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Sistema de Información Científica
Red de Revistas Científicas de América Latina y el Caribe, España y Portugal
Rev. Int. Contam. Ambient. 25 (3) 147-156, 2009
BACTERIAL POPULATION DYNAMICS AND SEPARATION OF ACTIVE DEGRADERS
BY STABLE ISOTOPE PROBING DURING BENZENE DEGRADATION IN A
BTEX-IMPACTED AQUIFER
Arturo ABURTO
1,2*
, and Andrew S. BALL
1,3
1
Department of Biological Sciences, University of Essex, Wivenhoe Park, Colchester CO4 3SQ, United
Kingdom. *Corresponding author: Arturo Aburto. Tel.: +52 55 58044600 ext 2677, Fax: +52 55 58046407.
E-mail address: aaburto@correo.cua.uam.mx
2
Present address: Universidad Autonoma Metropolitana, Artifcios 40, Miguel Hidalgo, Cuajimalpa, México.
3
Present address: School of Biological Sciences, Flinders University of South Australia, GPO Box 2100,
Adelaide, South Australia 5001
(Recibido agosto 2008, aceptado febrero 2009)
Key words: aerobic benzene degradation, SIP, groundwater, population dynamics
ABSTRACT
The activity and diversity of a groundwater bacterial community was studied during the
degradation of benzene in samples from a BTEX-contaminated aquifer (SIReN, UK)
through the use of denaturing gradient gel electrophoresis (DGGE), followed by excision
and sequencing of dominant bands. Rapid aerobic benzene degradation occurred in all
samples, with 60-70 % degradation of benzene. DGGE analysis revealed that unique,
stable bacterial communities were formed in each sample.
Pseudomonas putida
and
Aci-
dovorax delafeldii
were identifed in groundwater samples 308s and W6s respectively,
suggesting they are the important taxa involved in the degradation of benzene. Further
work based on stable isotope probing (SIP) of RNA using
13
C benzene was carried out.
Prominent bands were identifed as
Acidovorax
and
Malikia
genera; the latter is very
similar to the benzene-degrader
Hydrogenophaga
, which confrms the presence oF ac
-
tive benzene degraders in the groundwater samples. The identifcation oF the prominent
communities provides knowledge of the bioremediation processes occurring
in situ
and
the potential to enhance degradation. This study highlights the potential of combining
community fngerprinting techniques, such as DGGE, together with SIP.
Palabras clave: degradación aeróbica de benceno, SIP, agua subterránea, dinámica de poblaciones
RESUMEN
El presente estudio investiga la dinámica de poblaciones durante la degradación de ben-
ceno en muestras de un acuíFero contaminado con BTEX mediante la técnica de DGGE,
obtención de bandas y su posterior secuenciación. Los resultados indican degradación
aeróbica de benceno en todas las muestras y que las poblaciones son diferentes en cada
muestra, pero se mantienen estables durante la degradación del contaminante. Las prin-
cipales bandas obtenidas en el gel de poliacrilamida Fueron secuenciadas e identifcadas
como
Pseudomonas putida
y
Acidovorax delafeldii
en las muestras 308s y W6s respec-
tivamente, lo que sugiere que estos microorganismos son los principales degradadores
A. Aburto and A.S. Ball
148
INTRODUCTION
The contamination of groundwater represents a
major hazard in terms of both environmental damage
and public health; contaminants such as PAH (pol
-
yaromatic hydrocarbons), MTBE (methyl
tert
-butyl
ether), and BTEX (benzene, toluene, ethylbenzene
and xylenes) enter natural waters via wastewater ef-
Fuents from petroleum re±ning industries, rainwater
runoff from roads, accidental spills and leakages
from underground storage tanks. The most soluble
components of gasoline and diesel fuel are the BTEX
compounds, making them common contaminants in
a large number of aquifers. Among the BTEX com-
pounds, benzene requires special attention because
it is the most toxic, the most soluble (Sikkema
et al
.
1995) and carcinogenic (Dean 1985). Benzene is readi-
ly degraded aerobically, and recent studies have shown
it is also degraded in the absence of oxygen (Phelps
et al
. 1998, Rooney-Varga
et al
. 1999, Caldwell and
SuFita 2000, Villatoro-Monzón
et al
. 2003, Kasai
et al
.
2006). Further work is now required to elucidate the
microorganisms responsible for benzene degradation
in the various environments.
Stable isotope probing
Stable isotope probing (SIP) is used to identify
microorganisms that assimilate a speci±c growth
substrate. This is achieved by labelling the substrate
with a stable isotope (the most common is
13
C) that
will be incorporated into the microorganism s cellu-
lar biomarkers (e.g. lipids, DNA and rRNA) if they
utilize the substrate. The labelled nucleic acids can
then be resolved from the unlabelled ones by density
gradient ultracentrifugation, speci±c genes ampli±ed
and separated in a denaturing gel, and sequenced.
Thus, by sequencing the labelled nucleic acids, the
microorganisms responsible for the substrate uptake
can be identi±ed and therefore link function to mi
-
crobial identity. Although SIP can be performed with
DNA or rRNA, the latter presents some advantages
over DNA, such as a higher copy number and a higher
turnover rate, which is a reFection of cellular activity
independent of replication (Mane±eld
et al
. 2002a,
Mane±eld
et al
. 2002b, Lueders
et al
. 2004, Dumont
and Murrell 2005).
The aim of this study was to obtain a pro±le of
the bacterial community in several groundwater
samples during benzene degradation and also to link
the aerobic bacterial community in the groundwater
sample DW3d to a speci±c function, i.e. benzene
degradation. Sample DW3d was selected from the
SIReN site for this experiment because it is heavily
polluted (42.9 mgL
1
;
Table I
) with benzene (Earle
et
al
. 2001) and the microorganisms thriving in it have
been exposed to the contaminant for a long period,
TABLE I.
PHYSICAL AND CHEMICAL CHARACTERISTICS O² WELLS SAMPLED AT THE SIReN
Well
Class
a
pH
DO
b,c
(mgl
1
)
Benzene
c
Toluene
c
Ethyl
Benzene
c
m-
&
p-
xylene
c
o-
xylene
c
Total
PAH
Total
alkanes
W6s
Clean
5.3
0.00
<1
<1
<1
<1
<1
2.03
11.5
W18s
Clean
7.0
2.22
46
2
<1
<1
<1
0.56
12.2
308s
Low
10.2
0.16
15563
61
484
9
8
0.20
21.0
DW3s
Low
6.1
0.91
16009
171
142
89
120
1.24
27.6
DW3d
High
6.0
0.03
42969
1439
18390
1805
921
1.80
12.1
All values are in µg l
1
except where stated
a. Class = Classi±cation
b. DO = Dissolved Oxygen
c. Data from Shell Global Solutions measurements taken in April 2003 (G. Lethbridge, personal communication), except
for benzene measurement from DW3d which was measured on the second sampling session as part of this study.
en ciertas muestras. Mediante el uso de isótopos estables se comprobó la existencia de
microorganismos aerobios degradadores de benceno, los cuales fueron identi±cados
como miembros de los géneros
Acidovorax
y presumiblemente
Hydrogenophaga
en
otra de las muestras. La identi±cación de comunidades prominentes brinda información
sobre los procesos que ocurren in situ y del potencial para aumentar la degradación. El
presente estudio muestra el potencial de combinar técnicas como DGGE con isótopos
estables.
POPULATION DYNAMICS DURING BENZENE DEGRADATION
149
suggesting adaptation to the pollutant. Furthermore,
previous clone libraries indicate the presence of
aerobic hydrocarbon degraders such as
Polaromonas
naphthalenivorans
(Aburto
et al
. 2009).
MATERIALS AND METHODS
Study site and sampling
The study site is a BTEX-contaminated aquifer
located below an operational petrochemical plant
known as SIReN (Site for Innovative Research in
Natural Attenuation) in the UK. This study focuses on
fve wells showing diFFerent physicochemical char
-
acteristics and levels of contamination, nominally
classifed as high, low or clean (
Table I
). Details
about the site can be found elsewhere (Earle
et al
.
2001, Fahy
et al
. 2005).
The groundwater used to prepare the micro-
cosms was collected on the second sampling ses-
sion (19/05/05) as described previously (Aburto
et al
. 2009). Samples DW3s, 308s and W18s were
collected with a peristaltic pump, while a bladder
type was used for DW3d and W6s. Groundwater
was ±ushed From wells until pH, temperature and
conductivity were stable. Samples were collected in
1 L glass bottles. The groundwater was refrigerated
and protected from light during transportation to the
laboratory.
Microcosms and benzene monitoring
The microcosms were prepared approximately
eight hours after sampling by dispensing 60 mL of
groundwater samples DW3d, DW3s, W18s, W6s
and 308s in 110 mL serum bottles, and spiking with
13
C-benzene to a fnal concentration oF 25 mgL
1
,
which is about half of the last measurement
in situ
.
These groundwater samples were selected in order to
compare high contaminated, low contaminated, and
clean samples (
Table I
). Microcosms were incubated
in the dark at 12 °C (which is the
in situ
temperature)
for seven days. Autoclaved microcosms were used
as controls throughout the experiment. Standards
and controls all had the same liquid/headspace ratio,
and were held at the same temperature as the test
samples.
DNA extraction and PCR
DNA was extracted at four different time points
as described previously (Aburto
et al
. 2009) after the
preparation of the microcosms. The time points were
labelled: 0, 2, 5, and 7 days that correspond to a precise
sampling time of 12, 48, 120 and 168 h respectively.
Partial 16S rRNA gene fragments were obtained from
PCR amplifcation with bacterial Muyzer primers 1.
Forward with GC clamp (CGC CCG CCG CGC GCG
GCG GGC GGG GCG GGG GCA CGG GGG GCC
TAC GGG AGG CAG CAG) and 2 Reverse (ATT
ACC GCG GCT GCT GG) (
E. coli
position 341 to
534) (Muyzer 1993). The cycling conditions were as
follows: an initial denaturation step of 94 °C for 1
min, followed by 30 cycles of denaturation at 94 °C
for 1 min, annealing at 55 °C for 1 min, and extension
at 72 °C For 2 min. Cycling was completed by a fnal
elongation at 72 °C for 10 min.
DGGE
PCR products with a GC clamp were separated
without purifcation in an 8 % w/v polyacrylamide
gel with a linear denaturing gradient, increasing
from 40 % at the top of the gel to 60 % at the bottom
(100 % denaturants correspond to 7 M urea and 40 %
v/v formamide). The DGGE gel was run at 60 °C for
5 h in a D Code Universal Mutation Detection System
(Bio-Rad Hercules, CA, USA). Gels were stained
For 15 min with 15 µL oF SYBR Gold (Molecular
Probes BV, Leiden, Netherlands) diluted in 150 mL
of 1X TAE buffer. Stained gels were visualized in a
GelDoc System (Bio Rad). Dominant bands were
excised and incubated in 40 µL of elution buffer (0.5
M ammonium
acetate, 10 mM magnesium acetate, 1
mM EDTApH 8 and 0.1 % w/v SDS) For 4 h at 37 °C.
DNA was precipitated with two volumes of absolute
ethanol, washed with 70 % w/v ethanol and air dried.
The pellet was resuspended in 20 µL DEPC treated
water. Bands 1, 4, 9, 12, 13 (
Fig. 1
) were cloned
in order to avoid sequencing co-migrating DGGE
bands. The 16S rRNAgene Fragment was reamplifed
with primer Forward M13 and reverse M13 in a frst
instance to confrm presence oF the insert and later
with both forward and reverse Muyzer primers prior
to sequencing and identifcation.
Sequencing
Sequencing reactions contained 1 µL of 10 pmol
µL
1
of the reverse primer 1389R (Invitrogen), 2 µL
of Big Dye Terminator V2.0 Cycle Sequencing kit, 6
µL of 2.5
×
sequencing buFFer, 5 µL oF purifed PCR
product (approx 15 ng) and 6 µL oF water. Amplif
-
cation conditions were: 25 cycles of (96 °C, 15 sec;
60 °C, 15 sec; 60 °C, 4 min). After sodium acetate
and ethanol precipitation, the sequencing reaction
products were run on a Perkin Elmer ABI PRISM
310 capillary electrophoresis automated genetic
analyzer. Sequences were then processed with DNA
Sequencing Analysis Software version 3.3.
A. Aburto and A.S. Ball
150
Stable isotope probing
The protocol used in this study is an adaptation of
the method used by Manefeld
et al.
(2002a). Micro-
cosms containing 100 mL of groundwater (prepared
eight hours after sampling) from well DW3d were
spiked with the stable isotope
13
C-benzene (all carbon
atoms labelled) to 25 mgL
1
as fnal concentration
and incubated at 12 °C for 10 days. Nucleic acid
extractions were performed at time 0 (4 hours after
the
13
C benzene spike) and on the days 2, 5 and 7.
RNA was isolated and quantifed. 50 µL oF template
RNA (approx. 10 ng) was added to a mixture of 9.69
mL caesium tri±uoroacetate (2.0 g mL
1
), 0.39 mL
Formamide and 1.37 mL H
2
O in an 11.5 mL Sorvall
ultracentrifugation tube (supplied by Kendro labora-
tories). The mixture was centrifuged at 35,000 rpm
(119,500 x
g
), for 48 h in a Sorvall Discovery 90SE
Ultraspeed Centrifuge, with a TST 41.14 rotor. It was
later fractionated in 0.5 mL aliquots by piercing the
bottom of the centrifuge tube with a needle to collect
the Fraction, and injecting sterile H
2
O on top of the
tube with the help of a peristaltic pump at a rate of 10
µL per second. A total of 20 fractions were collected
per gradient, and refractive indices were measured.
The RNA of each fraction was precipitated with
isopropanol as follows: 1 mL isopropanol was added
to each fraction and kept at
20 °C for 1 hour, centri-
fuged at 13,000 rpm for 30 min at 4 °C; the pellet was
washed with 70 % ethanol, air dried and resuspended
in 25 μL oF DEPC treated H
2
O. Each fraction was
later subjected to reverse transcription and PCR am
-
plifcation with Muyzer primers containing the GC-
clamp. Appropriate fractions were selected and run
in an acrylamide gel For community fngerprinting.
Bands 1 to 7 were excised, cloned and reamplifed
with Muyzer primers (without the GC clamp) for
sequencing. Sequences were compared with those
in the EMBL database using a FastA search in order
to identify the closest relatives.
For clarity, from now on, a sample spiked with
13
C-benzene will be called heavy sample , while a
sample with background concentrations of benzene
will be reFerred as “light sample”. In order to confrm
the separation of labelled from the non-labelled RNA
in sample DW3d, heavy and light gradients obtained
from the isolate
Rhodococcus
Q71 were tested.
The
Rhodococcus
isolate Q71 was grown on
12
C
and
13
C-benzene separately. Gradients were prepared
with RNA extracted from both conditions. RNA
Fractions were reverse transcribed, amplifed with
Muyzer primers and run in an agarose gel.
In the same way as with the
Rhodococcus
iso-
late, microcosms prepared with sample DW3d were
amended separately with
12
C and
13
C-benzene. The
gradients were centrifuged, fractionated and the
refractive indices measured for each fraction (
Table
III
). The fractions were treated as described for the
Rhodococcus
strain.
RESULTS
Changes in bacterial community during benzene
degradation
Benzene degradation was detected from the sec-
ond measurement at day 2 for most of the samples
and until the fnal measurement on day 7 (
Fig. 2
)
when the last microcosm was sacrifced For nucleic
acid extraction. Generally, the rates of benzene
degradation were similar for all samples (except the
killed control), with a 60-70 % reduction in benzene
concentration remaining after the 7 d incubation.
The DGGE pattern (
Fig. 1
) obtained from the
different samples corresponds to each of the time
points of benzene monitoring by gas chromatogra-
phy (
Fig. 2
) and shows a stable community over
time in most of the samples as well as different com-
munities among the samples, which can also be seen
in a dendrogram (
Fig 3
). All of the prominent bands
in the gel were excised in order to identify them;
however, several bands (n = 7) had poor sequence
quality. Good quality sequences were retrieved from
DW3d
DW3s
W18s
W6s
308s
02 57
02 57
02 57
02 57
02 57
14
5
6
8
7
9
12
13
11
Fig. 1.
DGGE gel showing the profle oF the time series analysis
carried out on microcosms spiked with
13
C benzene in
the following samples: DW3d, DW3s, W18s, W6s and
308s. The numbers 0, 2, 5, and 7 indicate the day at which
the community was analyzed
POPULATION DYNAMICS DURING BENZENE DEGRADATION
151
the bands marked on the DGGE gel (
Fig. 1
) and their
closest relatives are shown in
table II
.
The sequences retrieved from sample DW3d
were closely related (97 % similar) to
Malikia gra-
nosa
and a bacterium isolated from soil (
Table II
).
The organisms retrieved from the other groundwater
samples (
Fig. 1
) included the following bacterial
species:
Aquaspirillum autotrophicum
(band 7) in
sample W18s,
Lactococcus lactis
(bands 5, 8, 11)
in samples W18s, W6s and 308s,
Bacteriovorax
stolpii
(previously known as
Bdellovibrio stolpii
;
Baer
et al
. 2000) (band 6) in sample W18s, and
Pseudomonas putida
strain KT 2440 (band 12) in
sample 308s. The sequences retrieved from bands
9 (sample W6s) and 13 (sample DW3d) matched
with the organisms
Acidovorax delafeldii
and a
bacterium found in a TEX mixture respectively;
however, they were distant to this organism with
16S rRNA sequence similarity below 90 % for both
sequences (
Table II
).
0
5
10
15
20
25
30
35
40
45
01234567
Time (days)
308s
W18s
W6s
DW3s
DW3d
Killed Control (DW3d)
Benzene concentration (mg l
-1
)
Fig. 2.
Degradation of
13
C benzene in samples 308s, W18s,
W6s, 308i, DW3s, DW3d. Error bars represent 1 SD of
two samples. One representative control is shown (i.e.
autoclaved groundwater from well DW3d)
Group average
W18s-0
DW3s-0
DW3d-7
DW3d-0
DW3d-2
W6s-5
W6s-7
W6s-0
W6s-2
W18s-7
W18s-2
W18s-5
DW3s-2
DW3s-5
DW3s-7
Samples
10
08
06
04
02
0
Similarity
Transform: Presence/absence
Resemblance: S17 Bray Curtis similarity
Fig. 3.
Dendrogram showing the relationship between bacterial communities
from Fve groundwater samples over seven days
. The numbers 0, 2, 5,
and 7 indicate the number of days at which the nucleic acids extraction
was performed. Some time points had to be left out of the analysis since
bands in the DGGE gel were very faint
TABLE II.
CLOSEST RELATIVES OF BAND SEQUENCES
EXCISED FROM DGGE IN
FIG 1
, # CLONED
BANDS
Sample Band
Closest relative
Accession
number
%
Similarity
DW3d
#1
Malikia granosa
AJ627188
97
#13
Bacterium isolated
from oil
AB081542
80
DW3s
#4
Malikia granosa
AJ627188
91
W18s
5
Lactococcus lactis
AE006288
94
6
Bacteriovorax stolpii
AJ288899
99
7
Aquaspirillum
autotrophicum
AB074524
99
W6s
8
Lactococcus lactis
AE006288
94
#9
Acidovorax delafeldii
AF078764
86
308s
11
Lactococcus lactis
AE006288
94
#12
Pseudomonas putida
KT2440
AE016776
96
A. Aburto and A.S. Ball
152
The use of stable isotope probing for detection of
RNA from isolate Q71,
Rhodococcus erythropolis
Stable isotope probing relies on the separation of
labelled and non-labelled nucleic acids; therefore, the
ultracentrifugation is critical and may be the most
important step in the process. Gradients were centri-
fuged, fractionated and the refractive indices (
Table
III
) indicated a good separation of the fractions.
The presence of amplicons in the Frst fractions
(1 to 5) of the heavy sample for the isolate Q71 and
their absence in the light sample suggests that the
ultracentrifugation achieved the goal of separating
the labelled from the non-labelled nucleic acids
(
Fig. 4A, B
). This is supported by the appearance of
bands only from the seventh fraction onwards in the
light sample, except for a faint band on the second
fraction. However, this band may be the product of
experimental contamination as has been described
previously (ManeFeld
et al
. 2002a), since no bands
are visible in the adjacent fractions (1, 3, 4, 5, 6 in
Fig. 4B
).
The use of stable isotope probing for detection of
RNA from environmental sample DW3d
On the basis of the successful results on the iso-
late Q71, it was decided to test the technique SIP
on the environmental sample DW3d. The sample,
designated as heavy, in fact contains a mixture of
13
C-
labelled benzene (25 mgL
1
) and unlabelled benzene
present in the groundwater, which is consistent with
the presence of amplicons from all the fractions of
the gradient. This is further supported by the lack of
amplicons in the Frst fractions of the light sample and
their presence from the sixth sample onwards (
Fig
5B
). Tentative evidence to support the separation of
labelled and non-labelled DNA comes from the faint
band in fraction 7, which is the zone where heavy
and light nucleic acids are separated.
The presence of amplicons in all the fractions of
the heavy sample was evident (
Fig. 5A
). This was
expected since this sample contains heavy and light
benzene; in contrast, amplicons are only visible from
the sixth fraction onwards in the light sample (
Fig.
5B
). This suggests that fractions 1 to 5 in the heavy
sample correspond to labelled DNA and is consistent
with the result observed with the isolate Q71.
Fractions 1 to 5 and 6 to 10 of the heavy DW3d
TABLE III.
RE±RACTIVE INDICES O± THE ISOLATE Q71
Isolate
13
C-
Rhodococcus
Q71
gmL
-1
Isolate
12
C-
Rhodococcus
Q71
gmL
-1
Fraction
Density
Density
1
2.0100
2.0100
2
1.9969
1.9969
3
1.9969
1.9969
4
1.9969
1.9805
5
1.9696
1.9561
6
1.9561
1.9534
7
1.9291
1.9291
8
1.9291
1.9291
9
1.9157
1.9024
10
1.9157
1.9024
11
1.9051
1.9024
12
1.899
1.8990
13
1.8759
1.8891
14
1.8733
1.8785
15
1.8621
1.8759
16
1.8575
1.8627
17
1.8496
1.8627
18
1.5981
1.7217
19
1.1584
1.5981
20
1.1584
1.1584
16
17
18
+
-
1Kb
a)
b)
123
4
567
8
9
10
11
12
13
14
15
1Kb
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
+
-
16
17
18
+
-
+
-
16
17
18
Fig. 4.
PCR products from fractions collected from two gradients
of isolate
Rhodococcus
Q71.
A
: the isolate was spiked
with
13
C benzene.
B
: the isolate was spiked with
12
C
benzene. Fraction 1 corresponds to the bottom of the
tube (most dense fraction); 18 corresponds to the top of
the tube (least dense fraction)
POPULATION DYNAMICS DURING BENZENE DEGRADATION
153
sample were assumed to represent the heavy and
light DNA respectively, and were subjected to
DGGE analysis. The profle obtained For both sets
of fractions was very similar, and bands 1 to 7 were
identical (
Fig. 6
). Based on 16S rRNA sequences
retrieved from the bands excised from the gel, spe-
cies like
Malikia granosa
and
Acidovorax
sp. KSP1
(
Table IV
), with sequence similarities of 92 and 91
%, respectively, were present.
DISCUSSION
Changes in bacterial community during benzene
degradation
Rapid aerobic benzene degradation was observed
for most of the groundwater samples (
Fig. 2
), which
confrms the presence oF adapted aerobic benzene
degraders in each of the samples, in spite of the low
amounts of oxygen
in situ
in wells DW3d, DW3s,
high pH (10.2) in 308s and low benzene concentra
-
tions in samples such as W18s and W6s (
Table I
;
Earle
et al
. 2001).
The DGGE profile (
Fig. 1
) suggests that the
communities remained quite stable over 7 days, yet
each groundwater community was different from
each other. The relationship of these communities
can also be observed in a dendrogram depicting
the dissimilarity among these groundwater samples
(
Fig. 3
). Most of the different time points cluster
according to the groundwater sample and not the
time point, confrming that each groundwater sample
harbours different communities and that they do not
change signifcantly over time. The only time point
that was out oF any cluster was W18s-0. However,
this is understandable because the DGGE profle For
this time point only shows one band along the lane
(
Fig. 1
, band 5) while the successive time points
for that sample include several bands in each lane.
Furthermore, the rate of benzene degradation was
the slowest for this sample. This suggests that the
community changed dramatically in this sample from
time 0 to time 2. However, it remained stable For
successive time points. Samples DW3d and DW3s
share fve bands; one oF those bands was identifed as
the microorganism
Malikia granosa
in both samples
(
Fig. 1
, Bands 1 and 4).
Malikia
granosa
and
Malikia spinosa
are poly-
16
17
18
19
20
+
-
16
17
18
+
-
1Kb
a)
1
23
4
567
8
9
10
11
12
13
14
15
1Kb
b)
1
23
4
567
8
9
10
11
12
13
14
15
16
17
18
19
20
+
-
16
17
18
+
-
Fig. 5.
PCR products from fractions collected from two gradients
of groundwater sample DW3d.
A
: the groundwater sam-
ple was spiked with
13
C benzene.
B
: groundwater sample
spiked with
12
C benzene. Fraction 1 corresponds to the
bottom of the tube (most dense fraction); 18 corresponds
to the top of the tube (least dense fraction)
Fig. 6.
DGGE gel showing the profle oF Fractions 1-10 From
groundwater sample DW3d obtained after gradient ultra centrifu-
obtained after gradient ultra centrifu-
gation in order to separate
13
C benzene from
12
C benzene, Time
point 0: 4 hours after the
13
C benzene spike. DNA in fractions
1-5 is suspected to have incorporated
13
C benzene. Far left and
right lanes contain a reference ladder
1
2 3
4
56
7
8 910
DW3d 2005
1
2
6
3
5
7
4
13
C
12
C
A. Aburto and A.S. Ball
154
hydroxyalkanoate and polyphosphate accumulat-
ing bacteria (Spring
et al
. 2005) and are related
to the
Hydrogenophaga
species; this is supported
by the previous detection of similar clones in this
groundwater sample (DW3d) (Aburto
et al
. 2009).
Furthermore, isolates of a benzene-degrading
Hy-
drogenophaga
strain in another location (308s) of
the same site have been obtained (Fahy
et al
. 2008).
And after sequences analysis, 99 % percent similar-
ity was shown between bands 1, 4 and the
Hydrog-
enophaga
isolate. One of the sequenced bands from
groundwater sample W18s was related, with a 16S
rRNA gene sequence similarity of 99 % to the organ-
ism
Aquaspirillum autotrophicum
. This is an aerobic
hydrogen-oxidizing bacterium that contains one or
more hydrogenase enzymes that bind hydrogen and
use it either to produce ATP or for reducing power
for autotrophic growth (Madigan 2000). This bacte-
rium has been previously isolated from a eutrophic
freshwater lake, and it can grow autotrophically or
heterotrophically (Aragno 1999).
A constant band appearing at time zero for sam-
ples W18s, W6s and 308s (bands 5, 8 and 11;
Fig.
1
) was related, with a 94 % 16S rRNA similarity,
to
Lactococcus lactis
. This organism, as indicated
by its name, produces lactic acid as a sole fermen-
tation product; it is an aerotolerant anaerobe able
to grow even in the presence of oxygen (available
in sample W18s;
Table I
). Another constant band
appearing throughout the second to the seventh day
in sample W18s was identifed with a 16S rRNA
gene sequence similarity of 99 % to
Bacteriovorax
stolpii
(band 6;
Fig. 1
), previously known as
Bdell-
ovibrio stolpii
(Baer
et al
. 2000). This organism is
known for its ability to invade other bacteria and
live parasitically on the host, until the host s death
when a new generation of the
B. stolpii
is released
(Kadouri and O Toole 2005, Lambert
et al
. 2006,
Rogosky
et al
. 2006). They can be found in diverse
environments such as marine and freshwaters, sew-
age and soil.
A good quality sequence was obtained from
sample 308s (
Fig. 1
; band 12); this band was closely
related to
Pseudomonas putida
strain KT2440,
which is a well known hydrocarbon degrader. This
organism contains a large number of dioxygenases
and at least four pathways for the degradation of
hydrocarbons (Jiménez
et al
. 2002). Therefore, this
bacterium is likely to carry out benzene degrada-
tion in spite oF the high pH (10.2) and this is also
supported by the isolation of benzene-degrading
Pseudomonas
species from the same well (Fahy
et
al
. 2008).
It can be concluded that each of the groundwater
samples harbours different communities that tend to
remain stable throughout the degradation of the pol-
lutant and that rapid benzene degradation suggests
the presence of acclimated aerobic degraders in the
groundwater samples studied here.
SIP
The community fingerprint of sample DW3d
obtained by DGGE resulted in an identical profle
for heavy and light nucleic acids (
Fig 6
). This is ex-
plained by the availability of both types of substrates
(heavy and light) in the sample and it is clear that
they were making use of both. Moreover, benzene
was by far the main growth substrate present in the
microcosm at concentrations far higher than any
other, ruling out the possibility of another substrate
for the microbes to feed on (
Table I
). The results in
fgure 6
demonstrate that those organisms appearing
in DGGE profles shortly aFter the onset (time point
0: 4 hours after the
13
C benzene spike) of benzene
degradation are likely to be directly involved in
benzene degradation.
The most prominent bands were related to the
strains
Malikia granosa
and
Acidovorax
sp. KSP1
(
Table IV
). Both species had been detected in dif-
ferent activated sludges elsewhere;
Malikia
granosa
is a polyhydroxyalkanoate and polyphosphate ac-
cumulating bacteria (Spring
et al
. 2005) closely
related to the
Hydrogenophaga
genera and, on this
basis and the fact that several
Hydrogenophaga
strains have been detected and isolated from this
and other groundwater samples of the same site,
we strongly suggest that these bands are in fact
benzene-degrading
Hydrogenophaga
strains identi-
fed as
Malikia
due to the shortness of the amplicon.
Acidovorax
strain KSP1 is a denitrifying bacterium
capable of degrading the polyhydroxyalkanoate:
poly (3-hydroxybutyrate-co-3-hydroxyvalerate)
(Khan
et al
. 2002).
TABLE IV.
CLOSEST RELATIVES OF BAND SEQUENCES
EXCISED FROM DGGE IN
FIG. 6
Band
Closest relative
Accession
number
%
Similarity
1
Malikia granosa
AJ627188
92.1
2
Malikia spinosa
AB077038
91.7
3
Acidovorax
sp.
KSP1
AB076842
91.0
4
Acidovorax
sp.
KSP1
AB076842
91.0
5
Malikia spinosa
AB077038
92.0
6
Beta proteobact Wuba 72
AF336361
89.0
7
Malikia spinosa
AB077038
92.0
POPULATION DYNAMICS DURING BENZENE DEGRADATION
155
CONCLUSIONS
Fast aerobic degradation of benzene con±rms the
presence of acclimated microbes
in situ
that readily
utilize the oxygen when it is available to degrade
benzene; in order to support this, a few strains have
been isolated from the same source (308s) (Fahy
et
al
. 2008) as well as detected in previous clone li-
braries (Fahy
et al
. 2006). The population dynamics
study shows different communities for each of the
samples.As for the identi±cation of benzene degrad
-
ers by SIP in one of the samples, we succeeded in
the separation of the labelled RNAand identi±cation
of the prominent bands from the heavy fractions;
however, it is necessary to increase the certainty in
the identi±cation of those degraders by obtaining
and sequencing a larger amplicon. Nevertheless, this
study shows that stable isotope probing was useful
in order to link microbial identity to function in a
contaminated sample.
REFERENCES
Aburto A., Fahy A., Coulon F., Lethbridge G., Timmis
K.N., Ball A.S. and McGenity T.J. (2009). Mixed
aerobic and anaerobic microbial communities in
benzene-contaminated groundwaters J. Appl. Micro-
biol. 106, 317-328.
Aragno M.H.G.S. (1999).
The mesophilic hydrocarbon-
oxidizing (knallgas) bacteria. The Prokaryotes, an
evolving electronic resource for the microbiological
community.
Springer-Verlag release 3.14, New York,
USA, 1120 p.
Baer M., Ravel J., Chun J., Hill R. and Williams H. (2000).
A proposal for the reclassi±cation of
Bdellovibrio
stolpii
and
Bdellovibrio starrii
into a new genus,
Bacte-
riovorax
gen. nov. as
Bacteriovorax stolpii
comb. nov.
and
Bacteriovorax starrii
comb. nov., respectively. Int.
J. Syst. Evol. Microbiol
.
50, 219-224.
Caldwell M.E. and Su²ita J.M. (2000). Detection of
phenol and benzoate as intermediates of anaerobic
benzene biodegradation under different terminal
electron-accepting conditions. Environ. Sci. Technol.
34, 1216-1220.
Dean B.J. (1985). Recent ±ndings on the genetic toxicol
-
ogy of benzene, toluene, xylenes and phenols. Mutat.
Res. 154, 153-181.
Dumont M.G. and Murrell J.C. (2005). Stable isotope
probing - linking microbial identity to function. Nat.
Rev. Microbiol. 3
,
499-504.
Earle R., Jones D., Lethbridge G., McCarthy P. and Thom-
son S. (2001). P2-208/TR/2. Project SIReN: Phase 2a.
Conceptual site model & groundwater model
.
R&D
technical report. Environment Agency, Bristol, UK
,
80 pp.
Fahy A., McGenity T.J., Timmis K.N. and Ball A.S. (2006).
Heterogeneous aerobic benzene-degrading communi
-
ties in oxygen-depleted groundwaters. FEMS. Micro-
biol.
Ecol. 58, 260-270.
Fahy A., Lethbridge G., Earle R., Ball A.S., Timmis
K.N. and McGenity T.J. (2005). Effects of long-term
benzene pollution on bacterial diversity and commu-
nity structure in groundwater. Environ. Microbiol. 7,
1192-1199.
Fahy A., Ball A.S., Lethbridge G., Timmis K.N. and
McGenity T.J. (2008). Isolation of alkalitolerant
benzene-degrading bacteria from a contaminated
sandstone aquifer. Lett. Appl. Microbiol. 47, 60-66.
Jiménez J.I., Minambres B., García J.L. and Díaz E.
(2002). Genomic analysis of the aromatic catabolic
pathways from
Pseudomonas putida
KT2440. Environ.
Microbiol. 4, 824-841.
Kadouri D. and O Toole G.A. (2005). Susceptibility of
bio±lms to
Bdellovibrio bacteriovorus
attack. Appl.
Environ. Microbiol
.
71, 4044-4051.
Kasai Y., Takahata Y., Mane±eld M. and Watanabe K.
(2006). RNA-based stable isotope probing and isola-
tion of anaerobic benzene-degrading bacteria from
gasoline-contaminated groundwater. Appl. Environ.
Microbiol. 72, 3586-3592.
Khan S.T., HoribaY.,Yamamoto M. and HiraishiA. (2002).
Members of the family
Comamonadaceae
as primary
poly(3-hydroxybutyrate-co-3-hydroxyvalerate)-
degrading denitri±ers in activated sludge as revealed
by a polyphasic approach. Appl. Environ. Microbiol.
68, 3206-3214.
Lambert C., Evans K.J., Till R., Hobley L., Capeness M.,
Rendulic S., Schuster S., Aizawa S. and Sockett E.
(2006). Characterizing the ²agellar ±lament and the
role of motility in bacterial prey-penetration by
Bdell-
ovibrio bacteriovorus
.
Mol. Microbiol
.
60, 274-286.
Lueders T., Mane±eld M. and Friedrich M.W. (2004).
Enhanced sensitivity of DNA- and rRNA-based stable
isotope probing by fractionation and quantitative
analysis of isopycnic centrifugation gradients. Environ.
Microbiol. 6, 73-78.
Madigan M.T.M. (2000).
Brock Biology of Microorgan-
isms
. Prentice Hall, Upper Saddle River, New Jersey,
992 pp.
Mane±eld M.,WhiteleyA.S., Grif±ths R.I. and Bailey M.J.
(2002a). RNA stable isotope probing, a novel means of
linking microbial community function to phylogeny.
Appl. Environ. Microbiol. 68, 5367-5373.
Mane±eld M., Whiteley A.S., Ostle N., Ineson P. and
Bailey M.J. (2002b). Technical considerations for
A. Aburto and A.S. Ball
156
RNA-based stable isotope probing: an approach to
associating microbial diversity with microbial com-
munity function. Rapid Commun. Mass Spectrom.
16, 2179-2183.
Muyzer G. (1993). Profiling of complex microbial
populations by denaturing gradient gel electrophoresis
analysis of polymerase chain reaction-ampliFed genes
coding for 16S rRNA. Appl. Environ. Microbiol. 59,
695-700.
Phelps C.D., Kerkhof L.J. andYoung L.Y. (1998). Molecu
-
lar characterization of a sulfate-reducing consortium
which mineralizes benzene. FEMS Microbiol.
Ecol
.
27, 269-279.
Rogosky A.M., Moak P.L. and Emmert E.A.B. (2006).
Differential predation by
Bdellovibrio bacteriovorus
109J. Curr. Microbiol.
52, 81-85.
Rooney-Varga J.N., Anderson R.T., Fraga J.L., Ringelberg
D. and Lovley D.R. (1999). Microbial communities
associated with anaerobic benzene degradation in a
petroleum-contaminated aquifer. Appl. Environ. Mi-
crobiol. 65, 3056-3063.
Sikkema J., Debont J.A.M. and Poolman B. (1995).
Mechanisms of membrane toxicity of hydrocarbons.
Microbiol. Rev. 59, 201-222.
Spring S., Wagner M., Schumann P. and Kampfer P. (2005).
Malikia granosa
gen. nov., sp. nov., a novel polyhy-
droxyalkanoate- and polyphosphate-accumulating
bacterium isolated from activated sludge, and reclas-
siFcation of
Pseudomonas spinosa
as
Malikia spinosa
comb. nov.
Int. J. Syst. Evol. Microbiol.
55, 621-629.
Villatoro-Monzón W.R., Mesta-Howard A.M. and Razo-
Flores E. (2003). Anaerobic biodegradation of BTEX
using Mn(IV) and Fe(III) as alternative electron ac-
ceptors. Water Sci. Technol. 48, 125-131.
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