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This is to certify that the
dissertation entitled
CHARACTERIZATION OF MICROBIAL POPULATIONS

IN ANAEROBIC FOOD WEBS THAT REDUCTIVELY
DECHLORINATE CHLOROPHENOLS

presented by

Robert Allan Sanford

has been accepted towards fulfillment
of the requirements for

Ph . D . degree in mm};

 

 

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Date July 30, 1996

 

MSU is an Affirmative Action/Equal Opportunity Institution 0-12771

 

 

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MSU Is An Nflnnatlvo Adlai/Equal Opponunity Institution
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CHARACTERIZATION OF MICROBIAL POPULATIONS
IN ANAEROBIC FOOD WEBS THAT REDUCTIVELY
DECHLORINATE CHLOROPHENOLS
BY

Robert Allan Sanford

A DISSERTATION

Submitted to
Michi an State University
in partial ful illment of the requirements
for the Degree of

DOCTOR OF PHILOSOPHY

Department of Microbiology
° 1996

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ABSTRACT

CHARACTERIZATION OF MICROBIAL POPULATIONS IN ANAEROBIC
FOOD WEBS THAT REDUCTIVELY DECHLORINATE
CHLOROPHENOLS

By
Robert Allan Sanford

In this study the anaerobic dechlorination activities of mono- and
dichlorophenols (CPS and DCPs) in nitrate-fed microcosms were characterized.
Denitrification was shown to not be coupled to the reductive dechlorination of
CPs or DCPs. Enrichment cultures derived from the microcosms exhibited
different substrate specificity for individual GP or DCP isomers depending on
the substrate fed to the enrichment. Four dechlorinating cultures with different
dechlorination patterns were studied further, with two new halorespiring
microbial isolates and two novel meta-dechlorinating enrichment cultures
characterized. The first isolates, strains ZCP-C and 20P-3, were identified as
Anaeromyxo dehalogenans, the first anaerobic myxobacteria characterized.
Strains ZCP-C and ZCP-3 are able to grow using acetate as an electron donor
and 2-CP as an electron acceptor. The second enrichment culture exhibited the
ortho-dechlorination of 2,3-DCP and yielded the new halorespiring isolate
Desulfitobacterium chlororespiricans strain 0023. Strain 0023 coupled the
partial oxidation of lactate to reductive dechlorination to gain energy for growth.
The third culture exhibited meta-dechlorination of 3-CP under methanogenic
conditions. Inhibitors for methanogenesis and eubacterial growth added to the
culture indicated that a eubacterial population was responsible for
dechlorination. The final enrichment culture derived from the microcosms
dechlorinated ortho- and meta- substituted chlorines from 2,5-DCP, 2- plus 3-

CP simultaneously. 62% of the available reducing equivalents were used for

reduci
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reductive dechlorination, indicating that this was likely to be a respiratory
process. H2 was observed in the headspace of the microcosm used for the OM
enrichment culture, which indicated that H2 was a very important intermediate
in this anaerobic ecosystem. The microbial population responsible for H2
production from formate in the OM enrichment culture was isolated and
characterized. Strain FOX1 is a novel sulfate-reducing bacterium with the novel
ability to grow on formate oxidation to H2 and 002 as the sole source of energy.
Energetically FOX1 is able to grow at the lowest AG' observed for any

microorganism and produces H2 at greater rates and to higher concentrations

than previously reported.

To my father Allan and in memory of my grandfather Roscoe

both who's passion and dedication to science inspired me.

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ACKNOWLEDGMENTS

The author expresses his sincere thanks to:

Dr. James Tiedje for accepting me into his lab so very long ago and for
having the patience to allow me to pursue my own pathway to enlightenment. I
feel very fortunate to have had the opportunity to work in his lab.

Dr. Joanne Ghee-Sanford, my wife, friend and colleague who has put up
with the graduate student life for way too long. She helped make it much
easier. My daughter Kelsey for bringing joy and adventure to my life. My dog
Vmax for being a loyal companion.

Members of my committee, Dr. John Breznak, Dr. Craig Criddle, Dr. Mike
Klug and Dr. Robert Hausinger (for pinch-hitting at the last moment).

Craig Criddle for introducing me to the world of environmental
engineering.

My parents for their continuous support.

Fellow lab colleagues, some of whom will be probably forgotten, but are
appreciated with all the others: Marcos Fries, Suzanne Thiem, Jorge Santo
Domingo, Arturo Massol-Deya, Frank Loffler, Jim Champine, Mike Apgar, Jim
Cole, Amy Cascarelli, Mary Ann Bruns, John Dunbar, Yuichi Suwa, Grace
Matheson, Jong-ok Ka, Jizhong Zhou, Nancy Sasaki, Al Rhodes, Roberta
Fulthorpe, Dave Demezas, Rick Ye, Cathy McGowan, Dave Harris, Joyce
Wildenthal and Inez Tore-Suarez.

The Center for Microbial Ecology and NSF for providing the financial
support for my graduate study.

TABLE OF CONTENTS

UST OF TABLES ............................................................................................................. ix
UST OF FIGURES ........................................................................................................... xi
CHAPTER I
INTRODUCTION ............................................................................................................... 1
Oven/iew ........................................................................................................................... 1
Background ...................................................................................................................... 3
Bicdegradation Under Denitrifying Conditions .............................................. 3
Naturally occurring halogenated compounds ................................................ 5
Reductive dehalogenation of chlorophenols ................................................. 6
Energy flow in anaerobic food webs: the role
of formate and H2 .............................................................................................. 12
References .................................................................................................................... 16
CHAPTER II
Chlorophenol dechlorination and subsequent degradation in
nitrate-limited denitrifying microcosms ...................................................................... 21
Abstract ............................................................................................................... 21
Introduction ......................................................................................................... 22
Materials and Methods .................................................................................... 23
Source of microcosm inocula ............................................................. 23
Microcosms ............................................................................................ 24
Medium formulation .............................................................................. 24
Enrichment cultures .............................................................................. 25
Chemical analysis ................................................................................. 25
Chemicals ............................................................................................... 26
Results .................................................................................................................. 26
Dechlorinating denitrifying microcosms ............................................ 26
Independence of dechlorination from nitrate reduction ................. 31
Characterization of enrichment activity ............................................. 34
Discussion .......................................................................................................... 35
Acknowledgments ............................................................................................. 41
References ......................................................................................................... 41
CHAPTER III
Isolation and preliminary characterization of the aryl-halorespiring
anaerobic myxobacteria strains 2CP-C and 20P-3 ............................................... 45
Introduction ......................................................................................................... 45
Materials and Methods ..................................................................................... 46

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Isolation ................................................................................................... 46

Growth medium ..................................................................................... 47
Electron donors and acceptors ........................................................... 47
Nitrate effect on dechlorination ........................................................... 48
Microscopy ............................................................................................. 49
Hydrogen uptake and threshold determination ............................... 49
Determination of fe ................................................................................ 49
Analytical procedures ........................................................................... 50
Results ................................................................................................................. 51
Isolation of 2CP strains ........................................................................ 51
Electron donors and acceptors ........................................................... 53
Dechlorination in the presence of nitrate .......................................... 57
Indicators of halorespiration ................................................................ 57
Discussion .......................................................................................................... 60
References ......................................................................................................... 67
CHAPTER IV

Characterization of Desulfitobacterium chlororespiricans sp. nov.
strain 0023, which grows by coupling the oxidation of lactate to

the reductive dechlorination of 3-chloro-4-hydroxybenzoate ............................... 70
Abstract ............................................................................................................... 71
Introduction ......................................................................................................... 72
Materials and Methods ..................................................................................... 73

Growth conditions ................................................................................. 73
Determination of electron donors and acceptors ............................ 74
Pasteurization and microscopy .......................................................... 76
Temperature effect on growth and dechlorination .......................... 76
16S rRNA gene isolation, sequencing and analysis ...................... 76
Chemical analyses ............................................................................... 78
Chemicals ............................................................................................... 79
Results ................................................................................................................. 79
Isolation of Desulfitobacterium chlororespiricans
strain C023 ............................................................................................. 79
Evidence for halorespiration ............................................................... 80
Range of electron donors and acceptors used ................................ 82
Growth and dechlorination rates ........................................................ 87
Phylogeny of strain C023 .................................................................... 92
Discussion .......................................................................................................... 92
Description of Desulfitobacterium chlororespiricans
sp. nov. .................................................................................................. 101
Acknowledgments .......................................................................................... 1 02
References ....................................................................................................... 102
CHAPTER V

Meta-dechlorination of 3—chlorophenol and
2,3-dichlorophenol in methanogenic enrichment cultures:

the effects of inhibitors and different electron donors .......................................... 107
Introduction ...................................................................................................... 107
Materials and Methods .................................................................................. 108

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Enrichment Cultures and growth conditions .................................. 108

Inhibition of meta-dechlorination ..................................................... 109
Electron donors for meta-dechlorination ........................................ 1 10
Induction of dechlorination ................................................................ 110
Utilization of 2,3-DCP ......................................................................... 110
Analytical procedures ........................................................................ 1 10
Results .............................................................................................................. 111
Characterization of reductive dechlorination activity ................... 111
Effect of inhibitors on reductive dechlorination ............................. 112
Effect of preincubation on meta-dechlorination lag time ............. 112
Dechlorination of 2,3-DCP ................................................................ 1 16
Electron donors for meta-dechlorination ........................................ 116
Discussion ........................................................................................................ 119
References ....................................................................................................... 125
CHAPTER VI
Evidence for halorespiration in an anaerobic enrichment culture that
simultaneously dechlorinates 2- and 3—chlorophenol to phenol ....................... 128
Introduction ...................................................................................................... 128
Materials and Methods .................................................................................. 129
Enrichment Cultures and growth conditions .................................. 129
Determination of electron donors and acceptors .......................... 130
Hydrogen uptake and threshold determination ............................ 131
Determination of fe ............................................................................. 132
Analytical procedures ........................................................................ 1 32
Results .............................................................................................................. 1 33
Enrichment culture activity ................................................................ 133
Electron acceptors and donors ........................................................ 135
H2 from formate oxidation supports dechlorination ..................... 138
Evidence for halorespiration ............................................................. 138
Discussion ........................................................................................................ 141
References ....................................................................................................... 147
CHAPTER VII
Anaerobic oxidation of formate to H2 + CO2 supports growth
of a microbial culture .................................................................................................. 150
Abstract ............................................................................................................. 151
References and Notes ................................................................................... 163
CHAPTER VIII
SUMMARY AND CONCLUSIONS .......................................................................... 166

viii

Table

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LIST OF TABLES

Table page

CHAPTER I
Table 1.1 Relevant half-reactions for reductive dechlorination of monochloro-
phenol and other electron acceptors. Half-reactions for formate, H2 and
butyrate as electron donors are also shown. AG°’ values shown where
known or calculated from theoretical thermodynamic calculations (Thauer et
al. 1977; Criddle et al. 1991; Dolfing and Harrison 1992) .............................. 10

CHAPTER II
Table 2.1. Summary of dechlorinating and denitrifying microcosms that were
active within a year of incubation. Microcosms received mixtures of MCPs
and DCPs and nitrate at 5 mM or 1 mM .............................................................. 27

Table 2.2 Characteristics of active enrichment cultures. At least five different
dechlorination patterns are observed in the enrichments. Shown are the
chlorophenolic substrate specificities of the different enrichment cultures.
Also indicated is the sensitivity of dechlorination to nitrate and whether

methane is detected in the enrichment cultures ............................................... 33
CHAPTER III
Table 3.1. Characteristics of 2CP strains. Data reflect general features
observed in the 2CP strains isolated and described in this study ................. 55

Table 3.2. Electron acceptors used by different 2CP strains. A (+) indicates
reductive dechlorination and acetate consumption occurred over at least
three additions of the halogenated substrate. This indicated growth had
occurred, which was confirmed by microscopic examination in those
cultures marked with an (*). (+I-) indicates that activity is observed but is
slow and complete degradation did not occur .................................................. 56

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CHAPTER IV
Table 4.1. Mass balance of electron donors (lactate, formate and butyrate) and
3CI-4-HBA consumed and yield of cells as a dry-weight ................................ 85

Table 4.2. Electron donors tested for use by strain 0023 in a minimal medium.
2,3-DCP 0r 3-Cl-4-OH-benzoate served as electron acceptors. A positive
score indicated growth which was monitored by measuring the depletion of
electron donor and acceptor. Cultures were grown at 30° C ....................... 86

Table 4.3. Electron acceptors tested with strain C023 with (+) indicating support
of growth and (-) indicating no growth. Growth was determined by
measuring the depletion of the electron donor and/or electron acceptor as
well as observing visual increase in culture turbidity. Lactate or butyrate
were used as electron donors. Cultures were incubated at 30° C ............... 89

Table 4.4. Pairwise similarity values for strain C023 and selected bacterial
species ...................................................................................................................... 93

Table 4.5. Comparison of strain C023 with two other isolated dehalogenating
strains. The data for Desulfitobacterl'um dehalogenans and strain DOB-2
are from Utkin et. al. (1994). and (Madsen and Licht, 1992) respectively....98

CHAPTER V
Table 5.1. The effect of different electron acceptors and inhibitors on the
efficiency of dechlorination, methanogenesis and the generation of acetate.
The 95% confidence intervals are shown in parentheses ............................ 120

CHAPTER VI
Table 6.1. Electron acceptors and donors utilized by OM enrichment culture. A
(+) indicates that both the electron donor and acceptor were depleted.
Electron donors were tested using 2-CP plus 3-CP as electron

acceptors ................................................................................................................ 137

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CHAPTER I

Figure 1.1. Several naturally occurring halogenated phenolic compounds.
Taken from Gribble (1992) and de Jong (1994) .................................................. 7

Figure 1.2. Anaerobic degradation pathways for trichlorophenols. Heavy
arrows denote reductive dechlorination reactions known to support growth
via halorespiration. Arrows with dotted lines indicate theoretical
transformations and arrows with thin-lines indicate reactions that have been
observed in anaerobic systems .............................................................................. 8

Figure 1.3. Schematic representation of H2 cycling mechanism in a sulfate
reducing microorganism using formate as an electron donor. FD = formate
dehydrogenase; Hyd = hydrogenase; SR = sulfate reductases; ecR =
electron carrier reductase and AT: ATP ase. Adapted from Odom and Peck
(1981) ........................................................................................................................ 15

CHAPTER II
Figure 2.1. Degradation of chlorophenol in the MCP compost soil microcosm
showing ortho—dechlorination activity. o-MCP dechlorination and
persistence of m-MCP and p-MCP (A). Denitrification activity and transient
appearance of phenol. Filled arrows indicate addition of o-MCP (8). Open
arrows indicate the addition of fresh nitrate. Nitrite was not detected in the
enrichment .............................................................................................................. .28

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Figure 2.2. Dechlorination of o-MCP and m-MCP in the 01C MCP microcosm.
(A) Concentrations of MCPs over 550 days of incubation. Closed arrows
indicating the addition of o-MCP and m-MCP. (8) Indicates the
concentrations of phenol and nitrate as they accumulate and dissappear

respectively .............................................................................................................. 30

Figure 2.3. Transfers of dechlorinating and denitrifying enrichments to media
with MCP plus 1 mM nitrate (A) or 1 mM acetate (0). Phenol (D) is shown
as a product of dechlorination or the substrate for degradation. (A)
Dechlorination activity inc-MCP enrichment from Trop I microcosm. (8)
Meta dechlorination in the DCP enrichment culture from the compost
microcosm. (C) Enrichment with phenol and nitrate (1 mM) from compost
microcosm Arrows denote addition of substrate .............................................. 32

Figure 2.4. Schematic representation that summarizes the different
dechlorination and degradation activities observed in the enrichments.
Large numbers indicate the predominant reaction mediated by a particular
enrichment and small numbers indicate additional dechlorination reactions
that are observed in these cultures. (Numbers correspond to the following
enrichments: reaction 1 = o-MCP; reaction 2 = 2,3-DCP; reaction 3 = o-
and m-MCP; reaction 4 = m-MCP and reaction 5 = 3,4-DCP) Reactions 1, 2
and 3 represent distinct ortho dechlorinating pathways. Cultures mediating
reaction 1 do not dechlorinate 2,3-DCP. Meta dechlorination is mediated

' by at least two different communities depicted by reactions 3 and 5. Para
dechlorination is only observed for 3,4-DCP as shown by reaction 4. The
fate of phenol is summarized in two possible pathways; denitrifying and

methanogenic through benzoate ......................................................................... 38
CHAPTER III
Figure 3.1. Dechlorination of 2-CP to phenol by strain 2CP-C. Reduction of
acetate by the same culture is also shown ........................................................ 52

Figure 3.2. (A) Vegetative cells of strain ZCP-C by phase constrast microscopy.
Bar is 13 um long. (8) Colony formed by 2CP-C on anaerobic agar, which
shows the formation of a central cell-mass ........................................................ 54

xii

Figure 3.3. Use of nitrate by strain 2CP-3 at concentrations of 2.5 and 5 mM.
Mass balance shows that most of the nitrate is converted to NH4 ................ 58

Figure 3.4. Dechlorination as indicated by phenol accumulation from 2-CP and
nitrate utilization by 2CP-3. Arrows indicate when 2-CP was completely
dechlorinated and when new additions of 2-CP were made ......................... 59

Figure 3.5. Fraction of electron (to) from acetate used for reductive
dechlorination of 2-CP by strain 2CP-C. The to is derived from the slope of
the regression line generated by plotting the acetate consumed as H2
equivalents versus the umoles 2-CP
dechlorinated ........................................................................................................... 61

Figure 3.6. Consumption of H2 by 2CP-C under electron acceptor limited
conditions. Arrows indicate when 2-CP was added to the medium ............. 62

Figure 3.7. Uptake of H2 by 2CP strains in the presence of excess 2-CP. Inset
shows the threshold concentration of H2 for this culture ................................. 63

CHAPTER lV
Figure 4.1. Phase-contrast photomicrograph of strain 0023 which shows
vegetative cells and the presence of spores and terminal swelling in cells
developing spores. Reference bar is 13 um ..................................................... 81

Figure 4.2. Growth of strain C023 as indicated by an increase in turbidity (OD
600 nm). Duplicate cultures with lactate and 3-Cl-4-HBA are shown
compared to a control culture containing lactate alone ................................... 83

Figure 4.3. Graphical determination of to, the fraction of electrons from the
electron donor going to the electron acceptor. Electrons from lactate are
plotted as umole equivalents of H2 generated in the incomplete oxidation to

acetate ...................................................................................................................... 84

xiii

 

Figure 4.4. Growth of strain C023 on different electron acceptors and lactate as
measured by increases in turbidity (OD 600 nm). The symbols are as
follows: sulfur (El), sulfite (A), thiosulfate (A), sulfate (X), 3Cl-4-
hydroxyphenylacetate (O), 3Cl-4-HBA (O), and lactate (7). Data are the
average of duplicate cultures ............................................................................... 88

Figure 4.5. Exponential growth of strain C023 on pyruvate at 30° C. Data are
averaged from triplicate cultures .......................................................................... 90

Figure 4.6. Temperature dependence for growth and dechlorination by strain
C023. Growth rates were determined by averaging the Ilmax for duplicate
cultures. Dechlorination rates were determined with resting cells grown on
pyruvate at 37° C and incubated at various temperatures with SCI-4-HBA
and pyruvate. Also shown are the umoles 0f 3Cl-4-HBA dechlorinated over
23 h ............................................................................................................................ 91

Figure 4.7. Maximum likelihood phylogenetic tree. Numbers at internal nodes
are the percent of 100 bootstrap samples in which the group to the right of
the node was monophyletic. Scale is in expected number of substitutions
per positions ............................................................................................................ 94

CHAPTER V
Figure 5.1. Schematic representation of the dechlorination pathway of 3—CP to
phenol and subsequent tranforrnation to benzoate and methane. The
ortho-dechlorination of 2,3—DCP is shown in the lower left and is converted
completely to 3-CP prior to further degradation .............................................. 113

Figure 5.2. Effect of inhibitors on dechlorination in a methanogenic enrichment
culture. (A) Cumulative 3-MCP dechlorination in cultures with VFA mixture
(A), VFA plus BESA (O) and VFA plus vancomycin(l:l). (B) Methane
production in the same cultures. Initial culture volume was 20 ml. Brackets
are 95% confidence intervals ............................................................................. 114

xiv

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Figure 5.3. The dechlorination of 3-CP in cultures incubated with (A) and
without (8) 3-CP for 57 days, and grown on VFAs. 3-CP with or without
BESA was added at 57 days of incubation t0 VFA- and phenol-fed cultures.
(A) shows the onset of dechlorination in the control culture grown on VFAs
plus 3-CP upon the addition of 3-CP as designated by the arrow. (B) shows
the onset of dechlorination in cultures incubated without 3-CP with the
arrow indicating the addition of 3—CP and BESA. Brackets are the 95 %
confidence intervals ............................................................................................. 115

Figure 5.4. Cumulative dechlorination of 2,3-DCP. Ortho dechlorination (El)
and ortho plus meta dechlorination (O). (A) has no BESA and (B) has
BESA added. Brackets are 95% confidence intervals ................................. 117

Figure 5.5. Effect of electron donor on the efficiency of reductive dechlorination
of 3-CP as measured by the proportion of reducing equivalents used for

dechlorination ....................................................................................................... 118
CHAPTER VI
Figure 6.1. Uptake of H2 by CM enrichment culture and cumulative production
of phenol from the dechlorination of 2- and 3-CP .......................................... 134

Figure 6.2. (A) Dechlorination 0f 2,5-DCP by OM enrichment culture. Phenol
accumulation is also shown. (8) Appearance of 2- and 3-CP as
intermediates of 2.5-DCP dechlorination and the cumulative H2 uptake in

the OM culture ....................................................................................................... 136

Figure 6.3. Production of H2 from formate-fed OM enrichment cultures and the
dechlorination of of 2.5-DCP as indicated by the appearance of phenol..139

Figure 6.4. The fraction of electrons (to) used for dechlorination in the OM
enrichment culture. The to is equivalent to the slope of the regression line
of the plotted data. The 95% confidence interval was calculated for six
replicate cultures. Dechlorination activity started after an average of 102
moles of H2 had been consumed (arrow) ..................................................... 140

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Figure 6.5. Hydrogen uptake and threshold concentrations by the OM
enrichment culture. Inset has log scale to show the lower final H2 threshold

concentration ......................................................................................................... 142

Figure 6.6 Schematic representation of dechlorination pathways of OM
enrichment culture. Heavier arrows represent the slightly favored reactions,
thus ortho-dechlorination occurs slightly faster than meta-
dechlorination ....................................................................................................... 143

CHAPTER Vll

Figure 7.1. The relationship between the H2 partial pressure and the AG' for
formate oxidation (CI) (HCOO' + H2O --> HCOa' + H2) and for
methanogenesis (Q) (4 H2 + HCOa' + H+ --> 3H2O + CH4). Initial
concentrations are: formate = 0.01 M, HCOa' = 0.01 M and CH4 = 10 kPa.
The central shaded area shows the H2 concentration range that would
support the growth of a formate oxidizing organism and a methanogen
consuming the H2 in a simple anaerobic community. The adjacent flanking
shaded regions indicate the extended H2 concentration range that
corresponded to the AG' associated with the growth of FOX1 alone .......... 154

Figure 7.2. A. Growth of anaerobic tomato oxidizing bacterium FOX1 in broth
culture at 30° C as measured by direct microscopic counts of acridine
orange stained cells. A known volume of culture was stained and passed
through a 0.2 micron filter. Stained cells were then observed under
fluorescent illumination and enumerated. B. Illustrates the loss of formate
from cultures and the subsequent stoichiometric production of H2. Culture
volume was 100 ml with a headspace of 60 ml. Error bars indicate the 95%
confidence intervals for triplicate cultures ........................................................ 156

Figure 7.3. Fluorescent microscopic image of acridine orange stained FOX1
bacterium after growth on formate. The average size of cells in the culture
is2umwideand3to10umlong ...................................................................... 158

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Figure 7.4. 16S rRNA phylogeny of FOX1 and related organisms. Near
complete (ca. 1500 nucleotide positions) 16S rRNA genes were aligned
using the primary and secondary structure with sequences of the nearest
relatives obtained from the Ribosome Database Project (RDP).
Phylogenetic relationships were inferred by maximum likelihood analysis
(15) .......................................................................................................................... 159

Figure 7.5. Hydrogen production and free energy change in duplicate cultures
started at three formate concentrations: 15 mM(O), 30 mM(Cl) and 60
mM(A). A. Graphically represented is the average partial pressure for H2
in duplicate cultures. Hydrogen concentration reached a steady state in all
cultures after 20 days. B. Free energy AG‘ is shown for the same cultures
with different initial formate concentrations. The AG' was calculated by
measuring the H2 partial pressure and determining by difference the
tomato and bicarbonate concentrations. Formate was measured initially
and at the final data point to verify the free-energy calculations. All three
cultures converge at thermodynamic equilibrium at 20 days. ..................... 161

xvii

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Chapter I
INTRODUCTION

Overvlew

During the mineralization of xenobiotic compounds, microbial
communities require a terminal electron acceptor in order to grow and capture
energy from catabolic reactions. In addition to oxygen; nitrate, ferric iron,
halogenated organic compounds, sulfate, carbon dioxide and other organic
compounds have been shown to act as electron acceptors for growth in
bacteria. In the environment microorganisms utilize these electron acceptors in
accordance with their relative abundance and the energy available from their
reduction (Lovley and Goodwin 1988; Lovley et al. 1994). Although
chlorophenolic compounds are common contaminants and their degradation
under aerobic, methanogenic and sulfidogenic conditions has been well
studied, only a few studies have investigated dechlorination and degradation
under denitrifying conditions (Genthner et al. 1989; Haggblom et al. 1993).
Since it has been shown that nitrate can act as a terminal electron acceptor
during growth on xenobiotic compounds under anoxic conditions (Braun and
Gibson 1984; Bossert et al. 1986; Hutchins et al. 1991; Kaiser and Bollag 1991;
Haggblom et al. 1993), it would be reasonable to look for denitrifiers able to
degrade chlorophenols. Since reductive dechlorination of chlorophenols is
energy yielding under anoxic conditions,it may provide a benefit to a particular

population of denitrifiers or to the microbial community.

torme

Thep

1
presenc
receiver
dichloro
these m
2.3.dichl
culture 5
MicrocoS
pathwayE
fOcuseu
analyzing
T cl
From The
Capable
haloresDir

 

Sucre Torr;

redUchVe;'
'l

2

The objectives of this study were rooted in the goals established for the
former Novel organisms thrust group within the Center for Microbial Ecology.
The primary questions that this study addressed are as follows:

0 Do denitrifying microbial populations exist that can dechlorinate and mineralize
chlorophenols?

0 What are the microbial populations responsible for reductive dechlorination of
chlorophenols?

- Of what energetic value is the dechlorination activity to the microbial populations
within the chlorophenol degrading anaerobic community?

0 What is the influence of electron donors on the flux of energy within the
anaerobic food-web? Particularly what is the energetic relationship between
formate and H2?

To study the potential for dechlorination under anoxic conditions in the
presence of nitrate, I set up microcosms containing different soils. Microcosms
received low concentrations of nitrate and mixtures of monochlorophenols or
dichlorophenols. Both dechlorination and denitrification were observed in
these microcosms, with complete degradation of 2-chlorophenol (CP), 3-CP,
2,3-dichlorophenol (DCP), and 2,5-DCP occurring. My results from enrichment
culture studies indicated a complex anaerobic food-web was operating in the
microcosms, with at least five different dechlorinating populations, and two
pathways of phenol degradation: methanogenic and denitrifying. I then
focused on characterizing the different dechlorinating populations and
analyzing the energy flow in this anaerobic food web.

I chose four different dechlorinating enrichment cultures for further study.
From these cultures I isolated and characterized two new microorganisms
capable of coupling growth to reductive dechlorination, referred to as
halorespiration. These consisted of two new strains of the 2-CP halorespiring
Anaeromyxo dehalogenans , the first anaerobic Myxobacteria, and a novel
spore forming microorganism, named Desulfitobacterium halorespiricans, that

reductively dechlorinated 2,3-DCP and 3-Cl-4-hydroxybenzoate. A third

«er-4 “a“

 

dechlorina'
dechlorinat
completed
and if the l
dechlorinati
ewdence r
dechlorinat.
Chlorines frt
source of er
but was st:
dechlorinattc
medium resU
Since
TOGO-web CO
microbes lik.
ecosiStem. I.
With acetate ;
to The pTOdUC4
FOXt Was t

demortslrated

 

TEVEaled TWO
IOTa mlCTOOrg

olhamic Tole f

Biodeg
lemmas are

3

dechlorinating culture I studied was an enrichment that exhibited meta-
dechlorination of 3-CP under methanogenic conditions. Experiments were
completed to evaluate what electron donors were important for dechlorination
and if the methanogen population was important for this activity. The fourth
dechlorinating enrichment culture I selected yielded no isolates but exhibited
evidence of halorespiration activity. This OM (for ortho- and meta-
dechlorinating) enrichment culture had the novel ability to remove both
chlorines from 2,5-DCP simultaneously while using H2 and CO2 as the sole
source of energy and carbon. Formate could also serve as an electron donor,
but was stoichiometrically converted to H2 before the onset of reductive
dechlorination. My efforts to cultivate the dechlorinating culture on solid
medium resulted in the isolation of a novel H2 producing microorganism FOX1.
Since H2 was potentially an important intermediate in the anaerobic
food-web containing the dechlorinating populations, l evaluated the role
microbes like the formate-oxidizing FOX1 might have in the anaerobic
ecosystem. I determined that FOX1 was actually able to grow on formate alone
with acetate added for incorporation into biomass. Formate oxidation coupled
to the production of H2 was the sole source of energy for this microorganism.
FOX1 was the first organism for which this type of growth has been
demonstrated. An analysis of the free-energy associated with this process
revealed two other novel features, FOX1 grew at the lowest AG' ever observed
for a microorganismand it catalyzed this reaction to thermodynamic equilibrium.
The existence of microbial populations like FOX1 suggests a more energetically

dynamic role for formate cycling in anaerobic ecosystems.

Background
Blodegradatlon Under Denltrlfylng Conditions. Biodegrading

denitrifiers are underinvestigated relative to their potential practical importance

in bicremeq
bowed
environmen
evidence th.
groundwater
Tradi
or aromatic.
Much of the
cxygenases
CO-lhetaboli
dichloroethe
Anaerobic t
dechlorinat;
observed in
Reineke and
T€$ult The DIE
tor aromatic
Tanslormed'
Studies have
mediate The
1992; Files
focused on t
WmOfls in
Chloru
hhsidered w
The mam 0r

 

4

in bioremediation. Nitrate is potentially an attractive alternative to oxygen
because of its solubility, mobility, e' accepting capacity, and low toxicity in the
environment. Recent field studies support this strategy and have yielded
evidence that nitrate addition stimulated removal of BTEX from contaminated
groundwater (Hutchins et al. 1991).

Traditionally, the degradation of xenobiotic compounds, chlorinated and
or aromatic, has been evaluated under aerobic or strictly low redox conditions.
Much of the aerobic microbial research has focused on oxygen requiring
oxygenases involved in aromatic or methane oxidative pathways as well as on
co-metabolism of chlorinated solvents like trichloroethene (TCE) and
dichloroethene (DCE) (Chaudry and Chapalamadugu 1991; Hardman 1991).
Anaerobic biodegradation research has primarily focused on reductive
dechlorination of highly substituted halogenated organic compounds, as
observed in methanogenic and sulfidogenic enrichments (Suflita et al. 1982;
Reineke and Knackmuss 1988; Hardman 1991; Mohn and Tiedje 1992). As a
result the prevailing scientific opinion had been that oxygenases were required
for aromatic ring cleavage and that highly chlorinated chemicals only were
transformed, but not mineralized, under strictly anaerobic conditions. Recently,
studies have identified denitrifying and sulfate-reducing populations that can
mediate the anaerobic oxidation of toluene (Evans et al. 1991; Better at al.
1992; Fries et al. 1994). This work has encouraged further research that
focused on the degradation potential of denitrifiers and the moderate redox
conditions in which they thrive.

Chlorinated aromatic compounds have properties that must be
considered when considering their degradability under denitrifying conditions.
The major one is the degree of chlorination and its impact on the catabolic

pathway. For example, can a moderately chlorinated (2-3 CI) aromatic

 

 

compound "
pathway u
compounds
as they cor.
requiring r
Knackmuss
and Tiedje
Phen
extensivelyl
pollutants. l
volatility.
Potential SI
biodegradai
halogenater
al. 1990; D
teasgmamE
competing
haloQEnatE
Nat
research '

5

compound be degraded via a pathway similar to the toluene degradation
pathway under denitrifying conditions? If this is possible, then these
compounds could be degraded by pure cultures under denitrifying conditions,
as they could presumably follow the dechlorination steps used by non-oxygen
requiring reactions found in aerobic or anaerobic pathways (Reineke and
Knackmuss 1988; Chaudry and Chapalamadugu 1991; Hardman 1991; Mohn
and Tiedje 1992).

Phenolic compounds, especially chlorinated ones, have been used
extensively in industry and therefore have become common environmental
pollutants. Most are relatively soluble under the right conditions and have low
volatility. This makes them very mobile in aqueous environments and a
potential source of groundwater pollution. Some of the early work on
biodegradation under denitrifying conditions has focused on similar, but non-
halogenated phenolic compounds, especially cresols and phenol (Haggblom et
al. 1990; Dangel et al. 1991; Rudolphi et al. 1991; Seyfried et al. 1991). It is
reasonable to predict that degradation pathways of chlorinated phenolic
compounds may be related to those previously identified in similar non-
halogenated compounds.

Naturally occurrlng halogenated compounds. Although much
research has focused on the biodegradation of xenobiotic chlorinated organic
compounds, there has been little awareness of the existence of natural
halogenated compounds. It is the naturally biogenic or geogenic halophenolic
compounds that are probably responsible for the microbial mediated catabolic
activities that are utilized today for the bioremediation of xenobiotic analogs to
those compounds. In a recent review Gribble (1992) described hundreds of
naturally occurring halogenated compounds. Some of these are halogenated

phenolic compounds, analogous to the chlorophenols that have been used for

 

extensive a
natural cor
environme'
mg kg'1 of
(de Jong e
diverse en
bromophen
tyrosine ha
mollusks(G
the reason;
evolved the

conditions.

 

organisms,
culture has.

Redt
dehalogena
anaerobic er
al. 1992), ;
isomers the
that aTiaerol
CO’llltTEIely
Show To Se
as Asap), an
Even TITOSE
Show” to Cc
a1'1995l. F

was Seleclet

6

extensive aerobic and anaerobic biodegradation research (Figure 1.1). These
natural compounds are sometimes present in fairly high concentrations in the
environment. For example, a white-rot fungus was shown to produce up to 70
mg kg'1 chlorinated anisyl metabolites (e.g. 3-chloroanisaldehyde) in soil litter
(de Jong et al. 1994). Compounds shown in Figure 1.1 are found in very
diverse environments, from the sex pheromone of the lone star tick to the
bromophenolic excretions of marine tube worms. Chlorinated and brominated
tyrosine has even been shown to be incorporated into certain proteins in
mollusks(Gribble 1992). The ubiquitous nature of these compounds leads to
the reasonable hypothesis that considerable microbial diversity would have
evolved the ability to degrade these compounds under anaerobic and aerobic
conditions. This has been demonstrated with bromophenoI-producing marine
organisms, from which anaerobic degradation has been observed and a pure
culture has been isolated (King 1988; Steward et al. 1995).

Reductive dehalogenatlon of chlorophenols. Reductive
dehalogenation of chloroaromatic compounds has been well studied in
anaerobic ecosystems (Suflita et al. 1982; Mohn and Tiedje 1992; Nicholson et
al. 1992). As an example, the dechlorination pathways for four trichlorophenol
isomers and six dichlorophenol isomers are shown in Figure 1.2. It is evident
that anaerobically these substrates have been observed to be dechlorinated
completely to phenol and 4-CP. Several of the chlorophenols have been
shown to serve as terminal electron acceptors in microbial cultures, illustrated
as heavy arrows in Figure 1.2 (Cole et al. 1994; Utkin et al. 1994) (this thesis).
Even those reactions that appear to support higher cell yields, have been
shown to cometabolically dechlorinate more complex chlorophenols (Utkin et
al. 1995). For example Desulfitobacterium dehalogenans strain JW/IU DC1

was selected in enrichment cultures that exhibited ortho-dechlorination of 2,4-

F "493.14
I

 

2,5

(nose

 

3-chlr

 

Takefiln

 

OH OH
Cl or CI
2,6-dichlorophenol . 9'
(lone star tick) 2,4-drchlorophenol
(Penicillium)
OH CH
Cl Br r
CI
2,5-dichlorophenol 2,6-dibromophenol
(grasshopper salivary excretions) (marine hemichordate)
OH O=CH
Br or
Cl Cl
NH2 OH
900' _ 3,5-dichloroanisaldehyde
3-chloro-5-bromotyrosrne (whimomngus)
(mollusce)

 

Figure 1.1. Several naturally occurring halogenated phenolic compounds.
Taken from Gribble (1992) and de Jong (1994).

 

 

2.6-DCP

 

2-CP

9‘"-

 

F'Tdte 12.

8

 

( 0H 0H 0H OH R
G 0 CI
0 G a do
0 a a
145-70" 2.4,5-TCP 3,4541”: 2.3.5739

\ /r .../L

\
ii)

OOOH
L MON-benzoate j

 

 

 

Figure 1.2. Anaerobic degradation pathways observed for trichlorophenols.
Heavy arrows denote reductive dechlorination reactions known to support
growth via halorespiration. Arrows with dotted lines indicate transformations
that are likely to occur and arrows with thin-lines indicate reactions that have
been observed in anaerobic systems.

 

   
  

DCP' an
dechlotlna
dechlorinh
renewed b:

the

ills guide“t
mmarate re
ability to ”l
been four“
the first TSC‘ .
reductive (35'
dee 199l
phylogenellcl
[Cole et al- I
exhibited ir
hydroxypherl
Non-aromatic]
support halCI
anaerobic PC!
when H2 WE'

elect

iOTl ace“-

l
Doha/ospr'ri/hll

stain exhibits

 

9

DCP, and it has been shown that when induced this strain can also
dechlorinate pentachlorophenol (PCP) to 3,4,5-TCP. In general ortho reductive
dechlorination of chlorophenols is the most common reaction observed
followed by meta- and para-dechlorination (Nicholson et al. 1992).

The free-energy, AG, associated with reductive dechlorination of
chloroaromatic compounds is significantly exergonic (Dolfing and Harrison
1992), very close to the AG°' of nitrate reduction to nitrite (Table 1.1). A
comparison of the AG°' associated with various electron-accepting reactions are
shown in Table 1.1. When coupled to the oxidation of H2 as an electron donor
it is evident that reductive dechlorination potentially provides more energy than
fumarate reduction, sulfate reduction or methanogenesis. Halorespiration is the
ability to utilize the energy from this reaction for growth. This process has only
been found in a few microorganisms. Desulfomonile tiedjei strain DCB-t was
the first isolated bacterium shown to grow and couple ATP generation to the
reductive dechlorination of 3-chlor0benzoate (3-CBA) (Dolfing 1990; Mohn and
Tiedje 1990). More recently an anaerobic 2-CP halorespiring microbe
phylogenetically related to the myxobacteria was isolated and characterized
(Cole et al. 1994). Desulfitobacterium dehalogenans strain JW/IU DC1
exhibited increased cell-yields when it was dechlorinating 3-chloro-4-
hydroxyphenylacetate with pyruvate as the electron donor (Utkin et al. 1994).
Non-aromatic compounds such as tetrachloroethene (PCE) are also known to
support halorespiration. For example, Holliger et al. (1993) described the
anaerobic PCE dechlorinating microbe Dehalobacter restrictus that only grew
when H2 was provided as an electron donor and PCE was added as an
electron acceptor (Holliger and Schumacher 1994). PCE was also utilized by
Dehalospirillum multivorans as a terminal electron acceptor, however this

strain exhibited a much broader range of substrates that would support growth

I " '4" 'l:

 

Table 1.1
phenol ar
butyrate al
or calcula:
Griddle et

 

e accept
N20 + 2H*
Q + 4H“ +
“03' +6H+
to + M
Cdvcou .
We? +
504'2 + 9H+
”003' + are
2l-c03- + 9H

”We ...

l
l
l
|

i donating-
it» 2H+
hoe . H20
Witches»

\

 

10

Table 1.1 Relevant half-reactions for reductive dechlorination of monochloro-
phenol and other electron acceptors. Half-reactions for formate, H2 and
butyrate as electron donors are also shown. AG°' values shown where known
or calculated from theoretical thermodynamic calculations (T hauer et al. 1977;

Criddle et al. 1991; Dolfing and Harrison 1992).

Hallzteacticn
e- acceptlng reactions:

 

 

N2O+2H++20-—->N2+H2O
02+4H++4e'-—-—>2H2O
N03'+6H++5e--->1/2N2+3H2O
N03'+2H++20--->N02'+H2°
Cal-I40IOI-I+2H++Ze-—>06H§OH+HCI

Funarate'2 + 2 H+ + 2 e' > succinate'z

 

304-2 + 9H+ + 8e' --—> HS‘ + 4H2O
I-I003‘+8H++8e‘ --->CH4+3H20

2H003’ + 9H+ + 8e' ---> Acetate' + 4H2O

2H" 4» 2e' -——> H2

e- donatlng reactions:

H2 -—> 2H+ + 29"

HOOO' + H20—>H003' + 2H+ + 2e'

CI-I3CH2CH2000' + 2H20 ---> ZCH3000‘ 4» SW” + 46'

+21

+23

+27

+40

-39.4

-28

 

 

(Neumar
The halo
diversity.
respirator
Ma
anaerobic
energetic
polychlori
methanog
equivalent
(Bedard ar
mediated j
predominar
ltductive
dechlorinati
and dechlor
was added
was TBSDOn.
inhibited her
Since bOlh
reClllctive de
Occurring in l

Was Diesen:

 

 

and acetcge

11

(Neumann et al. 1994; Neumann et al. 1995; Schlolz-Muramatsu et al. 1995).
The halorespiring strains described represent a wide-range of phylogenetic
diversity, suggesting that the ability to utilize halogenated compounds as
respiratory electron acceptors has perhaps evolved many times.

Many of the dechlorination reactions of chlorophenols observed in
anaerobic environments and enrichments do not appear to be linked to any
energetic benefit to a particular microbial population. The dechlorination of
polychlorinated biphenyls often has been associated with at least some
methanogenic activity, where methane accounts for most of the reducing
equivalents from electron donors consumed in these anaerobic cultures
(Bedard and Quensen Ill 1995). Even though some evidence of methanogen
mediated para-dechlorination of PCB congeners has been discovered, the
predominant energy yielding reaction in most PCB enrichment cultures was not
reductive dechlorination. One recent study showed that reductive
dechlorination of chlorophenols continues in the absence of methanogenesis
and dechlorination ceased when the eubacterial specific antibiotic vancomycin
was added (Perkins et al. 1994). This suggested that a eubacterial population
was responsible for dechlorination; however, even when methanogenesis was
inhibited homo-acetogenesis became the major electron sink for the ecosystem.
Since both methanogenesis and acetogenesis yield less free-energy than
reductive dechlorination it is apparent that halorespiration is not likely to be
occurring in these anaerobic enrichment cultures. If a halorespiring population
was present, it should have a competitive advantage over the methanogens
and acetogens present in the community and eventually would displace the
other microbial populations. When this doesn't occur, alternative mechanisms
should be considered. First it is possible that the reductive dechlorination is a

fortuitous activity that is beneficial to the anaerobic food-web by serving as a H2

sink.
energeti
channel
organic
halorest
is not a
could b
enrichm-
isomers
E
H2. T
syntrOpt
exergcm
caliable
SYHtrOpr
AG°' for
(Table 1
by COHSL
e<TUilibrii
f0”haste
1989; Sc
carrier at
thereiOre
H2 Drody
Wehtiar
are in eq

diseituilit

12

sink. For example, obligate H2 generating syntrophs would benefit
energetically if some of the excess reducing equivalents they produce were
channeled toward the dechlorination of chlorophenols or other halogenated
organic compounds instead of to H2 synthesis. This is similar to
halorespiration, except the energy from the reductive-dehalogenation reaction
is not available to the microorganism. Alternatively reductive dechlorination
could be a detoxification mechanism, however the specificity exhibited in
enrichment cultures for these reactions and the persistence of some chlorinated
isomers suggest that this might not be the case.

Energy flow In anaerobic food webs: the role of formate and
H2. The oxidation of small chain organic acids is largely mediated by
syntrophic microorganisms in anaerobic environments. These reactions are
exergonic only under low H2 concentrations, thus a microbial population
capable of consuming H2 to low concentrations must be present in order for the
syntroph to grow and survive (Schink and Friedrich 1994). For example the
AG°' for butyrate oxidation coupled to Hit reduction to H2 is + 12 kJ/ mole 9'
(Table 1.1), therefore butyrate will only support growth if the AG' can be lowered
by consuming the H2 produced. Fomato, however, is close to thermodynamic
equilibrium with H2 and it has been proposed that syntrophs may rely on
fomate as an electron carrier instead of H2 (Thiolo ot al. 1988; Boone ot al.
1989; Schink and Friedrich 1994). The arguments for fomate as this electron
carrier are that it is more soluble and has a higher diffusion coefficient than H2,
therefore energy will be dissipated more quickly from a tomato producer than a
H2 producer. A major assumption to the energetic equivalency of those two
potential electron carriers is that under physiological conditions tomato and H2
are in equilibrium. Anaerobic aquifers have been shown to have an obvious

disequilibrium between formate and H2 (McMahon and Chapollo 1991), with

 

tomato 01
not likely
microorga
Fort
and serve
acetogene
equated to
been some
two compc
experiment
produced b;
(Schauer at
shown to he
the Substrat.
1973i Hunt
CO2 by non.
are one of th
Suggested the
rumen (Hun 96
do not behave

 

13

fomate oxidation favored thermodynamically. In this type of environment it is
not likely that formate is the primary electron carrier for syntrophic
microorganisms.

Fomato is a common intermediate in anaerobic degradation processes,
and serves as a substrate for many terminal anaerobic processes such as
acetogenesis, methanogenesis and sulfidogonosis. Although fomate often is
equated to H2 in anaerobic microbial ecosystems (Stams 1994), there have
been some interesting observations made in the past that suggest that those
two compounds are not equivalent as energy substrates. In comparative
experiments using tomato or H2 + 002 the yield of cells per mole of methane
produced by Methanobactorium formicicum was found to be greater for fomate
(Schauor and Ferry 1980). The sulfate-reducer Dosulfovibrio vulgaris was
shown to have a higher coll-yield per mole 804‘2 reduced when fomate was
the substrate instead of H2 + 002 (Badziong and Thauor 1978; Magoo ot al.
1978). Hungato et al. (1970) suggested that formate was oxidized to H2 and
CO2 by non-methanogenic microorganisms in the rumen whore mothanogons
are one of the predominant groups of organisms present. In addition, it was
suggested that H2 and 002 were converted to methane and not fomate in the
rumen (Hungato 1967). All of these observations indicate that formate and H2
do not behave equivalently in the environment, and that there may be energetic
differences associated with their metabolism by microorganisms. Perhaps the
most convincing evidence of the differences between tomato and H2 is alluded
to in data presented by Stephenson and Stickland (1933). They report the
conversion of fomate into methane by a putative pure culture, however the
same culture is reported to reduce sulfate to sulfide with H2. Since there are no
known methanogons that are also sulfidogons it is likely this was a coculture

between a sulfate reducer and tho methanogen. Formate would then support

 

 

 

the g

CORSU

readily

shown

genera

Multiple

both cy
propose
periplasr
1992). '
involved

coupling i
iaCilltates
Out of the
can then ‘
earners, I
and are U:
A SCilemat
Suhidogen
Formaye is
have a 991
Will give a
membrane
elegymn dc
are “Sing t

590m in Till

14

the growth of two organisms in this community, where the methanogen
consumes the H2 produced by the sulfidogen during the oxidation of fomate.
Sulfate reducers, particularly Dosulfovibrio , are known to be able to
readily metabolize both formate and H2. This group of organisms has been
shown to possess a least three types of hydrogenases, involved in either H2
generation or consumption (Fauquo at al. 1988; Widdol and Hanson 1992).
Multiple hydrogenases have been identified in individual microorganisms, with
both cytoplasmic and periplasmic locations within the cell. It has been
proposed that cytoplasmic hydrogenases are involved in H2 production, while
periplasmic hydrogenases are involved in H2 uptake (Widdol and Hanson
1992). This positioning of hydrogenases in Dosulfovibrios is thought to be
involved in hydrogen cycling which serves as a general mechanism for energy
coupling in these microorganisms (Odom and Peck Jr. 1981). Hydrogen cycling
facilitates the formation of a membrane potential by utilizing the diffusion of H2
out of the cell as a proton transport mechanism. Hydrogenasos in the periplasm
can then oxidize the H2 to protons and reduce membrane associated electron
carriers. Electrons are then transported back across the membrane into the cell
and are used to reduce the teminal electron acceptor, for example thiosulfate.
A schematic of the enzyme localization and the electron flux across a theoretic
sulfidogen membrane demonstrating hydrogen cycling is shown in Figure 1.3.
Fomato is shown as the electron donor. Since at least some sulfate reducers
have a periplasmic location for their fomate dohydrogonaso, fomate oxidation
will give a not positive charge to the outside of the cell, thus enhancing the
membrane potential (Widdol and Hanson 1992). In the presence of suitable
electron donors, a considerable proton gradient can be established. If the cells
are using formate and producing H2, it is possible that H2-cycling could still

occur in the absence of electron acceptors, since high H2 concentrations will

15

INSIDE OUTSIDE

M H2 H2
@

K
j XH HT + 002
Ht
z-\ a

\

HCOO'

 

 

x H+
a; .. @
H S \
2 2H+
XHT‘ AT fi—J—HT'

x/3 ATP

3042-

 

 

 

 

membrane

Figure 1.3. Schematic representation of H2 cycling mechanism in a sulfate

reducing microorganism using fomate as an electron donor. FD = fomate
dehydrogenase; Hyd = hydrogenase; SR = sulfate reductases; ocR =
electron carrier reductase and AT: ATP ase. Adapted from Odom and Peck
(1981 ).

 

 

it.

not it
proton
some j
for slot
of bion

oxidatio

16

not inhibit fomate oxidation (Sanford, 1996, this dissertation). Therefore a
proton gradient from fomate oxidation can develop and even be enhanced with
some proton generating hydrogenaso activity outside the cell. One other sink
for electrons besides the cytoplasmic hydrogenaso, would be for the synthesis
of biomass, which implies that growth could possibly occur from formate

oxidation utilizing this type of mechanism.

References

Badzlong w. and R.K. Thauor. 1978. Growth yields and growth rates of
Dosulfovibrio vulgaris (Marburg) growing on hydrogen plus sulfate and
hydrogen plus thiosulfate as the sole energy sources. Arch. Microbiol.
117:209-214.

Bedard D.L. and J.F. Quensen Ill. 1995. Microbial reductive
dechlorination of polychlorinated biphenyls, p. 127-216. In L. Y. Young and
0. Corniglia (od.), Microbial transformation and degradation of toxic
organic chemicals. Wiley-Liss Division, John Wiley 8: Son, New York

Beller I-I.R., D. GrbIc-Gallc and M. Reinhard. 1992. Microbial
degradation of toluene under sulfate-reducing conditions and the influence
of iron on the process. Appl. Environ. Microbiol. 58:786-793.

Boone D.R., R.L. Johnson and Y. Llu. 1989. Diffusion of the intorspecies
electron carriers H2 and formate in methanogenic ecosystems and its

implications in the measurement of Km for H2 or fomate uptake. Appl.
Environ. Microbiol. 55:1735-1741 .

Bossert I.D., M.D. Rivera and L.Y. Young. 1986. p-Cresol
biodegradation undor denitrifying conditions: isolation of a bacterial
coculture. FEMS Microbiol. Ecol. 38:313-319.

Braun K. and DJ. Gibson. 1984. Anaerobic degradation of 2-
aminobonzoate (anthranilic acid) by denitrifying bacteria. Appl. Environ.
Microbiol. 48:102-7.

Chaudry G.R. and S. Chapalamadugu. 1991. Biodegradation of
halogenated organic compounds. Microbiol. Rev. 55:59-79.

Cole J.A., A. Cascarelli, w.w. Mohn and J.M. Tiedje. 1994. Isolation
and characterization of a novel bacterium growing via reductive
dechlorination of 2-chlorophonol. Appl. Environ. Microbiol. 60:3536-3542.

 

Cdddh
pm)
(ed
Dor

Dangel

Ddfing
ATP
hhcr

Dlon;
nah)
acce
2215

Evans |

degra
57A

FGUqUe
Derv

17

Criddle 0.5., L.M. Alvarez and P.L. McCarty. 1991. Microbial
processes in porous media, p. 641-691. In J. Bear and M. Y. Corapcioglu
(od.), Transport processes in porous media. Kluwor Academic Publishers,
Dordrocht

Dangel w., R. Brackman, A. Lack, M. Mohamed, J. Koch, B. Oswald,
B. Seyfrled, A. Tschech and G. Fuchs. 1991. Differential expression
of enzyme activities initiating anoxic metabolism of various aromatic
compounds via bonzoyl-CoA. Arch. Microbiol. 155:256-62.

de Jong E., J.A. Field, H.-E. Splnnler, J.B.P.A. lenberg and J.A.M.
de Bont. 1994. Significant biogenosis of chlorinated aromatics by fungi in
natural environments. Appl. Environ. Microbiol. 60:264-270.

Dolfing J. 1990. Reductive dechlorination of 3-chlorobonzoato is coupled to
ATP production and growth in an anaerobic bacterium, strain DOB-1. Arch.
Microbiol. 153:264-266.

Dolfing J. and B.I(. Harrison. 1992. Gibbs free energy of formation of
halogenated aromatic compounds and their potential role as electron
acceptors in anaerobic environments. Environ. Sci. Technol. 26:2213-
2218.

Evans P.J., D.T. Mang, K.S. Kim and L.Y. Young. 1991. Anaerobic
degradation of toluene by a denitrifying bacterium. Appl. Environ. Microbiol.
57:1139-1145.

Fauque G., l-I.D. Peck Jr., J.J.G. Moura, B.H. Huynh, Y. Berller, D.V.
DerVartanlan, M. Telxelra, A.E. Przybyla, P.A. Lesplnat, I. Moura
and J. LeGaII. 1988. The three classes of hydrogenases from sulfate-
roducing bacteria of the genus Dosulfovibrio. FEMS Microbiol. Rev.
54:299-344.

Fries M.R., J.-Z. Zhou, J.C. Choc-Sanford and J.M. Tiedje. 1994.
Isolation, characterization, and distribution of denitrifying toluene dograders
from a variety of habitats. Appl. Environ. Microbiol. 60:2802-2810.

Genthner B.R.S., WA. Price II and P.H. Prltchard. 1989. Anaerobic
degradation of chloroaromatic compounds in aquatic sediments under a
variety of enrichment conditions. Appl. Environ. Microbiol. 55:1466-1471.

Gribble G.W. 1992. Naturally occurring organohalogon compounds -- a
survey. J. Nat. Prod. 55:1353-1395.

Higgblom M.M., M.D. Rivera, I.D. Bossert, J.E. Rogers and L.Y.
Young. 1990. Anaerobic biodegradation of para-crosol under three
reducing conditions. Microbial Ecol. 20:141-50.

 

 

Hoiiige
res

Hungai
flue

Hungat
1974
Bad

Hutchin
DJ).
addi
demc

HUtChin:
Bicde
denhr

Kaiser J
hydmx

n

”"9 GM
SOUrce

Lovley D
caKNYZi

Way DJ:

'“dicato.
aquatic 5

18

Haggblom M.M., M.D. Rivera and L.Y. Young. 1993. Influence of
alternative electron acceptors on the anaerobic biodegradability of
chlorinated phenols and benzoic acids. Appl. Environ. Microbiol. 59:1162-
1167.

Hardman D.J. 1991. Biotransformation of halogenated compounds. Crit.
Rev. Biotechnol. 11:1-40.

Holliger 0., G. Schraa, A.J.M. Stams and A.J.B. Zehnder. 1993. A
highly purified enrichment culture couples the reductive dechlorination of
tetrachloroethene to growth. Appl. Environ. Microbiol. 59:2991-2997.

Holliger C. and w. Schumacher. 1994. Reductive dehalogenation as a
respiratory process. Antonie van Leeuwenhoek 66:239-246.

Hungato R.E. 1967. Hydrogen as an intermediate in the rumen fermentation.
Arch. Microbiol. 59:158-164.

Hungato R.E., w. Smith, T. Bauchop, I. Yu and J.C. Rabinowitz.
1970. Formate as an intermediate in the bovine rumen fermentation. J.
Bacteriol. 102:389-397.

Hutchins s.R., w.c. Downs, J.‘I'. Wilson, G.B. Smith, D.A. Kovacs,
D.D. Fine, R.H. Douglass and D.J. Hendrix. 1991. Effect of nitrate
addition on biorestoration of fuel-contaminated aquifer: Field
demonstration. Groundwater 29:571-580.

Hutchins s.R., G.W. Sewell, D.A. Kovacs and G.A. Smith. 1991.
Biodegradation of aromatic hydrocarbons by aquifer microorganisms under
denitrifying conditions. Environ. Sci. Technol. 25:68-76.

Kaiser J.P. and J.-M. Bollag. 1991. Metabolism of pyridine and 3-
hydroxypyridine under aerobic, denitrifying and sulfate-reducing conditions.
Experientia 47:292-296.

King G.M. 1988. Dehalogenation in marine sediments containing natural
sources of halophenols. Appl. Environ. Microbiol. 54:3079-3085.

Lovley D.R., P.H. Chapeile and J.C. Woodward. 1994. Use of
dissolved H2 concentrations to determine distribution of microbially

catalyzed redox reactions in anoxic groundwater. Environ. Sci. Technol.
28:1205-1210.

Lovley D.R. and S. Goodwin. 1988. Hydrogen concentrations as an
indicator of the predominant terminal electron-accepting reactions in
aquatic sediments. Geochim. Cosmochim. Acta 52:2993-3003.

 

Pefkins

RQineke
haIOa

RudDiphi
cr'980I

schaUer
Mei/ta

SW“ 3.
0dean

19

Magee E.L., B.D. Ensiey and LL. Barton. 1978. An assessment of
growth yields and energy coupling in Desulfovibrio. Arch. Microbiol.
117:21-26.

McMahon P.B. and P.H. Chapelle. 1991. Microbial production of organic
acids in aquitard sediments and its role in aquifer geochemistry. Nature
349:233-235.

Mohn W.W. and J.M. Tiedje. 1990. Strain DOB-1 conserves energy for
growth from reductive dechlorination coupled to formate oxidation. Arch.
Microbiol. 1 53:267-271 .

Mohn W.W. and J.M. Tiedje. 1992. Microbial reductive dehalogenation.
Microbiol. Rev. 56:482-507.

Neumann A., H. SchoIz-Muramatsu and G. Dlekert. 1994.
Tetrachloroethene metabolism of Dehalospirillum multivorans. Arch.
Microbiol. 162:295-301 .

Neumann A., G. Wohlfarth and G. Diekert. 1995. Properties of
tetrachloroethene and trichloroethene dehalogenase of Dehalospirillum
multivorans. Arch. Microbiol. 163:276-281.

Nicholson D.K., S.L. Woods, J.D. lstok and 0.0. Peek. 1992.
Reductive dechlorination of chlorophenols by a pentachlorophenol
acclimated methanogenic consortium. Appl. Environ. Microbiol. 58:2280-
2286.

Odom J.M. and H.D. Peck Jr. 1981. Hydrogen cycling as a general
mechanism for energy coupling in the sulfate-reducing bacteria,
Desulfovibrio sp. FEMS Microbiol. Lett. 12:47-50.

Perkins P.S., S.J. Komisar, J.A. Puhakka and J.P. Ferguson. 1994.
Effects of electron donors and inhibitors on reductive dechlorination of
2,4,6-trichlorophenol. Wat. Res. 28:2101-2107.

Reineke W. and H.J. Knackmuss. 1988. Microbial degradation of
haioaromatics. Annu. Rev. Microbiol. 42:263-287.

Rudolphi A., A. Tschech and G. Fuchs. 1991. Anaerobic degradation of
cresols by denitrifying bacteria. Arch. Microbiol. 155:238-48.

Schauer N.L. and J.G. Ferry. 1980. Metabolism of formate in
Methanobacferium fonnicicum. J. Bacteriol. 142:800-807.

Schink B. and M. Friedrich. 1994. Energetics of syntrophic fatty acid
oxidation. FEMS Microbiol. Rev. 15:85-94.

20

Schioiz-Muramatsu H., A. Neumann, M. MeBmer, E. Moore and G.
Dlekert. 1995. Isolation and characterization of Dehalospirillum
multivorans gen. nov., sp. nov., a tetrachloroethene-utilizing, strictly
anaerobic bacterium. Arch. Microbiol. 163:48-56.

Seyfrled B., A. Tchech and G. Fuchs. 1991. Anaerobic degradation of
phenylacetate and 4-hydroxyphenylacetate by denitrifying bacteria. Arch.
Microbiol. 155:249-55.

Stams A.J.M. 1994. Metabolic interactions between anaerobic bacteria in
methanogenic environments. Antonie van Leeuwenhoek 66:271-294.

Stephenson M. and L.H. Stickland. 1933. Hydrogenase. Ill. The
bacterial formation of methane by the reduction of one-carbon compounds
by molecular hydrogen. Biochem. J. 27:1517-1527.

Steward 0.0., ‘l'.0. Dixon, Y.P. Chen and 0.R. Lovell. 1995.
Enrichment and isolation of a reductively debrominating bacterium from the
burrow of a bromometabolite-producing marine hemichordate. Can. J.
Microbiol. 41:637-642.

Sufllta J.M., A. Horowitz, D.R. Shelton and J.M. Tiedje. 1982.
Dehalogenation: a novel pathway for the anaerobic biodegradation of
haioaromatic compounds. Science 218:1115-1117.

Thauer R.K., K. Jungermann and K. Decker. 1977. Energy
conservation in chemotrophic anaerobes. Bacterioi. Rev. 41:100-180.

Thiele J.H., M. 0hartraln and J.G. Zelkus. 1988. Control of interspecies
electron flow during anaerobic digestion: role of fioc formation in syntrophic
methanogenesis. Appl. Environ. Microbiol. 54:10-19.

Utkin l., 0.0. Dalton and J. Wlegel. 1995. Specificity of reductive
dehalogenation of substituted ortho-chlorophenols by Desulfitobacterium
dehalogenans JW/lU-DC1. Appl. Environ. Microbiol. 61:346-351.

Utkin l., 0. Woese and J. Wlegel. 1994. isolation and characterization of
Desulfitobacterium dehalogenans gen. nov., sp. nov., an anaerobic
bacterium which reductively dechlorinates chlorophenolic compounds. int.
J. Syst. Bacteriol. 44:612-619.

Wlddel F. and T.A. Hansen. 1992. The dissimiiatory sulfate- and sulfur-
reducing bacteria, p. 583-624. In A. Balows, H. G. Triiper, M. Dworkin, W.
Harder and K. H. Schleifer (ed.), The Prokaryotes. Springer Veriag, New
York

 

anaero
dichlorc
soils we
mM or 5
three di
a-DDFOpr
transfer
DEhEIOS

ShOwe-d

Chapter il

Chlorophenol dechlorination and subsequent degradation in
nitrate-limited denitrifying microcosms

Abstract

The potential of using nitrate as a terminal electron acceptor to stimulate
anaerobic degradation of mixtures of monochlorophenols (MCPs) or
dichlorophenois (DCPs) was evaluated. Contaminated and non-contaminated
soils were added to water-saturated anaerobic microcosms supplemented with 1
mM or 5 mM nitrate. Denitrification and dechlorination activities were present in
three diverse soil types and were maintained upon refeeding both nitrate and the
appropriate chlorophenol. However, dechlorination activity could only be serially
transferred in enrichments with an added electron donor such as acetate.
Dehalogenation activity in enrichments from four of the primary microcosms
showed at least five different dechlorination reactions, each mediated by different
microbial communities. Three of these are distinct ortho-dechlorinating paths;
two are meta-dechlorinating and one is the para-dechlorination of 3,4-DOP.
Simultaneous dechlorination and denitrification activities could be maintained in
microcosms only in the presence of low nitrate concentrations. These processes
were mediated by two separate microbial communities; one that dechlorinates
without use of nitrate and one that denitrifies while oxidizing the aromatic ring.
There was no evidence that dechlorination is mediated by the denitrifying
community, however the maintenance of a denitrification potential using low (< 1
mM) nitrate concentrations may be useful for completing the food chain by

stimulating the mineralization of phenol and benzoate.

21

 

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22
Introduction

Chlorophenolic compounds (CPs) have been used extensively in several
industries and there has been considerable effort to study the degradation of
these compounds under aerobic and strictly anaerobic conditions (Haggblom
1992; Mohn and Tiedje 1992), Several polychlorinated phenols and
monochlorophenols (MCPs) are degraded under aerobic and anaerobic
conditions (Boyd et al. 1983; Gibson and Suflita 1986; Hrudey et al. 1987;
Genthner et al. 1989; Dietrich and Winter 1990; Hale et al. 1990; Zhang and
Wiegel 1990; Madsen and Aamand 1992). The use of denitrifying conditions with
chlorophenols as substrates has been relatively underinvestigated, and none of
the previous studies have shown simultaneous dechlorination and nitrate
reduction (Genthner et al. 1989; Haggbiom et al. 1993). Relatively high nitrate
concentrations were used in these studies, which may have inadvertently
masked the potential of denitrifiers or dechlorinators acclimated to low nitrate
concentrations . Nitrate is attractive as an electron acceptor because of its
solubility and because denitrifying conditions promote complete mineralization of
some aromatic compounds at reasonably rapid rates. Also, reductive
dechlorination of chlorinated phenolic compounds, an exergonic reaction (Dolfing
and Harrison 1992), would not be inhibited under low nitrate concentrations as it
is by oxygen. Other studies have shown that dechlorination of MCPs does occur,
presumably once nitrate has been depleted, but efforts to transfer these cultures
to nitrate-containing media resulted in no dechlorination activity (Genthner et al.
1989). Nitrate-dependent chlorobenzoate-degrading enrichments, however,
have been observed (Genthner et al. 1989b; Haggblom et ai. 1993). This would
indicate that there may be differences between the microbial communities that

degrade these two types of compounds.

 

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SUbSeQUEn‘

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23

Anaerobic chlorophenol dechlorination studies have predominantly used
anaerobic sediments and sludges as inocula for enrichments or microcosms
(Boyd et al. 1983; Gibson and Suflita 1986; Genthner et al. 1989; Kohring et ai.
1989; Hale et ai. 1991; Hendriksen et ai. 1992; Haggblom et al. 1993). In
contrast, the best source of denitrifying populations may be surface soils where
the presence of nitrate and the fluctuating oxygen status provides regular
selection for denitrifiers. Soils high in litter residue may also harbor reservoirs of
naturally occurring chlorinated aromatic compounds. Several MOPs and
dichlorophenois (DCPs) are known to be produced naturally (Berger 1972; Siuda
and Debemardis 1973; Gribble 1992) and white rot fungi are known to produce
up to 75 mg/kg litter of chlorinated anisyl metabolites (0AMs) in nature(de Jong
et al. 1994).

We evaluated the degradation of mixtures of MCPs and DCPs under low
nitrate concentrations (< 5 mM) in microcosms seeded with contaminated and
relatively pristine soils. Mixtures were used to model the complex nature of
wastes that occur in contaminated material and to broaden the selective pressure
for dechlorinating microorganisms. Enrichment cultures were developed from
microcosms that exhibited both denitrification and dechlorination activity. Our
objectives were to determine the relationship of nitrate and denitrifiers to the
degradation activity and to evaluate the substrate range and specificity of the
dechlorination reactions mediated by these enriched populations. We observed
at least five different dechlorination reactions in these microcosms and

subsequent enrichments indicated they were carried out by different populations.

Materials and Methods
Source of microcosm inocula. Soils were obtained from several

sources. Material from contaminated sites was obtained from Remediation-

 

Technology
from a Pal
provided b]
Station wen
24-0. A c
compost oil.
the lowland
Mlcrc
ml of boiled
beads were
ofoMCP, m-
with 125 “M
and DCPs dr
125 NM. Niira
Nitrate was r
additions omy
were closed w
”Ediun
1992) was use

and per liter Ce

 

 

 

24

Technology and Ecova Corp. in Seattle, Washington. Sediment sludge samples
from a paper pulp mill effluent treatment system in Ontario, Canada were
provided by Dr. Roberta Fulthorpe. Soil samples from the Kellogg Biological
Station were obtained from agricultural plots being treated with different levels of
24-0. A compost soil, high in organic matter, was sampled at the base of a
compost pile in Lansing, Michigan. Two tropical soil samples were collected in
the lowland rainforest of equatorial Cameroon by Dr. Roland Weller.

Microcosms. Microcosms were started in 160 ml serum bottles with 100
ml of boiled degassed medium and approximately 10 g of soil or sludge. Glass
beads were used as a negative control. One microcosm set received a mixture
of o-MCP, m-MCP and p-MCP at 200 uM each. The other set was supplemented
with 125 pM each of 2,3-DCP, 2,4-DCP, 2.5-DCP and 3,4-DCP. Those MCPs
and DCPs degraded during incubation were replenished at a concentration of
125 pM. Nitrate (as KNOa) was provided at 5 mM or 1 mM to the microcosms.
Nitrate was replenished to initial levels once it was depleted. After several
additions only 1-2 mM nitrate was added to active microcosms; Serum bottles
were closed with Teflon lined butyi rubber stoppers and incubated at 30 ° 0.

Medium formulation. The following mineral salts medium (Stevens et al.
1992) was used for all cultures: 2 mM potassium phosphate buffer (pH 7.2 - 7.5),
and per liter CaCl2-2H20 (0.015 g), MgCl2-6H 20 (0.02 g.) FeSO4 -7H20 (0.007
g), and Naz 804 (0.005 g). A trace metals solution was added to give the
following final concentration per liter: MnCi2¢4H20 (5 mg), H3803 (0.5 mg),
ZnCl2 (0.5 mg), CoCl2-6H 20 (0.5 mg), NiSO4-6H20 (0.5 mg), CuCl2~2H 20 (0.3
mg), and NaMoO4°2H20 (0.1 mg). NH4CI was added to a concentration of 8
mM and 10 mM NaHCOa was added to buffer the headspace which contained a
N2:002 ratio of 95:5. A vitamin solution (Wolin et al. 1963) was provided after

cooling.

 

fresh i
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25

Enrichment cultures. Aliquots from microcosms were transferred to
fresh medium in serum bottles or anaerobic culture tubes with butyl rubber
stoppers for the purpose of enriching the dechlorinating and denitrifying activities.
Enrichments received nitrate at concentrations of 0, 1 or 5 mM. A subset of
those with and without nitrate received 1 mM acetate or a mixture of volatile fatty
acids (formate 625 (M, succinate 125 (M, propionate 125 pM, and butyrate 125
(M) as electron donors. The media were reduced with 0.2 mM cysteine and 0.2-
0.5 mM Na2 S. Resazurin was added as a redox indicator. Individual MCPs and
DCPs'were used as substrates at a concentration of 125 (M. Cultures were
incubated at 30 ° 0.

Chemical analysis. Nitrate and nitrite were analyzed by using a
Whatman Partisil 10 SAX column connected to a Schimadzu HPLC. The eluent
was 50 mM phosphate buffer (pH = 3.0) pumped at a rate of 1 ml/min. UV
adsorption at 210 nm was used for detection. Samples from primary enrichments
were diluted 1:100 in deionized H20 prior to nitrate and nitrite analysis.

MCPs, DCPs and aromatic products of dechlorination were analyzed on a
Hewlett Packard 1050 HPLC with a Chemstation analysis package. The eluent
was phosphoric acid (0.1 °/o) buffered methanol at 1.5 mein using a gradient
from 48% to 55% methanol. A Hibar RP-18 (10 um) column was used. Peaks
were quantified at 218 nm on a UV multiwavelength detector. Samples (1 mi)
from the enrichments were taken, made basic with 10 pl of 2N NaOH, centrifuged
for 6 min in a microfuge and filtered through Acrodisc L013 PVDF 0.45 pm filters
prior to HPLC analysis.

The headspaces of the microcosms and enrichments were analyzed for
CH4, N2, H2 and 002 using a Carie gas chromatograph equipped with a 1.83 m
Porapak 0 column and a thermal conductivity (T CD) detector. Headspace

pressures were normalized to atmospheric by venting with a needle prior to

 

 

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deté

Phei

micro
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contro.
simulta
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degrade
Comillet.
GC anal
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(Table 2'-
dechlorlna
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m' and or
Concentrali
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pfedOlTiinan.

26
removing 0.3 mi of gas for injection into the 00. N20 was quantified on a Perkin
Elmer 910 CC with a Porapak 0 column and 63Ni -eiectron capture (ECD)
detector. Denitrification products were measured in an Ar headspace.
Chemicals. MCPs and DCPs were obtained from Aldrich Chemical Co.

Phenol was purchased from Malinkrodt and benzoate from Sigma Chemical Co.

Results

Dechlorinating denitrifying microcosms. Only seven of the 30
microcosms had both dechlorination and denitrification activities after one year of
incubation (Table 2.1). No activity was observed with glass beads in the abiotic
control. Microcosms with MCPs exhibited either ortho-dechlorination alone or
simultaneously with meta-dechlorination. in contrast, DCP microcosms showed
considerable dechlorination from all positions, and this resulted in the
degradation of all the DCPs. Extensive gas production was observed indicating
complete reduction of nitrate to nitrogen gas (Table 2.1). This was confirmed by
GC analysis-of the headspace gases which showed increases of N2 and N20
after nitrate and nitrite depletion. Two sets of microcosms, Trop l and Trop II,
were very sensitive to the nitrate concentration, with both dechlorination and
denitrification occurring only when the nitrate concentration was 1 mM or lower
(T able 2.1). With the 5 mM Trop I microcosm, nitrate was slowly depleted and
dechlorination did not occur until nitrate had been removed.

The compost soil microcosm with 5 mM nitrate containing a mixture of o-,
m-, and p-MCP exhibited specificity for o-MCP and was sensitive to high nitrate
concentrations. This microcosm showed no evidence of m-MCP or p-MCP
disappearance over a 200 day period (Figure 2.1A). Phenol was the
predominant dechlorination product observed, but was rapidly removed in the

presence of nitrate (Figure 2.18). Benzoate was also detected at lower

Tabli
wrtm'r
nitratl

If

 

 

a *DC indlca
Occurred, aI
measuring l
f 0f - lndlca
pa’eilti‘leses

~03 . 1'5 91/9

 

 

27

Table 2.1. Summary of dechlorinating and denitrifying microcosms that were active
within a year of incubation. Microcosms received mixtures of MCPs and DCPs and
nitrate at 5 mM or 1 mM.

 

 

Sell only! sub-mm Denltrlflcatlon' Dechlorination" Products
Corrpost MCPS; N03'(5)° +DC ++(o-MCP) Phenol: benzoate
DCPs: N03'(5) +00 ++(aii DCPs) flagrficogm
Trop I MCPs; N03'(1) +DC ++(o-MCP) Phenol
MCPs; N03'(5) +D +(o-MCP) Phenol
Trop ll MCP8;N03’(1) +DC ++(o-MCP) Phenol
010 MCPs; N03‘(5) +0 ++(o-MOP, Phenol
m-MCP)
DCPB: N03'(5) +D +(aii DCPs) Phenol, m-MCP, p-MCP

 

 

 

 

 

 

 

‘ +00 indicates denitrification occurred concurrently with dechlorination. +D indicates that denitrification
occurred, and nitrate was depleted prior to any other activity. Denitrification was determined by

measuring N2 or N20 in the microcosm headspace after N03' and N02’ disappearance.

b + or - indicates presence of dechlorination acitivity. Substrates dechlorinated are indicated in
parentheses. ++ indicates a relatively higher level of dechlorination activity.

° N03“ is dven in (mM).

 

[phenol] (“Mi

28

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

180
-o-o-MCP
~+~m~MCP
135 --o--p-MCP
3.:
5 9° 1
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"'§ 50- w . 5 4.15
m l “ .
ft- 1 “ .. l l l l 3
0' _ WA) _4._‘_ \‘l .\.. 3.1. 0
o 5 100 150 m

Days

Figure 2.1. Degradation of chlorophenol in the MCP compost soil microcosm
showing ortho-dechlorination activity. o-MCP dechlorination and persistence of
m-MCP and p-MCP (A). Denitrification activity and transient appearance of
phenol. Filled arrows indicate addition of o-MCP (8). Open arrows indicate the
addition of fresh nitrate. Nitrite was not detected in the enrichment.

 

 

conc
nnra?
dech
addn
depka
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TM
dfBChlorin.
there Was

dechlorina

29

concentrations as a degradation intermediate in this microcosm. When excess
nitrate (7 mM) was amended to the microcosm on about day 55, the
dechlorination rate was considerably reduced. However, when subsequent
additions of only 2 mM nitrate were made to the microcosm after nitrate
depletion, both dechlorination and denitrification occurred concurrently. Other
microcosms exhibited similar sensitivity to nitrate, but at a lower concentration of
1 mM (data not shown). Nitrite was not detected in any of the denitrifying
microcosms.

The 010 microcosms with the MCP mixture dechlorinated both o-MCP
and m-MCP stoichiometrically to phenol (T able 2.1). This activity took between
140 and 200 days to develop and only occurred when nitrate was completely
depleted. Upon refeeding, both o-MCP and m-MCP were dechlorinated
simultaneously (Figure 2.2). The addition of nitrate after 380 days slowed the
dechlorination rate, but nitrate was rapidly consumed and N2 was detected in the
Ar-purged headspace. This indicated that an active denitrifying population was
present despite the absence of nitrate for nearly 300 days in the microcosm. The
persistence of p-MCP suggests that abiotic loss of CPs was not a factor in this
culture.

Two microcosms amended with mixtures of DCPs also showed
dechlorination activity and similar sensitivity to nitrate concentrations, however
there was little selectivity for which DCPs were dehalogenated. Several aromatic
dechlorination products were observed in the microcosms, including all of the
MCPs (T able 2.1). Phenol, o-MCP, m-MCP and benzoate appeared transiently
after several subsequent additions of the DCP mixture. Para-chlorophenol
continued to accumulate in the compost DCP microcosm, but to only 28% of the
theoretical concentration that would occur if no para-dechlorination of the isomers

had occurred.

 

"S

[Phenol] (HM)

 

30

150

 

 

120

 

[MCP] (1M)
8

8-

 

 

 

 

 

 

 

O

 

 

 

1200 ‘6

l
.5

(Wm) [612nm]

Nitrate \ 3
‘// ;

Phenol}

 

I
--l
N

 

 

 

 

 

+ fl-o-e—L—o—Q—o—J
. ~ 0
150 300 450 600
Days

 

[Phenol] (pM) ...
.° s as s s

Figure 2.2. Dechlorination of o-MCP and m-MCP in the 01C MCP microcosm.
(A) Concentrations of MCPs over 550 days of incubation. Closed arrows
indicating the addition of o-MCP and m-MCP. (B) indicates the concentrations
of phenol and nitrate as they accumulate and dissappear respectively.

 

 

 

 

of t
mine
teste
the a

wher

31

Independence of dechlorination from nitrate reduction. Since several
of the above experiments suggested dechlorination occurred prior to
mineralization and that nitrate above 5 mM was inhibitory to dechlorination, we
tested directly the effect of nitrate on dechlorination in enrichments derived from
the above microcosms. Degradation of MCPs to phenol occurred in enrichments
when acetate was supplied as an electron donor, however no degradation
occurred when nitrate was added. Figure 2.3A illustrates transfers of the 1 mM
nitrate microcosm to a fresh medium with and without nitrate. In the presence of
nitrate and no additional electron donors no degradation was seen. With acetate
as an electron donor, this o-MCP enrichment stoichiometrically dechlorinated o-
MCP to phenol. Phenol concentrations reached greater than 2 mM as long as
acetate was present. Similar results were obtained in enrichments inoculated
from other microcosms exhibiting o-MCP dechlorinating activity. When both
acetate and nitrate were present dechlorination activity also continued in these 0-
MCP enrichments (data not shown). In contrast, nitrate was inhibitory to all other
dechlorination activities, even in enrichments also containing acetate (Table 2.2).
Transfers of the DCP microcosms with meta-dechlorination activity to media
containing m-MCP with and without nitrate also showed the same response.
Meta-chlorophenol was dechlorinated to phenol with acetate present as an
electron donor. The culture with nitrate and m-MCP did not have any activity
(Figure 2.38). Meta dechlorination was considerably slower than ortho
dechlorination, taking almost 100 days. Similar results were obtained with
enrichments containing 2,3-DCP which exhibited ortho-dechlorination in the
presence of acetate. Subsequent meta-dechlorination was also observed in
these enrichments after a considerable lag time. In contrast, transfers done from
microcosms to a phenol plus nitrate medium showed complete phenol

degradation (Figure 2.30). These cultures exhibited extensive gas production in

E

[2-MOP] or [phenol] (pM)

 

[3-MOP] or [phenol] (11M)
8

[Phenol] (pM)
shame"

32

 

 

V V V

v I V

 

 

 

 

 

 

 

 

 

 

 

 

 

60
Days

   

 

100

 

 

 

 

120

Figure 2.3. Transfers of dechlorinating and denitrifying enrichments to media
with MCP plus 1 mM nitrate (A) or 1 mM acetate (0). Phenol (CI) is shown as a
product of dechlorination or the substrate for degradation. (A) Dechlorination
activity inc-MCP enrichment from Trop l microcosm. (B) Meta dechlorination in
the DCP enrichment culture from the compost microcosm. (0) Enrichment with
phenol and nitrate (1 mM) from compost microcosm Arrows denote addition of

substrate.

33

Table 2.2 Characteristics of active enrichment cultures. At least five different
dechlorination patterns are observed in the enrichments. Shown are the
chlorophenolic substrate specificities of the different enrichment cultures. Also
indicated is the sensitivity of dechlorination to nitrate and whether methane is detected
In the enrichment cultures.

 

 

 

 

 

Enrich mentsa
o-MCP 2, P o- and m- m-MCP 3,4- P
SubstratdSib culture I I MCP I
o-MCP + - + - -
m-MCP - + + + +
p-MCP - - - - -
2,3-DOP->m-MCP - + - - -
2,4-DCP—>p-MCP -I- - + - '-
2,5-DCP->phenol - - + - -
2,5-DOP-> m-MCP + - + - -
2,6-DOP->phenol + - - - -
2,6-DCP—> o-MCP + + - - -
3,4-DCP—> m-MCP - - - - +
Dechlorhafion with
N03‘° + - - - -
wheelsd - + - + +

 

 

a Enrichments are identified by the substrate used for dechlorination. Since m-MCP is a product in the 2,3-
DCP and 3,4-DCP cultures the m-MCP dechlorinating enrichment probably is a subset microbial population
of these other dechlorination enrichments.

b Substrates are chlorinated phenolic con'pounds monitored for dechlorination in cultures amended with
acetate as and electron donor.

° 5 mM nitrate was added to media containing the primary chlorinated suburate of the enrichment and
acetate. A positive result indicates that nitrate is not inhibitory to dechlorination.

d Positive indicates that CH4 was detected in the headspace of enrichment cultures showing dechlorination

 

 

 

each CUliU
capable 01
showed d
activity we:
Cha

activity in
electron do

added as
populations
To test this,
and DCPs (3
test substra
SDGCifiCity, ir
dechlorinatio
was Confirms
A”Willi all
enrichment 1
Source for a,
Exhibited higr

Were used as

    
   
 
 

diVGrSe mlCr
enrichments
dechlonnatiOr;
dechlorinatmr
Nitrate “’83 Sf)
2.3.0CP el'irl

reducibn.

34
each culture and nitrate was completely depleted, indicating that denitrifiers were
capable of degrading phenol (data not shown). Cultures with phenol alone also
showed degradation activity with the appearance of methane, however this
activity was considerably slower than was exhibited with nitrate.

Characterization of enrichment activity. Although dechlorination
activity in all enrichments was dependent on the presence of acetate as an
electron donor, there appeared to be different responses to the MCPs and DCPs
added as substrates. A possible explanation was that different microbial
populations were mediating reductive dechlorination in each enrichment culture.
To test this, each enrichment was tested for its ability to dechlorinate the MCPs
and DCPs originally used in the microcosms. in addition 2,6-DCP was used as a
test substrate. Each of the five enrichments showed a different substrate
specificity, indicating that five different microbial populations were responsible for
dechlorination (Table 2.2). This physiological diversity between the enrichments
was confirmed by testing a variety of electron donors for reductive dechlorination.
Although all enrichments used acetate as an electron donor, subsequent
enrichment transfers showed that this was not the best carbon and energy
source for all of the dechlorinating populations. Several of the enrichments
exhibited higher levels of dechlorination activity when formate, H2, or butyrate
were used as electron donors (data not shown). This would be expected among
diverse microbial populations. Methanogenesis also occurred in some
enrichments and it was not resolved whether this activity was correlated to the
dechlorination observed in these cultures. Nitrate (1 mM) was inhibitory to
dechlorination in all enrichments except the o-MCP enrichment (Table 2.2).
Nitrate was shown to be preferentially reduced to nitrite when it was added to the
2,3-DOP enrichment and no dechlorination was observed during nitrate

reduction.

"J

 

 

 

 

 

 

pres
snot
mhal
Show
dhon
medn
dudy
mmnx
did no
readdi
meint
ennchr
mainta
and se
nitrate
degrac
remove
Worth;
Sansim
effect c
Oiher S
aCtivity
conCen

”issu.

35
Discussion

Dechlorination of MCPs and DCPs in the microcosms occurred in the
presence of nitrate, but was not coupled to denitrification. This is clearly evident
since dechlorination activity in the enrichments was maintained in the absence of
nitrate as long as a suitable electron donor was present. Previous studies have
shown that denitrifying microcosms failed to sustain the dechlorination of
chlorophenols when amended with new substrates or transferred to fresh nitrate
media (Genthner et al. 1989; Haggblom et al. 1993). it was suggested‘in one
study that nitrate at levels of 15 mM was completely depleted in the original
microcosm before dechlorination occurred (Genthner et al. 1989). These authors
did not report monitoring the nitrate levels in these cultures, but did report that the
readdition of nitrate inhibited the dechlorination activity. This is consistent with
the inhibition of dechlorination activity we observed in both the microcosms and
enrichment cultures. The use of low initial nitrate concentrations along with
maintaining nitrate-limiting conditions were essential to our observed concurrent
and sequential denitrification and dechlorination. My results indicate that when
nitrate concentrations are high (> 5 mM) in the microcosms MCP and DCP
degradation rates are slowed significantly. However, as long as nitrate is
removed prior to its readdition at low concentrations, the degradation of the
chlorinated phenols proceeded uninhibited. Enrichment cultures are even more
sensitive to nitrate, suggesting that the microcosm environment has a buffering
effect on this inhibitory activity allowing dechlorination to continue. The reason
other studies have failed to observe concurrent dechlorinating and denitrifying
activity in chlorophenol enrichments may be because of the excessive nitrate
concentrations used, generally 15 mM or greater (Genthner et al. 1989;

Haggbiom et al. 1993). High nitrate concentrations are contrary to what would

 

liar -——

 

naturally
tolerant er
Ba:
dechlorina
al. 1982).
stoichiome
in the me
dechlorinai
chlorophen<
these cultur
available frc
(Dolfing and
of nitrate to
electron don
Occurring sini
reductive (19¢
DOB“ does .
gain (Dolfing
isolated an o_
decmon'liallon
The miCFObiai

When sUitabl

 

Ch[ororesifira‘
dechlorinating r

“Other gram

 

36
naturally be present in the environment, and low levels would provide a more
tolerant environment for these two processes.

Based on the appearance of phenol and chlorophenols as products of
dechlorination in the microcosms, indicates that this activity is reductive (Suflita et
al. 1982). This is further supported by the enrichment cultures where
stoichiometric concentrations of dechlorinated phenolic products were observed
in the medium if electron donors were provided. Whether individual
dechlorinating populations are gaining energy through the use of the
chlorophenols as physiological electron acceptors has yet to be determined in
these cultures. However it is a reasonable possibility since the free energy
available from the dechlorination of MCPs and DCPs is about -160 kJ/rxn
(Dolfing and Harrison 1992), which is similar to that available from the reduction
of nitrate to nitrite(Thauer et al. 1977). The ability for acetate to serve as an
electron donor in the enrichments also suggests that halorespiration may be
occurring since the oxidation of acetate is not exergonic unless it is coupled to
reductive dechlorination. It has been established that Desulfomonile tiedjei
DOB-1 does couple reductive dechlorination of 3-chlorobenzoate to energetic
gain (Dolfing 1990; Mohn and Tiedje 1990). Recently Cole and co-workers
isolated an o-MCP dechlorinating culture which also couples growth to reductive
dechlorination establishing that chlorophenol respirers exist (Cole et al. 1994).
The microbial populations in the current study mediate this dechlorination only
when suitable electron donors are present, a feature consistent with
chiororespiration. The isolation of a trichlorophenol (TCP) and DOP
dechlorinating anaerobic spore former was recently reported, although its ability
to grow with halorespiration was not established (Madsen and Licht 1992).
Another gram-positive 2,4-DCP dechlorinating bacterium, Desulfitobacterium

dehalogenans, was isolated from enrichments that had been heat-treated,

 

 

 

implying
shown to
electron a
The
form of a 1
these acti
considerab
recognize
focusing or
activities, ca
the MCPs
chlorOpheno
Specificities
environment
o'GClllorinatior
flied each ott
CleChlorinatiOI
microbial pop,
cultures are a
well as both
Step one, inv
i”dependent c
3,4.DCP (Ste

  

(EESOned the

DCPs, hOWev
DCP (T able 2.:

an ahemailVe

 

37
implying the presence of spores (Utkin et al. 1994). This strain also was not
shown to grow using chlorinated phenolic compounds as its sole physiological
electron acceptor.

The dechlorination activities observed in this study are summarized in the
form of a possible food web in Figure 2.4. Our evidence suggests that each of
these activities may be carried out by a different population, indicating
considerable diversity in chlorophenol dechlorinating organisms. It is important to
recognize the different dechlorinating activities of each community before
focusing on isolation, so that a broader group of isolates, some with novel
activities, can be sought. Figure 2.4 suggests possible degradation pathways for
the MCPs and DCPs in the enrichments studied. Even though some
chlorophenols are used by more than one enrichment culture, there are substrate
specificities that differentiate these cultures so that they could co-exist in an
environment where a mixture of substrates exist. Those specific reductive
dechlorinations mediated by different enrichment cultures would potentially cross-
feed each other in the overall microbial community. Enrichment culture-specific
dechlorination reactions are numbered to indicate each potentially distinct
microbial population. The compost soil microcosms and subsequent enrichment
cultures are able to mediate four of the five suggested dechlorination reactions as
well as both denitrifying and methanogenic degradation pathways for phenol.
Step one, involving o-MCP dechlorination to phenol, has been shown to be
independent of the dechlorination of m-MCP (step 5) or the dechlorination of the
3,4-DCP (step 4), which involves an initial para dechlorination. it might be
reasoned that the ortho dechlorinating population would dechlorinate all the
DCPs, however the o-MCP (step 1) cultures were not able to dechlorinate 2,3-
DOP (Table 2.2). As would be expected, a separate enrichment culture exhibits
an aitemative ortho dechlorination pathway which did dechlorinate 2,3-DCP (step

 

38

 

0|§m®
.55"! ----- o2. \:°

60' acetate

Ei::® 3&1.ng

 

 

 

 

 

 

 

 

 

OH / Vé ICH4 + 002
x...
C' or VFAs

Cl ‘s‘

OH ‘ F OH -
GC'SE, E:

c] L CI _

L J

 

Figure 2.4. Schematic representation that summarizes the different
dechlorination and degradation activities observed in the enrichments. Large
numbers indicate the predominant reaction mediated by a particular enrichment
and small numbers indicate additional dechlorination reactions that are observed
in these cultures. (Numbers correspond to the following enrichments: reaction

I = o-MCP; reaction 2 = 2,3-DOP; reaction 3 = o- and m-MCP; reaction 4 = m-
MCP and reaction 5 = 3,4-DCP) Reactions 1, 2 and 3 represent distinct OI'U'IO
dechlorinating pathways. Cultures mediating reaction 1 do not dechlorinate 2,3-
DCP. Meta dechlorination is mediated by at least two different communities
depicted by reactions 3 and 5. Para dechlorination is only observed for 3,4-DCP
as shown by reaction 4. The fate of phenol is summarized in two possible
pathways; denitrifying and methanogenic through benzoate.

 

 

 

 

nnc
cou
the
sub:
deci
The
head
drec
are i
dechl
dechk

”fixtur
dechk
dechlo
t dd

enflchr
Pahen
Observ
has be
hosel
1984;l
Winter
of meta

Mimi]

39

2). This 2,3-DOP enrichment is not able to use o-MCP, which indicates that the
microbial populations represented in enrichments mediating step one and two
could coexist when both substrates are present. Also included in Figure 2.4 is
the alternative pathway for simultaneously dechlorinating ortho- and meta-
substituted MCPs and 2.5-DCP (step 3). This represents a third ortho
dechlorination reaction, perhaps mediated by hydrogen as an electron donor.
This latter suggestion is derived from the transient appearance of H2 in the
headspace of enrichments in the absence of methanogenesis, suggesting a
direct coupling to dechlorination. In summary the evidence suggests that there
are three different ortho -dechiorinating populations, two different meta-
dechiorinating populations and at least one population capable of para-
dechlorination.

in contrast to previous studies using chlorinated phenolic compounds,
mixtures of MCPs and DCPs were used, which perhaps enabled a more diverse
dechlorinating community to be selected. For example although meta
dechlorination did not occur in the compost soil microcosm with the MCP mixture,
it did occur with the DCP mixture. it was even successfully transferred to
enrichments with m-MCP as a substrate. Despite these differences the overall
patterns of dechlorination in these cultures are similar to those previously
observed. Ortho dechlorination, the most prevalent activity in our microcosms,
has been observed frequently in many anaerobic cultures, although generally
those have been maintained under methanogenic conditions (Boyd and Shelton
1984; Hrudey et al. 1987; Genthner et al. 1989; Kohring et al. 1989; Dietrich and
Winter 1990; Hale et al. 1990; 1991; Haggblom et al. 1993). The dechlorination
of meta substituted phenols has also been previously observed with this activity
occurring at generally slower rates than ortho dechlorination (Boyd and Shelton

1984). The persistence of p-MCP is also common among other studies and our

 

 

 

mat
0516
the

COM

Of th

iii-311,;
”par

for Sir

40
study supports the contention that it is the least common activity observed in
methanogenic microbial enrichments (Genthner et al. 1989; Hale et al. 1990;
Parker et al. 1993). However the para-dechlorination of 3,4-DCP to m-MCP in
the enrichment transfer culture is the predicted activity based on the
eiectronegativity of the chlorine in the para position which makes it the better
leaving group (Cozza and Woods 1992). Previous studies, in contrast, have
shown that meta dechlorination of 3,4-DOP is the apparent predominant
biological dechlorination activity in some enrichments (Mikesell and Boyd 1985;
Woods et al. 1989).

One of the interesting aspects of the results is the influence of source
material on the presence of both denitrification and dechlorination. in contrast to
other studies, mostly surface soils were used in our microcosms in order to favor
the recovery of denitrifiers. In general, microcosms seeded with non-
contaminated soils exhibit more activity than those with contaminated soil. Four
of the five observed different dechlorination reactions were recovered from the
compost soil'microcosms. The reductive removal of chlorines from o-MCP was
recovered in both compost and tropical soil derived enrichments. A possible
selective basis for the innate dechlorination ability in these soils may be the
presence of naturally occurring chlorinated phenols (Siuda and Debemardis
1973; Gribble 1992). Recently de Jong et al. (1994) established that up to 70
mg/kg of chlorinated anisyl metabolites produced by white rot fungi can be found
in forest litter. They illustrated that these compounds can easily be transformed
by bacteria into chlorophenols, in this case 3-chioro-anisaldehyde and 3,5-
dichloroanisaldehyde to o-MCP and 2,6-DCP, respectively. This implies that high
organic matter soils with a potentially large fungal population could be reservoirs

for significant amounts of CPS.

 

 

 

 

”18

ad»
(:hl

the

9251;
work
9tooe

Berg]

B°Yd

B°ld

41

Although dechlorination and denitrification in the microcosms are not
mediated by the same bacterial population it is possible that mineralization may
be stimulated through the addition of low concentrations of nitrate. This could be
advantageous in formulating remediation strategies for contaminated sites.
Chlorophenolic compounds would continue to be anaerobically dechlorinated and
the products will be subsequently mineralized. by the denitrifiers present.

We have shown that both dechlorination and denitrification activity can be
maintained in an active microcosm, but that these processes are mediated by
physiologically different populations. We found no evidence for the existence of
denitrifiers that dechlorinate MCPs or DCPs with nitrate reduction. Anaerobic
dechlorination of MCPs and DCPs is complex, with at least five different types of
reactions observed in this study. it is, however, common and under appropriate

conditions, may be harnessed for use in bioremediation systems.

Acknowledgements
Tropical soils were collected by Roland Weller in Cameroon as part of:
Rad au Cines Foundation, ELF, Afique 91 sponsored by ELF-SEREPCA. This
work was supported by the Center for Microbial Ecology through grant No. BIR-

91006 from the National Science Foundation.

References

Berger R.S. 1972. 2,6-Dichlorophenol, sex pheromone of the lone star tick.
Science 177:704-705.

Boyd S.A., D.A. Shelton, D. Berry and J.M. Tiedje. 1983. Anaerobic
biodegradation of phenolic compounds in digested sludge. Appl. Environ.
Microbiol. 46:50-54.

Boyd SA. and D.R. Shelton. 1984. Anaerobic biodegradation of chlorophenols
in fresh and acclimated sludge. Appl. Environ. Microbiol. 47:272-277.

 

 

Cole J.A..
chal
ol 2-

Com C.l
sub
2£K

de Jong E
199«
envi

Dietrich G.
ennc

Dolfing J.
ATP
Anfll

Dolfing J.
halog
accer
2218.

Genthner l
degra
vane

Genthner l
anae
Env‘

G'Ibsons.
9w

 

42

Cole J.A., A. Cascarelli, w.w. Mohn and J.M. Tledle. 1994. Isolation and
characterization of a novel bacterium growing via reductive dechlorination
of 2-chlorophenol. Appl. Environ. Microbiol. 60:3536-3542.

Cozza C.L. and S.L. Woods. 1992. Reductive dechlorination pathways for
substituted benzenes: a correlation with electronic properties. Biodeg.
2:265-278.

de Jong E., J.A. Fleld, H.-E. Splnnler, J.B.P.A. Wllnberg and J.A.M. de Bont.
1994. Significant biogenesis of chlorinated aromatics by fungi in natural
environments. Appl. Environ. Microbiol. 60:264—270.

Dletrlch G. and J. Wlnter. 1990. Anaerobic degradation of chlorophenol by an
enrichment culture. Applied Microbiology and Biotechnology 34:253-8.

Dolflng J. 1990. Reductive dechlorination of 3-chlorobenzoate is coupled to
ATP production and growth in an anaerobic bacterium, strain DCB-1.
Arch. Microbiol. 153:264-266.

Dolflng J. and B.K. Harrlson. 1992. Gibbs free energy of formation of
halogenated aromatic compounds and their potential role as electron
acceptors in anaerobic environments. Environ. Sci. Technol. 26:2213-
2218.

Genthner B.R.S., W.A. Prlce II and P.H. Prltchard. 1989. Anaerobic
degradation of chloroaromatic compounds in aquatic sediments under a
variety of enrichment conditions. Appl. Environ. Microbiol. 55:1466-1471.

Genthner B.R.S., W.A. Prlce II and P.H. Prltchard. 1989b. Characterization of
anaerobic dechlorinating consortia derived from aquatic sediments. Appl.
Environ. Microbiol. 55:1472-1476.

Glbson S.A. and J.M. Sufllta. 1986. Extrapolation of biodegradation results to
groundwater aquifers: reductive dehalogenation of aromatic compounds.
Appl. Environ. Microbiol. 52:681-688.

Grlbble G.W. 1992. Naturally occurring organohalogen compounds - a survey.
J. Nat. Prod. 55:1353-1395.

Higgblom MM. 1992. Microbial breakdown of halogenated aromatic pesticides
and related compounds. FEMS Microbiol. Rev. 103:29—72.

Haggblom M.M., H.D. Rivera and LY. Young. 1993. Influence of alternative
electron acceptors on the anaerobic biodegradability of chlorinated
phenols and benzoic acids. Appl. Environ. Microbiol. 59:1162-1167.

l-lale D.D., J.E. Rogers and J. Wlegel. 1991. Environmental factors correlated
to dichlorophenol dechlorination in anoxic freshwater sediments. Environ.
Toxicol. Chem. 10:1255-1265.

 

 

Hat 0
l

Hendfi

\
'.
\

ltudey
t

Kohfing

 

 

43

Halo D.R., J.E. Rogers and J. Wlegel. 1990. Reductive dechlorination of
dichlorophenols by nonadapted and adapted microbial communities in
pond sediments. Microbial Ecol. 20:185-96.

Hendrlksen H.V., S. Larsen and B.K. Ahrlng. 1992. Influence of a
supplemental carbon source on anaerobic dechlorination of
pentachlorophenol in granular sludge. Appl. Environ. Microbiol. 58:365-
370.

Hrudey S.E., E. Knettlg, S.A. Dalgnault and PM. Fedorak. 1937. Anaerobic
biodegradation of monochlorophenols. Environ. Technol. Lett. 8:65-76.

Kohrlng c.-w., J.E. Rogers and J. Wlegel. 1989. Anaerobic biodegradation of
2,4-dichlorophenol in freshwater lake sediments at different temperatures.
Appl. Environ. Microbiol. 55:348-353.

Kohrlng G.-W., X. Zhang and J. Wlegel. 1989. Anaerobic dechlorination of
2,4-dichlorophenol in freshwater sediments in the presence of sulfate.
Appl. Environ. Microbiol. 55:2735-2737.

Madsen T. and J. Aamand. 1992. Anaerobic transformation and toxicity of
trichlorophenols in a stable enrichment culture. Appl. Environ. Microbiol.
58:557-561. '

Madsen T. and D. Llcht. 1992. Isolation and characterization of an anaerobic
chlorophenol-transforming bacterium. Appl. Environ. Microbiol. 58:2874-
2878.-

Mlkesell MD. and S.A. Boyd. 1985. Reductive dechlorination of the pesticides
2.4-D, 2,4,5-T and pentachlorophenol in anaerobic sludges. J. Environ.
Qual. 14:337-340.

Mohn w.w. and J.M. fledle. 1990. Strain DCB-t conserves energy for growth
from reductive dechlorination coupled to formate oxidation. Arch.
Microbiol. 153:267-271.

Mohn w.w. and J.M. Tledle. 1992. Microbial reductive dehalogenation.
Microbiol. Rev. 56:482-507.

Parker W.J., G.J. Farquhar and E.R. Hall. 1993. Removal of chlorophenolics
and toxicity during high-rate anaerobic treatment of segregated Kraft mill
bleach plant effluents. Environ. Sci. Technol. 27:1783-1789.

Sluda J.P. and J.P. Debemardis. 1973. Naturally occurring halogenated
organic compounds. ondia 36:107-143.

Stevens T.O., H.D. Watts, J.J. Walker and J.K. Fredrickson. 1992. Medium
formulation presented in a Poster Entitled: Optimization of solid growth

 

l;

 

 

 

medlur
subsur
1992.

Suflila J.M
Dehak
haloan

Thauer alt,
chemo.

Utkin l., C. l
Desu/f
bacteri
J. Syst

Wolin E.A.,
bacters

Woods S.L.
chlorc;
lreatm:

 

Zhang X. a
dichlo.
56:111

44

medium for isolation and culture of microorganisms from the terrestrial
subsurface, p. 366. In (ed.), Abstr. 92nd Gen. Meet. Am. Soc. Microbiol.
1992. American Society for Microbiology, Washington, D. C.

Sufllta J.M., A. Horowitz, D.R. Shelton and J.M. Tledle. 1982.
Dehalogenation: a novel pathway for the anaerobic biodegradation of
haloaromatic compounds. Science 218:1115-1117.

Thauor R.K., K. Jungermann and K. Decker. 1977. Energy conservation in
chemotrophic anaerobes. Bacteriol. Rev. 41 :100-180.

Utkin |., C. Woese and J. Wlegel. 1994. Isolation and characterization of
Desulfitobacterium dehalogenans gen. nov., sp. nov., an anaerobic
bacterium which reductively dechlorinates chlorophenolic compounds. Int.
J. Syst. Bacteriol. 44:612-619.

Wolln E.A., H.J. Wolln and R.S. Wolfe. 1963. Formation of methane by
bacterial extracts. J. Biol. Chem. 238:2882-2886.

Woods S.L., J.P. Ferguson and MM. Benjamin. 1989. Characterization of
chlorophenol and chloromethoxybenzene biodegradation during anaerobic
treatment. Environ. Sci. Technol. 23:62-68.

Zhang X. and J. Wlegel. 1990. Sequential anaerobic degradation of 2,4-
dichlorophenol in freshwater sediments. Appl. Environ. Microbiol.
56:1119-1127.

 

 

Isolation

The.
acceptors h.
and Tiedje
dechlorinatic
ability to hal
organism ca
aromatic cm
(Gribble 19s
lungi actua'
litter. Thus
'0 occur in

The
lmponant

contaminal

are femove

 

Chapter lll

Isolation and preliminary characterization of the aryl-haloresplrlng
anaeroblc myxobacteria strains ch-C and 2CP-3

Introduction

The ability to utilize haloaromatic compounds as physiological electron
acceptors has only been described in a few pure cultures (Dolfing 1990; Mohn
and Tiedje 1990; Cole et al. 1994; Sanford et al. 1996). Since reductive
dechlorination releases considerable energy (Dolfing and Harrison 1992), the
ability to halorespire in anaerobic environments is potentially beneficial to the
organism catalyzing this transformation. The types of natural halogenated
aromatic compounds that might be present in such an environment are many
(Gribble 1992), and recently de Jong and coworkers (1994) found that white rot
fungi actually generate up to 70 mg/kg of chlorinated anisyl metabolites in leaf
litter. Thus, not only is a halorespiratory metabolism potentially useful, it is likely
to occur in natural anaerobic ecosystems.

The reductive removal of chlorines from aromatic compounds may be
important to anaerobic bioremediation strategies where considerable
contamination with chlorophenolic compounds has occurred. Once chlorines
are removed, the resulting aromatic compounds are generally more amenable
to oxidative attack. This was observed in denitrifying microcosms in which 2-
chlorophenol (2-CP) was dechlorinated by one microbial population to phenol,
which was then degraded by denitrifiers (Chapter 2). Similar results have been
~ observed with anaerobic methanogenic systems that are fed pentachlorophenol

(PCP) or other poly-chlorinated phenolic compounds (Mikesell and Boyd 1986).

45

 

 

 

 

 

"u 3mm": .-'-

01

of

ob

be

An
o'el
ha»
and
this
anal
1992
loca
dive
grow
line |
Unde
Orga
hEre

Dies

Com;

Oin

46

Many studies have shown the reductive dechlorination of PCP to di- and mono-
CPs (Nicholson et al. 1992). These lesser-halogenated products of
dechlorination more easily degraded under aerobic environmental conditions.

In this study, two novel anaerobic myxobacteria strains that are capable
of growth using 2-CP as a physiological electron acceptor were isolated and
characterized. A similar anaerobic halorespiring myxobacteria was recently
describedby Cole and others (1994). These strains have been named
Anaeromyxo dehalogenans, for their novel anaerobic life-style and ability to
dehalogenate chlorophenols. As a group of microorganisms, the myxobacteria
have always been considered obligately aerobic microorganisms (Reichenbach
and Dworkin 1992; Reichenbach 1993). However the phylogenic placement of
this group, within the delta-Proteobacteria, would suggest the possibility of an
ancestral anaerobic life-style for these microorganisms (Shimkets and Woese
1992). For example, sulfate-reducers and other anaerobic microorganisms are
located within this group and exhibit considerable anaerobic physiological
diversity. It is possible that these facultatively anaerobic myxobacteria have
growth requirements that have made it difficult to isolate similar organisms in
the past. The findings presented here suggest a possible life-style previously
underinvestigated in the microbial world, that of the microrespirotroph. Such an
organism, as observed with the Anaeromyxo dehalogenans strains described
here, does not grow well when high concentrations of an electron acceptor are
present. Novel culturing strategies may be necessary for isolating these types
of microorganisms.

Materials and Methods

lsolatlon. Cultures were enriched from anaerobic microcosms

containing different soils and mixtures of mono-chlorophenols in the presence

of nitrate (Chapter 2). Enrichments were obtained with the ability to

47

dechlorinate 2-CP using acetate as the sole carbon and energy source. To
obtain pure cultures, the agar-shake technique was used with 10 ml of 2.0 %
low melting-point agarose containing 1 mM acetate and 0.2 mM 2-CP. After
several weeks of incubation colonies were removed using a sterile syringe, with
anoxic conditions maintained by continuous flushing of the culture tube with 02-
free N2. The harvested colonies were transferred to broth culture containing
100 uM 2-CP and 1 mM acetate. Broth cultures that exhibited dechlorination
activity were amended with more 2-CP once it was consumed. When phenol
concentrations exceeded 2 mM. cultures were transferred to fresh medium.
Broth cultures were used to inoculate agar plates that were degassed for at
least 24 hours in an anaerobic glove box containing an N2:H2 gas mixture of
97:3. Plates were incubated in the anaerobic glove box and single colonies
were transferred to new agar plates. After at least three transfers, colonies were
transferred into chlorophenol or fumarate (5 mM) broth for further testing.
Cultures were subsequently grown on plates at 30° C in an anaerobic growth
vessel with an atmosphere of 10-20% 002 and the balance N2.

Growth medium. Cultures were grown in 160 ml serum bottles with
100 ml of boiled degassed medium or in anaerobic culture tubes with 20 ml of
medium and closed with butyl rubber stoppers. The mineral salts medium and
vitamin supplemental solution were as previously described in Chapter 2.

The plate media used for culturing the 2CP strains was R2A (Difco) agar
medium supplemented with 10 mM fumarate or 5 mM nitrate as electron
acceptors.

Electron donors and acceptors. The ability of the 2CP strains to
reductively dechlorinate 2-CP with different electron donors was determined in
duplicate in 20 ml anaerobic culture tubes incubated at 30° C. The soluble

volatile fatty acids (VFAs) acetate, formate, butyrate, succinate, propionate,

 

 

 

48

fumarate, lactate, pyruvate and H2 were tested as potential electron donors.
The VFAs were added to concentrations of 0.5 or 2.0 mM. H2 was tested by
addingtB umoles (6.6 % v/v) to the headspace of the duplicate culture tubes. 2-
CP (100 uM) was added as the electron acceptor. A 0.5 % inoculum was added
to each tube. The disappearance of 2-CP and appearance of phenol were
monitored by HPLC. 2-CP was replenished at least three times at
concentrations of 100 uM to those cultures exhibiting dechlorination activity.
VFA utilization was monitored by HPLC and H2 was determined by GC. A
positive test was indicated if both dechlorination and depletion of electron donor
were observed.

The range of electron acceptors used by the ZCP strains was determined
using the same growth conditions as with the electron donor determination.
With acetate (1 mM) serving as an electron donor, 100 (M of 2,3-DCP; 2.4-DCP;
2,5-DCP; 2,6-DCP; 2-CP; 3-CI-4-HBA; 2-fluorophenol; 2-bromophenol; 2-
iodophenol; 2,3,5-TCP; 2,4,6-TCP; pentachlorophenol (PCP); or 3-
chloroanisaldehyde were tested as halogenated electron acceptors. Nitrate,
sulfate, sulfite, thiosulfate and fumarate were also tested at concentrations of 5
mM. Tubes were inoculated with 0.1 - 0.5 % of cultures grown on acetate and
2-CP. Halogenated substrates were monitored by HPLC and were replenished
once depleted. Nitrate and fumarate concentrations were measured by HPLC.
Use of sulfur compounds was determined by monitoring increases in turbidity
versus control cultures with no-cells. Acetate consumption was monitored by
HPLC to verify physiological activity.

Nitrate effect on dechlorination. The effect of nitrate (2 mM or 5
mM) on reductive dechlorination of 2-CP was tested in anaerobic culture tubes
with acetate (1 mM) as the electron donor. Duplicate cultures of the 2CP strains

were inoculated and monitored for the appearance of phenol, the

 

 

 

disa;
nitrat
anal)

melh

Orthc
bear
suspr
in 10
were
were

Photc

uptair
hydrc
caoac
100 rr
culturr
CCUil
amehc
lying
We r
CODCen

deiector

file l6du.

49

disappearance of nitrate and appearance of nitrite. In order to determine if
nitrate was reduced to N2 or ammonia, N2 and N20 were monitored by GC
analysis. Ammonia was determined by the indophenol blue colorimetric
method.

Microscopy. Microscopic observations were done using a Leitz
Crthoplan 2 microscope. Colonies on plates were illuminated with a focusing
beam of light and observed under the 5 X objective. Cells were observed by
suspending portions of colonies and centrifuged suspensions of broth cultures
in 10 ul of phosphate buffer spread on dry agarose coated slides. Cover slips
were sealed with a molten mixture of 50% paraffin and 50% vaseline. Cells
were observed under oil-immersion with a 60 X phase-contrast objective lens.
Photographs were taken using TMAX 100 black and white film.

Hydrogen uptake and threshold determination. Hydrogen
uptake was monitored in relation to dechlorination activity to test if the
hydrogenase activity was closely linked to the terminal electron-accepting
capacity of the 2CP strains. Hydrogen (5.0 %) was added to the headspace of
100 ml cultures in 160 ml serum bottles. 2-CP (100 (M) was added to these
cultures and the H2 concentration was monitored by GC as dechlorination
occurred. After the complete dechlorination of the 2-CP, an additional
amendment of 100 uM was made. To determine the threshold concentration for
hydrogen maintained by the 2CP strains while dechlorinating, triplicate cultures
were monitored for H2 depletion with excess 2-CP present. Threshold
concentrations of H2 were measured using a GC equipped with a reduction gas
detector from Trace Analytical, Menlo Park, CA.

Determination of fe. As an indication of energetic efficiency and
evidence of halorespiration, the fraction of electrons (to) from acetate used for

the reductive dechlorination of 2-CP was determined. Duplicate cultures of

 

 

 

each
receii
and a
reach
Nomi
pmoh
reduc
hydrc
hai-r

Thus,

are 9

uhng
was 5
adsor
ennci

ahahs

wee;
Packag
at 1.5;

“”U col

50

each strain were grown on acetate (1 mM) and 2-CP (125 uM). Control cultures
received acetate alone. Concentrations of 2-CP were monitored daily by HPLC
and amendments of new 2-CP (125 uM) were made prior to the concentration
reaching zero. Acetate was determined by HPLC. The to was calculated by
plotting the acetate concentration consumed as H2 equivalents versus the
umoles phenol produced. Hydrogen equivalents were used since each
reductive dechlorination requires one mole H2 per mole phenol formed. The
hydrogen equivalents from acetate were calculated according the following
half-reaction:

CH3COO' + 4H20 ---> 2 HC03' + H‘l' + 4 H2
Thus, for every mole of acetate consumed four moles of hydrogen equivalents
are generated.

Analytical procedures. Nitrate and nitrite were analyzed by HPLC
using a Whatman Partisil 10 SAX column on a Schimadzu HPLC. The eluent
was 50 mM phosphate buffer (pH = 3.0) pumped at a rate of 1 ml/min. UV
adsorption at 210 nm was used for detection. Samples from primary
enrichments were diluted 1:100 in deionized H2O prior to nitrate and nitrite
analysis. A 40 ul sample was injected by an Alcott 738 autosampler.

Chlorophenols, dichlorophenols and aromatic products of dechlorination
were analyzed on a Hewlett Packard 1050 HPLC with a Chemstation analysis
package. The eluent was phosphoric acid (0.1 %) buffered methanol pumped
at 1.5 mein using a gradient from 48% to 55% methanol. A Hibar RP-18 (10
um) column was used. Detection of eluted peaks was done at 218 nm, 230 nm
and 275 nm simultaneously on a UV multiwavelength detector. Samples (1 ml)
from the enrichments were taken, made basic with 10 ul of 2N NaOH,
centrifuged for 6 min in a microfuge and filtered through Acrodisc LC13 PVDF
0.45 pm filters prior to HPLC analysis.

 

 

Vda
BbRad Ar
OWSNH
0%NH2
was pump
reabr

rm
002 usir
column a
normalize
gas for ir

with a F

were me

Is
exhibiter
and the
Observe
mmew
“Eductivr
cultures
Wire is:
Isolation
of 2-CP
exhibited

lUmarale

51

Volatile fatty acids were analyzed using the Schimadzu HPLC with a
BioRad Aminex HPX-87H ion exclusion column heated to 60°C and using
0.005 N H2SO4 as the eluent. Previously filtered samples were acidified to
0.25 N H2SO4 by adding 100 pl of 2.5 N H2SO4 to 900 pl of sample. Eluent
was pumped at 0.6 ml/min and detection of VFAs was at 210 nm by a UV
detector.

The headspace of the incubation vessels was analyzed for N2, H2 and
CO2 using a Carle gas chromatograph equipped with a 1.83 m Porapak Q
column and a thermal conductivity (T CD) detector. Headspace pressure was
normalized to atmospheric by venting with a needle prior to removing 0.3 ml of
gas for injection into the GC. N2O was quantified on a Perkin Elmer 910 GO
with a Porapak O column and 63Ni -ECD detector. Denitrification products

were measured in an Ar headspace.

Results

Isolation of 2CP strains. Enrichment cultures from two soil types
exhibited dechlorination of 2-CP. One culture was of tropical origin (Cameroon)
and the other from a local compost pile. Several isolated colonies were
observed in agar shake cultures from both of these enrichments. Colonies,
some with a red pigment, were transferred to broth culture and showed
reductive dechlorination of 2-CP with acetate as an electron donor. Three
cultures designated ZCP-C (from Cameroon), 2CP-3 and 20P-5 (from compost)
were isolated. 2CP-C and 2CP-3 were selected for continued study after
isolation on agar plates. Figure 3.1 shows the typical pattern of dechlorination
of 2-CP and consumption of acetate exhibited by 2CP-C. The other strains
exhibited similar behavior. 2CP colonies grown on R2A agar medium with

fumarate as an electron acceptor developed a red pigmentation after several

52

 

     
 

 

 

 

3000 * 1200
’0 .
A .p I
E 2500 I - 1000
l': _O- 2'CP up ‘
8 2000 .. --9--phenol ‘ol L 800
2 r —O—acetate 6’. I
a 1
-- 1500 ~ ’ - 50°
2 - ‘
m a" ‘ 400
E:- 1000 :- I j
C? r .Q'fl 4
SE. 500 - 1’", . - 200
‘ ‘ _' ' _' ~_' ' _' ' _' .' ' _' ' ‘ O
0 60 120 180 240 300 360 420
hours

Figure 3.1. Dechlorination of 2-CP to phenol by strain 2CP-C. Reduction in
the level of acetate by the same culture is also shown.

(wfl) [erereoel

53

weeks of incubation. The red pigment was not extractable with acetone. Some
colonies appeared to change with time, becoming more dense in pigmentation,
and cells appeared to concentrate and build-up a mound at the center of the
colony. These masses formed small raised colonies on the surface of the agar
(Figure 3.2A). Microscopic observations of cells from this concentrated colony
mass showed evidence of refractile bodies, which are indicative of spores.
Vegetative cells from young cultures are shown in Figure 3.28 and some cells
at this growth stage were observed to have a gliding-like motility, however (no
motility was seen in most of our observations. The terminal ends of cells formed
hook-like structures and blebs. All ZCP strains stained Gram negative.

Electron donors and acceptors. Table 3.1 lists the general
characteristics of the 2CP strains and the electron donors utilized for the
reductive dechlorination of 2-CP. Acetate was the best electron donor for
combined growth and dechlorination activity, although hydrogen was used in
preference to acetate when both were present. The halogenated electron
acceptors utilized by 2CP-C, 2CP-3 and 2CP-1 (a strain isolated by Cole
(1994)) are summarized in Table 3.2. In general, all strains had the same
substrate range with one exception, 2CP-C dechlorinated 2,4-DCP and 2,4,6-
TCP much faster and more completely than 2CP-1. In addition to ortho-
substituted chlorophenols 2CP-C and 20P-1 were able to grow with 2-
bromophenol as an electron acceptor. Concentrations of chlorophenols above
250 uM were inhibitory to dechlorination activity. Iodo- and fluoro-substituted
halophenols were not utilized.

Nitrate, oxygen and fumarate were also used as electron acceptors by
the ZCP strains (T able 3.2). Plates that were incubated with 5 % oxygen and 10
% 002 added to the gas mixture exhibited colony formation within one to two

weeks. Aerobically incubated plates exhibited poor growth or colonies did not

54

 

Figure 3.2. (A) Vegetative cells of strain 2CP-C by phase contrast microscopy.
Bar is 13 um long. (B) Colony formed by 2CP-C on anaerobic agar, showing
the formation of a central cell-mass.

 

55

Table 3.1. Characteristics of 2CP strains. Data reflect general features
observed in the ZCP strains isolated and described in this study.

 

Waco
Gram stain -
Motility“ «1-
Cell morphologyb long slender rods with tapered ends
Uoux1u)
Colony morphologyb pink or red small round colonies
Sporesb 4-
Inhibition of dechlorination by -
"03"=
Temperature range 20° C - 37° C
W
Acetate +
H2 +
Succinate +
. Pyruvate +
Lactate +I-
Formate +
Butyrate -
Propionate -

 

 

' Motility was observed in cells from a solid agar medium placed on a wet-mount agarose coated slide.
Cells appeared to have gliding-type motility.

h Cells were grown on R2A agar plates anaerobically with fumarate as an electron acceptor. Cells
were examined microscopically as described for motility.

° Nitrate (5 mM) did not inhibit dechlorination of 2-CP as long as acetate

was present. 2-CP was used preferentially to nitrate as an electron acceptor.

5 Electron donors were tested with 2-CP as an electron acceptor. lf dechlorination was observed
over two feedings of chlorophenol the substrate was scored positive.

56

Table 3.2. Electron acceptors used by different 2CP strains. A (+) indicates
reductive dechlorination and acetate consumption occurred over at least three
additions of the halogenated substrate. This indicated growth had occurred,
which was confirmed by microscopic examination in those cultures marked
with an (*). (+l-) indicates that activity was observed but was slow and
complete degradation did not occur.

 

 

 

 

Halogenated
£33122, 2c P-C 2c P-1 2c P-3
*2-CP + + +
3-CP - - '-
4-CP - - -
2,3-DCP - - -
2.4-DCP + +I- +
2,5-DCP + + -
*2,6-DCP + + +
30l-4-OH-benzoate - nd nd
2,4,6-TCP ‘ + +l- nd
2,3,5-TCP - nd nd
PCP - - nd
2-Fluorophenol - - nd
*2-Bromophenol + + nd
2-lodophenol - - nd
3-Cl-anisaldehydea - nd nd
Other electron
acceptors
’N03' -> NH4+ + + +
'02 + + +
*Fumarate + + +

 

 

a 3-Cl-anisaldehyde is a natural product produced by white-rot fungi.
nd. not determined.

 

 

 

meas
cultu
2CP-

strair

micrr

USEd

57

grow at all. Fumarate and nitrate supported growth on plates and in broth
cultures. The oxidized sulfur compounds sulfate and thiosulfate did not support
growth in broth medium.

To determine if the 2CP strains were denitrifiers or were reducing nitrate
to nitrite or ammonia, the fate of nitrate in cultures grown with acetate was
quantified. After 21 days of incubation strain 2CP-3 converted most of the
nitrate at initial concentrations of 2.5 mM and 5.0 mM into ammonia (Figure 3.3).
Nitrite was also detected. Very small concentrations of N20 and N2 were
measured, but accounted for less than 5 % of total nitrate added in the 2.5 mM
culture and were insignificant in the 5 mM nitrate culture. Strains 2CP-C and
2CP-1 exhibited similar ammonia formation from nitrate, however both of these
strains were not as robust in their growth on nitrate.

Dechlorination In the presence of nitrate. Since the original
microcosms from which the 2CP strains were enriched contained both 2-CP
and nitrate as electron acceptors, the preference exhibited by 2CP-1, 2CP-C
and 2CP-3 for one or the other was determined. Figure 3.4 illustrates that
dechlorination and nitrate reduction occurred in the same culture, however the
preference of electron acceptors is not apparent since 2-CP was depleted
completely (arrows) on three occasions during incubation. After it was depleted
2-CP was replenished to the culture. In a parallel experiment with 2-CP
maintained in the cultures at all times, nitrate reduction was considerably
inhibited (data not shown). Reductive dechlorination in these same cultures
was not affected by the presence of nitrate at concentrations of 5 mM, indicating
a preference for 2-CP as an electron acceptor.

Indicators of halorespiration. The fraction of electrons from acetate

used for reducing the electron acceptor, fe, indicated that 64% of the electrons

 

58

 

80

 

 

5%

70:-

b

I
Z Z
I

60}

.13“ 1:]
:

 

 

 

 

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40

 

umoles

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t = 0 t = 21 t = Q = 2
nitrate = 2.5 mM nitrate = 5 mM

V
f
I
I
!

 

 

Figure 3.3. Use of nitrate by strain 2CP-3 at concentrations of 2.5 and 5 mM.
Mass balance shows that most of the nitrate is converted to NH4.

59

 

500

400 '

 

      

 

 

 

 

 

 

 

fix. Nitrate
. ‘~. Phenol
300 _ fi- 1
E
1 :
200_
100: ..
o It 11111 1.32.1...
o 5 10 15 20 25 30 35 40

Days

Figure 3.4. Dechlorination as indicated by phenol accumulation from 2-CP
and nitrate utilization by 2CP-3. Arrows indicate when 2-CP was completely
dechlorinated and when new additions of 2-CP were made.

 

60

from acetate were used for dechlorination by strain 2CP-C (Figure 3.5). Similar
results were obtained with the other ZCP strains.

Hydrogen uptake and the determination of a threshold concentration for
H2 were confirmatory indicators of a halorespiratory mechanism in the 2CP
strains. Figure 3.6 shows that H2 was only assimilated in the presence of 2-CP
and that the addition of this substrate immediately stimulated H2 uptake.
Threshold concentrations of hydrogen are theoretically associated with the
energetic redox couple controlling their growth. The uptake of hydrogen by the
2CP strains was monitored in the presence of excess 2,6-DCP, to determine the
threshold H2 concentration (Figure 3.7). 2,6-DCP was used since the ZCP
strains were able to reductively remove both chlorines and therefore an
equivalent concentration had twice the electron accepting capacity of 2-CP.
Within 25 hours the H2 concentration was approximately 1 ppm and still
decreasing. Final threshold concentrations after 48 hours of incubation, of less

than 0.5 ppm were measured for the ZCP strains (data not shown).

Discussion

Strains 2CP-C and 2CP-3 along with the previously isolated ZCP-I
represent a new group of facultatively anaerobic myxobacteria (Cole et al.
1994). Phylogenetically they are most similar to the Myxococcaceae, however
their 163 rRNA secondary structure shares features with Nanocystis and the
Chondomyces group (Cole, Sanford and Tiedje, manuscript in preparation).
This indicates the possible early evolutionary branching of the 2CP strains from
the well-characterized aerobic myxobacteria. Several physiological features of
the 2CP strains are shared with other myxobacteria. Bright red pigments were
produced by all 2CP isolates, which are common among the myxobacteria

(Reichenbach and Dworkin 1992). Cell shape, bleb formation and the ability to

61

 

 

 

 

soo,
» an
250 L ‘0'-
E g ‘5.
D. .b 12 = 0.985
E 150 ' 2‘\
O D
$01005 N.
R
C]
50 r “‘13..
'D
: - A A - I - - A g 1 A - - E l # - A - 1.3?‘1 A - -
0o 100 zoo 300 400” 500

Acetate equivalents as H2 (umoles)

Figure 3.5. Fraction of electron (fe) from acetate used for reductive
dechlorination of 2-CP by strain 2CP-C. The fe is derived from the slope of the
regression line generated by plotting the acetate consumed as H2 equivalents
versus the umoles 2-CP dechlorinated.

 

62

120

100 i

 

' r v v v

40

: 3Q l
20f Q

[H2] (ppm x 10")
8

 

 

 

r ‘0--------
0 - - - - l - - - A 1 - - - - I - -9 L 1 - - - Be - -
0 10 20 30 40 50 60
hours

Figure 3.6. Consumption of H2 by 2CP-C under electron acceptor limited
conditions. Arrows indicate when 2-CP was added to the medium.

63

 

 

 

 

 

 

 

 

 

 

 

 

 

3000
1000
2500
100 r- -------- -
E
2000 f g. 10
0' 1
O- 1500 — 1
1 t
a L 0.10 l l l 5 so
1000 _ 5 10 ho11frs 20 2
500 -
o 25 so

 

Figure 3.7. Uptake of H2 by 2CP strains in the presence of excess 2-CP. Inset
shows the threshold concentration of H2 for this culture.

 

iorr
19E
col
W
of

my

it 1
Th
co
rer
su

ce

elr

UF

SL

re

64

form spore-like structures are also commonly observed features (Reichenbach
1993). The colony morphology exhibited by 2CP-C, where cells appear to
collect together in a raised mound is very similar to the morphology shown by
Myxococcus xanthus fruiting-body deficient mutants (Zusman 1984). The loss
of the ability to form fruiting-bodies has been reported to be common in
myxobacteria, particularly if they have been maintained in broth cultures
(Reichenbach 1993). As all of the 2CP strains were enriched in liquid medium,
it would be reasonable to expect that well-formed fruiting bodies would not form.
The gliding-like motility of the 20P-C cells observed microscopically is also
consistent with the behavior of myxobacteria (Burchard 1984). Possible
reasons that this motility was not always observed is the lack of a suitable
surface and inappropriate environmental conditions such as pH, O2 tension or
cell-density.

The ability of the 2CP strains to grow anaerobically on a diverse group of
electron acceptors implies that the description of the myxobacteria as having
strictly aerobic life-styles needs to be reevaluated. It is not necessarily
unexpected to find anaerobic myxobacteria, since they are in the delta-
subdivision of the Protiobacteria. This group is known for sulfate reducers, ferric
reducers and other halorespirers which contribute to the group's considerable
anaerobic physiological diversity.

Perhaps the most unique feature of the 2CP strain physiology is the
preferential utilization of 2-CP as an electron acceptor over nitrate.
Energetically nitrate reduction to nitrite has a AG°' of -81.6 kJ mole electron'1
transferred (Thauer et al. 1977), while 2-CP reduction to phenol has a AG°' of
-78.5 kJ mole electron'1 (Dolfing and Harrison 1992). This suggests that nitrate
reduction should occur in preference to reductive dechlorination. However, the

AG°' of nitrate reduction to ammonia is only -75.0 kJ mole electron'1, which is

 

less

acce

ene
hair
gre
red

anI

ger
exr
evi
cor
up
the
ox

9V

65

less than the free-energy from the dechlorination of 2-CP. If the electron
acceptor of choice was only mediated by the thermodynamics of the reaction it
might be reasonable to expect the initial reduction of nitrate to nitrite, followed
by reductive dechlorination and finally nitrite reduction to ammonia. The reason
that the 2CP strains do not use nitrate before 2-CP may be that nitrite
accumulation would be inhibitory to their growth and therefore they must reduce
the nitrite further to ammonia. This apparent selective advantage of using the
energy available from reductive dechlorination is indirectly suggestive of a
halorespiratory metabolism for the 2CP strains. In order to gain an equal or
greater benefit energetically from reductive dechlorination over the respiratory
reduction of nitrate to ammonia, an electron transport chain most likely must be
involved.

The ability to use acetate as the sole carbon and energy source is
generally considered to be limited to the domain of respiring microbes, with the
exception of acetoclastic methanogens. Acetate utilization therefore provides
evidence of halorespiration in the 2CP strains. A respiratory mechanism is
confirmed when hydrogen provided as an electron donor is consumed. H2
uptake only occurs when 2-CP was present (Figure 3.6). As H2 could serve as
the electron donor for growth, the plausible explanation was that hydrogen
oxidation is coupled to 2-CP reduction via an electron transport chain. Other
evidence of halorespiration by the 2CP strains is apparent in the determination
of the fa and the measurement of hydrogen uptake correlated to the presence
of 2-CP. The fe accounts for the portion of electrons from the electron donor
used for the reduction of the electron acceptor (Criddle et al. 1991). The
remaining fraction of electrons is designated the fs; those that are incorporated
into biomass synthesis. Since fe + fs = 1, the fs is calculated by subtracting

the to from 1. In the case of the 2CP strains grown on acetate, this gave an fs

 

 

66

of 0.36, which is less than what is expected for denitrifiers, but greater than
expected for sulfate reducers (McCarty 1975; Criddle et al. 1991).

The hydrogen threshold concentration of less than 0.5 ppm also
suggests that halorespiratory-mediated growth was occurring in the 2CP
strains. The hydrogen threshold concentration reflects the minimum amount of
energy that an organism requires to grow (Cord-Ruwisch et al. 1988; Lovley
and Goodwin 1988). Hydrogen threshold concentrations are therefore
correlated with the energy available from the redox couples mediating their
growth. In the case of the 2CP strains, this is for H2 oxidation and 2-CP
reduction to phenol. The threshold concentration of H2 measured is
comparable to that observed for fumarate reduction and slightly greater than
found for nitrate respiration (Cord-Ruwisch et al. 1988). However, it is much
less than observed for sulfidogens, methanogens and acetogens. DeWeerd ,
Concannon and Suflita (1991) measured a comparable threshold concentration
of 0.7 ppm for hydrogen with the halorespirer Desu/fomonile tiedjei strain D08-
1 when grown on H2 and 3-chlorobenzoate. This indicates that hydrogen
threshold concentrations may be reliable indicators of halorespiratory
metabolism in pure and mixed cultures where a halogenated substrate is the
only electron acceptor provided other than CO2.

One interesting feature of the microcosms from which the 2CP strains
were enriched and isolated was the apparent sensitivity to nitrate
concentrations greater than 5 mM (Chapter 2). 2CP-C came from a tropical soil
microcosm which had no dechlorination activity above 1 mM nitrate. In growth
assays, nitrate concentrations were always 5 mM or less with the 2CP strains.
The use of low concentrations in enrichments and in culturing is in contrast to
previous strategies for isolating such organisms, but more reasonably models in

situ environmental conditions. With the exception of fumarate, the 2CP strains

67

only grew well when low concentrations (< 2.5 mM) of nitrate, 02 or 2-CP were
provided. It is possible that these anaerobic myxobacteria represent a
previously uncharacterized life-style, that of the microrespirotroph. Such a
microbial population would be sensitive to high concentrations of electron
acceptors much like an oligotroph is sensitive to high concentrations of carbon
substrate. The microrespirotroph life-style would also explain why microbes like
the 2CP strains have not been recovered before, since enrichment and isolation
methods used in the past usually provided excess electron acceptor, which by
definition would inhibit the growth of such organisms. The fortuitous use of low
chlorophenol concentrations, mainly for the purpose of toxicity reduction, may
have led to the discovery of this life-style in the 2CP microorganisms. This may
be of benefit for the development of new culturing strategies for uncovering

some of the untapped microbial diversity that exists in nature.

References

Burchard R.P. 1984. Gliding motility and taxes, p. 139-161. In E. Rosenberg
(ed.), Myxobacteria. Development and cell interactions. Springer-Veriag,
New York

Cole J.A., A. Cascarelli, W.W. Mohn and J.M. Tiedje. 1994. Isolation
and characterization of a novel bacterium growing via reductive
dechlorination of 2-chlorophenol. Appl. Environ. Microbiol. 60:3536-3542.

Cord-Ruwlsch R., l-I.-J. Seltz and R. Conrad. 1988. The capacity of
hydrogenotrophic anaerobic bacteria to compete for traces of hydrogen
depends on the redox potential of the terminal electron acceptor. Arch.
Microbiol. 149:350-357.

Criddle C.S., L.M. Alvarez and P.L. McCarty. 1991. Microbial
processes in porous media, p. 641-691. In J. Bear and M. Y. Corapcioglu
(ed.), Transport processes in porous media. Kluwer Academic Publishers,
Dordrecht

de Jong E., J.A. Fleld, H.-E. Splnnler, J.B.P.A. Wijnberg and J.A.M.
de Bont. 1994. Significant biogenesis of chlorinated aromatics by fungi
in natural environments. Appl. Environ. Microbiol. 60:264-270.

 

 

68

DeWeerd K.A., F. Concannon and J.M. Suflita. 1991. Relationship
between hydrogen consumption, dehalogenation, and the reduction of
sulfur oxyanions by Desulfomonile tiedjei. Appl. Environ. Microbiol.
57: 1 929-1 934.

Dolflng J. 1990. Reductive dechlorination of 3-chlorobenzoate is coupled to
ATP production and growth in an anaerobic bacterium, strain DCB-I.
Arch. Microbiol. 153:264-266.

Dolfing J. and B.K. Harrison. 1992. Gibbs free energy of formation of
halogenated aromatic compounds and their potential role as electron
acceptors in anaerobic environments. Environ. Sci. Technol. 26:2213-
2218.

Gribble G.W. 1992. Naturally occurring organohalogen compounds -- a
survey. J. Nat. Prod. 55:1353-1395.

Lovley D.R. and S. Goodwin. 1988. Hydrogen concentrations as an
indicator of the predominant terminal electron-accepting reactions in
aquatic sediments. Geochim. Cosmochim. Acta 52:2993-3003.

McCarty P.L. 1975. Stoichiometry of biological reactions. Prog. Water Tech.
7:157-172.

MIkeseIl M.D. and S.A. Boyd. 1986. Complete reductive dechlorination
and mineralization of pentachlorophenol by anaerobic microorganisms.
Appl. Environ. Microbiol. 52:861-865.

Mohn W.W. and J.M. Tiedje. 1990. Strain DCB-1 conserves energy for
growth from reductive dechlorination coupled to formate oxidation. Arch.
Microbiol. 153:267-271 .

Nicholson D.K., S.L. Woods, J.D. Istok and D.C. Peek. 1992.
Reductive dechlorination of chlorophenols by a pentachlorophenol
acclimated methanogenic consortium. Appl. Environ. Microbiol. 58:2280-
2286.

Reichenbach H. 1993. Biology of the Myxobacteria: ecology and taxonomy,
p. 13-62. In M. Dworkin and D. Kaiser (ed.), Myxobacteria II. American
Society for Microbiology. Washington, DC

Reichenbach H. and M. Dworkin. 1992. The myxobacteria, p. 3416-3487.
In A. Balows, H. G. Tri'rper, M. Dworkin, W. Harder and K. H. Schleifer
(ed.), The Prokaryotes. Springer Verlag, New York

Sanford R.A., J.R. Cole, F.E. Léffler and J.M. Tiedje. 1996.
Characterization of Desulfitobacterium halorespiricans sp. nov. strain
C023, which grows by coupling the oxidation of lactate to the reductive

69

dechlorination of 3-chIoro-4-hydroxybenzoate (3Cl-4-HBA). Submitted to
Bicdegradation .

Shlmkets L. and 0.R. Woese. 1992. A phylogenetic analysis of the
myxobacteria: basis for their classification. Proc. Natl. Acad. Sci. 89:9459-
9463.

Thauer R.K., K. Jungermann and K. Decker. 1977. Energy
conservation in chemotrophic anaerobes. Bacteriol. Rev. 41:100-180.

Zusman D.R. 1984. Developmental Program of Myxococcus xanthus, p. 185-
213. In E. Rosenberg (ed.), Myxobacteria. Development and cell
interactions. Springer-Veriag, New York

Chapter IV

Characterization of Desulfitobacterium chlororesplrans sp. nov.
strain 0023, which grows by coupling the oxidation of lactate to the
reductive dechlorination of 3-chIoro-4-hydroxybenzoate

The work described here was completed by Robert A. Sanford with the
exception of the 168 rRNA phylogenetic analysis completed by James R. Cole
and the deterination of the temperature stability of dechlorination activity done
by Frank E. L0ffler.

70

 

71

Abstract

Strain 0023, an anaerobic spore-forming microorganism, was
enriched and Isolated from a compost soil based on Its ability to
grow using 2,3-dlchlorophenol (DCP) as Its electron acceptor.
Ortho chlorines were removed from poly-substituted phenols, but
not from monohalophenols. Growth by chlororesplratlon was
Indicated by a growth yield of 3.24 9 cells per mole of H2
equivalents from lactate oxidatlon to acetate In the presence of 3-
chloro-4-hydroxybenzoate (30l-4-HBA), but no growth In the
absence of the halogenated electron acceptor. Other Indicators of
halorespiration were the fraction of electrons from the electron
donor used for dechlorination (0.67) and the H2 threshold
concentration of <1.0 ppm. Addltional electron donors utilized for
reductive dehalogenatlon were pyruvate, formate, butyrate,
crotonate and H2. Pyruvate supported homoacetogenlc growth In
the absence of an electron acceptor. Strain Co23 also used sulfite,
thlosulfate and sulfur as electron acceptors for growth, but not
sulfate, nitrate or fumarate. The temperature optimum for growth
was 37°C, however the rates of dechlorination were optimum at
45°C and activity persisted to temperatures as high as 55°C. The
16S rRNA sequence was determined and strain C023 was found to
be related to Desulfitobacterium dehalogenans, with a sequence
similarity of 97.2%. The phylogenetic and physiologlcal properties
exhibited by strain C023 place It as a new species designated

Desulfitobacterium chlororesplrans.

 

72

Introduction

Chlorophenolic compounds have been of environmental concern due to
their extensive use in several industries and their toxicity. This has led to
considerable efforts to study the degradation of these compounds under
aerobic and strictly anaerobic conditions (Haggblom 1992; Mohn and Tiedje
1992), Pentachlorophenol (PCP), a widely used wood preservative, is
degraded under methanogenic conditions via a series of reductive
dechlorinations (Mikesell and Boyd 1986; Mikesell and Boyd 1988; Nicholson
et al. 1992; Parker et al. 1993; Juteau et al. 1995) and under aerobic conditions
by several pure bacterial cultures via hydroquinone intermediates (Stanlake
and Finn 1982; Middeldorp et al. 1990). Other polychlorinated phenols and
monochlorophenols have been observed to degrade in a similar fashion under
anaerobic and aerobic conditions (Boyd et al. 1983; Gibson and Suflita 1986;
Hrudey et al. 1987; Genthner et al. 1989; Dietrich and Winter 1990; Hale et al.
1990; Zhang and Wiegel 1990; Madsen and Aamand 1992).

Anaerobic dechlorination has been widely observed in nature, although
few pure cultures capable of coupling growth to this process via halorespiration
have been isolated (Dolfing 1990; Mohn and Tiedje 1990; Holliger et al. 1993;
Cole et al. 1994). Halorespiration refers to the ability of a microorganism to
utilize the considerable energy released during reductive dehalogenation as
the terminal process in a respiratory electron transport chain. Free-energies
(AG°') available from reductive dechlorination have been reported in the range
of -140 to -160 kJ/reaction for chlorinated aromatic compounds (Dolfing and
Harrison 1992) and -130 to -170 kJ/reaction for chlorinated aliphatic
compounds (Dolfing and Janssen 1994). Desulfomonile tiedjei strain DOB-1

uses 3-chlorobenzoate (3-CBA) as a terminal electron acceptor and was the

73

first organism isolated capable of growing in this manner (Shelton and Tiedje
1984). The anaerobic myxobacteria strains, 2CP-1, 2CP-2, 2CP-C and 2CP-3,
all have been shown to grow at the expense of acetate oxidation and 2-
chlorophenol (2-CP) dechlorination (Cole et al. ; Cole et al. 1994). More
recently Desulfitobacterium dehalogenans , a gram-positive microorganism,
' was described with the ability to remove ortho-substituted chlorines from
polychlorinated phenols (Utkin et al. 1994). Direct evidence, however, for
growth associated with halorespiration was not presented for this strain,
although cell yields increased in the presence of a chlorinated substrate.
Tetrachloroethene (PCE) has also been shown to serve as an electron acceptor
for growth of two new genera, Dehalobacter restrictus and Dehalospirillum
multivorans (Holliger et al. 1993; SchIoIz-Muramatsu et al. 1995).

Previously we described microcosms and enrichment cultures with the
ability to dechlorinate 2,3-DCP to 3-CP and subsequently to phenol and
methane (Sanford and Tiedje 1996). Here we describe the isolation of strain
C023, a spore forming relative of Desulfitobacterium dehalogenans , with the
ability to grow using lactate as an electron donor and several haloaromatic

compounds as electron acceptors.

Materials and Methods
Growth conditions. Cultures were grown in 160 ml serum bottles with
100 ml of boiled degassed medium or in 30 ml anaerobic culture tubes with 20
ml of medium and closed with butyl rubber stoppers. The mineral salts medium
(Stevens et al. 1992) and vitamin solution compostion (Wolin et al. 1963) were
as described previously (Chapter 2) Cultures were incubated at 30 ° C unless

indicated otherwise.

 

 

74

The solid medium used for culturing strain 0023 was identical to the
basal medium with the following additions: 15 g/l Bactoagar, 10 mM pyruvate,
1.0 g/I yeast extract and 10 mM 30l-4-HBA. R2A agar medium (Difco) was also
used to grow Co23. Plates were incubated in an anaerobic glove-box under a
97% N2: 3 % H2 atmosphere.

Liquid cultures of strain C023 were grown on the mineral medium
amended with 20 mM lactate and 1 mM 30l-4-HBA. After dechlorination had
occurred two 1.0 % transfers were made to 20 ml of media, one with lactate and
3Cl-4-HBA and one containing lactate only. Additional amendments of SCI-4-
.HBA (2 mM each time) were made after its dechlorination to 4-hydroxybenzoate
(4-HBA) until the lactate had been depleted. In order to ensure continuous
growth, 3Cl-4-HBA was never allowed to be completely removed. Growth was
monitored by measuring absorbance in the anaerobic culture tubes at 600 nm.
The cell yield was determined by dry-weight measurements from 100 ml of
culture after cells had consumed 20 mM lactate. Cells were concentrated by
centrifugation in corex glass tubes to a volume of 5 ml, which was subsequently
filtered through a pre-weighed 0.2 pm filter. The filters were dried for 2 h at 104°
C, allowed to cool in a desiccator and weighed until a constant mass was
recorded.

Determination of electron donors and acceptors. The ability for
strain 0023 to reductively dechlorinate 2,3-DCP with different electron donors
was determined in duplicate 20 ml culture tubes incubated at 30° C. Acetate,
formate, butyrate, succinate, propionate, fumarate, lactate, pyruvate, crotonate
and H2 were tested as potential electron donors. The volatile fatty acids (VFAs)
were added to concentrations of 0.5 or 2.0 mM. H2 (18 moles or 6.6 % v/v)

was added to the headspace of the duplicate culture tubes. 2,3-DCP (100 uM)

75

'or 1 mM 3Cl-4-HBA were added as electron acceptors. A 0.5 % inoculum was
added to each tube from a butyrate, formate and 2,3-DCP grown culture. The
disappearance of 2,3-DCP and appearance of 3-CP were monitored by HPLC.
2,3-DCP was replenished three times at concentrations of 100 uM to those
cultures exhibiting dechlorination activity. 4-HBA appearance was monitored in
cultures receiving 3Cl-4-HBA as an electron acceptor. VFA utilization was
monitored by HPLC and H2 use was determined by GC. Dechlorination and
depletion of electron donor constituted a positive test for growth.

The range of electron acceptors used by strain 0023 was determined
using the same growth conditions as for the electron donor determination, but
with lactate (5 mM) or butyrate (5 mM) serving as the electron donors. The
halogenated electron acceptors tested were 100 pM of 2,3-DCP; 2,4-DCP; 2,5-
DCP; 2,6-DCP; 2-CP; 3-CP; 4-CP; 3Cl-4-HBA; 3-chlorobenzoate (CBA); o-
fluorophenol; o-bromophenol; o-iodophenol; 2,3,5-TCP; 2,4,6-TCP; 2,4,6-
tribromophenol; pentachlorophenol (PCP); 3-chloroanisaldehyde; 3-chloro-L-
tyrosine; and 3-Cl-4-hydroxyphenylacetate. Nitrate, sulfate, sulfite, thiosulfate
and fumarate were also tested at a concentration of 5 mM. Sulfur was tested as
an electron acceptor by adding 0.1 ml of a powdered S° slurry to one set of
tubes. Tubes were inoculated with a 0.1 to 0.5 % inoculum from cultures grown
on butyrate and 2,3-DCP or lactate and SCI-4-HBA. The concentrations of
halogenated substrates were monitored by HPLC and were replenished once
depleted. Nitrate and fumarate concentrations were measured by HPLC. Use
of sulfur compounds was determined by measuring growth as indicated by
absorption at 600 nm as compared to a lactate-only control culture. Lactate and

butyrate consumption were monitored by HPLC to verify physiological activity.

76

Pasteurization and microscopy. To test for the presence of heat-
resistant spores, fresh anaerobic medium containing lactate and SCI-4-HBA
washeated to a temperature of 80° C. A 1.0% inoculum from a one week old
C023 culture was added to duplicate bottles, which were immediately placed
back in an 80° C water bath for 10 min. The cultures were removed from the
water bath and placed in the 30° C incubator and monitored for dechlorination
activity and consumption of lactate.

Microscopic observations were done as previously described in Chapter

Temperature effect on growth and dechlorination. Cultures of
0023 were inoculated into anaerobic culture tubes at temperatures ranging
from 15° C to 75° C with 20 mM pyruvate and 1 mM 3Cl-4-HBA. Growth was
monitored by measuring absorbance in the anaerobic culture tubes at 600 nm.
To determine the temperature stability of the dechlorination activity in the
absence of growth, C023 was grown at 37° C on 20 mM pyruvate and 1 mM
3Cl-4-HBA until stationary phase had been reached, about 48 h. The cell
suspension was aliquoted (10 ml) into separate culture tubes which were
amended with 4 mM 3CI-4-HBA and 10 mM pyruvate and incubated at
temperatures ranging from 20° to 65° C. The rate of dechlorination in these
cultures was monitored over the next 3 days by HPLC analysis.

16S rRNA gene Isolation, sequencing and analysis. Cells of
C023 were grown in 100 ml of lactate and SCI-4-HBA medium. Total DNA was
isolated from C023 by a method shown to work on diverse bacteria
(Visuvanathan et al. 1989). The primers used to amplify near full length 168
rRNA gene sequences (5'AGAGTTTGATCCTGGCTCAGS' and
5'AAGGAGGTGATCCAGCC3') were modified from the primers FD1 and ROI of

77

Weisberg et al. (1991) by removing the linker sequences. The PCR reaction
consisted of 1.5 mM MgCI2, 0.2 mM of each dNTP, 0.25 uM of each primer, 1x
Taq polymerase buffer, 0.75 UTaq polymerase (Promega, Madison, WI), and
0.1 pg DNA in a volume of 30 ul. Amplification was carried out using a
GeneAmp PCR system 9600 thermocycler (Perkin Elmer, Norwak, CT) with a
program consisting of an initial denaturation at 92' C for 2 min 10 s, 30 cycles of
94' C for 15 s, 55' C for 30 s, and 72' C for 2 min 10 s, followed by a final
elongation cycle at 72' C for 6 min 10 s.

The resulting PCR product was purified by gel electrophoresis through a
1% agarose gel and recovered using Gene Clean purification resin according
to the manufacturer's suggestions (Bio 101, La Jolla, CA). The purified PCR
product was cloned in the vector pCRII, using a TA cloning kit (Invitrogen, San
Diego, CA). Plasmid DNA containing the 168 rRNA gene insert was isolated
from one clone using the Qiagen plasmid mini kit (Qiagen, Chatsworth, CA).

The DNA sequence of the insert was determined by automated
fluorescent dye terminator sequencing using an ABI Catalyst 800 laboratory
robot and ABI 373A Sequencer (Applied Biosystems, Foster City, CA). Primers
used corresponded to conserved regions of the 168 sequence (Woese 1987).
Approximately 95% of the insert sequence was determined in both directions.

Initial phylogenetic placement was obtained using the Ribosomal
Database Project (RDP) (Larsen et al. 1993) electronic mail server program
SIMILARITY_RANK. Related sequences and a preliminary alignment were
obtained using the RDP programs SUBALIGNMENT and ALIGN_SEQUENCE.
The alignment was completed using the sequence editor GDE obtained from
the RDP. A maximum-likelihood phylogenetic tree was created using the

fastDNAml (Olsen et al. 1994), using a weighting mask to include only

78

unambiguously aligned positions with all other program options at their default
values. This analysis was repeated on 100 bootstrap samples to obtain
confidence estimates on the branching order (Felsenstein 1985). The program
consense from PHYLIP (Felsenstein 1989) was used to determine the number
of times that each group shown in the final tree was monophyletic in the
bootstrap analysis. The final tree was then produced using the program
TreeTool from the RDP.

The 168 rRNA sequence of Clostridium sp. DMC ( EMBL accession no.
X86690) was obtained from the European Bioinformatics Institute web server
(www.ebi.ac.uk). The following sequences were obtained from the RDP
(GenBank accession number in parenthesis): Desulfotomaculum orienfis str.
Singapore I (M34417), Desulfitobacterium dehalogenans str. JW/lU-DC1
(L28946), Peptococcus niger (X55797), Heliobacterium chlorum (M11212),
Desulfotomaculum nigrificans (X62176), Desulfotomaculum ruminis str. DL
(M34418), Bacillus subtilis str. 168 (K00637 M10606 X00007), and Escherichia
coli (J01695).

Chemical analyses. Chlorophenols, dichlorophenols and aromatic
dechlorination products were analyzed on a Hewlett Packard 1050 HPLC with a
Chemstation analysis package. The eluent was phosphoric acid (0.1 %)
buffered methanol pumped at 1.5 mein using a gradient from 48 % to 55 %
methanol. A Hibar RP-18 (10 um) column was used. Peaks were quantified at
218 nm on a UV multiwavelength detector. Samples (1 ml) from the cultures
were taken, made basic with 10 ul of 2N NaOH, centrifuged for 6 min in a
microfuge and filtered through Acrodisc LC13 PVDF 0.45 pm filters prior to
HPLC analysis.

79

Nitrate and nitrite were analyzed by HPLC using a Whatman Partisil 10
SAX column on a Shimadzu HPLC system. The eluent was 50 mM phosphoric
acid buffer (pH = 3.0) pumped at a rate of 1 ml/min. UV adsorption at 210 nm
was used for detection.

Volatile fatty acids (VFAs) were analyzed using the Shimadzu HPLC with
a BioRad Aminex HPX-87H ion exclusion column heated to 60°C and using
0.005 N H2SO4 as the eluent (Guerrant et al. 1982). Previously filtered
samples were acidified to 0.25 N H2SO4 by adding 100 pl of 2.5 N H2SO4 to
900 pl of sample. Eluent was pumped at 0.6 ml/min and detection of VFAs was
at 210 nm by a UV detector.

The headspace of the cultures was analyzed for H2 and CO2 using a
Carle gas chromatograph equipped with a 1.83 m Porapak O column and a
thermal conductivity detector (T CD). Headspace pressure was normalized to
atmospheric by venting with a needle prior to removing 0.3 ml of gas for
injection into the GC. Threshold concentrations of H2 were measured using a
GC equipped with a reduction gas detector from Trace Analytical, Menlo Park,
CA

Chemicals. Chlorophenols, dichlorophenols, 3Cl-4-HBA, and 3-CI-4-
hydroxyphenylacetate were obtained from Aldrich Chemical Co. 4-HBA and 3-
Cl-L-tyrosine were purchased from Sigma Chemical Co. 3-Chloro-

anisaldehyde was provided by Roberta Fulthorpe, University of Toronto.

Results
Isolation of Desulfitobacterium chlororesplrans strain C023.
Enrichment cultures inoculated from microcosms of compost soil (Chapter 2)
that exhibited complete dechlorination of 2,3-DCP were fed with 2,3-DCP and

acetate and accumulated 3-CP. 2,3-DCP repeatedly added to this enrichment

80

culture was completely dechlorinated in the ortho position. This activity was
retained after transfer to new enrichment medium. Anaerobic agar shake
cultures inoculated with the secondary enrichments yielded isolated colonies.
Several colonies that were transferred to anaerobic broth medium containing
butyrate and formate dechlorinated 2,3-DCP. A survey of electron donors of
one of the cultures showed that acetate no longer supported dechlorination of
2,3-DCP. Since 3-0P was inhibitory to growth and dechlorination activity at
concentrations above 1 mM, additional chlorinated substrates were evaluated
which would not yield inhibitory products and therefore result in higher cell
yields. 30l-4-HBA was shown to be the optimum electron acceptor for growing
the culture since the product of dechlorination, 4-HBA, was not inhibitory at
concentrations below 20 mM. The culture was then purified by dilution of
dechlorination activity to extinction in medium containing 3Cl-4-HBA, butyrate
and formate. The purity of the resulting culture, designated strain 0023 (for
compost soil origin), was confirmed by streaking culture fluid onto agar medium.
Only one colony type was observed on both 3Cl-4-HBA and R2A agar media.
Representative isolated colonies were transferred back into broth culture and
shown to dechlorinate 30l-4-HBA.

The isolate is a motile, gram negative-staining, curved bacillus 3 to 4 pm
long by 1 pm wide (Figure 4.1). Terminal spores are present in some cells. To
evaluate whether the refractal bodies (Figure 4.1) were heat resistant spores,
duplicate cultures of 0023 were pasteurized. After exposure to 80° C,
dechlorination and growth resumed after one week of incubation, confirming
strain 0023's ability to survive the heat treatment.

Evidence for halorespiration. The ability of strain 0023 to couple

reductive dechlorination to growth was determined by measuring increases in

81

t" ‘1"): \-'\r’. ’\"'1"‘IF\Y’I C” .‘ {’7‘ o—‘r . (’3

"- . o "- I I“, v o , ~. ‘ k
. 2.;- e-td' ”4‘- W ens/17°“- .: 7&5:
"~:H1.‘r"‘ '5‘1fi514’

 

Figure 4.1. Phase-contrast photomicrograph of strain 0023. Vegetative cells
and the presence of spores and terminal swelling in cells developing spores
are shown. Reference bar is 13 um.

82

cell density in the presence and absence of 3Cl-4-HBA. The cell density
increased in lactate-fed cultures only when 30l-4-HBA was added (Figure 4.2).
Stoichiometric generation of 4-HBA and acetate was observed from the 3CI-4-
HBA and lactate, respectively. Lactate was not oxidized in the absence of an
electron acceptor. The pH was periodically adjusted with 2 N KOH to 7.2 during
growth because dechlorination activity was inhibited when the pH reached 6.2.

Additional evidence of halorespiration was provided in measurements of
the hydrogen threshold and the fraction of electrons from the electron donor (fe)
being used to reduce 3CI-4-HBA. Hydrogen was consumed within a week to a
threshold level of less than 1 ppm in duplicate cultures with excess 30l-4-HBA
present in the medium. The fe was determined by plotting the equivalent
lactate concentration as pmoles H2 used for oxidation to acetate versus the
umoles 4-HBA generated in the culture (Figure 4.3). The slope of the resulting
regression line is equivalent to the fa, which was determined to be 0.67.
Values of 0.69 were calculated with cultures that used butryate and formate as
electron donors.

The mass balance of substrate use and cell yield when grown on lactate
plus 30l-4-HBA is shown in Table 4.1. The cell-yield expressed as H2
equivalents used from growth on lactate was 3.23 9 cell dry weight/mole H2
equivalent and the yield expressed on the basis of 30l-4-HBA dechlorinated
was 6.90 9 cell dry weight/ mole 30l-4-HBA. With butyrate and formate as
electron donors the yield was 1.69 9 cells/ mole H2 equivalent.

Range of electron donors and acceptors used. Strain 0023 was
capable of using a wide range of electron donors for reductive dechlorination
(Table 4.2). Several substrates supported growth of strain 0023 with 30l-4-

HBA or 2,3-DCP as electron acceptors. Growth was indicated by the measured

83

 

 

 

 

0.15,
0.14}
0,12 ’. lactate + 3CI-4-HBA "
I a e
’ 0
g 0.1:—
§ 0.08} n
8 0.05} . ' 0' '
0.04- , ’ A ___
i :4 ‘ .,A----""" A
0.02 — , / _‘_f.9-"A lactate only
“- -a—-- a A
0M-1-W1
100 200 300 400 500

Figure 4.2. Growth of strain 0023 as indicated by an increase in turbidity (OD
600 nm). Duplicate cultures with lactate and 3-Cl-4-HBA are shown compared

to a control culture containing lactate alone.

84

 

500 ’1‘)
. _.-‘
. ..-"
l ,a‘
A _ ,-
3.3 : Q".
(E) 300 _ fe = 0.67 {.5
p 2 - I
E"; ’ r .. 0.997 .3"-
. .d'
$ 200- a."
I ,3"
<1 i 90'
100— 16¢.-
' I g.-‘b
9'... l-.--l----l----1-- 1+--1--1

 

 

 

O 100 200 300 400 500 600 700 800
umoles consumed as H2
Figure 4.3. Graphical determination of fe, the fraction of electrons from the

electron donor going to the electron acceptor. Electrons from lactate are plotted
as umole equivalents of H2 generated in the incomplete oxidation to acetate.

85

.000E0.0 0.0. 0200000000. .0000 00000.0 E00. 0000.020 .0 00000.. n 0. 0
.2000100 .0 00_._000E00 000E0.0 00.E0000 .0: .0 .0903 0.:E00. 0 00 00000 000E :00 05 .0 00.0220 0.0EE 0

0.00500000 0. .0220. 000000000 5000 000 000E 00 03000 00.0..» 0...... 00.02.00. 00...... 0

 

 

 

 

000 00.0 00.. 0.0000. 0.0 0... 00.. 0.05:0
000 0.0000“.

.0000. 0.0 00.0 00.. o 0.050..

0000.0 0.2.38.0 0.0.0. 0.0 .20 .000. 0.0 00.. 40.0 00.003
0.0000. 0 00.. 03.0 < 0.003

0. 0.00.100 .2000 01: 6.0551... 2880 (0:4 .23. 32.3... 23.00

0:00 E2. 0...
03003120003130.» «507.151.0050

 

00203-00 0 00

0:00 .0 0.0.» 000 00E:0000 $1.100 000 0.0500 000 0.0000. 0.080.. 0.0000 00000.0 .0 0000.00 000.2 .3. 0.00...

 

86

Table 4.2. Electron donors tested for use by strain 0023 in a minimal medium.
2,3-DCP or 3-0l-4-OH-benzoate served as electron acceptors. A positive
score indicated growth which was monitored by measuring the depletion of
electron donor and acceptor. Cultures were grown at 30° C.

 

 

 

 

h iroduct (moles per
Substrate(moles) 3CI-4-HBA 2,3-DCP mole substrate)
Acetate(1) ' ' ND
Formate(1) +3 +b none
Succinate(1) ' " N D
Propionate(1 ) ' " N D
Butyrate( 1 ) + + acetate (2)
Crotonate ”d + acetate
H2 -I-a +a none
Fumarate(1) 11d - N D
Pyruvate(1) +° I'd acetate(l .0)
Lactate(1) + nd acetate(1.0)

 

a Growthoocltredonlyifacetatewaspresentasasouceofcarbon.
b Formate utilization observed in the presence of butyrate.

° Pyruvate smported growth both fermentatively and in the presence of an electron acceptor.
ND -- not determined

 

87

consumption of both electron donor and acceptor over three successive
feedings. All of the organic acid electron donors, except formate, were
incompletely oxidized to acetate. Pyruvate also supported growth in the
absence of an electron acceptor and was fermented stoichiometrically to
acetate.

Growth of strain 0023 on lactate and several different electron acceptors,
as measured optical density, is shown in Figure 4.4. No increase in turbidity
occurred with lactate alone or with lactate plus sulfate. Sulfite, thiosulfate and
sulfur all supported growth as electron acceptors, but sulfate, nitrate and
fumarate did not. Several halogenated phenols supported growth of strain
0023 (Table 4.3). Ortho-substituted chlorines were removed from
polychlorinated phenols with two or greater halogen substituents, with the
exception of 2,4- and 2,5-DCP. Mono-substituted halogenated phenols were
not utilized by strain 0023. 30l-4-HBA and 3-0l-4-hydroxyphenyl-acetate were
the only mono-chlorinated substrates used as electron acceptors.

Growth and dechlorination rates. Due to the difficulty of
maintaining sufficient 30I-4-HBA in the culture medium to sustain exponential
growth of strain 0023, growth rates were measured using pyruvate as the sole
carbon and energy source. Additions of only 2 - 3 mM 3CI-4-HBA were
possible, since higher concentrations inhibited growth. Strain 0023 exhibited
typical exponential growth with a doubling time (td) of 15.4 h when grown on
pyruvate at 30° C (Figure 4.5). When tested at several temperatures, growth
was observed from 15° to 37° 0, with the optimal growth rate expressed as 1.1...

(1/td) equal to 0.17 h'1 at 37° C (Figure 4.6). No growth was observed above
45° C.

88

 

0.16

‘ olf'flv.’..&
0.14 7 .v'u—ruw'
012 ‘1' AKA.
. a A. .
g , .-
8 01 T .l .' __., —— ‘
o 0.08 '— ‘. a. w .
D
O

 

 

 

 

Figure 4.4. Growth of strain 0023 on different electron acceptors and lactate as
measured by increases in turbidity (OD 600 nm). The symbols are as follows:
sulfur (El), sulfite (A), thiosulfate (A), sulfate (X), 30l-4-hydroxyphenylacetate
(OI), 30l-4-HBA (O), and lactate (V). Data are the average of duplicate

cu tures.

89

Table 4.3. Electron acceptors tested with strain 0023 with (+) indicating
support of growth and H indicating no growth. Growth was determined by
measuring the depletion of the electron donor and electron acceptor as well
as observing visual increase in culture turbidity. Lactate or butyrate were used
as electron donors. Cultures were incubated at 30° C.

 

 

Electron icceptor fiGrowth
2-0P -
3-CP -
4-CP -

2-Fluorophenol -
2-Bromophenol -
240dophenol -
2,3-DCP (3-CP)a +
2,4-DCP ..
2.5-DCP -
2,6-DCP (2-CP) +
2,4,6-TCP (4-CP) +
2,4,6-Tribromophenol (4-BrP) +
2,3,5-TCP -
POP -
30l-4-HBA (4-H BA) 4.
3-CBA -
3-Cl -4-OH-phenyI-acetate (4-OH-

phony-acetate)
3-0l-L-tyrosine ..
3-Cl-anisaldehyde -

Mm:

Sulfate -
Sulfite +
Thiosulfate +
Sulfur +
Nitrate ..
Fumarate -

 

 

 

a Products of dechlorination are noted in parentheses.

90

 

 

0.2
. .- __________ O
015- 9-0-0
’ 00‘
a i 8' 011 0
c > d : o‘g
a l a » ‘98,
3 0'1 ” 8’0 : O ",0
8 ’ 8' 0'01: "00'
005- Ag 0 Doubling time =15.4 hrs
: 0.001----I----I---.J----I----I----I----
g 010200.040506070
-l----l----olu.r§--l-.--

 

 

 

 

 

 

009.---1----1--
0 50 100

150 200 250 300 350
hours

Figure 4.5. Exponential growth of strain 0023 on pyruvate at 30° 0. Data are

averaged from triplicate cultures.

91

 

 

 

 

 

 

0.2 r l i i i ‘ 7
l 3 -lu
_. 00
: .6 E‘z’
,.. +0000.
1- ‘ 5'9
I ’ j a:
5 j4 3'3
x 0.1- 1 3‘
13 :3 55
: ‘57;
’ ‘ 2 9.0
0.05- , :0:
C 3 x?
. ‘ 1 fig
, 1 3:.
'0-A--- A. U ‘o
10 20 3O 40 50 60 70

T (°C)

Figure 4.6. Temperature dependence for growth and dechlorination by strain
0023. Growth rates were determined by averaging the llmax for duplicate
cultures. Dechlorination rates were determined with resting cells grown on
pyruvate at 37° 0 and incubated at various temperatures with 30l-4-HBA and
pyruvate. Also shown are the umoles of 30l-4-HBA dechlorinated over 23 h.

92

Dechlorination rates were determined for resting cells incubated at
temperatures from 15°C to 65° C. Dechlorination activity had measureable
rates at temperatures as high as 55° C and the maximal rate was observed at
45° 0 (Figure 4.6). Complete dechlorination of 4 mM 30l-4-HBA occurred
within 23 h at temperatures ranging from 25° to 45° 0 (Figure 4.6). Although
initial dechlorination rates at 50° C were significant, no further activity was
observed after 11 h of incubation (data not shown). At a growth temperature of
30° C, resting cells dechlorinated 30l-4-HBA at a rate of 3.44 (+/- 1.09) mmol h'
19'1 cells dry weight.

Phylogeny of strain 0023. Comparison of available 168 rRNA
sequences to that of strain 0023 indicated that strain 0023 is a member of the
Desulfotomaculum group of the gram-positive bacteria (RDP release version 5).
Pairwise similarity values calculated between 0023 and the available near
complete 168 sequences from the Desulfotomaculum group produced
similarity values ranging from 82.2% identity with Peptococcus niger to 97.2%
identity with Desulfitobacterium dehalogenans, a bacterium that is also able to
reductively dechlorinate chlorophenols (Table 4.4). Partial 16$ sequences are
avaliable for several other closely related bacteria. These sequences were
included in the construction of a maximum likelihood phylogenetic tree (Figure
4.7). Bootstrap analysis of the maximum likelihood phylogenetic tree confirmed

the close specific relationship between 0023 and D. dehalogenans.

Discussion
The most distinctive feature of strain 0023 is that it was able to couple
growth to halorespiration. The term halorespiration has been suggested for
those organisms that are capable of utilizing the considerable energy released

from reductive dehalogenation for oxidative phosphorylation. The AG°' for

93

Table 4.4. Pairwise similarity values for strain 0023 and selected bacterial
speciesa.

 

 

1 2 3 4 5 6 7

 

 

1) D. dehalogenans 100

2) D. chlororespirans str. 0023 97.2 100

3) P.niger 82.6 82.2 100

4) H. chlorum 87 86.3 82.6 100

5) Desulfotomaculum nigrificans 84.9 84.2 82.8 85.0 100

6) B.subtilis 83.7 83.7 80.8 82.4 83.1 100

7) E. coli 78.8 78.9 76.8 78.8 77.7 78.8 100

 

a Similarity values were calculated for 1384 aligned residues. Species as in
Figure 4.7.

 

94

96 Desulfitobacterium dehalogenans
Desulfitobacterium chloromplrlcans 0023
Desulfotomaculum on'entis

Peptococcus niger

Clostridium sp. DMZ

Heliobacterium chlorum
100,—— Desulfofomawlum mmInis
1——— Desullofomaculum nlg'ilicms

Bacilhls wblilis
Escherichia coli

 

 

 

  

 

 

 

0.10

 

Figure 4.7. Maximum likelihood phylogenetic tree. Numbers at internal nodes
are the percent of 100 bootstrap samples in which the group to the right of the
node was monophyletic. Scale is in expected number of substitutions per
positions.

95

reductively dechlorinating 2,3-DCP to 3-CP is -144.3 kJ/reaction (Dolfing and
Harrison 1992). When coupled to the oxidation of lactate (AG°' -2.1 kJ/mol H2)
or butyrate (AG°' +24.1 kJ/mol H2) the overall reaction is strongly exergonic
(Thauer et al. 1977). Strain 0023 is one of only a few microorganisms
described to grow by this process. The anaerobic myxobacterium strains 2CP-
1, 2CP-2, 2CP-C and 2CP-3 were the first microorganisms described that are
able to couple acetate oxidation to the reductive dechlorination of 2-
chlorophenol (Cole et al. ; Cole et al. 1994). Desulfitobacterium dehalogenans
may also grow via halorespiration, however yeast extract was always present in
the medium and growth in the absence of this additive at the expense of
reductive dechlorination has not yet been clearly established (Utkin et al. 1994).

The evidence for strain 0023's ability to grow with halogenated
substrates as electron acceptors is as follows. Lactate was shown to support
considerable growth with 30l-4-H8A as an electron acceptor, but no growth
occurred when only lactate was present. Similar results occurred with butyrate
and formate. The reducing equivalents from the partial oxidation of lactate to
acetate are partitioned to 3Cl-4-H8A dechlorination and to biomass. The 10
was determined to be 0.67, implying that 67 % of the available electrons from
lactate are used for reductive dechlorination. From the cell yield data it was
possible to calculate a value for fs, the fraction of electrons from lactate going to
energy for cell synthesis (Table 4.1). Hence fs + fe equals 0.96, which is close
to the theoretical optimum of 1.0. Therefore the data account for 96% of the
electrons used from lactate oxidation with 67% going to electron acceptor
reduction, which is consistent with the partitioning expected in respiratory

processes (Criddle et al. 1991). Using both the fa and fa values the following

96

equation accounts for the mass balance of lactate oxidation, 3Cl-4-H8A

reduction and cell yield:

0.25 lactate + 0.064 H20 + 0.355 3Cl-4-HBA + 0.0145 NH3 --
-> 0.214 acetate 4» 0.25 CO2 + 0.0145 C5H7O2N + 0.355 4-HBA
+ 0.355 HCI

Additional evidence for halorespiration is the measured H2 concentration
in cultures incubated for one week. Cord-Ruwisch et al. (1988) have shown that
the steady-state concentration of hydrogen in a culture is largely dependent on
the available energy associated with the reduction of the terminal electron
acceptor. Thus the H2 threshold concentration is when H2 oxidation is no
longer energetically favorable to the cell for the reduction of an electron
acceptor. This is reflected in the 'critical' free energy (AGc), which is the
minimum energy required that will permit growth (Seitz et al. 1990). Given this
relationship, one would predict that threshold concentrations would decrease in
the following order: H2:002 acetogenesis > methanogenesis> sulfidogenesis
> fumarate reduction 2 halorespiration 2 N03' reduction to NH3 > Fe+3
reduction > denitrification > O2 reduction. Our data show a H2 threshold of less
than 1 ppm, which is in good agreement with what is expected if haloreSpiration
was the process responsible for H2 consumption. Other halorespiring strains
were shown to lower H2 concentrations to 0.7 ppm and to less than 0.5 ppm for
Desulfomonile tiedjei DOB-1 (DeWeerd et al. 1991) and 2CP strains (Cole et
al. ), respectively.

Although phylogenetically similar to Desulfitobacterium dehalogenans

(Utkin et al. 1994) and physiologically similar to strain DOB-2 (Madsen and

97

Licht 1992), strain 0023 has several features that delineate it from these two
microorganisms (Table 4.5). Pyruvate is fermented homoacetogenically by
0023 and DOB-2, in contrast to D. dehalogenans which produces considerable
amounts of lactate (Utkin et al. 1994). Nitrate and fumarate support growth of D.
dehalogenans and 008-2, but do not support growth of strain 0023. Butyrate is
used as an electron donor for dechlorination by strain 0023 in contrast to the
case forD. dehalogenans . Although strain 0023 stains gram negative, the
Desulfotomaculum group does not yield a reliable gram stain (Campbell and
Singleton Jr. 1986). Utkin et al. (1994) demonstrated a gram-positive cell wall
for D. dehalogenans by TEM and, since strain 0023 is phylogenetically closely
related to D. dehalogenans, 0023 is likely to have the same type of cell wall
structure. Neither D. dehalogenans nor DOB-2 have yet been shown to grow
via halorespiration, since both were always cultured with considerable amounts
of yeast extract. Strain 0023 is not able to utilize 2.4-DCP while the other two
strains did, although cell-free assays have shown that the dehalogenating
enzyme system does have some activity toward this compound (L0ffler et al. ).
Also, the same cell-free assays showed that 3,5-DOP, a compound utilized by
DOB-2 (Madsen and Licht 1992), is not dechlorinated by 0023. Finally the
rates of dechlorination of 30l-4-H8A are more than an order of magnitude
greater for strain 0023 than D. dehalogenans (Table 4.5) (Utkin et al. 1995).
Strain 0023's 168 rRNA sequence places it close to Desulfitobacterium
dehalogenans , with a similarity of 97.2 % (Table 4.4). In addition, secondary
structure features of the 0023 sequence are consistent with those of D.
dehalogenans. This would at least place 0023 within the Desulfitobacterium
genus, but the sequence differences and especially key phenotypic differences

suggest that 0023 is a different species. Recently it has been shown that

98

Table 4.5. Comparison of strain 0023 with two other isolated dehalogenating
strains. The data for Desulfitobacterium dehalogenans and strain DCB-2 are
from Utkin et. al. (1994). and (Madsen and Licht, 1992) respemively.

 

 

 

D.
Tralt C 023 dehalogenans DC B-2
Gram sta'n ' + 1'
Spores + " +
Motility + + +
Haloresp'rationa + nd nd
«1031307033309 3.44b 0.04b nd
(mmol h'1g cells-1)
Grows readily on solid
medium6 '1' ' +
Pym“ “menial“ Acetate Acetate Acetate
9mm“ lactate
Wire:
I l l . l'
Lactate + + nd
Pyruvate + + ”d
H2 '1' '1' nd
Formate + + nd
Butyrate '1' ' I'ld
Yeast extract nd + +
W
2,4-DCP - + +
3,5-DCP ' ' + .
N03’ ' + +
Fumarate ' '1' Rd

 

a Growth observed with electron donor, lactate, and a halogenated phenol as an electron
acceptor, with no addition of yeast extract as a supplemental carbon and energy source.

b Dehalogenation rate by resting cells of 30l-4-HBA is shown on a cell dry-weight basis. Rate
data for D. dehalogenans is from (Utkin et. al, 1995).

c Colonies are easily propagated on anaerobic plate media with no special modifications to the
medium or inctbation conditions.

 

99

strains at 168 rRNA evolutionary distances of 2.5 to 3.0 (97 to 97.5% sequence
identity) are unlikely to share 70% DNAzDNA homology and therefore are likely
to be different species (Stackebrandt and Goebel 1994). The key physiological
differences are the inability of strain 0023 to use fumarate and nitrate as
electron acceptors and homoacetogenic growth on pyruvate, all important
physiological and taxonomic traits for anaerobes. Although strain 0023 and D.
dehalogenans share very similar chlorinated substrate use patterns, this feature
seems insufficient to group these two strains into the same species since it may
reflect only one biochemical character. A phylogenetic comparison with DOB-2
is not possible since the 16S rRNA sequence of this organism is not available,
however this strain shares some physiological characteristics with 0023 that
are not shared with Desulfitobacterium dehalogenans . For example 008-2
was reported to grow homoacetogenically on pyruvate and produced visible
spores (Madsen and Licht 1992). Although many features of comparison have
not yet been measured for DCB-2, it is unique in its ability to dechlorinate 3,5-
DCP to 3-CP. I

The specificity that strain 0023 exhibits for removing ortho-substituted
halogens from poly-halogenated phenols and the coupling to respiratory growth
suggests that this capability has evolved for some period of time. One
hypothesis is that natural halogenated compounds are ubiquitous and
abundant enough to select for and support novel anaerobic microbial
populations. A possible natural source of halogenated aromatic compounds
are fungi. Recently a white rot fungus was reported to generate concentrations
of up to 75 mg kg'1 of litter of chlorinated anisyl metabolites, eg. 3-
chloroanisaldehyde (de Jong et al. 1994). Although strain 0023 could not

directly use this compound, 3-Cl-anisaldehyde could reasonably undergo

100

demethylation and oxidation reactions to produce 30l-4-H8A, the best growth
substrate for halorespiration by strain 0023. Another naturally occurring
aromatic amino-acid, 3-chlorotyroslne (Gribble 1992), also was not
dechlorinated by strain 0023. Again, however, it is possible that degradation of
this compound in an anaerobic environment could result in 30l-4-HBA or 2-CP.
The presence of either 3-chloroanisaldehyde or 3-chlorotyrosine in the original
compost-soil microcosm, however, has not been established. Bromophenolic
compounds are produced by marine hemichordates and have been used to
enrich for an anaerobic debrominating microorganism (Steward et al. 1995).
Several other naturally occurring chlorophenolic compounds that could
potentially serve as substrates for halorespiring microorganisms are discussed
by Gribble (1992).

Strain 0023 has some features desired for bioremediaton, such as
relatively rapid growth, compared to previously isolated aryl-halorespirers.
Since growth is selected by the chlorinated substrate, the process is easier to
manage than if it were by cometabolism. Strain 0023 has also been shown to
dechlorinate a broad range of halogenated substrates in cell-free assays,
generally being able to remove most ortho-substituted chlorines from
pentachlorophenol to dichlorophenol (Leffler et al. ). The temperature stability
of the dechlorination activity may also be exploited by designing treatment
systems that operate at higher temperatures. For example at 45° C the
dechlorination activity would be stable, uncoupled from growth and more rapid
compared to 37° 0. Also the spore-forming capability should provide for long
term survival of inoculum in a habitat.

Strain 0023 is also a good candidate for basic studies on the mechanism

of halorespiration because it has faster growth rates and higher cell yields than

101

Desulfomonile tiedje DCB-I and the anaerobic myxobacterium 2CP-1 (Cole et
al. 1994). Since it is a gram-positive microorganism it is an important model
system for comparison to the dechlorination process in gram-negatives.
Description of Desulfitobacterlum chlororespirans sp. nov.
Desulfitobacterium chlororespirans (chlor.o.resp.ir.i.cans' M. L. part. adj.;
chloro referring to the group VII element chlorine; fr. L respirare to blow,
breathe; chlororespirans , breathing chlorine, referring to the characteristic of
coupling oxidation of electron donors to reductive removal of chlorines from
various chlorophenolic compounds via a respiratory process used for obtaining
energy for growth). Cells are slightly curved motile rods 3 to 5 pm long and 0.5
to 0.7 pm wide. Terminally located spores appear in late growth and cultures
are resistant to heat-treatment at 80° 0. Cells stain gram negative, but
phylogenetically this organism is within the gram-positive Desulfotomaculum
group. Growth is obligately anaerobic and sulfite, thiosulfate, sulfur and ortho-
substituted polychlorophenolic compounds are used as electron acceptors. 3-
Chloro-4-hydroxybenzoate is the best substrate for halorespiration. Pyruvate,
lactate, formate, butyrate and H2 are used as electron donors. Cells grow by
partial oxidation of carbon substrates to acetate coupled to the reductive
dechlorination of ortho-substituted chlorophenolic compounds. Growth of cells
on pyruvate alone is homoacetogenic. The optimal growth temperature is 37°
C. The pH range for growth is 6.8 to 7.5. Colonies grown on R2A agar medium
are white, round and smooth with a 1-2 mm diameter after one week of growth.
This organism was isolated from a Michigan residential compost soil

microcosms fed dichlorophenols and nitrate. The type strain is 0023.

102

Acknowledgments
This work was supported by the Center for Microbial Ecology through
grant No. BlR-912 0006 from the National Science Foundation and by a
Feodor-Lynen-fellowship from the Alexander von Humboldt-Stiftung to F.E.L.

References

Boyd S.A., D.A. Shelton, D. Berry and J.M. Tiedje. 1983. Anaerobic
biodegradation of phenolic compounds in digested sludge. Appl. Environ.
Microbiol. 46:50-54.

Campbell LL. and R. Singleton Jr. 1986. Genus IV. Desulfotomaculum,
p. 1200-1201. In P. H. A. Sneath, N. S. Mair and E. Sharpe (ed.), Bergey's
manual of determinative bacteriology. Williams 8 Wilkens, Baltimore

Cole J.A., A. Cascarelli, w.w. Mohn and J.M. Tiedje. 1994. Isolation
and characterization of a novel bacterium growing via reductive
dechlorination of 2-chlorophenol. Appl. Environ. Microbiol. 60:3536-3542.

Cole J.A., R.A. Sanford and J.M. Tiedje. Manuscript in preparation.

Cord-Ruwlsch R., H.-J. Seltz and R. Conrad. 1988. The capacity of
hydrogenotrophic anaerobic bacteria to compete for traces of hydrogen
depends on the redox potential of the terminal electron acceptor. Arch.
Microbiol. 149:350-357.

Criddle C.S., L.M. Alvarez and P.L. McCarty. 1991. Microbial
processes in porous media, p. 641-691. In J. Bear and M. Y. Corapcioglu
(ed.), Transport processes in porous media. Kluwer Academic Publishers,
Dordrecht

de Jong E., J.A. Fleld, H.-E. Splnnler, J.B.P.A. lenberg and J.A.M.
de Bont. 1994. Significant biogenesis of chlorinated aromatics by fungi in
natural environments. Appl. Environ. Microbiol. 60:264-270.

DeWeerd K.A., F. Concannon and J.M. Suflita. 1991. Relationship
between hydrogen consumption, dehalogenation, and the reduction of
sulfur oxyanions by Desulfomonile tiedjei. Appl. Environ. Microbiol.
57:1929-1934.

Dletrlch G. and J. Wlnter. 1990. Anaerobic degradation of chlorophenol
by an enrichment culture. Applied Microbiology and Biotechnology
34:253-8.

103

Doetsch R.N. 1981. Determinative methods of light microscopy, p. 21-33. In
P. Gerhardt, R. G. E. Murray, R. N. Costilowet al (ed.), Manual of methods
for general microbiology. American Society for Microbiology, Washington,
D. C.

Dolflng J. 1990. Reductive dechlorination of 3-chlorobenzoate is coupled to
ATP production and growth in an anaerobic bacterium, strain DCB-I. Arch.
Microbiol. 153:264-266.

Dolflng J. and B.K. Harrison. 1992. Gibbs free energy of formation of
halogenated aromatic compounds and their potential role as electron
acceptors in anaerobic environments. Environ. Sci. Technol. 26:2213-
2218.

Dolflng J. and D.B. Janssen. 1994. Estimates of Gibbs free energies of
formation of chlorinated aliphatic compounds. Bicdegradation 5:21-28.

Felsensteln J. 1985. Confidence limits on phylogenies: an approach using
the bootstrap. Evolution 39:783-791.

Felsensteln J. 1989. Phylogeny inference package (Version 3.2). Cladistics
5:164-166. ‘

Genthner B.R.S., W.A. Prlce II and P.H. Pritchard. 1989. Anaerobic
degradation of chloroaromatic compounds in aquatic sediments under a
variety of enrichment conditions. Appl. Environ. Microbiol. 55:1466-1471.

Gibson S.A. and J.M. Suflita. 1986. Extrapolation of biodegradation
results to groundwater aquifers: reductive dehalogenation of aromatic
compounds. Appl. Environ. Microbiol. 52:681-688.

Gribble G.W. 1992. Naturally occurring organohalogen compounds -- a
survey. J. Nat. Prod. 55:1353-1395.

Guerrant G.D., M.A. Lambert and C.W. Moss. 1982. Analysis of short
chain acids from anaerobic bacteria by high performance liquid
chromotography. J. Clin. Microbiol. 16:355-360.

Haggblom MM. 1992. Microbial breakdown of halogenated aromatic
pesticides and related compounds. FEMS Microbiol. Rev. 103:29-72. .

Hale D.H., J.E. Rogers and J. Wlegel. 1990. Reductive dechlorination of
dichlorophenols by nonadapted and adapted microbial communities in
pond sediments. Microbial Ecol. 20:185-96.

Holliger C., G. Schraa, A.J.M. Stams and A.J.B. Zehnder. 1993. A
highly purified enrichment culture couples the reductive dechlorination of
tetrachloroethene to growth. Appl. Environ. Microbiol. 59:2991-2997.

104

Hrudey S.E., E. Knettlg, S.A. Dalgnault and PM. Fedorak. 1987.
Anaerobic biodegradation of monochlorophenols. Environ. Technol. Lett.
8:65-76.

Juteau P., R. Beaudet, G. McSween, F. Leplne, S. Mllot and J.-G.
Blsalllon. 1995. Anaerobic biodegradation of pentachlorophenol by a
methanogenic consortium. Appl. Microbiol. Biotechnol. 44:218-224.

Larsen N., G.J. Olsen, B.L. Maldak, M.J. McCaughey, R. Overbeek,
T.J. Macke, T.L. Marsh and 0.R. Woese. 1993. The ribosomal
database project. Nucleic Acids Res. 21 :3021-3023.

Lbffler F.E., R.A. Sanford and J.M. Tiedje. Initial characterization of a
reductive dehalogenase from Desulfitobacterium chlororespirans strain
0023. Submitted to Appl. Environ. Microbiol .

Madsen T. and J. Aamand. 1992. Anaerobic transformation and toxicity of
trichlorophenols in a stable enrichment culture. Appl. Environ. Microbiol.
58:557-561. '

Madsen T. and D. Llcht. 1992. Isolation and characterization of an
anaerobic chlorophenol-transforming bacterium. Appl. Environ. Microbiol.
58:2874-2878.

Middeldorp P.J.M., M. Briglla and M.S. Salklnoja-Salonen. 1990.
Biodegradation of pentachlorophenol in natural soil by inoculated
Rhodococcus chlorophenolicus. Microbial Ecol. 20:123-39.

Mikesell M.D. and S.A. Boyd. 1986. Complete reductive dechlorination
and mineralization of pentachlorophenol by anaerobic microorganisms.
Appl. Environ. Microbiol. 52:861-865.

Mikesell M.D. and S.A. Boyd. 1988. Enhancement of pentachlorophenol
degradation in soil through induced anaerobiosis and bioaugmentation
with anaerobic sewage sludge. Environ. Sci. Technol. 22:1411-1414.

Mohn W.W. and J.M. Tiedje. 1990. Strain DCB-1 conserves energy for
growth from reductive dechlorination coupled to formate oxidation. Arch.
Microbiol. 1 53:267-271 .

Mohn W.W. and J.M. Tiedje. 1992. Microbial reductive dehalogenation.
Microbiol. Rev. 56:482-507.

Nicholson D.K., S.L. Woods, J.D. Istok and D.C. Peek. 1992.
Reductive dechlorination of chlorophenols by a pentachlorophenol
acclimated methanogenic consortium. Appl. Environ. Microbiol. 58:2280-
2286.

105

Olsen G.J., H. Matsuda, R. Hagstrom and R. Overbeek. 1994.
fastDNAml: a tool for construction of phylogenetic trees of DNA sequences
using maximum liklihood. CABIOS 10:41-48.

Parker W.J., G.J. Farquhar and E.R. Hall. 1993. Removal of
chlorophenolics and toxicity during high-rate anaerobic treatment of
segregated Kraft mill bleach plant effluents. Environ. Sci. Technol.
27:1783-1789.

Sanford R.A. and J.M. Tiedje. 1996. Chlorophenol dechlorination and
subsequent degradation in nitrate-limited denitrifying microcosms.
Submitted for publication .

Schlolz-Muramatsu H., A. Neumann, M. MeBmer, E. Moore and G.
Dlekert. 1995. Isolation and characterization of Dehalospirillum
multivorans gen. nov., sp. nov., a tetrachloroethene-utilizing, strictly
anaerobic bacterium. Arch. Microbiol. 163:48-56. .

Seltz H.-J., B. Schink, N. Pfennig and R. Conrad. 1990. Energetics of
syntrophic ethanol oxidation in defined chemostat cocultures. 1. Energy
requirements for H2 production and H2 oxidation. Arch. Microbiol. 155:82-

88.

Shelton D.R. and J.M. Tiedje. 1984. Isolation and partial characterization
of bacteria in an anaerobic consortium that mineralizes 3-chlorobenzoic
acid. Appl. Environ. Microbiol. 48:840-848.

Stackebrandt E. and BM. Goebel. 1994. Taxonomic note: a place for
DNA-DNA reassoceation and 168 rRNA sequence analysis in the present
species definition in bacteriology. Int. J. Syst. Bacteriol. 44:846-849.

Stanlake G.J. and R.K. Finn. 1982. Isolation and characterization of a
pentachlorophenol-degrading bacterium. Appl. Environ. Microbiol.
44:1421-1427.

Stevens T.O., H.D. Watts, J.J. Walker and J.K. Fredrlckson. 1992.
Medium formulation presented in a Poster Entitled: Optimization of solid
growth medium for isolation and culture of microorganisms from the
terrestrial subsurface, p. 366. In (ad), Abstr. 92nd Gen. Meet. Am. Soc.
Microbiol. 1992. American Society for Microbiology, Washington, D. 0.

Steward C.C., T.C. Dlxon, Y.P. Chen and C.R. Lovell. 1995.
Enrichment and isolation of a reductively debrominating bacterium from the
burrow of a bromometabolite-producing marine hemichordate. Can. J.
Microbiol. 41:637-642.

106

Thauer R.K., K. Jungermann and K. Decker. 1977. Energy
conservation in chemotrophic anaerobes. Bacteriol. Rev. 41:100-180.

Utkln I., D.D. Dalton and J. Wlegel. 1995. Specificity of reductive
dehalogenation of substituted ortho-chlorophenols by Desulfitobacterium
dehalogenans JW/IU-DCI. Appl. Environ. Microbiol. 61:346-351.

Utkin l., C. Woese and J. Wlegel. 1994. Isolation and characterization of
Desulfitobacterium dehalogenans gen. nov., sp. nov., an anaerobic
bacterium which reductively dechlorinates chlorophenolic compounds. Int.
J. Syst. Bacteriol. 44:612-619.

Vlsuvanathan S., M.T. Moss, J.L. Stanford, J. Hermon-Taylor and J.
McFadden. 1989. Simple enzymic method for isolation of DNA from
diverse bacteria. J. Microbiol. Meth. 10:59-64.

Welsburg W.W., S.M. Barns, D.A. Pelletler and D.J. Lane. 1991. 168
ribosomal DNA amplification for phylogenetic study. J. Bacteriol. 173:697-
703.

Woese 0.R. 1987. Bacterial evolution. Microbiol. Rev. 51:221-271.

Wolln E.A., M.J. Wolln and R.S. Wolfe. 1963. Formation of methane by
bacterial extracts. J. Biol. Chem. 238:2882-2886.

Zhang X. and J. Wiegel. 1990. Sequential anaerobic degradation of 2,4-
dichlorophenol in freshwater sediments. Appl. Environ. Microbiol. 56:1119-
1127.

Chapter V

Meta-dechlorination of 3-chlorophenol and
2,3-dlchlorophenol in methanogenic enrichment cultures:
the effects of Inhibitors and different electron donors

Introduction

Reductive dechlorination has been known to support respiratory
metabolism in several microorganisms (Dolfing 1990; Mohn and Tiedje 1990;
Holliger et al. 1993; Cole et al. 1994), however evidence suggests that this
reaction is not energy yielding for all dechlorinating microorganisms. For
example, polychlorinated biphenyls (PCBs) are reductively dechlorinated in
enrichments, but the majority of reducing power from organic matter with a few
exceptions flows to methane (Bedard and Quensen III 1995) and enhanced
growth as measured by increased dechlorination activity is not apparent.
Similar results have been obtained in anaerobic reactors fed with
chlorophenols (Perkins et al. 1994; Juteau et al. 1995). Thus, it is probable that,
despite the potential energetic advantage from using reductive dechlorination,
some microbes carry out this transformation without capturing this energy. The
cell, however, may still benefit by detoxification if the substrate is toxic.

Chlorophenolic compounds have been used extensively in several
industries and there has been considerable effort to study the degradation of
these compounds under aerobic and strictly anaerobic conditions (Haggblom
1992; Mohn and Tiedje 1992), Several polychlorinated phenols and
monochlorophenols (CPs) are degraded under aerobic and anaerobic

conditions (Boyd et al. 1983; Gibson and Suflita 1986; Hrudey et al. 1987;

107

 

108

Genthner et al. 1989; Genthner et al. 1989b; Hale et al. 1990; Zhang and
Wiegel 1990; Madsen and Aamand 1992). In many of the anaerobic
polychlorophenol degrading communities studied, 3-chlorophenol (3-CP) and
other meta-substituted phenols accumulated. As a result there are few
enrichment cultures described that show meta-dechlorination activity of
chlorophenols, particularly 3-CP. In anaerobic microcosms we observed the
complete dechlorination of 2,3-DCP (Chapter 2). Enrichment cultures derived
from these microcosms were used to further study the meta-dechlorination
activity and to further simplify the consortium with the goal of obtaining a pure
culture.

The anaerobic microcosms were fed a mixture of dichlorophenols. They
exhibited ortho- followed by meta-dechlorination of 2,3-dichlorophenol (DCP)
(Chapter 2). The enrichment cultures derived from these microcosms
completely dechlorinated and degraded 2,3-DCP or 3-CP to CH4 and 002.
The objective of this work was to further characterize the type of dechlorination
expressed in these cultures and to determine whether methanogens were
responsible for the dechlorination. In addition, individual electron donors were
evaluated to determine which were optimal for dechlorination. Our results
suggest that a eubacterial population is responsible for the fortuitous reductive
dechlorination of 3-CP and that a mixture of electron donors was the best
choice for efficienct chlorine removal. The dechlorination activity also appeared

to be inducible.

Materials and Methods
Enrichment Cultures and growth conditions. Enrichment
cultures were inoculated with anaerobic compost microcosms that had been fed

nitrate and mixtures of dichlorophenols. These cultures exhibited the complete

109

dechlorination of 2,3-DCP (Chapter 2). The enrichment cultures were fed
acetate (1 mM) and 2,3-DCP or 3-CP (125 pM). Those enrichment cultures that
showed dechlorination activity by the removal of meta and/or ortho substituted
chlorines were transferred to fresh medium with either 2,3-DCP or 3-CP (125
uM). A mixture of 1.25 mM formate , 0.25 mM succinate , 0.25 mM propionate
and 0.25 mM butyrate (VFAs) were added as electron donors to this and all
subsequent enrichment cultures. Cultures were incubated at 30° C.

Cultures were grown in 160 ml serum bottles with 100 ml of boiled
degassed medium or in anaerobic culture tubes with 20 ml of medium and
closed with butyl rubber stoppers. The mineral salts medium (Stevens et al.
1992) and vitamin solution (Wolin et al. 1963) was provided as described
previously (Chapter 2).

lnhlbltlon of meta-dechlorination. Inhibitors specific for
methanogens or eubacteria were used to test which microbial population was
responsible for the meta-dechlorination of 3-CP to phenol. Bromoethane-
sulfonic acid (BESA 1 mM) a methanogen inhibitor or Vancomycin (150 mg/l), a
eubacteria inhibitor, were added to triplicate 20 ml cultures containing the VFA
mixture described above and 100 1.1M 3-CP. Controls consisted of triplicate
tubes with no inhibitors added and triplicate tubes that were autoclaved. One
ml (5% v/v) from the enrichment culture which exhibited complete
dechlorination and degradation of 3-CP to CH4 was used to inoculate the
tubes. Samples were taken at approximately three-week intervals and
analyzed by HPLC for 3-CP, phenol, benzoate and VFAs. Headspace gas was
analyzed for CH4, CO2 and H2 at the same time. Once 3-CP or the VFAs were
depleted in the culture they were replenished at the original concentrations by

transferring from degassed stock solutions to the medium with sterile needles

1 10
and syringes. Additional BESA (2 mM) was added as needed to inhibit
methanogenesis if CH4 was observed.

Electron donors for meta-dechlorination. Formate (2 mM), 1 mM
succinate, 1 mM propionate, or 1 mM butyrate were added individually to sets of
triplicate anaerobic culture tubes to determine which VFAs supported
dechlorination. Culture conditions were as described for the inhibition
experiment. Individual electron donors were replenished to the medium once
they had been depleted. 3-CP, phenol, benzoate, VFAs, and headspace gas
were monitored as previously described.

Induction of dechlorination. Parallel cultures were incubated with
the VFA mixture or phenol, but in the absence of 3-CP. Once methanogenesis
was well established, additions of 125 uM 3-CP were made to triplicate sets of
tubes with and without 4 mM BESA. A 3-CP plus VFA mixture culture served as
the positive control. The time between start of dechlorination in the already
active control culture and the onset of dechlorination in the test cultures was
defined as the induction period. Culture conditions and monitoring were as
previously described.

Utlllzatlon of 2,3-DCP. The effects oi inhibiting methanogenesis on
enrichment cultures capable of both the ortho- and meta-dechlorination of 2,3-
DCP were examined. Triplicate sets of tubes were incubated with the VFA
mixture plus 125 11M 2,3-DCP. One set of culture tubes received BESA (1 mM).
Cultures were replenished with substrate and monitored as in previous
experiments.

Analytical procedures. Chlorophenols, dichlorophenols and
aromatic products of dechlorination were analyzed on a Hewlett Packard 1050
HPLC with a Chemstation analysis package. The eluent was a phosphoric acid

(0.1 %) buffered methanol pumped at 1.5 mein using a gradient from 48% to

11 1

55% methanol. A Hibar RP-18 (10 um) column was used. Eluted peaks were
detected at 218 nm, 230 nm and 275 nm simultaneously on a UV
multiwavelength detector. Samples (1 ml) from the enrichments were taken,
made basic with 10 ul of 2N NaOH, centrifuged for 6 min in a microfuge and
filtered through Acrodisc LC13 PVDF 0.45 pm filters prior to HPLC analysis.

VFAs were analyzed using a modification of the method described by
Guerrant et al. (1982). A Schimadzu HPLC with a BioRad Aminex HPX-87H ion
exclusion column heated to 60°C and using 0.005 N H2SO4 as the eluent was
used for the analyses. Previously filtered samples were acidified to 0.25 N
H2804 by adding 100 pl of 2.5 N H2SO4 to 900 pl of sample. Eluent was
pumped at 0.6 ml/min and detection of VFAs was at 210 nm by a UV detector.

The headspace of the incubation vessels was analyzed for CH4, H2 and
002 using a Carle gas chromatograph equipped with a 1.83 m Porapak 0
column and a thermal conductivity (T CD) detector. Headspace pressure was
normalized to atmospheric by venting with a needle prior to removing 0.3 mm

gas for injection into the GC.

Resuns

Characterization of reductive dechlorlnation activity.
Enrichment cultures inoculated from the DCP compost microcosm and fed 3-CP
and 2,3-0P showed complete degradation of the chlorophenols under
methanogenic conditions. 2,3-DCP was dechlorinated to 3-CP within 10 to 12
days of the start of incubation. Meta-dechlorination of 3-CP did not occur until
six to eight weeks after inoculation and was inhibited by the presence of 2,3-
DCP. Phenol accumulated in cultures fed 2,3-DCP and 3-CP, but was
eventually converted to benzoate which appeared transiently. No 4-

hydroxybenzoate was detected. Methane and 002 were the final products

1 12
detected in these enrichment cultures. Cultures received at least three
additions of 2,3-DCP or 3-CP before being transferred. The data suggest the
pathway shown in Figure 5.1.

Effect of Inhibitors on reductive dechlorination. Reductive
dechlorination of 3-CP in the enrichment cultures exhibited a lag time of 21
days (Figure 5.2A). The presence of BESA had little effect on dechlorination
activity, although initial rates appeared to be slower. Final amounts of 3-CP
dechlorinated were similar in cultures with and without BESA. Vancomycin
totally inhibited dechlorination activity (Figure 5.2A), but had little effect on
methanogenesis (Figure 5.28). Some methanogenesis occurred in the BESA
inhibited culture, however this was stopped by amending cultures with more
BESA (Figure 5.28). The total methane evolved from the inhibited culture was
approximately 25 % of that produced by the uninhibited culture.

Effect Incubation time on meta-dechlorination lag period.
Since dechlorination activity required a considerable lag period, experiments
were done to determine the induction period of dechlorination in cultures
incubated in the absence of 3-CP. Prior to the addition of 3-CP, enrichments
fed VFAs or phenol were incubated for 57 days. All VFAs and phenol were
depleted in these cultures with concomitant production of methane. At the end
of the incubation period 3-CP was added to all cultures, including a positive
control which had been fed 3-CP plus VFAs throughout the 57 day incubation
period. BESA was added to one set of the VFA-fed cultures to inhibit further
methanogenesis. Meta-dechlorination commenced within 7 days in the control
3-CP + VFA and incubated VFA-fed cultures (Figure 5.3). The phenol-
incubated culture showed no dechlorination of 3-CP, even though phenol

degradation activity continued (data not shown). No further methane production

113

 

OH OH

Cl \ 000‘

OH

06'
0|
CH4+co2

L J

 

Figure 5.1. Schematic representation of the dechlorination pathway
of 3-CP to phenol and subsequent tranforrnation to benzoate and
methane. The ortho-dechlorination of 2,3-DCP is shown in the
lower left and is converted completely to 3-CP prior to further
degradation.

 

114

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

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Figure 5.2. Effect of inhibitors on dechlorination in a methanogenic enrichment
culture. (A) Cumulative 3-MCP dechlorination in cultures with VFA mixture (A),
VFA plus BESA (O) and VFA plus vancomycin(EI). (B) Methane production in
the same cultures. Initial culture volume was 20 ml. Brackets are 95%
confidence intervals.

umoles phenol

115

 

 
 

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VFA + BESA —
'

 

 

 

 

I"
. Phenol
0’ - ----1.A---T--J_i
50 75 100 125 150 175

Figure 5.3. The dechlorination of 3-CP in cultures Incubated with
(A) and without (8) 3-CP for 57 days, and grown on VFAs. 3-CP
with or without BESA was added at 57 days of incubation to VFA-
and phenol-fed cultures. (A) shows the onset of dechlorination in
the control culture grown on VFAs plus 3-CP upon the addition of
3-CP as designated by the arrow. (B) shows the onset of dechlori-
nation in cultures incubated without 3-CP with the arrow indicating
the addition of 3-CP and BESA. Brackets are the 95 % confidence
intervals.

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116
was observed in cultures with BESA and considerably less total dechlorination
activity occurred compared to uninhibited cultures.

Dechlorination of 2,3-DCP. Methanogenic enrichment cultures that
exhibited both ortho- and meta-dechlorination were treated with BESA to
compare the effects of this inhibitor to those observed with the 3-CP cultures.
Figure 5.4 A and B shows the ortho- and combined ortho— plus meta-
dechlorination of 2,3-DCP in cultures without and with BESA. Ortho-
dechlorination was not affected by the presence of BESA, as there was little
variance in the rate or the extent of dechlorination with uninhibited cultures.
Although there was apparently less total ortho- plus meta-dechlorination in the
BESA-inhibited culture, the difference was not statistically significant.
Considerable 3-CP remained in both treatments at the end of the experiment,
and meta-dechlorination was never observed to occur simultaneously with
ortho-dechlorination (data not shown). No methane production was observed
in the inhibited culture, however considerable acetate accumulated in contrast
to the uninhibited culture (data not shown).

Electron donors for meta-dechlorination. Formate, succinate,
pr0pionate and butyrate, as individual electron donors, supported reductive
dechlorination of 3-CP. Enrichment cultures with formate or butyrate showed
lag periods of less than 21 days before dechlorination occurred (Figure 5.5).
For comparative purposes, the efficiency of dechlorination as mmoles
dechlorinated per mole H2 equivalent consumed of each electron donor was
determined (Figure 5.5). At the end of the experiment the efficiency of
dechlorination had reached a steady-state for each electron donor, with formate
> propionate = succinate > butyrate. Early efficiencies were higher than at

steady state for all electron donors, except formate. For example, at the start of

FIQUre
and 0
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umoles dechlorinated

 

 

 

 

 

 

 

 

 

 

 

umoles dechlorinated
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11111
150 180

4

111.111
90 120

Figure 5.4. Cumulative dechlorination of 2,3-DCP. Ortho dechlorination (El)
and ortho plus meta dechlorination (O). (A) has no BESA and (B) has BESA
added. Brackets are 95% confidence intervals.

E
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Figure 5.5. Effect of electron donor on the efficiency of reductive
dechlorination of 3-CP as measured by the proportion of reducing equivalents
used for dechlorination.

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dechlorination, propionate and succinate had nearly double the efficiency
observed at the 'end of the experiment (Figure 5.5).

Data were compiled to calculate efficiencies of dechlorination for each
treatment (Table 5.1). In addition, the molar yields of methane and acetate per
mole H2 equivalent of electron donor were calculated to provide reference
indices for the different treatments. The most efficient dechlorination was
observed in the 2,3-DCP + BESA treatment, which showed almost three times
the value observed with any of the 3-CP treatments. Ortho-dechlorination
accounted for an average of 67 % of the total dechlorination observed in the
2,3-DCP treatments. The enrichments provided a VFA mixture exhibited a
dechlorination efficiency between that of succinate and formate alone,
depending on the specific treatment.

The fraction of electrons from the electron donors used for reductive
dechlorination was determined by calculating the slope of the linear regression
line of the electron donor consumed as H2 equivalents versus the equivalents
dechlorinated. In pure cultures the fraction of electrons from the electron donor
used to reduce the electron acceptor is referred to as the to. When 3-CP was
present, the apparent to for meta-dechlorination was 0.01. With 2,3-DCP as the

electron acceptor, the apparent fe was 0.05.

Discussion
Meta-dechlorination of chlorophenols has been observed in other
anaerobic enrichments (Nicholson et al. 1992), but the types of microorganisms
responsible for this activity have not been established. 3-CP often accumulates
transiently in enrichment cultures, indicating that meta-dechlorination of this
compound may not be common in the environment. Enrichment cultures

derived from 2,3-DCP fed microcosms were shown to mediate meta-

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dechlorination and thus were selected for further study. In the enrichment
cultures vancomycin completely inhibits meta-dechlorination and BESA does
not. This indicates that eubacteria are responsible for reductively
dechlorinating the 3-CP to phenol in the enrichment cultures. Similar
conclusions were reported for a methanogenic microbial community that
dechlorinated 2,4,6-TCP (Perkins et al. 1994). Dechlorination, however, is not
affected by methanogenesis, since uninhibited cultures have higher initial
dechlorination rates and higher meta-dechlorination than BESA-inhibited
cultures. Similar results are also found with 2,3-DCP fed cultures.

One interesting observation is that the incubation of the cultures on VFAs
alone exhibits an equivalent lag period for dechlorination to the cultures
incubated with 3-CP plus VFAs (Figure 5.3). This indicates that the microbial
populations responsible for dechlorination are growing in the absence of 3-CP.
It is also possible that the dehalogenation specific enzymes are induced in the
absence of 3-CP, although it seems more likely that dechlorination is
fortuitously mediated by a eubacterial population. The absence of
dechlorination activity in the cultures fed phenol also supports this conclusion,
since induction for specific dehalogenase activity might be expected with a
similar compound like phenol. Since only the VFA mixture was provided as a
electron donor source, there are only a limited number of energy yielding
mechanisms possible for this methanogenic microbial community. The
oxidation of each electron donor, except formate, would require a syntrophic
interaction with a H2 consuming methanogen or acetogen (Conrad et al. 1986;
Schink 1988). Syntrophic microorganisms might benefit from reductive
dechlorination of 3-CP by using it as a H2 scavenging mechanism. An

acetogen might also fill the Hz-scavenging niche, using the chlorophenol as an

auerna

proces

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122
alternative sink for reducing equivalents and perhaps gaining energy from the
process. _

Experiments with individual electron donors show that all compounds
tested could support meta-dechlorination activity, with formate being the most
efficient. It is interesting that the VFA mixture exhibits similar efficiencies of
dechlorination to formate. This indicates that the inclusion of formate as an
electron donor may be sufficient to raise the efficiency of dechlorination.
Formate is converted to H2 and CO2 in most anaerobic ecosystems and even
methanogens that can use formate will first oxidize it to H2 + CO2. The H2 from
this reaction could then be used for methanogenesis with some also available
to reductively dechlorinate 3-CP. It is possible that the microorganisms
responsible for dechlorination require H2 as an electron donor and therefore
reactions that favor more H2 generation would lead to more efficient microbial
dechlorination. Since formate oxidation is not as energetically constrained, (i.e.
requiring a syntrophic partnership, as would be the case for butyrate or
propionate oxidation) there would be significant H2 produced even in the
absence of a methanogen (Chapter 7).

There is also the possibility that a homoacetogen is responsible for the
dechlorination activity, since considerable acetate was observed in the formate-
fed enrichment cultures. Acetate is also formed with the other electron donors
although the mechanism of acetate generation differed in each culture.
Butyrate is oxidized to two acetates. Propionate is converted to acetate and
CO2. Succinate is decarboxylated to propionate and subsequently to acetate
and CO2. Acetate appears transiently in the cultures and is converted into

methane except in those cultures receiving BESA. Some H2 is also formed

from acetate during acetoclastic methanogenesis (Lovley and Ferrey 1985) and

123
therefore methanogens could be supplying reducing equivalents to
dechlorinating microorganisms in the enrichment cultures.

The presence of both methanogenesis and acetogenesis in formate-fed
enrichment cultures requires that H2 concentrations are high enough to
energetically support both of these activities. The threshold H2 concentration,
below which no energy can be obtained, is about 500 ppm for acetogens and
50 ppm for methanogens (Cord-Ruwisch et al. 1988). In order for both
methanogens and acetogens to coexist, the H2 concentration must be greater
than 500 ppm. At this relatively high concentration of H2 the dechlorinators may
be able to act as scavengers of reducing equivalents, which they might not be
able to do at lower H2 concentrations. It is possible that the lower efficiency of
dechlorination observed with butyrate is associated with a low H2 concentration
in that particular set of enrichment cultures. This may be due to the obligate
syntrophic life-style of the butyrate oxidizers, which require low H2
concentrations for growth (Schink and Friedrich 1994). In contrast, formate
oxidation would release the equivalent amount of energy to butyrate at a higher
threshold concentration of H2 and therefore may support both acetogenic and
methanogenic populations.

When the efficiencies of dechlorination are compared with the mmoles of
methane produced per mole of reducing equivalent (Table 5.1), it is apparent
that most of the reducing power in the enrichment cultures is not diverted to 3-
CP reduction. Even when BESA was added to inhibit methanogenesis, the
additional reducing equivalents end up in acetate rather than being used for
reductive meta-dechlorination. An exception was observed with 2,3-DCP +
BESA enrichment cultures, which have equal reducing equivalents routed to
dechlorination and acetate production. This may be due to a microbial

population in the enrichment culture using the energy from the ortho-

dechlo
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124

dechlorination of 2,3-DCP. Some evidence supporting this assertion was found
in strain C023 isolated from this enrichment, which was able to grow using 2,3-
DCP as an electron acceptor (Chapter 4). The amount of acetate serving as an
electron sink is likely to be underestimated, since acetate could serve as a
carbon source even in the presence of BESA. Therefore, it is possible that
acetogenesis efficiencies in the different treatments were close to
methanogenesis efficiencis. By calculating the ratio of CH4 + acetate
efficiencies to the dechlorination efficiency (Table 5.1), it is apparent that the
best meta-dechlorination of 3-CP occurred in enrichments with ratios between
12 and 15. This is valid for several different treatments and suggests that both
acetogenesis and methanogenesis may be important for meta-dechlorination
even though a methanogen is not directly responsible for the dechlorination
activity.

Since the meta-dechlorination activity is closely correlated to
methanogenesis and acetogenesis activity in the enrichment cultures, it may be
difficult to isolate the responsible microorganism. This is likely since the
dechlorination of 3-CP did not serve as a major electron sink and therefore was
not likely to act as a significant electron acceptor for a microorganism. The
presence of 3-CP in the enrichment culture therefore offers no selective
advantage to any microbial population beyond tolerance to its toxicity. Although
not very specific, one strategy would be to isolate as many component
populations as possible and screen them for the ability to mediate this reductive
dechlorination reaction. Even this may not be successful, since the reaction
may rely on the synergistic relationship between many microbial populations
within the community. It would probably not be practical to test all possible
combinations of isolates recovered. The desired microbial population could

also be non-culturable under the conditions being used for growth. Perhaps the

best:
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1991]

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125
best strategy is to characterize the community by analyzing the 168 rRNA
phylogeny of the component microbial populations, as has been done in
several studies of difficult to isolate cultures (Ward et al. 1990; Schmidt et al.
1991). Knowing the phylogeny of the component populations could provide
information for developing isolation strategies.

Results from these enrichment culture experiments suggest that certain
combinations of electron donors may be better suited for stimulating meta-
dechlorination of chlorophenols. This information would be useful for
developing bioremediation strategies for chlorophenolic waste material. It
appears that a mixture of VFAs containing formate is best suited for obtaining
the most efficient dechlorination. In contrast the use of butyrate resulted in the
lowest dechlorination efficiency and phenol showed no capacity to support
reductive dechlorination. It is possible that the interrnittant supply of formate to
the culture was responsible for keeping the H2 concentrations above the
threshold concentration for methanogenesis and therefore keeping sufficient
reducing equivalents available for reductive dechlorination. This suggests that
the best strategy for stimulating the anaerobic dechlorination of chlorophenols

is to use an intermittent feeding of formate and perhaps other electron donors.

References

Bedard D.L. and J.P. Quensen Ill. 1995. Microbial reductive
dechlorination of polychlorinated biphenyls, p. 127-216. In L. Y. Young and
C. Cerniglia (ed.), Microbial transformation and degradation of toxic
organic chemicals. Wiley-Liss Division, John Wiley & Son, New York

Boyd S.A., D.A. Shelton, D. Berry and J.M. Tiedje. 1983. Anaerobic
biodegradation of phenolic compounds in digested sludge. Appl. Environ.
Microbiol. 46:50-54.

Cole J.A., A. Cascarelll, w.w. Mohn and J.M. Tiedje. 1994. Isolation
and characterization of a novel bacterium growing via reductive
dechlorination of 2-chlorophenol. Appl. Environ. Microbiol. 60:3536—3542.

 

Co

Co

Dc

126

Conrad R., B. Schink and T.J. Phelps. 1986. Thermodynamics of H2-
consuming and Hz-producing metabolic reactions in diverse

methanogenic environments under in situ conditions. FEMS Microbiol.
Ecol. 38:353-360.

Cord-Ruwlsch R., H.-J. Seltz and R. Conrad. 1988. The capacity of
hydrogenotrophic anaerobic bacteria to compete for traces of hydrogen
depends on the redox potential of the terminal electron acceptor. Arch.
Microbiol. 149:350-357.

Dolflng J. 1990. Reductive dechlorination of 3-chlorobenzoate is coupled to .
ATP production and growth in an anaerobic bacterium, strain DCB-1.
Arch. Microbiol. 153:264-266.

Genthner B.R.S., W.A. Prlce II and P.H. Pritchard. 1989. Anaerobic
degradation of chloroaromatic compounds in aquatic sediments under a
variety of enrichment conditions. Appl. Environ. Microbiol. 55:1466-1471.

Genthner B.R.S., W.A. Prlce II and P.H. Pritchard. 1989b.
Characterization of anaerobic dechlorinating consortia derived from
aquatic sediments. Appl. Environ. Microbiol. 55:1472-1476.

Glbson S.A. and J.M. Suflita. 1986. Extrapolation of biodegradation
results to groundwater aquifers: reductive dehalogenation of aromatic
compounds. Appl. Environ. Microbiol. 52:681-688.

Guerrant G.D., M.A. Lambert and c.w. Moss. 1982. Analysis of short
chain acids from anaerobic bacteria by high performance liquid
chromatography. J. Clin. Microbiol. 16:355-360.

Haggblom MM. 1992. Microbial breakdown of halogenated aromatic
pesticides and related compounds. FEMS Microbiol. Rev. 103:29-72.

Hale D.H., J.E. Rogers and J. Wiegel. 1990. Reductive dechlorination of
dichlorophenols by nonadapted and adapted microbial communities in
pond sediments. Microbial Ecol. 20:185-96.

Holllger 0., G. Schraa, A.J.M. Stams and A.J.B. Zehnder. 1993. A
highly purified enrichment culture couples the reductive dechlorination of
tetrachloroethene to growth. Appl. Environ. Microbiol. 59:2991-2997.

Hrudey S.E., E. Knettlg, S.A. Dalgnault and PM. Fedorak. 1987.
Anaerobic biodegradation of monochlorophenols. Environ. Technol. Lett.
8:65-76.

Juteau P., R. Beaudet, G. McSween, F. Léplne, S. Mllot and J.-G.
Blsalllon. 1995. Anaerobic biodegradation of pentachlorophenol by a
methanogenic consortium. Appl. Microbiol. Biotechnol. 44:218-224.

127

Lovley D.R. and J.G. Ferrey. 1985. Production and consumption of H2

during growth of Methanosarcina spp. on acetate. Appl. Environ.
Microbiol. 49:247-249.

Madsen T. and J. Aamand. 1992. Anaerobic transformation and toxicity of
trichlorophenols in a stable enrichment culture. Appl. Environ. Microbiol.
56:557-561.

Mohn w.w. and J.M. Tledle. 1990. Strain DCB-1 conserves energy for
growth from reductive dechlorination coupled to formate oxidation. Arch.
Microbiol. 153:267-271 .

Mohn w.w. and J.M. Tiedje. 1992. Microbial reductive dehalogenation.
Microbiol. Rev. 56:482-507.

Nlcholson D.K., S.L. Woods, J.D. Istok and D.C. Peek. 1992.
Reductive dechlorination of chlorophenols by a pentachlorophenol
acclimated methanogenic consortium. Appl. Environ. Microbiol. 58:2280-
2286.

Perkins P.S., S.J. Komisar, J.A. Puhakka and J.P. Ferguson. 1994.
Effects of electron donors and inhibitors on reductive dechlorination of
2,4,6-trichlorophenol. Wat. Res. 28:2101-2107.

Schlnk B. 1988. Principles and limits of anaerobic degradation:
Environmental and technological aspects, p. 771-846. In A. J. B. Zehnder
(ed.), Biology of Anaerobic Microorganisms. Wiley and Sons, New York

Schink B. and M. Friedrich. 1994. Energetics of syntrophic fatty acid
oxidation. FEMS Microbiol. Rev. 15:85-94.

Schmldt T.M., E.F. DeLong and N.R. Pace. 1991. Analysis of a marine
picoplankton community by 168 rRNA gene cloning and sequencing. J.
Bacteriol. 173:4371 -4378.

Stevens T.O., H.D. Watts, J.J. Walker and J.K. Fredrlckson. 1992.
Medium formulation presented in a Poster Entitled: Optimization of solid
growth medium for isolation and culture of microorganisms from the
terrestrial subsurface, p. 366. In (ad), Abstr. 92nd Gen. Meet. Am. Soc.
Microbiol. 1992. American Society for Microbiology, Washington, D. C.

Ward G.M., R. Weller and MM. Bateson. 1990. 168 rRNA sequences
reveal numerous uncultured microorganisms in a natural community.
Nature 345:63-65.

Zhang x. and J. Wlegel. 1990. Sequential anaerobic degradation of 2,4-
dichlorophenol in freshwater sediments. Appl. Environ. Microbiol.
56:1119-1127.

Chapter VI

Evidence for haloresplratlon In an anaerobic enrlchment
culture that simultaneously dechlorlnates
2- and 3-chlorophenol to phenol

Introduction

Reductive dechlorination of chlorophenols has been observed in many
anaerobic microbial studies (Mohn and Tiedje 1992). The ability to couple the
energy released from this reaction to growth has only been verified in a few
microorganisms (Dolfing 1990; Mohn and Tiedje 1990; Cole et al. 1994;
Sanford et al. 1996). Since this reductive dechlorination reaction releases
considerable energy (Dolfing and Harrison 1992), the ability to halorespire in
anaerobic environments is potentially beneficial to the organism catalyzing this
transformation. The types of natural halogenated aromatic compounds that
might be present in the environment and support halorespiration are many
(Gribble 1992), and recently de Jong and coworkers (1994) found that white rot
fungi actually generate up to 70 mg/kg of chlorinated anisyl metabolites in leaf
litter. Thus, not only is a halorespiratory metabolism potentially useful, it is likely
to occur in natural anaerobic ecosystems.

The reductive removal of chlorines from aromatic compounds may be
important to anaerobic bioremediation strategies where considerable
contamination with chlorophenolic compounds has occurred. Once chlorines
are removed, the resulting aromatic compounds are generally more amenable
to oxidative attack. This was observed in denitrifying microcosms in which 2-

chlorophenol (2-CP) was dechlorinated by one microbial population to phenol,

128

129
which was then degraded by denitrifiers (Chapter 2). Often it is shown that
dechlorination of multi-chlorinated phenols is incomplete, with meta- and para-
substituted chlorophenols accumulating in anaerobic cultures (Juteau et al.
1995). Many studies have shown the reductive dechlorination of PCP to di- and
mono-chlorophenols (Nicholson et al. 1992). In most cases the order of
dechlorination is ortho, para and finally meta, although the latter two have been
known occur in the reverse order (Mikesell and Boyd 1986) (Chapter 2). These
lesser-halogenated products of dechlorination are often more easily degraded
under aerobic environmental conditions.

In this study an enrichment culture with the novel ability to simultaneously
reductively dechlorinate ortho- and meta- chlorines from monochlorophenols
and 2.5-DCP was characterized. Ortho-dechlorination of chlorophenols is
common in anaerobic environments, however, meta-dechlorination is relatively
rare and therefore its occurrence in an enrichment culture warranted further
study. The measurement of H2 threshold concentrations and the determination
of the fraction of electrons from the electron donor used for reductive
dechlorination, fa, suggested a halorespiring microbial population was
responsible for the dechlorination activity. This is the first evidence presented
that shows that meta-dechlorination of any chlorophenol could serve as a
terminal electron acceptor for growth. _

Materials and Methods

Enrichment Cultures and growth conditions. . Enrichment
cultures were inoculated from anaerobic soil microcosms that had been fed
nitrate and mixtures of monochlorophenols (CP) and exhibited the complete
dechlorination of 2- and 3-CP (Chapter 2). The enrichment cultures were fed
acetate (1 mM) and 2-CP and 3-MCP (125 uM each). Those enrichment

cultures that showed dechlorination activity by the removal of meta and ortho

130

substituted chlorines were transferred to fresh medium with either 2.5-DCP or
the 2- plus 3-CP mixture (125 uM). A mixture of 1.25 mM formate , 0.25 mM
succinate , 0.25 mM propionate and 0.25 mM butyrate (VFAs) was added as
electron donors to these enrichment cultures. Subsequent transfers contained
10 mM formate or H2 (5 % v/v) added to the headspace as the electron donors
for dechlorination. Acetate (0.5 - 1.0 mM) was added as a carbon source.
Chlorophenols and phenol concentrations were monitored by HPLC. Depletion
of VFAs was also monitored by HPLC. The composition of headspace gases
was evaluated by GC. The growth medium and conditions were as described in
Chapter 2 of this thesis. Cultures were incubated at 30° C.

Determination of electron donors and acceptors. The ability of
the enrichment culture to reductively dechlorinate 2- and 3-CP with different
electron donors was determined in duplicate 20 ml culture tubes incubated at
30° C. Acetate, formate, butyrate, succinate, propionate, fumarate, lactate,
pyruvate, crotonate and H2 were tested separately as potential electron donors.
The soluble 'VFAs were added to duplicate tubes to achieve concentrations of
0.5 or 2.0 mM. 18 umoles (6.6 % v/v) H2 was added to the headspace of the
duplicate culture tubes. 2- and 3-CP (100 (M each) were added as electron
acceptors. A 5 % inoculum was added to each tube from a VFAs and 2- plus 3-
CP grown culture. Chlorophenols were replenished three times at
concentrations of 100 uM to those cultures exhibiting dechlorination activity.
Dechlorination and depletion of electron donor were used as an indication of a
positive test for growth along with the ability to serially transfer the
dechlorination activity in the culture.

The range of electron acceptors used by this enrichment culture was
determined using the same growth conditions as for the electron donor

determination. With the four VFA mixture serving as electron donors, 100 uM of

131
2-CP; 3-CP; 4-CP; 2,3-DCP; 2,4-DCP; 2,5-DCP; 2,6-DCP; SCI-4-HBA; o-
fluorophenol; o-bromophenol; o-iodophenol; 2,3,5-TCP; 2,4,6-TCP;
pentachlorophenol (PCP); and 3-chloroanisaldehyde were tested separately as
halogenated electron acceptors. Nitrate, and fumarate were also tested at a
concentration of 5 mM. Tubes were inoculated with 5 °/o inoculum from cultures
grown on VFAs and 2- plus 3-CP. Halogenated substrates were monitored by
HPLC and were replenished once depleted. Nitrate and fumarate
concentrations were measured by HPLC. Consumption of VFAs was monitored
by HPLC to verify physiological activity.

Hydrogen uptake and threshold determination. Hydrogen
uptake was monitored in relation to dechlorination activity to test if the
hydrogenase activity was closely linked to the terminal electron accepting
capacity of the 2CP strains. Hydrogen (5.0 °/o) was added to the headspace of
100 ml cultures in 160 ml serum bottles. 2,5-DCP (100 uM) was added to these
cultures and the H2 concentration was monitored by GC as dechlorination
occurred. After the complete dechlorination of the 2,5-DCP, an additional
amendment of 100 uM was made. To determine the threshold concentration for
hydrogen maintained by the enrichment cultures while dechlorinating, duplicate
cultures were monitored for H2 depletion with excess 2.5-DCP present.
Threshold concentrations of H2 were measured using a GC equipped with a
reduction gas detector from Trace Analytical, Menlo Park, CA

The ability of the dechlorinating enrichment culture to produce H2 and
dechlorinate was tested by inoculating a culture with 20 mM formate as the sole
energy source and 100 uM 2.5-DCP. Acetate (1 mM) was added as a carbon
source. A 3% inoculum from the eighth serial transfer maintained on H2 + 002

and chlorophenols was used. These serum bottle cultures were incubated on a

132
shaker at 30° C. H2 was monitored by GC and dechlorination of 2.5-DCP was
monitored by HPLC. Formate depletion was checked by HPLC analysis.

Determination of fe. As an indication of energetic efficiency and that
halorespiration was occurring, the fraction of electrons (fe) from H2 used for the
reductive dechlorination of 2- and 3-CP or 2,5-DCP was determined. Several
replicates of the enrichment culture were grown on 5 °/o H2, 7% C02, 1 mM
acetate and 2.5-DCP (125 uM). Alternatively 2- and 3-CP (100 uM each) were
used as electron acceptors. Control cultures received H2 or 2.5-DCP alone.
Concentrations of chlorophenols were monitored by HPLC and amendments of
new substrate (125 uM) were made prior to the concentration reaching zero.
Acetate was determined by HPLC. The fe was calculated by plotting the H2
equivalents used versus the total umoles dechlorinated and determining the
slope of the regression line. For example the total umoles Cl' dechlorinated
from 2,5-DCP was equal to the umoles of phenol formed (X2) plus the umoles of
2-CP and 3-CP present in the medium at a particular time point.

Analytical procedures. ChlorOphenols, dichlorophenols and
aromatic products of dechlorination were analyzed on a Hewlett Packard 1050
HPLC with a Chemstation analysis package. The eluent was a phosphoric acid
(0.1 %) buffered methanol pumped at 1.5 mein using a gradient from 48% to
55% methanol. A Hibar RP-18 (10 um) column was used. Detection of eluted
peaks was done at 218 nm, 230 nm and 275 nm simultaneously on a UV
multiwavelength detector. Samples (1 ml) from the enrichments were taken,
made basic with 10 ul of 2N NaOH, centrifuged for 6 min in a microfuge and
filtered through Acrodisc LC13 PVDF 0.45 pm filters prior to HPLC analysis.

Volatile fatty acids were analyzed using the Schimadzu HPLC with a
BioRad Aminex HPX-87H ion exclusion column heated to 60°C and using

0.005 N H2SO4 as the eluent. Previously filtered samples were acidified to

133
0.25 N H2SO4 by adding 100 pl of 2.5 N H2SO4 to 900 pl of sample. Eluent
was pumped at 0.6 ml/min and detection of VFAs was at 210 nm by a UV
detector.

The headspace of the incubation vessels was analyzed for CH4, H2 and
CO2 using a Carle gas chromatograph equipped with a 1.83 m Porapak O
column and a thermal conductivity (TCD) detector. Headspace pressure was
normalized to atmospheric by venting with a needle prior to removing 0.3 ml of
gas for injection into the GC. N2O was quantified on a Perkin Elmer 910 CO
with a Porapak O column and 63Ni -ECD detector. Denitrification products
were measured in an Ar headspace.

Nitrate was analyzed by HPLC using a Whatman Partisil 1O SAX column
on a Schimadzu HPLC. The eluent was a 50 mM phosphate buffer (pH = 3.0)
pumped at a rate of 1 ml/min. UV adsorption at 210 nm was used for detection.
Samples from primary enrichments were diluted 1:100 in deionized H2O prior
to nitrate and nitrite analysis. A 40 pl sample was injected by an Alcott 738

autosampler.

Resuns

Enrichment culture activity. Enrichment cultures fed the VFA mixture
and monochlorophenols showed complete dechlorination of 2- and 3-CP to
phenol. As H2 was observed in the headspace of these initial enrichment
cultures, subsequent transfers were made to media containing either H2 or
formate as electron donors. Cultures maintained with H2 as an electron donor
exhibited simultaneous ortho- and meta-dechlorination activity with the
stoichiometric production of phenol (Figure 6.1). These enrichments are
subsequently referred to as OM, for ortho plus meta dechlorinating enrichment

culture. Hydrogen uptake was correlated to the appearance of phenol in these

134

 

umoles

 

 

 

 

 

Figure 6.1. Uptake of H2 by OM enrichment culture and cumulative
production of phenol from the dechlorination of 2- and 3-CP.

 

lillll

 

135
enrichment cultures, although initial consumption of several umoles of H2
occurred prior to the onset of dechlorination activity. Control cultures containing
only H2 or only chlorophenols did not exhibit any H2 uptake or dechlorination
activity, respectively (data not shown).

Since 2- and 3-CP were used as electron acceptors, 2.5-DCP was used
as an aitemative substrate for dechlorination. 2,5-DCP was stoichiometrically
dechlorinated to phenol in the presence of H2 (Figure 6.2A). The transient
appearance of 2- and 3-CP indicated that both ortho- and meta-chlorines were
removed from 2.5-DCP sequentially with either position removed prior to the
other (Figure 6.2B). The H2 consumed is shown in Figure 6.28. On day 110 of
incubation, H2 additions were suspended to determine the effects of no H2 on
dechlorination and to determine the threshold concentration of H2 (see below).
As the H2 concentration was reduced, the rate of 2.5-DCP dechlorination
slowed with 2- and 3-CP accumulating in the medium (Figure 6.2). This was
confirmatory evidence of the requirement for H2 for reductive dechlorination
activity.

Electron acceptors and donors. Several chlorinated phenolic
compounds were tested for dechlorination by the OM enrichment culture (Table
6.1). 3-CP was only dechlorinated when 2-CP was also present in the medium.
2-CP, however, was dechlorinated to phenol when supplied as the sole
electron acceptor. While 2,5-DCP was dechlorinated to phenol by the OM
enrichment culture; no dechlorination of 2,3-DCP or 2,6-DCP was observed.
Ortho-dechlorination of 2.4-DCP to 4-CP was supported by this culture, but no
TCPs or PCP were utilized. Nitrate and fumarate were not utilized as electron
acceptors and did not support growth as measured by VFA depletion.

The electron donors that supported the dechlorination of 2- and 3-CP in

the OM enrichment culture are listed in Table 6.1. Although acetate initially

136

200

 

A phenol

. C
150 ° '

1114

11111

[2.5-DCP] (uM)

 

 

 

 

ff

 

 

 

 

 

100 _- H‘ , 3.2,5-DCP3
i ‘ .
' 0
so?
: “mm ‘
o .9 "4‘...” 0... W0
0

 

 

 

 

2500

2000

1 500

1 000

500

800
700
600
500
400
300

~ 200

100

O 2 100 120 140 160
100 B
A 1' -El-i3-—::.
E. 30- . H afie-
v .. )2? ‘
E I
60—
:3 . 2‘32 3-CP
E 40- |.
‘5 i I -CP
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‘*O
0 2O 4O 60 80 100 120 140 160

Days

Figure 6.2. Dechlorination of 2,5-DCP by the OM enrichment culture. (A)
Shows the dechlorination of 2,5-DCP and the stoichiometric accumulation of

phenol. (B) shows the appearance of 2- and 3-CP as intermediates of 2,5-
DCP dechlorination and the cumulative H2 uptake in the OM culture.

(wfl) [Ioueud]

peumsuoo aH sejounf

137

Table 6.1. Electron acceptors and donors utilized by OM enrichment culture.
A (+) indicates that both the electron donor and acceptor were depleted.
Electron donors were tested using 2-CP plus 3-CP as electron acceptors.

 

 

 

Compound tested Use
Electron acceptors
2-CP +
3-CP -
2- & 3-CP +
4-CP -
2-Fluorophenol -
2-Bromophenol -
240dophenol -
2,3-DCP -
2,4-DCP--> p-CP +
2,5-DCP--> phenol +
2,6-DCP ‘ -
2,3,5-TCP .
2,4,5-TCP -
PCP -
3-Cl-4-hydroxybenzoate -
3-Cl-anisaldehydea -
Fumarate -
Nitrate '
Electron donors

Acetate '
H2 4-
Formate ---> H2b +
Succinate '
Propionate -
Butyrate -
Crotonate -

Pyruvatec +I-
Lactate +

 

 

 

a SCI-anisaldehyde is a natural product produced by white-rot fungi.

b Formate was stoichiometrically converted to H2 before dechlorination.

° Pyruvate was depleted prior to any dechlorination and supported growth in the absence of
chlorophenols.

138

supported dechlorination, after months of incubation in the original enrichment
cultures, the enrichment transfers showed no activity with acetate as an electron
donor (data not shown). Formate was as good an electron donor as H2, but
measurements of the headspace suggested that formate was converted to H2
before the start of dechlorination activity. Pyruvate and lactate also supported
dechlorination, however, H2 was measured in the headspace of both these
cultures and may have been responsible for the dechlorination activity.

H2 from formate oxidation supports dechlorination. Since H2
was observed in formate-ted OM enrichment cultures, the ability to oxidize
formate was tested in OM cultures that had been serially transferred eight times
with H2 as the electron donor. This would determine if H2 production from
formate was a stable feature of the OM enrichment culture. Upon feeding
formate to the serially-transferred cultures, extensive production of H2 was seen
followed by dechlorination of 2,5-DCP after a considerable lag time (Figure 6.3).
Stoichiometric production of hydrogen was observed from formate (data not
shown). Rates of dechlorination in formate-fed OM enrichment cultures were,
however, slower than H2-fed cultures, despite the presence of substantial H2 in
the former.

Evldence tor halorespiration. As an indicator of whether
halorespiration occurs in the OM enrichment culture, the fraction of electrons
from H2 used for reductive dechlorination, fe, was determined (Figure 6.4).
The to of 0.61 was observed in six replicate cultures. From Figure 6.4 it is
apparent that dechlorination did not start in the OM enrichment cultures until
102 pmoles (+/- 5) of H2 had been consumed. In autoclaved controls no loss of
H2 was observed, indicating that H2 consumption that occurred prior to

dechlorination was biologically mediated.

139

 

|oueud se|owrl

 

 

 

 

 

Days

Figure 6.3. Production of H2 from formate-fed OM enrichment cultures and the
dechlorination of of 2.5-DCP as indicated by the appearance of phenol.

140

 

500 _

  
 

 

400

 

 

 

 

 

.__ fe = 0.61 :

o . 95% conf. int. = +/- 0.022 :

a) 300 ,-

2 ~ " 2

CE) .

:2. 200_
1oob ‘

 

 

 

ilLl

 

o 100 200 300 400 500 600 700 800
umoles H2 consumed

Figure 6.4. The fraction of electrons (fe) used for dechlorination in the OM
enrichment culture. The fe is equivalent to the slope of the regression line of
the plotted data. The 95% confidence interval was calculated for six replicate
cultures. Dechlorination activity started after an average of 102 umoles of H2
had been consumed (arrow).

141
Additional evidence of halorespiration was provided by the determination
of the H2 threshold concentration. Cultures were starved for H2 while
maintaining the presence of 2.5-DCP as an electron acceptor. The H2
concentration was monitored until it was below the detection limit of the
instrument, <0.5 ppm (Figure 6.5). Rates of H2 uptake in the OM enrichment
cultures were slow, since the cultures were not shaken. Therefore 90 days

were required for the culture to approach a final H2 threshold concentration.

Discussion

Reductive dechlorination of chlorophenols often has exhibited a
preferential order for the removal of specific positions from the aromatic ring,
with ortho > para > meta (Cozza and Woods 1992; Juteau et al. 1995). The CM
enrichment culture, in contrast to these previous studies, is shown to
simultaneously remove ortho- and meta- chlorines from monochlorophenols
and 2.5-DCP. H2 serves as the sole electron donor for this culture. A
schematic of the dechlorination pathway exhibited by the OM enrichment
culture is shown in Figure 6.6. In addition to the dechlorination from both
positions occurring concurrently, two other features of the OM enrichment
culture are quite interesting. The culture exhibits no dechlorination activity with
3-CP alone, therefore meta-dechlorination activity was induced by the presence
of 2-CP. Second, 2.5-DCP, a substrate with chlorines located para to each
other, is completely dechlorinated to phenol while the ortho substituted
dichlorophenols 2,3-DCP and 2,6-DCP are not dechlorinated at all. This
suggests substrate specificity by the microbial population responsible for the
reductive dechlorination of 2,5-DCP. In contrast to the chlorophenol
halorespiring cultures, Anaeromyxo dehalogenans (20F strains) and

Desulfitobacterium halorespiricans strain C023, ortho-chlorines are not

142

 

 

 

 

 

 

 

 

 

 

 

 

8 .
1000 : :
7 Z 2
i

V — .
O 6 : 2
X 5 r 1
g- 4 E :
D. I 2
v .. ..
IN : 90 :
1 Z— -i
0 L1 l I L11 1 1 JJ 1 1 l l .1

0 1O 20 3O 4O 50 60 70

Figure 6.5. Hydrogen uptake and threshold concentrations by the OM
enrichment culture. Inset has log scale to show the lower final H2 threshold
concentration.

143

 

 

L

HCI OH
| HCI
OH Cl I OH
or
H2

H2

Figure 6.6. Schematic representation of dechlorination pathways of
OM enrichment culture. Heavier arrows represent the slightly favored
reactions, thus ortho-dechlorination occurs slightly faster than meta-

dechlorination.

 

144

removed from dichlorophenols with more than two adjacent (chlorine or
hydroxyl) substituents (e.g. 2,3-DCP and 2,6-DCP) (Chapters 3 and 4). Since
the OM enrichment culture removes meta-chlorines as well, it is possible that
2.5-DCP is the natural substrate for dechlorination by the responsible
microorganism. 2.5-DCP was identified as a natural compound occurring in the
salivary juices of certain grasshoppers (Eisner et al. 1971), so it is possible that
sufficient selection pressure has been presence to drive the evolution of
enzymes specific for the anaerobic dechlorination of 2.5-DCP.

Evidence that halorespiration occurrs in the OM enrichment culture is
apparent in the determination of the fa, and the measurement of hydrogen
uptake correlated to the presence of 2.5-DCP. The fe accounts for the portion
of electrons from the electron donor used for the reduction of the electron
acceptor (Criddle et al. 1991). A value of 0.61 is calculated for the OM
enrichment culture when grown with H2 and 2,5-DCP, which is similar to the
0.64 and 0.67 observed for A. dehalogenans strain ZCP-C and D.
halorespiricans strain 0023 respectively (Chapters 3 and 4). The to and
corresponding ts, the fraction of electrons from the electron donor used for
biomass, are correlated to the overall energy available from the metabolism of
the electron donor and acceptor (Criddle et al. 1991). The to and fa values
observed from the OM enrichment culture and the other halorespiring isolates
are in the range of predicted values for the AG°' from the overall reaction
(McCarty 1975). For example the AG°' from H2 oxidation and 2-CP
dechlorination to phenol is -236.9 kJ/mol H2 (Thauer et al. 1977; Dolfing and
Harrison 1992). Oxygen reduction, nitrate reduction to nitrite and sulfate
reduction to sulfide have AG°'s of -318 kJ/mol H2, -243 kJ/mol H2 and -118
kJ/mol H2 respectively, when coupled to H2 oxidation. It is apparent that the

AG°' associated with reductive dechlorination of chlorophenolic compounds is

145

less than 02, significantly higher than SO4'2, and very close to nitrate reduction
to nitrite. As theoretical fs values are based on AG°' calculations, the
corresponding to values for different electron acceptors increase in value as
follows: 02 < NO3' < 2-CP < 804'2 < CO2 (Criddle et al. 1991). McCarty
(1971) has shown that these theoretical values for to and fs are in general
agreement with observations made of different cultures. Therefore it can be
inferred from the fa data for the OM enrichment culture that the energy available
from reductive dechlorination is being coupled to growth.

Additional evidence for halorespiration in the OM enrichment culture is
provided in measurements of the apparent threshold concentration for H2. The
hydrogen threshold concentration reflects the minimum amount of energy that
an organism requires to grow (Lovley 1985; Lovley and Goodwin 1988).
Hydrogen threshold concentrations are therefore correlated with the energy
available from the redox couples mediating their growth. In the case of the OM
enrichment culture this is H2 oxidation and the reductive dechlorination of 2,5-
DCP reduction to phenol. The threshold concentration of < 0.5 ppm H2
measured is comparable to that observed for fumarate reduction and slightly
greater than found for nitrate respiration (Cord-Ruwisch et al. 1988). However,
it is much less than observed for sulfidogens, methanogens and acetogens.
The pure cultures Anaeromyxo dehalogenans (2CP strains) and
Desulfitobacterium halorespiricans strain C023 also had threshold
concentrations of H2 less than 1 ppm (Chapters 3 and 4). DeWeerd et al (1991)
measured the comparable threshold concentration of 0.7 ppm for hydrogen with
the halorespirer Desulfomonile tiedjei strain DCB-1 when grown on H2 and 3-
chlorobenzoate. This indicates that hydrogen threshold concentrations may be
reliable indicators of halorespiratory metabolism in pure and mixed cultures

where a halogenated substrate is the only electron acceptor provided other

146

than CO2. Hydrogen threshold concentrations have been shown to be more
correlated to the redox potential of the terminal electron acceptor rather than
individual microbial species (Cord-Ruwisch et al. 1988). Ninety days were
required by the OM enrichment culture to reach the H2 threshold concentration.
It would have been easy to misinterpret the final H2 concentration in the OM
enrichment culture due to the slow H2 consumption rate. Thus, it is important to
verify that a steady-state H2 threshold concentration has been achieved.

One interesting feature of the H2 consumption by the OM enrichment
culture is that approximately 100 pmoles was consumed prior to the beginning
of dechlorination (Figure 6.4). The low solubility of H2 in the media precludes
the liquid phase alone from being a significant sink for H2, a result which is
corroborated by the autoclaved control cultures. Therefore the initial H2 uptake
must be biologically mediated. One possible explanation is that non-
dechlorinating microorganisms, that provide essential co-factors to the
dechlorinating population, grow initially and consume most of the H2. Since the
non-methanogenic OM enrichment culture was fed only H2 + C02, the possible
types of non-dechlorinating coculture organisms is limited to one, a
homoacetogen. Acetate addition facilitated the onset of dechlorination when
added to the media, however, there was no less H2 consumed in these cultures
prior to dechlorination (data not shown). Other factors produced by acetogens
might also be important to the growth of the dechlorinating culture. Also since
H2 + CO2 conversion to formate is energetically exergonic, it is theoretically
possible that a microorganism could grow on these substrates in a non-
acetogenic process (Chapter 7). Such an organism might also provide
essential growth co-factors to the dechlorinating microorganism. It is interesting
that the OM enrichment culture exhibits the reverse reaction when formate was

provided as an electron donor, conversion to H2 + 002, which is also exergonic

147

under appropriate conditions. Formate was stoichiometrically converted to H2 4-
CO2 by the OM enrichment culture prior to the onset of dechlorination. It was
shown that the formate oxidation to CO2 with the production of H2 was
mediated by the novel sulfate reducer FOX1, which was not able to dechlorinate
2.5-DCP (Chapter 7). These results confirm that the OM enrichment culture has
at least two microbial populations: FOX1, perhaps responsible for initial H2
consumption, and the H2-consuming dechlorinating population. H2 may be the
only substrate that supports dechlorination in the OM enrichment, since it was
observed in the headspace of cultures amended with each of the organic
substrates tested as electron donors. This would be similar to Halobacter
restrictus which only uses H2 as an electron donor and PCE as an electron
acceptor (Holliger et al. 1993).

The prospects of isolating the population responsible for dechlorinating
2.5-DCP in the OM enrichment culture are good. The evidence that
halorespiration is occurring suggests that this mechanism could be utilized for
selective advantage in an enrichment. The best strategy may be to use a
slightly more complex medium (e.g. add yeast extract or rumen fluid) with
excess acetate. Any additional cofactors required by the dechlorinating
population would then be provided in the medium and the excess acetate
would provide a carbon source and possibly inhibit acetogenesis. The unique
ability of the OM enrichment culture to mediate simultaneous ortho- and meta-

dechlorination make it a very attractive candidate for isolation and further study.

References

Cole J.A., A. Cascarelli, w.w. Mohn and J.M. Tiedje. 1994. Isolation
and characterization of a novel bacterium growing via reductive
dechlorination of 2-chlorophenol. Appl. Environ. Microbiol. 60:3536-3542.

Cord-Ruwisch R., H.-J. Seitz and R. Conrad. 1988. The capacity of
hydrogenotrophic anaerobic bacteria to compete for traces of hydrogen

148

depends on the redox potential of the terminal electron acceptor. Arch.
Microbiol. 149:350-357.

Cozza C.L. and S.L. Woods. 1992. Reductive dechlorination pathways for
substituted benzenes: a correlation with electronic properties. Biodeg.
2:265-278.

Criddle C.S., L.M. Alvarez and P.L. McCarty. 1991. Microbial
processes in porous media, p. 641-691. In J. Bear and M. Y. Corapcioglu
(ed.), Transport processes in porous media. Kluwer Academic Publishers,
Dordrecht

de Jong E., J.A. Field, H.-E. Spinnler, J.B.P.A. lenberg and J.A.M.
do Bont. 1994. Significant biogenesis of chlorinated aromatics by fungi
in natural environments. Appl. Environ. Microbiol. 60:264-270.

DeWeerd K.A., F. Concannon and J.M. Suflita. 1991. Relationship
between hydrogen consumption, dehalogenation, and the reduction of
sulfur oxyanions by Desulfomonile tiedjei. Appl. Environ. Microbiol.
57:1929-1934.

Dolflng J. 1990. Reductive dechlorination of 3-chlorobenzoate is coupled to
ATP production and growth in an anaerobic bacterium, strain DOB-1.
Arch. Microbiol. 153:264-266.

Dolflng J. and B.K. Harrison. 1992. Gibbs free energy of formation of
halogenated aromatic compounds and their potential role as electron
acceptors in anaerobic environments. Environ. Sci. Technol. 26:2213-
2218.

Elsner T., LB. Hendry, D.B. Peakall and J. Meinwald. 1971. 2,5-
Dichlorophenol (from ingested herbicide?) in defensive secretion of
grasshopper. Science 172:277-278.

Grlbble G.W. 1992. Naturally occurring organohalogen compounds -- a
survey. J. Nat. Prod. 55:1353-1395.

Holliger C., G. Schraa, A.J.M. Stams and A.J.B. Zehnder. 1993. A
highly purified enrichment culture couples the reductive dechlorination of
tetrachloroethene to growth. Appl. Environ. Microbiol. 59:2991-2997.

Juteau P., R. Beaudet, G. McSween, F. Lépine, S. Mllot and J.-G.
Blsalllon. 1995. Anaerobic biodegradation of pentachlorophenol by a
methanogenic consortium. Appl. Microbiol. Biotechnol. 44:218-224.

Lovley D.R. 1985. Minimum threshold for hydrogen metabolism in
methanogenic bacteria. Appl. Environ. Microbiol. 49:1530-1531.

149

Lovley D.R. and S. Goodwin. 1988. Hydrogen concentrations as an
indicator of the predominant terminal electron-accepting reactions in
aquatic sediments. Geochim. Cosmochim. Acta 52:2993-3003.

McCarty P.L. 1971. Energetics and bacterial growth, p. 495-531. In J. Faust
and J. V. Hunter (ed.), Organic Compounds in Aquatic Environments.
Marcel Dekker, Inc., New York

McCarty P.L. 1975. Stoichiometry of biological reactions. Prog. Water Tech.
7:157-172.

Mlkesell M.D. and S.A. Boyd. 1986. Complete reductive dechlorination
and mineralization of pentachlorophenol by anaerobic microorganisms.
Appl. Environ. Microbiol. 52:861-865.

Mohn w.w. and J.M. Tiedje. 1990. Strain DOB-1 conserves energy for
growth from reductive dechlorination coupled to formate oxidation. Arch.
Microbiol. 153:267-271 .

Mohn w.w. and J.M. Tiedje. 1992. Microbial reductive dehalogenation.
Microbiol. Rev. 56:482-507.

Nlcholson D.K., S.L. Woods, J.D. Istok and D.C. Peek. 1992.
Reductive dechlorination of chlorophenols by a pentachlorophenol
acclimated methanogenic consortium. Appl. Environ. Microbiol. 58:2280-
2286.

Sanford R.A., J.R. Cole, F.E. Lbffler and J.M. Tiedle. 1996.
Characterization of Desulfitobacterium halorespiricans sp. nov. strain
0023, which grows by coupling the oxidation of lactate to the reductive
dechlorination of 3-chloro-4-hydroxybenzoate (SCI-4-HBA). Submitted to
Bicdegradation .

Thauer R.K., K. Jungermann and K. Decker. 1977. Energy
conservation in chemotrophic anaerobes. Bacteriol. Rev. 41:100-180.

Chapter VII

Anaerobic formate oxidation supports growth in
H2 producing microbial culture

All of the work described was done by Robert A. Sanford with the
exception of the 168 rRNA sequencing and phylogenetic analysis performed by
John W. Urbance. This chapter is formatted as a manuscript for submission to

the journal Science.

150

151

Abstract

Strain FOX1, a newly isolated microorganism obtains energy for growth
by anaerobic oxidation of formate to equimolar amounts of hydrogen and 002.
Such a reaction was not expected because the free-energy is so low, but
exponential growth of FOX1 continued to a AG' of -5 to -10 kJ, providing
evidence for a new lower limit of free-energy that supports growth. Furthermore,
FOX1 catalyzed the reaction to thermodynamic equilibrium. H2 was produced
at rates of 7300 ml h'1g'1 protein and to partial pressures in excess of 100 kPa.
suggesting that this bacterium may have economic value as a H2 source.
Analysis of the 168 ribosomal RNA places FOX1 within the family
Desulfovibrionaceae, in the 5-subdivision of the Proteobacteria. Our work also
suggests that the role of formate in anaerobic ecosystems should be

reevaluated, particularly its assumed equivalency to H2.

Hydrogen, a key intermediate in anaerobic ecosystems (1), has been
described as the fuel of the future (2) . It is an important feedstock in industrial
processes, as well as a source of clean energy (2). Although currently a
byproduct from petroleum refining, impurities from this source of H2 can prevent
its use in some processes, like fuel cells. Hydrogen is also produced from the
steam reformation of natural gas or methanol, which although a clean process,
requires high temperatures and therefore considerable energy input.
Biologically generated hydrogen may provide an environmentally sound,
energetically efficient, small scale production method that could be an attractive
renewable alternative to existing technologies. This is particularly true for H2
generated from industrial waste treatment. In the past biologically generated
hydrogen was not economical in part because of slow production rates and its

susceptibility to by-product inhibition (3). In addition light is required in many of

152

these biological processes, which may be impractical in some circumstances
(4). Other technologies being investigated are H2 from cyanobacterial
nitrogenase and the conversion of CO to H2 and CO2 (4). Although they show
promise these developing technologies either do not produce hydrogen at very
high rates or are limited by the solubility of the substrate, e.g. CO. Recently it
was shown that good rates of anaerobic fermentative H2 generation from
simulated carbohydrate wastes can be sustained (5). We have isolated strain
FOX1 (for formate oxidation), a bacterium that generates H2 from formic acid in
the dark in a energy-yielding process that sustains growth. Since formate is
common in anaerobic ferrnentations, FOX1 may also augment H2 production in
anaerobic waste treatment processes. Sufficient hydrogen was produced to
higher partial pressures by this culture and at sustained rates approaching an
order of magnitude higher than generally reported for other biological systems
(3, 4), and hence may be considered a new approach for hydrogen production.
Hydrogen is also important in anaerobic food webs that usually terminate
in methanogenic or acetogenic processes. The generation of methane or
acetate requires the presence of a suitable electron source, usually H2 or
formate. Obligate syntrophic microorganisms rely on methanogens to consume
the H2 they produce. This interspecies H2 transfer is a fundamental component
of anaerobic food webs terminating in methanogenesis (6, 7, 8). Many
researchers have proposed that formate transfer would be energetically
equivalent and more likely than H2 transfer mainly due to its greater solubility
and higher diffusion coefficient (6, 9). Microbiologists and engineers treated
these two electron donors as energetically equivalent, because their
interconversion is close to thermodynamic equilibrium under standard

conditions (10) (Equation 1).

153

HCOO' + H2O -—> HCOa' + H2 AG°' = +1.3 lereactlon Eq. 1

However, formate, H2 and CO2 are not likely to be in equilibrium in the
environment. At realistic environmental concentrations of reactants and
products, this slightly endergonic reaction becomes exergonic. The formate
oxidizing isolate we have discovered confirms this theoretical consideration,
since it grows using formate as its sole energy source. To calculate the AG' at

30° C we used the following formula:

AG'(kJ) .-. AG°' + 2.52ln([HC03'] . [H2]) - 2.52*In[HCOO'] Eq. 2

Figure 7.1 shows that at pH 7.0 with formate and HCOa' at 0.01 M this reaction
is exergonic for all H2 concentrations under 100 kPa (1 atm). The AG' is -21.9
kJ when the initial H2 partial pressure is 0.01 kPa. Under these initial
conditions it is clearly not energetically favorable to convert H2 and CO2 into
formate. Hence, interspecies H2 transfer should be favored over formate
transfer, when excess formate is present. This implies that there may be a
thermodynamic constraint on interspecies formate transfer, since if an organism
produces formate, it is energetically favorable to oxidize it to H2. In Figure 7.1
the energy available from CO2 reduction to methane with hydrogen is also
shown. The area shaded in the middle corresponds to H2 concentrations,
where formate oxidation would support both a formate oxidizing microorganism

(like FOX1) and a methanogen. Since this demonstrates that energy is

154

 

 

 

 

 

 

a: .
6 : ‘0. :
E . \\ .
E -50 ‘ .
3‘ ’ -10 kJ '20 “\I“.
(D ’ 5
<1 : \ :
0E . .
r ; Fox“: and/OR '3’3‘11'333925 FOX1‘I.‘ ‘
- , me anogen : ' on s‘ -
' 9 9”“ Igm ~ growth y
50 l l l 1 l l l l l l l l l 1 l l l l 1 l 1 J l l I l l l l l l l l 1
' 2 1 o -1 -2 -3 -4 -5
log H2 (kPa)

Figure 7.1. The relationship between the H2 partial pressure and the AG' for
formate oxidation (0) (HCOO' 4» H20 --> HCOa' + H2) and for
methanogenesis (Q) (4 H2 + HCOa' + H+ --> 3H2O + CH4). Initial
concentrations are: formate = 0.01 M, HCOa' = 0.01 M and CH4 = 10 kPa.
The central shaded area shows the H2 concentration range that would
support the growth of a formate oxidizing organism and a methanogen
consuming the H2 in a simple anaerobic community. The adjacent flanking
shaded regions indicate the extended H2 concentration range that
corresponded to the AG' associated with the growth of FOX1 alone.

155

available from formate oxidation, we sought and found a somewhat specialized
microorganism that has evolved the capability to couple growth to this process.

FOX1 was obtained from an anoxic enrichment culture fed volatile fatty
acids (VFAs), including formate, and that showed reductive dechlorination of
chlorOphenol. Analysis of the headspace gas of this enrichment showed
significant concentrations of H2 (3.0 kPa), but only trace amounts of methane,
which was expected to be the predominant product. Both the dechlorination
activity and H2-producing activity were serially transferred twice in enrichment
cultures that were supplemented with a vitamin mixture (450 pg/l) before an
effort was made to isolate colonies on anaerobic medium (11). Transfers of this
culture have shown that formate alone will support the production of hydrogen,
although this activity is stimulated when acetate is added to the medium. This is
not unusual for anaerobic cultures, particularly those that have so little energy
available. Presumably acetate is used for biosynthesis of cell material. FOX1
was purified from the original enrichment by serial dilution in the
formate/acetate medium (12).

To prove that anaerobic formate oxidation supported actual growth of
FOX1, direct microscopic counts of acridine orange stained cells were made by
taking samples at different time intervals (Figure 7.2 A). Direct counts were done
since the total number of cells were not high enough for a turbidity assay.
Controls without formate or with H2 and CO2 did not show appreciable
increases in cell numbers. The doubling time was 30 hours during exponential
growth. Autoclaved controls containing cells with formate and acetate exhibited
no H2 evolution(data not shown). Figure 7.2 B shows that as formate was
consumed stoichiometric quantities of H2 were evolved from the same culture.
No acetate or other VFAs were produced by this culture. Also shown in Figure

7.2 B is the AG' calculated at different points during the growth of this culture.

156

 

250’

 

   
   

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

t A
15 200}
‘; : + Formate\
g; 150}
a .
0 :
“5 1007
o' I No Formats
2 .
50: Hz-I-CO2
.9
“5 l>
E 62
O
",7, E
\
f3 .3.
E 5'
:3.
(E..1,4411...11L...3-35
0 so 100 150 200 250 300 350

Hours

Figure 7.2. A. Growth of anaerobic formate oxidizing bacterium FOX1 in broth
culture at 30° C as measured by direct microscopic counts of acridine orange
stained cells. A known volume of culture was stained and passed through a
0.2 micron filter. Stained cells were then observed under fluorescent
illumination and enumerated. B. Illustrates the loss of formate from cultures
and the subsequent stoichiometric production of H2. Culture volume was 100
ml with a headspace of 60 ml. Error bars indicate the 95% confidence
intervals for triplicate cultures.

157

Significant growth occurred at AG' values between -10 kJ and -5 kJ, at which
point exponential increase in cells stopped. This would be a lower growth
supporting AG than has been observed for any microorganism. A minimum AG
of -20 kJ had been proposed due to the theoretical quantum of energy required
to translocate one proton, which would correspond to one-third ATP (1).

The H2 evolution rate in batch cultures was 7300 ml h'1g'1 protein (0.33
mole h'1g'1) and the cell yield was 20.9 pg protein per mmol formate
consumed. Microscopic observations of cells in stationary phase showed that
FOX1 is pleomorphic after growth on formate (Figure 7.3), while little variation in
cell morphology was observed after growth on pyruvate. This may be the due to
the stress caused by the limited energy available from formate oxidation.

Preliminary identification and characterization of FOX1 was achieved by
phylogenetic analysis using the 168 ribosomal RNA gene sequence (13). Four
168 rDNA clones from the enrichment culture were examined and all had
essentially identical sequences. The identical sequences and unambiguous
sequence results from the uncloned PCR product suggested that the culture did
consist of a single organism. Phylogenetic analysis placed the formate-
oxidizing organism within the family Desulfovibrionaceae, in the delta-
subdivision of the Proteobacteria, as illustrated in Figure 7.4 (14, 15). Its
nearest relative was Desulfovibrio africanus (88.0% sequence similarity).
Preliminary experiments have shown that FOX1 is a sulfate reducer (12) and
therefore its physiology is consistent with its phylogeny. It is perhaps not
surprising to find this organism related to the delta-subgroup of the
Proteobacteria, since this group is particularly noted for its diversity of anaerobic
processes.

Since the energy apparently available from anaerobic formate oxidation

to hydrogen is minimal (Eq.1), we determined whether thermodynamic

05054533

 

Figure 7.3. Fluorescent microscopic image of acridine orange stained FOX 1
bacterium after growth on formats. The average size of cells in the culture is 2
pm wide and 3 to 10 um long.

159

 

Escherichia coli
Desulfobacter postgatei
Desulfo vibrio salexigens
Desulfovibn‘o desulfun'cans E1
Desulfovibn’o‘ sp. str. PlB
Desulfovibrio sp. str. Teske
Desulfovibn'o Iongus
Desulfovibrio gigas
FOX1
Desulfovibrio africanus
Desulfo vibn'o desulfuricans
Desulfovibrios vulgaris
Desulfovibrio Iongreachii
‘— Desulfovibrio sp. str. PT-2
, sym. Sus_sc (NCTC 12656)’
I sym. Mesoau“
.Desulfomicrobium baculatus
Desulfovibrio desulfuricans str. Norway 4
Desulfomicrobium escambium
Desulfurella acetivorans
-—[ Camp ylobacter fetus
Helicobacter pylori

 

  
   
  
 

 

 

 

 

 

 

 

 

 

 

 

 

 

Desulfomonile tiedjei
Geobacter metal/ireducens

 

 

 

.10

 

* Intracellular ileal symbiont of Suis scrofus
** Intracellular ileal symbiont of Mesocn‘cetus auratus

Figure 7.4. 168 rRNA phylogeny of FOX1 and related organisms. Near
complete (ca. 1500 nucleotide positions) 16S rRNA genes were aligned using
the primary and secondary structure with sequences of the nearest relatives
obtained from the Ribosome Database Project (RDP). Phylogenetic
relationships were inferred by maximum likelihood analysis (15).

160

equilibrium was reached or whether there was a H2 threshold concentration
beyond which no further formate oxidation would occur. A threshold may be
expected in some systems since energy may be required to initiate the process,

such as transport of formate. If this were the case the free-energy (AG’) would

be negative for the system favoring further formate oxidation even after H2
evolution ceases. To determine this, cultures of the H2 producer were initiated
with different initial formate concentrations: 15 mM, 30 mM and 60 mM. Results
showed that as the initial formate concentration increased the final H2
concentration increased to as high as 66 kPa with the 60 mM formate culture
(Figure 7.5 A). In fact H2 concentrations of greater than 100 kPa were obtained
with similar cultures that had been rated formic acid. Regardless of the initial
formate concentration the calculated AG' at the end of incubation was
approximately 0 kJ/mole, indicating that thermodynamic equilibrium had been
obtained (Figure 7.5 B). Most of the available free-energy was used during
initial formate oxidation when no H2 was present. For example, when equation
2 is calculated after the reaction has partially proceeded, with formate equal to
0.009 M, HCOa' at 0.011 M and H2 equal to 3.7 kPa, the AG' increases from
-21.9 kJ to 65 kJ (assuming a 100 ml culture with a 60 ml headspace). At
equilibrium under the same conditions, the H2 partial pressure reaches
approximately 18.5 kPa, illustrating that most of the H2 evolution exhibited by
this organism does not yield much energy. The ability to easily determine the
free-energy under initial conditions and at equilibrium is perhaps unique for a
biological process that yields energy for growth and hence may be a good
model for studying the coupling of free-energy to growth.

Our observations demonstrate the feasibility of formate serving as an
energy source for anaerobic growth, with stoichiometric production of hydrogen.

Since FOX1 produces H2 at sustained rates up to ten times those of other

161

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

'35 - - L l - a . I A L a L 4 - - l A ; - I - - -
0 4 8 12 16 20 24
Days

Figure 7.5. Hydrogen production and free energy change in duplicate
cultures started at three formate concentrations: 15 mM(O), 30 mM(D) and 60
mM(A). A. Graphically represented is the average partial pressure for H2 in
duplicate cultures. Hydrogen concentration reached a steady state in all
cultures after 20 days. B. Free energy AG' is shown for the same cultures with
different initial formate concentrations. The AG' was calculated by measuring
the H2 partial pressure and determining by difference the formate and
bicarbonate concentrations. Formate was measured initially and at the final
data point to verify the free-energy calculations. All three cultures converge at
thermodynamic equilibrium at 20 days.

162

biological systems (4), it may provide a more attractive method for industrial H2
production. Although many microorganisms, including E. coli, can convert
formate into H2, this reaction has not been described as energetically beneficial
and independent of other energy yielding substrates (16, 17). The discovery of
this culture suggests that some of the previously described H2-evolving,
formate-oxidizers should be reevaluated for the potential energetic benefit of
this process, particularly other sulfate-reducing 5-Proteobacteria. The
Desulfovibrio are known to contain a variety of hydrogenases and are known to
possess the low-potential electron carriers, like ferrodoxins, that would be
required for the energetically favorable generation of H2 (18), which may then
be coupled to cell growth. The positioning of hydrogenases across the
membrane is one mechanism that may explain growth of FOX1 at a AG' greater
than -10 kJ, since H2 diffusion out of a cell could serve as a type of proton
translocation with little energetic cost.

The role of formate in anaerobic food webs, particularly its possible role
in interspecies electron transfer, should be reevaluated. For example in a
methanogenic ecosystem with a significant formate flux, the formate oxidation to
methane and CO2 may actually support the growth of two microbial
populations: the FOX1-type H2-producer and the methanogen (Figure 7.1).
The only requirement for this type of consortium to function is to have formate
not in equilibrium with H2 and CO2. Such an environment was described by
McMahon and Chapelle when they found that high concentrations of formic and
acetic acid are produced in aquitard sediments and subsequently diffuse into
anaerobic sulfidogenic and methanogenic aquifer material (19). Since H2
concentrations in such anaerobic environments are kept very low by

methanogenesis and sulfate reduction, the oxidation of formate to H2 would be

163

thermodynamically favorable. Therefore reasonable conditions do exist in the

natural environment for the anaerobic oxidation of formate to hydrogen.

References and Notes

1. B. Schink, M. Friedrich, FEMS Microbiol. Rev. 15, 85-94 (1994).

2. J. A. Serfass, D. Nahmias, A. J. Appleby, Int. J. Hydrogen Energy16,
551-556 (1991).

3. A. M. KIibanov, B. N. Alberti, S. E. Zale, Biotechnol. Bioeng. 24, 25-36
(1982).

4. P. F. Weaver, P. -C. Maness, S. Martin, S. Muralidharan, A. J. Frank
Proceedings of the 1995 DOE Hydrogen Program Review, Volume ll., Coral
Gables, Florida (National Renewable Energy Laboratory--NREL/CP-430-
20036). 675-681 (1995). Y. Nemoto ibid 617-674 (1995). M. L. Ghirardi, S. P.
Toon, M. Seibert ibid 683-692 (1995). J. Woodward, S. M. Mattingly ibid 833-
845 (1995). A. Mitsui Proceedings of the 1994 DOE/NREL Hydrogen Program
Review, Liverrnore, California (National Renewable Energy Laboratory). 353-
395 (1994). P. Weaver, P.-C. Maness, A. Frank, S. Li, S. Toon ibid 397-406. M.
A. Bennett, H. H. Weetall, J. Solid-Phase Biochemi. 1, 137-142 (1976).

5. N. Ron, 8. Wang, F. Ma, Water Environ. Res. In press (1996). F.
Taguchi, N. Mizukami, T. Saito-Taki, K. Hasegawa, Can. J. Microbial. 41, 536-
540 (1995).

6. S. S. Ozturk, B. O. Palsson, J. H. Thiele, Biotechnol. Bioeng. 33, 745-757
(1989). A. J. M. Stams, Antonie van Leeuwenhoek 66, 271-294 (1994).

7. M. P. Bryant, E. A. Wolin, M. J. Wolin, R. S. Wolfe, Arch. Microbial. 59, 20-
31 (1967). M. J. Wolin, in Microbial interactions and communities, vol. 1‘ A. T.
Bull, J. H. Slater, Eds. (Academic Press, London, 1982) pp. 323-356. M. J.
Wolin, T. L. Miller, ASM News 48, 561-565 (1982).

a. R. Conrad, B. Schink, T. J. Phelps, FEMS Microbial. Ecol. 33, 353-360
(1986)

9. D. R. Boone, R. L. Johnson, Y. Liu, Appl. Environ. Microbial. 55, 1735-
1741 (1989).

10. R. K. Thauer, K. Jungermann, K. Decker, Bacterial. Rev. 41, 100-180
(1977)

11. T. O. Stevens, H. D. Watts, J. J. Walker, J. K. Fredrickson, in Abstracts of
the 92nd General Meeting of the American Society for Microbiology 1992.

164

(American Society for Microbiology, New Orleans, LA, 1992) pp. 366. E. A.
Wolin, M. J. Wolin, R. S. Wolfe, J. Biol. Chem. 238, 2882-2886 (1963). The
basic medium composition was obtained from Stevens et. al. and included the
following comonents: 2 mM potassium phosphate buffer (pH 7.2 - 7.5), and per
liter CaCI2-2H2O 0.015 g, MgCI2-6H20 0.02 g, FeSO4 -7H20 0.007 g,
Na2SO4 0.005 g, (trace metals solution) MnCI2-4H2O (5 m9). H3B03 (0.5 mg),
ZnCl2 (0.5 mg), CoCI2-6H2O (0.5 mg), NiSO4-6H2O (0.5 m9). CuCl2-2H2O
(0.3 mg), NaMoO4-2H2O (0.1 mg). NazSeO4 (0.003 mg) and Na2WO4 (0.008
mg). NH4CI was added to a concentration of 8 mM and 10 mM NaHC03 was
added to buffer the headspace which contained a N2:CO2 ratio of 95:5.
Vitamins were added as described by Wolin et. al. The medium used for plating
the culture contained formate, butyrate, succinate and propionate (VFA mixture)
at a concentration of 2 mM each. Colonies of several morphologies grew on
these plates and were subsequently transferred as a mixture back into anoxic
broth containing the VFAs and chlorophenols. Although no dechlorination
activity occurred over several months, significant hydrogen production was
measured along with the depletion of formate. The rate of hydrogen generation
was also significantly increased under shaking at 30° C.

12. Formate grown cultures were transferred eight times with 0.1 % (v/v) per
transfer. After this three successive serial dilution series to extinction of H2

generation were done to purify FOX1. Since so little energy is available from

formate oxidation, dilutions of 10'6 were sufficient to achieve purity. The purity
of FOX1 was verified microscopically by observing cells grown on pyruvate (a
common fermentation substrate) and lactate plus sulfate (a common substrate
for sulfidogens). Pyruvate was fermented to acetate and succinate while sulfate
was reduced to sulfide by FOX1. Observations verified the presence of a single
morphotype and both of these rich-media cultures were successfully transferred
back to a formate medium which showed H2 production activity. Also pyruvate
grown FOX1 was subjected to DNA extraction and PCR amplification of 16S
rDNA. Only one sequence, was observed which was identical to the sequence
found for FOX1 grown on formate. Efforts to grow FOX1 on agar medium have
been unsuccessful.

13. A replicate of the formate-oxidizing culture was harvested by
centrifugation. Cells were lysed by repeated freeze/thaw cycles and the nucleic
acids purified by phenol/chloroform extraction and alcohol precipitation. Near
complete (ca. 1500 bp) 16S rRNA genes were then PCR amplified using
previously described primers. J. Zhou, M. R. Fries, J. Chee-Sanford, J. M.
Tiedje, Int. J. Syst. Bacterial. 45, 500-506 (1995). A fraction of the amplified
product was cloned using a commercial kit (TA Cloning Kit. lnvitrogen. San
Diego, CA.) and four of the resulting clones were then randomly selected and
sequenced, along with the uncloned, amplified product.

14. R. Devereux, et al., J. Bacterial. 172, 3609-3619 (1990).

15. B. L. Maidak, et al., Nucleic Acids Research 22, 3485-3487 (1994). J.
Felsenstein, J. Mal. EvoI. 17, 368-376 (1981). G. J. Olsen, H. Matsuda, R.
Hagstrom, R. Overbeek. Camput. Appl. Biasci. 10, 41-48 (1994). Sequences

165

from the nearest relatives were identified and obtained from the Ribosome
Database Project (RDP) using the SIMILARITY_RANK and SUBALIGNMENT
programs of the RDP respectively and the FOX1 sequence brought into
alignment using both primary and secondary structure using the GDE program
obtained from the RDP.

16. G. Sawers, Antonie van Leeuwenhoek 66, 57-88 (1994).
17. C. T. Gray, H. Gest, Science 143, 186-192 (1965).

18. G. Fauque, et al., FEMS Microbial. Rev. 54, 299-344 (1988). J. M. Odom,
H. D. Peck Jr., FEMS Microbial. Left. 12, 47-50 (1981). J. Legall, G. Fauque, in
Biology of Anaerobic Microorganisms, A. J. B. Zehnder, Ed. (Wiley, New York,
1988), pp. 587-639. Hydrogenasos have been found in both the cytoplasm and
periplasmic space of sulfate reducers. One hypothesis for this arrangement of
hydrogenases is that they function in hydrogen cycling. The internal
hydrogenase produces H2, while the external hydrogenase consumes H2,
generating protons outside the cell. This facilitates the formation of a
membrane potential.

19. P. B. McMahon, F. H. Chapelle, Nature 349, 233-235 (1991).

20. This work was supported by the Center for Microbial Ecology through
grant No. BIR-91006 from the National Science Foundation.

Chapter VIII
SUMMARY AND CONCLUSIONS

In the concluding chapter I will summarize how my research has
answered my primary research questions and the importance of certain
concepts that have emerged from this work. In doing so, the contribution of
individual chapters to the synthesis of those concepts will become apparent.

Four research questions were addressed during this study (Chapter 1).
The answer to the first question appears to be that denitrifiers do not exist that
can dechlorinate and degrade chlorophenols. My results more thoroughly
evaluated this trend noted from other research (Chapter 2), although it is very
difficult to conclude that this is globally true. It is still a reasonable assertion that
some microorganism that is difficult to culture may be capable of both of these
activities and therefore an absolute statement cannot be made. At best such
organisms are rare or are difficult to enrich.

The second research question was to identify the microbial populations
responsible for reductive dechlorination of chlorophenols. Five different
dechlorinating activities were identified in enrichment cultures (Chapter 2).
Four of these dechlorinating cultures were chosen for further experimentation,
which resulted in the isolation of the anaerobic myxobacterium strain 2CP-C
(Chapter 3) and Desulfitobacterium chlororespirans strain C023 (Chapter 4).
These isolates exhibited the novel capability of halorespiration using
chlorophenolic compounds as a terminal electron acceptor. The other two

dechlorinating cultures exhibited distinctly different dechlorination patterns, but

166

167
did not yield isolates (Chapters 5 and 6). In the case of the meta-dechlorinating
methanogenic enrichment culture (Chapter 5), halorespiration did not appear to
occur since most of the available electrons from the added electron donor pool
were converted to methane. Since the energy from reductive dechlorination is
more favorable than methanogenesis, it is not likely that this process was
coupled to growth in these enrichment cultures. Of the five dechlorinating types
of cultures identified, four were enriched from the same type of microcosm, one
containing a Michigan compost soil. This suggests that considerable diversity
of microorganisms capable of reductive dehalogenation exists in non-polluted
and not strictly anaerobic environments.

The third research question was to determine the energetic value of
reductive dechlorination for the microbial populations in the enrichment
cultures. Cultures of strains ZCP-C and 0023 could be studied by monitoring
growth coupled to the reductive dechlorination of a chlorophenolic compound,
however this is more difficult to assess in enrichment cultures. I found that the
measurement of the H2 threshold concentration and the fraction of electrons
from the electron donor used for reductive dechlorination (fe), were good
indicators that growth was coupled to this process. For strains 0023, ZCP-C
and the OM enrichment culture the fa was between 0.6 and 0.7 (Chapters 3,4
and 6), while the fa of the methanogenic enrichment culture was only 0.01,
indicating a minor flux of electrons to reductive dechlorination (Chapter 5).
When compared to other electron accepting processes the fa for the reductively
dechlorinating pure cultures was between that seen for denitrification and
sulfate reduction. This is in the predicted range if halorespiration is occurring.
The H2 threshold concentration has been proposed as an indicator of the
microbial redox couple controlling particular environmental conditions. If

reductive dechlorination is supporting a respiratory process, 3 H2 threshold

168

range is predictable based on comparisons to other electron acceptors. Thus
H2 thresholds would increase for different electron acceptors in this order: O2 <
N03' < chlorinated organic compounds < 804‘2 < CO2 (methanogenesis).
Results of H2 measurements from strains ZCP-C, 0023 and the OM enrichment
culture all showed H2 concentrations less than 1 ppm (Chapters 3, 4 and 6).
This is in agreement with the predicted range of concentrations for
halorespiration. In addition, the demonstration of H2 serving as an electron
donor for reductive dechlorination is also indicative of a respiratory process. To
more thoroughly evaluate this principle, it would be useful to compare the to
and H2 threshold concentrations of other reductive dechlorinating cultures with
the cultures that I studied. This would more clearly establish these
measurements as good indicators of halorespiration.

My evidence suggests that the cultures that I have characterized have an
apparent preference for relatively low electron acceptor concentrations. In
Chapter 3 I introduce the term microrespirotrophy to describe this behavior. It is
possible that all the reductive dechlorinating populations are
microrespirotrophs, like strain C023 or the dechlorinators present in the
methanogenic and OM enrichment cultures (Chapters 4, 5 and 6). None of the
anaerobic dechlorinating cultures tolerated chlorophenol concentrations above
200 uM very well. Even other electron acceptors could not be supplied in
concentrations much above 1 mM. The main point I wish to stress is that the
use of excessive electron acceptor concentrations in enrichments or isolation
protocols may reduce the number of apparently culturable microorganisms. It
is quite likely that most microorganisms in nature are microrespirotrophs, simply
because electron acceptor concentrations are rarely excessive. This suggests
that strategies for culturing new microorganisms may be more successful if low

concentrations of electron acceptor are used. Even those organisms that can

 

169
use higher concentrations will still likely be able to grow at the lower
concentrations, so the benefit would be to maximize the recovery of culturable
organisms.

The final research question was to assess the influence of electron
donors an anaerobic food-webs terminating with reductive dechlorination and
particularly to investigate the relationship between formate and H2 in anaerobic
ecosystems. Specificity for different electron donors supporting reductive
dechlorination was evident among the different dechlorinating cultures
(Chapters 3,4,5 and 6). This suggests that the diversity of dechlorinating
microorganisms is not only dependent on the electron acceptor specificity but
also on the ability to use different electron donors. For example, strains 2CP-3
and 0023 both use 2,6-DCP as an electron acceptor and were isolated from the
same environment. As unique electron donors, strain ZCP-3 can use acetate
and strain 0023 can use butyrate, thus it is theoretically possible to maintain a
co-culture of these organisms on 2,6-DCP and butyrate, since 0023 oxidizes
butyrate to acetate and can only remove one chlorine. In the case of the meta-
dechlorinating methanogenic enrichment culture different electron donors
supported different efficiencies of reductive dechlorination with formate serving
as the most effective substrate for reductive dechlorination activity (Chapter 5).
Formate was also the best electron donor for the OM-enrichment culture, but
was converted to H2 and CO2 prior to reductive dechlorination, suggesting that
H2 was the true electron donor (Chapter 6). This led to the isolation of strain
FOX1, the microorganism capable of growing at the expense of formate
oxidation and H2 generation (Chapter 7). Strain FOX1 was shown to grow at
AG' in the range of -10 kJ to -5 kJ per mole of formate, higher than the reported
minimum AG' thought necessary for growth. Ecologically, this novel ability in

strain FOX1 implies that formate and H2 are not equivalent in the environment

170

and that some organisms may take advantage of the energy available from their
disequilibrium. In the OM-enrichment culture FOX1 was cross-feeding a
dechlorinating microorganism, however the cell-yields were always low.
Theoretically it should be possible to co-culture FOX1 with a non-formate-
utilizing methanogen and demonstrate an increase in cell-yield of FOX1. This
would demonstrate the potential importance of this type of activity in the an
anaerobic food-web.

Research in the future on this tapic should focus on two areas. First the
role of halorespiration in anaerobic environments needs to be more completely
investigated. Particularly important is the determination of what naturally
occurring chlorinated organic compounds serve as electron acceptors. By
focusing the research toward natural ecosystems the diversity of
microorganisms capable of halorespiration may be more thoroughly
determined. I would recommend the second area focus to be on formate
oxidation resulting in the generation of H2. From the preliminary studies with
FOX1 it is evident that a viable economic method for generating H2 may be
developed using these types of microorganisms. This is especially true if
anaerobic digestion of waste material can be channeled and enriched for
formate production leading subsequently to H2 generation. Studying the
mechanisms associated with formate oxidation by FOX1 and similar types of
organisms would also facilitate the development of these anaerobic H2

generating systems.

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