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Applied and Environmental Microbiology, November 1999, p. 5089-5099, Vol. 65, No. 11
0099-2240/99/$04.00+0
Copyright © 1999, American Society for Microbiology. All rights reserved.
Natural Communities of Achromatium
oxaliferum Comprise Genetically, Morphologically, and Ecologically
Distinct Subpopulations
N. D.
Gray,1,2
R.
Howarth,1,2
A.
Rowan,1,2
R. W.
Pickup,3
J. Gwyn
Jones,4 and
I. M.
Head1,2,*
Fossil Fuels and Environmental Geochemistry
Postgraduate Institute (NRG)1 and Centre
for Molecular Ecology,2 University of Newcastle,
Newcastle upon Tyne NE1 7RU, and Institute of Freshwater
Ecology3 and Freshwater Biological
Association,4 Windermere Laboratories, Far
Sawrey, Ambleside, Cumbria LA22 0LP, United Kingdom
Received 23 March 1999/Accepted 21 July 1999
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ABSTRACT |
The diversity and ecology of natural communities of the
uncultivated bacterium Achromatium oxaliferum were studied
by use of culture-independent approaches. 16S rRNA gene sequences were PCR amplified from DNA extracted from highly purified preparations of
cells that were morphologically identified as A. oxaliferum present in freshwater sediments from three locations in northern England (Rydal Water, Jenny Dam, Hell Kettles). Cloning and sequence analysis of the PCR-amplified 16S rRNA genes revealed that multiple related but divergent sequences were routinely obtained from the A. oxaliferum communities present in all the sediments
examined. Whole-cell in situ hybridization with combinations of
fluorescence-labelled oligonucleotide probes revealed that the
divergent sequences recovered from purified A. oxaliferum
cells corresponded to genetically distinct Achromatium
subpopulations. Analysis of the cell size distribution of the
genetically distinct subpopulations demonstrated that each was also
morphologically distinct. Furthermore, there was a high degree of
endemism in the Achromatium sequences recovered from
different sediments; identical sequences were never recovered from
different sampling locations. In addition to ecological differences that were apparent between Achromatium communities from
different freshwater sediments, the distribution of different
subpopulations of Achromatium in relation to sediment redox
profiles indicated that the genetically and morphologically distinct
organisms that coexisted in a single sediment were also ecologically
distinct and were adapted to different redox conditions. This result
suggests that Achromatium populations have undergone
adaptive radiation and that the divergent Achromatium
species occupy different niches in the sediments which they inhabit.
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INTRODUCTION |
Achromatium oxaliferum is
a large sediment-dwelling bacterium found principally in freshwater and
brackish environments (21). Notable for its large size, the
bacterium also precipitates intracellular calcium carbonate, a property
which makes it unique among bacteria.
Since it was first described in 1893 (32), the bacterium has
remained uncultivated; however, aspects of its ecology have been
inferred from its phylogenetic position, morphological characteristics, in situ activity measurements, and distribution in relation to geochemical gradients in sediments (3, 4, 14, 17, 21, 37).
For instance, the presence of sulfur inclusions within the bacterium
has suggested its involvement in the sulfur cycle of sediments
(21). Furthermore, by artificial manipulation of the
magnitude of A. oxaliferum populations in situ, it has been determined that the bacterium is capable of mediating the oxidation of
reduced sulfur species to sulfate (14). This finding
is consistent with phylogenetic studies that placed A. oxaliferum in the 
-subdivision of the class
Proteobacteria, most closely related to a number of
sulfur-oxidizing bacteria (13, 17).
Initial phylogenetic analysis of A. oxaliferum cells
purified from a freshwater sediment identified several distinct but
related 16S rRNA sequences from a population of cells previously
thought to be homogeneous (17). On this basis, it was
postulated that this A. oxaliferum population was
genetically heterogeneous, comprising several related A. oxaliferum-like organisms. More recently, Achromatium sequences have been obtained from cells collected from two freshwater lakes in Germany (13). In this report, only single sequences belonging to the Achromatium clade were recovered. However,
subsequent whole-cell in situ hybridization analysis with multiple
fluorescence-labelled oligonucleotide probes revealed that one of the
communities contained at least three genetically distinct subgroups of
A. oxaliferum-like bacteria, while the second was
genetically homogeneous (13).
In this study, we have explored the genetic diversity of three
Achromatium communities from freshwater sediments in
northern England by cloning and sequencing of 16S rRNA genes PCR
amplified from purified cell preparations. Whole-cell in situ
hybridization with fluorescence-labelled oligonucleotide probes
targeting sequences recovered from 16S rRNA gene clone libraries was
used to determine the structure and distribution of the
Achromatium subpopulations in relation to different sediment
redox zones.
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MATERIALS AND METHODS |
Sample sites and purification of cells from sediment.
Three
freshwater environments in northern England that contained populations
of Achromatium were studied: Jenny Dam, a shallow upland
tarn located near Windermere in the English Lake District, Cumbria,
United Kingdom (54°21'N, 2°51'W); a wetland area on the margins of
Rydal Water, Cumbria, United Kingdom (54°27'N, 3°00'W); and Hell
Kettles (38), located to the south of Darlington, County Durham, United Kingdom (54°29'N, 1°33'W). Hell Kettles, formed by
the dissolution and collapse of underlying limestone strata in the year
1179, comprise two ponds (Croft Kettle and Double Kettle) situated in
agricultural fenland (38). Samples were taken from the
margins of the more southerly Croft Kettle.
Sediment samples containing Achromatium cells were obtained
from all three sites by removing the top 3 to 5 cm of the sediment surface with a vacuum sampling device (29) or with sediment coring devices (17). Sediment samples were sequentially
filtered through 100- and 64-µm-mesh-size nylon meshes to remove
larger sediment particles. Crude cell suspensions obtained in this way were purified as described previously (8, 17). Briefly, the screened sediment was placed in a sterile flask. The flask was tilted
until the sediment settled to its base and a white line of
Achromatium cells could be observed just below the meniscus. Cells were aseptically removed with a micropipette and transferred to a
sterile microcentrifuge tube (1.5-ml capacity). Cells used for DNA
extraction were washed exhaustively in filter-sterilized distilled
water (usually four or five times) until preparations free of
contaminating bacteria, detectable by acridine orange direct counting,
were obtained (17).
DNA extraction, PCR amplification, cloning, and sequencing of 16S
rRNA genes.
The methods used for DNA extraction and PCR
amplification have been described previously (17). PCR
products were purified with either a SpinBind system (Flowgen,
Lichfield, United Kingdom) or a Qiagen PCR clean-up kit (Qiagen,
Crawley, United Kingdom). Cloning was carried out by two methods. TA
cloning was done with the pGEM-T vector and Escherichia coli
JM109 (Promega, Southampton, United Kingdom) as described previously
(17); however, for some samples, forced cloning
(30) was also used. PCR products were amplified with PCR
primers pA and pH' (9) modified to contain PstI
(pAR; 5'-GTGCTGCAGAGAGTTTGATCCTGGCTCAG-3') and
BamHI (pHR;
5'-CACGGATCCAAGGAGGTGATCCAGCCGCA-3') restriction sites (in bold and italic typeface) at their 5' ends. These PCR products were cloned with pUC18 (Boehringer Mannheim, Lewes, United Kingdom). PCR products and pUC18 DNA were digested with PstI
and BamHI and, following their purification with a Qiagen
PCR clean-up kit, were ligated with T4 DNA ligase (Life
Technologies/Gibco BRL, Paisley, United Kingdom). Ligated DNA was used
to transform Epicurian Coli SURE supercompetent E. coli cells (Stratagene, Cambridge, United Kingdom). All procedures
were carried out in accordance with supplier instructions.
White colonies containing inserts (22 from the Hell Kettles clone
library and 20 from the Jenny Dam clone library) were selected at
random, and sequence data were obtained for each with primer pE'
(9). The methods used for the PCR amplification and
sequencing of cloned 16S rRNA gene inserts were described previously
(17). Nearly complete 16S rRNA sequences were obtained from
selected clones with the primers of Edwards et al. (9). All
sequencing was conducted with the DyeDeoxy chain termination method and
an ABI Prism automated DNA sequencer (PE Applied Biosystems,
Warrington, United Kingdom). Sequences were aligned manually by use of
the genetic data environment sequence editor (33) with
reference to the E. coli secondary-structure model.
Phylogenetic distance analyses were conducted with the Jukes-Cantor
(
20) correction for multiple substitutions at a single
site
and the neighbor-joining method (
31) as implemented in
the
TREECON package (
35). Bootstrap resampling was conducted
with 100 replicates. Parsimony analysis was conducted with the
DNAPARS
program, and bootstrapped data sets were generated with
SEQBOOT.
Consensus trees were constructed with the CONSENS program.
All computer
programs used in these analyses were from the PHYLIP
software package
(
11). Maximum-likelihood analysis was conducted
with
fastDNAml (
28). The final alignment used comprised 32 sequences
and covered positions 100 to 202 and 217 to 1480 (
E. coli numbering).
Whole-cell hybridization with fluorescence-labelled 16S
rRNA-targeted oligonucleotide probes.
Oligonucleotide probes
specific for Achromatium-derived sequence clusters recovered
from different locations (Table 1) were designed from aligned 16S rRNA sequences obtained from
Achromatium-derived 16S rRNA gene clone libraries. In some
cases, Achromatium sequences obtained from different
sampling sites contained the same target sequence (Table 1).
High-pressure liquid chromatography-purified oligonucleotide probes 5'
end labelled with tetramethylrhodamine or fluorescein were purchased
commercially (Genosys, Cambridge, United Kingdom). Three
Achromatium 16S rRNA sequence clusters were identified at
each sampling location. The structure of the populations was analyzed
with (i) a cluster-specific, rhodamine-labelled probe, (ii) a
cluster-specific, fluorescein-labelled probe, and (iii) an unlabelled
competitor probe targeting the final cluster to ensure probe
specificity (22). Eub338 (5'-GCTGCCTCCCGTAGGAGT-3') and nonEub (5'-ACTCCTACGGGAGGCAGC-3') (2)
were used in positive and negative control probe hybridization
reactions, respectively.
Achromatium cells were fixed prior to hybridization as
described previously (
17). Fixed cells were pelleted (13,000 rpm,
1 min; MSE Microcentaur; Sanyo, Loughborough, United Kingdom),
washed once in phosphate-buffered saline (130 mM NaCl, 10 mM sodium
phosphate [pH 7.4]), and either resuspended in phosphate-buffered
saline for temporary storage at 4°C (no more than 2 days) or washed
in sterile distilled water and used immediately. Suspensions of
fixed
cells were added to sterile 1.5-ml microcentrifuge tubes
and pelleted
(13,000 rpm, 1 min; MSE Microcentaur), and the supernatant
was removed.
The cells were resuspended in 50 µl of hybridization
buffer
(
1) containing 40% formamide and the combinations of
oligonucleotide probes shown in Table
2.
Hybridization mixtures
contained 100 pmol of each oligonucleotide and
were incubated
at 42°C for 2 to 3 h. Hybridized cells were
pelleted, washed three
times in 50 µl of hybridization buffer for 15 min each time at
42°C, and washed once in filter-sterilized deionized
water (Milli-Q50;
Millipore, Watford, United Kingdom). Finally, cells
were resuspended
in 10 µl of an antifading agent (Citifluor,
Canterbury, United
Kingdom) and mounted on microscope slides.
Hybridized cells were
viewed with an Olympus BH-2-RFCA microscope
fitted with a high-pressure
mercury vapor lamp and blue and green
filter sets (BP545 and BP409).
Micrographs were taken with Kodak
Ektachrome Elite 400 film. Automatic
exposure was used for
phase-contrast micrographs, and exposure
times of 10 s (rhodamine
fluorescence) and 30 s (fluorescein fluorescence)
were used to
obtain double-exposure epifluorescence micrographs.
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TABLE 2.
Combinations of probes used to determine
Achromatium community compositions in three
freshwater sediments
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Determination of the composition of Achromatium
populations by whole-cell in situ hybridization.
Sediment samples
were obtained with a 50-ml syringe coring device (17). The
top 5 cm of the core was removed, and Achromatium cells were
purified from this sediment sample. Two methods of determining
community composition were used. (i) Purified cells were split into two
aliquots and fixed. The samples were hybridized with different
combinations of oligonucleotide probes to provide independent
measurements of the community composition (Table 2). For each aliquot,
200 to 400 cells from randomly selected fields of view were counted and
assigned to different Achromatium-like sequences identified
in clone libraries, based on their hybridization with the different
fluorescence-labelled probes. (ii) Purified cells were split into three
aliquots and fixed. The samples were hybridized with different
combinations of Eub338 and a single Achromatium
subpopulation-specific probe. For each aliquot, 200 to 400 cells from
randomly selected fields of view were counted and assigned to different
Achromatium-like sequences identified in clone libraries,
based on their hybridization with the different fluorescence-labelled
probes. A comparison of the two methods with replicate samples produced
similar results (unpublished data).
Determination of the size distribution of genetically distinct
Achromatium subpopulations by whole-cell
hybridization.
Crude cell preparations from bulk sediment samples
were hybridized with different combinations of oligonucleotide probes
(Table 2). Cells corresponding to the different Achromatium
lineages identified in comparative 16S rRNA analyses were photographed (Kodak Ektachrome Elite 400 film), and their dimensions were determined on projected photographic transparencies. The size distributions for
genetically distinct Achromatium subpopulations obtained
from this analysis were compared statistically. Analysis of variance was conducted on log10-transformed cell diameter data,
followed by a multiple pairwise comparison of group means (least
significant difference).
Determination of the depth distribution of genetically distinct
Achromatium populations by whole-cell hybridization.
A
Rydal Water sediment core (6-cm internal diameter; Perspex core tubes)
was obtained with a Jenkin surface mud sampler (27). The
core was sectioned at 5-mm intervals under oxygen-free nitrogen, and
1.5-cm3 subsamples were removed for the determination of
Achromatium community composition (see above). An additional
subsample (100 µl) was removed from each core section and fixed with
formaldehyde (2% [wt/vol] final concentration) for the determination
of total Achromatium cell counts (14). To
determine sediment redox conditions, the remainder of the sediment
slurry was sealed under nitrogen and stored at 
20°C prior to
analysis of Fe2+ and sulfate by previously described
methods (14). The Achromatium community
compositions in the oxidized (the top 3 cm of the sediment core) and
reduced (below a 3-cm depth) zones in the sediment were compared with a
Pearson chi-square analysis. The oxidized and reduced zones were
operationally defined on the basis of Fe2+ concentrations.
Nucleotide sequence accession numbers.
The 12 nearly
full-length 16S rRNA sequences determined in this study (4 from Jenny
Dam and 8 from Hell Kettles) have been deposited in the GenBank
database under accession numbers AF129548 to AF129559. The accession
numbers for Achromatium sequences from other studies
are L42543, L79966, L79967, L79968, AJ010593, and AJ010596
(13, 17, 18).
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RESULTS |
Phylogenetic diversity in Achromatium communities.
A total of 42 partial 16S rRNA sequences (ca. 480 bp) were obtained in
this study (20 from Jenny Dam and 22 from Hell Kettles). Nearly
complete 16S rRNA sequences were obtained for 13 selected 16S rRNA gene
clones (5 from Jenny Dam and 8 from Hell Kettles), and these were
compared with 6 nearly full-length sequences from previous studies
(13, 17, 18). Phylogenetic analyses with different regions
of the 16S rRNA sequence were conducted, and one of the sequences from
Jenny Dam (JD clone 5) appeared to be chimeric. This sequence was
omitted from subsequent analyses.
Phylogenetic analyses, including analyses of four
Achromatium-derived sequences previously obtained from Rydal
Water sediments
(
17,
18) and two sequences recovered from
Achromatium communities
inhabiting sediments in German
freshwater lakes (Lake Stechlin
and Lake Fuchskuhle) (
13),
confirmed that
Achromatium-like sequences
formed a strongly
supported monophyletic group within the
-subdivision
of the class
Proteobacteria (
13,
17). Only
Achromatium-like
sequences were recovered from the highly
purified cell preparations
obtained in this
study.
16S rRNA sequences recovered from
Achromatium cells from
several geographical locations demonstrated that within the cluster
defined by the
Achromatium-derived sequences, two strongly
supported
clades were present (Fig.
1).
This finding supported the proposal
that
Achromatium-derived
16S rRNA sequences can be accommodated
within two phylogenetic clusters
that have been termed cluster
A and cluster B (
13). This
bifurcation was strongly supported
by distance analysis (100%
bootstrap support for clusters A and
B) and maximum-likelihood analysis
(
P < 0.01). Support for cluster
A was also high in
parsimony analysis (100% bootstrap support),
while support for cluster
B was less pronounced (76% bootstrap
support). The two clusters could
also be distinguished on the
basis of secondary-structure elements in
the V6 region of the
16S rRNA molecule (Fig.
2). This unusual structural motif was
first noted in an
Achromatium 16S rRNA sequence from Rydal
Water
(
17) and was confirmed in subsequent studies
(
13). The V6
region typically contains three helical motifs
(helices 36, 37,
and 38, based on
E. coli secondary
structure). Sequences from
Achromatium cluster A all have a
characteristic deletion and lack
helix 38 (positions 1024 to 1036,
E. coli numbering); many
Archaea and
Eucarya sequences also lack this helix (Fig.
2)
(
16). Sequences
in cluster B, in contrast, have a V6 region
that resembles the
E. coli 16S rRNA secondary structure and
are more typical of the
majority of bacteria (Fig.
2) (
16).
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FIG. 1.
Phylogenetic distance tree based on the comparative
analysis of nearly full-length 16S rRNA gene sequences recovered from
Achromatium cells purified from Rydal Water (RY), Jenny Dam
(JD), and Hell Kettles (HK) sediments. Two additional
Achromatium-derived sequences, from Lake Stechlin and Lake
Fuchskuhle (13), were included in the analysis. The numbers
in parentheses indicate the percentages of clones from the three clone
libraries that were closely related to the annotated clusters or
sequences (RY, n = 7; JD, n = 19; HK,
n = 22). The sequences targeted by the oligonucleotide
probes used in this study are also labelled. The scale bar denotes 2%
sequence divergence, and the values at the nodes indicate the
percentages of bootstrap trees that contained the cluster to the right
of the node. Bootstrap values lower than 50 are not shown.
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FIG. 2.
Secondary structure of the V6 region of small-subunit
rRNA from a range of organisms (16). (a to d)
Bacteria. (e) Archaea. (f) Eucarya.
(a) Achromatium cluster A, RY clone 5. (b)
Achromatium cluster B, JD clone 2. (c) E. coli.
(d) Mycoplasma capricolum. (e) Methanobacterium
formicicum. (f) Drosophila melanogaster.
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Representative sequences from both clusters were identified at all
three sites investigated in the current study. Within clusters
A and B,
further structure was evident. Sequences recovered from
Jenny Dam and
Rydal Water formed monophyletic groups that were
distinct from the
clades harboring Hell Kettles sequences (Fig.
1). These relationships
were also evident in parsimony and maximum-likelihood
analyses, and the
monophyletic nature of the Rydal Water-Jenny
Dam and Hell Kettles
sequences within clusters A and B was supported
by bootstrap values in
the range of 94 to 100%. An
Achromatium sequence from Lake
Stechlin (
13) was most closely related to
the cluster A
sequences from Hell Kettles, whereas an
Achromatium sequence
from Lake Fuchskuhle (
13) may represent a distinct
lineage
affiliated with the cluster B sequences from Rydal Water
and Jenny Dam
(Fig.
1).
Occurrence of phylogenetically distinct subpopulations within
Achromatium communities.
There are a number of
possible explanations for the recovery of multiple related
Achromatium-like sequences in PCR-derived clone libraries of
16S rRNA genes produced from purified preparations of
Achromatium cells. The sequence diversity may correspond to genetically distinct populations of Achromatium cells
present in a single sample. Alternatively, the observed sequence
diversity may be due to the presence of multiple divergent 16S rRNA
genes present within individual cells of a single species of
Achromatium. Heterogeneous 16S rRNA operons are known to
occur in the genomes of many bacterial species (7, 23, 26),
and it has been suggested that heterogeneity in rRNA operons could, in
some cases, contribute to the diversity observed in bacterial
communities studied by rRNA-based techniques (12). A third
explanation is that the sequences came from bacteria that were
phylogenetically related to Achromatium but were not
morphologically identifiable as Achromatium cells.
Whole-cell hybridization with fluorescence-labelled oligonucleotide
probes specific for the different Achromatium-derived sequences recovered from the three geographical locations studied here
was used to establish which explanation was correct.
Oligonucleotide probes were designed to target the more deeply
branching lineages within the
Achromatium clades (i.e.,
those
exhibiting less than 97% sequence similarity) (Table
3 and Fig.
1). On this basis, three
probes were used to discriminate genetically
distinct
Achromatium subpopulations at each study site. For example,
to analyze the
Achromatium population in Rydal Water
sediments,
two different probes were used to identify cells containing
RY
clone 1 or RY clone 5 sequences (93.75% similarity), and a single
probe was used to identify both the RY clone 7 and the RY clone
8 sequences (97.26% similarity).
Simultaneous whole-cell hybridization with fluorescence-labelled
oligonucleotide probes targeting different
Achromatium-derived
sequences clearly demonstrated that the
divergent sequences recovered
in 16S rRNA gene clone libraries
represented phylogenetically
distinct subgroups present within natural
Achromatium populations
(Fig.
3). This finding was confirmed for all of
the sediments
investigated in this study. In whole-cell hybridization
experiments,
Achromatium populations comprised
fluorescein-labelled, rhodamine-labelled,
and unlabelled cells. The
binding of more than one probe to a
single cell was never observed with
any combination of probes
used. Consequently, different target
sequences are unlikely to
represent multiple rRNA operons occurring
within a single
Achromatium cell. Furthermore, all of the
Achromatium-specific probes bound
to cells that could be
identified, on the basis of their morphology,
as
Achromatium.
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FIG. 3.
Whole-cell in situ hybridization of
Achromatium cells purified from Rydal Water sediments. Cell
were simultaneously hybridized with three oligonucleotide probes:
ARY655a (rhodamine-labelled, RY clone 5-specific probe), ARY655b
(unlabelled [competitor] RY clone 7- or RY clone 8-specific probe),
and ARY655c (fluorescein-labelled, RY clone 1-specific probe). Phase
contrast (a) and epifluorescence (b) micrographs of the same
microscopic field are shown. Two RY clone 5 cells (red fluorescence)
and one RY clone 1 cell (green fluorescence) are shown. The scale bar
represents 20 µm and applies to both micrographs.
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Using different combinations of fluorescein-labelled,
rhodamine-labelled, and unlabelled probes (Table
2), we obtained direct
counts of specific subgroups of
Achromatium for cells
collected
from Rydal Water, Jenny Dam, and Hell Kettle sediments. The
compositions
of the
Achromatium communities were determined
independently with
different probe combinations (Table
4). For example, in a sample
of cells
collected from Jenny Dam (July 1998), the proportions
of cells
identified as containing JD clone 1-like (or JD clone
8-like) sequences
were very similar whether they were identified
on the basis of
hybridization with a JD clone 1-specific (or JD
clone 8-specific) probe
or on the basis of the fact that they
did not bind probes specific for
the other two groups analyzed
(Table
4). Furthermore, at least 98% of
Achromatium cells (
n = 200) identified by
phase-contrast microscopy hybridized with
fluorescence-labelled
Achromatium-specific probes. This value
is comparable to the
97.8% of cells (
n = 267) which bound the
general
bacterial probe Eub338. Similar values were obtained with
samples from
all three sites investigated; thus, the diversity
encompassed by the
majority of
Achromatium cells was characterized
with PCR
amplification and cloning of 16S rRNA genes.
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TABLE 4.
Community compositions of different
Achromatium subpopulations in Jenny Dam sediments (July
1998) determined with different combinations of labelled and
unlabelled competitor probesa
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Morphological differentiation of phylogenetically distinct
subpopulations within Achromatium communities.
The
frequency distribution of cell diameter in Achromatium
communities from different geographical locations was analyzed. Cell
diameter rather than cell volume was found to be a better descriptor of
differences in cell size, presumably because length or volume
measurements of Achromatium cells, which grow along their
long axis and divide perpendicular to it, include growth-related variations.
The range of cell diameters observed for
Achromatium
communities from Rydal Water, Jenny Dam, and Hell Kettles sediments
(Fig.
4) fell within the range of cell
dimensions reported for
Achromatium cells from other
habitats (
3,
5,
10,
24,
32,
36,
37). Moreover, the mean cell
diameters observed for
Achromatium communities from the
different study sites investigated here were
found to be significantly
different (
P < 0.05) (Fig.
4). This
finding suggested
that different species or strains of
Achromatium with
different mean cell diameters were present in the different
sediments
studied. However, while this conclusion may be true,
until now no
detailed analysis of the size distribution of
Achromatium cells has been reported, and the considerable overlap in the dimensions
of
Achromatium cells has confounded the use of cell size as
a
criterion for the differentiation of
Achromatium species
(
10).
This situation is well illustrated in the analysis of
Achromatium cells from Rydal Water, Jenny Dam, and Hell
Kettles sediments.
Although the mean cell diameters for the communities
were statistically
significantly different, the considerable overlap in
the size
distribution of each population would make it impossible to
assign
an individual cell to any particular population, except perhaps
the larger cells present in Hell Kettles sediments.
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FIG. 4.
Frequency distribution of the diameters of
Achromatium cells from sediment samples taken from Rydal
Water, Jenny Dam, and Hell Kettles. The Achromatium
communities present at each sampling site had a characteristic site
distribution, and the mean diameters of cells from these locations were
significantly different. CI, confidence interval.
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Phylogenetic analysis of cells from different geographical locations
(Fig.
1) confirmed not only that the
Achromatium communities
present in a range of freshwater sediments were genetically distinct
but that each community comprised several genetically distinct
subpopulations (Fig.
1 and
3). To determine if there was morphological
differentiation (on the basis of cell size) of
Achromatium
subpopulations
from a single sediment, identification of genetically
distinct
cells by whole-cell hybridization was combined with
measurements
of cell
diameter.
The size distribution of genetically distinct
Achromatium
subpopulations identified by whole-cell in situ hybridization indicated
that cells corresponding to the different
Achromatium
sequence
types fall into distinct size classes (Fig.
5). It should be noted
that the frequency
distributions of cell diameters presented were
obtained by measurement
of cells selected from the different subpopulations
until sufficient
cells had been measured to obtain a meaningful
distribution. Thus, the
sum of the frequency distributions is
not representative of a naturally
occurring
Achromatium community.
Multiple pairwise
comparisons (least significant difference) of
the mean cell diameters
of the genetically distinct
Achromatium subpopulations
indicated significant differences (
P < 0.01) between
almost all of the phylogenetically distinct
Achromatium subpopulations
identified by whole-cell in
situ hybridization. Statistically
significant differences in mean cell
diameters were evident both
in subpopulations coexisting in a single
sediment and among all
the subpopulations identified at different
geographical locations,
with the exception of cells identified as JD
clone 1 from Jenny
Dam. Comparisons of JD clone 1 with JD clone 8 and
of JD clone
1 with HK clone 13 (Fig.
5) revealed no significant
differences
in cell diameters. However, only five cells corresponding
to JD
clone 1 were identified in this analysis (Fig.
5c); thus, a much
smaller sample size was used in the statistical comparison. If
a larger
number of JD clone 1-like cells had been available for
cell diameter
measurements, a more definitive statistical analysis
of this
Achromatium subpopulation would have been possible.
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|
FIG. 5.
Frequency distributions of genetically distinct
Achromatium cells from Rydal Water (a), Hell Kettles (b),
and Jenny Dam (c) sediments. Cells were discriminated with
fluorescence-labelled oligonucleotide probes specific for different
Achromatium-derived sequences identified in clone libraries
from the different sediments (Tables 1 and 2). Cells corresponding to
JD clone 1 were rather rare, and only a small number of cells were
measured (n = 5); consequently, data from this
subpopulation were not plotted. CI, confidence interval.
|
|
Temporal variations in the compositions of Achromatium
communities.
Using whole-cell hybridization with specific
fluorescence-labelled oligonucleotide probes, we obtained direct counts
of Achromatium subpopulations from Rydal Water and Hell
Kettles sediment cores sampled at different times of the year. The
structure of the Achromatium communities within sediments
was dynamic, and changes in the numerical dominance of different
subpopulations were observed at different sampling dates (Fig.
6). Changes in community structure were
clearly demonstrated for Achromatium communities from Rydal
Water sediments. Between December 1997 and June 1998, there was a
relative decrease in the larger RY clone 1 cells, while there was a
relative increase in the RY clone 7 or RY clone 8 cells. Less
pronounced population changes were observed at Hell Kettles, where the
HK clone 1 cell type remained dominant throughout the period of
sampling (Fig. 6).
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FIG. 6.
Relative abundance of genetically distinct
Achromatium subpopulations in sediment samples taken at
different times of the year from Rydal Water and Hell Kettles. Cells
belonging to the different subpopulations were discriminated with
fluorescence-labelled oligonucleotide probes specific for different
Achromatium-derived sequences (Tables 1 and 2). The total
number of cells (the sum of cells counted from each aliquot) counted in
each sample was as follows: Rydal Water December 1997, n = 913, April 1998, n = 384, and June 1998, n = 395; Hell KettlesJanuary 1998, n = 300, March 1998, n = 300, and May 1998, n = 300.
|
|
Depth distributions of Achromatium subpopulations.
The depth distribution of Achromatium cells in a sediment
core from Rydal Water indicated that Achromatium cells were
present throughout the top 5 cm of the sediment (Fig.
7a). The majority of cells were located
in the top 3 cm, where high sulfate and low Fe2+
concentrations indicated oxidizing conditions (Fig. 7b) (6). Below 3 cm, where Achromatium cells were less abundant, low
sulfate and high Fe2+ concentrations indicated more
reducing conditions. The distribution of Achromatium
across both oxidized and reduced zones in sediments was consistent with
previous observations (14). Analysis of the composition of
the Achromatium population at each depth was conducted by
whole-cell in situ hybridization. This analysis demonstrated that each
subpopulation was present at all depths (Fig. 7c). However, while the
proportions of the RY clone 1 cell type remained similar regardless of
depth, a relative change in the proportions of RY clone 7 or RY clone 8 and RY clone 5 cell types corresponded with the transition between
oxidized and reduced zones. The RY clone 7 or RY clone 8 cell type,
which comprised 67.8% of cells (n = 1,137) in the
oxidized zone, represented a statistically significantly smaller
proportion of the population (50.1%, n = 373) in the
reduced zone (Pearson chi-square value, 56.69; df, 2; P <
0.001). Conversely, the RY clone 5 cell type represented 14.8%
(n = 1,089) of the Achromatium population in
the oxidized zone but represented a significantly larger proportion of
the population (34.6%, n = 399) in the reduced zone
(Pearson chi-square value, 72.55; df, 2; P < 0.001).
Thus, the subpopulations present within a single Achromatium
community exhibited different distributions under different redox
conditions, providing evidence of niche adaptation in the
Achromatium subpopulations.
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FIG. 7.
Depth profiles of Achromatium cells (a),
redox-sensitive chemical species (Fe2+ and
SO42 ) (b), and relative abundance of the
three main Achromatium lineages identified in a sediment
core from Rydal Water (c). The trend lines show a three-point moving
average of the data, and error bars represent 95% confidence intervals
for counts of the Achromatium subpopulations.
|
|
| |
DISCUSSION |
Phylogenetic diversity in Achromatium populations.
The presence of phylogenetic diversity in Achromatium
communities has now been demonstrated unequivocally. Of the five
Achromatium communities from different geographical
locations studied to date (three in this study and two studied by
Glöckner et al. [13]), four have been shown to
comprise several genetically distinct subgroups, while only one, the
community present in Lake Fuchskuhle sediments, appears to be
homogeneous. Moreover, the genetically distinct Achromatium
subpopulations present at each site were endemic to those lake
sediments (Fig. 1). Only Lake Fuchskuhle and Lake Stechlin potentially
share a common Achromatium subpopulation (13). A
total of 28% of the cells in Lake Stechlin bound two oligonucleotide
probes designed to target an Achromatium sequence recovered
from Lake Fuchskuhle (AFK192 [positions 192 to 211, E. coli
numbering] and AFK433 [positions 433 to 450, E. coli
numbering); nonetheless, a sequence corresponding to the genotype found
in Lake Fuchskuhle was not detected in 16S rRNA gene clone libraries obtained from cells purified from Lake Stechlin. However, as
Glöckner et al. (13) pointed out, the phylogenetic
affinity of the cells from Lake Fuchskuhle sediments and
AFK192/AFK433-positive cells in Lake Stechlin is only speculative,
since their identification was based on probe binding alone and the
presence of two identical target sequences in different 16S rRNA
molecules does not necessarily imply high similarity elsewhere in the
molecules (13). This point is well illustrated when the
divergent sequences obtained in this study are considered. If one were
to construct oligonucleotide probes targeting the same positions in the
16S rRNA molecules as AFK192 and AFK433 but specific for
Achromatium rRNA sequences obtained from Jenny Dam and Rydal
Water, cells containing the JD clone 1 and RY clone 1 sequences would
bind both probes, yet these sequences have only 97.6% sequence
identity (Table 3). Likewise, cells belonging to JD clone 2 or JD clone
13 and to RY clone 7 or RY clone 8 (97.3 to 97.9% similarity) would
also bind both probes but are clearly phylogenetically distinct (Fig. 1). It is also evident that much more divergent sequences also contain
common diagnostic oligonucleotide sequences; for example, RY clone 5 and HK clone 2 sequences have only 92.1% homology in their 16S rRNAs,
yet both contain the target sequence for probe ARY655a (Table 1).
The occurrence of genetically distinct subpopulations in
Achromatium communities provides evidence to support the
delineation
of
Achromatium into different taxonomic
groupings. For instance,
many of the 16S rRNA sequences obtained from
Achromatium cells
had less than 97.5% 16S rRNA sequence
homology. This level of
sequence divergence has been used to delineate
species (
34).
On this basis, most of the
Achromatium subpopulations identified
may reasonably be
defined as separate species. No members of the
genus
Achromatium have been isolated in culture; consequently,
relationships among different
Achromatium spp. can be
inferred
only by phylogenetic analysis of environmental samples,
morphology,
and possibly habitat. On these bases, the
Achromatium population
native to Lake Fuchskuhle sediments
has been described as a new
species, "
Candidatus
Acromatium minus" (
13). The genetic characterization
of
Achromatium and the discovery that genetically distinct
cells
are differentiated morphologically on the basis of cell size in
the current study indicate that several more
Achromatium
spp.
remain to be
described.
Significance of morphology (cell diameter) in the classification of
Achromatium.
From a historical perspective, the different
cell size distributions of phylogenetically distinct
Achromatium subpopulations observed in this study clarify
one of the earliest debates surrounding the taxonomy of
Achromatium. Without access to cultured organisms or
molecular techniques, early classification relied on morphological methods alone. These early studies (5, 10, 24, 32, 36, 37)
and some later contributions (e.g., 3) reported that Achromatium cells from different locations had different
dimensions. These early reports indicated that the bacterium exhibited
cell dimensions ranging from 7 to 36 µm in diameter and 7 to 102 µm in length. On this basis, Nadson and Visloukh (24) suggested that a number of different forms of A. oxaliferum existed
and proposed the additional epithets minus, medium, majus, elongatum, and gigas to describe them (see also reference (5)).
Classification based on cell size has been applied to filamentous
sulfur bacteria related to Achromatium. For example,
different Thioploca spp. and Beggiatoa spp. can
be distinguished on the basis of filament diameter, a characteristic
which can vary by as much as 1 order of magnitude within these genera
(25). However, in the genus Achromatium, there is
a considerable overlap in the cell size distributions of different
species. Given this continuous distribution of cell size and the
natural variations that populations of bacteria are likely to exhibit
when located in different environments, one author (10)
considered that delineation of different Achromatium species
on the basis of cell size was not valid. The data presented here
provide the first direct evidence that different Achromatium species are morphologically distinct and support the original proposal
of Nadson and Visloukh (24). However, although different Achromatium spp. have now been shown to have characteristic
cell sizes, in isolation cell size cannot be recommended as a criterion for their identification. Recovery of 16S rRNA sequences and whole-cell in situ hybridization provide the only reliable means of identifying new Achromatium species.
Ecological significance of genetically distinct subpopulations of
Achromatium.
Achromatium species from different
geographical locations show some degree of site specificity; i.e.,
Achromatium sequences recovered from Rydal Water and Jenny
Dam are more closely related to each other than they are to sequences
from Hell Kettles (Fig. 1). Furthermore, this relationship is apparent
within both cluster A and cluster B sequences and appears to correlate
with the nutritional characteristics of the Achromatium
communities found at the different sites. Recent studies on the uptake
of inorganic and organic substrates by Achromatium cells in
Rydal Water and Hell Kettles sediments, determined by
microautoradiography (15), have indicated that some cells in
both populations can assimilate simple organic compounds. However,
while the majority of cells from Rydal Water were shown to assimilate
inorganic carbon, none of the cells from Hell Kettles had this
ability. Furthermore, homologues of the ribulose-1,5-bisphosphate carboxylase/oxygenase large-subunit gene (rbcL), involved in
CO2 fixation, and adenosine-5'-phosphosulfate reductase
genes (aprBA), associated with energy-conserving sulfur
oxidation pathways (19), could be amplified from highly
purified preparations of Achromatium cells from Rydal Water
and Jenny Dam sediments but not from cells collected at Hell Kettles
(18). The presence of these genes in genomic DNA from
Achromatium species present at Rydal Water is consistent
with the autotrophic potential exhibited by at least some of the cells
in this population.
While different geographical locations appear to harbor genetically and
nutritionally distinct
Achromatium populations, presumably
as a result of ecosystem-imposed selection pressures, phylogenetically
and morphologically distinct
Achromatium subpopulations also
occur
together within the same ecosystem; at a single location, the
Achromatium population structure can be dynamic (Fig.
6).
These
results suggest that conditions which favor the proliferation
of
one
Achromatium subpopulation do not necessarily apply to
all.
In this study, it was not possible to determine physiological
differences among coexisting
Achromatium subpopulations;
however,
depth- and redox-related changes in community composition were
detected (Fig.
7c). This finding indicated that different
Achromatium subpopulations are better adapted to different
redox conditions
and probably occupy distinct ecological niches within
the sediment
environment. Thus, adaptive radiation in response to
environmental
heterogeneity may have resulted in the evolution of
several coexisting
but genetically and ecologically distinct
Achromatium subpopulations
that have optimal activities
under different redox conditions.
One source of heterogeneity in
sediments is the depth-related
succession of respiratory processes
(
6) that gives rise to
a number of physiochemical and
depth-defined niches in close proximity.
The presence of these niches
may have been the driving force behind
the diversification observed for
Achromatium communities.
| |
ACKNOWLEDGMENTS |
We thank Nick Davis for providing data on cell size distributions
from Rydal Water. We also thank Frank Oliver Glöckner and Rudolf
Amann for providing Achromatium sequence data from Lake Fuchskuhle and Lake Stechlin prior to publication and for stimulating discussion.
This work was supported by the IFE and the FBA. Financial support was
provided by the Natural Environment Research Council (grant GR3/9148 to
I.M.H., R.W.P., and J.G.J. and studentship GT4/95/235/F to R.H.).
| |
FOOTNOTES |
*
Corresponding author. Mailing address: Fossil Fuels and
Environmental Geochemistry Postgraduate Institute (NRG), University of
Newcastle, Newcastle upon Tyne NE1 7RU, United Kingdom. Phone: 44 (0)
191 222 7024. Fax: 44 (0) 191 222 5431. E-mail:
i.m.head{at}newcastle.ac.uk.
| |
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Top
Abstract
Introduction
