Evidence of the activity of dissimilatory sulfate-reducing
prokaryotes in nonsul¢dogenic tropical mobile muds
Vanessa M. Madrid1, Robert C. Aller1, Josephine Y. Aller1 & Andrei Y. Chistoserdov2
1
Marine Sciences Research Center, Stony Brook, NY, USA and 2Department of Biology, University of Louisiana at Lafayette, Lafayette, LA, USA
Correspondence: Andrei Y. Chistoserdov,
P.O. Box 42451, Department of Biology,
University of Louisiana at Lafayette, Lafayette,
LA 70504, USA. Tel.: 11 337 482 1330;
fax: 11 337 482 5660; e-mail:
[email protected]
Received 4 October 2005; revised 2 January
2006; accepted 3 January 2006.
First published online 27 April 2006.
doi:10.1111/j.1574-6941.2006.00123.x
Editor: Gary King
Keywords
dsr AB; sulfate reducing bacteria; sulfate
reduction; mobile sediments.
Abstract
In spite of the nonsulfidic conditions and abundant reactive iron(III) commonly
found in mobile tropical deltaic muds, genes encoding dissimilatory sulfite
reductase (dsr) were successfully amplified from the upper 1 m of coastal
deposits sampled along French Guiana and in the Gulf of Papua. The dsr sequences
retrieved were highly diverse, were generally represented in both study regions and
fell into six large phylogenetic groupings: Deltaproteobacteria, Thermodesulfovibrio
groups, Firmicutes and three groups without known cultured representatives. The
spatial and temporal distribution of dsr sequences strongly supports the contention that the sulfate-reducing prokaryote communities in mobile mud environments are cosmopolitan and stable over a period of years. The decrease in the
35
SO2
4 tracer demonstrates that, despite abundant reactive sedimentary iron(III)
(350–400 mmol g1), the sulfate-reducing prokaryotes present are active, with the
highest levels of sulfide being generated in the upper zones of the cores (0–30 cm).
Both the time course of the 35S-sulfide tracer activity and the lack of reduced sulfur
in sediments demonstrate virtually complete anaerobic loss of solid phase sulfides.
We propose a pathway of organic matter oxidation involving at least 5–25% of the
remineralized carbon, wherein sulfide produced by sulfate-reducing prokaryotes is
cyclically oxidized biotically or abiotically by metal oxides.
Introduction
The use of molecular tools in the study of microbial communities has revealed that the massive fluidized mud deposits
that occur along thousands of kilometers of tropical continental margins downdrift of major rivers, such as the
Amazon, are amongst the most phylogenetically diverse,
microbially dominated environments in the world (Kemp &
Aller, 2004). The relative roles and interactions between the
microbial groups mediating the high carbon remineralization
rates observed within these nonsulfidic deposits remain
unknown. Although diverse microbial communities are identifiable in the 16S rDNA gene libraries, one limitation of the
rRNA-based analysis used in library construction is that it
does not necessarily provide a direct link to physiological
capabilities, and metabolic features must be inferred from
genetically close relatives, which have been cultured. Sulfatereducing prokaryotes (SRPs) typically have a central role in
carbon mineralization and sulfur cycling in anoxic sedimentary environments (i.e. Jrgensen, 1982; Jrgensen & Bak,
1991). Apparent SRPs are also significant components of the
rDNA libraries obtained from tropical mobile muds, yet
evidence of sulfate reduction is not obvious in these deposits
FEMS Microbiol Ecol 57 (2006) 169–181
based on general geochemical properties, such as SO2
4
depletion or net HS production (Todorov et al., 2000;
Madrid et al., 2001). In the present study, we further evaluate
the diversity of SRPs by examining the diversity of the
dissimilatory sulfite reductase (dsr) gene necessary for respiratory sulfate reduction. Direct measurements of sulfate
reduction using 35SO2
4 carried out in the present study imply
an active underlying role for sulfate reduction in the remineralization of carbon in nonsulfidic, suboxic mud. We propose
a coupled reduction–oxidation pathway, dependent on sedimentary dynamics and metal oxide availability, through
which sulfate-reducing activity is in immediate counterbalance with abiotic and microbially mediated sulfide oxidation.
Two enzymes, adenosinemonophosphate sulfate (APS)
reductase (Friedrich, 2002) and dissimilatory sulfite reductase
(DSR) (Klein et al., 2001), mediate dissimilative sulfate
reduction. APS reductase reduces APS to sulfite and DSR
catalyzes the six-electron reduction of sulfite to sulfide and is
required by all SRPs. Both genes have been targeted for
studies of the diversity of SRPs (e.g. Klein et al., 2001;
Friedrich, 2002). However, the interpretation of phylogenetic
data generated using genes for these two enzymes is complicated as a result of numerous lateral gene transfer events.
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170
Klein et al. (2001) have investigated and identified the likely
transfers of the dsr gene amongst different lineages of bacteria
and archaea and, therefore, genes for DSR are most commonly used to describe the phylogenetic diversity of SRPs
(Minz et al., 1999; Chang et al., 2001; Joulian et al., 2001;
Castro et al., 2002; Dhillon et al., 2003; Fishbain et al., 2003;
Fukuba et al., 2003; Liu et al., 2003; Nakagawa et al., 2004a, b).
Tropical mobile deltaic deposits, such as those studied
here, differ from typical marine sulfidogenic sediments in
that they often exhibit no net sulfide production over
extensive depth intervals (Alongi, 1995; Aller, 1998;
Aller et al., 2004a, b). Although oxygen penetrates only a
few millimeters below the sediment surface, physical mixing
as a result of tidal currents, seasonal winds and local
geomorphologic features periodically re-oxidizes the sediments to a depth of 470 cm (Aller, 1998; Aller et al.,
2004a, b). During remobilization, large amounts of highly
weathered and reactive oxidized debris are entrained and
redox conditions reset. Because of the relative abundance of
oxidants, this re-oxidized layer passes through oxic and NO
3
reducing conditions within a day, followed by manganese
reduction stages for several weeks, and then remains in a
suboxic iron reduction stage for up to a year before becoming sulfidic (Aller et al., 2004b). Thus, despite high remineralization rates, repetitive remobilization and re-exposure
of thick sediment zones over annual timescales apparently
promote virtually continuous suboxic iron/manganese
cycling instead of sulfate reduction (Aller, 2004; Aller
et al., 2004b).
Materials and methods
Study sites
Sediments were collected from six different sampling stations: four off the Atlantic coast of French Guiana near
Sinnamary [sampling stations: KS98-3, 5123.108 0 N,
52150.935 0 W; KS98-9, 5124.204 0 N, 52150.912 0 W (in 1998);
KS01-1, 5123.615 0 N, 52155.368 0 W; KS01-4, 5128.63725 0 N,
51257.90385 0 W (in 2001)] and two off the Gulf of Papua,
Papua New Guinea [G19, 710.983 0 S, 14410.636 0 E; GS48,
810.037 0 S, 14410.787 0 E (in 2000)]. The French Guiana
stations were located in migrating mudwave deposits derived from the upstream Amazon River delta. These deposits
move north-westward at approximately 1–3 km per year
(Froidefond et al., 1988; Eisma & Vanderma, 1991). At the
time of sampling, site KS98-3 was on the inshore trailing
edge, KS98-9 was on the offshore trailing edge and KS01-4
was on the offshore leading edge of the mudwave. The Gulf
of Papua sites represent topset (G19) and upper foreset
(GS48) regions of a prograding clinoform delta system
formed from multiple river sources along the south coast of
Papua New Guinea (Walsh et al., 2004).
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V.M. Madrid et al.
Sampling
Samples off the French Guiana coast were obtained from an
individual mudwave at locations in approximately 1 m
water depth. Push cores were obtained by diving using
15-cm-diameter cellulose acetate butyrate tubing capable of
retrieving approximately 1 m cores. The lengths of the cores
from the KS98-3, KS98-9, KS01-1 and KS01-4 stations were
90, 90, 90 and 100 cm, respectively. In the laboratory, the
cores were subcored into 10 cm sections, which were stored
anoxically until processed. In the Gulf of Papua, cores were
obtained using a stainless steel kasten coring unit (Kuehl
et al., 1985, modified by Brunskill et al. (2002) able to
retrieve cores with a length of approximately 2.3 m. The
cores were sectioned onboard into 10 cm subcores, which
were stored until processed in sealed metallized plastic
pouches containing oxygen scrubbers (Walton Feed Inc.,
Montpelier, ID).
Sulfate reduction kinetics
Samples for sulfate reduction kinetic experiments were
obtained over 10 cm depth intervals in sediments from the
stations KS01-1 and KS01-4, French Guiana coast, and the
station GS48, the Gulf of Papua. Incubations with 35SO2
4
were carried out using a modification of the method
employed by Ulrich et al. (1997). Briefly, under a continuous
nitrogen stream, subcore samples from 10 cm depth intervals were collected in cut-off syringes in triplicate and
injected with carrier-free 35SO2
4 (25 mCi). The subcoring
procedure precluded the introduction of oxygen during
handling. Samples were incubated for 0–40 min in the dark
under anoxic conditions at room temperature. Spiking and
incubations were carried out in an anaerobic hood with a
slow flow of nitrogen in order to create a positive pressure
and, therefore, to exclude the possibility of oxygen introduction. At the end of each incubation period, samples were
fixed with 10 mL of 20% zinc acetate and frozen at
20 1C until ready for extraction. In order to measure the
quantities of 35S0 formed, 1 g of the zinc acetate-fixed
subsamples was extracted with organic solvents (cyclohexane, methanol) (Zopfi et al., 2004), mixed with 10 mL of
scintillation cocktail (Ultima Gold, Packard, CA) and
counted in a scintillation counter (LSC 1600TR, Canberra
Packard). Measurements of chromium-reducible sulfur
P
H2S) were made on 2 g of zinc
(CRS = FeS2, FeS,
acetate-fixed subsamples distributed in 125 mL nitrogenpurged airtight bottles containing a trap with 2 mL of 10%
zinc acetate (Fossing et al., 1989). After the addition of 8 mL
of 1 M CrCl2 in 0.5 M HCl solution, followed by 8 mL of
anoxic 12 M HCl, bottles containing the sediment slurry
were incubated at room temperature with agitation for
48 h. The trap contents were mixed with 10 mL of scintillation cocktail (Ultima Gold, Packard) and counted in a
FEMS Microbiol Ecol 57 (2006) 169–181
171
Diversity of sulfate reducers in nonsulfidic mobile sediments
scintillation counter (LSC 1600TR, Canberra Packard).
Counts were made for a sufficient period to obtain counting
uncertainties of less than 2%. Samples from station KS01-1
were also subjected to a two-step distillation procedure in
which acid-volatile sulfide (AVS, cold 1 M HCl) was distilled
in an initial step and CRS was determined on the residual
material in a second step.
Construction of dsr libraries
Ten centimeter sections from each core were used to isolate
genomic DNA. Total genomic DNA from 1 g of sediment for
surface samples and up to 10 g of sediment for sample
depths greater than 60 cm was extracted using an UltracleanTM Soil DNA Extraction Kit (MoBio Inc., CA) according to the manufacturer’s instructions. For each core, dsr
gene libraries were created from samples taken from the
10–30 and 460 cm depth intervals from each station. A
1.9 kb DNA fragment encoding most of the b and a subunits
of the DSR could be amplified by PCR from all recognized
lineages of sulfate-reducing prokaryotes with a single primer
set dsr1F and dsr4R, as described previously by Klein et al.
(2001). To minimize PCR artifacts, PCR amplification
conditions were optimized using genomic DNA of Desulfovibrio vulgaris ATCC29579, as suggested by Qiu et al. (2001).
The reaction mixture for PCR amplification (25 mL) contained a Ready Taq mix (Amersham, NJ), 21 mL of nucleasefree H2O, 1 mL of each primer (2.5 mM) and 50 ng mL1 of
DNA. PCR amplification was carried out in a Thermocycler
model iCycler (BioRad, CA) with the following conditions:
initial denaturation at 94 1C for 1 min, followed by 25 cycles
at 94 1C for 1 min, 60 1C for 1 min and 72 1C for 1 min. A
final extension step of 72 1C for 15 min was also used. The
1.9 kb PCR products were purified with the Wizard PCR
Clean-Up System (Promega, WI) in accordance with the
manufacturer’s instructions. Triplicate PCR mixtures for
each site were combined, ligated into the vector pGEM-Teasy (Promega) and transformed into commercially available competent cells of Escherichia coli strain JM109 according to the manufacturer’s instructions (Promega). Using
blue–white colony screening, the plasmids of selected colonies were screened with EcoRI for the detection of a 1.9 kb
insert. Restriction fragment length polymorphism (RFLP)
analysis of isolated plasmids was carried out by HinfI, HhaI,
DdeI and HaeIII (New England Biolabs, Beverly, MA)
digestions. DNA sequencing of selected clones representing
an individual banding pattern was performed with an ABI
PRISM BigDye Terminator Cycle Sequencing Ready Reaction Kit (Applied Biosystems, CA) and an ABI PRISM 3700
DNA analyzer at the SUNY DNA Sequencing Facility. Raw
sequencing data were edited, analyzed and translated using
the Sequencher program, version 4.0 (Gene Codes Corp.,
MI). Nucleotide sequences were aligned with relevant seFEMS Microbiol Ecol 57 (2006) 169–181
quences and sequences of closest relatives from the GenBank
database in CLUSTAL X (Thompson et al., 1997) and MEGALIGN
(DNAStar Corp., MI). Deduced amino acid sequences were
realigned using CLUSTAL X and MEGALIGN. Unambiguously
aligned nucleic acid and protein sequence regions were
employed to construct bootstrap-supported (100 resamplings) neighbor-joining phylogenies using PHYLIP (Felsenstein, 1995). A phylogenetic tree was constructed based on
the deduced amino acid sequence alignment of retrieved
sequences and those from the database using the neighborjoining algorithm by PHYLIP software. Statistical comparisons
of the clone sequence libraries for each of the sites
Rwere carried out with LIBSHUFF (Singleton et al., 2001) and
-LIBSHUFF (Schloss et al., 2004). Rarefaction analysis and
SRB community richness predictions were carried out
according to (Kemp & Aller, 2004) and (Martin, 2002). dsrB
sequences were deposited into GenBank with the accession
numbers AY753074–AY53142.
Results
dsrB gene diversity
Genes encoding DSR were successfully amplified from
sediments collected from all stations (data not shown). The
depth spans from which dsr was amplified were variable for
different cores. For the French Guiana KS98-3 and KS01-4
station cores, the dsr gene was amplifiable throughout the
whole length of the cores. For the KS98-9 station core, no dsr
sequences were detected deeper than 70 cm. For the Gulf of
Papua G19 and GS48 stations, the dsr gene could not be
amplified from depths greater than 80 and 160 cm, respectively. For each station, we chose a surface (10–30 cm) and a
deepest possible sample for the creation of dsr libraries (see
depth in Table 1).
Usually, two close bands of approximately 1.9 kb were
visualized in agarose gels after PCR amplification with dsr
primers (data not shown). The two amplicons were approximately the size generated in control amplifications of the dsr
gene of Desulfovibrio vulgaris. Separate cloning and sequencing of DNA from the two bands proved that they were
amplification products of dsr genes. Ten libraries were
generated, of which 172 clones representing an individual
HinfI/HhaI, DdeI/HaeIII pattern were analyzed by sequencing. The a and b subunits of recovered dsr gene fragments
were partially sequenced (representative clone names listed
in Table 1) and, on average, 650 nucleotides were determined for each subunit. Potentially chimeric artifacts and
non-dsr sequences were identified in some of the clone
libraries and were excluded from further analysis. Unambiguously aligned nucleic acid and amino acid data sets
for both subunits were estimated using distance matrix and
maximum-likelihood criteria to build phylogenetic dsr
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2006 Federation of European Microbiological Societies
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fg2d267, ng1d4, ng2d52, ng7d1022
fg1d1220, fg2d271, fg3d1056,
fg4d770, fg7d931, fg7d951,
fg8d1100, fg8d1113, fg11d1186,
ng2d836, ng3d853, ng3d996,
ng3d1004, ng3d1010, ng6d899,
ng7d1035
fg3d1061, fg3d1064, fg5d1271,
fg7d958, fg10d829, ng2d984,
ng7d1139, ng7d1027
ng2d87, ng3d1013, ng7d1034
ng3d856, ng3d1014
fg1d915, fg5d1087, fg8d1106,
fg11d960
fg3d1062, fg3d1048, fg7d920
fg2d283, fg3d1060, fg5d1145,
fg8d1104, fg11d963, ng1d16,
ng1d111, ng1d124, ng2d51,
ng2d991, ng3d851, ng3d1005,
ng4d870, ng4d868, ng7d1025
fg1d919, fg3d1044, ng2d64, ng2d70
fg3d1041, ng2d94, ng7d1033,
ng6d894
fg7d927, fg7d928, fg11d1196,
ng4d879, ng7d1029, ng7d1037
Representative clones
2
2
9
2
10–20 cm
4
1
5
8
80–90 cm
DSR group designation according to Thomsen et al. (2001) and Dhillon et al. (2003).
Thermodesulfovibrio
Group
Group V
Firmicutes
Group II
Group IV
Clade D
Clade E
Clade F
Clade C
Deltaproteobacteria
Clade A
Clade B (includes
Group I)
DSR affiliation
2
2
10
10–20 cm
2
2
9
2
2
60–70 cm
2
3
2
1
5
10–30 cm
1
1
3
7
3
2
90–100 cm
10
1
10–20 cm
2
1
2
2
2
1
5
70–80 cm
G19
KS01-4
KS98-3
KS98-9
Gulf of Papua
French Guiana
Table 1. Distribution and phylogenetic affiliation of characterized representative dsrB clones by sampling site, station and depth interval (cm)
3
2
15
7
10–20 cm
GS48
5
2
3
5
5
1
4
150–160 cm
172
V.M. Madrid et al.
FEMS Microbiol Ecol 57 (2006) 169–181
173
Diversity of sulfate reducers in nonsulfidic mobile sediments
gene-based trees. The two methods resulted in congruent
tree topologies for both dsrA/DsrA- and dsrB/DsrB-based
trees. Phylogenetic trees based on protein (DsrA and DsrB)
and nucleotide (dsrA and dsrB) sequences were also nearly
identical. These data are consistent with the observations of
Chang et al. (2001) that phylogenetic trees constructed with
different portions of the dsr genes reveal, in general,
consistent topologies for both a and b subunits of dsr.
Interestingly, Liu et al. (2003) noted that dsr/Dsr trees based
on combined DsrAB or DsrB sequences are slightly better
than those based on DsrA sequences in low-order branch
resolution. Therefore, a consensus distance matrix tree
based on amino acid sequences for the b subunit (i.e. DsrB),
generated by PHYLIP, is shown in Fig. 1.
The dsr sequences retrieved from mobile muds fall into
six large phylogenetic groups (Fig. 1). Based on data on
horizontal dsr transfer events by Klein et al. (2001), as well as
dsr trees published by other researchers (Thomsen et al.,
2001; Dhillon et al., 2003), we tentatively assign these groups
to Deltaproteobacteria, Thermodesulfovibrio groups, Firmicutes and three other groups, which currently lack cultured
representatives. We followed the nomenclature of Thomsen
et al. (2001) and Dhillon et al. (2003) for the groups without
cultured representatives: Groups I, II and III were first
described by Thomsen et al. (2001). Later, Dhillon et al.
(2003) showed that Group II sequences fall into at least two
distinct groups, Group II and Group IV. One group of DSR
sequences with no cultured representatives, which have not
been found in either Aarhus Bay or Guaymas Basin, was
designated Group V (see Fig. 1 and Table 2). Generally, each
of the large groups was represented at both locations (i.e.
French Guiana coast and the Gulf of Papua). Several smaller
rank clades were present only at one location: clades D and E
(Deltaproteobacteria) are composed solely of Gulf of Papua
sequences and clade F (Deltaproteobacteria) and Group II
are composed solely of French Guiana sequences. The most
diverse large group, comprising 78.7% of all sequences for
French Guiana and 64.1% for the Gulf of Papua, had
sequences closely related to the dsrAB genes of Deltaproteobacteria (Table 1). This group consists of the sequences
grouped into clades designated A, B, C, D, E and F in Fig. 1.
Clade A comprises seven representative sequences only distantly affiliated with incomplete oxidizers such as Desulfovibrio vulgaris. Sequences assigned to clades C, D and E are
related to sequences of both incomplete and complete oxidizers, such as Desulfobacter vibrioformis, Desulfacinum infernum and Desulfofustis glycolicus, respectively. Sequences
belonging to clade B are not affiliated with any dsr sequence
from cultured microorganisms, with the exception of
fg8d1113, which appears to be distantly related to Desulfonema limicola. Most of the sequences from clade B (with the
exception of fg8d1113 and ng3d853) apparently belong to
Group I according to Thomsen et al. (2001): they grouped
FEMS Microbiol Ecol 57 (2006) 169–181
together with sequences from the environmental bacterium
clones a-75 and a-G retrieved from the top 20 cm from Aarhus
Bay, Denmark sediments (Thomsen et al., 2001; Dhillon et al.,
2003). Clade E is affiliated with an environmental clone
B04P021 from Guaymas Basin (Dhillon et al., 2003).
The second most abundant and diverse group of sequences (9.5% from French Guiana and 23% from the Gulf
of Papua) was related and tentatively assigned to Group IV,
one of the major groups described by Dhillon et al. (2003)
for the Guaymas Basin. Amongst their closest relatives are
the environmental clones, uncultured bacterium KFY_322
(Moeslund et al., 1994), environmental clone from Guaymas
B04P004 (Dhillon et al., 2003) and uncultured bacterium
from Solar Lake Mat clone 917 (Minz et al., 1999).
The Thermodesulfovibrio group, Firmicutes, Groups II and
V account for a total of 6.4%, 2.3%, 2.3% and 2.9% of all
sequences, respectively. Group II sequences were exclusively
found in French Guiana sediments, whereas the three other
groups were represented at both sampling locations. Lacking
close cultured relatives, sequences comprising Group V were
related to sequences of environmental unclassified bacteria
from groundwater at a uranium mill tailing site (Chang
et al., 2001). Interestingly, no representatives belonging to
the so-called ‘deep branching’ group or Group III according
to Thomsen et al. (2001) were found in mobile sediments.
Thus, molecular analysis of dsr sequences amplified from
mobile mud samples from two different environments
indicates that SRP communities are not only present with
representatives from all major branches of SRP, but that they
are quite diverse.
Sulfide production kinetics
In order to elucidate whether SRP communities are actively
involved in sulfur cycle reactions in suboxic mobile muds,
we spiked sediments with 35SO2
4 . Following the introduc35
S-sulfide was
tion of 35SO2
4 , the initial production of
observed for samples from all sites (Fig. 2). In contrast with
classic sulfidic sediments (Jrgensen, 1982), however, the
time course of sulfide tracer production in the incubations
often showed major changes in the net rate of sulfide
production with time, consistent with simultaneous 35SO2
4
reduction and loss of sulfide by either biogenic or abiogenic
oxidation reactions (Moeslund et al., 1994; Fossing, 1995).
Because of the time-dependent sulfide production – loss
patterns, only short-term incubations (40 min) were considered in the estimation of 35SO2
4 reduction rates (Moeslund et al., 1994; Fossing, 1995). Unlike previous studies, in
which initial gross rates of 35SO2
4 reduction were followed
by lower but still positive net rates (Moeslund et al., 1994;
Fossing, 1995), longer incubations in many of the depth
intervals from the upper 50–70 cm at the French Guiana
sites did not result in progressive net production of 35S2006 Federation of European Microbiological Societies
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174
V.M. Madrid et al.
Fig. 1. Consensus neighbor-joining dendogram depicting the relationships between DsrB protein sequences. Numbers at the nodes represent the
percentage of bootstrap re-sampling based on 100 replicates; only values of more than 50 are presented. The clones from this study are in bold, named
according to the origin and depth interval.
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FEMS Microbiol Ecol 57 (2006) 169–181
175
1
1
1
1
1
1
1
1
1
1
1
FEMS Microbiol Ecol 57 (2006) 169–181
1, sequence type was detected; , sequence type was not detected.
DSR group designation according to Thomsen et al. (2001) and Dhillon et al. (2003).
Guaymas Basin libraries contain two additional small groups of sequences, which are not included in this table.
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
Deltaproteobacteria 1
(including Group I)
Group II
1
Group III
1
Group IV
Group V
Firmicutes
Thermodesulfovibrio group
DSR affiliation
KS98-3
KS98-9
KS01-4
G19
GS48
Groundwater from
Aarhus Bay a uranium mill
Guaymas Basin 10–20 cm 80–90 cm 10–20 cm 60–70 cm 10–30 cm 90–100 cm 10–20 cm 70–80 cm 10–20 cm 150–160 cm
Gulf of Papua
French Guiana
Table 2. Major dissimilatory sulfite reductase (DSR) lineages detected in Aarhus Bay sediments (Thomsen et al., 2001), Guaymas Basin sediments (Dhillon et al., 2003), groundwater from a uranium mill
tailing site (Chang et al., 2001) and French Guiana and Gulf of Papua sediments
Diversity of sulfate reducers in nonsulfidic mobile sediments
sulfide. Instead, the CRS initially produced showed a welldefined increase and then decrease (Fig. 3; KS01-1; KS01-4),
consistent with the lack of any net sulfide production in
these sediments. The Gulf of Papua 10–20 cm sample is
more typical of tracer patterns reported for deposits in
which both reduction and oxidation occur: sulfide production continuously increased after 40 min of incubation but
at a rate slower than that initially observed.
No accumulation of 35S0 was detected in organic solvent
tracerextractions (cyclohexane, methanol) of 35SO2
4
amended sediment, even with extended periods of incubation (up to 48 h). Although dissolved sulfide was not
separately measured during the tracer incubations, in previous long-term incubation experiments and analyses of in
situ pore water samples, dissolved sulfide was not detectable
at any time (o2 mM) (Aller et al., 2004b). A subset of
samples from French Guiana station KS01-1 was processed
through both a two-step AVS (1 M HCl) and CRS distillation and a single-step CRS distillation. No statistically
significant differences between the AVS and single-step CRS
pools were found. The lack of dissolved sulfide, the lack of
elemental sulfur and the correspondence between the twoand single-step CRS distillations imply that solid phase AVS
was the primary initially reduced sulfur product.
There was no correlation between sulfide production
patterns and pore water sulfate availability. Sulfate concentrations showed slight variations with depth in all three
cores but remained relatively high throughout. For example,
in core KS01-4, the surface concentration of sulfate was
18.1 mM, and that in the bottom depth interval was
15.9 mM, with a broad maximum of 22.3 mM at 20–40 cm.
The lowest measured sulfate concentration was 6.6 mM,
detected in core KS01-1 at a depth of 90–95 cm. The
maximum sulfate concentration of 15.9 mM in this core
was found at the surface.
Because of the rapid loss of 35S from the CRS pool, in
most cases, the tracer measurements were insufficient to
accurately resolve gross reduction rates during the first few
minutes of incubation. Moreover, the time courses often
differed in functional form within the different intervals and
stations, and it was not reasonable to treat them analytically
in exactly the same way. Rough estimates of initial sulfide
production rates were made as follows. In cases in which
there was a distinct maximum in 35S CRS count rates, a
second-order polynomial fit (parabolic) was made to the
first three to four time points (0–25 min), and the initial rate
was calculated from the slope extrapolated to time zero. In
all other cases, the initial linear rate was calculated using
time points up to the position at which there was a clear
decrease in slope (Fossing, 1995). In several incubations,
there was no obvious change in slope, and all points up to
1.5 h were used (GS48; Fig. 2). These calculations demonstrate that rates in the range 10–20 nmol cm3 day1 were
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176
V.M. Madrid et al.
French Guiana
station KS01-1
Sulfate reduction rates
Depth (cm)
0
0-10
10
10-20
30-40
40-50
60-70
80-90
200
150
20
30
40
50
Depth (cm)
CRS counts (average)
250
100
50
60
70
80
90
100
110
0
0.0
0.1
0.2
0.3
0.4
0.5
0.6
0.7
0.8
Incubation time (hours)
130
140
French Guiana
station KS01-4
250
Depth (cm)
0-10
CRS counts (average)
150
150
160
0
10-20
30-40
40-50
60-70
80-90
200
2
4
6
8
10 12 14 16 18 20 22 24 26 28
Sulfate reduction rates (nmol SO42– cm–3 d–1)
Fig. 3. Initial rates of sulfate reduction as a function of depth in
sediments at stations KS01-1 and KS01-4 (French Guiana) and GS48
(Gulf of Papua).
100
50
0
0.0
0.1
0.2
0.3
0.4
0.5
0.6
0.7
0.8
Incubation time (hours)
Gulf of Papua
station GS48
350
300
CRS counts (average)
Station KS01-1, French Guiana
Station KS01-4, French Guiana
Station GS48, Gulf of Papua
120
Depth (cm)
0-10
10-20
20-30
30-40
60-70
70-80
150-160
180-190
230-240
250
200
150
100
carbon cm3 day1) (Aller et al., 2004b). The apparent lack
of detectible sulfide production below 70–90 cm of the cores
may be explained by depletion of reactive organic carbon
reductant at depth and insufficient incubation times (o1 h)
to detect low SO2
4 reduction rates.
Our observations are consistent with earlier measurements of sulfate reduction by Alongi (1995) for Gulf of
Papua sediments, which showed measurable rates of sulfate
reduction to depths of 20 cm. Thus, direct measurements of
sulfide production show that sulfate reduction occurs in
mobile tropical sediments from both the French Guiana
coast and the Gulf of Papua, although the sediment remains
nonsulfidic (Figs 2 and 3).
Discussion
50
0
0.0
0.2
0.4
0.6
0.8
1.0
1.2
1.4
1.6
Incubation time (hours)
Fig. 2. 35SO2
4 radiotracer experiments for sediments collected at the
KS01-1 and KS01-4 (French Guiana) and GS48 (Gulf of Papua) stations.
Rapid sulfate reduction and concomitant disappearance of CRS are
consistent with fast re-oxidation of reduced sulfur species.
typical of the upper 0–30 cm (Fig. 3). Assuming a 2 : 1
stoichiometry of carbon remineralized to SO2
4 reduced, the
reduction
measured
in
this
way
could
account for
total SO2
4
roughly 5–25% of the total CO2 production commonly
measured in the surface mobile zone (200–600 nmol
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The French Guiana mobile belt system is characterized by
massive migrating mud banks which extend along the
north-east coast of South America for more than 1600 km
downdrift of the Amazon River (Faas, 1986; Froidefond
et al., 1988; Eisma & Vanderma, 1991; Allison et al., 1996,
2000; Kineke et al., 1996). The Gulf of Papua, located on the
south coast of Papua New Guinea, consists of a half-moonshaped shelf area of more than 50 000 km2, with a radius of
200 km, into which numerous rivers, the Fly River being the
largest, drain the highlands of New Guinea and transport
huge quantities of highly weathered material (Walsh et al.,
2004; Aller et al., 2004a). During energetic trade-wind
conditions, the fine-grained river-borne sediments containing terrestrial/mangrove-derived organic debris are
FEMS Microbiol Ecol 57 (2006) 169–181
177
Diversity of sulfate reducers in nonsulfidic mobile sediments
frequently suspended by tidal currents and waves, and
moved about as fluid mud (near-bed suspensions having
sediment concentrations of 10 g L1 to more than 100 g L1
(Faas, 1986; Kineke et al., 1996) shoreward of the 10 m
isobath in a north-easterly direction (Ogston et al., 2003).
During periods of the north-west monsoonal winds, the
fluid mud is temporarily deposited within the topset zone
(20 m shoreward) and stabilized for short (days to weeks)
or long (weeks to months) periods of time (Harris et al.,
1993; Wolanski & Alongi, 1995; Crockett et al., 2003).
Periodically, and presumably largely during the south-east
trade-wind period, these muds are re-fluidized, mobilized
and carried into the foreset region (40–60 m offshore),
where they accumulate (Walsh et al., 2004; Aller et al.,
2004a).
Biogeochemical cycling is closely coupled to sedimentary
dynamics and the refluxing of reduced components between
the seabed and well-oxygenated overlying water in mobile
mud belt systems (Aller, 1998, 2004). The unique biogeochemical features of these systems, which promote the
remineralization of tremendous quantities of organic carbon,
have been discussed in detail (Alongi, 1995; Aller, 1998; Aller
& Blair, 2004; Aller et al., 2004a, b). Although a generally
accepted model for early diagenetic reactions in metal oxiderich mobile sediments involves oxidation of organic matter
directly coupled with manganese(IV) and iron(III) reduction
(Fig. 4, panel A), an underlying interaction with reactions of
the sulfur cycle has been hinted at by 16S rDNA gene libraries
(Todorov et al., 2000; Madrid et al., 2001). These libraries
indicate that bacteria involved in both sulfate reduction and
sulfide oxidation are present in spite of the absence of
geochemical evidence of net sulfide production.
The amplification of the gene for DSR, the key enzyme for
the dissimilatory sulfate reduction pathway, is further evidence for the active participation of sulfur cycle reactions
during organic carbon remineralization in iron(III)-rich
coastal deposits (Fig. 4, panel B). The major, if not only,
large-scale process of sulfide production in nature is
mediated by SRP. The phylogenetic diversity of the dsr gene
in prokaryotic communities from French Guiana and the
Gulf of Papua mobile sediments was high, with dsr genes
from the members of all major groups of SRP recovered
from at least one mobile mud station. The fact that the
overall phylogenetic diversity of dsr genes increased with
depth at both sampling sites, with the greatest diversity
found in the deepest portion of the core from station GS48
(48 m water depth) from the Gulf of Papua, presumably
reflects a dampening of physical disturbance away from the
sediment–water interface, and increasingly favorable conditions for SO2
4 reducers. In addition, SRP communities in
mobile sediments appear to remain relatively stable over
time: a number of closely related and identical DSR
sequences were retrieved from samples collected in French
FEMS Microbiol Ecol 57 (2006) 169–181
(a)
FeIII(MnO2)
O2
Corg
FeII(MnII)
CO2
(b)
O2
FeII(MnII)
SO42–
Corg
HS–
CO2
S0
FeIII(MnO2)
Fig. 4. Intercalation of sulfur cycle with the manganese/iron and carbon
cycles as an intermediate oxidant/reductant. (a) Direct coupling of metal
cycles with oxygen and organic carbon (Corg) cycles. (b) Indirect coupling
of metals through sulfur reduction/oxidation with oxygen and Corg,
producing the same net reaction as in (a). Note that the reactions
coupling the sulfur and carbon cycles are reversible. For simplicity of
representation, the manganese and iron cycles are shown combined,
although, in reality, their cycles are not identical. Modified after Aller
(1994) and Burdige (1993).
Guiana 3 years apart (i.e. 1998 and 2001), and the overall
composition of SRP in these two temporally separated
sample sets appeared to be similar.
R
Statistical analyses of dsr libraries using the -LIBSHUFF
program (Schloss et al., 2004) suggest that libraries created
from the same core or within the same geographic location
(i.e. French Guiana coast and the Gulf of Papua) are
subsamples of each other (data not shown) and indicate a
high similarity of SRP communities. At least superficially,
SRP communities found in geographically distinct mobile
mud environments appear to have substantial overlap (i.e.
clades A and C and Group IV). However, some groups are
unique for each of the two environments, such as Clade F
and Group II dsr sequences, which were found only in
French Guinana sediments, and Clade D and E dsr sequences, which were found only in Gulf of Papua sediments.
There were no groups of SRP exclusively associated with the
upper portion of the cores. However, several groups (i.e.
Group II, Firmicutes and Thermodesulfovibrio group) were
exclusively found in deeper portions of the cores. As the dsr
sequences belonging to the thermophilic Thermodesulfovibrio group were recovered exclusively from deeper sections
of the cores, it is most likely that microorganisms belonging
to this group are inactive, because, at depths greater than
60 cm, the temperature is below the optimum (25 1C) and
sulfate reduction activity could not be detected (see below).
The diversity of the dsr genes in tropical mobile sediments
appears to be similar to that of most other sulfidogenic
environments. SRB assemblages from Aarhus Bay, Kysing
Fjord (Thomsen et al., 2001) and continental margin sediments (Liu et al., 2003) harbor two major groups [i.e. clade
B (Group I) and Group II] of the dsr gene observed in
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178
French Guiana and Gulf of Papua sediments (Table 2).
Remarkably, the SRP assemblages from thermophilic
[i.e. Guaymas Basin (Dhillon et al., 2003), hydrothermal
chimney vent structures (Nakagawa et al., 2004a), freshwater
(e.g. Florida Everglades Castro et al., 2002) and ground
waters of a uranium mill tailing site (Chang et al., 2001)]
share many major members of SRP communities (for
example, Group IV, see Table 2). Thus, at least three SRP
groups detected in our tropical mobile muds have been
found in these environments (Table 2). However, the overall
diversity of SRP communities in mobile sediments is still
considerably lower than that of a uranium mill tailing site
(Chang et al., 2001).
The physiological and ecological importance of the diversity of SRP in mobile sediments is not entirely clear;
however, the frequent input of tremendous quantities of
both labile and refractory organic matter in these environments probably leads to the availability of a range of electron
donors for both completely and incompletely oxidizing SRP,
consistent with our data showing that both physiological
groups of SRPs are present.
The major oxidants available to microbes in the mobile
zone (20–100 cm) of French Guiana and Gulf of Papua
deposits are ferric (hydro)oxides and sulfate (Aller et al.,
2004a, b). Oxygen, nitrite and nitrate are not detectable below
a few millimeters. Manganese(III,IV) is present at approximately 5–10 mmol g1 and, in the absence of mixing, is
reduced within a few days to weeks (Aller et al., 2004b).
Empirical models derived from measurements of decomposition pathways in temperate coastal and deep-sea deposits have
shown that, when iron(III) is sufficiently abundant (above
100 mmol g1), the direct coupling of iron(III) reduction with
organic carbon oxidation supports 100% of organic carbon
remineralization (Jensen et al., 2003). At the present study
sites, initial iron(III) exceeds 350–400 mmol g1 (Aller et al.,
2004a, b), and thus, based on existing models of diagenetic
respiration pathways, direct iron(III) reduction may be expected to exclusively dominate in the mobile suboxic zone of
both French Guiana and Gulf of Papua deposits.
In contrast with these expectations, the 35SO2
4 tracer
reduction patterns directly demonstrate that SO2
4 reducers
are metabolically active in the mobile suboxic zone, and the
loss of solid phase CRS following initial formation implies
rapid anaerobic re-oxidation. Therefore, based on the molecular biological dsr data, the 35SO2
4 tracer dynamics and the
general biogeochemical conditions, we propose that a significant proportion of the anaerobic oxidation of organic
matter in the suboxic mobile zone occurs through multiple,
closely coupled Fe–S–C pathways in series or in parallel (Fig.
4). Earlier work (Canfield et al., 1993; Kostka et al., 2002;
Thomsen et al., 2004) has shown that both metal oxide and
sulfate reduction may occur in the same sediment horizon in
lake, salt marsh or marine deposits; however, in these cases, a
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V.M. Madrid et al.
net build-up of solid phase reduced sulfur was observed, and
re-oxidation of sulfur apparently involved dissolved not
solid phase sulfide loss (Canfield et al., 1993; Moeslund
et al., 1994; Fossing, 1995; Kostka et al., 2002; Thomsen
et al., 2004). At least at the French Guiana sites, anaerobic
re-oxidation of sulfide appears to be 100% efficient, and
both solid phase sulfides and iron(III) are probable reactants. The oxidation of solid phase sulfide may also involve
manganese oxide intermediates which are known to completely oxidize iron sulfides (Schippers & Jorgensen, 2002).
In the present case, however, manganese is not sufficiently
abundant to sustain re-oxidation cycles for more than a few
weeks at most, and coupled Mn–S–C or Mn–Fe–S–C pathways cannot explain the lack of reduced sulfur build-up in
natural coastal deposits which can remain stable for months
between sediment reworking events (exposure, re-oxidation) (Aller et al., 2004b).
There may be several pathways leading to the re-oxidation
of sulfide back into sulfate. A portion of any sulfide formed
could be rapidly oxidized by a combination of abiotic
reactions with either manganese or iron oxides to S0 and
the biogenic disproportionation of S0 to SO2
4 and HS
(Thamdrup et al., 1993, 1994). As noted, manganese oxides
can also completely oxidize both dissolved and solid phase
sulfides (Schippers & Jorgensen, 2002), but the abundance
of manganese implies a relatively minor role under the
present circumstances. Previous studies have been unable
to demonstrate complete oxidation of sulfides by iron(III)
(Schippers & Jorgensen, 2002); however, based on the
present data, we hypothesize that a novel group or consortium of sulfide-oxidizing bacteria can directly oxidize
solid phase sulfide, most likely through reaction with
iron(III) species. We did not detect the formation of labeled
S0 in our experiments, which suggests that re-oxidation of
sulfide occurs biotically directly to sulfate. There is virtually
no reduced sulfur build-up in the solid phase of these
deposits, particularly in the French Guiana sites, where total
reduced sulfur is typically 14 7 mmol g1 dry sediment;
nor is there measurable depletion of pore water SO2
4 in the
suboxic zone, implying complete back oxidation under
natural conditions. The overall sulfide oxidation rates in situ
must therefore be of the same order of magnitude or higher
than the apparent sulfate reduction rates in at least
the surface zone muds where remineralization is intense,
with little lag in re-oxidation of the CRS pool (Fig. 2).
Although sulfide can potentially be oxidized faster than
it is produced, it is present in such low initial concentrations that oxidation rates are presumably not saturated and
the maximum sulfide-oxidizing potential may not be realized. As sulfide begins to accumulate in sediments, its
oxidation rate may proportionally increase and eventually
surpass the rate of sulfate reduction in sediments in which
no net CRS is formed.
FEMS Microbiol Ecol 57 (2006) 169–181
179
Diversity of sulfate reducers in nonsulfidic mobile sediments
Conclusions
A diverse set of dsr genes can be amplified from nonsulfidic,
suboxic mobile muds off coastal French Guiana and in the
Gulf of Papua, implying active sulfate reduction in these
deposits.
The SRP assemblages in French Guiana and Papua New
Guinea sediments appear to be similar in composition, with
representatives of the five major SRB groups based on dsr
gene phylogenies, and compare well with assemblage patterns reported from highly sulfidic environments.
Rapid reduction of introduced 35SO2
4 and subsequent
loss from the chromium reducible product pool in sediments, particularly from the upper 30 cm, indicate that
closely coupled sulfur cycle reactions occur in mobile suboxic sediments in spite of the lack of net sulfide production
and accumulation.
We hypothesize that sulfate-reducing activity is intimately coupled with abiotic and microbially mediated solid
phase sulfide oxidation, which is complete, 100% efficient
and utilizes iron oxides as oxidant. Our reduction rate
estimates (Fig. 3) indicate that the minimum relative contribution of the coupled Fe–S pathway (Fig. 4) to the total
oxidation of organic matter is 5–25%. Further work is
required to measure organic carbon and sulfide and elemental sulfur oxidation rates directly, and to compare these
rates with the expression levels of genes involved in sulfide
oxidation and sulfate and metal oxide reduction.
Acknowledgements
We would like to express gratitude to two anonymous
reviewers whose comments were very helpful in the preparation of this manuscript. The valuable advice by T.
Ferdelman on the technique for extraction with organic
solvents is greatly appreciated. We would like to thank
Frederic Baltzer and Mead Allison for field support in
French Guiana; ENGREF (Ecole Nationale du Génie Rural
des Eaux et des Forêts) and Hydreco, Laboratoire Environnement (Petit Saut) kindly provided laboratory facilities and
aided logistics. The expertise and efforts of the crew of the
RV/Cape Ferguson from the Australian Institute of Marine
Sciences were critical to the success of sample collection in
the Gulf of Papua, as was the assistance of Gregg Brunskill,
Irena Zagorskis, Paul Dixon and John Pfitzner. This research
was supported by NSF Grants OCE 9818574 and OCE0219919 to R.C.A. and J.Y.A., and the Louisiana Board of
Regents Grant LEQSF (2003-05)-RD-A-31 to A.Y.C.
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