MacDonald, L. Links between the iron and sulfur cycles: gradient

Links between the iron and sulfur cycles:
gradient tube analysis
Luke MacDonald
Princeton University
Microbial Diversity 2006, MBL
Abstract:
This study asks whether iron redox cycling can occur in microxic zones when
bacterially driven iron oxidation is coupled to reduction of ferric iron via sulfide
oxidation. We employ gradient tubes bearing anaerobic reduced sulfur and/or
iron agarose plugs, overlayed with growth media exposed to oxygen to create
oxygen, sulfide, and ferrous iron gradients. Two unidentified species tend to
dominate the observed growth bands, one possible Leptothrix or Gallionella, and
one possible Thiobacillus species. The development of a double growth band in
tubes bearing both ferrous iron and excess sulfide presents an interesting
mystery, where it is unclear what electron donor and acceptor pairs are used in
either band to obtain energy, and it is unclear whether iron redox cycling occurs.
Introduction:
The elemental cycles of sulfur and iron are important biogeochemical processes
in a variety of freshwater, groundwater, and saline environments. Several key
links exist between these cycles, including the reduction of ferric oxides by
hydrogen sulfide (H2S) to solid ferrous sulfide (FeS) in anaerobic environments.
Lab studies show that at circumneutral pH iron oxidizing bacteria can oxidize
iron(II) derived from FeS(s) and FeCO3(s) to form amorphous iron(III) oxides
(Emerson and Moyer, 1993), and it is well known that hydrogen sulfide gas will
react with amorphous iron(III) oxides to form elemental sulfur and FeS.
Therefore, iron oxidizing bacteria should posses the ability to continuously cycle
a finite Fe(II) source in the presence of sulfide (Figure 1), but to our knowledge
no study demonstrates such cycling.
One obstacle to a complete understanding of the links between sulfur and iron
cycles is a lack of information on iron oxidizing organisms, which is currently
insufficient to predict how the sulfur and iron cycles interact in natural microbial
communities. For example, it is not clear whether some of the so-called iron
oxidizers commonly found near Fe(II) seeps, e.g. Leptothrix, directly oxidize iron
chemolithoautotrophically or whether they derive energy from degrading humics,
thereby liberating Fe(II) for abiotic iron oxidation1. Consequently, we cannot
predict what effect sulfide driven iron reduction will have on the redox state of
iron. If Leptothrix mediate iron oxidation for metabolism, then sulfide driven iron
reduction could result in iron cycling from Fe(II) to Fe(III), transferring electrons to
biota, but if iron oxidation is an incidental result of humic degrading metabolism
then sulfide driven iron reduction will most likely result in Fe(II) as a terminal
electron acceptor. By collecting field samples and exploring the links between
iron and sulfur cycling in gradient tube enrichments we can expand our
understanding of iron oxidizing organisms.
Gradient tube microcosms offer a useful way of examining the links between iron
and sulfur cycles in microoxic conditions and to enrich for potentially novel iron
oxidizing organisms (e.g. Emerson and Moyer, 1993). These tubes consist of
anaerobic agarose plugs containing Fe(II) and/or sulfide, overlayed with slightly
viscous growth media exposed to oxygen. As oxygen diffuses down and Fe(II)
and/or sulfide diffuses upwards, bacteria existing within regions of overlap
between the two zones oxidize either iron or sulfide under microoxic conditions
more efficiently than abiotic oxidation. The aim of this project is to explore the
possibility of dissolved sulfide and iron-oxidizing bacteria mediating iron redox
cycling at the aerobic/anaerobic interface, and to identify iron-oxidizing
organisms inside gradient tube enrichments.
Materials and Methods:
Field sampling site: A bacterial mat on School Street in Woods Hole, MA, was sampled on three
separate occasions. This mat is characterized by high concentrations of flocculent iron,
overlayed with a purple to blue floating crystalline sheets that may be oxidized manganese
precipitates. The mat extends from a swampy area onto a paved road, where Fe(II) seeping out
of the groundwater oxidizes upon contact with the air. All samples were taken at the interface of
the vegetated and paved zon; milky bacterial mats were visible to the naked eye at this location.
Gradient tubes: 16x75 mm culture tubes (Fisher) were filled with three plugs of agarose and
nutrients (Figure 7). The bottom 0.5 mL consisted of 10 mM Na2S dissolved in a modified Wolfe’s
mineral media (MWMM) (Hanert, 1999) and mixed to 2% wt/v agarose; the middle 0.5 mL
consisted of saturated ferrous sulfide prepared according to (Hanert, 1999), mixed 1:1 with
MWMM and added to 2% wt/vol final concentration agarose; the top 2.5 mL consisted of MWMM
with vitamins and minerals added, buffered in bicarbonate, with 0.15% wt/v. Chemical control
tubes substituted one or both of the bottom 0.5 mL plugs for 2% wt/v agarose in MWMM. All
media was made and kept anaerobic until inoculation. Tubes were inoculated by inserting a pipet
tip to a consistent depth into the top agar and expelling 15 µL of field sample into the agar as the
tip was withdrawn.
Recent research demonstrates that Leptothrix species express an iron oxidizing protein, which
tends to support the argument that these organisms oxidize iron to meet their energy
requirements (Corstjens, et al, 1992.). However, finding a protein does not definitively
demonstrate that Leptothrix commonly oxidizes iron in the environment for metabolism, and, to
our knowledge, there have been no surveys of the abundance of this protein in sediments. Other
methods of obtaining energy may be far more favorable and more commonly used.
1
Microelectrodes: Unisense pH, O2, and H2S electrodes were used to obtain concentration profiles
as a function of depth. The probes were mounted on a stage and manipulated remotely via the
MC-232 micromanipulator connected to a PC running the Profix software by Unisense. The O2
probe was calibrated separately for each type of tube at 0% by extended the tip into the deepest
part of the tube where it was clearly anoxic, and at 100% by setting the tip only ~1µM below the
surface. The H2S probe was calibrated in H2S free water and in 6.67 mM sulfide kept at pH 4,
anaerobically stored in titanium citrate as a reducing agent under nitrogen gas. Because the
gradient tubes had an opening too narrow for the reference electrode to fit through, a Pasteur
pipette was bent to insert into the sample and filled with 2% agar. This allows for an electrical
connection between the reference electrode and the sample and accurate pH readings.
Microscopy: Samples were withdrawn from the visible growth bands inside the tubes using sterile
Pasteur pipettes, wet mounted, viewed by under a Zeiss microscope, and photographed with a
Zeiss auxiocam mounted into the scope.
DAPI: Samples from the field and from the growh bands inside gradient tubes were filtered under
low vacuum (<5 mm Hg) onto a 0.2 µm filter and stained with DAPI dissolved in mounting media.
Clone Library: 16S DNA sequences from field samples were PCR amplified, cloned into a
plasmid vector, transformed into E. coli (Invitrogen TOPO TA Cloning Kit), and plated onto LB
plates containing 50 µg/ml kanamycin, 40 µg/ml x-Gal. Successful transformations formed white
colonies, and 192 colonies were picked for sequencing. Of these, 158 had a quality index above
59 in 4-peaks software, and these were aligned in ARB with the greengenes PT server.
Results:
Visible growth bands appeared in the gradient tubes 7-8 days after inoculation
(Figure 8). Single bands grew in the tubes bearing only Na2S or FeS, while two
bands arose in tubes containing both Na2S and FeS separated by about 1/2 mm.
The oxygen profiles dropped to zero at the depth of all bands first encountered
(Figures 2-4). The sulfide profiles dropped to zero above the band in tubes
bearing only Na2S (Figure 5), and above the top band in tubes bearing both Na2S
and FeS (graphs not shown). pH remained stable throughout the tubes, varying
between 7 and 8 (Figure 6).
The slope of the oxygen profiles changed with time, such that after growth was
initiated in tubes with ferrous iron plugs the oxygen levels gradually dropped off
until they reached zero at the top of the growth band (Figures 2-4). In tubes
bearing only Na2S the growth band appeared nearer the top of the tube than in
any other tube type. In all cases, the oxygen profile dropped off more slowly in
abiotic tubes than in inoculated tubes, although the abiotic Na2S tubes consumed
oxygen more effectively than any of the inoculated tubes except inoculated Na2S
tubes. Orange to red Fe(III) was visibly precipitated above the growth in FeSonly tubes, while such visible iron oxidation was absent from FeS+Na2S tubes.
Organisms found in the growth bands of tubes bearing only FeS as an electron
donor include long filamentous bacteria that cluster with solid precipitates, which
are most likely iron(III) oxides. These filaments appear empty under light
microscopy (Figure 9), but DAPI staining reveals cellular structure within each
filament (Figure 10). Similar filamentous bacteria are found in the lower band of
the gradient tubes containing both FeS and Na2S, and are absent from the top
growth band. Organisms within the top growth band include a large number of
short, curved rods that exhibit a hopping movement in a wet mount (Figure 11).
There is a wide variety of other morphotypes in all tubes, ranging from clusters of
spherical cells to spirilla shaped organisms.
Discussion:
This study raises a number of important questions that remain unanswered,
namely what organisms are growing in the growth bands and what redox
reactions do they mediate? In tubes with only FeS or Na2S the possible redox
reactions are limited to only a few options. Given that oxygen and sulfide profiles
drop to nearly zero at the growth bands, tubes bearing Na2S plugs alone almost
certainly undergo H2S2 oxidation by oxygen, which may or may not be
biologically mediated due to the fast kinetics of chemical oxidation of H2S. If that
reaction is not biologically mediated then another way for microbes to gain
energy is to use sulfate as the electron acceptor, present in the media in low
levels, and dissolved organic carbon (DOC) carried into the tubes from the
inoculation as the electron donor. DOC is the best donor here, albeit present in
low concentrations relative to H2S. Although we do not expect a sulfate profile in
the top agar, and hence we expect homogenous growth, not bands, we expect a
DOC gradient that decreases with distance from the inoculation point, which may
lead to growth bands.
In tubes bearing FeS plugs alone, the potential electron donors near the growth
band are Fe(II) and DOC, while sulfate and O2 are the sole acceptors. Since O2
is thermodynamically a much better acceptor, I expect that the organisms within
the growth band directly consume O2. However, the actual growth may be off of
organic carbon carried with the inoculation. A reddish zone containing iron(III)
oxides appears just above the growth band, but this oxidized iron may be the
product of chemical oxidation as Fe(II) diffuses up from the bottom and not
directly due to biological activity.
In tubes bearing both FeS and Na2S we see two growth bands, and here the
electron donor/acceptor in the respective bands are not as apparent as tubes
bearing only one electron donor plug. In the top band, the electron acceptor is
oxygen, as shown by the oxygen profiles, which drop off at the growth band
(Figure 2). The donor, however, may be H2S, Fe(II), or DOC. and Fe(II).
Therefore, it seems likely that H2S is oxidized in the top growth band, although
2
Hereafter, H2S is used to denote both H2S and HS- sulfide species.
we must apply same considerations of the biological reaction discussed in the
section above on strict Na2S tubes. Fe(II) may be oxidized here, although there
are no orange colorations in this gradient tube.
The bottom band lacks good candidates for electron acceptors or donors,
however, and this mystery stems from an incomplete sampling of the chemistry
within the gradient tubes. Fe(II) and DOC are, again, potential donors, but it is
unlikely that Fe(II) is the donor, since we see no orange coloration near this band
and DOC is present in very small amounts, especially below the top growth band,
and consequently also seems an unlikely donor. Since oxygen is consumed in
the top band, then only sulfate is left within the media to act as an electron
acceptor.
One possible explanation is that the top band exhibits biotic or abiotic iron
oxidation from Fe(II) diffusing upwards from the plug, and ferric iron diffuses
downwards to act as an electron acceptor for sulfide oxidation. Indeed, this is
the reaction that this project set out to discover (Figure 1), but the lack of orange
precipitates near either growth band indicates that Fe(III) may not, in fact, be
generated in the tubes.
To resolve the issue and determine which redox reactions occur in the double
growth bands observed in tubes bearing FeS and Na2S plugs we need more
detailed chemical profiles. The H2S profile was inconclusive; due to an unknown
problem with the sulfide H2S was not detected in the top agar. Fe(II) and Fe(III)
measurements were not obtained near the growth bands, and these
measurements would greatly facilitate the redox analysis. Another approach to
resolve the issue would be to limit the DOC introduced into the tubes by spinning
and diluting the inoculum prior to injection.
The second major unanswered question asks which organisms are present in the
media. Filamentous bacteria observed in the light microscope to be associated
with some metallic precipitate were also observed with DAPI to possess cells
inside the sheath. These bacteria grew to large numbers in the growth bands of
FeS tubes, and in the bottom growth bands of FeS and Na2S tubes.
Morphologically, they could be either Leptothrix or Gallionella species. Serial
dilutions and transfer may help to isolate and sequence these filamentous
organisms to positively identify them, but initial attempts in this study were
unsuccessful (methods not presented here).
In the top band of the double banded tubes and in the Na2S alone tubes the
short, curved rods with bright spots on one end may be Thiobacillus species,
capable of oxidizing both iron and sulfide, although only a few species are
capable of growing at circumneutral pH. Again, isolation and sequencing are
appropriate techniques to positively identify the organisms.
Conclusions:
Double banding in gradient tubes bearing both FeS and Na2S presents a
puzzling redox mystery, the solution to which may change the way we think
about iron and sulfur cycling. Determining where H2S is oxidized and whether
iron cycles between redox states by migrating from band to band will help us to
piece together the connections between the iron and sulfur cycles as we might
see them in the environment. To solve this riddle, we need a complete
characterization of the bacteria and redox pairs involved, and, with luck, this will
expand our currently limited knowledge of how these processes couple in the
natural environment.
Figure 1: Geochemical cycling between iron and sulfur. In this diagram, we see that
oxygen can drive both sulfide and iron oxidation, and that sulfide will reduce oxidized
iron. All three of these reactions occur spontaneously abiotically, but may be bacterially
driven under microoxic conditions, since under low oxygen levels enzyme catalyzed
sulfide and iron oxidation can occur more rapidly than abiotic oxidation.
FeS and Na2S
1.5 days
2.5 days
7 days
%Oxygen
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Figure 2 Oxygen profile for gradient tube with FeS and Na2S : Oxygen diffuses into
the tube during days 1.5 and 2.5, and by day 7 drops to zero at the top of the growth
band.
Na2S tubes
1.5 days
2.5 days
7 days
%Oxygen
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Figure 3 Oxygen profile for gradient tube with Na2S alone: Oxygen diffuses in at day
1.5, at day 2.5 migrates downward, but by day 7 has migrated upwards and drops to zero
at the top of the growth band.
FeS tubes
0
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40 %Oxygen
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Figure 4 Oxygen profile for a gradient tube with FeS alone: Oxygen diffuses inwards
and at day 7 is completely consumed at the top of the growth band.
Na2S
1.5 days
7 days
mM Sulfide
0.0
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Figure 5 Sulfide profile in tube bearing Na2S alone: Sulfide levels are zero above the
growth band and increase moving down into the tube.
FeS
FeS Na2S"
Na2S
FeS Blank
Fes Na2S blank
Na2S Blank
pH at 7 days
4.0
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Figure 6 pH profiles: The pH is stable in all tubes, varying between 7 and 8.
Figure 7 Gradient tube setup: The bottom white zone contains sodium sulfide in 2%
agarose, the middle black zone contains FeS in 2% agarose, and the top portion is growth
media in 0.15% agarose.
Figure 8, comparing FeS alone gradient tubes at day 8. The inoculated tube on the left
displays a visible growth band and iron oxidation zone halfway up the tube, while the
uninoculated tube on the right lacks a growth band and displays an iron oxidation zone
near the bottom of the tube.
Figure 9, Light microscopy of samples from FeS bearing tube: Long filments
associated with iron oxides dominate the community in these samples.
Figure 10, DAPI staining from an FeS gradient tube: DAPI reveals cells within the
long filamentous sheaths associated with iron oxides.
Figure 11, light microscopy of sample from Na2S gradient tube: Short curved rods
dominate the microbiota in these samples. Notice that there are bright spots at the ends
of some these microbes, whose function is unclear.
References
Emerson, D., and C. Moyer. 1997. Isolation and characterization of novel ironoxidizing bacteria that grow at circumneutral pH. Appl. Environ. Microbiol.
63:4784-4792
P L Corstjens, J P de Vrind, P Westbroek, and E W de Vrind-de Jong. 1992.
Enzymatic iron oxidation by Leptothrix discophora: identification of an ironoxidizing protein. Appl Environ Microbiol. February; 58(2): 450–454.
Hanert H. H. 1999. The genus Gallionella In: Starr, M. P., Stolp, H., Trüper, H.
G., Balows, A., and Schlegel, H. G (ed.), The prokaryotes. Berlin, SpringerVerlag