Heterogeneous aerobic benzene-degrading communities in
oxygen-depleted groundwaters
Anne Fahy, Terry J. McGenity, Kenneth N. Timmis & Andrew S. Ball
Department of Biological Sciences, University of Essex, UK
Correspondence: Anne Fahy, Department of
Biological Sciences, University of Essex,
Wivenhoe Park, Colchester CO4 3SQ, UK.
Tel.: 144 01206 873370; fax: 144 01026
872592; e-mail: [email protected]
Present address: Andrew S. Ball, School of
Biological Sciences, Flinders University of
South Australia, GPO Box 2100, Adelaide,
South Australia 5001, Australia.
Received 17 January 2006; revised 12 April
2006; accepted 13 April 2006.
First published online 23 June 2006.
DOI:10.1111/j.1574-6941.2006.00162.x
Editor: Max Häggblom
Keywords
bacterial community composition; benzene
contamination; alkaline groundwater; DNA
fingerprinting; terminal restriction fragment
length polymorphism (T-RFLP).
Abstract
A sandstone aquifer beneath a petrochemicals plant (SIReN site, UK) is heterogeneously contaminated with benzene and oxygen-depleted. Despite low redox
potentials in three of the most contaminated groundwaters (benzene concentrations from 17.8 to 294 mg L1), we observed aerobic benzene degradation in
microcosms, indicating the presence in situ of a latent community of obligate
aerobic microorganisms or an active community of facultative aerobes responding
rapidly to oxygen ingress. Moreover, benzene degradation occurred at the ambient
pH of 8.9 and 9.4, considerably more alkaline conditions than previously reported.
16S rRNA analyses showed that the groundwater microcosm communities were
distinct from each other, despite sharing the function of aerobic benzene
degradation. From DNA fingerprinting, one consortium was dominated by
Acidovorax spp., another by Pseudomonas spp.; these benzene-degrading consortia
were similar to the in situ communities, perhaps indicating that these organisms
are active in situ and degrading benzene microaerophilically or by denitrification.
Conversely, in the third sample, benzene degradation occurred only after the
community changed from a Rhodoferax-dominated community to a mix of
Rhodococcus and Hydrogenophaga spp. Four of the main benzene-degrading strains
were brought into culture: Hydrogenophaga and Pseudomonas spp., and two strains
of Rhodococcus erythropolis, a ubiquitous and metabolically versatile organism.
Introduction
Benzene is a widely used chemical. In 2001, 110 tonnes were
estimated to be released into controlled waters and sewers in
the UK, mainly through vehicle emissions and the chemical
industry (Environment Agency, 2004). It is toxic and carcinogenic, and because of its relatively high solubility compared to other hydrocarbons, it is a mobile component in
water systems. Benzene is therefore a groundwater pollutant
of environmental and health concern.
Benzene is known to be metabolized aerobically by a
number of microorganisms, and this process can quickly
generate anaerobic conditions in aquifers. We previously
showed that in the SIReN aquifer (site for innovative research in natural attenuation) in north-west England, anoxia was the main environmental factor influencing bacterial
community structures in the groundwater rather than the
toxicity of benzene, the main contaminant (Fahy et al.,
2005). Anaerobic oxidation of benzene has been reported
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(Kazumi et al., 1997; Coates et al., 2002), but it can be a slow
process, particularly when other contaminants are present.
In the presence of oxygen, benzene degradation occurs more
readily, thus aerobic degradation can play an important role
in natural attenuation if oxygen is present or readily replenished: at high flow rates, at low benzene concentration, at
the early stages of a contamination incident or at the edges
of benzene plumes. Furthermore, the presence of a proficient aerobic benzene-degrading microbial consortium
offers options for engineered remediation with the addition
of oxygen and nutrients.
The upper shallow aquifer of the SIReN site has been subjected to long-term pollution, and the geochemical characteristics of groundwaters from 76 monitoring wells show
large variations, including in pH and in the concentrations
of contaminants (Jones et al., 2001). The aims of this study
were to investigate the influence of high benzene concentration and suboxic conditions on aerobic benzene-degrading
microbial communities, and to assess whether benzene
FEMS Microbiol Ecol 58 (2006) 260–270
261
Benzene-degrading communities in groundwaters
degradation could occur at the ambient pH of 8.9 and 9.4 of
two of these groundwaters, at which benzene degradation
has not previously been reported.
medium sensitivity. Measurements were calibrated against
control sterile microcosms at each GC session.
Isolation of benzene-degrading organisms
Materials and methods
Site and sample collection
The SIReN aquifer is situated below a chemical plant that
has been operational for 60 years; the main groundwater
contaminants are BTEX (benzene, toluene, ethylbenzene
and xylenes), of which benzene is the principal component
(Jones et al., 2001). Three monitoring wells from the most
impacted area of the site were chosen for this study, and
groundwater was collected on 12 February 2001 as previously described (Fahy et al., 2005). Oxygen ingress was minimized during the collection and transport of the
groundwater samples, and DNA was extracted within 36 h.
Primary groundwater characteristics are indicated in Table
1; further details of the site have been previously published
(Fahy et al., 2005), and comprehensive data are presented in
technical reports of the Environment Agency UK (Jones
et al., 2001; Lethbridge et al., 2003).
Microcosms and benzene measurements
Triplicate microcosms were prepared with the samples after
9 days of storage at 4 1C, by placing 18 mL of groundwater in
100 mL crimp top serum bottles, and adding 1 mL of ammonium nitrate or/and phosphate solution (1 g L1
NH4NO3; 1.11 g L1 Na2HPO4 and 0.25 g L1 KH2PO4 final
concentration). The bottles were spiked with benzene to
80 mg L1 above background concentration, sealed with
PTFE-lined butyl septa (Agilent) and incubated in the dark
at 12 1C. Benzene concentration was quantified by gas chromatography (GC) of the head space, using a Unicam 610
Series GC fitted with a 4 mm internal diameter, 17 cm outer
diameter glass packed column (10% apiezon on chromosorb W) and a flame ionization detector, at an injector temperature of 250 1C, column 155 1C and detector 250 1C,
Extinction dilutions were prepared in 20 mL serum bottles
with 80 mg L1 of benzene in 10 mL of minimal medium
(pH 7.2), and incubated at 12 1C. The bottles with the highest dilution factor at which growth occurred were then used
to isolate organisms on minimal medium/washed agar
plates with benzene supplied as vapour at 12 1C. The benzene-degrading property of each isolate was then confirmed
in liquid mineral medium with benzene as the sole carbon
source. The medium was prepared with 100 mL each of solutions A and B, 1 mL of solution C, and 800 mL of water.
Solution A: 5 g MgSO4 7H2O and 1 g CaCl2 H2O L1 (filtered). Solution B: 11.1 g Na2HPO4, 2.5 g KH2PO4 and 10 g
NH4NO3 L1 (autoclaved). Solution C: 10 g FeSO4 7H2O,
0.64 g Na2EDTA 3H2O, 0.1 g ZnCl2, 0.015 g H3BO3, 0.175 g
CoCl2 6H2O, 0.15 g Na2MoO4 2H2O, 0.02 g MnCl2 4
H2O and 0.01 g NiCl2 6 H2O L1 (filtered).
DNA extraction and PCR amplification
Community DNA was extracted from 40 mL aliquots of
groundwater or enrichment as previously described (Fahy
et al., 2005) using a guanidinium thiocyanate lysis buffer
(Boom et al., 1990) and recovering the DNA with Glassmilk
(Bio 101, Geneclean Spin Glassmilk). DNA from isolates was
similarly extracted from 20 mL of culture.
PCRs targeting the 16S rRNA gene were performed with a
Perkin Elmer Gene Amp PCR System 9700. Standard cycling
conditions were: 94 1C, 2 min, (94 1C, 1 min; 55 1C, 1 min;
72 1C, 2 min) 30, 72 1C, 10 min. Each 50 mL reaction contained: 1 mL DNA template (c. 0.5–5 ng), 2.5 U of Taq DNA
polymerase (Qiagen), 10 buffer (Qiagen), 5 Q solution
(Qiagen), 5 mmol dNTPs (Invitrogen), 20 pmol each of forward and reverse primer.
Fluorescently labelled primers for terminal restriction
fragment length polymorphism (T-RFLP) were obtained
from Applied Biosystems: FAM63F (5 0 -CAG GCC TAA
Table 1. Primary groundwater measurements
Well
Pi
Rs
Ri
Well codes
W18i
308s
308i
Depthw (m)
5.5
5.3
19.6
pHz
7.6
8.9
9.4
Benzenez (mg L1)
Redox potential‰ (mV)
5
188
128
74
2.94 10
1.78 104
2.08 104
BTEX trendk
Declining
Declining
Increasing
These are the codes used in the EA reports; for clarity, in this study both locations are designated by a single letter. The suffix indicates the relative depth
of clusters of three wells: s, shallow; i, intermediate.
w
R. Earle and G. Lethbridge (pers. commun.).
z
Measured on receipt of samples, February 2001.
‰
Measured in March 2000 (Jones et al., 2001).
z
Measured in March 2000 (R. Earle & G. Lethbridge, pers. commun.).
k
Trend of BTEX concentration over 4 years (Jones et al., 2001).
FEMS Microbiol Ecol 58 (2006) 260–270
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Published by Blackwell Publishing Ltd. All rights reserved
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262
CAC ATG CAA GTC-3 0 ) and HEX1389R (5 0 -ACG GGC
GGT GTG TAC AAG-3 0 ) (Marchesi et al., 1998). Primers for
clone libraries were supplied by Invitrogen: 27F (5 0 -AGA
GTT TGA TCM TGG CTC AG-3 0 ) and 1492R (5 0 -TAC GGY
TAC CTT GTT ACG ACT T-3 0 ) (Lane, 1991). PCR products
were viewed on 1% w/v agarose gels; duplicate PCR products were pooled and cleaned with Qiagen PCR purification
kit.
T-RFLP and cluster analyses
Ten microliters of fluorescent PCR product were digested
separately with 10 U of AluI and CfoI (Roche) for 3 h at
37 1C. T-RFLP profiles were obtained on an ABI Prism 310
genetic analyser (internal size marker Genescan 500 ROX,
POP4, filter set D, 36 cm capillary; ABI), and analyzed with
Genescan version 3.1 (ABI). Electropherograms were examined for any imperfections and rerun as necessary. The digital data for each primer and each enzyme were processed
separately: peak areas converted to relative abundance and
an alignment by fragment size produced, as previously described (Fahy et al., 2005). A dendrogram was generated
from this alignment by performing a hierarchical cluster
analysis using UPGMA (unweighted pair-group method using arithmetic averages) and a quantitative measure:
squared Euclidean distance (MVSP version 3.1, multivariate
statistical package, Kovach Computing Services).
Cloning, RFLP screening and sequencing
Cloning was carried out with TOPO TA Cloning Kit (Invitrogen) as per the manufacturer’s instructions. Positive
clones were screened by incubating the transformants on
LB-ampicillin-Xgal plates. PCR products were obtained
from positives clones by placing a few cells directly in a
50 mL reaction using primers T3 and T7 (5 0 -ATT AAC CCT
CAC TAA AGG GA-3 0 and 5 0 -TAA TAC GAC TCA CTA TAG
GG-3 0 ). The reactions were heated to 94 1C for 10 min before proceeding with standard cycling conditions.
PCR products of the correct size were grouped by RFLP
(restriction fragment length polymorphism) using CfoI and
AluI in succession. Clones with the same RFLP pattern for
both enzymes were assumed to belong to the same phylogenetic group, and one representative was sequenced.
Sequencing reactions were carried out using 1 mL of
10 pmol mL1 primer 518R (5 0 -CGT ATT ACC GCC GCT
GCT GG-3 0 ) (Invitrogen), 2 mL Big Dye Terminator V2.0
Cycle Sequencing Kit, 6 mL of 2.5 sequencing buffer,
2–8 mL of purified PCR product, and sdH2O to 20 mL. Cycling conditions were (96 1C, 15 s; 60 1C, 15 s; 60 1C,
4 min) 25. The products of the sequencing reaction were
precipitated with Na-acetate (pH 4.6) and ethanol, and sequenced in a Perkin Elmer ABI Prism 310 capillary electrophoresis automated genetic analyser as per manufacturer’s
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A. Fahy et al.
recommendations. The sequences were analyzed with DNA
Sequencing Analysis Software version 3.3 (reagents and software ABI Prism).
In order to link organisms with the community profiles,
T-RFLP profiles of the main clones were obtained by performing PCR reactions with FAM63F and HEX1389R primers from dilutions of the PCR products obtained with T3
and T7 primers, followed by restriction digestion and electrophoresis as described above. T-RFLP profiles of isolates
were also obtained.
Phylogenetic analyses
Sequences were subjected to an EMBL EBI Fasta33 search
(Pearson & Lipman, 1988). Chimeric sequences were eliminated using the Check Chimera application (Cole et al., 2003).
Preliminary alignments of partial 16S rRNA gene sequences
were obtained with the CLUSTALW tool (Thompson et al., 1994),
and the alignments were manually refined in GeneDoc (Nicholas & Nicholas, 1997). Phylogenetic trees were produced
with the Phylip interface (Felsenstein, 1993), using the Jukes
and Cantor distance and the neighbour-joining method.
Bootstrap analysis was carried out with 1000 replicates.
Nucleotide sequence accession numbers
The sequence data reported in this study were deposited in
the EMBL database under the accession numbers
AM110007–AM110077.
Results and discussion
Bacterial community structures
The relationship between the community structures from
the three sets of microcosms and the corresponding in situ
communities, as obtained by T-RFLP analysis, is illustrated
in a dendrogram (Fig. 1a). The three communities not
amended with nutrients (indicated by suffix ) cluster with
the in situ communities, showing that they had not changed
greatly during this experiment and the preceding 9 days of
storage at 4 1C. The benzene concentrations at the time of
collection of community DNA are shown in Fig. 1b. Benzene
degradation was generally, but not uniformly, initiated by
the presence of oxygen: for example, no benzene degradation occurred in Rs (no nutrient), Rs1P (with phosphate)
nor in Ri1N (with nitrate).
The communities from microcosms prepared with
groundwater from wells Ri and Pi each form a discrete cluster, B and D respectively, and both clusters include the communities sampled in situ. In contrast, a large shift in
community structure is observed between microcosms
prepared with groundwater from Rs, reflecting whether benzene degradation occurred (cluster C), or not (cluster A,
FEMS Microbiol Ecol 58 (2006) 260–270
263
Benzene-degrading communities in groundwaters
(a)
o
Rs +P
Rs–
Rs
# o
A
in situ
Ri+N
o
Ri+NP
+
Ri+P
+
#
Ri–
B
+
Ri in situ
Rs +NP
+
#
Rs +N
Pi+NP
+
#
Pi–
Pi+N
D
+
+
Pi in situ
2
1.6
1.2
0.8
0.4
0
squared Euclidean distance
(b)
Clone libraries
In order to characterize the communities and to distinguish
between benzene-degrading and nondegrading communities, representatives of clusters A, B, C and D (Fig. 1a) were
chosen to generate 16S rRNA gene clone libraries: Pi , Ri
and Rs from microcosms not amended with nutrients,
and Rs1N (with added nitrate). Benzene degradation was
recorded in these microcosms except Rs .
day 12
day 12
Ri+NP
Ri+P
Ri+N
Ri–
Ri day0
#
Rs+NP
Rs+N
#
Rs+P
Rs–
Rs day0
#
Pi+NP
Pi+P
Pi+N
Pi–
200
180
160
140
120
100
80
60
40
20
0
which includes the in situ community). It is worth noting
that such a shift in community structure is not observed between Ri microcosms: the T-RFLP profile from Ri1N (no
degradation) is marginally different from other Ri microcosms where degradation occurred (Fig. 1a).
As with all PCR-based techniques, T-RFLP analysis and
clone libraries have inherent biases and are not strictly quantitative; however all samples were treated identically
throughout each procedure, thus allowing between-sample
comparisons within this study.
FEMS Microbiol Ecol 58 (2006) 260–270
+
Pi+P
Pi day0
Fig. 1. Bacterial community dynamics in
groundwater microcosms enriched with benzene. (a) Relationship between bacterial community structures represented by a dendrogram
(UPGMA cluster analysis). Pi, Ri and Rs indicate
the location and relative depth of the groundwater samples. The suffix in situ indicates the
communities on receipt of groundwater samples.
Other suffixes indicate the treatment of benzene-spiked microcosms: , no treatment; N,
addition of ammonium nitrate; P, addition of
phosphate buffer; NP, addition of ammonium nitrate and phosphate buffer. Communities from
Ri and Pi microcosms each form a discrete cluster,
B and D. Communities from Rs form two distinct
clusters A and C, separating benzene-degrading
from nondegrading consortia. # = communities
chosen for clone libraries. 1 = benzene degradation, o = no benzene degradation, (b) Benzene
concentrations at the time communities were
harvested ( s.d.; n = 3). # = communities chosen for clone libraries.
C
+
#
day 28
A summary of the bacterial phylogenetic groups determined from 16S rRNA gene libraries further highlights the
distinct compositions of the four communities (Fig. 2). Betaproteobacteria dominate both Pi and Rs1N libraries,
while Gammaproteobacteria form the majority of Ri . Actinobacteria and Betaproteobacteria are evenly represented in
Rs . Details of each clone library are displayed in Table 2.
Most sequences are closely related to sequences from organisms previously detected in the following environments: water, primarily freshwater (e.g. groundwater, mineral water,
river, lake or glacier) with a few exceptions from marine or
brackish environments; soil or rhizosphere; contaminated
environments, including petroleum, chlorinated solvents
and heavy metal pollution, and activated sludge.
The library from Pi is dominated by Comamonadaceae
(Betaproteobacteria) including three distinct strains of Acidovorax: an Acidovorax sp. isolated from aerobic activated
sludge (Khan et al., 2002), A. delafieldii isolated from soil
(Sang et al., 2002) and an Acidovorax strain previously
named ‘Pseudomonas’ P51. The latter was isolated from a
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264
A. Fahy et al.
Pi–
Alpha,
Ri–
Beta,
Bacteroidetes;
Rs+N
Gamma,
Rs–
Delta Proteobacteria;
A Actinobacteria;
others.
Fig. 2. Bacterial divisions represented in the four clone libraries. Clones
with identical RFLP patterns for both AluI and CfoI were assumed to belong to the same group. No benzene degradation occurred in Rs .
mixture of sediment from the River Rhine and sand from a
Dutch water works (van der Meer et al., 1987) and can degrade chlorinated benzenes.
The Ri library is mainly comprised of a Pseudomonas sp.
(Gammaproteobacteria) (Table 2), related to a strain isolated
from a noncontaminated aquifer in the French Vosges (Ranjard et al., 2003). This strain, or close relatives thereof deposited in the database, is not specifically associated with the
degradation of petroleum compounds, however the propensity for a wide variety of pseudomonads to perform such
activities is well known.
The library from the benzene-degrading consortium Rs1N
consists mainly of organisms related to Rhodococcus erythropolis (Actinobacteria), and to an uncultured Betaproteobacterium
from a naphthalene-contaminated sediment (Jeon et al., 2003)
related to Hydrogenophaga flava (Betaproteobacteria, Comamonadaceae). Both types of organism are of environmental
significance: Hydrogenophaga spp. are frequent members of
water treatment communities (Kämpfer et al., 1993; Lemmer
et al., 1997) and R. erythropolis is a ubiquitous organism
(Brandão et al., 2002) with great catabolic versatility.
In contrast, the Rs library, generated from microcosms
where no aerobic benzene degradation occurred, is dominated by organisms related to Rhodoferax antarcticus (97.9%
similarity), a Betaproteobacterium isolated from a microbial
mat from the Antarctic (Madigan et al., 2000). R. antarcticus
is not, to the authors’ knowledge, associated with the degradation of hydrocarbons. These dominant Rhodoferax-like
clone sequences were more than 99% similar to the predominant uncultivated population detected in an anaerobic
underground reactor treating chlorobenzene-contaminated
groundwater (Alfreider et al., 2002). It is conceivable therefore that the organisms represented by clone Rs121 are also
from an anaerobic population; certainly no aerobic benzene-degrading strains related to this organism were isolated
in this study. Ongoing anaerobic studies may yield information on the function of this population.
In three of the clone libraries, several distinct ribotypes of
the dominant organism have been detected: the Acidovorax
and Rhodoferax spp. represented by clones Pi102 and Rs121
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respectively show seven ribotypes, and the Pseudomonas sp.
represented by clone Ri137 four ribotypes. Generally, the
clones from this study are 95% or more similar to sequences
of isolates. There are a few intriguing exceptions: for example clones Pi175 and Rs127 cluster with clones from a petroleum treatment unit and activated sludge but have
sequences 80% similar to the closest isolated organism (Table 2). Likewise, Rs134 sequence is similar to a clone sequence from an organism detected in drinking water (Table
2), but not to sequences of cultured organisms. The phylogenetic tree in Fig. 3 illustrates the relationship between the
organisms detected in the groundwater microcosms and reference strains; a detailed relationship within the Comamonadaceae (Betaproteobacteria), is shown in Fig. 4.
Seven different families of the Alphaproteobacteria group
were detected, many represented by a single clone (Fig. 3
and Table 2). However, several clones of Rhodobacter spp.
were found in the benzene-degrading communities Ri and
Rs1N; these organisms are closely related to R. capsulatus, a
bacterium capable of phototrophic and chemotrophic
growth using various alcohols as carbon sources (Pantazopoulous & Madigan, 2000). It is reasonable to speculate
that the Rhodobacter spp. may be involved in the metabolism
of intermediates of benzene degradation.
Isolates
Four benzene-degrading strains were isolated by extinction
dilution: one strain each from groundwaters Pi and Ri, and
two strains from Rs (Table 3). The dilution from which the
organisms were isolated gives some indication of their high
abundance in situ. A peak consistent with strain Ri71 is
dominant in the in situ T-RFLP profile from groundwater Ri
(Fig. 5), but strains Pi71, Rs71 and Rs73 are barely detected
in the corresponding in situ profiles.
Strain Ri71 is closely related to a Pseudomonas sp. isolated
from a Korean salt marsh, and clusters with P. anguilliseptica
(Fig. 3). Strain Rs71 clusters with Hydrogenophaga flava
(Fig. 4). Both isolates are closely related to organisms from
the corresponding clone libraries (Figs 3 and 4). The remaining two strains Pi71 and Rs73 were isolated from different wells and the sequenced portion of their 16S rRNA
gene is identical. They also share this sequence with 29
Rhodococcus erythropolis isolates, and their sequences differ
from clone Rs207 by just one base pair.
Furthermore, these Pseudomonas and Rhodococcus spp. were
the most abundant organisms in two of the benzene-degrading
consortia (Table 2 and Fig. 5, peaks 1 and 4, respectively). In
the benzene-degrading consortium Pi , the dominant benzene-degrading Acidovorax spp. were not isolated by extinction
dilution, in spite of their high numbers in clone libraries and
their predisposition to cultivation (Table 2). Instead, the
Rhodococcus strain Pi71 was isolated by this approach.
FEMS Microbiol Ecol 58 (2006) 260–270
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Benzene-degrading communities in groundwaters
Table 2. Details of clone libraries
Group
Clone
Freq
Pi (benzene-degrading community)
b, Comam
Pi101
12
Pi111
10
Pi102
8
Pi195
Pi155
Pi191
Pi176
Pi166
1
1
1
1
2
Pi163
1
Pi113
6
Bacteroid
Pi161
1
Actin, Nocar
?
Pi185
Pi184
Pi175
1
2
1
Ri (benzene-degrading community)
a, Rhodob
Ri203
5
a, Acetob
Ri231
Ri215
Ri244
Ri247
Ri207
3
1
1
1
1
b, Comam
Ri201
1
Ri208
3
Ri222
1
b, Oxalob
Ri216
Ri241
1
1
g, Pseudom
Ri137
23
Firm, Clostr
Ri134
Ri236
Ri220
Ri232
1
1
1
1
a, Rhizob
Firm, Strept
Ri246
1
Rs1N (benzene-degrading community)
a, Rhodob
Rs216
2
a, Rhizob
Rs262
2
b, Comam
Rs211
8
Rs223
1
Rs227
Rs254
2
1
FEMS Microbiol Ecol 58 (2006) 260–270
Closest relative (EMBL); notes
Accession number
% sim
Acidovorax delafieldii B7-7; polymer degrader
Acidovorax sp. KSP1; activated sludge; polymer deg
‘Pseudomonas’ sp. P51; chlorinated benzenes deg
Acidovorax delafieldii
As Pi102
As Pi102
As Pi102
As Pi102
‘Pseudomonas’ sp. P51
Acidovorax sp. BSB421; activated sludge
Clone; bioreactor treating contaminated soil
‘Pseudomonas’ sp. P51
Clone; benzene-contaminated groundwater
strain PB7; anaerobic propylbenzene culture
Rhodoferax antarcticus; Antarctic microbial mat
Clone; freshwater lake, Sweden
Flavobacterium ferrugineum
As Pi161
Rhodococcus erythropolis; dichloropropene reactor
Clone; petroleum land treatment unit
Wolinella succinogenes (e, Helicobacteria)
AF332182
AB076842
AF015487
AF332182
AF015487
AF015487
AF015487
AF015487
AF015487
Y18617
AY699582
AF015487
AY214181
AY686732
AF084947
AY509322
M62798
AY509322
AJ237967
AY154390
M26636
99.8
97.7
100
98.7
100
99.8
99.8
99.6
99.6
99.6
98.5
98.3
99.8
97.3
95.4
95.2
89.4
94.0
100
99.8
79.6
Clone; Mammoth Hot Springs, Yellowstone
Rhodobacter sphaeroides
Clone; Mammoth Hot Springs, Yellowstone
Agrobacterium sp. PB; soil
As Ri215
As Ri215
Clone; Rocky Mountain alpine soil
clone; PCB polluted soil
Acidisphaera rubrifaciens HS-AP3
Clone; reactor and GW with chlorobenzene
Rhodoferax antarcticus; Antarctic microbial mat
Clone; drainage water from magnesite mine, Austria
Hydrogenophaga taeniospiralis ATCC 49656
Clone; rape rhizoshere
‘Pseudomonas’ sp. P51
As Ri222
Clone; soil, uranium mine waste
Herbaspirillum sp. G8A1; denitrifyer
Pseudomonas sp. Hsa.28
Pseudomonas anguilliseptica B1
As Ri137
As Ri137
As Ri137
Clone; groundwater
Acidaminobacter hydrogenoformans; estuarine mud
Streptococcus uberis HN1; raw milk
AF446309
D16425
AF446309
AF482682
AF482682
AF482682
AY192273
AJ292602
D86512
AF407413
AF084947
AJ536813
AF078768
AJ295481
AF015487
AJ295481
AJ582194
AJ012069
AY259121
AF439803
AY259121
AY259121
AY259121
AY651823
AF016691
AB023576
99.3
96.5
99.5
100
100
99.8
95.0
92.7
90.0
100
99.2
99.4
97.5
99.6
99.6
99.1
99.2
92.8
99.0
98.7
99.0
99.1
98.6
95.6
95.3
99.6
Rhodobacter sp. Jip03; rotten rice straw
Rhodobacter capsulatus ATCC11166
Clone; grassland soil
Devosia neptuniae J1
Clone; naphthalene-contaminated sediment
Hydrogenophaga flava
Clone; Mammoth Hot Springs, Yellowstone
Hydrogenophaga atypica; activated sludge
As Rs223
AB122032
D16428
AF078292
AF469072
AY250107
AF078771
AF445679
AJ585992
AF445679
AF407413
96.7
95.8
99.0
95.8
99.2
98.1
99.2
98.7
99.2
99.8
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A. Fahy et al.
Table 2. Continued.
Group
Clone
Freq
Rs232
Rs257
1
3
Rs261
2
d?
Rs259
3
Actin, Nocar
Rs207
14
Actin, Microc
Rs264
3
Rs233
1
Rs (nonbenzene-degrading community)
a, Holosp
Rs134
1
a, Caulob
a, Sphing
Rs155
Rs178
1
1
a, Rhizob
Rs184
1
a, Hyphom
Rs154
1
b, Comam
Rs121
17
Rs197
Rs118
Rs152
Rs148
Rs159
Rs188
Rs169
1
1
1
3
1
1
1
Rs173
2
b, Alcalig
Rs112
1
b, Methylop
Rs200
1
g, Legion
g, Xanthom
Rs103
Rs195
1
1
?
Rs127
1
Bacteroid
Rs156
Rs191
1
1
Actin, Microc
Rs185
1
Verrucom
Rs111
2
Closest relative (EMBL); notes
Accession number
% sim
clone; reactor and GW with chlorobenzene
Rhodoferax antarcticus; Antarctic microbial mat
As Rs254
Clone; trichloroethene-contaminated site
Ramlibacter sp. HTCC332; trichloethene contam GW
Curvibacter delicatus
Clone; grassland soil
Acidovorax sp.; pupa of Diptera
Clone; Lake Fuchskuhle
Bacteriovorax sp. PNEc1
Clone; rainbow trout skin
Rhodococcus erythropolis EK5; dichloropropene biofilm
R. erythropolis; non-saline alkaline groundwater
Strain CanDirty7; Canada Glacier, Antarctica
Arthrobacter globiformis
Strain 12-8; aerobic copiotrophic bacterium, urban soil
Frigoribacterium sp. GWS-SE; sediment, Wadden Sea
AF084947
97.7
AF407413
AF422684
AY429716
AF078756
AY177768
AJ400840
AJ290009
AY294221
AJ631301
AJ237967
AJ717371
AF479339
M23411
AB008510
AY332185
98.7
97.7
96.0
94.8
99.8
99.4
97.6
96.7
100
99.8
99.8
98.8
98.3
99.
497.6
Clone; drinking water
‘Odyssella thessalonicensis’
Caulobacter crescentus CB15
Clone; drinking water
Sphingomonas sp. BN6; xenobiotic compounds deg
Strain MB12; drinking water
Methylosinus sporium
Strain PI_GH2.1.D7; tropical forest soil
Devosia neptuniae
Clone; reactor and GW with chlorobenzene
Rhodoferax antarcticus; Antarctic microbial mat
As Rs121
As Rs121
As Rs121
As Rs121
As Rs121
As Rs121
Clone; mineral water
Hydrogenophaga atypica strain BSB 41.8
Clone; contaminated sediment
Hydrogenophaga flava
Clone; Lake Inba, Japan
Alcaligenes sp. L6; 3-chlorobenzoate degrader
Clone; uranium and nitrate contaminated sediment
Methylophilus sp. strain ECd5
Legionella waltersii strain 2074-AUS-E
Clone; water downstream of manure
Nevskia ramosa strain OL1
Clone; activated sludge foam
Wolinella succinogenes (e, Helicobacteria)
Sphingobacterium sp. AC74
Clone; bacterioplankton, Lake Fuchskuhle
Flexibacter filiformis strain IFO 15056
Strain AH13; bacterioplankton, Lake Fuchskuhle
Arthrobacter sulfureus DSM 20167
Clone; activated sludge
Prosthecobacter sp. FC2
AY328755
AF069496
AE005930
AY328610
X94098
AY328843
Y18946
AY162048
AF469072
AF407413
AF084947
AF407413
AF407413
AF407413
AF407413
AF407413
AF407413
AF523049
AJ585992
AY250107
AF078771
AB195760
X92415
AY527746
AY436794
AF122886
AY212685
AJ001011
AF513093
M26636
AJ717393
AJ290011
AB078049
AJ289962
AB046358
Z94005
U60013
98.6
83.9
99.1
99.1
98.6
99.8
91.7
98.4
95.8
100
97.9
100
99.8
99.6
99.4
99.2
97.9
99.6
99.2
98.5
97.5
99.9
95.0
99.2
96.6
96.1
99.8
98.7
95.7
80.0
99.4
93.5
85.3
99.5
99.3
97.9
95.2
a, Alphaproteobacteria; Acetob, Acetobacteraceae; Caulob, Caulobacteraceae; Holosp, Holosporaceae; Hyphom, Hyphomicrobiaceae; Rhizob,
Rhizobiaceae; Rhodob, Rhodobacteraceae; Sphing, Sphingomonadaceae; b, Betaproteobacteria; Alcalig, Alcaligenaceae; Comam, Comamonadaceae;
Methylop, Methylophilaceae; Oxalob, Oxalobacteraceae; g, Gammaproteobacteria; Legion, Legionellaceae; Pseudom, Pseudomonadaceae; Xanthom,
Xanthomonodacaceae; d, Deltaproteobacteria; e, Epsilonproteobacteria; Bacteroid, Bacteroidetes; Actin, Actinobacteria; Microc, Micrococcaceae;
Nocard, Nocardiaceae; Firm, Firmicutes; Clostr, Clostridiaceae; Strept, Streptococcaceae; Verrucom, Verrucomicrobia.
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FEMS Microbiol Ecol 58 (2006) 260–270
267
Benzene-degrading communities in groundwaters
Fig. 3. Phylogenetic relationship of organisms detected in groundwaters Pi, Rs and Ri, based on partial sequences of 16S rRNA gene. Clones and
isolates from this study are in bold; the number of
similar clones detected is indicated in parentheses.
Jukes and Cantor distance (430–480 nucleotides);
neighbour-joining method; outgroup: Methanothrix
soehngenii; scale bar: average nucleotide substitution per base; bootstrap values (1000 samples)
4 60% are indicated. A detailed tree of Comamonadaceae is shown in Fig. 4.
Community profiles
The T-RFLP profiles of the microcosm communities chosen
for clone libraries are compared to the in situ community
profiles in Fig. 5. These electrophoresis patterns highlight
differences between the benzene-degrading and nondegrading consortia from groundwater Rs. The T-RFs corresponding to the most abundant clones detected in the microcosms
are indicated by numbered arrows. The T-RFLP analysis and
the clone libraries were generated from PCR products obtained with different sets of primer pairs, 63F-1389R and
27F-1492R, respectively, and it is worth noting that both
procedures give a consistent description of the communities.
Concluding remarks
The groundwaters chosen for this study have very low oxygen levels in situ caused by their long-term exposure to, and
degradation of, contaminants (principally benzene). In spite
FEMS Microbiol Ecol 58 (2006) 260–270
of this, aerobic benzene degradation occurred in all three
samples, and was carried out by highly distinct bacterial
communities (Figs 1a and 5, Table 2). The community profiles of Pi and Ri microcosms are similar to their in situ
profiles (Figs 1a and 5), with Acidovorax spp. abundant in
groundwater Pi, and Pseudomonas spp. in Ri. Given the excess of benzene over other possible sources of carbon and
energy, it is reasonable to expect that mainly microorganisms involved in benzene degradation would appear in these
T-RFLP profiles, suggesting that these aerobic benzene-degrading microorganisms are active in situ. It is understood
that benzene can be degraded with very low levels of oxygen
(Yerushalmi et al., 2001) or when coupled to nitrate reduction (Coates et al., 2001); it is also known that some pseudomonads can grow microaerophilically and by
denitrification (Chayabutra & Ju, 2000). In addition, denitrifying Acidovorax spp. have been isolated from sludge reactors (Etchebehere et al., 2001). It is therefore feasible that
these are the mechanisms by which these species degrade
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268
A. Fahy et al.
benzene in situ, and they switch to aerobic growth when oxygen is available.
In contrast, the benzene-metabolizing community from
Rs1N is very different from Rs where no degradation occurred, and from the in-situ community (Rs in situ; Figs 1a
50
150
2000
1000
200
250
300
Pi in situ
Pi71
0
3000
1500
0
2
3
3000
1500
0
Ri71
3000
1500
0
1
4
Pi–
+
Ri in situ
3000
1500
0
3000
1500
0
3000
1500
0
Fig. 4. Phylogenetic relationship of Comamonadaceae detected in
groundwaters Pi, Rs and Ri, based on partial sequences of 16S rRNA
gene. Clones and isolates from this study are in bold; the number of similar clones detected is indicated in parentheses. Jukes and Cantor distance (480 nucleotides); neighbour-joining method; outgroup:
Pseudomonas putida; scale bar: average nucleotide substitution per
base; bootstrap values (1000 samples) 4 60% are indicated.
100
5
Ri71
Rs–
+
Ri in situ
Ri73
3
Rs–
4
o
4
Rs+N
2
+
Fig. 5. T-RFLP profiles of the bacterial communities in situ and from
the communities chosen for clone libraries with primer 63F and
restriction enzyme AluI; T-RFs (terminal restriction fragments) from
primer 1389R are not shown. The vertical scale of the electropherograms represents the relative fluorescence, and the horizontal scale
the T-RF length in nucleotides. Several groups of organisms may
share the same T-RF length. 1, benzene degradation; o, no degradation.
The numbered peaks correspond to the T-RF profiles of the following
clones: 1, clone Ri137 (Pseudomonas spp.); 2, clones Pi101, Pi102,
Pi111, Rs173 (Acidovorax and Hydrogenophaga spp.); 3, clones Pi113,
Rs121 (Rhodoferax sp.); 4, clones Pi184, Rs207 (Rhodococcus erythropolis); 5, clones Ri203, Ri231 (Rhodobacter spp.). Peaks corresponding
to the T-RF profiles of isolates are indicated in the in situ community
profiles: Pi71, Rs73 (R. erythropolis), Ri71 (Pseudomonas sp.), Rs71
(Hydrogenophaga sp.).
Table 3. Details of benzene-degrading isolates
Group
GW
Strain
Dilution factor
Closest relative (EMBL database)
Accession number
% simw
b, Comam
Rs
Rs71
5
g, Pseudom
Ri
Ri71
5
Actin, Nocard
Pi
Pi71
6
Rs
Rs73
5
Clone Rs211 (this study)
Hydrogenophaga sp. YED1-18; arsenite oxydising biofilm
Hydrogenophaga flava
Clone Ri137 (this study)
Pseudomonas sp. GC06, salt marsh soil, Korea
Pseudomonas anguilliseptica B1;
Clone Pi184 (this study)
Rhodococcus erythropolis; 1,3-dichloropropene reactor
Clone Rs207 (this study)
Rhodococcus erythropolis; 1,3-dichloropropene reactor
AM110076
AY168753
AF078771
AM110075
AY690672
AF439803
AM110074
AJ237967
AM110077
AJ237967
99.4
99.2
98.3
99.2
100
97.9
100
100
99.8
100
All strains were isolated from extinction dilutions in minimal medium pH 7.2.
Dilution factor from which each organism was isolated.
w
Percentage similarity over 455–482 nt of the 16S rRNA gene.
b, Betaproteobacteria; Comam, Comamonadaceae; g, Gammaproteobacteria; Pseudom, Pseudomonadaceae; Actin, Actinobacteria; Nocard,
Nocardiaceae.
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FEMS Microbiol Ecol 58 (2006) 260–270
269
Benzene-degrading communities in groundwaters
and 5). Notably, Rhodoferax spp. dominated in situ, while
Rhodococcus erythropolis and Hydrogenophaga spp. co-dominated in the aerobic benzene-degrading microcosm. This
suggests that, while well Rs also harbours aerobic benzenedegraders, these are not necessarily active in situ, and that
the long-term decline in benzene concentration (Table 1) in
well Rs is not due to aerobic benzene degradation.
In this study, we observed aerobic benzene degradation at
the ambient pH of groundwater from wells Rs and Ri, respectively, 8.9 and 9.4. To the authors’ knowledge, benzene
degradation has not been reported to occur in such alkaline
conditions. Furthermore, we isolated several benzene-degrading strains closely related to the organisms detected by
cloning and sequencing of environmental DNA, despite the
fact that few organisms present in most environments are
readily culturable (Amann et al., 1995; Zengler et al., 2002).
Consequently we are investigating the pH, oxygen and benzene range for growth of these important isolates.
Acknowledgements
This work was funded by the BBSRC and the Environment
Agency (studentship award to AF), the Scottish Executive
Environment and Rural Affairs Department (UEX/001/03
Programme: project ISMoNACh) and the DTI within the
Bioremediation LINK Programme; a contribution from the
EU project COMMODE is also acknowledged. The authors
wish to thank Gordon Lethbridge and David Jones for coordinating access to the SIReN site and collecting the
groundwater samples.
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