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Studies on the influence of bacteria and carbon source on the products of dissimilatory iron reduction

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Studies on the influence of bacteria and carbon source on the products of dissimilatory iron reduction

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Content STUDIES ON THE INFLUENCE OF BACTERIA AND CARBON SOURCE ON
THE PRODUCTS OF DISSIMILATORY IRON REDUCTION
by
Everett Cossio Salas
A Dissertation Presented to the
FACULTY OF THE GRADUATE SCHOOL
UNIVERSITY OF SOUTHERN CALIFORNIA
In Partial Fulfi llment of the
Requirements for the Degree
DOCTOR OF PHILOSOPHY
(GEOLOGICAL SCIENCES)
December 2008
Copyright 2008 Everett Cossio Salas
ii

Nothing can take the place of persistence. Talent will not; nothing is more common
than unsuccessful men with great talent. Genius will not; unrewarded genius is almost
a proverb. Education will not; the world is full of educated derelicts. Persistence and
determination alone are omnipotent.
- Calvin Coolidge
EPIGRAPH
iii
DEDICATION
And so the dark night of the soul is at long last over, giving way to a dawn full
of opportunity and possibility. This long, often diffi cult, often invigorating, and yes,
sometimes seemingly quixotic journey could not have been undertaken nor completed
without the help of several individuals, who either by providence or fortuitousness,
where present at critical junctures along this journey. From those days of wandering
through the undergraduate desert at Cal Poly Pomona, where I seemingly did my best
to become academically disqualifi ed every quarter, and often wondered what the point
of it all was, to my haphazard ride through a Master’s program at the Keck School of
Medicine, where I was at best an underwhelming and undistinguished student, to this
Ph.D. program, where I struggled constantly to overcome years of self-doubt, I have
been aided by professors and advisors who, for whatever reason, thought I was worth
the effort and not beyond help.
To my family: Everardo, Rosamaria and Brenda. You helped shape me and gave
me the tenacity and stubbornness that has kept me going. It was your support and
encouragement that has counter-balanced those ever-present feelings that I could not
cut it at this level. I look at your experiences and feel a debt of gratitude. You left
home and came to this strange place, with a different culture and a different language.
Although you did not have much, you left it behind and started over; you sacrifi ced,
worked multiple jobs and held high expectations for us, so that we could have
opportunities you could not even dream of. Knowing where you started, and knowing
where you are today, if I can accomplish a fraction of what you have, I will be wildly
successful.
iv
To my wife, Liliana: I lack the diction to express my love and affection in words that
do you justice. This effort has been fi lled with emotional hardships, of which you
have borne the brunt. You have endured the long nights, my constant preoccupation
with research and my stress-induced short temper and detachment. For this I can
only say thank you. Thank you for being there, and thank you for putting up with the
disappointments and frustrations that come with being married to someone whose
thoughts are always preoccupied with science. As we start this next phase of our
journey, I hope you will continue to show patience as I attempt to gain perspective
and balance in life.
v
ACKNOWLEDGEMENTS

I would like to thank my committee: Kenneth Nealson, William Berelson, Steve
Finkel, Doug Hammond, Frank Corsetti of USC, and Tony Kampf of the Natural
History Museum of Los Angeles County. Thank you for your patience as I learned
how to think and act like a scientist and for providing me with the guidance that
has allowed me to successfully earn this degree. I would also like to thank Rohit
Bhartia, Lonne Lane, Bill Abbey, Sasha Tsapin, Bill Hug and Ray Reid of the Jet
Propulsion Laboratory and Photon Systems, Inc. for all of the support during those
turbulent years at JPL. Finally, I would like to acknowledge my fellow graduate
students, particularly Lisa Collins, Tim Riedel and Orianna Bretschger, for all of your
constructive critisism and insight throughout my time at USC.
vi
TABLE OF CONTENTS
Epigraph ii
Dedication iii
Acknowledgements v
List of Tables vii
List of Figures ix
Abstract xi
Chapter One: Iron Oxides and Dissimilatory Iron Reducers 1
Chapter Two: The Infl uence of Bacterial Strain Specifi city on Secondary
Iron Mineral Formation 23
Chapter Three: The Impact of Bacteria and Carbon Source on the
Formation of Reduced Iron Minerals 48
Chapter Four: How Important Are Shewanellae to the Cycling of Iron and
Organic Carbon in Sediments?
82
Bibliography 109
Appendix A: Aerobic Growth Curves and O
2
Respiration Rates for
Shewanella oneidensis MR-1 Using Different Carbon Sources 128
Appendix B: Thermodynamic Data Used for Stability Diagram
Calculations
135
Appendix C: Iron Reduction and Carbon Consumption Data Sets 139
Appendix D: Susceptibility Values for the Products of Ferric Iron
Reduction Coupled to Lactate Oxidation
163

vii
LIST OF TABLES
Table 2.1 Bacterial strains used in dissimilatory iron reduction
experiments
26
Table 2.2 Medium Composition 27
Table 2.3 Summary of iron reduction rates and biomineral distribution 38
Table 3.1 Medium Composition 51
Table 3.2 Organic Carbon Sources 52
Table 3.3 Mineral phases associated with each organic carbon source 59
Table 3.4 Stoichiometry for reactions with lactate, pyruvate and uridine 71
Table 4.1 Bacterial strains used in respiration rate calculations 93
Table 4.2 Fe(III) reduction rates for Shewanella strains 95
Table 4.3 Fe(III) Reduction rates for some sedimentary environments 96
Table B1 Equations used to construct stability diagrams 135
Table B2 Thermodynamic Constants Used in Calculations 136
Table C1 Lactate oxidation and Ferric Iron Reduction by Shewanella
putrefaciens W3-18-1
138
Table C2 Pyruvate oxidation and Ferric Iron Reduction by Shewanella
putrefaciens W3-18-1
141
Table C3 Uridine oxidation and Ferric Iron Reduction by Shewanella
putrefaciens W3-18-1
145
Table C4 Lactate oxidation and Ferric Iron Reduction by Shewanella
putrefaciens CN32
148
Table C5 Pyruvate oxidation and Ferric Iron Reduction by Shewanella
putrefaciens CN32
151
Table C6 Uridine oxidation and Ferric Iron Reduction by Shewanella
putrefaciens CN32
153
viii
Table C7 Lactate oxidation and Ferric Iron Reduction by Shewanella
oneidensiss MR-4
155
Table C8 Pyruvate oxidation and Ferric Iron Reduction by Shewanella
oneidensiss MR-4
158
Table D1 Shewanella putrefaciens W3-18-1 Susceptibility 162
Table D2 Shewanella putrefaciens CN32 Susceptibility 164
Table D3 Shewanella oneidensis MR-4 Susceptibility 166

ix
LIST OF FIGURES
Figure 1.1 Conceptual Model of Iron Biomineralization 12
Figure 2.1 Environmental scanning electron micrographs of reduced iron
oxide biominerals
32
Figure 2.2 XRD patterns for bominerals and standards. 33
Figure 2.3 Total HCl-extractable reduced iron-oxide produced by the
tested Shewanella strains using HFO as the electron acceptor
and lactate as the electron donor.
34
Figure 2.4 Concentrations of reduced aqueous phase iron. 35
Figure 2.5 Semi-log plot of susceptibility values 36
Figure 2.6 Box plot of solid phase inorganic carbon 37
Figure 2.7 Thermodynamic stability fi eld of hydrous ferric oxide as a
function of Fe
2+
concentration and pH
40
Figure 2.8 Thermodynamic stability fi eld of hydrous ferric oxide as a
function of Fe
2+
concentration and HCO
3
-
concentration
42
Figure 3.1
Environmental scanning electron micrographs of reduced iron
oxide biominerals produced by strain W3-18-1
57
Figure 3.2
Siderite concretions (arrow) produced by W3-18-1 when incu-
bated with pyruvate
58
Figure 3.3
Iron oxide reduction through time by strain W3-18-1 using
various carbon sources
60
Figure 3.4
XRD patterns for biominerals and standards using lactate and
uridine
61
Figure 3.5 XRD pattern for pyruvate incubation experiments 62
Figure 3.6 Aqueous phase Fe
2+
production by strain W3-18-1 over time 63
Figure 3.7
Consumption of organic carbon and production of metabolic
byproducts using hydrous ferric oxide as the electron acceptor
65
Figure 3.8 pH of cultures over time 66
x
Figure 3.9
Inorganic carbon production by strain W3-18-1 using three dif-
ferent carbon sources
67
Figure 3.10
Thermodynamic stability fi eld of hydrous ferric oxide as a
function of Fe
2+
concentration and pH
68
Figure 4.1
log-log plot of ferric iron respiration rates for Shewanella
strains using lactate as the carbon source
97
Figure A1
Aerobic exponential growth curves for MR-1 using different
carbon sources
129
Figure A2
Figure A2. Growth curves for MR-1 using different carbon
sources
130
Figure A3
Average oxygen respiration rates per cell through exponential
growth 133

xi
ABSTRACT
The aim of my work is to elucidate the interactions between microbes and minerals.
The work I present here describes studies I have done investigating factors that can
infl uence the identity of reduced iron oxide minerals.
There is no doubt that microbes play an important role in the redox cycling of iron
oxides in soils and sediments. The microbially mediated reduction of iron oxides can
lead to the remobilization of iron as well as the formation of reduced iron minerals.
Beyond acting as a source of Fe2+, the bacteria themselves are thought not to play a
role with respect to the nature of the resulting reduced iron minerals. Several factors,
such as the carbon substrate used for energy, the strain of bacteria used in experiments
and the rates at which the bacteria reduced iron, were investigated under laboratory
conditions to determine their importance in iron mineral formation.
Shewanella putrefaciens CN32, Shewanella putrefaciens W3-18-1 and Shewanella
oneidensis MR-4 all produced magnetite when given lactate as the carbon source.
However, they differed in the amount of magnetite each produced. Strain W3-18-1
also produced a mixture of magnetite and iron carbonate hydroxide, although most of
the reduced iron remained in an amorphous phase. When strain W3-18-1 was given
pyruvate, the major biomineral was siderite, while incubations with uridine produced
pure iron carbonate hydroxide. Although the bacteria reduced iron at comparable
rates, the products of this reduction were all quite different, suggesting that rate, while
important, is not the primary determinant in what phases of iron minerals will be pro-
xii
duced in laboratory studies. The results presented here suggest that chemistry appears
to be the important factor in determining the identity of the resultant iron minerals.
However, the chemistry itself was extensively altered by the metabolic activity of the
bacteria, making them much instrumental to iron mineral formation than merely act-
ing as a source of reduced iron.
1
CHAPTER ONE
IRON OXIDES AND DISSIMILATORY IRON REDUCERS
1.1 Iron Minerals and Iron Redox Chemistry
1.1.1 Distribution of Iron Containing Minerals
Iron (Fe) is an element of great biogeochemical importance. On the earth's surface, it
is generally found in two valence states, ferrous (Fe
2+
) and ferric (Fe
3+
), The reduced
form, Fe
2+
, is usually found in suboxic environments, and is rapidly oxidized at pH
> 5, forming insoluble iron oxides and oxyhydroxides. Fe
2+
is a component of many
minerals including pyrite (FeS
2
), vivianite (Fe(PO
4
)2) and siderite (FeCO
3
). It is also
found in olivines (Fe
2
SiO
4
) and pyroxenes (FeCaSi
2
O
6
, MgFeSi
2
O
6
, Fe
2
Si
2
O
6
), the
major components of peridotites. Both Fe
2+
and Fe
3+
are found in the mixed oxide
magnetite (Fe
3
O
4
). Fe
3+
is also found in several iron oxides and hydroxides such as
hematite (Fe
2
O
3
), goethite ( α-FeO(OH)), akageneite ( β-FeO(OH)) and ferrihydrite
(HFO), in addition to several iron-rich clays.
Under standard temperature and pressure, siderite formation requires near neutral pH,
low Eh, a high concentration of dissolved carbonate, a high Fe
2+
/Ca
2+
ratio and low
dissolved sulfi de concentrations (1). However, siderite concretions have been reported
from high sulfate reduction environments, suggesting that siderite can be formed in
the presence of sulfi de provided the production of Fe
2+
exceeds the production of
2
sulfi de (2). It has been proposed that siderite formation requires biological mediation
to provide both Fe
2+
and carbonate via the oxidation of organic carbon coupled to dis-
similatory iron reduction (3, 4). However, other studies have demonstrated the possi-
bility of siderite formation via abiotic means (5, 6).
Favorable conditions for the precipitation of vivianite include high concentrations of
phosphate, high ferrous iron concentrations, low Eh and low sulfi de concentrations
(1, 7). Vivianite is widespread in lacustrine environments and environments where the
concentrations of sulfate and sulfi de are generally low (1, 7, 8), although the origin
and conditions of its formation are not well understood (7).
Magnetite is a ubiquitous feature in almost all metamorphic and igneous rocks, typi-
cally occuring as millimeter-sized grains and comprising less than 5% of the host
rock (9-13). Magnetite formation in low temperature settings has been described as
occurring during the breakdown of smectite (14), oxidation of pyrites (15, 16) and at
the sediment-water interface by magnetotactic bacteria (17-19). Although magnetite
is a common product of laboratory studies with dissimilatory iron reducing bacte-
ria, (20-24), very few examples exist in sedimentary environments in which there is
strong evidence for a link between iron reducing bacteria and the presence of magne-
tite (25, 26).
Iron oxides are present in almost all environments (27) and can be found as poorly or-
dered minerals like green-rust and ferrihydrite (HFO) or more crystalline forms such
as goethite, hematite, magnetite or lepidocrosite (28). Ferrihydrite occurs in soils
3
undergoing rapid weathering and which contain soluble silicate and organic acids that
can inhibit the formation of more crystalline iron oxide forms (29-32). Iron hydrox-
ides are readily dehydroxylated to their oxide counterparts. Both forms are highly
stable, although under appropriate environmental conditions, almost every iron oxide
can undergo conversion into at least two other phases (27).
1.1.2 Iron Redox Chemistry
Iron can exist in redox states that allow it to easily transform under conditions found
in natural sedimentary and aquatic environments (33-36). In sediments and soils,
the availability of Fe
3+
frequently exceeds that of other electron acceptors, such as
oxygen, nitrate and sulfate (28, 37-42). The redox cycling of iron can exert a strong
infl uence on the behavior of organic and inorganic compounds in anoxic sedimentary
environments (35). Under anoxic conditions, microbes may oxidize organic com-
pounds via reduction of Fe
3+
. The reduced iron can then become reoxidized by atmo-
spheric oxygen, or some biotic process, making it available for further organic matter
oxidation.
Under certain conditions, higher oxidation states of iron can be formed, such as Fe
4+
,
Fe
5+
and Fe
6+
(43-45). Fe
4+
and Fe
5+
are unstable and quickly dismutate to Fe
3+
and
Fe6+ (46). However, certain Fe4+ and Fe5+ porphyrins are known to play important
roles in biological hydroxylation reactions (43, 44, 47). Fe
6+
salts have been known
for well over a century and are very strong oxidants, although they are not stable un-
4
der present conditions found on earth’s surface (46, 48, 49). Fe
6+
can, however, exist
in solution under high alkaline, high pH (>10) conditions (46).
The importance of solid phase Fe
3+
species as electron acceptors for microbial respi-
ration in anaerobic sediments is well established (50, 51). Iron reducers inhabit both
freshwater and marine sediments and can have a signifi cant impact on the cycling of
carbon in these sediments (50-52). As such, an extensive amount of work has been
done by several researchers investigating the degree to which dissimilatory Fe
3+
re-
ducers are able to couple carbon source utilization to the redox cycling of iron phases
found in sediments. There has been considerable interest in understanding the factors
that may play a role in controlling the rate and extent of Fe
3+
reduction in sediments.
Several factors have been shown to infl uence the rate, extent and products of iron
reduction, including solution chemistry, iron oxide surface area, electron donor and
acceptor ratios, adsorption of reduction products and co-precipitated ion activity.
1.1.3 Secondary Mineral Formation
While a fair amount of work has been done investigating the rate of Fe
3+
reduction
by both Shewanella and Geobacter strains, the role that reduction kinetics may play
in determining the kinds of minerals formed as a result of dissimilatory iron reduc-
tion is not fully understood. Researchers have postulated that the amount of Fe
2+
in
a system during the early stages of reduction can have an impact on the subsequent
mineralization of iron. Hansel, et al. (22) reported that in the presence of ferrihydrite,
initial concentrations of Fe
2+
may have an infl uence on magnetite formation. Low
5
initial concentrations of Fe
2+
can promote goethite accumulation and inhibit magnetite
precipitation, even if Fe
2+
concentrations increase at a later time. Zachara et al. (24)
postulate that the primary factor controlling the nature of secondary minerals appears
to be the Fe
2+
supply rate and magnitude of Fe
2+
produced. However, experiments cor-
relating Fe
2+
production rates per cell to the physical properties of iron mineral prod-
ucts have not been done. Because the type of secondary minerals produced by dis-
similatory iron reduction will have an impact on subsequent biogeochemical cycling,
it is important to understand the degree to which iron reduction kinetics affect mineral
dissolution and re-precipitation as ferrous iron diffuses throughout the sedimentary
environment.
1.2 Dissimlatory Iron Reducers
1.2.1 Shewanella and Geobacter
It has been recognized that bacteria play an important role in iron cycling and as such,
extensive work has been done by numerous researchers attempting to understand the
nature of microbially mediated Fe
3+
reduction. While iron reducing organisms have
been found within several groups of bacteria, the majority of this work has been done
using strains from two groups within the delta and gamma proteobacteria: Geobacter
(Geobacteraceae) and Shewanella, respectively (51-53). Geobacter metallireducens
strain GS-15 was isolated from iron-rich sediments of the Potomac River (54). This
strain reduces iron using acetate as an electron donor, making it a potentially impor-
tant organism in anoxic environments where acetate is considered to be a key inter-
6
mediate in the degradation of complex organic matter. Little is currently known about
the genes required for dissimilatory iron reduction in Geobacter species. However,
strain GS-15 possesses two genes, ppcA and omcB, that code for a periplasmic and
membrane bound c-type cytochrome, respectively, and both of these genes have been
shown to be required for Fe
3+
reduction in GS-15 (55, 56).
The Shewanellae genus encompasses several strains that have been shown to be
capable of dissimilatory Fe
3+
reduction. Shewanella are gram-negative, facultative
anaerobes belonging to the γ-proteobacteria. They have a rapid generation time under
aerobic conditions, making them ideal for genetic and physiological studies. Although
Shewanella species are thought to be incapable of acetate utilization under anaero-
bic conditions, they are able to use a myriad of other carbon compounds including
lactate, pyruvate, some sugars, amino acids and nucleic acids. They are also very
versatile with respect to the number of electron acceptors they can utilize. In addition
to being able to use oxygen, nitrate, nitrite, thiosulfate, trimethylamine N-oxide and
fumarate, these bacteria can use a wide range of metal oxides and metalloids such as
As, Co, Cr, Fe, Mn, Tc, U and V when given lactate as a carbon source (42, 57-63).
They have also been shown to posses the capacity for metal reduction using hydrogen
as an electron donor (24, 64).
A set of genes has been identifi ed as being involved in the metal reduction pathway
of Shewanella (65-70). Of these, omcA and omcB code for outer membrane decaheme
c-type cytochromes (65, 68). However, sequence analysis suggests that these proteins
are not related (53). OmcB mutants have been shown to be defective in solid phase
7
Fe
3+
and manganese reduction (67) while omcA mutants appear to be defective in Mn
reduction (68, 71).
S. oneidensis strain MR-1 has been used most extensively for genetic characteriza-
tion of Fe
3+
reduction. Strain MR-1 was isolated from anaerobic sediments in Oneida
Lake, New York, and was fi rst reported as having the ability to couple anaerobic
growth to the reduction of manganese dioxide (72). It has since been reported to have
the ability to couple anaerobic growth to a variety of electron acceptors: Fe
3+
, nitrate,
nitrite, sulfate, Cr(VI), U(VI) and fumarate, among others (42, 51, 57, 58, 61). Strain
MR-1 is also diverse with respect to the range of carbon sources it is able to utilize. In
addition to lactate, strain MR-1 is able to aerobically respire acetate, N-acetyl-D-glu-
cosamine, α-keto-butyrate, glutamate, uridine, adenosine, propionate, inosine, pyru-
vate, methyl-pyruvate, leucine and isoleucine, among others. Under anaerobic con-
ditions, the variety of carbon sources available to MR-1 is not as great. The carbon
source can also infl uence the ability of strain MR-1 to use different electron accep-
tors. For instance, MR-1 can couple reduction of Co(III) to oxidation of lactate, but
not to the oxidation of N-acetyl-glucosamine (Obraztsova, pers. comm.). Strain MR-1
can grow at a pH range of 6.3 to 9, and can tolerate temperature ranges between 3
0
C
and 37
0
C. Optimal growth occurs between 24
0
C and 37
0
C, although comparable
population densities can be reached at colder temperatures over longer periods of time
(73). The ability of MR-1 to reduce metal oxides (including iron) has been attributed
to the presence of several decaheme c-type cytochromes (53, 74), two of which are
high molecular weight c-type cytochromes located on the outer membrane (57, 68).
omcA encodes an 83kDa decaheme c-type cytochrome that has been localized to the
8
outer membrane and has been found to be a lipoprotein (75). A second outer-mem-
brane decaheme c-type cytochrome, OmcB, was also found to be a lipoprotein (76).
Gene disruption studies on omcA and omcB indicate that the cytochromes encoded by
both genes play a role in metal oxide reduction (67, 68).
A third decaheme c-type cytochrome, MtrB, was found to localize in the outer mem-
brane (66). MR-1 mutants lacking the mtrB gene show a diminished capacity for the
reduction of both soluble and solid phase Fe
3+
as well as MnO
2
(66, 68). The outer
membrane cytochromes OmcA and OmcB are localized to the periplasmic space in
these mutants, implying that MtrB may have some role as a scaffold or in the local-
ization of OmcA and OmcB to the outer membrane. mtrA codes for a periplasmic
40 kDa decaheme cytochrome c that is associated with MtrB. Mutations of the mtrA
gene result in a defect in soluble Fe
3+
reductive ability (66). Expression of MtrA in E.
coli, with a plasmid containing cytochrome c maturation genes, imparts E. coli with
the ability to reduce soluble Fe
3+
species, demonstrating the ability of MtrA to receive
electrons from a range of electron donors (77).
1.2.2 Regulation of Fe3+ Reduction
In Escherichia coli and Salmonella enterica, expression of genes that facilitate
growth under anaerobic conditions is controlled by FNR (for fumarate and nitrate
reduction) (78). S. oneidensis MR-1 contains an analogue to FNR known as EtrA
(electron transport regulator A), which was suggested as a possible regulator of genes
required for anaerobic electron transport (79). However, subsequent work has shown
9
that EtrA is not required for anaerobic growth on various substrates, including fer-
ric citrate (80). This suggests that the FNR system does not appear to be required by
MR-1 for anaerobic respiration of minerals.
Mutation studies on the cyclic AMP (cAMP) receptor protein (CRP) gene, crp, sug-
gest that CRP may play a role in regulating the induction of metal reducing genes in
MR-1 (79). CRP is known to play a role in the regulation of a myriad of biological
functions (81). Strains carrying mutations in the crp gene were defective in their abil-
ity to reduce both ferric citrate and MnO
2
. Addition of cAMP to aerobic cultures of
wild-type MR-1 resulted in an increase of soluble Fe
3+
reduction activity, although the
increased rates were still much lower than anaerobic cultures (79).
1.2.3 Nanowires
Although both Shewanella and Geobacter strains contain omcB genes, the genes do
not appear to be related (53). This fact has been used to suggest that the genes encode
enzymes that utilize different mechanisms for Fe
3+
reduction. However, recent work
has indicated that both Shewanella and Geobacter produce monolateral pili, common-
ly known as nanowires, that can be used to establish contact with substrates capable
of acting as electron acceptors (82, 83). Although Reguera et al. have suggested that
the pili are responsible for mediation of electron transfer from the cell to the acceptor,
in Shewanella it appears that the outer membrane-bound cytochromes are responsible
for electron transfer (83), raising the possibility that both GS-15 and Shewanella used
the same mechanism with different cytochromes to accomplish the same function.
10
1.2.4 Factors Infl uencing Dissimilatory Iron Reduction
Because of the importance of iron redox cycling in sedimentary environments, much
work has been done trying to understand the factors involved in dissimilatory iron re-
duction. A number of factors have been shown to infl uence the rate and extent of iron
reduction as well as the secondary minerals resulting from biologically mediated iron
reduction. Under laboratory conditions, solution chemistry has been found to exert
a considerable effect on the products of biologically mediated ferrihydrite reduction
(21, 24). A bicarbonate buffer can promote the formation of siderite (21, 84). High
N
2
:CO
2
ratios lead to the formation of magnetite (24). The addition of phosphates
can lead to the formation of vivianite (21), although Zachara et al. (24) postulate that
phosphate used in conjunction with PIPES buffer can inhibit mineralization, leading
to the precipitation of amorphous green rust.
Electron acceptor to donor ratios can also have an infl uence on the rate and prod-
ucts of iron reduction. Fredrickson et al. (20) have shown that iron reduction rates
decrease with increasing hydrous ferric oxide (HFO):lactate ratios. High ratios of
HFO:lactate also resulted in a greater amount of HFO recrystallizing to goethite, due
possibly to interactions with Fe. This recrystallization may have contributed to de-
creased reduction rates, as less crystalline iron oxides are more available to dissimila-
tory iron reducers (85). Lactate can also adsorb strongly to HFO at neutral pH (20).
This interaction may have the effect of reducing both the availability of HFO and of
lactate. When the electron donor is in excess, ferrihydrite is almost completely trans-
formed into fi ne-grained magnetite (20).
11
The adsorption of Fe to cell surfaces can interfere with the bioreduction of aqueous
Fe
3+
(86). For example, incubation of S. putrefaciens strain CN32 with Fe prior to ini-
tiation of bioreduction experiments had a marked impact on both Fe
3+
reduction and
CN32 growth rates. Both growth rates and Fe
3+
reduction rates showed a prolonged
lag phase of about 20 hours after 24 hours of incubation with FeCl
2
. Lower initial
concentrations of cells lead to a longer lag phase than higher initial concentrations.
Initial concentrations of Fe
2+
and iron oxide species have been shown to infl uence
subsequent HFO remineralization pathways (20, 22, 87). Low initial concentrations
of Fe
2+
can promote goethite precipitation, while high initial concentrations tend to
favor precipitation of magnetite (22, 87). When the initial concentration of HFO is
very low (<10mM), the relative rate of Fe
2+
production exceeds the rate of recrystal-
lization such that no secondary remineralization is observed (20). However, Hansel
et al (88) have also shown that while initial concentrations of Fe can infl uence subse-
quent mineralization products, the ratio of various iron oxides can also play a role in
iron reduction products. They reported that the rate of magnetite precipitation was a
function of the proportion of goethite to lepidocrosite, with higher concentrations of
goethite slowing down magnetite formation.
Although the majority of experiments using HFO are done with relatively pure syn-
thetic ferrihydrite, the presence of other cations can also have an effect on rates and
mineral speciation. While both Ni and Co prevent HFO recrystallization to goethite,
cells incubated with HFO containing Co are able to reduce Fe
3+
, while HFO contain-
ing Ni resists bacterial reduction (24, 89). Anions such as phosphate and silicate are
12
also known to inhibit recrystallization of HFO to goethite (90). The use of phosphates
in media in conjunction with PIPES buffer is not thought to inhibit iron reduction
rates, but it is thought to inhibit crystallization of reduced iron to magnetite, instead
promoting the formation of green rust (21, 24).
Zachara et al. (24) have proposed a conceptual model where the rate of supply and
total concentration of Fe is the primary determinant of secondary mineralization prod-
ucts (Figure 1.1). Under this concept, low rates of Fe fl ux promote recrystallization
of HFO into goethite, hematite or other more thermodynamically stable iron oxide
phases. Higher reduction rates can lead to the formation of magnetite and siderite,
while phosphate and other metals, such as Ni and Co, can act as inhibitors of recrys-
tallization.
Figure 1.1. Conceptual model of iron biomineralization. The arrows represent the magnitude and rate
of Fe2+ fl ux. From Zachara et al., 2002.
13
1.3 Geological Signifi cance of Dissimilatory Iron Reduction
Most sedimentary environments contain a dry weight iron concentration of 3-5%
(39), making iron one of the more important elements in terms of concentration. Iron
(hydr)oxides are important sorbents of trace metals such as Al, Cd, Co, Pb, Zn, As
and U (53, 91, 92). As such, any changes in the state of iron oxides in soils can have
an important impact on the mobility of trace metals, many of which are considered
to be pollutants (93, 94). Understanding the extent to which dissimilatory iron reduc-
ers either dissolve or produce minerals that are able to immobilize these metals can
provide further insights into the impact that iron reducers may have in contaminated
subsurface environments.
On a longer geologic time scale, dissimilatory iron reduction is considered to be one
of the earliest forms of microbial respiration (95). If this is the case, it would stand
to reason that the oxidation of iron that is thought to have occurred during the late
Archean-Paleoproterozoic and again in the Neoproterozoic should have been impact-
ed by the activity of dissimilatory iron reducers. The role that iron reducers may have
played in the deposition of what are now known as banded iron formations (BIFs) has
been given some consideration in concept (96, 97). However, the majority of experi-
mental work that has been done in trying to understand the role of biology in BIFs
has been done with an eye towards understanding the signifi cance of iron oxidation
(97-102). Recently work has been done attempting to correlate the isotopic signature
of Fe produced by iron reducers to the signature found in BIF deposits (23). However,
these results are somewhat ambiguous. Trying to understand the role of iron reducers
14
in BIF deposition is extremely diffi cult due to the lack of knowledge concerning the
amount of Fe in the Precambrian ocean, the rates and location of Fe
3+
deposition and
the relative importance of iron reduction with respect to other metabolic strategies.
Still, understanding the rate at which a cell can reduce iron and the products that are
formed as a function of these rates can begin to provide some insight into what iron
reducers may have been capable of and responsible for with respect to BIF deposi-
tion.
15
Chapter 1 Endnotes
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23
CHAPTER TWO
THE INFLUENCE OF BACTERIAL STRAIN SPECIFICITY ON
SECONDARY IRON MINERAL FORMATION
2.1 Abstract
It has been known for some time that microbes play an important role in the redox
cycling of iron, and while a considerable amount is known about the various factors
that affect the rate and extent of solid phase iron reduction, little is known about the
factors that control secondary mineral formation. To this end, this chapter describes
experiments conducted using 3 different strains of Shewanella. Strains CN32, MR-4
and W3-18-1, all well known for their ability to reduce HFO (hydrous ferric oxide),
using lactate as the organic carbon and energy source. Mineral products of iron
reduction were analyzed using X-ray diffraction, electron microscopy, coulometry
and susceptometry. Under similar nutrient loadings, iron reduction rates between
strains W3-18-1 and CN32 were similar and about twice as fast as MR-4. Qualitative
and quantitative assessment of biomineral products indicated that strains with
similar rates (CN32 and W3-18-1) produced different end products (secondary
minerals) and strains with different iron reduction rates (CN32 and MR-4) produced
similar end products. The major product of iron reduction by strains CN32 and
MR-4 was magnetite, while for W3-18-1, it was a mixture of magnetite and iron
carbonate hydroxide hydrate, a precursor to fougerite. Another notable difference
was that strains CN32 and MR-4 converted all of the starting ferric iron material
24
into magnetite, while W3-18-1 did not convert most of the Fe
3+
into a recognizable
crystalline material. These results suggest that the relative abundance of mineral end
products and the relative distribution of these products are strongly dependent on the
bacterial species or strain catalyzing iron reduction.
2.2 Introduction
Iron is one of the most abundant elements on earth, making up about 5 weight
percent of the earth’s crust. Under surface conditions found in the modern earth,
iron can occur as one of several oxyhydroxides, carbonates, phosphates or silicates.
These minerals can form a major constituent of some rocks and sediments (1-3).
The structure of these minerals can infl uence their susceptibility to reduction and
oxidation reactions, thus infl uencing the biogeochemical cycling of iron in soils and
sediments (1, 4).
It is well known that bacteria can play an important role in iron biogeochemical
cycling (5-11). The process by which bacteria can couple the oxidation of organic
carbon to the external reduction of iron oxide is known as dissimilatory iron
reduction. Dissimilatory iron reducers are ubiquitous in soils and sediments (10). The
secondary mineral products of dissimilatory iron reduction can have an impact on
the mobilization or sequestration of organic and inorganic pollutants, as well as the
subsequent geochemical cycling of iron in soil and sedimentary environments (12-
15).
25
There are several factors thought to control the products of dissimilatory iron
reduction. Researchers have postulated that the amount of Fe
2+
in a system during
the early stages of reduction can have an impact on the subsequent mineralization
(16-18). Low initial concentrations of Fe
2+
can promote goethite accumulation and
inhibit magnetite precipitation, even if Fe
2+
concentrations increase at a later time
(17). Under laboratory conditions, it is thought that the type of buffer used is largely
responsible for the identity of the reduced iron oxide biominerals (13, 19). Under
these conditions, carbonate buffered media produce largely siderite, phosphate
buffered media produce largely vivianite, while media buffered with organics produce
magnetite. A conceptual model has also been proposed where the rate of supply and
total concentration of Fe
2+
is the primary determinant of secondary mineralization
products (13). According to this model, low rates of Fe
2+
production promote
recrystallization of HFO into goethite, hematite or other more thermodynamically
stable iron oxide phases. Higher Fe
3+
reduction rates can lead to the formation of
magnetite and siderite. It is believed that bacteria have no infl uence on the formation
of secondary iron oxide minerals beyond producing a supply of Fe
2+
(13).
While dissimilatory iron reducers are ubiquitous in nature, most laboratory-based
work has been done on strains from Geobacter and Shewanella (9, 14, 20-26). The
Shewanella genus encompasses several strains that have been shown to be capable
of dissimilatory iron reduction (19, 24, 27-29). Shewanella are gram-negative,
facultative anaerobes belonging to the γ-proteobacteria. They have a rapid generation
time under aerobic conditions, making them ideal for physiologic studies. The
majority of work done investigating the factors that affect the formation of secondary
iron minerals within Shewanella has been done on S. putrefaciens CN32 (12, 13,
26
17, 19, 21, 30-32). In this chapter, we compare the secondary iron mineral products
of different Shewanella strains using lactate in order to understand the infl uence of
bacterial strain on mineral formation.
2.3 Materials and Methods
2.3.1 Bacterial Cultures
The Shewanella strains used in this study are listed in table 2.1. Strains were provided
by the Pacifi c Northwest National Lab. The bacteria were inoculated directly from
frozen stocks into Luria-Bertani (LB) broth and grown overnight in a 15
0
C incubator
shaking at 125rpm. The cultures were washed with carbonate buffered medium to
remove traces of LB. Defi ned medium is adapted from Fredrickson et al, 1998 (19).
Medium composition is listed in table 2.2 and has been shown to support the growth
of several strains of Shewanella.
Table 2.1 Bacterial strains used in dissimilatory iron reduction experiments
Strain Strain Origin
Biofi lm
Formation
Lipopolysacharide
Layer
Putative Metal
Reducing genes*
Shewanella
putrefaciens CN32
Uranium Mine,
New Mexico
Weak Short, rough 11 heme omcB
Shewanella sp. MR-4
Black Sea Water
Column
Weak Smooth
omcA1, mtrD,
mtrE, mtrF
Shewanella
putrefaciens W3-18-1
Pacifi c Ocean
Marine Sediments
Strong ND 11 heme omcB
* In addition to mtrA, mtrB and mtrC; ND = No Data
27
Table 2.2 Medium Composition
Chemical Description Concentration (moles/L)
NaHCO
3
3.1x10
-2
Ammonium chloride 2.84x10
-2
Potassium chloride 1.34x10
-3
Sodium phosphate monobasic 8.32x10
-4
Sodium chloride 3.0x10
-2
Sodium Lactate 2.0x10
-2
Biotin (d-biotin) 8.19x10
-8
Folic acid 4.53x10
-8
Pyroxine HCl 4.86x10
-7
Ribofl avin 1.33x10
-7
Thiamine HCl 1.0 H2O 1.41x10
-7
Nicotinic acid 4.06x10
-7
d-pantothenic acid, hemicalcium salt 2.10x10
-7
B12 7.38x10
-10
p-aminobenzoic acid 3.64x10
7
Thioctic acid 2.42x10
-7
L-glutamic acid 1.36x10
-4
L-arginine 1.15x10
-4
DL-serine 1.90x10
-4
Nitrilotriacetic acid 7.85x10
-5
Magnesium sulfate heptahydrate 1.21x10
-4
Manganese sulfate monohydtrate 2.96x10
-8
Sodium chloride 1.71x10
-4
Ferrous sulfate heptahydrate 3.60x10
-6
Calcium chloride dihydrate 6.80x10
-6
Cobalt chloride hexahydtrate 4.20x10
-6
Zinc chloride 9.54x10
-6
Cupric sulfate pentahydrate 4.01x10
-7
Aluminum potassium disulfate
dodecahydtrate
2.10x10
-7
Boric acid 1.62x10
-6
Sodium molybdate dihydrate 1.03x10
-6
Nickel chloride hexahydrate 1.01x10
-6
Sodium tungstate 7.58x10
-7
pH is not adjusted, and typically stabilizes at approximately 8.6
The medium was fi lter-sterilized with a .2 μm PES vacuum fi ltration system (Nalgene)
prior to inoculation. Medium (18mL) was then added to serum bottles along with
2mL of 0.2 M hydrous ferric oxide (2x10
-2
M fi nal concentration). The bottles were
28
gassed with pure N
2
, plugged with butyl-rubber stoppers and crimp sealed. Washed
cells were injected into the sealed serum bottles with a syringe using a 21-gauge
needle to achieve a fi nal cell concentration of approximately 1 X 10
8
cells/ml. The
cultures were incubated at 15
0
C and sampled at defi ned intervals for up to 1000 hours.
2.3.2 Preparation of Hydrous Ferric Oxide
A stock solution of 0.2M hydrous ferric oxide (HFO) was prepared according to the
method of Cornell and Schwertmann (33). Ferric chloride hexahydrate (54g of FeCl
3
.
6H
2
O) was dissolved into 18M Ω water (2L). NaOH pellets were added to bring
the pH up to approximately 7, causing precipitation of the dissolved ferric iron. The
precipitated iron slurry was washed repeatedly with 18M Ω water to remove any trace
salts and brought to a fi nal volume of 1L. The resulting material was analyzed via
X-ray diffraction to confi rm the production of HFO.
2.3.3 Fe
2+
Analysis
At selected time points, bottles were removed from the incubator and placed in an
anaerobic glove box (Coy Labs) under N
2
/H
2
headspace. Aliquots (250 μl) were
extracted with a 21-gauge syringe needle and placed into micro-centrifuge tubes.
These samples were centrifuged at 10600xg rcf for 2 minutes. The supernatant was
placed into a separate tube containing 250 μl of 1N HCl. This sample was used to
determine the soluble Fe(II) concentration. The pellet was re-suspended with 250
μl of anaerobic 18M Ω water and 250 μl of 1N HCl. This solution was used for
29
determining the HCL-extractable solid phase, or adsorbed Fe(II). Both portions of the
reduced iron were quantifi ed using the ferrozine spectophotometric assay as described
by Stookey (34).
2.3.4 Mineralogical Analysis
Samples were prepared in a similar fashion for all analyses done to determine mineral
identity, concentration and morphology. All samples were collected in the glove box.
Approximately 2ml of material was collected from the anaerobic serum bottles using
21-gauge needles. In an attempt to remove organics and salts from the collected
material, samples were washed 2-3x with anaerobic 18M Ω water.
2.3.4.1 Environmental Scanning Electron Microscopy
Approximately 10 μl of the washed samples were placed onto a polycarbonate
fi lter and allowed to air dry under anaerobic conditions. Samples were kept under
anaerobic conditions until the time of analysis. These samples were used to determine
biomineral morphology by use of an environmental scanning electron microscope
(ESEM, Hitachi TM-1000 Table Top Microscope).
2.3.4.2 Susceptometry
Magnetic susceptibility was utilized in order to determine the relative amount of
magnetite among the collected samples. This technique has been applied to the
analysis of magnetite produced by magnetotactic bacteria and by dissimilatory
30
iron reducers (35), suggesting that this type of technique can yield interpretable
results for analyzing reduced iron oxide biominerals. The remaining material from
the washed samples was collected for analysis. Samples were transferred into
micro-centrifuge tubes and allowed to dry under anaerobic conditions. The dried
samples were then weighed, placed into small plastic containers and analyzed on a
susceptometer (Kappabridge KLY-4S). The resulting values for the tested biominerals
were compared to a synthetic magnetite (Sigma-Aldrich) in order to derive a mass
for the amount of magnetite found in the biological samples. After analysis, these
samples were placed in serum bottles, sealed and evacuated of any air for coulometric
analysis.
2.3.4.3 Coulometry
Coulometry was used to quantify the amount of inorganic carbon produced by the
tested bacterial strains. The samples used for susceptometry measurements were
reweighed, placed into serum bottles, stoppered and sparged with 100% N2. 400 μl of
20% sulfuric acid was added to each sample to dissolve the solid material and release
any solid inorganic carbon as carbon dioxide. The acidifi ed samples were placed on
a shaker rotating at 105 rpm and left overnight to ensure complete dissolution of the
sample. Once the samples had dissolved, the serum bottles were connected in-line to
a coulometer (UIC, Inc. Coulometrics model 5012) via two 21-gauge needles serving
as an inlet and outlet. The gas in the serum bottles was displaced by nitrogen carrier
gas.
31
2.3.4.4 XRD
At the end of the experiment, samples were collected in order to characterize the
major biomineral products produced by the tested bacterial strains. An aliquot of
3mL was collected, washed repeatedly with anaerobic 18M Ω water to remove any
organics and salts, and air dried under anaerobic conditions. Samples remained under
anaerobic conditions until the time of analysis. A small amount of dried material was
loaded onto a sample holder and analyzed using a Rigaku R-Axis Spider diffraction
system equipped with a goniometer and Molybdenum X-ray source.
2.4 Results
2.4.1 Reduced Iron Oxide Biominerals
ESEM images comparing the morphology of the biomineral products produced
by the three bacterial strains tested in these experiments are shown in Figure 2.1.
In all cases, the end product was different from the starting material. Both CN32
and MR-4 produced nanocrystalline grains. In the case of CN32, these grains are
morphologically similar to those previously reported (13). Strain W3-18-1 produced
nanocrystalline material, as well as hexagonal plates (Fig 2.1B).
XRD analysis revealed that all three strains produced magnetite (ig 2.2), and that that
the degree of crystalinity for the magnetite products was different for the three strains.
Both CN32 and MR-4 produced more crystalline magnetite than did W3-18-1. In the
32
Figure 2.1 Environmental scanning electron micrographs of reduced iron oxide biominerals produced
by the strains used in this experiment. A. Magnetite produced by strain CN32 (white arrow, and
surrounding fi eld of view); B. Magnetite/Iron carbonate hydroxide hydrate mixture produced by strain
W3-18-1. Iron carbonate indicated by white arrows.
Figure 2.1A. CN32
Figure 2.1B. W3-18-1 20 μm
20 μm

case of W3-18-1, XRD results indicated that a second mineral was formed. Peak
patterns for this second product are consistent with iron carbonate hydroxide hydrate
(green rust), a precursor to fougerite (Fe
2+
6(1-x)
Fe
3+
6x
(OH)
4(4-3x)
(OOH)
2(3x-1)
CO
3
) (36).
33
CN32
MR-4
W3-18-1
Iron Carbonate Hydroxide Hydrate
Magnetite
Figure 2.2 XRD patterns for bominerals and standards. X-axes are values in two-theta.
34
0 100 200 300 400 500 600 700 800 900
0
2
4
6
8
10
W3-18-1
CN32
MR-4
Control
2.4.2 Comparison of HFO Reduction

Reduction of HFO through 1000 hours is shown in Figure 2.3. HCl (0.5N) is
considered to be an effective solvent for most biogenic solid phase Fe
2+
, including
magnetite (16). As such, measurements of HCl-extractable Fe
2+
are used here as an
indicator of total reduced iron production. The rate of reduction by S. putrefaciens
CN32 appeared to be constant through 800 hours. Rates of reduction by strain W3-
18-1 were to be similar to CN32, although W3-18-1 showed a slight increase between
100 and 200 hours. There was a lag in the reduction rate for strain MR-4 between 100
and 300 hours, after which the rate of reduction increased.
HCl-extractable Fe
2+
(mM)
Time (hours)
Figure 2.3 Total HCl-extractable reduced iron-oxide produced by the tested Shewanella strains using
HFO as the electron acceptor and lactate as the electron donor. Error bars indicate the standard for each
time point.
35
The amount of soluble Fe
2+
liberated during HFO reduction was different for the
three strains (Fig. 2.4). Strains CN32 and W3-18-1 showed a similar pattern of
increase through 400 hours, at which point the amount of aqueous phase Fe
2+
began
to decrease for strain CN32. The concentration of aqueous phase reduced iron
continued to increase steadily for W3-18-1 through the duration of the experiments.
Strain MR-4 also showed a steady increase in aqueous phase Fe
2+
, although it was
approximately 5 times lower than the other two strains. Although the concentration of
aqueous Fe
2+
was four times higher for W3-18-1 compared to CN32 and MR-4, in all
cases the concentration of Fe
2+
found in solution constituted less than 5% of the total
reduced iron in the samples.
0 100 200 300 400 500 600 700 800 900
0
100
200
300
400
500
600
700
CN32
MR-4
W3-18-1
Control
Figure 2.4 Concentrations of reduced aqueous phase iron. Data for each sample is based on 4 to 9
independent replicates. Error bars indicate the standard error for each time point.
Time (hours)
Aqueous Fe
2+
(μM)
36
2.4.3 Relative Magnetite Concentrations
The pattern of magnetic susceptibility for strain CN32 showed a rapid increase
followed by a plateau after 400 hours (Fig. 2.5). Magnetic susceptibility of the
products of iron reduction by strain MR-4 showed an exponential increase with time,
reaching a level similar to strain CN32 by 1000 hours. In contrast, strain W3-18-
1 produced materials that showed no signifi cant change in response to the applied
magnetic fi eld beyond 100 hours. However, even in this case, the susceptibility values
were larger than that of the controls, indicating that a transformation of some of the
starting ferric iron oxide material has taken place.
Figure 2.5 Semi-log plot of susceptibility values ( χ) for the tested Shewanella strains. Strains are
represented by the following symbols: ( S ) S. putrefaciens CN32; ( ) S. sp. MR-4; (Q ) S. putrefaciens
W3-18-1; ( ) negative control. Error bars indicate the standard deviation of the mean for each time
point.
0 100 200 300 400 500 600 700 800 900
1.0x10
-6
1.0x10
-5
1.0x10
-4
1.0x10
-3
1.0x10
-2
Time (hours)
χ
(m
3
/kg)
37
2.4.4 Inorganic Carbon Concentrations
Solid phase inorganic carbon present in the samples at the end of the experiment
are shown in Figure 2.6. Values for strains CN32 and MR-4 are within the range of
the background values produced by the zero control samples. Strain W3-18-1, on
the other hand, produced almost fi ve times as much particulate inorganic carbon.
Conditions in these cultures are such that the only expected solid-phase carbonate
would be an iron carbonate of some kind. Based on the information gathered from
XRD analysis, the iron carbonate is in the form of iron carbonate hydroxide hydrate.
By using the formula weight of iron carbonate hydroxide (Fe
6
(OH)
12
CO
3
*2H
2
0) and
an iron to carbon ratio of 6, we conclude that approximately one-third of the starting
ferric iron material has been incorporated into iron carbonate.
CN32 MR-4 W3-18-1 Control
0
25
50
75
100
125
150
Figure 2.6.Box plot of solid phase inorganic carbon, in micrograms, produced by the bacterial strains
used in these experiments. The error bars represent the standard deviation.
Inorganic Carbon ( μg)
38
2.5 Discussion
The information derived from these experiments can be used to construct a profi le of
the relative mineral distribution produced by the tested strains (Table 2.3). Magnetite
concentrations were derived from comparisons to magnetite standards analyzed
on the susceptometer. In the case of CN32 and MR-4, this procedure produced a
calculated mass of magnetite that was greater than the actual mass of the tested
sample. There are various factors that can infl uence the susceptibility of a material,
such as mineral concentration, mineral composition, crystal size and crystal shape
(37). While grain size can play a role in the degree to which a sample will respond
to an applied magnetic fi eld, microscopy does not suggest large differences in the
morphology of the magnetite products.
Table 2.3 Summary of iron reduction rates and biomineral distribution
S. putrefaciens CN32 S. species MR-4 S. putrefaciens W3-18-1
Major Reduced
Iron Products
Magnetite (Fe
3
O
4
) Magnetite (Fe
3
O
4
)
Magnetite (Fe
3
O
4
) /
Iron Carbonate Hydroxide
Hydrate Fe
6
(OH)
12
CO
3
*2H
2
O
Distribution (mg) Fe
3
O
4
: 30.9 +/- 4.6 Fe
3
O
4
: 10.8 +/- 1.2
Fe
3
O
4
: 3.3 +/- 0.8
Fe
6
(OH)
12
CO
3
*2H
2
O: 7 +/- 0.2
Reduction Rate
(μmol/L*day)
134-566 100-370 210-623
Final pH 8.5 8.4 8.2
39
For CN32 and MR-4, the products of dissimilatory iron reduction consist almost
entirely of magnetite. This is in agreement with previous results in which Shewanella
strains were placed in cultures buffered in carbonate with pure N
2
as the headspace
and given lactate as the electron donor (12, 16, 19). In these studies, the major
biomineral formed for both CN32 and MR-4 cultures was magnetite. Based on the
amount of reduced iron produced (Figure 2.3), CN32 converted all of the starting
material into crystalline magnetite, while MR-4 converted approximately one third
of the hydrous ferric oxide into magnetite. Strain W3-18-1 also produced magnetite,
however, the amount of magnetite produced was much smaller, the major product of
iron reduction being iron carbonate hydroxide. Additionally, unlike strains CN32 and
MR-4, a majority of the reduced Fe
2+
was not remineralized into any form detectible
by the methods used in these experiments.
Thus, under similar conditions, only strain W3-18-1 produced a detectable iron
carbonate mineral product. The production of iron carbonate material by iron
reducing bacteria under laboratory conditions has been reported by other researchers
(13, 19, 38, 39). However, the conditions under which these minerals were produced
involved large amount of CO
2
in the headspace; under conditions applied here, in
which the headspace was initially composed of pure nitrogen gas, the typical iron
biomineral product of dissimilatory HFO reduction was magnetite (13, 19).
It has been shown that increased concentrations of Fe
2+
can have an inhibitory
effect on the bioreduction of ferric iron (40, 41). This inhibition would infl uence
the remineralization and subsequent identity of mineral products by reducing the
amount of reduced iron being introduced into solution (41). However, the starting
40
amounts of Fe
2+
used in those experiments are larger than the concentration of
aqueous Fe
2+
reported here. Given this, and the fact that the aqueous Fe
2+
constitutes
such a small fraction of the overall reduced iron oxide pool, it seems unlikely that the
observed differences in mineral products reported here are due to differences in the
concentration of soluble Fe
2+
.
It is known that organics can interact strongly with both ferric and ferrous forms of
iron (42-47). As such, it is possible that differences in the metabolism of lactate might
lead to the formation of secondary metabolites that would chelate either form of iron.
This might lead to infl uences in the degree of remineralization based on the types of
metabolites produced by individual bacteria. Genetic information, however, suggests
that processing of lactate occurs via identical metabolic pathways, although such
information is putative. However, work done on HPLC (data not shown) does not
suggest that there are major differences in the production of secondary metabolites.
6 7 8 9 10 11 12 13 14
0
2
4
6
8
10
12
14
16
18
20
22
24
W3-18-1
Fe
3
O
4
Fe
6
(OH)
12
CO
3
*2H
2
O
FeCO
3
Fe(OH)
3
CN32
MR-4
pH
-log [Fe
2+
] (M)
Figure 2.7. Thermodynamic
stability fi eld of hydrous
ferric oxide (assumed
stoichiometry: Fe(OH)
3
),
magnetite (Fe
3
O
4
), green rust
(Fe
6
(OH)
12
CO
2
*2H
2
O) and
siderite (FeCO
3
) as a function
of Fe
2+
concentration and pH.
A [HCO
3
-
] of 0.03M was
assumed, based on buffer
concentration.
41
A stability diagram for magnetite, siderite and green rust indicates that the major
product of iron reduction under the conditions at the end of the experiments should
be magnetite (Figure 2.7). The results observed by strains CN32 and MR-4 are
in agreement with what would be predicted under these conditions. In the case of
W3-18-1, however, there is a signifi cant amount of green rust present, despite its
apparent instability under these conditions. Although there is a large concentration
of bicarbonate in these experiments (Table 2.2), the presence of green rust is still
not predicted under these pH conditions (Figure 2.8). Green rust is considered to
be metastable with respect to magnetite and siderite (48), and it is possible that the
material produced by strain W3-18-1 will eventually transform into magnetite or
siderite. However, it is has been reported in some soils (36, 49), indicating that it does
posses at least some temporal stability. Nonetheless, its presence is still noteworthy,
as it is a precursor to lepidocrosite in environmental settings (50). Thus, while the
primary products of dissimilatory iron reduction my undergo further transformation
as environmental conditions change, the identity of succeeding iron minerals may be
infl uenced by these early stage precipitates.

Based on the results presented here, there is a fair degree of overlap between the
rates of iron reduction by strains CN32 and W3-18-1, and between CN32 and MR-4
(Table 2.3). It has been suggested that the rate of Fe
2+
production can play a role in
determining the identity of the products of dissimilatory iron reduction (13), although
the relationship between Fe
3+
reduction and specifi c biomineral products has not been
quantifi ed. While the initial rates of Fe
3+
reduction are similar, there is a deviation at
42
0 1 2 3 4 5 6 7 8 9 10
0
2
4
6
8
10
12
14
16
18
20
Fe
3
O
4
Fe(OH)
3
FeCO
3
about 100 hours (Figure 2.3). This suggests that reduction rate may not be the
principal factor controlling mineral nucleation, and other factors may play a role.
The results presented here indicate that different bacterial strains can produce
different mineral end products under virtually identical starting conditions. The
factor(s) responsible for these differences remain to be elucidated, but our results
Figure 2.8. Thermodynamic stability fi eld of hydrous ferric oxide
(assumed stoichiometry: Fe(OH)
3
), magnetite (Fe
3
O
4
), and siderite (FeCO
3
) as a function of Fe
2+

concentration and HCO
3
-
concentration. A pH of 8.4 was assumed, based on the average from data
gathered in these experiments (table 2.3). The ( z ) symbol represents a HCO
3
-
concentration of 0.03M
and a Fe
2+
concentration of 8.7mM for W3-18-1, 7.1mM for CN32 and 4.4mM for MR-4
-log [Fe
2+
] (M)
-log [HCO
3
-
] (M)
43
suggest they do not involve differences in rates of Fe
3+
reduction, types of excreted
metabolites, medium pH, medium Fe
2+
concentration, or CO
2
concentration in the
medium headspace. Other factors might include differences in extracellular proteins
or polysaccharides, neither of which were studied here. It is also possible that biofi lm
formation is intimately involved with secondary mineral formation. Results from
our laboratory (51) indicate that there are comparative differences in the degree to
which these strains produce biofl ims. In the case of W3-18-1, biofi lm formation
is robust. On the other hand, biofi lm formation is much weaker for strains CN32
and MR-4. It may be that this difference manifests itself in the types of minerals
produced by iron reducing bacteria. Extra cellular polysaccharide (EPS) layers might
retard the diffusion of reduced iron into solution, preventing its interaction with the
outside medium and thus, preventing reprecipitation of iron minerals. Thus, while
the reduced iron might be found in the solid phase, it would be in a non-crystalline
form, bound within the biofi lm of the iron reducing bacteria. It would be of great
utility to investigate the degree to which biofi lms or EPS layers might infl uence the
formation of reduced iron oxide biominerals and the infl uence that this might have in
the movement of reduced iron through pore waters.
44
Chapter 2 Endnotes
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48
CHAPTER THREE
THE IMPACT OF BACTERIA AND CARBON SOURCE ON THE
FORMATION OF REDUCED IRON MINERALS
3.1 Abstract
We have examined the infl uence of carbon source on both the rate of iron reduction
and the mineralogy of the reduction products with Shewanella putrefaciens strain
W3-18-1. When pyruvate is the carbon source, the secondary products were spherules
composed of siderite. When uridine was used as the carbon source, the products
were hexagonal plate-like structures identifi ed as iron carbonate hydroxide hydrate,
also known as carbonate green rust, a precursor to fougerite. When lactate was
used as the carbon source, products were a mixture of iron carbonate hydroxide and
magnetite. In terms of reaction stoichiometry, there were differences in the amount
of acetate produced depending on the starting organic carbon source. Incubation
with pyruvate produced a relatively large amount of acetate compared to incubation
with uridine and lactate. There were also differences in the fi nal pH of the cultures.
While the pH for incubations with lactate started at 8.6 and ended between 8.0-8.3,
the pH of cultures incubated with uridine was found to be almost a full unit lower at
the conclusion of the experiment (~7.4). Solubility diagrams based on the chemistry
found in our experiments predict that the production of Fe
2+
(aq)
should always lead to
49
the formation of magnetite. However, strain W3-18-1 produced different minerals
depending on the carbon source utilized as the electron acceptor.
3.2 Introduction
The coupling of iron oxide reduction to organic carbon oxidation via microbial
activity is thought to be important in certain natural subsurface environments (1-5).
This activity, commonly known as dissimilatory iron reduction, can in turn infl uence
the formation and distribution of reduced iron minerals in soils and sediments. The
secondary mineral products of dissimilatory iron reduction can have an impact on
the mobilization or sequestration of organic and inorganic pollutants, as well as the
subsequent geochemical cycling of iron in soils and sediments (6-9)
A vast body of work has been generated since the fi rst metal reducing organisms
were isolated, attempting to understand the various factors that play a role in the
reduction of solid phase iron (10, 11). Variables identifi ed as infl uencing both the
reduction of sedimentary iron oxides and the subsequent formation of reduced iron
minerals include the type of iron oxide available for reduction (9, 12-16), medium
composition (17, 18), surface area of the starting material (19, 20), ferrous iron
concentration (21) and trace metal composition (9, 22, 23). Researchers have also
reported the dependence of secondary iron mineral formation on the identity of the
low molecular weight organic acid used as the carbon source (24). It is thought that
the microbes themselves play no role in the formation of reduced iron minerals other
than to supply aqueous Fe
2+
to the system (9).
50
While low molecular weight organic acids are thought to have a wide distribution in
soils and sediments and thus be an important source of energy (25-27), there have
been limited studies investigating the reduction of iron using more complex organic
compounds as energy sources (24, 28). The aim of this study was to determine [1]
what importance the oxidation of more complex organic compounds might have on
the products of dissimilatory iron reduction, when compared to resuts using more
traditional low molecular weight organic acids, and [2] to compare these products to
the minerals that would be predicted based on pure chemistry.
3.3 Methods
3.3.1 Bacterial Cultures
Shewanella putrefaciens strain W3-18-1 is a facultatively anaerobic, gram-negative
psychrophilic bacteria. This bacterium was isolated off the coast of Washington State,
in sediments underlying 670m of water (29). Like the majority of other Shewanella
strains, W3-18-1 is capable of metal reduction. Strain W3-18-1 was inoculated
directly from frozen stocks into Luria-Bertani (LB) broth and grown overnight in
a 15
0
C incubator shaking at 125rpm. The culture was then washed with carbonate
buffered media to remove traces of LB. Medium composition is listed in Table 1. The
medium was fi lter-sterilized with a 0.2 μm PES vacuum fi ltration system (Nalgene)
prior to inoculation. 14mL of media was added to serum bottles along with 2mL of
0.2 M hydrous ferric oxide (2x10
-2
M fi nal concentration). Additionally, 4mL of a
fi lter-sterilized stock solution
51
Table 3.1. Medium Composition
Chemical Description Concentration (moles/L)
NaHCO
3
3.1x10
-2
Ammonium chloride 2.84x10
-2
Potassium chloride 1.34x10
-3
Sodium phosphate monobasic 8.32x10
-4
Sodium chloride 3.0x10
-2
Biotin (d-biotin) 8.19x10
-8
Folic acid 4.53x10
-8
Pyroxine HCl 4.86x10
-7
Ribofl avin 1.33x10
-7
Thiamine HCl 1.0 H2O 1.41x10
-7
Nicotinic acid 4.06x10
-7
d-pantothenic acid, hemicalcium salt 2.10x10
-7
B12 7.38x10
-10
p-aminobenzoic acid 3.64x10
7
Thioctic acid 2.42x10
-7
L-glutamic acid 1.36x10
-4
L-arginine 1.15x10
-4
DL-serine 1.90x10
-4
Nitrilotriacetic acid 7.85x10
-5
Magnesium sulfate heptahydrate 1.21x10
-4
Manganese sulfate monohydtrate 2.96x10
-8
Sodium chloride 1.71x10
-4
Ferrous sulfate heptahydrate 3.60x10
-6
Calcium chloride dihydrate 6.80x10
-6
Cobalt chloride hexahydtrate 4.20x10
-6
Zinc chloride 9.54x10
-6
Cupric sulfate pentahydrate 4.01x10
-7
Aluminum potassium disulfate dodecahydtrate 2.10x10
-7
Boric acid 1.62x10
-6
Sodium molybdate dihydrate 1.03x10
-6
Nickel chloride hexahydrate 1.01x10
-6
Sodium tungstate 7.58x10
-7
pH is not adjusted, and typically stabilizes at approximately 8.6
containing lactate, pyruvate or uridine (Table 3.2) were added to yield bottles with an
organic carbon concentration of approximately 20mM. The organic carbon sources
were chosen to study the effects of carbon compounds with different carbon oxidation
52
states and different carbon numbers on iron oxide reduction. The bottles were gassed
with pure N
2
, plugged with butyl-rubber stoppers and crimp sealed. Washed cells
were injected into the sealed serum bottles with a syringe using a 21-gauge needle
to achieve a fi nal cell concentration of approximately 1 X 10
8
cells/ml. The cultures
were incubated at 15
0
C and sampled at defi ned intervals for up to 1000 hours.
Table 3.2. Organic Carbon Sources
Carbon Source Lactate Pyruvate Uridine
Structure
Average Carbon
Oxidation State
0 +2 +2/3
3.3.2 Analysis of Organic Carbon Consumption
Organic carbon consumption and metabolic byproducts were analyzed via High
Pressure Liquid Chromatography (HPLC). Samples were extracted from the serum
bottles using a 21-gauge needle. This solution was then pushed through a 0.2 μM
syringe fi lter to remove both cells and any solid phase iron. A drop of each fi ltrate
was used to measure the pH of the solutions (EMD colorphast strips). The remaining
fi ltered solution was added to a microcentrifuge tube containing 10 μL of a 2.5 M
solution of H
2
SO
4
and mixed. The sulfuric acid was used to bring the pH of the
samples in line with the pH of the mobile phase in the HPLC. The samples were
stored at -80
0
C until analysis. Samples were run on an Agilent 1100 Series HPLC
equipped with a C18 column (Phenomenex, Luna 5u). Samples were analyzed at
53
room temperature with 1.25mM H
2
SO
4
mobile phase and a fl ow rate of 0.5mL per
minute.
3.3.3 Ferric Iron Reduction Analysis
3.3.3.1 Preparation of hydrous ferric oxide
Stock solution of 0.2M hydrous ferric oxide (HFO) was prepared according to the
method of Cornell and Schwertmann (30). 54 grams of ferric chloride hexahydrate
(FeCl
3
.
6H
2
O) was dissolved into 18M Ω water. NaOH pellets were added to bring
the pH up to approximately 7, causing precipitation of the dissolved ferric iron. The
precipitated iron slurry was washed repeatedly with 18M Ω water to remove any trace
salts and brought to a fi nal volume of 1L. The resulting material was analyzed via
X-ray diffraction to confi rm the production of HFO.
3.3.3.2 Ferrous iron analysis
At selected time points, bottles were removed from the incubator and placed in
an anaerobic glove box (Coy Labs) under N
2
/H
2
headspace. 250 μl aliquots were
extracted with a 21-gauge syringe needle and placed into micro-centrifuge tubes.
Preliminary work indicated that under acidic conditions, Fe
3+
was readily reduced to
Fe
2+
in the presence of pyruvate (data not shown). As such, it was necessary to wash
the solid phase reduced iron prior to analysis. Although this effect was not observed
with samples incubated with lactate or uridine, all samples were treated the same to
maintain consistency.
54
The 250 μl samples drawn were centrifuged at 10600xg rcf for 2 minutes. The
supernatant was placed into a separate microcentrifuge tube containing 250 μl of
1N HCl. This sample was used to determine the concentration of soluble Fe
2+
. The
pellet was re-suspended with 250 μl of anaerobic 18M Ω water and 250 μl of 1N HCl.
This solution was used for determining the HCL-extractable solid phase, or adsorbed
Fe
2+
. Both portions of the reduced iron were quantifi ed using the ferrozine assay as
described by Stookey (31). Once combined, this data provided the total amount of
iron reduced by W3-18-1 using each carbon source.
3.3.4 Mineral Analysis
Samples were prepared in a similar fashion for all analyses done to determine mineral
identity, concentration and morphology. All samples were collected in the anaerobic
glove box. Approximately 2ml of material was collected from the anaerobic serum
bottles using 21-gauge needles. In an attempt to remove organics and salts from the
collected material, samples were washed repeatedly with anaerobic 18M Ω water.
3.3.4.1 ESEM
Approximately 10 μl of the washed samples were placed onto a polycarbonate
fi lter and allowed to dry under anaerobic conditions. Samples were kept under
anaerobic conditions until the time of analysis. These samples were used to determine
biomineral morphology by use of an environmental scanning electron microscope
(ESEM, Hitachi TM-1000 Table Top Microscope).
55
3.3.4.2 Coulometry
The amount of inorganic carbon produced by strain W3-18-1 using the selected
carbon sources was determined by coulometric analysis. The samples were weighed
and placed into serum bottles, stoppered and sparged with 100% N2. 400 μl of 20%
sulfuric acid was added to each sample to release any solid inorganic carbon as
carbon dioxide. The acidifi ed samples were placed on a shaker rotating at 105rpm and
left overnight to ensure complete dissolution of the sample. Following dissolution,
the serum bottles were connected in-line to a coulometer (UIC, Inc. Coulometrics
model 5012) via two 21-gauge needles serving as an inlet and outlet. The gas in the
serum bottles was displaced by N
2
carrier gas. The coulometer gives a reading in
micrograms of carbon, which can then be used to determine the weight percent of
solid phase inorganic carbon in each sample.
3.3.4.3 XRD
At the end of the incubations, samples were collected in order to characterize the
major biomineral products produced by W3-18-1 using the tested carbon sources. 3ml
of material was collected, washed repeatedly with anaerobic 18M Ω water to remove
any organics and salts, and dried under anaerobic conditions. Samples remained under
anaerobic conditions until the time of analysis. A small amount of dried material
was loaded onto a sample holder and analyzed using a Rigaku R-Axis Spider curved
imaging plate microdiffractometer employing monochromatized MoK α radiation.
56
3.4 Results
3.4.1 Reduced Iron Oxide Mineral Products
3.4.1.1 Biomineral morphology
ESEM images comparing the morphology of the biomineral products produced
by strain W3-18-1 are shown in Figures 3.1 and 3.2. When lactate was used as
the carbon source, the products of iron oxide reduction were a mixture of fl ocular,
nanocrystalline material and hexagonal plates (Figure 3.1a). These plates tended to be
less than 5 μm in diameter. Incubation with uridine also produced hexagonal plates
similar to those seen with lactate (Figure 3.1b). However, they were more abundant
and larger in size, ranging between 5-10 μm in diameter and 1-2 μm in thickness.
Using pyruvate as the carbon source produced the nodular structures seen in fi gure
3.2. However, coupling iron reduction to pyruvate oxidation also produced globular
structures as seen in Figure 3.2b. These globular structures have been reported
previously (32). In that report, the authors concluded that the structures were siderite.
Elemental analysis of these structures indicated that they indeed have a similar
stoichiometry to siderite; however, they appear to produce no distinct crystal pattern
(data not shown). Treatment of these structures with hydrogen peroxide, did not lead
to dissolution, suggesting that the identifi ed carbon is inorganic.

57
3.4.1.2 Biomineral identifi cation
The collected XRD patterns were compared to the known database in order to
identify the products of iron reduction by strain W3-18-1. The patterns indicate
that all three of the tested carbon sources produce different materials. In the case of
A
B
Figure 3.1. Environmental scanning
electron micrographs of reduced
iron oxide biominerals produced
by strainW3-18-1; A. fl occular
magnetite (hashed arrow) and
hexagonal iron carbonate hydrox-
ide hydrate(white arrows) mixture
produced by incubation with lactate;
B. Iron carbonate hydroxide hydrate
produced by incubation with uridine.
Scale bars represent 10 μm.
58
lactate, XRD analysis showed that there were two major mineral products: magnetite
(Fe
3
O
4
) and iron carbonate hydroxide hydrate ([(Fe
2+
)
4
(Fe
3+
)
2
](OH)
12
CO
3
*2H
2
O;
Figure 3a). The broadness of the magnetite peaks suggests that the material is poorly
crystalline. XRD analysis indicated that the products of iron reduction with uridine
were consistent with iron carbonate hydroxide hydrate (fi gure 3b). This compound is
more commonly known as carbonate green rust (9, 33, 34). Green rusts appear to be
precursors to the mineral fougerite (35).
Figure 3.2. A: Siderite concretions
(arrow) produced by W3-18-1
when incubated with pyruvate; B.
siderite concretions, along with large
carbonate spheres (arrow) produced
by incubation with pyruvate; scale
bars represent 10 μm.
A
B
59
XRD patterns for the products of reduction by W3-18-1 with pyruvate as the carbon
source are shown fi gure 4. In both cases, the diffraction pattern produced peaks in
line with those of siderite (FeCO
3
). However, there was also a large peak occurring at
approximately 15 degrees that was not identifi ed. This peak occurred in all samples,
although the intensity and broadness of this peak was variable.
Table 3.3. Mineral phases associated with each organic carbon source
Organic Carbon Identifi ed Mineral Phases
Lactate (Fe
2+
4
Fe
3+
2
)(OH)
12
CO
3
*2H
2
O; (Fe
2+
2
Fe
3+
)O
4
Uridine (Fe
2+
4
Fe
3+
2
)(OH)
12
CO
3
*2H
2
O
Pyruvate FeCO
3
3.4.2 Comparison of Ferric Iron Oxide Reduction
Reduction of Fe
3+
through time under the conditions tested is shown in Figure 3.5.
HCl (0.5N) is considered to be an effective dissolving agent for most biogenic solid
phase Fe
2+
, including magnetite (18). Therefore, we consider measurements of HCl-
extractable Fe
2+
to be indicators of total ferrous iron oxide production. Relative to the
other samples, iron reduction coupled to pyruvate oxidation was rapid during the fi rst
100 hours before slowing down and reaching a plateau after about 400 hours. About
8mM of Fe
3+
were reduced before all of the pyruvate was exhausted (Fig 3.9b). Over
the course of the experiments, approximately 11mM of Fe
3+
were reduced by strain
W3-18-1 when given pyruvate as the carbon source.
Rates of reduction with lactate and uridine appeared to be similar to each other. There
was a slight lag in the lactate samples during the fi rst 100 hours. The amount of iron
60
reduced with lactate appeared to plateau at about 800 hours, despite the fact that there
was still plenty of lactate available for consumption at this point in the experiments
(Figure 3.9a). Iron reduction with uridine continued to take place until the end of the
experiment, reaching the same levels as the pyruvate samples.
Figure 3.3. Iron oxide reduction through time by strain W3-18-1 using various carbon sources. Fe2+
represent the total HCl-extractable ferrous iron in each culture. Error bars denote standard error from
the mean.
0 200 400 600 800 1000
0
2
4
6
8
10
12
14
Lactate
Pyruvate
Uridine
Control
Time (hours)
61
Iron Carbonate Hydroxide Hydrate
A
B
Figure 3.4. XRD patterns for biominerals and standards using lactate and uridine. X-axes are values in
2-theta degrees. A: XRD pattern for lactate incubation; B: XRD pattern for uridine incubation. X-axis
is in 2-theta degrees.
Magnetite
Iron Carbonate Hydroxide Hydrate
2θ
2θ
Iron Carbonate Hydroxide Hydrate
62
Figure 3.5. XRD pattern for pyruvate incubation experiments. X-axis is in 2-theta degrees. The large
peak occurring at approximately 15 degrees is unidentifi ed.
Siderite
2θ
Pyruvate Experiment
Pyruvate Experiment
63
3.4.2.1 Aqueous Fe
2+
Aqueous phase Fe
2+
is graphed in Figure 3.6. Reduction of ferric iron by W3-18-1
appears to have produced differing amounts of aqueous ferrous iron. The highest
concentrations were reached during incubations in which pyruvate was the organic
carbon source. In this case, a maximum aqueous phase Fe
2+
concentration of
approximately 2.5mM was reached 200 hours into the experiments. This maximum
was followed by a substantial decrease over the remaining time points. Lactate,
on the other hand, showed a steady increase in aqueous Fe
2+
but never achieved
concentrations of greater than 0.5mM. In the case of the uridine incubations, the
ferrous iron in solution built up gradually over 800 hours before decreasing in
concentration at 1000 hours. However, even in this case, the amount of aqueous phase
Fe
2+
never reached the levels seen with the pyruvate incubations.
0 100 200 300 400 500 600 700 800 900 1000
0.0
0.5
1.0
1.5
2.0
2.5
Lactate
Pyruvate
Uridine
Control
Time (hours)
Figure 3.6. Aqueous phase Fe
2+
production by strain W3-18-1 over time. Error bars denote the standard
deviation from the mean
64
3.4.3 Carbon Consumption and Metabolic Byproducts
Organic carbon consumption and the production of metabolites were monitored
throughout the duration of the experiments in order to understand the relative
amounts of iron reduced by various organic compounds (Figure 3.7). Lactate
is present in excess as indicated by the observation that it is never completely
exhausted, which is in agreement with work done previously (18). We detected
production of acetate, formate and succinate, which is consistent with what is known
about the anaerobic metabolism of Shewanella (36, 37). Although Shewanella are
known to oxidize lactate to pyruvate, no pyruvate was detected.
Pyruvate wascompletely consumed by 200 hours (Figure 3.7b). However, after
the pyruvate was exhausted, there was still in increase in the concentration of total
ferrous iron in the cultures subsequent to 200 hours. Most likely, this continuation
of iron reduction was due to the oxidation of formate, which initially increased
but was consumed after all of the pyruvate was used up. Acetate concentrations
rose to approximately 13mM and then remained constant through the duration of
the experiments. Uridine metabolism is shown in Figure 3.7c. Uridine was never
completely used up over the time of the experiments. However, based on the curves
for iron reduction and uridine consumption, it appeared that these reactions were still
proceeding at the time the experiments were concluded (Figure 3.5). HPLC data also
indicated that there was steady acetate production during these incubations.
65
Figure 3.7. Consumption of organic carbon and production of metabolic byproducts using hydrous
ferric oxide as the electron acceptor. A: lactate incubations; B: pyruvate incubations; C: uridine incuba-
tions. Error bars represent standard deviations from the mean. The controls are experiments with the
same initial condition, but no bacteria added. Controls showed only the starting carbon substrates.
66
3.4.4 Chemistry
3.4.4.1 pH
There appeared to be three different patterns of pH change in these experiments
(Figure 3.8). The initial pH for all samples was between 8.5 and 8.7. In all cases,
there was a decrease in pH. However, the magnitude of this change varied depending
on the carbon source utilized. For the samples incubated with pyruvate, the pH
dropped to approximately 7.4 within the fi rst 100 hours of incubation. However, after
400 hours, the pH increased to approximately 7.8. In the case of uridine, there was a
linear decrease in pH over the course of 300 hours, stabilizing at approximately 7.4.
0 200 400 600 800 1000
7.0
7.5
8.0
8.5
9.0
Lactate
Pyruvate
Uridine
Control
Time (hours)
Figure 3.8. pH of cultures over time. Error bars represent standard deviation from the mean.
67
3.4.4.2 Inorganic carbon
Figure 3.9 represents concentrations of solid phase inorganic carbon present in the
samples at the end of the experiment. In all cases, the values for each sample were
above the background values produced by the control samples. In the case of lactate,
inorganic carbon made up approximately 0.3 weight percent of the solid phase
sample. Pyruvate samples produced the most solid phase inorganic carbon, yielding
approximately 2.4 weight percent. Uridine samples contained approximately 0.5
weight percent inorganic carbon. Conditions in these cultures are such that the only
expected solid-phase carbonate would be an iron carbonate of some kind. Based
on the information gathered from XRD analysis of products from the lactate and
uridine experiments indicate that the iron carbonate is in the form of iron carbonate
hydroxide hydrate, or carbonate green rust. For pyruvate, we assume that the iron
carbonate is in the form of siderite, based on XRD analysis.
Figure 3.9. Inorganic carbon production by strain W3-18-1 using three different carbon sources. Error
bars represent the standard deviation from the mean.
68
3.5 Discussion
3.5.1 Consumption of Organic Carbon
Shewanella are known to grow well when given lactate as the carbon and energy
source (11, 17). Consequently, the majority of work done, investigating dissimilatory
iron reduction with these bacteria has been done using lactate as the electron donor (8,
9, 17, 18, 21, 38). While pyruvate is a metabolic byproduct of lactate oxidation (Scott,
1994 #36), no pyruvate was detected via HPLC. It is possible that either the amount
of pyruvate produced was below detection limits, or that it was rapidly utilized so
its concentration did not increase. The latter explanation is more likely as it appears
that strain W3-18-1 can consume pyruvate quite rapidly (Figure 7b). Abiotic control
cultures showed no decrease in lactate concentration. Previous work has shown that
signifi cant amounts of lactate are not consumed when starting concentrations of
lactate are at 20mM (18). We see similar results here, in that only a modest amount of
the available lactate was consumed by W3-18-1 (Figure 7a). It has been hypothesized
that the transformation of the starting oxide material from HFO to more crystalline
forms like goethite or hematite would reduce the amount of bioavailable Fe
3+
(18).
However, other studies suggest that the presence of organic acids, including lactate,
can inhibit the recrystallization of amorphous iron oxide phases (39, 40). If this is
true, the presence of lactate, acetate and formate should have prevented crystallization
from occurring in the lactate cultures; thus keeping the HFO in an amorphous, more
bioavailable state. Additionally, it is unlikely that the change in pH or production of
waste products like acetate served to inhibit further iron reduction; compared to both
the pyruvate and uridine samples, the production of acetate and the change in pH
69
were modest in the cultures given lactate as the carbon source (Figures 3.7a and 3.8).
Previous studies have also demonstrated that an increased pool of aqueous Fe
2+
can
inhibit iron reduction (21). While there was very little aqueous Fe
2+
built up in the
lactate incubations (Figure 3.6), it is possible that a signifi cant amount of reduced iron
is adsorbed to either the cells, or the remaining HFO. The incomplete oxidation of
lactate despite the availability of ferric iron is still not well understood.
In contrast to experiments with lactate, all of the pyruvate was consumed within the
fi rst few hundred hours. Unlike lactate, which must fi rst undergo oxidation, pyruvate
can be split into Acetyl-CoA and either formate or CO
2
, feeding directly into the
Tricarboxylic Acid (TCA) cycle or various other carbon assimilation pathways (36,
37). Providing the bacteria with pyruvate would essentially save a step by giving
them a substrate that they could directly shunt into this pathway. Shewanella can also
convert pyruvate directly to acetate and formate. The ability to direct pyruvate into
various metabolic pathways may explain its rapid utilization. Current understanding
is that Shewanella are incapable of acetate utilization under anaerobic conditions
(2). Other strains in this genus have been reported to produce high concentrations
of acetate under anaerobic conditions (41). This inability to utilize acetate would
explain the build up of acetate in these cultures over the course of the experiments
(Figure 3.7b). Although pyruvate was consumed within the fi rst two hundred hours,
iron reduction continued beyond that time point. This was probably due to the
consumption of formate (Figure 3.7b). While they are unable to utilize formate for
growth, Shewanella are able to couple its oxidation to the reduction of ferric iron
for energy production. Control cultures also showed a decrease in pyruvate, but no
70
increase in metabolites. Organic acids are known to interact with iron, adsorbing to
particulate iron oxides (42-44). In this case, we surmise that the missing pyruvate
may have been adsorbed to the HFO.
Oxidation of uridine by Fe
3+
produced relatively large quantities of acetate (fi gure
3.7c). The genome for W3-18-1 has been sequenced, allowing for construction of
putative metabolic pathways for various organic carbon sources (KEGG Database;
http://www.genome.jp/kegg/). In the case of uridine metabolism, acetate is not
predicted to be a direct metabolic byproduct of uridine metabolism. However, the
genomic data for strain W3-18-1 predicts that the products of uridine metabolism can
be sent into pathways, such as the pentose phosphate pathway, which might produce
acetate as a metabolic byproduct. HPLC chromatograms indicated the presence
of additional metabolites that were not identifi ed (data not shown). Further work
is needed to understand the pathway(s) used by W3-18-1 to process uridine under
anaerobic conditions.
3.5.3 Reaction Stoichiometry
Based on the results of anaerobic respiration by strain W3-18-1 using lactate,
pyruvate and uridine, the proposed stoichiometry for each reaction is listed in table
3.4. While there was cell growth during the fi rst few hundred hours of incubations,
by the end of the experiments cell populations had decreased such that cell growth
was negligible (data not shown). Therefore, the amount of organic carbon accounted
for by biomass would fall into the uncertainty of the equations, but it is likely to
71
be insignifi cant to the overall Stoichiometry of the reactions. Because of this, the
reactions do not describe biomass production.
Table 3.4. Stoichiometry for reactions with lactate, pyruvate and uridine
Experimental Data
Lactate
Pyruvate
Uridine
Our results indicate that under these conditions, strain W3-18-1 reduced
approximately half of the available HFO, which is in agreement with previous studies
(17, 18). The remaining ferric iron was incorporated into magnetite, iron carbonate
green rust, or remained in a hydrated ferric oxide form. The ratio for carbonate green
rust in the lactate equation was derived from the fraction of solid material composed
of inorganic carbon (Figure 3.9). The assumption is that all of the insoluble carbonate
is in the form of iron carbonate hydroxide. Other researchers have reported the
production of iron carbonate minerals in laboratory settings (9, 17, 32, 45). In most
cases, the conditions under which these minerals are produced involve providing the
cultures with a large amount of CO
2
both in the media buffer and in the headspace.
5C
3
H
5
O
3
−
+11.3Fe(OH )
3
→
0.7Fe
3
O
4
+ 0.6Fe
6
(OH )
12
CO
3
•2H
2
O + 0.5Fe
2+
(aq) + 5.1Fe(OH )
2
+4.3C
2
H
3
O
2
−
+ C
4
H
6
O
4
+1.8HCO
3
−
+ 7.7H
2
O + 3H
+
+ 2.9e
−
12C
3
H
3
O
3
−
+11Fe(OH )
3
+ 3.6H
2
O → 3.4FeCO
3
+ 0.3Fe
2+
(aq)
+7.3Fe(OH )
2
+12.5C
2
H
3
O
2
−
+ 7.6HCO
3
−
+16.5H
+
+ 9e
−
7C
9
H
12
N
2
O
5
+ 11Fe (OH )
3
+ 102 .2H
2
O →
0.6Fe
6
(OH )
12
CO
3
• 2H
2
O + 0.1Fe
2 +
( aq ) + 6.4 Fe (OH )
2
+10C
2
H
3
O
2
−
+ 42.4 HCO
3
−
+ 14 NH
4
+
170 .6H
+
+ 132 .3e
−
72
O’Laughlin et al (46) have reported the production of iron carbonate hydroxide using
argon headspace and formate as the electron donor; while Lee et al (15) have reported
the production of siderite using N
2
gas and organic buffered media when using
pyruvate as the electron donor. However, under conditions in which the headspace is
composed of pure nitrogen gas, the typical iron biomineral product of dissimilatory
HFO reduction is magnetite (9, 17, 45). The amount of magnetite produced in these
reactions was estimated from previous work comparing the products of lactate
oxidation coupled to iron reduction by different strains of Shewanella (Chapter 2).
In all the carbon sources tested, a large portion of the reduced iron was neither in
aqueous phase or tied to an identifi able mineral. In this case, we describe this fraction
of solid ferrous iron as a generic ferrous hydroxide Fe(OH)
2
. It is possible that this
material was in the form of an amorphous, solid phase iron. It could also be that the
material was adsorbed to the cells, either directly or via a biofi lm, or to the unreduced
HFO. Further work is required in order to gain a better understanding the nature of
the remaining ferrous iron.
Because not all of the metabolic byproducts of the uridine incubations were
identifi ed, the proposed equation is an approximation of the reaction involved in
uridine oxidation by strain W3-18-1. However, even in this rudimentary form, there
is the suggestion that uridine metabolism produces a signifi cant number of protons,
which would account for the 1+ unit drop in pH (Figure 3.8).
73
3.5.4 Solution Chemistry and the Infl uence of Organic Carbon
The interaction between organic carbon and iron oxides is important not only in
terms of redox chemistry, but also in the infl uence that organics have on the solubility
and crystallinity of iron solids in soils and sediments (39, 40, 42, 44). In terms of
dissimilatory iron reduction, there is not a good understanding on how different types
of organics can infl uence the rates of iron reduction, the amount of iron reduced, or
the identity of the secondary mineral products. There has been work done by previous
researchers to understand the role that specifi c organic substrates play in determining
the formation of particular reduced iron biominerals (24). These workers have
reported differences in the products of iron reduction depending on the type organic
acid used as the energy source and electron acceptor. In this case, we report similar
results with the exception that strain W3-18-1 produced a mixture of magnetite and
iron carbonate hydroxide, rather than pure magnetite, when incubated with lactate
(fi gure 3.4a). While the concentration of aqueous phase Fe
2+
increased over time, the
concentrations relative to the total amount of reduced iron were modest, indicating
that most of the iron remained on the solid phase. It is possible that the reduced
iron is being immobilized by the cells, preventing the Fe
2+
from going into solution.
However, the behavior of the aqueous phase iron produced as a result of dissimilatory
iron reduction coupled to lactate oxidation is different from what is seen with
pyruvate and uridine (fi gure 3.6).
W3-18-1 produced siderite when iron reduction was coupled to pyruvate oxidation.
This is in agreement with reports published for other strains of Shewanella when
given pyruvate as the electron donor (24). However, in this case, XRD patterns
74
indicate that strain W3-18-1 generated a signifi cant signal at 15 degrees which was
not readily identifi able via the known crystallographic databases (Figure 3.5). The
nature of this signal is unkown. Pyruvate samples reduced slightly more than half
of the starting hydrous ferric oxide material, although the rate of reduction began
to decrease after 200 hours (Figure 3.7). This may be accounted for by the fact that
pyruvate had been completely consumed by this point, forcing the bacteria to begin
utilizing the formate that was produced during pyruvate oxidation. This may also
account for the changes in pH during the course of the experiments. Although the
oxidation of pyruvate liberates protons, the oxidation of formate consumes them.
The data collected indicated that the pH of the solutions experienced a precipitous
drop of over one unit during the fi rst 200 hours, upon which the pH began a gradual
rise back to 8. The oxidation of formate coupled to iron reduction has been shown
to lead to increase in pH (46). If W3-18-1was indeed coupling formate oxidation
to iron reduction, which appears to be the case in the pyruvate samples, this would
explain the pH increase of the pyruvate incubations. The uridine samples accounted
for approximately the same amount of reduced iron as the pyruvate samples at the
conclusion of the experiments (Figure 3.7). The pH in these samples decreased
steadily, stabilizing at approximately 7.4-7.5 after 400 hours. The utilization of
uridine as an electron donor led to the formation of large, well defi ned hexagonal
plates (Figure 3.1b). XRD analysis determined that these structures were iron
carbonate hydroxide, more commonly known as green rust. Green rust is a precursor
to a mineral known as fougerite, which is common in anoxic, waterlogged soils (35).
There were differences in the amount of aqueous Fe
2+
. It is not entirely clear what
changes in physiology brought about by the use of specifi c organic carbon sources
75
would lead to differences in the buildup of aqueous Fe
2+
present in each incubation.
It is worth noting, however, that the buildup of aqueous ferrous iron corresponds to
the large increase in acetate concentration for both the pyruvate and uridine samples.
Incubations of lactate and uridine with elevated levels of acetate did not lead to
differences in the buildup of aqueous ferrous iron (Figure 3.8). While the faster initial
rates of Fe
2+
production by the pyruvate samples might explain the early buildup of
ferrous iron in solution, they do not explain the buildup in the uridine samples, as
those rates of iron reduction were quite similar to the rates from the lactate group.
Further work is needed to determine what role acetate has to play in determining
the identity of reduced iron biominerals. It has been postulated that the rate of Fe
2+

production as a result of dissimilatory iron reduction would impact the identity of
the resulting secondary minerals (9).Faster rates of reduction would favor magnetite,
while slower rates would favor green rust (14). However, in this case, incubations
with pyruvate yielded faster rates of Fe
2+
production than incubations with either
uridine or lactate. Yet, the products of iron reduction coupled to pyruvate oxidation
were siderite (Figure 3.5). Additionally, while the rates of reduction for lactate and
uridine were quite similar (Figure 3.7), in one case, a mixture of magnetite and green
rust were formed; while in another, green rust and some goethite were the observed
mineral products (Figure 3.4).
There was a slight drop in pH with the lactate incubations; however this change
was not as pronounced as the changes seen with the other organic carbon sources
(Figure 3.8). The oxidation of lactate coupled to iron reduction is a proton consuming
process (18). This would lead to a prediction that lactate oxidation would result in an
increase in pH. However, the processes of magnetite and iron carbonate hydroxide
76
6 7 8 9 10 11 12 13 14
0
2
4
6
8
10
12
14
16
18
20
22
24
Lactate
Fe
3
O
4
Fe
6
(OH)
12
CO
3
*2H
2
O
FeCO
3
Fe(OH)
3
Pyruvate
Uridine
formation both liberate protons (18), potentially accounting for the modest decrease
in pH. Additionally, typical heterotrophic metabolism is a CO
2
generating process.
While aqueous inorganic carbon was not measured, there was a detectable amount
of inorganic carbon in solid phase (Figure 3.9). It is unlikely that all of the CO
2

produced by W3-18-1 would be locked up in solid material.
-log [Fe
2+
] (M)
Figure 3.10. Thermodynamic stability fi eld of hydrous ferric oxide (assumed stoichiometry: Fe(OH)
3
),
magnetite (Fe
3
O
4
), green rust (Fe
6
(OH)
12
CO
2
*2H
2
O) and siderite (FeCO
3
) as a function of Fe
2+

concentration and pH. A [HCO
3
-
] of 0.03M was assumed, based on buffer concentration. Points show
location of incubation experiments after 1000 hours.
pH
77
The stability fi elds of magnetite, siderite and green rust indicate that the predicted
products for all three carbon sources should be magnetite (Figure 3.10). However, in
none of the cases observed here was magnetite the sole product of dissimilatory iron
reduction by strain W3-18-1. In contrast, the major product of iron reduction coupled
to pyruvate oxidation was siderite, while the major product of uridine oxidation was
iron carbonate hydroxide. This would suggest that that solution chemistry alone is not
enough to predict the nature of reduced iron biominerals. The conditions presented
in Figure 3.10 describe solution chemistry for the entire sample. It may be that on a
smaller scale (i.e. at the microbe-mineral interface) the conditions are such that the
production of the observed precipitates is favored. This might point to physiologic
difference in strain W3-18-1 depending on the carbon substrate used for energy.
A more thorough investigation needs to be done to ascertain the degree to which
physiologic differences in the metabolism of these carbon sources impacts the solid-
phase products of dissimilatory iron reduction. Additionally, given the apparent
differences of the reduced iron, in its distribution between mineral types, unidentifi ed
amorphous phases and aqueous concentrations, further work needs to be done using a
wider range of organic substrates. Shewanella species are capable of utilizing a wide
range of organic carbon sources (11, 28, 47). It would be of some utility to understand
the range of iron minerals produced by these bacteria when given a variety of carbon
sources. Also, as HFO is not the only available form of ferric iron in the environment,
it would be of some interest to explore the impact of using a variety of carbon sources
with more crystalline material.
78
Chapter 3 Endnotes
1. Lovley, D. R. & Phillips, E. J. P. (1986) Applied and Environmental
Microbiology 51, 683-689.
2. Lovley, D. R. (1991) Microbiological Reviews 55, 259-287.
3. Nealson, K. H., P., W., & Ew, d. J. (1983) in Biomineralization and Biological
Metal Accumulation, ed. Anonymous (D. Reidel Publishing Company), pp.
459-479.
4. Nealson, K. H. & Saffarini, D. (1994) Annual Review of Microbiology 48,
311-343.
5. Canfi eld, D. E. & Marais, D. J. D. (1993) Geochimica Et Cosmochimica Acta
57, 3971-3984.
6. Lovley, D. R. (1995) Advances in Agronomy, V ol 54 54, 175-231.
7. Cooper, D. C., Picardal, F. F., & Coby, A. J. (2006) Environmental Science &
Technology 40, 1884-1891.
8. Zachara, J. M., Fredrickson, J. K., Smith, S. C., & Gassman, P. L. (2001)
Geochimica Et Cosmochimica Acta 65, 75-93.
9. Zachara, J. M., Kukkadapu, R. K., Fredrickson, J. K., Gorby, Y . A., & Smith,
S. C. (2002) Geomicrobiology Journal 19, 179-207.
10. Lovley, D. R. & Phillips, E. J. P. (1988) Applied and Environmental
Microbiology 54, 1472-1480.
11. Myers, C. R. & Nealson, K. H. (1988) Science 240, 1319-1321.
12. Roden, E. E. & Zachara, J. M. (1996) Environmental Science & Technology
30, 1618-1628.
13. Kostka, J. E., Haefele, E., Viehweger, R., & Stucki, J. W. (1999)
Environmental Science & Technology 33, 3127-3133.
79
14. Ona-Nguema, G., Abdelmoula, M., Jorand, F., Benali, O., Gehin, A., Block, J.
C., & Genin, J. M. R. (2002) Environmental Science & Technology 36, 16-20.
15. Lee, S. H., Lee, I. S., & Roh, Y . (2003) Geosciences Journal 7, 217-226.
16. Glasauer, S., Weidler, P. G., Langley, S., & Beveridge, T. J. (2003)
Geochimica Et Cosmochimica Acta 67, 1277-1288.
17. Fredrickson, J. K., Zachara, J. M., Kennedy, D. W., Dong, H. L., Onstott, T.
C., Hinman, N. W., & Li, S. M. (1998) Geochimica Et Cosmochimica Acta
62, 3239-3257.
18. Fredrickson, J. K., Kota, S., Kukkadapu, R. K., Liu, C. X., & Zachara, J. M.
(2003) Biodegradation 14, 91-103.
19. Roden, E. E. (2006) Comptes Rendus Geoscience 338, 456-467.
20. Jaisi, D. P., Kukkadapu, R. K., Eberl, D. D., & Dong, H. L. (2005)
Geochimica Et Cosmochimica Acta 69, 5429-5440.
21. Liu, C. G., Zachara, J. M., Gorby, Y . A., Szecsody, J. E., & Brown, C. F.
(2001) Environmental Science & Technology 35, 1385-1393.
22. Cooper, D. C., Picardal, F., Rivera, J., & Talbot, C. (2000) Environmental
Science & Technology 34, 100-106.
23. Parmar, N., Gorby, Y . A., Beveridge, T. J., & Ferris, F. G. (2001)
Geomicrobiology Journal 18, 375-385.
24. Lee, J. H., Roh, Y ., Kim, K. W., & Hu, H. G. (2007) Geomicrobiology Journal
24, 31-41.
25. Finke, N., Vandieken, V ., & Jorgensen, B. B. (2007) Fems Microbiology
Ecology 59, 10-22.
26. Lovley, D. R. & Phillips, E. J. P. (1989) Applied and Environmental
Microbiology 55, 3234-3236.
27. Mcmahon, P. B. & Chapelle, F. H. (1991) Nature 349, 233-235.
80
28. Pinchuk, G. E., Ammons, C., Culley, D. E., Li, S. M. W., McLean, J. S.,
Romine, M. F., Nealson, K. H., Fredrickson, J. K., & Beliaev, A. S. (2008)
Applied and Environmental Microbiology 74, 1198-1208.
29. Stapleton, R. D., Sabree, Z. L., Palumbo, A. V ., Moyer, C. L., Devol, A. H.,
Roh, Y ., & Zhou, J. Z. (2005) Aquatic Microbial Ecology 38, 81-91.
30. Cornell, R. M. & Schwertmann, U. (1996) The Iron Oxides: Structure,
Properties, Reactions, Occurrence and Uses (VCH, Weinheim).
31. Stookey, L. L. (1970) Analytical Chemistry 42, 779-781.
32. Roh, Y ., Gao, H. C., Vali, H., Kennedy, D. W., Yang, Z. K., Gao, W. M.,
Dohnalkova, A. C., Stapleton, R. D., Moon, J. W., Phelps, T. J., et al. (2006)
Applied and Environmental Microbiology 72, 3236-3244.
33. Genin, J. M. R., Refait, P., Bourrie, G., Abdelmoula, M., & Trolard, F. (2001)
Appl Geochem 16, 559-570.
34. Kukkadapu, R. K., Zachara, J. M., Fredrickson, J. K., Kennedy, D. W.,
Dohnalkova, A. C., & McCready, D. E. (2005) American Mineralogist 90,
510-515.
35. Genin, J. M. R., Aissa, R., Gehin, A., Abdelmoula, M., Benali, O., Ernstsen,
V ., Ona-Nguema, G., Upadhyay, C., & Ruby, C. (2005) Solid State Sci 7, 545-
572.
36. Scott, J. H. & Nealson, K. H. (1994) Journal of Bacteriology 176, 3408-3411.
37. Tang, Y . J., Meadows, A. L., Kirby, J., & Keasling, J. D. (2007) Journal of
Bacteriology 189, 894-901.
38. Hansel, C. M., Benner, S. G., Neiss, J., Dohnalkova, A., Kukkadapu, R. K., &
Fendorf, S. (2003) Geochimica Et Cosmochimica Acta 67, 2977-2992.
39. Cornell, R. M. & Schwertmann, U. (1979) Clays and Clay Minerals 27, 402-
410.
40. Schwertmann, U. (1966) Nature 212, 645-646.
81
41. Lovley, D. R., Phillips, E. J. P., & Lonergan, D. J. (1989) Applied and
Environmental Microbiology 55, 700-706.
42. Cheah, S. F., Kraemer, S. M., Cervini-Silva, J., & Sposito, G. (2003)
Chemical Geology 198, 63-75.
43. Essington, M. E., Nelson, J. B., & Holden, W. L. (2005) Soil Science Society
of America Journal 69, 996-1008.
44. Gu, B. H., Schmitt, J., Chen, Z., Liang, L. Y ., & McCarthy, J. F. (1995)
Geochimica Et Cosmochimica Acta 59, 219-229.
45. Mortimer, R. J. G. & Coleman, M. L. (1997) Geochimica Et Cosmochimica
Acta 61, 1705-1711.
46. O’Loughlin, E. J., Larese-Casanova, P., & Cook, R. (2007) Geomicrobiology
Journal 24, 211-230.
47. Wang, Y ., Kan, J., T.V ., K., Obraztsova, A., & Nealson, K. H. (2008) in
American Society for Microbiology 108th General Meeting (Boston, MA.).
82
CHAPTER FOUR
HOW IMPORTANT ARE SHEWANELLA TO THE CYCLING OF
IRON AND ORGANIC CARBON IN SEDIMENTS?
4.1 Introduction
The previous two chapters described laboratory studies investigating the potential
importance of using specifi c strains Shewanella and specifi c carbon sources on
both the rates and products of dissimilatory iron reduction. In this chapter, I will
attempt to place this work into a larger environmental context. While these bacteria
enjoy widespread use as laboratory models for investigating various aspects of
environmental microbiology, attempts to determine the distribution and relative
abundance of Shewanellae in the environment have provided mixed results. This
has lead many researchers to question their importance in the context of global
biogeochemistry. Before delving into the question of the preceding work’s relevance,
this chapter will discuss some general background on iron chemistry, sedimentary
organic carbon and the distribution of dissimilatory iron reducers.
4.2 Iron
Iron has been described as the most important metal in the universe (1, 2). While this
might be interpreted as a facetious comment, it is worth considering the properties
possessed by this element that make it an important factor to biology. Indeed, our
83
planet’s magnetic fi eld, which life depends on for its existence, is a result of iron’s
ferromagnetic properties. In mammals, iron is important enough that it is recycled
and stored when not in use; outside of bleeding, there is no way for mammals to lose
any iron (2). Iron is the fourth most abundant element, by weight percent, found in
the earth’s crust. Of the eight elements that account for approximately 98% of the
earth’s crust, iron is the only transition metal to be found in the group; it is 10 times
more abundant than the next most common transition metal (titanium).
Transition metals possess a chemistry that allows them to engage in more complex
redox reactions than several of the other elements found on earth. They are redox
sensitive compounds that form more than one cation (3). While this characteristic is
not unique to transition metals (i.e. sulfur, nitrogen, cholorine), it is far less common
outside of the transition elements. Iron, for instance, is able to exist in two valence
states (Fe
2+
and Fe
3+
) under conditions found in the modern earth. Other oxidation
states of iron are possible, such as Fe
4+
, Fe
5+
and Fe
6+
, however these require
specialized conditions and are subject to rapid dismutation to Fe
3+
and Fe
2+
(4-6).
The changes in redox state between Fe
2+
and Fe
3+
can occur quite easily in nature.
This seems to make it ideal for mediating electron transfer, and it is a major
component of the ferredoxins and cytochromes that make up the electron transport
chain, which every organism on the planet requires for energy, and thus, life. In
environmental settings, iron redox changes typically take place at the interface
between oxic and suboxic environments (1, 7-9). The more reduced form, Fe
2+
, is
typically seen in suboxic environments and has a higher solubility than the ferric ion
form (7). It is often seen in a solid phase as a component of pyrite (FeS
2
), siderite
84
(FeCO
3
), olivine (Fe
2
SiO
4
), pyroxenes (FeCaSi
2
O
6
, MgFeSi
2
O
6
, Fe
2
Si
2
O
6
) and
mixed valence minerals like magnetite (Fe
3
O
4
). Fe
2+
is readily oxidized to Fe
3+
under
aerobic, circumneutral pH conditions. Under these conditions, Fe
3+
forms insoluble
iron oxides and oxyhydroxides such as amorphous ferrihydrite (HFO), goethite
( α-FeO(OH)), lepidocrosite ( γ-FeO(OH)) or hematite ( α−Fe
2
O
3
), among others.
In soils and sediments, Fe
3+
concentrations frequently exceed that of other electron
acceptors, such as oxygen, nitrate and sulfate (10-16). This, coupled with its redox
sensitivity, means that iron can both be impacted by, and have a signifi cant impact
on, organic and inorganic compounds like carbon, sulfur, phosphorous and a variety
of trace metals (1, 7, 17). Iron oxyhydroxides are good adsorbers of phosphates (10,
18-20), which impact their concentrations in sediment pore waters (12, 21, 22). In the
Santa Barbara Basin, Reimers et al. (22) have observed that increases in pore water
phosphate concentrations correlate with the horizon at which iron oxide reduction
occurs. In soils, the pore water concentration of phosphate is typically below 20 μM,
despite the fact that soils are a large reservoir for P (23). In goethite-rich materials,
phosphate adsorption is so strong it is partly irreversible (7). These interactions
appear to result from ligand exchange with OH groups (24, 25). Interactions between
ferric iron and phosphorous can inhibit ferrihydrite transformation into goethite or
hematite (18, 26). The nature of the iron oxides present in soils and sediments is of
great importance from the perspective of dissimilatory iron reduction. Laboratory
studies have demonstrated that differences in the degree of crystallinity of the iron
oxide substrate can affect the overall rate and amount of iron reduction. Microbes
have been shown to have the capacity of reducing goethite, hematite, lepidocrocite,
akaganeite and magnetite under laboratory conditions (27-32). However, the rate and
85
extent of reduction can be much less than when iron reducers are given HFO (31, 33).
Additionally, incubations with mixtures of HFO and goethite have shown that iron
reducers will preferentially reduce HFO (15).
The most signifi cant abiotic reductant of iron oxides is hydrogen sulfi de (17).
The following reactions can proceed rapidly with poorly crystalline iron oxides,
generating solid phase iron monosulfi des (FeS) or FeS
2
(11, 17, 34):
2FeO(OH ) + H
2
S + 4H
+
→ 2Fe
2+
+ S
(O )
+ 4H
2
O (1)
Fe
2+
+ H
2
S → FeS + 2H
+
(2)
FeS + H
2
S → FeS
2
+ 2H
+
(3)
It is believed that when sulfate is present in large quantities, the diffusion of sulfi de,
generated via sulfate reduction, into the iron reduction zone will out-compete any
bacterially mediated iron reduction due to the differences in kinetics (34, 35).
However, this view is not universally accepted (11, 17, 36, 37). Although attempts
have been made to quantify the rates of iron reduction in sulfate-rich sediments
(38-40), these techniques are indirect and require extrapolating dissimilatory iron
reduction by subtracting the carbon oxidation rates attributed to sulfate reduction
from the total carbon oxidation rates (41). This is typically done under laboratory
settings and assumes that all other forms of anaerobic respiration such as nitrate
reduction, nitrite reduction, manganese reduction, etc., are negligible. There is one
undeniable impact that sulfi de will have on iron cycling in sediments: the presence of
sulfi des in appreciable quantities inevitably leads to the burial of reduced iron in the
86
form iron sulfi des. This would prevent the diffusion of Fe
2+
into the oxygenic zone
where it could be recycled back to Fe
3+
, thus sequestering this iron from the pool
available for biogeochemical cycling.
Iron oxides are good sorbents of trace metals. As such, any changes in the oxidation
state of iron will have an impact on the mobility of these trace metals, several of
which are considered to be contaminants. This has generated extensive research into
understanding the impact of iron reduction of sediment metal chemistry (31, 42-48).
Reduction of HFO has been shown to release arsenate (As
5+
), arsenite (As
3+
) and
Ni
2+
under laboratory conditions (42, 44). Zachara et al (46) have demonstrated that
microbial reduction of Co
2+
substituted goethite can cause its release into solution. In
environmental settings, dissolution of goethite and hematite-rich soils has been shown
to release signifi cant quantities of Co, Cr, Cu, Mn, Ni and Zn (49). The examination
of pore-water profi les from a contaminated harbor indicates that the release of Co, Ni,
Cu and Cd is associated with the reductive dissolution of iron oxide (50). Conversely,
the generation of iron carbonate green rusts can remove several of these compounds
from pore-waters (43, 45, 51). In these instances, trace metals like Ni, U and Tc were
incorporated into the sites normally occupied by Fe
2+
.
From what we know based on the rock record, iron redox cycling has been a globally
important phenomenon throughout Earth’s history. The most observable indicators
of this are massive iron-rich deposits dating to the Precambrian. These deposits
are more commonly known as banded iron formations (BIF). Iron formations are
typically defi ned as “a chemical sediment, typically thin bedded or laminated, whose
principal chemical characteristic is an anomalously high content of iron, commonly
87
but not necessarily containing layers of chert (52).” Included in this defi nition was
a quantitative lower limit of 15 wt%, but subsequent authors consider this too be
arbitrary and restrictive (53). While wt% of iron can vary, typical values for Archean
deposits average a little less than 30 wt%, while values of up to 40 wt% have been
reported for iron formations from the Rapitan and Urucum Groups (53). Although
iron formations are commonly referred to as BIFs—banded iron formations—most
well-banded iron formations are older than about 2.0 Ga (53). BIFs have occurred
sporadically throughout the late Archean and Paleoproterozoic, but disappeared as
oxygen levels rose to a concentration allowing for changes in the redox state of the
ocean (54).
By ca. 1.8 Ga, widespread BIF deposition had ceased. However, 1 billion years
later, they return, in association with the Neoproterozoic glacial deposits. Economic
Neoproterozoic banded iron formations are found in northwestern Canada, Southern
Australia (55), Namibia (56), Saudi Arabia (57), China (58), and Brazil (59-61).
These deposits are associated with glaciogenic lithologies such as diamictites and
dropstones, and most are associated with rift basins. However, the use of the term
“banded” is relative; most do not display the fi ne laminations of alternating chert and
iron rich layers characteristic of the Archean and Paleoproterozoic examples. Rather,
most are iron-enriched siliciclastic sediments with little banding (the Rapitan Iron
Formation of NW Canada is a notable exception). Although worldwide glaciations
are regularly called upon to explain the presence of iron formation in this time (53,
62-64), the true cause of their deposition remains a mystery.
88
4.3 Organic Carbon in Sediments
The majority of organic carbon produced in marine environments is remineralized
in the water column. While exact values are subject to spatial and temporal
considerations, in general only a small fraction of organic carbon ever reaches the
sediment (65-67). As stated, these values are highly variable depending on the when
and where, and can range from less than 1 %
dry weight
gram of organic carbon per gram
of sediment in pelagic settings, to almost 20% in coastal and estuarine environments
(68). The extent to which organic carbon is remineralized in sediments is subject
to both the ‘quality’ and the fl ux of the organic carbon (69-72). The term ‘quality’
is poorly defi ned, but it appears to be used by researchers to describe the degree to
which organic carbon is susceptible to degradation. Within sediments, the depth and
rate at which organic carbon is remineralized will impact both the availability of
carbon to other geochemical cycles, as well as the amount of organic carbon that is
buried in sediments. The extent of organic carbon burial in sediments is important, as
it has implications for the levels of O
2
and CO
2
both in the water column and in the
atmosphere (68). Because of this, there has been great interest in trying to understand
the nature of both the particulate organic carbon (POC) and dissolved organic carbon
(DOC) that manages to make its way into sedimentary environments.
By far, the largest body of work trying to understand the nature and degradation of
sedimentary organic matter has involved volatile fatty acids (VFA). VFA are low
molecular weight carbon compounds, containing 6 carbons or fewer, which are the
byproducts of fermentative processes (73-75). Some typical examples are compounds
like lactate (C
3
H
5
O
3
-
), acetate (C
2
H
3
O
2
-
), pyruvate (C
3
H
3
O
3
-
) and propionate
89
(C
2
H
5
O
2
-
). In marine sediments, sulfate reduction and iron reduction are considered
to be the most signifi cant terminal electron acceptors in the upper anoxic zones (41,
76). Investigations aimed at identifying the substrates used by sulfate reducers have
shown that VFA and hydrogen are the major organic substrates utilized (72, 73, 77-
79), although recently there has been some work indicating that this may not be true
everywhere (39). Nevertheless, these compounds appear to have a near universal
presence in sedimentary environments. Because of the diffi culties in directly
quantifying iron reduction rates in sulfate-rich sediments (41), no studies exist
identifying the major substrates utilized by iron reducers. In freshwater sediments,
however, there have been assertions that acetate is the major organic carbon source
used by iron reducing bacteria (80-82).
In addition to VFA, work has been done attempting to characterize the potential for
larger compounds like sugars, amino acids, nucleic acids and hydrocarbons to act
as substrates for anaerobic respiration. Characterization of amino acids in sediments
indicates that they can make up anywhere from 10%-20% of the organic carbon that
is remineralized (83, 84). Extracellular DNA is widespread in both the DOC and POC
pools of lacustrine and marine environments (85, 86). Laboratory and fi eld studies
have shown that DNA is not only a source of carbon, but also a potentially important
source of nitrogen and phosphorus (87). While workers have shown that up to 70%
of extracellular DNA is hydrolysable (86), it has also been shown that microbes will
show a preference towards low molecular weight nutrients over DNA when presented
with both (87). Additionally, it is not clear how much of the DNA that is taken up is
metabolized and how much is incorporated into the microbe’s genome.
90
Most of the techniques used to identify organic matter composition and concentration
require a processing step that destroys the macromolecular structure (84, 88-90). As
such, while we have an understanding of the individual components that make up
organic matter—VFA, amino acids, nucleotides, etc.—we lack a clear understanding
of the macromolecular nature of POC. Conventional wisdom states that
remineralization of organic carbon follows a series of steps: POC is broken down into
high molecular weight DOC, which is then broken down into low molecular weight
DOC. These compounds are fermented into VFA, which are fi nally remineralized into
CO
2
. However, not all POC is eventually remineralized; some portion is ultimately
buried. The fraction of organic carbon that is buried is variable, and ultimately
depends on factors we do not yet understand.
4.4 Dissimilatory Iron Reducers
The widespread distribution of iron oxides in sediments, and the favorable energetics
of iron reduction under anaerobic conditions have lead to the exploitation of this
redox reaction by biology. Dissimilatory iron reducers are organisms that have
developed the ability to use iron as a terminal electron acceptor in their electron
transport chain. What makes this different from more mainstream forms of respiration
(i.e. aerobic respiration) is that it requires the cell to transfer electrons to a substrate
that is solid and external. Despite this apparent diffi culty, dissimilatory iron reducers
have been found within several groups of bacteria and have proven to be ubiquitous
in iron-rich sedimentary environments (82, 91-93).
91
The majority of the work that has been done studying the physiology, genetics and
mineralogy of iron reduction has been done on organisms from two groups within
the delta and gamma proteobacteria: Geobacter (Geobacteraceae) and Shewanella
(81, 91, 94a). Geobacter metallireducens strain GS-15 was isolated from iron-rich
sediments of the Potomac River (95). GS-15 is an obligate anaerobe that reduces iron
using acetate as an electron donor. This makes it a potentially important organism
in anoxic environments where acetate is considered to be a key intermediate in the
degradation of complex organic matter. Geobacteraceae contains several bacteria that
are able to couple Fe
3+
reduction to the oxidation of a variety of organic compounds,
including hydrogen (82).
The Shewanella genus encompasses several strains that have been shown to be
capable of dissimilatory Fe
3+
reduction. Shewanella are gram-negative, facultative
anaerobes belonging to the γ-proteobacteria. They have a rapid generation time under
aerobic conditions, making them ideal for genetic and physiological studies. Although
Shewanella species are thought to be incapable of acetate utilization under anaerobic
conditions, they are able to use a myriad of other carbon compounds including
lactate, pyruvate, some sugars, amino acids and nucleic acids. They have also been
shown to posses the capacity for metal reduction using hydrogen as an electron donor
(95, 96).
While these two groups of bacteria are the best studies iron reducers, other iron
reducers have been isolated among the Acidobacteria, Thermodesulfobacteria,
Firmicutes and Euryarchaea, among others. (97-99). Although their global
signifi cance to iron biogeochemical cycling is still not well understood, their presence
92
and the subsequent identifi cation of 16s rDNA sequences most similar to these
groups in subsurface sediments suggests that they could be globally signifi cant (92).
However, thus far, many of the putative iron reducers identifi ed by molecular biology
techniques have yet to be cultured in the laboratory.
4.5 The Potential Importance of Shewanella in Sedimentary Environments
Shewanella are widely used in laboratory settings to investigate basic physiology
(100, 101), enzymology (102), gene expression (103), metabolic diversity (9, 104-
106), microbe-mineral interactions (96, 107) and bioremediation (44, 108). These
organisms have been isolated from a wide variety of locations: fresh-water lake
sediments (109), brackish waters (110), meromictic seas (111), tidal marshes (112),
river deltas (113), deep-sea sediments (114), hydrothermal vents (115), uranium
mines (107) and other creatures (116). These organisms are relatively easy to isolate,
have rapid generation times and are diverse with respect to both carbon source and
electron acceptor utilization. However, despite their near ubiquity both in the lab and
in the environment, their relevance to biogeochemical processes on a global scale has
been called into question practically since they were fi rst isolated (80-82).
These objections stem from work done attempting to characterize microbial
communities from a variety of locations. In these settings, conclusions concerning the
relative abundance of Shewanella in specifi c locations has been mixed. For example,
in some deep-sea sediments they have been found to comprise a signifi cant portion
of the population (117, 118), while in others they have not (119). While they have
been found present in some contaminated aquifers (120), they have not been detected
93
in others (121). The argument that has been put forth is that because an organism is
easy to isolate does not make it the dominant organism for that environment (82).
However, the reverse would also apply in that ease of isolation does not exclude an
organism from importance.
The second objection to their potential importance stems from their inability to utilize
acetate as a carbon substrate under anaerobic conditions (122). VFA have received
widespread attention due to their near ubiquity in sedimentary environements. Acetate
has been found in detectable quantities in several locations (39, 75, 79, 88), and both
laboratory and fi eld-based studies have indicated that acetate is consumed by other
iron reducing bacteria (14, 123-125). However, the assumption is that all of the VFA
in these environments is acetate, and that all of this acetate is of fermentative origin.
5 strains of Shewanella that were used to study the relationship between iron oxide
respiration rate per attached cell as a function of cell density (Table 4.1). The aim here
was to understand how much iron a cell was capable of reducing, and whether that
would translate into something reasonable when applied to the environment. There
have been studies suggestive of the possibility that Shewanella produce extra-cellular
Table 4.1. Bacterial strains used in respiration rate calculations
Bacterial Strain Origin
MR-1 Lake Oneida Sediments, New York
CN32 Uranium Mine, New Mexico
MR-4 Black Sea Water Column
OS155 Baltic Sea Water Column
W3-18-1 Pacifi c Ocean Sediments
94
organic compounds that can be used as electron shuttles (126-128). This would
remove the need for direct attachment to the ferric iron in order to use it as a terminal
electron acceptor. However, the exact impact of these shuttles on overall reduction
rates by Shewanella, as well as their distribution in other strains beyond MR-1, has
not been demonstrated. As such, the assumption here will be that the Shewanella
strains tested require direct attachment to the iron oxides in order to respire them.
A comparison of Fe
3+
reduction rates gathered from various studies is shown in
Table 4.2. Included in this table are data from work described in chapter 2, chapter
3 and appendix A of this work. As can be seen, the majority of work that has been
done investigating iron reduction has been done using strain CN32. However, this
is starting to change as more and more strains are isolated. In general, there appears
to be a wide distribution of reduction rates with almost 1000-fold differences in
the more extreme cases. Although the rates for the work presented here appear
to be on the lower end, they are within a reasonable distance of most of the work
done by other researchers. Table 4.3 shows Fe
2+
production for a few sedimentary
environments. According to the authors, the major forms of anaerobic metabolism
are these locals are iron, sulfate and, to a lesser extent, manganese reduction (11,
129). A quick comparison between tables 4.2 and 4.3 indicates that the amount of
iron produced in the sediments is far less than what is observed under laboratory
conditions. This is not surprising, given that all of these laboratory studies were done
in batch culture, with a high density of bacteria and an excess of nutrients. However,
there are differences in the information that each table is providing. The data for
Table 4.2 is describing total reduced iron. This is both solid phase Fe
2+
as well as
aqueous phase Fe
2+
. Table 4.3 describes only aqueous phase Fe
2+
. The stoichiometric
95
Table 4.2. Fe(III) reduction rates for Shewanella strains
Shewanella
Strain
Iron oxide type
and concentration
Organic
carbon source
Fe(III)
reduction
(mM/day)
Incubation
Temperature Reference
CN32 45mM HFO 18mM lactate 2.0 30
0
C Fredrickson
1998
CN32 50mM HFO 27mM lactate 0.17 30
0
C Zachara
1998
CN32 25mM HFO 30mM lactate 1.2 30
0
C Fredrickson
2003
CN32 4mM HFO 20mM lactate 7.2x10
-5
25
0
C Glasauer
2003
CN32 50mM HFO 30mM lactate 0.5 30
0
C Kukkadapu
2001
MR-1 Smectite clay 10mM lactate 0.9 25
0
C Kostka
2002
W3-7-1 70mM akaganeite 10mM lactate 9.3x10
-2
14
0
C Roh 2003
CN32 80mM
lepidocrosite
75mM formate 2.5 25
0
C O’laughlin
2007
MR-1 80mM
lepidocrosite
75mM formate 0.5 25
0
C O’laughlin
2007
CN32 20mM HFO 20mM lactate 0.17 15
0
C This study
CN32 20mM HFO 20mM uridine 0.18 15
0
C This study
MR-4 20mM HFO 20mM lactate 0.10 15
0
C This study
MR-4 20mM HFO 20mM
pyruvate
0.36 15
0
C This study
MR-4 20mM HFO 20mM uridine 0.19 15
0
C This study
W3-18-1 20mM HFO 20mM lactate 0.21 15
0
C This study
W3-18-1 20mM HFO 20mM
pyruvate
0.27 15
0
C This study
W3-18-1 20mM HFO 20mM uridine 0.27 15
0
C This study
equations derived in chapter 3 indicate that about 5% of the Fe
2+
produced by
Shewanella remains in solution. Compiling the reduction rate data in table two gives
an average reduction rate of approximately 500 μM Fe
3+
per day. 5% of this value
would be approximately 25 μM per day; this is within the range of the data for the
upper anoxic layers in Table 4.3.
96
Table 4.3. Fe(III) Reduction rates for some sedimentary environments
Location Depth (cm)
Fe(III) Reduction
(μM/day)
Reference
FOAM Site, Long Island
Sound
0-2 37.5 Canfi eld 1989
2-4 5 “ “
4-6 1.25 “ “
4-9 1 “ “
Mississippi Delta 0-3 18 “ “
3-6 3 “ “
6-9 1.6 “ “
9-12 1.26 “ “
Danish Coast; Station 4 0-2 20 Canfi eld 1993
2-4 12.5 “ “
4-6 5 “ “
Danish Coast; Station 6 0-2 -- “ “
2-4 30 “ “
4-6 60 “ “
Another interesting relationship between cell density and reduction rates per cell is
suggested by Figure 4.1. While the reduction rates per cell decrease as cell density
increases, the relationship is a power function. Essentially, the amount of iron that is
reduced per day changes very little over the cell densities described in the fi gure. This
seems to imply that only a small fraction of the cells in culture were actually reducing
iron at any given point in time. It is worth mentioning that if these microbes were
producing an extracellular shuttle, one would expect the rates to remain unchanged,
as there would be no surface area issues to contend with. One caveat is that these data
were derived from incubations that lasted upwards of 1000 hours. The conditions
within these cultures certainly changed, perhaps affecting respiration rates.
Nonetheless, what these data imply is that a population of Shewanella at cell
densities of 10
6
will reduce on the order of 300 μM Fe
3+
per day, 15 μM of which
97
would be in solution. Cell densities in sedimentary environments have been described
as being on the order of 10
8
to 10
9
cells/ml (130, 131). This implies that a population
of Shewanella comprising less than 1% of a total microbial community might be
able to produce enough aqueous ferrous iron to yield a detectable signal. This data
suggests that there is at least the potential that a small community of bacteria could
be active enough to support subsequent biogeochemical cycling. If this is true, then
the relative abundance of a particular bacteria within a microbial community would
not give an indication of its importance to that community.
While this would go against the wishes of some, it is unlikely that one organism will
be the dominant force in any environment, polluted or otherwise human-impacted
sites notwithstanding. More likely, dominance will be defi ned by the biogeochemistry
Fe Reduction Rate ( μmol/cell*day)
Figure 4.1. log-log plot of ferric iron respiration rates for Shewanella strains using lactate as the carbon
source. R
2
= 0.94.
Cells/mL
98
that is occurring in a particular locale. High levels of nitrate will probably mean that
nitrate reducers are dominant in anoxic settings. Locations in which Mn has been
focused will probably mean that Mn-reduction is just as, or more, signifi cant than Fe-
reduction.
Shewanella’ s fl exibility in terms of what it can eat and what it can breathe makes
it a good candidate to occupy environments that are dynamic, such as oxic-anoxic
transition zones. In these settings, the type of carbon source available to an organism
might change quite rapidly, as would the available terminal electron acceptors. In
these settings it is more advantageous to have metabolic options. While they may
not be as good at specifi c metabolisms as more specialized microbes, they appear to
be good enough. Shewanella’ s near ubiquity in the environment can be seen as the
successful implementation of this more generalized approach.
99
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APPENDIX A
AEROBIC GROWTH CURVES AND O
2
RESPIRATION RATES
FOR SHEWANELLA ONEIDENSIS MR-1 USING DIFFERENT
CARBON SOURCES
Abstract
The following works represents preliminary data for a project done in conjunction
with William Berelson and Tim Riedel. The aim was to assess the growth rates and
O
2
respiration effi ciency of MR-1 using different carbon sources with the intention of
then comparing these rates to iron oxide respiration rates.
Methods
Frozen stocks of MR-1 were grown up in LB at 24
0
C for 17 hours. Cells were
transferred into defi ned medium to achieve an initial concentration of approximately
10
5
cells/ml. Cultures were grown on a shaker at 24
0
C. The defi ned media
contained one of the following carbon sources: acetate, lactate, isoleucine, N-acetyl
glucosamine or uridine. Growth rates were determined by counting colony forming
units (CFUs) over time. CFUs were determined by spot plating serial dilutions. O
2

respiration during exponential growth phase was measured via microelectrodes.
129
Results
The growth rates through exponential are shown in fi gure 1. The slopes for each
carbon source were as follows: acetate: 0.41 (r = 0.99), lactate: 0.70 (r = 0.99),
isoleucine: 0.43 (r = 0.95), N-acetyl glucosamine: 0.52 (r = 0.99), uridine: 0.60
(r = 0.98). Although some cultures experienced a longer lag phase than others,
exponential growth lasted for approximately 12-15 hours for all carbon sources
tested.
1.E+04
1.E+05
1.E+06
1.E+07
1.E+08
1.E+09
1.E+10
1.E+11
1.E+12
1.E+13
0 5 10 15 20 25 30
Time (hours)
Lactate
N-Acetyl Glucosamine
Uridine
Isoleucine
Acetate

Figure A1. Aerobic exponential growth curves for MR-1 using different carbon sources.
130
1.E+04
1.E+05
1.E+06
1.E+07
1.E+08
1.E+09
0 100 200 300 400 500 600 700
Time (hours)
1.E+04
1.E+05
1.E+06
1.E+07
1.E+08
1.E+09
1.E+10
0 100 200 300 400 500 600 700
Time (hours)
Acetate
Lactate
131
1.E+04
1.E+05
1.E+06
1.E+07
1.E+08
1.E+09
1.E+10
0 100 200 300 400 500 600 700
Time (hours)
1.E+04
1.E+05
1.E+06
1.E+07
1.E+08
1.E+09
1.E+10
0 100 200 300 400 500 600 700
Time (hours)
Isoleucine
N-Acetyl Glucosamine
132
1.E+04
1.E+05
1.E+06
1.E+07
1.E+08
1.E+09
1.E+10
0 100 200 300 400 500 600 700
Time (hours)
Figure A2. Growth curves for MR-1 using different carbon sources. With the exception of
N-acetyl glucosamine, cultures appeared to grow exponentially until they reached a cell density
of approximately 10
8
CFUs/ml. In the case of N-acetyl glucosamine, MR-1 continues to grow
exponentially until the cultures reached a density of 10
9
cells/ml. Except for acetate, cultures using all
of the other carbon substrates continued to grow after log phase to a population size of approximately
10
9
CFUs/ml.
Uridine
133
1.E-13
1.E-12
1.E-11
12 3
Time (hours)
1.E+08
1.E+09
2.E+09
3.E+09
4.E+09
5.E+09
O2 Respiration
CFUs
1.E+06
1.E+07
1.E+08
1.E+09
69 12
Time (hours)
1E-12
1E-11
1E-10
CFUs
O2 Respiration
B
A
134
1E-13
1E-12
1E-11
1E-10
1.E+06 1.E+07 1.E+08 1.E+09 1.E+10
CFUs/ml
Figure A3 Average oxygen respiration rates per cell through exponential growth. lactate (A) and
N-acetyl glucosamine (B) during exponential phase. Figure (2C) is a log-log plot of average oxygen
consumption per CFU as a function of cell density (r = 0.97). The average oxygen respiration rate
appears to be a function of cell density.
C
135
APPENDIX B
THERMODYNAMIC DATA USED FOR STABILITY DIAGRAM
CALCULATIONS
Table B1. Equations used to construct stability diagrams
logk
1
Fe(OH )
3
+ H
+
+ e
−
= Fe
3
O
4
+ 5H
2
O
1.71
− log Fe
2+
= log k + 2 pH
2
Fe
3
O
4
+ 3HCO
3
−
+ 2Fe
2+
+ 2H
2
O = 3FeCO
3
+ H
+
+ 2Fe(OH )
3
1.17
− log Fe
2+
=
1
2
pH +
3
2
log HCO
3
−
−
1
2
log k
3
6FeCO
3
+ 8H
2
O + 2Fe(OH )
3
= Fe
6
(OH )
12
CO
3
• 2H
2
O + 5HCO 3
−
+ H
+
+ 2Fe
2+
-25.01
− log Fe
2+
=
5
2
log HCO
3
−
−
1
2
log k +
1
2
pH
These equations were used to construct the stability diagrams in chapters 2 and 3. HCO
3
-
concentration
was assumed to be 0.03M, based on defi ned medium composition. Hydrous ferric oxide is represented
by Fe(OH)
3
. Thermodynamic data used to calculate logk values are listed in table B2.
136
Table B2. Thermodynamic Constants Used in Calculations
Species ΔG
o
f
(KJ/mol) Reference
Solids
Fe(OH)
3
-699 Stumm and Morgan, 1996
Fe
3
O
4
-1012.6 “ “
Fe
6
(OH)
12
CO
3
*2H
2
O
-4059.9 Drissi H, et al. 1995
FeCO
3
-666.7 Stumm and Morgan, 1996
Liquid
H
2
O
-238 “ “
Aqueous
Fe
2+
-78.87 “ “
HCO
3
-
-586.8 “ “
137
APPENDIX C
IRON REDUCTION AND CARBON CONSUMPTION DATA SETS
Appendix C contains ferric iron reduction data sets for Shewanella putrefaciens
CN32, Shewanella putrefaciens W3-18-1 and Shewanella oneidensis MR-4 using
lactate, pyruvate or uridine as the carbon source. These data were collected as
described in chapters 2 and 3.
138
Time
(hours)
pH
Solid
Fe
2+

(mM)
Aqueous
Fe
2+

(mM)
Total
Fe
2+

(mM)
Lactate
(mM)
Acetate
(mM)
Pyruvate
(mM)
Formate
(mM)
Succinate
(mM)
Total
Cells/ml
Unattached
Cells/ml
Attached
Cells/ml
0 8.5 18.421 0.000 0.000 0.000 0.000
8.5 18.223 0.000 0.000 0.000
8.7 17.941 0.000 0.000 0.000 0.000
8.1 0.089 0.000 0.089 2.20E+08
8.7 0.000 0.000 22.168 0.000 0.000 0.000 2.65E+08 5.85E+07 2.07E+08
8.7 0.135 0.014 0.149
50 0.657 0.043 0.700
0.667 0.048 0.715
0.648 0.043 0.690
0.589 0.004 0.593 9.09E+08 2.37E+07 8.85E+08
100 8.3 1.542 0.056 1.599 17.950 1.126 0.000 0.289
8.5 1.990 0.040 2.030 18.002 1.290 0.000 0.210
8.5 1.566 0.055 1.621 18.156 1.135 0.000 0.209
8.7 2.250 0.036 2.285 22.286 0.000 3.997 0.122 3.15E+08 1.55E+07 3.00E+08
Table C1. Lactate oxidation and Ferric Iron Reduction by Shewanella putrefaciens W3-18-1
139
120 1.118 0.043 1.162 1.52E+09 1.90E+07 1.50E+09
8.7 0.531 0.035 0.566
8.7 0.722 0.036 0.759
150 2.438 0.053 2.491 8.33E+08 1.35E+07 8.20E+08
200 8.5 4.245 0.151 4.396 17.288 2.059 0.000 0.287 0.463
8.5 4.614 0.199 4.813 17.070 2.156 0.000 0.964 0.486
8.5 4.426 0.120 4.545 17.136 1.938 0.000 0.512 0.529
8.5 3.852 0.057 3.909 21.950 0.000 4.317 0.456 4.35E+08 4.00E+06 4.31E+08
300 8.7 1.275 0.014 1.289
8.7 3.326 0.041 3.367
400 8.3 6.547 0.146 6.693 15.894 3.125 0.000 1.250 0.611
8.1 6.264 0.257 6.521 16.376 3.171 0.000 1.379 0.646
7.9 6.138 0.145 6.283 15.954 2.342 0.000 0.605 0.687
8.5 7.285 0.145 7.430 21.720 0.000 5.748 0.863 2.05E+08 1.05E+07 1.95E+08
Time
(hours)
pH
Solid
Fe
2+

(mM)
Aqueous
Fe
2+

(mM)
Total
Fe
2+

(mM)
Lactate
(mM)
Acetate
(mM)
Pyruvate
(mM)
Formate
(mM)
Succinate
(mM)
Total
Cells/ml
Unattached
Cells/ml
Attached
Cells/ml
Table C1 Continued
140
800 8.1 8.783 0.564 9.347 13.962 4.366 0.000 1.313 0.938
8.1 8.463 0.582 9.045 13.990 4.101 0.000 1.170 0.984
8.1 8.766 0.388 9.154 13.951 4.004 0.000 0.000 0.997
8.5 9.249 0.212 9.462 21.970 0.000 6.813 0.000
1000 8.5 9.642 0.211 9.853 1.50E+08 7.00E+06 1.43E+08
8.1 7.736 0.617 8.353 14.040 4.144 0.000 0.000 1.077
8.1 7.689 0.433 8.122 13.718 4.440 0.000 0.000 0.981
8.1 7.893 0.626 8.519 13.870 4.179 0.000 0.000 1.023
1250 8.5 9.116 0.087 9.203
8.5 7.089 0.146 7.234
8.5 7.466 0.136 7.602
2100 8.5 11.805 0.296 12.102
Time
(hours)
pH
Solid
Fe
2+

(mM)
Aqueous
Fe
2+

(mM)
Total
Fe
2+

(mM)
Lactate
(mM)
Acetate
(mM)
Pyruvate
(mM)
Formate
(mM)
Succinate
(mM)
Total
Cells/ml
Unattached
Cells/ml
Attached
Cells/ml
Table C1 Continued
141
Time
(hours)
pH
Solid Fe
2+

(mM)
Aqueous
Fe
2+
(mM)
Solid Fe
2+

(mM)
Pyruvate
(mM)
Acetate
(mM)
Formate
(mM)
Total Cells/
ml
Unattached
Cells/ml
Attached
Cells/ml
0
8.3 0.227 3.15E+08
8.3 0.181 0.000 0.181 18.145 0.000 0.000 1.25E+08 3.15E+07 9.35E+07
8.7 2.90E+08
8.7 0.237 0.000 0.237 17.990 0.055 0.000 1.85E+08
8.7 0.238 0.000 0.238 19.207 0.055 0.000 6.50E+07
8.7 0.256 0.000 0.256 18.058 0.055 0.000 2.10E+08
8.7 18.506 0.000 0.000
8.7 17.700 0.000 0.000
9 18.363 0.000 0.000
20 3.145 3.00E+08 2.00E+07 2.80E+08
Table C2. Pyruvate oxidation and Ferric Iron Reduction by Shewanella putrefaciens W3-18-1
142
50
5.148 1.338 6.487 2.65E+08 2.40E+07 2.41E+08
0.786 0.056 0.842
0.945 0.068 1.013
0.873 0.065 0.939
8.3 2.829 0.058 2.888 13.292 0.055 0.000
8.3 2.550 0.051 2.600 13.326 0.055 0.257
8.3 2.066 0.062 2.128 14.298 0.055 0.000
8.3 2.312 0.106 2.419
6.272 2.037 8.309 3.53E+08 2.25E+07 3.31E+08
100
7.4 2.752 0.718 3.470 3.221 13.562 0.000
7.4 3.027 0.763 3.790 2.995 13.802 0.000
7.4 3.043 0.789 3.832 3.043 13.988 0.000
6.303 2.540 8.843 4.04E+08 3.65E+07 3.68E+08
7.4 5.125 1.959 7.084 0.000 12.190 3.398 4.40E+08 2.30E+07 4.17E+08
Table C2 Continued
Time
(hours)
pH
Solid Fe
2+

(mM)
Aqueous
Fe
2+
(mM)
Solid Fe
2+

(mM)
Pyruvate
(mM)
Acetate
(mM)
Formate
(mM)
Total Cells/
ml
Unattached
Cells/ml
Attached
Cells/ml
143
150
7.1 5.321 1.935 7.257
6.727 2.886 9.613 1.01E+09 2.50E+07 9.85E+08
200
7.1 4.371 2.045 6.416 0.428 11.084 6.529
7.1 4.991 2.077 7.068 0.569 11.254 6.312
7.4 5.148 2.030 7.178 0.536 11.545
7.4 5.800 2.799 8.600 0.000 14.205 3.426 4.50E+08 1.90E+07 4.31E+08
7.4 6.107 2.462 8.568
400
7.4 7.725 1.849 9.574
7.4 8.079 0.623 8.702 0.000 12.140 1.710
7.4 8.652 0.647 9.299 0.000 12.403 1.636
7.1 9.595 0.608 10.203 0.000 12.741 1.961
7.4 9.658 0.663 10.320 0.416 15.658 2.953 5.80E+08 2.50E+07 5.55E+08
800
7.9 7.608 0.461 8.068
7.9 9.932 0.476 10.408
7.9 10.010 0.413 10.423
8.3 11.033 0.183 11.216 0.077 18.378 0.963 3.60E+08 2.25E+07 3.38E+08
Time
(hours)
pH
Solid Fe
2+

(mM)
Aqueous
Fe
2+
(mM)
Solid Fe
2+

(mM)
Pyruvate
(mM)
Acetate
(mM)
Formate
(mM)
Total Cells/
ml
Unattached
Cells/ml
Attached
Cells/ml
Table C2 Continued
144
1000
7.9 9.323 0.386 9.709
7.9 10.501 0.338 10.839
7.9 12.339 0.322 12.662
8.3 11.601 0.254 11.855 0.000 11.516 0.000 7.00E+07 5.50E+06 6.45E+07
1100
7.9 10.600 0.136 10.737
8.3 11.915 0.184 12.099 0.061 0.406
2100
8.7 11.852 0.144 11.996
Table C2 Continued
Time
(hours)
pH
Solid Fe
2+

(mM)
Aqueous
Fe
2+
(mM)
Solid Fe
2+

(mM)
Pyruvate
(mM)
Acetate
(mM)
Formate
(mM)
Total Cells/
ml
Unattached
Cells/ml
Attached
Cells/ml
145
Time
(hours)
pH
Solid Fe
2+

(mM)
Aqueous Fe
2+

(mM)
Total Fe
2+

(mM)
Uridine
(mM)
Acetate
(mM)
Total Cells/ml
Unattached
Cells/ml
Attached
Cells/ml
0 8 0.089 0.089 2.20E+08
8.5 0.084 0.000 0.084 2.30E+08 2.05E+07 2.10E+08
8.5 5.25E+08
8.7 22.752 0.000
8.7 22.400 0.000
8.7 23.804 0.000
50 1.011 0.044 1.056
1.027 0.052 1.079
0.652 0.029 0.682
8.5 2.014 0.029 2.043
0.589 0.004 0.593 9.09E+08 2.37E+07 8.85E+08
100 8.1 3.342 0.201 3.543 18.969 2.181
8.1 3.216 0.186 3.402 19.864 2.170
8.3 2.894 0.205 3.099 20.265 3.527
8.3 0.927 0.004 0.931 3.05E+08 1.50E+07 2.90E+08
120 1.118 0.043 1.162 1.52E+09 1.90E+07 1.50E+09
Table C3. Uridine oxidation and Ferric Iron Reduction by Shewanella putrefaciens W3-18-1
146
150 8.1 5.101 0.339 5.440
2.438 0.053 2.491 8.33E+08 1.35E+07 8.20E+08
200 7.7 4.308 0.755 5.063 18.748 4.112
7.9 5.038 0.759 5.798 18.685 4.167
7.9 4.583 0.505 5.088 18.208 5.200
350 7.4 8.817 1.017 9.834
400 7.4 4.818 1.770 6.589 16.768 5.747
7.4 5.580 1.495 7.076 16.708 5.837
7.4 4.504 1.095 5.599 17.025 6.270
8.1 5.266 0.218 5.484 3.90E+08 1.20E+07 3.78E+08
800 7.4 6.190 1.794 7.984
7.4 8.420 1.762 10.182
7.4 6.234 1.558 7.792
1000 7.4 9.087 1.495 10.582 15.515 7.212
7.4 12.151 1.205 13.356 15.384 7.283
7.4 8.191 1.378 9.569 14.697 9.719
7.9 9.147 0.846 9.993
Table C3 Continued
Time
(hours)
pH
Solid Fe
2+

(mM)
Aqueous Fe
2+

(mM)
Total Fe
2+

(mM)
Uridine
(mM)
Acetate
(mM)
Total Cells/ml
Unattached
Cells/ml
Attached
Cells/ml
147
1100 7.4 8.691 0.170 8.861
2000 7.9 10.108 7.434 17.543
Table C3 Continued
Time
(hours)
pH
Solid Fe
2+

(mM)
Aqueous Fe
2+

(mM)
Total Fe
2+

(mM)
Uridine
(mM)
Acetate
(mM)
Total Cells/ml
Unattached
Cells/ml
Attached
Cells/ml
148
Time
(hours)
pH
Solid Fe
2+

(mM)
Aqueous Fe
2+

(mM)
Total Fe
2+

(mM)
Lactate (mM) Acetate (mM)
Pyruvate
(mM)
Succinate
(mM)
0 8.7 17.229
8.7 16.514
8.7 17.049
8.7 0.106 0.004 0.111
0.130 0.130
70 0.919 0.006 0.925
100 8.7 0.828 0.102 0.930 17.475 0.660 4.439
8.5 0.820 0.054 0.874 16.773 1.272
8.5 1.566 0.065 1.632 16.411 1.312
120 1.982 0.054 2.037
1.192 0.050 1.242
1.112 0.036 1.148
150 3.067 0.091 3.158
Table C4. Lactate oxidation and Ferric Iron Reduction by Shewanella putrefaciens CN32
149
200 8.5 2.312 0.051 2.364 17.574 0.833 0.450
8.5 1.786 0.042 1.828 16.950 0.688 0.426
8.3 3.719 0.090 3.808 16.278 1.601 0.000
4.127 0.108 4.235
3.192 0.066 3.259
400 8.7 5.243 0.255 5.497 17.149 1.001 1.081
8.7 4.905 0.225 5.130 16.831 0.839 1.069
8.3 5.117 0.344 5.461 15.871
800 8.3 5.274 0.095 5.369
8.5 7.112 0.155 7.267
8.3 9.053 0.109 9.162
1000 8.5 6.445 0.132 6.577
8.3 6.146 0.210 6.356
8.5 7.599 0.139 7.738
Time
(hours)
pH
Solid Fe
2+

(mM)
Aqueous Fe
2+

(mM)
Total Fe
2+

(mM)
Lactate (mM) Acetate (mM)
Pyruvate
(mM)
Succinate
(mM)
Table C4 Continued
150
1200 7.9 8.801 0.426 9.228
8.479 0.205 8.685
8.354 0.204 8.557
Time
(hours)
pH
Solid Fe
2+

(mM)
Aqueous Fe
2+

(mM)
Total Fe
2+

(mM)
Lactate (mM) Acetate (mM)
Pyruvate
(mM)
Succinate
(mM)
Table C4 Continued
151
Time
(hours)
pH
Solid Fe
2+

(mM)
Aqueous Fe
2+
(mM)
Total Fe
2+

(mM)
Pyruvate (mM) Acetate (mM) Formate (mM)
0 8.7 19.480 0.000 0.000
8.7 18.801 0.000 0.000
8.7 18.257 0.000 0.000
100 8.5 1.542 0.076 1.619 16.273 2.102 0.000
8.5 1.645 0.087 1.732 15.804 2.361 0.000
8.3 1.817 0.076 1.893 14.705 1.702 0.000
200 8.5 1.770 0.047 1.817 14.025 3.024 0.000
8.3 3.389 0.098 3.486 13.284 2.710 0.000
8.3 2.147 0.083 2.230 12.553 2.964 0.000
400 8.3 3.923 1.967 5.890 12.107 2.741 0.000
8.3 4.653 2.540 7.194 11.682 2.559 0.000
8.3 7.733 1.872 9.605 11.380 2.968 0.000
800 8.3 1.755 0.081 1.836
8.3 1.739 0.080 1.819
8.3 2.061 0.083 2.144
Table C5. Pyruvate oxidation and Ferric Iron Reduction by Shewanella putrefaciens CN32
152
1000 8.3 1.817 0.115 1.932 0.000 0.000 0.000
8.3 1.802 0.133 1.935 0.000 0.000 0.000
7.9 2.108 0.137 2.245 0.000 0.000 0.000
Table C5 Continued
Time
(hours)
pH
Solid Fe
2+

(mM)
Aqueous Fe
2+
(mM)
Total Fe
2+

(mM)
Pyruvate (mM) Acetate (mM) Formate (mM)
153
Time (hours) pH Solid Fe
2+
(mM) Aqueous Fe
2+
(mM) Total Fe
2+
(mM) Uridine (mM)
0 8.5 22.767
8.5 24.328
8.5 23.555
100 8.5 2.352 0.100 2.452 30.497
8.5 2.894 0.104 2.998 24.440
8.3 2.619 0.102 2.721 34.387
200 7.9 4.009 0.292 4.301 26.594
7.9 4.418 0.274 4.692 29.367
7.9 4.245 0.278 4.523 28.230
400 7.7 3.844 0.867 4.711
7.9 5.337 1.857 7.194
7.7 4.567 1.268 5.835
800 7.7 6.963 0.113 7.076
7.7 6.987 0.050 7.037
7.7 4.567 1.268 5.835
Table C6. Uridine oxidation and Ferric Iron Reduction by Shewanella putrefaciens CN32
154
1000 7.7 7.301 0.607 7.908
7.9 7.395 0.375 7.770
7.9 5.855 0.416 6.271
Table C6 Continued
Time (hours) pH Solid Fe
2+
(mM) Aqueous Fe
2+
(mM) Total Fe
2+
(mM) Uridine (mM)
155
Time
(hours)
pH
Solid
Fe
2+

(mM)
Aqueous
Fe
2+

(mM)
Total
Fe
2+

(mM)
Lactate
(mM)
Acetate
(mM)
Pyruvate
(mM)
Formate
(mM)
Succinate
(mM)
Total
Cells/ml
Unattached
Cells/ml
Attached
Cells/ml
0 8.5 16.934 0.000 0.000 0.000
8.5 17.305 0.000 0.000 0.000
8.7 16.404 0.000 0.000 0.000
8.1 0.123 0.123 1.85E+08
8.7 21.925 0.000 0.000 0.000 2.45E+08 1.70E+07 2.28E+08
8.7 0.117 0.023 0.140
50 0.824 0.034 0.858
0.809 0.036 0.845
0.828 0.031 0.859
0.852 0.010 0.862 9.34E+08 6.37E+07 8.70E+08
100 8.5 1.165 0.025 1.190 15.814 1.297 0.469 0.000
8.5 1.323 0.041 1.364 16.710 1.313 0.399 0.000
8.5 1.346 0.051 1.397 16.148 1.256 0.517 0.000
8.7 0.256 0.008 0.264 22.749 2.628 0.110 0.000 3.55E+08 2.15E+07 3.34E+08
Table C7. Lactate oxidation and Ferric Iron Reduction by Shewanella oneidensiss MR-4
156
120 1.110 0.038 1.148 1.36E+09 3.25E+07 1.33E+09
8.7 0.599 0.032 0.631
8.7 0.519 0.029 0.547
200 1.425 0.058 1.483 1.04E+09 8.40E+07 9.56E+08
8.5 0.511 0.000 22.839 2.816 0.000 0.122 4.10E+08 8.00E+06 4.02E+08
300 8.7 0.758 0.020 0.778
8.7 0.703 0.025 0.727
400 8.5 1.967 0.054 2.021 23.494 3.146 0.000 0.258 3.25E+08 5.50E+06 3.20E+08
8.3 2.163 0.047 2.211 16.384 1.519 0.468
8.3 2.415 0.061 2.475 16.799 1.548 0.417
8.3 2.242 0.069 2.310 16.604 1.567 0.547
800 8.3 3.671 0.094 3.765 15.003 1.955 0.282 0.000 0.000
8.1 3.546 0.086 3.632 15.426 2.178 0.151 0.000 0.445
8.3 3.530 0.087 3.617 15.711 2.038 0.334 0.000 0.457
8.5 5.337 0.123 5.460 23.003 3.452 0.085 0.000 0.000
Time
(hours)
pH
Solid
Fe
2+

(mM)
Aqueous
Fe
2+

(mM)
Total
Fe
2+

(mM)
Lactate
(mM)
Acetate
(mM)
Pyruvate
(mM)
Formate
(mM)
Succinate
(mM)
Total
Cells/ml
Unattached
Cells/ml
Attached
Cells/ml
Table C7 Continued
157
1000 8.7 6.350 0.100 6.450 2.70E+08 6.00E+06 2.64E+08
8.3 3.428 0.071 3.499 15.390 2.172 0.202 0.500
8.1 3.742 0.087 3.830 15.649 2.187 0.260 0.455
8.3 3.530 0.083 3.613 15.088 2.097 0.260 0.000
1300 8.7 6.759 0.087 6.846
1224 7.4 5.628 1.700 7.327
2100 8.5 8.676 0.185 8.861
Table C7 Continued
Time
(hours)
pH
Solid
Fe
2+

(mM)
Aqueous
Fe
2+

(mM)
Total
Fe
2+

(mM)
Lactate
(mM)
Acetate
(mM)
Pyruvate
(mM)
Formate
(mM)
Succinate
(mM)
Total
Cells/ml
Unattached
Cells/ml
Attached
Cells/ml
158
Table C8. Pyruvate oxidation and Ferric Iron Reduction by Shewanella oneidensiss MR-4
Time
(hours)
pH
Solid
Fe
2+

(mM)
Aqueous
Fe
2+

(mM)
Total
Fe
2+

(mM)
Lactate
(mM)
Acetate
(mM)
Pyruvate
(mM)
Formate
(mM)
Succinate
(mM)
Total
Cells/ml
Unattached
Cells/ml
Attached
Cells/ml
0 8.5 16.934 0.000 0.000
8.5 17.305
8.7 16.404
8.1 0.123 0.123 1.85E+08
8.7 21.925 0.000 0.000 0.000 2.45E+08 1.70E+07 2.28E+08
8.7 0.117 0.023 0.140
50 0.824 0.034 0.858
0.809 0.036 0.845
0.828 0.031 0.859
0.852 0.010 0.862 9.34E+08 6.37E+07 8.70E+08
100 8.5 1.165 0.025 1.190 15.814 1.297 0.469
8.5 1.323 0.041 1.364 16.710 1.313 0.399
8.5 1.346 0.051 1.397 16.148 1.256 0.517
8.7 0.256 0.008 0.264 22.749 2.628 0.110 0.000 3.55E+08 2.15E+07 3.34E+08
159
Table C8 Continued
120 1.110 0.038 1.148 1.36E+09 3.25E+07 1.33E+09
8.7 0.599 0.032 0.631
8.7 0.519 0.029 0.547
200 1.425 0.058 1.483 1.04E+09 8.40E+07 9.56E+08
8.5 0.511 0.000 22.839 2.816 0.000 0.122 4.10E+08 8.00E+06 4.02E+08
300 8.7 0.758 0.020 0.778
8.7 0.703 0.025 0.727
400 8.5 1.967 0.054 2.021 23.494 3.146 0.000 0.258 3.25E+08 5.50E+06 3.20E+08
8.3 2.163 0.047 2.211 16.384 1.519 0.468
8.3 2.415 0.061 2.475 16.799 1.548 0.417
8.3 2.242 0.069 2.310 16.604 1.567 0.547
800 8.3 3.671 0.094 3.765 15.003 1.955 0.282
8.1 3.546 0.086 3.632 15.426 2.178 0.151 0.445
8.3 3.530 0.087 3.617 15.711 2.038 0.334 0.457
8.5 5.337 0.123 5.460 23.003 3.452 0.085 0.000
Time
(hours)
pH
Solid
Fe
2+

(mM)
Aqueous
Fe
2+

(mM)
Total
Fe
2+

(mM)
Lactate
(mM)
Acetate
(mM)
Pyruvate
(mM)
Formate
(mM)
Succinate
(mM)
Total
Cells/ml
Unattached
Cells/ml
Attached
Cells/ml
160
1000 8.7 6.350 0.100 6.450 2.70E+08 6.00E+06 2.64E+08
8.3 3.428 0.071 3.499 15.390 2.172 0.202 0.500
8.1 3.742 0.087 3.830 15.649 2.187 0.260 0.455
8.3 3.530 0.083 3.613 15.088 2.097 0.260 0.000
1300 8.7 6.759 0.087 6.846
2100 8.5 8.676 0.185 8.861
1200 7.4 5.628 1.700 7.327
Time
(hours)
pH
Solid
Fe
2+

(mM)
Aqueous
Fe
2+

(mM)
Total
Fe
2+

(mM)
Lactate
(mM)
Acetate
(mM)
Pyruvate
(mM)
Formate
(mM)
Succinate
(mM)
Total
Cells/ml
Unattached
Cells/ml
Attached
Cells/ml
Table C8 Continued
161
APPENDIX D
SUSCEPTIBILITY V ALUES FOR THE PRODUCTS OF FERRIC
IRON REDUCTION COUPLED TO LACTATE OXIDATION
Appendix D contains susceptibility data for Shewanella putrefaciens W3-18-1,
Shewanella putrefaciens CN32 and Shewanella oneidensis MR-4 using lactate as the
carbon source. These data were collected as described in chapter 2.
162
Blanks
Cube -2.185E-06
Tube -1.050E-07
100
hours
1 2 3 Average w/o blanks χ
Calculated
Fe3O4
(mg)
Sample
Mass (mg)
%
1 -6.91E-07 -6.66E-07 -6.88E-07 -6.82E-07 1.61E-06 4.86E-05 0.1304 3.31 3.938
2 -2.22E-07 -1.98E-07 -2.03E-07 -2.08E-07 2.08E-06 6.23E-05 0.1401 3.34 4.193
3 -8.56E-07 -7.55E-07 -7.49E-07 -7.87E-07 1.50E-06 4.45E-05 0.1283 3.38E 3.798
400
hours
1 2.09E-06 2.08E-06 2.10E-06 2.09E-06 4.38E-06 1.56E-04 0.1870 2.814 6.644
2 5.99E-07 5.60E-07 5.11E-07 5.57E-07 2.85E-06 1.04E-04 0.1556 2.737 5.687
3 -1.27E-06 -1.27E-06 -1.29E-06 -1.28E-06 1.01E-06 3.64E-05 0.1183 2.781 4.253
Table D1. Shewanella putrefaciens W3-18-1 Susceptibility
163
800
hours
1 -1.83E-06 1.83E-06 -1.90E-06 -6.34E-07 1.66E-06 1.77E-04 0.131 0.937 14.020
2 -1.41E-06 -1.43E-06 -1.43E-06 -1.42E-06 8.66E-07 4.42E-05 0.115 1.958 5.887
3 -1.84E-06 -1.86E-06 -1.95E-06 -1.89E-06 4.05E-07 4.61E-05 0.1059 0.877 12.0696
1 2 3 Average w/o blanks χ
Calculated
Fe3O4
(mg)
Sample
Mass (mg)
%
Table D1 Continued
164
Blanks
Cube -2.185E-06
Tube -1.050E-07
100 hours 1 2 3 Average w/o blanks χ
Calculated
Fe3O4
(mg)
Sample
Mass(mg)
%
1 -4.28E-07 -3.98E-07 -3.96E-07 -4.07E-07 1.88E-06 1.12E-04 0.136 1.687 8.060
2 -2.65E-07 -3.03E-07 -3.43E-07 -3.04E-07 1.99E-06 6.51E-05 0.138 3.05 4.528
3 7.81E-06 7.81E-06 7.83E-06 7.81E-06 1.01E-05 4.16E-04 0.304 2.43 12.493
Control -2.01E-06 -2.04E-06 -2.08E-06 -2.04E-06 2.49E-07 8.89E-06 2.798
200 hours
1 1.15E-05 1.15E-05 1.15E-05 1.15E-05 1.38E-05 5.96E-04 0.378 2.307 16.386
2 7.20E-05 7.27E-05 7.39E-05 7.29E-05 7.52E-05 5.02E-03 1.630 1.498 108.842
3 6.85E-07 6.48E-07 7.12E-07 6.82E-07 2.97E-06 1.09E-04 0.158 2.723 5.810
Control -1.86E-06 1.90E-06 -1.90E-06 -6.20E-07 1.67E-06 4.93E-05 3.389
Table D2. Shewanella putrefaciens CN32 Susceptibility
165
800 hours
1 5.02E-05 5.03E-05 5.03E-05 5.03E-05 5.26E-05 3.45E-03 1.1693 1.525 76.675
2 1.19E-04 1.17E-04 1.18E-04 1.18E-04 1.20E-04 6.90E-03 2.549 1.742 146.323
3 1.27E-04 1.27E-04 1.25E-04 1.26E-04 1.28E-04 7.01E-03 2.714 1.831 148.2301
Control -1.88E-06 -2.00E-06 -1.93E-06 -1.94E-06 3.55E-07 1.16E-05 3.059
400 hours
1 5.68E-05 5.70E-05 5.74E-05 5.70E-05 5.93E-05 3.86E-03 1.3071 1.538 84.984
2 8.58E-05 8.57E-05 8.57E-05 8.57E-05 8.80E-05 5.11E-03 1.893 1.723 109.835
3 1.05E-04 1.05E-04 1.05E-04 1.05E-04 1.07E-04 6.48E-03 2.279 1.65 138.128
Control -2.32E-06 -1.95E-06 -1.99E-06 -2.09E-06 2.02E-07 6.41E-06 3.153
Table D2 Continued
1 2 3 Average w/o blanks χ
Calculated
Fe3O4
(mg)
Sample
Mass(mg)
%
166
Blanks
Cube -2.185E-06
Tube -1.050E-07
100 hours 1 2 3 Average w/o blanks χ
Calculated
Fe3O4
(mg)
Sample
Mass(mg)
%
1 -1.03E-06 -9.51E-07 -9.75E-07 -9.85E-07 1.30E-06 4.20E-05 0.124 3.104 4.001
2 -1.08E-06 -9.82E-07 -1.01E-06 -1.02E-06 1.27E-06 4.47E-05 0.123 2.833 4.357
3 -8.88E-07 -8.94E-07 -8.91E-07 -8.91E-07 1.40E-06 4.47E-05 0.126 3.132 4.027
Control -1.97E-06 -1.95E-06 -1.99E-06 -1.97E-06 3.20E-07 1.11E-05 0.104 2.882 3.613
400 hours
1 2.03E-06 2.06E-06 1.97E-06 2.02E-06 4.31E-06 1.59E-04 0.185 2.706 6.853
2 4.28E-06 4.27E-06 4.35E-06 4.30E-06 6.59E-06 2.65E-04 0.232 2.482 9.345
3 3.14E-06 3.11E-06 3.15E-06 3.13E-06 5.42E-06 2.24E-04 0.208 2.419 8.607
Control -2.07E-06 -2.08E-06 -2.11E-06 -2.09E-06 2.03E-07 9.15E-06 0.102 2.215 4.593
Table D3. Shewanella oneidensis MR-4 Susceptibility
167
800 hours
1 3.87E-05 3.87E-05 3.87E-05 3.87E-05 4.10E-05 1.89E-03 0.933 2.171 42.993
2 2.87E-05 2.86E-05 2.86E-05 2.86E-05 3.09E-05 1.39E-03 0.728 2.225 32.731
3 2.81E-05 2.81E-05 2.80E-05 2.81E-05 3.04E-05 1.26E-03 0.717 2.4 29.857
Control -1.99E-06 -1.99E-06 -1.99E-06 -1.99E-06 3.02E-07 1.13E-05 0.104 2.666 3.892
Table D3 Continued
1 2 3 Average w/o blanks χ
Calculated
Fe3O4
(mg)
Sample
Mass(mg)
%

Abstract (if available)

Abstract The aim of my work is to elucidate the interactions between microbes and minerals. The work I present here describes studies I have done investigating factors that can influence the identity of reduced iron oxide minerals.

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University of Southern California Dissertations and Theses

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Asset Metadata

Creator Salas, Everett Cossio (author)

Core Title Studies on the influence of bacteria and carbon source on the products of dissimilatory iron reduction

Contributor Electronically uploaded by the author (provenance)

School College of Letters, Arts and Sciences

Degree Doctor of Philosophy

Degree Program Geological Sciences

Publication Date 11/14/2008

Defense Date 08/01/2008

Publisher University of Southern California (original), University of Southern California. Libraries (digital)

Tag biominerals,dissimilatory iron reduction,fougerite,geomicrobiology,green rust,iron minerals,magnetite,microbe-mineral interactions,OAI-PMH Harvest,Shewanella,siderite

Language English

Advisor Nealson, Kenneth H. (committee chair), Berelson, William M. (committee member), Corsetti, Frank A. (committee member), Finkel, Steven E. (committee member), Hammond, Douglas E. (committee member), Kampf, Anthony R. (committee member)

Creator Email ecssce@gmail.com,everetts@usc.edu

Permanent Link (DOI) https://doi.org/10.25549/usctheses-m1770

Unique identifier UC165023

Identifier etd-Salas-2452 (filename),usctheses-m40 (legacy collection record id),usctheses-c127-136839 (legacy record id),usctheses-m1770 (legacy record id)

Legacy Identifier etd-Salas-2452.pdf

Dmrecord 136839

Document Type Dissertation

Rights Salas, Everett Cossio

Type texts

Source University of Southern California (contributing entity), University of Southern California Dissertations and Theses (collection)

Repository Name Libraries, University of Southern California

Repository Location Los Angeles, California

Repository Email cisadmin@lib.usc.edu