

To obtain the TRFLP profiles, DNA was extracted from 

the same amount of sediment for all samples, re-suspended 

in the same volume of buffer, and the same volume was 

subsequently used to amplify and digest the samples. Sample TRFLP profiles, therefore, were normalized to equal 

amounts of sediment. DNA was extracted from microcosm 

slurries or surficial sediments using the Bio101 FastDNA 

spin kit for soil according to manufacturer’s instructions 

(Bio101 Inc., Vista, California). PCR was performed to 

amplify a portion of the bacterial 16S rDNA using the primers 46 forward with a fluorescent tag (FAM) and 519 reverse 

(Brunk and others, 1996) at a concentration of 0.2 μΜ. PCR 

conditions were as follows: one cycle of 94 °C for 5 minutes 

followed by 30 cycles of 94 °C for 30 seconds, 56 °C for 

30 seconds, 72 °C for 90 seconds, and a final extension for 

7 minutes at 72 °C. McrA DNA amplifications were performed using the degenerate mcrA oligonucleotide primer 

pair described by Luton (1996) at 0.2 μM, except the forward primer was labeled with the fluorescent tag FAM. PCR 

conditions were as follows: one cycle of 94 °C for 5 minutes 

followed by 38 cycles of 94 °C for 45 seconds, 56 °C for 45 

seconds, and 72 °C for 2 minutes, with the first four cycles 

employing a slow (50 percent) ramp between the 56°C and 

72 °C incubation.

After amplification, bacterial 16S rDNA amplicons were 

digested with the restriction enzyme MnlI, whereas mcrA 

amplicons were digested with RsaI (New England Biolabs, 

Beverly, Massachusetts) at 37 °C overnight. Digests were 

precipitated in ethanol, centrifuged, dried, and resuspended 

in Tris-EDTA buffer. Samples were electrophoresed on 

an ABI310 sequencer (Applied Biosystems, Foster City, 

California), generating a plot called an electropherogram of 

the size (using internal standards TAMRA 500) and the relative intensity of each fluorescent fragment. Undigested 

mcrA amplicons also were analyzed to determine the proportion of Methanosarcinaceae in the methanogen population on 

the basis of fragment length heterogeneity. Biomass in the 

microcosm slurry was estimated from total DNA extracted, 

the intensity of PCR amplifications, and TRFLP peak intensities.

Calculation of Degradation Rates Biodegradation rates 

were calculated assuming first-order kinetics, as in earlier 

experiments (Lorah and others, 1997). The apparent biodegradation rate constant, k, was determined from the slope of a 

linear regression of the natural logarithm of the aqueous 

TeCA or 112TCA concentration [ln(TeCA) or ln(112TCA)] 

remaining in the live microcosms against time of incubation. 

The rate constants calculated for the killed controls were 

subtracted from the rate constants in the live microcosm if 

they were greater than the standard error calculated for the 

rate constants in the live microcosms. Biotic daughter 

products generally were not detected in the killed controls 

during the time periods used to calculate rate constants. 
14 TeCA Degradation and Microbial Communities in a Freshwater Tidal Wetland, APG, Md.

After the rate constant k was estimated, biodegradation rates 

of TeCA or 112TCA (units of μM/day) were calculated by 

multiplying k (units of per day) by the initial TeCA or 

112TCA concentration (units of μM) in the experiment.

Laboratory Enrichment Experiments

Enrichment Preparation and Incubation Two types of 

enrichment experiments were done: (1) enrichment culture 

experiments that were used to select for members of the 

microbial community with the ability to produce VC, and 

(2) VC/sediment enrichments that were used to study the 

potential of the sediment to degrade VC under various 

geochemical and physiological conditions (table 2). For the 

enrichment culture experiments, sediment slurry from the 

October–November 2000 microcosms that were amended 

with TeCA (no MeOH added) was used as the inoculum to 

examine VC production from different precursors (table 2). 

After sampling the microcosm bottles on day 39, which 

required removal of about 7 percent of the liquid, the microcosm bottles were resealed with an anaerobic headspace of 

95 percent nitrogen and 5 percent carbon dioxide and preincubated for an additional 6 weeks at room temperature 

(microcosms were methanogenic and depleted of VOCs by 

this time). After this preincubation, 10 mL of microcosm 

slurry from WB23 and WB30 treatments was added to 

100 mL of a bicarbonate-buffered (pH of 6.8) anaerobic 

enrichment medium in a 160-mL serum bottle. The remaining 50 mL of headspace in the bottles initially contained 

80 percent nitrogen and 20 percent carbon dioxide. The 

enrichment medium contained nutrients and minerals 

(500 mg/L of NH4Cl, 600 mg/L of NaH2PO4, and 100 mg/

L of KCl), a suite of vitamins and trace minerals, yeast 

extract (50 mg/L), and cysteine HCl (250 mg/L) as a reductant. Each enrichment culture was amended with one of 

three potential precursors of VC (112TCA, cDCE, and 

tDCE) as the electron acceptor (table 2). These compounds 

were added as neat liquids. The enrichment cultures were 

given a non-fermentable electron donor of either acetate (5 

mM) or hydrogen (H2) (10mL overpressure in the headspace). Bottles were inverted, incubated at room temperature, and a small subsample of the headspace was removed 

periodically to monitor VOC concentrations.

 Three sets of VC/sediment enrichment experiments 

were conducted (table 2). The first two VC/sediment enrichments were designed to evaluate the effects of redox condition and spatial variability in the wetland sediment on VC 

degradation, whereas the third experiment was designed to 

evaluate the effect of pre-exposure of the sediment to VOCs 

on VC degradation (table 2). The concentration of VC provided to the enrichments represented a concentration about 

7 times higher than that observed during the peak accumulation of VC in the microcosms. The first VC/sediment 

enrichment was constructed using sediment slurries from the 

TeCA-amended (no MeOH added) microcosms constructed 

in October–November 2000. The microcosm slurries were 

preincubated as described above to deplete sulfate, iron, and 

VOCs. After preincubation, sediment slurry (10 mL) was 

placed in a pressure tube (28 mL), and 1 mL of 1 percent VC 

gas was added. Some treatments also received ferric iron 

(Fe(III)) in the form of amorphous ferric oxyhydroxide 

(FeOOH) (1 mL of a suspension resulting in a final addition 

of 10 mM Fe(III)) to induce iron-reducing conditions or BES 

(20 or 50 mM) to inhibit methanogenesis. No excess electron donor (such as acetate or H2) was added. Tubes were 

incubated on their sides in order to maximize the surface 

area for exchange of VC across the interface between the 

headspace and the sediment slurry. The enrichments were 

analyzed for VC, Fe(III), and CH4 concentrations.

For the second VC/sediment enrichment, a test of the 

potential effect of iron-reducing conditions on VC degradation was repeated, this time using a soluble chelated form of 

ferric iron (Fe(III)-NTA) that is reported to be more readily 

available to iron-reducing bacteria than FeOOH (table 2) 

(Bradley and Chapelle, 1996). The second VC/sediment 

enrichment was prepared transferring 3 mL of proven VC 

degrading sediment from site WB23 into three tubes. The 

tubes were resealed, and the headspace was purged with 

nitrogen before amending again with VC.

In April 2001, the third VC/sediment enrichment experiment was performed to evaluate the effect of pre-exposure to 

TeCA on VC degradation (table 2). Fresh sediment was collected from sites WB23 and WB30 to prepare three sets of 

VC/sediment enrichments for this experiment (table 2). One 

set of VC/sediment enrichments was amended with VC 

5 days after collection, and this sediment was referred to as 

“fresh.” A second set of VC/sediment enrichments, referred 

to as “pre-exposed,” was preincubated for 42 days with 

TeCA (7.1 μM added at day 0 to give an initial aqueous concentration of about 3.0 μM), using the same methods to prepare and incubate anaerobic microcosms as reported in the 

preceding section on Laboratory Microcosm Experiments. 

A third set of VC/sediment enrichments, referred to as “preincubated,” also was preincubated for 42 days but was not 

amended with TeCA or any other compounds. Each set of 

VC/sediment enrichments was tested for VC degradation 

(initial VC concentration of 44.6 μM) in triplicate tubes 

of three treatments: (1) no amendments other than VC; 

(2) amended with amorphous FeOOH; and (3) amended with 

BES (table 2). In addition, VC was added to killed controls 

to monitor abiotic loss of VC. The killed controls for the 

“fresh” experiment were poisoned with 1 percent formaldehyde. Sediment for the “preincubated” and “pre-exposed” 

killed controls was autoclaved three times on 3 consecutive 

days for 1 hr each. Individual tubes were sacrificed over 

time to obtain geochemical and microbial samples.

Geochemical and Microbial Community Analyses In all 

the enrichment experiments, VC and methane were analyzed 

by injecting 0.3 mL subsample of the headspace onto a 

packed column and measuring with a GC/FID at the USGS 

in Reston, Virginia. For the April 2001 enrichment experiments, the anaerobic microcosms that were constructed prior 

to preparation of the preincubated and pre-exposed enrichment cultures were monitored periodically for VOCs by 

GC/MS analysis at the on-site laboratory at APG as reported 

in the preceding section for other microcosm experiments. 
Methods and Data Analysis 15

 

Ferrous and ferric iron concentrations available for microbial 

reduction in the sediment slurries were monitored in the 

April 2001 microcosms using the extraction and ferrozine 

assay technique detailed in Lovley and Phillips (1987). 

Sediment slurry samples were collected at selected time 

intervals for all enrichment experiments for molecular analysis of the microbial communities (as detailed in the preceding section on Laboratory Microcosm Experiments); 

however, these analyses have not been completed yet.

Field Data Collection and Analyses

Surficial Wetland Sediment Samples To evaluate spatial 

and seasonal variability, grab samples of the surficial wetland sediment were collected from 0 to 7.6 cm below land 

surface near piezometer sites along the A-A′ and C-C′

transects in March and August 1999. The sediment samples 

were frozen immediately and analyzed later for microbial 

communities as detailed previously in the section on Laboratory Microcosm Experiments. Biomass was estimated from 

the TRFLP procedure and also from direct cell counts on 

stained slides. Cores of the wetland sediment also were 

Table 2. Treatments used in enrichment experiments with anaerobic daughter products of

1,1,2,2-tetrachloroethane (TeCA) degradation

 [112TCA, 1,1,2-trichloroethane; cDCE, cis-1,2-dichloroethene; tDCE, trans-1,2-dichloroethene; VC, vinyl chloride; FeOOH, ferric 

oxyhydroxide; Fe(III)-NTA; BES, 2-bromoethanesulfonic acid. Concentrations given in millimolar (mM) or micromolar (μM)]

Sediment Daughter compound added Other amendments

Enrichment Cultures (October–November 2000 Microcosm Slurry)

WB23 (microcosm 23TeCA.10/00) 112TCA (10.8 mM), cDCE (8 mM), or tDCE (14.9 mM) acetate (5 mM)

WB23 (microcosm 23TeCA.10/00) 112TCA (10.8 mM), cDCE (8 mM), or tDCE (14.9 mM) hydrogen

WB30 (microcosm 30TeCA.10/00) 112TCA (10.8 mM), cDCE (8 mM), or tDCE (14.9 mM) acetate (5 mM)

WB30 (microcosm 30TeCA.10/00) 112TCA (10.8 mM), cDCE (8 mM), or tDCE (14.9 mM) hydrogen

VC/Sediment Enrichment Experiment 1 (October–November 2000 Microcosm Slurry)

WB23 (microcosm 23TeCA.10/00) VC (44.6 μM) none

WB23 (microcosm 23TeCA.10/00) VC (44.6 μM) amorphous FeOOH

WB23 (microcosm 23TeCA.10/00) VC (44.6 μM) BES (20 mM)

WB30 (microcosm 30TeCA.10/00) VC (44.6 μM) none

WB30 (microcosm 30TeCA.10/00) VC (44.6 μM) amorphous FeOOH

WB30 (microcosm 30TeCA.10/00) VC (44.6 μM) BES (50 mM)

VC/Sediment Enrichment Experiment 2 (October–November 2000 Microcosm Slurry)

WB23 from Enrichment 1 VC (44.6 μM) none

WB23 from Enrichment 1 VC (44.6 μM) Fe(III)-NTA

WB23 from Enrichment 1 VC (44.6 μM) BES (20 mM)

VC/Sediment Enrichment Experiment 3 (April 2001 Microcosm Slurry)

WB23, fresh, preincubated, and pre-exposed VC (44.6 μM) none

WB23, fresh, preincubated, and pre-exposed VC (44.6 μM) amorphous FeOOH

WB23, fresh, preincubated, and pre-exposed VC (44.6 μM) BES (20 mM)

WB30, fresh, preincubated, and pre-exposed VC (44.6 μM) none

WB30, fresh, preincubated, and pre-exposed VC (44.6 μM) amorphous FeOOH

WB30, fresh, preincubated, and pre-exposed VC (44.6 μM) BES (20 mM)
16 TeCA Degradation and Microbial Communities in a Freshwater Tidal Wetland, APG, Md.

collected at sites WB23 and WB30 (fig. 1) in August 2000 

by hammering a 1.2 m-long, 0.076-m-diameter polyvinyl 

chloride (PVC) pipe with a clear plastic liner into the sediment. Depths to the top of the sediment core were measured 

every 30 cm to determine the amount of sediment compaction. The PVC pipes then were capped before manually 

pulling the PVC pipe from the sediment. Sediment samples 

from selected depths in these cores were analyzed at the 

USGS laboratory in Reston, Virginia for bioavailable ferrous 

and ferric iron concentrations using the extraction and 

ferrozine assay technique (Lovley and Phillips, 1987).

Ground-Water Samples Ground-water-quality data collected during 1995 and 2000 from selected piezometers and 

multi-level samplers are used in this report to estimate contaminant degradation rates and to compare daughter product 

distributions and redox characteristics in the laboratory 

microcosms to field observations. Sampling and analytical 

methods for the 1995 data are reported in Lorah and others 

(1997) and Olsen and others (1997). Sampling and analytical methods for the 2000 data are reported in Spencer and 

others (2002). The analytical techniques used in 1995 were 

the same as those used in 2000.

Calculation of Degradation Rates TCE degradation rates 

that were estimated using field data collected along or near 

section A-A′ during June–October 1995 were reported previously in Lorah and others (2001). For this report, TeCA 

degradation rates were calculated for the same sites using the 

data reported in Lorah and others (1997) and Olsen and 

others (1997), and the same rate estimation method for TCE 

reported in Lorah and others (2001). Data from the multilevel sampler at site WB30 that were collected in June–July 

2000 (Spencer and others, 2002) also are used to calculate 

TeCA and TCE degradation rates using the same estimation 

method. Field rates of TeCA and TCE degradation were 

estimated using TeCA, TCE, and chloride concentrations 

along upward flowpaths from the shallow aquifer to the top 

of the wetland surface. At sites selected for rate calculations, 

there were sufficient screened intervals and detectable TeCA 

and TCE concentrations in the wetland porewater to give at 

least two points (usually at least three points) along the flowpath to estimate rates. Measured TeCA and TCE concentrations were corrected for the effects of advection, dispersion, 

and dilution by normalization with the conservative tracer 

chloride, using the correction method outlined by Wiedemeier and others (1998). Ravi and others (1998) found 

excellent agreement between rate constants estimated with 

this tracer-corrected method and a method that utilizes a onedimensional analytical model. Because TCE degradation 

rates estimated using both chloride and bromide as conservative constituents at the same sites were essentially the same 

(Lorah and others, 2001), only chloride was used in this 

study. Because chloride concentrations in the aquifer are in 

the range of 50 to 100 mg/L and the VOC concentrations are 

in the μg/L range (Lorah and others, 1997; 2001), the effect 

of dechlorination reactions on the chloride concentrations 

would be negligible.

The tracer-corrected rates can be assumed to represent 

biodegradation rates if volatilization and sorption effects are 

assumed to be negligible. Volatilization was shown to be a 

negligible transport mechanism for the volatile contaminants 

in the wetland study area by using sulfur hexafluoride as a 

conservative gas in a ground-water tracer test (L.D. Olsen, 

U.S. Geological Survey, written commun., 2002). Sorption 

also is assumed to have a negligible effect on the estimated 

field rates because the plume is believed to be at steady state 

(Lorah and others, 1997; Chapelle and Bradley, 1998). The 

first-order biodegradation rate at each site was estimated by 

linear regression of the natural logarithm of the normalized 

TCE concentration against traveltime along the flowpath. 

The traveltime was calculated using the linear ground-waterflow velocity in the wetland sediments and the travel distance measured between the mid-point of each screened 

interval. The piezometers used for the 1995 calculations 

have 15-cm-long screens, whereas the multi-level sampler 

has 7.5-cm-long screens at each sampling point (Spencer and 

others, 2002). Linear ground-water-flow velocities between 

0.6 and 0.9 m/yr were estimated using flow-net analyses 

along sections A-A′ and C-C′, measured horizontal hydraulic conductivities, and estimated vertical hydraulic conductivities (Lorah and others, 1997). The maximum of this 

estimated velocity range (0.9 m/yr) was selected for use in 

the original field-estimated TCE degradation rates (Lorah 

and others, 2001) and was used in this report for consistent 

comparison. Variations of ground-water-flow velocities in 

time and space because of the heterogeneity of the wetland 

sediments and the seasonal and tidal fluctuations in the 

hydraulic gradient still are large, undefined sources of 

uncertainty.

Laboratory Experiments on Anaerobic 

Degradation of 1,1,2,2-Tetrachloroethane and 

Association with Microbial Communities

The following sections discuss the anaerobic degradation 

of TeCA in the laboratory microcosm and enrichment experiments that were conducted during 1998–2001, and the association between degradation pathways and the microbial 

communities in the microcosm sediment. TeCA degradation 

rates in all the microcosm experiments that were conducted 

at this site, including the previously reported experiment 

(Lorah and others, 1997; Lorah and Olsen, 1999a), are discussed in the first subsection, providing a partial overview 

of the results. Understanding the degradation pathways of 

TeCA requires an understanding of the production and 

degradation of the many possible daughter products (fig. 2). 

Ultimately, the feasibility of natural attenuation as a remediation method for the chlorinated VOC plumes discharging to 

the wetland sediment depends on the fate of these daughter 

products. Discussion of the many laboratory experiments 

that were conducted (tables 1 and 2), therefore, focuses on 

the daughter products and the possible factors that could 

control their distribution and degradation in the wetland 
Laboratory Experiments on Anaerobic Degradation of TeCA and Association with Microbial Communities 17

sediments. Laboratory microcosm and enrichment culture 

experiments that provide information for each factor are 

discussed together and not in chronological order. Some 

experiments are discussed in more than one section because 

the results have a bearing on more than one factor. Results 

of the microcosm and enrichment experiments are presented 

as averages of duplicate samples, unless otherwise noted. 

After the first 1 to 3 days of incubation, reproducibility in the 

microcosm experiments generally was within or slightly 

greater than the relative percent difference of the analysis, 

which ranged from 5 to 13 percent for TeCA and its potential 

daughter compounds.

Degradation Rate of 1,1,2,2-Tetrachloroethane

Rapid removal of TeCA from solution occurred in all live 

(no formaldehyde added) microcosm experiments with wetland sediment and ground water, regardless of the site at 

which the sediment was collected (table 3; figs. 5–10). The 

TeCA biodegradation rate was affected substantially, however, by the addition of MeOH to microcosms and in some 

cases by the season during which the sediment was collected 

(fig. 5a). Apparent first-order biodegradation rate constants, 

which were corrected for losses in killed (formaldehyde 

added) controls to account for abiotic losses such as sorption 

and volatilization, ranged from 0 to 0.50 per day in the 

WB23 microcosms and from 0.06 to 0.63 per day in the 

WB30 microcosms (table 3). Figure 5a shows the initial 

apparent rate of TeCA degradation, which is calculated by 

multiplying the initial first-order rate constant estimated for 

each experiment by the initial substrate (TeCA) concentration (table 3) in each of the microcosm experiments. The 

slope of the best-fit line through the plotted TeCA degradation rates against the initial TeCA concentration (using only 

those experiments that did not contain amendments other 

than TeCA) gives the first-order rate constant that best fits 

the studied range of concentrations (from about 1.0 to 

13.0 μmol/L) (fig. 5a). It should be noted that for many 

experiments, there was a second period of faster TeCA degradation (table 3) after the initial period of degradation that is 

shown in figure 5a. For the July–August 1999 experiment 

with sediment from site WB23, for example, the initial 

TeCA degradation rate (day 3 to 12) was 0. Substantial 

TeCA degradation did occur after the initial period of day 

3 to 12, as evidenced by the production of daughter compounds. A degradation rate was not estimated for this later 

period, however, because a slight rise in TeCA concentrations (that most probably is due to desorption) occurs 

between day 12 and 21 (see fig. 8a). For other experiments, 

a distinct change in slope in plots of the natural log of TeCA 

concentrations against time was observed and a second rate 

was calculated (table 3).

Plotting the initial TeCA degradation rate for each microcosm gives two distinct lines, with the first-order rate constant about two times higher in the MeOH-amended 

microcosms than in those constructed without MeOH 

(fig. 5a). Both data groups show a linear increase in degradation rate with initial TeCA concentration, but the trend is 

more evident in the MeOH-amended microcosms because 

they span a larger concentration range. The 1998 microcosms, which were amended with MeOH in the TeCA stock 

solution, were designed to include three initial TeCA levels, 

whereas the other experiments only targeted one initial concentration level (table 1) that was thought to be most representative of the average ground-water TeCA concentrations 

in the aquifer (Lorah and others, 1997; Lorah and Olsen, 

1999b). The first-order rate constant for TeCA biodegradation in the MeOH-amended microcosms was 0.12 per day, 

corresponding to a half-life of 5.8 days (fig. 5a). Compared 

to the other microcosms that contained MeOH, the TeCA 

degradation rate was relatively high in the microcosm constructed with sediment from the background site WB19, and 

relatively low in the microcosms amended with sulfate and 

BES (fig. 5a). The first-order rate constant of 0.053 per day 

(half-life of 13 days) estimated for the microcosms without 

MeOH (fig. 5a) is more applicable to natural field conditions. It is unknown, however, if this first-order estimate 

is applicable to initial TeCA concentrations above about 

3 μmol/L (500 μg/L) (fig. 5a). The linear increase in biodegradation rate indicates that TeCA is not toxic to the degrading microorganisms up to an initial TeCA concentration of 

13 μmol/L, and that a maximum degradation rate has not 

been reached as described by Monod kinetic theory for utilization of a single rate limiting substrate (fig. 5a). A greater 

range in initial concentrations, however, should be investigated to fully describe TeCA degradation kinetics.

The higher degradation rates with addition of MeOH 

may be due to increased bioavailability of the TeCA because 

MeOH increases the solubility of TeCA and decreases sorption by a cosolvent effect (Schwarzenbach and others, 1993, 

p. 96; Pankow and Cherry, 1996, p. 255). The higher bioavailability and degradation rates with MeOH are indicated 

by the high rebound in daughter compound concentrations 

(especially 12DCA) near the end of experiments 23TeCA.2 

and 23TeCA.3 (figs. 6b and 6c). It is not possible to determine if this effect occurred in experiment 23TeCA.1, which 

also contained added MeOH, because a shorter incubation 

and sampling period was used (fig. 6a). After daughter 

compound concentrations decreased to below detection 

levels by day 40 in experiments 23TeCA.2 and 23TeCA.3, 

12DCA concentrations again increased to between 1.5 and 

2.0 μmol/L by day 45, without any rebound in TeCA concentrations above the detection level (figs. 6b and 6c). These 

concentrations indicate desorption of TeCA and subsequent 

rapid biodegradation. This high of a rebound in concentrations was not evident in the microcosms that were not 

amended with MeOH (figs. 7–9). In addition, the availability of MeOH as a substrate for microorganisms may have 

affected biodegradation rates by altering degradation pathways and composition of the microbial community, as discussed in the section on Daughter Compound Production and 

Degradation.

Seasonally, the lowest initial TeCA biodegradation rate 

was observed in the one set of microcosms conducted during 

the summer (July–August 1999) (table 2; fig. 5a). In the first 

12 days of the July–August 1999 experiment, the TeCA 
18 TeCA Degradation and Microbial Communities in a Freshwater Tidal Wetland, APG, Md.

 

Table 3. First-order rate constants and half-lives for degradation of 1,1,2,2-tetrachloroethane

 (TeCA) or 1,1,2-trichloroethane (112TCA) in anaerobic microcosm experiments

 conducted using wetland sediments, May 1996–November 2000

 [Rate constants (± standard error for a 95-percent confidence interval) are presented for the average of duplicate microcosms for each 

experiment. Ci

 is the concentration (in micromoles per liter, or μmol/L) of TeCA or 112TCA remaining at the first day of the time period 

used to calculate the first-order degradation rate. Concentrations were corrected for losses in killed controls before determining rates, 

except for BES-TeCA.2, which did not have killed controls. Dashes indicate not applicable]

Treatment

name

Ci

(μmol/L)

Rate

constant

(day-1 )

Half-life

(days)

Correlation

 coefficient,

 r2

Time

period

(days)

May–June 1996:

30E4 1.2 0.25 ± 0.03 2.8 0.99 4-8

February–April 1998:

23TeCA.1 1.4 0.50 ± 0.14 1.4 .86 2-8

23TeCA.2 3.6 0.28 ± 0.04 2.5 .98 3-9

23TeCA.3 13 0.16 ± 0.04 4.3 .85 3-16

Bck-TeCA.2 3.3 0.39 ± 0.11 1.8 .86 2-11

23SO4-TeCA.2 3.1 0.25 ± 0.04 2.8 .96 2-10

23BES-TeCA.2 3.2 0.15 ± 0.02 4.6 .95 1-12

23TCA.1 2.4 0.38 ± 0.13 1.8 .82 1-9

23TCA.2 6.3 0.46 ± 0.07 1.5 .91 1-16

October 1998:

23TeCA.2 (methanol) 2.8 0.36 ± 0.03 1.9 .98 2-11

23TeCA.2 (no methanol) 2.0 0.16 ± 0.02 4.3 .96 2-14

23TCA.1 (methanol) 1.6 0.58 ± 0.05 1.2 .99 2-8

23TCA.1 (no methanol) 1.9 0.22 ± 0.03 3.2 .95 2-14

March–April 1999:

23TeCA.3/99 2.8 0.15 ± 0.02 4.6 .98 1-9

.73 0.41 ± 0.02 1.7 .99 12-25

30TeCA.3/99 2.2 0.15 ± 0.02 4.6 .92 1-12

.39 0.63 ± 0.02 1.1 1.0 16-25

23TeCA.3/99 3.2 0.22 ± 0.02 3.2 .93 1-35

30TeCA.3/99 2.8 0.32 ± 0.01 2.2 .99 16-25

July–August 1999:

23TeCA.7/99 4.3 0 ± 0.03 A -- -- 3-12

30TeCA.7/99 4.0 0.06 ± 0.03 12 .96 3-9

1.9 0.55 ± 0.17 1.3 .91 21-30

23TCA.7/99 2.4 0.13 ± 0.03 5.3 .92 21-35

October–November 2000:

23TeCA.10/00 2.0 0.16 ± 0.05 4.3 .83 1-19

30TeCA.10/00 2.5 0.10 ± 0.01 6.9 .98 3-10

1.3 0.24 ± 0.01 2.9 .99 10-21

30MeOH.10/00 3.1 0.10 ± 0.05 6.9 .94 1-6

1.9 0.39 ± 0.05 1.8 1.0 6-10

30TCA.10/00 2.6 0.07 ± 0.02 9.6 .85 1-10

1.3 0.27 ± 0.003 2.6 1.0 10-18

A Substantial TeCA degradation does occur after the initial period of day 3 to 12, as evidenced by the production of daughter compounds. A degradation rate 

was not estimated, however, because a slight rise in TeCA concentrations (most likely caused by desorption) occurs between day 12 and 21 (see fig. 8a).
Laboratory Experiments on Anaerobic Degradation of TeCA and Association with Microbial Communities 19

 

biodegradation rate in the WB23 microcosm was insignificant after the rate of TeCA loss in the killed controls was 

subtracted from that of the live microcosms (the difference 

was within the standard error for the rate constant of 0.02 per 

day). The lower concentrations of biotic daughter products 

(all except TCE) in the first 12 days of the July–August 1999 

experiments compared to the March–April 1999 experiments 

also shows the lower TeCA degradation rate in the July–

August 1999 experiments (figs. 7 and 8). The similar loss of 

TeCA in the killed and live microcosms for the July–August 

1999 experiments constructed from WB23 and WB30 sediments is evident in figure 8. For the WB30 microcosms in 

July–August 1999, however, the initial TeCA biodegradation 

rate was significant compared to loss in the killed controls, 

although the rate is relatively low compared to the other 

microcosms (table 3 and fig. 5a).

Comparison of the decrease in TeCA concentrations in 

the live microcosms to the killed controls indicates that 

abiotic processes caused substantial decrease in TeCA concentrations at times (figs. 6–9). During the first 4 to 7 days 

of incubation, some loss of TeCA often was observed in the 

killed microcosms without concurrent production of biotic 

daughter products and without corresponding losses in the 

water controls. Sorption to the organic-rich wetland sediments is the likely cause of most of this abiotic loss of 

 

TeCA in the microcosms. Volatilization from the microcosm 

bottle or sorption to the microcosm bottle or stopper was 

determined to be insignificant because losses in water 

controls, which were constructed with most experiments 

(table 1), were insignificant (within the relative percent difference of the analyses) (data on file at the USGS office in 

Baltimore, Maryland). TCE, which is an abiotic daughter 

product of TeCA, commonly was observed in the killed 

controls, although concentrations were less than 6 percent of 

the initial added TeCA (data on file at the USGS office in 

Baltimore, Maryland). TCE concentrations in live and killed 

controls were highest in the July–August 1999 microcosms 

and the microcosms incubated at 5 °C, as discussed in the 

section on Seasonal Variability. As much as 2 percent of 

the TCE observed in all microcosms may have resulted from 

contamination by the neat TeCA used to amend the microcosms. Analyses of the neat TeCA showed that TCE was 

an impurity, comprising up to 2 percent of the solution. 

The only other daughter compounds observed in the killed 

TeCA-amended microcosms were cDCE and tDCE. The 

total concentration of these compounds remained below 

10 percent of the initial TeCA concentration during the time 

that complete degradation of TeCA was observed in the 

corresponding live microcosms (generally the first 12 to 

25 days of incubation; figs. 6–9). Concentrations of 
20 TeCA Degradation and Microbial Communities in a Freshwater Tidal Wetland, APG, Md.

12DCE sometimes did increase greatly after this time, possibly indicating the recovery of microorganisms from the 

effect of formaldehyde or the occurrence of abiotic dichloroelimination of TeCA (Lorah and Olsen, 1999a). TRFLP 

profiles of the microbial communities in the killed controls 

sometimes did indicate the recovery of microorganisms 

near the end of incubations, as discussed in the section on 

Microbial Communities and Associations with Degradation 

Pathways. Of all the killed microcosm controls conducted, 

the most rapid decrease in TeCA (figs. 6–9) and earliest 

increase in 12DCE were observed in the February–April 

1998 experiments that were amended with TeCA mixed in a 

MeOH solution. After day 16 in the February–April 1998 

experiments, all the loss of TeCA in the killed controls can 

be attributed to 12DCE production, and over 70 percent of 

the initial TeCA was degraded to 12DCE by the end of the 

experiments (fig. 6a). Daughter compounds were not 

detected in any 112TCA-amended killed controls, but sorption losses were similar to those in the TeCA-amended killed 

controls. 

Mass balances between the aqueous TeCA concentration 

and the sum of daughter products produced provide evidence 

that the decrease in TeCA concentrations over time in the 

live microcosms was due largely to degradation of the TeCA, 

rather than to sorption. In all experiments, production of 

daughter compounds in the live microcosms showed that 

biodegradation of TeCA by microorganisms indigenous to 

the wetland sediments was occurring. Figure 10 shows the 

sum of daughter compounds (maximum sum at any one time 

step in the experiment) compared to the initial (day 1) TeCA 

concentration for each live microcosm experiment. Most 

of these points fall close to the 1:1 line that indicates parent 

compound concentration equals the sum of daughter compounds (fig. 10). The three points that are well below the 1:1 

mass-balance line are the two TeCA-amended microcosm 

experiments in July–August 1999 (23TeCA.7/99 and 

30TeCA.7/99) and the microcosm that was incubated at 

5 °C. These three microcosm experiments also had the lowest TeCA degradation rates (table 3), and the lack of mass 

balance reflects the fact that the TeCA was not completely 

degraded within the incubation time of the experiments. 

None of the live controls (microcosms with sediment and 

water that were not amended with TeCA and were incubated 

in the same manner as the amended microcosms) had detectable TeCA or daughter compound concentrations throughout 

the experiments. Desorption and degradation of background 

contaminants in the wetland sediment, therefore, did not 

affect the mass-balance calculations.

In the microcosms that were amended with 112TCA 

rather than TeCA, 112TCA degradation rates were as high 

or higher than TeCA degradation rates (table 3; fig. 5). The 

first-order rate constant estimated for the 112TCA-amended 

microcosms that contained MeOH was 0.45 per day (corresponding to a half-life of 1.5 days), which was about four 

times faster than TeCA degradation in comparable MeOHamended microcosms (fig. 5). As observed for the TeCAamended microcosms, 112TCA degradation rates were about 
Laboratory Experiments on Anaerobic Degradation of TeCA and Association with Microbial Communities 21

 
22 TeCA Degradation and Microbial Communities in a Freshwater Tidal Wetland, APG, Md.

twice as high in microcosms that contained MeOH compared 

to those that did not. No significant difference was observed 

between the 112TCA degradation rates for microcosms constructed with sediment from site WB23 compared to those 

constructed with sediment from site WB30, but few data 

points are available for comparison (fig. 5b). 

Daughter Compound Production and Degradation

Spatial Variability Although TeCA degradation rates 

were similar in the microcosms constructed with WB23 and 

WB30 sediments (fig. 5), the daughter product distributions 

in the microcosms with the two sediments did show significant differences. In particular, the relative proportions of 

VC produced and degraded differed with the location from 

which the sediment was collected (figs. 7–9; fig. 11). 

Daughter product distributions as molar percentages were 

calculated by dividing the maximum measured concentration 

(in molar units) of each daughter product over the incubation 

period by the initial TeCA concentration in the microcosm 

(fig. 11). The spatial variability effect on daughter product 

distributions is most clearly observed by comparing the 

WB23 and WB30 microcosms in the March–April 1999 and 

the October–November 2000 (without MeOH added) experiments (fig. 11). Because the February–April 1998 experiments were constructed only with WB23 sediment and with 

the addition of MeOH (table 1), these experiments will be 

discussed in the section on Substrate Type. In the July–

August 1999 microcosms, the concentrations of some 
Laboratory Experiments on Anaerobic Degradation of TeCA and Association with Microbial Communities 23

daughter compounds still were rising at the end of the 

incubation period because of the slower TeCA degradation 

rates (figs. 5 and 10). Therefore, the spatial variability in 

daughter compound distributions in the July–August 1999 

microcosms could not be evaluated by comparing peak concentrations, as was done for the March–April 1999 and the 

October–November 2000 experiments. 

Spatial Variability of Vinyl Chloride Production—The microcosms constructed with WB23 sediment had lower concentrations of VC and 112TCA than those constructed with 

WB30 sediment (fig. 11). Peak production of VC in the 

microcosms constructed with WB23 sediment was 43 and 

32 percent of the initial TeCA concentrations in the March–

April 1999 and the October–November 2000 experiments, 

respectively. The peak production of VC in the WB30 

microcosms was about twice as high as that in the WB23 

microcosms (fig. 11). Similarly, peak production of 112TCA 

was about twice as high in the WB30 microcosms, indicating 

that the higher VC production resulted from a greater predominance of the 112TCA dichloroelimination pathway in 

these microcosms (pathway 1B in fig. 2). Because production of TCE and 12DCE was about the same in the microcosms constructed with sediment from WB23 and WB30 

(fig. 11), it is unlikely that the higher VC production in the 

microcosms with WB30 sediment resulted from greater 

hydrogenolysis of 12DCE, which can be produced through 

either dichloroelimination or dehydrochlorination of TeCA 

(pathways 2 or 3 in fig. 2). Further evidence of the pathway 

through which the higher VC is produced in the microcosms 

constructed with WB30 sediments is provided by the distribution of daughter products in the microcosms amended 

with 112TCA, which could produce VC only through dichloroelimination (fig. 12). The 112TCA-amended microcosms 

constructed with sediment from WB30 also showed twice as 

much VC production as those constructed with sediment 

from WB23 (fig. 12).

A possible alternative explanation for the higher peak 

VC concentrations in the WB30 microcosms compared to 

the WB23 microcosms is that VC degradation was more 

rapid in the WB23 microcosms. This explanation was discounted, however, by the higher VC degradation that also 

was noted in the WB30 microcosms, as discussed in the 

section on Spatial Variability of VC Degradation.

Although VC production was higher in WB30 microcosms than the WB23 microcosms, it is important to note 

that VC was the predominant daughter compound produced 

from all microcosms in 1999 and 2000 that did not contain 

added MeOH (figs. 7–9; fig. 11). The high production of 

this compound clearly indicates that VC results from multiple degradation pathways. Dichloroelimination of 112TCA 

and hydrogenolysis of 12DCE produced through TeCA 

dichloroelimination appear to be the predominant pathways 

of VC production because TCE concentrations generally 

were less than 10 percent of the initial TeCA concentration. 

In all microcosms, 12DCE and 112TCA were produced 

simultaneously and immediately from the beginning of the 

experiment, regardless of the site from which the sediment 

was collected (figs. 7–9). The spatial variability observed in 

the daughter product distributions results from the predominant pathway of 112TCA degradation, with the production 

of VC being predominant in the WB30 sediment (fig. 11). 

A correspondingly higher production of 12DCA from 
24 TeCA Degradation and Microbial Communities in a Freshwater Tidal Wetland, APG, Md.

112TCA in the WB23 sediment compared to the WB30 sediment sometimes was observed (fig. 11). The higher production of 12DCA in the WB23 sediment was most evident in 

the March–April 1999 microcosms amended with TeCA 

(fig. 11). It is possible that the peak concentration of 

12DCA was missed in the October–November 2000 microcosm with WB23 sediment because fewer sampling points 

were obtained during this experiment than during other 

experiments (figs. 9a–b). 

The spatial variability in production of VC in the microcosms shows some association with ferrous iron production 

(from reduction of ferric iron) and CH4 production (figs. 

7–9; fig. 13). In the March–April 1999 and October–

November 2000 microcosms, iron reduction occurred 

throughout the experiments with WB30 sediment, whereas 

iron concentrations remained relatively low and constant in 

the microcosms with WB23 sediment (figs. 13a–b). The 

early occurrence of iron reduction in the microcosms with 

WB30 sediment appeared to cause a delay in the onset of 

methanogenesis in these microcosms (figs. 13b and 13d). 

Methanogenesis did not begin in the microcosms with WB30 

sediment until days 16 and 21 in the March–April 1999 and 

October–November 2000 microcosms, respectively (fig. 

13d). In contrast, methanogenesis was evident from day 1 

in the WB23 microcosms in March–April 1999 (fig. 13c). 

Similarly, the onset of VC production was delayed in the 

microcosms with WB30 sediment compared to those with 

WB23 sediment (figs. 7 and 9). The onset of 12DCA production also was delayed in the microcosms with WB30 sediment compared to those with WB23 sediment, although 

12DCA production generally began before VC production 

(figs. 7 and 9). The delay in the onset of methanogenesis in 

the WB30 microcosms was associated with a greater accumulation of 112TCA. When methanogenic conditions did 

occur, 112TCA dichloroelimination to VC may have been 

more rapid than the 112TCA hydrogenolysis to 12DCA, 

resulting in the relatively higher VC production in these 

WB30 microcosms than the WB23 microcosms (fig. 11). 

Alternatively, the WB30 sediment may have a higher prevalence of microorganisms that directly or indirectly mediate 

112TCA dichloroelimination to VC than those that can 

mediate 112TCA hydrogenolysis to 12DCA. 

The microcosms that were amended with 112TCA provide further evidence of the relation between VC and 

12DCA production and methanogenic conditions (figs. 14 

and 15). In the March–April 1999 and October–November 

2000 microcosms, onset of VC production from 112TCA 

dichloroelimination generally coincided with the onset of 

methanogenesis in microcosms with WB23 and WB30 sediments (figs. 14 and 15). In contrast, 112TCA hydrogenolysis to 12DCA occurred prior to the onset of methanogenesis 

in all the microcosms (figs. 14 and 15). Comparison of the 

steepest slopes of the 12DCA and VC concentration lines 

indicates that 12DCA production is faster or about the same 

as VC production in these microcosms. Therefore, the doubling of VC production in the 112TCA-amended microcosms constructed with WB30 sediment compared to those 

constructed with the WB23 sediment (fig. 12) is most likely 

due to differences in the microbial communities in the two 

sediments. 

Enrichment cultures amended with 112TCA and 12DCE 

were used to select for members of the microbial community 

with the ability to produce VC and to examine the apparent 

differences in VC production in WB30 wetland sediment 

compared to WB23 wetland sediment (table 2). Two important differences were observed in the biogeochemical reactions in the microcosm experiments compared to those in the 

enrichment culture experiments. First, although the starting 

microcosm slurry used to inoculate the enrichment cultures 

was methanogenic, methane production was not observed 

during the enrichment culture experiments with either WB23 

or WB30 microcosm slurry. Because 112TCA dichloroelimination to VC occurred without production of CH4 in these 

enrichment culture experiments, in contrast to the microcosm experiments (figs. 14 and 15), methanogens were not 

necessarily directly involved in this reaction pathway. Second, 12DCA was not produced in any of the 112TCAamended enrichment cultures, whereas hydrogenolysis of 

112TCA to 12DCA was an important reaction in the microcosms, especially in those constructed with WB23 sediment. 

These differences between the microcosms and the enrichment cultures are not surprising because the enrichment 

cultures reflect a selection from the original microbial 

community. Selectivity results from a combination of 

effects, including dilution of the microbial community, culturing conditions differentially affecting growth or activity 

of organisms, and differential sensitivity to VOC toxicity. 
Laboratory Experiments on Anaerobic Degradation of TeCA and Association with Microbial Communities 25
26 TeCA Degradation and Microbial Communities in a Freshwater Tidal Wetland, APG, Md.

VOC concentrations added to the enrichment cultures were 

substantially greater than the VOC concentrations added to 

the microcosms (tables 1 and 2).

In enrichment cultures amended with 112TCA, linear 

production of VC began without a lag in enrichments inoculated with either WB23 or WB30 sediments and provided 

with either acetate or H2 as an electron donor (fig. 16a). 

This similar behavior in 112TCA metabolism shows that the 

enrichment cultures reflect selection of a similar sub-component of the microbial community from both the WB23 and 

WB30 wetland sediments. Enrichment cultures with the 

WB23 microcosm slurry produced about twice as much VC 

from 112TCA after 16 days as cultures with the WB30 

microcosm slurry (fig. 16a). This contrasts with the higher 

VC production from 112TCA dichloroelimination observed 

in the microcosm experiments with WB30 sediment (figs. 11 

and 12). Apparently, the wetland sediment at site WB23 has 

a higher potential than sediment at site WB30 to degrade 

112TCA to VC that is not expressed in situ. 

The cDCE-amended enrichment cultures inoculated with 

WB30 or WB23 microcosm slurry, respectively, produced 

VC after an initial lag of 6 or 19 days (fig. 16b). After this 

lag, the amount of VC production in the cDCE-amended 

enrichment cultures surpassed that observed in the 112TCAamended enrichment cultures. Enrichment cultures with the 

WB23 and WB30 microcosm slurries that were amended 

with tDCE also produced VC, but the lag periods before 
Laboratory Experiments on Anaerobic Degradation of TeCA and Association with Microbial Communities 27

increasing VC concentrations were substantially longer than 

those in the cDCE-amended cultures (fig. 16b). After the lag 

period, VC production rates in the tDCE-amended cultures 

were similar to those in the cDCE-amended cultures with 

WB30 sediment. VC concentrations increased exponentially 

in 12DCE-amended enrichment cultures, as opposed to the 

linear increase observed in the 112TCA-amended enrichment cultures (fig. 16). The exponential increase suggests 

the growth of microbial species that could utilize cDCE with 

either acetate or H2 as the electron donor (fig. 16b). The 

shorter lag before VC production began in the cDCEamended cultures inoculated with WB30 microcosm slurry 

compared to those with WB23 microcosm slurry indicates 

that the WB30 inoculation provided a higher number of 

these microorganisms or a more active population.

To summarize, the enrichment culture results indicated 

that the WB30 sediment has a higher capacity to produce VC 

from cDCE than the WB23 sediment does, whereas the 

WB23 sediment has a higher capacity to produce VC from 

112TCA than the WB30 sediment does. In contrast, microcosm results indicated that the WB30 sediment has a higher 

capacity to produce VC from 112TCA than the WB23 sediment does, whereas there was no apparent difference in the 

capacity of the two sediments to produce VC from 12DCE 

(at least in the March–April 1999 and October–November 

2000 microcosms discussed here). The combined microcosm and enrichment culture results, therefore, indicate that 

both sediments have the microorganisms necessary to carry 

out these VC production reactions, but their population size 

or activity varies with conditions such as redox or toxicity 

effects. For example, the apparent difference in the capacities of the sediment to produce VC from dichloroelimination 

of 112TCA in the microcosm compared to the enrichment 

cultures could be associated with the difference in methane 

production in the two types of experiments. The immediate 

and linear increase in VC concentrations from 112TCA degradation suggests that the enrichment cultures did not select 

for the growth of a microorganism with 112TCA-degrading 

metabolic capability, unlike the cDCE degradation reaction 

in the enrichment cultures. The linear increase and implied 

lack of growth indicates that the organisms may not be deriving energy from the 112TCA dichloroelimination reaction 

and that the reaction is cometabolic under the non-methanogenic conditions in the enrichment cultures. It is possible 

that a cometabolic 112TCA dichloroelimination reaction was 

enhanced in the methanogenic microcosms with WB30 

sediment compared to WB23 sediment because a specific 

methanogen species or group that enhances this dichloro- 
28 TeCA Degradation and Microbial Communities in a Freshwater Tidal Wetland, APG, Md.

elimination reaction was more prominent at site WB30. 

Alternatively, a microorganism capable of utilizing 112TCA 

for growth may be active in the WB30 sediment under methanogenic conditions, adding to the cometabolic reaction and 

causing the approximate doubling of VC production in the 

112TCA-amended microcosms constructed with WB30 sediment compared to those constructed with the WB23 sediment (fig. 12).

Spatial Variability of Vinyl Chloride Degradation—The greater 

production of VC in surficial wetland sediment from site 

WB30 could be of concern because of the highly toxic nature 

of this daughter compound. However, all microcosms constructed with WB30 sediment also showed a greater capacity 

to degrade VC than those constructed with WB23 sediment. 

For example, in the March–April 1999 microcosms constructed with WB23 sediment, VC concentrations increased 

beginning at day 9 and then remained approximately constant from day 30 through day 42 (end of the experiment) 

(fig. 7a). In the March–April 1999 microcosms constructed 

with WB30 sediment (and without MeOH), VC concentrations increased rapidly between days 16 and 25 and then 

decreased by approximately 50 percent over the next 5 days 

(fig. 7b). The greater VC degradation in wetland sediment 

from WB30 was a consistent characteristic of all microcosms conducted during 1996–2000 (fig. 17). The percent 

of VC degraded, calculated using the peak VC concentration 

detected during each experiment and the VC concentration 

remaining at the end of each experiment, was greater than 30 

percent and as high as 96 percent in the microcosms constructed with WB30 sediment. In contrast, the percent of VC 

degraded was between 0 and 14 percent in the microcosms 

with WB23 sediment without MeOH added (fig. 17). 

The spatial variability in VC degradation that was 

observed in microcosms constructed with wetland sediment 

from sites WB23 and WB30 (fig. 17) also was investigated 

further with VC/sediment enrichment experiments (table 2). 

The removal rates of VC and the production of methane in 

enrichments with the two sediment slurries were similar after 

extended incubation under methanogenic conditions (fig. 

18). The WB23 and WB30 wetland sediment, therefore, 

showed an equally large capacity to degrade high concentrations of VC (44.6 μM, or 2,750 μg/L) after pre-exposure to 

the contaminants and prolonged incubation under methanogenic conditions (the effects of added BES or iron are discussed in the section on Redox Conditions). VC degradation 

was rapid under methanogenic conditions, with 90 percent of 

the initial VC removed in 15 days (fig. 18).

Although VC production and degradation clearly are 

important processes in the wetland microcosms, the pathways of VC degradation could not be determined definitively from the microcosm or enrichment experiments. 

Degradation of VC by hydrogenolysis would result in 

production of ethene and/or ethane. Ethene and ethane, 

however, were detected infrequently in the microcosm 

experiments and in trace concentrations. Ethene and ethane 

also were not detected in the VC/sediment enrichment experiments. Either these compounds were not daughter products 

of the TeCA and 112TCA degradation, or ethene and/or 

ethane degradation rates were much faster than the VC 

degradation rate so that they did not accumulate in the 

microcosms. In preliminary enrichment experiments, ethene 

was added to sediment slurries from WB23 and WB30 that 

had been exposed previously to VC in VC/sediment enrichment experiments. VC degradation was much faster than 

ethene degradation in these enrichments, and no substantial 

accumulation of ethene was observed (data on file at USGS, 

Reston, Virginia). Ethene was degraded to ethane in the 

VC-adapted, ethene-amended sediment with WB23 sediment but was not degraded (or showed very limited degradation) in the amended WB30 sediment. These results indicate 

that the primary pathway of VC degradation does not produce ethene in these wetland sediments, but duplicate experiments with sediment spiked with VC and ethene at the same 

time would need to be conducted in order to directly compare the rates of degradation. Studies by Bradley and 

Chapelle (1996, 1998, 1999a,b, 2000) indicated that anaerobic oxidation of VC can occur, producing CO2 or both CH4

and CO2 as the end products (fig. 2). The micromolar concentrations of CH4 and CO2 that would be produced from 

this VC oxidation could not be discerned from the ambient 

CH4 and CO2 produced by other natural organic carbon 

substrates in the wetland sediment, unless experiments with 

radiolabeled VOCs were conducted. 
Laboratory Experiments on Anaerobic Degradation of TeCA and Association with Microbial Communities 29

Seasonal Variability The greatest seasonal variability 

in the microcosm experiments incubated under the same 

temperature of 19 °C was the substantially lower TeCA 

degradation rates and lower daughter compound production 

in microcosms constructed with wetland sediment collected 

in the summer (July 1999) compared to those constructed 

with sediment collected in the late winter/early spring (early 

March 1999) or fall (October 1999) (figs. 5a and 10). Except 

for the abiotic daughter product TCE, the peak daughter 

product concentrations in the July–August 1999 microcosms 

generally were lower and occurred after a longer incubation 

time than those in the March–April 1999 experiments (figs. 

7 and 8). TCE was present between day 1 and 16 in the 

March–April 1999 experiments, but comprised only about 

3 percent of the initial TeCA added (figs. 7 and 11a). In 

contrast, TCE was present from days 1 through 35 in the 

July–August 1999 experiment and comprised 7.2 percent of 

the initial TeCA, which is more than twice the TCE observed 

in March–April 1999 (fig. 8). TCE also was a more prominent daughter product in the October–November 2000 

experiment than in the March–April 1999 experiment (fig. 

11). The broader peaks for the daughter products over time 

indicate that production and degradation of the daughter 

compounds also was slower for the October–November 

2000 experiment, although not as slow as for the July–

August 1999 experiment (figs. 7–9). It should be noted that 

the October–November 2000 microcosm experiment with 

sediment from WB23 had fewer sampling times than the 

other experiments and, as a consequence, peak concentrations of some of the daughter products may have been 

missed.

There were two other differences between the July–

August 1999 microcosm experiments and the March–April 

1999 and October–November 2000 microcosm experiments. 

First, the July–August 1999 microcosms amended with 

TeCA and 112TCA did not show a relation between VC 

production and methane production. VC production was 

observed by day 6 in the WB30 microcosms in the July– 

August 1999 experiment (fig. 8b), although methane production was minimal by day 6 and remained low throughout 

incubation (figs. 13 c–d). In contrast, VC production in the 

WB30 microcosms in March–April 1999 coincided with an 

increase in methane concentrations from day 16 to 21 (figs. 

7b and 13d). Second, VC production began earlier in the 

microcosms constructed with WB30 sediment than those 

constructed with WB23 sediment in the July–August 1999 

experiment, whereas VC production was delayed in the 

WB30 microcosms compared to WB23 microcosms in the 
30 TeCA Degradation and Microbial Communities in a Freshwater Tidal Wetland, APG, Md.

March–April 1999 and October–November 2000 experiments (figs. 7–9).

To evaluate the effect of seasonal changes in temperature 

on degradation, one set of microcosms was constructed with 

sediment from site WB23 in early March 1999, when ice 

was present on the surface of the wetland, and incubated at 

5 °C. Substantial but slower degradation of TeCA was 

observed at the cold temperature of 5 °C compared to 19 °C. 

About 50 percent of the initial TeCA in solution was converted to daughter products within the 35-day incubation 

period at 5 °C, whereas complete loss of the added TeCA 

occurred within 21 days in duplicate microcosms incubated 

at 19 °C (table 4; figs. 7a and 10). Of the remaining 50 percent of the initial TeCA in the 5 °C microcosms, 20 percent 

was in solution at day 35, while the rest most likely was 

sorbed to the sediment based on comparison to killed controls (data not shown). The daughter product distributions 

over time (data not shown) and the maximum percentages of 

each daughter product (table 4) in the 5 °C microcosms were 

similar to the July–August 1999 experiment that was conducted at 19 °C. The abiotic daughter product TCE comprised 7.4 percent of the initial TeCA in the microcosms that 

were incubated at 5 °C. Methane production was negligible 

in the 5 °C microcosms, and also was low in the July–August 

1999 experiment that was conducted at 19 °C compared to 

the March–April 1999 experiment (fig. 13c).

The relatively slow degradation rate in July–August 1999 

microcosms incubated at 19 °C compared to other microcosms incubated at this temperature and the similarities 

between the summer experiment (July–August 1999) and 

microcosms incubated at 5 °C were unexpected. Because 

microbial growth and activity generally increases with 

increasing temperature, higher microbial biomass and higher 

degradation rates were expected with sediment collected in 

the summer compared to sediment collected during late winter/early spring. Instead, seasonal variation in TeCA biodegradation in the surficial wetland sediments did not seem to be 

associated with temperature effects. The trend observed was 

highest degradation in the winter/early spring and lowest 

degradation in the summer, followed by increasing degradation in the fall. The degradation of TeCA and the production 

and degradation of daughter compounds in the microcosms 

constructed in March 1999 with sediment from site WB30 

was similar to that in the May 1996 microcosms (figs. 3 and 

7b), indicating some consistency in this seasonal trend from 

year to year.

Substrate Type Production and degradation of VC 

varied with the substrate type added to microcosms and 

enrichment experiments. The effect of MeOH as a substrate 

during TeCA degradation was tested inadvertently in the 

February–April 1998 experiments when TeCA and 112TCA 

stock solutions used to amend the microcosms were mixed in 

MeOH. The February–April 1998 experiments that were 

amended with TeCA and prepared with sediment from site 

WB23 (except the background experiment Bck-TeCA.2), 

had substantially higher 12DCA and lower VC concentrations than experiments constructed without MeOH (figs. 6, 

11, and 19a). Methane production also was substantially 

higher in these MeOH-amended microcosms. Methane concentrations increased above 1,500 μmol/L after 20 days of 

incubation in the MeOH-amended experiments 23TeCA.2 

and 23TeCA.3 (fig. 20), compared to less than 800 μmol/L in 

microcosms without MeOH and amended with similar initial 

TeCA concentrations (fig. 13 c).

 

 

 

Table 4. Percentage of each daughter product observed in the TeCA-amended microcosms constructed 

with sediment from site WB23 and incubated at 5 degrees Celsius (experiment 23TeCA5.3/99) 

compared to those incubated at 19 degrees Celsius in March–April 1999 (experiment 

23TeCA.3/99) and July–August 1999 (experiment 23TeCA.7/99) A

 [112TCA, 1,1,2-trichloroethane; 12DCA, 1,2-dichloroethane; CA, chloroethane; TCE, trichloroethene; 12DCE, cis-1,2-dichloroethene and 

trans-1,2-dichloroethene; VC, vinyl chloride]

Experiment name 112TCA 12DCA CA TCE 12DCE VC Total B

23TeCA.3/99 (19 °C) 16 29 0 3.1 25 43 116

23TeCA.7/99 (19 °C) 21 3.8 0 7.2 16 16 64

23TeCA5.3/99 (5 °C) 20 6.3 0 7.4 14 1.8 50

A To calculate percentages, maximum molar concentrations were used for each daughter product observed over the course of the experiment and divided by 

the initial TeCA molar concentration. Note that as much as 2 percent of the TCE may have resulted from contamination by the neat TeCA used to 

amend the microcosms.

B Sum of the percentages of daughter products observed over the course of the experiment.
Laboratory Experiments on Anaerobic Degradation of TeCA and Association with Microbial Communities 31

The February–April 1998 experiment was designed 

largely to elucidate whether VC, which was one of the predominant daughter products observed in the first experiment 

conducted in 1996, was produced mainly from dichloroelimination of 112TCA or from hydrogenolysis of 12DCE. VC, 

however, was a relatively minor daughter product of TeCA 

and 112TCA degradation in the February–April 1998 experiment compared to other microcosm experiments that did not 

contain MeOH (figs. 11, 12, and 19). It appears that the differences between the two experiments resulted primarily 

from differing pathways of 112TCA degradation, rather than 

from differences in the initial TeCA degradation pathway. 

Similar percentages of 12DCE were observed in all TeCAamended microcosms (figs. 11 and 19a), indicating that the 

proportion of TeCA degrading by hydrogenolysis and 

dichloroelimination was similar in all experiments. The 

112TCA produced from hydrogenolysis of TeCA subsequently may have degraded predominantly by continued 

hydrogenolysis to 12DCA in the MeOH-amended microcosms and predominantly by dichloroelimination to VC in 

microcosms without MeOH (figs. 11 and 19). VC production was lower in 112TCA-amended microcosms with 

WB23 sediment when MeOH was added (fig. 19b). VC production accounted for about 20 percent of the 112TCA 

degraded in the March–April 1999 experiment when MeOH 

was not added, and only 4 percent in the February–April 

1998 when MeOH was added. It is possible that the differences in the peak VC concentrations are due partly to differences in VC degradation rates, rather than VC production 

rates. Faster VC degradation in the MeOH-amended microcosms could result in lower peak VC concentrations than 

in those without MeOH. In fact, all the VC measured in 

the TeCA-amended microcosms with MeOH from the 

February–April 1998 experiment was degraded by the end 

of the incubation period, and the percent VC degraded was 

higher in these experiments than in other microcosms that 

did not contain added MeOH (fig. 17). As discussed earlier 

for the microcosms without MeOH, ethene and ethane were 

not detected or were detected only at trace concentrations in 

the microcosms. 

The addition of MeOH to microcosms with sediment 

from site WB23 enhanced the spatial variability effect that 

was discussed in a previous section. A short microcosm 

experiment that was conducted in October 1998 was 

designed to test the TeCA degradation in microcosms constructed with WB23 sediment with and without added 

MeOH (table 1). These results indicated that added MeOH 

alone did not cause the high 12DCA and low VC concentrations observed in the February–April 1998 microcosms with 

WB23 sediment (Lorah-Devereux, 1999). The 1999 experi-
32 TeCA Degradation and Microbial Communities in a Freshwater Tidal Wetland, APG, Md.

ments comparing degradation in WB23 and WB30 sediments confirmed that WB23 sediments produced more 

12DCA and less VC even without the addition of MeOH 

(figs. 11 and 19). The use of MeOH increased VC degradation, however, whereas the sediment from site WB23 

showed a lower ability to degrade VC than sediment from 

site WB30 did in the microcosms without MeOH (fig. 17). 

In October–November 2000, microcosms with and without 

added MeOH also were compared for WB30 sediment (fig. 

11b). These results showed that MeOH also increased 

12DCA production and decreased observed VC production 

in WB30 surficial wetland sediment (fig. 11b). Methanol 

addition increased methane production in microcosms with 

WB23 (Lorah-Devereux, 1999) and WB30 sediment (fig. 

13d).

Concentrations of 12DCA decreased in the 112TCAamended microcosms after about 15 days of incubation, 

whether or not MeOH was added (Lorah-Devereux, 1999; 

fig. 14a). In the 1998 experiments with MeOH, however, the 

12DCA hydrogenolysis product chloroethane (fig. 2) was 

observed whereas it was not detected in the experiments 

without MeOH (fig. 19b). Possibly, the dichloroelimination 

degradation pathway for 12DCA (fig. 2) is predominant in 

the microcosms without MeOH, as it is for 112TCA degradation. Dichloroelimination of 12DCA produces ethene, 

which also was not detected in any of these microcosms, 

however.

The effect of hydrogen and acetate as available substrates 

during VC production was examined in enrichment culture 

experiments amended with 112TCA, cDCE, or tDCE (table 

2). VC production from all three precursors occurred without methane production in the enrichment culture experiments. VC was produced from 112TCA immediately and 

linearly without significant difference between enrichment 

cultures incubated with the different substrates (fig. 16a). In 

contrast, there was a lag in VC production from cDCE and 

tDCE followed by an exponential increase (fig. 16b). 

Although VC was produced in 12DCE-enrichment cultures 

with acetate as a substrate, VC concentrations after 20 days 

of incubation were more than three times higher when 

hydrogen was the added substrate. Ethene was produced in 

high concentrations only in enrichment cultures supplied 

with hydrogen. These results indicate that the pathway and 

efficiency of VC production and degradation varied with 

substrate type.

Redox Conditions The availability of different terminal 

electron acceptors under different redox conditions causes 

varying pathways and efficiencies of TeCA and daughter 

compound production and degradation. The effect of methanogenic and sulfate-reducing conditions on TeCA degradation was investigated in the February–April 1998 microcosm 

experiments by addition of BES as a methanogenic inhibitor 

or by addition of sulfate as an alternate electron acceptor 

(table 1; figs. 21 and 22). Methanol was added to all micro-
Laboratory Experiments on Anaerobic Degradation of TeCA and Association with Microbial Communities 33

cosms in the February–April 1998 experiment, and its possible effect on the BES- and sulfate-amended results is 

unknown. The effect of BES as a methanogenic inhibitor 

and of iron as a terminal electron acceptor on VC degradation was evaluated in VC/sediment enrichment experiments 

(table 2). 

Addition of BES to February–April 1998 microcosms 

did inhibit methanogenic activity significantly, although a 

slight increase in methane concentrations was observed over 

the 41-day incubation (fig. 22b). Of the MeOH-amended 

microcosms, TeCA degradation rates were lowest in those 

amended with BES (fig. 5a). The half-life of 4.6 days for 

TeCA in the BES-amended microcosms still was low, however, indicating that methanogenic activity was not necessary for TeCA degradation to occur. Substantial production 

of 112TCA, 12DCA, and 12DCE occurred in the BESamended microcosms, indicating that methanogens were not 

directly involved in the production of these compounds (fig. 

21). Peak concentrations of 112TCA, 12DCA, and 12DCE 

in the BES-amended microcosms were about the same or 

higher than those in the treatment without BES that had a 

similar initial TeCA concentration (23TeCA.2) (fig. 19a). 

VC and chloroethane (CA) production, however, were substantially lower in the BES-amended microcosms than in 

23TeCA.2 (fig. 19a). The lower VC production in the BESamended microcosms is consistent with the association 

noted between onset of methane production and of VC production from 112TCA dichloroelimination in the microcosms (figs. 14 and 15).

Methanogenic activity also was important in degradation 

of the daughter products 12DCE and VC, whereas 12DCA 

degradation was unaffected by BES addition (fig. 21). 

12DCE and VC were degraded to below detection levels by 

day 40 in the methanogenic microcosms, but their concentrations remained approximately constant from days 25 through 

40 in the BES-amended microcosms (figs. 6b and 21). In the 

sulfate-reducing microcosms where methane production was 

not evident after day 16, the approximately constant 12DCE 

and VC concentrations from day 16 through 46 indicated 

that degradation of 12DCE and VC also was negligible (fig. 

22). In previous TCE-amended microcosms with wetland 

sediment from site WB30, similar inhibition of 12DCE and 

VC degradation in BES-amended microcosms and under sulfate-reducing conditions was observed (Lorah and others, 

1997; Lorah and others, 2001). Other laboratory and field 

studies have shown that reductive dechlorination of 12DCE 

and VC is partially or completely inhibited under sulfatereducing conditions compared to methanogenic conditions 

(Bagley and Gossett, 1990; Kästner, 1991; Pavlostathis and 

Zhuang, 1991; Semprini and others, 1995).

Early experiments by Bradley and Chapelle (1996) indicated that oxidation of VC occurred under iron-reducing 

conditions. Because the WB30 sediments that showed a 

greater capacity to degrade VC in the microcosm experiments were initially iron-reducing (figs. 13 and 17), this possible oxidation pathway was investigated in the VC/sediment 

enrichment experiments by adding FeOOH to induce ironreducing conditions in the methanogenic sediments. Iron 

reduction occurred and methanogenesis was inhibited in 

these FeOOH-amended cultures (fig. 18). VC degradation 

was inhibited in the iron-reducing enrichments, indicating 

that anaerobic oxidation of VC by iron-reducing bacteria did 

not occur (fig. 18). The addition of a more bioavailable, 

chelated form of iron (Fe(III)-NTA) also did not stimulate 

degradation under iron-reducing conditions (data on file at 

USGS, Reston, Virginia). Substantial degradation of VC in 

the VC/sediment enrichments was observed only under 

methanogenic conditions. Inhibition of methanogenesis by 

BES inhibited VC degradation in the enrichment experiments, as observed in the microcosm experiments (figs. 18 

and 21). None of these sediment incubation experiments, 

however, can discern between an oxidation reaction (production of CH4 and CO2) and a reduction reaction (production 

of ethene or ethane) for the VC degradation observed under 

methanogenic conditions.

Toxicity Although the linear increase in TeCA degradation rates indicates that TeCA concentrations up to 

13 μmol/L were not noticeably toxic to the microorganisms 

responsible for the degradation (fig. 5a), methanogenic 

activity appeared to be partially inhibited in microcosms 

with the highest initial TeCA concentration (23TeCA.3), as 

shown by a slower increase in methane concentrations than 

observed at lower initial TeCA concentrations (23TeCA.1 

and 23TeCA.2) (fig. 20a). Methane concentrations of about 
34 TeCA Degradation and Microbial Communities in a Freshwater Tidal Wetland, APG, Md.

2,000 μmol/L were reached by day 15 at the lower initial 

TeCA concentrations and by later than day 22 at the highest 

initial TeCA concentration (fig. 20a). In addition, at the 

highest initial TeCA concentration (23TeCA.3), the greatest 

time lag was observed between the peak concentrations of 

112TCA and 12DCA, and the peak 112TCA concentration 

was substantially higher than the peak 12DCA concentration 

(fig. 6). Microcosm experiment 23TeCA.3 with the highest 

initial TeCA concentration also showed the lowest percentage of CA (fig. 19a). These results indicate that hydrogenolysis of 112TCA to 12DCA and CA was inhibited at the 

highest initial TeCA concentration, at least in microcosms 

amended with MeOH.

For the two initial concentrations of 112TCA that were 

tested in February–April 1998 (2.4 and 6.3 μmol/L), no toxic 

effect was observed on methanogenic activity, and rapid 

112TCA degradation rates occurred (figs. 5b and 20b). In 

enrichment cultures that were amended with much higher 

concentrations of 112TCA (10.8 millimolar, see table 2), 

degradation of 112TCA to VC did occur but without CH4 

production (fig. 16). Thus, the methanogenic community in 

the original microcosm slurry used for the enrichment cultures may have been inhibited by the 112TCA, although 

selective effects of dilution and the enrichment medium also 

could have affected methanogen growth in the enrichment 

cultures. Microcosms in the October–November 2000 

experiment that were constructed using sediment from site 

WB30 and amended with 3.6 μmol/L of 12DCA showed no 

degradation or methanogenic activity. Decreases in 12DCA 

concentrations in the live microcosms were not different 

from those in the killed controls throughout the 30-day incubation, and production of CA or ethane was not observed 

(data on file at the USGS office in Baltimore, Maryland). 

Therefore, 12DCA seemed to have a greater toxic effect than 

either TeCA or 112TCA. Other studies also have shown that 

chlorinated alkanes, including 12DCA, chloroform, and carbon tetrachloride, inhibit methanogenic activity at relatively 

high concentrations (Belay and Daniels, 1987; Liang and 

Grbic ́-Galic ́, 1993). It is important to note, however, that 

apparent toxicity effects in the batch microcosm experiments 

may not occur in a flowing system.

Pre-Exposure to Contaminants Pre-exposure of the wetland sediment to the presence of TeCA was not required for 

degradation of TeCA. In microcosms constructed with wetland sediment and ground water from background site 

WB19, the TeCA degradation rate was slightly higher than 

the other TeCA-amended microcosms that contained MeOH 

(fig. 5a). In addition, no lag occurred before degradation of 
Laboratory Experiments on Anaerobic Degradation of TeCA and Association with Microbial Communities 35

TeCA commenced in the microcosms constructed from the 

contaminated or the uncontaminated sites (figs. 6 and 23). 

Thus, acclimation to the presence of TeCA by sediment 

microorganisms was not needed for degradation to occur by 

the pathways observed in the MeOH-amended microcosm 

experiment. 

To test the effect of pre-exposure to TeCA on VC degradation in the wetland sediments, replicate VC/sediment 

enrichment experiments were constructed using three sets of 

sediments—“fresh” sediment that was amended with VC 

within 5 days of collection, “pre-exposed” sediment that was 

incubated with added TeCA for 40 days in the same manner 

as the microcosm experiments, and “preincubated” sediment 

that was incubated in the same manner as the “pre-exposed” 

sediment, but without the addition of TeCA (table 2). VC 

was not degraded in killed controls for any of these enrichments (data on file at USGS, Reston, Virginia), but all live 

and killed treatments showed an abiotic loss of VC in the 

first 2 or 3 days that probably was caused by sorption (fig. 

24a). In the live VC/sediment ennrichments, VC degradation was fastest in the pre-exposed enrichments with both 

WB23 and WB30 sediment, and slowest in the fresh sediment (fig. 24a). VC was degraded completely by day 6 in 

the pre-exposed enrichments (fig. 24a). VC degradation in 

the fresh WB23 sediment enrichments lagged for the first 

2 weeks, and then VC concentrations decreased at a rate 

similar to that observed in previous VC/sediment enrichment 

experiments (fig. 18). Complete removal of the VC was 

observed by day 28 in the fresh WB23 sediment enrichments. In the fresh WB30 sediment enrichments, degradation lagged for at least 3 weeks, and then VC concentrations 

decreased at a rate similar to that in fresh WB23 sediment 

cultures (fig. 24a). Methane production was greater in the 

three sets of enrichment cultures with WB23 sediment than 

in those with WB30 sediment (fig. 24b). The longer lag 

before VC degradation in the cultures with fresh and preincubated WB30 sediment could be associated with the lower 

methane production in these cultures compared to the preexposed WB30 enrichments and to the WB23 VC/sediment 

enrichments. In the pre-exposed enrichments, VC degradation was equally rapid in WB30 and WB23 sediment, 

although methane production was lower in the WB30 preexposed culture (fig. 24). 

Microbial Communities and Associations with Degradation 

Pathways

A microbial consortium, rather than one microbial species or functional group, apparently is involved in the degradation of TeCA, as indicated by the variability in daughter 

product distributions in the microcosms with the site or season from which the sediment was collected, and by the variable effect of redox conditions on different steps of the 

degradation pathways. The microbial community analyses, 

combined with the geochemical data, assist in evaluation of 

the reaction pathways of TeCA degradation in the anaerobic 

microcosms, and lead to identification of the possible microbial species or groups involved, either directly or indirectly, 

in different steps of the reaction pathways. To identify possible critical microbial species or groups, shifts in the TRFLP 

profiles (DNA fingerprints) of the microbial communities in 

the microcosm sediments over time and between different 

experiments were compared to shifts in the daughter product 

distributions to note consistent associations. In the discussion below, changes in degradation pathways associated with 

spatial and seasonal variability are linked to changes in the 

microbial communities. Other factors affecting degradation 

pathways and microbial communities, such as substrate type, 

are discussed where molecular data are available. In this 

report, the potential critical microbial species or groups are 

identified primarily by base pair sizes for the specific peaks 

on the DNA fingerprint. Future analyses of these specific 

peaks in archived frozen samples should provide definitive 

identification by using primers for specific microbial functional groups, such as iron-reducing bacteria, sulfate-reducing bacteria, or acetogens. The planned completion of 

genetic analyses of the enrichment experiments should assist 

in these identifications because enrichment decreases the 

number of TRFLP peaks to be identified by selecting for 

community members involved in the reaction of interest and 

diluting out others. Enrichment greatly decreases the diversity of microbial species and increases the intensity of peaks 

associated with the selected members of the microbial community. The following sections discuss the degradation 

reactions of TeCA for which possible indicator microbial 
36 TeCA Degradation and Microbial Communities in a Freshwater Tidal Wetland, APG, Md.

species or groups have been identified on the basis of the 

microcosm experiments.

Degradation of 1,1,2,2-Tetrachloroethane Simultaneous 

production of 12DCE and 112TCA from dichloroelimination 

and hydrogenolysis of TeCA, respectively, occurred without 

a lag in all live microcosm experiments that were conducted 

(figs. 6–9), including the experiments where low degradation 

rates of TeCA were observed—the July–August 1999 microcosms incubated at 19 °C and the March–April 1999 microcosms incubated at 5 °C (fig. 5; table 4). Because methane 

production often did not occur when 12DCE and 112TCA 

production occurred, bacteria (rather than methanogens) are 

most likely involved in these TeCA degradation reactions. 

On a molar percentage basis, the proportion of 12DCE produced was between 16 to 25 percent of the initial TeCA in 

microcosms constructed with wetland sediment from three 

different sites and under all observed redox conditions (figs. 

11 and 19a; table 4), indicating the involvement of metabolically versatile anaerobic bacteria or a cometabolic process in 

the dichloroelimination of TeCA. Although the percentage 

of 112TCA produced was more variable (figs. 11 and 19a; 

table 4), hydrogenolysis of TeCA to 112TCA also seems to 

involve metabolically versatile anaerobic bacteria. Identification of the microorganisms that potentially are involved in 

these early TeCA degradation reactions can be best examined by using the molecular data from microcosms where 

production of daughter compounds other than 12DCE and 

112TCA was minimal.

For dichloroelimination of TeCA to 12DCE, the molecular data for killed controls that showed production of 12DCE 

provides the best information because 12DCE (both cis- and 

trans- isomers) was the only daughter compound observed in 

these microcosms. Production of 12DCE in killed controls 

was greatest in the February–April 1998 microcosms that 

contained MeOH, as discussed in the preceding section on 

Degradation Rate of 1,1,2,2-Tetrachloroethane. Recovery of 

metabolic activity in the killed controls was indicated by 

decreasing TeCA concentrations (fig. 6) and corresponding 

increasing 12DCE concentrations by the end of the experiments (Lorah-Devereux, 1999; Lorah and Olsen, 1999a). 

Killed controls for experiments 23TeCA.1 and 23SO4-

TeCA.2 had the highest production of 12DCE of all microcosm experiments; the maximum molar percentage of 

12DCE produced was greater than 80 percent. The extensive metabolic activity in the killed controls for the MeOHamended experiments indicates that microbes involved in 
Laboratory Experiments on Anaerobic Degradation of TeCA and Association with Microbial Communities 37

TeCA dichloroelimination can utilize MeOH, and possibly 

formaldehyde, as substrates. Although DNA was not 

recovered from most killed microcosm samples from early 

time steps, DNA was recovered and analyzed by TRFLP 

from later time points where 12DCE production occurred. 

Bacterial TRFLP profiles in sediment from killed controls in 

the 1998 experiment show a substantial increase over the 

course of the experiment in intensity of several peaks, 

including peaks at 170, 198, 235, and 238 bp (base pairs) 

(fig. 25). There were other less prominent peaks, but none 

that consistently appeared in all microcosms that showed 

12DCE production from TeCA dichloroelimination. The 

bacteria TRFLP profiles for the TeCA-amended microcosms 

in July–August 1999 are shown for comparison of prominent 

peaks (fig. 26). Microbial biomass and diversity were substantially lower in the live July–August 1999 microcosms 

compared to all other microcosms, as discussed in more 

detail in the following section on Seasonal Variability (figs. 

26 to 28). Prominent bacterial peaks, therefore, were fewer 

and more distinct. At day 3 in the TeCA-amended live 

microcosms in July–August 1999, the daughter compound 

present in the highest concentrations was 12DCE, except for 

the abiotic daughter product TCE (fig. 8). Of the prominent 

bacterial peaks in the killed controls that produced 12DCE 

(fig. 25), the 198-bp peak was the most prominent at day 3 in 

the July–August 1999 microcosms (fig. 26). A role for the 

bacterial species or groups represented by 170, 235, and 238 

in TeCA dichloroelimination, however, can not be discounted as more than one bacterial group may be involved. 

TeCA dichloroelimination produced both the 12DCE isomers, although production of cDCE commonly was three or 

more times greater than production of tDCE in these and 

previously reported microcosms at this site (Lorah and 

Olsen, 1999a). Different bacteria likely are involved in production of the two 12DCE isomers. 

The February–April 1998 microcosms with WB23 sediment also were used to help identify bacteria potentially 

involved in hydrogenolysis of TeCA to 112TCA because 

both the use of MeOH and the use of WB23 sediment 

seemed to enhance this hydrogenolysis pathway, as shown 

by the predominance of 112TCA and 12DCA compared to 

VC in these microcosms (discussed previously in the section 

on Substrate Type). As an example, figure 25 shows the 

bacterial TRFLP profiles from experiment 23TeCA.1 after 

2 days of incubation, compared to bacteria profiles in the 

WB23 sediment prior to any microcosm preparations and to 

the live control microcosms that did not contain added TeCA 

or MeOH. 112TCA and 12DCE were the only biotic daughter products observed at day 2 in 23TeCA.1 (fig. 6a). The 

bacterial peaks at 90 and about 215 bp were enhanced in 

23TeCA.1 compared to the live control microcosm (fig. 25). 
38 TeCA Degradation and Microbial Communities in a Freshwater Tidal Wetland, APG, Md.
Laboratory Experiments on Anaerobic Degradation of TeCA and Association with Microbial Communities 39
40 TeCA Degradation and Microbial Communities in a Freshwater Tidal Wetland, APG, Md.
Laboratory Experiments on Anaerobic Degradation of TeCA and Association with Microbial Communities 41

Although other peaks were enhanced in 23TeCA.1, they 

were discounted as having a role in 112TCA production 

because they also were present in the killed controls (fig. 

25). The killed controls only showed production of 12DCE 

and not 112TCA. The bacterial group represented by the 

90 bp peak is more likely to be involved in TeCA hydrogenolysis to 112TCA than the 215 bp peak because the 90 bp 

peak was observed in all live TeCA-amended microcosm 

sediment samples (figs. 26–28). The presence of the 215 bp 

peak was not associated consistently with 112TCA production, sometimes occurring in extremely low intensity or not 

at all when substantial 112TCA production was observed 

(figs. 8 and 26).

Daughter Compound Production and Degradation

Spatial Variability of Vinyl Chloride Production—One of the 

greatest spatial variability effects observed in the microcosms was in the production of VC. At least three distinct 

VC production reactions were noted from the microcosm 

and enrichment culture experiments: (1) 112TCA dichloroelimination (possibly cometabolic) that was enhanced under 

methanogenic conditions and resulted in approximately 

twice as much VC production in WB30 microcosms compared to WB23 microcosms amended with either TeCA or 

112TCA, (2) cometabolic 112TCA dichloroelimination that 

occurred without methane production and at a similar rate in 

cultures inoculated with sediment slurry from WB23 and 

WB30, and (3) growth-linked hydrogenolysis of 12DCE that 

occurred without methane production and more rapidly in 

WB30 enrichments than in WB23 enrichments (see earlier 

section on Spatial Variability of VC Production). Thus, at 

least three different microbial species/groups potentially are 

involved in VC production in these wetland sediments. The 

combined microcosm and enrichment culture results indicate 

that both sediments have the microorganisms necessary to 

carry out these VC production reactions, but that their population size or activity vary with conditions such as redox or 

toxicity effects. In the discussion below, shifts in the TRFLP 

profiles that correspond to the variable timing of production 

of VC observed in the WB30 microcosms compared to the 

WB23 microcosms are used to identify possible microbial 

species/groups that are involved in VC production reactions, 

either directly or indirectly. Thus, spatial variability effects 

are discussed primarily in the context of clarifying degradation pathways and associated microorganisms, instead of 

identifying the causes of spatial variability. The underlying 

causes of spatial variability likely are complex and result 

from a combination of interconnected factors, including 

redox conditions, substrate types, and toxicity effects.

 In March–April 1999, the onset of VC production was 

delayed in the TeCA- and 112TCA-amended microcosms 

constructed with WB30 sediment compared to those constructed with WB23 sediment (figs. 7 and 14), but VC production was greater in the WB30 microcosms (figs. 11 and 

12). Conditions in the sediment at WB30 seemed to favor 

the dichloroelimination pathway of 112TCA degradation 

over the hydrogenolysis pathway, as discussed in the preceding section. Because methane production enhanced but was 

not required for 112TCA dichloroelimination to VC, bacteria 

rather than methanogens seem to be directly involved in this 

degradation reaction. Comparison of bacterial TRFLP profiles from both WB23 and WB30 microcosms in March–

April 1999 reveal development of a specific peak concomitant with the onset of VC production. This peak, a 198 bp 

fragment, appeared early in the experiments and grew in 

prominence as VC production increased (figs. 27a–b). VC 

production began by day 10 in the WB23 TeCA-amended 

microcosm, and the 198 bp peak was prominent by day 3 

(figs. 7a and 27a). In contrast, VC production began at day 

16 in the WB30 TeCA-microcosm, and the 198 bp peak also 

became prominent by day 16 (figs. 7b and 27b). The live 

control that was not amended with TeCA did not show an 

increase in intensity in the 198 bp peak, indicating that 

growth of this bacterial species or group was stimulated in 

the TeCA-amended microcosms (fig. 29a). This same 198 

bp peak possibly is associated with another dichloroelimination reaction--TeCA to 12DCE (fig. 25). 

The microbial profiles for the 112TCA-amended microcosms did not show as clear a relation between VC production and the 198 bp peak (fig. 30), but these microcosms may 

have been affected by toxicity effects of 112TCA (see previous section on Toxicity). The peak intensity and diversity 

generally was lower in the TRFLP profiles of the 112TCAamended microcosms compared to the TeCA-amended 

microcosms, as might be expected from toxicity effects (figs. 

27 and 30). The 112TCA-amended microcosm with sediment from site WB23 in March–April 1999, in which VC 

concentrations first increased between days 21 and 25 (fig. 

14a), did show a distinct increase in the 198 bp peak by day 

25 (fig. 30a). In the 112TCA-amended microcosm for site 

WB30, however, the 198 bp peak initially was prominent but 

then decreased substantially by the end of the experiment. It 

is possible that another bacterial species or group, such as 

the 255 and 258 bp peaks in fig. 30b, became involved in 

112TCA dichloroelimination under 112TCA toxicity conditions. A change in the bacterial species or group involved in 

VC production from 112TCA also was observed seasonally, 

as discussed in the following section on Seasonal Variability. 

A change in slope was observed in VC production in the 

TeCA-amended microcosms in March–April 1999, particularly those constructed with WB23 sediment (fig. 7a). The 

first increase in VC coincided with decreasing 112TCA concentrations and the second increase in VC coincided with 

decreasing 12DCE concentrations (fig. 7a). The second 

increase in VC production in the TeCA-amended microcosms in March–April 1999 began at day 25 (fig. 7a). A corresponding change in the bacterial community was not 

evident with the onset of this second major pathway for production of VC—hydrogenolysis of 12DCE (fig. 27a). A 

change in the methanogen community, however, was evident 

(fig. 31a). The methanogen TRFLP profile showed an 

increase by day 25 in a 307 bp peak, which is from a methanogen in the Methanococcales or Methanobacteriales family, 

methanogens that predominantly utilize CO2 and hydrogen 

to produce CH4 (fig. 31a). By day 30, the peak intensity of 
42 TeCA Degradation and Microbial Communities in a Freshwater Tidal Wetland, APG, Md.
Laboratory Experiments on Anaerobic Degradation of TeCA and Association with Microbial Communities 43
44 TeCA Degradation and Microbial Communities in a Freshwater Tidal Wetland, APG, Md.
Laboratory Experiments on Anaerobic Degradation of TeCA and Association with Microbial Communities 45

this 307 bp methanogen had decreased, and VC concentrations had decreased below 0.15 μmol/L (9.4 μg/L). Similarly, there was an increase in a 190 bp peak between days 16 

and 25, which also is believed to be from a methanogen in 

the Methanococcales or Methanobacteriales family and may 

be involved in the hydrogenolysis of 12DCE to VC. The 

intensity of this peak also decreased by day 30 (fig. 31a). 

The methanogen profiles for the March–April 1999 microcosms with WB30 sediment may indicate that the 307 bp 

methanogen is involved in 12DCE hydrogenolysis to VC, 

rather than the 190 bp methanogen (fig. 31b). In the WB30 

microcosms, CH4 production began between days 16 and 21 

(fig. 13d). Substantial VC production also began between 

days 16 and 21, as both 112TCA and 12DCE concentrations 

decreased (fig. 7b). The methanogen profiles for this experiment show that the 307 bp peak was prominent at day 16, 

whereas the 190 bp peak did not become prominent until day 

25, when VC production already was occurring (fig. 31b). 

The live control that was not amended with TeCA did not 

show an increase in intensity in the 307 or 190 bp peaks, 

indicating that growth of these CO2

- utilizing methanogens 

was stimulated in the TeCA-amended microcosms (fig. 29b).

In the 12DCE-amended enrichment culture experiments, 

the exponential increase in VC concentrations with addition 

of 12DCE indicated growth of a microbial population that 

can utilize 12DCE in non-methanogenic conditions (fig. 16). 

Bacteria, rather than methanogens, mediated hydrogenolysis 

of 12DCE in these cultures, and the WB30-inoculated cultures degraded cDCE more readily than the WB23-inoculated cultures (fig. 16). Molecular analyses of WB30 

sediment from the March–April 1999 microcosms using 

primers specific for dehalorespiring bacteria revealed the 

presence of Dehalococcoides ethenogenes and members 

of Desulfuromonas throughout the experiment. Similar 

molecular analyses targeting these dehalorespiring bacteria 

were unsuccessful with the WB23 sediment. It should be 

noted, however, that the detection limits of the nested PCR 

methodology for these dehalorespiring bacteria are 

unknown. Löffler and others (2000) recommend that enrichments be done to prevent false negative results. In addition, 

there could be unidentified dehalorespiring bacteria at this 

site that these specific primers would not detect. The eventual production of VC from 12DCE in the WB23-inoculated 

enrichment cultures after a long incubation (fig. 16) indicated that dehalorespiring bacteria were present at site 

WB23, but that their population size likely was much lower 

than at site WB30. Completion of the TRFLP analyses of 

the enrichment cultures and completion of molecular analyses of the microcosm slurry targeting the dehalorespiring 

bacteria (these analyses have been completed only for one 

set of microcosm experiments) could allow more definitive 

identification of the different bacteria and factors controlling 

VC production in the wetland sediment. 

Spatial Variability of Vinyl Chloride Degradation—The higher 

VC degradation that consistently occurred in the microcosms 

constructed with WB30 wetland sediment compared to those 

with WB23 sediment (fig. 17) was associated with an 

increase in the proportion of Methanosarcinaceae, a group 

that includes all methanogens that can utilize acetate to generate CH4 (acetotrophic methanogens). In the March–April 

1999 microcosms with WB30 sediment, the proportion of 

Methanosarcinaceae in the total methanogen community 

doubled between days 16 and 25, increasing from about 10 

to 20 percent (fig. 32). At day 25, VC concentrations began 

to decrease in these microcosms (fig. 7b). Similarly, in the 

October–November 2000 microcosms with WB30 sediment 

(and no added MeOH; the MeOH effect is discussed later), 

the proportion of Methanosarcinaceae tripled from 6 to 

19 percent between days 18 and 26 (fig. 33), and VC concentrations first began to decrease at day 26 (fig. 9b). These 

acetotrophic methanogens, therefore, may be linked either 

directly or indirectly to VC degradation at site WB30. In 

contrast, the Methanosarcinaceae proportional abundance 

remained about the same in the WB23 microcosms (fig. 32), 

and VC concentrations did not decrease substantially within 

the 42-day experiment (fig. 7a). It also is interesting to note 

that while the proportion of Methanosarcinaceae increased 

when CH4 production began and VC was degrading in the 

WB30 microcosms (figs. 7b, 13d, and 32), the relative abundance of total methanogens actually decreased in WB30, 

both between days 3 and 16 as well as days 16 and 25 in the 

March–April 1999 experiment (fig. 32). Note that the proportion of Methanosarcinaceae shown in figure 32 was calculated as a percentage of the total methanogen peak area on 

a given sampling day, whereas the relative abundance of 

total methanogens was calculated by comparing the total 
46 TeCA Degradation and Microbial Communities in a Freshwater Tidal Wetland, APG, Md.

mcrA peak from timepoint to timepoint over the incubation 

period. 

The Methanosarcinaceae could increase in proportion 

and potential importance by outcompeting the other methanogens in some way, such as the ability to use a substrate for 

methanogenesis such as acetate that the other methanogen 

groups cannot use. The initial dominance of iron-reducing 

conditions in the WB30 microcosms (fig. 13) was the only 

noticeable geochemical difference between the WB30 and 

WB23 microcosms. The higher availability of ferric iron in 

the surficial wetland sediments at site WB30 than at site 

WB23 accounts for the higher iron reduction observed in the 

WB30 microcosms (fig. 34). The initial iron reduction may 

have stimulated the growth of acetate-utilizing methanogens 

compared to CO2-utilizing methanogens, although the mechanism by which this enhancement might have occurred is 

unknown. Possibly, iron reduction released phosphorus 

from the sediment, and the increased availability of this 

nutrient in the WB30 microcosms stimulated the acetateutilizing methanogens. Because phosphorus can be retained 

in wetland sediments as ferric phosphate compounds and as 

oxides and hydroxides of iron and aluminum, the iron and 

aluminum content of wetland sediments affects nutrient 

availability (Mitsch and Gosselink, 1993, p. 605). Although 

onset of CH4 production was delayed in the WB30 microcosms compared to WB23 microcosms, the CH4 production 

rate was much higher in the WB30 microcosms once it 

began (fig. 13d). Higher nutrient availability in the WB30 

microcosms could have increased metabolism of the Methanosarcinaceae compared to the CO2-utilizers that were predominant in the WB23 microcosms by the end of the 

experiment (fig. 32). Alternatively, other factors such as 

toxicity or substrate type could have caused the decline in 

CO2-utilizing methanogens. Although the cause is unclear, 

it is clear that the non-acetate utilizing methanogens 

decreased in proportional abundance in the WB30 microcosms, whereas proportional abundance of acetate-utilizers 

was correlated with both the CH4 production and the VC 

degradation. 

The apparent link between VC degradation and acetotrophic methanogens observed in these microcosm experiments may support the theory that VC mineralization under 

anaerobic conditions begins with oxidation by acetogens and 

that acetotrophic methanogens enhance degradation, possibly by converting the acetate to CO2 and CH4 (Bradley and 

Chapelle, 2000) or by excretion of some biomolecule that 

stimulates degradation (Novak and others, 1998). Although 

the experiments reported here clearly show involvement of 

acetotrophic methanogens, involvement of acetogens is 

purely speculative. Evidence for the role of acetogens possibly could be obtained by conducting TRFLP analyses of 

archived microcosm samples with acetogen primers. 

Another possibility is that dehalorespiring bacteria (bacteria 

that can utilize VC as a terminal electron acceptor for 
Laboratory Experiments on Anaerobic Degradation of TeCA and Association with Microbial Communities 47

energy) actually degrade VC in the WB30 microcosm and 

subsequently cause low hydrogen levels compared to the 

WB23 microcosms. Hydrogen is a favored substrate of 

some dehalorespiring bacteria identified to date, and Fennel 

and Gossett (1998) have shown that low hydrogen levels 

driven by dechlorination reactions in a dehalorespiring 

bacteria culture favor acetotrophic methanogens over CO2-

utilizing methanogens. Molecular analyses of WB30 sediment from the March–April 1999 microcosms revealed the 

presence of the dehalorespiring bacteria Dehalococcoides 

ethenogenes and Desulfuromonas throughout the experiment, whereas they were not detected in the WB23 microcosm sediment. VC/sediment enrichment experiments, 

however, showed similar rapid VC degradation rates in 

WB23 and WB30 sediment after prolonged incubation under 

methanogenic conditions, indicating that the presence or 

absence of dehalorespiring bacteria was not a critical factor 

in the occurrence or rate of VC degradation in these wetland 

sediments (fig. 18). In addition, ethene is the end product of 

reductive dechlorination by Dehalococcoides ethenogenes

(Maymó-Gatell and others, 1997, 1999), and no ethene was 

detected in these microcosm experiments (although it had 

been detected in non-methanogenic enrichment cultures, as 

described in preceding sections). Possibly, ethene was not 

detected because it was degraded faster than VC, as discussed in the preceding section on Spatial Variability of 

Vinyl Chloride Degradation.

In any case, the activity of dehalorespiring bacteria 

would not discount a role of the acetotrophic methanogens 

in VC degradation. Maymó-Gatell and others (1995) concluded that Dehalococcoides requires growth factors such as 

vitamin B12 from other anaerobic bacteria or methanogens to 

sustain dechlorination of VC to ethene. In other words, VC 

utilization by Dehalococcoides ethenogenes can be considered cometabolic (Maymó-Gatell and others, 2001). It is 

possible that VC can be degraded by more than one reaction 

pathway in the wetland sediments, and the dominant pathway could change with changing redox conditions, substrate 

type, or other factors that affect the microbial community.

Seasonal Variability—The low TeCA degradation rates in 

the July–August 1999 microcosms (fig. 5) seem to be related 

to low microbial biomass. The microbial community analyses consistently showed that bacterial biomass was lower in 

the July–August 1999 microcosm sediment samples than in 

the March–April 1999 microcosms (figs. 26 and 27). The 

relative intensity of the peaks in the TRFLP profiles is 

related to the biomass present for each fragment length. The 

relative intensities were markedly lower in July–August 

1999 microcosms than in March–April 1999, especially in 

the bp size range of about 160 to 240 (figs. 26 and 27). Sediment samples collected from sites WB23 and WB30 and 

analyzed for microbial communities prior to microcosm 

preparation and incubation also show a lower intensity of 

peaks in the summer, indicating that the low microbial biomass in the July–August 1999 microcosms did not result 

from sediment manipulation (fig. 35). The methanogen biomass also was lower in the July–August 1999 microcosms, 

which is consistent with the low methane production in 

these microcosms (figs. 13c–d). This low biomass was contrary to the expected high microbial biomass and activity 

with high summertime temperatures. The October–November 2000 microcosm sediment samples, which had removal 

rates of TeCA and daughter products that were higher than 

those in July–August 1999 and lower than those in March–

April 1999, had biomass levels that similarly were intermediate of those in March–April and July–August 1999. The 

low microbial biomass observed using the TRFLP profiles, 

which measures the amount of DNA amplifiable by PCR 

from 1 g (gram) of sediment, was confirmed on selected sediment samples using direct cell counts (data on file at USGS, 

Reston, Virginia). Epifluorescent cell counts are difficult in 

sediment samples because of the natural fluorescence of the 

soil material. Although biomass estimates based on the concentration of PCR-amplifiable DNA are only semi-quantitative, these estimates probably are more sensitive measures of 

biomass than cell counts from soil matrices. 

Although the low microbial biomass and TeCA degradation rates seem to be a summertime occurrence, it is unknown if this is a recurrent phenomenon, or if the effect 

occurs at depths greater than the 0 to 25 cm from which 

surficial wetland sediments were collected for this study. 

Possibly, the drought conditions that occurred in the summer 

of 1999 caused unusual stress on the microbial communities 

in July–August 1999, resulting in unusually low degradation 

rates in the summer 1999 microcosms. No drought effects, 

however, were obvious in the wetland. Wetland sediments 

remained saturated close to or at land surface at sites WB23 

and WB30 throughout this study, and diurnal tidal flooding 

occurred as usual at these sites. A possible recurrent event 

that could cause low microbial biomass in the summer is the 

growth of vegetation beginning in May and continuing 

through the summer in the wetland. Phragmites (common 

reed) shows a marked increase in growth by July, commonly 

reaching heights of 3 to 5 m in the wetland area. Competition from this macrophyte growth could cause a decreased 

availability of the nutrients that are required for microbial 

growth.

One effect of the low microbial biomass in the surficial 

wetland sediments in the summer was the doubling in percentage of TCE produced by abiotic TeCA degradation in 

July–August 1999 compared to March–April 1999 microcosms (table 4). The TCE production in the July–August 

1999 microcosms was equally high (about 7 percent) in the 

microcosms that were incubated at 5 °C, where low microbial activity was expected. The October–November 2000 

TeCA-amended microcosms that had intermediate biomass 

conditions also had high TCE production (8 to 11 percent; 

fig. 11b). During experiments with anaerobic municipal 

digester sludge, Chen and others (1996) found that concentrations of this abiotic daughter product increased under low 

biomass conditions. In experiments with digester sludge, 

TCE comprised about 9 to16 percent of the initial TeCA 

added in four tests with sludge-seeded cultures that had high 

biomass and about 49 percent in diluted cultures with low 
48 TeCA Degradation and Microbial Communities in a Freshwater Tidal Wetland, APG, Md.

biomass (Chen and others, 1996). The higher TCE production observed in these low biomass sludge-seeded cultures 

than in the low biomass wetland sediment microcosms could 

be caused partly by the degree of dilution of the biomass and 

partly by the initial TeCA concentration. The initial TeCA 

concentrations that were used by Chen and others (1996) 

ranged from about 60 to 100 μmol/L, compared to the maximum of 13 μmol/L used in the microcosms presented in this 

report (table 1). It is possible that abiotic dehydrochlorination becomes more significant with extremely high TeCA 

concentrations, although a substantial difference in the percentage of TCE produced was not observed over the moderate concentration range used in the wetland sediment 

microcosms (fig. 19a).

Seasonal changes in the predominant microbial species 

or group involved in VC production were indicated by the 

lack of association between methane and VC production 

in all July–August 1999 microcosms. In addition, VC concentrations increased by days 6 and 21, respectively, in the 

WB30 and WB23 microcosms in July–August 1999 (fig. 8). 

The largest increase in VC in the WB23 microcosms in July–

August 1999 occurred between days 25 and 30, whereas it 

was the WB30 microcosms that showed this delay in VC 

production in March–April 1999 (figs. 7–8). The 198 bp 

bacteria peak that seemed linked to VC production from 

112TCA dichloroelimination in the March–April 1999 

microcosms was prominent in both WB23 and WB30 microcosms from the beginning of incubation in July–August 

1999 (fig. 26). These bacteria, therefore, do not seem to be 

associated with VC production in the July–August 1999 

microcosms. Comparing the times of onset of VC production and the increase in intensity of bacteria peaks for the 

July–August 1999 experiment, peaks at 86 bp and 170 bp 

seem to show the best link to VC production for both the 

WB23 and WB30 microcosms (fig. 26). Increases in intensity of these peaks also coincided with VC production in 

the TeCA- and 112TCA-amended microcosms in October–

November 2000 (fig. 28). Although the 198 bp peak also 

was evident in the October–November 2000 microcosms, its 

intensity was lower than the 86 and 170 bp peaks and 

decreased over the incubation time (fig. 28). Note that the 

170 bp peak, along with the 198 bp peak, was present in the 

killed controls that showed 12DCE production from TeCA 

dichloroelimination (fig. 25). The same bacterial species or 

groups, therefore, may be involved in both TeCA and 

112TCA dichloroelimination reactions. The CO2-utilizing 

methanogens that were associated with VC production from 

12DCE hydrogenolysis in the March–April 1999 microcosms also were observed in the July–August 1999 microcosms, indicating little seasonal change in this reaction (figs. 

31 and 36). 

Although the microorganisms involved in VC production 

may change seasonally, the wetland sediment from site 

WB30 consistently showed greater VC production than sediment from site WB23 (figs. 11 and 12). The involvement of 

Methanosarcinaceae in VC degradation and the greater percentage of VC degradation observed with WB30 sediment 

also seemed to be consistent seasonally on the basis of the 

March–April 1999 and October–November 2000 data (figs. 

32 and 33). Although data on the proportion of Methanosarcinaceae are not available for the July–August 1999 microcosms, substantial VC degradation was observed in the 

WB30 microcosms within the 35-day incubation period, 

whereas none was observed in the WB23 microcosms (fig. 

8). The presence of Dehalococcoides and Desulfuromonas

dehalorespiring bacteria in WB30 sediment samples, as discussed in the preceding section on Spatial Variability, could 

be at least partly responsible for the apparently greater 

capacity of these sediments to produce and degrade VC. 

Because methanogens compete for the supply of H2 as a substrate (Maymó-Gatell and others, 1995; Smatlak and others, 

1996), these dehalorespiring bacteria may be more active, or 
Laboratory Experiments on Anaerobic Degradation of TeCA and Association with Microbial Communities 49

more noticeably active, in the microcosms constructed in the 

summer when methanogen activity and biomass were lower 

(figs. 13). The presence of Dehalococcoides ethenogenes, 

which are known to obtain energy from 12DCE hydrogenolysis to VC, may explain why VC production unrelated to 

methane production was observed early in the WB30 microcosms in July–August 1999. Dehalococcoides ethenogenes

also is known to utilize 12DCA reductive dechlorination for 

growth (Maymó-Gatell and others, 1997, 1999). The presence of Dehalococcoides ethenogenes and its potentially 

greater activity in the summer also could explain the undetectable concentrations of 12DCA throughout the July–

August 1999 microcosm with WB30 sediment (fig. 8b). 

12DCA was observed in WB30 microcosms in February–

April 1998, March–April 1999, and October–November 

2000 (figs. 11 and 19). Despite the presence of the dehalorespiring bacteria at site WB30, TeCA degradation rates were 

lowest in the July–August 1999 microcosms, although the 

July–August 1999 rates were higher in the WB30 microcosms than the WB23 microcosms (fig. 5).

Substrate Type—Both VC production and VC degradation 

showed some variability with the predominant available 

substrate in the laboratory experiments. VC production from 

12DCE was substantially higher with addition of hydrogen 

as a substrate than with acetate in non-methanogenic enrichment cultures (fig. 16). In addition, enrichments that were 

amended with cDCE and hydrogen produced ethene in addition to VC, whereas enrichments that were amended with 

cDCE and acetate produced only VC (data on file at USGS, 

Reston, Virginia). These differences with substrate type are 

consistent with previous studies of two separate groups 

of dehalorespiring bacteria. One group, Dehalococcoides, 

utilized hydrogen as the favored substrate for reductive 

dechlorination of cDCE to VC and ethene (Maymó-Gatell 

and others, 1997, 1999), whereas a second group, Desulfuromonas, utilized acetate as a substrate to produce VC as 

the most reduced end product of reductive dechlorination 

(Löffler and others, 2000). VC production in the hydrogenamended enrichments was about three times higher than the 

acetate-amended enrichments (fig. 16), indicating that dehalorespiring bacteria such as Dehalococcoides that prefer 
50 TeCA Degradation and Microbial Communities in a Freshwater Tidal Wetland, APG, Md.

hydrogen as a substrate were dominant in the wetland sediment.

The microcosm experiments with added MeOH showed 

a substantial increase in VC degradation and an associated 

increase in the percentage of Methanosarcinaceae compared 

to microcosms without added MeOH (figs. 17 and 33). 

Methanol can be utilized directly as a substrate by Methanosarcinaceae, and MeOH also favors growth of acetogens. 

Because acetogens produce acetate that can be metabolized 

by Methanosarcinaceae, their growth on MeOH can stimulate growth of Methanosarcinaceae compared to CO2-utilizers (Maymó-Gatell and others, 1995). This increase 

occurred for both WB23 and WB30 sediments, indicating 

that substrate type could be an important factor in determining the methanogen type that is predominant. The enhanced 

growth of acetotrophic methanogens could be another factor 

causing the higher TeCA degradation rates in the MeOHamended microcosms (fig. 5).

Comparisons to Field Data on Anaerobic 

Degradation of 1,1,2,2-Tetrachloroethane and 

Association with Microbial Communities

Degradation Rate of 1,1,2,2-Tetrachloroethane

Using normalized concentrations from 1995 piezometer 

samples, estimated rate constants for TeCA degradation in 

the wetland porewater ranged from 0.0065 to 0.0085 per 

day at a flow velocity of 0.91 m/yr (table 5). These fieldderived degradation rates correspond to the estimated minimum and maximum flow velocities, respectively, estimated 

for the wetland sediments using two different methods. Estimated half-lives for TeCA based on the field data range from 

82 to 107 days (table 5). These rate estimates were based on 

summer 1995 data from four piezometer nests where three or 

more samples from piezometers screened in the wetland sediment had detectable concentrations of TeCA, including sites 

WB23 and WB30 that were used for sediment collection for 

the laboratory microcosm experiments (table 5; fig. 1). Late 

winter/spring sampling periods generally had lower VOC 

concentrations in the wetland porewater, and insufficient 

data were available to calculate degradation rates (Lorah and 

others, 1997; Olsen and others, 1997; Spencer and others, 

2002). These field observations of low porewater VOC concentrations in the winter/early spring are consistent with the 

higher microbial biomass and degradation rates observed in 

the March–April 1999 microcosms compared to the July–

August 1999 microcosms.

The rate constants for TeCA degradation are slightly 

lower than TCE degradation rates estimated previously using 

1995 data from the same sites (Lorah and others, 2001) 

(table 5). Because TCE can be an abiotic daughter product 

of TeCA, the field-derived rate constants for TCE degradation could be underestimated. This effect probably is small, 

however, because TCE accounted for only 3 to 11 percent of 

the TeCA degraded in laboratory microcosm experiments 

with the wetland sediment (table 4; figs. 11 and 19). Therefore, TCE concentrations used for calculating field degradation rates are predominantly from TCE present as a result of 

disposal activities rather than as a result of TeCA degradation.

The rate constants estimated based on the 1995 field data 

(table 5) agree well with the rate constants estimated from 

anaerobic laboratory microcosms that were amended with 

TCE (Lorah and others, 1997, 2001) and TeCA (fig. 5), 

although laboratory-derived rates were consistently higher. 

The laboratory-derived TCE rate constants ranged from 

0.032 to 0.37 per day (half-life of 2 to 22 days) in the laboratory experiments compared to field estimates of 0.009 to 

0.012 per day (half-life of 58 to 77 days). The best-fit line 

for TeCA degradation in the microcosms that did not contain 

MeOH gave a rate constant of 0.053 per day (half-life of 13 

days) (fig. 5) compared to estimated half-lives of 82 to 107 

days from the field data (table 5). The agreement between 

the laboratory- and field-derived rate constants can be considered good considering the inherent differences in a laboratory batch experiment and an uncontrolled field system. In 

addition, the low summer degradation rates were excluded 

from the regression line to calculate the laboratory-derived 

rate constant (fig. 5), but summer data were used to estimate 

field-derived degradation rate constants. The laboratoryderived rate constant, therefore, that was used for comparison to the field-derived rates was a maximum rate, whereas 

the field-derived rate constants in table 5 may represent 

minimum rates.

The laboratory- and field-derived estimates show similar 

TeCA degradation rates in the wetland sediments at sites 

WB23 and WB30 (table 5 and fig. 5). Rate constants for 

TeCA and TCE degradation also were estimated for site 

WB30 using porewater samples collected from a multi-level 

sampler that was added in 1999 as part of the ESTCP project 

(fig. 1 and table 5). Although a multi-level sampler also was 

installed at site WB23, chloride data were insufficient to correct TeCA and TCE concentrations and calculate degradation rate constants. Data collected from the multi-level 

sampler at WB30 in June–July 2000 allowed rate constants 

to be estimated using concentrations from twice as many 

depths in the wetland sediment as the two to three depths 

available from the drive-point piezometers in 1995 (table 5; 

figs. 3 and 37). For site WB30, the TeCA and TCE rate 

constants estimated from the piezometer and multi-level 

samplers generally are consistent, especially for TeCA 

with half-lives of 107 and 91 days using a flow velocity of 

0.9 m/yr (table 5). This comparison of summer 1995 and 

summer 2000 data indicates that degradation rates are consistent over time, at least when comparing samples collected 

during the same season, as was observed with the TeCAamended microcosms conducted in May–June 1996 and 

March–April 1999 with sediment collected from site WB30 

(figs. 4 and 7b).
Comparisons to Field Data on Anaerobic Degradation of TeCA and Association with Microbial Communities 51

 

 

Daughter Compound Production and Degradation

Laboratory microcosm experiments can assist in understanding the observed distributions of chlorinated VOCs in 

the wetland porewater. It should be noted that microcosms 

were constructed using sediment from relatively shallow 

depths in the wetland, and the microbial communities and 

biodegradation processes could vary with depth. Changes in 

redox conditions in the microcosms, however, generally 

reflected the observed changes in porewater chemistry along 

an upward flow gradient in the wetland sediment (figs. 13 

and 38). Along upward flowpaths, concentrations of the 

reduced form of dissolved iron (ferrous iron) generally 

reached a peak before methane did (fig. 38). Methane concentrations then peaked as ferrous iron concentrations 

decreased or remained about the same (fig. 38). Similarly, 

an early increase in iron concentrations in the microcosms 

resulted in higher dissolved iron than methane concentrations, whereas methane concentrations were higher at the 

end of the experiment (fig. 13). Thus, the changes in the 

microcosms over incubation time may be representative of 

redox conditions and biodegradation along a flowpath. 

Distributions of TeCA and its daughter compounds in wetland porewater along vertically upward flowpaths are shown 

 

at sites WB23 and WB30 (fig. 39). These distributions are 

shown as percentages of the initial TeCA present in the aquifer immediately underlying the wetland sediment at each 

site. The aquifer concentrations were obtained from multilevel sampler points WBM23G and WBM30G, screened at a 

depth of 3.9 m (figs. 1 and 3). Although TCE can be an abiotic daughter product of TeCA, TCE also was used and disposed of as a waste in the study area (Lorah and others, 

1997). The majority of the TCE in the wetland porewater 

probably is the result of its disposal as a waste product 

because the microcosms indicated that only 3 to 11 percent 

TCE resulted from TeCA degradation in these sediments 

(table 4 and fig. 11b). 

The most obvious difference in the distributions of 

daughter compounds in the wetland sediments at sites WB23 

and WB30 is the higher percentage of VC at all depths at site 

WB30 (fig. 39). When calculated relative to the amount of 

TeCA present in the aquifer, the TCE concentrations were 

about 30 percent higher at site WB30 than at site WB23 

(fig. 39). The higher percent TCE at site WB30, however, 

does not account for the difference in VC, which was as high 

as 55 percent. The distribution of daughter products 

observed in the field, therefore, is consistent with the pattern 

Table 5. Degradation rates for 1,1,2,2-tetrachloroethane (TeCA) and trichloroethene (TCE) estimated 

from field data in July–August 1995 and June–July 2000 along upward flowpaths in the 

wetland sediments, using TeCA and TCE concentrations normalized with chloride and an 

estimated linear flow velocity of 0.9 meters per year

 [TCE degradation rates in 1995 with a linear flow velocity of 0.9 meters per year (m/yr) were reported previously in Lorah and others 

(2001); r2, correlation coefficient]

TeCA Degradation Rate TCE Degradation Rate

Site Flowpath A

Rate

constant

(day-1 )

Half-life

(days) r2

Rate 

constant 

(day-1 )

Half-life 

(days) r2

July–August 1995 Piezometers

WB23 WB23D to -23B 0.0071 ± .0005 98 0.99 0.012 58 1.00

WB26 WB26F to -26C 0.0067 ± .0017 103 .94 0.011 ± .007 63 .68

WB27 WB27D to -27A 0.0085 ± .0007 82 .99 0.009 ± .0009 77 .94

WB30 WB30E to -30C 0.0065 ± .0040 107 .69 0.011 ± .002 63 .99

June–July 2000 Multi-level Sampler

WBM30 WBM30E to -30A 0.0076 ± .0003 91 .99 0.0065 ± .0008 107 .97

A Sequence of piezometers used to calculate rates at each site. See figures 1 and 3.
52 TeCA Degradation and Microbial Communities in a Freshwater Tidal Wetland, APG, Md.
Comparisons to Field Data on Anaerobic Degradation of TeCA and Association with Microbial Communities 53

of higher VC production observed in the microcosms constructed with sediment from site WB30. In addition, VC 

decreased from over 55 percent to less than 5 percent near 

the surface of the wetland sediments at site WB30 

(WBM30C to WB30B in fig. 39), which is consistent with 

the greater capacity for anaerobic degradation of VC that 

was observed in the WB30 microcosms (figs. 17 and 18). In 

contrast, the percentage of VC remained constant along the 

upward flowpath at site WB23, although the percentage of 

TeCA removed before a depth of 46 to 69 cm was reached 

was about the same at the two sites (fig. 39). The amount of 

TeCA lost along the flowpath at site WB23 was not accounted for by an equivalent increase in daughter products (fig. 

39). If a higher percentage of 12DCA was produced at this 

site from TeCA degradation as observed in the microcosms, 

the 12DCA was degraded as rapidly as it was produced. 

Overall, field measurements indicated a lower prevalence of 

12DCA in the wetland porewater than of 12DCE or VC 

(Lorah and others, 1997). Molecular analyses of the microbial communities with depth in the wetland sediment at these 

sites would provide a better understanding of the spatial heterogeneity in the biodegradation processes and of the correlation between laboratory and field results. 

Microbial Communities

All the bacterial peaks that were identified from the 

microcosm analyses as potentially involved in the degradation pathways of TeCA were present in the surficial wetland 

sediment samples collected from the contaminated sites 

along transects A-A′ and C-C′ (fig. 40) and at the background site WB19 (fig. 41). These include bacterial species 

or groups at base pair sizes 86, 90, 170, and 198 (figs. 40 and 

41). Bacterial diversity and biomass were substantially 

higher in transect A-A′ compared to transect C-C′ in March 

and August 1999 (fig. 40). Concentrations of chlorinated 

VOCs in shallow wetland porewater were lower in transect 

A-A′ compared to transect C-C′ , whereas methane concentrations were higher in transect A-A′ (Lorah and others, 

1997). Microcosm experiments showed that higher microbial biomass and higher methanogen activity were associated 

with higher TeCA and daughter compound degradation rates 

in the wetland sediments, indicating that the lower VOC 

concentrations along transect A-A′ could be at least partly 

due to higher degradation rates. Hydrogeologic factors, 

including the thinner wetland sediments and the occurrence 

of a focused discharge area that cause lower residence time 

of the contaminants, likely contribute to the higher VOC 

concentrations along transect C-C′ (Lorah and others, 1997; 

Lorah and Olsen, 1999b). It is possible that toxic effects of 

the generally higher contaminant concentrations along 

transect C-C′ than transect A-A′ reduced the biomass and 

diversity of the microbial community in this area of the wetland (fig. 40). 

The frequency and depth of tidal flooding on the wetland 

sediments also could affect the microbial diversity and 

biomass along the transects. Restriction of oxygen influx 

and other physiochemical effects on the sediment with more 
54 TeCA Degradation and Microbial Communities in a Freshwater Tidal Wetland, APG, Md.
Comparisons to Field Data on Anaerobic Degradation of TeCA and Association with Microbial Communities 55
56 TeCA Degradation and Microbial Communities in a Freshwater Tidal Wetland, APG, Md.

frequent and higher flooding levels could decrease microbial 

biomass and diversity. In addition, relatively low concentrations of VOCs (maximum of 50.2 μg/L) that could have a 

toxic effect on the microbial community were measured in 

the surface water (Phelan and others, 2001). Chloroform, 

the most common surface-water contaminant at this site, has 

been shown to be highly toxic to methanogenic and dehalorespiring microorganisms (Scholten and others, 2000; 

Weathers and Parkin, 2000; Maymó-Gatell and others, 

2001). Except for site WB26, the land-surface elevation of 

the wetland was higher along transect A-A′ than transect 

C-C′ (Olsen and others, 1997; Spencer and others, 2000), 

which would result in less frequent and lower tidal inundation along transect A-A′ (fig. 40). Field observations have 

shown that transect C-C′ is more consistently flooded at all 

piezometer sites in the wetland. Site WB26, which is on the 

bank of the creek channel and has the lowest land-surface 

elevation, was the only site along transect A-A′ that showed 

bacterial biomass and diversity as low as those measured 

along transect C-C′ (fig. 40).

The proportion of Methanosarcinaceae, the group that 

includes all acetate-utilizing methanogens, was between 10.8 

and 14.9 percent of the total methanogens in surficial wetland sediment samples collected along transect A-A′ in 

March 1999 (table 6). Increases in the proportion of Methanosarcinaceae in microcosms constructed with WB30 sediment compared to those constructed with WB23 sediment 

were linked to greater VC degradation. As observed in the 

microcosm sediments, the percent of Methanosarcinaceae 

was higher in the surficial wetland sediment samples from 

site WB30 than site WB23 (table 6). The percent Methanosarcinaceae was similar at site WB30 and the background 

site WB19, indicating that the relative proportion of these 

methanogens is not related to the presence of the contaminants. Although data are available for only two sites along 

transect C-C′, it is interesting to note that the lowest percent 

Methanosarcinaceae measured for both transects occurred at 

one of these sites (table 6). Site WB36 had 6.1 percent 

 

Methanosarcinaceae, compared to 10.8 to 14.9 percent in 

other samples from transects A-A′ and C-C′ (table 6). It is 

possible that the generally higher concentrations of daughter 

products, especially VC, observed in the wetland porewater 

along transect C-C′ compared to transect A-A′ (Lorah and 

others, 1997; Olsen and others, 1997; Spencer and others, 

2002) could be due in part to the lower proportion of Methanosarcinaceae in the methanogen community. Additional 

molecular analyses of sediment samples along transect C-C′

would be needed to confirm this association, however.

Implications for Natural Attenuation and Remediation

Understanding TeCA and TCE degradation pathways is 

necessary in determining whether natural attenuation, or 

in situ bioremediation, of the chlorinated VOCs discharging 

to the wetlands sediments is a feasible ground-water remediation method. The rapid TeCA degradation rates in wetland 

sediments observed in this study indicate that natural attenuTable 6. Abundance of Methanosarcinaceae as a 

percentage of the total methanogen 

community in surface grab samples of 

wetland sediment, March 1999

 [Transect and site locations are shown in fig. 1]

Transect

Site

number

Percent

Methanosarcinaceae

Background site WB19 14.1

A-A′ WB23 10.8

A-A′ WB24 11.5

A-A′ WB30 14.9

C-C ′ WB35 13.2

C-C ′ WB36 6.1
Comparisons to Field Data on Anaerobic Degradation of TeCA and Association with Microbial Communities 57

ation is a feasible remediation method in the anaerobic wetland sediments. The general consistency between TeCA and 

TCE degradation rates calculated from laboratory microcosm experiments and from field data indicates that the rapid 

anaerobic degradation rates measured in the laboratory are 

reasonable. The first-order rate constant of 0.053 per day 

(half-life of 13 days) estimated for the TeCA-amended 

microcosms without MeOH is high compared to biodegradation rates reported in the literature for chlorinated solvents in 

sand aquifers. Although no other TeCA degradation rates 

from field measurements are reported in the literature, degradation rates are available for TCE under a range of conditions. For example, anaerobic TCE biodegradation in 

microcosms constructed with sandy aquifer sediments at 

Picatinny Arsenal, New Jersey, and at a Superfund site at 

St. Joseph, Michigan, ranged from 0.0001 to 0.003 per day, 

which corresponds to half-lives of 230 to 6,930 days (Rifai 

and others, 1995). Because of the commonly slow degradation of chlorinated solvents in sand aquifers, Wiedemeier 

and others (1998) recommend that batch microcosm studies 

should last from 12 to 18 months to evaluate rates for natural 

attenuation studies, compared to the 1 to 1.5 months used in 

this study with wetland sediments. The high degradation 

rates in the wetland sediments reported from the initial study 

at this APG wetland site led to the construction or plans for 

construction of wetlands at several sites for the purpose of 

remediation of chlorinated VOCs (Richard and others, 2002; 

Kassenga and Pardue, 2002; Lorah and others, 2002).

Although both laboratory and field data showed that degradation rates are slower in the wetland sediments in the 

summer compared to late winter/early spring and fall, TCE 

and TeCA degradation rates that were calculated using wetland porewater samples collected in the summer gave halflives in the range of 60 to 100 days, which still is considerably faster degradation than reported for TCE in sand aquifers (Rifai and others, 1995). The observation of lower 

summer degradation rates is contrary to the previously 

hypothesized lower degradation rates in the winter because 

of the colder temperatures (Lorah and others, 1997). Field 

measurements of temperature and VOC concentrations with 

depth in the wetland sediment indicate that winter temperature decreases are substantial only in the upper 0.15 m, 

whereas concentrations of TeCA and its anaerobic daughter 

compounds have decreased to below or near detection levels 

before this shallow depth is reached (Lorah and others, 

2000). Temperature, therefore, apparently is not a major factor affecting degradation rates in these wetland sediments. 

Based on field measurements, the higher concentrations of 

daughter VOCs in the summer (without increase in breakthrough of parent VOCs to the wetland porewater) is due to a 

higher mass flux of parent VOCs from the aquifer to the wetland sediments, which occurs as a delayed effect of high 

spring-time recharge to the aquifer upgradient of the wetland 

(Lorah and others, 2000). The seasonal laboratory microcosm experiments presented earlier in this report indicate 

that lower biodegradation rates, caused by a lower microbial 

biomass, also could be a factor in the relatively high concentrations of VOCs in the wetland porewater in the summer. 

Because the microcosm experiments and microbial community analyses were conducted using surficial wetland sediments (from the upper 25 cm), it is unknown whether the 

lower biodegradation rates also occur at greater depths. 

However, the relatively slow degradation rates, lower microbial biomass, and higher daughter compound concentrations 

observed in the summer compared to late winter/early spring 

and fall indicate that monitoring during the summer months 

would be especially crucial in long-term monitoring plans.

The microbial groups or species represented by the 

TRFLP peaks that were identified as potentially involved in 

degradation of TeCA and its daughter products in the microcosm experiments were present at all sites where wetland 

sediment samples were collected for molecular analyses (fig. 

40). Although microbial biomass and diversity were substantially lower along transect C-C′ than transect A-A′, all of 

the identified microbial groups or species were present along 

transect C-C′. Field data along both transects have shown 

that natural attenuation is effective in reducing VOC concentrations to below detection levels before the wetland surface 

is reached (Lorah and others, 1997; Lorah and Olsen, 1999a, 

b). In addition, degradation of TeCA in microcosms constructed with sediment from the background site (WB19, 

fig. 23) occurred without a lag and at a similar rate as microcosms constructed with sediment from the two contaminated 

sites (fig. 5). Field data showed that the same identified 

TRFLP peaks were present at the uncontaminated and contaminated sites (figs. 40 and 41). The combined microcosm, 

field, and microbial analyses, therefore, indicate that the 

capacity to degrade TeCA is widespread in these wetland 

sediments and that similar degradation could be expected if 

contaminants discharged to a previously uncontaminated 

area of the wetland. In previous studies, the abundance of 

organic matter (and thus of substrates to support dechlorination) has been hypothesized to be a relatively ubiquitous 

characteristic of wetland sediments that could result in an 

ubiquitous ability of wetland sediments at APG and elsewhere to degrade chlorinated solvents like TeCA and TCE 

(Lorah and others, 1997; Lorah and Olsen, 1999a, b). However, TeCA and TCE degradation rates in wetland sediments 

from this site at APG were compared to freshwater wetland 

sediments in a forested swamp in the New Jersey Pine 

Barrens in a recent field and laboratory study (Lorah and 

others, 2002). In anaerobic laboratory microcosms amended 

with the same initial TeCA or TCE concentration, the TeCA 

or TCE degraded to below the detection level in less than 

25 days in the APG wetland sediment, whereas degradation 

was negligible over a 35-day period in the New Jersey wetland sediment (Lorah and others, 2002). Although 12DCE 

was detected in field samples of the New Jersey wetland 

porewater, VC was not detected. TeCA and TCE degradation, therefore, were substantially lower and incomplete in 

the New Jersey wetland sediments, which contained a higher 

organic carbon content than the APG wetland sediments 

(Lorah and others, 2002). Differences in the microbial communities are the likely cause of the differences in degrada-
58 TeCA Degradation and Microbial Communities in a Freshwater Tidal Wetland, APG, Md.

tion rates and completeness (Lorah and others, 2002). 

Although the abundance of organic substrates is an important factor in increasing the rates of degradation compared to 

sand aquifers, the occurrence of degradation is dependent on 

the presence of specific microorganisms. Similarly, addition 

of organic substrates and nutrients to an anaerobic sand 

aquifer at a test site at Dover Air Force Base, Delaware 

could not stimulate the indigenous microorganisms to 

degrade TCE beyond cDCE (Harkness and others, 1999; 

Ellis and others, 2000). TCE dechlorination beyond cDCE 

was achieved only after injection to the aquifer test site of a 

microbial enrichment culture that was originally started with 

sediment from another site where dechlorination to ethene 

did occur (Harkness and others, 1999; Ellis and others, 

2000). Evaluation of microbial communities by the molecular techniques used in this study would be an effective 

method of evaluating the feasibility of natural attenuation at 

new wetland sites that are natural discharge areas for chlorinated solvents, and in designing constructed wetlands for 

remediation of these contaminants. Definitive identification 

of the important species/groups noted in this study and construction of gene probes would simplify use of these molecular techniques.

The feasibility of natural attenuation as a remediation 

method depends on the fate of 12DCE and VC, the toxic 

compounds that laboratory experiments and field measurements show are the predominant, persistent daughter products (figs. 7–9 and 39). Although substantial spatial 

variation in degradation rates of TeCA was not observed in 

the laboratory experiments or in field estimates, both laboratory and field data indicate that spatial variation in the production and degradation of VC does occur. The substantially 

higher production of VC at site WB30 compared to site 

WB23 could hinder the use of natural attenuation as a remediation method, if the conditions leading to higher VC 

production were widespread in the wetland. The fact that the 

same sediments that produced high VC also showed a 

greater capacity to degrade VC, however, alleviates this 

problem. Geochemical analyses of the microcosm experiments indicated that VC production from both 112TCA and 

12DCE is linked to methanogenic activity, although microbial community analysis indicated that bacteria rather than 

methanogens were associated with the 112TCA degradation 

reaction. Microcosm experiments amended with BES and 

enrichment culture experiments also indicated that VC production from 112TCA and 12DCE could occur without 

methane production. However, because the enrichment 

cultures were inoculated with methanogenic microcosm 

slurry, the possibility that methanogens enhance degradation 

of these compounds without producing methane cannot be 

excluded. VC degradation seemed to be more strongly associated with methanogenesis than was VC production, as 

shown by the negligible VC degradation in the BESamended microcosms and the rapid VC degradation in methanogenic VC/sediment enrichments compared to BES- and 

FeOOH-amended enrichments. The strong relation 

observed between VC degradation and relatively high abundance of acetotrophic methanogens suggests that complete 

degradation of TeCA requires methanogenesis. However, 

during periods of lower methanogenic activity, there was an 

increase in significance of other VC degradation pathways. 

This is evidenced by the fact that VC degradation occurred 

in the summer when low methanogenic activity was 

observed, although TeCA degradation rates were lower (figs. 

5, 8, and 13). This non-methanogenic VC degradation most 

likely was through the action of dehalorespiring bacteria, 

such as the Dehalococcoides or Desulfuromonas species that 

were detected at this site using specific primers. Seasonal 

shifts in the microbial species or groups that seem to be 

involved in degradation also occurred for the production of 

VC from 112TCA dichloroelimination. Thus, the adaptable 

and diverse microbial community of the wetland sediments 

seems to provide compensation for changing spatial and seasonal conditions and allows for continued effective natural 

attenuation throughout the year.

The diversity of microorganisms involved in degradation 

of TeCA in these wetland sediments provides options to 

increase biodegradation rates in areas such as seeps, where 

high concentrations of VOCs (up to 50,000 μg/L) have been 

measured near the wetland surface (Phelan and others, 

2001). Possible enhancements to increase biodegradation 

include “biostimulation,” which involves adding substances 

such as nutrients to stimulate indigenous microorganisms 

present at a site, and “bioaugmentation,” which involves 

adding microorganisms to a site that is lacking in one or 

more microbial species/groups needed for biodegradation. It 

is unclear from the laboratory and field data presented here 

whether the acetrotrophic methanogens are directly involved 

in VC degradation or are indirectly involved by enhancing 

the degradation capability of dehalorespiring bacteria or of 

acetogens, as Bradley and Chapelle (2000) suggested. If 

acetotrophic methanogens are indirectly involved, enhancing 

either their population or enhancing the bacteria that they are 

supporting could be a possible remediation option. Molecular analysis of the microbial communities in the VC-degrading enrichment experiments could provide definitive 

evidence of whether their role is direct or indirect. VC/sediment enrichment experiments showed that WB23 and WB30 

wetland sediments had equally great capacities to degrade 

high concentrations of VC after pre-exposure to the contaminants and prolonged incubation under methanogenic conditions. The presence of microorganisms necessary for 

degradation, therefore, may be widespread in the wetland 

sediment, but populations may need to be enhanced in some 

cases. Enhancing the acetotrophic methanogen population 

in the wetland sediments where enhanced degradation rates 

might be needed could be more practical than enhancing the 

population of dehalorespiring bacteria.
Summary and Conclusions 59

Summary and Conclusions

Defining biodegradation rates and processes is a critical 

part of assessing the feasibility of monitored natural attenuation as a remediation method for ground water containing 

organic contaminants because biodegradation is commonly 

the primary destructive process for these contaminants. 

During 1998–2001, the U.S. Geological Survey conducted a 

microbial study at a freshwater tidal wetland at Aberdeen 

Proving Ground (APG), Maryland, where an ongoing investigation of natural attenuation of chlorinated volatile organic 

compounds (VOCs) in the wetland sediments began in 1992. 

In this microbial study, a combination of geochemical analyses and molecular biology techniques were used to investigate factors controlling the occurrence of the different 

anaerobic degradation pathways of 1,1,2,2-tetrachloroethane 

(TeCA) and to characterize the microbial communities that 

potentially are important in these degradation reactions. 

Laboratory microcosm and enrichment experiments were 

used to investigate the effects of spatial and seasonal variability, redox conditions, substrate type, and toxicity on the 

degradation reactions. Field measurements then were used 

to compare degradation rates, distributions of daughter products, and microbial community profiles to laboratory results 

and to discuss the implications on natural attenuation in the 

wetland. Geochemical analyses included measurements of 

VOCs and redox-sensitive constituents. Microbial analyses 

included terminal-restriction fragment length polymorphism 

(TRFLP) analysis of PCR (polymerase chain reaction)-

amplified bacterial 16S rDNA and mcrA (methylcoreductase 

gene) to obtain profiles of the bacterial and methanogen 

communities, respectively. These genetic analyses commonly are referred to as “DNA fingerprinting.” An intense 

microbial study to support the natural attenuation investigation at this site was warranted because of the expected complex pathways of TeCA degradation, the limited previous 

information available on degradation of this contaminant in 

ground water, and the expected large variability in biodegradation rates and processes in a dynamic, surficial wetland 

system. A better understanding of factors controlling production and degradation of toxic intermediate daughter compounds, particularly vinyl chloride (VC), was needed to 

evaluate the effectiveness of natural attenuation in the wetland sediments.

Anaerobic microcosm experiments conducted with wetland sediment collected from two different sites (WB23 and 

WB30) and during three different seasons (March–April 

1999, July–August 1999, and October–November 2000) 

showed little spatial variability but high seasonal variability 

in TeCA degradation rates. Seasonal evaluations of degradation rates included microcosms incubated at 19 °C (degrees 

Celsius) during the three periods and a duplicate set of 

microcosms in March–April 1999 that was incubated at 5 °C. 

For the March–April 1999 and October–November 2000 

microcosms incubated at 19 °C, initial first-order rate constants (generally calculated from TeCA loss in the first 12 

days) ranged between 0.10±0.01 and 0.16±0.05 per day 

(half-lives of 4.3 to 6.9 days) for degradation of TeCA, 

whereas rate constants of 0±0.03 and 0.06±0.03 per day 

were obtained in the July–August 1999 microcosms. Microbial community profiles showed that low microbial biomass 

and microbial diversity in the summer could account for 

these unexpectedly low degradation rates. Although it is 

unknown if these low summertime rates are a recurrent 

event, competition for nutrients by growth of the Phragmites

(common reed) that dominates wetland vegetation could 

reduce microbial biomass during the summer. Degradation 

of TeCA also was observed in the March–April 1999 microcosms incubated at 5 °C, indicating that biodegradation 

would continue during cold winter temperatures in the wetland sediment. About 50 percent of the initial TeCA in solution was converted to daughter products within the 35-day 

incubation period at 5 °C, whereas complete loss of the 

added TeCA occurred within 21 days in duplicate microcosms incubated at 19 °C.

Simultaneous production of 1,2-dichloroethene (12DCE) 

and 1,1,2-trichloroethane (112TCA) from dichloroelimination and hydrogenolysis of TeCA, respectively, occurred 

without a lag in all live microcosm experiments that were 

conducted. Although this initial TeCA degradation occurred 

at a similar rate in the microcosms constructed with WB23 

and WB30 sediments, VC production and degradation in the 

microcosms showed substantial spatial variability (variability associated with the location from which the sediment was 

collected for microcosm construction). The two major pathways of VC production were from dichloroelimination of 

112TCA and from hydrogenolysis of 12DCE that was produced by dichloroelimination of TeCA. Abiotic production 

of trichloroethene (TCE) was not higher than 11 percent, 

indicating that hydrogenolysis of TCE is not a substantial 

source of VC in the wetland porewater. The TeCA- and 

112TCA-amended microcosms constructed with WB30 sediment showed approximately twice as much VC production 

from 112TCA dichloroelimination as those constructed with 

WB23 sediment. VC degradation, however, also was substantially higher in the TeCA-amended microcosms constructed with WB30 sediment than those constructed with 

WB23 sediment. In microcosms constructed with WB30 

sediment, between 30 to 96 percent of the VC was degraded 

within the 35-day incubation period, whereas 0 to 14 percent 

of VC was degraded in the microcosms constructed with 

WB23 sediment. Addition of methanol to microcosms 

enhanced VC degradation in both WB23 and WB30 wetland 

sediments. Enrichment experiments amended with VC 

showed that the spatial difference in VC degradation was 

negligible after prolonged incubation under methanogenic 

conditions at high initial VC concentrations. Production of 

VC in enrichment cultures with 112TCA was linear, whereas 

production of VC in enrichment cultures with 12DCE was 

exponential and occurred after a lag.

Addition of an inhibitor of methanogenic activity (BES, 

or 2-bromoethanesulfonic acid) slightly lowered TeCA deg-
60 TeCA Degradation and Microbial Communities in a Freshwater Tidal Wetland, APG, Md.

radation rates but did not inhibit production of 112TCA, 

12DCA, and 12DCE. VC and chloroethane production, 

however, were substantially inhibited in the BES-amended 

microcosms. The lower VC production in the BES-amended 

microcosms is consistent with the association noted between 

onset of methane production and of VC production from 

112TCA dichloroelimination in many of the microcosms. 

Inhibition of methanogenesis by addition of BES also completely inhibited 12DCE and VC degradation in the microcosms and inhibited VC degradation in enrichment 

experiments with VC-amended sediment. VC degradation 

in enrichment experiments with VC-amended sediment also 

was inhibited under iron-reducing conditions.

Pre-exposure of the wetland sediment to TeCA was not 

required for degradation of TeCA, as shown by equally rapid 

TeCA degradation rates in microcosms constructed with 

wetland sediment from the contaminated sites WB23 and 

WB30, and from an uncontaminated site WB19. Pre-exposure of sediments to VC, however, substantially increased 

VC degradation rates in enrichment experiments.

A microbial consortium, rather than one individual 

microorganism or microbial group, is involved in the degradation of TeCA, as indicated by the occurrence of multiple 

degradation pathways and the variability in daughter product 

distributions in the microcosms with the site or season from 

which the sediment was collected. To identify possible critical microbial species or groups involved in degradation of 

TeCA, shifts in the TRFLP profiles (DNA fingerprints) of 

the microbial communities in the microcosm sediments over 

time and among different experiments were compared to 

shifts in the daughter product distributions to note consistent 

associations. Changes in degradation pathways observed 

from spatial and seasonal variability were linked to changes 

in the microbial communities. A bacterial peak at 90 base 

pair (bp) fragment length was linked to TeCA hydrogenolysis to 112TCA. Increases in intensity of a few bacterial species were observed in killed controls that showed recovery 

from the formaldehyde by the end of the experiment. The 

only daughter product observed in these killed controls was 

12DCE from TeCA dichloroelimination, and growth of species represented by 198 and 170 bp fragment lengths were 

associated with the production of 12DCE. Dichloroelimination of 112TCA to VC also was associated with growth 

of this 198 bp bacterial peak in March–April 1999 and 

October–November 2000 microcosms, whereas an 86 bp or 

the 170 bp bacterial peak was associated with 112TCA 

dichloroelimination in the July–August 1999 experiment. 

Hydrogenolysis of 12DCE to VC was associated with a 

CO2-utilizing methanogen at 307 bp in the March–April 

1999 and October–November 2000 microcosm experiments, 

whereas methanogen biomass was low in the July–August 

1999 experiments.

Production of VC that occurred without methane production in the July–August 1999 microcosms constructed with 

sediment from WB30 and in 112TCA-amended and 12DCEamended enrichment cultures indicated the involvement of 

other microorganisms in VC production in the wetland sediments. The immediate, linear production of VC in the 

112TCA-amended enrichment cultures indicated a cometabolic reaction. In contrast, the exponential increase of VC in 

12DCE-amended enrichment cultures after an initial lag 

indicated growth of a microbial species or group, possibly 

the dehalorespiring bacteria. Molecular analyses showed the 

presence of known dehalorespiring bacteria of the Dehalococcoides group (Dehalococcoides ethenogenes and Dehalococcoides sp. strain FL2) and of the acetate-oxidizing 

Desulfuromonas group (Desulfuromonas sp. strain BB1 and 

Desulfuromonas chloroethenica) in microcosm slurry from 

WB30, but not WB23. The lower activity of methanogens in 

the July–August 1999 microcosms may have allowed the 

activity of the dehalorespiring bacteria to increase because 

methanogens compete with dehalorespiring bacteria for 

hydrogen as a substrate. Both hydrogen, which is a favored 

substrate for Dehalococcoides, and acetate, which is a 

favored substrate for Desulfuromonas, stimulated VC production in enrichment experiments with 12DCE-amended 

cultures. Three times as much VC, however, was produced 

with hydrogen than with acetate in these enrichment experiments, indicating a greater population of hydrogen-utilizing 

than acetate-utilizing bacteria in the sediment. Ethene, 

which is an end product of dechlorination of VC by Dehalococcoides ethenogenes, was produced only in enrichments 

supplied with hydrogen.

The higher VC degradation in microcosms constructed 

with WB30 sediment compared to WB23 sediment and in 

microcosms amended with methanol showed a marked association with an increase of acetotrophic methanogens, a 

group of methanogens that can utilize acetate as a substrate. 

The apparent link between VC degradation and acetotrophic 

methanogens observed in these microcosm experiments may 

support the theory that VC mineralization under anaerobic 

conditions begins with oxidation by acetogens and that acetotrophic methanogens enhance degradation, although the 

involvement of acetogens in the experiments reported here is 

purely speculative. The microcosm and enrichment culture 

experiments indicate that VC can be produced and degraded 

by more than one reaction pathway and associated microorganisms in the wetland sediments, and the pathway that is 

dominant can change with changing seasons, location in the 

wetland, redox conditions, or substrate type.

Estimates of TeCA and TCE degradation rates using field 

data of porewater concentrations and flow velocity in the 

wetland sediments showed that the rapid anaerobic degradation rates measured in the laboratory are reasonable. Halflives for TeCA and TCE estimated from field data were in 

the range of 60 to100 days, which agrees well with laboratory estimates considering the inherent differences in the laboratory and field systems. The relative proportions of 

daughter products in the wetland porewater also are consistent with laboratory microcosm experiments, which showed 

12DCE and VC were the predominant, persistent daughter 

compounds from TeCA degradation. In addition, the porewater chemistry showed higher production of VC in the wetland sediment at site WB30 than at site WB23, as was 
Summary and Conclusions and References Cited 61

observed in the microcosm experiments. Molecular analyses 

of grab samples of surficial wetland sediment showed that all 

of the microbial species or groups linked to TeCA degradation in the microcosm experiments were present in all the 

sediment samples. Microbial biomass and diversity were 

lowest in an area of the wetland where porewater VOC concentrations are highest (transect C-C′), indicating that the 

higher VOC concentrations could result from lower degradation rates. The lower microbial biomass and diversity along 

transect C-C′ could be caused from toxicity effects of the 

contaminants, or possibly from differences in frequency and 

duration of tidal inundation. Land-surface elevation is lower 

along transect C-C′ than along A-A′, causing greater and 

more frequent tidal inundation, which would affect redox 

and other physiochemical conditions in the surficial wetland 

sediments. A toxicity effect also could be associated with 

tidal inundation because low concentrations of VOCs, 

including chloroform, were measured in the surface water.

The rapid TeCA and daughter product degradation measured in the wetland sediments observed in this study confirm that natural attenuation is a feasible remediation method 

in the anaerobic wetland sediments. The adaptable and 

diverse microbial community that appears to be involved in 

TeCA degradation in the wetland sediments provides compensation for changing spatial and seasonal conditions and 

allows natural attenuation to be effective throughout the 

year. The lower degradation rates in the summer, however, 

indicate that ground-water and surface-water monitoring 

during the summertime is especially crucial. Definitive 

identification of the microbial species involved in TeCA 

degradation by completion of genetic analyses of the enrichment experiments would allow development of gene probes 

that could be used for long-term monitoring of the presence 

of critical microbial species over time in the wetland sediment, and for evaluating the potential for natural attenuation 

at other contaminated sites. Definitive identifications also 

would assist in evaluating possible enhancements, including 

“biostimulation” or “bioaugmentation,” that could be used to 

increase biodegradation rates in areas such as seeps where 

higher concentrations of VOCs have been measured at the 

wetland surface. Because anaerobic TeCA degradation 

encompasses chlorinated ethane and ethene pathways, 

results of this study are transferable to other sites at APG and 

elsewhere that have TeCA, TCE, or their anaerobic degradation products as contaminants.

References Cited

Baedecker, M.J., and Cozzarelli, I.M., 1992, The 

determination and fate of unstable constituents in 

contaminated groundwater, in S. Lesage and R.E. 

Jackson (eds.), Groundwater Contamination and 

Analysis at Hazardous Waste Sites: New York, Marcel 

Dekker, p. 425–461.

Bagley, D.M., and Gossett, J.M., 1990, Tetrachloroethene 

transformation to trichloroethene and cis-1,2-

dichloroethene by sulfate-reducing enrichment cultures: 

Applied and Environmental Microbiology, v. 56, no. 8, 

p. 2,511–2,516.

Belay, Negash, and Daniels, Lacy, 1987, Production of 

ethane, ethylene, and acetylene from halogenated 

hydrocarbons by methanogenic bacteria: Applied 

and Environmental Microbiology, v. 53, no. 7, 

p. 1,604–1,610.

Bradley, P.M., and Chapelle, F.H., 1996, Anaerobic 

mineralization of vinyl chloride in Fe(III)-reducing 

aquifer sediments: Environmental Science and 

Technology, v. 30, no. 6, p. 2,084–2,086.

_____ 1998, Microbial mineralization of VC and DCE under 

different terminal electron accepting conditions: 

Anaerobe, v. 4, p. 81–87.

_____ 1999a, Methane as a product of chloroethene 

biodegradation under methanogenic conditions: 

Environmental Science and Technology, v. 33, no. 4, 

p. 653–656.

_____ 1999b, Role for acetotrophic methanogens in 

methanogenic biodegradation of vinyl chloride: 

Environmental Science and Technology, v. 33, no. 19, 

p. 3,473–3,476.

_____ 2000, Acetogenic microbial degradation of vinyl 

chloride: Environmental Science and Technology, v. 34, 

no. 13, p. 2,761–2,763.

Bradley, P.M., Chapelle, F.H., and Lovley, D.R., 1998, 

Humic acids as electron acceptors for anaerobic 

microbial oxidation of vinyl chloride and dichloroethene: 

Applied and Environmental Microbiology, v. 64, no. 8, 

p. 3,102–3,105.

Brunk, C.F., Avaniss-Aghajani, E., and Brunk, C.A., 

1996, A computer analysis of primer and probe 

hybridization potential with bacterial small-subunit 

rRNA sequences: Applied and Environmental 

Microbiology, v. 62, p. 872–879.

Capone, D.G., and Kiene, R.P., 1988, Comparison of 

microbial dynamics in marine and freshwater sediments: 

Contrasts in anaerobic catabolism: Limnology and 

Oceanography, v. 33, p. 725–749.

Chapelle, F.H., 1993, Ground-water microbiology and 

geochemistry: New York, John Wiley & Sons, 424 p.

Chapelle, F.H., and Bradley, P.M., 1998, Selecting 

remediation goals by assessing the natural attenuation 

capacity of groundwater systems: Bioremediation 

Journal, v. 2, nos. 3 & 4, p. 227–238.
62 TeCA Degradation and Microbial Communities in a Freshwater Tidal Wetland, APG, Md.

Chen, Chun, Puhakka, J.A., and Ferguson, J.F., 1996, 

Transformations of 1,1,2,2-tetrachloroethane under 

methanogenic conditions: Environmental Science and 

Technology, v. 30, no. 2, p. 542–547.

Clement, B.G., Kehl, L.E., DeBord, K.L. and Kitts, C., 

1998, Terminal restriction fragment patterns (TRFPs), a 

rapid, PCR-based method for the comparison of complex 

bacterial communities: Journal of Microbiological 

Methods, v. 31, p. 135–142.

Dyer, Linda Jo, Lorah, M.M., and Burris, D.R., 2002, 

Effect of sampling method on measured porewater 

concentrations in a wetland contaminated by chlorinated 

solvents, in K.W. Nehring and S.E. Brauning (eds.), 

Proceedings of the Second International Conference on 

Wetlands & Remediation, September 5–6, 2001, 

Burlington, Vermont: Columbus, Ohio, Battelle Press, 

p. 33–40.

Ellis, D.E., Lutz, E.J., Odom, J.M., Buchanan, R.J., Jr., 

Lee, M.D., Bartlett, C.L., Harkness, M.R., and 

Deweerd, K.A., 2000, Bioaugmentation for accelerated 

in situ anaerobic bioremediation: Environmental Science 

and Technology, v. 34, no. 11, p. 2,254–2,260.

Fathepure, B.Z., and Boyd, S.A., 1988, Dependence of 

tetrachloroethylene dechlorination on methanogenic 

substrate consumption by Methanosarcina sp. strain 

DCM: Applied and Environmental Microbiology, v. 54, 

p. 2,976–2,980.

Fennel, D.E., Carroll, A.B., Gossett, J.M, and Zinder, 

S.H., 2001, Assessment of indigenous reductive 

dechlorinating potential at a TCE-contaminated site 

using microcosms, polymerase chain reaction analysis, 

and site data: Environmental Science and Technology, 

v. 35, no. 9, p. 1,830–1,839.

Fennel, D.E., and Gossett, J.M., 1998, Modeling the 

production of and competition for hydrogen in a 

dechlorinating culture: Environmental Science and 

Technology, v. 32, p. 2,450–2,460.

Flynn, S.J., Löffler, F.E., and Tiedje, J.M., 2000, 

Microbial community changes associated with a shift 

from reductive dechlorination of PCE to reductive 

dechlorination of cis-DCE and VC: Environmental 

Science and Technology, v. 34, no. 6, p. 1,056–1,061.

Gantzer, C.J., and Wackett, L.P., 1991, Reductive 

dechlorination catalyzed by bacterial transition-metal 

coenzymes: Applied and Environmental Microbiology, 

v. 55, p. 2,144–2,151.

Harkness, M.R., Bracco, A.A., Brennan, M.J., Jr., 

Deweerd, K.A., and Spivack, J.L., 1999, Use of 

bioaugmentation to stimulate complete reductive 

dechlorination of trichloroethene in Dover soil columns: 

Environmental Science and Technology, v. 33, no. 7, 

p. 1,100–1,109.

Holliger, Christof, Schraa, Gosse, Stams, A.J.M., and 

Zehnder, A.J.B., 1993, A highly purified enrichment 

culture couples the reductive dechlorination of 

tetrachloroethene to growth: Applied and Environmental 

Microbiology, v. 59, no. 9, p. 2,991–2,997.

Jafvert, C.D., and Wolfe, N. Lee, 1987, Degradation of 

selected halogenated ethanes in anoxic sediment-water 

systems: Environmental Toxicology and Chemistry, v. 6, 

p. 827–837.

Kassenga, G.R., and Pardue, J.H., 2002, Fate of TCE in 

constructed and natural wetland mesocosms, in K.W. 

Nehring and S.E. Brauning (eds.), Proceedings of the 

Second International Conference on Wetlands & 

Remediation, September 5–6, 2001, Burlington, 

Vermont: Columbus, Ohio, Battelle Press, p. 25–32.

Kästner, Matthias, 1991, Reductive dechlorination of 

tri- and tetrachloroethylenes depends on transition 

from aerobic to anaerobic conditions: Applied and 

Environmental Microbiology, v. 57, no. 7, p. 2,039–

2,046.

Klein, A., Allmansberger, R., Bokranz, M., Knaub, S., 

Muller, B. and Muth, E. 1988, Comparative analysis of 

genes encoding methylcoenzyme M reductase in 

methanogenic bacteria: Molecular and General 

Genetics, v. 213, p. 409–420.

Krumholz, L.R., Sharp, Richard, and Fishbain, S.S., 

1996, A freshwater anaerobe coupling acetate oxidation 

to tetrachloroethylene dehalogenation: Applied and 

Environmental Microbiology, v. 62, no. 11, p. 4,108–

4,113.

Liang, Li-Nuo, and Grbic ́-Galic ́, D., 1993, 

Biotransformation of chlorinated aliphatic solvents in the 

presence of aromatic compounds under methanogenic 

conditions: Environmental Toxicology and Chemistry, 

v. 12, p. 1,377–1,393.

Löffler, F.E., Sanford, R.A., and Tiedje, J.M., 1996, Initial 

characterization of a reductive dehalogenase from 

Desulfitobacterium chlororespirans Co23: Applied 

and Environmental Microbiology, v. 62, no. 10, 

p. 3,809–3,813.

Löffler, F.E., Sun, Qing, Li, Jieran, and Tiedje, J.M., 

2000, 16S rRNA gene-based detection of 

tetrachloroethene-dechlorinating Desulfuromonas and

Dehalococcoides species: Applied and Environmental 

Microbiology, v. 66, no. 4, p. 1,369–1,374.

Lorah, M.M. and Clark, J.S., 1996, Contamination of 

ground water, surface water, and soil, and evaluation 

of selected ground-water pumping alternatives in the 

Canal Creek area of Aberdeen Proving Ground, 

Maryland: U.S. Geological Survey Open-File Report 

95–282, 318 p.

Lorah, M.M., Olsen, L.D., Smith, B.L., Johnson, M.A., 

and Fleck, W.B., 1997, Natural attenuation of 

chlorinated volatile organic compounds in a freshwater 

tidal wetland, Aberdeen Proving Ground, Maryland: 

U.S. Geological Survey Water-Resources Investigations 

Report 97–4171, 95 p.

Lorah, M.M., and Olsen, L.D., 1999a, Degradation of 

1,1,2,2-tetrachloroethane in a freshwater tidal wetland: 

Field and laboratory evidence: Environmental Science 

and Technology, v. 33, no. 2, p. 227–234.
References Cited 63

_____ 1999b, Natural attenuation of chlorinated volatile 

organic compounds in a freshwater tidal wetland: Field 

evidence of anaerobic biodegradation: Water Resources 

Research, v. 35, no. 12, p. 3,811–3,827.

Lorah, M.M., Olsen, L.D., and Johnson, J.C., 2000, 

Temporal dynamics of biogeochemical processes in a 

contaminated wetland, Aberdeen Proving Ground, 

Maryland: American Geophysical Union 2000 Spring 

Meeting, Washington, D.C., May 30–June 3, 2000. 

Eos, Transactions, American Geophysical Union, 

v. 81, no. 19, p. S197.

Lorah, M.M., Olsen, L.D., Capone, D.G., and 

Baker, J.E., 2001, Biodegradation of trichloroethylene 

and its anaerobic daughter products in freshwater 

wetland sediments: Bioremediation Journal, v. 5, no. 2, 

p. 101–118.

Lorah, M.M., Burris, D.R., and Dyer, L.J., 2002, 

Efficiency of natural attenuation of chlorinated solvents 

in two freshwater wetlands, in K.W. Nehring and 

S.E. Brauning (eds.), Proceedings of the Second 

International Conference on Wetlands & Remediation, 

September 5–6, 2001, Burlington, Vermont: Columbus, 

Ohio, Battelle Press, p. 9–16.

Lorah-Devereux, M.M., 1999, Natural attenuation of 

chlorinated volatile organic compounds in a freshwater 

tidal wetland: College Park, Maryland, University of 

Maryland, Ph.D. Dissertation, 210 p.

Lovley, D.R., and Phillips, E.J.P., 1987, Rapid assay for 

microbially reducible ferric iron in aquatic sediments: 

Applied and Environmental Microbiology, v. 53, no. 7, 

p. 1,536–1,540.

Luton, P.E., 1996, A study of landfill methanogens: 

Liverpool, England, Liverpool John Moores University, 

PhD Dissertation, [variously, paged].

Maymó-Gatell, Xavier, Anguish, Timothy, and Zinder, 

S.H., 1999, Reductive dechlorination of chlorinated 

ethenes and 1,2-dichloroethane by “Dehalococcoides 

ethenogenes” 195: Applied and Environmental 

Microbiology, v. 65, no. 7, p. 3,108–3,113.

Maymó-Gatell, Xavier, Chien, Yueh-Tyng, Gossett, J.M., 

and Zinder, S.H., 1997, Isolation of a bacterium that 

reductively dechlorinates tetrachloroethene to ethene: 

Science, v. 276, p. 1,568–1,571.

Maymó-Gatell, Xavier, Nijenhuis, Ivonne, and Zinder, 

S.H., 2001, Reductive dechlorination of cis-1,2-

dichloroethene and vinyl chloride by “Dehalococcoides 

ethenogenes”: Environmental Science and Technology, 

v. 35, no. 3, p. 516–521.

Maymó-Gatell, Xavier, Tandoi, Valter, Gossett, J.M., and 

Zinder, S.H., 1995, Characterization of an H2-utilizing 

enrichment culture that reductively dechlorinates 

tetrachloroethene to vinyl chloride and ethene in the 

absence of methanogenesis and acetogenesis: Applied 

and Environmental Microbiology, v. 61, no. 11, 

p. 3,928–3,933.

McCarty, P.L., and Semprini, Lewis, 1994, Ground-water 

treatment for chlorinated solvents, in R.D. Norris and 

others, Handbook of bioremediation: Ann Arbor, 

Michigan, Lewis Publishers, p. 87–116.

Mitsch, W.J., and Gosselink, J.G., 1993, Wetlands (2d

Edition): New York, John Wiley & Sons, 722 p.

National Research Council, 2000, Natural attenuation for 

groundwater remediation: Washington, D.C., National 

Academy Press, 274 p.

Novak, P.J., Daniels, L., and Parkin, G.F., 1998, Rapid 

dechlorination of carbon tetrachloride and chloroform 

by extracellular agents in cultures of Methanosarcina 

thermophila: Environmental Science and Technology, 

v. 32, no. 20, p. 3,132–3,136.

Olsen, L.D., Lorah, M.M., Marchand, E.H., Smith, B.L., 

and Johnson, M.A., 1997, Hydrogeologic, waterquality, and sediment-quality data for a freshwater tidal 

wetland, West Branch Canal Creek, Aberdeen Proving 

Ground, Maryland, 1992–96: U.S. Geological Survey 

Open-File Report 97–560, 267 p.

Oremland, R.S., and Capone, D.G., 1988, Use of "specific" 

inhibitors in biogeochemistry and microbial ecology, in

K.C. Marshall (ed.), Advances in microbial ecology: 

New York, Plenum Press, v. 10, p. 285–383.

Pankow, J.F., and Cherry, J.A., 1996, Dense chlorinated 

solvents: Portland, Oregon, Waterloo Press, 522 p.

Pavlostathis, S.G., and Zhuang, Ping, 1991, 

Transformation of trichloroethylene by sulfate-reducing 

cultures enriched from a contaminated subsurface soil: 

Applied Microbiology and Biotechnology, v. 36, 

p. 416–420.

Peijnenburg, Willie, Eriksson, Lennart, de Groot, 

Arthur, Sjostrom, Michael, and Verboom, Hans, 1998, 

The kinetics of reductive dehalogenation of a set of 

halogenated aliphatic hydrocarbons in anaerobic 

sediment slurries: Environmental Science & Pollution 

Research, v. 5, no. 1, p. 12–16.

Phelan, D.J., Olsen, L.D., Senus, M.P., and Spencer, T.A., 

2001, Assessment of volatile organic compounds in 

surface water at Canal Creek, Aberdeen Proving Ground, 

Maryland, November 1999–September 2000: U.S. 

Geological Survey Open-File Report 01–292, 49 p.

Ravi, V., Chen, J., Wilson, J.T., Johnson, J.A., 

Gierke, W., and Murdie, L., 1998, Evaluation of natural 

attenuation of benzene and dichloroethanes at the KL 

Landfill: Bioremediation Journal, v. 2, nos. 3 & 4, 

p. 239–258.

Richard, D.E., Connell, Doug, Berns, J.J., and Hirl, P.J., 

2002, Assessment of a reconstructed wetland for 

remediation of chlorinated solvents, in K.W. Nehring and 

S.E. Brauning (eds.), Proceedings of the Second 

International Conference on Wetlands & Remediation, 

September 5–6, 2001, Burlington, Vermont: Columbus, 

Ohio, Battelle Press, p. 117–124.
64 TeCA Degradation and Microbial Communities in a Freshwater Tidal Wetland, APG, Md.

Rifai, H.S., Borden, R.C., Wilson, J.T., and Ward, C.H., 

1995, Intrinsic bioattenuation for subsurface 

restoration, in R.E. Hinchee and others (eds.), Intrinsic 

bioremediation: Columbus, Ohio, Battelle Press, 

p. 1–29.

Rose, D.L., and Schroeder, M.P., 1995, Methods of 

analysis by the U.S. Geological Survey National WaterQuality Laboratory—Determination of volatile organic 

compounds in water by purge and trap capillary gas 

chromatography/mass spectrometry: U.S. Geological 

Survey Open-File Report 94–708, 26 p.

Schanke, C.A., and Wackett, L.P., 1992, Environmental 

reductive elimination reactions of polychlorinated 

ethanes mimicked by transition-metal coenzymes: 

Environmental Science and Technology, v. 26, no. 4, 

p. 830–833. 

Scholten, J.C.M., Conrad, R., and Stams, A.J.M., 2000, 

Effect of 2-bromo-ethane sulfonate, molybdate and 

chloroform on acetate consumption by methanogenic 

and sulfate-reducing populations in freshwater sediment: 

FEMS Microbiology Ecology, v. 32, p. 35–42.

Schwarzenbach, R.P., Gschwend, P.M., and Imboden, 

D.M., 1993, Environmental Organic Chemistry: 

New York, John Wiley & Sons, 681 p.

Semprini, Lewis, Kitanidis, P.K., Kampbell, D.H., and 

Wilson, J.T., 1995, Anaerobic transformation of 

chlorinated aliphatic hydrocarbons in a sand aquifer 

based on spatial chemical distributions: Water Resources 

Research, v. 31, no. 4, p. 1,051–1,062.

Smatlak, C.R., Gossett, J.M, and Zinder, S.H., 1996, 

Comparative kinetics of hydrogen utilization for 

reductive dechlorination of tetrachloroethene and 

methanogenesis in an anaerobic enrichment culture: 

Environmental Science and Technology, v. 30, no. 9, 

p. 2,850–2,858.

Spencer, T.A., Olsen, L.D., Lorah, M.M., and 

Mount, M.M., 2000, Water-quality and water-level data 

for a freshwater tidal wetland, West Branch Canal Creek, 

Aberdeen Proving Ground, Maryland, October 1998–

September 1999: U.S. Geological Survey Open-File 

Report 00–282, 184 p.

Spencer, T.A., Phelan, D.J., Olsen, L.D., and 

Lorah, M.M., 2002, Water-quality data for a freshwater 

tidal wetland, West Branch Canal Creek, Aberdeen 

Proving Ground, Maryland, November 1999—

May 2001: U.S. Geological Survey Open-File Report 

01-420, 295 p.

Tomassoni, Guy, 2000, A federal statutory/regulatory/

policy perspective on remedial decision-making with 

respect to ground-water/surface-water interaction, in

Proceedings of the ground-water/surface-water 

interactions workshop: EPA/542/R–00/007, p. 13–14.

U.S. Environmental Protection Agency, 1998, 

Announcement of the drinking water contaminant 

candidate list: Federal Register, v. 63, no. 40, 

March 2, 1998, p. 10,273–10,287.

_____ 2001, National Primary Drinking Water Regulations, 

Office of Water, March 2001: Available at http://

www.epa.gov/safewater/mcl.html.

Vogel, T. M., Criddle, C.S., and McCarty, P.L., 1987, 

Transformations of halogenated aliphatic compounds: 

Environmental Science and Technology, v. 21, no. 8, 

p. 722–736.

Vogel, T.M., and McCarty, P.L., 1985, Biotransformation 

of tetrachloroethylene to trichloroethylene, 

dichloroethylene, vinyl chloride, and carbon dioxide 

under methanogenic conditions: Applied and 

Environmental Microbiology, v. 49, p. 1,080–1,083.

Weathers, L.J., and Parkin, G.F., 2000, Toxicity of 

chloroform biotransformation to methanogenic bacteria: 

Environmental Science and Technology, v. 34, no. 13, 

p. 2,764–2,767.

Wiedemeier, T.H., Swanson, M.A., Moutoux, D.E., 

Gordon, E.K., Wilson, J.T., Wilson, B.H., Kampbell, 

D.H., Hansen, J.E., Haas, P., and Chapelle, F.H., 1998, 

Technical protocol for evaluating natural attenuation of 

chlorinated solvents in ground water: United States 

Environmental Protection Agency, EPA/600/R–98/128, 

September 1998, Available at http://www.epa.gov/ada/

reports.html.

Yager, R.M., Bilotta, S.E., Mann, C.L., and Madsen, E.L., 

1997, Metabolic adaptation and in situ attenuation of 

chlorinated ethenes by naturally occurring 

microorganisms in a fractured dolomite aquifer near 

Niagara Falls, New York: Environmental Science and 

Technology, v. 31, no. 11, p. 3,138–3,147.