Evaluation of the Role of 

Dehalococcoides

Organisms in the Natural 

Attenuation of Chlorinated 

Ethylenes in Ground Water

EPA/600/R-06/029

July 2006

Evaluation of the Role of Dehalococcoides

Organisms in the Natural Attenuation of 

Chlorinated Ethylenes in Ground Water

Xiaoxia Lu

National Research Council Post Doctoral Associate

tenable at the

U.S. Environmental Protection Agency 

Office of Research and Development 

National Risk Management Laboratory

Ada, Oklahoma 74820

Donald H. Kampbell, and John T. Wilson

U.S. Environmental Protection Agency 

Office of Research and Development 

National Risk Management Laboratory

Ada, Oklahoma 74820

Support from the U.S. Air Force Center for 

Environmental Excellence through 

Interagency Agreement # RW-57939566

Project Officer

John T. Wilson

Ground Water and Ecosystems Restoration Division

National Risk Management Research Laboratory

Ada, Oklahoma 74820

National Risk Management Research Laboratory

Office of Research and Development

U.S. Environmental Protection Agency

Cincinnati, Ohio 45268
ii

Notice

The U.S. Environmental Protection Agency through its Office of Research 

and Development funded the research described here. This work was 

conducted under in-house Task 3674, Monitored Natural Attenuation of 

Chlorinated Solvents, and in association with and with support from the 

U.S. Air Force Center for Environmental Excellence through Interagency 

Agreement # RW-57939566, Identification of Processes that Control Natural 

Attenuation at Chlorinated Solvent Spill Sites. Mention of trade names and 

commercial products does not constitute endorsement or recommendation 

for use. 

All research projects making conclusions and recommendations based on 

environmentally related measurements and funded by the U.S. Environmental Protection Agency are required to participate in the Agency Quality 

Assurance Program. This project was conducted under a Quality Assurance 

Plan prepared for Task 3674, Monitored Natural Attenuation of Chlorinated 

Solvents. Work performed by U.S. EPA employees or by the U.S. EPA 

on-site analytical contractor followed procedures specified in these plans 

without exception. Information on the plan and documentation of the quality 

assurance activities and results is available from John T. Wilson.
iii

Foreword

The U.S. Environmental Protection Agency is charged by Congress with protecting the Nation’s land, air, 

and water resources. Under a mandate of national environmental laws, the Agency strives to formulate 

and implement actions leading to a compatible balance between human activities and the ability of natural 

systems to support and nurture life. To meet this mandate, EPA’s research program is providing data 

and technical support for solving environmental problems today and building a science knowledge base 

necessary to manage our ecological resources wisely, understand how pollutants affect our health, and 

prevent or reduce environmental risks in the future.

The National Risk Management Research Laboratory is the Agency’s center for investigation of technological and management approaches for preventing and reducing risks from pollution that threatens human 

health and the environment. The focus of the Laboratory’s research program is on methods and their 

cost-effectiveness for prevention and control of pollution to air, land, water, and subsurface resources; 

protection of water quality in public water systems; remediation of contaminated sites, sediments and 

ground water; prevention and control of indoor air pollution; and restoration of ecosystems. NRMRL 

collaborates with both public and private sector partners to foster technologies that reduce the cost of 

compliance and to anticipate emerging problems. NRMRL’s research provides solutions to environmental 

problems by: developing and promoting technologies that protect and improve the environment; advancing scientific and engineering information to support regulatory and policy decisions; and providing the 

technical support and information transfer to ensure implementation of environmental regulations and 

strategies at the national, state, and community levels.

This publication has been produced as part of the Laboratory’s strategic long-term research plan. It is 

published and made available by EPA’s Office of Research and Development to assist the user community 

and to link researchers with their clients.

Chlorinated solvents such as tetrachloroethylene and trichloroethylene are an important category of contaminants in ground water at hazardous waste sites. Frequently, these compounds are subject to natural 

anaerobic biodegradation in ground water. During anaerobic biodegradation they undergo a sequential 

biological reductive dechlorination to produce cis-dichloroethylene, then vinyl chloride, and finally ethylene 

or ethane. Although cis-dichloroethylene is less hazardous than trichloroethylene or tetrachloroethylene, 

vinyl chloride is more hazardous. In contrast, ethylene or ethane is not hazardous to humans. If the biological reductive dechlorination is complete, with ethylene or ethane as the final product, then monitored 

natural attenuation can be used a remedy for the ground water contamination.

In recent years, bacteria that can dechlorinate dichloroethylene to ethylene or ethane have been isolated 

and characterized. All the strains that can dechlorinate vinyl chloride to ethylene or ethane belong to the 

genus Dehalococcoides. A biochemical assay for DNA specific to the genus Dehalococcoides is commercially available. This report provides technical recommendations on the interpretation of the biochemical 

assay and on the contribution of bacteria in the Dehalococcoides group to monitored natural attenuation 

of chlorinated solvents in ground water.

Stephen G. Schmelling, Director

 Ground Water and Ecosystems Restoration Division

National Risk Management Research Laboratory

v

Contents

Foreword .................................................................................................................................... iii

Figures ................................................................................................................................... vii

Tables .................................................................................................................................... ix

Acknowledgments .......................................................................................................................... xi

Abstract ...................................................................................................................................xiii

Section 1. Role of Biotransformation in Evaluation of MNA of Chlorinated Solvents ...................1

Intended Use of the Report ...............................................................................................2

State of Practice and Emerging State of the Science........................................................3

Section 2. Ecology of Microorganisms that Transform Chlorinated Solvents ..............................5

Bacteria that Gain Energy from Reductive Dechlorination (Halorespiring Bacteria) ........6

The Place of Dehalorespiring Bacteria in the Diversity of Life...........................................6

Organisms that Oxidize Chlorinated Ethylenes under Anaerobic Conditions ....................7

Organisms that Co-Metabolize Chlorinated Ethylenes.......................................................7

Aerobic Growth on Chlorinated Ethylenes ........................................................................7

Section 3. Tools to Assay Microorganisms that Completely Transform Chlorinated Solvents ...11

Polymerase Chain Reaction (PCR) Assays for Genetic Analysis of the Microbial 

Communities ....................................................................................................................11

Detection of Dehalococcoides Species by the PCR Assay..............................................13

Limitations of the PCR Assay for Dehalococcoides DNA ................................................14

Current State of Practice of PCR Tools to Evaluate Biotransformation of 

Chlorinated Solvents ........................................................................................................14

Section 4. Dehalococcoides DNA and Rates of Natural Attenuation ........................................17

A Definition of “Generally Useful” Rates of Biological Reductive Dechlorination.............17

Site Selection....................................................................................................................17

Groundwater Sampling.....................................................................................................19

Chemical Analysis ...........................................................................................................19

Detection of Dehalococcoides by Polymerase Chain Reaction Analysis ........................20

Calculation of Dechlorination Rates from Monitoring Data .............................................21

Calculation of Dechlorination Rates in Conventional Plumes .........................................21

Relationship Between Dehalococcoides DNA and Dechlorination Rates at 

Conventional Plumes .......................................................................................................23

Dehalococcoides DNA and Dechlorination Rates over Time in Particular Wells ............28

Rates of Natural Attenuation and Density of PCR Products from 

Dehalococcoides DNA......................................................................................................31

Biotransformation and Dominant Terminal Electron Accepting Processes .....................34

Example of Calibration of BIOCHLOR with Distance along a Flow Path.........................35

Example of Calibration of BIOCHLOR with Time in a Single Monitoring Well.................38

Conclusions .....................................................................................................................39
vi

Section 5. Geochemical Parameters and Occurrence of Dehalococcoides ..............................41

Synopsis ..........................................................................................................................41

Calibration of Computer Models to Evaluate MNA ..........................................................42

Sampling Sites ................................................................................................................42

Ground Water Sampling, Assay for Dehalococcoides DNA, and Chemical Analyses ....44

Detection of Dehalococcoides DNA .................................................................................44

Biogeochemistry of Ground Water with Detectable Dehalococcoides DNA ....................44

Comparison of Geochemistry where Dehalococcoides DNA is Present or Absent.........50

A Predictive Model for the Presence of Dehalococcoides DNA ......................................51

Summary and Conclusions ..............................................................................................52

Section 6. Presence of Dehalococcoides DNA and the Extent of Biodegradation ....................55

Enrichment Culture Preparation ......................................................................................55

Sampling and Analysis of Enrichment Cultures ..............................................................56

Biodegradation of Chlorinated Ethylenes in the Enrichment Cultures ............................56

Association of Dechlorination in Enrichment Cultures with Dehalococcoides DNA .......57

Section 7. Recommendations to Evaluate Biotransformation of Chlorinated Solvents .............61

Recommendations for Interpreting Data on Density of DNA in Ground Water ...............61

Recommendations for Interpreting Geochemistry of Ground Water ...............................63

Recommendations for Selecting Wells for Sampling .......................................................63

Recommendations for Sampling and Shipping of Samples ............................................64

Section 8. Data Quality ..............................................................................................................67

Analysis of Chemical Concentrations ..............................................................................67

Analysis of DNA Concentrations .....................................................................................69

Section 9. References ................................................................................................................95
vii

Figures

Figure 2.1. A phylogenetic tree based on comparisons of sequences in the 16s Ribosomal RNA. ................8

Figure 3.1. Separation of DNA by Denaturing Gradient Gel Electrophoresis (DGEE). ..................................12

Figure 4.1. The frequency distribution of the maximum concentration of chlorinated solvents and their 

transformation products at Department of Defense sites in the United States (from 

McNab et al., 2000). ......................................................................................................................18

Figure 4.2. Location of sites used to survey the relationship between Dehalococcoides DNA in ground 

water and the rate of natural attenuation of chlorinated ethylenes in field scale plumes...............18

Figure 4.3. Location of monitoring wells and distribution of cis-DCE at the Western Processing Site in 

Kent, Washington, fourth quarter 1994...........................................................................................24

Figure 4.4. Location of monitoring wells and distribution of cis-DCE in the Upper Saturated Zone Aquifer 

at the Landfill 3 Site, Tinker AFB, OK, in 2000...............................................................................26

Figure 4.5. Comparison of the locations of monitoring wells at the North Beach Site at the U.S. Coast Guard 

Support Center in Elizabeth City, North Carolina, to the distribution of PCE in ground water.......26

Figure 4.6. Location of monitoring wells and distribution of cis-DCE in the Intermediate Ground –Water 

Zone at Spill Site-4, the former England AFB, Louisiana, March 2002. .......................................28

Figure 4.7. Relationship between the density of Dehalococcoides cells as determined by quantitative 

PCR and the first order rate of attenuation of cis-DCE in ground water. ......................................32

Figure 4.8. Comparison of the density of Dehalococcoides cells in ground water as determined by 

quantitative PCR (real time PCR) to the density of Dehalococcoides DNA as determined 

by the semi-quantitative technique that uses the density of the band produced by Gel 

Electrophoresis as an estimate of the concentration of amplified DNA. ........................................33

Figure 4.9. Input screen to BIOCHLOR with calibration parameters for the North Beach Site........................34

Figure 4.10. Correspondence between the measured values for PCE and TCE at the North Beach Site in 

2002 and the concentrations that were predicted by calibrating BIOCHLOR using three 

different values for the first order rate constant for biotransformation. ..........................................36

Figure 4.11. Correspondence between the measured values for cis-DCE and vinyl chloride at the North 

Beach Site in 2002 and the concentrations that were predicted by calibrating BIOCHLOR 

using three different values for the first order rate constant for biotransformation.........................37

Figure 4.12. Input screen to BIOCHLOR with calibration parameters for the well A39L010PZ at Area 2500 

at England Air Force Base, Alexandria, Louisiana. .......................................................................38

Figure 4.13. Correspondence between predicted and actual values for the chlorinated ethylenes performed 

by calibrating BIOCHLOR to field data sampled in different events at the well. ...........................39

Figure 5.1. Location of contaminated sites used to compare presence or absence of Dehalococcoides

DNA to the geochemistry of the ground water. .............................................................................42

Figure 6.1. Relationship between the production of ethylene in the enrichment cultures and the 

concentration of dissolved oxygen in the corresponding ground water used for the inoculums 

of the enrichment cultures. ............................................................................................................59

ix

Tables

Table 2.1. Diversity of Bacteria that can Reductively Dechlorinate Ethylenes ..................................................7

Table 3.1 Comparison of Advantages and Disadvantages of PCR Tools to Evaluate Biotransformation 

of Chlorinated Solvents in Ground Water. .....................................................................................15

Table 3.2 Applications of PCR Tools to Evaluate Biotransformation of Chlorinated Solvents in 

Ground Water. ................................................................................................................................15

Table 4.1. Location of Specific Sites in the Survey and the Number of Wells Sampled at Each of the 

Sites ...............................................................................................................................................19

Table 4.2. Calibration Parameters for BIOCHLOR .........................................................................................22

Table 4.3. Relationship between the Concentrations of Chlorinated Ethylenes in Ground Water and 

Their Apparent Rates of Dechlorination along an Inferred Flow Path in the Aquifer ....................23

Table 4.4. Distribution of Chlorinated Ethylenes and Dehalococcoides DNA at Sites that Form 

Conventional Plumes .....................................................................................................................25

Table 4.5. Relationship between the Apparent Rates of Dechlorination along a Flow Path in the Aquifer 

and the Detection of Bacterial DNA and Dehalococcoides DNA in Water Samples from a 

Monitoring Well in the Plume .........................................................................................................27

Table 4.6. Distribution of Chlorinated Ethylenes and Dehalococcoides DNA at Sites at England AFB, 

which do not Form Conventional Plumes ......................................................................................29

Table 4.7. Relationship between the Concentrations of Chlorinated Ethylenes in Ground Water and 

Their Apparent Rates of Dechlorination over Time in Water Samples from Monitoring Wells 

at England AFB, Louisiana ............................................................................................................29

Table 4.8. Relationship between the Apparent Rates of Dechlorination over Time and the Detection of 

Bacterial DNA and Dehalococcoides DNA in Water Samples from Monitoring Wells at 

England AFB, Louisiana ................................................................................................................30

Table 4.9. Comparison of Rates on Attenuation to the Overall Geochemical Environment of the Sites 

in the Survey ..................................................................................................................................34

Table 5.1. Location of Sites Included in the Survey Comparing the Presence or Absence of 

Dehalococcoides DNA to the Geochemistry of the Ground Water ...............................................43

Table 5.2 The Intensity Scores of Dehalococcoides DNA and the Concentrations of Chlorinated 

Ethylenes and Ethylene in the Wells where Dehalococcoides DNA was Detected ......................45

Table 5.3. The Concentrations of Nitrate plus Nitrite Nitrogen, Methane, and the ORP Meter Reading 

in the Wells where Dehalococcoides DNA was Detected .............................................................46

Table 5.4. The Concentrations of Oxygen, Ferrous Iron, and Sulfate in the Wells where Dehalococcoides 

DNA was Detected ........................................................................................................................48

Table 5.5. The Concentration of Dissolved Molecular Hydrogen and Total Organic Carbon, and the 

Oxidation/Reduction Potential in the Wells where Dehalococcoides DNA was Detected .............49

Table 5.6. The Probability (p) that the Distribution of the Measured Values for Selected Geochemical 

Parameters between Ground Water where Dehalococcoides DNA was Present and Ground 

Water where Dehalococcoides DNA was not Present is not Statistically Different .......................51

Table 5.7. Comparisons between the Observed Presence or Absence of Dehalococcoides DNA and the 

Concentrations of Nitrate plus Nitrite Nitrogen, Methane, and the ORP Meter Reading in the 

Wells ..............................................................................................................................................53
x

Table 5.8. Comparisons between the Observed Presence or Absence of Dehalococcoides DNA and the 

Predicted Probabilities for the Presence of Dehalococcoides DNA ..............................................54

Table 6.1. Composition of the Basal Medium (pH 7) .....................................................................................55

Table 6.2. Comparisons of Biotransformation of Chlorinated Ethylenes in Enrichment Cultures to the 

Corresponding Presence or Absence of Amplifiable Dehalococcoides DNA in the Ground 

Water Sample Used to Inoculate the Enrichment Cultures ...........................................................58

Table 6.3. Comparison of Biotransformation of Chlorinated Ethylenes in Enrichment Cultures to the 

Corresponding Geochemistry of the Ground Water used for Inoculation of the Enrichment 

Cultures .........................................................................................................................................60

Table 7.1. Recommendations for Use of PCR Assays to Evaluate Biotransformation of Chlorinated 

Solvents .........................................................................................................................................62

Table 8.1. Typical Quality Performance Data for Analysis of TCE in Water ...................................................70

Table 8.2. Typical Quality Performance Data for Analysis of cis-DCE in Water .............................................74

Table 8.3. Typical Quality Performance Data for Analysis of Vinyl Chloride in Water ....................................78

Table 8.4. Typical Quality Performance Data for Analysis of Ethylene in Water or in Gas ............................82

Table 8.5. Typical Quality Performance Data for Analysis of Methane in Water or in Gas ............................86

Table 8.6. Typical Quality Performance Data for Analysis of Hydrogen in Gas .............................................90

Table 8.7. Typical Quality Performance Data for Analysis of Nitrate Plus Nitrite Nitrogen in Water. ..............93
xi

Acknowledgments

Formal peer reviews were provided by Dr. Carolyn Acheson with the U.S. 

EPA Office of Research and Development, National Risk Management 

Research Laboratory; Dr. Ned Black with U.S. EPA Region 9; Dr. Mitch 

Lasat with U.S. EPA Office of Research and Development, Office of 

Science Policy; Dr. Donna Fennell at Rutgers University, New Brunswick, 

NJ; Dr. James Gossett at Cornell University, Ithaca, NY; Dr. David Ellis with 

DuPont Company, Wilmington, DE; Dr. Guy Sewell at East Central State 

University, Ada, OK; and Dr. Andrea Leeson, a SERDP/ESTCP Cleanup 

Program Manager, Arlington, VA. Courtesy technical reviews were provided 

by Mr. Philip Dennis, SIREM Operations Manager, GeoSyntec Consultants, 

Guelph, Ontario, Canada, and by Dr. Margaret Findlay and Dr. Sam Fogel 

of Bioremediation Consulting, Inc., Watertown, MA.

xiii

Abstract

At most hazardous waste sites where monitored natural attenuation (MNA) of chlorinated solvents in 

ground water is successful as a remedy, the chlorinated solvents are biologically degraded to harmless 

end products such as ethylene or ethane. Many organisms can degrade chlorinated solvents such as 

tetrachloroethylene or trichloroethylene, to dichloroethylene and vinyl chloride. This contributes little to 

risk reduction because vinyl chloride is more toxic and more carcinogenic than tetrachloroethylene or 

trichloroethylene. The only organisms known to degrade dichloroethylenes and vinyl chloride to ethylene 

or ethane are members of the Dehalococcoides group. As a result, these organisms have a critical role 

in the evaluation of MNA at chlorinated solvent sites. In recent years, biochemical assays for the presence of DNA from the organisms have become commercially available. These assays are based on the 

polymerase chain reaction (PCR) for the amplification of DNA extracted from ground water. They are 

very sensitive and can be very specific. 

This report is designed for technical staff in the EPA Regions and in state agencies that require information 

on the contribution of Dehalococcoides bacteria to MNA of chlorinated solvents, and information on the 

proper application and interpretation of the assays in an evaluation of MNA. This report includes sections 

on the role of biotransformation in evaluation of MNA of chlorinated solvents, the ecology of microorganisms 

that transform chlorinated solvents, tools to assay microorganisms that transform chlorinated solvents, 

the relationship between Dehalococcoides DNA in ground water and rates of natural attenuation at field 

scale, the relationship between geochemical parameters and the occurrence of Dehalococcoides DNA 

in ground water, and the relationship Dehalococcoides DNA in ground water and behavior of chlorinated 

solvents in laboratory treatability studies or microcosm studies done with water from the plume.

1

Section 1. 

Role of Biotransformation in Evaluation of MNA of Chlorinated Solvents

Perchloroethylene (PCE) and trichloroethylene (TCE) 

are among the most pervasive chlorinated solvents in 

ground water at hazardous waste sites. PCE and TCE 

are subject to biological and chemical processes that 

may further transform them to cis-dichloroethylene 

(cis-DCE), trans-dichloroethylene (trans-DCE), 1,1-

dichloroethylene (1,1-DCE), vinyl chloride, ethylene, 

and ethane. Vinyl chloride is the most hazardous of 

the transformation products. Ethylene and ethane 

are essentially harmless. A wide variety of microorganisms can transform PCE or TCE to produce the 

dichloroethylenes. To date, only microorganisms 

from a specific and relatively uncommon group (the 

Dehalococcoides group) have been shown to have the 

capability to transform cis-DCE to vinyl chloride, and 

then transform vinyl chloride to ethylene. Complete 

transformation of PCE or TCE to ethylene can be 

an important process contributing to the monitored 

natural attenuation of PCE and TCE at hazardous 

waste sites. 

In the past ten years, very sensitive and specific 

biochemical tools have been developed that can 

recognize DNA from Dehalococcoides organisms. To 

carry out an assay for Dehalococcoides organisms, 

DNA is isolated from ground water or sediment. Then 

short pieces of DNA that are complementary to gene 

sequences that are unique to Dehalococcoides organisms are added to the extract. These short pieces 

bind to the Dehalococcoides DNA, and allow the DNA 

to be copied by an enzyme called DNA polymerase. 

The short pieces that bind to the Dehalococcoides

DNA are often referred to as primers for the DNA 

polymerase reaction. The process is repeated for 

a number of cycles. Each time the DNA that was 

synthesized in the previous reaction becomes the 

template for the subsequent reaction. In each cycle 

of the chain reaction, the concentration of Dehalococcoides DNA is doubled. Finally, the concentration of 

Dehalococcoides DNA is high enough that it can be 

identified and analyzed by other molecular biology 

procedures. The entire process is referred to as the 

polymerase chain reaction (PCR). 

A recurrent feature in the journal, Remediation, is 

the Monitored Natural Attenuation Forum: A Panel 

Discussion. In a recent issue (Borden et al., 2003), 

a panel of three experts responded to the question: 

Recently, there has been a lot of discussion 

about the need for Dehalococcoides ethenogenes to completely break down chlorinated 

solvents such as TCE in groundwater. Should 

natural attenuation studies automatically include a test for the microbe? Does the presence of cis-DCE and vinyl chloride preclude 

the need for microbe testing? Is suitable 

groundwater geochemistry or the detection of 

this bacterium in groundwater more valuable 

in natural attenuation studies?

Their responses are a good summary of the state of 

knowledge with respect to the use of PCR tools to 

evaluate the contribution of Dehalococcoides bacteria to natural attenuation of chlorinated ethylenes in 

ground water (Borden et al., 2003).

One panel member noted that due to the complexity 

of most field sites, we rely on multiple lines of evidence to evaluate the behavior of plumes. The panel 

member observed that: 

“While geochemical data are usually the best 

indicators of the potential for complete dechlorination in an aquifer, definitive information 

on the presence or absence of Dehalococcoides can provide very useful information in 

determining whether MNA is an appropriate 

approach for a specific site. At present, we do 

not understand enough about the distribution 

and activity of the currently available assays 

to use these results as a primary indicator for 

the presence/absence of complete reductive 

dechlorination at a site.”

A second panel member noted the assay as currently practiced has a high rate of false positives and 
2

false negatives. Based on the cost of the PCR assay 

compared to the cost for conventional geochemical 

parameters, the panel member concluded, 

“But clearly the cost benefit analysis of DSA 

[Dehalococcoides specific PCR assays] makes 

it comparatively less attractive, and due to limitation in sensitivity and interpretation, it should 

not be a required analysis at this time.” 

The third panel member noted that the significance 

of PRC data depended on the specific procedure that 

was used. She also noted that: 

“Another reason that limits the usefulness of 

microbial characterization at field sites has 

to do with the difficulty associated with using 

these data for assessing the efficacy of natural 

attenuation. Understanding the microbial population, its activity, and its diversity provides 

insight into the biodegradation processes and 

pathways but does not yield data that can be 

used to estimate concentration declines, generation of by-products, of clean-up times.”

Intended Use of the Report

This report is intended to facilitate the use of DNA 

analysis to document the role of Dehalococcoides

organisms in the natural attenuation of PCE, TCE, and 

their transformation products in ground water. The 

report contains seven sections. This section describes 

several innovative genetic tools for evaluating microbial 

communities that degrade chlorinated hydrocarbons. 

It also describes the relationship between the new 

genetic tools and previous technical recommendations 

published by EPA/ORD in the Technical Protocol for 

Evaluating Natural Attenuation of Chlorinated Solvents 

in Ground Water (Wiedemeier et al., 1998). 

A short review of the nutritional ecology and physiological diversity of organisms that can degrade 

chlorinated solvents in ground water is provided in 

Section 2. Ecology of Microorganisms that Transform 

Chlorinated Solvents. The new techniques in genetic 

analysis that are available to identify and enumerate 

specific microorganisms in ground water or aquifer 

sediment are described in Section 3. Tools to Assay 

Microorganisms that Transform Chlorinated Solvents. 

These sections provide background and context for the 

detailed discussion of Dehalococcoides organisms in 

the remainder of the report. A reader who is familiar 

with these topics can skip Sections 2 and 3.

An evaluation of Monitored Natural Attenuation (MNA) 

as defined by U.S. EPA in the OSWER Directive (U.S. 

EPA, 1999) requires a quantitative understanding of 

the behavior of the plume of contamination over time 

and space. The critical parameter is the rate of attenuation of concentration of the contaminant over time 

and with distance away from the source. To contribute 

to an evaluation of MNA, an assay for the presence 

or activity of microorganisms must be associated with 

the field scale behavior of the plume containing the 

microorganisms. Section 4. Dehalococcoides DNA 

and Rates of Natural Attenuation at Field Scale compares the achieved rates of natural attenuation of PCE, 

TCE, cis-DCE, and vinyl chloride at several chlorinated 

solvent plumes to estimates of the concentration of 

Dehalococcoides DNA in ground water provided by the 

polymerase chain reaction (PCR) assay. In general, 

concentrations of Dehalococcoides DNA that were 

high enough to be detected by a commercially available assay were associated with rates of attenuation 

that are useful for MNA. 

At most field scale plumes where MNA has been proposed as a remedy for chlorinated solvents in ground 

water, biotransformation of the solvents to harmless 

end products is an important part of the remedy. 

The expected contributions of other processes (such 

as sorption or dilution and dispersion) are usually 

not adequate to be protective of human health and 

the environment. As a consequence, the Technical 

Protocol for Evaluating Natural Attenuation of Chlorinated Solvents in Ground Water (Wiedemeier et al., 

1998) put a heavy emphasis on biotransformation as 

a process to achieve natural attenuation. 

The Technical Protocol (1998) used a number of geochemical parameters in a scoring system to predict 

whether ground water contained microorganisms that 

could biologically transform chlorinated solvents. The 

scoring system was criticized by the Committee on 

Intrinsic Remediation of the National Research Council (NRC, 2000). The Committee recommended that 

the scoring system should not be used to evaluate 

prospects for MNA. 

In Table 2.3 of the Technical Protocol (Wiedemeier, 

1998), scores are assigned based in part on analyses 

for oxygen, nitrate, iron II, sulfate, methane, molecular 

hydrogen, oxidation reduction potential (ORP), pH, 

total organic carbon, chloride, BTEX compounds, 

temperature, and alkalinity. An assay for the presence 

of Dehalococcoides DNA provides direct evidence of 

an organism that can completely transform chlorinated 

ethylenes. The presence or absence of Dehalococcoides DNA can be used to evaluate the information 

provided by the geochemical parameters. 
3

In Section 5. Geochemical Parameters and Occurrence of Dehalococcoides DNA, statistics are used 

to identify the conventional geochemical parameters 

that are associated with the presence of Dehalococcoides DNA in ground water. The distribution of the 

values for ORP, and the concentrations of nitrate and 

methane were significantly different between samples 

of ground water where Dehalococcoides DNA was 

detected and samples where Dehalococcoides DNA 

was not detected. As for the other parameters, there 

were no significant differences. In Section 5, a statistical technique is used to derive a formula for the probability that Dehalococcoides is present in ground water 

knowing the concentrations of nitrate and methane, 

and the oxidation/reduction potential. The formula is 

offered as a replacement for the scoring system in the 

Technical Protocol (Wiedemeier et al., 1998).

The process of evaluating MNA as an option at chlorinated solvent sites often involves a laboratory study 

to determine whether the contaminated aquifer harbors microorganisms that can entirely transform the 

contaminants. These studies are often referred to as 

engineering treatability studies or microcosm studies. 

Laboratory enrichment studies were conducted with 

ground water from plumes where the long-term monitoring data clearly indicated that biotransformation 

processes were responsible for the observed natural 

attenuation of the plume. In Section 6. Dehalococcoides DNA and Laboratory Studies of Biodegradation, 

the results of the laboratory studies were compared 

to the distribution of Dehalococcoides DNA in the 

contaminated aquifer. 

If the presence of Dehalococcoides DNA indicates 

the presence of organisms that will dechlorinate contaminants to ethylene, then all the cultures that were 

established with ground water containing amplifiable 

Dehalococcoides DNA would be expected to dechlorinate PCE or TCE to ethylene. If false positives 

for the assay are defined as water samples where 

Dehalococcoides DNA was detected, but ethylene 

was not detected in the enrichment culture, then the 

proportion of false positive predictions as evaluated 

against the results of all the enrichment cultures that 

were constructed was 55%. 

An absence of Dehalococcoides DNA would suggest 

that these organisms were absent from the ground 

water. Because the only organisms known to dechlorinate cis-DCE and vinyl chloride belong to the Dehalococcoides group, the absence of Dehalococcoides

DNA would indicate that dechlorination would not 

proceed to vinyl chloride or ethylene. If false negatives 

for the assay are defined as water samples where 

Dehalococcoides DNA was not detected, but vinyl 

chloride or ethylene was detected in the enrichment 

culture, then the proportion of false negative predictions as evaluated against the results of all the enrichment cultures that were constructed was 15%. The 

proportion of false negative predictions as evaluated 

against the results of only those enrichment cultures 

that showed dechlorinating activity was 43%. 

Although the number of false determinations was 

high, the assay can be useful, particularly when the 

overall evaluation of natural attenuation is based on 

a variety of tests and conditions. Not all strains of 

Dehalococcoides can dechlorinate cis-DCE to vinyl 

chloride or ethylene (Duhamel et al., 2004). However, 

the unequivocal presence of Dehalococcoides DNA in 

a ground water sample strongly suggests, although it 

does not prove, that chlorinated ethylenes are being 

dechlorinated to ethylene in the aquifer. The determination is much stronger if it is supported by other 

information that would be consistent with dechlorination to ethylene. A failure to detect Dehalococcoides

DNA in a sample of ground water should not be taken 

to mean that dechlorination in the aquifer will stop at 

the level of dichloroethylene, and that cis-DCE and 

vinyl chloride will not be degraded. 

Section 7. Recommendations to Evaluate Biotransformation of Chlorinated Solvents provides recommendations on the interpretation of data on the concentration 

of DNA in ground water samples, and on the sampling 

protocol.

State of Practice and Emerging State of the 

Science

Laboratory research and collection of field data to 

prepare this report on the Evaluation of the Role of 

Dehalococcoides Organisms in the Natural Attenuation of Chlorinated Ethylenes in Ground Water began 

in December 2002. At that time, the literature suggested that organisms in the Dehalococcoides group 

were primarily responsible for complete dechlorination 

of chlorinated solvents to ethylene, both in the field 

and in laboratory cultures. This report is intended to 

facilitate the use of DNA analysis to document the role 

of Dehalococcoides organisms in the natural attenuation of PCE, TCE, and their transformation products 

in ground water at hazardous waste sites. 

Future evaluations at U.S. EPA enforcement actions 

will be carried out by contractors and consultants to 

the responsible parties using commercially available 

services for the assay for Dehalococcoides DNA. To 

make our research findings consistent with results that 

would be obtained at other field sites, we obtained the 
assay for Dehalococcoides DNA from a commercial 

vendor, instead of doing the assay in-house with EPA 

research staff. To generate data that met our data 

quality objectives for comparability, all the assays for 

Dehalococcoides DNA reported in this study were 

conducted by the same commercial vendor using the 

same protocol. In December 2002, we were aware 

of only one commercial laboratory in the world that 

could assay ground water for the presence of Dehalococcoides. Since that time, a number of laboratories 

have entered the commercial market. 

The assay provided by our vendor was based on 

the polymerase chain reaction, and the assay used 

primers for the 16S-rRNA gene. This gene codes for 

a structural component of the ribosome. As a consequence the nucleotide sequences in the gene tend 

to be conserved as the organisms evolve over time. 

This makes it possible to recognize organisms that are 

currently widely distributed, but which had a common 

ancestor. However, the 16S-rRNA gene is not directly 

related to metabolism of chlorinated ethylenes, and 

it is possible that the assay detected organisms that 

belonged to the Dehalococcoides group, but were 

not able to metabolize cis-DCE and vinyl chloride. 

Genes have been identified in the Dehalococcoides

group for enzymes that dechlorinate cis-DCE and vinyl 

chloride (Krajmalnik-Brown et al., 2004; Magnuson 

et al., 1998; Müller et al., 2004), and PCR assays 

for genes encoding for a Vinyl Chloride Reductase 

enzyme are now commercially available. There may 

be many Vinyl Chloride Reductase genes, and an 

assay for only one of the Vinyl Chloride Reductase 

genes might fail to identify the capacity to reduce vinyl 

chloride to ethylene in the mixed microbial community 

in a contaminated plume. However, if a PCR assay 

detects DNA for a Vinyl Chloride Reductase, it is very 

likely that the microbial community has the capacity 

to degrade vinyl chloride.

The U.S. Federal Government has provided substantial funding for research to development new tools 

for genetic analysis of organisms that are capable of 

degrading chlorinated organic contaminants in ground 

water. Many talented people work in this field, their 

research is bearing fruit, and new applications are 

coming into the market place. This report is based on 

the state of commercial practice in 2004. It is already 

out of date as far as new applications or potential 

applications of genetic analysis. However, use of the 

polymerase chain reaction to assay for the 16S-rRNA 

gene of Dehalococcoides is still widespread, and there 

is need of a report that documents the performance of 

the assay to evaluate natural attenuation of chlorinated 

solvents in ground water. 
5

Section 2. 

Ecology of Microorganisms that Transform Chlorinated Solvents

The ecology of organisms that biologically transform 

chlorinated ethylenes has recently been reviewed 

(Bradley, 2003). This Section is a short summary of 

the available literature. It is intended to provide the 

necessary background for subsequent sections of this 

report. Bradley (2003) is recommended to any reader 

who is interested in more information on the ecology 

of organisms that degrade chlorinated ethylenes. 

Chlorinated solvents such as PCE and TCE are 

xenobiotic compounds. They have low molecular 

weight and are lipophilic, volatile, and nonflammable. 

This makes them effective solvents for oil and grease. 

The commercial use and production of PCE and TCE 

began in the 1920s and peaked in the 1970s. Use 

declined thereafter because they were suspected to 

be carcinogens.

The evolutionary origin of microbes capable of 

transforming chlorinated solvents is uncertain. Many 

chlorinated hydrocarbons occur naturally in the environment, including TCE and PCE (Gribble, 1994) 

and vinyl chloride (Keppler et al., 2002). As a result, 

microbial enzymes may have evolved that are specialized for degrading organochlorine compounds (Lee et 

al., 1998). However, the concentrations of chlorinated 

ethylenes in plumes of contaminated ground water 

are much higher than their natural concentrations. 

Chlorinated solvents such as PCE and TCE have been 

in the environment at high concentrations for only a 

short time. The dechlorinating bacteria may have 

evolved to degrade other substrates and by chance 

have the capability to degrade chlorinated solvents. 

Alternatively, in only a few decades, enzymes that 

were originally evolved to degrade other substrates 

may have adapted to degrade chlorinated solvents. 

The mechanisms involved in microbial metabolism of 

chlorinated solvents can be broadly classified into two 

categories: oxidation reactions and reductive dechlorination. Oxidation is a process where the chlorinated 

solvents are oxidized to carbon dioxide or other benign 

compounds. Reductive dechlorination is a process 

where a chlorine atom is removed and replaced with 

a hydrogen atom. Typically, reductive dechlorination 

occurs under anaerobic conditions. An electron donor 

is required to carry out reductive dechlorination. 

PCE and TCE are highly oxidized compounds and 

therefore are most susceptible to reductive dechlorination. In general, reductive dechlorination of PCE or 

TCE occurs by sequential dechlorination from PCE to 

TCE to DCE isomers to VC to ethylene. Depending 

upon environmental conditions, this sequence may be 

interrupted and not go all the way to ethylene (Wiedemeier et al., 1998). Reductive dechlorination may be 

performed by bacteria that couple their growth with 

the dechlorination of the chloroethylene or by bacteria that do not benefit from the dechlorination. In the 

former case, the process is known as halorespiration 

or dehalorespiration, and the bacteria are referred 

to as halorespiring bacteria. In the latter case, the 

process is a co-metabolic reaction where the growth 

of the bacteria is supported by metabolism of other 

compounds. 

During reductive dechlorination, all three isomers of 

DCE can theoretically be produced. However, Bouwer 

(1994) reported that under the influence of biotransformation, cis-DCE is a more common intermediate 

than trans-DCE and 1,1-DCE. Compared with PCE 

and TCE, the dichloroethylene isomers and vinyl 

chloride are not as highly oxidized. Conventional 

wisdom holds that they are not as readily reduced in 

chemical reactions. As a consequence, the biological 

reductive dechlorination of dichloroethylene and vinyl 

chloride should be slower and less extensive than is 

the case for PCE and TCE. This perspective was 

based on a comparison of the Gibbs free energy for 

complete dechlorination of each chlorinated ethylene; 

however, the chlorinated ethylenes are dechlorinated 

in a step-wise fashion. A better way to evaluate the 

energy yield is to compare the yield of each separate 

dechlorination reaction. The Gibbs free energy for 

reductive dechlorination of PCE to TCE, of TCE to 

DCE, of DCE to vinyl chloride, and vinyl chloride to 

ethylene is -171.8, -166.1, -144.8, and -154.5 kilo 

Joules per mole of chlorinated ethylene, respectively 

(Dolfing, 2000). The energy yields for each successive dechlorination are essential equivalent. The rates 
6

of reductive dechlorination of PCE, TCE, cis-DCE 

and vinyl chloride should be equivalent. Cupples et 

al. (2004b) compared dechlorination of cis-DCE and 

vinyl chloride by Dehalococcoides strain VS. The 

maximum growth rates were equivalent, the maximum 

rate of utilization of vinyl chloride was within 75% of 

the maximum rate of utilization of cis-DCE, and half 

saturation constants were equivalent.

The energy yield for reductive dechlorination is high 

relative to other anaerobic processes. The Gibbs free 

energy for nitrate reduction, sulfate reduction and 

methanogenesis is -112, -38, and -33 kilo Joules per 

mole of substrate consumed (Dolfing, 2000). 

The DCE isomers and vinyl chloride also have the 

potential to undergo oxidation to carbon dioxide 

or acetate (Bradley and Chapelle, 1998). Aerobic 

growth on PCE and TCE as an electron donor has 

never been reported. However, Ryoo et al. (2000) 

reported that PCE can be degraded co-metabolically 

under aerobic conditions by oxygenase enzymes, 

and the co-metabolic oxidation of TCE has been well 

demonstrated (Wilson and Wilson, 1985; Nelson et 

al., 1988; Ensley, 1991; Chang and Alvarez-Cohen, 

1994). In ground water, different organisms with different metabolic pathways may share the responsibility 

for natural attenuation of chlorinated solvents. The 

following sections describe the microorganisms that 

may be involved in biotransformation of chlorinated 

ethylenes. 

Bacteria that Gain Energy from Reductive 

Dechlorination (Halorespiring Bacteria) 

The halorespiring bacteria couple reductive dechlorination to growth. They were first described by Holliger and his co-workers (1993) who obtained a highly 

purified enrichment culture that was able to grow by 

the reduction of PCE to cis-DCE using hydrogen as 

the electron donor. The active organism was named 

Dehalobacter restrictus. Soon after, many different 

genera of bacteria capable of respiring chlorinated 

ethylenes were isolated. Table 2.1 lists many of the 

organisms that have been reported in the literature.

Some of the halorespiring organisms can only grow 

on a very limited range of substrates. They use only 

hydrogen as the electron donor and couple growth 

only to the reduction of chlorinated compounds. 

Examples are Dehalobacter restrictus, Strain TEA, 

Dehalococcoides ethenogenes strain 195, and Dehalococcoides sp. strain BAV1. Other halorespiring 

organisms are less restricted. They are able to use 

a number of different electron donors and acceptors 

for growth. Examples are Dehalospirillum multivorans

(Sulfospirillum) and various Desulfitobacterium strains 

including Desulfitobacterium strain PCE1, Desulfitobacterium strain PCE-S, and Desulfitobacterium 

frappieri TCE1.

The Place of Dehalorespiring Bacteria in the 

Diversity of Life

To understand the diversity of microbial communities 

and the evolutionary relationships between various 

kinds of microbes, microbiologists have compared the 

DNA sequences of genes that encode for enzymes 

that carry out the metabolic processes of interest 

and the DNA sequences of genes that encode for 

important structural components of the microbial cell. 

The gene sequence that encodes for 16S rRNA is 

commonly used to compare evolutionary relationships 

between bacteria. All living cells on earth contain an 

organelle called a ribosome that assembles proteins 

from amino acids. Because 16S rRNA is an important 

component of ribosomes, portions of the 16S rRNA 

sequence are highly conserved due to the pivotal role 

of protein synthesis in cell metabolism. Based on the 

similarity of 16S rRNA sequences, a phylogenetic tree 

of microbial groups has been constructed (Figure 2.1). 

All life on earth is divided into three great domains; the 

organisms with a cell nucleus (eukarya), the bacteria, 

and the archaea. The archaea are single-celled and of 

similar size as the bacteria but are more closely related 

to the eukarya in their genetics and biochemistry. The 

archaea include the methanogens and other members 

that live in extreme environments. Most common 

soil and ground water bacteria that degrade organic 

contaminants are bacteria and fall into the division 

Proteobacteria, the Bacteroides/Cytophaga group, 

and the Gram positive bacteria. The cell wall of the 

Gram positive bacteria has a characteristic structure 

that can be recognized using a specific staining technique, the Gram stain. The Gram positive bacteria are 

divided into a high C+G group and a low C+G group 

based on the relative proportion of the bases guanine 

and cytosine in their DNA.

Many halorespirers (Dehalobacter, Desulfitobacterium) fall within the group of low C+G Gram positives; 

however, some halorespirers are found within very 

distantly related phylogenetic groups. This distribution indicates that the ability to dechlorinate is not 

restricted to one phylogenetic cluster and may have 

evolved separately. On the other hand, no halorespiring organisms have yet been isolated that belong to 

the archaeal domain (Middeldorp et al., 1999). 

Of the organisms described to date, only Dehalococcoides species, which are phylogenetically located 

within the green non-sulfur bacteria, are capable of 
7

dechlorinating lower chlorinated ethylenes (i.e., dichloroethylene and vinyl chloride) and coupling growth 

with the dechlorination. For instance, Dehalococcoide ethenogenes strain 195 obtains energy from all 

dechlorination steps except the final step from vinyl 

chloride to ethylene (Maymó-Gatell et al., 1999, 2001). 

Dehalococcoides sp. strain BAV1 grows on vinyl 

chloride and all the dichloroethylene isomers (He et 

al., 2003a, 2003b). All laboratory mixed cultures that 

dechlorinate PCE or TCE beyond cis-DCE have been 

found to contain organisms in the Dehalococcoides

phylogenetic group (Adamson and Parkin, 2000; Ellis 

et al., 2000; Fennell et al., 2001; Duhamel et al., 2002; 

Richardson et al., 2002; Cupples et al., 2003; Dennis 

et al., 2003). Certain stains of Dehalococcoides can 

also partially degrade polychlorobiphenyls (PCBs), 

chlorobenzenes and dioxins (Fennell et al., 2004) 

and dichloroethane and dibromoethane (Thomson 

and Vidumsky, 2003). There is a group of Chloroflexi 

bacteria that is closely related to Dehalococcoides 

(but is not the same genus) that also dechlorinates 

PCBs, chlorobenzenes, and PCE (Miller et al., 2005; 

Watts et al., 2005; Wu et al., 2002a , 2002b).

If natural attenuation is to be a remedy for ground water contamination with chlorinated ethylenes, the chloTable 2.1. Diversity of Bacteria that can Reductively Dechlorinate Ethylenes*

Isolate or Culture Closest Phylogenetic 

Affiliation

Dechlorination 

Steps Performed Reference

Dehalobacter restrictus Low G+C Gram positive 

bacteria PCE to cis-DCE Holliger et al., 1993

Dehalospirillum 

multivorans, renamed 

Sulfospirillum

Proteobacteria, ε subdivision PCE to cis-DCE Scholz-Muramatsu et 

al., 1995

Desulfitobacterium strain 

PCE1

Desulfitobacterium

a Gram positive bacterium PCE to TCE Gerritse et al., 1996

Desulfuromonas 

chloroethenica Geobacter PCE to cis-DCE Krumholz et al., 1996

strain MS-1 Enterobacteriaceae PCE to cis-DCE Sharma and McCarty 

1996

strain TEA Gram positive bacteria

Low G+C PCE to cis-DCE Wild et al., 1996

Desulfitobacterium sp. 

strain PCE-S

Desulfitobacterium

a Gram positive bacterium PCE to cis-DCE Miller et al., 1997

Dehalococcoides 

ethenogenes strain 195 Green, nonsulfur bacteria PCE to ethylene Maymó-Gatell et al., 1997

Desulfitobacterium 

frappieri TCE1

Desulfitobacterium

a Gram positive bacterium PCE to cis-DCE Gerritse et al., 1999

Clostridium bifermentans

strain DPH-1 Clostridium PCE to cis-DCE Chang et al., 2000

Dehalococcoides sp. 

strain CBDB1

Dehalococcoides 

ethenogenes

PCE to transDCE Adrian et al., 2000

Desulfitobacterium sp. 

strain Y51

Desulfitobacterium

a Gram positive bacterium PCE to cis-DCE Suyama et al., 2002

Desulfitobacterium 

metallireducens

Desulfitobacterium

a Gram positive bacterium PCE to cis-DCE Finneran et al., 2003

Desulfuromonas 

michiganenis Geobacter PCE to cis-DCE Sung et al., 2003

Dehalococcoides sp. 

strain BAV1

Dehalococcoides 

etheneogenes

cis-DCE to 

ethylene He et al., 2003a,b

*

 Extracted from Major et al. (2003).
8

rinated ethylenes must be completely dechlorinated 

to harmless products. Because the Dehalococcoides

group is the only known group of organisms that can 

grow by carrying out the reductive dechlorination of 

dichloroethylene or vinyl chloride, it has a critical role 

in any evaluation of monitored natural attenuation in 

anaerobic ground water. 

Section 3 of this report discusses the use of techniques for genetic analysis to recognize members of 

Figure 2.1. A phylogenetic tree based on comparisons of sequences in the 16s Ribosomal RNA. The length of the 

thin lines is roughly proportional to the evolutionary distance between groups of organisms. The thick 

lines represent the range within a particular group. Redrawn and simplified from Figure 1 in Hugenholtz 

et al., 1998. 

the Dehalococcoides group in ground water and aquifer sediment. The Dehalococcoides group is unusual 

in that its sequences of ribosomal RNA group with 

sequences shared by the green non-sulfur bacteria 

(i.e., the Chloroflexi ). The green non-sulfur bacteria 

are green because they contain a form of chlorophyll. 

They are phototrophic, meaning that they gain energy 

from light and grow in the presence of light, but they 

do not produce oxygen during photosynthesis. 
9

Organisms that Oxidize Chlorinated 

Ethylenes under Anaerobic Conditions

Oxidation usually occurs in the presence of molecular oxygen; however, oxidation of cis-DCE and vinyl 

chloride may also occur under some anaerobic conditions (Bradley and Chapelle, 1998, 2000a, 2000b; 

reviewed in Bradley, 2003). Bradley and Chapelle 

(2000b) showed that vinyl chloride can be oxidized to 

acetate by an interesting class of anaerobic bacteria 

called acetogens. They oxidize the organic compound 

solely as part of their energy metabolism. They do 

not use the chlorinated ethylene as a substrate to 

build microbial cell constituents. Although their role 

in natural attenuation of chlorinated ethylenes in anaerobic aquifers may be as significant as that of the 

Dehalococcoides group, very little is known about 

them, and there are no techniques that are currently 

commercially available to estimate their contribution. 

Organisms that Co-Metabolize Chlorinated 

Ethylenes

Under some circumstances, dechlorination of chlorinated compounds is not coupled to growth. The 

reaction is catalyzed by an enzyme or cofactor that 

is fortuitously produced by the microbes for other 

purposes. When this occurs, the process is called 

co-metabolism. Chlorinated ethylenes can be cometabolized under aerobic conditions and under 

anaerobic conditions.

PCE and TCE can be co-metabolically dechlorinated 

by many types of anaerobic organisms, including 

certain species of methanogens such as Methanosarcina mazei (Fathepure and Boyd, 1988) and certain 

acetogens such as Acetobacterium woodii (Egli et 

al., 1988) and Sporomusa ovata (Terzenbach and 

Blaut, 1994); and sulfate-reducing bacteria (Bagley 

and Gossett, 1989). Only a small fraction of the total 

reducing equivalents derived from the oxidation of 

electron donors is used to reduce the chlorinated 

compounds.

Some bacteria contain certain enzymes that can fortuitously catalyze the oxidation of partially chlorinated 

compounds. These monoxygenase or dioxygenase 

enzymes require molecular oxygen as a substrate. 

Oxygen and a primary substrate are required for 

growth of the organisms. When these bacteria grow on 

primary substrates like methane, propane, propene, 

methanol, toluene, or phenol, they can co-metabolically oxidize partially chlorinated ethylenes such as 

TCE, cis-DCE, or vinyl chloride (Wilson and Wilson 

1985; McCarty and Semprini, 1994; Reij et al., 1995; 

Fitch et al., 1996). The only plausible co-substrates 

in a contaminated aquifer are methane or aromatic 

hydrocarbons such as toluene. If oxygen is available, 

bacteria can grow rapidly on the primary substrates, 

and exhaust supplies of either the primary substrate or 

oxygen. Ground water in a contaminated aquifer can 

be expected to have the primary substrate or oxygen, 

but not both. Aerobic co-oxidation might occur at the 

fringe of an anaerobic plume where contaminated 

ground water containing methane mixes with ground 

water containing oxygen. However, the process is 

unlikely to make a substantial contribution to natural 

attenuation of chlorinated ethylenes. 

Aerobic Growth on Chlorinated Ethylenes

There is no report on microbial growth supported by 

the oxidation of PCE or TCE. However, DCE and vinyl 

chloride can be directly oxidized by some bacteria. For 

instance, aerobic bacteria such as Actinomycetales sp. 

(Phelps et al., 1991), Mycobacterium sp. (Hartmans 

and de Bont, 1992), Rhodococcus sp. (Malachowsky 

et al., 1994), Pseudomonas sp. (Verce et al., 2000), 

and Nocardioides sp. (Coleman et al.; 2002a, 2002b) 

can grow on vinyl chloride as the sole carbon source. 

Similarly, cis-DCE has been shown to be utilized as the 

sole carbon source by some microorganisms (Bradley 

and Chapelle, 2000a, 2000b; Coleman et al., 2002a, 

2002b; Olaniran et al., 2004). 

Coleman et al. (2002b) determined the distribution of 

aerobic bacteria that could metabolize vinyl chloride 

in ground water from monitoring wells at chlorinated 

solvent spill sites. They were able to isolate vinyl chloride oxidizing strains from 15 of 24 samples of ground 

water or aquifer sediment. Aerobic organisms that 

degrade vinyl chloride are widely distributed. Reducing conditions are required to produce vinyl chloride 

or cis-DCE from TCE or PCE in ground water, yet 

oxygen is required for the further aerobic metabolism 

of cis-DCE or vinyl chloride. Degradation of cis-DCE 

and vinyl chloride by aerobic microorganisms can only 

be important at the fringe of a plume where DCE and 

vinyl chloride, produced in the plume by anaerobic 

microbial processes, are mixed with uncontaminated 

ground water containing oxygen. Aerobic biodegradation is probably not important in ground water plumes, 

unless the plume is very long. However, conditions 

conducive to aerobic biodegradation occur in the 

bed sediments of streams and lakes where plumes 

of contaminated ground water discharge to aerobic 

surface water. 

11

Section 3. 

Tools to Assay Microorganisms that 

Completely Transform Chlorinated Solvents

The only organisms that are known to completely 

transform chlorinated ethylenes to harmless products 

are members of the Dehalococcoides group of bacteria. This section discusses an analytical procedure 

called the polymerase chain reaction (PCR) that is 

used to amplify DNA from the Dehalococcoides group 

of bacteria when they are present in samples of ground 

water or aquifer sediment, and procedures used to 

detect and measure the amplified DNA produced by 

PCR. This section discusses the application of PCR 

technology to evaluate the presence and distribution 

of Dehalococcoides bacteria in ground water or aquifer 

sediments, and discusses limitation of the PCR assays 

for detecting Dehalococcoides bacteria.

The identification and enumeration of the members of a 

microbial community have traditionally been achieved 

by cultivation techniques such as plate counting. However, it has been estimated that the portion of microbes 

obtained by traditional cultivation techniques amounts 

to only 0.1 to 1 % of the total diversity (Amann et al., 

1995). As a result, non-plating techniques, including 

molecular genetic methods, have been developed 

that can detect and identify microbes in their natural 

environment based on variations in the sequences of 

base pairs in their DNA and RNA (e.g. Akkermans et 

al., 1995). Most of these techniques are based on 

amplification of DNA that encodes the gene for 16S 

rRNA, using the polymerase chain reaction (PCR). 

The amplified DNA is detected or characterized by a 

variety of techniques including denaturing gradient gel 

electrophoresis (DGGE) (Muyzer et al., 1993) and real 

time PCR (Lee et al., 1993; Livak et al., 1995).

Polymerase Chain Reaction (PCR) Assays 

for Genetic Analysis of the Microbial 

Communities

The development of PCR is a major step forward in the 

study of microorganisms in the environment (Erlich, 

1989). To prepare for the assay, a target organism 

is identified or selected. It is necessary to know the 

DNA sequences from the target organism. Based on 

that knowledge, short sequences of DNA are synthesized (“primers”) that are complementary to the 

gene sequences in the target organism. To carry out 

a PCR assay using the primers, DNA is isolated from 

ground water or sediment samples. Then, the primers 

are combined in a reaction mixture and bind to the 

target DNA, if it is present, and allow the DNA to be 

copied by an enzyme called DNA polymerase. The 

process is repeated for a number of cycles, usually 

30 to 40. Each time the DNA that was synthesized 

in the previous reaction becomes the substrate for 

the subsequent reaction. In each cycle of the chain 

reaction, the concentration of target DNA is doubled. 

Finally, the concentration of target DNA (the amplicon) 

is high enough that it can be identified and analyzed 

by optical or chemical procedures. The entire process 

is referred to as a polymerase chain reaction (PCR). 

Electrophoresis is a technique to separate large 

molecules such as DNA based in part on their ionic 

charge. If a direct current is imposed across a gel 

containing the large molecules in a solution, the 

molecules will migrate to one electrode or the other 

depending on their charge. 

The migration of the large molecules through the 

electric field is controlled by the strength of the field, 

the ionic charge of the large molecules, and the size 

and shape of the molecules. Because DNA has a 

net negative charge, DNA will migrate through the 

gel toward the positive electrode.

In cells, DNA molecules occur as a duplex or double 

helix of two separate strands of DNA. The two strands 

of the double helix are held together by hydrogen 

bonds between the two strands. The DNA duplex 

or double helix is folded into a three-dimensional 

structure that is held together by hydrogen bonding 

and interactions between hydrophobic regions in the 

DNA molecule. The two strands of DNA in the double 

helix can be separated from each other by chemicals 

and heat. 

DNA molecules are commonly separated by a variation of electrophoresis called denaturing gradient gel 

electrophoresis (DGGE). The gel contains a gradient 

of chemicals that cause the DNA double helix to unfold 

into single strands. As a DNA molecule moves through 
12

the gradient in the gel, the two strands of DNA start to 

separate or unfold. The unfolded DNA assumes a different shape, which produces more drag and reduces 

the mobility of the DNA molecule through the gel. 

Eventually, the partially unfolded DNA molecule will 

become so entangled in the gel matrix that it will stop 

moving. Small differences in the genetic sequence 

have a strong effect on the position in the gradient 

where the DNA molecule unfolds. As a result, small 

differences can cause the DNA molecule to take up 

different positions along the gradient in the gel. The 

fragments of DNA separate from each other and take 

up unique positions along the electrical gradient in 

the gel. When the DNA in the electrophoresis gel is 

stained, it shows up as bands (See Figure 3.1). 

Figure 3.1. Separation of DNA by Denaturing Gradient Gel Electrophoresis (DGGE). Wells are cut into a rectangular piece of gel. A different extract containing DNA is placed in each well. The DNA dissolves from 

the extract into the gel. An electrical current is imposed across the gel, causing the DNA molecules to 

migrate. Based on their ionic charge, size, and three-dimensional shapes, different pieces of DNA move 

different distances, causing the pieces of DNA to separate.

To identify the organism that supplied the DNA that 

was extracted from an environmental sample or an 

enrichment culture, the position of the band of unknown DNA is compared to the positions of the bands 

of DNA from known organisms. Because the DNA 

from a particular organism has been amplified by PCR 

before analysis by DGGE, the combination of the two 

techniques can detect very sparse populations of 

individual organisms in mixed microbial communities. 

The DNA from bands can be excised, reamplified by 

PCR, and sequenced, and the source of the DNA 

can be identified by comparing the sequences of the 

DNA isolated from the environmental sample to the 

sequence in known organisms. 
13

Detection of Dehalococcoides Species by 

the PCR Assay

Although many microbial groups are involved in 

reductive dechlorination of chlorinated ethylenes, 

Dehalococcoides is the only group known to completely 

dechlorinate PCE/TCE to ethylene (Maymó-Gatell et 

al., 1997; Major et al., 2003). A recent paper reports 

that 17 putative reductive dehalogenases and five 

hydrogenase complexes are encoded in the genes 

of Dehalococcoides ethenogenes, indicating this 

organism is highly evolved to utilize halogenated 

organic compounds and hydrogen (Seshadri et al., 

2005). Hence, detection of Dehalococcoides species 

may be a useful tool for assessing the efficiency of 

natural attenuation or engineered bioremediation at 

chlorinated solvent contaminated sites. 

Currently (as of 2005), a polymerase chain reaction 

(PCR) assay based on primers for genes encoding 

for 16S-rRNA is the major tool for the detection of 

Dehalococcoides species (Löffler et al., 2000; Fennell et al., 2001; Hendrickson et al., 2002; He et al., 

2003a, 2003b). 

Löffler et al. (2000) were the first to publish an assay 

for the density of Dehalococcoides bacteria based on 

PCR. In their study, they used a two-step nested primer 

PCR approach using universal bacterial primers 

followed by a second PCR with the Dehalococcoidestargeted primers. The Dehalococcoides-targeted 

primers were designed based on the 16S rRNA 

sequence of Dehalococcoides sp. strain FL2, which 

was closely related to Dehalococcoides ethenogenes 

(96.9% sequence similarity). Since Dehalococcoides

sp. strain FL2 was not available in pure culture, the 

sensitivity of the Dehalococcoides-targeted primers 

was evaluated using serially diluted plasmid DNA 

containing the 16S rRNA gene of strain FL2. One 

to 10 copies of the 16S rRNA gene of strain FL2 

were found to be sufficient to yield the expected 

PCR product. This nested PCR approach was used 

to detect Dehalococcoides populations in river and 

aquifer sediments, and the results were confirmed by 

microcosm studies.

Fennell and her co-workers designed a Dehalococcoides primer set based on the 16S rRNA gene of 

Dehalococcoides ethenogenes strain 195 and used 

it for the detection of Dehalococcoides at a TCEcontaminated site (Fennell et al., 2001). In their 

study, separate PCR reactions were performed with 

universal primers and with Dehalococcoides primers. 

If no product was obtained directly with the specific 

primers, a nested approach was performed using the 

products from the universal PCR as the template for 

PCR with the specific primers. The detection limits 

were found to be approximately 103 cells per 0.5 g soil 

for the direct PCR and 5 to 10 cells per 0.5 g soil for 

the nested approach. The results of PCR analysis were 

supported by field data and microcosm studies. 

Hendrickson and his co-workers developed a PCR 

assay to conduct an extensive survey for the presence of Dehalococcoides at multiple chloroethylenecontaminated sites (Hendrickson et al., 2002). In 

their study, seven Dehalococcoides primer sets were 

designed based on the 16S rRNA gene sequence 

of Dehalococcoides ethenogenes strain 195, which 

was originally isolated at Cornell University, and Dehalococcoides group sequences found in enrichment 

cultures originally isolated from sites at Victoria, Texas, 

and Pinellas, Florida. The sensitivities of the primer 

sets ranged from 10 to 1,000 copies of the gene per 

reaction mixture. The developed PCR assay was applied to ground water samples and soil samples collected from 24 sites. Positive results were obtained at 

21 sites where full dechlorination of chloroethylenes 

to ethylene occurred. Phylogenetic analysis of the amplicons confirmed that Dehalococcoides sequences 

formed a unique 16S rRNA group, which could be 

divided into three subgroups (Pinellas, Victoria, and 

Cornell) based on specific base substitution patterns 

in variable regions 2 and 6 of the Dehalococcoides

16S rRNA gene sequence. 

In general, the direct PCR technique using gel electrophoresis to detect the amplified gene product is a 

qualitative or semi-quantitative method. More recently, 

real time PCR (RT-PCR) has been developed to precisely quantify the density of the Dehalococcoides

population (Lendvay et al., 2003; He et al., 2003a, 

2003b). RT-PCR does not involve the use of gel 

electrophoresis to detect the amplified DNA, but it 

still makes use of the same principles of amplification as standard PCR. In RT-PCR amplification, a 

fluorescently labelled probe targeting 16S rRNA gene 

sequences of Dehalococcoides was included in the 

reaction. The fluorescence of the dye attached to the 

amplified DNA is measured during the “extension” 

phase of the polymerase chain reaction, allowing “realtime” monitoring of the accumulation of the replicated 

DNA by the instrument during each cycle of the polymerase chain reaction. The number of cycles that are 

required to accumulate a certain amount or so-called 

“threshold” concentration of copied DNA is used to 

quantitatively determine the starting concentration of 

DNA. This technique has been applied to measure 

the abundance of a Dehalococcoides population in 

a chloroethylene-contaminated aquifer undergoing 

active remediation (Lendvay et al., 2003). 
14

Limitations of the PCR Assay for 

Dehalococcoides DNA

Although Dehalococcoides organisms are closely related phylogenetically, their capacity to transform chlorinated ethylenes can be quite different (Duhamel et 

al., 2004; He et al., 2003a, 2003b ). Dehalococcoides 

strain CBDB1 can dechlorinate chlorobenzenes, but 

cannot grow on PCE or TCE (Adrian et al., 2000). 

Dehalococcoides ethenogenes strain CBDB1 and 

Dehalococcoides sp. strain 195 have 98% identity over 

1,422 nucleotides of the 16S rRNA gene sequence. 

However, strain CBDB1 can only convert PCE to transDCE, while strain 195 converts PCE to vinyl chloride 

and ethylene (Fennell et al. 2004). Dehalococcoides

ethenogenes 195 and Dehalococcoides strain FL2 

both grow on TCE or dichloroethylenes, but cannot 

grow on vinyl chloride, while Dehalococcoides sp. 

strain BAV1 can grow on vinyl chloride (He et al., 

2003b). Although these strains are closely related 

and are capable of dechlorinating some of the same 

substrates, they did not share the capacity to grow 

using vinyl chloride, which is critical to complete 

transformation of chlorinated ethylenes to harmless 

products. 

The PCR primers used in commercially available 

assays are designed to detect as many Dehalococcoides strains as possible. They are not designed 

to distinguish between the different strains. It would 

be nearly impossible to design primers for 16S rRNA 

genes that could distinguish strains that dechlorinate 

dichloroethylenes and vinyl chloride from strains that 

do not. Even if the PCR products were cloned and 

sequenced from strains that did or did not dechlorinate dichloroethylenes and vinyl chloride, it is unlikely 

that the relatively short amplicons that are typically 

obtained would contain enough information to distinguish between strains (personal communication, 

Donna Fennell, Rutgers University). 

One alternative to searching for a structural gene 

associated with the Dehalococcoides group is to 

create primers for the genes for the dehalogenase 

enzymes that actually carry out the transformation 

of TCE or vinyl chloride to ethylene. Some strains of 

Dehalococcoides are known to express an enzyme 

that can dechlorinate TCE to ethylene (Magnuson et 

al., 1998, Fennell et al., 2004) or an enzyme that can 

dechlorinate vinyl chloride to ethylene (Müller et al., 

2004, Krajmalnik-Brown et al., 2004). Genes for these 

enzymes have been sequenced, and a quantitative 

PCR assay for these gene sequences is commercially 

available. This is an area of active research, and new 

dehalogenase primers are being developed. Until a 

comprehensive catalogue of primers for dehalogenase 

genes can be developed, there is a strong chance 

of a false negative in interpreting the absence of the 

dehalogenase gene. Although a particular gene may 

be absent, other dehalogenase genes that were not 

recognized by the primer may be expressed in the microbial population. However, the interpretation of the 

detection of PCR product from dehalogenase genes 

is straightforward. If the product is amplified by the 

primer, the gene is present in the population.

Current State of Practice of PCR Tools 

to Evaluate Biotransformation of 

Chlorinated Solvents

In 2005, the Strategic Environmental Research and 

Development Program (SERDP) and the Environmental Security Technology Certification Program 

(ESTCP) of the U.S. Department of Defense organized 

a workshop to evaluate the application of molecular 

biological tools to environmental remediation. The 

report of the workshop is a useful summary of the 

state of practice and needs for technology development (Alleman et al., 2005). Table 3.1 presents the 

consensus opinion of the experts that participated 

in the workshop concerning the advantages and 

disadvantages of PCR tools that might be used to 

evaluate biotransformation of chlorinated solvents in 

ground water, and Table 3.2 presents the consensus 

opinion on the applications of the PCR tools and the 

frequency at which they are used.
15

Table 3.1 Comparison of Advantages and Disadvantages of PCR Tools to Evaluate Biotransformation of Chlorinated Solvents in Ground Water. Summarized from Table 2 of Alleman et al. (2005)

Tool Perceived Advantage Perceived Disadvantage

Direct PCR Easy to perform False negatives

Nested PCR Unsurpassed sensitivity Requires two PCR steps

Quantitative PCR 

for 16s rRNA 

gene

Provides information on presence/ absence/

abundance of organisms of interest; nearly 

reaches the sensitivity of nested PCR; commercially available for few key organisms 

(e.g. Dehalococcoides spp.); estimates of 

total bacterial numbers are possible

Does not provide confirmation of 

activity; sampling, handling, and 

analysis are not standardized

Quantitative PCR 

for functional 

genes

Provides information on presence/ absence/

abundance of functional gene of interest; 

commercially available for few key genes 

(e.g. reductase dehalogenase genes)

For DNA, does not provide conformation of activity; sampling, handling, and analysis are not standardized

Quantitative PCR 

for messenger 

RNA

Provides information on gene expression 

(i.e. activity); quantitative approaches under 

development

Relative instability of RNA presents 

sampling and preservation challenges; not commercially available 

to a significant extent; sampling, 

handling, and analysis are not standardized

Table 3.2 Applications of PCR Tools to Evaluate Biotransformation of Chlorinated Solvents in Ground Water. Summarized from Table 2 of Alleman et al. (2005)

Tool Current Applications

Current 

Relative 

Frequency 

of Use

Comments

Direct PCR

Screening tool for presence/absence [of DNA for putative active 

organisms]

Moderate Replaced by quantitative PCR

Nested PCR

Screening tool for presence/absence [of DNA for putative active 

organisms]

Moderate Replaced by quantitative PCR

Quantitative 

PCR for 16s 

rRNA gene

Screening tool for presence/absence of desired or indicator organisms; monitoring of growth and 

distribution of individual organisms

High

[Would benefit from] expansion to 

wider range of organisms; standardized procedures; availability 

of standards

Quantitative 

PCR for functional genes

Screening tool for presence/absence of target functional genes; 

monitoring of distribution and proliferation of individual genes

Low

Needs wider range of functional 

genes; extension to mRNA; standardized procedures; availability 

of standards

Quantitative 

PCR for messenger RNA

A few experimental applications 

for confirming expression of functional genes.Low

Needs wider range of genes of 

interest; standardization of approach; clarification of how mRNA 

abundance relates to activity

17

Section 4. 

Dehalococcoides DNA and Rate of Natural Attenuation 

Many evaluations of chlorinated solvent contamination 

in ground water use mathematical models to project 

the future behavior of the plume. The models are very 

sensitive to the rate constants for degradation (Newell 

et al., 2002). Biological reductive dechlorination can 

be an important mechanism for the removal of chlorinated solvents from many anoxic aquifers; however, 

there appears to be a significant variation in the rates 

and extent of dechlorination from one plume to the 

next (Suarez and Rifai, 1999). 

Hendrickson et al. (2002) reported that there was 

a strong association between the presence of 

Dehalococcoides DNA and complete dechlorination to 

ethylene. Lendvay et al. (2003) further demonstrated 

a quantitative relationship between Dehalococcoides

DNA and ethylene production. To determine if there 

is a valid association between Dehalococcoides

DNA in ground water and the observed rates of 

dechlorination at field scale, a survey was conducted 

at selected field sites. Rate constants for attenuation 

of chlorinated solvents at field scale were extracted 

from the monitoring data, and then the rate constants 

were compared to the presence or density of 

Dehalococcoides DNA in water from monitoring wells 

at the sites.

As is discussed under the subsection titled, Rates of 

Natural Attenuation and Density of PCR Products 

from Dehalococcoides DNA, the monitoring wells did 

not efficiently sample the Dehalococcoides organisms 

in the aquifer, and the number of Dehalococcoides 

cells recovered in a liter of well water was a small 

fraction of the number of cells that were exposed to a 

liter of ground water in the aquifer. Most of the Dehalococcoides cells were probably attached to sediment 

particles. As a result, there was not a quantitative 

relationship between the rates of natural attenuation 

at field scale and the density of Dehalococcoides cells 

in ground water from monitoring wells.

A Definition of “Generally Useful” Rates of 

Biological Reductive Dechlorination

There is no legally mandated time frame for monitored 

natural attenuation of contaminants in ground water 

(U.S. Environmental Protection Agency, 1999). For 

purposes of discussion, a time frame for remediation of 

30 years will be assumed. The Maximum Contaminant 

Levels (MCLs) for PCE and TCE in drinking water are 

5 μg/L. McNab et al. (2000) evaluated the distribution 

of contaminants in more than 200 chlorinated solvent 

plumes. Figure 4.1 shows the cumulative frequency 

distribution of the maximum concentrations of each 

separate chlorinated hydrocarbon in each of the 

plumes and the first order rates of degradation that 

are necessary to reduce the maximum concentrations 

to meet a MCL of 5 μg/L in 30 years. 

Fifty percent of the plumes had maximum concentrations 

of chlorinated hydrocarbons greater than 8,000 μg/L. 

The rate of natural biodegradation necessary to reduce concentrations from 8,000 μg/L to 5 μg/L in 30 

years would be 0.23 per year (calculated by a firstorder kinetic model). Similarly, the rates of degradation 

necessary to reduce 10% and 90% of the plumes in 

the survey of McNab et al. (2000) from their maximum 

concentrations to their MCLs within 30 years would be 

0.043 per year and 0.32 per year, respectively. For 

purposes of evaluating the data at our study sites, a 

rate of 0.3 per year can be considered a “generally 

useful” rate constant for monitored natural attenuation 

of chlorinated ethylenes in ground water. This “generally useful” rate should not be applied at other sites 

without due consideration of site specific conditions. 

If the initial concentrations of chlorinated ethylenes are 

low, or if the time allowed to reach MCLs is long, a 

slower rate might be acceptable at a particular site. 

Site Selection

Eight sites at six locations were selected for the 

survey because they had good records of long-term 

monitoring. This data made it possible to extract rate 

constants for attenuation of the chlorinated solvents. 

These sites are the Western Processing Site at Kent, 

Washington; Landfill Number 3 (LF3) and Fire Training Area Number 2 (FTA2) at Tinker Air Force Base, 

Oklahoma; the North Beach Site at the U.S. Coast 

Guard Support Center in Elizabeth City, North Carolina; Spill Site Number 17 (SS-17) at Altus Air Force 

Base, Oklahoma; the Target Area 1 Site at Dover Air 

Force Base, Delaware; and Area 800 and Area 2500 

at the former England Air Force Base, Louisiana. See 

Figure 4.2 for a map showing the locations of the sites 

within the United States.
18

Figure 4.1. The frequency distribution of the maximum concentration of chlorinated solvents and their transformation 

products at Department of Defense Sites in the United States (from McNab et al., 2000). The arrows 

identify the first order rates of biotransformation that are required to reach a Maximum Contaminant 

Level of 5 μg/L within thirty years at 10% of sites, at 50% of sites, and at 90% of sites.

Figure 4.2. Location of sites used to survey the relationship between Dehalococcoides DNA in ground water and 

the rate of natural attenuation of chlorinated ethylenes in field scale plumes.
19

The plume at the Western Processing Site was extracted with a pump and treat system from September 

1988 through April 2000. The plume at site LF3 has 

been extracted by a pump and treat system since early 

1999. The plume at site SS-17 has been extracted 

with a two-phase vacuum extraction system since 

September 1996. At the North Beach Landfill Site, 

the source was removed in 1999. There are no engineered remediation actions at the other sites. To avoid 

errors in fitting rate constants for natural attenuation, 

the rate constants were fit to data that were collected 

prior to initiation of any engineered remedies.

Ground Water Sampling

 At least two monitoring wells were sampled in the 

plume at each site. One well was located at the “hot 

spot” and one down gradient. See Table 4.1 for the 

number of wells sampled at each site. In most cases, 

the water samples were collected with a peristaltic 

pump at the well head using a polyethylene plastic 

tube inserted into the well. Occasionally, the water 

samples were collected using the dedicated submersible pump in the wells. Each well was purged for approximately one-half hour; at least two casing volumes 

were purged before samples were collected. 

The dissolved hydrogen in ground water was sampled 

using a Microseeps Cell following the bubble stripping method (McInnes and Kampbell, 2000). After 

sampling for dissolved hydrogen, the effluent from 

the pump was directed to pass through an over-flow 

cell for measuring pH, temperature, conductivity, and 

oxidation reduction potential (ORP) against a Ag/AgCl 

reference electrode. Then, the effluent from the pump 

was collected to determine the concentrations of 

dissolved oxygen and ferrous iron using colorimetric 

field test kits. Finally, the effluent from the pump was 

directed to fully fill various sample containers for analysis of Dehalococcoides DNA, chlorinated solvents, 

dissolved gases (methane and ethylene), inorganic 

ions (nitrate plus nitrite, sulfate and chloride), and total 

organic carbon (TOC) in the laboratory. 

The samples for chlorinated solvents and the samples 

for dissolved gases were preserved with 1% trisodium 

phosphate. The samples for nitrate plus nitrite were 

preserved with acid (five drops of 50% sulfuric acid to 

50 ml of water sample). The samples for Dehalococcoides DNA were packed in ice and stored in coolers 

prior to shipment for analysis. 

Chemical Analysis 

Ground water samples for the analysis of chlorinated 

solvents were prepared in an automatic static headspace sampler (U.S. EPA, 1988). The samples were 

analyzed by gas chromatography with a mass spectrometer detector. The reporting limits were 1.0 μg/L 

for all the analytes. The concentrations of ethylene and 

methane in ground water samples were determined 

using a headspace equilibration technique (Kampbell 

and Vandegrift, 1998). The gaseous components in 

the headspace were separated by gas chromatography and then measured with a thermal conductivity 

detector. The reporting limits were 1 μg/L in the original aqueous phase for both gases. The concentration 

of dissolved H2 was measured on a RGA3 Reduction 

Gas Analyzer equipped with a 60/80 molecular sieve 

5A column and a reduction gas detector. The reporting limit was 1.0 nM in the water originally sampled. 

Nitrate plus nitrite were analyzed using Lachat Flow 

Injection Analyses. The reporting limit was 0.1 mg/L 

as nitrogen. Sulfate and chloride were analyzed 

using Waters Capillary Electrophoresis. The reporting limits were 0.5 mg/L for sulfate and 1.0 mg/L for 

chloride. Total Organic Carbon was determined by 

a Dohrman DC-80 Carbon Analyzer with a reporting 

Table 4.1. Location of Specific Sites in the Survey and the Number of Wells Sampled at Each of the Sites 

Location Site No. Wells Sampled

Altus Air Force Base, Oklahoma Spill Site 17 (SS-17) 6

Dover Air Force Base, Delaware Target Area 1 8

U.S. Coast Guard Support Center, 

Elizabeth City, North Carolina The North Beach Landfill Site 6

England Air Force Base, Louisiana 

Area 800 4

Area 2500 6

Tinker Air Force Base, Oklahoma Landfill Number 3 (LF-3) 5

Fire Training Area Number 2 (FTA-2) 2

Kent, Washington The Western Processing NPL Site 6
20

limit of 0.5 mg/L. The dissolved oxygen, temperature, 

pH, ORP, and conductivity were measured by corresponding electrodes and meters. The concentration 

of ferrous iron, hydrogen sulfide, and alkalinity were 

determined in the field using a Chemetrics Kit Model 

K-6010D for ferrous iron, a Hach Kit Model HS-C for 

sulfide, and a Hach Kit Model ALAPMG-L for alkalinity. The reporting limits for ferrous iron, sulfide, and 

alkalinity were 0.1, 0.5, and 20 mg/L, respectively.

Detection of Dehalococcoides by Polymerase Chain Reaction Analysis

Dehalococcoides DNA in ground water samples was 

analyzed by SiREM (Guelph, Ontario) using their 

Gene-Trac Test. The test employs the polymerase 

chain reaction using primer sets specific to DNA sequences in the 16S rRNA gene of the Dehalococcoides group (see Section 3 for a discussion). Each test 

used four primer sets, three of which were designed 

to target various sequences that were specific to the 

Dehalococcoides 16S rRNA gene. A fourth primer 

set amplified sequences that were shared by most 

members of the True Bacteria (see Figure 2.1) and 

was used as a control to confirm that DNA extracted 

from the sampled ground water could be amplified 

successfully by the polymerase chain reaction. 

Bacteria were filtered from 1.0 liter of the ground 

water sample using a 0.45 μm sterile nylon filter, and 

DNA was extracted from the bacteria. The DNA was 

extracted and prepared in 55 μL of water that was free 

of DNAse and pyrogens. Then, 1 μL of the DNA extract was used in the PCR reaction mixture. The PCR 

product was separated by gel electrophoresis and then 

stained to visualize the PCR products produced from 

the amplification specific for Dehalococcoides 16S 

rRNA gene and from the amplification for the general 

(universal) bacterial 16S rRNA gene. 

The presence of Dehalococcoides DNA in the samples 

was assessed as either “Detected” or “Not Detected” 

based on interpretation of an electronic image of the 

stained band of DNA in the electrophoresis gel (see 

Figure 3.1). Detects (gel bands) were quantified 

using densitometry software and assigned a “band 

intensity percentage” using the relative intensity of 

the strongest bands obtained to the intensity of the 

positive control containing 105 gene copies. Workers 

with experience with PCR are generally reluctant to 

make a quantitative association between the quantity 

of PCR product and the number of gene copies in the 

original sample. As a consequence, SiREM assigned 

a “test intensity score” as follows: if the value was 0% 

of positive control, a score of (-) was assigned; if the 

value was smaller than 3% of positive control, a score 

of (+/-) was assigned; if the value was in the range 

of 4% to 33% of positive control, a score of (+) was 

assigned; if the value was in the range of 34% to 66% 

of positive control, a score of (++) was assigned; if 

the value was in the range of 67% to 100% of positive 

control, a score of (+++) was assigned; and if the value 

was larger than 100% of positive control, a score of 

(++++) was assigned.

SiREM has compared the semi-quantitative “intensity 

score” with the results of quantitative PCR analyses 

done on the same samples. When the score was (+), 

the approximate range of gene copies was 103 to 105

per liter; when the score was (++), the approximate 

range of gene copies was 104 to 106 per liter; when 

the score was (+++), the approximate range of gene 

copies was 105 to 106 per liter; and when the score 

was (++++), the approximate range of gene copies 

was 106 to 108 per liter. The effective detection limit 

of the PCR reaction for Dehalococcoides DNA using 

gel electrophoresis was 500 to 5,000 gene copies 

of Dehalococcoides DNA per liter of ground water 

extracted. 

The density of Dehalococcoides cells in the water 

samples was determined using Quantitative Real-time 

PCR of 16S rRNA genes. The assays were performed 

in the laboratory of Elizabeth Edwards at the University 

of Toronto, Ontario, Canada. The quantitation was 

performed using primers identical to those used in 

a commercial test (Quantitative Gene-Trac, SiREM, 

Guelph, Ontario). The primers are specific for variable 

regions of the Dehalococcoides 16S rRNA gene and 

produce an amplicon of 512 base pairs in length and 

are similar to those described by Hendrickson et al. 

(2002) and protected under US patent US6894156B2 

(Hendrickson and Ebersole). Real-time quantitative 

PCR (q-PCR) reactions (50 μl) were performed in 

duplicate using 25 μl of 2X DyNAmo SYBR Green 

qPCR Master Mix (MJ Research Inc., MA), 1.0 μl of 

water containing 25 pmol of each primer and 19 μl 

of DNase and RNase-free water (Sigma) and 4 μl of 

template DNA which were gently mixed at room temperature and transferred into a 96 well plate (OpticonTM 

Systems) and sealed with 8-strip Ultraclear caps (MJ 

research Inc., Waltham, MA). 

Real-time PCR was performed with a DNA Engine 

Opticon 2 System (MJ Research Inc., MA) with initial 

denaturation at 94 °C for 10 min; 45 cycles of 94 °C for 

45 seconds, annealing at 60 °C for 45 seconds, and 

extension of 72 °C for 50 seconds. Standard curves 

of Ct versus log10 16S rRNA gene copy number were 

produced using known quantities of cloned Dehalococcoides 16S rRNA genes. The standard curves 

were used to estimate the number of 16S rRNA gene 

copies in the ground water samples. Verification of the 
21

specificity and identity of the PCR products was determined by melting curve analysis performed between 

72 °C and 95 °C using the Opticon Monitor Software. 

The detection limit is near 2,000 gene copies per liter 

of ground water extracted. 

As mentioned above, SiREM uses primers patented by 

DuPont. Hendrickson et al. (2002) sequenced amplicons of the primers to determine the specificity of the 

primers for Dehalococcoides. Occasionally SiREM will 

sequence the amplicons at the request of their clients. 

In every case, the amplicons have been highly similar 

(percent similarity >90%) to known Dehalococcoides

organisms (personal communication, Philip Dennis, 

SiREM, Guelph, Ontario, Canada). Amplicons from 

samples provided in this study were not sequenced. 

Purified sterile water was processed with every ten 

samples to serve as negative controls or “DNA blanks.” 

If the controls produced a visible band, the test results 

were repeated or invalidated. In some cases where 

the bands from the controls were very weak, that fact 

was noted in the case narrative, and the test results 

were reported. 

If no Dehalococcoides DNA was recovered, PCR 

was conducted with a universal primer to determine 

if amplifiable concentrations of bacterial DNA were 

present in the sample. Genomic DNA from E. coli was 

used as the positive control for the assays with the 

universal primer. The sensitivity of the assay with the 

universal primer has not been explicitly determined, 

but it should be the same or slightly lower than the 

assay with Dehalococcoides primers based on the 

fact that fewer cycles of PCR are performed and that 

the amplification products, for the most part, are longer (personal communication, Philip Dennis, SiREM, 

Guelph, Ontario, Canada). The amplification of the 

bacterial primer was scored as “detected,” “trace,” or 

“not detected.” 

Calculation of Dechlorination Rates from 

Monitoring Data 

The BIOCHLOR decision support system was used 

to calculate the rates of reductive dechlorination of 

chlorinated ethylenes (available at http://www.epa.

gov/ada/csmos.html) (Aziz et al., 2000). The BIOCHLOR software simulates remediation of dissolved 

solvents by natural attenuation at chlorinated solvent 

release sites. It is based on the Domenico analytical 

solutions to the solute transport equation and has 

the ability to simulate one-dimensional advection, 

three-dimensional dispersion, linear adsorption, and 

sequential biotransformation of chlorinated ethylenes 

by reductive dechlorination. It assumes biotransformation follows a pseudo-first order rate law. Parameters 

used to calibrate BIOCHLOR to the plumes are listed 

in Table 4.2. A detailed example of one of the calibrations is provided at the end of this section. 

At six sites in five locations (see Table 4.3), the contamination in ground water formed a conventional 

plume. Each plume had a region of high contamination 

associated with the source and a region with lower 

concentrations extending away from the source in 

the direction of ground water flow. BIOCHLOR was 

calibrated to field data on contaminant concentrations 

from single sampling events. Site-specific information 

was collected from reports or papers on the study 

sites (Acree and Ross, 2003; Altus Air Force Base, 

2002; Dover Air Force Base, 2003; Landau Associates, 1995, 2002; Tinker Air Force Base, 1999, 2002; 

Wilson et al., 1997). To avoid errors in fitting rate 

constants for natural attenuation, the rate constants 

were fit to data that were collected prior to initiation 

of any engineered remedies. 

At two sites (Area 800 and Area 2500) at England AFB, 

Louisiana, there was no discernable overall direction 

of ground water flow in the plumes. The sites were 

overlaid by a bayou that communicated with the contaminated aquifer. Ground water in the aquifer flowed 

toward the bayou or away from the bayou depending 

on the seasons and recent precipitation events. As 

a result, concentration isopleths of TCE, cis-DCE, 

and vinyl chloride were arranged in concentric circles 

that were centered about the source area. Therefore, 

time series data on contaminant concentrations and 

dummy variables for hydrologic properties (seepage 

velocity 100 ft/yr, longitudinal dispersivity 10 ft and 

retardation factor 1) were inserted into BIOCHLOR to 

extract the rate constants of dechlorination over time 

in particular wells. 

Calculation of Dechlorination Rates in 

Conventional Plumes 

Table 4.3 shows the relationship between the concentrations of chlorinated ethylenes in the ground water 

plumes and their apparent rates of dechlorination 

along a flow path in the aquifer. A wide variety of concentrations were represented in the survey; however a 

relatively narrow range of rate constants was deduced. 

There did not appear to be any consistent relationship 

between the concentration of the chlorinated ethylenes 

and their first order rate of biotransformation.

At the Western Processing Site, field data before 

implementation of a pump and treat system were 

used for calibrating BIOCHLOR. The rate constants 

compared very favorably with the rate constants de-
22

rived from that site by other methods (Lehmicke et 

al., 2000, data cited in Table 4.3). 

At the other sites, when possible, field data from 

independent sampling events were used to calibrate 

BIOCHLOR. This was done to evaluate the variability 

over time of the rate constants that were extracted 

from the field data. There was agreement in the rate 

constants extracted for the LF3 plume from data collected in 1997 before the operation of an extraction 

system and in 2002 after the operation of the extraction system. The rate constant we extracted from data 

on the FTA2 plume was faster than a rate constant 

Table 4.2. Calibration Parameters for BIOCHLOR

Parameter Kent, WA Tinker AFB, OK Altus AFB, 

OK

USCG Support 

Center, Elizabeth City, NC

Dover 

AFB, DE

Western 

Processing Landfill 3 FTA2 SS17 North Beach Target Area 1

1988 Sept.

1997

Nov.

2002

Aug.

1997

March

2003 1997 Oct.2002

July

1997

Seepage 

Velocity

(m/yr)

20 29 29 29 8.3 8.5 37

Coefficient 

Dispersion

(m)

2.4 7.6 7.6 45 45 0.6 0.8 7.6

Ratio Lateral 

to Longitudinal 

Dispersion

0.2 0.1 0.1 0.1 0.1 0.2 0.2 0.2

Ratio Vertical 

to Longitudinal 

Dispersion

0.2 none none none none 0.2 0.2 0.2

Retardation 

Factor 1.1 3 3 3 1.3 1.1 1.1 1.0

Simulation Time

(years) 30 60 60 30 30 60 60 60

Modeled Area 

Width

(meters)

120 150 150 150 180 120 120 300

Modeled Area 

Length

(meters)

240 760 760 460 460 60 76 760

Source 

Thickness

(meters)

9 6 6 6 6 9 9 15

Source Width

(meters) 60 76 76 76 60 60 60 150

extracted at the same site in 1999 by a contractor for 

the U.S. Air Force (Parsons, Inc., cited in Tinker Air 

Force Base, 1999). There was agreement in the rate 

constants extracted from the North Beach Site using 

data collected in 1997 before source removal efforts 

and data collected in 2002 after source removal. 

The lower plume in the Target Area 1 is currently attenuating more rapidly near the source areas than 

in the far field. As a result, the current profile of 

concentrations with distance from the source is inverted. Therefore, we chose to calibrate BIOCHLOR 

to historical data collected in 1997. There was good 
23

Table 4.3. Relationship between the Concentrations of Chlorinated Ethylenes in Ground Water and Their Apparent 

Rates of Dechlorination along an Inferred Flow Path in the Aquifer

Facility/ Location Date

PCE TCE cisDCE VC PCE TCE cisDCE VC

Concentration near source

(μg/L)

Pseudo-first order 

rate constant for 

dechlorination

(per year)

Western Processing

Kent, WA

1988 a 10000 460 0.6 3

1999 26 44 0.6 b 1 b

4/2003 0.34 1.59

LF3 (landfill)

Tinker AFB, OK 

9/1997 c 5.3 97 38000 23000 1 3

11/2002 8 28 28400 20400 1 3

North Beach, USCG Support 

Center

Elizabeth City, NC

1997 d 2000 105 74 30.8 0.1 1 0.3 1

10/2002 561 52 25 <1 0.1 1 0.3 1

FTA2 (fire training)

Tinker AFB, OK

8/1997 6.1 9440 1200 1.7 0.1 e

11/2002 7.5 9330 977 2.9 0.3

SS17 Site

Altus AFB, OK 3/2003 38.5 8160 264 4.28 0.01

Target Area 1

(Lower Plume) 

Dover AFB, DE

7/1997 680 2400 560 <1 0.1 0.1 0.1 0.3

1997 f

Transect TA to TB 0.26 0.23 0.31 0.3

Transect TB to TC 0.18 0.07 0.27

Wells TA to TB 0.14 0.06 0.08 0.2

Wells TB to TC 0.48 0.22 0.28

a data collected before the operation of pump and treat system; b data calculated from the half-lives for cis-DCE (1.1 years) and VC 

(0.55 year) obtained by Lehmicke et al. (2000); c data collected before the operation of extraction system, except that data near 

source were from the earliest date available (August 2001); d data collected in May or December of 1997 before source removal; e

rate estimate obtained by Parsons (Tinker Air Force Base, 1999); f data obtained by Ei et al. (2002).

agreement between the rate constants extracted using 

BIOCHLOR and rate constants previously extracted 

by Ei et al. (2002) using a transect approach or wellto-well comparisons (Table 4.3).

Relationship between Dehalococcoides 

DNA and Dechlorination Rates at 

Conventional Plumes 

Dehalococcoides DNA was measured and determined 

to be present in 24 contaminated wells at six sites 

where the contamination in ground water formed conventional plumes. In nine wells at three of the sites, 

Dehalococcoides DNA was unequivocally detected. 

Table 4.4 compares the concentration of Dehalococcoides DNA as determined by the semi-quantitative 

PCR test using gel electrophoresis (Gene-Trac Test), 

and the concentration determined by quantitative 

real time PCR, to the concentrations of chlorinated 

ethylenes and ethylene in individual wells at the three 

sites. 

Figure 4.3 compares the locations of the monitoring 

wells at the Western Processing Site in Kent, Washington, to the distribution of cis-DCE in the plume 

in 1994. At the Western Processing Site, reductive 

dechlorination was essentially complete when the 

wells were sampled for Dehalococcoides DNA in 

2003 (Table 4.4). Wells at the original source area 

of the plume and a well down gradient in the plume 

had detectable concentrations of ethylene (Table 4.4). 

One of the source area wells and the down gradient 

well had detectable concentrations of vinyl chloride. 
24

The wells in the former source area and the down 

gradient well had intermediate concentrations of Dehalococcoides DNA based on the PCR assay with 

gel electrophoresis, and concentrations of Dehalococcoides cells ranging from 105

 to 107 per liter based on 

quantitative real time PCR. Ethylene, vinyl chloride, 

and Dehalococcoides DNA were not detected in the 

three background wells that had never experienced 

the contaminants.

Figure 4.4 compares the location of the monitoring 

wells at the Landfill 3 Site at Tinker AFB, Oklahoma, 

to the distribution of cis-DCE in 2000. Four of the 

wells were originally considered to be in the plume 

(wells 2-259B, 83BR, 2-299B, and 2-292B), and well 

2-304B was originally considered to be a background 

well. Dechlorination was less extensive than at the 

Western Processing Site, but the concentrations of 

chlorinated ethylenes were reduced to a major extent 

along the flow path (Table 4.4). When sampled as 

part of this survey, the background well 2-304B had 

low concentrations of cis-DCE and vinyl chloride. 

Dehalococcoides DNA was detected in all the wells, 

including the background well. The estimated cell 

Figure 4.3. Location of monitoring wells and distribution of cis-DCE at the Western Processing Site in Kent, Washington, fourth quarter 1994.

density varied from 108 per liter in the most contaminated well at the source to 106

 per liter in the down 

gradient wells (Table 4.4). 

Figure 4.5 compares the distribution of wells at the 

North Beach Site at the U.S. Coast Guard Support 

Center at Elizabeth City, North Carolina, to the distribution of PCE in 1997, before excavation of a portion 

of the source. In 1996 and 1997, the concentrations 

of PCE in the most contaminated well (GW 3-30 in 

Figure 4.5) varied from 2,000 μg/liter to 2,900 μg/liter 

(data not shown). When the plume was sampled in 

2002 for Dehalococcoides DNA, the concentration of 

PCE in well GW 3-30 was 561 μg/liter (Table 4.4).

Dechlorination at the North Beach Site was less extensive than at the Western Processing Site or the 

Landfill 3 Site (Table 4.4). Vinyl chloride was only 

detected in one well, and ethylene was not detected 

in any of the wells. The well where vinyl chloride 

was detected (MW1) was down gradient of the well 

in the source area. Dehalococcoides DNA was only 

detected in the down gradient well where vinyl chloride 

was detected. 
25

Table 4.4. Distribution of Chlorinated Ethylenes and Dehalococcoides DNA at Sites that Form Conventional 

Plumes

Well Position PCE TCE cisDCE

Vinyl

Chloride Ethylene DNA Score* Cell Density

μg/L cells/L

Western Processing Site 4/29/2003

15T2B Source <1 <1 <1 <MDL 8 ++ (63%) 2.3 x 106

6M6B Source <1 <1 <1 <1 3 ++ (54%) 3.4 x 107

15T4B Down 

Gradient <1 <1 <1 1.6 6 + (25%) 2.5 x 105

15M17B Back-ground <1 <1 <1 <1 <1 -

15M39B Back-ground <1 <1 <1 <1 <1 -

15M45B Back-ground <1 <1 <1 <1 <1 -

Landfill 3 Site, Tinker AFB, Oklahoma 11/27/2002

2-259B Source 7.95 28.4 28,400 20,400 5.3 ++ (46%) 2.0 x 108

83BR Mid-point 0.52 267 13.1 4.0 <1 ++++ (158%) 2.7 x 106

2-299B Down 

Gradient 0.42 234 0.75 <1 <1 + (24%) 4.8 x 106

2-292B Down 

Gradient 0.43 224 0.97 0.68 <1 + (21%) 1.3 x 106

2-304B Back-ground ND 0.48 0.47 <1 <1 + (15%) 6.7 x 105

North Beach Site, USCG Support Center, Elizabeth City, North Carolina 10/22/2002

GM 3-30 Source 561 52.1 25.3 <1 <1 -

GP23D Mid- Point 397 74.3 27.9 <1 <1 -

MW6 Down 

Gradient 208 18 14.5 <1 <1 -

MW1 Down 

Gradient 382 80.5 79.3 12 <1 ++++ (166%) 1.8 x 105

MW5 Back-ground <1 <1 <1 <1 <1 -

*

 maximum score detected by any of the Dehalococcoides primers. 0% of positive control = (-), <3% = (+/-), 4% to 33% = (+), 34% 

to 66% = (++), 67% to 100% = (+++), >100% of positive control = (++++). 

Table 4.5 shows the relationship between the apparent rates of dechlorination along the flow path and the 

detection of bacterial DNA and Dehalococcoides DNA 

in water samples from a monitoring well in the plume. 

At sites where Dehalococcoides DNA was detected 

in at least one monitoring well (Western Processing, 

LF3, and North Beach), the dechlorination rates of 

cis-DCE and vinyl chloride were equal to 0.3 per year, 

or were greater than 0.3 per year. 

“Generally useful” rates of dechlorination also occurred in a plume where Dehalococcoides DNA was 

not detected. In the lower plume at Target Area 1, 

the extracted rate of attenuation of vinyl chloride 

was 0.3 per year, but Dehalococcoides DNA was not 

detected (Table 4.5). Bacterial DNA was detected in 

water from this plume even though Dehalcoccoides

DNA was not detected.

The SS-17 Site was oxic, and reductive dechlorination was not expected. This site was included in the 

survey as a control. The extracted rate constant was 

very low, 0.01 per year. As would be expected, Dehalococcoides DNA was not detected.
26

Figure 4.4. Location of monitoring wells and distribution of cis-DCE in the Upper Saturated Zone Aquifer at the 

Landfill 3 Site, Tinker AFB, OK, in 2000.

Figure 4.5. Comparison of the locations of monitoring wells at the North Beach Site at the U.S. Coast Guard Support Center in Elizabeth City, North Carolina, to the distribution of PCE in ground water.
27

Table 4.5. Relationship between the Apparent Rates of Dechlorination along a Flow Path in the Aquifer and the 

Detection of Bacterial DNA and Dehalococcoides DNA in Water Samples from a Monitoring Well in the 

Plume

Facility/

Location Date

PCE TCE cisDCE VC Bacterial

DNA 

Primer

Dehalococcoides 

DNA

Rate of Dechlorination

(per year)

Primers

Detected

Maximum 

Score*

Western Processing

Kent, WA

1988 a 0.6 3

4/2000 Not Used +++ b

4/2003 Not Used 3 of 3 ++

LF3 (landfill)

Tinker AFB, OK 

9/1997 c 1 3

11/2002 1 3 Not Used 3 of 3 ++++

North Beach, USCG 

Support Center

Elizabeth City, NC

1997 d 0.3 1 1 3

10/2002 0.1 1 0.3 1 Not Used 2 of 3 ++++

FTA2 (fire training)

Tinker AFB, OK

8/1997 0.1 e

11/2002 0.3 Detected 0

SS17 Site

Altus AFB, OK 3/2003 0.01 Trace 0

Target Area 1

(Lower Plume) 

Dover AFB, DE

7/1997 0.1 0.1 0.1 0.3 Detected 0

*

 maximum score detected by any of the Dehalococcoides primers. 0% of positive control = (-), <3% = (+/-), 4% to 33% = (+), 34% 

to 66% = (++), 67% to 100% = (+++), >100% of positive control = (++++). 

a

 data collected before the operation of pump and treat system; b data obtained by Hendrickson et al. (2002, personal communication); c

 data collected before the operation of extraction system, except that data near source were from the earliest date available 

(August 2001); d data collected in May or December of 1997 before source removal; e rate estimate obtained by Parsons (Tinker 

Air Force Base, 1999); f data obtained by Ei et al. (2002).
28

Figure 4.6. Location of monitoring wells and distribution of cis-DCE in the Intermediate Ground –Water Zone at Spill 

Site-4, the former England AFB, Louisiana, March 2002. 

Dehalococcoides DNA and Dechlorination 

Rates over Time in Particular Wells 

At the two sites at England Air Force Base, there 

was no detectable net direction of ground water flow. 

The site is in the floodplain of the Red River, and 

the direction of flow varies widely with the seasons. 

The direction of flow also varies with the elevation of 

water in a body of surface water (Le Tig Bayou) that 

lies above the contaminated ground water. The distribution of contaminant concentrations forms a “bull’s 

eye” pattern around the two sources of contamination. 

Figure 4.6 compares the location of water wells in 

two separate plumes to the distribution of cis-DCE in 

March, 2002. Wells at one site (Area 2500) showed 

extensive dechlorination, and wells at the other site 

(Area 800) showed very limited dechlorination (Table 

4.6). However, Dehalococcoides DNA was detected 

in both plumes. 

Table 4.7 compares the rates of attenuation over time 

in individual wells to the concentrations of chlorinated 

ethylenes. Two values are entered under “Date Collected” in Table 4.7. The first date is the earliest date 

in the data set used to extract rate constants. The 

second date is the last date used to extract rate constants and the date that the samples were collected 

to assay for Dehalococcoides DNA. In many wells, it 

was not possible to calibrate BIOCHLOR and extract 

rate constant because the chlorinated ethylene was 

absent, or because concentrations increased over 

time. When it was possible to extract rate constants, 

there was not a consistent relationship between the 

concentrations of chlorinated ethylenes in ground water and the rates of dechlorination over time that were 

extracted by calibrating BIOCHLOR to the monitoring 

data (Table 4.7).

Seven wells were sampled for analysis of Dehalococcoides DNA (Tables 4.7 and 4.8). There was also 

no consistent relationship between the presence of 

Dehalococcoides DNA and the trend in concentrations 

of chlorinated ethylenes over time (Table 4.8). Well 

A39L010PZ at Area 2500 behaved like the wells in 

the conventional plumes discussed above. Dehalococcoides DNA was present, and the dechlorination 

rates of cis-DCE and vinyl chloride were both 0.3 per 

year. In contrast, Well A39L011PZ at Area 2500 

contained Dehalococcoides DNA, but the concentration of vinyl chloride increased over time rather than 

decreased. Well A39L009PZ at Area 800 contained 

Dehalococcoides DNA, but the rate of degradation 

of TCE and cis-DCE was very slow, and Well #23 at 

Area 2500 contained Dehalococcoides DNA, but the 

concentrations of both cis-DCE and vinyl chloride were 

increasing over time. 
29

Table 4.6. Distribution of Chlorinated Ethylenes and Dehalococcoides DNA at Sites at England AFB, which do not 

Form Conventional Plumes

Well PCE TCE cis-DCE Vinyl

Chloride Ethylene DNA Score*

Cell 

Density

μg/L cells/L

Area 800

SS45L001MW <1 249 126 4.46 0.02 -

A39L009PZ <1 20.8 52.6 5.21 <1 + (10%) 6.7 x 105

WELL # 17 <1 20.5 30.1 3.88 <1 -

WELL # 19 <1 <1 <1 3.85 <1 -

Area 2500

WELL # 005 <1 <1 0.58 166 0.014 -

A39L011PZ <1 <1 <1 102 0.02 +++ (88%) 4.0 x 106

A39L010PZ <1 <1 <1 95.5 0.895 ++ (37%) 1.2 x 106

WELL # 23 <1 <1 92 60.6 <1 + (30%) 4.8 x 105

WELL #3 1.3 <1 25.6 39.1 0.007 -

A39L015PZ <1 <1 <1 0.395 <1 -

*

 maximum score detected by any of the Dehalococcoides primers. 0% of positive control = (-), <3% = (+/-), 4% to 33% = (+), 34% 

to 66% = (++), 67% to 100% = (+++), >100% of positive control = (++++).

Table 4.7. Relationship between the Concentrations of Chlorinated Ethylenes in Ground Water and Their Apparent 

Rates of Dechlorination over Time in Water Samples from Monitoring Wells at England AFB, Louisiana

Location Well Date

Collected

TCE cis-DCE VC TCE cis-DCE VC

Concentration

(μg/L)

Rate of Dechlorination

(per year)

Area 2500 A39L011PZ 9/1997

4/2003

<1

<1

189

<1

59.2

102

No 

TCE

1 VC ↑

Area 2500 A39L010PZ 6/1997

4/2003

<1

<1

9.5

<1

549

95.5 0.3 0.3

Area 2500 Well #23 5/2000

4/2003

<1

<1

<1

92

<1

60.6 cis-DCE ↑ VC↑

Area 800 A39L009PZ 6/1997

4/2003

35.8

20.8

80.3

52.6

<1

5.2 0.1 0.1 1

Area 800 6/1997

4/2003

451

249

4.3

126

0.5

4.5 0.1 1 3

Area 800 Well #17 3/1999

4/2003

9.8

20.5

<1

30.1

<1

3.9 TCE ↑ cis-DCE ↑ VC ↑

Area 800 Well #19 3/1999

4/2003

<1

<1

<1

<1

<1

3.8

No 

TCE

No cisDCE VC ↑
30

 Again, “generally useful” rates of dechlorination also 

occurred in some wells where Dehalococcoides DNA 

was not detected. In Well SS45L001MW at Area 800, 

the dechlorination rates of cis-DCE and vinyl chloride 

were rapid (1 and 3 per year, respectively), but no 

Dehalococcoides DNA was detected even though 

bacterial DNA was detected. This can be considered 

a false negative for the PCR assay. The assay may 

have failed to detect Dehalococcoides bacteria that 

were present in the aquifer, or the transformation was 

carried out by a strain of Dehalococcoides bacteria 

that was not recognized by the PCR primer, or the 

transformation may have been carried out by other 

bacteria all together.

As would be expected, Dehalococcoides DNA was 

absent in oxygenated ground water where chlorinated 

ethylene concentrations were for the most part below 

detection limits (Well #19 at area 800). 

The rate of attenuation of concentrations of chlorinated 

ethylenes over time in monitoring wells is strongly 

influenced by the rate of physical and chemical weathering of the residual contamination in the source area 

of the plume (Newell et al., 2002). This residual can 

be TCE present as a non-aqueous phase liquid, or 

TCE that is sorbed to aquifer material. Apparently, the 

concentration of chlorinated ethylenes in the wells at 

England AFB was controlled by the rate of dissolution or desorption of the chlorinated ethylenes from 

the source material, and not by biotransformation of 

the chlorinated ethylenes dissolved in ground water. 

When the concentrations of chlorinated ethylenes are 

controlled by dissolution and sorption, a PCR assay 

for Dehalococcoides DNA should not be expected to 

predict the rate of attenuation in concentration over 

time. The PCR assay is not useful to predict the rate 

of natural attenuation in the source areas of plumes.

Table 4.8. Relationship between the Apparent Rates of Dechlorination over Time and the Detection of Bacterial 

DNA and Dehalococcoides DNA in Water Samples from Monitoring Wells at England AFB, Louisiana

Plume Well Date 

Collected

TCE cis-DCE VC

Bacterial 

DNA 

Primer

Dehalococcoides

DNA

Rate of Dechlorination

(per year)

Cell 

Density

cells/L

Area 

2500 A39L011PZ 9/19974/2003

No TCE

1 VC↑ Not Used +++ 3 

of 3

4.0 x 106

Area 

2500 A39L010PZ 6/19974/2003 0.3 0.3 Not Used ++ 3 of 3 1.2 x 106

Area 

2500 Well #23 5/20004/2003

cis-DCE 

↑ VC↑ Not Used + 2 of 3 4.8 x 105

Area 800 A39L009PZ 6/1997

4/2003 0.1 0.1 1 Not Used + 1 of 3 6.7 x 105

Area 800 6/1997

4/2003 0.1 1 3 Detected 0

Area 800 Well #17 3/1999

4/2003 TCE ↑ cis-DCE ↑ VC↑ Detected 0

Area 800 Well #19 3/1999

4/2003 No TCE No cisDCE VC↑ Detected 0

*

 maximum score detected by any of the Dehalococcoides primers. 0% of positive control = (-), <3% = (+/-), 4% to 33% = (+), 34% 

to 66% = (++), 67% to 100% = (+++), >100% of positive control = (++++).
31

Rates of Natural Attenuation and Density of 

PCR Products from Dehalococcoides

DNA

This section compares the rates of natural attenuation at field scale to the density of PCR products 

from Dehalococcoides DNA in monitoring wells. The 

comparison indicates that samples of ground water 

from wells do not adequately represent the density of 

Dehalococcoides bacteria in aquifers.

For a number of reasons, any association between 

Dehalococcoides DNA and rates of natural attenuation 

at field scale must be purely empirical. As discussed 

in the sections above, Dehalococcoides cells may be 

present in the aquifer, but not sampled in the ground 

water produced by a monitoring well. Dehalococcoides cells may be present in the aquifer, but they 

may be dead or inactive. Dehalococcoides DNA for 

the 16S rRNA gene may be extracted from stains of 

Dehalococcoides that are not capable of biological 

transformation of cis-DCE or vinyl chloride. 

To evaluate the effects of any sampling bias caused 

by sampling monitoring wells as opposed to sampling 

the aquifer sediment, the density of Dehalococcoides

DNA in monitoring wells and rates of attenuation at 

field scale were compared to rates in a field study 

of active anaerobic bioremediation (Lendvay et al., 

2003) and the rates in a laboratory culture of Dehalococcoides (Cupples et al., 2004b). Lendvay et al. 

(2003) compared the density of Dehalococcoides

DNA in sediment as measured by the quantitative real 

time polymerase chain reaction (q-PCR) to rates of 

removal of chlorinated ethylenes during in-situ bioremediation of a PCE plume. In a demonstration plot 

that was inoculated with a Dehalococcoides-containing culture, the initial concentration of cis-DCE was 

near 970 μg/L. This concentration is higher than the 

half saturation constant of 320 μg/L determined for 

the Victoria culture of Dehalococcoides (Cupples et 

al., 2004a). However, the removal of cis-DCE in the 

data presented by Lendvay is roughly first order on 

concentration. The plume of PCE was converted to 

a plume that was almost entirely cis-DCE in 8 days, 

and then 36% of the cis-DCE plume was converted to 

ethylene in an additional 12 days, 70% was converted 

to ethylene in 28 days, and 92% was converted to 

ethylene in 35 days. This corresponds to pseudofirst order rate constants for degradation of cis-DCE 

of 13.6, 15.7, and 26.3 per year, respectively. The 

average rate was near 19 per year. 

Lendvay et al. (2003) acquired sediment samples 

from three depths in the center of demonstration plot 

(the injection zone) and from three depths near the 

extraction wells at the periphery of the demonstration 

plot. Averaged across the six samples, the aquifer 

sediment in the plot that was inoculated contained 8 x 

105 Dehalococcoides cells per gram wet sediment. If 

the sediment contained 0.11 ml of pore water per gram 

wet weight (water filled porosity 25%), the density of 

Dehalococcoides DNA in the sediment corresponds 

to 7 x 109 cells in or exposed to a liter of pore water. 

Substrate was supplied to a control plot, but the ground 

water was not inoculated with Dehalococcoides organisms. However, the aquifer contained native strains of 

Dehalococcoides, and cis-DCE was degraded to ethylene. The initial concentration of cis-DCE was near 

1,500 μg/L. Transformation of cis-DCE began after a 

period of acclimation lasting 87 days. There was 27% 

conversion of cis-DCE to ethylene in the subsequent 

27 days and 76% conversion in the subsequent 34 

days, corresponding to first order rates of 8.2 and 15.3 

per year. The average rate was near 12 per year. The 

aquifer sediment in the plot that was not inoculated 

contained on average 3.6 x 104 Dehalococcoides cells 

per gram which corresponds to 3.2 x 108 cells in or 

exposed to a liter of pore water. 

The first order rates of attenuation of cis-DCE are 

compared to the density of Dehalococcoides cells in 

Figure 4.7. There was a little more than one-order 

of magnitude difference in the density of Dehalococcoides cells that achieved a rate of degradation 

near 20 per year in the in-situ bioremediation study 

of Lendvay et al. (2003). This is reasonably good 

agreement between two independent field-scale estimates of the effect of the density of Dehalococcoides

cells on the rate of transformation of cis-DCE in the 

aquifer. This estimate will be compared to the rate of 

natural biotransformation at the natural attenuation 

sites. It is important to note that the estimate is for 

only two pilot-scale plots at one site. We could not 

find another study in the literature that compared the 

rates of biodegradation of cis-DCE to the density of 

Dehalococcoides cells in the sediment. Until other 

studies are reported, there is no benchmark to determine whether the rates achieved by Lendvay et al. 

(2003) are typical.

If the rate of biotransformation of cis-DCE by an individual cell of Dehalococcoides is pseudo-first order 

on concentration, then the overall rate of transformation should be proportional to the number of active 

cells. Two cells would degrade cis-DCE twice as fast 

as one cell, and so on. The grey shape in Figure 4.7 

bounds the area where the rate of biotransformation 

is proportional to the cell density in the plot that was 

bioaugmented with an active culture of Dehalococ-
32

coides, and the plot that relied on the native strains 

of Dehalococcoides. The density of Dehalococcoides

cells in two of the monitoring wells at the MNA sites 

fell within the grey shape, indicating that the performance of the native Dehalococcoides stains in the 

plumes undergoing natural attenuation was roughly 

equivalent to the performance of the Dehalococcoides

stains at the active in-situ bioremediation plots. Based 

on the concentration of cells in the water from the 

monitoring wells, the performance in the other wells 

was substantially better than in the bioremediation 

demonstration plots. The rates were up to one order 

of magnitude faster than would be expected from the 

density of Dehalococcoides cells.

The solid square in Figure 4.7 is the first order rate of 

transformation of cis-DCE by a laboratory culture of 

Dehalococcoides strain VS (Cupples et al., 2004b). 

The culture was growing at 20° C with optimal concentrations of molecular hydrogen. The first order rate 

of cis-DCE transformation was calculated by dividing 

the maximum rate of transformation of cis-DCE by the 

half saturation constant for cis-DCE transformation. 

The dotted line in Figure 4.7 extrapolates the performance of the laboratory culture to cell densities that 

Figure 4.7. Relationship between the density of Dehalococcoides cells as determined by quantitative PCR and the 

first order rate of attenuation of cis-DCE in ground water. The data points with an open symbol are from 

ground water samples collected at natural attenuation sites as presented in Tables 4.3 and 4.6. The 

data points with a solid diamond symbol or a solid triangle symbol are from sediment samples from a 

site where biological reductive dechlorination was used to clean up a PCE spill (Lendvay et al., 2003). 

The data point with a solid square symbol is from a laboratory study of cis-DCE transformation by Dehalococcoides strain VS growing under optimum conditions (Cupples et al., 2004b).

were determined by quantitative PCR in monitoring 

wells at the study sites. Based on the concentration 

of Dehalococcoides cells in ground water from monitoring wells, the Dehalococcoides organisms in the 

aquifer at the field sites performed as well or better 

than the culture growing under optimal conditions in 

the laboratory. 

It is possible, but unlikely, that the native Dehalococcoides organisms at our survey sites were more efficient than the organisms reported by Lendvay et al. 

(2003) and Cupples et al. (2004b). It is possible, but 

unlikely, that other organisms were primarily responsible for natural attenuation of chlorinated ethylenes 

at our study sites. The most likely explanation is that 

the monitoring wells did not efficiently sample the 

Dehalococcoides organisms in the aquifer, and that 

the number of Dehalococcoides cells recovered in a 

liter of well water was a small fraction of the number 

of cells that were exposed to a liter of ground water in 

the aquifer. Most of the Dehalococcoides cells were 

probably attached to sediment particles.

If the relationship between the density of Dehalococcoides cells and the rates of cis-DCE degradation in 
33

Figure 4.8. Comparison of the density of Dehalococcoides cells in ground water as determined by quantitative PCR 

(real time PCR) to the density of Dehalococcoides DNA as determined by the semi-quantitative technique 

that uses the density of the band produced by Gel Electrophoresis as an estimate of the concentration 

of amplified DNA.

the bioremediation study of Lendvey et al. (2003) is extrapolated to a “generally useful” rate of bioremediation 

for natural attenuation of 0.3 per year, then a density 

of near or greater than 1 x 107 Dehalococcoides cells 

per liter is necessary to produce a useful rate of natural attenuation (compare Figure 4.7). Dennis (2005) 

reported that ethylene as a transformation product 

was detected in 78% of ground water samples where 

the density of Dehalococcoides cells was greater than 

1 x 104

 cells per liter but less than 9.9 x 105 cells per 

liter. When the density of Dehalococcoides cells varied between 1 x 106

 cells per liter and 9.9 x 107 cells 

per liter, ethylene was detected in 83% of the wells. 

The detection limit for the quantitative real time PCR 

assay for Dehalococcoides is near 2 x 103 cells per 

liter. The assay can easily detect Dehalococcoides

cells if they are present at densities that cause the 

accumulation of ethylene, or that can produce “generally useful” rates of natural attenuation. At its present 

level of development, the PCR assay has adequate 

sensitivity. False negative reports from the assay 

are most likely caused by a failure of a ground water 

sample to adequately represent the true density of 

Dehalococcoides cells in the aquifer. 

In addition to problems associated with sampling bias 

with water from monitoring wells, the interpretation of 

the semi-quantitative PCR assay based on gel electrophoresis is further complicated by variability in the 

estimate of the amplified DNA. Figure 4.8 compares 

the actual density of Dehalococcoides gene copies to 

the intensity score reported by the semi-quantitative 

test in Tables 4.4 and 4.6. For a given intensity score, 

the number of gene copies varied by as much as two 

orders of magnitude. When the intensity scores were 

greater than (++), there was no observed increase 

in the number of Dehalococcoides gene copies with 

an increase in the intensity score. When an intensity 

score was assigned, the density of Dehalococcoides

gene copies was uniformly greater than 1 x 105 cells 

per liter. However, there was no quantitative relationship between the different intensity scores. 
34

Biotransformation and Dominant Terminal 

Electron Accepting Processes 

Table 4.9 compares the geochemistry of the ground 

water at the studied sites. Based on the concentration of soluble electron acceptors and reduced 

metabolic products, the ground water at each site was 

classified and assigned to categories based on the 

inferred dominant electron accepting process. Most 

of the water samples fell into more than one category. 

This may reflect spatial heterogeneity in the aquifer 

with water from different geochemical environments 

contributing to the water sampled from the well. It 

may also reflect concurrent geochemical processes 

occurring in the aquifer. 

The SS-17 site at Altus AFB and the Area 800 site at 

the former England AFB were included in the survey 

as controls where reductive dechlorination and the 

presence of Dehalococcoides DNA were not expected. 

With the exception of well A39L009PZ at the Area 800 

site, the water at these two sites is oxygenated, and 

the concentration of cis-DCE and vinyl chloride are a 

Table 4.9. Comparison of Rates on Attenuation to the Overall Geochemical Environment of the Sites in the 

Survey

Site Rate Geochemistrya O2

NO3+NO2-

N Fe(II) SO4

-2 CH4 H2

per year mg/L nM

Well 

SS45L001MW 

at Area 800

cis-DCE

1

oxic, iron 

reducing 3.8 0.22 2.5 35.7 0.505 -

Well 2-259B at 

LF3

cis-DCE

1 iron reducing 0.2 0.03 9 <0.1 3.1 -

Well T4 at Western Processingcis-DCE

0.6

iron reducing and 

methanogenic 0.2 0.04 7.5 <0.1 20.7 2.38

Well GM3-30 at 

North Beach 

cis-DCE

0.3 sulfate reducing 0.2 <0.1 0.5 59.7 0.060 <0.4

Well MW-5 at 

North Beach

cis-DCE

0.3 sulfate reducing 0.3 <0.1 0.2 5.72 0.055 7.97

Well 2-62B

at FTA2 at Tinker AFB

TCE

0.3

nitrate reducing, 

sulfate reducing, 

methanogenic

0.1 5.32 - 138 0.938 7.69

Well MW212D 

at Target Area 1

cis-DCE

0.1

nitrate reducing, 

methanogenic 0.4 2.17 <0.1 0.71 3.26 1.19

Well DM353D at 

Target Area 1

cis-DCE

0.1

no dominant 

process 0.7 0.53 <0.1 0.36 0.247 10.47

Well 

A39L009PZ at 

Area 800

cis-DCE

0.1

iron reducing, 

sulfate reducing 0.1 0.08 7 18.6 0.19 6.54

Well WL080 at 

SS17 site,

Altus AFB

TCE

0.01

oxic or nitrate 

reducing 0.7 1.58 <0.1 1670 0.003 <0.4

Well #23 at 

Area 2500 No NA iron reducing, methanogenic 0.1 0.08 12 <0.1 3.11 2.27

a

 oxic: dissolved oxygen was greater than 0.5 mg/L; iron-reducing: iron II was greater than 0.5 mg/L; methanogenic: methane was near 

or greater than 1 mg/L; nitrate-reducing: nitrate plus nitrite –N was greater than 0.5 mg/L, and oxygen was not available; sulfate 

reducing: oxygen was not available, and the concentration of sulfate was greater than 20 mg/L.
35

Figure 4.9. Input screen to BIOCHLOR with calibration parameters for the North Beach Site.

small fraction of the concentration of TCE. Water in the 

lower plume at Target Area 1 is reducing in wells near 

the source (MW212D in Table 4.9), but oxygenated in 

a down gradient well (DM353D in Table 4.9). Water in 

the other plumes is iron-reducing, sulfate-reducing, or 

methanogenic. The rate of attenuation of chlorinated 

ethylenes in the ground water plumes in this survey 

could not be correlated with a unique dominant terminal electron accepting process (Table 4.9).

Example of Calibration of BIOCHLOR with 

Distance along a Flow Path 

The plume of PCE at the North Beach Landfill Site at 

the U.S. Coast Guard Support Center in Elizabeth City, 

North Carolina, will be used to illustrate the process 

of extracting rate constants for natural attenuation 

along a flow path in the aquifer. Figure 4.5 is a map 

showing the location of monitoring wells in the plume. 

The plume of contamination is contained in a shallow 

confined aquifer extending from depths of six to ten 

meters below land surface. The plume is crossed by 

a shallow agricultural drainage ditch that does not 

seem to communicate with the plume. 

Monitoring wells (GM3-30, GP23D, GM6-30, MW1, 

MW6, MW10, and MW7) along the centerline of the 

plume were used to extract the rate constants (some 

of the wells including GM3-30, GP23D, MW1, and 

MW6 were sampled for analysis of Dehalococcoides

DNA). To calculate the rate constants, site-specific 

estimates of the hydrological parameters of the aquifer 

(Wilson et al., 1997) and field data on contaminant 

concentrations in 2000 were entered into the BIOCHLOR software. Figure 4.9 shows the BIOCHLOR 

input screen. 

The first order rate constants were estimated by calibrating BIOCHLOR to field data following a forward, 

trial-and-error process until all the predicted concentrations of all the chlorinated ethylenes best matched 

their field data. 

The output of BIOCHLOR compares the measured 

concentrations of chlorinated ethylenes along a flow 

path to the expected concentrations along the centerline of the plume. At a real field site, the distribution 

of contaminants in monitoring wells may not match 
36

Figure 4.10. Correspondence between the measured values for PCE and TCE at the North Beach Site in 2002 and 

the concentrations that were predicted by calibrating BIOCHLOR using three different values for the 

first order rate constant for biotransformation. The dotted line was considered the best calibration to 

the field data.

the assumptions of the mathematical model. The 

available wells often do not lie along the centerline. 

The screened intervals of the monitoring wells may 

not match the vertical distribution of the contaminant 

in the aquifer. The flow direction of ground water can 

vary over time, moving a well into or out of a flow path 

from the source. The concentrations of contaminants 

in different regions of the source area are usually variable, and different monitoring wells may sample water 

that originated from different regions in the source 

area. The actual rates of biotransformation probably 

vary from one location to the next in an aquifer. 

As would be expected, there was scatter in the field 

data when concentrations were plotted against distance along the inferred flow path (Figure 4.10 and 

Figure 4.11). A sensitivity analysis revealed that if the 

rates of transformation varied by a factor of three, it 

was possible to clearly identify a rate that was the best 

fit to the data in the calibrations. Calibrations were 

examined at rate constants of 0.01 per day, 0.03 per 

day, 0.1 per day, 0.3 per day, 1 per day, 3 per day, 

and 10 per day. The rate constant producing the best 

match to the field data for each chlorinated ethylene 

was selected as the best estimate of the rate. 
37

Figure 4.11. Correspondence between the measured values for cis-DCE and vinyl chloride at the North Beach Site in 

2002 and the concentrations that were predicted by calibrating BIOCHLOR using three different values 

for the first order rate constant for biotransformation. The dotted line was considered the best calibration 

to the field data.
38

Figure 4.12. Input screen to BIOCHLOR with calibration parameters for the well A39L010PZ at Area 2500 at England 

Air Force Base, Alexandria, Louisiana. 

Calibration started with PCE, then proceeded to TCE, 

then to cis-DCE, and then to vinyl chloride. Then each 

calibration was checked again to see if calibration of 

the down stream transformation products had affected 

the calibration of the parent chlorinated ethylenes. 

Figure 4.10 and Figure 4.11 display the correspondence between the measured concentrations and 

concentrations predicted by the BIOCHLOR software. 

The obtained rate constants for reductive dechlorination of PCE, TCE, cis-DCE, and vinyl chloride were 

0.1, 1, 0.3, and 1 per year, respectively. 

Example of Calibration of BIOCHLOR with 

Time in a Single Monitoring Well

To extract rate constants for natural attenuation over 

time in a particular well, data from a series of sampling events were used. Data from the first sampling 

event were used as the source data. Data from other 

sampling events were arranged in an order according to their time interval to the first sampling date, 

similar to the way data were arranged according to 

their distance to the source along a flow path in a 

plume. Dummy variables for hydrologic properties 

were inserted into BIOCHLOR to extract the rate 

constants of dechlorination over time in the particular 

well, as opposed to over distance along a flow path. 

Well A39L010PZ at Area 2500 at the former England 

AFB in Alexandria, Louisiana was used to illustrate 

the process. Figure 4.12 shows the BIOCHLOR input 

screen for this well. 

The rate constants were estimated by calibrating BIOCHLOR to field data from a series of sampling events 

following a forward, trial-and-error process until all the 

predicted concentrations best matched their field data. 

Figure 4.13 displays the correspondence between 

the predicted and actual values performed by the 

BIOCHLOR software. The obtained rate constants for 

reductive dechlorination of cis-DCE and vinyl chloride 

were both about 0.3 per year. 
39

Figure 4.13. Correspondence between predicted and actual values for the chlorinated ethylenes performed by calibrating BIOCHLOR to field data sampled in different events at the well. A39L010PZ at England Air Force 

Base, Alexandria, Louisiana.

chlorinated solvents are controlled by biotransformation of chlorinated solvents already in solution. 

The field scale rates of natural attenuation were 

faster than would be expected from the density of 

Dehalococcoides DNA in ground water samples from 

monitoring wells, suggesting that monitoring wells 

failed to adequately sample the Dehalococcoides

cells in the aquifer. If assays for Dehalococcoides

DNA are to be used to make quantitative predictions 

of the rates of natural attenuation, it will generally be 

necessary to acquire and analyze samples of the 

aquifer sediment.

Conclusions

The commercially available assay for Dehalococcoides

DNA had adequate sensitivity to detect concentrations 

of Dehalococcoides DNA that could sustain “generally 

useful” rates on natural attenuation of cis-DCE and 

vinyl chloride. 

The assay is not appropriate in the source area of 

plumes where the concentrations of chlorinated solvents are controlled by dissolution and desorption of 

chlorinated solvents from residual contamination. The 

assay is more appropriate for those areas of a plume 

outside the source area where the concentrations of 
40
41

Section 5. 

Geochemistry of Ground Water and Occurrence of Dehalococcoides

Synopsis

The Technical Protocol for Evaluating Natural Attenuation of Chlorinated Solvents in Ground Water (Wiedemeier et al., 1998) used a number of geochemical 

parameters in a scoring system to predict whether 

ground water could reasonably be expected to contain 

microorganisms that could cause biotransformation of 

chlorinated solvents. The scoring system was based 

on professional judgment. It was not validated against 

any data set that compared selected geochemical 

parameters against the presence or absence of microorganisms that might be capable of transformation of chlorinated solvents. The scoring system was 

criticized by the Committee on Intrinsic Remediation 

of the National Research Council (NRC, 2000). The 

Committee recommended that the scoring system 

should not be used to evaluate prospects for MNA.

Because the Dehalococcoides group is the only known 

group of organisms that can grow by carrying out 

the reductive dechlorination of DCE or vinyl chloride, 

it plays a critical role in the evaluation of monitored 

natural attenuation of chlorinated ethylenes in anaerobic ground water. To determine the association 

between the presence of Dehalococcoides and the 

critical biogeochemical parameters that define the 

habitat of these organisms, various contaminated sites 

across the United States were sampled. This Section 

compares the distribution of selected geochemical 

parameters to the distribution of Dehalococcoides

DNA in ground water at the sites. A commercially 

available polymerase chain reaction (PCR) assay 

was used to detect Dehalococcoides organisms in 

the water samples. The presence or absence of Dehalococcoides DNA was compared to the values for 

selected geochemical parameters. 

Not every strain of Dehalococcoides can dechlorinate 

every chlorinated ethylene. As discussed in Section 

3, many strains cannot dechlorinate the dichloroethylenes or vinyl chloride to ethylene. However, every organism that has been identified that can dechlorinate 

the dichloroethylenes and vinyl chloride to ethylene is 

a member of the Dehalococcoides group. An assay 

for the presence of Dehalococcoides DNA provides 

direct evidence for the presence of a strain of bacteria 

that might be capable of completely dechlorinating 

chlorinated ethylenes. 

Knowledge of the growth of Dehalococcoides species 

is derived primarily from studies in the laboratory. 

Dehalococcoides are strict anaerobes. To date, they 

are known to use only chlorinated organic compounds 

as electron acceptors and molecular hydrogen (H2) 

as an electron donor. They do not use nitrate, nitrite, 

fumarate, ferric iron, sulfate, sulfite, thiosulfate, sulfur, 

or oxygen as electron acceptors (Maymó-Gatell et al., 

1997; He et al., 2003a; Adrian et al., 2000). In a contaminated aquifer, multiple terminal electron accepting 

processes may occur at the same time. In contaminated aquifers, Dehalococcoides organisms may have 

to compete with other H2-consuming bacteria for the 

shared electron donors (Lee et al., 1998). Obviously, 

the growth of Dehalococcoides can be influenced by 

the geochemical environment. 

Concentrations of nitrate, methane, and the oxidation/

reduction potential were associated with the presence 

or absence of Dehalococcoides DNA with a statistical 

confidence of 95%. The association of the other geochemical parameters with the presence or absence 

of Dehalococcoides DNA was not significant at 95% 

confidence. Logistical regression was applied to the 

data set to develop a formula (equation 5.1) that uses 

the concentrations of nitrate, methane, and the oxidation/reduction potential to calculate the probability that 

Dehalococcoides is present in ground water. 

The formula has several potential applications. A 

calculated probability would be useful to screen wells 

at a site in order to decide which wells should be 

sampled for a PCR assay. As discussed below, it can 

be used to improve the calibration of computer models 

of natural attenuation in ground water. 

The formula provides a simple and rapid way to 

calculate the probability that a specific dechlorinating 

organism (Dehalococcoides) is present at the site, as 

opposed to the scoring system in the Technical Protocol

(Wiedemeier et al., 1998). However, a calculated 
42

probability that an organism exists in ground water is 

not equivalent to a PCR assay for its presence, and 

probabilities calculated from geochemical parameters 

should not be used to replace PCR assays. 

Calibration of Computer Models to Evaluate 

MNA

Often the computer models that are used to evaluate monitored natural attenuation are distributed 

parameter models. In these models, different rates 

of biotransformation can be assigned to each cell 

of the model, based on the conditions pertaining to 

that cell. BIOCHLOR (Aziz et al., 2000), a simple 

screening model for monitored natural attenuation of 

chlorinated solvents, allows the user to assign two 

rates of natural biotransformation, depending on the 

local conditions. To properly calibrate these computer 

models, it is necessary to know the distribution of the 

capacity to transform chlorinated ethylenes throughout the aquifer. Data from the PCR assay may not 

Figure 5.1. Location of contaminated sites used to compare presence or absence of Dehalococcoides DNA to the 

geochemistry of the ground water. The dashed lines identify sites under monitored natural attenuation. 

The solid lines identify sites under active remediation. 

be available from every well at a site. The calculated 

probability of Dehalococcoides organisms based on 

the geochemistry of ground water in a well could be 

used to assign rate constants to cells in a model. If 

Dehalococcoides organisms are expected, a rate 

constant characteristic of reductive dechlorination 

at that site would be assigned. If Dehalococcoides

organisms are not expected, the rate constant for 

reductive dechlorination would be set to zero in that 

particular region of the aquifer. 

Sampling Sites

Samples were collected from fifteen plumes at ten 

locations across the United States as shown in Figure 5.1 and listed in Table 5.1. The contamination 

occurred in a variety of geological matrices including 

consolidated sandstone and siltstone; unconsolidated 

sandy clay; silty to fine sand; fine to course sand; 

and sands and gravels. The contamination sources 

included industrial landfills, airbase fire training areas, 
43

surface impoundments for industrial wastes, and accidental spills of solvents. At most sites, the major 

contaminants were TCE, cis-DCE, and vinyl chloride. 

In three sites (Target Area 1, FF-87, and North Beach), 

PCE was one of the major contaminants. 

A variety of remediation technologies were applied at 

these sites. An emulsion of soybean oil, surfactant, 

yeast extract, and lactate was injected into a barrier 

in the source zone at one site. A permeable reactive 

barrier composed of shredded tree mulch, cotton gin 

compost, and sand was installed across a plume 

Table 5.1. Location of Sites Included in the Survey Comparing the Presence or Absence of Dehalococcoides DNA 

to the Geochemistry of the Ground Water

Location Site

Geochemistrya

Treatment 

Area

Background 

Geochemistry Remedial Technology

No. 

Wells 

Sampled

Altus AFB, 

Oklahoma 

SS-17 I,S,M O,N

Soybean oil, surfactant, 

yeast extract and 

lactate

6

OU-1 I,S,M O,N Plant mulch permeable 

reactive barrier 5

Carswell AFB, 

TX Area 6 I,S,M O,N Iron permeable reactive barrier 5

Dover AFB, 

DE 

Target Area 

1 I,S,M O,N Soybean oil injection 8

Vandenberg 

AFB, CA Site 35 I,S,M O,N Molasses injection 3

USCG 

Elizabeth City, 

NC 

Building 79 I,S,M O,N Iron permeable reactive 

barrier 11

North 

Beach Site I,S O,N MNA 6

England AFB, 

LA

Area 800 O, N O,N MNA 4

Area 2500 I, M O MNA 6

Rickenbacker 

AFB, OH Site 41 S, M S,M MNA 4

Newark AFB, 

OH FF-87 I,S,M I Soybean oil injection 3

Tinker AFB, 

OK 

LF-3 O,N,S,M O MNA 5

FTA-2 N,S,M O,N MNA 2

OFTA O,N O,N Chemical oxidation 6

Kent, WA Western 

Processing I,M I,M MNA attenuation 6

a

 O=oxic: dissolved oxygen was greater than 0.5 mg/L; N=nitrate-reducing: nitrate plus nitrite –N was greater than 0.5 mg/L, and 

oxygen was not available; I=iron-reducing: iron II was greater than 0.5 mg/L; S=sulfate reducing: oxygen was not available, and the 

concentration of sulfate was greater than 20 mg/L; M=methanogenic: methane was near or greater than 1 mg/L

of TCE in ground water at another site. Permeable 

reactive barriers composed of zero-valent iron and 

sand were constructed across the plumes at two sites. 

Soybean oils were injected into the plume at two sites. 

Molasses was injected into a barrier in the source 

zone at one site. An oxidizing reagent (a mixture of 

potassium permanganate and Fenton’s reagent) was 

injected into the plume at one site. Seven other sites 

were managed through monitored natural attenuation. 

At the LF-3 site at Tinker AFB, Oklahoma, ground 

water has been extracted from the down gradient 

portion of the plume in recent years. 
44

Ground Water Sampling, Assay for 

Dehalococcoides DNA, and Chemical 

Analyses 

These procedures and analyses were conducted as 

described in Section 4.

Detection of Dehalococcoides DNA

A total of 81 monitoring wells were sampled from the 

15 sites. Most of these wells were contaminated with 

chlorinated ethylenes at concentrations above their 

maximum contaminant levels (MCLs) in drinking water. 

In 26 wells, Dehalococcoides DNA was unequivocally 

detected. These 26 wells were distributed over 12 

sites as shown in Table 5.2. 

Table 5.2 compares the concentrations of chlorinated 

ethylenes and ethylene to the intensity score of Dehalococcoides DNA in the wells where Dehalococcoides DNA was detected. The extent of reductive 

dechlorination was generally associated with the 

intensity score of Dehalococcoides DNA. That is, in 

10 of the 12 wells that had higher intensity scores 

of Dehalococcoides DNA (scores of ++ or more), 

complete dechlorination to ethylene was observed. 

In contrast, in 9 of the 14 wells that had lower intensity scores of Dehalococcoides DNA (scores of +), 

dechlorination only proceeded to cis-DCE or vinyl 

chloride. However, it should be noted that complete 

dechlorination to ethylene was also observed in four 

wells that had lower intensity scores of Dehalococcoides DNA (scores of +). 

A monitoring well is not an ideal instrument to sample 

bacteria in aquifers. In many aquifers, the bacteria 

are primarily associated with surfaces; they are not 

planktonic. If the water produced by a well has slight 

turbidity from silt or clay-sized particles, bacteria associated with these particles will be sampled with the 

ground water. If the screen of the well has a sand 

pack and if the well has been properly developed, 

turbidity will be low, and bacteria in the aquifer may 

not be sampled effectively. Detectable concentrations of bacterial DNA were recovered from 86% of 

the wells sampled, and detectable concentrations of 

Dehalococcoides DNA were recovered from 32% of 

the wells sampled.

In 53 samples, Dehalococcoides DNA was not detected unequivocally and in two ground water samples, 

the assay for Dehalococcoides DNA was inconclusive 

(intensity of the band in the electrophoresis gel was 

less than 3% of the positive control, intensity scores 

of +/-). For the 53 negative samples, the control using 

a universal bacterial PCR primer was performed to 

determine whether bacterial DNA of any type could 

be extracted from the water sample. The absence 

of bacterial DNA may indicate that sampling of the 

biomass from the aquifer was ineffective, as opposed 

to indicating that Dehalococcoides was not present in 

the aquifer. Most of the samples contained bacterial 

DNA; only 11 of the 53 samples of ground water did 

not contain enough bacterial DNA to allow the amplified DNA to be detected in the electrophoresis gel. 

Biogeochemistry of Ground Water with 

Detectable Dehalococcoides DNA

The geochemistry of the water in the 26 wells where 

Dehalococcoides DNA was detected was evaluated 

to determine the geochemical environment that was 

associated with the presence of Dehalococcoides 

DNA. It is important to remember that a monitoring 

well can produce ground waters from different portions of an aquifer and may mix and blend ground 

waters that have different geochemical properties. 

The presence or absence of a particular biogeochemical parameter, such as molecular oxygen, in 

well water containing Dehalococcoides DNA does not 

necessarily mean that the Dehalococcoides cells in 

the aquifer occupied ground water with the average 

biogeochemical characteristics of the water produced 

from the monitoring well. 

Table 5.3 presents data on concentrations of nitrate 

plus nitrite, concentrations of methane, and the meter 

reading for oxidation/reduction potential for the 26 

wells. As will be discussed later in this section, these 

were the parameters with the strongest association 

with the presence of Dehalococcoides DNA. In general, the concentrations of nitrate plus nitrite were low; 

most were below the detection limit. Concentrations 

of methane were high, usually above 2 mg/L, and the 

oxidation/reduction electrode potential was low, usually less than -50 mV against the Ag/AgCl reference 

electrode.

The associations are reasonable based on the current 

understanding of biological reductive dechlorination. 

Nitrate can serve as a direct inhibitor of reductive 

dechlorination (Nelson et al., 2002) and may prevent 

the colonization of the aquifer by Dehalococcoides

organisms. More importantly, nitrate may also serve 

as a competing terminal electron acceptor, and deplete concentrations of H2

 to concentrations that are 

below the concentration that is necessary to sustain 

the growth of Dehalococcoides bacteria (Fennell and 

Gossett. 1998; Yang and McCarty, 1998). The primary 

cause for the association of the absence of nitrate 

with the presence of Dehalococcoides DNA may be 
45

Table 5.2 The Intensity Scores of Dehalococcoides DNA and the Concentrations of Chlorinated Ethylenes and 

Ethylene in the Wells where Dehalococcoides DNA was Detected

Site Wells Dhc DNA 

Score*

Concentration (μg/L)

PCE TCE cis-DCE VC Ethylene

SS-17, Altus 

AFB, OK

TSMW-5

WL410

++++

+

<0.3

<0.3

64

41

250

15

1300

<1.0

91

<1.5

OU-1, Altus 

AFB, OK PESMP09 + <0.3 25 227 1.7 <1.5

Area 6, Carswell 

AFB, TX

WHGLTA071

WHGLTA072

++++

+

<0.3

<0.3

2.4

2.4

1.7

17

1.0

1.2

58

36

Target Area 1, 

Dover AFB, DE MW236S + <0.3 <0.4 <0.2 11 <1.5

Site 35, 

Vandenberg 

AFB, CA

MW20

MW13

++

+

<0.3

<0.3

210

250

140

13

170

<0.2

2.9

<1.5

Building 79, 

USCG Support 

Center, Elizabeth 

City, NC

ML22.5-0

ML22.5-8

++

+

1.3

1.2

1300

36

22

41

5.8

20

9.0

na

Area 800, 

England AFB, 

LA

A39L009PZ + <0.3 21 53 5.2 <1.5

Area 2500, 

England AFB, 

LA

A39L011PZ

A39L010PZ

Well#23

+++

++

+

<0.3

<0.3

<0.3

<0.4

<0.4

<0.4

<0.2

<0.2

92

100

96

61

20

90

<1.5

Site 41, 

Rickenbacker 

AFB, OH

MW103

MW104

MW105

++

+

+

<0.3

<0.3

<0.3

19

180

<0.4

5400

1700

3.0

940

250

10

41000

1500

1700

LF-3, Tinker 

AFB, OK

83BR

2-259B

2-299B

2-292B

2-304B

++++

++

+

+

+

0.5

8.0

0.4

0.4

<0.3

270

28

230

220

0.5

13

28000

0.8

1.0

0.5

4.0

20000

<0.2

0.7

<0.2

<1.5

5.3

<1.5

<1.5

<1.5

Western 

Processing, 

Kent, WA

6M6B

T2

T4

++

++

+

<0.3

<0.3

<0.3

<0.4

<0.4

<0.4

<0.2

<0.2

0.3

<0.2

<0.2

1.6

3.0

8.0

6.0

North Beach, 

USCG Support 

Center, Elizabeth 

City, NC

MW1 ++++ 380 81 79 12 <1.5

 na: not analyzed 

*

 maximum score detected by any of the Dehalococcoides primers. 0% of positive control = (-), <3% = (+/-), 4% to 33% = (+), 34% 

to 66% = (++), 67% to 100% = (+++), >100% of positive control = (++++). 
46

Table 5.3. The Concentrations of Nitrate plus Nitrite Nitrogen, Methane, and the ORP Meter Reading in the Wells 

where Dehalococcoides DNA was Detected 

Site/Location Well Dhc DNA 

Score*

NO3+NO2-N

(mg/L)

CH4

(mg/L)

Meter ORP

(mV)

SS-17, Altus AFB, OK

TSMW-5 ++++ <0.1 1.7 -210

WL 410 + <0.1 0.7 -82

OU-1, Altus AFB, OK PESMP 09 + <0.1 4.3 -52

Area 6, Carswell AFB, TX

WHGLTA071 ++++ <0.1 4.5 -180

WHGLTA072 + <0.1 3.6 -200

Target Area 1, Dover AFB, DE MW236S + 1.3 0.001 270

Site 35, Vandenberg AFB, CA

MW 20 ++ 0.4 4.5 -150

MW 13 + 8.6 8.3 -61

Building 79, USCG Support 

Center, Elizabeth City, NC

ML22.5-0 ++ 0.1 1.7 -19

ML22.5-8 + <0.1 na -64

Area 800, England AFB, LA A39L009PZ + <0.1 0.2 -77

Area 2500, 

England AFB, LA

A39L011PZ +++ 0.1 2.0 -94

A39L010PZ ++ <0.1 3.8 -140

WELL # 23 + <0.1 3.1 -110

Site 41, Rickenbacker AFB, 

OH

MW 103 + <0.1 1.2 na

MW 104 + <0.1 0.019 na

MW 105 + <0.1 0.97 na

LF-3, Tinker AFB, OK

83BR ++++ 0.2 0.03 150

2-259B ++ <0.1 3.1 -160

2-299B + 2.6 <0.001 330

2-292B + 2.9 <0.001 210

2-304B + 0.2 <0.001 280

Western Processing, Kent, WA

6M6B ++ <0.1 13 180

T2 ++ <0.1 19 -98

T4 + <0.1 21 -100

North Beach, USCG Support 

Center, Elizabeth City, NC MW1 ++++ 0.1 0.1 na

na: not analyzed 

*

 maximum score detected by any of the Dehalococcoides primers. 0% of positive control = (-), <3% = (+/-), 4% to 33% = (+), 34% 

to 66% = (++), 67% to 100% = (+++), >100% of positive control = (++++). 
47

an indirect association with dissolved oxygen. The 

absence of nitrate or nitrite is generally associated 

with the absence of dissolved oxygen. The presence 

of Dehalococcoides DNA in oxygenated ground water 

is not expected based on laboratory studies that show 

that Dehalococcoides organisms are strict anaerobes 

(Maymó-Gatell et al., 1997; He et al., 2003a, He et 

al., 2003b). 

The accumulation of methane would be expected 

in anoxic ground water with high concentrations of 

dissolved hydrogen. The accumulation of methane 

would suggest that supplies of dissolved hydrogen 

are also adequate to allow proliferation of Dehalococcoides organisms. The low electrode potentials (ORP) 

reflect reducing conditions that would favor growth of 

strict anaerobic bacteria such as Dehalococcoides

organisms. 

Data on the association of Dehalococcoides DNA 

and concentrations of oxygen, iron (II), and sulfate 

are presented in Table 5.4. As mentioned above, the 

presence of Dehalococcoides DNA is not expected 

in oxic conditions. In general, ground water that contained Dehalococcoides DNA had low concentrations 

of dissolved oxygen. The four oxic wells that contained 

Dehalococcoides DNA (83BR, 2-299B, 2-292B, and 

2-304B) were all located in site LF-3 at Tinker AFB. 

For several years, this plume has been managed by 

extracting ground water from the down gradient portion 

of the plume. Monitoring wells 83BR, 2-299B, 2-292B, 

and 2-304B were located adjacent to the pumped 

wells. The source area of the plume (represented by 

well 2-259B) was strongly anaerobic and had a high 

intensity score of Dehalococcoides DNA. The influence of the pumped wells may have mixed oxygenated 

ground water with the anaerobic plume that contained 

Dehalococcoides DNA. The PCR assay can detect 

DNA from non-viable Dehalococcoides cells. 

At three sites, Dehalococcoides DNA was not detected 

in any of the wells that were sampled (site FF-87 in 

Newark AFB, Ohio, and sites FTA-2 and OFTA in 

Tinker AFB, Oklahoma). At these three sites, the 

dissolved oxygen level was generally high (data not 

shown) with the median concentrations ranging from 

0.8 mg/L to 7.7 mg/L.

In most cases, the presence of Dehalococcoides

DNA was associated with detectable concentrations 

of iron (II) (Table 5.4). The scoring system presented 

in the Technical Protocol for Evaluating Natural Attenuation of Chlorinated Solvents in Ground Water

(Wiedemeier et al., 1998) suggested that reductive 

dechlorination should not be expected in ground water 

with sulfate concentrations above 20 mg/L. In this 

survey, Dehalococcoides DNA was found in several 

ground waters with concentrations of sulfate above 

100 mg/L. In this survey, there is no evidence that 

sulfate inhibits growth of Dehalococcoides organisms 

or prevents reductive dechlorination in contaminated 

aquifers (Table 5.4).

Biological reductive dechlorination requires a source 

of reducing power either as dissolved hydrogen or as 

an organic substrate. Table 5.5 compares the presence of Dehalococcoides DNA to the concentrations 

of dissolved hydrogen and total organic carbon (TOC). 

Yang and McCarty (1998) found that a mixed culture 

growing on benzoate with cis-DCE available as an 

electron acceptor (at 28° C) poised the hydrogen 

concentration at 2 nanomolar (nM). They interpreted 

this concentration as the minimum concentration of 

hydrogen that would support utilization by organisms 

carrying out reductive dechlorination. Fennell and 

Gossett (1998) reported that the lowest hydrogen concentration that would support dechlorination (at 35° C) 

was as low as 1.5 nM. Following their approach, the 

Technical Protocol for Evaluating Natural Attenuation 

of Chlorinated Solvents in Ground Water (Wiedemeier 

et al., 1998) predicts that reductive dechlorination of 

solvents will occur if the concentration of hydrogen exceeds 1 nM. The concentration of dissolved hydrogen 

was measured in 18 of the 26 wells that contained 

detectable concentrations of Dehalococcoides DNA. 

Of the 18 wells, 14 had dissolved hydrogen equal to 

or greater than 1.0 nM (Table 5.5).

The Technical Protocol for Evaluating Natural 

Attenuation of Chlorinated Solvents in Ground 

Water (Wiedemeier et al., 1998) suggests that 

biological reductive dechlorination is favored 

when concentrations of TOC are greater than 

20 mg/L. Detectable concentrations of Dehalococcoides DNA were found in ground water with much lower 

concentrations of TOC, in the range of 1 to 2 mg/L.

In Table 5.5, wells were assigned to redox conditions 

based on the concentrations of methane, sulfate, 

ferrous iron, and oxygen as follows: conditions were 

considered to be methanogenic when methane was 

greater than 1.0 mg/L; sulfate and iron reducing when 

methane was less than 1.0 mg/L, sulfate was greater 

than 20 mg/L, and ferrous iron was detected; and oxic 

when dissolved oxygen was greater than 1.0 mg/L, 

and ferrous iron and methane were not detected. 

This assignment was done to facilitate comparisons 

between general redox conditions and the presence 

or absence of Dehalococcoides DNA. Of the 26 wells 

where Dehalococcoides DNA was detected, 17 had 
48

Table 5.4. The Concentrations of Oxygen, Ferrous Iron, and Sulfate in the Wells where Dehalococcoides DNA was 

Detected 

Site/Location Well Dhc DNA 

Score*

O2

(mg/L)

Fe(II)

(mg/L)

SO4

-2

(mg/L)

SS-17, Altus AFB, OK

TSMW-5 ++++ 0.2 3.0 880

WL 410 + 0.2 0.2 520

OU-1, Altus AFB, OK PESMP 09 + 0.3 0.7 1100

Area 6, Carswell AFB, TX

WHGLTA071 ++++ 0.1 1.5 <5

WHGLTA072 + 0.1 0.4 <5

Target Area 1, Dover AFB, DE MW236S + 2.3 0.8 56

Site 35, Vandenberg AFB, CA

MW 20 ++ <0.1 30 23

MW 13 + 0.1 0.3 740

Building 79, USCG Support 

Center, Elizabeth City, NC

ML22.5-0 ++ 0.2 3.0 280

ML22.5-8 + 0.1 <0.1 21

Area 800, England AFB, LA A39L009PZ + 0.1 7.0 19

Area 2500, 

England AFB, LA

A39L011PZ +++ 0.5 14 <5

A39L010PZ ++ 0.2 16 <5

WELL # 23 + 0.1 12 <5

Site 41, Rickenbacker AFB, 

OH

MW 103 + na na 470

MW 104 + na na 110

MW 105 + na na 260

LF-3, Tinker AFB, OK

83BR ++++ 1.5 <0.1 42

2-259B ++ 0.2 9.0 <5

2-299B + 2.9 <0.1 64

2-292B + 4.9 <0.1 52

2-304B + 3.8 <0.1 21

Western Processing, Kent, WA

6M6B ++ 0.5 0.5 <5

T2 ++ 0.1 2.3 <5

T4 + 0.2 7.5 <5

North Beach, USCG Support 

Center, Elizabeth City, NC MW1 ++++ 0.1 0.9 77

na: not analyzed 

*

 maximum score detected by any of the Dehalococcoides primers. 0% of positive control = (-), <3% = (+/-), 4% to 33% = (+), 34% 

to 66% = (++), 67% to 100% = (+++), >100% of positive control = (++++). 
49

Table 5.5. The Concentration of Dissolved Molecular Hydrogen and Total Organic Carbon, and the Oxidation/Reduction Potential in the Wells where Dehalococcoides DNA was Detected 

Site/Location Well Dhc DNA 

Score* Redox Condition H2(nM)

TOC

(mg/L)

SS-17, Altus AFB, OK

TSMW-5 ++++ Methanogenic 4.6 8.7

WL 410 +

Sulfate and iron 

reducing 1.0 9.9

OU-1, Altus AFB, OK PESMP 09 + Methanogenic 1.0 15

Area 6, Carswell AFB, TX

WHGLTA071 ++++ Methanogenic >10 0.8

WHGLTA072 + Methanogenic 0.4 9.6

Target Area 1, Dover AFB, 

DE MW236S + Oxic 6.7 4.6

Site 35, Vandenberg AFB, 

CA

MW 20 ++ Methanogenic >10 910

MW 13 + Methanogenic >10 4.8

Building 79, USCG Support Center, Elizabeth City, 

NC

ML22.5-0 ++ Methanogenic 3.7 2.1

ML22.5-8 + 0.9 5.6

Area 800, 

England AFB, LA A39L009PZ +

Sulfate and iron 

reducing 6.5 1.1

Area 2500, 

England AFB, LA

A39L011PZ +++ Methanogenic >10 1.9

A39L010PZ ++ Methanogenic 0.8 2.1

WELL # 23 + Methanogenic 2.3 1.4

Site 41, Rickenbacker 

AFB, OH

MW 103 + Methanogenic na na

MW 104 + Methanogenic na na

MW 105 + Methanogenic na na

LF-3, Tinker AFB, OK

83BR ++++

Sulfate and iron 

reducing na 8.8

2-259B ++ Methanogenic na 92

2-299B + Oxic na <0.5

2-292B + Oxic na <0.5

2-304B + Oxic na <0.5

Western Processing, Kent, 

WA

6M6B ++ Methanogenic 6.4 13

T2 ++ Methanogenic 2.0 21

T4 + Methanogenic 2.4 16

North Beach, USCG Support Center, Elizabeth City, 

NC

MW1 ++++

Sulfate and iron 

reducing 0.2 na

na: not analyzed 

*

 maximum score detected by any of the Dehalococcoides primers. 0% of positive control = (-), <3% = (+/-), 4% to 33% = (+), 34% 

to 66% = (++), 67% to 100% = (+++), >100% of positive control = (++++). 
50

methanogenic conditions, 4 had sulfate and iron reducing conditions, and 4 had oxic conditions. There 

was a strong association between a high intensity 

score for Dehalococcoides DNA and low redox potential. With two exceptions, all the wells with higher 

intensity scores of Dehalococcoides DNA (scores of 

++ or more) had methanogenic conditions. 

In general, the geochemical habitat of Dehalococcoides, as defined by the parameters measured, was 

wide. In the 26 wells where Dehalococcoides DNA 

was detected, the pH ranged from 4.7 to 7.4, the 

temperature ranged from 13.4° C to 33.4°C, the concentration of BTEX ranged from not detected to more 

than 800 mg/L, the alkalinity ranged from 60 mg/L to 

680 mg/L, the electrical conductivity was in the range 

of 200 μs/cm to 5430 μs/cm, and the concentration of 

chloride was in the range of 5 to 1270 mg/L. 

Comparison of Geochemistry where 

Dehalococcoides DNA is Present or 

Absent

The geochemical parameters discovered as significant 

by the multiple testing were used as the explanatory 

variables in a statistical model using logistic regression. The prediction accuracy of the model was 

evaluated by comparing the observed response and 

the predicted probability for the presence of Dehalococcoides DNA. 

To determine whether a particular biogeochemical 

parameter could predict the presence or absence 

of Dehalococcoides DNA in ground water, the distribution of values for that parameter in water where 

Dehalococcoides DNA was detected was compared 

to the distribution in water where Dehalococcoides

DNA was not detected. The samples that did not have 

enough DNA to be amplified by the PCR assay and 

the samples with inconclusive test results for Dehalococcoides DNA were excluded from the comparison. 

A two-sample Kolmogorov-Smirnov test was used to 

compare the data distributions for each geochemical 

parameter. 

Fourteen geochemical parameters were compared. 

To deal with the problem of multiple comparisons, the 

False Discovery Rate (FDR) developed by Benjamini 

and Hochberg (1995) was used to control the proportion of significant results that were in fact type I errors 

(false positive). The two-sample Kolmogorov-Smirnov 

test was performed for each biogeochemical parameter to generate a probability (p) that there was no 

difference between the distribution of the parameter 

in wells where Dehalococcoides DNA was detected 

and the distribution in wells where Dehalococcoides

DNA was not detected. This is the probability of a false 

positive, the probability that the distributions appear to 

be different when in fact they are not different. 

A False Discovery Rate Ø of 0.05 (that is, in every 

20 significant results for different parameters, the expected number of false positives is one) was used in 

this study. The fourteen separate p-values were sorted 

in ascending order p(1), p(2), ...p(14). All geochemical 

parameters with p-values p ≤ p( j ) were rejected where 

j was the largest index for which

p j

( )j ≤ 0 05•

14

.

Table 5.6 shows the probability (p) that the distributions of each biogeochemical parameter are not different between water with and water without Dehalococcoides DNA as predicted by the Kolmogorov-Smirnov 

test. Table 5.6 also shows the FDR threshold for the 

values of p. To be protected from false positives, the 

value of p must be less than the FDR threshold. When 

the FDR threshold was set at a probability Ø of 0.05, 

significant test results were obtained for three of the 

biogeochemical parameters presented in Table 5.5. 

The distribution of the values of the concentration of 

NO3

-1 plus NO2-1-N, of ORP, and of the concentration 

of CH4 was significantly different between the samples 

of ground water where Dehalococcoides DNA was 

detected and samples where Dehalococcoides DNA 

was not detected. As for the other parameters, (O2, 

H2, Fe (II), SO4

-2, TOC, Cl-1, BTEX compounds, alkalinity, electrical conductivity, pH and temperature) there 

were no significant differences in the distribution of 

the values of the parameters at the specified FDR 

controlling level (Table 5.6).

A series of Chi-square tests were performed on each 

of these three parameters. It was found that the 

proportion of NO3

-1 plus NO2-1-N below 0.5 mg/L, the 

proportion of ORP below 0 mV, and the proportion of 

CH4 over 0.5 mg/L were significantly different between 

the water samples where Dehalococcoides DNA was 

either present or absent at 95% confidence level.

The two-sample Kolmogorov-Smirnov test was performed using SPSS software. The Chi-square test was 

performed using the Data Analysis Tool in Microsoft 

Excel. The logistic regression was performed using 

SAS software. 
51

Table 5.6. The Probability (p) that the Distribution of the Measured Values for Selected Geochemical Parameters 

between Ground Water where Dehalococcoides DNA was Present and Ground Water where Dehalococcoides DNA was not Present is not Statistically Different. To be Accepted as Statistically Significant, 

the Value of ( p) for a Parameter must be Less than the Value for the Threshold for the False Discover 

Rate (FDR) 

Parameter

Sample Size

(No. Wells where

Dehalococcoides DNA)

Z value a p-value

(2-tailed)

Index 

Number j

Threshold 

for FDR b

Detected Not 

Detected

Parameters found to be statistically significant at 90% confidence or greater

NO3

-1+NO2-1 -N 26 39 1.874 0.002 1 0.004

ORP 20 30 1.848 0.002 2 0.007

CH4 25 40 1.687 0.007 3 0.011

Parameters not found to be statistically significant at 90% confidence

Fe(II) 23 39 1.544 0.017 4 0.014

TOC 22 32 1.457 0.029 5 0.018

SO4

-2 26 42 1.196 0.114 6 0.021

O2 23 40 1.080 0.194 7 0.025

Alkalinity 15 18 1.017 0.252 8 0.029

Temperature. 17 24 0.943 0.336 9 0.032

BTEX 19 26 0.919 0.367 10 0.036

pH 23 37 0.761 0.608 11 0.039

Electrical Conductivity 17 24 0.704 0.705 12 0.043

Cl-1 18 32 0.636 0.813 13 0.046

a

 Z value in the Kolmogorov-Smirnov test. 

b Threshold for False Discovery Rate, defined as Ø•j

n , where j is the index number of each parameter, and n is the total number of 

parameters compared. In this study, the Ø was 0.05, and n was 14. 

A Predictive Model for the Presence of 

Dehalococcoides DNA 

It would be useful to have a predictive model for the 

presence of Dehalococcoides DNA based on the 

three simple geochemical parameters (methane, 

nitrate plus nitrite nitrogen, and oxidation reduction 

potential) that were found to be statistically correlated 

with the presence or absence of Dehalococcoides

DNA. For this purpose, a logistic regression with 

a binary response for the presence or absence of 

Dehalococcoides DNA was used to model the probability of the presence of Dehalococcoides DNA as a 

function of the biogeochemical parameters expressed 

as continuous variables. In the model, the response 

variable (Y) had two values: Y=1 for the presence of 

Dehalococcoides DNA and Y=0 for the absence of 

Dehalococcoides DNA. The three biogeochemical 

parameters (ORP, CH4, and NO3

-1 plus NO2-1-N) were 

the explanatory variables. The model was as follows 

(Equation 5.1): 

logit p

p

p

ORP CH

( ) 



 

 =  − =

− − × + × −

log

. . . .

1

0 1370 0 0050 0 1328 0 046 4 8 3

1

2

1

× − − −NO plus NO N

where p is the probability of the presence of Dehalococcoides DNA, ORP is the value of oxidation reduction potential against a silver/silver chloride reference 

electrode in millivolts, CH4 is the concentration of 

methane in mg/L, and NO3

-1 plus NO2-1-N is the concentration of nitrate plus nitrite nitrogen in mg/L. 

A total of 45 cases (wells) from ten sites were used 

to develop the model. Of the 45 cases, 20 had response value of 1, and 25 had response value of 0. 

The generalized coefficient of determination (R2) of 

the model was 0.2497, and the Max-rescaled R2 was 

0.3343. The parameter estimates indicated that the 

probability for the presence of Dehalococcoides DNA 

increased with the increase of CH4 or the decrease of 

ORP or NO3

-1 plus NO2-1-N. Table 5.7 compares the 
52

geochemical data from the wells used to develop the 

model to the presence or absence of Dehalococcoides

DNA. Table 5.8 compares the presence or absence of 

Dehalococcoides DNA to the predicted probability for 

the presence of Dehalococcoides DNA and the lower 

and upper 95% confidence limits for the predicted 

probability. Wells are listed in Table 5.8 from the well 

with the highest probability that Dehalococcoides DNA 

is present to the well with the lowest probability that 

Dehalococcoides DNA is present. 

The model accurately predicted the observed presence or absence of Dehalococcoides DNA in most 

of the wells. In 15 of the 20 wells that had observed 

response of 1, the predicted probability for the presence of Dehalococcoides DNA was greater than 0.50. 

In 23 of the 25 wells that had observed response of 

0, the predicted probability for the presence of Dehalococcoides DNA was less than 0.50.

There were seven “outlier” wells where the predicted 

probabilities for the presence of Dehalococcoides DNA 

were greatly different from the observed responses. 

They were well PESMP07 at site OU-1 on Altus AFB, 

OK; well MW236S at the Target Area 1 site at Dover 

AFB, DE; well 15M45B at the Western Processing 

site in Kent, WA; and wells 83BR, 2-292B, 2-299B, 

and 2-304B at the LF-3 site at Tinker AFB, OK. The 

behavior of the wells at the LF-3 site at Tinker AFB, 

OK, may be a result of the ground water extraction 

system, which may have blended anaerobic water 

from the plume bearing the Dehalococcoides DNA 

with clean aerobic ground water. The discrepancy of 

the other sites cannot be explained.

It is possible that the high concentration of methane, 

low concentrations of nitrate, and low values for redox 

potential are better predictors of biological reductive 

dechlorination than the presence or absence of Dehalococcoides DNA. However, we did not attempt a 

statistical comparison of the achieved rates of attenuation at field scale to the values of the geochemical 

parameters. Fitted rates of attenuation of cis-DCE 

and vinyl chloride were available from only three 

sites (Table 4.5). These are too few data for a robust 

comparison. 

Summary and Conclusions

For the evaluation of natural attenuation of chlorinated 

solvents, particularly in an anoxic aquifer, it’s important 

to know if Dehalococcoides organisms are present. 

The presence of Dehalococcoides DNA may be used 

to directly demonstrate the occurrence of biotransformation at the site. The predictive model established 

in this report provides a simple and rapid way to estimate the presence of Dehalococcoides organisms 

based on geochemical condition. This is very useful 

considering the cost associated with the analysis of 

Dehalococcoides DNA. 

The predictive model can be used to screen samples 

to be submitted for the biochemical assay. Fennell and 

her coworkers discovered that Dehalococcoides organisms are heterogeneously distributed in the aquifer 

(Fennell et al., 2001). To obtain useful DNA data from 

the field, it is a good practice to calculate the probability that Dehalococcoides is present based on the 

more easily available geochemical information. The 

water samples that most likely have Dehalococcoides

can be submitted to the biochemical assay. 

It should be noted that the predictive model is not 

intended to replace the biochemical assay. A calculated probability that Dehalococcoides exists in ground 

water is not equivalent to a direct biochemical assay 

for its presence. However, under conditions where 

data from a direct biochemical assay are not available, a calculated probability could be used to properly 

calibrate computer models of natural attenuation. In 

particular, a rate constant for biotransformation of 

lower chlorinated ethylenes such as cis-DCE and vinyl 

chloride is not appropriate to be applied to regions of 

the aquifer where the geochemical parameters suggest that Dehalococcoides organisms should not be 

expected.

Any well that blends ground water from different geochemical environments will produce confusing data. 

The predictions based on Equation 5.1 will work best 

from monitoring wells with relatively short screens. 

The short screens minimize the blending of ground 

water from different geochemical environments. The 

predictions of Equation 5.1 should not be applied to 

pumped treatment wells if the wells produce water that 

is obviously not in geochemical equilibrium.
53

 Table 5.7. Comparisons between the Observed Presence or Absence of Dehalococcoides DNA and the Concentrations of Nitrate plus Nitrite Nitrogen, Methane, and the ORP Meter Reading in the Wells 

Site/Location Well Observed 

Response

Nitrate+ Nitrite

N (mg/L)

CH4

(mg/L)

ORP

(mV)

Western Processing, Kent, WA T4 present 0.04 21 -100

Western Processing, Kent, WA T2 present 0.04 19 -98

Western Processing, Kent, WA 15M45B absent 0.04 12 -93

OU-1, Altus AFB, OK PESMP 07 absent 0.07 4.6 -200

Area 6, Carswell AFB, TX WHGLTA071 present 0.04 4.5 -180

Area 6, Carswell AFB, TX WHGLTA072 present 0.01 3.6 -196

Site 35, Vandenberg AFB, CA MW 20 present 0.35 4.5 -150

SS-17, Altus AFB, OK TSMW-5 present 0.07 1.7 -210

LF-3, Tinker AFB, OK 2-259B present 0.03 3.1 -160

Area 2500, England AFB, LA A39L010PZ present 0.07 3.8 -140

Site 35, Vandenberg AFB, CA MW 13 present 8.6 8.3 -61

Area 2500, England AFB, LA WELL # 23 present 0.08 3.1 -110

OU-1, Altus AFB, OK PESMP 09 present 0.04 4.3 -52

Western Processing, Kent, WA 6M6B present 0.04 13 180

Area 2500, England AFB, LA A39L011PZ present 0.12 2.0 -94

SS-17, Altus AFB, OK WL 410 present 0.05 0.7 -82

Area 800, England AFB, LA A39L009PZ present 0.08 0.2 -77

Area 800, England AFB, LA WELL # 19 absent 0.8 0.002 -24

Western Processing, Kent, WA 15M39B absent 0.03 1.8 73

Target Area 1, Dover AFB, DE MW236D absent 0.04 0.2 51

Area 6, Carswell AFB, TX WHGLFE002 absent 1.2 0.3 54

Area 800, England AFB, LA WELL # 17 absent 0.1 0.02 82

Western Processing, Kent, WA 15M17B absent 0.04 2.1 170

Area 6, Carswell AFB, TX LF04-4E absent 0.23 2.4 180

Area 800, England AFB, LA SS45L001MW absent 0.22 0.5 150

FTA-2, Tinker AFB, OK 2-62B absent 5.3 0.9 110

OU-1, Altus AFB, OK WL 019 absent 1.6 0.09 130

OU-1, Altus AFB, OK WL 250 absent 0.7 0.005 140

LF-3, Tinker AFB, OK 83BR present 0.2 0.03 150

FTA-2, Tinker AFB, OK 2-393B absent 0.5 <0.001 170

OFTA, Tinker AFB, OK 2-440B absent 3.10 <0.001 150

OFTA, Tinker AFB, OK 2-144B absent 0.61 0.9 220

OFTA, Tinker AFB, OK 2-394B absent 2.0 0.06 210

LF-3, Tinker AFB, OK 2-292B present 2.9 0 210

SS-17, Altus AFB, OK WL 082 absent 2.0 0 230

SS-17, Altus AFB, OK WL 094 absent 2.6 0.001 230

Target Area 1, Dover AFB, DE DM353D absent 0.5 0.3 270

SS-17, Altus AFB, OK WL 090 absent 3.7 <0.001 240

Target Area 1, Dover AFB, DE MW236S present 1.3 0.001 270

LF-3, Tinker AFB, OK 2-304B present 0.2 <0.001 280

OFTA, Tinker AFB, OK 2-143B absent 3.4 <0.001 260

Target Area 1, Dover AFB, DE MW101S absent 1.1 0.003 290

OFTA, Tinker AFB, OK 2-329B absent 7.3 0.007 240

SS-17, Altus AFB, OK WL 080 absent 1.6 0.003 330

LF-3, Tinker AFB, OK 2-299B present 2.6 <0.001 330

a

 LCL: lower 95% confidence limit for the predicted probability; b UCL: upper 95% confidence limit for the predicted probability.
54

Table 5.8. Comparisons between the Observed Presence or Absence of Dehalococcoides DNA and the Predicted 

Probabilities for the Presence of Dehalococcoides DNA 

Site/Location Well Observed 

Response

Predicted 

Probability LCLa UCLb

Western Processing, Kent, WA T4 present 0.958 0.315 0.999

Western Processing, Kent, WA T2 present 0.949 0.334 0.999

Western Processing, Kent, WA 15M45B absent 0.867 0.449 0.981

OU-1, Altus AFB, OK PESMP 07 absent 0.811 0.526 0.943

Area 6, Carswell AFB, TX WHGLTA071 present 0.799 0.519 0.936

Area 6, Carswell AFB, TX WHGLTA072 present 0.789 0.491 0.936

Site 35, Vandenberg AFB, CA MW 20 present 0.765 0.500 0.914

SS-17, Altus AFB, OK TSMW-5 present 0.756 0.407 0.934

LF-3, Tinker AFB, OK 2-259B present 0.746 0.461 0.909

Area 2500, England AFB, LA A39L010PZ present 0.743 0.477 0.901

Site 35, Vandenberg AFB, CA MW 13 present 0.705 0.087 0.984

Area 2500, England AFB, LA WELL # 23 present 0.692 0.440 0.865

OU-1, Altus AFB, OK PESMP 09 present 0.668 0.436 0.84

Western Processing, Kent, WA 6M6B present 0.651 0.108 0.966

Area 2500, England AFB, LA A39L011PZ present 0.643 0.387 0.837

SS-17, Altus AFB, OK WL 410 present 0.591 0.320 0.815

Area 800, England AFB, LA A39L009PZ present 0.567 0.292 0.807

Area 800, England AFB, LA WELL # 19 absent 0.487 0.259 0.721

Western Processing, Kent, WA 15M39B absent 0.435 0.254 0.634

Target Area 1, Dover AFB, DE MW236D absent 0.407 0.219 0.628

Area 6, Carswell AFB, TX WHGLFE002 absent 0.394 0.226 0.592

Area 800, England AFB, LA WELL # 17 absent 0.366 0.191 0.584

Western Processing, Kent, WA 15M17B absent 0.329 0.150 0.576

Area 6, Carswell AFB, TX LF04-4E absent 0.327 0.149 0.574

Area 800, England AFB, LA SS45L001MW absent 0.306 0.151 0.522

FTA-2, Tinker AFB, OK 2-62B absent 0.306 0.075 0.707

OU-1, Altus AFB, OK WL 019 absent 0.301 0.162 0.491

OU-1, Altus AFB, OK WL 250 absent 0.296 0.152 0.496

LF-3, Tinker AFB, OK 83BR present 0.287 0.135 0.508

FTA-2, Tinker AFB, OK 2-393B absent 0.268 0.126 0.482

OFTA, Tinker AFB, OK 2-440B absent 0.260 0.109 0.501

OFTA, Tinker AFB, OK 2-144B absent 0.246 0.105 0.475

OFTA, Tinker AFB, OK 2-394B absent 0.220 0.099 0.422

LF-3, Tinker AFB, OK 2-292B present 0.207 0.083 0.432

SS-17, Altus AFB, OK WL 082 absent 0.203 0.086 0.408

SS-17, Altus AFB, OK WL 094 absent 0.194 0.077 0.410

Target Area 1, Dover AFB, DE DM353D absent 0.183 0.062 0.434

SS-17, Altus AFB, OK WL 090 absent 0.182 0.059 0.441

Target Area 1, Dover AFB, DE MW236S present 0.174 0.063 0.399

LF-3, Tinker AFB, OK 2-304B present 0.174 0.053 0.442

OFTA, Tinker AFB, OK 2-143B absent 0.171 0.057 0.416

Target Area 1, Dover AFB, DE MW101S absent 0.162 0.054 0.398

OFTA, Tinker AFB, OK 2-329B absent 0.154 0.017 0.655

SS-17, Altus AFB, OK WL 080 absent 0.132 0.037 0.372

LF-3, Tinker AFB, OK 2-299B present 0.128 0.037 0.363

a

 LCL: lower 95% confidence limit for the predicted probability; b UCL: upper 95% confidence limit for the predicted probability 
55

Section 6. 

Presence of Dehalococcoides DNA and the Extent of Biodegradation

Presence of Dehalococcoides DNA in ground water 

may not indicate that the bacteria are active and 

degrading the chlorinated solvents. On the other 

hand, the failure to detect Dehalococcoides using a 

particular PCR primer does not prove that the aquifer does not contain Dehalococcoides organisms or 

other bacteria that can degrade chlorinated solvents 

to ethylene. Laboratory microcosm studies conducted 

with sediment or enrichment cultures conducted with 

ground water from a contaminated site play a key 

role in determining if the bacteria at a site have the 

potential for complete dechlorination (David Ellis, 

DuPont Company, Wilmington, Delaware, Personal 

Communication, September 16, 2005). To determine 

if dechlorination of chlorinated ethylenes could be carried out by indigenous microorganisms in the samples 

of ground water, ground water was used to establish 

enrichment cultures. The results of laboratory studies 

were compared to the distribution of Dehalococcoides DNA in the contaminated ground water at field 

scale. These experiments were intended to establish 

a connection between the extent of observed activity 

and the presence of Dehalococcoides as detected 

by PCR.

Enrichment Culture Preparation 

Ground Water samples for inoculation were collected 

from PCE and TCE contaminated wells at the SS-17 

Site at Altus AFB, OK; the OU-1 Site at Altus AFB, OK; 

Area 6 at Carswell AFB, TX; Target Area 1 at Dover 

AFB, DE; Site 35 at Vandenberg AFB, CA; Building 

79 at the USCG Support Center, Elizabeth City, NC; 

the North Beach Site at the USCG Support Center, 

Elizabeth City, NC; Area 800 at England AFB, LA; 

Area 2500 at England AFB, LA; the LF-3 Site at Tinker 

AFB, OK; the OFTA Site at Tinker AFB, OK; and the 

Western Processing Site at Kent, WA (described in 

Table 5.1). A total of 76 ground water samples were 

collected for constructing the enrichment cultures. 

Ground water samples were collected in sterile 40 ml 

VOA vials with no preservatives. The samples were 

collected with no headspace in the vials. 

Enrichment cultures were prepared in 160 ml serum 

bottles in an anaerobic glove box. For each ground 

water sample, two enrichment cultures were constructed in the same manner except that propionate 

(5 mM) was added as an additional elector donor in 

one enrichment culture but not in the other. Each 

enrichment culture contained 100 ml autoclaved basal 

medium and 25 ml ground water (as an inoculum) and 

was amended with 10 ml of a PCE stock solution in 

water or 1.5 ml of a TCE stock solution in water to 

produce a nominal concentration of 100 μM in the 

enrichment culture (nominal concentration corrects 

for headspace-liquid partitioning equilibrium within a 

bottle). The enrichment culture was sealed with a Teflon-lined butyl rubber septum and an aluminum crimp 

cap. The composition of the basal medium is shown 

in Table 6.1. The enrichments contained 0.1 g/L yeast 

extract, which is adequate to completely dechlorinate 

100 μM of PCE. 

Five rounds of enrichment cultures were constructed 

depending on the date the ground water samples were 

collected. In each round of enrichment cultures, two 

controls (with and without propionate) were prepared 

in the same manner except that an additional 25 ml 

of basal medium were added to each vial instead of 

ground water. The controls were preserved with 1% 

trisodium phosphate and autoclaved for 45 minutes 

at 121°C. All the enrichment cultures were incubated 

statically on their sides in an anaerobic glove box at 

room temperature (roughly 20°C).

The enrichment cultures are similar to the microcosm 

studies described in the RABITT protocol (Battelle Memorial Institute et al., 2002; Morse et al., 1998), and to 

microcosm studies conducted commercially (Findlay 

and Fogel, 2000). The concentrations of chlorinated 

solvents, of yeast extract, and of primary substrate 

are similar. An important exception is the absence 

of aquifer sediment in the enrichment cultures in this 

study. The RABITT protocol suggests 50 g sediment 

and 50 ml of ground water. At many hazardous waste 
56

Sampling and Analysis of Enrichment 

Cultures

Resazurin was used as a visual indicator of redox 

condition in the enrichment cultures (purple indicating an oxidizing condition, and the absence of color 

indicating a reduced condition). Enrichment cultures 

were sampled periodically (nondestructively) using 

small samples of liquid or headspace. The samples 

were analyzed for chlorinated ethylenes (PCE, TCE, 

DCE isomers, and vinyl chloride), dissolved gases 

(methane, ethylene, ethane, and hydrogen), and organic acids (propionate and acetate). 

For analysis of chlorinated ethylenes, as well as for 

analysis of fatty acids, 1 ml of liquid was taken from 

each enrichment culture using a 1 ml glass syringe 

and injected to the bottom of a 40 ml VOA vial containing 40 ml RO (reverse osmosis) water and 2 drops 

of concentrated hydrochloric acid. The vial was immediately sealed after sample collection. Analysis 

of chlorinated ethylenes was performed using automated purge and trap gas chromatography. Organic 

acids were analyzed using high performance liquid 

chromatography. 

For analysis of methane, ethylene, and ethane, 0.2 ml 

of gas was taken from the headspace of each enrichment culture into a 2.0 ml gas tight syringe; the sample 

was diluted to 2.0 ml with nitrogen and then injected 

into a GC for analysis. The high purity nitrogen used 

to dilute the sample was free of detectable concentrations of methane, ethylene, and ethane. Henry’s 

Law was used to calculate the original concentrations 

of methane, ethylene, and ethane in the water in the 

enrichment cultures. 

For analysis of hydrogen, a gas sample of 0.2 ml was 

taken from the headspace of each enrichment culture 

into a 2.0 ml gas tight syringe; the sample was diluted 

to 2.0 ml with nitrogen and then injected into a RGA3 

Reduction Gas Analyzer. The high purity nitrogen 

used to dilute the sample was free of detectable 

concentrations of molecular hydrogen. The original 

concentrations of dissolved molecular hydrogen in 

the water in the enrichment cultures were calculated 

using Henry’s Law. 

Biodegradation of Chlorinated Ethylenes in 

the Enrichment Cultures 

All the enrichment cultures were incubated for at 

least 20 months. Some enrichment cultures were 

incubated for more than 30 months. Out of 152 

enrichment cultures from ground water, ethylene as 

Table 6.1. Composition of the Basal Medium (pH 7)

Composition Concentration Unit

NH4Cl 1.0 g/L

MgSO4∙7H2O 0.1 g/L

CaCl2∙2H2O 0.05 g/L

yeast extract 0.1 g/L

resazurin 1.0 mg/L

KH2PO4 0.0066 Molar

Na2HPO4 0.013 Molar

EDTA 1.0 μg/L

FeSO4∙7H2O 2.0 μg/L

CaCl2∙6H2O 0.2 μg/L

MnCl2∙4H2O 0.03 μg/L

NiCl2∙6H2O 0.02 μg/L

ZnSO4∙7H2O 0.1 μg/L

H2SeO3 0.02 μg/L

H3BO3 0.3 μg/L

CuCl2∙2H2O 0.01 μg/L

NaMoO4∙2H2O 0.033 μg/L

CoCl2∙6H2O 0.2 μg/L

p-aminobenzoic 

acid 100 μg/L

folic acid 50 μg/L

lipoic acid 100 μg/L

nicotinic acid 200 μg/L

riboflavine 100 μg/L

thiamine 200 μg/L

panthotenic acid 100 μg/L

pyridoxamine 500 μg/L

vitamin B12 100 μg/L

biotine 20 μg/L

sites, the site owner cannot afford to collect sediment 

for the laboratory studies, and the studies are often 

done with ground water from established monitoring 

wells (Margaret Findlay, Bioremediation Consulting 

Incorporated, Watertown, MA, Personal Communication, November 3, 2005).
57

a dechlorination end product was observed in 28 

enrichments. These 28 enrichments originated from 

17 ground water samples (15 had been spiked with 

propionate and 13 had not). Of the 28 enrichments, 

ethylene as the only end product (100% recovery of 

the total chlorinated ethylenes on a molar basis) was 

observed in eight cases; ethylene as the major end 

product (68%-99% recovery of the total chlorinated 

ethylenes) was observed in four cases; and a smaller 

amount of ethylene (1%-16% recovery of the total 

chlorinated ethylenes) was observed in 16 cases. 

Vinyl chloride as a dechlorination end product was 

observed in nine enrichments. These nine enrichments 

originated from six ground water samples (2 had been 

spiked with propionate and 7 had not). The observed 

molar recovery of total chlorinated ethylenes as vinyl 

chloride ranged from 1% to 24%.

Dichloroethylenes (mainly cis-DCE) as a dechlorination end product, were observed in 28 enrichments. 

These 28 enrichments originated from 15 ground 

water samples (14 had been spiked with propionate 

and 14 had not). The observed molar recovery of 

total chlorinated ethylenes as total DCE (sum of three 

isomers) ranged from 1% to 74%.

In three enrichment cultures amended with PCE, 

a trace of TCE was detected as a transformation 

product. No transformation of the added PCE or 

TCE was observed in the other 84 ground water 

amended enrichment cultures. No transformation 

of the added PCE or TCE was observed in the ten 

sterile controls. 

For most ground water samples, the spike of propionate as an additional electron donor did not seem to 

greatly increase the rate and extent of dechlorination 

in the enrichments. Complete dechlorination to ethylene or partial dechlorination to DCE or vinyl chloride 

frequently occurred in the enrichments without the 

spike of propionate. This observation was not surprising because a significant amount of yeast extract 

(100 mg/L) was contained in the basal medium in the 

enrichment cultures. Acetate in the range of 0.02 to 

3.7 mM (1.2 to 220 mg/L) was detected in the enrichments without the spike of propionate, probably due 

to fermentation of yeast extract in the basal medium 

and organic compounds in the ground water. 

The measurement of dissolved molecular hydrogen 

also confirmed the availability of molecular hydrogen 

as an electron donor. In nearly all of the enrichment 

cultures (96%), the dissolved hydrogen concentrations were greater than 1 nM, a condition favorable 

for reductive dechlorination. The concentration of 

molecular hydrogen was greater than 9 nM in 75% 

of the enrichments, greater than 130 nM in 50% of 

the enrichments, and greater than 3000 nM in 25% of 

the enrichments. There was no statistically significant 

difference in the distribution of dissolved hydrogen 

concentration between the enrichment cultures spiked 

with propionate and the enrichment cultures without 

the spike of propionate (significance level α=0.05). 

Association of Dechlorination in Enrichment 

Cultures with Dehalococcoides DNA 

Prior to constructing the enrichment cultures, ground 

water samples collected from the same wells were 

analyzed for the presence or absence of Dehalococcoides DNA. Table 6.2 compares the extent of biotransformation of the chlorinated ethylenes obtained 

in the enrichment cultures to the presence or absence 

of Dehalococcoides DNA in the field samples used to 

establish the enrichments. 

Dehalococcoides DNA was detected in water samples 

from 22 wells out of a total of 72 sampled. The water 

was used to establish 44 enrichment cultures (one 

with propionate, one without for each of the 22 wells). 

Of the 44 enrichment cultures, 20 showed complete 

biotransformation to ethylene. The 20 microcosms 

corresponded to 11 of the 22 ground water samples 

where Dehalococcoides DNA was detected. In two 

ground water samples where Dehalococcoides DNA 

was detected, dechlorination proceeded only as far as 

DCE in the four corresponding enrichment cultures. 

Vinyl chloride as an end product was not detected in 

enrichment cultures that were inoculated with ground 

water that contained amplifiable Dehalococcoides

DNA. If dechlorination proceeded past dichloroethylene, it went all the way to ethylene.

Dehalococcoides DNA was not detected in water 

samples from 50 wells out of the 72 wells that were 

used to establish the enrichment cultures. Complete 

dechlorination to ethylene was detected in only 7 of the 

100 enrichment cultures that were constructed from 

the ground water samples that did not have amplifiable Dehalococcoides DNA. Dechlorination to vinyl 

chloride was detected in 8 enrichment cultures and 

dechlorination to DCE was detected in 20 of the 100 

enrichment cultures. If Dehalococcoides DNA was 

not detected, the majority of the cultures that showed 

activity stopped at the production of DCE.

The capacity to transform PCE or TCE to cis-DCE is 

common in anaerobic bacteria. The PCR assay for 

Dehalococcoides DNA is conventionally interpreted as 

an assay for organisms that might have the capabil-
58

ity to completely transform chlorinated ethylenes to 

ethylene. Out of 44 enrichment cultures established 

with ground water with detectable concentrations of 

Dehalococcoides DNA, 24 cultures failed to produce 

ethylene. If false positives for the assay are defined 

as water samples where Dehalococcoides DNA was 

detected, but ethylene was not detected in the enrichment culture, then 55% of the cultures were false positives. As will be discussed later, it is possible that the 

community of dechlorinating organisms was damaged 

during collection, transport, or storage of the sample 

prior to construction of the enrichment cultures. It is 

also possible that conditions in the enrichment cultures 

failed to support growth of organisms that could and 

did grow in the aquifer, and could dechlorinate PCE or 

TCE to ethylene. If the proportion of false positives is 

calculated on the basis of cultures that produced any 

transformation product, then 4 out of 24 enrichment 

cultures failed to produce ethylene. On this basis, the 

proportion of false positives is 17%. 

It is also possible that the Dehalococcoides DNA detected by the PCR assay belonged to strains that could 

not transform chlorinated ethylenes to ethylene. 

Although the number of false positives in our survey 

is high, the PCR assay can still be useful to evaluate 

monitored natural attenuation. This is particularly true 

when the overall determination is based on a variety 

of tests and conditions. The interpretation of the PCR 

assay is more straightforward if it is supported by other 

information that would be consistent with on-going 

dechlorination to ethylene in the aquifer. Additional 

information might include geochemical conditions that 

are conducive to the growth of Dehalococcoides species, a reduction in the concentration of vinyl chloride 

in ground water over time, and the accumulation of 

significant concentrations of ethylene. 

An absence of Dehalococcoides DNA would suggest 

that Dehalococcoides organisms were absent from 

the ground water and that dechlorination would not 

proceed to vinyl chloride or ethylene. If false negatives 

for the assay are defined as water samples where 

Dehalococcoides DNA was not detected, but vinyl 

chloride or ethylene was detected in the enrichment 

culture, then the proportion of false negatives is 15 

out of 100 enrichments, or 15%. When Dehalococcoides DNA was not detected in the ground water used 

to establish the cultures, 35 out of 100 enrichments 

showed production of some transformation product 

(dichloroethylene, vinyl chloride, or ethylene). The 

cultures transformed PCE or TCE to ethylene in 7 out 

of the 35 enrichments, to vinyl chloride in 8 of the 35 

enrichments, and stopped at the level of DCE in 20 

of the 35 enrichments. The proportion of false negative predictions, as evaluated against the number of 

enrichment cultures that showed production of at least 

one of the transformation products, was 15 out of 35 

cultures, or 43%. A failure to detect Dehalococcoides

DNA in a sample of ground water using the PCR assay should not be taken to mean that dechlorination 

in the aquifer will stop at the level of DCE. 

Several interactions might account for the complete 

lack of dechlorination activity in roughly half of the 

microcosms. First, the activity of dechlorinating bacteria might be suppressed by competition from other 

hydrogen utilizing bacteria such as the methanogens. 

Yang and McCarty (1998) reported that dechlorinators 

competed best against methanogens and acetogens 

when the hydrogen level was maintained between 2 

Table 6.2. Comparisons of Biotransformation of Chlorinated Ethylenes in Enrichment Cultures to the Corresponding Presence or Absence of Amplifiable Dehalococcoides DNA in the Ground Water Sample Used to 

Inoculate the Enrichment Cultures

Biotransformation 

Product Detected on the 

Last Sampling Date

Number of Enrichment 

Cultures where the Presence 

of Dehalococcoides DNA 

was detected in the Field

Number of Enrichment 

Cultures where the Presence 

of Dehalococcoides DNA 

was not detected in the Field

No Product 20 of 44 65 of 100

Dichloroethylene 4 of 44 20 of 100

Vinyl Chloride 0 of 44 8 of 100

Ethylene 20 of 44 7 of 100

The test results for the presence of Dehalococcoides DNA in two ground water samples were inconclusive (score of +/-); therefore, the 

enrichment cultures constructed from these two samples were not included in the comparison. Twelve enrichment cultures prepared 

using material from the North Beach Site were amended with PCE as the source contaminant. Of the 12 enrichment cultures, three 

had TCE as the biotransformation end product on the last sampling date (not shown in Table 6.2). 
59

and 11 nM. The basal medium in the microcosms is 

a much richer nutritional environment than the contaminated ground water at the sites. In the enrichment 

cultures, the hydrogen level was greater than 11 nM 

in 72% of the enrichment cultures. This may have 

posed a competitive advantage to other bacteria over 

dechlorinating bacteria.

Another possibility is that the living dechlorinating 

bacteria in the ground water samples that were returned to the laboratory for the enrichment study were 

killed by oxygen before the enrichment cultures were 

constructed. Many of the water samples had detectable concentrations of dissolved oxygen. The oxygen 

may have entered the well water in the monitoring well 

when oxygenated ground water in uncontaminated 

portions of the aquifer was mixed with the contaminant plume. The oxygen may also have entered the 

well water from the atmosphere when the well was 

sampled. 

A plot of the relationship between the production of 

ethylene in the enrichment cultures against the dissolved oxygen concentration measured in the field in 

the corresponding ground water used to inoculate the 

Figure 6.1. Relationship between the production of ethylene in the enrichment cultures and the concentration of 

dissolved oxygen in the corresponding ground water used for the inoculums of the enrichment cultures. 

Diamond symbols are ground water samples where Dehalococcoides DNA was detected, and square 

symbols are ground water samples where Dehalococcoides DNA was not detected. 

culture indicated that the dechlorination activity was 

strongly influenced by the dissolved oxygen concentration (Figure 6.1). The production of ethylene was 

expressed as the molar ratio of the final concentration 

of ethylene to the initial concentration of PCE or TCE 

supplied to the enrichments. In all 20 enrichment 

cultures where Dehalococcoides DNA was detected 

in the ground water used to establish the enrichment 

cultures, and the enrichment cultures had accumulated ethylene, the corresponding dissolved oxygen 

concentration in the well water sample used to establish the enrichment culture was below 0.5 mg/L. This 

observation agrees with reports that Dehalococcoides

organisms are strict anaerobes. The same relationship held for the enrichments where Dehalococcoides

DNA was not detected in the ground water used to 

establish the enrichment culture. With two exceptions, 

no ethylene was produced in any enrichment that was 

inoculated with ground water that had dissolved oxygen concentrations higher than 0.5 mg/L. In the two 

exceptional enrichments, only trace concentrations 

of ethylene were detected (the molar ratio of the final 

concentration of ethylene to the initial concentration 

of TCE was less than 0.01).
60

Table 6.3 compares the extent of dechlorination in the 

enrichment cultures with the geochemical character of 

the ground water used to inoculate the culture. (See 

Section 5 for the definition of the geochemical categories.) Dechlorination of PCE and TCE to ethylene 

or vinyl chloride occurred most frequently and most 

extensively in the enrichment cultures inoculated with 

methanogenic ground water. Dechlorination to vinyl 

chloride or ethylene was limited in cultures inoculated 

with water that was sulfate-reducing, iron-reducing, or 

oxic. However, dechlorination to DCE was frequent 

and extensive when the cultures were inoculated 

with ground water that was sulfate-reducing or ironreducing. 

Table 6.3. Comparison of Biotransformation of Chlorinated Ethylenes in Enrichment Cultures to the Corresponding 

Geochemistry of the Ground Water used for Inoculation of the Enrichment Cultures

Geochemistry

Number of Enrichment Cultures in Category

Mole Percent of Final Ethylene to Initial PCE or TCE

No Ethylene <1% 1%-50% >50%

Methanogenic 29 5 9 9

Sulfate and/or iron reducing 52 0 0 2

 Oxic 44 2 0 0

Mole Percent of Final VC to Initial PCE or TCE

No VC <1% 1-50% 50%

Methanogenic 39 1 8 4

Sulfate and/or iron reducing 51 3 0 0

Oxic 44 1 1 0

Mole Percent of Final DCE to Initial PCE or TCE

No DCE <1% 1-50% 50%

Methanogenic 25 18 9 0

Sulfate and/or iron reducing 29 18 7 0

Oxic 38 7 1 0
61

Section 7. 

Recommendations to Evaluate Biotransformation of Chlorinated Solvents 

Based on the performance of PCR assays at the 

sites in this study that were undergoing natural attenuation of chlorinated solvents, the authors offer a 

number of recommendations for collecting samples 

for PCR assays and interpreting the data provided. 

The recommendations are summarized in Table 7.1 

and are discussed in detail in the remainder of this 

section.

Recommendations for Interpreting Data on 

Density of DNA in Ground Water

The OSWER Directive on MNA (U.S. EPA 1999) identifies three lines of evidence that can be used to support the selection of MNA as a remedy. The first line 

of evidence is historical monitoring data that provide a 

clear and meaningful trend of decreasing concentrations or contaminant mass over time. The second line 

of evidence is hydrogeologic and geochemical data 

that can be used to demonstrate indirectly the types 

of natural attenuation processes active at the site, 

and the rate at which such processes will reduce contaminant concentrations to required levels. The third 

line of evidence is data from the field or microcosm 

studies which directly demonstrate the occurrence 

of a particular natural attenuation process at the site 

and its ability to degrade the contaminant of concern. 

The presence of Dehalococcoides DNA in an aquifer 

can contribute to the third line of evidence.

As specified in the OSWER Directive (U.S. EPA 1999), 

unless EPA or the overseeing regulatory authority determines that historical data (the first line of evidence) 

are of sufficient quality and duration to support a decision to use MNA, data characterizing the nature and 

rates of natural attenuation processes at the site (the 

second line of evidence) should be provided. Where 

the latter are inadequate or inconclusive, data from 

microcosm studies (or genetic analysis, the third line 

of evidence) may also be necessary.

Data provided from analysis of DNA in water samples 

from wells are a semi-quantitative lower boundary 

on the density of organisms in the aquifer. The 

microorganisms may be attached to aquifer solids, 

and as a consequence, not adequately sampled by 

ground water from a monitoring well. As a practical 

matter, many evaluations of the distribution of Dehalococcoides DNA will be done with ground water 

samples from permanent wells. These evaluations 

done with samples of ground water will be subject 

to false negatives.

As a consequence, the density of Dehalococcoides cells in ground water does not provide direct 

evidence for a particular rate of biotransformation of 

chlorinated solvents. An assay for Dehalococcoides 

DNA in ground water does not provide the second 

line of evidence for natural attenuation. However, an 

assay for Dehalococcoides DNA in ground water can 

readily provide the third line of evidence for natural 

attenuation. 

Although the PCR assay for Dehalococcoides DNA 

in ground water can only provide the third line evidence, the assay has two desirable features. The 

assays are relatively inexpensive, and they can be 

preformed in a short period of time. The PCR assay 

can reduce uncertainty in the role and contribution 

of biological reductive dechlorination to monitored 

natural attenuation in plumes of chlorinated ethylenes 

in ground water.

 The strong possibility of false negatives for the presence of Dehalococcoides DNA makes it important 

that no interpretation be put on a failure to recover 

Dehalococcoides DNA from a water sample. The 

fact that Dehalococcoides DNA is not detected in a 

sample of well water does not mean that Dehalococcoides organisms are absent in the aquifer. This is 

particularly true if DNA corresponding to a universal 

bacterial primer is absent from the water sample. If 

the assay fails to detect Dehalococcoides DNA, it simply fails to contribute to the third line of evidence. 

If the third line of evidence is critical to accepting 

MNA as a remedy, then other means to provide the 

third line of evidence are necessary, or MNA should 

be rejected. If the third line of evidence is not critical, 
62

Table 7.1. Recommendations for Use of PCR Assays to Evaluate Biotransformation of Chlorinated Solvents

Concern Recommendation

Does PCR for Dehalococcoides

DNA provide the Second Line of 

Evidence for MNA?

PCR should not be expected to provide the Second Line of Evidence 

when applied to water samples from monitoring wells.

Does PCR for Dehalococcoides

DNA provide the Third Line of 

Evidence for MNA?

The presence of DNA from Dehalococcoides provides the Third Line 

of Evidence. However, the absence of DNA from Dehalococcoides

should not be interpreted as the absence of biological natural attenuation.

Can PCR for Dehalococcoides

DNA provide the Second Line of 

Evidence in the future?

It will be necessary to extract DNA from sediment samples, and 

compare data on density of Dehalococcoides cells to rates attained 

in benchmark field studies.

How should sites be “scored” to 

determine whether site characterization of biological processes is justified?

The scoring system in the EPA Technical Protocol to Evaluate Natural Attenuation at Chlorinated Solvents in Ground Water should be 

replaced with Equation 5.1 in Section 5 of this report.

How can limited PCR data from 

a few wells be extrapolated to 

other wells in a contaminated 

aquifer?

Rather than assume that PCR data from a few wells apply to an 

entire aquifer, use Equation 5.1 to estimate whether it is likely that 

Dehalococcoides is present in ground water from a particular well.

Which wells should have the 

highest priority for a PCR assay 

for Dehalococcoides?

Sample wells screened in material with high hydraulic conductivity 

compared to the rest of the aquifer, and wells with high concentrations of transformation products. Sample an equal number of wells in 

the source area, in the region with intermediate concentrations, and 

at the toe of the plume.

What precautions are 

needed to sample monitoring 

wells for a PCR assay for 

Dehalococcoides?

Avoid cross contamination. Use dedicated sampling tubing. The EPA 

low flow procedure is not optimal for sampling for Dehalococcoides. 

Pump as rapidly as possible.

What precautions are needed to 

collect water samples for a PCR 

assay for Dehalococcoides?

Collect one liter samples into plastic bottles that have never been 

used for another purpose. Collect spare samples from each well. 

Store on ice in the field before transportation to the laboratory. Protect samples from cross contamination during storage and transportation. Prepare shipping container to keep samples cool during transportation to the laboratory.

What controls are needed to 

document data quality?

Prepare field blanks and field duplicates as specified in the Quality 

Assurance Plan for the site. Include positive and negative controls 

for amplification of DNA in the PCR assay, and include a control for 

extraction of bacterial DNA from the samples. 
63

then the presence of Dehalococcoides DNA merely 

strengthens a decision that was based on the first two 

lines of evidence. The absence of Dehalococcoides

DNA fails to strengthen a decision that was made on 

the first two lines of evidence.

Whenever possible, aquifer solids should be sampled 

and assayed for Dehalococcoides DNA. This will 

reduce the chance of false negative results for the 

presence of Dehalococcoides DNA as determined 

by the qualitative assay using electrophoresis, or 

the density of Dehalococcoides cells as determined 

by the quantitative real time PCR assay. If aquifer 

solids are extracted and analyzed, there is at least 

a possibility that at some time in the future it will be 

possible to relate the density of active organisms as 

revealed by a PCR assay to the achieved rate of reductive dechlorination at field scale. At many sites, it 

is possible to recover core samples quickly and at low 

cost with push technology (e.g. GeoProbe® tools). At 

other sites, the cost of acquiring core samples may 

be prohibitively expensive. 

Recommendations for Interpreting 

Geochemistry of Ground Water

The scoring system in the Technical Protocol for Evaluating Natural Attenuation of Chlorinated Solvents in 

Ground Water (Wiedemeier et al., 1998) was offered 

as a screening mechanism to identify ground water 

where biological reductive dechlorination is likely to occur; however, the scoring system has been criticized. 

Equation 5.2 as discussed in Section 5 provides a 

simple and rapid way to calculate the probability that 

a specific dechlorinating organism, Dehalococcoides, 

is present at the site. The probability that Dehalococcoides DNA occurs in the ground water, as calculated 

by Equation 5.2, should replace the scoring system. 

However, a calculated probability that an organism 

exists in ground water is not equivalent to a PCR 

assay for its presence, and probabilities calculated 

from geochemical parameters should not be used to 

replace PCR assays. 

Often the computer models that are used to evaluate 

monitored natural attenuation are distributed parameter models. To properly calibrate these computer 

models, it is necessary to know the distribution of the 

capacity to transform chlorinated ethylenes throughout the aquifer. Data from the PCR assay may not 

be available from every well at a site. The calculated 

probability of Dehalococcoides organisms based on 

the geochemistry of ground water in a well could be 

used to assign rate constants to cells in a model. If 

Dehalococcoides organisms are expected, a rate 

constant characteristic of reductive dechlorination 

at that site would be assigned. If Dehalococcoides

organisms are not expected, the rate constant for 

reductive dechlorination would be set to zero in that 

particular region of the aquifer. 

Recommendations for Selecting Wells for 

Sampling 

At many chlorinated solvent sites, computer models 

have been used to describe the previous behavior of 

the plume and make future projections of its natural attenuation over time. Often these computer projections 

are an important part of the conceptual model of a site. 

Frequently, the calibration of the computer models will 

assume a uniform rate constant for biotransformation 

across the entire plume or major portions of the plume. 

The calibration of the model frequently assumes that 

the rate constant will be sustained into the future. If 

biotransformation carried out by Dehalococcoides

organisms is the primary process for natural attenuation, an assay for Dehalococcoides DNA can be used 

to test these assumptions about uniform distribution 

and sustainability of biotransformation. Dehalococcoides organisms should be uniformly present in those 

portions of the aquifer where the computer model 

projects a rate constant for biotransformation, and the 

populations of Dehalococcoides organisms should be 

sustained during long-term monitoring. 

At most sites, there is little value in sampling every 

well for analysis of Dehalococcoides DNA. Wells 

with relatively high hydraulic conductivity should be 

sampled from the more permeable portions of the 

aquifer. These are the portions of the aquifer with 

the greatest capacity to transport contaminated water 

and may be the portions that provide the most risk 

of impacting a receptor. Wells with higher hydraulic 

conductivity should be selected for assays for Dehalococcoides DNA. 

Wells with higher relative concentrations of transformation products provide circumstantial evidence that 

biotransformation has occurred at some point along 

the flow path from the source to the well. However, 

the high concentrations of transformation products 

do not prove that the transformation occurred in the 

portion of the aquifer immediately proximate to the 

monitoring well. Nonetheless, the wells with higher 

relative concentrations of vinyl chloride and ethylene 

should be sampled.

Wells should be sampled at various positions along the 

flow path from the source to the most down gradient 

wells containing detectable concentrations of chlorinated solvents. For the specific purpose of evaluating 

in situ biotransformation of chlorinated solvents, there 
64

end of the sampling tube at the center of the screened 

interval of the monitoring well. 

Water samples for analysis of Dehalococcoides DNA 

should be collected in a one liter plastic bottle constructed of polypropylene (e.g. Nalgene®) or high 

density polyethylene. The bottles should be new and 

should never have been used for another purpose. 

Use the bottles as supplied by the manufacturer. Do 

not attempt to clean the bottles before use. If the 

original shipping container has been opened, the 

empty bottles should be shipped to the field site with 

the lid firmly attached to the bottle. 

Fill the bottle in the field and screw the lid back on 

the bottle to make a tight seal. As far as possible, 

avoid exposing the water sample to the atmosphere. 

Fill the bottle with the fill tube at the bottom of the 

bottle. Do not allow the water sample to flow down 

the inside of the bottle. 

Label the sample with a permanent marker on a strip 

of labeling tape that entirely circles the bottle. Provide, 

at a minimum, the complete name of the well being 

sampled, the name of the facility or location where 

the well is installed, the date and time the sample 

was collected, and the name or initials of the person 

collecting the samples. If a chain of custody form is 

required, fill in the required information at the same 

time. Seal the sample bottle in a plastic bag with a 

ZipLock® closure or equivalent. Remove excess air 

from the bag before the closure is sealed. The purpose of the bag is to prevent cross contamination of 

the samples in case one of the sample bottles leaks 

during shipment. Place the bottles on water ice immediately after they are collected, labeled, and sealed 

in a plastic bag. Keep the bottles on ice until they are 

packaged for shipment for analysis. 

Collect a sample and a spare sample from each well 

sampled. Ship both samples for analysis. Identify 

which is the sample and which is the spare. The 

spare will be available to the laboratory analyst if 

there is any problem with preparation or analysis of 

the sample. If a field duplicate is desired, collect a 

third sample from the well and label the third sample 

as the field duplicate.

Prepare water to make a field blank by filtering distilled water or deionized water through a 0.22 micron 

filter into a sterile container. This is best done in the 

laboratory before going to the field. To prepare a trip 

blank, take the filtered water to the well head of the 

well at the site that is most likely to contain Dehalococcoides and pour the previously prepared water 

is little value in sampling background wells. Equation 

5.2 as discussed in Section 5 can be used to identify 

wells where the geochemical environment is favorable 

for Dehalococcoides species, and to select wells for 

a PCR assay for Dehalococcoides species. Sample 

an equal number of wells in the source area, in the 

region with intermediate concentrations, and in the 

distal portion of the plume where concentrations are 

within a factor of ten to one hundred of the Maximum 

Contaminant Level (MCL) or other relevant clean-up 

goals.

There is little value in sampling for Dehalococcoides

DNA in every round of sampling. A better strategy 

is to perform a comprehensive baseline assessment 

across a plume as part of the selection of MNA as 

a remedy, and perform a second comprehensive 

assessment before the performance of the remedy 

goes for review. The biogeochemical parameters, 

particularly concentrations of nitrate, methane, and 

ORP should be determined at the same time ground 

water is collected for determination of Dehalococcoides DNA. 

Recommendations for Sampling and 

Shipping of Samples

To avoid any cross-contamination, equipment to purge 

and sample ground water should not be moved from 

one well to another even if an attempt is made to 

sterilize or decontaminate the equipment. The PCR 

assay can amplify DNA, even though the organisms 

have been killed. Water samples should be obtained 

using a dedicated pump or pump tubing, freshly 

installed pump tubing, a disposable pump that has 

not been used on another well, or a new clean bailer 

and line. Neither the bailer nor the line should have 

been used on another well. If possible, avoid using 

bailers. If water is produced with a peristaltic pump at 

the well head, the peristaltic pump tubing and effluent 

tubing should be replaced immediately before a well 

is purged and sampled. A minimum of one casing 

volume should be purged before the sample is taken. 

It is better to purge the casing volume plus a volume 

equal to the porosity of the sand pack around the well 

screen, if one is present.

The EPA low-flow sampling protocol is designed to 

produce a sample that is free of turbidity and suspended solids. If Dehalococcoides cells are sorbed 

to sediment particles in the aquifer, they may not be 

sampled efficiently. Sediment and turbidity in the water 

sample do not interfere with the extraction of bacterial 

DNA. To increase the turbidity and suspended solids 

in the water sample, purge and sample the well at the 

fastest rate the pump will allow. Set the pump or the 
65

into a one liter plastic bottle just as would be done for 

any other sample. Seal the lid, label the bottle, seal 

in a plastic bag, store on water ice, and ship to the 

laboratory along with the other samples.

Collect and ship the number of trip blanks and field 

duplicates that are specified in the quality assurance 

project plan for the site.

Package the samples in a plastic cooler with an adequate number of bricks of frozen brine sealed in a 

plastic cover (Blue Ice® or equivalent) to keep the 

samples at or below 10° C for two days. Do not ship 

the samples with water ice. The samples should arrive at the laboratory within five days after they were 

originally collected in the field. If the field site is remote from the analytical laboratory, ship the samples 

by air freight for overnight delivery. Avoid shipping on 

a Friday or the day before a holiday if the receiving 

laboratory will not be open for business and not available to accept and properly store the samples.

The laboratory should provide information in their 

report on the procedures used to prepare the water 

samples for the polymerase chain assay, on the 

number and types of primers used, and the results 

with each primer. The laboratory should also provide information on the number of gene copies that 

would be required to provide the minimal density of 

Dehalococcoides DNA that can be detected by the 

assay. The minimum number of gene copies should 

be expressed in gene copies per liter of well water 

or gene copies per kilogram sediment, whichever is 

appropriate.

Dehalococcoides DNA may be absent in a water 

sample because bacteria of any kind were absent in 

the sample. To be able to determine if Dehalococcoides organisms are not present in the microbial community of the aquifer being sampled, it is necessary to 

include a primer for a gene that is essentially universal 

in bacteria in the polymerase chain reaction assay. 

The absence of bacterial DNA suggests that the microbial community in the aquifer was not effectively 

sampled, and that Dehalococcoides organisms may 

have been present but were not sampled. Positive 

and negative controls should be included in the PCR 

assay to ensure that the reaction is working properly 

and that contamination of reagents has not occurred. 

Amplicons (DNA amplified by the PCR assay using 

the Dehalococcoides primers) should be cloned and 

sequenced periodically to ensure that the assay is 

working as intended.

67

Section 8. 

Data Quality

Analysis of Chemical Concentrations 

Laboratory analyses for data presented in Table 4.2, 

Table 4.4, Table 4.6, Table 5.2, Table 5.3, Table 5.4, 

Table 5.5, Table 5.7, and Table 6.4 were conducted 

at the R.S. Kerr Environmental Research Center in 

accordance with a Quality Assurance Project Plan 

prepared for in-house task 3674 (Monitored Natural Attenuation of Chlorinated Solvents). Concentrations of 

chlorinated solvents, dissolved gases (methane, ethylene, ethane, and hydrogen), inorganic compounds 

(nitrate plus nitrite, sulfate, and chloride), fatty acids 

(propionate and acetate), and total organic carbon 

(TOC) were determined following in-house Standard 

Operating Procedures (SOPs). Chlorinated solvents 

were analyzed by automated headspace gas chromatography and mass spectrometry. Methane, ethylene, 

and ethane were analyzed by gas chromatography 

with a thermal conductivity detector. Hydrogen was 

analyzed by RGA3 Reduction Gas Analyzer with a 

reduction gas detector. Nitrate plus nitrite was analyzed by Lachat flow injection analysis. Sulfate and 

chloride were analyzed by waters capillary electrophoresis. Total Organic Carbon (TOC) was analyzed by 

Dohrman DC-80 Carbon Analyzer. Propionate and 

acetate were analyzed by high performance liquid 

chromatography. 

Major quality assurance and quality control (QA/QC) 

evaluations for the analyses included method blank 

(MB), continuing calibration check (CCC), second 

source check (QC) using a sample obtained from 

the second source as identified by their designated 

names, laboratory duplicates (LD), and matrix spike 

(MS). Method blank was analyzed in the beginning 

and end of sample set. Calibration check standards 

were analyzed every ten samples as well as in the 

beginning and end of sample set. QC checks were 

analyzed every ten samples. Lab duplicates were 

analyzed every ten samples. Matrix spikes were 

analyzed every twenty samples. The data quality 

objectives were as follows: The target analyte in the 

method blank would be below method detection limit. 

The reported concentration of continuing calibration 

check standard, QC check standard, and matrix spike 

would agree with the expected concentration plus or 

minus 20% of the known concentration (i.e., recovery 

of the expected value would be in the range of 80-

120%). Laboratory duplicates would agree with each 

other plus or minus 20%. All the samples were held 

less than thirty days before analysis. 

Table 8.1 summarizes typical data quality for TCE in 

the ground water samples and in the water samples 

from the enrichment cultures. Two out of 38 of the 

calibration check standards did not meet the goal of 

± 20% of the nominal values (One calibration check 

analyzed on 4/2/03 was reported as 122% of the 

nominal value, and one calibration check analyzed on 

6/26/03 was reported as 69% of the nominal value.) 

One out of 43 of the QC check standards did not 

meet the goal of ± 20% of the nominal values (One 

QC check analyzed on 4/2/03 was reported as 126% 

of the nominal value.) One out of 36 laboratory duplicates did not agree within 20% (The relative percent 

difference for the duplicates analyzed on 4/2/03 was 

21%.) Four out of 50 method blanks had TCE concentration above method detection limit (The values 

were 0.99 and 0.675 on 4/2/03, and 1.03 and 0.89 on 

4/30/03.) All 25 matrix spikes met the goal of ± 20% 

of the expected values. 

Several quality controls did not meet the criteria on 

date 4/2/03 in the beginning of the analysis. The instrument was recalibrated, and the subsequent checks 

met the criteria. All the data for TCE were determined 

to be of acceptable quality, and the data were used 

in the report. 

Table 8.2 summarizes typical data quality for cisDCE in the ground water samples and in the water 

samples from the enrichment cultures. Five out of 38 

of the calibration check standards did not meet the 

goal of ± 20% of the nominal values. (Two calibration 

checks analyzed on 3/9/03 were reported as 127% 

and 129% of the nominal values, one calibration 

check on 4/30/03 was reported as 136% the nominal value, one calibration check analyzed on 6/26/03 

was reported as 78% of the nominal value, and one 
68

calibration check analyzed on 1/20/04 was 123% of 

the nominal value.) 

Four out of 43 of the QC check standards did not meet 

the goal of ± 20% of the nominal values (One QA 

check analyzed on 4/10/03 was reported as 129% of 

the nominal value, one QA check analyzed on 4/30/03 

was reported as 153% of the nominal value, and two 

QA checks analyzed on 2/20/04 were reported as 

169% and 145% of the nominal values.) All other 

quality controls met the objectives, including all 36 

duplicates, which agreed with each other within 20% 

difference, all 50 method blanks did not have detectable cis-DCE, and all 25 matrix spikes met the goal 

of ± 20% of the expected values. 

The chromatographic response for cis-DCE in the two 

QA checks analyzed on 2/20/04 showed evidence 

of peak asymmetry suggesting coelution of another 

compound. The previous and subsequent recoveries 

for cis-DCE in the continuing calibration checks met 

the data quality objectives. On 4/30/03, the reported 

values for cis-DCE might be slightly elevated over 

the true values, as shown in the continuing calibration checks and QC checks. Not all of the calibration 

checks met the goal of ± 20% of the nominal values. 

When the calibration checks did not meet the goal, 

they were still within 36% of the nominal values. An 

error of 36% would not change the interpretation 

placed on the data. Therefore, all the data for cis-DCE 

were determined to be of acceptable quality, and the 

data were used in the report. 

Table 8.3 summarizes typical data quality for vinyl 

chloride in the ground water samples and in the water 

samples from the enrichment cultures. Compared 

to TCE and cis-DCE, vinyl chloride is more volatile 

and more easily lost, resulting in more missed data 

quality objectives. Three out of 38 of the calibration 

check standards did not meet the goal of ± 20% of the 

nominal values (One calibration check analyzed on 

3/9/03 was reported as 78% of the nominal value, one 

calibration check analyzed on 3/30/03 was reported 

as 76% of the nominal value, and one calibration 

check analyzed on 6/5/03 was reported as 131% of 

the nominal value.) Four out of 43 of the QC check 

standards did not meet the goal of ± 20% of the 

nominal values (One QC check analyzed on 4/10/03 

was reported as 77% of the nominal value, one QC 

check analyzed on 6/26/03 was reported as 49% of 

the nominal value, and two QC checks analyzed on 

10/28/03 were reported as 51% and 62% of the nominal values.) Three out of 36 laboratory duplicates did 

not agree within 20% (The relative percent differences 

were 43.5% for the duplicates analyzed on 3/21/03, 

84.1% for duplicates analyzed on 6/26/03, and 42.9% 

for duplicates analyzed on 6/26/03.) One out of 50 

method blanks had vinyl chloride concentration above 

method detection limit. Five out of 25 matrix spikes 

did not meet the goal of ± 20% of the expected values 

(See Table 8.3.). 

Most of the missed data quality objectives for vinyl 

chloride were due to the problem of preparing the 

check standards, not the problem of the instrument. 

Subsequent samples using the analysis applied a 

modified standard preparation technique to minimize 

vinyl chloride losses. All the data for vinyl chloride 

were determined to be of acceptable quality, and the 

data were used in the report. 

If there is no data provided for calibration check 

controls (CCCs) in Tables 8.1, 8.2, and 8.3, that information was not included in the report provided by 

the analyst.

Table 8.4 summarizes typical data quality for ethylene 

in the ground water samples and in the gas phase 

of the enrichment cultures. All 98 calibration checks 

met the goal of ± 20% of the nominal values. All 24 

method blanks did not have detectable ethylene. All 

16 laboratory duplicates agreed with each other within 

20% difference. All seven matrix spikes met the goal 

of ± 20% of the expected values. 

All the data for ethylene were determined to be of 

acceptable quality, and the data were used in the 

report. 

Table 8.5 summarizes typical data quality for methane 

in the ground water samples and in the gas phase of 

the enrichment cultures. All 119 calibration checks 

met the goal of ± 20% of the nominal values. All 24 

method blanks did not have detectable methane. All 

16 laboratory duplicates agreed within 20% except 

the duplicates analyzed on 8/25/03 where the relative 

percent difference was 23.0%. All seven matrix spikes 

met the goal of ± 20% of the expected values. 

All the data for methane were determined to be of 

acceptable quality, and the data were used in the 

report. 

Table 8.6 summarizes typical data quality for hydrogen in the gas samples stripped from ground water 

and in the gas phase of the enrichment cultures. All 

106 calibration checks met the goal of ± 20% of the 

nominal values. Two out of 22 method blanks had 
69

hydrogen concentration above detection limit. Three 

out of 12 laboratory duplicates did not agree within 

20% (See Table 8.6.). 

All the data for hydrogen were determined to be of 

acceptable quality, and the data were used in the 

report. 

Table 8.7 summarizes typical data quality for nitrate 

plus nitrite nitrogen in the ground water samples. 

All 26 calibration checks met the goal of 20% of the 

nominal values. All seven QC checks met the goal 

of ± 20% of the nominal values. All ten laboratory 

duplicates agreed with each other within 20% difference. All 16 method blanks had nitrate plus nitrite 

nitrogen concentration below quantification limit. All 

ten matrix spikes met the goal of ± 20% of the expected values. 

All the data for nitrate plus nitrite nitrogen were determined to be of acceptable quality, and the data were 

used in the report. 

Analysis of DNA Concentrations 

The analyses of Dehalococcoides DNA were performed by SiREM (Guelph, Ontario) using their 

Gene-Trac® Test. The QA/QC was maintained by 

implementing clean techniques and control PCR 

reactions. 

DNA was extracted using a single-use sterile filter unit 

and single-use DNA extraction kit. Prior to PCR, all 

micropipettes and other equipments used in setting 

up reactions were swabbed with DNA AWAYTM, and 

10% bleach or 70% ethanol to ensure cleanliness and 

sterility. PCR reaction mixtures were assembled in a 

Forma HEPA flow cabinet to prevent the introduction 

of particles and bacteria/DNA that might produce 

false positives. 

Three types of control reactions were used in the 

Gene-Trac procedure: a negative control, a positive 

control, and a DNA extraction control. The controls 

were conducted and interpreted by SiREM, the 

vendor for the PCR assays. The negative control 

involved processing sterile water through the same 

DNA extraction procedure as the sample. It ensured 

that contamination of samples did not occur via the 

DNA extraction process, PCR setup, or performance 

of the reactions. In the data reported to EPA, a 

sample was flagged when DNA was amplified in the 

negative control. There was no DNA amplified in the 

negative control in the data reported in Tables 4.5, 

4.8, or 5.4. The positive control consisted of a PCR 

assay containing a cloned Dehalococcoides 16S rRNA 

gene. It ensured that all reagents and equipment were 

performing properly. If the PCR procedure failed to 

amplify the Dehalococcoides 16S rRNA gene, the 

data were discarded by SiREM and were not reported 

to U.S. EPA. The problem with the PCR procedure 

was corrected by SiREM, and the DNA extracted 

from the ground water sample was assayed and 

reported. The DNA extraction control was performed 

when the Dehalococcoides test was negative. In the 

DNA extraction control, PCR was performed on the 

sample using a universal bacterial PCR primer set. 

It was used to determine whether bacterial DNA was 

present and extracted from the sample. The results of 

the DNA extraction control are provided in Tables 4.5 

and 4.8, and are interpreted in Section 4. PCR data 

from the Dehalococcoides primers from three wells 

sampled at the Area 2500 Site at the former England 

AFB, Louisiana, were discarded because the bacterial 

DNA primer was not detected. 
70

Table 8.1. Typical Quality Performance Data for Analysis of TCE in Water. All Values are μg/L Unless Otherwise 

Indicated

Date Collected 1/24/03 2/4/03

Date Analyzed 11/7/02 1/7/03 2/11/03 3/6/03

CCC Standard Nominal 10

CCC Standard Measured 10.1

Percent of Check Standard 101%

CCC Standard Nominal 100

CCC Standard Measured 96

Percent of Check Standard 96%

CCC Standard Nominal 250

CCC Standard Measured 235

Percent of Check Standard 94%

QC Standard Nominal 20 20 50 20 20

QC Standard Measured 19.3 22.1 51.0 21.1 21.3

Percent of Check Standard 96.4% 111% 107% 106% 107%

QC Standard Nominal 200 200 50 200 200

QC Standard Measured 189 219 48.1 225 194

Percent of Check Standard 94.4% 110% 96.2% 113% 97%

Blank 1 <0.5 <0.23 <0.39 <0.3 <0.23

Blank 2 <0.5 <0.23 <0.39 <0.3 <0.23

Sample Analysis 1 80.5 2180 227 263 <0.23

Laboratory Duplicate 1 75.2 2130 225 235 <0.23

Relative Percent Difference 6.8% 2.3% 0.9% 11.2% -

Sample Analysis 2 231

Laboratory Duplicate 2 240

Relative Percent Difference 3.8%

Spike Concentration 1 200 200 50 200 200

Sample Concentration 1 <0.5 28.4 <0.39 215 10.4

Spike Recovery (Percent) 97% 108% 83.6% 100% 105%

Spike Concentration 2 50

Sample Concentration 2 232

Spike Recovery (Percent) 87.2%

Date Collected 2/24/03 3/6/03 3/11/03 3/26/03 4/8/03

Date Analyzed 3/9/03 4/2/03 3/21/03 4/10/03 4/30/03

CCC Standard Nominal 50 10 10 10

CCC Standard Measured 44.2 12.2 10.6 11.2

Percent of Check Standard 88% 122% 106% 112%

CCC Standard Nominal 200 100 250 50

CCC Standard Measured 168 113 259 45

Percent of Check Standard 84% 113% 104% 90%

CCC: Continuing Calibration Check; QC: Second Source Check
71

CCC Standard Nominal 50 10 250 100

CCC Standard Measured 42 8.44 253 108

Percent of Check Standard 84% 84.4% 101% 108%

QC Standard Nominal 20 25 10 100 50

QC Standard Measured 23.4 26.8 9.7 111 58.1

Percent of Check Standard 117% 107% 97% 111% 116%

QC Standard Nominal 200 50 200

QC Standard Measured 211 63.0 202

Percent of Check Standard 105% 126% 101%

Blank 1 <0.39 0.99 <0.07 <0.39 1.03

Blank 2 <0.39 0.675 <0.07 <0.39 0.89

Sample Analysis 1 204 63.9 96.7 158 <0.39

Laboratory Duplicate 1 203 51.7 102 157 <0.39

Relative Percent Difference 0.5% 21.1% 5.3% 0.6% -

Sample Analysis 2 212 160

Laboratory Duplicate 2 192 156

Relative Percent Difference 9.9% 2.5%

Spike Concentration 1 200 100 100

Sample Concentration 1 234 36.4 37.2

Spike Recovery (Percent) 100% 81% 95%

Spike Concentration 2

Sample Concentration 2

Spike Recovery (Percent)

Date Collected 4/21/03 4/29/03 5/16/03 6/23/03 7/23/03

Date Analyzed 5/20/03 5/5/03 6/5/03 6/26/03 8/5/03

CCC Standard Nominal 10 25

CCC Standard Measured 11.4 24

Percent of Check Standard 114% 96%

CCC Standard Nominal 50 100

CCC Standard Measured 51.2 93.9

Percent of Check Standard 102% 93.9%

CCC Standard Nominal 100 250

CCC Standard Measured 98 173

Percent of Check Standard 98% 69.2%

QC Standard Nominal 50 20 20 50 20

QC Standard Measured 52.3 21.3 20.3 44.8 22.3

Percent of Check Standard 105% 106% 102% 89.6% 112%

QC Standard Nominal 200 50 200

CCC: Continuing Calibration Check; QC: Second Source Check

Table 8.1. Typical Quality Performance Data for Analysis of TCE in Water. All Values are μg/L Unless Otherwise 

Indicated continued
72

QC Standard Measured 206 48.4 218

Percent of Check Standard 103% 97% 112%

Blank 1 <0.39 <0.23 <0.07 <0.39 <0.23

Blank 2 <0.39 <0.23 <0.07 <0.39 <0.23

Sample Analysis 1 344 <0.23 185 241 <0.23

Laboratory Duplicate 1 348 <0.23 182 231 <0.23

Relative Percent Difference 1.2% - 1.6% 4.2% -

Sample Analysis 2 182 188

Laboratory Duplicate 2 179 202

Relative Percent Difference 1.7% 7.2%

Spike Concentration 1 200 200 100

Sample Concentration 1 <0.23 190 247

Spike Recovery (Percent) 102% 91% 84%

Date Collected 8/7/03 8/20/03 10/3/03 10/7/03 10/24/03

Date Analyzed 8/12/03 8/22/03 10/22/03 11/3/03 10/28/03

Spike Concentration 2 200

Sample Concentration 2 168

Spike Recovery (Percent) 84%

CCC Standard Nominal 500 100 25

CCC Standard Measured 547 98.6 24.5

Percent of Check Standard 109% 98.6% 98%

CCC Standard Nominal 50

CCC Standard Measured 49.9

Percent of Check Standard 99.8%

CCC Standard Nominal 100

CCC Standard Measured 97.6

Percent of Check Standard 97.6%

QC Standard Nominal 20 20 20 100 5

QC Standard Measured 19.3 20.5 21 99.6 5.48

Percent of Check Standard 96% 103% 105% 100% 110%

QC Standard Nominal 200 200 200 50

QC Standard Measured 214 210 178 46.4

Percent of Check Standard 107% 105% 89% 92.8%

Blank 1 <0.23 <0.23 <0.07 <0.07 <0.39

Blank 2 <0.23 <0.23 <0.07 <0.07 <0.39

Sample Analysis 1 127 1410 155 60.0 147

Laboratory Duplicate 1 119 1370 155 57.0 143

Relative Percent Difference 6.5% 2.9% 0.0% 5.1% 2.8%

Sample Analysis 2 106 186 150

Laboratory Duplicate 2 105 176 151

CCC: Continuing Calibration Check; QC: Second Source Check

Table 8.1. Typical Quality Performance Data for Analysis of TCE in Water. All Values are μg/L Unless Otherwise 

Indicated continued
73

Relative Percent Difference 0.9% 5.5% 0.7%

Spike Concentration 1 200 200 100 100

Sample Concentration 1 111 9.3 163 59.3

Spike Recovery (Percent) 102% 106% 91% 94.5%

Spike Concentration 2 200 200

Sample Concentration 2 108 3.3

Spike Recovery (Percent) 105% 93%

Date Collected 1/14/04 2/19/04 6/4/04 9/7/04 7/14/05

Date Analyzed 1/20/04 2/20/04 6/17/04 9/17/04 7/15/05

CCC Standard Nominal 10 10 20 20 20

CCC Standard Measured 8.03 10.8 19.9 20.6 17.8

Percent of Check Standard 80% 108% 100% 103% 89%

CCC Standard Nominal 100 50 20 200

CCC Standard Measured 92.5 50.4 20 193

Percent of Check Standard 93% 101% 100% 96%

CCC Standard Nominal 100 100 20

CCC Standard Measured 92.8 103 21

Percent of Check Standard 93% 103% 105%

QC Standard Nominal 50 10 20 20

QC Standard Measured 46.3 11.0 22.7 21.4

Percent of Check Standard 93% 110% 114% 107%

QC Standard Nominal 100 100 200

QC Standard Measured 96.0 99.1 200

Percent of Check Standard 96% 99% 100%

Blank 1 <0.39 <0.39 <0.23 <0.23 <0.23

Blank 2 <0.39 <0.39 <0.23 <0.23 <0.23

Sample Analysis 1 1.20 80.8 149 148

Laboratory Duplicate 1 1.17 83.1 148 139

Relative Percent Difference 2.5% 2.8% 0.7% 6.3%

Sample Analysis 2 91.5 110 152 72.6

Laboratory Duplicate 2 93.2 108 154 63.8

Relative Percent Difference 1.8% 1.8% 1.3% 12.9%

Spike Concentration 1 100 100 100

Sample Concentration 1 <0.23 159 62.9

Spike Recovery (Percent) 105% 105% 95%

Spike Concentration 2 100 100 100

Sample Concentration 2 134 132 117

Spike Recovery (Percent) 99% 103% 89%

CCC: Continuing Calibration Check; QC: Second Source Check

Table 8.1. Typical Quality Performance Data for Analysis of TCE in Water. All Values are μg/L Unless Otherwise 

Indicated continued
74

Table 8.2. Typical Quality Performance Data for Analysis of cis-DCE in Water. All Values are μg/L Unless Otherwise 

Indicated

Date Collected 10/23/02 11/25/02 12/23/02 1/24/03 2/4/03

Date Analyzed 11/7/02 12/19/02 1/7/03 2/11/03 3/6/03

CCC Standard Nominal 10

CCC Standard Measured 8.71

Percent of Check Standard 87.1%

CCC Standard Nominal 100

CCC Standard Measured 100

Percent of Check Standard 100%

CCC Standard Nominal 250

CCC Standard Measured 233

Percent of Check Standard 93.2%

QC Standard Nominal 20 20 50 20 20

QC Standard Measured 21.1 21.9 54.8 21.3 21.6

Percent of Check Standard 106% 110% 110% 107% 108%

QC Standard Nominal 200 200 50 200 200

QC Standard Measured 187 216 51.1 219 206

Percent of Check Standard 93.6% 108% 102% 110% 103%

Blank 1 <0.2 <0.2 <0.36 <0.2 <0.2

Blank 2 <0.2 <0.2 <0.36 <0.2 <0.2

Sample Analysis 1 79.3 57.1 111 0.48 <0.2

Laboratory Duplicate 1 75.7 54.8 110 0.48 <0.2

Relative Percent Difference 4.6% 4.1% 0.9% 0.0% -

Sample Analysis 2 <0.2

Laboratory Duplicate 2 <0.2

Relative Percent Difference -

Spike Concentration 1 200 200 50 200 200

Sample Concentration 1 <0.2 28400 <0.36 0.48 72

Spike Recovery (Percent) 101% 108% 95% 105% 104%

Spike Concentration 2 50

Sample Concentration 2 <0.36

Spike Recovery (Percent) 106%

Date Collected 2/24/03 3/6/03 3/11/03 3/26/03 4/8/03

Date Analyzed 3/9/03 4/2/03 3/21/03 4/10/03 4/30/03

CCC Standard Nominal 50 10 10 10

CCC Standard Measured 48 9.58 10.6 11.0

Percent of Check Standard 95% 95.8% 106% 110%

CCC Standard Nominal 200 100 250 50

CCC Standard Measured 254 105 258 68.2

Percent of Check Standard 127% 105% 103% 136%

CCC: Continuing Calibration Check; QC: Second Source Check
75

CCC Standard Nominal 50 100 250 100

CCC Standard Measured 64 109 260 117

Percent of Check Standard 129% 109% 104% 117%

QC Standard Nominal 20 25 10 100 50

QC Standard Measured 22.1 24.4 9.4 129 76.3

Percent of Check Standard 110% 97.6% 94% 129% 153%

QC Standard Nominal 200 50 200

QC Standard Measured 211 57.7 185

Percent of Check Standard 105% 115% 92%

Blank 1 <0.36 <0.14 <0.06 <0.14 <0.14

Blank 2 <0.36 <0.14 <0.06 <0.14 <0.14

Sample Analysis 1 <0.36 249 56.7 <0.14 <0.14

Laboratory Duplicate 1 <0.36 <0.14 57.7 <0.14 <0.14

Relative Percent Difference - - 1.7% - -

Sample Analysis 2 <0.36 0.42

Laboratory Duplicate 2 <0.36 0.46

Relative Percent Difference - 9.1%

Spike Concentration 1 200 100 100

Sample Concentration 1 <0.36 40.7 18.8

Spike Recovery (Percent) 109% 88% 95%

Spike Concentration 2

Sample Concentration 2

Spike Recovery (Percent)

Date Collected 4/21/03 4/29/03 5/16/03 6/23/03 7/23/03

Date Analyzed 5/20/03 5/5/03 6/5/03 6/26/03 8/5/03

CCC Standard Nominal 10 25

CCC Standard Measured 10.3 19.5

Percent of Check Standard 103% 78%

CCC Standard Nominal 50 100

CCC Standard Measured 51.1 91.8

Percent of Check Standard 102% 91.8%

CCC Standard Nominal 100 250

CCC Standard Measured 102 264

Percent of Check Standard 102% 106%

QC Standard Nominal 50 20 20 50 20

QC Standard Measured 51.8 18.7 19.7 46.6 21

Percent of Check Standard 104% 93% 99% 93.2% 105%

QC Standard Nominal 200 50 200

QC Standard Measured 195 46.6 206

CCC: Continuing Calibration Check; QC: Second Source Check

Table 8.2. Typical Quality Performance Data for Analysis of cis-DCE in Water. All Values are μg/L Unless Otherwise 

Indicated continued
76

Percent of Check Standard 98% 93% 103%

Blank 1 <0.14 <0.2 <0.06 <0.14 <0.2

Blank 2 <0.14 <0.2 <0.06 <0.14 <0.2

Sample Analysis 1 17.5 <0.2 0.2 <0.14 <0.2

Laboratory Duplicate 1 19.4 <0.2 0.2 <0.14 <0.2

Relative Percent Difference 10.3% - 0.0% - -

Sample Analysis 2 1.2 140

Laboratory Duplicate 2 1.2 105

Relative Percent Difference 0.0% 28.6%

Spike Concentration 1 200 200 100

Sample Concentration 1 0.34 <0.06 12.9

Spike Recovery (Percent) 104% 94% 103%

Spike Concentration 2 200

Sample Concentration 2 <0.06

Spike Recovery (Percent) 87%

Date Collected 8/7/03 8/20/03 10/3/03 10/7/03 10/24/03

Date Analyzed 8/12/03 8/22/03 10/22/03 11/3/03 10/28/03

CCC Standard Nominal 500 100 25

CCC Standard Measured 518 93.7 21.8

Percent of Check Standard 104% 93.7% 87.2%

CCC Standard Nominal 50

CCC Standard Measured 51

Percent of Check Standard 102%

CCC Standard Nominal 100

CCC Standard Measured 95.6

Percent of Check Standard 95.6%

QC Standard Nominal 20 20 20 100 5

QC Standard Measured 20.4 20.8 20.2 96.2 5.59

Percent of Check Standard 102% 104% 101% 96% 112%

QC Standard Nominal 200 200 200 50

QC Standard Measured 204 200 191 46.3

Percent of Check Standard 102% 100% 95% 92.6%

Blank 1 <0.2 <0.2 <0.06 <0.06 <0.14

Blank 2 <0.2 <0.2 <0.06 <0.06 <0.14

Sample Analysis 1 <0.2 99.7 0.18 127 <0.14

Laboratory Duplicate 1 <0.2 99.0 0.18 122 <0.14

Relative Percent Difference - 0.7% 0.0% 4.0% -

Sample Analysis 2 <0.2 16.6 1.62

CCC: Continuing Calibration Check; QC: Second Source Check

Table 8.2. Typical Quality Performance Data for Analysis of cis-DCE in Water. All Values are μg/L Unless Otherwise 

Indicated continued
77

Laboratory Duplicate 2 <0.2 15.7 1.58

Relative Percent Difference - 5.6% 2.5%

Spike Concentration 1 200 200 100 100

Sample Concentration 1 <0.2 250 <0.06 0.2

Spike Recovery (Percent) 103% 97% 94% 88%

Spike Concentration 2 200 200

Sample Concentration 2 <0.2 1.2

Spike Recovery (Percent) 100% 91%

Date Collected 1/14/04 2/19/04 6/4/04 9/7/04 7/14/05

Date Analyzed 1/20/04 2/20/04 6/17/04 9/17/04 7/15/05

CCC Standard Nominal 10 10 20 20 20

CCC Standard Measured 12.3 11.5 20.7 19.4 20.2

Percent of Check Standard 123% 115% 103% 97% 101%

CCC Standard Nominal 100 50 20 200

CCC Standard Measured 104 57.7 19.8 210

Percent of Check Standard 104% 115% 99% 105%

CCC Standard Nominal 100 100 20

CCC Standard Measured 113 112 19.3

Percent of Check Standard 113% 112% 97%

QC Standard Nominal 50 10 20 20

QC Standard Measured 46.9 16.9 20.8 21.1

Percent of Check Standard 94% 169% 104% 105%

QC Standard Nominal 100 100 200 200

QC Standard Measured 95.0 145 217 207

Percent of Check Standard 95% 145% 109% 103%

Blank 1 <0.14 <0.14 <0.2 <0.2 <0.2

Blank 2 <0.14 <0.14 <0.2 <0.2 <0.2

Sample Analysis 1 <0.14 <0.2 <0.2 0.73

Laboratory Duplicate 1 <0.14 <0.2 <0.2 0.73

Relative Percent Difference - - - 0.0%

Sample Analysis 2 <0.14 <0.2 <0.2 <0.2

Laboratory Duplicate 2 <0.14 <0.2 <0.2 <0.2

Relative Percent Difference - - - -

Spike Concentration 1 100 100 100

Sample Concentration 1 <0.2 <0.2 1.31

Spike Recovery (Percent) 107% 105% 94%

Spike Concentration 2 100 100 100

Sample Concentration 2 <0.2 <0.2 <0.2

Spike Recovery (Percent) 105% 103% 91%

CCC: Continuing Calibration Check; QC: Second Source Check

Table 8.2. Typical Quality Performance Data for Analysis of cis-DCE in Water. All Values are μg/L Unless Otherwise 

Indicated continued
78

Table 8.3. Typical Quality Performance Data for Analysis of Vinyl Chloride in Water. All Values are μg/L Unless 

Otherwise Indicated

Date Collected 10/23/02 11/25/02 12/23/02 1/24/03 2/4/03

Date Analyzed 11/7/02 12/19/02 1/7/03 2/11/03 3/6/03

CCC Standard Nominal 10

CCC Standard Measured 8.78

Percent of Check Standard 88%

CCC Standard Nominal 100

CCC Standard Measured 90.5

Percent of Check Standard 91%

CCC Standard Nominal 250

CCC Standard Measured 219

Percent of Check Standard 88%

QC Standard Nominal 20 20 50 20 20

QC Standard Measured 20.1 21.6 50.8 21.3 20.8

Percent of Check Standard 101% 108% 102% 107% 104%

QC Standard Nominal 200 200 50 200 200

QC Standard Measured 180 208 44.3 211 196

Percent of Check Standard 90.1% 104% 88.6% 106% 98%

Blank 1 <0.5 <0.3 <0.22 <0.3 <0.3

Blank 2 <0.5 <0.3 <0.22 <0.3 <0.3

Sample Analysis 1 12 0.47 78.9 <0.3 <0.3

Laboratory Duplicate 1 9.93 <0.3 92.0 <0.3 <0.3

Relative Percent Difference 18.9% - 15.3% - -

Sample Analysis 2 <0.3

Laboratory Duplicate 2 <0.3

Relative Percent Difference -

Spike Concentration 1 200 200 50 200 200

Sample Concentration 1 <0.5 20400 <0.22 <0.3 3.69

Spike Recovery (Percent) 98% 87% 50% 101% 103%

Spike Concentration 2 50

Sample Concentration 2 <0.22

Spike Recovery (Percent) 56%

Date Collected 2/24/03 3/6/03 3/11/03 3/26/03 4/8/03

Date Analyzed 3/9/03 4/2/03 3/21/03 4/10/03 4/30/03

CCC Standard Nominal 50 10 10 10

CCC Standard Measured 41 10.9 8.89 7.62

Percent of Check Standard 82% 109% 89% 76.2%

CCC Standard Nominal 200 100 250 50

CCC Standard Measured 156 113 288 42.3

Percent of Check Standard 78% 113% 115% 84.6%

CCC: Continuing Calibration Check; QC: Second Source Check
79

CCC Standard Nominal 50 100 250 100

CCC Standard Measured 40 113 236 87.9

Percent of Check Standard 80% 113% 94% 87.9%

QC Standard Nominal 20 25 10 100 50

QC Standard Measured 23.3 25.5 9.0 76.7 55.6

Percent of Check Standard 117% 102% 90% 77% 111%

QC Standard Nominal 200 50 200

QC Standard Measured 192 58.4 180

Percent of Check Standard 96% 117% 90%

Blank 1 <0.22 <0.22 <0.49 <0.22 <0.22

Blank 2 <0.22 <0.22 <0.49 <0.22 <0.22

Sample Analysis 1 <0.22 1290 8.4 0.26 0.41

Laboratory Duplicate 1 <0.22 1290 5.4 0.26 0.38

Relative Percent Difference - 0.0% 43.5% 0.0% 7.6%

Sample Analysis 2 <0.22 <0.22

Laboratory Duplicate 2 <0.22 <0.22

Relative Percent Difference - -

Spike Concentration 1 200 100

Sample Concentration 1 <0.22 3.3

Spike Recovery (Percent) 103% 125%

Spike Concentration 2 100

Sample Concentration 2 20.3

Spike Recovery (Percent) 81%

Date Collected 4/21/03 4/29/03 5/16/03 6/23/03 7/23/03

Date Analyzed 5/20/03 5/5/03 6/5/03 6/26/03 8/5/03

CCC Standard Nominal 10 20 25

CCC Standard Measured 10 23.5 20.4

Percent of Check Standard 100% 118% 81.6%

CCC Standard Nominal 50 50 100

CCC Standard Measured 44.4 65.9 93.5

Percent of Check Standard 88% 131% 93.5%

CCC Standard Nominal 100 250

CCC Standard Measured 98.0 252

Percent of Check Standard 98% 101%

QC Standard Nominal 50 20 50 20

QC Standard Measured 56.7 22.3 24.4 21.2

Percent of Check Standard 113% 111% 49% 106%

QC Standard Nominal 200 200

CCC: Continuing Calibration Check; QC: Second Source Check

Table 8.3. Typical Quality Performance Data for Analysis of Vinyl Chloride in Water. All Values are μg/L Unless 

Otherwise Indicated continued
80

QC Standard Measured 220 227

Percent of Check Standard 110% 113%

Blank 1 <0.22 <0.3 <0.49 <0.22 <0.3

Blank 2 1.37 <0.3 <0.49 <0.22 <0.3

Sample Analysis 1 5.28 <0.3 <0.49 0.22 <0.3

Laboratory Duplicate 1 4.56 <0.3 <0.49 0.34 <0.3

Relative Percent Difference 14.6% - - 42.9% -

Sample Analysis 2 <0.49 32.1

Laboratory Duplicate 2 <0.49 78.7

Relative Percent Difference - 84.1%

Spike Concentration 1 200 200 100

Sample Concentration 1 1.59 <0.49 <0.3

Spike Recovery (Percent) 101% 144% 113%

Spike Concentration 2 200

Sample Concentration 2 <0.49

Spike Recovery (Percent) 141%

Date Collected 8/7/03 8/20/03 10/3/03 10/7/03 10/24/03

Date Analyzed 8/12/03 8/22/03 10/22/03 11/3/03 10/28/03

CCC Standard Nominal 500 100 50

CCC Standard Measured 470 89.4 41.7

Percent of Check Standard 94% 89.4% 83.4%

CCC Standard Nominal 

CCC Standard Measured

Percent of Check Standard

CCC Standard Nominal 

CCC Standard Measured 

Percent of Check Standard

QC Standard Nominal 20 20 20 100 5

QC Standard Measured 22.1 22.9 22.7 117 2.57

Percent of Check Standard 110% 115% 113% 117% 51%

QC Standard Nominal 200 200 200 50

QC Standard Measured 212 233 183 30.3

Percent of Check Standard 106% 116% 92% 62%

Blank 1 <0.3 <0.3 <0.49 <0.49 <0.22

Blank 2 <0.3 <0.3 <0.49 <0.49 <0.22

Sample Analysis 1 <0.3 0.42 <0.49 97 <0.22

Laboratory Duplicate 1 <0.3 0.35 <0.49 91.2 <0.22

Relative Percent Difference - 18.2% - 6.2% -

Sample Analysis 2 <0.3 <0.49 <0.22

CCC: Continuing Calibration Check; QC: Second Source Check

Table 8.3. Typical Quality Performance Data for Analysis of Vinyl Chloride in Water. All Values are μg/L Unless 

Otherwise Indicated continued
81

Laboratory Duplicate 2 <0.3 <0.49 <0.22

Relative Percent Difference - - -

Spike Concentration 1 200 200 100 100

Sample Concentration 1 <0.3 2.62 <0.49 4.6

Spike Recovery (Percent) 113% 111% 93% 89.5%

Spike Concentration 2 200 200

Sample Concentration 2 <0.3 3.4

Spike Recovery (Percent) 116% 87%

Date Collected 1/14/04 2/19/04 6/4/04 9/7/04 7/14/05

Date Analyzed 1/20/04 2/20/04 6/17/04 9/17/04 7/15/05

CCC Standard Nominal 10 10 20 20 20

CCC Standard Measured 9.97 10.7 19.8 20.8 19.9

Percent of Check Standard 100% 107% 99% 104% 100%

CCC Standard Nominal 100 50 20 200

CCC Standard Measured 105 48.9 20.8 169

Percent of Check Standard 105% 97.8% 104% 84%

CCC Standard Nominal 100 100 20

CCC Standard Measured 108 103 19.4

Percent of Check Standard 108% 103% 97%

QC Standard Nominal 50 10 20 20

QC Standard Measured 57.2 10.9 18.0 20.6

Percent of Check Standard 114% 109% 90% 103%

QC Standard Nominal 100 100 200 200

QC Standard Measured 113 101 192 188

Percent of Check Standard 113% 101% 96% 94%

Blank 1 <0.22 <0.22 <0.3 <0.3 <0.3

Blank 2 <0.22 <0.22 <0.3 <0.3 <0.3

Sample Analysis 1 <0.22 <0.3 <0.3 <0.3

Laboratory Duplicate 1 <0.22 <0.3 <0.3 <0.3

Relative Percent Difference - - - -

Sample Analysis 2 <0.22 <0.3 <0.3 <0.3

Laboratory Duplicate 2 <0.22 <0.3 <0.3 <0.3

Relative Percent Difference - - - -

Spike Concentration 1 100 100 100

Sample Concentration 1 <0.3 <0.3 <0.3

Spike Recovery (Percent) 101% 105% 100%

Spike Concentration 2 100 100 100

Sample Concentration 2 <0.3 <0.3 <0.3

Spike Recovery (Percent) 91% 105% 83%

CCC: Continuing Calibration Check; QC: Second Source Check

Table 8.3. Typical Quality Performance Data for Analysis of Vinyl Chloride in Water. All Values are μg/L Unless 

Otherwise Indicated continued
82

Table 8.4. Typical Quality Performance Data for Analysis of Ethylene in Water or in Gas. The Values for Check 

Standard Nominal and Check Standard Measured are ppm (v/v). All Other Values are mg/L Unless 

Otherwise Indicated

Date Collected 10/23/02 11/25/02 12/23/02 1/22/03 2/4/03

Date Analyzed 11/13/02 12/4/02 12/23/02 1/22/03 2/7/03

Check Standard Nominal 10 10 10 10 10

Check Standard Measured 11 10.5 10.9 10.1 10.9

Percent of Check Standard 110% 105% 109% 101% 109%

Check Standard Nominal 10 100 10 10 100

Check Standard Measured 10.7 108 11.1 10.4 106

Percent of Check Standard 107% 108% 111% 104% 106%

Check Standard Nominal 100 100 10 100 100

Check Standard Measured 104 106 10.9 109 108

Percent of Check Standard 104% 106% 109% 109% 108%

Check Standard Nominal 10000 10 100

Check Standard Measured 10200 11.1 107

Percent of Check Standard 102% 111% 107%

Check Standard Nominal 100 1000

Check Standard Measured 106 1010

Percent of Check Standard 106% 101%

Blank 1 <0.00028 <0.00028 <0.28* <0.28* <0.00028

Blank 2 <0.28*

Sample Analysis 1 <0.00028 <0.00028 <0.00028

Laboratory Duplicate 1 <0.00028 <0.00028 <0.00028

Relative Percent Difference - - -

Sample Analysis 2 <0.00028

Laboratory Duplicate 2 <0.00028

Relative Percent Difference -

Spike Concentration 0.0275 0.275 2.71

Sample Concentration <0.00028 <0.00028 <0.00028

Spike Recovery (Percent) 104% 113% 99%

Date Collected 2/20/03 3/6/03 3/11/03 3/25/03 4/8/03

Date Analyzed 2/20/03 3/11/03 3/19/03 3/25/03 4/14/03

Check Standard Nominal 10 10 10 10 10

Check Standard Measured 10.5 10.1 10.4 10.7 10.3

Percent of Check Standard 105% 101% 104% 107% 103%

Check Standard Nominal 10 100 100 10 100

Check Standard Measured 10.2 107 106 10.4 98

Percent of Check Standard 102% 107% 106% 104% 98%

Check Standard Nominal 10 100 10 100

Check Standard Measured 10.3 102 10.5 106

*The values are ppm (v/v).
83

Percent of Check Standard 103% 102% 105% 106%

Check Standard Nominal 100 10 1000

Check Standard Measured 106 10.4 1030

Percent of Check Standard 106% 104% 103%

Check Standard Nominal 1000 10000

Check Standard Measured 1080 10800

Percent of Check Standard 108% 108%

Blank 1 <0.00057 <0.00028 <0.00057

Blank 2

Sample Analysis 1 <0.00057 <0.00028 <0.00057

Laboratory Duplicate 1 <0.00057 <0.00028 <0.00057

Relative Percent Difference - - -

Sample Analysis 2 <0.00057 <0.00057

Laboratory Duplicate 2 <0.00057 <0.00057

Relative Percent Difference - -

Spike Concentration 0.964 2.68 1.94

Sample Concentration <0.00057 <0.00028 <0.00057

Spike Recovery (Percent) 116% 99% 100%

Date Collected 4/21/03 4/29/03 5/15/03 6/20/03 7/23/03

Date Analyzed 5/1/03 5/6/03 5/15/03 6/20/03 7/31/03

Check Standard Nominal 10 10 10 10 10

Check Standard Measured 11 10.2 10.3 10.3 9.92

Percent of Check Standard 110% 102% 103% 103% 99%

Check Standard Nominal 100 100 10 10 100

Check Standard Measured 109 108 10.4 10.4 107

Percent of Check Standard 109% 108% 104% 104% 107%

Check Standard Nominal 10 10

Check Standard Measured 10.2 10.6

Percent of Check Standard 102% 106%

Check Standard Nominal 10 10

Check Standard Measured 10.5 10.3

Percent of Check Standard 105% 103%

Check Standard Nominal 100 10

Check Standard Measured 107 10.4

Percent of Check Standard 107% 104%

Blank 1 <0.00057 <0.00057 <0.00057

Blank 2 <0.00057

*The values are ppm (v/v).

Table 8.4. Typical Quality Performance Data for Analysis of Ethylene in Water or in Gas. The Values for Check 

Standard Nominal and Check Standard Measured are ppm (v/v). All Other Values are mg/L Unless 

Otherwise Indicated continued
84

Sample Analysis 1 <0.00057 0.003 <0.00057

Laboratory Duplicate 1 <0.00057 <0.00057 <0.00057

Relative Percent Difference - - -

Sample Analysis 2

Laboratory Duplicate 2

Relative Percent Difference

Spike Concentration 1.96

Sample Concentration <0.00057

Spike Recovery (Percent) 92%

Date Collected 8/6/03 8/20/03 10/2/03 10/7/03 10/22/03

Date Analyzed 8/6/03 8/25/03 10/2/03 10/16/03 10/22/03

Check Standard Nominal 10 10 10 100 10

Check Standard Measured 10.9 9.44 10.4 101 10.1

Percent of Check Standard 109% 94.4% 104% 101% 101%

Check Standard Nominal 10 100 10 1000 100

Check Standard Measured 10.3 106 9.9 1030 106

Percent of Check Standard 103% 106% 99% 103% 106%

Check Standard Nominal 100 100 10000 1000

Check Standard Measured 106 105 10100 1080

Percent of Check Standard 106% 105% 101% 108%

Check Standard Nominal 100 100 100000

Check Standard Measured 108 106 97000

Percent of Check Standard 108% 106% 97%

Check Standard Nominal 10000 1000

Check Standard Measured 10900 1080

Percent of Check Standard 109% 108%

Blank 1 <0.57* <0.00057 <0.57* <0.0003 <0.57*

Blank 2

Sample Analysis 1 <0.00057 <0.0003

Laboratory Duplicate 1 <0.00057 <0.0003

Relative Percent Difference - -

Sample Analysis 2 <0.00057 0.039

Laboratory Duplicate 2 <0.00057 0.037

Relative Percent Difference - 5.3%

Spike Concentration 

Sample Concentration 

Spike Recovery (Percent)

*The values are ppm (v/v).

Table 8.4. Typical Quality Performance Data for Analysis of Ethylene in Water or in Gas. The Values for Check 

Standard Nominal and Check Standard Measured are ppm (v/v). All Other Values are mg/L Unless 

Otherwise Indicated continued
85

Date Collected 1/14/04 2/18/04 6/1/04 9/9/04 7/13/05

Date Analyzed 1/14/04 2/18/04 6/1/04 9/9/04 7/13/05

Check Standard Nominal 10 10 10 10 10

Check Standard Measured 9.98 10.3 10.1 10.3 10.8

Percent of Check Standard 100% 103% 101% 103% 108%

Check Standard Nominal 100 10 10 10 10

Check Standard Measured 104 9.82 10.4 10.4 10.1

Percent of Check Standard 104% 98% 104% 104% 101%

Check Standard Nominal 100 10 10 100 100

Check Standard Measured 106 10.4 10.2 112 105

Percent of Check Standard 106% 104% 102% 112% 105%

Check Standard Nominal 1000 10 100 100 100

Check Standard Measured 1070 10.5 107 110 104

Percent of Check Standard 107% 105% 107% 110% 104%

Check Standard Nominal 100 100 1000 1000

Check Standard Measured 106 108 1100 1020

Percent of Check Standard 106% 108% 110% 102%

Blank 1 <0.57* <0.57* <0.57* <0.57* <0.57*

Blank 2 <0.57*

Sample Analysis 1

Laboratory Duplicate 1

Relative Percent Difference

Sample Analysis 2

Laboratory Duplicate 2

Relative Percent Difference

Spike Concentration 

Sample Concentration 

Spike Recovery (Percent)

*The values are ppm (v/v).

Table 8.4. Typical Quality Performance Data for Analysis of Ethylene in Water or in Gas. The Values for Check 

Standard Nominal and Check Standard Measured are ppm (v/v). All Other Values are mg/L Unless 

Otherwise Indicated continued
86

Table 8.5. Typical Quality Performance Data for Analysis of Methane in Water or in Gas. The Values for Check 

Standard Nominal and Check Standard Measured are ppm (v/v). All Other Values are mg/L Unless 

Otherwise Indicated

Date Collected 10/23/02 11/25/02 12/23/02 1/22/03 2/4/03

Date Analyzed 11/13/02 12/4/02 12/23/02 1/22/03 2/7/03

Check Standard Nominal 100 10 10 10 10

Check Standard Measured 104 10.9 10.7 10.4 10.6

Percent of Check Standard 104% 109% 107% 104% 106%

Check Standard Nominal 100 100 10 10 100

Check Standard Measured 106 106 10.6 9.95 110

Percent of Check Standard 106% 106% 106% 100% 110%

Check Standard Nominal 1000 100 10 10 100

Check Standard Measured 1056 109 10.8 9.88 106

Percent of Check Standard 105% 109% 108% 99% 106%

Check Standard Nominal 10000 10000 100 100 1000

Check Standard Measured 9340 10000 107 102 1060

Percent of Check Standard 93% 100% 107% 102% 106%

Check Standard Nominal 100000 100 100

Check Standard Measured 99800 107 106

Percent of Check Standard 100% 107% 106%

Blank 1 <0.00003 <0.0003 <0.3* <0.3* <0.0003

Blank 2 <0.3*

Sample Analysis 1 0.2 <0.0003 <0.0003

Laboratory Duplicate 1 0.198 <0.0003 <0.0003

Relative Percent Difference 1.0% - -

Sample Analysis 2 0.0551

Laboratory Duplicate 2 0.0548

Relative Percent Difference 0.5%

Spike Concentration 0.0098 0.0978 0.971

Sample Concentration 0.0009 <0.0003 0.0015

Spike Recovery (Percent) 96% 114% 106%

Date Collected 2/20/03 3/6/03 3/11/03 3/25/03 4/8/03

Date Analyzed 2/20/03 3/11/03 3/19/03 3/25/03 4/14/03

Check Standard Nominal 10 10 10 10 10

Check Standard Measured 10.6 10.4 10.5 10.8 9.56

Percent of Check Standard 106% 104% 105% 108% 96%

Check Standard Nominal 10 100 100 10 100

Check Standard Measured 9.99 109 109 9.85 97.9

Percent of Check Standard 100% 109% 109% 99% 98%

Check Standard Nominal 10 1000 1000 100 1000

Check Standard Measured 10.2 1040 1050 105 1020

*The values are ppm (v/v).
87

Percent of Check Standard 102% 104% 105% 105% 102%

Check Standard Nominal 10 10000 10000 1000 10000

Check Standard Measured 10.1 10100 10300 1060 9900

Percent of Check Standard 101% 101% 103% 106% 99%

Check Standard Nominal 100 100000 10000 100000

Check Standard Measured 107 100000 9790 95900

Percent of Check Standard 107% 100% 98% 96%

Blank 1 <0.00042 <0.00003 <0.00042

Blank 2

Sample Analysis 1 0.086 0.007 0.002

Laboratory Duplicate 1 0.0854 0.007 0.002

Relative Percent Difference 0.7% 0.0% 0.0%

Sample Analysis 2 <0.00042 0.147

Laboratory Duplicate 2 <0.00042 0.145

Relative Percent Difference - 1.4%

Spike Concentration 0.964 0.964 0.957

Sample Concentration 0.007 <0.00003 0.019

Spike Recovery (Percent) 93% 108% 89%

Date Collected 4/21/03 4/29/03 5/15/03 6/20/03 7/23/03

Date Analyzed 5/1/03 5/6/03 5/15/03 6/20/03 7/31/03

Check Standard Nominal 10 100 10 10 100

Check Standard Measured 11 102 10.1 10.4 101

Percent of Check Standard 110% 102% 101% 104% 101%

Check Standard Nominal 100 10000 10 10 100

Check Standard Measured 104 10200 10.5 10.3 98.8

Percent of Check Standard 104% 102% 105% 103% 99%

Check Standard Nominal 1000 100000 100 100 1000

Check Standard Measured 1070 97300 102 100 1030

Percent of Check Standard 107% 97% 102% 100% 103%

Check Standard Nominal 10000 1000 1000 10000

Check Standard Measured 10300 1070 1060 9850

Percent of Check Standard 103% 107% 106% 99%

Check Standard Nominal 100000 10000 10000 100000

Check Standard Measured 98200 10300 10200 95300

Percent of Check Standard 98% 103% 102% 95%

Blank 1 <0.00042 <0.00042 <0.00042

Blank 2 <0.00042

*The values are ppm (v/v).

Table 8.5. Typical Quality Performance Data for Analysis of Methane in Water or in Gas. The Values for Check 

Standard Nominal and Check Standard Measured are ppm (v/v). All Other Values are mg/L Unless 

Otherwise Indicated continued
88

Sample Analysis 1 0.265 12.3 7.28

Laboratory Duplicate 1 0.229 12.9 7.02

Relative Percent Difference 14.6% 4.8% 3.6%

Sample Analysis 2

Laboratory Duplicate 2

Relative Percent Difference

Spike Concentration 0.964

Sample Concentration 0.001

Spike Recovery (Percent) 97%

Date Collected 8/6/03 8/20/03 10/2/03 10/7/03 10/22/03

Date Analyzed 8/6/03 8/25/03 10/2/03 10/16/03 10/22/03

Check Standard Nominal 100 100 10 100 10

Check Standard Measured 102 101 9.6 107 10.3

Percent of Check Standard 102% 101% 96% 107% 103#

Check Standard Nominal 100 100 100 10 100

Check Standard Measured 99.9 95.8 101 10.1 103

Percent of Check Standard 100% 96% 101% 101% 103%

Check Standard Nominal 1000 1000 100 100 1000

Check Standard Measured 1060 1020 102 103 1070

Percent of Check Standard 106% 102% 102% 103% 107%

Check Standard Nominal 10000 10000 1000 1000 10000

Check Standard Measured 10300 9810 1070 1050 10300

Percent of Check Standard 103% 98% 107% 105% 103%

Check Standard Nominal 100000 100000 10000 100000

Check Standard Measured 97800 94700 10300 98500

Percent of Check Standard 98% 95% 103% 99%

Blank 1 <0.42* <0.00042 <0.42* <0.0001 <0.42*

Blank 2

Sample Analysis 1 1.70 2.15

Laboratory Duplicate 1 1.35 2.15

Relative Percent Difference 23.0% 0.0%

Sample Analysis 2 2.44 3.85

Laboratory Duplicate 2 2.78 3.36

Relative Percent Difference 13.0% 13.6%

Spike Concentration 

Sample Concentration 

Spike Recovery (Percent)

*The values are ppm (v/v).

Table 8.5. Typical Quality Performance Data for Analysis of Methane in Water or in Gas. The Values for Check 

Standard Nominal and Check Standard Measured are ppm (v/v). All Other Values are mg/L Unless 

Otherwise Indicated continued
89

Date Collected 1/14/04 2/18/04 6/1/04 9/9/04 7/13/05

Date Analyzed 1/14/04 2/18/04 6/1/04 9/9/04 7/13/05

Check Standard Nominal 100 10 10 10 10

Check Standard Measured 98.4 10.7 10.9 10.7 9.88

Percent of Check Standard 98% 107% 109% 107% 98.8%

Check Standard Nominal 100 100 100 100 100

Check Standard Measured 101 102 107 105 101

Percent of Check Standard 101% 102% 107% 105% 101%

Check Standard Nominal 1000 1000 1000 1000 1000

Check Standard Measured 1070 1070 1080 1110 1040

Percent of Check Standard 107% 107% 108% 111% 104%

Check Standard Nominal 10000 10000 10000 10000 10000

Check Standard Measured 10200 10300 10500 11200 10300

Percent of Check Standard 102% 103% 105% 112% 103%

Check Standard Nominal 100000 100000 100000 100000 100000

Check Standard Measured 97700 98000 99200 99800 99800

Percent of Check Standard 98% 98% 99.2% 99.8% 99.8%

Blank 1 <0.42* <0.42* <0.42* <0.42* <0.42*

Blank 2 <0.42*

Sample Analysis 1

Laboratory Duplicate 1

Relative Percent Difference

Sample Analysis 2

Laboratory Duplicate 2

Relative Percent Difference

Spike Concentration 

Sample Concentration 

Spike Recovery (Percent)

*The values are ppm (v/v).

Table 8.5. Typical Quality Performance Data for Analysis of Methane in Water or in Gas. The Values for Check 

Standard Nominal and Check Standard Measured are ppm (v/v). All Other Values are mg/L Unless 

Otherwise Indicated continued
90

Table 8.6. Typical Quality Performance Data for Analysis of Hydrogen in Gas. All Values are ppm Unless Otherwise 

Indicated

Date Collected 10/23/02 12/23/02 2/20/03 3/6/03 3/11/03

Date Analyzed 11/5/02 12/23/02 2/20/03 3/11/03 3/19/03

Check Standard Nominal 1 0.5 0.5 0.5 1

Check Standard Measured 1.037 0.59 0.49 0.54 0.98

Percent of Check Standard 103% 118% 99% 108% 98%

Check Standard Nominal 5 1.19 0.5 0.5 0.5

Check Standard Measured 5.479 1.26 0.5 0.486 0.52

Percent of Check Standard 110% 92.4% 100% 97% 104%

Check Standard Nominal 0.5 2.5 1 1 1

Check Standard Measured 0.46 2.45 1.07 1.094 1.06

Percent of Check Standard 96% 98% 107% 109% 106%

Check Standard Nominal 1 10.1 5 5 5

Check Standard Measured 1.03 9.34 5.6 5.489 5.32

Percent of Check Standard 103% 98% 110% 109% 106%

Check Standard Nominal 5 10.1 10

Check Standard Measured 5.42 11.0 9.96

Percent of Check Standard 108% 109% 99%

Blank 1 <0.19 <0.19 <0.19 <0.19 <0.19

Blank 2

Sample Analysis 1 0.348 0.74 0.564

Laboratory Duplicate 1 0.351 2.67 0.601

Relative Percent Difference 0.9% 113.2% 6.4%

Sample Analysis 2 2.847 0.33

Laboratory Duplicate 2 2.042 0.36

Relative Percent Difference 32.9% 8.7%

Date Collected 3/25/03 4/8/03 4/21/03 4/29/03 5/15/03

Date Analyzed 3/25/03 4/14/03 5/1/03 5/6/03 5/15/03

Check Standard Nominal 0.5 0.5 1 0.5 0.5

Check Standard Measured 0.51 0.473 0.943 0.534 0.533

Percent of Check Standard 102% 95% 94% 107% 94%

Check Standard Nominal 1 0.5 1 1 1

Check Standard Measured 0.99 0.534 0.899 1.04 0.939

Percent of Check Standard 99% 107% 90% 104% 94%

Check Standard Nominal 1 1 5 5 5

Check Standard Measured 1 1.03 5.05 5.06 5.01

Percent of Check Standard 100% 103% 101% 101% 100%

Check Standard Nominal 5 1 10 10 10

Check Standard Measured 4.98 1.05 10.4 10.5 10.6

The values are ppm (v/v).
91

Percent of Check Standard 99% 105% 104% 105% 106%

Check Standard Nominal 5 5 20 20 20

Check Standard Measured 4.9 4.56 21.3 19.5 20.5

Percent of Check Standard 98% 91% 106% 98% 103%

Blank 1 <0.19 <0.19 <0.19 <0.19

Blank 2

Sample Analysis 1 2.08 <0.5 2.5

Laboratory Duplicate 1 2.59 <0.5 2.1

Relative Percent Difference 21.8% - 17.4%

Sample Analysis 2 0.669

Laboratory Duplicate 2 0.744

Relative Percent Difference 10.6%

Date Collected 6/20/03 7/23/03 8/6/03 8/20/03 10/2/03 10/7/03

Date Analyzed 6/20/03 7/31/03 8/6/03 8/25/03 10/2/03 10/16/03

Check Standard Nominal 0.5 0.5 0.5 0.5 0.5 1

Check Standard Measured 0.5 0.523 0.461 0.51 0.585 1.02

Percent of Check Standard 100% 105% 92% 102% 117% 102%

Check Standard Nominal 1 1 1 1 1 5

Check Standard Measured 1.04 1.17 0.959 0.96 0.964 4.6

Percent of Check Standard 104% 117% 96% 96% 96% 92%

Check Standard Nominal 10 5 5 20 5 10

Check Standard Measured 10.5 4.4 5.45 21.6 5.55 9.52

Percent of Check Standard 105% 88% 109% 108% 111% 95%

Check Standard Nominal 10 10 10 20

Check Standard Measured 11.1 11 8.89 17.6

Percent of Check Standard 111% 110% 89% 88%

Check Standard Nominal 20 20 20

Check Standard Measured 18.8 18.5 16.9

Percent of Check Standard 94% 93% 85%

Blank 1 <0.19 <0.19 <0.19 <0.19 0.232 <0.19

Blank 2

Sample Analysis 1 13.9 0.48 134

Laboratory Duplicate 1 13.1 0.42 136

Relative Percent Difference 5.9% 13.3% 1.5%

Sample Analysis 2

Laboratory Duplicate 2

Relative Percent Difference

The values are ppm (v/v).

Table 8.6. Typical Quality Performance Data for Analysis of Hydrogen in Gas. All Values are ppm Unless Otherwise 

Indicated continued
92

Date Collected 10/22/03 1/14/05 2/18/04 6/1/04 9/9/04 7/13/05

Date Analyzed 10/22/03 1/14/05 2/18/04 6/1/04 9/9/04 7/13/05

Check Standard Nominal 0.5 1 1 0.5 0.5 0.5

Check Standard Measured 0.444 1.12 1.07 0.586 0.465 0.459

Percent of Check Standard 113% 112% 107% 117% 93% 92%

Check Standard Nominal 1 5 5 1 1 1

Check Standard Measured 0.991 4.30 5.13 0.931 1.13 0.937

Percent of Check Standard 99% 116% 103% 93.1% 113% 93.7%

Check Standard Nominal 5 10 10 5 5 5

Check Standard Measured 5.23 9.45 9.72 4.42 5.67 4.88

Percent of Check Standard 105% 95% 97.2% 88.4% 113% 97.6%

Check Standard Nominal 10 20 10 10 10 10

Check Standard Measured 8.98 18.4 9.9 11.5 11.1 10.1

Percent of Check Standard 90% 109% 99% 115% 111% 101%

Check Standard Nominal 20 20 20 20 20

Check Standard Measured 19.5 19 18.1 18.6 19.2

Percent of Check Standard 98% 95% 90.5% 93% 96%

Blank 1 0.244 <0.19 <0.19 <0.19 <0.19 <0.19

Blank 2 <0.19

Sample Analysis 1

Laboratory Duplicate 1

Relative Percent Difference

Sample Analysis 2

Laboratory Duplicate 2

Relative Percent Difference

The values are ppm (v/v).

Table 8.6. Typical Quality Performance Data for Analysis of Hydrogen in Gas. All Values are ppm Unless Otherwise 

Indicated continued
93

Table 8.7. Typical Quality Performance Data for Analysis of Nitrate Plus Nitrite Nitrogen in Water. All Values are 

mg/L Unless Otherwise Indicated

Date Collected 10/23/02 2/4/03 3/6/03 3/11/03 4/8/03

Date Analyzed 11/6/02 2/7/03 3/20/03 3/26/03 4/14/03

CCC Standard Nominal 0.5 2.5 2.5 0.5 0.5

CCC Standard Measured 0.49 2.36 2.53 0.51 0.45

Percent of Check Standard 98% 94% 101% 102% 90%

CCC Standard Nominal 1 1 1 1

CCC Standard Measured 0.97 0.97 1 0.93

Percent of Check Standard 97% 97% 100% 93%

CCC Standard Nominal 2.5

CCC Standard Measured 2.45

Percent of Check Standard 98%

QC Standard Nominal 

QC Standard Measured 

Percent of Check Standard

QC Standard Nominal 7.06 2.99 2.99 2.99

QC Standard Measured 7.06 3.02 3.10 3.05

Percent of Check Standard 100% 101% 104% 102%

Blank 1 <0.1 <0.1 <0.1 <0.1 <0.1

Blank 2 <0.1

Sample Analysis <0.1 0.62 1.58 1.25 <0.1

Laboratory Duplicate <0.1 0.59 1.57 1.29 <0.1

Relative Percent Difference - 5.0% 0.6% 3.1% -

Spike Concentration 10 10 10 10 10

Sample Concentration <0.1 2.04 1.97 0.03 0.06

Spike Recovery (Percent) 107% 103% 110% 106% 116%

Date Collected 4/21/03 4/29/03 7/23/03 8/20/03 10/7/03

Date Analyzed 5/13/03 5/13/03 8/6/03 8/25/03 10/28/03

CCC Standard Nominal 1 1 2.5 0.5 0.5

CCC Standard Measured 0.95 0.95 2.39 0.48 0.5

Percent of Check Standard 95% 95% 95.6% 96% 100%

CCC Standard Nominal 5 1 5 1 1

CCC Standard Measured 4.99 0.96 4.78 0.92 0.93

Percent of Check Standard 100% 96% 96% 92% 93%

CCC Standard Nominal 1 5 5 5

CCC Standard Measured 0.96 4.74 4.91 5

Percent of Check Standard 96% 95% 98.2% 100%

QC Standard Nominal 10 10

QC Standard Measured 9.8 9.87

Percent of Check Standard 98% 98.7%

CCC: Continuing Calibration Check; QC: Second Source Check
94

QC Standard Nominal 13.3 13.3 21.3

QC Standard Measured 12.7 12.7 18.9

Percent of Check Standard 95% 95% 89%

Blank 1 <0.1 <0.1 <0.1 <0.1 <0.1

Blank 2 <0.1 <0.1 <0.1 <0.1 <0.1

Sample Analysis 3.39 <0.1 11.2 <0.1 <0.1

Laboratory Duplicate 3.49 <0.1 11.1 <0.1 <0.1

Relative Percent Difference 2.9% - 0.9% - -

Spike Concentration 10 10 5 10 5

Sample Concentration 1.26 0.04 <0.004 1.86 0.04

Spike Recovery (Percent) 116% 109% 84.6% 109% 101%

CCC: Continuing Calibration Check; QC: Second Source Check

Table 8.7. Typical Quality Performance Data for Analysis of Nitrate Plus Nitrite Nitrogen in Water. All Values are 

mg/L Unless Otherwise Indicated continued
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