giovanni gualandi
Università degli Studi di Bologna
Dipartimento di Ingegneria Chimica, Mineraria e delle Tecnologie Ambientali
Dottorato di Ricerca
in Ingegneria Chimica, dell’Ambiente e della Sicurezza
XIX Ciclo
CHLORINATED ALIPHATIC AND AROMATIC HYDROCARBONS
BIODEGRADATION:
BIOAUGMENTATION TESTS IN SLURRY MICROCOSMS AND STUDY
OF THE CATABOLIC POTENTIAL OF MICROBIAL COMMUNITY IN
THE INTERFACE BETWEEN GROUNDWATER AND SURFACE
WATER
Tesi di Dottorato di:
Ing. GIOVANNI GUALANDI
Relatore:
Prof. Ing. MASSIMO NOCENTINI
Coordinatore del corso di Dottorato:
Prof. Ing. GIULIO CESARE SARTI
Sigla Settore Scientifico Disciplinare:
ING-IND/26 TEORIA DELLO SVILUPPO DEI PROCESSI CHIMICI
TABLE OF CONTENTS
Page
INTRODUCTION AND PROBLEM DEFINITION………………………….…1
1 VOLATILE ORGANIC COMPOUNDS………………………………………..6
1.1 Introduction…………………………………………………………………....6
1.2 Volatile Oganic Compounds Detected in Groundwater……………………….7
1.3 Chlorinated Aliphatc Hydrocarbons (CAHs)……………………...…………10
1.3.1 Sources of CAHs……………………………………………………..10
1.3.2 Physical and Chemical Properties…………………………………....15
1.3.3 Transport Processes…………………………………………………..15
1.3.1 Transformation of CAHs……………………………………………..18
1.3.4.1 Abiotic Degradation Processes…………………………...20
1.3.4.2 Biotic Degradation Processes………………………….…24
1.3.4.2.1 Aerobic Biodegradation……………………...…25
1.3.4.2.2 Anaerobic Biodegradation……………………...30
1.4 Clorinated Benzenes………………………………………………………….37
1.4.1
Sources of Chlorinated Benzenes…………………............……...37
1.4.2
Biodegradation of Chlorinated Benzenes………...............…...….37
1.4.3
Biodegradation of Chlorobenzene………………............………..42
1.4.3.1Degradation via the Ortho-Cleavage Pathway……
...........43
1.4.3.2Degradation via the Meta-Cleavage Pathway...........…
......45
1.4.3.3 Biodegradation of Mixtures of Substituted Benzenes........47
1.4.3.4 Degradation under Oxygen-Limited Conditions................47
2
BIODEGRADATION TECHNOLOGIES FOR REMEDIATION OF
CONTAMINATED
SITES.................................................................................50
ii
2.1 Introduction......................................................................................................50
2.2 In Situ Bioremediation.....................................................................................53
2.2.1 Intrinsic In Situ Bioremediation.....................................................54
2.2.2 Enhanced In Situ Bioremediation...................................................55
2.2.2.1 Vadose Zone Soil Remediation..........................................56
2.2.2.2 Surficial Soil
Remediation..................................................59
2.2.2.3 Groundwater and Saturated Soil
Remediation....................60
2.3 Ex-Situ Bioemediation……………………………………………………….68
2.3.1
Solids……………………………………………………………..68
2.3.1.1 Land treatment……………………………………………
68
2.3.1.2 Composting………………………………………………
.69
2.3.1.3 Biopiles…………………………………………………
...70
2.3.2
Solid-Liquid Mixtures……………….…………………………...71
2.3.2.1 Slurry Bioreactors………………………….……………72
3
2.3.3
Liquids……………………………………………………………72
2.3.4
Constructed Wetlands…………………………………………….73
KINETIC MODELS FOR COMETABOLISM
3.1 Enxymatic Reaction: Michaelis Menten Kinetics…………………………....76
3.2 Substrate Inhibition…………………………………………………………..79
3.3 CAHs Degradation Product toxicity…………………………………………82
iii
4
MOLECULAR BIOLOGY TECHNIQUES USED IN THE ANALYSIS OF
MICROBIAL DIVERSITY IN CONTAMINATED SOILS…………….…..88
4.1 Microbial Ecology of Contaminated Soils…………………………………...89
4.1.1
Phenotypic Analysis of Soil Bacterial Communities………...…..89
4.1.2
Culture-independent Analysis of Microbial Communities by
Lipid-Based Tools………………………………………………..90
4.1.3
Culture-independent Analysis of the Bacterial Communities by
Nucleic Acid-based Tools………………………………..………91
5
GROWTH OF CHLORINATED SOLVENT-DEGRADING MICROBIAL
CONSORTIA IN METHANE- AND PROPANE-FED BIOREACTORS AND
TESTING OF THEIR EFFECTIVENESS AS INOCULA FOR THE
BIOAUGMENTATION OF DIFFERENT TYPES OF AQUIFERS………105
Abstract…...………………………………………………………………...105
5.1 Introduction…………………………………………………………………106
5.2 Materials and Methods……………………………………………………...110
5.2.1
Overview of the Experimental Scheme……….….……………..110
5.2.2
Growt Bioreactors Setup and Operation………………………...117
5.2.3
Microcosms Setup and Operation……………………………….120
5.2.4
Estimation of Lag Times, Degradation Rates and Rate/concentration
Ratios………………………………………122
5.2.5
Analytical Methods……………………………………………...125
5.3 Results………………………………………………………………………129
5.3.1
CAH Depletion in The Sterilized Controls……………………...129
5.3.2
Behaviour of The Duplicate Slurry Microcosms………………..130
5.3.3
Effect of Inoculum Growth Time and Condition (presence/absence
of CAHs)………………………………………………………..130
5.3.4
Effect of The Type of Aquifer Material Contained in The Inoculated
Microcosms…………………………………………135
5.3.5
Effect of The Type of Microbial Consortium Inoculated and Growth
Substrate Supplied……………………………………..139
5.3.6
Results Relative to The Liquid-phase Microcosms……………..143
5.4 Discussion and Conclusion………………...……………………………….144
iv
6
BIODEGRADATION OF CHLOROBENZENE:
STUDY OF THE CATABOLIC POTENTIAL AND THE STRUCTURE OF
THE MICROBIAL COMMUNITY IN THE INTERFACE BETWEEN
GROUNDWATER AND SURFACE WATER…………………………….147
6.1 Introduction…………………………………………………………………148
6.2 Materials and Methods……………………………………………………...151
6.2.1
Test Site and Sampling………………………………………….151
6.2.2
Retrieval of The Samples……………………………………….154
6.2.3
Batch Degradation Tests of Monochlorobenzene……………....158
6.2.4
GC-analysis of Monochlorobenzene Concentration in The
Microcosms……………………………………………………..160
6.2.5
Analysis of Dissolved Oxygen Concentration in The Microcosms…
…………………………………………………..160
6.2.6
Methanol Extraction of VOCs and GC-MS Analyses………….161
6.2.7
DNA extraction from sediment and water samples…………….161
6.2.8
PCR Amplification…...………………………………………....162
6.2.9
Denaturing Gradient Gel Electrophoresis………………………166
6.2.10 Cloning, Sequencing and Analysis of PCR Amplified 16S rRNA,
TmoA and DsrB gene…………………………………………...168
6.3 Results………………………………………………………………………169
6.3.1 Monitoring of The In Situ Physico-chemical Parameters……….169
6.3.2
Pollutants Concentration at Different Depths in The Sediment
Cores……………………………………………………………174
6.3.3
Degradation of Monochlorobenzene under Oxygen–limited
Conditions in Batch Degradation Tests…………………………175
6.3.4
Detection of Catabolic Genes at Different Depths of The Interface…
………………………………………………………181
6.3.5
Study of the Structure of The Eubacterial Community by 16S rRNA
Gene PCR - DGGE Analysis…………………………….186
v
6.3.6
Study of the Structure of The Sulfate-Reducing Bacterial
Community by PCR - DGGE Analysis…………………………195
6.3.7 PCR-DGGE Analysis of TmoA-like Genes……………………..199
6.3.8
Batch Degradation Tests: Study of The Evolution of The Microbial
Community During The Incubation Period……………………..203
6.3.9 Cloning and Sequencing of TmoA Gene Fragments……………207
6.3.10 Cloning and Sequencing of DsrB and 16SrRNA Gene Fragments…
…………………………………………………….211
6.4 Discussion…………………………………………………………………..213
6.5 Conclusions…………………………………………………………………219
CONCLUSIONS………………………………………………………………..221
BIBLIOGRAPHY………………………………………………………………225
vi
INTRODUCTION AND PROBLEM DEFINITION
______________________________________________________________________________________________________________________________________________________________
Especially in the last 50 years of industrial development, the amount and
variety of hazardous substances has drastically increased. It is estimated that the
human-made chemicals presently in use amount to 100.000, and hundreds of new
ones are produced every year. Due to the increase in industrial and agricultural
activities and exports of wastes, not only the traditional industrialized countries,
but all nations are confronted with widespread soil pollution. A significant
number of synthetic compounds persist in the environment, particularly those with
no relation to natural ones.
Essentially, there are three major categories of sites with polluted soils: (a)
sites that have been polluted by either spillage or leakage during production,
handling or use of industrial material (including mining and oil drilling); (b)
locations that have been used as disposal sites for diverse waste; (c) farmlands
that have been excessively exposed to pesticides.
Contaminated land sites are seriously dangerous for human beings and
therefore unsuitable for housing or agriculture. The downward migration of
pollutants from the soil into the groundwater is especially problematic in
developing countries, where groundwater is often directly drunk, without any
prior treatment.
The halocarbons, both halogenated compounds and solvents, are
widespread air, water, soil, and sediment pollutants; they are recalcitrant
molecules resistant to mineralization due to the stability of their carbon-halogen
bond. The stability and chemical inertness of many halogenated compounds is
part of their appeal in many industrial processes, but it also makes their
degradation extremely slow.
Halogenated compounds have been used for a variety of purposes for
hundreds of different industrial processes over the last 50 years, although they are
dangerous for human health and include known toxins and potential carcinogens
Introduction and Problem Definition
as dioxins, pesticides and PCBs. One prevalent example of a halogenated organic
compound is the widely used pesticide DDT, which has been shown to
bioaccumulate in animal fat tissue, disrupt hormone function, and damage
ecosystems. PCBs, polychlorinated biphenyls, are another type of halogenated
organic compound widely used for industrial applications as coolants, lubricants,
plasticizers, and dies. PCBs are toxic chemicals and their carcinogenicity has been
shown through laboratory studies. PCBs may also adversely affect human health
by contributing to neurological, immune system, reproductive system, and other
organs damage.
Among halogenated compounds, chlorinated solvents and their natural
transformation products are prevalent ground water organic contaminants in the
U.S. These solvents, consisting primarily of chlorinated aliphatic hydrocarbons
(CAHs), have been widely used in factory processing as fumigants, pesticides,
degreasing agents and solvents. They may pollute the environment through
accidental spills and leaks or illegal dumping. Their relative solubility in water
and their poor sorption, causes them to migrate downward through soils, thus
contaminating groundwater. Many CAHs and their transformation products are
defined as possible human carcinogens by the United States Environmental
Protection Agency (U.S. EPA).
A major problem with halogenated compounds is that they belong to a
class of molecules known as persistant organic pollutants, which tend to
biodegrade very slowly. It was originally thought that there were no natural
sources of halogenated compounds in the environment, hence no organisms had
evolved to exploit them. On the contrary, it has recently been shown that this
assumption was uncorrect (organisms as well as volcanic eruptions can produce
these compounds), and that natural production of chlorinated phenols may
actually be greater than anthropogenic sources. Since these compounds have in
fact existed for millions of years, there are naturally occurring strains of bacteria
which have evolved to break down halogenated compounds, thus opening up the
possibility for bioremediation treatment of contaminated sites.
2
Introduction and Problem Definition
Bioremediation principles
Biological cleaning procedures make use of the fact that most organic
chemicals are subject to enzymatic attack by living organisms. These activities are
summarized under the term biodegradation. However, the end products of these
enzymatic processes might differ drastically. For instance, an organic substance
might be mineralized (i.e. transformed to carbon dioxide and water), but it might
also be converted to a product that binds to natural materials in the soil, or to a
toxic substance.
Bioremediation refers to the productive use of micro-organisms to remove
or detoxify pollutants contaminating soils, water or sediments and threatening
public health. Bioremediation is not new: micro-organisms have been used to
remove organic matter and toxic chemicals from domestic and manufacturing
waste discharge for many years. Indeed, micro-organisms are frequently the only
means, biological or non-biological, to convert synthetic chemicals into inorganic
compounds. What is new is the emergence of bioremediation as an industry that is
driven by its particular usefulness for sites contaminated with petroleum
hydrocarbons.
Over the past decade, opportunities for applying bioremediation to a much
broader set of contaminants have been identified. Indigenous and enhanced
organisms have been shown to degrade industrial solvents, polychlorinated
biphenyls (PCBs), explosives, and many different agricultural chemicals. Pilot,
demonstration, and full-scale applications of bioremediation have been carried out
on a limited basis. Equally importantly, microorganisms that transform and
sequester heavy metals and radionuclides have been identified and employed, to a
limited extent, for in situ bioremediation. However, the full benefits of
bioremediation have not been realized, because processes and organisms effective
in controlled laboratory tests are not always equally effective in full-scale
applications. The failure to perform optimally in the field setting stems from a
lack of predictability due, in part, to inadequacies in the fundamental scientific
understanding of how and why these bioremediation processes work.
3
Introduction and Problem Definition
Content and objectives
Chlorinated aliphatic and aromatic hydrocarbons are among the most
common contaminants of soils, groundwaters and sediments, and most of them
are known or suspected carcinogen. Several studies showed that most of these
pollutant can be biodegraded by single bacterial strains or mixed microbial
populations via aerobic direct metabolism or cometabolism using aromatic and
aliphatic hydrocarbons such as methane, propane, butane, phenol or toluene as
growth substrates. In general, applications of in situ bioremediation can be
grouped into enhanced bioremediation and intrinsic bioremediation. In enhanced
bioremediation amendment such as oxygen, nutrients or even exogenous
microorganisms (bioaugmentation) are provided to manipulate the microbial
environment and facilitate biodegradation of contaminants. Conversely, intrinsic
bioremediation depends on indigenous microflora to degrade contaminants
without any amendments, exclusively relying on natural physical, chemical, and
biological processes to reduce or attenuate contaminant concentrations.
In this thesis, two studies have been carried out concerning different situations
where bioremediation processes of chlorinated hydrocarbons were involved. The
first one dealt with an enhanced bioremediation situation while the second was
related to intrinsic bioremediation.
1) The first experimental work (enhanced bioremediation) consisted in the study
of microbial consortia able to degrade a mixture of 6 CAHs (chlorinated
aliphatic hydrocarbons) via aerobic cometabolism. Several aspects of the longterm growth of these consortia were investigated. Biomass was grown in batch
bioreactors for 150 days and, during this period, the maintenance of the ability
to degrade the 6 CAHs mixture was tested. Furthermore the effectiveness of
these consortia as inocula for the bioaugmentation of different types of
aquifers was investigated. The reason why we were interested in the
characterization of the behavior of these consortia was their potential
usefulness as inocula in bioaugmentation treatment (in field scale or in pilot
scale).
2) The second study (intrinsic bioremediation) dealt with monochlorobenzene
biodegradation in the interface between groundwater and surface water.
4
Introduction and Problem Definition
It aimed at investigating the natural pollutant degradation capacity of the
aquifer zone representing this interface. The interface can be considered a
zone with changing redox conditions characterized by specific degradation
potential for pollutants passing through as a result of steep physico-chemical
gradients. Molecular techniques (PCR-DGGE) were also applied to
characterize the structure of the microbial community harboured in the
interface.
5
CHAPTER 1
1. VOLATILE ORGANIC COMPOUNDS
______________________________________________________________________________________________________________________________________________________________
1.1 INTRODUCTION
Within a physico-chemical context volatile organic compounds (VOCs)
are defined as “any chemical compound based on carbon chains or rings (and also
containing hydrogen) with a vapour pressure greater than 2 mm Hg at 25°C.
These compounds may contain oxygen, nitrogen and other elements. Substances
that are specifically excluded are: carbon dioxide, carbon monoxide, carbonic
acid, carbonate salts, metallic carbides and methane” (Australian Department of
Environment and Heritage, 2003). Within a regulatory context, USEPA provides
this definition (under the Clean Air Act, published in the Code of Federal
Regulation): “any compound of carbon, excluding carbon monoxide, carbon
dioxide, carbonic acid, metallic carbides or carbonates, and ammonium carbonate,
which participates in atmospheric photochemical reactions”. The IUPAC naming
convention identifies two classes of VOCs: aliphatic hydrocarbons, having an
open chain of carbon atoms (alkanes and alkenes) and aromatic hydrocarbons,
characterized by an alternating carbon-carbon single and double bonds arranged in
a ring structure. An alkane is a straight chain or cyclic (ring-like, such as
cycloalkane) structure that consists of carbon-carbon and carbon-hydrogen single
bonds. An alkene is typically a straight-chain structure that contains at least one
carbon-carbon double bond. These double bonds impart more stability to the
compound than the single bond in an alkane compound. A chlorinated alkane or
alkene also contains at least one chlorine-carbon single bond while chlorinated
aromatic compounds also contain one chlorine-carbon single bond (for example,
chlorobenzene). Aromatic compounds are typically more resistant to degradation
(more stable) than the alkane and alkene compounds.
Chapter 1 Volatile Organic Compounds
The aliphatic and the aromatic hydrocarbons are commonly sub grouped even
further based on the presence of attached halogen atoms (chlorine as chloro,
bromine as bromo, or fluorine as fluoro) or functional groups including, but not
limited to, alkyl radicals. The VOC subgroups include the alkyl benzenes (such as
methylbenzene), chlorinated alkanes (such as 1,2-dichloroethane), chlorinated
alkenes (such as 1,1-dichloroethene), and the chlorinated aromatics (such as 1,2dichlorobenzene).
Halogenated or alkylated aromatics such as chlorobenzene or toluene are more
easily degraded than benzene in aerobic and anaerobic ground water because the
stability of the benzene ring is reduced and the ring is weakened (Borden and
others, 1997). Adding halides or alkyl groups to the ring structure disperses the
electrical charges from the carbon-carbon bonds on the ring and weakens that
bond.
1.2 VOLATILE ORGANIC COMPOUNDS DETECTED IN GROUNDWATER
A relatively large amount of literature exists that describes VOCs in
groundwater at specific, known areas of contamination. Few documents, however,
describe VOC contamination in a regional or national context. One report by
Arneth and others (1989) lists the top 15 VOCs detected in ground water near
landfills in the United States and in Germany (table 1.1). This list shows that the
VOCs contaminating groundwater near landfills are similar in both countries.
Most of these VOCs are chlorinated solvents and gasoline compounds.
Furthermore, the frequency of VOCs detected in representative studies completed
on national, regional, and site-specific scales in the United States show a
remarkable similarity to those in table 1.1 (table 1.2; Delzer and Ivahnenko, 2003;
Moran, 2006; Zogorski and others, 2006). Although the number of VOCs
analysed in ground-water samples is large for national and regional studies, the
most commonly detected compounds, primarily chlorinated solvents and gasoline
compounds, are similar to those at site-specific studies completed at U.S.
Department of Defence installations (table 1.3).
7
Chapter 1 Volatile Organic Compounds
The ten most commonly detected VOCs in the studies summarized in tables 1.2
and 1.3 are methyl tert-butyl ether (MTBE), tetrachloroethene (PCE), 1,1,2trichloroethene (TCE), methylbenzene (toluene), 1,1,1-trichloroethane, (1,1,1TCA), benzene, cis-1,2-dichloroethene (1,2-cDCE), 1,1-dichloroethane (1,1DCA), trans-1,2-dichloroethene (1,2-tDCE), the dimethylbenzenes (m-, o-, pxylenes).
Table 1.1. Volatile organic compounds ranked by those frequently detected in groundwater
near landfills and hazardous waste dumps in the United States and the Federal Republic of
Germany1.
8
Chapter 1 Volatile Organic Compounds
Table 1.2. Volatile organic compounds detected in regional and national groundwater studies in the United States.
9
Chapter 1 Volatile Organic Compounds
Table 1.3. Volatile organic compounds detected in groundwater case studies at selected U.S.
Department of Defence installations.
1.3 CHLORINATED ALIPHATIC HYDROCARBONS (CAHs)
1.3.1 Sources of CAHs
CAHs are manmade organic compounds, typically manufactured from
methane, ethane, ethene and chlorine through various processes that substitute one
or more hydrogen atoms with a chlorine atom, or selectively dechlorinate
chlorinated compounds to a less chlorinated state.
CAHs are typically used in the manufacturing of industrial, chemical,
electronic, and consumer goods (Smith and others, 1988; U.S. Environmental
Protection Agency, 2005b). In addition, these compounds are heavily used as
solvents in cleaning and degreasing products. For example, 1,1,1-TCA is used as a
solvent for adhesives and in metal degreasing, pesticides, textile processing,
cutting fluids, aerosols, lubricants, cutting oil formulations, drain cleaners, shoe
polishes, spot cleaners, printing inks, and stain repellents. Carbon tetrachloride
10
Chapter 1 Volatile Organic Compounds
(CTET) was used as feedstock for the production of chlorofluorocarbon gases,
such as dichlorodifluoromethane (F-12) and trichlorofluoromethane (F-11), which
were used as aerosol propellants in the 1950s and 1960s (Holbrook, 1992). During
1974, the U.S. Food and Drug Administration (FDA) banned the sale of CTET in
any product used in the home and the USEPA regulated the use of
chlorofluorocarbon gases as aerosols or propellants. By 2000, CTET production
for no feedstock purposes was phased-out completely.
Chemical manufacturing is the largest use of 1,1-DCA and 1,2dichloroethane (1,2-DCA). Both compounds serve as an intermediate during the
manufacture of chloroethene (vinyl chloride, VC), 1,1,1-TCA, and to a lesser
extent high-vacuum rubber. Both DCA isomers also are used as a solvent for
plastics, oils, and fats, and in cleaning agents and degreasers (Agency for Toxic
Substances and Disease Registry, 1990c, p. 51; 2001, p.160). About 98% of the
1,2-DCA produced in the United States is used to manufacture VC. Smaller
amounts of 1,2-DCA are used in the synthesis of vinylidene chloride, TCE, PCE,
aziridines, and ethylene diamines, and in other chlorinated solvents (U.S.
Environmental ProtectionAgency, 1995).
1,1,1-TCA was initially developed as a safer solvent to replace other
chlorinated and flammable solvents. The compound is used as a solvent for
adhesives (including food packaging adhesives) and in metal degreasing,
pesticides, textile processing, cutting fluids, aerosols, lubricants, cutting
formulations, drain cleaners, shoe polishes, spot cleaners, printing inks, and stain
repellents, among other uses (Agency for Toxic Substances and Disease Registry,
2004, p. 181). The other TCA isomer, 1,1,2-trichloroethane (1,1,2-TCA), has
limited use as a common, general-use solvent but is used in the production of
chlorinated rubbers (Archer, 1979). In some cases, 1,1,2-TCA may be sold for use
in consumer products (Agency for Toxic Substances and Disease Registry, 1989,
p.59).
Before 1979, the single largest use of chloroethane was in the production
of tetraethyl lead. As recently as 1984, the domestic production of tetraethyl lead
accounted for about 80 percent of the chloroethane consumed in the United States,
whereas about 20 percent was used to produce ethyl cellulose, and used in
11
Chapter 1 Volatile Organic Compounds
solvents, refrigerants, topical anaesthetics, and in the manufacture of dyes,
chemicals, and pharmaceuticals. Since the 1979 ban on tetraethyl lead in gasoline
and its subsequent phase out in the mid-1980, the production of chloroethane in
recent years has declined substantially in the United States (Agency for Toxic
Substances and Disease Registry, 1998, p. 95).
Among the chloroethenes, PCE and TCE are two of the most
widely used and distributed solvents in the United States and Europe. The textile
industry uses the largest amount of PCE during the processing, finishing of raw
and finished textiles, and for industrial and consumer dry cleaning (U.S.
Environmental
Protection
Agency,
2005b,
Web
page:
http://www.epa.gov/opptintr/chemfact/f_perchl.txt, accessed May 23, 2006). Most
of the TCE used in the United States is for vapour degreasing of metal parts and
some textiles (U.S. Environmental Protection Agency, 2005b, Web page:
http://www.epa.gov/OGWDW/dwh/t-voc/trichlor.html, accessed May 23, 2006).
Other uses of PCE and TCE include manufacturing of pharmaceuticals, other
organic compounds, and electronic components, and in paint and ink formulations
(Smith and others, 1988).
Historical management of wastes containing CAHs has resulted in
contamination of soil and groundwater, with CAHs present at many contaminated
groundwater sites in the United States. TCE and PCE are the most prevalent of
those contaminants (U.S. Air Force 1998). In addition, CAHs and their
degradation products, including dichloroethane (DCA), dichloroethene (DCE),
and vinyl chloride (VC) tend to persist in the subsurface. Table 1.4 lists the CAHs
more commonly identified as environmental contaminants, their abbreviations,
their common names, and the types of waste from which they commonly
originate. Figure 1.1 presents the molecular structure of those CAHs.
12
Chapter 1 Volatile Organic Compounds
Table 1.4. CAHs commonly identified as environmental contaminants
13
Chapter 1 Volatile Organic Compounds
Chlorinated Ethenes
Chlorinated Ethanes
Chlorinated Methanes
Figure 1.1. Molecular structures of common CAHs
14
Chapter 1 Volatile Organic Compounds
1.3.2 Physical and chemical properties
The physical and chemical properties of CAHs govern their fate and
transport in the subsurface environment; the number of chlorine atoms directly
affects the physical and chemical behaviour of the compound: as the number of
substituted chlorine atoms increases, molecular weight and density generally
increases, and vapour pressure and aqueous solubility generally decreases. In table
1.5 the major physical and chemical data for the CAHs commonly identified as
subsurface contaminants are listed.
1.3.3 Transport processes
The extent of the contaminant spreading into the environment is affected
by the physical and chemical properties of the compound (in particular solubility,
volatility and density), besides the specific characteristics of the site.
A CAH released to the subsurface as a pure organic liquid (NAPL – Non
Aqueous Phase Liquid) will reach phase equilibrium, and it will remain as a
NAPL , adsorb to soil, dissolve in groundwater, or volatilise to soil gas to the
extent defined by the physical and chemical properties of the individual CAH and
the subsurface environment. Figure 1.2 shows the mechanisms by which CAHs
transfer phases in the attempt to reach equilibrium conditions.
Figure 1.2. Phase equilibrium mechanisms and defining properties of CAHs
15
Chapter 1 Volatile Organic Compounds
In particular it can be observed that:
•
partition coefficient (related to the hydrophobicity and the solubility)
define the extent to which a CAH will partition between NAPL and soil,
and NAPL adsorbed to soil and the groundwater;
•
the aqueous solubility of a CAH defines the equilibrium between NAPL
and groundwater;
•
CAHs dissolved in the groundwater will partition themselves between the
dissolved phase and the vapour phase, as defined by the Henry’s constant;
•
vapour pressure describes the equilibrium between NAPL or NAPL
adsorbed to soil and the soil gas.
Most of the CAHs are denser than water (referred to as dense non-aqueous phase
liquids - DNAPLs) and tend to sink through both unsaturated and saturated
permeable soils until they reach the impermeable layer at the bottom of the
aquifer. Those kinds of free phases are located in the deepest layers and are
therefore extremely difficult to identify and locate, thus making both biological
and physico-chemical remediation more difficult. Moreover the dense nonaqueous phase (NAPL o DNAPL) acts as a local polluting source gradually
releasing the contaminant in the aquifer. The contaminant concentrations observed
next to NAPL (next to water solubility) could then be toxic for microorganisms,
thus making a biological intervention more difficult or even impossible.
In addition to transferring phases in an attempt to reach equilibrium
conditions, CAHs can migrate in the subsurface in their non-aqueous, aqueous and
vapour phase by both active and passive processes. In active processes such
advection and dispersion , CAHs migrate along with the flow of the groundwater
or soil gas to which they are partitioned. Passive processes, such as diffusion, are
the result of concentration gradients, which cause the CAH to seek phase and
concentration equilibrium with its surrounding environment. Typically, releases of
CAH to the groundwater result in the formation of a plume and advection is one of
the most important processes affecting the transport of the contaminants.
16
Chapter 1 Volatile Organic Compounds
Table 1.5. Chemical and physical properties of CAHs.
17
Chapter 1 Volatile Organic Compounds
1.3.4 Transformation of CAHs
In the natural environment CAHs may undergo chemical and biological
transformations. The chlorinated alkanes can be degraded by abiotic processes
through hydrolysis or dehydrohalogenation (with no external transfer of electrons)
or by biotic processes through reductive dechlorination or (direct and
cometabolic) aerobic oxidation; (oxidation-reduction reactions, requiring an
external transfer of electrons). These degradation processes can proceed under
either aerobic or anaerobic conditions (Vogel and McCarty, 1987a; Vogel, 1994).
According to McCarty (1997), 1,1,1-TCA is the only chlorinated compound that
can be degraded in groundwater within 20 years under all likely groundwater or
aquifer conditions. Oxidation-reduction reactions are the dominant mechanisms
driving VOC degradation and most of these reactions are catalyzed by
microorganisms (Wiedemeier and others, 1998; Azadpour-Keeley and others,
1999). Substitution reactions that can remove chlorine atoms, such as hydrolysis,
can degrade some chlorinated alkanes (trichloroethane) to nonchlorinated alkanes
(ethane) with or without a microbial population catalyzing the reaction (Vogel and
others, 1987; Olaniran and others, 2004).
While aerobic oxidation and anaerobic reductive dechlorination can occur
naturally under proper conditions, enhancements such as the addition of electron
donors, electron acceptors, or nutrients help to provide the proper conditions for
aerobic oxidation or anaerobic reductive dechlorination to occur. In general,
highly chlorinated CAHs degrade primarily through reductive reactions, while
less chlorinated compounds degrade primarily through oxidation (Vogel and
others 1987b). Highly chlorinated CAHs are reduced relatively easily because
their carbon atoms are highly oxidized. During direct reactions, the
microorganism causing the reaction gains energy or grows as the CAH is
degraded or oxidized. During cometabolic reactions, the CAH degradation or
oxidation is caused by an enzyme or cofactor produced during microbial
metabolism of another compound. CAH degradation or oxidation does not yield
any energy or growth benefit for the microorganism mediating the cometabolic
reaction. The degradation mechanisms that typically occur in the degradation of
18
Chapter 1 Volatile Organic Compounds
each CAH are summarized in table1.6 while table 1.7 shows biological
degradation mechanisms.
Table 1.6. Common abiotic and biotic reactions involving halogenated aliphatic hydrocarbons.
19
Chapter 1 Volatile Organic Compounds
Table 1.7. Chemical and physical properties of CAHs.
1.3.4.1 Abiotic degradation processes
The abiotic processes occurring most frequently under either aerobic or anaerobic
conditions are hydrolysis and dehydrohalogenation. Abiotic transformations
generally result only in a partial transformation of the compounds that are either
more readily or less readily biodegraded by microorganisms.
Hydrolysis and dehydrohalogenation are two abiotic processes that may
degrade chlorinated ethanes under either aerobic or anaerobic conditions. The
tendency for a chlorinated ethane to degrade by hydrolysis depends on the ratio of
chlorine to carbon atoms (figure 1.3) or the location of chlorine atoms on the
number 2 carbon in the compound. Chlorinated alkanes are more easily
hydrolyzed when the chlorine-carbon ratio is less than two or when chlorine atoms
are only located on the number 1 carbon atom (Vogel and McCarty, 1987b; Vogel,
20
Chapter 1 Volatile Organic Compounds
1994). For example, chloroethane and 1,1,1-TCA have half-lifes that are measured
in days or months (Vogel and others, 1987; Vogel, 1994; table 1.8). Conversely,
the more chlorinated ethanes such as 1,1,1,2-tetrachloroethane (PCA) and those
with chlorine atoms on the number 2 carbon tend to have half-lifes measured in
decades or centuries (table 1.8). Dehydrohalogenation is the removal of one or two
halogen
atoms
from
an
alkane
(Vogel
and
McCarty,
1987a).
The
dehydrohalogenation of two chlorine atoms is called dichloroelimination.
Figure 1.3. Relation between degree of chlorination and anaerobic reductive-dechlorination,
aerobic degradation and sorption onto subsurface material. Degree of chlorination is number
of chloride atoms divided by number of carbon atoms.
Table 1.8. Laboratory half-lives and by-products of the abiotic degradation (hydrolysis or
dehydrohalogenation) of chlorinated alkane compounds detected in groundwater
21
Chapter 1 Volatile Organic Compounds
Chen and others (1996) show that PCA can be abiotically transformed to
TCE under methanogenic conditions (figure 1.4). In addition, the abiotic
degradation of 1,1,1-TCA has been well studied in the scientific literature (figure
1.5; Jeffers and others, 1989; McCarty and Reinhard, 1993; Chen and others,
1996; McCarty, 1997). McCarty and Reinhard (1993) indicate that the
transformation of 111-TCA by hydrolysis is about four times faster than by
dehydrochlorination. During abiotic degradation, about 80 percent of 1,1,1-TCA is
transformed to acetic acid by hydrolysis (McCarty, 1997), and the remaining 20
percent is transformed to 1,1-DCE by dehydrochlorination (Vogel and McCarty,
1987b; McCarty, 1997). The presence of 1,1-DCE in contaminated groundwater is
probably the result of the dehydrochlorination of 1,1,1-TCA (McCarty, 1997).
Figure 1.4. Laboratory-derived pathway for the abiotic degradation, anaerobic and
methanogenic biodegradation of 1,1,2,2-tetrachloroethane; 1,1,2-trichloroethane; and 1,1,2trichloroethane.
22
Chapter 1 Volatile Organic Compounds
Figure 1.5. Laboratory-derived pathway for the abiotic degradation, anaerobic and
methanogenic biodegradation of 1,1,1-trichloroethane.
23
Chapter 1 Volatile Organic Compounds
1.3.4.2 Biotic degradation processes
Bacteria transform environmentally available nutrients to forms that are
useful for incorporation into cells and synthesis of cell polymers. Biotic
transformations occur through reactions involving a transfer of electrons between
the chlorinated solvents and an external agent; energy is made available when an
electron donor transfers its electrons to a terminal electron acceptor. The energy
gained is stored as high energy compounds, such as ATP and low-energy
compounds, such as nicotinamide adenine dinucleotide (NAD). A portion of the
stored energy is used to conduct to biological processes necessary for cell
maintenance and reproduction. In addition, cell building-block materials are
required in the form of carbon and other nutrients (such nitrogen and phosphorus).
The terminal electron acceptor used during metabolism is important for
establishing the redox conditions, and therefore the type of zone that will
dominate in the subsurface. Common terminal electron acceptors include oxygen
under aerobic conditions and nitrate, Mn(IV), Fe(III), sulphate and carbon dioxide
under anaerobic conditions.
The typical electron-acceptor classes of bacteria are listed in table 1.9 in
the order of those causing the largest energy generation during the redox reaction
to those causing the smallest energy generation during the redox reaction. A
bacteria electron acceptor class causing a redox reaction generating relatively
more energy will dominate over a bacteria electron acceptor class causing a redox
reaction generating relatively less energy (Table 1.9). Aerobic biotic
transformations generally are oxidations: they are classified as “hydroxilations”,
in the case of a substitution of a hydroxyl group on the molecule, or
“epoxidations”, in the case of unsaturated CAHs. The anaerobic biotic processes
generally are reductions that involve either hydrogenolysis, the substitution of a
hydrogen atom or chlorine on the molecule, or dehaloelimination, where two
adiacent chlorine atoms are removed, leaving a double bond between the
respective carbon atoms.
24
Chapter 1 Volatile Organic Compounds
Table 1.9. Typical electron-acceptor classes of bacteria
1.3.4.2.1 Anaerobic biodegradation
Organic compounds can be transformed by microorganisms through two
basically different processes: direct metabolism and cometabolism. In the first
process the organism consumes the organic compound as a primary substrate to
satisfy its energy and carbon needs. The compound serves as an electron donor
and as a primary growth substrate for the microbe mediating the reaction.
Electrons that are generated by the oxidation of the compound are transferred to
an electron acceptor such as oxygen. In addition a microorganism can obtain
energy for cell maintenance and growth from the oxidized compound. In general
only the less chlorinated CAHs (with one or two chlorine atoms) can be used
directly by microorganism as electron donors. The CAHs are oxidized into carbon
25
Chapter 1 Volatile Organic Compounds
dioxide, water, chlorine and electrons, in conjunction with the reduction of
oxygen to water.
Few CAHs have been shown to serve as primary substrates for energy and
growth. Pure cultures have been isolated that can grow aerobically on
dichloromethane (DM) as sole carbone and energy source. VC and 1,2-DCA have
also been shown to be available as primary substrates under aerobic conditions
(Hartmans et al, 1992, Verce et al, 2000, Klier et al, 1998). Other CAH that can
oxidized directly include DCE, DCA, CA, MC and CM (Bradley 1998; RTDF
1997; Harknessvand others 1999). These few exceptions suggest that only the less
halogenated one- and two-carbon CAHs might be used as primary substrates, and
that the organisms that are capable of doing this are not necessarily widespread in
the environment. Figure 1.6.shows an example of aerobic oxidation of a CAH.
Figure 1.6. Aerobic oxidation (direct).
Most of the CAHs can be biologically transformed by the process of
cometabolism. Cometabolism is the fortuitous transformation of an organic
compound by non specific enzymes, produced for other purposes during microbial
metabolism of another compound. Most of the enzymes involved in the
degradation of chlorinated solvents determine a sequence of reactions that oxidize
NADH to NAD+, but do not catalyse the opposite reduction process of NAD+ to
NADH. Thus, in the cometabolic transformation the microorganisms do not get
energy or carbon from the process; The transformation does not provide the
26
Chapter 1 Volatile Organic Compounds
organisms any direct benefit, indeed it may be harmful to them, resulting in
increased maintenance requirements and decay rates (Criddle, 1992). A primary
growth substrate must be at least intermittently available to prevent the depletion
of energy and maintain a viable microbial population.
The CAHs that have been observed to be oxidized cometabolically under
aerobic conditions include TCE, DCE, VC, TCA, DCA, CF and MC (MunakataMarr 1997; McCarty and others 1998; RTDF 1997; Edwards and Cox 1997;
McCarty 1997a; Bradley and Chapelle 1998; Travis and Rosenberg 1997). The
electron donors observed in aerobic cometabolic oxidation include methane,
ethane, ethene, propane, butane, aromatic hydrocarbons (such as toluene and
phenol), and ammonia Under aerobic conditions a moooxygense enzyme mediates
the electron donation reaction. That reaction has the tendency to convert CAHs
into unstable epoxides (Anderson and Lovley 1997). Unstable epoxides degrade
rapidly in water to alcohols and fatty acids, which are readily degradable. Figure
1.7 shows an example of aerobic cometabolic oxidation of a CAH.
Figure 1.7. Aerobic oxidation (cometabolic).
Aerobic biodegradation of chlorinated alkanes. According to the degradation
pathway constructed by Sands and others (2005) and Whittaker and others (2005),
the dichloroethanes are not a by-product of 1,1,1-TCA or 1,1,2-TCA
biodegradation under aerobic conditions (figure 1.5). Apparently, the only source
of 1,1-DCA and 1,2-DCA via a degradation pathway is the reductive
27
Chapter 1 Volatile Organic Compounds
dechlorination of 1,1,1-TCA and 1,1,2-TCA, respectively, under anaerobic
conditions (figures 1.4 and 1.5). Under aerobic conditions, however, 1,2-DCA can
be degraded when used as a carbon source by microorganisms. The intermediate
by-product of this degradation is chloroethanol, which is then mineralized to
carbon dioxide and water (figure 1.8; Stucki and others, 1983; Janssen and others,
1985; Kim and others, 2000; Hage and others, 2001).
Figure 1.8. Laboratory-derived pathway for the aerobic biodegradation of 1,2dichloroethane.
Aerobic biodegradation of chlorinated alkenes. Several studies have shown that
chlorinated ethenes, with the exception of PCE, can degrade under aerobic
conditions by oxidation (Hartmans and De Bont, 1992; Klier and others, 1999;
Hopkins and McCarty, 1995; Coleman and others, 2002) and by co-metabolic
processes (Murray and Richardson, 1993; Vogel, 1994; McCarty and Semprini,
1994). Studies describing the degradation of PCE under aerobic conditions were
not found in the peer-reviewed literature. In one study, aerobic biodegradation of
28
Chapter 1 Volatile Organic Compounds
PCE was not measurable beyond analytical precision after 700 days of incubation
(Roberts and others, 1986). Furthermore, Aronson and others (1999) indicate that
PCE is not degraded when dissolved oxygen (DO) is greater than 1.5 mg/L, the
approximate boundary between aerobic and anaerobic conditions (Stumm and
Morgan, 1996). Chen and others (1996) suggest the structure and oxidative state
of PCE prevents its aerobic degradation in water.
According to the aerobic biodegradation pathway constructed by
Whittaker and others (2005), the dichloroethenes are not a by-product of TCE
degradation under aerobic conditions (figure. 1.5). Rather, TCE is degraded along
three different pathways by different microorganisms (figure. 1.9). These
pathways do not form any of the dichlorothene compounds and the only apparent
source of 1,2-DCE is by the reductive dechlorination of TCE under anaerobic
conditions (figures. 1.4 and 1.12). The compounds 1,2-DCE and VC, however,
can be degraded under aerobic conditions by microorganisms utilizing the
compounds as a primary carbon source (figure. 1.8; Bradley and Chapelle, 1998).
Although PCE is not known to degrade through cometabolism under aerobic
conditions, co-metabolism is known to degrade TCE, the dichloroethenes, and
VC. The rate of cometabolism increases as the degree of chlorination decreases on
the ethene molecule (Vogel, 1994). During aerobic cometabolism, the chlorinated
alkene is indirectly dechlorinated by oxygenase enzymes produced when
microorganisms use other compounds, such as BTEX compounds, as a carbon
source (Wiedemeier and others, 1998). The co-metabolic degradation of TCE,
however, tends to be limited to low concentrations of TCE because high
concentrations in the milligram per litre range are toxic to microbes catalyzing
this reaction (Wiedemeier and others, 1998). In field studies by Hopkins and
McCarty (1995), VC is shown to degrade by co-metabolism under aerobic
conditions when phenol and toluene were used as a carbon source.
29
Chapter 1 Volatile Organic Compounds
Figure 1.9. Laboratory-derived pathway for the aerobic biodegradation of trichloroethane.
1.3.4.2.2 Anaerobic biodegradation
Under anaerobic conditions, reductive dechlorination mechanisms can
effectively biodegrade CAHs. This process generally involves a series of
decarboxylations and oxidation-reduction (redox) reactions catalyzed either by
single microorganisms or by a consortium of microorganisms (Dolfing, 2000). In
direct anaerobic reductive dechlorination the mediating bacteria use the CAH
directly as an electron acceptor in energy-producing redox reactions.
Cometabolic anaerobic reductive dechlorination occurs when bacteria
30
Chapter 1 Volatile Organic Compounds
incidentally dechlorinate a CAH in the process of using another electron acceptor
to generate energy.
Theoretically, reductive dechlorination is the sequential replacement of
one chlorine atom on a chlorinated compound with a hydrogen atom. The
replacement continues until the compound is fully dechlorinated. For example,
PCE can undergo reductive dechlorination to less-chlorinated compounds, such as
TCE or 1,2-DCE, or to nonchlorinated compounds such as ethene, ethane, or
methane (methanogenesis). Each successive step in the dechlorination process is
theoretically slower than the preceding step. The dechlorination process slows
because as chlorines are removed the energy costs to remove another chlorine
atom increases (free energy of the reaction decreases; Dolfing, 2000). As a result,
biodegradation may not proceed to completion in some aquifers leaving
intermediate compounds (for example, dichloroethenes and vinyl chloride) to
accumulate in ground water (Azadpour-Keeley and others, 1999). Other
constraints on biodegradation such as a reduction in or loss of primary substrate,
or microbial suppression also can play a role in the accumulation of intermediate
compounds. This is a particular concern with VC because it is a known human
carcinogen (Agency for Toxic Substances and Disease Registry, 2005) and its
accumulation may create a health issue that might not be a concern during the
early stages of groundwater contaminated by TCE.
Reductive dechlorination theoretically is expected to occur under most
anaerobic conditions, but has been observed to be most effective under sulfatereducing and methanogenic conditions (EPA 1998). As in the case of aerobic
oxidation, the direct mechanism may biodegrade CAHs faster than cometabolic
mechanism (McCarthy and Semprini, 1994).
In direct anaerobic reductive dechlorination bacteria gain energy and grow
as one or more chlorine atoms on a chlorinated hydrocarbon are replaced with
hydrogen. In that reaction, the chlorinated compound serves as electron acceptor,
and hydrogen as electron donor (Fennel and others 1997). Hydrogen used in the
reaction typically is supplied indirectly through the fermentation of organic
substrates (lactate, acetate, methanol, glucose, toluene). The reaction is also
referred to halorespiration or dehalorespiration (Gosset and Zinder 1997). Direct
31
Chapter 1 Volatile Organic Compounds
anaerobic reductive dechlorination has been observed in anaerobic systems in
which PCE, TCE, DCE, VC and DCA are used directly by a microorganism as an
electron-acceptor in their energy-producing redox reactions. The mechanism
generally results in the sequentiql reduction of a chlorinated ethene or chlorinated
ethane to ethene or ethane. Figure 1.10 shows the step-by-step dechlorination of
PCE.
Figure 1.10. Anaerobic reductive dechlorination of PCE.
Several CAHs have been observed to be reductively dechlorinated by
cometabolic mecanisms. In those instances, the enzymes that are intended to
mediate the electron-accepting reaction “accidentally” reduce and dehalogenate
the CAH. Cometabolic anaerobic reductive dechlorination has been observed for
PCE, TCE, DCE, VC, DCA and CT under anaerobic conditions (Fathepure 1987;
Workman 1997; Yager and others 1997).
Anaerobic biodegradation of chlorinated alkanes. While researching the
scientific literature for their report, Wiedemeier and others (1998) did not find
published studies describing anaerobic biodegradation of chlorinated ethanes in
ground water. Since the publication of Wiedemeier and others (1998), however,
numerous published studies describe the anaerobic biodegradation of chlorinated
ethanes. McCarty (1997) indicates that carbon tetrachloride was transformed to
chloroform under denitrifying conditions and mineralized to carbon dioxide and
water under sulfate-reducing conditions (figure 1.11). Adamson and Parkin (1999)
show that under anaerobic conditions, carbon tetrachloride and 1,1,1-TCA tend to
inhibit the degradation of each other. Adamson and Parkin (1999) also show that
32
Chapter 1 Volatile Organic Compounds
carbon tetrachloride was rapidly degraded by cometabolism when acetate was the
carbon source.
Chen and others (1996) describe how methanogenic conditions in a
municipal sludge digester allowed the degradation of TeCA to 1,1,2-TCA, and
1,1,2-TCA to 1,2-DCA through dehydrohalogenation (figure 1.4). De Best and
others (1999) report that cometabolic transformations of 1,1,2-TCA will occur
under methanogenic conditions. In this study, 1,1,2-TCA was degraded to
chloroethane when sufficient amounts of the carbon source were present (figure
1.4). This transformation was inhibited by the presence of nitrate, but not nitrite.
Dolfing (2000) discusses the thermodynamics of reductive dechlorination
during the degradation of chlorinated hydrocarbons and suggests that fermentation
of chloroethanes to ethane or acetate may be energetically more favorable than
“classic” dechlorination reactions. Moreover, polychlorinated ethanes may
degrade preferentially by reductive dechlorination under strongly reducing
conditions. Dichloroelimination, however, may actually be the dominant
degradation reaction for polychlorinated ethanes because more energy is available
to microorganisms than is available during reductive dechlorination (Dolfing,
2000). During anaerobic biodegradation, the mean half-lifes of the chloroethane
compounds can be as short as three days, in the case of 1,1,1-TCA, or as long as
165 days, in the case of 12-DCA (table 1.10).
Table 1.10. Mean half-life in days for the anaerobic biodegradation of selected chlorinated
alkane and alkene compounds.
33
Chapter 1 Volatile Organic Compounds
Figure 1.11. Laboratory-derived
tetrachloromethane.
pathway
for
the
anaerobic
biodegradation
of
Anaerobic biodegradation of chlorinated alkenes. Many laboratory and field
studies have shown that microorganisms degrade chlorinated ethenes under
anaerobic conditions (Bouwer and others, 1981; Bouwer, 1994, Dolfing, 2000).
Groundwater is considered anoxic when the dissolved oxygen concentration falls
below 1.0–1.5 mg/L (Stumm and Morgan, 1996; Christensen and others, 2000).
Under anoxic conditions, anaerobic or facultative microbes will use nitrate as an
electron acceptor, followed by iron (III), then sulphate, and finally carbon dioxide
(methanogenesis; Chapelle and others, 1995; Wiedemeier and others (1998). As
the concentration of each electron acceptor sequentially decreases, the redox
34
Chapter 1 Volatile Organic Compounds
potential of the ground water becomes greater (more negative) and biodegradation
by reductive dechlorination is favoured.
Anaerobic conditions in ground water can be determined by measuring the
vertical and spatial concentrations of oxygen, iron (II), manganese (II), hydrogen
sulfide or methane in groundwater and using that data as a qualitative guide to the
redox status (Stumm and Morgan, 1996; Christensen and others, 2000). Other
measurements of anaerobic conditions involving microorganism biomarkers
include volatile fatty acids, ester-linked phospholipid fatty acid (PLFA),
deoxyribonucleic acid (DNA), and ribonucleic acid (RNA) probes, and TEAP
bioassay (Christensen and others, 2000). The reduction of iron (III) to iron (II),
manganese (IV) to manganese (II), sulfate to hydrogen sulfide, and carbon
dioxide to methane during the microbial reduction of CAHs can have a major
influence on the distribution of iron (II), manganese (II), hydrogen sulfide, and
methane concentrations in ground water (Stumm and Morgan, 1996; Lovley,
1991; Higgo and others, 1996; Braun, 2004).
The highly chlorinated alkenes are commonly used as electron acceptors
during anaerobic biodegradation and are reduced in the process (Vogel and others,
1987). The primary anaerobic process driving degradation of CAHs, except VC, is
reductive dechlorination (figures 1.4 and 1.12; Bouwer and others, 1981; Bouwer,
1994). Tetrachloroethene and TCE are the most susceptible to reductive
dechlorination because they are the most oxidized of the chlorinated ethenes;
however, the more reduced (least oxidized) degradation by-products such as the
dichloroethenes and vinyl chloride are less prone to reductive dechlorination. The
main by-product of anaerobic biodegradation of the polychlorinated ethenes is VC
(figure 1.12), which is more toxic than any of the parent compounds (Agency for
Toxic Substances and Disease Registry, 2004). The rate of reductive
dechlorination tends to decrease as the reductive dechlorination of daughter
products proceeds (Vogel and McCarty, 1985; Bouwer, 1994). Murray and
Richardson (1993) suggest that the inverse relation between the degree of
chlorination and the rate of reductive dechlorination may explain the
accumulation of 1,2-DCE and VC in anoxic groundwater contaminated with PCE
and TCE. In addition, the anaerobic reduction of VC to ethene is slow and
35
Chapter 1 Volatile Organic Compounds
inefficient under weak reducing conditions, which favours the persistence of VC
in anoxic groundwater (Freedman and Gossett, 1989).
Reductive dechlorination has been demonstrated under nitrate- and
iron-reducing
conditions
(Wiedemeier
and
others,
1998).
Reductive
dechlorination of the CAHs, however, may be more rapid and more efficient when
oxidation-reduction (redox) conditions are below nitrate-reducing levels
(Azadpour- Keeley and others, 1999). Sulfate-reducing and methanogenic
groundwater conditions create an environment that facilitates not only
biodegradation for the greatest number of CAHs, but also more rapid
biodegradation rates (Bouwer, 1994). Reductive dechlorination of DCE and VC is
most apparent under sulfate reducing and methanogenic conditions (Wiedemeier
and others, 1998). Anaerobic biodegradation rates for the chlorinated alkenes can
be as short as 45 minutes, in the case of VC, to as long as 9 years for PCE (table
1.10).
36
Chapter 1 Volatile Organic Compounds
Figure 1.12. Laboratory-derived
tetrachloroethene.
pathway
for
the
anaerobic
biodegradation
commonly
detected
in
of
1.4 CHLORINATED BENZENES
1.4.1 Sources of chlorinated benzenes
Four
chlorinated
benzenes
groundwater
contamination studies include chlorobenzene (CB), 1,2-dichlorobenzene (1,2DCB), and two isomers of trichlorobenzene, 1,2,3-trichlorobenzene (1,2,3-TCB)
and 1,2,4-trichlorobenzene (1,2,4-TCB; tables 1.2 and 1.3). Chlorobenzene is
commonly used as a solvent for pesticide formulations, in the manufacturing of
di-isocyanate, as a degreaser for automobile parts, and in the production of
nitrochlorobenzene. Solvent uses accounted for about 37 percent of chlorobenzene
consumption in the United States during 1981 (Agency for Toxic Substances and
Disease Registry, 1990a, p. 45). The compound 1,2-DCB is used primarily to
produce 3,4-dichloroaniline herbicides (Agency for Toxic Substances and Disease
Registry, 1990b, p. 263). The two trichlorobenzene isomers are primarily used as
dye carriers in the textile industry. Other uses include septic tank and drain
cleaners, the production of herbicides and higher chlorinated benzenes, as wood
preservatives, and in heat-transfer liquids (U.S. Environmental Protection Agency,
2005b).
1.4.2 Biodegradation of chlorinated benzenes
Several studies have shown that chlorinated benzene compounds
containing up to four chlorine atoms can be degraded by microorganisms under
aerobic conditions (Reineke and Knackmuss, 1984; Spain and Nishino, 1987;
Sander and others, 1991). Under aerobic conditions, 1,2,4-trichlorobenzene
(1,2,4-TCB; Haigler and others, 1988) and chlorobenzene (CB; Sander and others,
1991) are used as a primary carbon source during biodegradation by
microorganisms such as Burkholderia and Rhodococcus species (Rapp and
Gabriel-Jürgens, 2003). During biodegradation, these compounds are completely
mineralized to carbon dioxide (CO2) (van der Meer and others, 1991). Rapp and
Gabriel-Jürgens (2003) also indicate that all of the dichlorobenzene isomers were
37
Chapter 1 Volatile Organic Compounds
biodegraded by the Rhodococcus bacterium. The biodegradation pathways for
1,2,4-TCB, 1,4-DCB, 1,2-DCB, and CB, under aerobic conditions are shown in
figures 1.13 to 1.15, respectively. These pathways are similar to that of benzene,
except that one chlorine atom is eventually eliminated through hydroxylation of
the chlorinated benzene to form a chlorocatechol, then ortho cleavage of the
benzene ring (Van der Meer and others, 1998).
Calculated and published degradation half-lives for the chlorobenzenes
under aerobic conditions are shown in table 1.11. The compounds 1,2,4-TCB, 1,2DCB, and CB lose 50 percent of their initial mass within 180 days (table 1.11).
Conversely, Dermietzel and Vieth (2002) show that chlorobenzene was rapidly
mineralised to CO2 in laboratory and in situ microcosm studies, with complete
mineralisation ranging from 8 hours to about 17 days. In addition, the compound
1,4-DCB was completely mineralised within 25 days. Nevertheless, under the
aerobic conditions of Dermietzel and Vieth (2002) study, 1,2,4-TCB, 1,2-DCB,
and 1,3-DCB were only partially degraded after 25 days. In another laboratorymicrocosm study by Monferran and others (2005), all isomers of DCB were
mineralised to CO2 within 2 days by the aerobe Acidovorax avenae.
Although Wiedemeir and others (1998) indicate that few studies existed
that described the anaerobic degradation of the chlorobenzene compounds, a study
by Ramanand and others (1993) did suggest that 1,2,4-TCB could be biodegraded
to chlorobenzene with 1,4-DCB as an intermediate compound under anaerobic
conditions. Moreover, Middeldorp and others (1997) show that 1,2,4-TCB was
reductively dechlorinated to 1,4-DCB, then to chlorobenzene in a methanogenic
laboratory microcosm in which chlorobenzene-contaminated sediment was
enriched with lactate, glucose, and ethanol. These compounds served as carbon
sources. Furthermore, the microbial consortia facilitating the dechlorination of
1,2,4-TCB also was able to degrade isomers of tetrachlorobenzene to other
isomers of TCB and 1,2-DCB. More recent studies show that a strain of
Dehalococcoides, can reductively dechlorinate 1,2,4-TCB under anaerobic
conditions (Holscher and others, 2003; Griebler and others, 2004a). In addition,
Adrian and others (1998) suggest that fermentation is the primary degradation
process for the chlorobenzenes under anaerobic conditions. This study also
38
Chapter 1 Volatile Organic Compounds
showed that the cometabolism of 1,2,4-TCB was inhibited by the presence of
sulfate, sulfite and molybdate.
Furthermore, Ramanand and others (1993) show that 1,2,4-TCB had
declined by 63% within 30 days under anaerobic conditions. Dermietzel and Vieth
(2002) show that the anaerobic biodegradation of 1,4-DCB was markedly slower
under iron-reducing conditions than under aerobic conditions. In general, it
appears that the biodegradation of the chlorinated benzenes is slower under
anaerobic than under aerobic conditions.
Figure 1.13. Laboratory-derived pathway for the aerobic and anaerobic biodegradation of
1,2,4-tricholorobenzene.
39
Chapter 1 Volatile Organic Compounds
Figure 1.14. Laboratory-derived pathway for the aerobic biodegradation of 1,4dicholorobenzene.
Table 1.11. Laboratory or environmental half-lives and by-products for the aerobic and
anaerobic biodegradation of selected chlorinated benzene compounds detected in
groundwater.
40
Chapter 1 Volatile Organic Compounds
Figure 1.15. Laboratory-derived pathway for the aerobic biodegradation of chlorobenzene
and 1,2-dicholorobenzene.
41
Chapter 1 Volatile Organic Compounds
1.4.3 Bioegradation of monochlorobenzene
In four billion years, micro-organisms have evolved an extensive range of
enzymes and control mechanisms to be able to degrade a wide array of naturally
occurring aromatic compounds including chlorobenzene. The rate of naturally
occurring biodegradation is often limited by either the concentration of an
appropriate electron-acceptor or the availability of nutrients for cell growth.
Although chlorobenzene can be metabolyzed aerobically (Nishino et al., 1992;
Reineke et al., 1984; Rochkind et al., 1986; Van der Meer et al., 1992), it has not
been reported to be degraded through the use of other electron acceptors (Bouwer
et al., 1983; Nishino et al., 1992; Reineke et al., 1984). Molecular oxygen appears
necessary for ring fission. Some removal through reductive dechlorination may
occur in conducive environments with excess chlorobenzene (Montgomery et al.,
1994). In general, intermediary metabolites of chlorobenzene appear similar to
those documented for unhalogenated aromatic compounds. Biodegradation under
microaerophilic conditions has also been reported (Vogt et al., 2003).
The main enzymes involved in chlorobenzene’s catabolic reactions are
oxygenases. Oxygen can be incorporated immediately into organic products by
reactions catalyzed by enzymes such as oxygenases or hydroxylases (Gibson et
al., 1982; Harayama et al., 1992). These enzymes use metals to activate dioxygen
that is not reactive in its original state. During these processes oxygen is
metabolized into very reactive forms like singlet oxygen and hydroxyl radicals.
Oxygenase enzymes play an important role in aromatic catabolic pathways. They
initiate the degradation of aromatic compounds by hydroxylation of the aromatic
ring for preparation of the ring fission and are involved in ring fission.
Oxygenases of different organisms catalyzing similar reactions share similar
features, structures, and reaction mechanisms. By means of the amino acid
sequence, oxygenases can be divided into different families. Comparisons of the
amino acid sequence of related enzymes can give information on amino acids of
essential function including active sites, and on the evolution of the enzymes.
Oxygenases involved in initial attack of the aromatic compound can be
divided in either mono-oxygenases or dioxygenases. Oxygenases that incorporate
42
Chapter 1 Volatile Organic Compounds
only one oxygen atom into the structure of the substrate are called monooxygenases. In that case, the remaining oxygen atom of the oxygen molecule is
reduced into H2O. Mono-oxygenases are multi-component enzyme complexes.
They consist of a combination of the following components, i.e., a hydroxylase
component consisting of an -, - and -subunit, a ferredoxin component, a
small oxygenase subunit, and a flavo-iron-sulfur NADH-oxidoreductase
component. Some mono-oxygenases contain additional polypeptides with
unknown function. The hydroxylase component is the protein that activates
molecular oxygen and binds it to the substrate.
Oxygenases that incorporate two oxygen atoms into the substrate are
called dioxygenases. They are also multi-component enzyme complexes, and in
most cases consist of four components including an iron-sulfur oxidase large asubunit, a ferredoxin component, an iron-sulfur oxidase small b-subunit and a
reductase component.
1.4.3.1 Degradation via the ortho-cleavage pathway
The majority of the microorganisms able to mineralize chlorinated
aromatics do not posses enzyme systems capable of initial dechlorination. They
transform chloroaromatics to chlorocatechols, which are further metabolyzed via
the enzyme of the ortho-cleavage pathway, and dechlorination occurs after ringcleavage (Schlömann, 1984).
In 1983, Reineke et al. isolated a bacterium (strain WR136) able to grow
on monochlorobenzene and proposed a degradative pathway on the basis of the
enzyme activities found (Figure 1.16). 3-Chlorocatechol is subject to ortho
cleavage with formation of 2-chloro-cis,cis-muconic acid. This is cycloisomerized
with
coincident
or
subsequent
elimination
of
chloride
yielding
4-
carboxymethylenebut-2-en-4-olide, which is further converted by use of a
hydrolase. The resulting maleylacetate is reduced in an NADH-dependent reaction
to 3-oxoadipate. This modified ortho pathway is the only pathway currently
known for the aerobic degradation of catechol formed from chlorobenzene.
Analogous pathways have been described for the dichlorocatechols derived by the
transformation of dichlorobenzenes (de Bont et al., 1986; Haigler et al., 1988;
43
Chapter 1 Volatile Organic Compounds
Schraa et al., 1986; Spain et al., 1987). In each instance, the initial attack is by a
dioxygenase. The initial oxidation results in the formation of a cis-dihydrodiol.
Subsequent ring fission and elimination of chloride leads to the detoxification and
mineralization of these compounds. The key enzime is the pyrocatechase II (Dorn
et al., 1978; Reineke et al., 1984) that converts chlorocatechols to chloro-cis,cismuconic acids. Absence of this enzyme in organisms with initial oxygenases with
broad substrate specificities may lead to the accumulation of chlorocatechols or to
the misrouting of chlorocatechol down the meta cleavage pathway, ultimately
resulting in cell death.
Figure 1.16. Modified ortho pathway.
44
Chapter 1 Volatile Organic Compounds
In another study (Nishino et al., 1992), bacterial isolates were obtained
from groundwater and soils contaminated with chlorobenzene. The isolates were
tested to determine whether the natural community could remove the groundwater
contaminants. These isolates were identified and characterized as to their ability to
grow on chlorobenzene and related aromatic compounds. The complete
consortium could mineralize approximately 54% of the chlorobenzene within 7
days, with no accumulation of 3-chlorocatechol. Metabolic pathways were
evaluated for several isolates. One phenotype was characterized by the ability to
degrade chlorobenzene by the modified ortho pathway. One strain also degraded
p-dichlorobenzene by using the same pathway. Isolates exhibiting a second
phenotype degraded p-cresol, benzene, and phenol by the classical ortho pathway
and accumulated 3-chlorocatechol when grown in the presence of chlorobenzene.
Strains of the third phenotype grew on complex media in the presence of
chlorobenzene but did not transform any of the aromatic compounds tested.
1.4.3.2 Degradation via the meta-cleavage pathway
It is generally accepted that degradation of chloroaromatics does not
proceed via the meta-cleavage pathway (Knackmuss, 1981; Pettigrew et al., 1991;
Rojo et al., 1987). An explanation for this has been found in the production of an
acylchloride from 3-chlorocatechol by the catechol 2,3-dioxygenase of the metacleavage pathway, which leads to rapid suicide inactivation of the enzyme
(Bartels et al., 1984). Therefore, meta-cleavage is considered to be unsuitable for
the mineralization of haloaromatics that are degraded via halocatechols. Whereas
chlorocatechols are mineralized via ortho–cleavage pathways, methylaromatics
are commonly mineralized via meta-cleavage routes. Simultaneous metabolism of
chloro- and methylcatechols often creates biochemical anarchy. Meta-cleavage
leads to substrate misrouting in the case of 4-chlorocatechol or formation of a
suicide product in the case of 3-chlorocatechol. Formation of dead-end
methyllactones can occur when the ortho-cleavage pathway is dealing with
methylcatechols.
Consequently, only a few strains which can grow on mixtures of
methylated and chlorinated aromatics are known (Haigler et al., 1992; Pettigrew
45
Chapter 1 Volatile Organic Compounds
et al., 1991). They all use a modified ortho-cleavage pathway for the conversion
of the chlorinated substrate. Pseudomonas putida GJ31 (Oldenhuis et al., 1989)
and Pseudomonas sp. strains JS6 (Pettigrew et al., 1991) and JS150 (Haigler et
al., 1992) are the only strains known to grow on a mixture of chlorobenzene and
toluene. Mars et al. (1997) showed that Pseudomonas putida GJ31 grows on
chlorobenzene via a meta-cleavage pathway which allows the simultaneous
utilization of toluene (Figure 1.17). In addition, 3-chlorocatechol was found to be
the ring cleavage substrate formed from chlorobenzene, which was dehalogenated
during ring cleavage to produce 2-hydroxymuconic acid. The authors observed
that the enzymes of the modified ortho-cleavage pathway were never present,
while the enzymes of the meta-cleavage pathway were detected in all cultures.
Apparently, Pseudomonas putida GJ31 has a meta-cleavage enzyme (catechol
2,3-dioxygenase) which is resistant to inactivation by the acylchloride, providing
this strain with the exceptional ability to degrade both toluene and chlorobenzene
via the meta-cleavage pathway.
Figure 1.17. Proposed catabolic pathway of chlorobenzene by P.putida GJ31 by analogy to
the known meta-cleavage pathway. Enzymes: 1, chlorobenzene dioxygenase; 2,
chlorobenzene dihydrodiol dehydrogenase; 3, catechol 2,3-dioxygenase; 4, oxalocrotonate
isomerase; 5, oxalocrotonate decarboxylase; 6, 2-oxopent-4-enoate hydratase; 7, 4-hydroxy2-oxovalerate aldolase.
46
Chapter 1 Volatile Organic Compounds
1.4.3.3 Biodegradation of mixtures of substituted benzenes
The degradation of a wide range of substituted aromatic compounds by a
strain of Pseudomonas has been observed (Haigler et al., 1992). Pseudomonas sp.
strain JS150 was isolated as a nonencapsulated variant of Pseudomonas sp. strain
JS1 that contains the genes for the degradative pathways of a wide range of
substituted aromatic compounds. Pseudomonas sp. strain JS150 grew on phenol,
ethylbenzene, toluene, benzene, naphthalene, benzoate, p-hydroxybenzoate,
salicylate, chlorobenzene, and several 1,4-dihalogenated benzenes. Enzyme assays
with cell extracts showed that the enzymes of the meta, ortho, and modified ortho
cleavage pathways can be induced in strain JS150. Strain JS150 contains a
nonspecific toluene dioxygenase with a substrate range similar to that found in
strains of Pseudomonas putida. Chlorobenzene-grown cells of strain JS150
degraded
mixtures
of
chlorobenzene,
benzene,
toluene,
naphthalene,
trichloroethylene, and 1,2- and 1,4-dichlorobenzenes in continuous culture.
Results indicated that induction of appropriate biodegradative pathways in strain
JS150 permits the biodegradation of complex mixtures of aromatic compounds.
1.4.3.4 Degradation under oxygen-limited conditions
Vogt et al. (2002) studied the monochlorobenzene degradation at low
oxygen concentration by five bacterial strains (Acidovorax facilis B517,
Cellulomonas turbata B529, Pseudomonas veronii B547, Pseudomonas veronii
B549, and Paenibacillus polymyxa B550) isolated on chlorobenzene as the sole
source of carbon and energy. These strains were screened for the accumulation of
the putative metabolic intermediate 3-chlorocatechol during growth on
chlorobenzene under oxygen-limited conditions in the presence and absence of
nitrate (1 mM). 3-Chlorocatechol accumulated in the growth media of all five
strains, but accumulation was significantly less in cultures of A. facilis B517
compared to the other four strains. The presence of nitrate did not influence the
biological conversion pattern. For P. veronii B549, a clear relationship between
the presence of 3-chlorocatechol in the medium and low oxygen concentrations
was demonstrated. The authors made the assumption that accumulation of 3chlorocatechol was due to the low enzymatic turnover of the 3-chlorocatechol
cleaving enzyme, catechol-1,2-dioxygenase, at low oxygen concentrations.
47
Chapter 1 Volatile Organic Compounds
Other strains able to degrade monochlorobenzene as a sole carbon source
include Escherichia Hermanii (Kiernicka et al.,1999) and Acidovorax avenae
(Monferràn et al., 2005). Escherichia Hermanii was isolated from sludge of an
industrial wastewater treatment plant. High chlorobenzene concentrations (up to
394 mg l-1) had low toxic effects towards this strain, which was able to degrade
chlorobenzene without any previous adaptation. Acidovorax avenae was isolated
in a polluted site of Suquìa River (Argentina) from a subsurface microbial
community acclimatated during 15 days using 1,2-dichlorobenzene as the sole
carbone source (aerobic conditions). Acidovorax avenae was able to perform the
complete biodegradation of 1,2-dichlorobenzene in two days affording
stoichiometric amounts of chloride. This pure strain was also tested for
biodegradation of chlorobenzene, 1,3- dichlorobenzene and 1,4- dichlorobenzene,
giving similar results to the experiments using dichlorobenzene. The aromaticring-hydroxylating dioxygenase (ARHDO) α-subunit gene core, encoding the
catalytic site of the large subunit of chlorobenzene dioxygenase, was detected by
PCR amplification and confirmed by DNA sequencing. These results suggest that
the isolated strain of A. avenae could use a catabolic pathway, via ARHDO
system, leading to the formation of chlorocatecols during the first steps of
biodegradation, with further chloride release and subsequent paths that showed
complete substrate consumption.
Cometabolic biodegradation has also been reported (Jeckorek et al.,
2002). The degradation of chlorobenzene was investigated with the specially
chosen strain Methylocystis sp. GB 14 DSM 12955, in 23 ml headspace vials and
in a soil column filled with quaternary aquifer material from a contaminated
location in Bitterfeld (Germany). A long-term experiment was carried out in this
column: groundwater polluted by chlorobenzene was continuously fed through the
column, bubbled with a 4% CH4-96% air mixture. Chlorobenzene was oxidized
by up to 80% under pure culture conditions in the model experiments and was
completely degraded under the mixed culture conditions of the column
experiments. The enzyme responsible for this ability was the sMMO (soluble
methane monooxygenase)
48
Chapter 1 Volatile Organic Compounds
49
CHAPTER 2
2. BIODEGRADATION TECHNOLOGIES FOR REMEDIATION
OF CONTAMINATED SITES
___________________________________________________________________________________________________
_
2.1 INTRODUCTION
Bioremediation is a grouping of technologies that use microbiota to degrade or
transform hazardous contaminants to compounds such as carbon dioxide, water,
inorganic salts, microbial biomass, and other byproducts that may be less
hazardous than the parent compounds. Numerous application of bioremediation
are nowadays widely accepted as a remedial alternative and are in wide use at site
contaminated with petroleum products and/or hazardous wastes. Some
bioremediation technologies, such as cometabolic bioventing, are still in
development and should be considered innovative. Other bioremediation
technologies, such as anaerobic bioventing, are current topic of research.
The following contaminants have been bioremediated succesfully at many sites:
•
Halogenated and non-halogenated volatile organic compounds (VOCs)
•
Halogenated and non-halogenated semi-volatile organic compounds
(SVOCs).
Contaminants with a more limited bioremediation performance include:
•
Polycyclic aromatic hydrocarbons (PAHs)
•
Organic pesticides and herbicides
•
Polychlorinated biphenils (PCBs).
Bioremediation remains an active field of technology research and development at
both the laboratory and field scale. For example, applications to chlorinated
aliphatic hydrocarbons (CAHs), perchlorate and methyl-tert-butyl ether (MTBE)
were developed rapidly in recent years.
Chapter 2 Biodegradation Technologies for Remediation of Contaminated Sites
The field of bioremediation can be divided in to several broad categories. For
example, bioremediation technologies may be applied to in situ or ex situ media.
In situ processes treat soils and groundwater in place without removal while ex
situ processes involve the removal of the contaminated media to a treatment area.
Another way to divide the bioremediation field is based on additives to
environmental media. Intrinsic bioremediation depends on indigenous microflora
to degrade contaminants (EPA, 2000). This approach is used in situ and takes
advantage of pre-existing processes to degrade hazardous wastes. Intrinsic
bioremediation require careful site assessment and monitoring to make sure that
the ongoing processes are protective of environmental receptors. Alternatively,
enhanced bioremediation facilitates biodegradation by manipulating the microbial
environment, typically by supplying chemical amendments such as air, organic
substrates, electron donors, nutrients and other compounds that affect metabolic
reactions (EPA, 2000). Enhanced bioremediation may also called biostimulation
when only chemical amendments are added. Examples include bioventing, land
farming, biopiles, composting. Biostimulation can be applied in situ or ex situ, to
treat soil and other solids, groundwater and surface water. Sometimes
bioaugmentation (addition of microbial cultures) is used to enhance biotreatment.
Bioaugmentation (almost always performed in conjunction with biostimulation)
may be needed for specific contaminants that are not degraded by the indigenous
microorganisms.
In bioremediation, fundamental biological activities are exploited to degrade or
transform contaminants of concern. The biological activity to be exploited
depends on the specific contaminants of concern and the media where the
contamination is located. For example in aerobic environments many microbes
are able to degrade organic compounds, such as hydrocarbons. These microbes
gain energy and carbon for building cell materials from these biochemical
reactions. At many sites with fuel contamination, the amount of oxygen present
limits the extent of biotreatment. Thus, by adding oxygen in the form of air,
contaminant degradation proceeds directly. In cometabolism, microbes do not
gain energy or carbon from degrading a contaminant. Instead, the contaminant is
degraded via a side reaction. Cometabolic bioventing is an example, where
51
Chapter 2 Biodegradation Technologies for Remediation of Contaminated Sites
microbes may be fed with propane and degrade trichloroethyline (TCE) or less
chlorinated ethenes.
Depending on the contaminant of concern and the media, a technology may
exploit aerobic or anaerobic metabolism. Aerobic metabolism is more commonly
exploited and can be effective for hydrocarbons and other organic compounds.
Many organisms are capable of degrading hydrocarbons using oxygen as the
electron acceptor and the hydrocarbons as carbon and energy sources. In some
cases, contaminants are aerobically degraded to carbon dioxide and water, but in
other cases the microbes do not completely degrade contaminants. Aerobic
technologies may also change the ionic form of metals. If a site contains mixed
metal and organic wastes, it is necessary to consider whether theoxidized forms of
the metal species will be environmentally acceptable.
Anaerobic metabolism involves microbial reactions occurring in the absence of
oxygen,
and
encompasses
many
processes
including
fermentation,
methanogenesis, reductive dechlorination, sulfate-reducing activities, and
denitrification. Depending on the contaminant of concern, a subset of these
activities could be cultived.
In anaerobic metabolism, nitrate, sulfate, carbon dioxide, oxidized metals, or
organic compounds may replace oxygen as the electron acceptor. For example, in
anaerobic reductive dechlorination, chlorinated solvents may serve as the electron
acceptor.
When selecting a bioremediation technology, it is important to consider the
contaminants of concern, contaminated matrix, potential biological pathways to
degrade a contaminant, and current condition of a site. For example, TCE can be
degraded via aerobic and anaerobicmechanisms. If groundwater is contaminated
with TCE current groundwater conditions may be helpful in deciding which
biological mechanism to exploit. If groundwater is already anaerobic, then
anaerobic reductive dechlorination may be the best approach. However, if the
TCE plume is diffuse and the groundwater is aerobic, it may be possible to use
cometabolic technologies.
A key concept in evaluating all bioremediation technologies is microbial
availability: if the contaminant is so tightly bound up in the solid matrix that
52
Chapter 2 Biodegradation Technologies for Remediation of Contaminated Sites
microorganism cannot access it, then it cannot be bioremediated. However, low
microbial availability does not imply an absence or risk; compounds may be
available to environmental receptors depending on the receptors and routes of
exposure.
Thus, when selecting a bioremediation technology for a specific site, it is prudent
to consider the contaminants of concern, potential degradation intermediates and
residual of the contaminants, co-contaminants, environmental receptors, routes of
exposure, and buffer zones between contamination and receptors. Bioremediation
technologies have proven to be protective and cost-effective solutions at many
sites. However, conditions at a specific site may not be appropriate.
2.2 IN SITU BIOREMEDIATION
There are two major types of in situ bioremediation: intrinsic and enhanced. Both
rely on natural processes to degrade contaminants with (enhanced) or without
(intrinsic) amendments.
In recent years, in situ bioremediation concepts have been applied in treating
contaminated soil and groundwater. Removal rates and extent vary based on the
contaminant of concern and site-specific characteristics. Removal rates also are
affected by variables such as contaminant distribution and concentrations;
indigenous microbial populations and reaction kinetics; and parameters such as
pH, moisture content, nutrient supply and temperature. Many of these factors are a
function of the site and the indigenous microbial community and, thus, are
difficult to manipulate.
When in situ bioremediation is selected as a treatment, site monitoring activities
should demonstrate that biologically mediated removal is the primary route of
contaminant removal. Sampling strategies should consider appropriate analytes
and tests, as well as site heterogeneity. In some cases, extensive sampling may be
required to distinguish bioremediation from other removal mechanisms or
statistical variations. Small-scale treatability studies using samples from the
contaminated site may also be useful in the demonstrating the role that biological
activity plays in contaminant removal (EPA, 1995B; EPA, 1998a; EPA, 2000).
53
Chapter 2 Biodegradation Technologies for Remediation of Contaminated Sites
2.2.1 Intrinsic in situ bioremediation
Intrinsic bioremediation relies on natural processes to degrade contaminants
without
altering
current
conditions
or
adding
amendments.
Intrinsic
bioremediation may play a role in monitored attenuation (MNA) sites. Natural
attenuation (NA) relies on natural physical, chemical and biological processes to
reduce or attenuate contaminant concentrations. Under favorable conditions, NA
will reduce the concentrations, mass, toxicity, mobility, and/or volume of
contaminants in soil and groundwater. Natural processes in NA include dilution,
dispersion, sorption, volatilization, chemical reactions such as oxidation and
reduction, biological reactions and stabilization. Some processes have undesirable
results, such creation of toxic degradation products or the transfer of contaminants
to other media.
Implementing natural attenuation requires a thorough site assessment and
development of a conceptual model of the site. After determining the presence of
a stable shrinking plume, site-specific, risk-based decisions using multiple lines of
evidence may facilitate implementation of MNA at a site. While MNA is
somewhat passive in that nothing is being added to the contamination zone, it
requires active monitoring , which should be included as part of the design plan
for a site. In some cases, such long-term monitoring may be more expensive than
active remediation. MNA is only applicable to carefully controlled and monitored
sites and must reduce contaminant concentrations to levels that are protective of
human health and the environment in reasonable time frames (EPA, 1998a).
Depending on site-specific conditions, MNA may be a reasonable alternative for
petroleum hydrocarbons as well as chlorinated and non chlorinated VOCs and
SVOCs (EPA, 1999a; EPA, 1999b).
Important observations related to the performance of natural attenuation
technology are:
•
it is a relatively simple technology compared to other remediation
technologies;
•
it can be carried out with little or no site disruption;
•
it often requires more time to achieve cleanup goals than other
conventional remediation methods;
54
Chapter 2 Biodegradation Technologies for Remediation of Contaminated Sites
•
it requires a long-term monitoring program; program duration affects
the costs;
•
if natural attenuation rates are too slow, the plume could migrate;
•
it is difficult to predict with high reliability the performance of natural
attenuation;
Sites must meet one ore more of the following criteria:
•
it must be located in an area with little risk to human health or the
environment;
•
the contaminated soil or groundwater must be located an adequate
distance from potential receptors;
•
there must be evidence that natural attenuation is actually occurring at
the site.
2.2.2 Enhanced in situ bioremediation
Enhanced in situ bioremediation can be applied to groundwater, vadose
zone soils or, more rarely, aquatic sediments. Exogenous microorganisms may be
added where organisms able to degrade specific contaminants are absent
(bioaugmentation). Additives such as oxygen (or other electron acceptors),
nutrients, biodegradable carbonaceous substrates, bulking agents, and/or moisture
are added to enhance the activity of natural occurring or indigenous microbial
populations:
Bioaugmentation: involves the addition of supplemental microbes to the
subsurface where organisms able to degrade specific contaminants are deficient.
Microbes may be “seeded” from populations already present at a site and grown in
aboveground reactors or from specially cultivated strains of bacteria known to
degrade specific contaminants. The application of bioaugmentation technology is
highly site-specific and highly dependent on the microbial ecology and
physiology of the subsurface (EPA 1998).
Nutrient addition: involves the addition of key biological building blocks,
such as nitrogen and phosphorus and other trace nutrients necessary for cell
growth. Addition of nutrients generally is applied as a supplement to
55
Chapter 2 Biodegradation Technologies for Remediation of Contaminated Sites
bioaugmentation or addition of electron donors or electron acceptors, so that
concentrations of nutrients in the subsurface do not become a limiting factor for
an in situ bioremediation application.
Electron donor addition: involves the addition of a substrate that acts as a
reductant in the redox reaction used by the CAH-degrading microbe to produce
energy. A substrate such as toluene, propane, or methane may be added to act as a
cometabolic oxidant, when the CAH also is oxidized. A substrate such as
hydrogen, a source of hydrogen, or a hydrogen release compound may be added to
act as a direct reductant, when the CAH is reduced.
Electron acceptor addition: involves the addition of oxygen (for aerobic
mechanisms) or an anaerobic oxidant such as nitrate (for anaerobic mechanisms),
which is used by the CAH-degrading microbes present in the subsurface.
2.2.2.1 Vadose zone soil remediation
While the funadamental biological activities exploited by in situ bioremediation
may occur naturally, many sites will require intervention to facilitate cleanup. For
example the addition of organic substrates, nutrients or air will provide the
appropriate environment for specific microbial activities or enhanced removal
rates. In general, hydrocarbons and lightly chlorinated contaminants may be
removed through aerobic treatment while highly chlorinated species are degraded
primarily through anaerobic treatment. Both anaerobic and aerobic treatment may
occur through direct or cometabolic pathways (see 1.3.4.2).
The primary in situ biological technology applicable is bioventing (aerobic,
cometabolic, or anaerobic).
Aerobic bioventing is useful in treating aerobically degradable
contaminants such as fuels. Contaminated unsaturated soils with low oxygen
concentrations are treated by supplying oxygen to facilitate aerobic microbial
biodegradation. Oxygen is typically introduced by air injection wells that push air
into the subsurface (figure 2.1); vacuum extraction wells , which draw air through
the surface, may also be used. Extracted gases may require treatments since
volatile compounds may be removed from the ground. Compared with soil vapor
extraction bioventing employs lower air flow rates that provide only the amount
ofoxygen required to enhance removal. Operated properly the injection of air does
56
Chapter 2 Biodegradation Technologies for Remediation of Contaminated Sites
not result in the release of the contaminants to the atmosphere through
volatilization because of these low flow rates. Bioventing is designed primarily to
trea aerobically degradable contaminants, such as non-chlorinated VOCs and
SVOCs, that are located in the vadose zone of the capillary fringe. In addition to
fuels treatment, aerobic bioventing has treated a variety of other contaminants
including non-halogenated solvents such as benzene, acetone ,toluene, and
phenol; lightly halogenated solvents such as 1,2-dichloroethane, dichloromethane,
and chlorobenzene; and SVOCs such as low-molecular-weight PAHs.
Nevertheless bioventing has some limitations involving the ability to deliver
oxygen to the contaminated soil. For example, soils with extremely high moisture
content may be difficult to biovent because of reduced soil gas permeability.
While it is relatively inexpensive, bioventing can take a few years to clean
up a site depending on contaminant concentrations and site-specific removal rates.
Figure 2.1. Aerobic bioventing in injection mode.
Cometabolic bioventing has been used at a few sites to treat chlorinated
solvents
such
as
trichloroethilene
(TCE),
trichloroethane
(TCA)
and
dichloroethene (DCE). Similar to bioventing, cometabolic bioventing involves the
injection of gases into the subsurface; however cometabolic bioventing injects
both air and a volatile organic substrate, such as propane. This technology exploits
competitive reactions mediated by monooxygenase enzymes which catalyze the
oxidation of hydrocarbons, often through epoxide intermediates. These enzyme
can also catalyze the dechlorination of chlorinated hydrocarbons. Thus, by
57
Chapter 2 Biodegradation Technologies for Remediation of Contaminated Sites
supplying an appropriate organic substrate and air, production of monooxygenases
may be stimulated resulting in the contaminants breakdown (see 1.3.4.2.1).
In addition to the variable discussed for the aerobic bioventing, the degradation
rate and design of cometabolic bioventing systems are dependent on many factors
including soil gas permeability, organic substrate concentration, type of organic
substrate selected, and oxygen supply and radius of influence. As with aerobic
bioventing, difficulty in distributing gases in the subsurface may make the
application of this technology more complicated.
Anaerobic bioventing. While aerobic and cometabolic bioventing are useful for
degrading many hydrocarbons and lightly chlorinated compounds, some
chlorinated species are not effectively treated aerobically. Microbes may degrade
these contaminants directly via anaerobic reductive dechlorination or through
anaerobic cometabolic pathways. Anaerobic reductive dechlorination is a
biological mechanism typically marked by sequential removal of chlorine from a
molecule (see 1.3.4.2.2). Microbes possessing this pathway do not gain energy
from this process. Anaerobic cometabolism is similar to aerobic cometabolism in
that microbes fortuitously degrade contaminants while reducing other compounds
(cometabolites). Anaerobic bioventing may use both biological mechanisms to
destroy the contaminants of concern.
Anaerobic bioventing uses the same type of gas delivery system as the
other bioventing technologies, but injects nitrogen and an electron donor, instead
of air, to establish reductive anaerobic conditions. The nitrogen displaces the soil
oxygen, and small amounts of an electron donor gas (such as hydrogen and carbon
dioxide) produce reducing conditions in the subsurface, thereby facilitating
microbial dechlorination. Volatile and semi-volatile compounds may be produced
during anaerobic bioventing. Some of these compounds may be slow to degrade
under anaerobic conditions. These compounds may be treated in two ways.
Volatile compounds may diffuse into the soils surrounding the treatment zone,
where aerobic degradation may occur. SVOCs and VOCs remaining in the
treatment zone may be treated by following anaerobic bioventing with aerobic
bioventing. Since aerobic and anaerobic bioventing share similar gas delivery
systems, the switch can be made by simply changing the injected gas.
58
Chapter 2 Biodegradation Technologies for Remediation of Contaminated Sites
Anaerobic bioventing is an emerging technology that has been
demonstrated in several laboratory and field studies. This process may be useful
in treating highly chlorinated compounds such as tetrachloroethene (PCE), TCE,
RDX,
pentachlorophenol,
and
dichlorodiphenyltrichloroethane
pesticides
(DDT).
As
such
with
the
as
lindane
other
and
bioventing
technologies, the ability to deliver gases to the subsurface is important. Soils with
high moisture content or low gas permeability may require careful system design
to deliver appropriate levels of nitrogen and the electron donor. Sites with shallow
contamination or nearby buildings are also a challenge since this technology is
operated by injecting gases. In addition, anaerobic bioventing can take a few years
to clean up a site depending on the contaminant concentrations and site-specifi c
removal rates.
2.2.2.2 Surficial soil remediation
If contamination is shallow, soil may be treated in place using techniques similar
to land treatment or composting. Variations of these technologies involve tilling
shallow soils and adding amendments to improve aeration and bioremediation.
Since these treatments do not include an impermeable sublayer, contaminant
migration may be a concern depending on the contaminants of concern and
treatment amendments. A more prudent approach would be to excavate soils and
treat them in lined beds.
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Chapter 2 Biodegradation Technologies for Remediation of Contaminated Sites
2.2.2.3 Groundwater and saturated soil remediation
In situ bioremediation techniques applicable to ground water and saturated
soil include dechlorination using anaerobic reducing conditions, enhanced aerobic
treatment, biological reactive barriers that create active remediation zones, and
bioslurping/biosparging techniques that promote aerobic degradation.
Anaerobic reductive dechlorination has been used at many sites where the
ground water has been contaminated with chlorinated solvents, such as TCE or
PCE. In this treatment, organic substrates are delivered to the subsurface where
they are fermented. The fermentation creates an anaerobic environment in the area
to be remediated and generates hydrogen as a fermentation byproduct. The
hydrogen is used by a second microbial population to sequentially remove
chlorine atoms from chlorinated solvents (see 1.3.4.2.1). If PCE were degraded
via reductive dechlorination, the following sequential dechlorination would be
observed: PCE would be converted to TCE, then to DCE, vinyl chloride (VC),
and/or dichloroethane (EPA, 1998a).
Anaerobic dechlorination may also occur via cometabolism where the
dechlorination is incidental to the metabolic activities of the organisms. In this
case, contaminants are degraded by microbial enzymes that are metabolizing other
organic substrates. Cometabolic dechlorination does not appear to produce energy
for the organism. At pilot- or full-scale treatment, cometabolic and direct
dechlorination may be indistinguishable, and both processes may contribute to
contaminant removal. The microbial processes may be distinguished in the more
controlled environment of a bench-scale system (EPA, 1998a).
Anaerobic reductive dechlorination is primarily used to treat halogenated
organic contaminants, such as chlorinated solvents. As well as the variables
discussed initially, the treatment rate and system design are dependent on several
factors including site hydrology and geology, type and concentration of organic
substrates, and site history. As with cometabolic bioventing, the selection of
organic substrate and the concentration used are controllable and can be important
to the removal rate. Treatability or bench-scale testing can be useful in selecting
the best organic substrate and concentration for a site. In addition, small-scale
testing can demonstrate that full dechlorination is possible at a site. In some cases,
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Chapter 2 Biodegradation Technologies for Remediation of Contaminated Sites
dechlorination may stall at DCE despite the presence of sufficient electron donors.
If a site does not demonstrate full dechlorination (either as part of site assessment
or in microcosm testing), a combined treatment strategy, such as anaerobic
treatment followed by aerobic treatment, may be successful. Alternatively,
bioaugmentation may improve the dechlorination rate.
Aerobic Treatment. Similar to bioventing, enhanced in situ aerobic ground water
bioremediation processes are used in situations where aerobically degradable
contaminants, such as fuels, are present in anaerobic portions of an aquifer. In
these situations, air or other oxygen sources are injected into the aquifer near the
contamination (figure 2-2). As the oxygenated water migrates through the zone of
contamination, the indigenous bacteria are able to degrade the contaminants
(EPA, 1998a; EPA, 2000).
Aerobic treatment may also be used to directly or cometabolically degrade
lightly chlorinated species, such as DCE or VC. In the direct aerobic pathway, air
is injected into the aquifer. The microbes appear to generate energy by oxidizing
the hydrocarbon backbone of these contaminants, resulting in the release of
chloride (EPA, 2000). This process has been used to complete contaminant
removal following anaerobic treatment at several sites (EPA, 1998a; EPA, 2000).
Cometabolic aerobic treatment is founded on the same biological
principles as cometabolic bioventing and involves the addition of oxygen and
organic substrates, such as methane, to the aquifer. As with other cometabolic
processes, these organic substrates are metabolized by enzymes that incidentally
degrade the contaminant. In this treatment, sufficient oxygen must be present to
fuel the oxidation of both the substrate and contaminant.
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Chapter 2 Biodegradation Technologies for Remediation of Contaminated Sites
Figure 2.2. Aerobic treatment.
Amendment Delivery. In situ groundwater treatment, either aerobic or
anaerobic, may be configured as direct injection of air or aqueous streams or as
ground water recirculation. In direct injection, amendments, such as organic
substrates, oxygen sources, or nutrients, are directly injected into the aquifer. For
example, oxygen may be sparged into the aquifer as a gas. Lactate or hydrogen
peroxide may be injected as a liquid stream; when using hydrogen peroxide,
caution should be used as it may act as a disinfectant. In some cases, both liquids
and gases are added. The ground water recirculation configuration involves
extracting ground water, amending it as needed, and then re-injecting it back into
the aquifer. Recirculation may also be conducted below the ground surface by
extracting ground water at one elevation, amending it in the ground, and reinjecting it into another elevation (EPA, 1998a; EPA, 2000).
In addition to the variables discussed initially, the treatment rates and
system design are the result of several factors including site hydrology and
geology, amendment to be added, solubility of air or oxygen sources, and site
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Chapter 2 Biodegradation Technologies for Remediation of Contaminated Sites
history. The low solubility of air in water often limits reaction rates and may make
this process impractical if cleanup time is short.
Biological reactive barriers consist of an active bioremediation zone
created in the contamination zone. The use of in situ treatment walls for
remediation is an emerging technology that has been developed and implemented
only within the last few years. Treatment walls are structures installed
underground to treat the contaminated groundwater found at hazardous waste sites
(figure 2.3).Treatment walls rely on the natural movement of water to carry the
contaminants through the wall structure. As contaminated groundwater passes
through the treatment wall, the contaminants are either trapped by the treatment
wall or transformed into harmless substances that flow out of the wall (USEPA,
1996d). Target contaminant groups for passive treatment walls are VOCs,
SVOCs, and inorganics. The specific filling chosen for the wall is based on the
contaminant found at the site. Wall fillings work through different chemical
processes, of which the three most common are (USEPA, 1996d; Birke et al.,
2003):
•
Sorption barriers contain fillings that remove contaminants from
the groundwater by physically removing contaminants from the
groundwater and holding them on the barrier surface. Zeolites and
activated carbon are two examples of sorption barriers.
•
Precipitation barriers contain fillings that react with contaminants
in the groundwater as they pass through the treatment wall. The
reactions cause the contaminants dissolved in groundwater to
become insoluble and to precipitate out. The barrier traps the
insoluble products and clean groundwater flows out the other side.
•
Degradation barriers cause reactions that break down the
contaminants in the groundwater into harmless products. Filling
walls with iron granules helps ton degrade certain VOCs, and walls
filled with a mixture of nutrients and oxygen sources can stimulate
the activity of the microorganisms found in the groundwater.
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Chapter 2 Biodegradation Technologies for Remediation of Contaminated Sites
Two main types of treatment walls exist:
•
Permeable reactive trench: this is the simplest form of treatment
walls and it consists of a trench that extends across the entire width
of the plume. The system is installed by digging a trench and filling
it with permeable material. As the contaminant plume moves
through the wall, contaminants are removed by various mass
transfer processes such as air stripping, SVE, and adsorption.
•
Funnel and gate systems: used primarily when contaminated
plumes are too large or too deep to dig a trench across its width. To
overcome this problem, a system consisting of low permeability
cut-off walls are installed to funnel contaminated groundwater to a
smaller reactive wall to treat the plume. When dealing with funnel
and gate systems, the gate is used to pass contaminated
groundwater through the reactive wall, and the funnel is integrated
into the system to force water through its gates. Plumes which
contain a mixture of contaminants are funnelled through a gate
with multiple reactive walls in series
Important observations related to the performance of passive/reactive
treatment technology are:
•
It is limited to a subsurface lithology that has a continuous aquitard
at a depth that is within the vertical limits of the trenching
equipment.
•
Passive treatment walls have a tendency to lose their reactive
capacity over time, and require replacement of the reactive
medium.
•
Large and deep plumes are more difficult to remediate than small
and shallow plumes. The complete cost of using treatment walls to
remediate contaminated groundwater is not available. However, the
cost is believed to be dependent on the reactive media and the
contaminant concentration in the groundwater.
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Chapter 2 Biodegradation Technologies for Remediation of Contaminated Sites
Figure 2.3. Permeable reactive barrier.
Biosparging and bioslurping. Biosparging (similar to air sparging)
involves the injection of a gas (usually air or oxygen) and occasionally gas-phase
nutrients, under pressure, into the saturated zone to promote aerobic
biodegradation. In air sparging, volatile contaminants also can be removed from
the saturated zone by desorption and volatilization into the air stream. Emphasis
on the biological degradation rate over physical removal, as well as lower rates of
air injection, are what distinguishes this technology from air sparging.
Typically, biosparging is achieved by injecting air into a contaminated
subsurface formation through a specially designed series of injection wells. The
air creates an inverted cone of partially aerated soils surrounding the injection
point. The air displaces pore water, volatilizes contaminants, and exits the
saturated zone into the unsaturated zone. While in contact with groundwater,
oxygen dissolution from the air into the groundwater is facilitated and supports
aerobic biodegradation.
A number of contaminants have been successfully addressed with
biosparging technology, including gasoline components such as benzene, toluene,
ethylbenzene, and xylenes (BTEX) and SVOCs. Biosparging is most often
recommended at sites impacted with mid-weight petroleum hydrocarbon
contaminants, such as diesel and jet fuels. Lighter contaminants, such as gasoline,
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Chapter 2 Biodegradation Technologies for Remediation of Contaminated Sites
tend to be easily mobilized into the unsaturated zone and physically removed.
Heavier contaminants, such as oils, require longer remedial intervals because of
reduced microbial bioavailability with increasing carbon chain length (EPA,
2004b). Care must be taken to determine whether contaminant concentrations in
soil gas and released vapors resulting from biosparging require treatment. For this
reason, biosparging may be implemented along with SVE or bioventing as a
remedy for increased contaminant concentrations in the unsaturated zone. The
SVE wells are designed to capture the introduced air and contaminant vapors
(EPA, 2004b). Figure 2.4 depicts a typical biosparging system with optional SVE
system. Alternatively, a lower-flow bioventing system may be added to facilitate
bioremediation of volatilized contaminants in the vadose zone.
One specialized form of biosparging involves the injection of organic
gases into the saturated zone to induce cometabolic biodegradation of chlorinated
aliphatic hydrocarbons (analogous to cometabolic bioventing). The injection of
gases below the water table distinguishes biosparging from bioventing. In contrast
to cometabolic bioventing, the solubility of organic gases in water limits delivery
of the primary substrate during cometabolic biosparging applications.
Figure 2.4. Biosparging system.
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Chapter 2 Biodegradation Technologies for Remediation of Contaminated Sites
Bioslurping (also known as multi-phase extraction) is effective in
removing free product that is floating on the water table (Battelle, 1997).
Bioslurping combines the two remedial approaches of bioventing and vacuumenhanced free-product recovery. Bioventing stimulates aerobic bioremediation of
contaminated soils in situ, while vacuum-enhanced free-product recovery extracts
light, nonaqueous-phase liquids (LNAPLs) from the capillary fringe and the water
table. A bioslurping tube with adjustable height is lowered into a ground water
well and installed within a screened portion at the water table (see Figure 2.5). A
vacuum is applied to the bioslurping tube and free product is “slurped” up the tube
into a trap or oil water separator for further treatment. Removal of the LNAPL
results in a decline in the LNAPL elevation, which in turn promotes LNAPL flow
from outlying areas toward the bioslurping well. As the fluid level in the
bioslurping well declines in response to vacuum extraction of LNAPL, the
bioslurping tube also begins to extract vapors from the unsaturated zone. This
vapor extraction promotes soil gas movement, which in turn increases aeration
and enhances aerobic biodegradation (Miller, 1996).
Figure 2.5. Bioslurping technology.
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Chapter 2 Biodegradation Technologies for Remediation of Contaminated Sites
2.3 EX SITU BIOREMEDIATION
Ex situ bioremediation technologies can most easily be classified by the
physical state of the medium to which they are typically applied: solids, solid–
liquid mixtures, and liquids. Also common to the ex situ remediation technologies
are the processes for removing contaminated materials for treatment.
Contaminated media are excavated or extracted (e.g., ground water removal by
pumping) and moved to the process location, which may be within or adjacent to
the contamination zone.
2.3.1 Solids
The most common types of solids bioremediation are (1) land farming or
land treatment, (2) composting, and (3) biopiles, cells, or mounds.
2.3.1.1 Land Treatment
Land treatment, also called land farming, is useful in treating aerobically
degradable contaminants. This process is suitable for non-volatile contaminants at
sites where large areas for treatment cells are available. Land treatment of sitecontaminated soil usually entails the tilling of an 8-to 12-inch layer of the soil to
promote aerobic biodegradation of organic contaminants. The soils are
periodically tilled to aerate the soil, and moisture is added when needed. In some
cases, amendments may be added to improve the tilth of the soil, supply nutrients,
moderate pH, or facilitate bioremediation. Typically, full-scale land treatment
would be conducted in a prepared-bed land treatment unit (see Figure 2.6)—an
open, shallow reactor with an impermeable lining on the bottom and sides to
contain leachate, control runoff, and minimize erosion and with a leachate
collection system under the soil layer (EPA, 1993). In some cases, hazardous
wastes (such as highly contaminated soils) or process wastes (such as distillate
residues) may be treated in land treatment units. In these cases, the waste may be
applied to a base soil layer.
The performance of land treatment varies with the contaminants to be
treated. For easily biodegradable contaminants, such as fuels, land treatment is
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Chapter 2 Biodegradation Technologies for Remediation of Contaminated Sites
inexpensive and effective. Contaminants that are difficult to degrade, such as
PAHs, pesticides, or chlorinated organic compounds, are topics of research.
Figure 2.6. Land treatment.
2.3.1.2 Composting
Composting is a controlled biological process that treats organic
contaminants using microorganisms under thermophilic conditions (40°–50°C).
For some practitioners, the creation of thermophilic conditions is the primary
distinction between composting and biopiles (which operate at less than 40°C),
although others use composting as a term that encompasses both temperature
ranges.
In composting, soils are excavated and mixed with bulking agents and
organic amendments, such as wood chips and vegetative wastes, to enhance the
porosity of the mixture to be decomposed. Degradation of the bulking agent heats
up the compost, creating thermophilic conditions. Oxygen content, moisture
levels, and temperatures are monitored and manipulated to optimize degradation.
Oxygen content usually is maintained by frequent mixing, such as daily or weekly
turning of windrows. Surface irrigation often is used to maintain moisture content.
Temperatures are controlled, to a degree, by mixing, irrigation, and air flow, but
are also dependent on the degradability of the bulk material and ambient
conditions.
There are three designs commonly applied for composting:
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Chapter 2 Biodegradation Technologies for Remediation of Contaminated Sites
•
aerated static piles: compost is formed into piles and aerated with
blowers or vacuum pumps;
•
mechanically agitated in-vessel composting: compost is placed in a
reactor vessel, in which it is mixed and aerated;
•
windrow composting: compost is placed in long, low, narrow piles
(i.e., windrows) and periodically mixed with mobile equipment.
Windrow composting is the least expensive method, but has the potential
to emit larger quantities of VOCs. In-vessel composting is generally the most
expensive type, but provides for the best control of VOCs. Aerated static piles,
especially when a vacuum is applied, offer some control of VOCs and are
typically in an intermediate cost range, but will require offgas treatment. Berms
may also be needed to control runoff during composting operations. Runoff may
be managed by retention ponds, provision of a roof, or evaporation.
Composting has been successfully applied to soils and biosolids
contaminated with petroleum hydrocarbons (e.g., fuels, oil, grease), solvents,
chlorophenols, pesticides, herbicides, PAHs, and nitro-aromatic explosives (EPA,
1998b; EPA, 1997; EPA, 2004b). Composting is not likely to be successful for
highly chlorinated substances, such as PCBs, or for substances that are difficult to
degrade biologically (EPA, 1998b).
2.3.1.3 Biopiles
Biopiles involve the mixing of excavated soils with soil amendments, with
the mixture placed in a treatment area that typically includes an impermeable
liner, a leachate collection system, and an aeration system. Biopiles are typically
2–3 meters high, and contaminated soil is often placed on top of treated soil (see
figure 2.7). Moisture, nutrients, heat, pH, and oxygen are controlled to enhance
biodegradation. This technology is most often applied to readily degradable
species, such as petroleum contaminants. Surface drainage and moisture from the
leachate collection system are accumulated, and they may be treated and then
recycled to the contaminated soil. Nutrients (e.g., nitrogen and phosphorus) are
often added to the recycled water. Alkaline or acidic substances may also be
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Chapter 2 Biodegradation Technologies for Remediation of Contaminated Sites
added to the recycled water to modify or stabilize pH to optimize the growth of
select microbes capable of degrading the contaminants of concern.
An air distribution system is buried in the soil as the biopile is constructed.
Oxygen exchange can be achieved utilizing vacuum, forced air, or even natural
draft air flow. Low air flow rates are desirable to minimize contaminant
volatilization. If volatile constituents are present in significant concentrations, the
biopile may require a cover and treatment of the offgas.
Biopile treatment lasts from a few weeks to a few months, depending on
the contaminants present and the design and operational parameters selected for
the biopile. Biopiles are typically mesophilic (10°–45°C).
Figure 2.7. Typical biopile system.
2.3.2 Solid–liquid mixtures
Solid-liquid mixtures consist of materials such as slurries and sludges. One
technology for treating such mixtures is discussed below.
2.3.2.1 Slurry Bioreactors
Slurry bioreactors are utilized for soil, sediments, sludge, and other solid
or semi-solid wastes. Slurry bioreactors are costly and, thus, are likely to be used
for more difficult treatment efforts.
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Chapter 2 Biodegradation Technologies for Remediation of Contaminated Sites
Typically, wastes are screened to remove debris and other large objects,
then mixed with water in a tank or other vessel until solids are suspended in the
liquid phase. If necessary, further particle size reduction can be accomplished
before the addition of water (by pulverizing and/ or screening the wastes) or after
the addition of water (through use of a sheering mixer). Suspension and mixing of
the solids may increase mass transfer rates and may increase contact between
contaminants and microbes capable of degrading those contaminants (EPA,
1990). Mixing occurs in tanks or lined lagoons. Mechanical mixing is generally
conducted in tanks. Typical slurries are 10–30% solids by weight. Aeration, with
submerged aerators or spargers, is frequently used in lagoons and may be
combined with mechanical mixing to achieve the desired results. Nutrients and
other additives, such as neutralizing agents, surfactants, dispersants, and cometabolites (e.g., phenol, pyrene) may be supplied to improve handling
characteristics and microbial degradation rates. Indigenous microbes may be used
or microorganisms may be added initially to seed the bioreactor or may be added
continuously to maintain proper biomass levels. Residence time in the bioreactor
varies with the matrix as well as the type and concentration of contaminant (EPA,
1990).
Once contaminant concentrations reach desired levels on a dry-weight
basis, the slurry is dewatered. Typically, a clarifier is utilized to dewater the slurry
by gravity. Other dewatering equipment may be used depending on slurry
characteristics and cost considerations (Olin et al., 1999). Water, air emissions
from all process steps, and oversize materials may require additional treatment.
2.3.3 Liquids
Liquids, such as surface water, groundwater, mine drainage, and effluent
from other treatment operations, can undergo ex situ bioremediation in
constructed wetlands. Note that surface water and groundwater have important
differences, such as concentrations of contaminants and degradable organic
material, than may be found in waste streams from other treatment operations.
2.3.3.1 Constructed wetlands
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Chapter 2 Biodegradation Technologies for Remediation of Contaminated Sites
Constructed wetlands provide for biological assimilation, breakdown, and
transformation of contaminants; chemical breakdown and transformation of
contaminants; and physical sedimentation and filtration (USDA and EPA 1994a),
as shown in figure 2.8. Biological processes associated with wetlands include
bioremediation (microbially-based remediation) and phytoremediation (plantbased remediation). Microbes attached to the surfaces of plants, plant litter, and
the wetland substrate degrade and/or sorb the organic substances present in the
water undergoing treatment (USDA and EPA, 1994a). Phytoremediation uses
plants to remove, transfer, stabilize, or destroy contaminants through biological,
chemical, and physical processes that are influenced by plants and their roots (i.e.,
rhizosphere) that include degradation, extraction through accumulation in plant
roots/shoots/leaves, metabolism of contaminants, and immobilization of
contaminants at the interface of roots and soil (EPA, 2004a).
Wetlands inherently have a higher rate of biological productivity/activity
than many other natural ecosystems and are thus capable of efficiently and
economically transforming many common contaminants to harmless byproducts
(Kadlec and Knight, 1996). Constructed wetlands have been applied successfully
to remove contaminants such as metals, petroleum hydrocarbons, and glycols; to
decrease metal concentrations via chemical or microbial precipitation; and to
neutralize acidity. Recent research also has demonstrated applicability to
explosive-contaminated water (Bader, 1999). However, wetlands are sensitive to
high ammonia levels, herbicides, and contaminants that are toxic to the plants or
microbes.
Constructed wetlands are well suited for the treatment of contaminated
groundwater emerging from surface and mine seeps, pump-and-treat waste
streams with low concentrations of easily biodegradable contaminants, and
contaminated surface waters (EPA, 2001c). Constructed wetlands may also be
used to pretreat contaminated water prior to conventional treatment or to further
treat a waste stream prior to disposition or discharge (USDA and EPA, 1994b).
However, applicability to highly acidic waste streams may not be cost-effective
(USDA and EPA, 1994b). Discharges must meet applicable effluent limitations
and related regulatory requirements. Discharges that do not meet these
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Chapter 2 Biodegradation Technologies for Remediation of Contaminated Sites
requirements may be required to undergo further treatment or may be found
suitable for recycling into the wetland as a supplemental water source.
There are various types of constructed wetlands, depending on the type of
flow (surface or subsurface), contaminant of concern, or type of substrate, which
can include limestone, organic material such as compost, or gravel. The chemical
and microbial processes may proceed either in an anaerobic or aerobic
environment.
Since constructed wetlands function both as macroscopic and microscopic
ecosystems to promote contaminant treatment, the biological characteristics of the
system must be taken into account during the design phase. The chemistry of the
waste stream and how the passive chemical, physical, and biological processes
affect this or are, in turn, affected by the waste stream are important design
factors. The chemical characteristics of the waste stream can affect sizing of the
system for adequate retention time and whether the waste stream may require
pretreatment to (1) address concentration, ammonia, nutrient, and organic loads
that may damage vegetation, or (2) remove solids or materials, such as grease, that
may clog the wetland (USDA and EPA, 1994a). In addition, pH adjustment may
be necessary, either prior to waste stream treatment or through use of limestone
substrate (USDA and EPA, 1994b). Climatic and seasonal circumstances as well
as waste stream characteristics are important considerations when selecting the
types of plants to use in a constructed wetland. Salinity, either in the waste stream
or as a result of treatment, can harm or destroy the wetland vegetation if the plants
are not salt tolerant. In addition, cold weather can reduce microbial activity, and
hail or other weather events can damage the plants (USDA and EPA, 1994a).
The low cost, passivity (i.e., lack of dependence on power or mechanical
components), and efficacy for treating many common contaminants are key
advantages of constructed wetland treatment systems. Constructed wetlands are
often visually attractive, but can require more space than other remedial systems.
The wetlands should be sized with an understanding that both plant-based and
bacterial-based remediation will decline during colder seasons. A key design
element is sizing to achieve adequate retention time to enable the biological,
chemical, and physical processes to be effective (USDA and EPA, 1994a).
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Chapter 2 Biodegradation Technologies for Remediation of Contaminated Sites
Seasonal, climatological, and waste stream factors that control the water balance
in the wetland also must be considered during design to achieve project goals.
Constructed wetlands require a continuous supply of water. While tolerant
of fluctuating flows, constructed wetlands cannot withstand complete drying. A
slow water flow must be maintained to prevent the development of stagnant water
that can lead to performance and vector difficulties. Recycling wetland water can
supplement inflow, but this can increase salinity over time, which can affect
design and cost (USDA and EPA, 1994a).
Figure 2.8. Constructed wetland.
75
CHAPTER 3
3. KINETIC MODELS FOR COMETABOLISM
____________________________________________________________________________________________________
Cometabolic biotransformation models most often stem from Michaelis
Menten and Monod enzyme kinetics. These expressions have been expanded to
include processes such as substrate inhibition (Broholm at al., 1992; Ely et al.,
1995a; and Kim et al., 2002a), product toxicity (Chang and Criddle, 1997;
Alvarez-Cohen and McCarty, 1991; Kim et al., 2000) and reducing energy
limitations (Chang and Alvarez-Cohen, 1995a; Sipkema et al., 2000). AlvarezCohen and Speitel (2001) provided a review and discussion of these processes and
the models representing them.
3.1 ENZYMATIC REACTIONS: MICHAELIS-MENTEN KINETICS
In 1913 Michaelis and Menten proposed a kinetic model to describe
enzyme catalyzed reactions: with some simplifications, this model can be adopted
to simulate most of the reactions occurring in the biological systems with a
limited number of parameters. The model is based on the hypothesis that the free
enzyme (E) and ths substrate (S) bind in an activated complex (ES), generating
the product (P) and the enzyme (E):
k1
k2
E + S ⇔ ES → E + P
k-1
The parameters k1, k2, k-1 represent the reaction constants; the first reaction is
assumed at equilibrium, while the second is considered irreversible.
Chapter 3 Kinetic Models for Cometabolism
Assuming that the process of formation and disruption of the
activatedcomplex follow a second order and a first order kinetics respectively, the
reaction rates can be expressed by thr following equations:
(3.1)
(3.2)
where the expressions in the square brackets represent the concentrations of the
corresponding reagents.
The enzyme mass balance assumes the following form, where E 0 is the
total enzyme concentration:
[E] + [ES] = E0 = const
(3.3)
The system can be analitically solved with the hypothesis that the mass of
the activated complex is conservative, as expressed by the following equation:
(3.4)
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Chapter 2 Biodegradation Technologies for Remediation of Contaminated Sites
This assumption, known as quasi-stationary state approximation, is
correct if the ratio of the total enzyme concentration E0 to the initial substrate
concentration S0 is sufficiently low. This condition is satisfied in most of the
practical applications, with the exception of a short initial transitory state. With
these hypotheses, the analytical solution for the product formation rate,
corresponding to the substrate degradation rate, assumes the form:
(3.5)
where KS, known as “affinity constant” or “half saturation constant, is defined:
(3.6)
For the application of this model to the microbial mediated reactions , [E0]
can be assumed as a constant among different microorganisms; thus, [E0] is
proportional to the microbial concentration X. The substrate degradation velocity
can be rewritten, omitting the brackets for simplicity, as:
(3.7)
The substrate specific degradation velocity per unit of biomass, also
known as “degradation rate” (qs = r s / X), is expressed by the following equation:
(3.8)
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Chapter 2 Biodegradation Technologies for Remediation of Contaminated Sites
The parameter kmax,s accounts for both the constant k2 and the factor of
proportionality between E0 and X; this constant is generally recognized as
“maximum specific degradation rate” of substrate consumption, and corresponds
to the maximum degradation velocity that can be achieved by a unitary mass of
microorganisms. This condition, expressed by Eq. 2.9, is verified when the
affinity constant Ks can be neglected in comparison to the substrate concentration
S; in this concentration range, the substrate degradation follow a zero order
kinetic:
(3.9)
The kinetic parameters are specific for the microbial culture and the
substrate. When the degradation is sustained by a microbial consortium, the
kinetic constants kmax and Ks are obtained from a weighed average of the
characteristic values of each species.
The following equation represents the microbial growth (mg L−1 d−1)
according to the Monod model, and introduces the cell growth yield Y (mg
cell/mg substrate) and the cell decay coefficient b (d-1), representing the
endogenous cell inactivation:
(3.10)
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Chapter 2 Biodegradation Technologies for Remediation of Contaminated Sites
3.2 SUBSTRATE INHIBITION
Microorganisms capable of transforming CAHs through aerobic
cometabolism have catabolic oxygenases that catalyze the initial oxidation step of
their respective primary growth substrates and have potential for initiating the
oxidation of CAHs. The oxygenases are often non specific and fortuitously initiate
oxidation of a variety of compounds including most of the CAHs. In general,
oxygenases act on unsaturated CAHs such as TCE by adding oxygen across the
double bond to form an epoxide. With saturated CAHs such as CF or TCA, a
hydroxyl group is generally substituted for one of the hydrogen atoms in the CAH
molecule. Frequently, the resulting products from CAHs oxidation are chemically
unstable: they decompose yielding products that are further metabolized by other
microorganisms present in nature.
Because a single enzyme is responsible for the oxidation of both types of
substrates, the presence of the growth substrate can inhibit the oxidation rate of
the non beneficial substrate and vice versa. Substrate inhibition describes the
hindrance of substrate transformation or utilization due to the competition for, or
alteration of degradative enzymes. There are several types of inhibition, including
self , competitive, noncompetitive, and mixed-inhibition. Self inhibition may
result when the growth substrate itself is inhibitory at high concentrations. When
an enzyme lacks specificity, competitive inhibition may occur in which one
substrate binds to the catalytic site of the enzyme, thus preventing another
substrate from reacting. A substrate may also bind to a non-reactive site on the
enzyme, altering its conformation and creating noncompetitive inhibition which
reduces the utilization of another substrate. Competitive and noncompetitive
inhibition may occur simultaneously, causing a condition termed mixed inhibition
(Rittman and McCarty, 2001). Competition between the growth substrate and the
cometabolic substrate for oxygenase enzymes may significantly affect
cometabolic degradation rates.
A single enzyme may thus be able to catalyze the degradation of two or
more substrates; this scenario, which includes the cometabolic processes, can be
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Chapter 2 Biodegradation Technologies for Remediation of Contaminated Sites
described by the following reaction scheme, where A and B represent distinct
substrates:
(3.11)
With the same assumptions as presented in the case of one substrate, the
following degradation rates are obtained for the two substrates:
The half-saturation portion of each equation becomes a function of the
inhibitor competitive inhibition constant (Kc,A and Kc,B respectively). This model is
known as “competitive inhibition” and describes the most frequent inhibition type
included in mathematical modeling of cometabolic biotransformations (Broholm
et al., 1992; Chang and Alvarez-Cohen, 1995; Chang and Criddle, 1997; Lee et
al., 2000). The competitive inhibition constant has often been approximated with
the competing substrate half-saturation constant. Kim et al. (2002), however,
noted various studies where this appeared to be an incorrect assumption and, in
response, presented a method for determining the inhibition type and the
respective constants. Kim focused on butane utilization by a mixed culture with
cometabolic transformation of 1,1-DCE, 1,1-DCA, and 1,1,1-TCA and observed
that competitive and mixed inhibition occurred. CAHs competitively inhibited the
81
Chapter 2 Biodegradation Technologies for Remediation of Contaminated Sites
degradation of the other CAHs and butane, while butane showed mixed inhibitory
effects toward the CAHs.
Noncompetitive inhibition more specifically influences the maximum
degradation rate, and equation 2.7 may be transformed to:
(3.12)
where Iu = aqueous concentration of noncompetitive inhibitor (mg/L)
KI,u = constant for noncompetitive inhibition (mg inhibitor/L)
In the case of mixed inhibition, the equation assumes a combined form of 2.11
and 2.12, resulting in:
(3.13)
Competitive and non competitive inhibition may or may not be caused by
the same inhibitor. Terms for competitive and non competitive inhibition are
additive and equation 2.13 may be extended to include several inhibitors.
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Chapter 2 Biodegradation Technologies for Remediation of Contaminated Sites
3.3 CAHs DEGRADATION PRODUCT TOXICITY
The oxidation of chlorinated organics by oxygenase enzymes generates
short lived toxic intermediate products that may damage cells, causing cellular
inactivation: this phenomenon is known as “product toxicity”. Alvarez-Cohen and
McCarty verified that some oxidation products of chlorinated aliphatics, such as
fosgene in the oxidation of chloroform, and epoxides in oxidation of chlorinated
ethenes, irreversibly bind to proteins and lipids, thus inactivating some cellular
functions.
The capability of microbial cultures to oxidize compounds which exert
product toxicity can be quantified using the “transformation capacity” (TC)
parameter, defined as the maximum mass of solvent that can be transformed by a
given amount of cells before they are completely inactivated.
In the cometabolic processes, after the initial oxidation step, growth
substrates are further degraded to regenerate reducing energy (NADH), which
promotes more substrate oxidation. In the absence of growth substrates, methane
oxidizers are capable of using both internal energy sources, such as polyhydroxylbutirate (PHB), and methane catabolic intermediates, such as formate, to
regenerate NADH; oxidation of non beneficial substrates can be carried out in the
absence of growth substrate as long as some source of NADH regeneration is
available. As oxygenase expressing cultures, such as propane and phenol
oxidizers, may also exhibit similar responses since they have similar enzyme
mechanisms. However, the oxidation of non beneficial substrates in the absence
of growth substrates can cause the depletion of NADH in cells, since NADH is
not regenerated. Organisms in the absence of growth subtrate are referred as
“resting cells”.
When cometabolic substratessuch as chlorinated organics are oxidized by
resting cells, the degradation may be limited by both the depletion of endogenous
cellular reducing energy and the product toxicity; it follows that T C measured in
these conditions is a function of both NADH level and toxicity. If the external
reducing energy runs out before the cells are completely inactivated by toxicity,
the reaction is interupted because of the absence of NADH, resulting in an
artificially low TC.
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Chapter 2 Biodegradation Technologies for Remediation of Contaminated Sites
To estimate the effects of toxicity alone, TC should be measured in the
presence of a NADH regenerant; a comparison of TC measured in the two
conditions may reflect the amount of endogenous reducing energy available to
cells. However some NADH regenerant may exert additional effect on cells which
may also bias the measured TC: for example, growth substrates would promote
additional enzyme production, whereas other amendments may be toxic
themselves.
Chang and Alvarez-Cohen (1995b) tested the effect of amendments on the
tranformation capacities of four oxidizing cultures. The T C values of chlorinated
organics could be significantly increased by the addition of low concentrations of
growth substrates; this effect was overcome at higher concentrations for two of
the cultures, pressumably by toxicity of the growth substrates. No better results
were obtained providing as amendments catabolic intermediates of the growth
substrate. These results suggest that although TC may be a good tool for the
comparison of the toxic effects of chlorinated organic degradations, care must be
taken to minimize the effects of reducing energy limitations and amendment
interference. The measurement of TC in the presence of growth substrate avoids
errors due to energy limitations, but the potential for confounding factors such as
substrate toxicity and enzyme regeneration should be cosidered. The use of
nontoxic NADH regenerant which is not a growth substrate may result in
TC.measurements which more directly reflect the effect of degradation toxicity.
Further research performed by Chang and Alvarez-Cohen gave deeper
insight into the phenomenon of toxicity:
•
All the experiments demonstrated that toxicity is caused by the
degradation products rather than the solvent themselves; this was
verified in batch microcosms with four CAHs and three different
substrates. In the first phase, the solvent degradation was inhibited,
thus exposing the cells to the solvent in the absence of their
degradation products; in the second phase the solvent were
stripped. The growth substrate degradation was evaluated prior and
after the exposure to the solvent: the measured value did not differ
significantly;
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Chapter 2 Biodegradation Technologies for Remediation of Contaminated Sites
•
the transformation capacity of a given chlorinated solvent changes
with the primary substrate used for growing the culture; thus TC is a
good parameter for comparing the potential of different substrates
to address a specific contaminant;
•
in mixed bacterial consortia, the transformation capacity may
change in time, following shifts in the composition of the microbial
population;
•
the transformation capacity relative to a specific chlorinated
solvent and a growth substrate is independent from the initial
concentrations of both biomass and contaminant;
•
the transformation capacity of cells grown on a specific substrate
decreases with the chlorine- to carbon- atoms numerical ratio in the
solvent molecule;
•
the transformation yield Ty is proportional to the transformation
capacity TC through the growth yield Y: Ty = Tc x Y.
Alvarez-Cohen and Speitel (2001) reviewed the interpretations for approximating
inactivation, separating them into two classes. One class (1) represents loss of full
cellular function, while the other class (2) assumes the loss of specific enzyme
activity. Among the second class models, important contributions were provided
by Ely in 1995. The model incorporates enzyme inhibition, caused by the
presence of a cometabolic compound, inactivation, resulting from toxicity of a
cometabolic product, and recovery associated with bacterial sinthesis of new
enzyme in response to inactivation. The first class is the most commonly used in
modelling biodegradation.
In 1991 Alvarez-Cohen and McCarty proposed the kinetic model
introducing the transformation capacity in order to account for the toxic effects of
the degradation products on the biomass. The transformation capacity Tc was thus
defined as the quantity of a compound that a specific mass of microorganisms can
degrade before it is inactivated by toxicity fromtransformation products. Units of
transformation capacities are tipically mass of degraded substrate per mass of cell.
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Chapter 2 Biodegradation Technologies for Remediation of Contaminated Sites
Assuming that the degradation of the chlorinated solvents follows the
Michaelis-Menten kinetics, as described in 3.1, and that the biomass concentration
decreases in time as a consequence of both product toxicity and endogenous
decay, the following model can be applied in the absence of growth substrate
(Alvarez-Cohen and McCarty, 1991):
(3.14)
where:
rx (mg protein/L/h), cell decay rate
rc (g/mg protein/h), contaminant degradation rate
b (h-1), cell endogenous decay coefficient
Tc (mg/mg), contaminant transformation capacity
X (mg protein/L), specialized cell concentration
Successive studies (Chang and Alvarez, 1995a, b) evidenced that the
oxidation of chlorinated organics by resting cells is also limited by the depletion
of reducing energy. Cometabolic degradation rates were observed to increase with
the addition of external energy sourcrs; on the other hand, the degradation
performances were found to be affected by substrate inhibition, when the external
energy was provided through the addition of growth substrate.
A modification of Michaelis-Menten/Monod kinetics was proposed to
describe the kinetics of cometabolic degradation, incorporating the effects of
product toxicity, depletion of oxygen and reducing energy, competition between
growth substrate and cometabolic substrate. The model, summarized in the
equations 3.15 – 3.17, was able to predict the experimental results. The factors
R/(R+KR) and O2/(KO2+ O2), included in the growth substrate and contaminant
degradation rate equations, take into account oxygen and reducing energy sources
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Chapter 2 Biodegradation Technologies for Remediation of Contaminated Sites
as possibly limiting reactants. When they are supplied in large excess, such that R
>> KR and O2 >> KO2, the corresponding Monod terms approach 1 and the specific
degradation rates become functions of substrate, contaminant and biomass
concentration alone.
Where:
87
CHAPTER 4
4. MOLECULAR BIOLOGY TECHNIQUES USED IN THE ANALYSIS OF
THE MICROBIAL DIVERSITY IN CONTAMINATED SOILS
____________________________________________________________________________________________________
Identification of microorganisms by conventional methods requires the
isolation of pure cultures followed by laborious characterization experiments.
These procedures are therefore inadequate for study of the diversity of a natural or
engineered ecosystem. A new set of molecular techniques developed during the
1990s revolutionized microbial ecology research. Among these techniques,
cloning and the creation of a gene library, denaturant gradient gel electrophoresis
(DGGE) and fluorescent in situ hybridization with DNA probes (FISH) stand out.
Cloning provides very precise taxonomical information, but it is time consuming
and requires specialized personnel whereas DGGE is a rapid and simple method
that provide characteristic band patterns for different samples, allowing quick
sample profiling, while retaining the possibility of a more thorough genetic
analysis by sequencing of particular bands. FISH makes possible to identify
microorganisms at any desired taxonomical level, depending on the specificity of
the probe used. It is the only quantitative molecular biology technique, although
quantification is either complex or tedious and subjective. Combination with a
confocal laser-scanning microscope allows the visualization of three-dimensional
microbial structures. These methods have deepened our understanding of the
microbiology of contaminated soils. Both DGGE and FISH have been extensively
employed.
Chapter 4 Molecular Techniques in The Analysis of Microbial Diversity in Contaminated Soils
4.1 MICROBIAL ECOLOGY OF CONTAMINATED SOILS
4.1.1 Phenotypic analysis of soil bacterial communities
The diversity of bacterial communities in contaminated soils has been
based in the 1980-1990’s on CFU counting and colony morphology typing.
Colony morphology typing relies on grouping of bacterial colonies cultured on
plates containing relevant media according to their colony appearance (Haldeman
and Amy, 1993). Latest years, community-level physiological profiles (CLPPs) or
sole-carbon-source utilization profiles have been used as an indicator of
community structure and function (Becker and Stottmeister, 1998; Degens, 1998;
Gamo and Shoji, 1999; Garland and Mills, 1991; Garland and Mills, 1994; Smalla
et al., 1998; Wünsche et al., 1995). CLPP is based on metabolic response patterns
of communities extracted from environmental samples, inoculated into a 96 wells
BIOLOG plate. The 96-well microtiter plates contain nutrients and a tetrazolium
dye. When a bacterial community is capable of oxidizing the carbon substrate, the
dye turns purple, and a spectrophotometric plate reader quantifies the response.
However, these patterns not always reflect the organisms directly involved in the
mainstream energy flux of the ecosystem (Boon, 2002). Wünsche et al. (1995)
applied these techniques to examine the effect of hydrocarbon contamination on
microbial community structure and function and found that characteristic shifts of
the substrate utilization patterns followed changes in hydrocarbon content in soils.
Furthermore, the altered patterns of substrate utilization corresponded to similar
changes in abundance of hydrocarbon-utilizing bacteria determined by plate
counts. Strong-Gunderson and Palumbo (1994) adapted this CLPP method for
rapidly screening the metabolic potential of bacteria to oxidize semi-volatile and
volatile compounds as a sole carbon source.
Although the CLPP assay has been proposed as a measure of functional
diversity, assay responses are attributed mainly to a small subset of heterotrophic
bacteria in the tested environmental sample (Ibekwe et al., 2001). Difficulties in
analyzing the complexity of bacterial communities by classic methods of
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Chapter 4 Molecular Techniques in The Analysis of Microbial Diversity in Contaminated Soils
cultivation and subsequent physiological characterization have necessitated the
development of new approaches for community and functional analysis.
4.1.2 Culture-independent analysis of microbial communities by lipid-based
tools
White and Findlay (1988) developed a community-level approach to
characterize microbial community structure by evaluating shifts in phospholipid
fatty acids (PLFA) from environmental samples. Different groups of bacteria are
characterized by specific PLFA profiles. Therefore, a change in the phospholipid
pattern in soil would indicate a change in the bacterial composition of that soil.
The polar lipid fraction of environmental samples is composed primarily of PLFA
of “viable” micro-organisms present in the sample (assuming rapid degradation of
intact phospholipids after cell death) (Ringelberg et al., 2001; White and Findlay,
1988). For complex matrices such as soil, PLFA analysis has been shown to be a
valuable tool for detecting changes in microbial communities in response to
pollution of alkanes (Ringelberg et al., 1989) and chlorinated hydrocarbons
(Phelps et al., 1988). Recently, analysis of
13
C-labeled PLFAs resulting from
incorporation of 13C during cell growth from 13C-labeled C substrates has been
used to define the groups of organisms utilizing those substrates (Boschker et al.,
1998; Padmanabhan et al., 2003). Therefore, 13C-labeled substrates are introduced
into the soil where bacteria use them as a C-source and after PLFA extraction the
active micro-organisms are identified by analyzing 13C-labeled PLFA. Hanson et
al. (1999) identified the indigenous population(s) responsible for toluene
degradation in Yolo silt loam, by employing both traditional culture-based
approaches and PLFA and 13C-PLFA analysis. After 119 h of incubation with 13Ctoluene, 96% of the incorporated
13
C was detected in only 16 of the total
59 PLFAs (27%) extracted from the soil. Of the total 13C-enriched PLFAs, 85%
were identical to the PLFAs contained in a toluene-metabolizing bacterium
isolated from the same soil, showing that this strain was one of the main toluene
degraders in that soil. In contrast, the majority of the soil PLFAs (91%) became
labeled when the same soil was incubated with
90
13
C-glucose. In laboratory
Chapter 4 Molecular Techniques in The Analysis of Microbial Diversity in Contaminated Soils
microcosms, Pelz et al. (2001a, 2001b) incubated sediments from a petroleum
hydrocarbon (PHC) contaminated aquifer and a nearby pristine aquifer under
anoxic sulfate-reducing conditions with methyl-14C-toluene to determine the 14Cmass balances and with methyl-13C-toluene to follow the flow of carbon from
toluene into PLFA.
14
C quantification revealed that 61.6% of the methyl-14C-
toluene was mineralized and 2.7% was assimilated, while
13
C-labeled PLFA
analysis linked toluene degradation to the metabolic activity of Desulfobacter-like
populations. These populations could play an important role in the clean-up of
aromatic PHC contaminated aquifers.
4.1.3 Culture-independent analysis of the bacterial communities by nucleic
acid-based tools
Conventional microbiological techniques, based on isolation of pure
cultures and morphological, metabolic, biochemical and genetic assays, have
provided extensive information on the biodiversity of microbial communities in
natural and engineering systems. However the drawbacks of the existing
conventional methods, such as incomplete knowledge about their physiological
needs and the complex syntrophic and symbiontic relations, which are abundant in
nature, make impossible to obtain pure cultures of most microorganisms in natural
environments. Moreover, most culture media tend to favor the growth of certain
groupso of microorganisms, whereas others that are important in the original
sample do not proliferate. It is therefore generally accepted nowadays that the
number of known prokaryotic species (including the two domains Bacteria and
Archaea) is very small compared to the diversity of microorganisms and
illustrates how difficult it is to get a full picture of the bacterial diversity of an
ecosystem by relying only on conventional methodology. At present, about 7000
bacterial species have been described, but according to molecular and ecological
estimates, the real number must be several order of magnitude higher (Amman et
al, 1995). This small known fraction does not reflect the composition and
diversity of a microbial community.
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Chapter 4 Molecular Techniques in The Analysis of Microbial Diversity in Contaminated Soils
One suitable solution to this problem is to use molecular biology
approaches. The techniques are based on the RNA of the small ribosomial subunit
(16SrRNA for prokaryotes) or their corresponding genes, considering it as a
“molecular clock” or “evolutionary chronometer”. This molecule was chosen
because of its universality and abundance in al living beings (103 to 105
ribosomes/cell) and the fact that it is a highly conserved molecule throughout
evolution although bears some highly variable regions. These features allow
comparison of organisms within the same domain, as well as differentiation of
strains of the same species. Moreover, the gene sequence is sufficiently long to
generate statistically relevant data and can be easily sequenced with current
technology.
As a consequence of the necessity to also address non-culturable members
of the bacterial community, more and more molecular gene probe methods have
been developed that are based on the analysis of nucleic acids extracted from soil
(Akkermans et al., 1995; Amann et al., 1995; Holben and Harris, 1995; Sayler and
Layton, 1990; Shi et al., 1999; Stapleton et al., 1998; Trevors and van Elsas,
1995). The development of methods for directly extracting DNA from
environmental samples bypassed the need to culture organisms, thus providing a
more representative sampling of microbial constituents within a complex
community (Akkermans et al., 1995; Leahy and Colwell, 1990; Shi et al., 1999;
Stapleton et al., 1998; Trevors, 1992; Trevors and van Elsas, 1995).
Cloning and sequencing of the gene that codes for 16S rRNA is still the
most widely used molecular technique in the field of microbial ecology. This
methodology implies the extraction of nucleic acids, amplification and cloning of
the 16S rRNA genes, followed by sequencing and, finally, identification and
affiliation of the isolated clone with the aid of phylogenetic software. While
amplicons generated from pure cultures of bacteria could be sequenced directly, in
the case of genomic DNA extracts from microbial communities, the cloning step
has to be included. This is necessary in order to separate the different copies of
16SrDNA, as a mixed template cannot be sequenced. Because this aproach is so
widwspread, half of the approximately 240000 sequences deposited in the 16S
rDNA NCBI-database (April 2006), belong to non-cultured and unknown
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Chapter 4 Molecular Techniques in The Analysis of Microbial Diversity in Contaminated Soils
organisms, that is, organisms detected by 16S rDNA cloning. This illustrates how
extensively and succesfully the cloning strategy has been employed since its
introduction in the beginning of the 1990s (Ward et al, 1990). However cloning is
time consuming and so less apt for analyzing larger sets of samples, for example,
when monitoring changes in natural or engineered microbial communities over
time, particularly if several time points are required. The main advantages and
disadvantages of thi approach can be summarized as follows:
•
Advantages:
−
complete 16S rRNA sequencing allows:
very precise taxonomic studies and phylogenetic trees of high
resolution to be obtained;
design of primers (for PCR) and probes (for FISH);
−
if time and effort is not a limiting factor, the approach cover most
microorganisms, including minority groups, which would be hard to detect
with genetic fingerprinting methods;
−
identification of microorganisms that have not been yet cultured or
identified.
•
Disadvantages:
−
very time consuming and laborious, making it unpractical for high samples
throughput;
−
extraction of a DNA pool representative of the microbial community can
bedifficult when working with certain sample types (e.g. soil, sediments).
−
many clones have to be sequenced to ensure most of individual species in
the sample are covered;
−
it is not quantitative. The PCR step can favor certain species due to
differences in DNA target site accessibility.
Denaturing gel electrophoresis (DGGE) is based on the different
mobility on a gel of denaturated DNA-fragment of the same size but with
different nucleic acid sequences, thus generating band patterns that directly reflect
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Chapter 4 Molecular Techniques in The Analysis of Microbial Diversity in Contaminated Soils
the genetic biodiversity of the sample. The number of bands corresponds to the
number of dominant species. Coupled with sequencing and phylogenetic analysis
of the bands, this method can give a good overview of the composition of a given
microbial community. DGGE is the method of choice when the desired
information does not have to be as phylogenetically exhaustive as that provided
by cloning, but still relatively precise to determine the dominant members of a
microbial community with medium phylogenetic resolution.
The most important application of DGGE is monitoring dynamic changes
in microbial communities, especially when many samples have to be processed.
For community analysis, the variable areas of the 16S rRNA gene are often used
as bioindicator (Maidak et al., 1997; Stackebrandt et al., 1993; Woese, 1987). A
great number of 16S rRNA gene sequences are accessible via databases and
sequence comparisons allows the identification of areas with unique sequences for
specific bacterial groups (Maidak et al., 1997; van Elsas et al., 1998).
As such, primers can be designed for PCR detection of specific groups of
bacteria by amplifying the corresponding 16S rRNA gene. The obtained PCR
fragments can then be cloned and sequenced (Amann et al., 1995; Hugenholtz and
Pace, 1996) or they can be separated and visualized by fingerprinting techniques
allowing direct diversity analysis (Dejonghe et al., 2001). Community fingerprints
are generated by separating the amplified nucleic acid fragments based on their
sequence variability as in Denaturing Gradient Gel Electrophoresis/Temperature
Gradient Gel Electrophoresis (DGGE/TGGE) (Muyzer et al., 1993) and Single
Strand Conformation Polymorphism (SSCP) analysis (Lee et al., 1996), or by
their size as in Terminal Restriction Fragment Length Polymorphism (T-RFLP)
(Liu et al., 1997) and Amplified Ribosomal DNA Restriction Analysis (ARDRA)
(Massol-Deya et al., 1995). Using DGGE/TGGE, Muyzer et al. (1993) showed
that it is possible to identify constituents of the microbial population which
represent only 1% of the total population. There are unfortunately some constrains
related to these techniques, i.e., the occurrence of multiple bands relating to one
bacterium (e.g. multiple 16S rDNA sequences) or one band corresponding to
more than one strain. Ralebitso et al. (2000) enriched and isolated in the presence
of different selection pressures, particularly based on pH and electron donor
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Chapter 4 Molecular Techniques in The Analysis of Microbial Diversity in Contaminated Soils
concentration, indigenous microbial associations which catabolize selected
petroleum hydrocarbon components (benzene, toluene and o-, m- and p-xylene
(BTX), from a petroleum hydrocarbon contaminated sandy soil. PCR and 16S
rDNA fingerprinting by DGGE were employed to explore the diversities and
analyse the structures of the isolated microbial associations. Pearson productmoment correlation indicated that different, but chemically similar, petroleum
hydrocarbon molecules, effected the isolation of different associations. However,
some similar numerically-dominant bands characterized the associations. A 30%
similarity was evident between the m- and o-xylene catabolizing associations
regardless of the molecule concentration and the enrichment pH. PCR-DGGE was
also used to complement conventional culture-based microbiological procedures
for environmental parameter optimisation. Band pattern differences indicated
profile variations of the isolated associations, which possibly accounted for the
growth rate changes recorded in response to pH and temperature perturbations.
Currently, no reports are available concerning TGGE, SSCP or T-RFLP
application to follow up the microbial diversity at petroleum hydrocarbon
contaminated sites polluted with prevalently BTEX. Massol-Deya et al. (1997)
used ARDRA to compare community composition, succession, and performance
in fluidized bed reactors (FBR) treating BTX contaminated water. One reactor
was inoculated with the toluene degrading strains P. putida mt-2 (PaW1), B.
cepacia G4, and B. pickettii PKO1. Strain mt-2 was found to outcompete the other
two strains. When groundwater strains were allowed to challenge the steady-state
biofilm developed by the inoculated strains, they readily displaced the inoculated
strains and further reduced the toluene effluent concentration. ARDRA of 16S
rRNA gene amplicons from the reactor community showed a succession of
populations into a pattern that was stable for at least 4 months of operation. The
convergence of communities to the same composition from three different starting
conditions and their constancy over several months suggested that a rather stable
community was selected.
Alternatively, 16S rDNA group specific probes can be designed for
DNA:DNA hybridization targeting 16S rRNA. However, the polymerase chain
reaction technique is much more sensitive (by 3 orders of magnitude), permitting
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Chapter 4 Molecular Techniques in The Analysis of Microbial Diversity in Contaminated Soils
the detection of 1 cell per g of sediment sample (Steffan and Atlas, 1988; Leahy et
al., 1990). To quantify hybridization signals and determine the amount of DNA in
each sample, a regression equation is generated from hybridization signal
intensities of known DNA standards, included on each vacuum blot. Signal
intensities are obtained by computer-aided analysis of autradiogram images (Guo
et al., 1997; Stapleton et al., 1998; Shi et al., 1999). The relative abundance of
domains or subgroups is determined by normalizing hybridization signals to the
signal generated from hybridization to a universal 16S rDNA probe (Zheng et al.,
1996). Shi et al. (1999) used phylogenetic probes in hybridization analysis to (i)
determine in situ microbial community structures in regions of a shallow sand
aquifer that were oxygen depleted and fuel contaminated (FC) or aerobic and noncontaminated (NC) and (ii) examine alterations in microbial community structures
resulting from exposure to toluene and/or electron-acceptor supplementation
(nitrate). The latter objective was addressed by using the NC and FC aquifer
materials for anaerobic microcosm studies in which phylogenetic probe analysis
was complemented by microbial activity assays. Domain probe analysis of the
aquifer samples showed that the communities were predominantly Bacteria,
Eucarya and Archaea were not detectable. At the phylum and subclass levels, the
FC and NC aquifer material showed similar relative abundance distributions of 43
to 65% β- and γ-Proteobacteria (B+G), 31 to 35% α-Proteobacteria (ALF), 15 to
18% sulfate-reducing bacteria, and 5 to 10% high G+C gram positive bacteria.
Compared to that of the NC region, the community structure of the FC material
differed mainly in an increased abundance of B+G relative to that of ALF. The
microcosm communities were similar to those of the field samples. Addition of
nitrate and/or toluene stimulated microbial activity in the microcosms, but only
supplementation of toluene alone significantly altered community structure. For
the NC material, the dominant subclass shifted from B+G to ALF, while in the FC
microcosms 55 to 65% of the Bacteria community was no longer identifiable by
the phylum or subclass probes used. The latter result suggested that toluene
exposure fostered the proliferation of phylotype(s) that were otherwise minor
constituents of the FC aquifer community. These studies demonstrated that
alterations in aquifer microbial communities resulting from specific anthropogenic
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Chapter 4 Molecular Techniques in The Analysis of Microbial Diversity in Contaminated Soils
perturbances can be inferred from microcosm studies integrating chemical and
phylogenetic probe analysis and in the case of hydrocarbon contamination may
facilitate the identification of organisms important for in situ biodegradation
processes.
The main advantages and disadvantages of this technique are summarizing
as follows:
•
Advantages:
−
Permits rapid and simple monitoring of the spatial-temporal variability of
microbial populations if just band patterns are considered;
−
it is relatively easy to obtain an overview of the dominant species of an
ecosystem;
−
it is adequate for analysis of a large number of samples (far more than
cloning).
•
Disadvantages:
−
depending on the nature of the sample, extraction and amplification of
representative genomic DNA can be difficult (as in cloning);
−
after the PCR amplification, the DNA copy number – which depends on
abundance of a particular microorganism and the ease of amplification of
the 16SrRNA – can be very different (as in cloning). The intensity the
bands obtained on a DGGE gel may therefore vary (not quantitative);
−
the number of detected bands is usually small, which implies:
the number of identified species is also small;
the bands correspond, although not ncessarily, to the predominant
species in the original sample;
−
the sequences of the bands obtained from a gel correspond to short DNA
fragments (200 – 600 bp), and so phylogenetic relations are less reliably
established than with cloning of the whole 16SrRNA gene. In addition,
short sequences are less useful for designing new specific primers and
probes.
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Chapter 4 Molecular Techniques in The Analysis of Microbial Diversity in Contaminated Soils
The latest advance in molecular technology is the use of nucleic acid
microarrays or DNA chips in which the probes are immobilized (Blohm and
Guiseppi-Elie, 2001; Cho and Tiedje, 2001; Koizumi et al., 2002; Sergei et al.,
2001; Small et al., 2001; Urakawa et al., 2003; Wilson et al., 2002). This method
allows the simultaneous study of thousands of genes or messenger RNAs under
various physiological states. Both methods place a variety of single strand DNA
probes of interest on glass computer chips (or microscope slides). However the
use of this technique for environmental samples is still limited. Recently, Koizumi
et al. (2002) characterized a mesophilic, sulfate-reducing, toluene-degrading
consortium (TDC) and an ethylbenzene-degrading consortium (EDC) by DGGE
fingerprinting of PCR amplified 16S rRNA gene fragments, followed by
sequencing. The sequences of the major bands were affiliated with the family
Desulfobacteriaceae. Another major band from EDC was related to an uncultured
non-sulfate-reducing soil bacterium. Oligonucleotide probes specific for the 16S
rRNAs of target organisms corresponding to the major bands were designed, and
hybridization conditions were optimized for two analytical formats, membrane
and DNA microarray hybridization. Both formats were used to characterize the
TDC and EDC, and the results of both were consistent with the DGGE analysis.
In order to assess the utility of the microarray format for analysis of
environmental samples, oil-contaminated sediments from the coast of Kuwait
were analyzed. The DNA microarray successfully detected bacterial nucleic acids
from these samples, but probes targeting specific groups of sulfate-reducing
bacteria did not give positive signals.
Application of quantitative PCR techniques such as competitive PCR
and real-time PCR allow to obtain a quantitative picture of the specific groups of
bacteria. Recently, due to the development of methods for total extraction of RNA
from environmental samples of different origin RT-PCR can be used to amplify
cDNA derived from 16S rRNA from a RNA extract. This allows to amplify 16S
rRNA from metabolically active populations in a community and to get
information about the active members of a community. However, no RT-PCR
studies concerning 16S rRNA diversity to detect the active bacterial community
present in BTEX contaminated soils are reported.
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Chapter 4 Molecular Techniques in The Analysis of Microbial Diversity in Contaminated Soils
In case sequence information exist about functional genes involved in
degradation of xenobiotic compounds, this information can be used to design
specific primers or DNA probes for direct PCR detection of or hybridization with
the corresponding genes. Application of such techniques can provide a more
detailed picture of the catabolic gene structure and sequence diversity in
environmental samples, which will increase significantly our knowledge of the
functional potential of the microbial community in the studied environment.
Moreover, shifts in catabolic gene structure allow the deduction of the
evolutionary fitness of catabolic genes, operons and their respective hosts (Junca
and Pieper, 2004). Recently, many studies reported the design of PCR primers to
detect and/or quantify by PCR the presence of genotypes, encoding key steps in
bacterial BTEX biodegradation pathways in soil DNA extracts. Moreover, RTPCR on RNA extracts allows to see if the genes are actively transcribed in the
sample. Ogram et al. (1995) designed primer pairs to specifically detect the genes
encoding the α-subunit of the hydroxylase component (TmoA) of the toluene 4mono-oxygenase of P. mendocina KR1 and the iron-sulfur oxidase α-subunit
(TodC1) of the toluene dioxygenase of P. putida F1. The primers were used for
the detection of the expression of these genes and hence their activity in lowbiomass deep subsurface BTEX contaminated sediments by employing RT-PCR
on mRNA extracted from the aquifer. They detected tmoA homologous RNA
transcripts. Recently, Baldwin et al. (2003) designed degenerate primers to
specifically detect genes encoding two groups of α-subunits of the diiron
hydroxylase component of different multi-component mono-oxygenases and the
gene encoding the hydroxylase component (XylM) of the side chain xylene monooxygenase. The primer sets were used for detection and enumeration of aromatic
oxygenase genes by multiplex and real-time PCR on pure cell DNA, but they have
no data available on aquifer samples. Primer sets for the detection of C23O genes
specific for the meta-cleavage of the aromatic catechol structure in fluorescent
Pseudomonas or Sphingomonas were reported by Hallier-Soulier et al. (1996),
Okuta et al. (1998), Meyer et al. (1999), Mesarch et al. (2000) and Junca and
Pieper (2004). Cavalca et al. (2003) analyzed the functional and phylogenetic
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Chapter 4 Molecular Techniques in The Analysis of Microbial Diversity in Contaminated Soils
biodiversity of bacterial communities in a BTEX-polluted aquifer treated by airsparging. Five months of air injection reduced species diversity in the cultivable
community (as calculated by the Shannon-Weaver index), while little change was
noted in the degree of biodiversity in the total bacterial community, as monitored
by DGGE analysis of PCR amplified 16S rRNA genes. BTEX-degrading isolates
belonged to the genera Pseudomonas, Microbacterium, Azoarcus, Mycobacterium
and Bradyrhizobium. The degrading capacities of three strains in batch liquid
cultures were also studied. In some of these micro-organisms different pathways
for toluene degradation seemed to operate simultaneously. Pseudomonas strains
of the P24 operational taxonomic unit, able to grow only on catechol and not on
BTEX, were the most abundant, and were present in the groundwater community
at all stages of treatment, as evidenced both by cultivation approaches and by
DGGE profiles. The presence of different tmo-like genes in phylogenetically
distant strains of Pseudomonas, Mycobacterium and Bradyrhizobium suggested
recent horizontal gene transfer in the groundwater. Junca et al. (2004) used PCRSSCP for determining the diversity of C23O genes in environmental samples.
These PCR-SSCP results were assessed by comparing sequence data from PCRDNA clone libraries and C23O sequences and metabolic performance of microorganisms exhibiting C23O activity. PCR-SSCP was demonstrated to be a reliable
and rational tool to rapidly determine sequence diversity within a catabolic gene
family in environmental samples obtained from a BTEX contaminated site. In
another study, Junca et al. (2003) used an approach identical to ARDRA or
amplified functional DNA restriction analysis (AFDRA) to rapidly characterize
C23O subfamily I.2.A genes, known to be of crucial importance for aromatic
degradation. Restriction of the genes by Sau3A1 theoretically produced
characteristic profiles from each subfamily I.2.A member and their similarities
reassembled the main divergent branches of C23O gene phylogeny. Cluster
analyses of the restriction fragment profiles obtained from isolates from a BTEX
contaminated site showed patterns with distinct similarities to the reference strain
profiles, allowing to distinguish four different groups. Sequences of PCR
fragments from isolates were in close agreement with the phylogenetic
correlations predicted with the amplified functional DNA restriction analysis
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Chapter 4 Molecular Techniques in The Analysis of Microbial Diversity in Contaminated Soils
(AFDRA) approach. AFDRA thus provided a quick assessment of C23O diversity
in a strain collection and insights of its gene phylogeny affiliation among known
family members, but may also define the predominant polymorphism of a
functional gene present in environmental DNA extracts. This approach may be
useful to differentiate functional genes also for many other gene families.
The microbial diversity or presence and activity of functional genes may
also be assessed by techniques like hybridization by application of specific DNA
probes using the whole microbial DNA, DNA amplified by PCR or even at the
rRNA level. Guo et al. (1997) studied concentrations of selected genes, including
several involved in the degradation of BTEX (tmoABCDE, todC1C2BA, xylE) by
quantitative DNA:DNA hybridization on DNA extracted from subsurface soil
samples collected along a gradient of BTEX concentrations at a fuel oilcontaminated site. DNA from contaminated samples was significantly enriched in
most of the catabolic genes, relative to DNA from the non-contaminated site.
Hybridization of tmo and xylE were significantly higher in the contaminated
samples than in the non-contaminated samples. The level of hybridization was, in
descending order, as follows: xylE > tmoABCDE. No hybridization was observed
with todC1C2BA. Hybridization of xylE increased with increasing aromatic
concentration up to approximately 100 mg aromatics g-1 soil. Above that
concentration, the hybridization of xylE generally decreased. In an assessment of
the microbiological potential for the natural attenuation of petroleum
hydrocarbons in a shallow aquifer system, Stapleton and Sayler (1998) performed
a quantitative DNA:DNA hybridization molecular analysis on 60 uncontaminated
aquifer samples (only 15 had previous exposure to low levels of hydrocarbons)
using DNA probes targeting genes encoding degradative enzymes such as toluene
dioxygenase (todC1C2), toluene mono-oxygenase (tomA), and xylene monooxygenase (xylA). Each target sequence was present in nearly all samples.
Hydrocarbon degrading genotypes from previously exposed samples did not differ
from the other 45 samples that had no prior contaminant exposure, suggesting that
the microbial community of previously exposed sediments had re-equilibrated.
The level of hybridization was, in descending order, as follows: todC1C2 > tomA
> xylA. The genotype consistently found in the lowest abundance was xylA. From
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Chapter 4 Molecular Techniques in The Analysis of Microbial Diversity in Contaminated Soils
the same samples, they isolated and characterized 26 indigenous micro-organisms
capable of biodegrading fuel-related compounds such as BTEX (Stapleton et al.,
2000). Only one isolate hybridized with the todC1C2 gene probe and two isolates
hybridized with the xylA gene probe, while no isolates hybridized with the tomA
gene probe. To monitor changes in the molecular microbial ecology as well as
stimulation of natural biodegradative processes under transient field study
conditions, Stapleton and Sayler (2000) introduced a large, synthetic “model” jet
fuel mixture containing BTEX compounds and naphthalene in a decane carrier
into the subsurface. Over time they took subsurface samples at different places of
the spreading petroleum hydrocarbon plume and monitored changes in subsurface
catabolic gene frequencies during natural attenuation of the petroleum
hydrocarbons by quantitative DNA:DNA hybridization using the same DNA
probes. Each of the target genotypes showed significant responses to hydrocarbon
exposure. At first only significant enrichment for degradative micro-organisms
was seen for the todC1C2 genotype. Then both degradative genotypes todC1C2
and xylA significantly increased in samples collected from the source. After
reaching a certain peak population level, both genotypes underwent significant
decreases followed by a stabilization of both the plume front and degradative
genotypes.
Methods allowing the direct (whole cell hybridization, applied on the
sample without nucleic acid extraction as preceding step) characterization of
microbial communities and specific nucleic acid sequences have been long
awaited and include in situ (RT-)PCR (Amann and Kühl, 1998), and fluorescent
in situ hybridization (FISH) (Amann et al., 2001). FISH is a technique where
fluorescent oligonucleotides (16-20 nucleotides) recognize 16S rRNA sequences
in fixed cells and hybridize with them in situ (DNA-RNA matching). It involves:
(i) fixation and permeabilisation of the sample, (ii) hybridization by fluorescently
labeled, rRNA targeted oligonucleotide probes, (iii) washing steps to remove
unbound probe, and (iv) detection of labeled cells by microscopy (epifluorescence
or confocal laser scanning microscopy) or flow cytometry (Boon, 2002). The
fluorescence signal emitted by a cell also indicates the physiological state of the
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Chapter 4 Molecular Techniques in The Analysis of Microbial Diversity in Contaminated Soils
cell. The more active a cell, the more ribosomes are present that can serve as a
target for the oligonucleotides.
Microorganisms can be identified, localized and quantified in almost every
ecosystem with hybridization (Amann et al, 1990). The specifity of the probe
enables detection/identification of any desired taxonomic level, from domain
down to a resolution suitable for differentiating between individual species. The
main shortcoming of this technique lies in the lack of availability of probes
targeting the desired bacterial taxon or group. Although it is possible, in theory, to
design the most apt probe for each application thanks to the growing rRNA
sequence database (16/18S and 23/28S rRNA), it may be impossible to develop a
probe that specifically detects certain groups of microorganisms that share
metabolic properties (for example, sulfate-reduction or halo-respiration).
Furthermore, some previous knowledge of the expected microorganisms in the
sample is often required to apply this method succesfully. To target a particular
species, a specific probe must be ready or its 16S rRNA sequence must be
available. FISH is exclusively a taxonomic method that is most commonly used to
examine whether members of a specific phylogenetic affiliation are present in a
sample. It cannot, however, reveal information about the function or metabolic
features of the microorganisms detected with phylogenetically-related bacteria.
In situ (RT-)PCR involves amplification of specific nucleic acid
sequences inside intact prokaryotic cells followed by color or fluorescence
detection of the localized PCR product via bright-field or epifluorescence
microscopy (Chen et al., 2000; Hodson et al., 1995). Chen et al. (2000) coupled
prokaryotic in situ RT-PCR with flow cytometry to detect mRNA transcripts of
the toluene dioxygenase (todC1) gene in intact cells of the bacterium P. putida F1.
The combination of flow cytometry and a prokaryotic in situ RT-PCR approach
allowed the rapid detection and enumeration of functional populations of
microbial cells. Tani et al. (2002) injected Ralstonia eutropha KT1, which
degrades TCE, into an aquifer after activation with toluene, and then monitored
the number of bacteria by in situ PCR targeting the phenol hydroxylase gene and
by FISH targeting 16S rRNA. Recently, a combination of FISH and
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Chapter 4 Molecular Techniques in The Analysis of Microbial Diversity in Contaminated Soils
microautoradiography was developed to determine in situ the identities, activities
and specific substrate uptake profiles of individual bacterial cells within complex
microbial communities, but to our knowledge, this has not been applied for
detection of BTEX degrading bacteria in environmental samples (Lee et al.,
1999).
As an alternative to DGGE as a community profiling method, terminal
restriction fragment lenght polymorphism (tRFLP) can be applied when trating
complex, species-rich samples. This technique is also PCR based but the further
procedure differs from PCR/DGGE or PCR/cloning. In tRFLP the 16S gene is
amplified with universal primers, one of them being fluorescently labelled, and
the product is digested with frequently cutting restriction enzymes. Given that
each species in the sample has differences in the amplified gene sequences , the
terminal restriction fragment will differ in size, so can be separated
electrophoretically. Furthermore, it is possible to sequence and identify the
generated fragments via comparison with a sequence database. The strenght of the
fluorescent signal yields additional information on the abundance of different
species, though this feature should be regarded with caution, just like the band
intensity in patterns of a DGGE gel.
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CHAPTER 5
5. GROWTH OF CHLORINATED SOLVENT-DEGRADING MICROBIAL
CONSORTIA IN METHANE- AND PROPANE-FED BIOREACTORS AND
TESTING OF THEIR EFFECTIVENESS AS INOCULA FOR THE
BIOAUGMENTATION OF DIFFERENT TYPES OF AQUIFERS
______________________________________________________________________________________________________________________________________________________________
ABSTRACT
In this work we studied the long-term growth process of two microbial consortia
effective in the aerobic cometabolic biodegradation of a mixture of 6 chlorinated
aliphatic hydrocarbons (CAHs), and the effectiveness of these consortia as inocula
for the bioaugmentation of different types of microcosms. The main goals of the
study were to verify the maintenance of the consortia’s capacity to degrade a
CAH mixture during a prolonged growth process in the absence of the CAHs, and
to verify the consortia’s effectiveness in CAH biodegradation upon inoculation in
slurry microcosms set up with different types of aquifer materials. The propaneutilizing consortium generally proved the most effective one, being able to
biodegrade vinyl chloride, cis- and trans-1,2-dichloroethylene, trichloroethylene,
1,1,2-trichloroethane and 1,1,2,2-tetrachloroethane at all the CAH concentrations
tested. Both consortia maintained unaltered CAH degradation capacities during a
300-day growth period in the absence of the CAHs and were effective in inducing
the rapid onset of CAH depletion upon inoculation in slurry microcosms set up
with 5 types of aquifer materials. A consortium developed in microcosms supplied
with both methane and propane combined the best degradation capacities of the
two single-substrate consortia. The degree of conversion of the organic Cl to
chloride ion was equal as an average to 90%.
Growth of CAH-degrading Consortia in Methane- and Propane-fed Bioreactors
5.1 INTRODUCTION
Chlorinated solvents, or chlorinated aliphatic hydrocarbons (CAHs), are
among the most common contaminant of soils, groundwaters and wastewaters,
and most of them are known or suspected carcinogens (Gossett, 2005). Of over
1200 hazardous waste sites included in the U.S. EPA 2002 National Priority List,
47% are contaminated by trichloroethene (TCE), 42% by perchloroethylene
(PCE) and 37% by 1,1,1-trichloroethane (1,1,1-TCA) (U.S. EPA, 2002). Since the
1980s, numerous literature studies documented the successful biodegradation of
CAHs by means of both aerobic and anaerobic processes. In particular, research
on aerobic CAH cometabolism tested primarily the utilization of methane
(Andersen and McCarty, 1996; Chang and Alvarez-Cohen, 1996), toluene, phenol
(Hopkins and McCarty, 1995; McCarty et al., 1998) and ammonia (Ely et al.,
1997; Keener and Arp, 1993) as growth substrates, whereas a limited number of
studies focused on propane. These studies showed that propane-grown singlestrains (Wackett et al., 1989; Wilcox et al., 1995) and mixed cultures can trasform
TCE,
cis-
and
trans-1,2-dichloroethylene
(cis-
and
trans-DCE),
1,1-
dichloroethylene (1,1-DCE), vinyl chloride (VC), 1,1,2,2-tetrachloroethane
(1,1,2,2-TeCA), 1,1,2- and 1,2,2-trichloroethane (1,1,2- and 1,2,2-TCA), 1,1- and
1,2-dichloroethane (1,1- and 1,2-DCA) and chloroform (CF).
Despite the encouraging results of the experimental studies, practitioners
are still reluctant to utilize aerobic cometabolism for the full-scale remediation of
CAH-contaminated sites (Semprini, 2001). One of the reasons for this is
represented by thr long lag-time that is sometimes required for the onset of the
aerobic cometabolic process by the indigenous biomass of CAH-contaminated
sites (Frascari et al.,2006). As a result of an extended lag period, a fraction of the
contaminated plume may pass through the treatment zone and reach sensitive
targets. When preliminary lab-scale investigations indicates the presence of long
lag-phases, bioaugmentation, consisting of the introduction of a suitable microbial
inoculum into the treatment system, can represent a very effective tool (Gentry et
al., 2004; Vogel, 1996). Several studies of lab-scale and in-situ biodegradation of
CAHs report the successful application of this technology (Jitnuyanont et al.,
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Growth of CAH-degrading Consortia in Methane- and Propane-fed Bioreactors
2001; Harkness et al.,1999; Munakata et al.,1996; Steffan et al.,1999).
Bioaugmentation can be performed both with a single bacterial strain and with a
microbial consortium; if a site is polluted with a complex contaminant mixture,
the latter solution may be preferred, the combined action of different
microorganisms being required to achieve a complete clean-up. In this case, in
order to perform a full-scale bioaugmentation, it is necessary to develop a suitable
initial microbial inoculum, and to grow it in a bioreactor so as to produce the
amount of biomass required to colonize an appropriate fraction of the
contaminated area. The inoculum growth process should be operated in an
economically feasible way, but at the same time it must guarantee the stability of
the microbial consortium, and consequently the maintenance of its degradation
capacities. The characteristics required by the growth process in order to satisfy
these two conditions represent an important theme of investigation. For example,
growing the inoculum on the primary substrate in the absence of the target
contaminants would represent a significant simplification of the production plant
and a reduction of the fixed and operational costs; however, this solution might
lead to a loss of the consortium’s degradation abilities. Moreover, inoculated
strains often survive poorly and may loose their in mixed microbial ecosystems.
Several factors have been implicated in the survival, activity and maintenance of
introduced strains. Some factors are of physicochemical nature, such as presence
or absence of oxygen, pH or temperature. Other factors reflect the physiological
adaptability of the bacterium, such as kinetics of substrate utilization, nutrients
and trace elements scavenging. Some bacteria may be particularly prone to
predation by protozoa when they maintain a freely suspended state rather than
attach themselves easily to surfaces or form sticky material (McClure et al.,
1991). Most inoculation studies have shown that the population size of introduced
strains decline strongly in mixed microbial ecosystems (McClure et al., 1989;
Watanabe et al., 1998). On the other hand, some strains were shown to be
particularly effective in colonizing and maintaining themselves (McClure et al.,
1991; Megharaj et al., 1997), Which was attributed to the fact that the strains were
“preadapted” to the prevailing conditions or originating from the same
environment. Some concern also exists that under some conditions the genetic
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Growth of CAH-degrading Consortia in Methane- and Propane-fed Bioreactors
information for degradation of the pollutants may not be stable in the introduced
organism and be lost or altered after some time (Kuar and Schugerl, 1990;
Proctor, 1994). Thus, effective assessment of the capabilities of bacteria requires
the study of their maintenance and activity. Therefore we were very interested in
evaluating the maintainance of degradation abilities of mixed consortia after longterm growth in batch bioreactors and in assessing their capacity to quickly start
and maintain the contaminant’s degradation process in environments differing
from the one they had been isolated from.
This study represents the continuation of a previous microcosm study
(Frascari et al., 2006) where we had developed a methane-utilizing and a propaneutilizing microbial consortium able to perform the long-term biodegradation of a
mixture of VC, trans- and cis-DCE, TCE, 1,1,2-TCA and 1,1,2,2-TeCA (a highchlorinated solvent generally considered non-biodegradable in aerobic conditions)
via aerobic cometabolism in slurry conditions.
The goals of the study were (i) to study the long-term aerobic biodegradation
of the CAH mixture above mentioned; (ii) to investigate the efficacy of
bioaugmentation with two types of internal inocula obtained from the indigenous
biomass of the studied site; (iii) to identify the CAH-degrading bacteria. VC,
methane and propane were utilized as growth substrates. The biodegradation
process was investigated at both 25 and 17°C by means of bioaugmented and nonbioaugmented
sediment-groundwater
slurry
microcosm
tests.
The
non-
bioaugmented microcosms were characterized, at 25 ◦C, by an average 18-day
lag-time for the direct metabolism of VC (accompanied by the cometabolism of
cis- and trans-DCE) and by long lag-times (36–264 days) for the onset of methane
or propane utilization (associated with the cometabolism of the remaining CAHs).
In the inoculated microcosms the lag-phases for the onset of growth substrate
utilization and CAH cometabolism were significantly shorter (0–15 days at 25
°C). Biodegradation of the 6-CAH mixture was successfully continued for up to
410 days. The low-chlorinated solvents were characterized by higher depletion
rates. The composition of the microbial consortium of a propane-utilizing
microcosm was determined by 16s rDNA sequencing and phylotype analysis.
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Growth of CAH-degrading Consortia in Methane- and Propane-fed Bioreactors
This study showed that the indigenous biomass of the investigated aquifer
material proved able to grow on VC 18–43 days (depending on the temperature)
after the establishment of aerobic conditions, and to degrade via cometabolism
cis- and trans-DCE but not TCE, 1,1,2-TCA or 1,1,2,2-TeCA present. Conversely,
the supply of methane or propane led to the biodegradation of the entire 6-CAH
mixture. Moreover the bioaugmentation treatments, performed with internal
inocula obtained from the site’s indigenous biomass, were highly effective in
reducing the long and variable lag-phases required for the onset of propane or
methane uptake in the non-augmented microcosms. In all the propane- or
methane-fed microcosms the biodegradation of each CAH rapidly reached a
stationary condition with higher rates in the low-chlorinated solvents. Besides this
was, to the best of our knowledge, the first study that documented the long-term
aerobic biodegradation of 1,1,2,2-TeCA.
In this work we investigated several aspects relative to the feasibility of
utilizing the two above-mentioned microbial consortia as inocula for
bioaugmentation treatments of CAH-contaminated sites with different physicalchemical characteristics. In particular, the goals of the study were:
i) to verify the maintenance of the consortia’s capacity to degrade the 6CAH mixture during a prolonged process of microbial growth in the presence as
well as in the absence of the 6-CAH mixture;
ii) to verify the consortia’s ability – after a prolonged growth process - to
lead to the rapid onset of biodegradation of the CAH mixture upon inoculation in
slurry microcosms set up with aquifer materials taken from sites with different
physical-chemical characteristics;
iii) to develop a third consortium able to combine the best characteristics
of the methane-utilizing and of the propane-utilizing consortia object of the study:
in fact, the previous study had shown that, while both consortia were effective in
the aerobic cometabolic biodegradation of VC and cis-DCE, the methane-utilizing
biomass had a higher capacity to transform trans-DCE, whereas the propaneutilizing one was more effective towards 1,1,2-TCA, 1,1,2,2-TeCA and,
secondarily, TCE;
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Growth of CAH-degrading Consortia in Methane- and Propane-fed Bioreactors
iv) to characterize in terms of specific CAH depletion rates and degree of
mineralization of the organic Cl the best methane-utilizing and the best propaneutilizing consortium obtained as a result of the inoculation in the microcosms set
up with different aquifer materials.
In order to evaluate the consortia’s ability and the maintenance of their
degradation capacities during the prolonged growth process , we chose to start
with a “black box” empirical method consisting in the inoculation – at different
times during the growth process – of small amounts of the wo consortia in slurry
microcosms containing aquifer materials from different sites, and in the
subsequent evaluation of the lag-times for the onset of biodegradation of the CAH
mixture and of the long-term CAH depletion rates obtained.
5.2 MATERIALS AND METHODS
5.2.1 Overview of the experimental scheme
This paragraph provides a short description of the experimental scheme,
whereas the details relative to the set-up and operation of each microcosm and
growth reactor are presented in the following paragraphs. The experimental
scheme of the study is shown – limitedly to the bioaugmented microcosms and
growth reactors – in Figure 5.1. In addition, a non-bioaugmented control
microcosm was set up for each type of aquifer material and for each growth
substrate, and two sterile control microcosms were set up to monitor abiotic
reactions, losses through caps and losses due to the microcosm sampling
procedures.
As explained afterwards the study involved the following steps:
- inoculation of growth bioreactors and set up of Time 0 slurry
microcosms in order to characterize the biodegradation abilities of the
initial inoculum;
- inoculation at different times of slurry microcosms with biomass
sampled from the growth bioreactors in order to verify the maintenance of
the consortia’s capacity to degrade the 6-CAH mixture during the
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Growth of CAH-degrading Consortia in Methane- and Propane-fed Bioreactors
prolonged growth process in the presence as well in the absence of the 6CAH mixture;
- inoculation of slurry microcosms set up with aquifer materials taken
from sites with different physical-chemical-biological characteristics
with biomass sampled from the growth reactors, in order to evaluate the
effectiveness of bioaugmentation in different types of aquifers;
- development of a new consortium able to combine the best
characteristics of the two studied consortia by inoculation of slurry
microcosms with both the methane-utilyzing and the propane-utilyzing
biomass;
- inoculation of liquid-phase microcosms in order to characterize in terms of
specific CAH depletion rates and degree of mineralization of the organic Cl
the best consortia obtained in the microcosms set up with different aquifer
materials.
As explained in section 5.2.4, the performace of the studied consortia was
evaluated in terms of:
- lag-time required for the onset of the aerobic cometabolic biodegradation of
the 6-CAH mixture;
- long term degradation rate of the 6-CAH mixture.
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Growth of CAH-degrading Consortia in Methane- and Propane-fed Bioreactors
Figure 5.1. Graphical representation of the experimental scheme of the bioaugmented
microcosms. The first group of letters indicates the type of bioreactor: GB, growth
bioreactor; S, slurry microcosm; L, liquid-phase microcosm. The second group of letters
indicates the type of biomass initially inoculated as well as the growth substrate supplied: M,
inoculation of methane-utilizing biomass and supply of methane; P, inoculation of propaneutilizing biomass and supply of propane; MP: inoculation of both types of biomass and
supply of both substrates. The third group of letters indicates the condition of biomass
growth (in the case of the growth bioreactors) or the type of inoculum introduced (in the
case of the slurry microcosms)(this group of letters is not included in the labelling of the
liquid-phase microcosms): NC, inoculum growth in the absence of the CAH mixture; C,
inoculum growth in the presence of the CAH mixture. The number indicates the time the
microcosm was set up and inoculated (the time is not specified for the growth bioreactors,
which were set up only at time zero). The last letter – present only in the labelling of the
slurry microcosms – indicates the type of aquifer material and groundwater utilized for
microcosm set up. Duplicate microcosms are indicated with the subscript “1,2”.
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Growth of CAH-degrading Consortia in Methane- and Propane-fed Bioreactors
With regard to the methane-utilizing consortium, the initial inoculum
object of the study was obtained by sampling 25 mL of suspension from each of
three methane-fed slurry microcosms of the previous study (Frascari et al., 2006)
with a sterile syringe after 30 seconds of intense agitation, whereas the propaneutilizing initial inoculum was obtained from three propane-fed microcosms of the
above-mentioned study, following the same procedure. The initial concentration
of both inocula was equal to 2.7 x 107 colony forming units (CFU) per mL.
At time zero (onset of this study), the methane-grown inoculum was
introduced into two liquid-phase methane-fed growth bioreactors where the
consortium was grown for 150-300 days in the presence (bioreactor GB-M-C) and
in the absence (bioreactor GB-M-NC) of the same 6-CAH mixture object of the
previous study (VC, trans- and cis-DCE, TCE, 1,1,2-TCA and 1,1,2,2-TeCA).
The growth process was continued in reactor GB-M-C for 150 days and in reactor
GB-M-NC for 300 days, by supplying methane pulses corresponding to an
average feed of 37 mmolC/week. In addition, in order to characterize the initial
methane-grown inoculum in terms of its ability to lead to the rapid onset of
biodegradation of the 6-CAH mixture upon inoculation in slurry microcosms set
up with the same aquifer material object of the previous study (soil + groundwater
type A), two duplicate slurry microcosms (S-M-C-0A) set up with the same
aquifer material were bioaugmented with the same inoculum introduced in the
growth bioreactors. These microcosms were then spiked with methane and with
the 6-CAH mixture, at the same initial concentrations utilized in the previous
study and typical of aquifer A (methane 125 µM; VC 25 µM; trans-DCE 3.4 µM;
cis-DCE 3.1 µM; TCE 1.9 µM; 1,1,2-TCA 0.30 µM; 1,1,2,2-TeCA 0.15 µM).
Similarly, at time zero the propane-grown inoculum was introduced into
two propane-fed bioreactors where the consortium was grown in the presence
(bioreactor GB-P-C) and in the absence (bioreactor GB-P-NC) of the 6-CAH
mixture, and two slurry microcosm (S-P-C-0A) set up with soil and groundwater
type A were bioaugmented with the same inoculum, spiked with propane and with
the 6-CAH mixture (propane 46 µM; same CAH initial concentrations as in
methane-fed microcosms S-M-C-0A) and subsequently operated by adding
113
Growth of CAH-degrading Consortia in Methane- and Propane-fed Bioreactors
consecutive propane and CAH pulses to evaluate the propane and CAH lag-times
and the CAH depletion rates.
After 30 days of growth of the two consortia in the four bioreactors (time
1), four slurry microcosms were set up with soil and groundwater from site A and
spiked with propane and with the 6-CAH mixture (at the same initial
concentrations utilized for the previous propane-fed microcosms). Of these, two
duplicates (S-P-NC-1A) were augmented with 2 mL of biomass suspensions
sampled from bioreactor GB-P-NC, and two (S-P-C-1A) were augmented with 2
mL from bioreactor GB-P-C.
After 150 days of growth of the two consortia (time 2), six slurry
microcosms were set up with aquifer material from site A and spiked with the 6CAH mixture (at the same initial concentrations utilized for the previous
microcosms). Of these, two (S-M-NC-2A and S-M-C-2A) were spiked with
methane (125 µM) and augmented respectively from bioreactors GB-M-NC and
GB-M-C, and two (S-P-NC-2A and S-P-C-2A) were spiked with propane (46 µ
M) and augmented from bioreactors GB-P-NC and GB-P-C; finally, in the attempt
to develop a third consortium able to combine the best characteristics of the
methane-utilizing and of the propane-utilizing consortia object of the study, two
duplicate microcosms (S-MP-NC-2A) were spiked with both methane (62.5 µM)
and propane (23 µM) and augmented from both GB-M-NC and GB-P-NC.
All the slurry microcosms set up at times 0, 1 and 2 with aquifer material
type A were operated by adding consecutive pulses of methane or propane (or
both in the case of S-MP-NC-2A) and of the 6-CAH mixture, at the same
concentration supplied for each compound in the initial pulse, for a total of 5-7
CAH pulses. As an example, the plot of CAH aqueous phase concentration versus
time relative to the first 4 days of operation of one of the two duplicate
microcosms S-M-C-0A is shown in Figure 5.2. The growth substrate and CAH
concentrations measured in these microcosms were utilized to evaluate the lagtimes for the onset of substrate utilization and of biodegradation of the entire
CAH mixture and the long-term CAH depletion rates, as explained in detail in
section 5.2.4. The results were compared with those obtained respectively in the
methane-fed and propane-fed non-bioaugmented slurry microcosms set up with
114
Growth of CAH-degrading Consortia in Methane- and Propane-fed Bioreactors
the same aquifer material, described in the previous study (Frascari et al., 2006)
and utilized as the source of the initial inoculum. With the exception of S-MPNC-2A, the microcosms of time 2, as well as the following ones, were not set up
as duplicates, as the results from the microcosms set up at times 0 and 1 indicated
a high reproducibility of the data obtained in the duplicate microcosms (as
reported in more detail in section 5.3.2 and in Figure 5.3). Conversely S-MP-NC2A, as well as the double-substrate microcosms set up at later times, was set up in
duplicate, in order to have the possibility, after a consistent number of CAH
pulses sustained by the supply of both substrates, to perform a final period of
operation characterized by the supply of only methane in one duplicate and only
propane in the other one.
On the basis of the positive results obtained in the microcosms set up at
times 1 and 2 with regard to the possibility to grow either consortium in the
absence of the CAH mixture and to utilize it to bioaugment slurry microcosms
identical - in terms of type of aquifer material as well as type and concentration of
contaminants - to those where the two consortia had initially been developed (see
section 5.3.3), at time 3 (300 days of consortium growth in the bioreactors) we set
up a further group of microcosms (S-M-NC-3A, B, C, D and E; S-P-NC-3A, B, C,
D and E) aimed at investigating the potential of utilization the two consortia as
inocula for the bioaugmentation of different CAH-contaminated aquifers. These
microcosms, inoculated respectively from growth reactors GB-M-NC and GB-PNC (consortia grown in the absence of the CAH mixture), were constructed with
soil and groundwater taken from different sites: aquifers C and D, similarly to A,
are CAH-contaminated sites containing sandy/silty soils, whereas aquifers B and
E are uncontaminated sites containing respectively a sandy soil and a humic soil
characterized by a high fraction of organic carbon (1.5%). The purpose of
utilizing aquifer materials from uncontaminated sites was to simulate a
bioremediation treatment conducted right after the occurrence of the
contamination with the CAH mixture: in this condition, the site’s indigenous
biomass has not been subjected to the selective pressure of the chlorinated
compounds. In order to give a more general scope to this section of the work, the
microcosms of time 3 were spiked both at set-up and in the subsequent pulses
115
Growth of CAH-degrading Consortia in Methane- and Propane-fed Bioreactors
with an equal molar concentration (set to 4 µM) of the different CAHs. Besides,
VC was not included in the CAH mixture, as numerous studies showed that this
compound is easily biodegraded by several bacteria via both direct metabolism
and cometabolism (Frascari et al., 2006; Verce et al., 2002; Coleman et al., 2005;
6
30
4
20
2
10
0
0
0
10
20
Time (d)
trans-1,2 DCE
TCE
1,1,2,2 - TeCA
30
VC (µM)
Trans-1,2 DCE, cis-1,2 DCE, TCE,
1,1,2-TCA, 1,1,2,2-TeCA (µ M)
Hartmans et al., 1992).
40
cis-1,2 DCE
1,1,2-TCA
VC
Figure 5.2. Aqueous phase concentrations of the 6 CAHs versus time during the first 40 days
of operation of microcosm S-M-C-0A1. For higher clarity the methane pulses, supplied daily
at 125 µM and rapidly consumed, are not represented.
In addition to the ones above-listed, a further microcosm (S-MP-NC-3E)
was set up in duplicate at time 3 with aquifer material from site E, bioaugmented
from both GB-M-NC and GB-P-NC and spiked with the 5-CAH mixture (no VC)
at 4 µM as well as with both methane (62.5 µM) and propane (23 µM). Besides, a
non-bioaugmented control microcosm was set up for each substrate and for each
aquifer material, for a total of 10 tests (K-M-A, B, C, D and E; K-P-A, B, C, D
and E), and additioned with the 5-CAH mixture at 4 µM and with methane (125 µ
M) or propane (46 µM).
The microcosms of time 3, similarly to the previous ones, were operated
by addition of consecutive pulses of methane or propane (or both in the case of S-
116
Growth of CAH-degrading Consortia in Methane- and Propane-fed Bioreactors
MP-NC-3E) and of the 6-CAH mixture, at the same concentration supplied for
each compound in the initial pulse, for a total of 3 CAH pulses (7 for the E-type
microcosms). The results, elaborated in terms of substrate and CAH mixture lagtime and of CAH depletion rates, were compared to those obtained in the
corresponding non-bioaugmented control microcosms.
Finally, at time 4 (390 days: 300 days of consortium growth in bioreactors
GB-M-NC and GB-P-NC in the absence of CAHs + 90 days of consortium
growth in the slurry microcosms of time 3 in the presence of the 5-CAH mixture),
four liquid-phase microcosms were set up with a chloride-free mineral medium
and spiked with the 5-CAH mixture (4 µM for each compound): one (L-M-4) was
inoculated from slurry microcosm S-M-NC-3E and spiked with methane (125 µ
M), one (L-P-4) was inoculated from S-P-NC-3E and spiked with propane (46 µ
M), whereas two duplicates (L-MP-4) were augmented from S-MP-NC-3E and
spiked with both methane (62.5 µM) and propane (23 µM). The purpose of
operating microcosms in the absence of soil was to obtain an evaluation of the
long-term CAH specific depletion rates (in the slurry microcosms, given the
difficulty of performing a precise valuation of active biomass, specific depletion
rates cannot be evaluated with high accuracy). The reason why the liquid-phase
microcosms were inoculated from the slurry microcosms containing soil from a
site not characterized by a historical CAH contamination (site E) is that we were
interested in investigating the characteristic of the consortia obtained from the
interaction between the CAH-degrading methane-utilizing or propane-utilizing
inoculated consortium and an indigenous biomass not affected by a previous
selective pressure due to CAH contamination. The liquid-phase microcosms,
similarly to the slurry ones, were operated by addition of consecutive pulses of
growth substrate (methane or propane or both) and of the 5-CAH mixture, at the
same concentration of each compound as in the initial pulse.
5.2.2 Growt bioreactors set up and operation
Growth bioreactors (Figure 5.3) consisted of 5-l glass bottles sealed with Teflonlined septa and containing 1 L of sterile mineral medium (Table 5.1).
117
Growth of CAH-degrading Consortia in Methane- and Propane-fed Bioreactors
(a)
(b)
Figure 5.3. Batch growth reactors with the two different biomasses: a) metanotrophs; b)
propanotrophs
Table 5.1. Mineral medium composition
Compound
Concentration (µM)
(NH4)2SO4
797
MgSO4·7H2O
244
CaCl2
132
K2HPO4
8902
NaH2PO4·H2O
5355
FeSO4·7H2O
22.6
NaNO3
9000
MnCl2·4H2O
1.52
ZnSO4·7H2O
0.510
H3BO3
1.00
Na2MO4·2H2O
0.450
NiCl2·2H2O
0.144
CuCl2·2H2O
0.100
CoCl2·6H2O
0.100
118
Growth of CAH-degrading Consortia in Methane- and Propane-fed Bioreactors
As shown in Table 5.2, each bioreactor was inoculated with 30 ml of the
starting inoculum (2.7 x 107 CFU/mL). The primary substrate (methane or
propane) was supplied in consecutive spikes (at the initial liquid-phase
concentration in each pulse of 54 µM for methane and 20 µM for propane.)
corresponding to an average feed rate of 37 mmolC/week. The 6 chlorinated
solvents were introduced into bioreactors GB-M-C and GB-P-C by spiking 7.4 ml
of gaseous VC and 900 µL of a concentrated aqueous solution of trans-DCE (14.8
mM), cis-DCE (8.1 mM), TCE (7.9 mM), 1,1,2-TCA (0.83 mM) and 1,1,2,2TeCA (0.12 mM) each time all of them had been completely degraded. The CAH
initial concentrations in each pulse were : VC 25 µM; trans-DCE 3.4 µM; cisDCE 3.1 µM; TCE 1.9 µM; 1,1,2-TCA 0.30 µM; 1,1,2,2-TeCA 0.15 µM. As a
result of this procedure, the CAH overall feed rate was equal to 0.1 mmol/week.
Aerobic conditions were maintained by adding pure oxygen each time the primary
substrate was added, and the reactor aqueous phase was periodically air-stripped
in order to remove CO2 and possible volatile toxic degradation products A nutrient
solution containing ammonium (as NH4Cl 918 mM) and phosphate (as KH2PO4:
38 mM and K2HPO4: 45 mM) was periodically provided so as to maintain a C:N:P
molar ratio in the bioreactors feed equal to 220:11:1. Every 120 days the culture
medium was centrifuged and biomass resuspended in fresh medium.
The growth reactors, equipped with baffles in order to increase the gas-liquid
mass transfer rate, were kept in continuous agitation in an orbital shaker (130
rpm) at 25 °C.
Table 5.2. Growth reactors set-up and operational data
Growth substrate
Reactor
Type
Initial inoculum
Initial
CAH
Volume
Concentration
aq.concentration
mixture
(ml)
(107 CFU/ml)
µM)
GB-M-NC
GB-M-C
GB-P-NC
GB-P-C
Methane
54
Propane
20
5.2.3 Microcosms set up and operation
119
Not present
30
2,7
Present
30
2,7
Not present
30
2,7
Present
30
2,7
Growth of CAH-degrading Consortia in Methane- and Propane-fed Bioreactors
Slurry and liquid phase microcosms were set up using 119 ml amber serum bottles
sealed with Teflon-lined rubber septa. Each microcosm contained 20 g of soil and
50 ml of groundwater. In the microcosms set up with soil from aquifers B and E,
in the absence of groundwater from the site, we introduced the same mineral
medium utilized for the growth bioreactors (Table 5.3). The liquid-phase
microcosms contained 50 ml of a Cl-free mineral medium having the same
composition as that of the bioreactors, but containing CaSO4 instead of CaCl2
(Table 5.4). Each inoculated microcosm was augmented with 2 mL of biomass
suspension. The biomass concentrations in the growth bioreactors at the time of
each inoculation are reported in Table 5.5. The primary substrate was supplied in
consecutive spikes (initial liquid-phase concentration in each pulse: methane 125
µM, propane 46 µM, methane 62.5 µM + propane 23 µM in the double-substrate
tests), corresponding to an average feed rate of 1.1 mmol C/week. The CAH were
introduced by spiking gaseous VC (except for the tests of times 3 and 4) and an
aqueous solution of the remaining 5 CAHs (solution utilized for the tests of time
0, 1 and 2: trans-DCE 11.1 mM, cis-DCE 9.1 mM, TCE 5.9 mM, 1,1,2-TCA 1.8
mM, 1,1,2,2-TeCA 0.32mM; solutions for the tests of times 3 and 4: trans-DCE
(7.9 mM), cis-DCE (6.5 mΜ), TCE (8 mΜ), 1,1,2-TCA (5.9 mΜ) and 1,1,2,2TeCA (5.7 mΜ)
Aerobic conditions were maintained by adding pure oxygen (9 ml) with a
frictionless glass syringe prior to each primary substrate supply and the
microcosms aqueous phase was air-stripped each time a new pulse of CAHs was
added, in order to remove CO2 and possible volatile toxic degradation products. A
nutrient solution containing ammonium (as NH4Cl 289 mM) and phosphate (as
KH2PO4: 12 mM and K2HPO4: 14 mM) was periodically provided so as to
maintain a C:N:P molar ratio in the feed equal to 220:11:1. The microcosms were
maintained in agitation in a roller (3.3 rpm) at 25 ° C. To evaluate abiotic CAH
depletion rates and losses through caps, four control microcosms were set up and
sterilized with NaN3 57 mM: two (ST-A1,2) contained E-type aquifer materials and
were spiked with the CAH mixture at the initial concentrations similar to those
typical of site A (and utilized in the microcosms of times 0, 1 and 2), whereas two
120
Growth of CAH-degrading Consortia in Methane- and Propane-fed Bioreactors
(ST-E1,2) contained E-type aquifer materials and were spiked with TCE, 1,1,2TCA and 1,1,2,2-TeCA at 4 µM.
Table 5.3. Set up data of the slurry microcosms
Time
T0
T1 (30 d)
T2 (150 d)
Non inoculated microcosms
T3 (300 d)
Label
Composition
Growth substrate
Soil A + water A
Methane
Propane
Soil A + water A
Propane
Parent bioreactor
Duplicates
2
2
2
2
1
1
1
1
2
1
1
1
1
1
1
1
1
1
1
2
S-M-C0A
S-P-C0A
S-P-NC-1A
S-P-C-1A
S-M-NC-2A
S-M-C-2A
S-P-NC-2A
S-P-C-2A
S-MP-NC-2A
S-M-NC-3A
S-M-NC-3B
S-M-NC-3C
S-M-NC-3D
S-M-NC-3E
S-P-NC-3A
S-P-NC-3B
S-P-NC-3C
S-P-NC-3D
S-P-NC-3E
S-MP-NC-3E
Soil A + water A
Soil B + mineral medium
Soil C + water C
Soil D + mineral medium
Soil E + mineral medium
Soil A + water A
Soil B + mineral medium
Soil C + water C
Soil D + mineral medium
Soil E + mineral medium
Soil A + water A
Methane + Propane
GB-P-NC
GB-P-C
GB-M-NC
GB-M-C
GB-P-NC
GB-P-C
GB-P-NC/GB-M-NC
GB-M-NC
GB-M-NC
GB-M-NC
GB-M-NC
GB-M-NC
GB-P-NC
GB-P-NC
GB-P-NC
GB-P-NC
GB-P-NC
GB-P-NC/GB-M-NC
KMA
KMB
KMC
KMD
KPA
KPB
KPC
KPD
Soil A + water A
Soil B + mineral medium
Soil C + water C
Soil D + mineral medium
Soil A + water A
Soil B + mineral medium
Soil C + water C
Soil D + mineral medium
Methane
Methane
Methane
Methane
Propane
Propane
Propane
Propane
-
2
2
2
2
2
2
2
2
KMPE
Soil E + mineral medium
Methane + propane
-
2
Methane
Soil A + water A
Propane
Methane + propane
Methane
Propane
Table 5.4. Set up data of the liquid-phase microcosms
Label
Growth substrate
Parent microcosm
Duplicates
L-MP-4
Methane + propane
S-MP-NC-3E
2
L-M-4
Methane
S-M-NC-3E
1
L-P-4
Propane
S-P-NC-3E
1
121
Growth of CAH-degrading Consortia in Methane- and Propane-fed Bioreactors
Table 5.5. Viable biomass concentrations in the growth bioreactors at the times of inoculation of the
slurry microcosms
5.2.4 Estimation of lag times, degradation rates and rate/concentration ratios
The lag-times for the onset of growth substrate consumption and CAH
degradation were obtained by the intersection of the maximum slope line of the
concentration-time curve with the horizontal line passing through the initial
concentration value. The lag-times are reported in the Results in terms of substrate
lag-time and additional lag-time relative to the CAH mixture, measured from the
onset of substrate consumption and defined as the longest of the additional lagtimes relative to the single CAHs.
Each CAH pulse was characterized by the maximum degradation rate
relative to each compound, calculated by dividing the maximum slope of the
mass-time curve by the volume of the liquid phase. Each degradation rate was
associated with the aqueous phase concentration corresponding to the initial value
of the portion of the mass-time curve utilized to calculate the degradation rate.
The CAH depletion rates were elaborated according to the method described in
detail in the previous study (Frascari et al., 2006): the rates obtained for each
compound in each microcosms were plotted versus the mass of growth substrate
consumed; Figure 5.4 shows as an example the plot degradation rate versus
consumed carbon mass for the first 5 pulses of VC in duplicate microcosms S-MC-0-A1 and S-M-C-0-A2. This elaboration indicated that, after about 4 - 5 pulses
of CAH mixture depletion, each microcosm reached a roughly stationary
condition in terms of CAH depletion rates; therefore we calculated the average of
the depletion rates evaluated – for each compound and for each microcosm – in
122
Growth of CAH-degrading Consortia in Methane- and Propane-fed Bioreactors
the stationary phase; this average rate was divided by the average of the
concentrations associated with the depletion rates included in the average. The
rate/concentration ratio obtained – indicate in the following with k* - was utilized
as an index to compare the depletion rates obtained in a given microcosm
relatively to the different CAHs, and in different microcosms relatively to the
same CAH. This simplified approach allowed to compare, within each
microcosm, the depletion rates obtained for the different CAHs at different initial
concentrations; it also allowed to compare the rates obtained for a given CAH at
time 3 (when all the CAHs were supplied at 4 M) with those obtained for the
same CAH at earlier times (when the different CAHs were supplied at the
concentrations typical of site A).
S-M-C0-A1
S-M-C0-A2
0,9
degradation rate (mM/d)
0,8
0,7
0,6
0,5
0,4
0,3
0,2
0,1
0,0
0
1
2
3
4
5
6
7
8
9
consumed carbon mass (mmol)
Fig 5.4. Plot of the degradation rate-growth substrate consumed for the first 5 VC pulses in
duplicate microcosms S-M-C-0-A1, S-M-C-0-A2.
123
Growth of CAH-degrading Consortia in Methane- and Propane-fed Bioreactors
The ratio of depletion rate to concentration (k*) can be considered a
pseudo first-order constant that includes biomass concentration), in the hypothesis
that the biodegradation of each CAH followed a first-order kinetic model within
the concentration ranges tested for each compound. This hypothesis was
confirmed – although within concentration ranges lower than those of this study,
in particular for 1,1,2-TCA and 1,1,2,2-TeCA – by the results of the previous
study (Frascari et al., 2006), where it was shown that, in each slurry microcosm,
the depletion rates obtained for each CAH at a roughly constant biomass
concentration were proportional to the corresponding initial concentrations in the
pulses. The assumption of first-order kinetic is in agreement with the observation
that the CAH half-saturation constants reported in the literature are typically
higher than the concentration ranges investigated in this study (Alvarez-Cohen et
al., 2001; Arp et al., 2001; Oldenhuis et al., 1991).
The only exception to the above-described procedure for the elaboration of
the CAH depletion rates was made for the slurry microcosms set up at time 3 with
aquifer materials from sites A, B, C and D: because these tests were operated only
for 3 CAH pulses, the plots of the depletion rates versus substrate mass consumed
indicated that the stationary condition had not been achieved. Therefore, to
characterize the rates achieved in these microcosms we chose, for each compound,
the ratio of the depletion rate of the third CAH pulse to the corresponding
aqueous-phase concentration. Consequently, the microcosms set up at time 3 with
aquifer material from site E (which were operated for 7 CAH pulses) were
characterized according to this criterion in the elaborations where they were
compared with the corresponding microcosms containing materials from sites A,
B, C and D, whereas in the elaborations where they were compared with other
microcosms that had achieved the stationary condition, they were characterized
according to the general criterion of the stationary CAH depletion rate.
In the case of the liquid-phase microcosms, the procedure described with
regard to the slurry microcosms was applied to the CAH specific degradation
rates, obtained by dividing each CAH rate by the corresponding biomass
concentration measured at the beginning of the CAH pulse. The ratio of specific
depletion rate to concentration is indicated in this study with ksp*
124
Growth of CAH-degrading Consortia in Methane- and Propane-fed Bioreactors
The lag-times and depletion rates reported in the Results relatively to the
duplicate microcosm are averages of the corresponding values estimated in the
two duplicates.
5.2.5 Analytical methods
Gas Chromatography Analysis
The gas-phase concentrations of methane, propane and CAHs were
measured with a HP6890gaschromatograph equipped with a capillary HP-VOC
column connected to a Flame Ionisation Detector (FID) for the analysis of
methane, propane and VC and to a micro Electron Capture Detector (-ECD) for
the analysis of the remaining CAHs; the instrument and the method characteristics
are reported in Table 5.6. Detection limits were (M in the aqueous phase):
methane and propane, 0.007; VC, 1.2; trans-DCE, 0.08; cis-DCE, 0.15; TCE,
4x10-6; 1,1,2-TCA, 0.01; 1,1,2,2-TeCA, 0.02.
Table 5.6. GC Method for methane, propane and the 6 CAHs mixture’s analysis
Instrument
Column
Column I.D.
Column length
Liner
Split ratio
Front detector
Back detector
Carrier gas
Make up gas
Flow
Injection volume
Pressure
Injector temperature
Detector temperature
Initial temperature
Ramp
Final temp
Run time
HP 6890 serie II plus
HP-VOC capillary column
0.32 mm
30 m
Splitless
10:1
µECD
FID
Helium
Nitrogen
0.9 ml/min
500 μL
1.6 Bar
250°C
250°C
60°C (3 min)
20°C/min to 230°C
230°C (5 min)
16.5 min
Total masses and aqueous phase concentrations in standards and slurry
microcosms were calculated utilizing the gas/liquid and solid/liquid equilibrium
constants estimated at 25°C. The following equations describe the equilibrium:
125
Growth of CAH-degrading Consortia in Methane- and Propane-fed Bioreactors
mi = (cL,i • VL + cG,i • VG )
(5.1)
cG,i = Hi • cL,i
(5.2)
that implies:
cL,i = mi /(VL + Hi • VG)
(5.3)
where:
VL, VG: liquid and gas volume in the standard
Hi : dimensionless Henry constant for the compound i (Sanders, 1999)
mi: mass of the compound i (mg)
cG,i, gas and liquid concentration of compound i
Concerning the evaluation of the compound concentration into the microcosms,
these have been based on the gas-phase concentration by headspace analysis. Soil
and liquid-phase concentration have been evaluated by assuming equilibrium
between the phases. The calculation has been based on equation 5.2 and on the
following linear absorption isotherm (Semprini, 2000):
cS,i = Kd,i • cL,i
(5.4)
where:
cS,i : concentration of compound i in the solid phase (soil) related to dry soil mass
(mg/kg dry soil)
Kd,i : adsorption constant of compound i (L/kg)
The adsorption constant has been evaluated on the basis of the soil organic carbon
content (foc, dimensionless) and of the value of the carbon-water partition constant
(Koc, L/kg):
Kd,i = foc • Koc,i
(5.5)
126
Growth of CAH-degrading Consortia in Methane- and Propane-fed Bioreactors
The amount of compound (substrate or contaminant) in the microcosm has then
been measured (starting from the gas-phase concentration) through the following
relation:
mi = cL,i • (VL + Hi • VG + foc • Koc,i • MT)
(5.6)
where MT (kg) is the soil mass in the microcosm, while cL,i has been evaluated
through the relation (5.2).
Bacterial counts
Serial dilution of biomass suspension sampled from growth bioreactors
were plated on Petri dishes containing R2A medium. Bacterial colonies grown on
R2A agar plates were grouped in different clusters and were counted on the basis
of their different morphologies after a 5-7 days incubation at 30°C; the viable
cells concentration was expressed as colonies forming units per ml of suspension
(CFU/ml). R2A medium contained: yeast extract (500 mg/L), caseine hydrolyzed
(500 mg/L), thiotone/peptone (500 mg/L), Glucose (500 mg/L), Sodium Piruvate
(300 mg/L), Na2HPO4 (300 mg/L), MgSO4·7H2O (30 mg/L); Agar (gelificant
agent, 15 g/L).
Measure of CO2 and O2 concentration in the headspace
In order to evaluate the consumption of O2 and the release of CO2
correlated with the consumption of primary substrate, 1 ml of microcosms
headspace gas has been sampled and analysed in a VARIAN 3300 gas
chromatograph equipped with a TCD detector (Thermal Conductivity Detector)
and a packed column CARBONSIEVE SII SS. Injector temperature was 150°C ,
filament temperature, 250°C and detector temperature, 220°C. The temperature
program was as follows (Figure 5.5): 5 min at 60 °C; ramp to 220°C at 10°C/min;
14 min at 220°C.
127
Growth of CAH-degrading Consortia in Methane- and Propane-fed Bioreactors
Figure 5.5. Temperature oven ramp for CO2 and O2 analyses
TEMPERATURE OVEN RAMP [GC VARIAN]
250
225
200
T (°C)
175
150
125
100
75
50
0
5
10
15
20
25
30
35
t (min)
Total protein concentration was evaluated as described by Peterson (1977).
Concentration of Cl- was measured by Ion Chromatography with the method
described by Frascari et al.(2006).
128
Growth of CAH-degrading Consortia in Methane- and Propane-fed Bioreactors
5.3 RESULTS
5.3.1 CAH depletion in the sterilized controls
The ratios of CAH initial abiotic depletion rate to initial concentration (kst*)
evaluated in the sterile controls are reported in Table 5.7. The comparison
between these rates and the corresponding rates obtained in the viable microcosms
is reported and commented in sections 5.3.3 and 5.3.4. CAH depletion in the
sterilized controls followed a first-order kinetic. The best estimates of the firstorder constants, reported in Table 2, correspond to abiotic half-lives varying
between 2 months (for TCE and 1,1,2,2-TeCA) and 2.5 years (for cis-DCE).
Table 5.7. Abiotic rate/concentrations ratios (kst*) esatimated in the sterilized controls STA1,2 and ST-E1,2 operated at 25°C (average values).
129
Growth of CAH-degrading Consortia in Methane- and Propane-fed Bioreactors
5.3.2 Behaviour of the duplicate slurry microcosms
All the duplicate slurry microcosms led to equal results (with deviations <
10%) in terms of both lag-times for the onset of CAH biodegradation and CAH
depletion rates. As an example, the plot of CAH concentration versus time relative
to the first 13 days of operation of microcosms S-P-NC-1A1 and S-P-NC-1A2 is
shown in Figure 5.6.
25
20
3
15
2
VC (µM)
Trans-1,2 DCE, cis-1,2 DCE, TCE,
1,1,2-TCA, 1,1,2,2-TeCA (µ M)
4
10
1
5
0
0
0
2
4
6
8
Time (d)
10
12
Trans-1,2 DCE (duplicate 1)
Trans-1,2 DCE (duplicate 2)
Cis-1,2 DCE (duplicate 1)
Cis-1,2 DCE (duplicate 2)
TCE (duplicate 1)
1,1,2-TCA (duplicate 1)
TCE (duplicate 2)
1,1,2-TCA (duplicate 2)
1,1,2,2-TeCA (duplicate 1)
1,1,2,2-TeCA (duplicate 2)
VC (duplicate 1)
VC (duplicate 2)
Figure 5.6. Aqueous phase concentrations of the 6 CAHs versus time during the first 13 days
of operation of duplicate microcosms S-P-NC-1A1 and S-P-NC-1A2. For higher clarity the
propane pulses, supplied daily at 46 M and rapidly consumed, are not represented.
5.3.3 Effect of inoculum growth time and condition (presence/absence of
CAHs)
The lag-times for the onset of substrate utilization and CAH mixture
biodegradation obtained in the parent non-bioaugmented microcosms relative to
the previous study (Frascari et al., 2006) and in the microcosms set up at times 0,
130
Growth of CAH-degrading Consortia in Methane- and Propane-fed Bioreactors
1 and 2 with aquifer materials from site A are reported in Figure 5.7, whereas the
corresponding k* are shown in Figure 5.8 in normalized form, having equalized to
1 the values corresponding to the parent non-bioaugmented microcosms. The
actual ratios k* relative to these microcosms are reported in the left-hand part of
Table 5.8.
Figure 5.7 shows in the first place that the both inocula utilized at time
zero to set up the growth bioreactors led to a drastic decrease of the lag-time for
the onset of biodegradation of the entire 6-CAH mixture upon introduction in the
microcosms containing aquifer materials form site A (microcosms S-M-C-0A and
S-P-C-0A: from 100-200 days in the non-bioaugmented tests to 2-4 days in the
inoculated ones). This important result indicates that, in case of an in-situ
cometabolic bioremediation of site A, the utilization of either inoculum is
fundamental in order to rely on a fast onset of the remediation process and on a
significant saving on substrate costs. Figure 5.7 also shows that, for both
consortia, the 150-day growth process in the presence as well as in the absence of
the selective pressure exerted by the CAH mixture did not lead to any significant
loss of the capacity to induce the rapid onset of biodegradation of the CAH
mixture, indicating that an eventual process of production of large amounts of
biomass to utilize for a real-scale bioaugmentation treatment can be operated in
the absence of the CAHs, with a significant simplification of the production plant
and a reduction of the fixed and operational costs. A possible explanation for this
experimental result is the fact that the inocula initially supplied to the growth
bioreactors had previously been subjected to a prolonged period of CAH
biodegradation in the slurry microcosms operated within the previous study (410
days for the methanotrophs, 310 days for the propanotrophs), during which they
had been strongly stabilized by the CAH selective pressure. Lastly, it can be
observed in Figure 5.7 that in the non-bioaugmented microcosms the lag-time
basically coincides with the time required by the site’s indigenous biomass to start
growing on the primary substrate, whereas in the inoculated tests the short lagtime is in some cases a substrate lag, and in others a CAH lag.
131
Growth of CAH-degrading Consortia in Methane- and Propane-fed Bioreactors
1000
1000
a - Metanotrophs
b - Propanotrophs
CAH mixture lag-time
10
CAH mixture lag-time
100
Substrate lag-time
Lag-time (d)
Lag-time (d)
100
Substrate lag-time
10
1
1
0,1
0,1
Time 2 – Inoculum grown in the
absence of CAHs (S -P-NC -2A)
Time 1 – Inoculum grown in the
absence of CAHs (S -P-NC -1A)
Time 2 – Inoculum grown in the
presence of CAHs (S -P-C -2A)
Time 1 – Inoculum grown in the
presence of CAHs (S -P-C -1A)
Time 0 – Initial inoculum
(S-P-C-0A)
Parent non -bioaugmented
microcosms
Time 2 –
Inoculum grown in the
absence of CAHs
(S-M -NC -2A)
Time 2 –
Inoculum grown in the
presence of CAHs
(S-M -C-2A)
Time 0 –
Initial inoculum
(S-M-C-0A)
Parent non bioaugmented
microcosms
Figure 5.7. Lag-times for the onset of primary substrate utilization and CAH mixture
biodegradation by methane-utilizing (a) and propane-utilizing (b) biomasses in the parent
non-bioaugmented microcosms, in the microcosms bioaugmented with the initial inoculum
(time 0) and in those bioaugmented with biomass sampled from the growth reactors at times
1 (30 days) and 2 (150 days). All the data refer to microcosms containing aquifer materials
from site A, and to the CAH initial concentration typical of site A (Table 5.8).
The k* reported in Figure 5.8 show that, for both consortia, the prolonged
growth process in the presence as well as in the absence of the CAH mixture did
not lead to any decrease of the long-term CAH depletion rates obtained in the
inoculated microcosms set up with materials from site A, in comparison with the
rates obtained inoculating the same type of microcosms with the initial inoculum
(tests set up at time zero: S-M-C-0A and S-P-C-0A). Besides, with regard to
trans- and cis-DCE, TCE and 1,1,2-TCA, in the microcosms inoculated at time
zero we obtained depletion rates higher than those measured in the parent nonbioaugmented microcosms: this result may be explained by considering that each
of the initial inocula was obtained by mixing biomass samples from three slurry
microcosms, which may have resulted in the formation of two consortia
combining the best degradation capacities of the biomasses they originated from.
132
Growth of CAH-degrading Consortia in Methane- and Propane-fed Bioreactors
6
a - Methanotrophs
k* / knb*
5
4
3
2
1
0
VC
transDCE
cis-DCE
TCE
1,1,2TCA
1,1,2,2TeCA
P arent no n-bio augmented micro co sms
Time zero - Initial ino culum (S-M -C-0A )
Time 2 - Ino culum gro wn in the presence o f CA Hs (S-M -C-2A)
Time 2 - Ino culum gro wn in the absence o f CA Hs (S-M -NC-2A)
6
b - Propanotrophs
k* / knb*
5
4
3
2
1
0
VC
transDCE
cis-DCE
TCE
1,1,2TCA
1,1,2,2TeCA
P arent no n-bio augmented micro co sms
Time zero - Initial ino culum (S-M -C-0A )
Time 1- Ino culum gro wn in the presence o f CA Hs (S-P -C-1A )
Time 1- Ino culum gro wn in the absence o f CA Hs (S-P -NC-1A )
Time 2 - Ino culum gro wn in the presence o f CA Hs (S-P -C-2A)
Time 2 - Ino culum gro wn in the absence o f CAHs (S-P -NC-2A )
Figure 5.8. Normalized ratios of depletion rate to concentration(k*) relative to the
biodegradation of the 6 CAH mixture by methane-utilizing (a) and propane-utilizing (b)
biomasses in the parent non-bioaugmented microcosms, in the microcosms bioaugmented
with the initial inoculum (time 0) and in those bioaugmented with biomass sampled from the
growth reactors at times 1 (30 days) and 2 (150 days). The k*relative to the parent nonbioaugmented microcosms (knb*) were equalized to 1, and their actual value is reported in
the left-hand part of Table 5.8. All the data refer to microcosms containing aquifer materials
from site A, and to the CAH initial concentrations typical of site A (Table 5.8).
133
Growth of CAH-degrading Consortia in Methane- and Propane-fed Bioreactors
Combining the results shown in Figures 5.7 and 5.8, it can be stated that the
prolonged growth process of both inocula in the absence of CAHs allowed to
attain, as a result of the inoculation in type-A slurry microcosms, CAH
biodegradation rates equal or higher than those obtained in the non-bioaugmented
microcosms, with drastically shorter lag-times for the onset of the bioremediation
process.
As evidenced in our previous study (Frascari et al., 2006), the k* obtained
in the non-bioaugmented microcosms and reported in the left-hand part of Table
5.8 are characterized by a tendency to a decrease as the number of chlorine atoms
in the solvent increases, with – in the case of the methanotrophs – a difference of
two order of magnitude passing from VC to 1,1,2,2-TeCA. This tendency was
maintained during the consortia growth process in the four bioreactors.
Table 5.8. CAH depletion rate / concentration ratios relative to the microcosms utilized as
references for the normalized data reported in Figures 5.8 and 5.10 (day-1).
Slurry microcosms set up at time 3
Parent non-bioaugmented
with soil E and inoculated from
a
microcosms (Frascari et al., 2006) growth bioreactors GB-M-NC or
CAH
GB-P-NCb,c
S-M-NC-3E
S-P-NC-3E
Methane-fed
Propane-fed
(methane-fed)
(propane-fed)
d
d
VC
29
24
trans-DCE
10
0.59
1.7
1.1
cis-DCE
3.1
8.3
1.6
12
TCE
0.40
1.6
1.8
6.1
1,1,2-TCA
0.44
5.7
1.5
9.2
1,1,2,2-TeCA
a
0.29
1.1
0
1.0
Initial concentration in the pulses: VC 25 µM; trans-DCE 3.4 µM; cis-DCE 3.1 µM; TCE 1.9 µ
M; 1,1,2-TCA 0.30 µM; 1,1,2,2-TeCA 0.15 µM.
b
Initial concentration in the pulses: 4 µM for all the 5 CAHs.
c
The CAH depletion rates utilized for the microcosms set up at time 3 are - unlike those utilized
for the parent non-bioaugmented tests - those of the third CAH pulse, when the microcosms were
not yet in a stationary conditions in terms of CAH rates.
d
VC was not included in the microcosms set up at time 3.
134
Growth of CAH-degrading Consortia in Methane- and Propane-fed Bioreactors
5.3.4 Effect of the type of aquifer material contained in the inoculated
microcosms
Figure 5.9 reports the substrate and CAH mixture lag-times obtained in the
non-bioaugmented methane-fed and propane-fed microcosms containing aquifer
materials from sites A, B, C, D and E (and spiked with the 5-CAH mixture, with
each compound at 4 M) and in the corresponding microcosms inoculated at time
3 from bioreactors GB-M-NC and GB-P-NC (300 days of consortia growth in the
absence of CAHs). The CAH lags relative to the methanotrophs (Figure 5.9a) do
not include 1,1,2,2-TeCA as, after a monitoring time of at least 100 days, the
biodegradation of this solvent (at the initial concentration of 4 M) was observed
neither in the non-bioaugmented microcosms nor in the inoculated ones. Figure
5.9 shows that the inoculation of the two consortia led to drastic lag-time
reductions in the case of aquifers A and E, and to less marked – but not negligible
– reductions in the case of aquifers B, C and D (in the worst case, occurred with
the inoculation of the propanotrophs in aquifer material type B, we observed a 2fold lag-time reduction). The short lag-times obtained in the inoculated
microcosms indicate that the prolonged growth process of the two consortia in the
absence of CAHs did not lead - with the exception of 1,1,2,2-TeCA for the
methanotrophs - to any significant loss of their ability to induce the rapid onset of
CAH biodegradation. The lack of 1,1,2,2-TeCA biodegradation in the methanefed inoculated microcosms is probably to be ascribed to the higher initial
concentration of this compound in the microcosms of times 3 with respect to those
inoculated at previous times (4 versus 0.15 M), rather than to a loss of 1,1,2,2TeCA degradation capacity of the methane-utilizing consortium during the growth
process in GB-M-NC. This hypothesis, although not fully demonstrated within
this study, is partly supported by the observation that, while in the non-inoculated
microcosms containing A-type aquifer material and 1,1,2,2-TeCA at 0.15 M we
observed the biodegradation of this compound shortly after the onset of methane
consumption, in the non-inoculated tests containing the same aquifer material and
1,1,2,2-TeCA at 4 M no biodegradation of this solvent occurred, indicating that
the microbial capacity to transform 1,1,2,2-TeCA is affected by its concentration.
135
Growth of CAH-degrading Consortia in Methane- and Propane-fed Bioreactors
The results shown in Figure 5.9 relatively to the non-bioaugmented
microcosms containing soil E confirm the observation that when particularly long
overall lag-times are observed, they are due mainly to the time required by the
site’s indigenous biomass to start growing on the primary substrate.
1000
CAH mixture lag-time
Substrate lag-time
a - Metanotrophs
Lag-time (d)
100
10
1
0,1
Aquif er material E–
Inoculum
Aquif er material E–
Indigenous biomass
Aquif er material D–
Inoculum
Aquif er material D–
Indigenous biomass
Aquif er material C–
Inoculum
Aquif er material C–
Indigenous biomass
Aquif er material B–
Inoculum
Aquif er material B–
Indigenous biomass
Aquif er material A–
Inoculum
Aquif er material A–
Indigenous biomass
1000
CAH mixture lag-time
b - Propanotrophs
Substrate lag-time
Lag-time (d)
100
10
1
Aquif er material E–
Inoculum
Aquif er material E–
Indigenous biomass
Aquif er material D–
Inoculum
Aquif er material D–
Indigenous biomassv
Aquif er material C–
Inoculum
Aquif er material C–
Indigenous biomass
Aquif er material B–
Inoculum
Aquif er material B–
Indigenous biomass
Aquif er material A–
Inoculum
Aquif er material A–
Indigenous biomass
0,1
Figure 5.9. Lag-times for the onset of primary substrate utilization and CAH mixture
biodegradation by methane-utilizing (a) and propane-utilizing (b) biomasses in nonbioaugmented microcosms set up with five different aquifer materials and in the
corresponding microcosms bioaugmented with biomass sampled in growth reactors GB-MNC and GB-P-NC (microbial growth in the absence of CAHs) at time 3 (300 days of biomass
growth). VC was not spiked in the microcosms of time 3. In part (a) the CAH lag-times refer
136
Growth of CAH-degrading Consortia in Methane- and Propane-fed Bioreactors
to the time for the onset of the transformation of trans- and cis-DCE, TCE and 1,1,2-TCA, as
no 1,1,2,2-TeCA biodegradation was observed in the methane-fed tests of time 3.
Figure 5.10 reports the k* relative to the methane-fed and propane-fed
microcosms set up with the different aquifer materials and inoculated at time 3
from GB-M-NC and GB-P-NC. Given the wide ranges of variations obtained, the
rate/concentration ratios relative to the microcosms containing E-type materials
were made equal to 1, and their actual value is reported in the right-hand side of
Table 5.8. As explained in section 5.2.4, the CAH depletion rates utilized to build
Figure 5.10 are - unlike those of Figure 5.8 - those of the third CAH pulse, when
the microcosms were not yet in a stationary condition in terms of CAH rates. The
data summarized in Figure 5.10 and in Table 5.3 show that – with the exception of
1,1,2,2-TeCA for the methanotrophs – the two inocula, after the prolonged growth
process in the absence of CAHs, were able to induce in all the aquifer materials
tested CAH depletion rates analogous to those obtained in the parent nonbioaugmented microcosms. Overall, the bioaugmented microcosms containing Etype materials resulted in the highest CAH degradation rates; for this reason, as
well as for our interest in site E as a non historically contaminated site, these
microcosms were operated for a significantly longer time than the other
microcosms of time 3 (7 versus 3 CAH pulses), and were eventually utilized as
the source of the inocula for the liquid-phase microcosms of time 4.
137
Growth of CAH-degrading Consortia in Methane- and Propane-fed Bioreactors
1,4
1,2
a - Metanotrophs
k* / kE*
1
0,8
0,6
0,4
0,2
0
transDCE
cis-DCE
TCE
Aquifer material A
Aquifer material C
Aquifer material E
1,1,2TCA
1,1,2,2TeCA
Aquifer material B
Aquifer material D
1,4
1,2
b - Propanotrophs
k* / kE*
1
0,8
0,6
0,4
0,2
0
transDCE
cis-DCE
TCE
Aquifer material A
Aquifer material C
Aquifer material E
1,1,2TCA
1,1,2,2TeCA
Aquifer material B
Aquifer material D
Figure 5.10. Normalized depletion rate to concentration ratios (k*) for the biodegradation of
the 5-CAH mixture by methane-utilizing (a) and propane-utilizing (b) biomasses in the
microcosms set up with different aquifer materials and bioaugmented with biomass sampled
from growth reactors GB-M-NC and GB-P-NC at time 3. The k* relative to the E-type
microcosms (kE*) were equalized to 1 and their actual value is reported in the right hand
part of Table 5.8.
138
Growth of CAH-degrading Consortia in Methane- and Propane-fed Bioreactors
The ratio of the kst* evaluated in sterile controls ST-E1,2 (spiked with TCE,
1,1,2-TCA and 1,1,2,2-TeCA at 4 M) to the corresponding k* obtained in the
microcosms of time is < 1% for 1,1,2-TCA, < 3% for 1,1,2,2-TeCA and < 5% for
TCE. This result confirms the minor contribution of abiotic reactions and losses
through the caps to the depletion rates observed in the viable microcosms.
The combined results shown in Figures 5.9 and 5.10 indicate that both the
inocula object of this study have a high potential for the bioaugmentation of
CAH-contaminated sites, even in cases where the site’s indigenous biomass has
not been previously affected by the selective pressure due to a historical CAH
contamination. The lack of 1,1,2,2-TeCA biodegradation at 4 M by the
methanotrophs indicates, in agreement with numerous literature studies (Kim et
al., 2000; U.S. E.P.A, 2000; Chen et al., 1996), that the aerobic biodegradation of
TeCA is a problematic process and that propane is more effective than methane in
inducing the microbial degradation of this compound.
5.3.5 Effect of the type of microbial consortium inoculated and growth
substrate supplied
Figure 5.11a shows the k* relative to the three types of slurry microcosms
set up with type-A aquifer material and inoculated at time 2 with biomass grown
in the absence of the CAH mixture: S-M-NC-2A (inoculated with methane-grown
biomass and fed with methane), S-P-NC-2A (inoculated with propane-grown
biomass and fed with propane) and S-MP-NC-2A (inoculated with both methanegrown and propane-grown biomass and fed with both substrates). Similarly, Fig.
11b shows the ratios of depletion rate to concentration relative to the three types
of slurry microcosms set up with type-E aquifer material and inoculated at time 3
with biomass grown in the absence of the CAH mixture: S-M-NC-3E, S-P-NC-3E
and S-MP-NC-3E. The CAH rates utilized to build Figure 5.11b, unlike those
utilized for Figure 5.10 and for the right-hand part of Table 5.8, are long-term
rates obtained when the microcosms were in a stationary condition in terms of
CAH depletion rates.
139
Growth of CAH-degrading Consortia in Methane- and Propane-fed Bioreactors
It can be observed in the first place that the “single-substrate” consortia
(methane-fed and propane-fed) are characterized by significantly different
biodegradation capacities with respect to the various CAHs. In particular, while
the methanotrophs, at both inoculation times and with both aquifer materials, were
14 times more effective on trans-DCE than the propanotrophs, the latter were
significantly more effective than the methanotrophs on 1,1,2-TCA (with a 9-10
fold advantage), 1,1,2,2-TeCA (with a 2-fold advantage at time 2, when this
compound was supplied at 0.15 µM, and with the lack of TeCA biodegradation by
the methanotrophs at time 3, when it was supplied at 4 µM) and – limitedly to the
A-type microcosms – TCE (with a 5-fold advantage). As for VC, while it was not
supplied in the microcosms of time 3, at time 2 it was characterized by the similar
ratio of depletion rate to concentration in the methane-fed and in the propane-fed
consortia, as can be evinced by comparing the last VC bars in Figure 5.8a and b.
The different but complementary degradation capacities of the two singlesubstrate consortia suggested the idea to test the characteristics of a doublesubstrate consortium. It can be observed in both Figure 5.11a and 5.11b that, for
all the CAHs characterized by a different degradation ability of the two singlesubstrate consortia, the mixed propane/methane-fed consortium behaved like the
best single-substrate consortium.
Interestingly, the k* obtained, for each CAH and for each type of substrate,
by inoculating A-type microcosms at time 2 (Figure 5.11a) and E-type
microcosms at time 3 (Figure 5.11b), are approximately the same, with the
exception of TCE and 1,1,2,2-TeCA for the methanotrophs (a probable
consequence of the increase of concentration, as discussed for TeCA in section
5.3.4). This result indicates that neither the additional 150-day growth period of
the inocula in the absence of the CAH mixture, nor the different type of aquifer
material (and consequently of indigenous biomass) contained in the inoculated
microcosms led – in particular for the propane-fed and methane/propane-fed
consortia – to any significant difference in the type of consortium obtained as a
result of the interaction between the inoculum and the microcosms’ indigenous
biomass. Besides, considering that the A-type microcosms were spiked with CAH
initial concentrations in the 0.15-3.4 µM range (excluding VC), while the E-type
140
Growth of CAH-degrading Consortia in Methane- and Propane-fed Bioreactors
tests contained each CAH at 4 µM, the similarity of the k* represents a
confirmation of the validity of the utilization of the pseudo first-order constant k*
as an index of CAH degradation capacity of the different consortia.
In order to further investigate the effect of the type of growth substrate on
the CAH degradation capacity, in microcosms S-M-NC-3E, S-P-NC-3E, S-MPNC-2A1,2 and S-MP-NC-3E1,2, after 7 pulses of biodegradation of the CAH
mixture we operated a change of the type of substrate supplied: in particular, in
the first two microcosms we inverted the growth substrate, feeding S-M-NC-3E
with propane (46 µM) and S-P-NC-3E with methane (125 µM), whereas in each
double-substrate test we started to feed one duplicate (S-MP-NC-2A1 and S-MPNC-3E1) with only methane (125 M), and the other (S-MP-NC-2A2 and S-MPNC-3E2) with only propane (46 µM). In each of these microcosm, after one week
of feed with the new substrate in the absence of CAHs, we monitored the
biodegradation of one further pulse of the 5- or 6-CAH mixture (supplied at the
same concentration of each compound as in the previous pulses). In the doublesubstrate microcosms (S-MP-NC-2A and S-MP-NC-3E), the supply of only one
substrate did not lead - limitedly to the single CAH pulse monitored - to any
significant change of the CAH depletion rates (data not shown). Conversely, in
microcosms S-M-NC-3E and S-P-NC-3E the inversion of substrate led each
consortium to attain CAH depletion rates analogous to those observed in the
corresponding double-substrate microcosm S-MP-NC-3E: in other words, the
supply of methane in S-P-NC-3E resulted in a marked increase of the trans-DCE
rate and in the maintenance of the depletion rates relative to the other CAHs,
whereas the supply of propane in S-M-NC-3E led to a marked increase of the
1,1,2-TCA rate, to the onset of 1,1,2,2-TeCA biodegradation and, as in S-P-NC3E, to no significant loss of the degradation capacity relative to the other CAHs
(data not shown). While the rapid development of new degradation capacities as a
result of the inversion of substrate represents an interesting result, the
maintenance of the degradation abilities characteristic of the previous substrate
cannot be considered a definitive result: in fact, because it was observed only
relatively to one CAH pulse, it might be a residual of the CAH-transformation
capacities acquired during the period of feed with the initial substrate.
141
Growth of CAH-degrading Consortia in Methane- and Propane-fed Bioreactors
k* (d-1)
100
a - Time 2, aquifer material A
10
1
0,1
transDCE
cis-DCE
TCE
1,1,2TCA
1,1,2,2TeCA
Methane (S-M-NC-2A)
Propane (S-P-NC-2A)
Methane+propane (S-MP-NC-2A)
k* (d-1)
100
b - Time 3, aquifer material E
10
1
0,1
transDCE
cis-DCE
TCE
1,1,2TCA
1,1,2,2TeCA
Methane (S-M-NC-3E)
Propane (S-P-NC-3E)
Methane+propane (S-MP-NC-3E)
Figure 5.11. Depletion rate to concentration ratios (k*) for the biodegradation of the 5-CAH
mixture by biomasses grown on methane, propane and methane + propane in microcosms
set up at time 2 with aquifer material A (a) and at time 3 with aquifer material E (b), and
bioaugmented with biomass sampled from growth reactors GB-M-NC and GB-P-NC
(microbial growth in the absence of CAHs).
142
Growth of CAH-degrading Consortia in Methane- and Propane-fed Bioreactors
5.3.6 Results relative to the liquid-phase microcosms
Fig. 5.12 shows the ratios of specific CAH rate to concentration (ksp*)
relative to the methane-, propane- and methane/propane-fed liquid-phase
microcosms, inoculated with biomass suspension sampled from the corresponding
E-type microcosms after 90 days of CAH biodegradation and spiked with the 5CAH mixture at 4 µM for each compound. The results obtained in the liquidphase tests are highly similar to those of the E-type microcosms. In particular,
with regard to each compound, the mixed propane/methane-fed consortium (LMP-4) behaved like the best single-substrate consortium. Besides, for each CAH,
the ratio of the ksp* of propane-fed L-P-4 to the corresponding ksp* of methane-fed
L-M-4 is about equal to the same ratio calculated for slurry microcosms S-P-NC3E and S-M-NC-3E, as well as for microcosms S-P-NC-2A and S-M-NC-2A, in
terms of non-specific depletion rates: 0.07–0.11 for trans-DCE, about 1 for cisDCE and TCE, 9-10 for 1,1,2-TCA, infinite for 1,1,2,2-TeCA (except the A-type
slurries, possibly due to the lower TeCA concentration). This result suggests that
the different degradation abilities shown in slurry tests – for any given CAH – by
the methane-utilyzers in comparison with the propane-utilyzers are not the mere
result of the attainment of different concentrations of active biomass in the two
types of microcosms. On the contrary, they reflect actual differences in the
specific CAH transformation capacities of the two consortia.
In the liquid-phase microcosms, the utilization of a Cl-free mineral
medium allowed a precise evaluation of the increase in Cl concentration as a
result of CAH dechlorination. The resulting ratios of the Cl - moles actually
produced to the Cl- moles corresponding to the complete dechlorination of the
total amount of CAHs depleted, evaluated in correspondence of the degradation of
6 CAH pulses, are equal to 0.88 for L-MP-4, 0.92 for L-P-4 and 0.90 for L-M-4.
The missing 10% of Cl- moles indicates that further research is needed to evaluate
the type of degradation products as a result of the aerobic cometabolism of CAHs.
143
Growth of CAH-degrading Consortia in Methane- and Propane-fed Bioreactors
k sp* (L / (mgprot. d))
0,1
0,01
0,001
0,0001
trans-DCE
cis-DCE
TCE
1,1,2-TCA
Methane (L-M-4)
Propane (L-P-4)
Methane+propane (L-MP-4)
1,1,2,2TeCA
Figure 5.12. Ratios of specific depletion rate (ksp*) for the biodegradation of the 5-CAH
mixture by biomasses grown on methane, propane and methane + propane in the liquidphase microcosms,
5.4 DISCUSSION AND CONCLUSIONS
In this work we studied the long-term growth process of two CAHdegrading microbial consortia under different experimental conditions, and we
investigate the effectiveness of these consortia as inocula for the operation of
bioaugmentation treatments in different types of aquifers.
Our results show in the first place that methane and propane are effective
growth substrates for the aerobic cometabolism of CAH mixtures. In particular,
the propane-grown biomass proved able to degrade VC, trans- ans cis-DCE, TCE,
1,1,2-TCA and 1,1,2,2-TeCA at all the concentrations tested, whereas the
methanotrophs failed to deplete 1,1,2,2-TeCA when its concentration was raised
from 0.15 to 4 µM.
We consider the long-term biodegradation of 1,1,2,2-TeCA up to 4 µM by
the propane-utilyzing biomasses a result of particular significance. In fact, this
high-chlorinated compound as been generally considered in the literature as non-
144
Growth of CAH-degrading Consortia in Methane- and Propane-fed Bioreactors
biodegradable by means of aerobic processes (U.S. E.P.A., 2000; Chen et al.,
1996). Its aerobic cometabolic biodegradation by methane-oxidizing cultures was
evidenced for the first time by Chang and Alvarez-Cohen (1996), whereas
Frascari et al. (2006) had documented in a previous study its long-term
biodegradation by methane-fed and propane-fed biomasses in the 0-0.65 µM
range.
This study also evidenced that, in the case of a large-scale bioremediation
with bioaugmentation, the production of large amounts of biomass starting from
the two inocula object of the investigation can be operated in the absence of the
CAH mixture, with a significant simplification of the production plant and a
reduction of the fixed and operational costs. A possible explanation of this
experimental result id the fact that the inocula initially supplied to the growth
bioreactors had previously been subjected to a prolonged period of CAH
biodegradation in the slurry microcosms operated within the previous study (410
days for the methanotrophs, 310 days for the propanotrophs), during which they
had been strongly stabilized by the CAH selective pressure.
The CAH lag-times and depletion rates obtained in the non-bioaugmented
microcosms set up with different types of aquifer materials indicate that the
indigenous biomasses of different sites can have significantly diverse capacities to
grow on the primary substrate supplied and to start degrading the CAH mixture:
in fact, out of five aquifer materials tested, two resulted in lag-phases of over three
months for the onset of substrate utilization (both with methane and with
propane), whereas in the remaining three the overall lag-times (substrate + CAH
mixture) were equal to two weeks at the most. Conversely, the introduction of
either inoculum led in all the five types of aquifers to very short lag-times (< 4
days) for the onset of CAH degradation. This result indicates, in agreement with
the findings of the previous study (Frascari et al., 2006), that bioaugmentation can
play a crucial role in the successful bioremediation of CAH-contaminated sites
and that, consequently, further research on the production and the stability of
CAH-degrading inocula and on their interaction with the indigenous biomasses of
different aquifers is needed.
145
Growth of CAH-degrading Consortia in Methane- and Propane-fed Bioreactors
Interestingly, in the inoculated microcosms, while the lag-times were
almost independent of the type of aquifer material utilized, the depletion rates
obtained for each CAH with the different aquifer materials differed in some cases
by one order of magnitude, indicating that the chemical, physical and biological
characteristics of the bioaugmented site play a significant role in the long-term
CAH depletion rates achieved. Besides, the short lag-times and the high CAH
depletion rates obtained in the inoculated E-type microcosms show that
bioaugmentation can be successful even in sites whose indigenous biomass has
not been exposed to the selective pressure due to a previous CAH contamination
and is not capable to grow on any of the primary substrates supplied.
Lastly, in the microcosms supplied with both methane and propane we
obtained a microbial consortium combining the degradation capacities of the two
single-substrate consortia. This result suggests that the double-substrate approach
– a novel technique not previously reported to the best of our knowledge in any
study of cometabolic biodegradation – can find useful applications for the
degradation of complex CAH mixtures.
146
CHAPTER 6
6. BIODEGRADATION OF CHLOROBENZENE: STUDY OF
THE CATABOLIC POTENTIAL AND THE STRUCTURE OF
THE
MICROBIAL
COMMUNITY
IN
THE
INTERFACE
BETWEEN GROUNDWATER AND SURFACE WATER
_______________________________________________________________________________
ABSTRACT
The catabolic potential and the structure of the microbial community
present in the interface between groundwater and surface water were studied. The
main goal of this study was to find out whether bacteria present in the interface
are involved in pollutants degradation. Therefore batch degradation tests and
molecular analyses (PCR-DGGE analysis of 16S rRNA gene, catabolic genes,
dsrA gene) were carried out on aquifer material extracted at different depths in the
interface in three locations characterized by different monochlorobenzene
contamination levels. Batch tests were performed under oxygen-limited
conditions in order to study chlorobenzene degradation under the in situ
conditions. The position in the interface did not have any effect on the process and
biodegradation was exclusively limited by a lack of oxygen. Up to 50 mg/l of
monochlorobenzene were consumed in 20 days in both aquifers, and also in
groundwater and surface water, when sufficient oxygen was available (1.5–2
mg/l). 16S rRNA PCR-DGGE analysis were carried out on undisturbed sediment
cores extracted from the three studied locations in different seasons. Results
indicated that the structure of the microbial community changed in function of
depth. Moreover the structure of the community appeared different in the three
locations while significant similarities were observed in samples extracted in each
location in different seasons. Cloning and sequencing allowed to identify the
dominant bands in the DGGE pattern as belonging to the group of Proteobacteria.
It is still unclear if bacteria corresponding to these bands play a role in
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
chlorobenzene degradation. The only degradative gene detected until now is the
mono-oxygenase tmoA. This gene is involved in the degradation of BTEX, which
are structurally similar to monochlorobenzene, thus being probably involved in its
degradation. This hypothesis seemed to be confirmed by the observation that,
after 200 days of incubation, some bands became more visible in tmoA-DGGE
analyses of samples taken from the batch degradation tests.
6.1 INTRODUCTION
The widespread use of chlorobenzenes during the last decades led to their
common occurrence in the environment. Chlorobenzenes are of great concern
because of their toxicity, persistence and accumulation in the food chain (Aelion
et al., 1987). Monochlorobenzene has been identified as priority pollutant by the
U.S. Environmental Protection. Chlorinated aromatic compounds are nondegradable or slowly degradable by microorganism (van der Meer, 1997), thus
being considered among the most problematic categories of environmental
pollutants. Nevertheless, bacteria that are able to use these compounds as sole
source of carbon and energy have been isolated from polluted environments
(Schraa et al., 1986; Spain and Nishino, 1987; van der Meer et al., 1987; Haigler
et al., 1988; Sander et al., 1991; Spiess et al., 1995). Chlorobenzenes are readily
mineralized under appropriate conditions in the laboratory by bacteria isolated
from soil and water (de Bont et al., 1986; Haigler et al., 1988; Reineke et al.,
1984; Schraa et al., 1986; Spain and Nishino, 1987). Field studies on
contaminated sites have shown that river sediments exposed to chlorobenzenes
degraded them faster than sediments from unpolluted sites (Aelion et al., 1987).
Furthermore, chlorinated benzenes are chemically stable in nature, their
photochemical degradation does not play an important role in soil and aquatic
environment. Biological degradation could therefore be considered a feasible
process to eliminate these compounds.
Polluted groundwater in urban and industrial areas often represents a
continuous source of (diffuse) contamination of surface waters. However there are
148
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
strong indications that the interface between groundwater and surface water plays
an important role in the natural degradation of organic contaminants. This is
especially the case for mobile contaminants (such as monochlorobenzene) that are
persistent in anaerobic subsurface environment, but relatively easily mineralised
under more oxidized environmental conditions (Figure 6.1).
anaerobic
Ground water
interface
aerobic
Aerobic
surface water
H+, Cl-, CO2, H2O
micro
(mobile
aliphatics)
HO
Cl
aerobic biodegradation
interface
Cl
Cl
(chlorophenol)
(monochloro- (benzene)
benzene)
(vinylchloride)
anaerobic
Ground water
in-flux of organic chemicals that slowly attenuate under anaerobic
conditions, including natural biodegradation intermediates
Figure 6.1. Natural attenuation at the reactive interface between groundwater and surface
water
The interface is a dynamic ecotone where active exchanges of water and
dissolved material between the stream and groundwater in many porous sand- and
gravel-bed rivers occur (Karaman, 1935; Orghidan, 1959; Sabater and Vila,
1991). It contains a unique invertebrate fauna (Williams and Hynes, 1974) next to
many forms of fungi and microbes that transfer, release, and stabilise different
forms of transient nutrients (Hendricks, 1993). Interfaces are important storage
zones for organic carbon (Bretschko and Moser, 1993) and are generally
characterised by sharp physical and chemical gradients (Fraser and Williams,
1998), thus enabling a broad spectrum of metabolic processes to occur within
small spatial scales. As a consequence the interfaces are often hot spots in
productivity and diversity of organisms (Pusch et al., 1998) and may substantially
149
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
contribute to the carbon, nutrient and energy flow through the river system
(Naegeli and Uehlinger, 1997).
Although nitrogen (Valett et al., 1997) and dissolved organic carbon
biogeochemistry (Vervier and Naiman, 1992) have received much attention in
relation to streambed hydrologic retention and surface/subsurface exchange, not
much is known about the effect of the interface on the degradation of pollutants.
Lendvay and Adriaens (Lendvay and Adriaens, 1999) observed, using multilevel
arrays, that concentrations of methane and chloroethene decreased as groundwater
became increasingly oxidised along the groundwater-surface water interface in
sample
points
impacted
by
infiltration
of
oxygenated
surface
water.
Schwarzenbach et al. (1983) observed the enhanced removal of alkylated and
chlorinated benzenes in the interface. Since under the conditions typical for the
groundwater environment these aromatic compounds do not undergo chemical
reactions at significant rates and since these compounds are also weakly sorbed,
these authors presumed that any elimination must be attributed to biological
transformation and/or mineralisation. Fuller and Harvey (Fuller and Harvey,
2000) observed an enhanced metal uptake in the interface by the analysis of
dissolved-metal streambed profiles and conservative solute tracers.
Although previous studies (Schwarzenbach et al., 1983; Lendvay and
Adriaens, 1999) presumed that the decrease in the concentration of pollutants was
due to the activity of the microbial community present in the interface, abiotic
processes (such as dispersion, adsorption, convection) may as well have been
responsible for this decrease, which is in fact only a dilution of the pollutant, not
resulting in a reduced risk for humanity and ecosystems.
This study aims at investigating the presumed involvement of bacteria
present in the interface in monochlorobenzene degradation in order to understand
if interface can act as a biobarrier towards the infiltrating contaminants. If such
condition really exists in the field and residence times of the polluted groundwater
in the transitional zones are sufficient, the “naturally occurring biobarrier” could
provide a valuable guarantee that pollution is dealt with adequately, thus not
requiring “active” and expensive remediation technology (such as pump and
threat or air-sparging).
150
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
To investigate these issues we therefore carried out batch degradation tests using
sediment material extracted at two different depths in the interface, from three
sampling locations characterized by different level of contamination. Tests were
operated under oxygen-limited conditions similar to those observed in situ.
Furthermore we used different molecular techniques (Polymerase chain reactionDenaturing Gradient Gel Electrophoresis (PCR-DGGE)) to study the structure of
the interface microbial community in three different locations throughout the year
(analysis of seasonal data). We chose three sampling locations with different
levels of contamination to investigate the effect of the presence/absence of the
contaminant on the microbial community, and we analyzed samples extracted in
different times of the year to evaluate the effect of seasonal variations. Due to the
structural similarity between chlorobenzene and BTEX compounds we chose to
investigate if BTEX catabolic genes were present in our samples. In addition we
studied the diversity of sulfate-reducing bacteria in the same samples. Cloning and
sequencing of the dominant DGGE-bands eventually enabled us to identify the
dominant species in the microbial community.
6.2 MATERIALS AND METHODS
6.2.1 TEST SITE AND SAMPLING
We used three sampling places located in an industrial site in the Port of
Amsterdam (The Netherlands), mainly polluted with monochlorobenzene at
different contamination levels (Figure 6.2). The first one is a ditch constructed by
TNO (Toegepast Natuurwetenschappelijk Onderzoek, The Netherlands) and
characterized by high monochlorobenzene concentration in groundwater (5–10
mg/l), the second one (Leendertgracht) is a canal representing a natural interface
situation with lower monochlorobenzene concentration in groundwater (0,3 mg/l),
the third one (Vijver) is a pond chosen as negative control, due to the lack of
monochlorobenzene in the sediment of the interface and in surface water.
151
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
(a)
(b)
(c)
Figure 6.2. The three sampling places in the industrial site in the Port of Amsterdam (The
Netherlands); (a), Artificial ditch; (b), Leendertgracht; (c), Vijver.
152
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
Groundwater was sampled as close as possible to the interface sampling
point. Three different boreholes were used: Q-32 (artificial ditch), A-31/1
(Leendertgracht) and A-18/2 (Vijver). Due to the lack of a closer one we chose
borehole A-18/2 for the Vijver; nevertheless this is one of the wells beside the
canal.
Sediment and water samples were used for molecular tests and batch
degradation tests; physico-chemical parameters (pH, redox potential, dissolved
oxygen concentration, conductivity, temperature) were also measured and water
was analysed to determine the concentration of chlorobenzenes, benzene,
chlorides, electron acceptors and metals.
Samplings were performed on 24/11/2004, 22/03/2005, 31/05/2005,
31/08/2005, 20/04/2006, 22/11/2006 in order to have sediment samples
representative for each season in the year and to study the microbial community
seasonal variations. Table 6.1 shows the tests carried out for each sampling.
Table 6.1. Tests carried out on sediment cores
Sampling point
Date
Artificial ditch
Leendertgracht
Vijver
24/11/2004
(Autumn)
Molecular tests
-
-
22/03/2005
(Winter)
Molecular tests +
batch degradation
tests
Molecular tests +
batch degradation
tests
Molecular tests +
batch degradation
tests
31/05/2005
(Spring)
Molecular tests
Molecular tests
Molecular tests
31/08/2005
(Summer)
Molecular tests
Molecular tests
Molecular tests
20/04/2006
(Spring)
Batch degradation
tests (only water)
Molecular tests +
batch degradation
tests
Molecular tests +
batch degradation
tests
22/11/2006
-
-
-
153
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
6.2.2 Retrieval of the samples
Undisturbed sediment samples were obtained by hammering a 4 cm
diameter plastic tube into the interface (Figure 6.3). Fresh aquifer material
obtained from these cores was used in batch degradation tests: they were cut into
two parts (top and bottom) which were homogenized and used in the setup of the
microcosms. Some of the cores were frozen on dry ice immediately after sampling
in the field in order to be used for molecular analyses. In the laboratory, these
undisturbed frozen samples were cut into slices of approximately 1 cm by using
an electrical saw. Great care was taken in the sterilisation of the saw between each
different cut using ethanol. From each core all the slices from the first 7–10 cm
were selected and those from 10 cm downwards were analysed every 4–5 cm.
From each selected slice sub-samples were used for molecular tests (Figures 6.4
and 6.5) and for the analysis of chlorobenzenes concentration (extraction with
methanol followed by GC-MS analysis). Groundwater was withdrawn by means
of a peristaltic pump (Eijkelkamp, Agrisearch Equipment BV, Giesbeek, The
Netherlands); dissolved oxygen, pH, redox, conductivity and temperature were
measured on site using a multimeter (WTW Multiline P4, Weilheim, Germany) in
a flow through cell (Eijkelkemp Agrisearch Equipment BV, Giesbeek, The
Netherlands) (Figure 6.6). The groundwater level was monitored by means of an
Interface Meter (Eijkelkemp Agrisearch Equipment BV, Giesbeek, The
Netherlands). Groundwater and surface water were collected in 2,5 l bottles and in
40 ml tubes containing 2 g of ascorbic acid. Chlorobenzenes and benzene
concentrations were measured in the laboratory by GC-MS headspace analysis by
pouring 5 g of water from the 40 ml tubes in 10 ml vials and adding 100 l of
85% ortho-phosphoric acid, as explained in paragraph 6.2.5. In the laboratory, the
pellets of a 150 ml sample obtained after centrifugation during 20 min at 7500
rpm were re-suspended in 2 ml of the same water and used in the molecular
analyses described in paragraph 6.2.6.
154
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
Figure 6.3. Extraction of an undisturbed sediment sample
155
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
DNA
extraction
PCR
amplification
total DNA
Amplified
fragments
Agarose
gel
Figure 6.4. Presentation of the strategy followed to study the presence of catabolic genes, 16S
rRNA gene and dsrB gene at different depths in the interface.
Figure 6.5. Extraction of sub-samples for PCR-DGGE analyses from frozen sediment slices.
156
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
(a)
(b)
(c)
(d)
Figure 6.6. Sampling of surface water (a) and groundwater (b) and measure of pH,
temperature, redox potential, dissolved oxygen and conducibility in groundwater (c) and
surface water (d).
157
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
6.2.3 Batch degradation tests of monochlorobenzene
Two groups of microcosms were set up using aquifer material and water
sampled on 22/03/2005 and on 20/04/2006. Tests were performed in 160 ml glass
bottles. 37,5 g of aquifer material were suspended in 70 ml of surface water or
groundwater (Figure 6.7). Both kinds of aquifer materials (top and bottom,
obtained as explained above) were used in order to evaluate the effect of the
position in the interface on the monochlorobenzene degradation potential. Each
type of sediment tests were performed in two different conditions using surface
water or groundwate. Degradation by the community present in groundwater and
in the surface water was investigated in bottles containing just water (85 ml) and
no aquifer material. In the first group of tests (22/03/2005) microcosms flasks
were filled in addition with filter-sterile or non-filter-sterile groundwater to study
monochlorobenzene degradation by bacteria present in the aquifer only. In order
to consider the a-biotic removal of monochlorobenzene control tests were set up
by adding 35 ml of groundwater and 35 ml of surface water to 19 g of each kind
of sediment (top and bottom) and by poisoning bacteria with 800 l of
formaldehyde. Monochlorobenzene was added to the microcosms where a
concentration lower than 1 mg/l was detected, in order to reach a final
concentration of 3 to 4 mg/l (the in situ monochlorobenzene concentration) and
the degradation process was followed in function of time by GC-analysis of the
headspace. Microcosms were set up using materials sampled from each of the
studied locations and incubated statically at room temperature; all tests were
performed in duplicate. Details of the setup are showed in Table 6.2.
Microcosm tests were carried out under anaerobic conditions in an
anaerobic glove box thus avoiding oxygen concentration raising in water and
sediment and starting the test at the same conditions present in the field.
Flasks of the first group of microcosms were incubated for 215 days. Since
monochlorobenzene concentration was constant in all the bottles (except the ones
containing only water) oxygen was added to try to stimulate biodegradation and
the process was monitored for 200 further days. The second group of microcosms
158
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
(20/04/2006) was set up to study the degradation process more in detail
(specifically regarding the effect of oxygen concentration). No tests were
performed with sediment from the Artificial Ditch, since on 20/04/06 it was
impossible to take samples from the interface; from this location tests were carried
out using only surface water or groundwater. Monochlorobenzene and oxygen
concentrations were monitored during the whole incubation period and, since the
former remained stable for more than 30 days, small amounts of oxygen were
added to one of the two duplicate flasks in order to reach the level of 2,5–3 mg/l,
whereas the second duplicate was used as negative control.
Each time monochlorobenzene was completely consumed, microcosms
were re-spiked at the initial concentration. Concentration was then gradually
increased up to 50 mg/l to test the microbial community capacity to degrade high
contaminant concentration. At the end of the experiment DNA extraction
followed by PCR–DGGE analysis was performed on sediment material present in
the bottles and on groundwater/surface water (for tests containing only water), as
explained below. The results obtained were compared with the ones from PCR –
DGGE analysis of sediment and water used to set up the microcosms (Time 0) in
order to evaluate the differences in catabolic genes and changes in the microbial
community, due to the exposure and degradation of monochlorobenzene during
the 200 days.
Table 6.2. Setup of the batch degradation tests
Aquifer
Top
Bottom
Top
Bottom
Top + bottom
Water
Surface water
Groundwater
Surface water
Groundwater
Surface water
Groundwater
Filter-sterile groundwater
(only in 23/03/2005 tests)
Groundwater + surface water
+ formaldehyde
Figure 6.7. Batch degradation test
159
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
6.2.4 GC-analysis of monochlorobenzene concentration in the microcosms
Monochlorobenzene concentrations were measured by headspace analysis
on a CP 3800 Varian gas chromatograph connected with a flame ionization
detector (FID) (type 1079 at a temperature of 250°C) equipped with a Rtx502,2:30 m x 0,53 mm x 3µm and a DB-1:30 m x 0,53 x 5 µm column. Split
injection was implemented at an inlet temperature of 250 °C. Helium was used as
carrier gas at a constant flow rate of 11,9 ml/min. Analyses were carried out using
the following temperature gradient: 2 min at 50°C, ramp to 155°C at 10°C/min,
ramp to 190°C at 20°C/min, 3 min at 190°C. An external standard calibration
curve (one point) was used to calculate the concentrations of the analytes.
6.2.5 Analysis of dissolved oxygen concentration in the microcosms
Dissolved oxygen was monitored by injecting a 300 µL sample in an
oxygen meter (Strath Kelvin Instruments, Glasgow, Scotland) and waiting 2 min
for the stabilization of the instrument (Figure 6.8).
Figure 6.8. Analysis of dissolved oxygen concentration
160
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
6.2.6 Methanol extraction of VOCs and GC-MS analyses
2,5 g of sediment were suspended in 2,5 g of methanol in 10 ml vials;
internal standard (D4-1,2-dichloroethane, D8-toluene, D6-benzene, D4-1,2dichlorbenzene) was added to the vials before putting them into an ultrasonic bath
for 30 min. Methanol containing the extracted VOCs was then diluted by putting
0,5 g of supernatant in 4,5 g water in new 10 ml vials. 100 l of 85 % orthophosphoric acid were also added to kill biomass.
Chlorobenzenes and benzene concentration was determined by GC-MS
headspace-analysis using a Thermo GC-MS equipped with a DB-5ms 60 m x 0,25
mm x 0,25 µm column. The samples were injected into a split/splitless injector at
220°C and put on to the column with a constant flow of 9 ml/min Helium. The
GC-oven program works as follows: 3 min at 38°C, ramp to 175°C at 5°C/min.
The analytes are then detected by the MS-detector. The concentration of each
analyte was calculated through internal standard calibration curve (including eight
different concentrations).
6.2.7 DNA extraction from sediment and water samples
Total genomic DNA from soil and water samples was extracted and
purified as previously described by Hendrickx et al. (2006). Two g of sediment or
the pellet of 2 ml watersample was suspended in 4 ml Tris-glycerol buffer (10
mM Tris, 15 % glycerol, pH = 7). The cells were mechanically lysed by beating
with glass beads (diameter: 0,10 – 0,11 mm) for 2 x 30 sec in a MK4 bead beater
apparatus (Braun Biotech International GmbH, Melsugen, Germany). Before lysis
with proteinase K (32 µl, 20 mg/ml) and 20 % sodium dodecyl sulfate (120 µl)
during 30 min at 50°C, cells were subjected to an enzymatic lysing step with
lysozyme (160 µl, 50 mg/mL) in Tris-glycerol buffer (30 min at 37°C). This was
followed by an addition of 2 ml of NaKPO 4 buffer (conc.: 1,12 M, pH = 8) and a
second bead-beating step for 2 x 30 sec. Glass beads, soil and cell debris were
removed by centrifugation (10 min at 7000 rpm) and the DNA contained in the
aqueous phase was extracted twice with 5 ml phenol/chloroform/isoamylalcohol
(25:24:1), and purified with 5 ml chloroform/isoamylalcohol (24:1). 0,1 g of
polyvinylpyrrolidone (Sigma-Aldrich, Germany) were added to the DNA solution
161
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
and vials were rotated on a rotating shaker for 30 min. Polyvinylpyrrolidone was
eliminated by centrifugation (10 min at 6000 rpm) and the DNA was precipitated
with 2 volumes of 100% ethanol (Merck KgaA, Darmstadt, Germany) at – 20 °C
overnight. The crude DNA pellet was suspended in 500 µL of sterile water and
purified over a Wizard column (Wizard DNA Clean-Up System, Promega
Corporation, Madison, USA). The purified DNA was recovered in 50 µL of TE
buffer (10 mM Tris, 50 mM EDTA, pH 9) and stored at –20 °C
6.2.8 PCR amplification
Polymerase chain reaction on the extracted DNA was performed in a
volume of 50 µL. A 495 bp eubacterial 16S rRNA gene fragment was amplified
using the primer set GC-63F/518R, described by Marchesi et al. (1998). 1 µL of
1:10 or 1:50 dilution of template DNA was added to 49 µl of PCR mix consisting
of 5 µL of 10x exTaq reaction buffer (20 mM MgCl2), 0,25 µL exTaq Polymerase
(5 U µL-1), 4 µl dNTP (deoxynucleoside triphosphate; 2,5 mM each), 0,25 µL of
both primers and 39,25 µl sterile demineralised water. The exTaq Polymerase,
dNTPs and PCR reaction buffer were purchased from TaKaRa (TaKaRa Shuzo
Co., Biomedical Group, Japan). The PCR profile consisted of an initial
denaturation of 5 min at 94°C, followed by 35 further denaturation cycles of 1
min at 94 °C, annealing of 1 min at 55°C, and elongation for 1 min at 65°C. The
last step included an extension for 5 min at 65°C.
The primer sets for detection of the genotypes tmoA, xylE, todC1-like
genes, cdo, tbuE, and todE are reported in Table 6.3 and were applied in PCR as
described by Hendrickx et al., (2006). Το allow DGGE analysis of the amplicons,
PCR products obtained with the catabolic primer set tmoA-F/tmoA-R were
submitted to a semi nested PCR by using the same primer with a GC clamp at the
forward primer (tmoA-F). These primer sets are used for the detection of catabolic
genes involved in the aerobic degradation of BTEX compounds. Due to the
homology between the structure of the BTEX compounds and the structure of
monochlorobenzene the same primers were also used to detect genes involved in
the degradation of the last one. Primer sets GC-P2060F/DSR4R, described by
162
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
J.Geets et al. (2006), amplifies a 350 bp fragment of the β-subunit dissimilatory
sulfite reductase (dsrB) gene of sulfate-reducing bacteria. The PCR profile
consisted of an initial denaturation of 5 min at 94°C, followed by 40 further
denaturation cycles of 1 min at 94 °C, annealing of 1 min at 55°C, and elongation
for 1 min at 72°C. The last step included an extension for 8 min at 72°C. PCR was
performed on a Biometra thermocycler (Biometra, Göttingen, Germany). 10 µL of
the PCR products were analysed by agarose gel electrophoresis to evaluate their
size and quality, (1,5 % agarose (Invitrogen, Paisley, Scotland, UK), 1 x EY
running buffer (10 x EY-buffer: 0,4 M Tris, 0,02 M EDTA, on pH = 7,9 with
acetate in H2O), 1 hour at 85 V). DNA bands were visualized by ethidium
bromide staining (1 mg/l) (Figure 6.9).
163
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
Table 6.3. PCR primer sets used in this study.
*
A 40 bp GC clamp (5’- CGCCCGCCGCGCGCGGCGGGCGGGGCGGGGGCACGGGGGG-3’) was attached to the 5’ end of forward primers TMOA-F, DSRp2060F and 63F (Muyzer et al., 1993)
Primer pair
Proteins targeted
Sequence
Amplicon
size (bp)
TBMD-F/TBMD-R
Subfamily 1 of α-subunits of hydroxylase
component of multi-component monooxygenases
Subfamily 2 of α-subunits of hydroxylase
component of multi-component monooxygenases
Subfamily 5 of hydroxylase component
of two-component side chain monooxygenases
Electron transfer component of twocomponent side chain mono-oxygenases
Subfamilies D.1.B + D.1.C + D.2.A +
D.2.B + D.2.C of α-subunits of Type D
iron-sulfur multi-component aromatic
dioxygenases
Subfamily I.2.A of catechol extradiol
dioxygenases
Subfamily I.2.B of catechol extradiol
dioxygenases
cdo (U01826) of subfamily I.2.C of
catechol extradiol dioxygenases
tbuE (U20258) of subfamily I.2.C of
catechol extradiol dioxygenases
todE (Y18245), todE (Y18245), tobE
(AF180147) of subfamily I.3.B of
catechol extradiol dioxygenases
Eubacterial 16S rRNA gene
5’-GCCTGACCATGGATGC(C/G)TACTGG-3’
5’-CGCCAGAACCACTTGTC(A/G)(A/G)TCCA-3’
(GC-)TMOA-F/TMOA-R*
TOL-F/TOL-R
XYLA-F/XYLA-R
TODC1-F/TODC1-R
XYLE1-F/XYLE1-R
XYLE2-F/XYLE2-R
CDO-F/CDO-R
TBUE-F/TBUE-R
TODE-F/TODE-R
(GC-)63F/518R*
Reference
640
PCR
annealing
temp (°C)
65.5
5’-CGAAACCGGCTT(C/T)ACCAA(C/T)ATG-3’
5’-ACCGGGATATTT(C/T)TCTTC(C/G)AGCCA-3’
505
61.2
(Hendrickx et al., 2006a)
5’-TGAGGCTGAAACTTTACGTAGA-3’
5’-CTCACCTGGAGTTGCGTAC-3’
475
55
(Baldwin et al., 2003)
5’-CCAGGTGGAATTTTCAGTGGTTGG-3’
5’-AATTAACTCGAAGCGCCCACCCCA-3’
5’-CAGTGCCGCCA(C/T)CGTGG(C/T)ATG-3’
5’-GCCACTTCCATG(C/T)CC(A/G)CCCCA-3’
291
64
(Hendrickx et al., 2006)
510
66
(Hendrickx et al., 2006)
5’-CCGCCGACCTGATC(A/T)(C/G)CATG-3’
5’-TCAGGTCA(G/T)CACGGTCA(G/T)GA-3’
5’-GTAATTCGCCCTGGCTA(C/T)GTICA-3’
5’-GGTGTTCACCGTCATGAAGCG(C/G/T)TC-3’
5’-CATGTCAACATGCGCGTAATG-3’
5’-CATGTCTGTGTTGAAGCCGTA-3’
5’-CTGGATCATGCCCTGTTGATG-3’
5’-CCACAGCTTGTCTTCACTCCA-3’
5’-GGATTTCAAACTGGAGACCAG-3’
5’-GCCATTAGCTTGCAGCATGAA-3’
242
61.5
(Hendrickx et al., 2006)
906
64
(Hendrickx et al., 2006)
255
58
(Hendrickx et al., 2006)
444
60
(Hendrickx et al., 2006)
246
58
(Hendrickx et al., 2006)
5’-CAGGCCTAACACATGCAAGTC-3’
5’-TTACCGCGGCTGCTGG-3’
455
55
Marchesi et al. (1998)
164
(Hendrickx et al., 2006)
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
Figure 6.9. Loading of an agareose gel
6.2.9 Denaturing gradient gel electrophoresis
165
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
Bacterial
diversity
was
examined
by
denaturing
gradient
gel
electrophoresis. In DGGE analysis DNA fragments of the same length but with
different base-pair sequences can be separated, thus obtaining a band pattern in a
denaturing polyacrylamide gel in which each band theoretically corresponds to
one type of bacterium. Eubacterial 16S rRNA gene PCR products obtained with
the primer set GC-63F/518R were analysed in 8% polyacrylamide gels with a
denaturing gradient of 35% to 65% urea-formamide (100% denaturant gels
contain 7 M urea and 40% formamide). PCR products obtained with the GCP2060F/DSR4R primer set were analysed in 8% polyacrylamide gels with a
denaturing gradient (40% to 70%). In both cases, DGGE was performed at a
constant voltage of 120 V for 15 h in 1 x TAE (Tris-acetate-EDTA) running
buffer at 60°C. PCR products obtained with the GC-TMOA-F/TMOA-R primer
set were analysed in 6% polyacrylamide gels with a denaturing gradient (40% to
70%) at a constant voltage of 110 V for 16 h 40 min in1 x TAE running buffer at
60°C. In all cases electrophoresis was performed on an INGENY phorU-2 DGGE
apparatus (INGENY International BV, Goes, The Netherlands). After
electrophoresis, the gels were stained in a 1 x TAE buffer containing 1 x SYBR
Gold nucleic acid stain (molecular Probes Europe BV, Leiden, The Netherlands)
and photographed under UV light with a Pharmacia digital camera system with
Liscap Image Capture 1.0, Pharmacia Biotech, UK). Photo files were processed
and analysed with Bionumerics software (version 2.5, Applied Maths, Kortrijk,
Belgium) (Figure 6.10).
166
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
Figure 6.10. DGGE (Denaturing Gradient Gel
polyacrylamide gel and capture of the DGGE image.
167
Electrophoresis).
Electrophoresis,
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
6.2.10 Cloning, sequencing and analysis of PCR amplified 16S rRNA, tmoA
and dsrB gene
PCR products obtained with the 16S rRNA gene primer set 63F/518R and
with primer sets TMOA-F/TMOA-R and P2060F/DSR4R were cloned into
plasmid vector pCR2.1-TOPO using the TOPO TA cloning kit with the TOP10
One Shot Electrocompetent cells (N.V. Invitrogen SA, Merelbeke, Belgium) as
described in the kit’s protocol. Clones containing recombinant vectors (blue
colonies, Figure 6.11) and forming white colonies on selective agar medium (LB
plates containing 50 – 100 g/ml ampicillin and 40 mg/ml X-gal 2%), were
examined for the presence of the exact insert by PCR using first M13 primer
(delivered with the kit) which confirms whether the exact insert had been found,
followed by a semi nested primer with GC-63F/518R or TMOA-F/TMOA-R or
GC-P2060F/DSR4R.
Cloned fragments were compared with the original soil sample fingerprint
by using DGGE. A selection of clones with different DGGE patterns was
sequenced by VIB Genetic Service Facility (University of Antwerp, Belgium).
Figure 6.11. White and blue colonies on selective agar medium. White colonies contain the
exact insert
168
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
6.3 RESULTS
6.3.1 Monitoring of the in situ physico-chemical parameters
Tables 6.4, 6.5 and 6.6 show the physico-chemical characteristics of
groundwater and surface water in the three studied locations measured at each
sampling. Most of the parameters, except water temperature and oxygen
concentration, remained quite stable in the three locations.
The temperature of the surface water was the lowest in November 2004 in
the Artificial Ditch (5.3°C) while the maximum value was reached in August
2005 (20°C). At this time the difference was also maximum between the three
locations (3°C between the Leendertgracht and the Vijver) while only very slight
differences were noticed in Autumn (Figure 6.12a). In groundwater the lowest
temperature was measured in Winter while in Summer values around 20°C were
measured in all the three locations (Figure 6.12b). In groundwater anaerobic or
microaerophilic conditions prevailed: the highest oxygen concentration was
measured in November 2006 in the Vijver (2.33 mg/l) and in the Artificial Ditch
(2 mg/l). In surface water the oxygen level ranged from 2.15 mg/l (November
2004) to 7.56 mg/l (April 2006) in the Artificial Ditch, from 1.76 mg/l (November
2006) to 9.64 mg/l (March 2005) in the Leendertgracht and from 3.75 mg/l
(November 2006) to 13.09 mg/l (May 2005) in the Vijver. A very high chloride
concentration (up to 3000 mg/L) was measured in the Artificial Ditch and in the
Vijver (surface water). No significant concentrations of N were present in any of
the locations (often behind the detection limit), while significant amounts of SO42
(up to 450 mg/l) were found in the Artificial Ditch and in the Vijver (surface
water). Tables 6.4 and 6.5 show that the main pollutant was monochlorobenzene
(up to 10000 g/L in the Artificial Ditch’s groundwater) and, to a lesser extent,
1,4-dichlorobenzene and benzene. No VOC’s contamination was present in
surface water in the Vijver.
169
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
(a)
25
Temperature (°C)
20
15
10
5
0
11/2004
05/2005
artificial ditch
12/2005
leendertgracht
07/2006
01/2007
vijver
20
(b)
Temperature (°C)
16
12
8
4
0
11/2004
05/2005
artificial ditch
12/2005
leendertgracht
07/2006
01/2007
vijver
Figure 6.12. Monitoring of the temperature in surface water (a) and in groundwater (b) in
different seasons in the three studied locations
170
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
Table 6.4. Physico-chemical parameters of groundwater and surface water in the Artificial Ditch. ND: not detected.
Artificial Ditch
Groundwater (borehole Q-32)
Parameter
Surface water level (m)
Groundwater level (m-mv)
O2 concentration (mg L-1)
Temperature (°C)
pH
Redox (mV)
Conductivity (µS cm-1)
Cl- (mg L-1)
NO32-(mg L-1)
NO22-(mg L-1)
SO42-( mg L-1)
SO32-( mg L-1)
Fe (µg L-1)
monochlorobenzene (µg L-1)
1,4-dichlorobenzene (µg L-1)
benzene (µg L-1)
November
24, 2004
2
0.84
11.8
6.46
-33
8020
2370
1.45
<0.02
399
<0.1
3580
5661
88
788
March
22, 2005
0.23
14.3
6.97
-129
2400
3.6
<0.02
390
0.34
3240
7251
83
271
May
31, 2005
1.83
0.95
11.5
7.08
-158
1644
1800
<0.23
<0.02
450
<0.1
2460
10041
ND
ND
August
31, 2005
1.55
1.57
18.5
6.84
7830
2200
<0.23
<0.02
410
<0.1
1040
ND
ND
Surface water
April
20, 2006
0.88
11.2
7.1
-131
1680
1100
<0.23
<0.02
240
<0.1
<50
4493
ND
ND
November
22, 2006
1
2
13.1
7.37
1810
280
<0.23
<0.02
150
<0.1
380
-
171
November
24, 2004
0.3
2.15
5.3
7.13
89
2070
3590
0.31
<0.02
580
<0.1
192
6
ND
ND
March
22, 2005
0.3
6.1
10.6
6.90
-58
2490
1700
0.56
<0.02
409
<0.1
1220
162
ND
ND
May
31, 2005
4.25
14.5
7.89
3
2060
2600
<0.23
<0.02
360
<0.1
845
209
ND
ND
August
31, 2005
0.4
7.42
20
7.42
8490
2400
<0.23
<0.02
370
<0.1
2050
ND
ND
April 20,
2006
>1
7.56
12.2
8.49
3
1709
810
<0.23
<0.02
280
<0.1
548
ND
ND
November
22, 2006
>1
3.4
9.1
7.65
2220
360
<0.23
<0.02
200
<0.1
1000
-
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
Table 6.5. Physico-chemical parameters of groundwater and surface water in the Leendertgracht.
Leendertgracht
Groundwater (borehole A-31/1)
Parameter
Surface water level (m)
Groundwater level (m-mv)
O2 concentration (mg L-1)
Temperature (°C)
pH
Redox (mV)
Conductivity (µS cm-1)
Cl- (mg L-1)
NO32-(mg L-1)
NO22-(mg L-1)
SO42-( mg L-1)
SO32-( mg L-1)
Fe (µg L-1)
monochlorobenzene (µg L-1)
1,4-dichlorobenzene (µg L-1)
benzene (µg L-1)
November
24, 2004
0.1
<0.02
<0.1
0
-
March
22, 2005
<0.02
<0.1
0
86
-
May
31, 2005
1.18
13.4
7.3
-104
2250
780
<0.23
<0.02
5.7
<0.1
536
287
-
August
31, 2005
0.63
18.7
7.15
2170
410
<0.23
<0.02
26
<0.1
<50
-
Surface water
April
20, 2006
0.63
0.15
8.8
7.59
-203
1151
170
<0.23
<0.02
34
<0.1
1870
15
-
November
22, 2006
0.99
0.62
14.5
7.29
1787
260
<0.23
<0.02
62
<0.1
4400
-
172
November
24, 2004
0.5
3.35
6.6
7.06
50
1390
175
<0.23
<0.02
43.2
<0.1
1030
159
0.7
2.2
March
22, 2005
0.7
9.64
12.5
8.07
108
1052
120
<0.23
<0.02
53
<0.1
587
ND
ND
ND
May
31, 2005
14.2
7.56
-108
1415
200
<0.23
<0.02
100
1.25
7620
1
ND
-
August
31, 2005
4.13
18.8
7.3
1672
260
<0.23
<0.02
4.9
<0.1
3660
ND
ND
-
April 20,
2006
2.8
10.3
7.4
93
1415
210
<0.23
<0.02
28
<0.1
1480
2
ND
-
November
22, 2006
0.35
1.76
8.6
7.53
1760
200
<0.23
0.027
58
<0.1
3700
ND
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
Table 6.6. Physico-chemical parameters of groundwater and surface water in the Vijver
Vijver
Surface water
Groundwater (borehole A-18/2)
Parameter
Surface water level (m)
Groundwater level (m-mv)
O2 concentration (mg L-1)
Temperature (°C)
pH
Redox (mV)
Conductivity (µS cm-1)
Cl- (mg L-1)
NO32-(mg L-1)
NO22-(mg L-1)
SO42-( mg L-1)
SO32-( mg L-1)
Fe (µg L-1)
monochlorobenzene (µg L-1)
March
22, 2005
0.06
9.9
7.46
-222
1678
160
<0.23
<0.02
15
<0.1
97
86
May
31, 2005
13.7
7.4
-143
1604
180
<0.23
<0.02
3.1
<0.1
71
62
August
31, 2005
0.27
19.2
7.15
0
1833
170
<0.23
<0.02
3.7
<0.1
110
-
April
20, 2006
0.52
11.2
7.19
-162
1555
202
<0.23
<0.02
2.4
<0.1
233
92
November
22, 2006
0.4
2.33
13.7
7.15
1567
130
<0.23
<0.02
11
<0.1
360
-
173
March
22, 2005
7.2
13.5
7.78
<0.23
<0.02
<0.1
ND
May
31, 2005
0.86
13.09
17.7
8.11
-10
2420
2800
<0.23
<0.02
440
<0.1
852
ND
August
31, 2005
0.55
9.47
21.6
7.93
7940
2400
<0.23
<0.02
370
<0.1
1060
ND
April 20,
2006
0.43
8.08
11.2
7.83
60
1834
2000
1.4
<0.02
330
<0.1
379
ND
November
22, 2006
0.65
3.75
8.6
7.81
8780
2600
<0.23
<0.02
410
<0.1
450
ND
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
6.3.2 Pollutants concentration at different depths in the sediment cores
High concentrations of monochlorobenzene were measured in cores
extracted from the Artificial ditch. Concentration ranged from 50 to 1000 µg/kg in
November 2004, from 60 to 6000 µg/kg in March 2005, from 500 to 18000 µg/kg
in May 2005 and from 20 to 10000 µg/kg in August 2005. The lowest values were
observed in slices close to the surface, and concentration increased depending on
depth (Figure 6.13a). Not negligible but significantly lower concentrations of 1,4dichlorobenzene were also observed (up to 1000 µg/kg), showing the same trend
in function of depth. Other chlorobenzenes (1,2- 1,3-dichlorobenzene; 1,2,3- 1,2,4
and 1,3,5-trichlorobenzene) were present in low concentrations (mostly one order
of magnitude lower). In March and May low concentrations of benzene were also
measured
(up
to
µg/kg).
100
In
the
Leendertgracht (Figure 6.13b)
monochlorobenzene level generally decreased from the top to the bottom of the
core. The lowest values were measured in cores sampled in August 2005 (0-150 µ
g/kg), while highest ones were observed in May 2005 (10 – 10000 µg/kg). 1,4dichlorobenzene did not exceed 400 µg/kg while the other compounds (1,2- 1,3dichlorobenzene; 1,2,3- 1,2,4 and 1,3,5-trichlorobenzene and benzene) ranged
between 10 and 50µg/kg. In general the other pollutant followed the same trend as
monochlorobenzene, thus increasing in function of depth. No chlorobenzenes or
benzene were detected in sediments cores extracted from the Vijver.
(a)
(b)
1,E+05
1,E+04
1,E+04
1,E+03
1,E+03
1,E+02
1,E+02
1,E+01
1,E+01
1,E+00
1,E+00
0
10
20
30
40
50
60
0
depth (cm)
24/11/2004
22/03/2005
31/05/2005
22/03/2005
31/08/2005
10
20
depth (cm)
31/05/2005
30
31/08/2005
40
50
20/04/2006
Figure 6.13. Monochlorobenzene concentration in undisturbed sediment cores extracted
from the Artificial Ditch (a) and the Leendertgracht (b).
174
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
A direct relation was moreover observed between the concentration of
monochlorobenzene adsorbed to the sediment and the total organic matter. As an
example Figure 6.14 shows organic matter and monochlorobenzene concentration
in function of depth in two cores extracted from the Artificial Ditch and the
Leendertgracht.
MCB concentration (ug/kg)
(a)
(b)
1,E+05
3,0
1,E+04
2,5
1,E+03
2,0
1,E+04
30
25
1,E+03
20
1,E+02
1,E+02
1,5
15
1,E+01
1,E+01
1,0
1,E+00
0,5
1,E+00
0,0
1,E-01
1,E-01
0
10
20
30
40
50
60
5
0
0
depth (cm)
chlorobenzene
10
10
20
depth (cm)
30
40
50
organic matter
Figure 6.14 Total organic matter and monochlorobenzene concentration at different depths
in cores extracted from the Artificial Ditch on 31/08/05 (a) and from the Leendertgracht on
31/05/05 (b).
6.3.3 Degradation of monochlorobenzene under oxygen–limited conditions in
batch degradation tests
Monochlorobenzene degradation was studied under the in situ conditions
by bringing the aquifer material (top or bottom) in contact with groundwater or
surface water sampled in situ.
Tests set up with material sampled on 23/03/2005
Figure 6.15 shows the time-concentration profiles of monochlorobenzene
and oxygen for the microcosms constructed with sediment and water from the
Artificial ditch.
A small decrease in monochlorobenzene concentration was noticed in
these microcosms during the first 30 days of incubation both in the living and
dead control conditions; after this initial period the degradation stopped and the
concentration did not change significantly for 215 days. On day 215, after
microcosms were spiked with oxygen, biodegradation readily started again in all
175
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
living tests but not in the dead controls. The pollutant was in some cases (aquifer
bottom + surface water and aquifer bottom + filter-sterile groundwater) even
completely consumed. In the following 250 days no further oxygen was added
and the contaminant’s concentration remained stable.
Analogous results were observed in the tests set up with material sampled
monochlorobenzene concentration (µg/l)
monochlorobenzene concentration (µg/l)
oxygen concentration (mg/l)
oxygen concentration (mg/l)
from the other two locations (Leendertgracht and Vijver).
16000
14000
T=215 days:
oxygen spiking
12000
10000
8000
6000
4000
2000
0
0
50
100
150
200
250
300
350
400
450
3
2,5
2
1,5
1
Figure 0,5
6.15. Monochlorobenzene and oxygen aqueous phase concentration in the batch tests
carried out with sediment and water sampled in the Artificial Ditch on 23/03/2005.
0
0
50
100
150
200
250
300
350
400
time (days)
aquifer top + surface water
aquifer bottom + surface water
aquifer top + filter-sterile groundwater
aquifer bottom + filter-sterile groundwater
aquifer top + groundwater
aquifer bottom + gro undwater
dead control
176
450
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
Tests set up with material sampled on 20/04/2006
The initial oxygen concentration in the microcosm tests containing both
water and sediment ranged between 2 and 2,5 mg/l. Figure 6.16 shows
monochlorobenzene concentration in function of time in the tests carried out with
material sampled from the Leendertgracht, during the first 35 days of incubation.
Limited monochlorobenzene biodegradation took place during the first 6 days in
all microcosm, then it stopped; from day 6 to day 36 monochlorobenzene
concentration remained stable and oxygen concentration stabilized at 1,5 – 2 mg/l.
5000
4000
3000
2000
1000
0
0
5
10
15
0
5
10
15
20
25
30
35
40
20
25
30
35
40
3
2
1
0
Time (d)
aquifer top + surface w ater
aquifer bottom + surface w ater
aquifer top + groundw ater A/31-1
aquifer bottom + groundw ater A/31-1
dead control
Figure 6.16. Monochlorobenzene and oxygen aqueous phase concentration in the batch tests
carried out with sediment and water sampled in the Leendertgracht on 20/04/2006 (first 35
days of incubation).
177
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
On day 37 oxygen was therefore added (2.5-3 mg/l) to the first duplicate of
each type of microcosm (duplicate 1), resulting in an immediate onset of the
biodegradation, which stopped when the oxygen concentration dropped to 1,5–2
mg/l. Each time degradation stopped the spiking with a low volume of oxygen (to
increase the concentration of 1 ppm) proved to be effective to stimulate
biodegradation. Surprisingly, in the condition “aquifer top+groundwater” we
observed a slow but total degradation also in duplicate 2, with oxygen
concentrations
below
1.5
mg/l.
The
complete
degradation
of
the
monochlorobenzene originally present required the addition of 17 mg (average
value of the 4 microcosms aquifer top/bottom + groundwater, aquifer top/bottom
+ surface water) of oxygen altogether. Considering the total amount of
monochlorobenzene provided throughout the 200 days of incubation, 16 mg of
oxygen were needed for eachmg of monochlorobenzene added (average value of
the
microcosms
mentioned
above).
During
this
incubation
period
monochlorobenzene was re-spiked each time it was completely degraded and
concentrations were gradually increased up to 45–50 mg/l. Even at these
concentrations the contaminant was consumed with no evident inhibition effects.
Oxygen
concentration
was
apparently
the
only
factor
limiting
biodegradation: the process stopped each time oxygen concentration dropped
under 1,5–2 mg/l, regardless the position in the interface (top or bottom of the
sediment) and the type of water (groundwater or surface water). On the contrary,
no degradation was observed in the living microcosms without oxygen spiking,
where the same behaviour as in the a-biotic controls was observed. (Figure 6.17).
Similar results were obtained in tests performed using samples from the
non polluted location (Vijver), except for the smaller oxygen amount required to
biodegrade monochlorobenzene: 3.3 mg of oxygen were necessary to oxidize the
initial amount of monochlorobenzene and 2.5 mgoxygen/mgmonochlorobenzene were
consumed on the whole. In the condition “aquifer top+groundwater” degradation
only started when oxygen concentration was increased to 3.5-4 mg/l.
178
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
monochlorobenzene concentration
(µg/l)
duplicate 1
5000
5000
4000
4000
3000
3000
2000
2000
1000
1000
0
0
oxygen concentration
(mg/l)
20
40
60
80
100
3,0
3,0
2,5
2,5
2,0
2,0
1,5
1,5
1,0
1,0
0,5
0,5
0
20
40
60
0
20
40
80
100
0,0
0,0
0
20
40
60
80
100
60
80
100
Time (d)
Time (d)
monochlorobenzene concentration
(µg/l)
a-biotic control
aquifer top+groundwater
aquifer top+surface water
0
aquifer bottom+groundwater
aquifer bottom+surface water
5000
5000
4000
4000
3000
3000
2000
2000
1000
1000
0
0
0
oxygen concentration
(mg/l)
duplicate 2
20
40
60
80
0
100
3,0
3,0
2,5
2,5
2,0
2,0
1,5
1,5
1,0
20
40
60
80
20
40
60
80
100
1,0
0,5
0,5
0,0
0
20
40
60
80
0,0
100
0
Time (d)
Time (d)
Figure 6.17. Monochlorobenzene and oxygen aqueous phase concentration in the duplicate
batch tests carried out with sediment and water sampled in the Leendertgracht on
20/04/2006 (100 days of incubation).
179
100
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
Tests carried out with groundwater or surface water (with no sediment),
show that initial oxygen concentration was higher than in microcosms with
sediment (2,5–5,5 mg/l) and oxygen was consumed more slowly, thus inducing
very fast degradation of the added monochlorobenzene in water sampled from all
the studied locations: 5 mg/l of monochlorobenzene were degraded in less than 5–
6 days; in most of the cases several monochlorobenzene pulses were degraded
before the process stopped, with oxygen concentration of 1,5–2 mg/l. When
monochlorobenzene concentration was increased to 40–45 mg/l degradation was
fast anyway: it took 20-30 days, if enough oxygen was present to support
biodegradation. (Figure 6.18). 1.3 mgoxygen/mgmonochlorobenzene were in average
necessary to oxidize monochlorobenzene.No remarkable differences were noticed
depending on the location (Artificial ditch, Leendertgracht and Vijver) or the type
of water used (groundwater or surface water).
monochlorobenzene concentration (µg/l)
50000
oxygen
addition
5000
40000
4000
oxygen
addition
oxygen
addition
30000
3000
20000
2000
10000
1000
0
100
0
0
20
40
60
80
100
120
140
160
180
160
180
200
oxygen concentration (mg/l)
5
4
3
2
1
0
0
20
40
60
80
100
120
140
Time (d)
artificial ditch - surface water
leendertgracht - surface water
vijver - surface water
artificial ditch - groundwater Q-32
leendertgracht - groundwater A-31/1
vijver - groundwater A-18/2
Figure 6.18. Monochlorobenzene and oxygen aqueous phase concentration in the batch tests
carried out with groundwater and surface water sampled in the Artificial Ditch,
Leendertgracht and Vijver on 20/04/2006. No degradation was observed in any of the dead
controls (data not shown).
180
200
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
A high monochlorobenzene degradation potential seems to be present in
both sediment and water: apparently groundwater and surface water and both
aquifer materials (top and bottom) contained a microbial community able to
degrade monochlorobenzene very quickly. The aquifer material sampling depth in
the interface did not influence the biodegradation of monochlorobenzene, and,
during the 200 incubation days, the only limiting factor appeared to be the oxygen
concentration.
6.3.4 Detection of catabolic genes at different depths of the interface
DNA extracts were used to investigate the spread of catabolic genes over
the interface in the three studied locations, at different times of the year. Among
all the studied catabolic genes (tmoA, xylE, todC1-like genes, cdo, tbuE, and todE)
only the tmoA gene was systematically detected. This gene encodes the initial
attack in the degradation of toluene and benzene, and has already been detected in
soils polluted with BTEX compounds (Hendrickx et al., 2005).
As an example Figure 6.19 shows the gel image of the PCR products
obtained from a sediment core sampled on 24/11/2004 in the Artificial ditch. The
fragment of the expected length (505 bp) is present with a bright signal down to
approximately 10 cm; from 12 to 16 cm the signal becomes weaker and
disappears completely at higher depths. The bright bands corresponding to the
positive controls (+) confirmed the success of the reaction; the absence of signal
corresponding to the negative controls shows that no interference had biased the
amplification process. The universal marker (100 bp ladder) is necessary to know
the exact size of the amplified amplicon.
181
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
M
41 38 35 32 30 28 26 24
+
-
M
22 20 18 16 14 12 10 9
+
-
+
-
M
8 7 6 5 4 3 2 1
Figure 6.19, Gel image of the PCR products obtained with TMOA primer from a sediment
core sampled in the artificial ditch on 24/11/2004. Numbers represent the depth of the
analysed slice in the interface (cm), M indicates the universal marker, + and – the positive
and negative controls.
Figures 6.20, 6.21, 6.22 shows the results of the PCR analyses carried out
on sediment cores extracted respectively from the Artificial ditch, Leendertgracht
and Vijver in the different seasons. In most cases, the tmoA gene is present at least
down to 15-20 cm in the interface and is not detected at greater depths. Hoewever
in some cases depths of even 35–40 cm were reached (in particular in Vijver in all
seasons, except Winter).
Apparently a relation also exists between the presence of the gene and the
season, since in Winter tmoA has been detected in the Artificial ditch only down
to the depth of 10 cm and is completely absent in any other location. The highest
depths are apparently reached in the samples collected in Summer. These results
are summarized in Table 6.7 PCR targeting the tmoA gene never gave positive
results for groundwater samples, whereas in surface water the gene was present
only in two cases (both sampled in Summer 2005).
Among the other catabolic genes, only tbmD, cdo and tbuE were
occasionally detected in a few samples (data not shown).
182
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
Autumn
2004
Winter
2005
Spring
2005
Summer
2005
Surface water
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
Groundwater
Surface water
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
Surface water
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
Groundwater
Surface water
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
Groundwater
Groundwater
Figure 6.20. PCR results of the detection of the tmoA gene in sediment cores extracted from
the Artificial ditch at different sampling times. Boxes represent sediment slices obtained
from the frozen sediment core, groundwater or surface water; White: not analysed samples,
blue: strong amplification, light blue: weaker amplification, shaded: no amplification.
183
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
Winter
2005
Spring
2005
Summer
2005
Spring
2006
Surface water
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
Groundwater
Surface water
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
Groundwater
Surface water
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
Surface water
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
Groundwater
Groundwater
Figure 6.21. PCR results of the detection of the tmoA gene in sediment cores extracted from
the Leendertgracht at different sampling times. Boxes represent sediment slices obtained
from the frozen sediment core, groundwater or surface water; White: not analysed samples,
blue: strong amplification, light blue: weaker amplification, shaded: no amplification.
184
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
Winter
2005
Spring
2005
Summer
2005
Spring
2006
Surface water
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
Groundwater
Surface water
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
Groundwater
Surface water
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
Groundwater
Surface water
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
Groundwater
Figure 6.22. PCR results of the detection of the tmoA gene in sediment cores extracted from
the Vijver at different sampling times. Boxes represent sediment slices obtained from the
frozen sediment core, groundwater or surface water; White: not analysed samples, blue:
strong amplification, light blue: weaker amplification, shaded: no amplification.
185
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
Table 6.7. Maximum depth of the tmoA gene in sediment cores from the three studied
locations at different sampling times; NA: not analysed, ND: not detected.
Autumn 2004
Winter 2005
Spring 2005
Summer 2005
Winter 2006
Artificial ditch
16 cm
10 cm
15 cm
29 cm
NA
Leendertgracht
NA
ND
10 cm
27 - 32 cm
14 cm
Vijver
NA
ND
35 – 45 cm
34 cm
29 cm
6.3.5 Study of the structure of the eubacterial community by 16S rRNA gene
PCR - DGGE analysis
Diversity of the eubacterial community at different depths in the interface:
Figure 6.23 shows the PCR–DGGE results of a sediment core sampled on
20/04/2006 from the Vijver as an example.
A complex DGGE pattern was obtained for slices from the different
positions in the interface, thus indicating that different types of Eubacteria were
present and a high diversity of species exists in these sediments; however the
degree of dominance of the bacteria seemed to be related with depth: a DGGE
pattern mostly observed in the first 10 – 15 cm slightly changed in function of
depth.
Some bands appeared starting from the depth of 3 cm, sometimes
becoming brighter as the depth increased (for example band “a”) down to 19 cm
and suddenly disappearing. None of the bands seemed to be present at every depth
of the interface. In addition, the dominance of certain bacterial species of the
microbial community diminished depending on depth and a DGGE pattern with a
smaller number of bright and so dominant bands occurred in the slices obtained
from a deeper position in the interface (from 19 cm to the bottom) and profiles
differ from each other. Surface water showed a less complex DGGE pattern
compared to that of sediment slices, characterized by the presence of 3 – 4 bright
bands. Some of the bands present in these fingerprints were also observed in the
underlying sediment slices. For instance band “b” was also present in slices 1 and
2, thus indicating that surface water infiltrates down to 2 – 3 cm. On the other
hand bands “c” and “d” seemed to be present only in surface water, thus
indicating the difference between the two microbial communities present in the
186
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
surface water and the interface. In groundwater only a few and faded bands were
observed, showing that this microbial community is much less diverse than those
of the sediment and surface water with a very limited presence of bacteria.
L
S
1
2
3
4
5
6
7
L
8
10
12
14
16
19
c
d
b
a
25
L
29
33
37
41
45
49
53
G
L
Figure 6.23. Eubacterial 16S rRNA gene PCR-DGGE profiles of the surface water (S),
groundwater (S) and slices of an undisturbed sediment core sampled in the Vijver on
20/04/2006. Numbers indicate the depth (cm) in the interface; L: DGGE marker.
Study of the microbial community in different locations.
Very low similarities were observed in the DGGE patterns obtained from
sediment samples taken at the same time from the three different sampling
locations Artificial ditch, Leendert gracht and Vijver. In Figure 6.24 the results
are shown from sediment cores sampled on 22/03/2005.
A high diversity was present in slices from the Artificial ditch (Figure
6.24a), we observed the appearance and disappearance of bands going from the
187
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
top to the bottom of the core. Band “a” was an example of a very bright band
present down to 5 cm and disappearing downwards; band “b” appeared at the
depth of 3 cm and was present down to 7 – 8 cm but its abundance increased
suddenly at 7 cm; band “c” was present at a depth from 3 to 10 cm, while a very
bright band (“d”) was observed only at 16 cm. This band could correspond to the
“d” band of the surface water. Apparently more dominant bands could be
observed in the first 10 cm of the core. From 20 cm to the bottom, a few dominant
bands were observed. Several faint bands have been observed over the total core
thus demonstrating considerable diversity.
In the core from Leendertgracht (Figure 6.24b), two bands (“e” and “f”)
were observed almost throughout the whole sample, while a third one (“g”)
disappeared at 7 cm and returned at 13 cm; the dominance of a few bacteria was
higher in the first layers where 8 – 9 dominant bands were present and decreased
starting from the depth of 5–6 cm; in the bottom layers only very thin bands
seemed to be present. Two bands (“h” and “i”) were unique at depths of 6 and 13
cm respectively.
The observations we did in the two previous locations were confirmed in
the third one (Vijver). The bacterial dominance in the DNA samples from Vijver
appeared very low in the first 10 cm (only very thin bands) and slightly increased
from 19 cm to the bottom (Figure 6.24c).
Results of PCR-DGGE analyses of undisturbed sediment cores
demonstrated the very low similarity observed on 16SrRNA level when the three
different sampling locations are considered. The highest microbial diversity was
observed in samples from the Artificial ditch, whereas a much less diverse
microbial community was present in the Leendertgracht, but almost present in the
whole core. On the other hand samples from the Vijver showed in general a quite
high diversity from a certain depth downwards.
188
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
(a) Microbial community of Artificial Ditch
1
1
2
a
2
L
3
4
5
6
7
10
13
16
L
(b) Microbial community of Leendertgracht
b
3
4
L
5
6
7
8
10
c
d
13
16
19
L
19
22
26
e
g
f
h
i
30
34
38
(c) Microbial community of Vijver
1
2
3
4
5
6
L
7
10
13
16
19
22
26
30
L
34
38
42
46
Figure 6.24. Eubacterial 16S rRNA gene PCR-DGGE profiles of undisturbed sediment cores
slices obtained from Artificial ditch (a), Leendertgracht (b) and Vijver (c) on 22/03/2005.
Numbers represent the depth in the interface (cm); L: DGGE marker.
189
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
The cluster analysis of the community fingerprints (Figure 6.25) confirms
that these communities are mostly different in the three places. Three main groups
can thus be observed. The greatest differences can be observed between Artificial
Ditch and Leendertgracht while a less marked difference can be observed between
bacterial communities of Leendertgracht and Vijver. Moreover, Figure 6.25 shows
that, for each location, the samples can be divided into two groups: the first one is
composed by the slices from the first 5 – 10 cm, the second one is made up of the
deeper slices, thus confirming that the structure of the community changes clearly
depending on depth.
190
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
Cosine coefficient (T ol 1.0%- 1.0%) ( H>0.0% S>0.0%) [0.0%-100.0%]
100
98
96
94
16 S
92
90
16 S
.
22/03/05
artific ial ditch - A34 (13 cm )
.
22/03/05
artific ial ditch - A37 (10 cm )
.
22/03/05
artific ial ditch - A28 (19 cm )
.
22/03/05
artific ial ditch - A31 (16 cm )
.
22/03/05
artific ial ditch - A40 (7 c m)
.
22/03/05
artific ial ditch - A41 (6 c m)
.
22/03/05
artific ial ditch - A45 (2 c m)
.
22/03/05
artific ial ditch - A46 (1 c m)
.
22/03/05
artific ial ditch - A42 (5 c m)
.
22/03/05
artific ial ditch - A44 (3 c m)
.
22/03/05
artific ial ditch - A43 (4 c m)
.
22/03/05
artific ial ditch - A21 (26 cm )
.
22/03/05
artific ial ditch - A17 (30 cm )
.
22/03/05
artific ial ditch - A13 (34 cm )
.
22/03/05
artific ial ditch - A05 (42 cm )
.
22/03/05
artific ial ditch - A01 (46 cm )
.
22/03/05
artific ial ditch - A09 (38 cm )
.
22/03/05
artific ial ditch - A25 (22 cm )
.
22/03/05
vijver - P05 (42 c m)
.
22/03/05
vijver - P01 (46 c m)
.
22/03/05
vijver - P13 (34 c m)
.
22/03/05
vijver - P17 (30 c m)
.
22/03/05
vijver - P21 (26 c m)
.
22/03/05
vijver - P28 (19 c m)
.
22/03/05
leendertgrac ht - L18 (2 cm)
.
22/03/05
vijver - P31 (16 c m)
.
22/03/05
leendertgrac ht - L19 (1 cm)
.
22/03/05
vijver - P25 (22 c m)
.
22/03/05
vijver - P40 (7 c m)
.
22/03/05
vijver - P41 (6 c m)
.
22/03/05
vijver - P37 (10 c m)
.
22/03/05
vijver - P34 (13 c m)
.
22/03/05
vijver - P09 (38 c m)
.
22/03/05
vijver - P42 (5 c m)
.
22/03/05
vijver - P45 (2 c m)
.
22/03/05
vijver - P44 (3 c m)
.
22/03/05
vijver - P43 (4 c m)
.
22/03/05
leendertgrac ht - L12 (8 cm)
.
22/03/05
leendertgrac ht - L13 (7 cm)
.
22/03/05
leendertgrac ht - L10 (10 cm)
.
22/03/05
leendertgrac ht - L04 (16 cm)
.
22/03/05
leendertgrac ht - L01 (19 cm)
.
22/03/05
leendertgrac ht - L17 (3 cm)
.
22/03/05
leendertgrac ht - L16 (4 cm)
.
22/03/05
leendertgrac ht - L14 (6 cm)
.
22/03/05
leendertgrac ht - L15 (5 cm)
.
22/03/05
leendertgrac ht - L07 (13 cm)
.
22/03/05
vijver - P46 (1 c m)
Figure 6.25. Eubacterial 16S rRNA gene; UPGMA clustering of the DGGE fingerprints of
undisturbed sediment core slices obtained from Artificial ditch (a), Leendertgracht (b) and
Vijver (c) on 22/03/2005. Numbers represent the depth in the interface (cm); L: DGGE
marker. Clustering was performed using the Cosine similarity coefficient (processed by
Bionumerics, Applied Maths, Belgium).
191
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
Study of the microbial community at different times.
The comparison between PCR-DGGE profiles of sediment cores taken
from the Artificial ditch throughout the year (Autumn, Winter, Spring, Summer)
offered the chance to follow the evolution of the microbial community structure
over time. For example, by comparing banding patterns referring to Autumn and
Winter (Figure 6.26 a and b) some similarities can be observed: in both situations
a dominant band is present from the surface to the depth of 5 cm; also profiles
comprised between 5 and 10 cm showed significant similarities: bands b and c
and region d appeared in both situations approximately at the same depth and a
less complex pattern occurred in the slices from the approximate depth of 15 cm
to 40 cm. In addition the same band “a” seemed to be present also in the profiles
from sediment cores sampled in Spring and Summer. On the other hand also in
Figure 6.26b (Winter) and c (Spring) similar elements are certainly present: the
fingerprints corresponding to the first 10 cm in Figure 6.26c (rectangle and region
d) showed a considerable similarity even with the samples referring to Autumn
and Winter, but the latter appeared brighter. In the deeper slices (starting from 15
cm) diversity was much higher and more bands absent in the two other seasons
clearly appeared. In Autumn, Winter and Spring the most important activity was
certainly at the top of the core. In Spring, for example, we observed two blocks of
bands: in DGGE fingerprints derived from the top of the core the high GC%
bands were especially missing (lower denaturation range of the gel), while they
were present in the cores from 15 cm to 39 cm ( approximatly 50% denaturation
range).
In sediment cores extracted in Summer (Figure 6.26d) we observed only a
few bright bands, but probably the diversity is so high that we only found thin
bands. The “d” group of the summer sample is unique, but when we compared the
gel with the others it was quite clear that the migration of this gel was different
and probably we lost that group in the other cases. Even at the depth of 60 cm
there was an important population present during Summer. Here the bands are
dominant but fewer.
192
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
In undisturbed sediment cores from the Leendertgracht interesting
analogies between the different seasons were also observed and at least 3-4
common bands (sometimes at similar depths) were present in cores extracted in
different periods. The dominance of a couple of bacteria in the microbial diversity
was in general lower than in the Artificial ditch. In particular in Spring samples
very few and thin bands were detected, whereas microbial community was very
diverse in Summer samples, mostly down to the depth of 10 cm.
In cores extracted in Spring 2005 in the Vijver a low dominance of certain
bacterial species (presented by very bright bands) was observed in the superficial
layers and it increased from 15 cm downwards, while in Spring 2006 microbial
dominance seemed lower in the first 3-4 cm than in the deeper layers and a
sudden change in the banding pattern was observed from 20 cm downwards.
Moreover a greater degree of microbial dominance was detected in Summer: a
very bright band was observed in the first slice (the same band, probably coming
from surface water, was also present in Spring 2005), and only slight changes
down to 5-6 cm. From this depth downwards the DGGE profile changed very
frequently (at least every 4 cm) and diversity remained very high down to 54 cm,
with an extremely high number of very thin bands.
193
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
(a)
(b)
L
1
2
3
4
5
6
7
8
L
9
10
14
16
18
20
22
24
L
26
28
30
32
35
38
41
a
b
1
d
a
2
L
3
4
5
c
b
d
c
6
7
10
13
16
L
19
22
26
30
34
38
L
(d)
(c)
1
L
2
3
4
5
6
7
8
10
15
20
25
30
35
39
L
a
d
1
2
3
4
5
L
6
9
14
19
24
29
34
39
44
49
54
58
L
a
Figure 6.26. Eubacterial 16S rRNA gene PCR-DGGE profiles of undisturbed sediment cores
slices obtained from Artificial ditch in Autumn 2004 (a), Winter 2005 (b), Spring 2005 (c)
and Summer 2005(d). Numbers represent the depth in the interface (cm); L: DGGE marker.
194
d
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
6.3.6 Study of the structure of the sulfate-reducing bacterial community by
PCR - DGGE analysis
Study of the microbial community in different locations.
In the first place PCR-DGGE analysis showed that sulfate-reducing
bacteria were present in all sediment cores extracted from the studied locations,
regardless of their depth in the interface or of the sampling time.
The comparison between the results obtained from the three sampling
locations on 22/03/05 shows that a very diverse community was present especially
in the Artificial Ditch (Figure 6.27a) and mostly down to 7-10 cm, whereas at
greater depths different fingerprints were observed and diversity significantly
decreased. Moreover the structures of the bacterial communities observed in the
three positions appeared markedly different from each other. In the sediment core
extracted from the Leendertgracht (Figure 6.27b) 3-4 dominant bands were
present throughout the core with only slight changes in the DGGE profile (for
example bands “a” and “b” disappeared from 6 cm downwards). Several very
faint bands were observed in DGGE patterns obtained from the core sampled in
the Vijver (Figure 6.27c); even if the profile did not change significantly
depending on depth some very dominant bands appeared starting from 16 cm
(groups “c” and “d”), while other bands became more important (for example
“e”). All these bands suddenly disappeared at the depth of 34 cm.
Similar results were obtained - from samples extracted in the other seasons
too: it was clear that sediments from the three studied locations harboured
different sulfate-reducing bacterial communities.
195
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
(a))
(b))
2
L
3
4
5
6
7
10
13
16
L
19
22
26
30
32
35
38
41
1
2
3
4
L
5
6
a
b
8
10
13
16
19
L
(c))
1
2
3
4
5
6
L
7
10
13
16
19
22
26
30
L
34
38
42
46
c
d
e
Figure 6.27. Sulfate reducing bacteria PCR-DGGE profiles of undisturbed sediment cores
slices obtained from Artificial ditch (a), Leendertgracht (b) and Vijver (c) on 22/03/2005.
Numbers represent the depth in the interface (cm); L: DGGE marker.
196
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
Study of the microbial community at different times:
Figure 6.28 shows the comparison between sulfate reducing bacteria
DGGE profiles obtained from sediment cores taken from the Artificial Ditch in
different seasons. Microbial diversity was very high in Autumn and Winter and
gradually decreased in Spring and Summer, when fewer and thinner bands were
observed. In all these cores the banding patterns suddenly changed (sometimes
thoroughly) from 10-15 cm downwards and the same happened in the cores
extracted in the other two locations. Two or three dominant bands were present in
the surface layers and they completely disappeared when the above cited depths
were reached. Several similar bands were present in the profiles from cores
extracted in different seasons, mainly concerning those extracted in Winter and
Autumn. For example band “a” was present in both Autumn and Winter, but also
in Spring, from 1 to 10 cm deep. Also band “b” could be observed in Autumn and
Winter. Band “c” appeared in Autumn, in Winter and in Spring, band “d” in
Winter, in Spring and in Summer, “e” was present in Spring and in Summer only
and “f” in Winter and in Autumn, while “g” appeared in Spring, in Summer ,in
Autumn and inWinter throughout the whole core. Data regarding the other two
locations were similar (data not shown).
The comparison between sulphate reducing bacteria PCR-DGGE profiles
of undisturbed sediment cores slices obtained from Artificial ditch over four
seasons indicated that a large diversity in bacteria exists, and several species were
present during the whole year.
The selected regions represented our findings. Band “c” for example was
present in nearly every slice, even in the surface water.
Sometimes their appearance was more visible in one season than in
another (bands “g“ and “d“). During Winter we observed the clearest diversity
with the presence og strong bands in the upper part of the core. In the rest of the
year fainter fingerprints were found in the upper part and some bright bands
appeared downwards.
197
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
Winter 2005
dsr
g
f
a
b
c
Autmn 2004
g
f
a
b
Spring 2005
d
e
a
c
g
Summer 2005
d
e
d
g
.
22/03/05
artificial ditch - A01 (46 cm)
.
.
22/03/05
artificial ditch - A05 (42 cm)
.
.
22/03/05
artificial ditch - A09 (38 cm)
.
.
22/03/05
artificial ditch - A13 (34 cm)
.
.
22/03/05
artificial ditch - A17 (30 cm)
.
.
22/03/05
artificial ditch - A21 (26 cm)
.
.
22/03/05
artificial ditch - A25 (22 cm)
.
.
22/03/05
artificial ditch - A28 (19 cm)
.
.
22/03/05
artificial ditch - A31 (16 cm)
.
.
22/03/05
artificial ditch - A34 (13 cm)
.
.
22/03/05
artificial ditch - A37 (10 cm)
.
.
22/03/05
artificial ditch - A40 (7 cm)
.
.
22/03/05
artificial ditch - A41 (6 cm)
.
.
22/03/05
artificial ditch - A42 (5 cm)
.
.
22/03/05
artificial ditch - A43 (4 cm)
.
.
22/03/05
artificial ditch - A44 (3 cm)
.
.
22/03/05
artificial ditch - A45 (2 cm)
.
.
24/11/04
artificial ditch - 01 (41 cm)
.
.
24/11/04
artificial ditch - 04 (38 cm)
.
.
24/11/04
artificial ditch - 07 (35 cm)
.
.
24/11/04
artificial ditch - 10 (32 cm)
.
.
24/11/04
artificial ditch - 12 (30 cm)
.
.
24/11/04
artificial ditch - 14 (28 cm)
.
.
24/11/04
artificial ditch - 16 (26 cm)
.
.
24/11/04
artificial ditch - 18 (24 cm)
.
.
24/11/04
artificial ditch - 20 (22 cm)
.
.
24/11/04
artificial ditch - 22 (20 cm)
.
.
24/11/04
artificial ditch - 24 (18 cm)
.
.
24/11/04
artificial ditch - 26 (16 cm)
.
.
24/11/04
artificial ditch - 28 (14 cm)
.
.
24/11/04
artificial ditch - 30 (12 cm)
.
.
24/11/04
artificial ditch - 32 (10 cm)
.
.
24/11/04
artificial ditch - 33 (9 cm)
.
.
24/11/04
artificial ditch - 34 (8 cm)
.
.
24/11/04
artificial ditch - 35 (7 cm)
.
.
24/11/04
artificial ditch - 36 (6 cm)
.
.
24/11/04
artificial ditch - 37 (5 cm)
.
.
24/11/04
artificial ditch - 38 (4 cm)
.
.
24/11/04
artificial ditch - 39 (3 cm)
.
.
24/11/04
artificial ditch - 40 (2 cm)
.
.
24/11/04
artificial ditch - 41 (1 cm)
.
.
24/11/04
artificial ditch - positive
.
.
31/05/05
artificial ditch 01 (cm 39)
.
.
31/05/05
artificial ditch 05 (cm 35)
.
.
31/05/05
artificial ditch 10 (cm 30)
.
.
31/05/05
artificial ditch 15 (cm 25)
.
.
31/05/05
artificial ditch 20 (cm 20)
.
.
31/05/05
artificial ditch 25 (cm 15)
.
.
31/05/05
artificial ditch 30 (cm 10)
.
.
31/05/05
artificial ditch 32 (cm 8)
.
.
31/05/05
artificial ditch 33 (cm 7)
.
.
31/05/05
artificial ditch 34 (cm 6)
.
.
31/05/05
artificial ditch 35 (cm 5)
.
.
31/05/05
artificial ditch 36 (cm 4)
.
.
31/05/05
artificial ditch 37 (cm 3)
.
.
31/05/05
artificial ditch 38 (cm 2)
.
.
31/05/05
artificial ditch 39 (cm 1)
.
.
31/08/05
artificial ditch 00-groundwater
.
.
31/08/05
artificial ditch 01 (cm 58)
.
.
31/08/05
artificial ditch 05 (cm 54)
.
.
31/08/05
artificial ditch 10 (cm 49)
.
.
31/08/05
artificial ditch 15 (cm 44)
.
.
31/08/05
artificial ditch 20 (cm 39)
.
.
31/08/05
artificial ditch 25 (cm 34)
.
.
31/08/05
artificial ditch 30 (cm 29)
.
.
31/08/05
artificial ditch 35 (cm 24)
.
.
31/08/05
artificial ditch 40 (cm 19)
.
.
31/08/05
artificial ditch 45 (cm 14)
.
.
31/08/05
artificial ditch 50 (cm 9)
.
.
31/08/05
artificial ditch 53 (cm 6)
.
.
31/08/05
artificial ditch 54 (cm 5)
.
.
31/08/05
artificial ditch 55 (cm 4)
.
.
31/08/05
artificial ditch 56 (cm 3)
.
.
31/08/05
artificial ditch 57 (cm 2)
.
.
31/08/05
artificial ditch 58 (cm 1)
.
.
31/08/05
artificial ditch 59-surface water
.
Figure 6.28. Comparison between sulfate reducing bacteria PCR-DGGE profiles of undisturbed sediment cores
slices obtained from Artificial ditch in Autumn 2004, Winter 2005, Spring 2005 and Summer 2005 (processed
by Bionumerics, Applied Maths, Belgium). Depths are indicated in brackets
.
198
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
6.3.7 PCR - DGGE analysis of tmoA-like genes
Study of the microbial community in different locations:
Figure 6.29 shows that in Summer 2005 the communities carrying tmoAlike genes were characterized by a rather low microbial diversity. In addition
significant differences were observed in the three sampling locations: in the
Artificial Ditch two dominant bands were present, starting from the top of the core
down to the depth of 24 and 59 cm (rectangle). These bands seemed to be present
also in the Leendertgracht but only at the depths of 5, 7 and 42 cm. It also
appeared at 14 cm in the Vijver. In the analysis of tmoA-like gene band (or group
of bands) “a” seemed very important and it was interestingly present in the Vijver
too and very feebly in the Artificial ditch, in all cases approximately down to 30
cm. In addition some further thin bands appeared in Leendertgracht, down to a
depth of 7-8 cm.
In the Vijver a series of very close bands can be noticed, while three
further isolated ones were observed at the depth of 14 cm.
Despite the similarities among the dominant bands of these communities
we observed important differences between their fingerprints.
In undisturbed sediment cores extracted in other seasons a very low
diversity of tmoA-like genes and only slight differences between the sampling
places were observed. In both Spring 2005 and Spring 2006 band “a” was the only
significant one (present in all the locations) while in Winter 2005 tmoA gene was
absent in the Leendertgracht and Vijver (data not shown).
199
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
(a)
1
2
4
5
6
L
9
14
19
24
29
34
39
44
L
49
54
59
(b)
1
2
3
4
a
L
5
6
7
8
12
17
22
27
L
32
37
42
(c)
1
L
a
2
3
4
5
6
9
14
19
24
29
L
34
39
44
49
Figure 6.29. TmoA gene PCR-DGGE profiles of undisturbed sediment cores slices obtained
from Artificial ditch (a), Leendertgracht (b) and Vijver (c) on 31/08/2005. Numbers
represent the depth in the interface (cm); L: DGGE marker.
200
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
Study of the microbial community at different seasons:
Comparing the tmoA results obtained from the Artificial Ditch in cores
extracted in different seasons we observed that two dominant bands which
appeared in Autumn (Figure 6.30a) were also present in Winter (Figure 6.30b)
and in Summer (Figure 6.30c), but not in Spring. In the Spring samples another
double band appeared and it was different from the one we observed in the other
seasons.
These observations, concerning the tmoA-like genes in function of the
seasons showed us a very poor bacterial activity with some single extra bands in
Summer and an extra double band in Autumn at the depths of 24 and 26 cm.
In the Leendertgracht and Vijver the diversity was higher in Summer 2005
than in Spring 2005 and in Spring 2006, when the only dominant band was “a”,
already shown in Figure 6.29 and only few and faint bands were observed (data
not shown).
201
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
(a)
1
L
2
3
4
5
6
7
8
L
9
10
12
14
16
18
20
22
L
24
26
28
30
32
35
38
41
L
(b)
1
2
L
3
4
5
6
7
10
13
16
19
L
22
26
30
34
38
1
2
L
1
2
3
4
5
6
4
5
6
L
9
14
19
24
29
34
39
44
L
49
54
59
8
10
15
L
20
25
30
35
40
(c)
(d)
Figure 6.30. TmoA gene PCR-DGGE profiles of undisturbed sediment cores slices obtained
from Artificial ditch in Autumn 2004 (a), Winter 2005 (b), Spring 2005 (c) and Summer
2005(d). Numbers represent the depth in the interface (cm); L: DGGE marker.
202
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
6.3.8 Batch degradation tests: study of the evolution of the microbial
community during the incubation period
Figure 6.31 shows 16S rRNA gene DGGE profiles obtained from sediment
and water samples, taken from the microcosms at the end of the incubation,
compared with samples used in the set up at time 0. It was clear that in the tests
where monochlorobenzene biodegradation occurred, new DGGE bands appeared.
Moreover the intensity of the band was often related to the mass of
pollutant consumed. For example in tests set up with surface water from the
Artificial Ditch (1A and 1B), four very strong bands appeared at the end of the
incubation, after 7 and 5 mg of monochlorobenzene were degraded respectively.
These bands were apparently absent in the time 0 water sample, thus
indicating that these bacteria probably grew on monochlorobenzene. The same
has been observed in groundwater samples (2A and 2B: 4.5 and 6.5 mg of
monochlorobenzene consumed) where only two very faint bands were detected in
the time 0 sample, while 7 – 8 bands appeared at the end. Also in surface water
and groundwater samples from the Leendertgracht (3A and 3B: 6.5 and 4.5 mg of
monochlorobenzene; 4A and 4B: 5.5 and 3.5 mg of monochlorobenzene) new
bands appeared. On the other hand in tests containing also sediment, some faint
bands appeared in microcosms where oxygen was added to stimulate
biodegradation (6A and 8A: 1.2 and 1.7 mg of monochlorobenzene), while no
bands or even weaker ones (compared to time 0 samples) appeared where
biodegradation was not stimulated (6B and 8B: 0.2 and 0.3 mg of
monochlorobenzene). Batch tests carried out with sediment and water sampled in
the Vijver gave DGGE patterns different from both the Artificial Ditch and the
Leendertgracht. New bands were detected in microcosms set up with surface
water (10A: 5 mg of monochlorobenzene) and groundwater (11A and 11B: 11 mg
and 8 mg) and one band appeared in the conditions “aquifer bottom+surface
water” and “aquifer bottom+groundwater” (13B and 15B: 6 mg of
monochlorobenzene), where degradation was stimulated with oxygen, while no
differences were observed with time 0 sediment samples, where degradation was
not stimulated (13A and 15A: 1.5 mg of monochlorobenzene). The new bands
203
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
observed in the samples analysed at the end of the incubation (especially where
considerable amounts of monochlorobenzene had been consumed) were mostly
different from the ones observed in time 0 samples, where, in some cases, the
signal was even absent.
These findings could indicate that species different from the ones
dominant in time 0 samples probably grew utilising monochlorobenzene as
growth substrate. No positive signals were obtained in the PCR performed with
DNA extracted from the a-biotic control, thus indicating the inhibition of the
L
Time 0 -surface water
10A
10B
Time 0 -groundwater
11A
11B
Time 0 -aquifer top
12A
12B
14A
14B
Time 0 -aquifer bottom
13A
L
13B
15A
15B
Time 0 -surface water
1A
1B
Time 0 -groundwater
2A
2B
Time 0 -surface water
L
3A
3B
Time 0 -groundwater
4A
4B
Time 0 -aquifer top
5A
L
5B
7A
7B
Time 0 -aquifer bottom
6A
6B
8A
8B
microbial activity by formaldehyde.
Figure 6.31. 16S rRNA gene PCR-DGGE profiles of sediment samples obtained from batch
degradation tests at the end of the incubation period. Numbers represent the labels of the
microcosms (Table 6.8); L: DGGE marker.
204
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
Label
1A
1B
2A
2B
3A
3B
4A
4B
5A
5B
6A
6B
7A
7B
8A
8B
10A
10B
11A
11B
12A
12B
13A
13B
14A
14B
15A
15B
Description
artificial ditch – surface water
artificial ditch – groundwater
leendertgracht – surface water
leendertgracht – groundwater
leendertgracht – aquifer top + surface water
leendertgracht – aquifer bottom + surface water
leendertgracht – aquifer top + groundwater
leendertgracht – aquifer bottom + groundwater
vijver – surface water
vijver – groundwater
vijver – aquifer top + surface water
vijver – aquifer bottom + surface water
vijver – aquifer top + groundwater
vijver – aquifer bottom + groundwater
Table 6.8. Labels used for batch degradation tests in Figures 6.31 and 6.32
Figure 6.32 shows the fingerprints of tmoA-like genes in the time 0
samples used to set up the microcosms (water and sediment) and in samples
collected from the flasks at the end of the incubation.
•
Artificial ditch: in surface water the gene was not present neither at the
beginning nor at the end of the incubation, while in groundwater a faint band
is present and became very thick at the end (band “a”). Besides two new
strong bands appeared after the 200 days incubation (samples 2A and 2B,
band “b”).
•
Leendertgracht: band “a” appeared in one duplicate in surface water (3A)
while band “b” appeared in the second (3B). Band “a” was also present in
groundwater at the end of the incubation. Band “b” was observed again in the
time 0 top sediment and apparently became less important at the end of the
205
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
experiment while it was almost absent in the bottom sediment (end of
incubation and time 0).
•
Vijver: band “b” appeared in surface water (10B) and groundwater (11A) at
the end of the experiment and two dominant bands were noticed in two cases
in microcosms set up with the top sediment (12A and 14B). Apparently the
diversity in the sediment from the Vijver was greater than the one of the
Leendertgracht.
These results showed that in most cases new and thick bands appeared at
the end of the incubation in the flasks where considerable amount of
monochlorobenzene had been consumed (2A: 4.5 mg, 2B and 3A: 6.5 mg, 4B: 3.5
mg, 12A: 11.5 mg, 12B: 6 mg), while very limited differences were observed
when comparing the time 0 samples with the ones where small amounts of
monochlorobenzene had been degraded (5B: 0.3mg; 6B, 8B, 14A: 0.2 mg; 15A:
b
Artificial ditch
Leendertgracht
Vijver
206
13B
15A
15B
Time 0-aquifer bottom
a
13A
L
12A
12B
14A
14B
Time 0-aquifer top
Time 0-groundwater
11A
11B
10A
10B
L
Time 0-surface water
Time 0-aquifer bottom
6A
6B
8A
8B
Time 0-aquifer top
5A
5B
7A
L
7B
4A
4B
Time 0-groundwater
Time 0-surface water
3A
L
3B
Time 0-groundwater
2A
2B
1A
1B
L
Time 0-surface water
0.1 mg).
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
Figure 6.32. TmoA gene PCR-DGGE profiles of sediment samples obtained from batch
degradation tests at the end of the incubation period. Numbers represent the labels of the
microcosms (Table 6.8); L: DGGE marker.
207
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
6.3.9 Cloning and sequencing of tmoA gene fragments
Among the obtained samples extracted on 31/05/2005 and 31/08/2005 we selected
the following ones for further research on cloning and sequencing of the tmoA
gene:
- 31/05/05 – Artificial Ditch – 6 cm deep;
- 31/05/05 – Leendertgracht – 7 cm deep;
- 31/08/05 – Leendertgracht – 17 cm deep;
- 31/08/05 – Vijver – 4 cm deep.
TmoA gene amplicons were then cloned to determine the gene sequences
corresponding to the DGGE bands and those clones were matched with the
corresponding DGGE profiles obtained from sediment slices. For all samples,
almost all dominant bands were recovered in the clone libraries.
Figure 6.33 shows DGGE profiles obtained with sample “31/08/05 Leendertgracht – 17 cm deep” (rectangles) and fingerprints resulting from the
cloning. Clones E3, E5, E11 and E12 were selected for sequencing and the
deduced nucleotides sequences were blasted against the NCBI bank.
Dechloromonas
aromatica RCB
Uncultured
bacterium clone
Dechloromon
as aromatica
RCB
Uncultured bacterium clone
A4Z/3 alpha subunit
monooxygenase protein gene
E3
E5
E12
E11
Figure 6.33. DGGE profiles obtained with sample “31/08/05 - Leendertgracht – 17 cm deep”
(rectangles) and fingerprints resulting from the cloning.(tmoA gene).
208
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
As an example Figure 6.34 shows the result of the blast analysis for clone
E5. Lines represent the sequences producing the best alignment.
Dechloromona
s
aromatica
Figure 6.34. Result of the blast analysis for clone E5
Table 6.9 summarizes the results of blast analysis for our sequences and
shows the nearest nucleotide matches based on blast analysis of the cloned
sequences.
In the Artificial Ditch samples sequenced clones were similar to Pseudomonas
Mendocina KR1 (a Pseudomonas strain able to metabolise toluene) while in the
samples from Leendertgracht tmoA sequences were similar to those of Ralstonia
Pickettii PKO1 (clone E3), also able to grow on BTEX. In several cases, clones
obtained from samples taken in the Leendertgracht and in the Vijver seemed
related to uncultured bacteria clones obtained from samples recovered from
209
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
BTEX contaminated sites (clones E3, E12, F5, F6, F8, F10). Clone E3 was also
related with Pseudomonas JS150, which is able to degrade a wide range of
substituted aromatic compounds, including chlorobenzene. TmoA sequences of
clones E5, E11 and F2 were related (90% of similarity) to the sequence of
Dechloromonas aromatica strain RCB.
Dechloromonas aromatica strain RCB is the only organism in pure culture
that can oxidize benzene in the absence of oxygen. It can also oxidize aromatics
such as toluene, benzoate, and chlorobenzoate. D. aromatica couples growth and
benzene oxidation to the reduction of either O2, or chlorate, or nitrate. These
results showed that clones obtained in the three locations were mostly different.
Moreover, also in the not contaminated location (Vijver) sequences related
with the metabolism of aromatics were found. Clones strongly similar to
Dechloromonas aromatica RCB were recovered from both the Leendertgracht and
the Vijver.
210
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
Origin
31/05/05 – Artificial Ditch – 6 cm
Clone
designation
11
12
31/05/05 – Leendertgracht – 7 cm
31/08/05 – Leendertgracht – 17 cm
16
E3
E5
E11
E12
31/08/05 – Vijver – 4 cm
F2
F4
F5
F6
F7
F8
F10
Nearest matches in BLAST analysis
Nucleotide
(Accession no.)
Toluene-4-monooxygenase gene
cluster (AY552601)
Monooxygenase alpha subunit
gene (AY504976)
Toluene-4-monooxygenase gene
cluster (AY552601)
Monooxygenase alpha subunit
gene (AY504976)
seq. too short
Alpha subunit monooxygenase
protein gene (AY450333)
Toluene-para-monooxygenase
gene cluster (AY541701)
Tbc2 gene cluster (AF282898)
CP000089
Gene for putative benzene
monooxygenase (AB274231)
CP000089
Gene for putative benzene
monooxygenase (AB274231)
Alpha subunit monooxygenase
protein gene (AY450315)
Alpha subunit monooxygenase
protein gene (AY450318)
CP000089
(CP00431) seq. too short (36 bp)
Alpha subunit monooxygenase
protein gene (AY450323)
Alpha subunit monooxygenase
protein gene (AY450322)
Toluene-4-monooxygenase gene
cluster (AY552601)
Alpha subunit monooxygenase
211
protein gene (AY450323)
Alpha subunit monooxygenase
protein gene (AY450323)
Host
Pseudomonas mendocina KR1
Nucleot.
identities
79%
Arthrobacter polychromogenes A169
Pseudomonas mendocina KR1
80%
Pseudomonas mendocina KR1
80%
Uncultured bacterium clone A4Z/3
99%
Ralstonia pickettii strain PKO1
87%
80%
Burkholderia cepacia JS150
Dechloromonas aromatica RCB
Uncultured bacterium
87%
90%
93%
Dechloromonas aromatica RCB
Uncultured bacterium
90%
92%
Uncultured bacterium clone A1Z/7
100%
Uncultured bacterium clone A1Z/10
99%
Dechloromonas aromatica RCB
Rhodococcus RHA1
Uncultured bacterium clone A3Z/3
91%
96%
97%
Uncultured bacterium clone A3Z/2
94%
Pseudomonas mendocina KR1
99%
Uncultured bacterium clone A3Z/3
98%
Uncultured bacterium clone A3Z/3
92%
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
Table 6.9. Sequences from clones isolated from sediment slices (tmoA gene).
212
6.3.10 Cloning and sequencing of dsrB and 16SrRNA gene fragments
DsrB and 16SrRNA gene amplicons obtained from sediment cores
extracted from the Leendertgracht on 31/08/2005 (7 cm and 32 cm) were cloned.
Concerning dsrB gene amplicons, at least 40 different clones were obtained.
Among these clones, the ones corresponding to the most dominant bands of the
original samples were sequenced. Therefore 11 samples were selected: 7 for
“31/08/05 – Leendertgracht – 32 cm deep” (clones C1, C5, C9, C10, C12, C19,
C20 – Figure 6.35) and 4 for “31/08/05 – Leendertgracht – 7 cm deep” (clones
D5, D7, D8 and D12). Rectangles indicate fingerprints of the original samples the
clones came from.
C1
C1
9
C5
C9
C1
0
C1
2
C2
0
Figure 6.35. DGGE profiles obtained with sample “31/08/05 - Leendertgracht – 32 cm deep”
(rectangles) and fingerprints resulting from the cloning.(dsrB gene).
The results of the sequencing are summarized in Table 6.10. All the
recovered clones were related to uncultured sulfate-reducing bacteria.
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
Due to the high diversity of bands present in the original DNA sample,
several different clones were obtained resulting in a great differentiation of
sequences (except clones C20 and C10).
Table 6.10. Sequences from clones isolated from sediment slices (dsrB gene).
Origin
Clone
Nearest match in BLAST analysis
designation
Nucleotide
31/08/05
Leendertgrachtde
pth: 32 cm
C1
C5
C9
Host
(Accession no.)
Dissimilatory sulfite reductase beta
subunit (dsrB) gene (EF064998)
Dissimilatory sulfite reductase beta
subunit (AY753141)
Dissimilatory sulfite reductase alpha
subunit (dsrA) and dissimilatory sulfite
Uncultured
sulfate-reducing
bacterium isolate DGGE gel
band 09
Uncultured bacterium clone
ng7d1139
Uncultured
sulfate-reducing
bacterium clone LGWG24
Nucleot.
identities
94%
83%
97%
reductase beta subunit (dsrB) genes
C10
C12
C19
C20
31/08/05
Leendertgracht
depth: 7 cm
D5
D7
D8
D12
(EF065029)
Dissimilatory sulfite reductase alpha
subunit (dsrA) and dissimilatory sulfite
reductase beta subunit (dsrB) genes
(EF065024)
Seq. too short for sequencing
Dissimilatory sulfite reductase alpha
subunit-like (dsrA) gene (DQ250756)
Sulfite reductase alpha subunit (dsrA)
and dissimilatory sulfite reductase beta
subunit (dsrB) genes (EF065024)
Dissimilatory sulfite reductase alpha
subunit (dsrA) and dissimilatory sulfite
reductase beta subunit (dsrB) genes
(EF065066)
Dissimilatory sulfite reductase subunit
B (dsrB) gene (AY015596)
Dissimilatory sulfite reductase alpha
subunit-like (dsrA) gene (DQ250781)
Seq. too short for sequencing
214
Uncultured
sulfate-reducing
bacterium clone LGWG08
94%
Uncultured
sulfate-reducing
bacterium clone G-77
92%
Uncultured
sulfate-reducing
bacterium clone LGWG08
93%
Uncultured sulfate-reducing
bacterium clone LGWK15
95%
Uncultured
sulfate-reducing
bacterium
clone
UMTRAdsr624-8
Uncultured
sulfate-reducing
bacterium clone I-70
88%
91%
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
With regard to 16SrRNA gene amplicons, 6 among the more than 40
obtained clones were sequenced (Table 6.11). Apparently Proteobacteria were the
main group of resident bacteria, with 96% similarity (clone B15) and 98% (clone
B19).
Table 6.11. Sequences from clones isolated from sediment slices (16S rRNA gene).
Origin
Clone
designation
Closest relative in blast analysis (Accession
no.) (Class)
Similarity
31/08/05
Leendertgracht
depth: 32 cm
A7
Uncultured Desulfosarcina
sp.
clone
CBII140 (DQ831553)(Proteobacteria)
90%
A16
A18
B15
uncultured bacterium (AY711541)
uncultured actinobacterium (AY307865)
Uncultured beta proteobacterium clone JG36GS-10 (AJ582037)
93%
95%
96%
B16
Uncultured delta proteobacterium
Hyd89-52 (sequence too short)
AY221613.1
31/08/05
Leendertgracht
depth: 7 cm
B19
clone
98%
6.4 DISCUSSION
This work deals with the study of monochlorobenzene degradation in the
interface between groundwater and surface water. The catabolic potential of the
microbial community present in the interface was studied in batch degradation
tests setup with sediments sampled from three different location operated at low
oxygen concentrations. The diversity of the microbial community was also
studied using molecular techniques (PCR/DGGE) and the presence of catabolic
genes was investigated.
Our results indicate that aquifer material (top and bottom) from the three
studied
locations
are
characterised
by
a
high
degradation
potential.
Monochlorobenzene biodegradation was only limited by a lack of oxygen: in the
batch tests the indigenous microorganisms were able to degrade up to 50 mg/l
monochlorobenzene when sufficient oxygen was available, with no need to add
215
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
nutrients or other amendments. Apparently, no degradation occurred below
oxygen concentrations of 1.5-2 mg/l. Moreover, the catabolic potential seemed
not to be related with depth: degradation took place indifferently in the
microcosms containing both top and bottom aquifer material. The fact that
significantly different amounts of oxygen were required to degrade approximatly
the same amounts of monochlorobenzene in tests setup with different aquifer
materials (from the Leendert gracht and the Vijver) could be explained
considering that the total organic matter content was rather different in the two
situations. Sediment material used to setup microcosms contained on average
2.45% organic matter in the Leendertgracht while only 0.95% in the Vijver. The
probable presence of other potential carbon sources and their preferential use by
microorganisms could have resulted in a fast oxygen consumption preceding
monochlorobenzene degradation. For the same reason oxygen concentrations in
microcosms containing only water remained higher and monochlorobenzene
degradation went on quickly, requiring only a few oxygen spikings. The specific
amount of oxygen (mgoxygen/mgmonochlorobenzene) needed was thus even lower
compared to the sediment tests.
Surprisingly, a high degradation potential was present in sediment material
from the Vijver too. This is in contrast with other studies (Van der Meer et al.,
1998; Dermietzel and Vieth, 2001), isolating CB-degrading bacteria only within
the contaminated zone, while degradation potential was very low or even absent
in uncontaminated subsurface material. Possibly, CB-degrading bacteria arose
elsewhere and was transported to the Vijver zone because of the closeness of the
two locations.
Unfortunately, the chloride concentration of groundwater and surface
water samples was very high (up to 3000 mg/l) making it impossible to use
chloride release analysis to determine whether complete monochlorobenzene
mineralization had occurred. Furthermore, a study (Vogt et al., 2003) reporting
chlorobenzene degradation in microaerophilic conditions by five strains isolated
on chlorobenzene as sole carbon source, showed the accumulation of the toxic
intermediate 3-chlorocatechol. In one of these strains a clear relationship was
demonstrated between the presence of 3-chlorocatechol in the medium and low
216
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
oxygen concentration. However, no 3-chlorocatechol was detected under oxygenlimited reactor operation in liquid samples of the Bitterfeld pilot plant in situ
reactor (Vogt et al., 2004).
TmoA gene-PCR-DGGE analysis carried out at the end of the microcosms
incubation on sediment contained in the flasks showed a different DGGE pattern
between the aquifer materials from the two locations (Leendertgracht and Vijver),
but also the presence of some common bands (band”b”-Figure 6.32). New bands
appeared at the end of the incubation in some microcosms probably as a result of
monochlorobenzene degradation by these microorganisms. Some of these bands
are absent or not dominant in DNA extracted from the analyzed cores from the
Leendertgracht and the Vijver. This could indicate that microorganisms present in
situ are not active in monochlorobenzene degradation. Nevertheless this is not
surprising, because no chlorobenzene was detected in the Vijver, while its
concentration in the Leendertgracht was maybe too low to stimulate and sustain
the growth of microorganisms. These findings seem confirmed by the observation
that also in 16S rRNA DGGE pattern new bands (absent or very faint at Time 0)
appeared after the incubation period were, thus probably indicating that these
bacteria were present at concentrations below the detection limit and significantly
grew using chlorobenzene as energy source.
Besides, the low oxygen concentration measured in groundwater and
surface water seems to suggest that chlorobenzene degradation is scarcely
probable in the sediment: results from batch degradation tests indicated that at
least 1.5-2 mg/L of oxygen are required to sustain the degradation process but the
contribution given by surface water is probably unsufficient due to its low oxygen
concentration (often below 4 mg/l). Furthermore oxygen consumption by
sediments has to be taken into account. This consumption, also depending on the
organic matter concentration, can reach high levels and significantly reduce the
oxygen amount available for chlorobenzene degradation. The prevailing redox
conditions at different depths in the interface need investigation through further
research. The very low chlorobenzene concentrations detected in surface water
compared to groundwater are probably a result of dilution and sorption in the
organic matter of the sediment (up to 15 mg/kg of MCB; Figures 6.13 and 6.14).
217
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
Cloning and sequencing of tmoA sequences showed that cloned sequences
were mostly related with microorganisms involved in the metabolism of BTEX.
One sequence was related in third similarity with Pseudomonas JS150, which can
grow on a wide variety of aromatic compounds, including monochlorobenzene.
Possibly, these microorganisms have monoxygenases catilizing the first step of
the reaction, but unable to completely mineralize chlorobenzene. Initial steps of
chlorobenzene degradation give rise to 3-chlorocatechol which is usually
degraded via the ortho pathway described by Reineke and Knackmuss (1984).
The initial attack is by a dioxigenase acting like toluene dioxygenase and benzene
dioxygenase. The initial oxidation results in the formation of a cis-dihydrodiol.
Subsequent ring fission and elimination of chloride leads to the mineralization of
these compounds. The key enzyme is the pyrocatechase II (Dorn et al., 1978;
Reineke et al., 1984) that converts chlorocatechols into chloro-cis,cis-muconic
acids. The absence of this enzyme in organisms with initial oxygenases with
broad substrate specificities may lead to the accumulation of chlorocatechols or to
the misrouting of chlorocatechol down the meta cleavage pathway, ultimately
resulting in cell death. Further research should focus on isolating and
characterizing chlorobenzene-degraders present in the interface.
218
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
PCR-DGGE analysis of the eubacterial 16S rRNA gene showed, in the
first place, that in general the structure of the microbial community is very diverse
and changes depending on depth., significant changes were frequently observed
from 5-10 cm downwards. Koizumi et al. (2003) also observed vertical changes of
a bacterial community structure in a mesophilic lake sediment by DGGE analysis
of amplified 16S rDNA and reversely transcribed 16S rRNA fragments. They
noticed that the diversity indices obtained from the 16S rDNA-based DGGEprofiles were greater than those obtained from the 16S rRNA-based DGGE
profiles. The diversity of inactive bacteria (DNA level) did not change drastically
in function of depth since they were only influenced by bacteria that accumulated
in association with sedimentation. In contrast, the diversity of active bacteria
(RNA level) decreases with sediment depth. More specifically, the rRNA-based
dendrogram showed a significant difference between the upper layers (0-2, 2-5,
and 5-8 cm) and the lower ones (8-11, 11-14, 14-17, and 17-20 cm).
In most cases a huge number of very faint bands was observed together
with some dominant thick bands, thus confirming that a very diverse bacterial
community, dominated by a few species, is harboured in the interface. The
depending-on-depth relevant changes in the community’s structure are probably
related with the gradient in the redox conditions. The microbial communities from
the three sampling locations appeared considerably different although some
similar bands were observed. In the Artificial ditch diversity seemed higher and
more dominant bands were present; this could be caused by the high concentration
of chlorobenzene. The presence of high concentration of the contaminant could
have stimulated the development and growth of microorganisms able to use it as
carbon and energy source (evidenced by the presence of the dominant bands).
This is in contrast with another study carried out in a BTEX-contaminated site
(Hendricks et al., 2005), demonstrating that the uncontaminated area was
characterized by a much more diverse bacterial community than the contaminated
one. Alfreider et al., (2002) studied the microbial diversity in an in situ reactor
system treating MCB-contaminated groundwater. They observed that the
significance of specific pollutants for the structure within the bacterial
assemblages in contaminated groundwater ecosystems is hard to assess, because
219
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
various physical, chemical and biological factors may often mask anthropogenic
effects. They also observed differences in the microbial community structure
between sediments and groundwater samples (as we did) and they explained it
referring to the differences between these two habitats, attached and free-living
bacteria. Very little is known about the differences in the microbial community
structure between original sediment and groundwater. Roling et al., (2001) found
that pollution in a landfill leachate–contaminated aquifer did not affect the
particle-bound microorganisms, but groundwater community structure was clearly
affected by pollution and redox processes, thus supporting the hypothesis that
bacteria attached to sediment particles and forming biofilms usually consists of
stable communities which are less influenced by changing environmental factors.
Conversely the Leendertgracht and the Vijver were characterized by a
great diversity and a more uniform distribution of vanishing bands.
The characterization of the bacterial community revealed the presence of
sequences related to Proteobacteria in agreement with Alfreider et al., (2002)
who found this bacteria in chlorobenzene-contaminated groundwater and
sediment samples from their in situ reactor.
PCR-DGGE analyses of 16S rRNA gene and tmoA gene showed a greater
diversity in cores extracted in Summer. This is probably due to the more intense
microbial activity depending on higher temperature (Figures 6.12, 6.26 and 6.30).
DsrB gene fragments cloning and sequencing confirmed that a very high
diversity of sulfate-reducing bacteria is present in each of the three locations, in
every season, regardless of depth. This seems in contrast with the presence of
bacteria carrying tmoA like genes in the first 10-20 cm sediment layers. The
presence of anaerobic obligate sulfate reeducing bacteria can be explained
assuming that the sediment might contain both aerobic and anaerobic microniches. The co-occurrence of sulfate-reducing bacteria and aerobic organisms has
been shown before in aerobic wastewater biofilms (Ito et al., 2002b; Ito et al.,
2002a; Kühl and Jorgensen, 1992; Okabe et al., 1999). Recently, Shi et al. (1999)
showed the co-occurrence of these organisms in a fuel contaminated aquifer,
where the conditions were micro-aerophilic to anaerobic.
220
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
Anaerobic bacteria can be responsible of the reduction of highly
chlorinated
benzenes
to
chlorobenzene
and
of
the
accumulation
of
monochlorobenzene in the aquifer. This could also provide an explanation for the
detection of di- and especially tri-chlorobenzenes concentrations which were
significantly lower if compared to monochlorobenzene.
Studies on the effect of the interface between groundwater and surface
water on the degradation of pollutants is limited. Lendvay et al. (1998, 1999)
studied the biogeochemical effects of a large surface water on a chloroethene
contaminated anaerobic groundwater at the groundwater/surface water interface
(GSI) using spatially discretized multilevel arrays. Concentrations of methane and
chloroethene decreased as the groundwater became increasingly oxidized along
the GSI in shallow sample points impacted by infiltration of oxygenated lake
water. Cis-1,2-dichloroethene remained unchanged or increased at the same
locations indicating that the decrease in methane and chloroethene was not due to
dilution effects from lake water infiltration. Schwarzenbach et al. (1983)
investigated, by the installation of a network of observation wells, the transport
and fate of chlorinated hydrocarbons, alkylated benzenes, and chlorinated phenols
during natural infiltration of river water to groundwater. Biotransformation was
observed in the interface for all alkylated C1-C4-benzenes, naphthalene, the
methylnaphthalenes, and 1,4-dichlorobenzene. Alkylated benzenes were always
eliminated within the first few meters of infiltration, even at temperatures below
5°C. The biotransformation of 1,4-dichlorobenzene occurred at a lower rate while
chloroform, 1,1,1-trichloroethane, trichloroethylene, and tetrachloroethylene were
not degraded in the interface. With respect to these last compounds, bank
infiltration is thus ineffective as a first step in the treatment of river water for
water supplies. Feris et al. (2003) investigated through the use of microbial
techniques (DGGE, 16S rRNA phylogeny, phospholipid fatty acid analysis, direct
microscopic enumeration, and quantitative PCR) the effect of a range of sediment
metal loads on the microbial community inhabiting the hyporheic zone of six
different rivers. They found that metal stress in fluvial environments does not
reduce biomass, diversity, or productivity rather the structure of microbial
communities changes. It appeared that the hyphoreic-zone communities exhibited
221
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
a decrease in -proteobacteria but no significant change in -proteobacteria and
an increase in -proteobacteria with increasing metal contamination. They
concluded from their study that more studies are needed to define or describe the
taxa that comprise the communities in the interface that are affected by pollutants
and that this could be mainly achieved by the use of molecular techniques.
6.5 CONCLUSIONS
In this study we investigated the biodegradation of chlorobenzene in the
interface between groundwater and surface water. Main goal was to understand if
the microbial community present in the interface was involved in the degradation
of pollutants passing through the sediment layer and if the interface can therefore
have an active role in the breakdown of pollutant in groundwaters reaching
surface waters.
Therefore we carried out batch degradation tests and we applied molecular
techniques (PCR-DGGE), in order to study the catabolic potential of the microbial
community present in the interface, using sediment material extracted from the
interface in three different locations and in different seasons.
Results of batch tests carried out at low oxygen concentrations (such as the
presumed in situ conditions) showed that a high chlorobenzene degradation
potential is present, regardless of depth in the interface. Biodegradation of
chlorobenzene was only limited by a lack of oxygen. Furthermore we found the
catabolic gene tmoA (involved in the initial step of the degradation of BTEX
compounds, similar to monochlorobenzene) everywhere. This gene was only
present down to a depth of 10-20 cm, indicating that a shortage of oxygen
possibly prevents its presence in the microbial population of the deeper layers.
Further research will have to investigate whether mineralization of chlorobenzene
is complete or if it is a partial transformation, possibly resulting in the
accumulation of the toxic metabolite chlorocatechol. Further research will also
have to investigate the real in-situ situation in order to assess the presence of the
right conditions for biodegradation.
222
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
Molecular tests also indicated that the structure of the microbial
community significantly changed depending on depth. Besides, communities from
the three studied locations were mostly different, while similiraties were observed
in each location throughout the whole year.
223
CONCLUSIONS
______________________________________________________________________________________________________________________________________________________________
Especially in the last 50 years of industrial development, the amount and
variety of hazardous substances has drastically increased. Among them,
halogenated compounds, are widespread air, water, soil, and sediment pollutants;
they are recalcitrant molecules resistant to mineralization due to the stability of
their carbon-halogen bond. Since these compounds have existed for millions of
years, there are naturally occurring strains of bacteria which have evolved to
break down halogenated compounds, thus opening up the possibility for
bioremediation treatment of contaminated sites.
Chlorinated aliphatic and aromatic hydrocarbons are among the most
common contaminants of soils, groundwaters and sediments. Several studies
showed that most of these pollutant can be biodegraded by single bacterial strains
or mixed microbial populations via aerobic direct metabolism or cometabolism.
In this thesis, two studies have been carried out concerning different situations
where bioremediation processes of chlorinated hydrocarbons were involved.
The first experimental work consisted in the study of microbial consortia
able to degrade a mixture of 6 CAHs (chlorinated aliphatic hydrocarbons) via
aerobic cometabolism. We studied the long-term growth process of two microbial
consortia using different primary substrates (methane and propane) and effective
in the aerobic cometabolic biodegradation of a mixture of 6 chlorinated aliphatic
hydrocarbons (CAHs), and the effectiveness of these consortia as inocula for the
bioaugmentation of different types of aquifer materials. The main goals of the
study included:
•
to verify the maintenance of the consortia’s capacity to degrade the 6CAH mixture during a prolonged process of microbial growth in the
presence as well as in the absence of the 6-CAH mixture;
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
•
to verify the consortia’s ability – after a prolonged growth process - to lead
to the rapid onset of biodegradation of the CAH mixture upon inoculation
in slurry microcosms set up with aquifer materials taken from sites with
different physical-chemical characteristics;
•
to develop a third consortium able to combine the best characteristics of
the methane-utilizing and of the propane-utilizing consortia object of the
study: in fact, a previous study had shown that, while both consortia were
effective in the aerobic cometabolic biodegradation of VC and cis-DCE,
the methane-utilizing biomass had a higher capacity to transform transDCE, whereas the propane-utilizing one was more effective towards 1,1,2TCA, 1,1,2,2-TeCA and, secondarily, TCE;
•
to characterize in terms of specific CAH depletion rates and degree of
mineralization of the organic Cl the best methane-utilizing and the best
propane-utilizing consortium obtained as a result of the inoculation in the
microcosms set up with different aquifer materials.
The propane-utilizing consortium generally proved the most effective one,
being able to biodegrade vinyl chloride, cis- and trans-1,2-dichloroethylene,
trichloroethylene, 1,1,2-trichloroethane and 1,1,2,2-tetrachloroethane at all the
CAH concentrations tested.
Both consortia maintained unaltered CAH degradation capacities during a 300day growth period in the absence of the CAHs and were effective in inducing the
rapid onset of CAH depletion upon inoculation in slurry microcosms set up with 5
types of aquifer materials.
A consortium developed in microcosms supplied with both methane and
propane combined the best degradation capacities of the two single-substrate
consortia.
The degree of conversion of the organic Cl to chloride ion was equal as an
average to 90%.
These results indicated that a large amount of inoculum potentially useful in
bioaugmentation tratments of CAH-contaminated sites could be grown in liquid-
225
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
gas bioreactors in the absence of CAHs and soil, starting from small amounts of
biomass suspension. The use of a methane-propane-utilizing consortium would
result in the best degradative performances.
The second study dealt with monochlorobenzene biodegradation in the
interface between groundwater and surface water. Soil remediation in practice
often consists of the application of extensive techniques for the active removal of
the contamination source and remediation of the plume. Objects of risk are often
surface water systems. There are strong indications that the interface between
groundwater and surface water plays an important role in the natural degradation
of organic contaminants. This is especially the case for mobile contaminants that
are persistent in anaerobic subsurface environment, but mineralized relatively
easy under more oxidized environmental conditions (e.g. chlorobenzene or vynil
chloride).
Main goal was investigating the natural pollutant degradation capacity of the
aquifer zone representing this interface. The interface can be considered a zone
with changing redox conditions characterized by specific degradation potential for
pollutants passing through as a result of steep physico-chemical gradients. Thus
the catabolic potential and the structure of the microbial community present in the
interface between groundwater and surface water were studied to find out whether
bacteria present in the interface are involved in pollutants degradation. Therefore
batch degradation tests and molecular analyses (PCR-DGGE) were carried out on
aquifer material extracted at different depths in the interface in three locations
characterized
by
different
monochlorobenzene
contamination
levels.
Chlorobenzene degradation was studied in batch tests under oxygen-limited
conditions in order to simulate the in situ conditions. 16S rRNA PCR-DGGE
analysis were carried out on undisturbed sediment cores extracted from the three
studied locations in different seasons to detect the presence of catabolic genes at
different depth in the interface and to study the structure of the microbial
community.
226
Biodegradation of Monochlorobenzene in The Interface Between Groundwater and Surface Water
Results from batch degradation tests indicated that the position in the
interface did not have any effect on the chlorobenzene degradation and the
process was exclusively limited by a lack of oxygen. Up to 50 mg/l of
monochlorobenzene were consumed in 20 days in both aquifers, and also in
groundwater and surface water, when sufficient oxygen was available (1.5–2
mg/l)
The structure of the microbial community changed in function of depth.
Moreover the structure of the community appeared different in the three locations
while significant similarities were observed in samples extracted in each location
in different seasons. Cloning and sequencing allowed to identify the dominant
bands in the DGGE pattern as belonging to the group of Proteobacteria. Bacteria
carrying tmoA-like genes were mostly related to BTEX degraders: Pseudomonas
Mendocina KR1, Ralstonia Pickettii PKO1, Dechloromonas aromatica strain
RCB.
It is still unclear if bacteria corresponding to these DGGE bands play a role in
chlorobenzene degradation. The only degradative gene detected until now is the
mono-oxygenase tmoA (involved in the degradation of BTEX, structurally similar
to monochlorobenzene) thus being probably involved in its degradation.
227
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