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Applied and Environmental Microbiology, March 2001, p. 1328-1334, Vol. 67, No. 3
0099-2240/01/$04.00+0 DOI: 10.1128/AEM.67.3.1328-1334.2001
Copyright © 2001, American Society for Microbiology. All rights reserved.
Suboxic Deposition of Ferric Iron by Bacteria in
Opposing Gradients of Fe(II) and Oxygen at Circumneutral pH
Dmitri
Sobolev and
Eric E.
Roden*
Department of Biological Sciences, The
University of Alabama, Tuscaloosa, Alabama
Received 10 August 2000/Accepted 11 December 2000
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ABSTRACT |
The influence of lithotrophic Fe(II)-oxidizing bacteria on patterns
of ferric oxide deposition in opposing gradients of Fe(II) and
O2 was examined at submillimeter resolution by use of an
O2 microelectrode and diffusion microprobes for iron. In
cultures inoculated with lithotrophic Fe(II)-oxidizing bacteria, the
majority of Fe(III) deposition occurred below the depth of
O2 penetration. In contrast, Fe(III) deposition in abiotic
control cultures occurred entirely within the aerobic zone. The
diffusion microprobes revealed the formation of soluble or colloidal
Fe(III) compounds during biological Fe(II) oxidation. The presence of
mobile Fe(III) in diffusion probes from live cultures was verified by
washing the probes in anoxic water, which removed ca. 70% of the
Fe(III) content of probes from live cultures but did not alter the
Fe(III) content of probes from abiotic controls. Measurements of the
amount of Fe(III) oxide deposited in the medium versus the probes
indicated that ca. 90% of the Fe(III) deposited in live cultures was
formed biologically. Our findings show that bacterial Fe(II) oxidation is likely to generate reactive Fe(III) compounds that can be
immediately available for use as electron acceptors for anaerobic
respiration and that biological Fe(II) oxidation may thereby promote
rapid microscale Fe redox cycling at aerobic-anaerobic interfaces.
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INTRODUCTION |
Neutrophilic bacteria associated
with Fe(III) oxide precipitation have been known for a long time
(26). Some species were shown to precipitate Fe(III)
oxides during heterotrophic metabolism (6, 7). However,
neutrophilic autotrophic iron oxidation has only recently been reliably
demonstrated (8, 10). The role of these bacteria in
biogeochemical iron cycling, however, has not been extensively studied.
Such lack of study can be attributed to the fact that Fe(II) is highly
unstable in oxic environments at circumneutral pH, resulting in the
widely held belief that Fe(II) will be rapidly oxidized regardless of
the presence or absence of microbial catalysis (24).
Furthermore, Emerson and Moyer (8) have convincingly
demonstrated that Fe(II)-oxidizing bacteria do not alter the rate of
Fe(III) oxide accumulation in diffusion-limited opposing-gradient systems.
Regardless of the possible influence of Fe(II)-oxidizing bacteria on
rates of Fe(II) oxidation, these organisms have the potential to affect
Fe(III) oxide precipitation processes by altering the spatial
relationship between O2 and Fe(II) gradients. Emerson and
Moyer (8) suggested that bacterial Fe(II) oxidation might promote coupling between iron oxidation and reduction by producing amorphous (9) or poorly crystalline Fe(III) oxides which
are readily available for Fe(III)-reducing bacteria. The potential for
a tight, microbially mediated coupling between iron oxidation and
reduction has important environmental implications, given the critical
influence which iron cycling exerts on the behavior of various organic
and inorganic compounds in aquatic systems (24).
In this study we examined O2 and Fe(II) gradients together
with bacterial numbers and patterns of Fe(III) oxide deposition at
submillimeter resolution in Fe(II)-oxidizing gradient cultures, using
organisms enriched from iron-rich freshwater wetland sediments. The
goal was to determine the positioning of Fe(II)-oxidizing bacteria with
respect to Fe(II) and O2 gradients and to examine how these
organisms influence patterns of Fe(III) oxide deposition within these gradients.
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MATERIALS AND METHODS |
Enrichment and isolation.
A neutrophilic Fe(II)-oxidizing
enrichment culture (TW1) was obtained from iron-rich surficial
sediments of a freshwater wetland in the Talladega National Forest in
north central Alabama by use of gradient cultures (15)
with FeS as an Fe(II) source. After several passages on FeS (low iron)
medium, the culture was transferred to medium with 50 mM PIPES
[piperazine-N,N'-bis(2-ethenesulfonic acid)]-buffered FeCl2 · 2H2O as an iron
source (high iron), the same medium used in the experiments reported
here. The culture was shown to contain heterotrophic satellite bacteria
capable of growth on rich medium (50% strength tryptic soy agar
[TSA]). PCR-denaturing gradient gel electrophoresis (DGGE) analysis
of ca. 200-bp 16S rRNA gene fragments was performed as described elsewhere (18) and revealed two strains, one presumably
the lithotrophic Fe(II) oxidizer, and the other presumably a
heterotrophic satellite. However, as described below, we found no
evidence that the latter organism had a major impact on the results of
our Fe(II) oxidation experiments. Another, apparently pure culture
(TW2) used in this study was enriched and isolated by repeated passages on high-iron medium. This culture failed to grow on rich medium, and
DGGE analysis revealed only a single genome. A BLAST search (2) on a 1,485-bp fragment of the 16S rRNA gene sequenced
suggested that our organism is closely related (94% similarity) to
Dechlorisoma suilla, a dissimilatory perchlorate-reducing
bacterium (1, 4). Further molecular and physiological
characterization of TW2 is under way.
Gradient cultures.
The gradient culture system consisted of
two layers in 250-ml beakers: a bottom layer containing the Fe(II)
source (50 mM FeCl2 in anaerobic 10 mM PIPES buffer with
2% [wt/vol] Noble agar [pH 7.0]), and a top layer consisting
of mineral medium (NaHCO3, 30 mM; NH4Cl, 10 mM;
KH2PO4, 1 mM) supplemented with vitamins and
minerals (17) and stabilized with 0.25% Noble agar. Layer volumes were 25 ml (bottom) and 125 ml (top), resulting in layer depths
of 12 and 60 mm, respectively. This concentration of Fe(II) was
necessary to sustain Fe(II) flux over the course of the experiment. Although the Fe(II) concentration in the bottom layer was
unrealistically high relative to natural systems, diffusion within the
agar column resulted in environmentally relevant Fe(II) concentrations
of 1 to 2 mM (22) close to the oxic-anoxic boundary. Prior
to initiation of the experiment, beakers were covered with aluminum
foil and autoclaved. The foil cover was kept on all the time except
when components were added. After the bottom layer was poured, probes for Fe(II) and bacterial numbers (see below) were inserted, and the
agar was allowed to solidify under an anaerobic atmosphere (about an
hour under approximately 95:5 [vol/vol]
N2-H2). The top layer, which had been degassed
with 80:20 (vol/vol) N2-CO2, autoclaved in
crimp-sealed bottles, and cooled to approximately 30°C, was then
added to the system. The beakers were incubated overnight in an
anaerobic chamber in order to allow a supply of Fe(II) to diffuse into
the top layer prior to inoculation. Inoculation was achieved by
inserting a pipette into the top layer and ejecting about 0.1 ml of the
microaerobic inoculum (surface layer of a high-iron culture of TW1 or
TW2) as the pipette was withdrawn; this procedure was repeated four to
six times per culture. Although this procedure could have enhanced
O2 penetration into the surface layer, our "time zero"
O2 measurements revealed no difference between
O2 profiles in inoculated and uninoculated cultures.
Additionally, in later experiments with TW2, possible O2
introduction was controlled for by stabbing uninoculated controls with
a sterile pipette in the manner similar to the inoculation procedure.
Fe(II) measurements.
We employed a diffusion microprobe
technique modified from that of Davison et al. (5) to
determine Fe(II) concentrations at submillimeter resolution in the
gradient cultures. The probe consisted of a 0.1-mm-thick 5% (vol/vol)
agar film attached to a glass microscope slide. The film was cast
between hot (ca. 70°C) glass slides autoclaved in an aluminum foil
boat, and the agar was allowed to solidify. Excess agar was trimmed
from the sides of the slides, and the slides were separated, leaving
the film attached to one slide. The probe was inserted into the culture beaker with the agar film facing the center. All of the above procedures were accomplished under aseptic conditions. Probes were
retrieved immediately after the final O2 measurements (see below), dipped into 1% (wt/vol) potassium ferricyanide
[K3Fe(CN)6] solution [which forms an
insoluble blue complex with Fe(II)] for about 1 s, retrieved, and
allowed to react for ca. 3 min after being removed from the solution.
This fixed the Fe(II) within the agar film and provided a colored
substance whose abundance could be quantified. Probes were then soaked
in distilled water for ca. 5 min to remove excess potassium
ferricyanide. Images of the probes were collected under a dissecting
microscope with an attached camera. The images were then digitized and
converted to black and white by use of Adobe PhotoShop. The optical
density of the images, presumably representing the density of
Fe3[Fe(CN)6]2, was measured by
the NIH-Image software. For TW1 cultures and controls, three depth
profiles from a single probe were collected and measured. For TW2, a
single profile was recorded for each of the probes from triplicate
cultures. Fe(II) concentrations in the probes were quantified against a
calibration curve obtained by measuring the optical density of the agar
strips of the same thickness incubated overnight in anoxic Fe(II)-EDTA
solutions of known concentration. This yielded linear standard curves
with an R2 of 0.8 or greater within the range
from 1 to 25 mM Fe(II).
In order to validate the diffusion probe technique, culture systems
were cored with a detipped 1-ml plastic syringe immediately after
O2 measurements but before probe retrieval. The core was sectioned anoxically, and Fe(II) in several depth intervals was extracted with 0.5 M HCl and measured by the Ferrozine
(23) method. To ensure complete extraction of Fe(II) from
the agar, tightly closed vials containing 0.5 M HCl and sample were
gently heated in a waterbath (ca. 80°C) until the agar dissolved.
To determine the abundance of Fe(III) in probes from control and live
culture systems, high-iron cultures were set up as described
above and
incubated for 6 days at 20°C. Core samples of the medium
which
included the whole depth of the top layer were taken anaerobically
and
immediately placed into preweighed vials containing 0.5 M
HCl. Probes
were retrieved immediately after taking the core,
and the agar film
(exposed to the medium) was scraped off into
preweighed vials
containing 0.5 M HCl. The portion of the film
exposed to the atmosphere
above the medium dried out and was firmly
attached to the slide,
allowing collection of only the portion
submerged in the medium. Acid
extracts were analyzed using Ferrozine
for Fe(II) and total Fe, from
which Fe(III) content was
calculated.
To estimate what percentage of the Fe(III) trapped in the probes
comprised mobile (soluble and/or colloidal) compounds, triplicate
sets
of probes were retrieved from live and abiotic control cultures
and
washed three times in 100 ml of anoxic distilled water, transferring
the probes each time into a fresh beaker of water. Results were
compared to the Fe(III) content of triplicate unwashed probes
retrieved
from parallel cultures. The difference was assumed to
represent the
diffusionally mobile Fe(III)
content.
Oxygen microelectrode measurements.
A Clark-style
O2 microelectrode with guard cathode (21)
(Diamond General Corp, Ann Arbor, Mich.), attached to an
electronically controlled micromanipulator (National Aperture model
MM33CR), was used to determine O2 profiles in the gradient
cultures. The microelectrode was calibrated to indicate percent air
saturation of O2, with a detection limit of approximately
0.1% saturation, which, under our conditions, was equivalent to 0.279 µM. Zero depth was set by manually lowering the electrode to the agar
surface and identifying the moment of contact by the formation of a
visible meniscus around the electrode tip. The electrode was raised
until the tip separated from the surface and the meniscus disappeared, and slowly lowered until the meniscus formed again. At this point, the
manipulator counter was set at zero. Oxygen was considered depleted
when three consecutive measurements (covering a distance of 50 to 200 µm) below the detection limit (ca. 0.28 µM) were obtained.
In one experiment, O
2 microprofiles were measured
repeatedly over a 7-day course. In this case, uninoculated culture
systems
were measured first to minimize contamination. Between
sampling,
the electrode was allowed to stand in distilled water. The
electrode
was occasionally soaked in dilute HCl to remove oxide
precipitates.
In some experiments, pH gradients were measured using a
microcombination pH electrode (Orion) according to the manufacturer's
instructions. Since this electrode has a relatively large tip
(ca. 1 mm
diameter), pH was measured at 1-mm
intervals.
Bacterial numbers.
The Rossi-Cholodny buried slide technique
(20) was employed to enumerate bacteria (in units of cells
per unit area of slide) at submillimeter resolution in our cultures.
This technique is based on colonization of glass slides inserted into
stratified bacterial communities. After colonization, slides are
retrieved, fixed, and stained, and the bacteria attached to them are
enumerated. Although this technique enumerates only those bacteria
which attach to the slide, it provides a better depth resolution (0.2 mm or better) than coring and slicing the culture (1 to 2.5 mm). A
basic assumption of our application of this technique is that an equal percentage of bacteria attach to the slides at each depth, which is not
unreasonable for a pure culture. Slides were inserted into the culture
systems for the duration of the experiment, and bacterial growth was
allowed to occur. Slides were then removed, fixed with 4% formaldehyde
solution, and stained with acridine orange, and bacteria were counted
under an epifluorescent microscope. Five fields were counted at each of
the depth intervals spaced at 0.2 mm or greater. These area counts were
used as a proxy for actual number of bacteria per unit volume.
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RESULTS |
Oxygen and Fe(III) oxide distributions.
Oxygen gradients were
much steeper in culture systems inoculated with Fe(II)-oxidizing
bacteria than in sterile controls (Fig. 1). After 5 days of incubation,
O2 penetrated to 12 mm in the controls, versus 1.5 mm in
the TW1 culture (average of three separate profiles in a single system
for both control and live cultures). A distinct bacterial plate formed
at the oxic-anoxic interface in the live cultures, whereas in the
control system only low bacterial numbers (>10-fold lower) were
detected, and only at the surface (Fig. 1A and B). A similar experiment
conducted with the presumably pure TW2 culture produced nearly
identical results (Fig. 1C and D).
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FIG. 1.
Distribution of Fe(II), particulate Fe(III) oxides,
O2, and bacteria in two Fe(II)-oxidizing cultures. (B and
D) TW1 and TW2 cultures, respectively. (A and C) Abiotic controls for
the cultures shown in B and D, respectively. Note that Fe(II) profiles
determined by densitometry in diffusion probes are confounded by the
presence of Fe(III) compounds in the probe (see text). O2
profiles are averages of triplicate measurements. (A and B)
Representative Fe(II) profile from triplicate profiles obtained from a
single probe. (C and D) Fe(II) profiles are averages of single
measurements from probes in triplicate cultures. No error bars are
shown. Bacterial numbers in A and B are averages of counts on
triplicate slides from a single culture (A and B) or averages of counts
from a single slide from each of triplicate cultures (C and D); error
bars were omitted for clarity. In control cultures, bacterial numbers
were never significantly different from zero.
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Fe(III) oxides were deposited below the O
2
penetration zone in the bacterial cultures as opposed to the
controls, in which
the entire zone of oxide deposition was aerobic
(Fig.
1).
Control experiments.
Since TW1 was not pure, control
experiments in iron-free cultures were conducted to account for
possible heterotrophic growth of the satellite organisms on impurities
in the agar, because such growth could influence O2
gradients and thus confound interpretation of the oxide band data. TW1
formed a visible bacterial plate when inoculated into Fe-free gradient
cultures and changed O2 profiles compared to uninoculated
controls (data not shown). However, this was not the case for TW2,
which failed to grow or alter the O2 profile in an
identical Fe(II)-free system (data not shown).
The possible effect of the growth of heterotrophic bacteria in TW1 on
the O
2/Fe(II) relationship in our experimental systems
was
tested by inoculating a culture of satellite organisms isolated
and
grown on 50% strength TSA into Fe(II)-containing gradient
systems.
After 10 days of incubation, no significant differences
between
O
2 profiles in inoculated and uninoculated control were
detected (data not shown). These results suggest that the effect
of the
heterotrophic bacteria on Fe(II) oxidation, independent
of their
possible synergistic interaction with Fe(II)-oxidizing
bacteria, was
minor.
We were concerned that deposition of oxide below the depth of
O
2 penetration in the TW1 and TW2 cultures was an artifact
caused
by the existence of a deeper oxic-anoxic boundary early in the
experiment. To test this possibility, measurements of O
2
and Fe(III)
oxide band positions were obtained at daily intervals over
6 days
in cultures inoculated with TW1. In the abiotic control systems,
the bottom boundary of the oxide band was always observed above
the
oxic-anoxic boundary. This is illustrated in Fig.
2; where
the depth of the oxide band
lower boundary and the depth of O
2 penetration are plotted
versus time; O
2 always penetrated to depths
below the lower
boundary of the oxide band (Fig
2A) in the controls.
In contrast, in
the TW1-inoculated cultures, the depth of O
2 penetration
(as defined above) was above the bottom boundary of the oxide
band at
all times except for the first two measurements (Fig.
2B). In addition,
the depth of O
2 penetration increased steadily
during the
experiment, which argues against the possibility that
the oxides were
deposited in association with an upward-retreating
O
2
front.
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FIG. 2.
Relationship between O2 penetration depth
and the base of the oxide band in control and TW1-inoculated cultures.
Data represent averages of triplicate cultures, and error bars show
95% confidence intervals.
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Iron distributions.
Fe(II) measurements obtained from the
diffusion microprobes agreed well with Ferrozine analysis at depth in
the cultures (Fig. 3). However, near the
surface, the probes indicated Fe(II) concentrations several times
higher than found by the Ferrozine analyses. This effect was likely due
to (i) the presence of heavy oxide deposits in the probe from the
control culture and to (ii) the presence of a dark green band in the
probe from the live TW1 and TW2 cultures (Fig.
4). In both cases, the dark bands were
detected by densitometric analysis of the images of the probes, leading
to erroneously high Fe(II) concentration estimates. The green band
observed in the probe from the live cultures was not observed in the
untreated probes and was likely due to the presence of soluble or
colloidal Fe(III) compounds which reacted with the potassium
ferricyanide reagent. We infer this from the fact that potassium
ferricyanide is known to form a green precipitate with Fe(III) at
neutral pH (Fig. 4, standards) and that no other components in the
medium were present in quantities sufficient to produce such a colored precipitate. Because no such bands were evident in the control cultures, the formation of soluble or colloidal Fe(III) can be attributed to bacterial activity.
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FIG. 3.
Comparison of soluble Fe(II) concentration determined by
microprobe technique and Ferrozine in control (A) and TW1-inoculated
(B) cultures. Ferrozine data are averages of triplicate cores, and
error bars represent the 95% confidence interval; they are omitted if
smaller than the size of the symbol. For the microprobes, a single
representative profile from three different measurements from a single
probe is shown.
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FIG. 4.
Photo of K3Fe(CN)6-fixed
microprobes from control and Fe(II)-oxidizing organism-inoculated
cultures shown in Fig. 1. Superimposed black bars indicate the
positioning of Fe(III) oxide bands, measured independently, in the
cultures; white bars denote oxic zones. Arrows indicate the approximate
position of the bacterial number peaks (live cultures only). Standards
were photographed under different light conditions and may not be
directly comparable with the microprobes.
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Although copious amounts of Fe(III) oxide deposits, detected by their
brownish color, were observed in probes from the sterile
control
cultures, much smaller quantities of such oxides were
evident in the
probes from the live cultures (Fig.
4). Similar
observations were
obtained consistently with both the mixed (TW1)
and pure (TW2)
cultures. As discussed further below, the lower
abundance of
particulate oxide deposits in live than in control
cultures indicates
that bacterial catalysis was the dominant mechanism
for Fe(II)
oxidation in the live cultures. In order to verify
quantitatively the
lower abundance of particulate oxides in probes
from the live cultures,
we measured the concentrations of Fe(III)
in probes as well as in cores
of whole medium in three replicate
live and control cultures. We also
examined how washing affected
Fe(III) concentrations in probes from
live cultures, which, as
mentioned above, appeared to contain soluble
and/or colloidal
Fe(III). The concentration of Fe(II) was nearly
identical in probes
and whole medium in both culture systems (Fig.
5A). The same was
true for Fe(III)
concentrations in the control systems. In contrast,
the concentration
of Fe(III) was about threefold higher in whole
medium than in probes
from the live culture. Since oxides from
the control culture did not
dissolve completely in 0.5 M HCl (discussed
below), the actual ratio is
probably higher. These findings quantitatively
confirm our consistent
visual observation of lower Fe(III) oxide
abundance in probes from live
versus control cultures.
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FIG. 5.
Fe(II) and Fe(III) in whole medium samples and diffusion
probes. (A) Comparison among probes and the medium in TW2 and
control cultures; (B) comparison between washed and unwashed
probes from TW2 and control cultures. Data represent averages of
triplicate cultures, and error bars show 95% confidence intervals.
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Washing of probes from the live culture resulted in removal of
approximately 70% of their Fe(III) content (Fig.
5B, right).
These
results verified the presence of soluble Fe(III), as indicated
by the
green band in potassium ferricyanide-fixed diffusion probes.
In
contrast, identical washing of probes from the abiotic control
cultures
resulted in no change in Fe(III) content, whereas the
amount of Fe(II)
was significantly decreased (Fig.
5B, left).
This observation is
consistent with the potassium ferricyanide
analysis and further
indicates that the presence of mobile Fe(III)
forms is a biologically
mediated phenomenon. The concentration
of Fe(III) in whole medium from
live cultures was more than twice
as high as in probes from the same
cultures (Fig.
5A). In addition,
we can attribute most of the Fe(III)
present in the probe to soluble
or colloidal Fe(III) compounds
(detected visually with potassium
ferricyanide) (Fig.
4), since washing
with anoxic distilled water
removed a large portion (ca. 70%) of the
Fe(III) from these probes
(Fig.
5B).
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DISCUSSION |
Oxygen distribution and locus of oxide deposition.
One of the
most important observations in this study is the inversion of the locus
of oxide deposition in relation to the depth of O2
penetration in biotic and abiotic opposing-gradient systems. In the
abiotic systems, the whole oxide band resided within the oxic zone
(Fig. 1A and C). However, when Fe(II)-oxidizing bacteria were present,
a significant part of the band was located below the depth of
O2 penetration, which was substantially shallower than in
abiotic control cultures (Fig. 1B and D). This observation held true
for both mixed and pure cultures. The control experiments demonstrated
that the satellite bacteria present in TW1 did not alter O2
gradients in the presence of Fe(II). Since TW2 failed to grow or alter
O2 gradients in the absence of Fe(II), we can safely assume
that in all our experiments, O2 gradients were controlled by bacterially mediated Fe(II) oxidation rather than heterotrophic metabolism.
A certain amount of Fe(III) oxide could have been deposited below the
apparent O
2 penetration boundary by means of Fe(II)
reacting with O
2 at concentrations below the detection
limit of
our electrode. However, several lines of evidence argue that
this
phenomenon had no significant confounding effect on our
observations.
First, the pronounced suboxic Fe(III) oxide deposition
observed
in the live cultures (up to 60% of the oxide band width, or
up
to approximately 4 mm, was found below the O
2
penetration boundary),
together with our conservative estimate of the
O
2 depletion depth
(see Materials and Methods), suggests
that even if O
2 was present
at concentrations below the
detection limit (<1 µM), it would
have been depleted before reaching
the bottom of the oxide band.
In addition, no suboxic deposition was
detected in abiotic cultures,
in which gradients were not as sharp as
in the live system, which
suggests that interaction between Fe(II) and
O
2 at subdetectable
concentration did not play a
significant role in suboxic deposition
of Fe(III) compounds. Finally,
the postulated presence of mobile
forms of Fe(III) (discussed below)
suggests a plausible mechanism
responsible for the observed
effects.
Previous studies in opposing-gradient systems similar to our low-iron
system demonstrated decreased O
2 penetration depth in
biotic versus abiotic cultures (
8). However, no inversion
of
the oxide band relative to O
2 gradient position was
reported.
In the study of Emerson and Moyer (
8), the use
of FeS as an
Fe(II) source provided a submillimolar equilibrium
concentration
of Fe
2+ at circumneutral pH
(
24), far lower than the concentration
of Fe(II) present
in our culture systems. This low iron content
resulted in lesser oxide
deposition over the course of the experiment
relative to that in the
experiments presented here. Consequently,
there was less material to be
observed, so oxide deposition below
the O
2 penetration zone
(if any) would probably not have been
as apparent as in our study. A
recent study of bacterial Fe(II)
oxidation by Benz et al.
(
3), in which millimolar concentrations
of soluble Fe(II)
were employed in an opposing-gradient system,
failed to demonstrate
suboxic deposition of Fe(III) alone or in
the presence of
microaerophilic nitrate-reducing, Fe(II)-oxidizing
bacteria. The
contrast between these findings and our own suggests
that suboxic
Fe(III) deposition may depend on the type of bacteria
involved in
Fe(II) oxidation
specifically, perhaps, on the organism's
ability to
produce a mobile form of Fe(III) as the initial end
product of Fe(II)
oxidation (see
below).
Biotic and abiotic oxidation.
Our original intent in deploying
the diffusion probes was to try to detect the influence of bacterial
activities on Fe(II) microgradients at the aerobic-anaerobic interface,
in a manner analogous to the analysis of the influence of
Beggiatoa spp. on H2S oxidation done by Nelson
et al. (19). Unfortunately, due to Fe(III) interference
with the densitometric measurements, the diffusion probes did not
provide satisfactory measurements of dissolved Fe(II) microgradients at
the aerobic-anaerobic interface. However, they revealed several effects
of biologically catalyzed Fe(II) oxidation which have not been
described previously.
The relative scarcity of particulate oxides in the diffusion probes
from the live cultures suggests that Fe(II) oxidation
was dominated by
biological processes. Since the 5% agar content
of the probe would be
expected to effectively exclude bacteria,
only chemical oxidation would
be expected to occur within the
agar film. Equilibrium is achieved very
rapidly in films as thin
as our probes (
h = 0.1 mm).
Assuming a diffusion coefficient (
D)
of 10
5
cm
2 s
1 for Fe(II) (
5), the
characteristic diffusion time (
h2/
D)
is only 10 s, which is essentially instantaneous on the time
scale
of a week-long experiment. The low abundance of oxide deposits
in
probes from the live cultures therefore suggests that the
Fe(II)-oxidizing
bacteria scavenged Fe(II) rapidly enough to strongly
depress Fe(II)
diffusion into the probe and subsequent abiotic
oxidation and
oxide precipitation within the
film.
The above findings suggest that the diffusion probes may provide a tool
for distinguishing quantitatively between biological
and abiotic
oxidation of Fe(II) in opposing-gradient systems.
Simply comparing
total Fe(III) oxide deposition in biotic and
abiotic systems will not
be adequate, because the limiting step
in Fe(II) oxidation is often
diffusional transport of the reduced
compounds rather then the reaction
itself (
8). However, by
applying a diffusion probe to the
system and quantifying the amount
of ferric oxide deposited per unit
volume of the probe [corrected
for the presence of soluble Fe(III)],
it should be possible to
estimate the amount of oxide which was
deposited abiotically.
The difference between the concentration of
oxide accumulated
within the probe and that in the system as a whole
will reflect
the amount of oxide deposition that was biologically
catalyzed.
Applying this approach to our live cultures, we can estimate
the
percentage of Fe(III) oxide deposited abiotically as
(Fe(III)
Pprobe)/Fe(III)
Tmedium,
where Fe(III)
Pprobe is the Fe(III) remaining in
the probe
after anoxic wash and Fe(III)
Tmedium
is the total Fe(III)
in the medium. Since Fig.
5B indicates that ca.
70% of the Fe(III)
present in the probe from the live culture is lost
during wash
and Fig.
5A shows that there was ca. 2.5 times more total
Fe(III)
in the live culture medium than in the probe, we can estimate
that ca. 90% of total Fe(III) in the live cultures was generated
via
biological processes. In fact, this fraction might be higher
in
steady-state systems, since in our situation some Fe(II) oxidation
inevitably occurred abiotically before a plate of Fe(II)-oxidizing
bacteria was established. The presence of similar Fe(III) oxide
concentrations in the probes and the whole medium in control cultures
suggested that Fe(II) oxidation and Fe(III) oxide deposition proceeded
at similar rates and by the same mechanism in both the medium
and the
probes.
Soluble Fe(III) formation.
The observation of a green
band in the live system (Fig. 4) suggested the presence of soluble
and/or colloidal forms of Fe(III). The presence of such compounds was
verified by analysis of washed and unwashed probes (Fig. 5B). Although
the pH decrease associated with Fe(II) oxidation at the
O2-Fe(II) boundary (8) might potentially account for the Fe(III) remaining in solution, in our experiments such
a decrease was far less than would be required to stabilize any
significant amount of Fe(III) (the lowest pH value observed was 6.6 in
the zone of oxide deposition, compared to 7.2 at the surface; data not
shown). It is possible that the Fe(III) was kept in solution by a
chelator excreted by the bacteria specifically for the purpose of
retarding or at least delaying cell surface encrustation with oxide
precipitates. Encrustation of the bacterial cells with particulate
oxides, leading to their eventual entombment, has been suggested as one
of the possible environmental challenges which gradient-dwelling
solid-phase oxide-producing organisms have to overcome
(8). Formation of soluble Fe(III) compounds and eventual
remote deposition of the oxides may reduce or delay such encrustation.
We hypothesize that, as soluble ferric iron complexes diffuse away from
the Fe(II)-oxidizing bacteria, they become destabilized, resulting in
precipitation of Fe(III) oxides within as well as below the zone of
O2 penetration.
Davison et al. (
5), using diffusion microprobe techniques
in lake surface sediments, identified a soluble iron peak in
their gels
at a depth of approximately 8 mm beneath the sediment-water
interface.
The authors hypothesized that this peak represented
accumulation of
Fe(II) as a result of localized Fe(III) oxide
reduction activity.
However, their analysis could not distinguish
between mobile forms of
Fe(III) and Fe(II). Our results suggest
a possible alternative
explanation, i.e., that the activity of
the iron-oxidizing bacteria
caused a local accumulation of soluble
Fe(III). Interestingly, a green
band was observed at a depth of
ca. 2 mm when a diffusion probe was
applied to a sediment core
from the freshwater wetland from which the
organisms used in this
study were obtained (not shown). A recent study,
employing voltammetric
electrodes, demonstrated the presence of soluble
organic complexes
of Fe(III) in marine surface sediments
(
25). These observations
suggest that the presence of
bacterially generated dissolved or
colloidal Fe(III) might be more
widespread than has previously
been
recognized.
Biogeochemical implications.
Measurements of Fe(III)
abundance in the diffusion probes and the whole medium showed
that the vast majority of Fe(II) was oxidized biologically in the
presence of Fe(II)-oxidizing bacteria. These results suggest that
bacteria can compete successfully with the abiotic oxidation process
and that the biological oxidation of Fe(II) might be the predominant
process leading to the formation of Fe(III) oxides in surficial aquatic
sediments. Emerson and Revsbech (10) found that organisms
from a natural Fe(III)-depositing bacterial mat accelerated Fe(II)
oxidation in a reactor system designed to simulate the in situ
conditions. Since their system was not diffusionally limited,
acceleration of Fe(II) oxidation when bacteria were present suggested a
significant involvement of those organisms in the process. Our
experiments show that Fe(II)-oxidizing bacteria could be similarly
involved in Fe(III) generation in a diffusion-limited system.
Our results indicate that Fe(II)-oxidizing bacterial activity can lead
to suboxic deposition of reactive Fe(III) oxides. An
important
implication of these findings is that they suggest the
possibility for
a rapid coupling between Fe(II) oxidation and
Fe(III) oxide reduction
within millimeters of the oxic-anoxic
interface. Furthermore, Emerson
and Revsbech (
9) found active
Fe(III)-reducing bacteria in
a natural Fe(II)-oxidizing mat in
an iron seep, indicating tight
coupling of Fe(II) oxidation and
Fe(III)
reduction.
Fe(III) produced by Fe(II)-oxidizing bacteria has traditionally been
considered to be represented by immobile, solid-phase
compounds.
Our findings suggest, however, that, at least immediately
after formation, Fe(III) compounds can be treated as soluble ions
which
are subject to diffusive transport. As mobile Fe(III) compounds
diffuse
away from the bacterial plate, they are likely to become
destabilized,
resulting in hydrolysis and precipitation of amorphous
Fe(III) oxides.
Formed in the anoxic zone, these oxides would
be immediately available
for reduction by Fe(III)-reducing bacteria,
completing the microscale
Fe redox cycle. In addition to sediment-water
interface environments,
other environments where microscale bacterial
Fe redox coupling might
occur include subsurface sediments, where
Fe(II) and O
2 may
coexist in "patchy" redox environments, forming
a dynamic network
of oxic-anoxic interfaces (
14), and the rhizosphere
of
aquatic plants, in which rapid Fe cycling is known to occur
(
12,
13,
22) and in which the presence of both Fe(II)-oxidizing
and
Fe(III)-reducing bacteria in close association with plant
roots has
recently been demonstrated (
11,
16).
| |
ACKNOWLEDGMENTS |
This research was supported by grants from the National Science
Foundation (DEB 94-7233), the U.S. Department of Energy, Office of
Energy Research, Environmental Management Science Program
(DE-FG07-96ER62321), and the School of Mines and Energy Development,
University of Alabama.
We thank D. Emerson, W. C. Ghiorse, and R. G. Wetzel for
review of an earlier version of the manuscript.
| |
FOOTNOTES |
*
Corresponding author. Mailing address: Department of
Biological Sciences, University of Alabama, Tuscaloosa, AL 35487-0206. Phone: (205) 348-0556. Fax: (205) 348-1403. E-mail:
eroden{at}bsc.as.ua.edu.
| |
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Applied and Environmental Microbiology, March 2001, p. 1328-1334, Vol. 67, No. 3
0099-2240/01/$04.00+0 DOI: 10.1128/AEM.67.3.1328-1334.2001
Copyright © 2001, American Society for Microbiology. All rights reserved.
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Top
Abstract
Introduction
