Interactions of Fe(II) with the iron oxidizing bacterium
Rhodopseudomonas palustris TIE-1
by
Lina J. Bird
Submitted to the Department of Biology in Partial Fulfillment of the Requirements for
the Degree of
DOCTOR OF PHILOSOPHY IN BIOLOGY AT THE MASSACHUSETTS INSTITUTE OF
ARtm
INSTTE
SSAONUSETTSNOLOGY
TECHNOLOGY
F TECHU
JUNE 2013
BRA RES
@ Lina J. Bird. All rights reserved
The author hereby grants to MIT permission to reproduce and to distribute publicly
paper and electronic copies of this thesis document in whole or in part in any medium
now known or hereafter created.
.-
...................
Signature of Author...
-
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Department of Biology
[anticipated May 1st, 2013]
.-
Certified by..-
-.
- - -- - -- -Dianne Newman
Professor of Biology
Thesis Supervisor
Accepted by ......... .
..
.--------.-.--
.---
------......-------------
[Biology graduate committee chair]
1
Interactions of Fe(II) with the iron oxidizing bacterium
Rhodopseudomonaspalustris TIE-1
by
Lina J. Bird
Submitted to the Department of Biology on May 24, 2013
in Partial Fulfillment of the Requirements
for the Degree of Doctor or Philosophy in Biology
Abstract
Microbial anaerobic iron oxidation has long been of interest to biologists and
geologists, both as a possible mechanism for the creation of banded iron formations
before the rise of oxygen, and as a model system for organisms able to accept electrons
from an external, inorganic source. Previous work with the purple photoferrotroph
Rhodopseudomonas palustris TIE-1 showed that three genes were required for
phototrophic growth with Fe(Il): PioA, a decaheme cytochrome, PioB, an outer
membrane porin, and PioC, a high potential iron protein (HiPIP). These proteins
suggested a model of Fe(II) oxidation that ends with transfer of electrons to the
photosynthetic reaction center. The goal of this thesis was to test and extend this
model through characterization of the electron transfer proteins PioA and PioC.
In the course of our experiments, we discovered that Fe(II) could also delay
growth under certain conditions. We then broadened our focus to encompass several
facets of the interaction of TIE-1 with Fe(II) under anaerobic conditions:
The first portion describes how low amounts of Fe(II) cause a growth delay in
TIE-1 cultures growing anaerobically on other substrates - a surprising result for an
organism that grows on millimolar concentrations of iron. The cause of this toxicity was
found to be dependent on copper, which istoxic to TIE-i at fairly low concentrations.
Our results indicate the copper toxicity is synergistically increased by Fe(II) under strictly
anaerobic conditions.
2
The second part of this work describes characterization of the HiPIP PioC and a
second HiPIP in the TIE-1 genome. The results showed that PioC is capable of reducing
the reaction center, as expected, though at a slower rate than is usually found for this
kind of interaction. The second HiPIP cannot reduce the reaction center and likely serves
an alternate function in the cell unrelated to photosynthesis, possibly involving
detoxification of metals.
The final section redefines our understanding of the Fe(II) oxidation pathway by
putting it in the context of reverse electron transfer, a process that is not well
understood in photosynthetic bacteria. Evidence from whole cell experiments using
flash induced spectrometry indicated that electrons from Fe(II) may, rather than going
to the reaction center, enter the quinone pool through the bc1 complex. This model is
significantly different from previous preferred models of phototrophic oxidation, but is
similar to the reverse electron transfer system described in acidophilic lithotrophic iron
oxidizing bacteria.
Taken together, the experiments described in this thesis highlight the complex
and interconnected nature of a bacterial cell's interactions with iron under anoxic
conditions. It also suggests future avenues of study for phototrophic reverse electron
transfer, a poorly understood process that is vital to anoxygenic photoautotrophic
growth.
Thesis Supervisor: Dianne K. Newman
Title: Professor of Biology and Geobiology, California Institute of Technology and
Investigator, Howard Hughes Medical Institute
3
Acknowledgements
So many people have helped me on this path that it would be impossible to
thank them all.
First and foremost, I would like to thank my advisor, Dianne Newman. I still
remember the first time I heard her present. I was completing my masters and
beginning to think about graduate school, when I heard her give a talk on arsenate
reduction. Her enthusiasm for science was contagious, and I immediately knew that I
wanted to work with her. That decision proved the right one time and again - her
teaching, support, and mentorship has been unwavering.
I thank my committee (Sean Elliot, Bob Sauer, and Graham Walker) for their
advice and support through the many twists and turns my project took. I learned a great
deal from Graham and Bob's teaching at MIT; I still refer to my class notes. Sean was a
lifesaver as my project steered to the more specialized areas of metals biology, and was
a pleasure to have in the lab during his sabbatical. I feel privileged to have had such an
excellent committee.
I thank my collaborators, Ricardo Louro, Ivo Saraiva, and Wolfgang Nitschke;
their expertise in their respective fields made this work possible.
I thank every Newman Lab member I have worked with. Their friendship,
support, scientific advice, and discussions, enriched me in more ways than I can count.
Special thanks to my year mate Suzanne Kern, who has been a wonderful friend and has
kept me from missing more than a few deadlines!
I thank my classmates, and the rest of the MIT community. Special thanks to
Frank Solomon for his support and encouragement.
I also thank my Duquesne University mentors John Stolz, Partha Basu, and Nancy
Trun - their training, guidance and encouragement prepared me for graduate school
and convinced me to apply.
Finally, I thank my family for - everything.
4
Table of Contents
Abstract .......................................................................................................................
2
Acknowledgements..................................................................................................
4
Table of Contents...................................................................................................
5
List of Figures...............................................................................................................6
List of Tables ................................................................................................................
7
Chapter 1: Introduction...........................................................................................
8
Chapter 2: Bioenergetic challenges of microbial iron metabolisms .........................
13
Chapter 3: Iron and copper act synergistically to delay anaerobic growth in bacteria.45
Chapter 4: Non-redundant roles for cytochrome c2 and two HiPIPs in the
photoferrotroph Rhodopseudomonas palustris TIE-1 ............................................
83
Chapter 5: Visualizing photosynthesis in whole cells ................................................
117
Chapter 6: A larger perspective ................................................................................
145
Appendix A: Attempted purification of PioA ............................................................
150
5
List of Figures
19
Figure 2.1. Electron transport in At. ferrooxidans. ......................................................
23
Figure 2.2. Electron transfer in phototrophic iron oxidizers. ......................................
Figure 2.3. Model of Fe(III) reduction in Geobactersulfurreducens.............................28
29
Figure 2.4. Electron transfer to Fe(Ill) in Shewanella ...................................................
Figure 3.1. Effect of Fe(II) and copper on growing R. palustris cells............................55
Figure 3.2. Anaerobic growth curves of R. palustris with Cu(II) in the presence of Fe(II),
58
Co (II), o r Ni(ll) ...........................................................................................................
60
Figure 3.3. Effect of Cu(II) and Fe(II) on other bacteria ..............................................
62
Figure 3.4. Abiotic reduction of Cu(II) by Fe(II) and ascorbate .....................................
Figure 3.5. Effect of Cu(l) vs. Cu(II) on growth of R. palustris.......................................63
Figure 3.6. Q-RT-PCR showing fold change after 15 minute shock (T15/TO) by Cu(II) and
65
Cu(II) + Fe (II) .............................................................................................................
Figure 3.7. Growth curves of the parent and the mutant strain LEM59 in the presence of
67
m eta ls .......................................................................................................................
Figure 4.1. WT TIE-1 and Acyc2 growing photoheterotrophically with acetate and
94
photoautotrophically w ith hydrogen.. ................................................................
95
Figure 4.2. Fe(II) oxidation rates of TIE-1 and mutant strains .....................................
Figure 4.3. Rpal_4085 and pioC transcription and localization in strains TIE-1 and
pioC-+ Rpal 4085 ................................................................................................
. . 96
Figure 4.4. X-Band (9.66 GHz) EPR spectra of oxidized purified PioC and Rpal_4085......98
Figure 4.5. X-Band (9.39 GHz) EPR spectra of PioC incubated with a suspension of TIE-1
membranes in the dark (top) or under light (bottom) .........................................
99
Figure 4.6. Cyclic voltammograms of PioC and Rpal_4085.............................................100
Figure 4.7. re-reduction of the reaction center in membrane fragments......................103
Figure 4.8. Transcriptional reponse of Rpal_4085 to metals.........................................104
Figure 4.9. electron flow chart in R. palustris TIE-1........................................................108
Figure 5.1. Difference spectra of oxidized vs reduced photosynthetic components ..... 122
Figure 5.2. Model of the electron transfer system components and their light induced
abso rb ance sh ifts....................................................................................................124
Figure 5.3. Absorbance changes in acetate grown cultures (red) vs. cultures growing
129
rapidly (green) or slow ly (blue) on hydrogen .........................................................
Figure 5.4. The effect of acetate and Fe(II) on oxidized cultures ................................... 131
Figure 5.5. Effect of antim ycin on cells...........................................................................134
Figure 5.6. Response of A) ApioA, and B) wild type TIE-1 to the addition of Fe(II) ........ 136
139
Figure 5.7. A new possible m odel for Fe(II) oxidation. ...................................................
Figure A.1. PioA expression constructs made in this study ............................................ 157
159
Figure A.2. Spectra of E. coli BL21 cells expressing PioA ................................................
Figure A.3. Gel of PioA-pFCM 21 expression in E. coli.....................................................160
6
List of Tables
Table 2.1. Reduction potentials and free energies of relevant compounds and proteins.
..................................................................................................................................
35
Table 3.1: Strains used in this w ork.............................................................................. 73
Table 3.2. Genes upregulated more than 5-fold following a 15-minute shock with 5 mM
Fe(II) and approxim ately 260 nM copper. ...........................................................
74
Table 3.3. Comparison of the growth delays of the metal treated cultures from Figure
3.7.............................................................................................................................
77
Table 4.1: Strains used in this w ork................................................................................110
Table 4.2: Prim ers used in this w ork. .............................................................................
111
Table A.1: Prim ers used in making constructs................................................................165
Table A.2: induction conditions for heterologous expression ........................................ 167
Table A.3: buffers tested for processing heterologous expression ................................ 169
Table A.4: Conditions tested for Homologous expression ............................................. 170
7
Chapter 1: Introduction
8
Motivation
Iron is an essential element that can nevertheless be toxic when present in
excess. Most research on iron's interactions with bacteria - particularly on its toxic
effects - has been done in the presence of oxygen. Iron can be expected to behave
quite differently under anoxic conditions for two reasons:
1. Significant levels of soluble iron can build up as Fe(II), which would be
oxidized and precipitated when oxygen is present.
2. Iron's main presumed mechanism of toxicity, the production of oxygen
radicals through the Fenton reaction (H20 2 + Fe2+ -+ Fe3+ + HO + OH-) (1,
2), does not occur under anoxic conditions.
Because of the lack of Fenton chemistry, anoxic iron has not generally been
considered a highly toxic metal; however, there are examples of anaerobic Fe(II) toxicity
(3, 4). The mechanism behind this toxicity is unknown.
Despite the toxic effects of iron, a number of bacteria can utilize high iron
concentrations anaerobically by oxidizing ferrous iron to gain energy and/or reducing
power. One type of anaerobic iron oxidation is photoferrotrophy (5), in which bacteria
grow using light as the energy source, CO2 as the carbon source, and Fe(II) as the
electron source.
Rhodopseudomonas palustris TIE-1, which was isolated as the first genetically
tractable photoferrotroph (6), is an excellent model organism for studying different
facets of the anaerobic bacterial iron interaction. Because it can grow using Fe(II) as the
sole electron source, TIE-1's iron oxidation pathway can be studied in detail using a
variety of techniques. In addition, TIE-i is a versatile bacterium that grows well
anaerobically using organic substrates and will generally not oxidize iron if an organic
substrate is available. This makes it a good model in which to explore the anaerobic
effects of iron during growth on other substrates.
Previous work with TIE-i identified the pio operon as essential to growth on
Fe(II) (7). This work set out to build on that knowledge and further explore iron
oxidation as a growth substrate. Our findings completely redefine our initial model of
9
Fe(Il) oxidation, and suggest that the Pio proteins are not the only ones involved. Along
the way, we also examine the detrimental effects of Fe(ll) on TIE-1, highlighting the fact
that bacteria respond very differently to the same compound when environmental
conditions are slightly altered.
Overview
Chapter 2 provides background on the anaerobic iron cycle and the bacteria that
take part in it. There are various bacteria that can oxidize or reduce iron depending on
their needs and on the other elements of the environment.
Chapter 3 describes the effects of iron on TIE-1 cultures that are growing on
acetate and not oxidizing Fe(II). Surprisingly, as TIE-1 grows photoferrotrophically with
millimolar concentrations of iron, the growth of acetate cultures was delayed by
micromolar levels of Fe(II). This seeming paradox was resolved by the discovery that the
inhibitory effect was the result of a synergy between the added iron and nanomolar
concentrations of copper. The synergistic phenomenon is also more broadly relevant;
not only does the synergy occur at environmentally relevant metal concentrations, but
Escherichia coli, Rhodobacter sphaeroides, and Shewanella oneidensis are also affected
synergistically by copper and iron.
Chapter 4 investigates the role of two high potential iron proteins (HiPIPs) in TIE1's response to iron. HiPIPs are often used as electron donors to the reaction center
during anaerobic photosynthesis in purple bacteria. TIE-i is unusual in that a) it has two
HiPIPs encoded in its genome, and b) neither one is sufficient for photosynthetic
growth, as the main donor to the reaction center is cytochrome c2. Previous work
showed that PioC is important in Fe(II) oxidation (7)and is up-regulated both by low
oxygen tension and during growth on iron (8). The second HiPIP in the genome,
Rpal_4085, has an unknown function, but was hypothesized to be partially redundant
with PioC. This chapter shows that this is not the case - while both proteins are
unequivocally HiPIPs, as demonstrated by their characterization, only PioC is capable of
10
being oxidized by the reaction center and participating in iron oxidation. Studies with
flash induced absorbance spectrometry indicate that the rate of reaction center
reduction by PioC is measurable, but much slower than reduction by cytochrome c2.
Though the function of Rpal_4085 is still not determined, it is transcriptionally upregulated in acetate cultures by several divalent metals, including Fe(II) - leading to the
intriguing possibility that these two seemingly similar proteins have unique and nonoverlapping functions that nevertheless both touch on the cell's various responses to
Fe(II).
In Chapter 5, Flash-induced spectrometry on whole cells helped us make some
very surprising discoveries; first, that although previous work (7) has shown PioA, PioB
and PioC (encoded by the pio operon) to be essential to rapid iron oxidation, the
quinone pool in mutants of these proteins was still reduced by iron, indicating that there
is an alternate pathway for electrons, at least on a small scale. Even more surprising,
reduction of the quinone pool by iron was completely blocked by the bci inhibitor
antimycin. This result indicates that, rather than traveling through the reaction center,
electrons from Fe(ll) reach the quinone pool through the bc1 complex. This result
directly contradicts our original model of Fe(II) oxidation, and forced us to develop a
new model based on the reverse electron transport chain in acidophilic Fe(ll) iron
oxidizers.
Chapter 6 considers some of the broader questions raised by this work, and
suggests future directions.
Some of the questions raised by this work could likely be addressed by
characterizing PioA, a decaheme protein that is likely the main iron oxidase. Appendix A
describes several unsuccessful attempts to purify PioA using various over-expression
systems and suggests options for continuing this attempt.
11
References
1. Cornelis P, Wei Q,Andrews SC, Vinckx T. 2011. Iron homeostasis and management
of oxidative stress response in bacteria. Metallomics. 3(6):540-549.
2. Touati D. 2000. Iron and oxidative stress in bacteria. Arch. Biochem. Biophys.
373(1):1-6.
3. Dunning JC, Ma Y, Marquis RE. 1998. Anaerobic killing of oral streptococci by
reduced, transition metal cations. Appl. Environ. Microbiol. 64(1):27-33.
4. Poulain AJ, Newman DK. 2009. Rhodobacter capsulatus catalyzes light-dependent
Fe(ll) oxidation under anaerobic conditions as a potential detoxification mechanism.
Appl. Environ. Microbiol. 75(21):6639-6646.
5. Widdel F, Schnell S, Heising S, Ehrenreich A, Assmus B, Schink B. 1993. Ferrous Iron
Oxidation by Anoxygenic Phototrophic Bacteria. Nature. 362(6423):834-836.
6. Jiao Y, Kappler A, Croal LR, Newman DK. 2005. Isolation and characterization of a
genetically tractable photoautotrophic Fe(II)-oxidizing bacterium,
Rhodopseudomonas palustris strain TIE-1. Appl Environ Microbiol. 71(8):4487-4496.
7. Jiao Y, Newman DK. 2007. The pio operon is essential for phototrophic Fe(II)
oxidation in Rhodopseudomonas palustris TIE-1. J Bacteriol. 189(5):1765-1773.
8. Bose A, Newman DK. 2011. Regulation of the phototrophic iron oxidation (pio)
genes in Rhodopseudomonas palustris TIE-i is mediated by the global regulator,
FixK. Mol Microbiol. 79(1):63-75.
12
Chapter 2: Bioenergetic challenges of microbial iron
metabolisms
Lina J. Bird, Violaine Bonnefoy, and Dianne K. Newman
This chapter was adapted from the published manuscript: Bird, L. Bonnefoy, V. and
Newman, D. (2011) Bioenergetic challenges of microbial iron metabolisms. Trends in
microbiology vol. 19(7) pp. 330-40
Contribution:
Dr. Bonnefoy wrote the section on acidophilic iron oxidation and made Figure 2.1. I
wrote the remaining portions and prepared the remaining figures.
13
Abstract
Before cyanobacteria invented oxygenic photosynthesis and 02 and H20 started
cycling between respiration and photosynthesis, redox cycles between other elements
were used to sustain microbial metabolism on a global scale. Today, these cycles
continue to occur in more specialized niches. In this review, we focus on the
bioenergetic aspects of one of these cycles-the iron cycle-because iron presents
unique and fascinating challenges for cells that use it for energy. While iron is an
important nutrient for nearly all life forms, we restrict our discussion to energy-yielding
pathways that use ferrous iron [Fe(II)] as an electron donor or ferric iron [Fe(Ill)] as an
electron acceptor. Here, we briefly review general concepts in bioenergetics, focusing
on what is known about the mechanisms of electron transfer in Fe(ll)-oxidizing and
Fe(Ill)-reducing bacteria, and highlighting aspects of their bioenergetic pathways that
are poorly understood.
Bioenergetics and diversity of iron metabolism
Billions of years ago, microbial iron metabolisms likely drove the carbon cycle
and catalyzed the global deposition of massive sedimentary ore deposits known as
banded iron formations [1]. Today, these microbial metabolisms remain highly relevant
in a variety of environments, ranging from anaerobic aquifers [2] to acid mines [3] to the
deep sea [4]. Because of their importance to modern biogeochemical cycles and their
potential usefulness in bioremediation and biotechnology, a number of informative
reviews have been written on the ecology, physiology, and diversity of these organisms
[5-13]. However, with a few exceptions [14], relatively little attention has been paid to
the bioenergetic underpinnings of these metabolisms, which isthe focus of this review.
Electron transfer metabolisms allow organisms to capture, store, and release
energy. Not only do these metabolisms profoundly impact the environments in which
they occur, they are also fascinating at the molecular level. Substrates that are oxidized
(give up electrons) are called electron donors (i.e. reductants); substrates that are
reduced (gain electrons) are called electron acceptors (i.e. oxidants). It is well known
14
that oxygenic phototrophs, such as plants, harvest energy from the sun by transferring
electrons from H2 0 to CO 2 , producing 02 and reduced carbon compounds - sugars, fats,
proteins, DNA, and numerous small metabolites. The details of this remarkable
bioenergetic feat have been studied for decades, and much is now understood about
how water is oxidized by the photosynthetic reaction center [15] and the path electrons
take to fix CO2 [16, 17]. Similarly, it iswell appreciated that heterotrophs, such as
animals, can obtain energy by oxidizing organic material to CO2 and transferring
electrons to 02 to make H2 0; mechanistic studies of the respiratory electron transport
chain date back nearly a century [18]. While important details regarding electron
transport in oxygenic photosynthesis and aerobic respiration remain unknown, it is fair
to say that the depth of understanding of these systems is orders of magnitude greater
than that of bioenergetic pathways involving iron.
One of the most exciting traits of bacteria and archaea is their ability to extract
energy from sources that are inaccessible to other life forms. A minimal constraint for
any catabolic (i.e. energy-yielding) pathway is the generation of a proton motive force
(PMF) across the cytoplasmic membrane that can be harnessed to synthesize ATP,
energize membrane transporters, and drive flagellar rotation. Microbes have diverse
ways of doing this, ranging from using coupling sites in the electron transport chain to
running the ATP synthase in reverse [16]. Nowhere isthis better exemplified than in the
case of microbial iron metabolisms. For example, rather than obtaining electrons from
water as described above, some photosynthetic bacteria oxidize Fe(ll) to fuel CO2
fixation (anoxygenic photosynthesis); other bacteria transfer electrons from organic
carbon to Fe(lll) instead of 02 (heterotrophic respiration or fermentation); still others
obtain energy by oxidizing Fe(ll) and reducing 02 or NO3 (lithotrophic respiration). While
the general principles of energy conservation are the same, the mechanisms vary.
Knowledge of what controls microbial iron metabolisms at a cellular scale is
necessary to predict their impact in bioremediation and mining environments.
Moreover, there is currently much interest in biofuel cells that use bacteria to generate
electricity or other forms of fuel. In many cases, the systems that bacteria use to
15
transfer electrons to and from electrodes are the same ones that they use to grow on
solid substrates, such as iron minerals [19]. Understanding the mechanisms of these
systems is vital if we wish to optimize them for the production of electricity or fuel. In
the following sections, we will review what is understood about: (i) coupling Fe(II)
oxidation to 02 reduction at acidic pH, (ii) coupling Fe(II) oxidation to CO2 reduction in
photosynthesis, and (iii) coupling Fe(Ill) reduction to organic carbon oxidation.
Fe(II) oxidation
Fe(II) oxidation is performed by many different types of bacteria, including
autotrophs and heterotrophs, phototrophs and chemotrophs, and aerobes and
anaerobes. Because Fe(II) oxidizes rapidly in the presence of 02 at neutral pH, Fe(Il)oxidizing bacteria are limited to environments with low or no 02, or to highly acidic
environments where abiotic oxidation is much slower.
LithotrophicFe(II) oxidation
Lithotrophic iron oxidizers obtain energy by coupling Fe(II) oxidation to the
reduction of a compound with a more positive reduction potential. For acidophilesmicroorganisms living at acidic pH-this terminal electron acceptor is 02. There are
some advantages to acidophilic Fe(II) oxidation. The main benefit is that Fe(II)
autooxidation by 02 is minimized, rendering Fe(ll) stable and readily available as an
electron donor for bioenergetic processes. Interestingly, while, the natural substrates
for most of the acidophilic iron oxidizers are minerals, such as pyrite (FeS 2) or
chalcopyrite (CuFeS 2), these Fe(II) minerals are considerably more soluble than Fe(Ill)
(hydro)oxides, thus acidophiles likely encounter Fe2' as their substrate. Another
advantage is that the midpoint potential (Em) of the 0 2/H 20 couple increases at low pH,
increasing the potential energy available from the Fe(II) oxidation (see below). However,
Fe(Il)-oxidizing acidophiles face several specific challenges as described below.
General challenges of acidophily. Several reviews on pH homeostasis in
acidophiles have been published [20-24]. Briefly, acidophiles maintain circumneutral
16
cytoplasmic pH by maintaining an inverted transmembrane electrical potential, Alp
(positive inside). The external medium acidity provides a large favorable chemical
potential of protons (H*) between the periplasm and cytoplasm, ApH. Protons enter the
cytoplasm through leakage, secondary H+pumps, or H*-translocating ATP synthases
leading to ATP synthesis. The PMF is created by the topography of the electron transport
chain components and is maintained by the removal of cytoplasmic H' by the reduction
of 02 to H20.
Specific challenges of iron oxidation. In the neutral pH of the cytoplasm, Fe(II) is
rapidly autooxidized, producing free radicals that damage macromolecules, leading to
cell death. Furthermore, the Fe(Ill) produced will clog and acidify the cytoplasm through
ferric oxyhydroxide precipitation. Acidophilic Fe(ll)-oxidizing bacteria appear to avoid
these problems by oxidizing Fe(II) outside the cell.
Energetics of Fe(ll) oxidation. The Em of Fe(II)/Fe(Ill) is about +0.77 V while
0 2/H 20 is +0.82 V at neutral pH. However, the involvement of H+in 02 reduction
(Equation 1) confers a pH dependence to the Em of the 0 2/H 20 couple, which increases
to +1.12 V at pH 2, making more energy available (Figure 2.1) [25].
2e +YO 2 + 2H* 4H 20 Equation 1
The localization of the 02 reduction site near the periplasmic face of the
membrane where the local environment is acidic allows the Em of 0 2/H 20 to be
increased, even though the H* used in the reaction comes from the neutral cytoplasm.
Fe(Il)-oxidizing autotrophs obtain both energy and reducing power from Fe(ll)
oxidation to fix CO2 and, if necessary, N2. Fe(II) oxidation must therefore provide not
only ATP but also reduced NAD*. Since the Em of NAD*/NADH is -0.32 V at cytoplasmic
pH, some electrons coming from Fe(ll) oxidation are pushed 'uphill' against the
unfavorable redox potential (Figure 2.1). This 'reverse' electron transport is driven by
the PMF [14, 25]. While this electron pathway has been described in the Gram-negative
bacterium Acidithiobacillusferrooxidans (see below), how the electron flow switches
between 02 and NAD* is not understood [14].
17
Electron pathway from Fe(l) to 02 in At. ferrooxidans. The electron transfer chain
between Fe(ll) and 02 was first proposed [25] and investigated [26] in At. ferrooxidans.
It has since been extensively studied ([27-30], and references therein). A schema of the
electron pathway based on diverse datasets is shown in Figure 2.1.
18
A
2 Fe(II)
pH 1.6-3
2 Fe(Ill)
Outer
Membrane
Periplasm
H+
H+
Inner
Membrane
H+
Alas*
-'
Cytoplasm
(pH 6.5)
NAD+
NADH
H20
1120+
ADP +F
H+H+H+
H+
B
Upl electron
transfer
-0.4
NAD'NADH
0
c,
+0.2
+0.4
+0.6 -
Cyc2
CycI
/Cyc
+0.8 - Fe(Ill)I Fe(II)
+1
+1.2 -2
H20
Downhill"
electron transfer
Figure 2.1. Electron transport in At.ferrooxidans. (A) Model of Fe(ll) oxidation in At.
ferrooxidans. Electrons extracted from Fe(ll) on the side facing the outside medium of
the outer membrane are transferred either to 02 or to NAD* on the cytoplasmic side of
the inner membrane. Electron pathway is indicated as dotted lines. Proton fluxes are
indicated as arrows. Cytochromes c are represented in cyan, blue copper protein in light
purple, bci complex in dark blue, terminal oxidase in red, NADHI1 complex in green and
ATP synthase in yellow. (B)Schematic of reduction potentials of the At. ferrooxidans
electron transfer pathway. The redox tower on the left represents the Em values under
physiological pH conditions [pH 2 of the periplasm for the Fe(ll)/Fe(lll) and the 0 2/H 20
couples and pH 6.5 of the cytoplasm for the NAD*/NADH couple]. Note that specific
potentials are reported at specific pH and substrate/product concentrations, and that
changing either parameter can have significant effects on the reduction potential; this
applies to all subsequent figures. Proteins with an undetermined Em are not shown.
Abbreviations: ATPase, ATP synthase; UQ, ubiquinone; Rus, Rusticyanin.
19
Electrons from Fe(ll) and metal sulfides, which are the natural substrate of At.
ferrooxidans, are conducted 'downhill' to 02 through an 'electron wire' spanning both
the outer and inner membranes. This wire consists of the outer membrane cytochrome
c type protein Cyc2, the periplasmic blue copper protein rusticyanin, the membranebound cytochrome c4 Cyc1, and the integral cytoplasmic membrane cytochrome oxidase
CoxBACD, where 02 is reduced to H20 (Figure 2.1). Proton translocation through the
oxidase combined with the consumption of protons in the cytoplasmic reduction of 02
helps the cell maintain a large difference in proton concentration between the
cytoplasm (pH 6.5) and the periplasm (pH 2).
Driven by the PMF, protons enter the cytoplasm through the ATP synthase and
also through membrane associated transport processes such as the antiport or symport
of solutes, the efflux or influx of metal(oid)s, etc. The PMF is also thought to provide the
energy to push the electrons 'uphill' to NAD*. In this pathway, electrons are transferred
from rusticyanin via the cytochrome c4 CycAl, the cytochrome bc 1 complex, and the
membrane associated quinones to the NADH dehydrogenase (NADHI1) complex. The
split of electron flow to NAD' ('uphill') or 02 ('downhill') has been proposed to occur at
the rusticyanin level. By adjusting the electron flow at the rusticyanin branch point, At.
ferrooxidans could balance NAD* and 02 reduction.
Fe(ll) oxidation in other acidophiles. The Fe(ll) oxidation pathways of some other
acidophiles have been proposed, mainly from functional genomics data. These pathways
have been reviewed recently ([27, 28] and references therein). The various organisms
differ in the components involved, even between phylogenetically related species [3134]. However, though the components of the pathway might differ significantly, the
bioenergetics of the different systems are presumed to be similar at least in Gramnegative bacteria: they all conserve energy by transferring electrons from Fe(II) to 02
through an outer membrane cytochrome c, a periplasmic protein, a membrane bound
cytochrome c and an integral cytoplasmic membrane terminal oxidase; in addition, they
pump electrons 'uphill' to NAD* through a bc1 complex.
20
Photosyntheticiron oxidation
Fe(ll) oxidation can supply electrons to fix CO2 in anoxygenic photosynthesis
according to the general reaction [35] given in Equation 2.
4Fe2+ + CO 2 + 11H 2 0
+
hv -> [CHO] + 4Fe(OH) 3 + 8H+ Equation 2
Phototrophic Fe(II) oxidation was first postulated by Hartman [36] and first
described in purple non-sulfur bacteria in 1993 [37]. Several other Fe(Il)-oxidizing purple
sulfur and non-sulfur bacteria and one green sulfur bacterium have since been isolated
[35, 38-42]. These bacteria all live at circumneutral pH.
Challenges of photosynthetic Fe(l) oxidation. Similar to their acidophilic
counterparts, photosynthetic iron-oxidizing bacteria face several challenges: (i) they
must be able to oxidize the various forms of Fe(ll) found at circumneutral pH, including
free ions, ligand bound Fe(II), and Fe(II) that istrapped in minerals, all of which have
widely varying reduction potentials (Table 2.1); (ii) they must transfer electrons uphill to
NAD*; and (iii) they must deal with the product of Fe(II) oxidation, Fe(lll), which
precipitates rapidly as ferric (hydr)oxide [Fe(OH) 3] at pH 7. Although genes catalyzing
phototrophic Fe(II) oxidation have been identified in two purple non-sulfur bacteria, the
details of how their protein products function are poorly understood. No studies have
yet described the genes or gene products involved in Fe(ll) oxidation by green sulfur
bacteria, but it is reasonable to assume that they will be different from those in purple
bacteria because their photosynthetic reaction centers are significantly different.
21
The pio operon. Rhodopseudomonas palustris strain TIE-1 is the only genetically
tractable phototrophic Fe(II) oxidizer known to-date and has been most explored at the
molecular level. An operon containing three genes is required for phototrophic iron
oxidation (the pio operon). These genes (pioA, pioB, and pioC) encode a periplasmic
decaheme cytochrome c type protein, an outer membrane porin, and a periplasmic high
potential iron protein (HiPIP), respectively. Both cytochromes and HiPIPs are often
involved in electron transfer reactions, and it is therefore likely that PioA and PioC
transfer electrons from Fe(II) to their destination in the cell, while the outer membrane
protein PioB could be involved in Fe(II) transport into or Fe(Ill) transport out of the cell.
In a genetic screen, an inner membrane ATP binding cassette (ABC) transport protein
and a protein homologous to CobS (a cobaltochelatase) were also found to be required
for phototrophic Fe(II) oxidation [40]. It is interesting to note that the cobS gene found
is a secondary copy of this gene; TIE-1 also has a full cob operon, suggesting that the
CobS detected in the screen serves a different function.
Thefox operon. Rhodobacter sp. SW2 is not yet genetically tractable, so genes
from this organism that stimulate phototrophic Fe(II) oxidation (the fox genes) were
identified by heterologous expression in Rhodobacter capsulatus SB1003 [43]. The genes
in this operon contain a diheme cytochrome c FoxE, a predicted quinoprotein FoxY, and
a predicted inner membrane transport protein FoxZ. While there are some similarities
to the R.palustris system in the sense that the redox proteins are predicted to be
periplasmic, these two systems are not homologous; the cytochromes differ
significantly, and the other proteins are not related.
Based on these limited studies and the reduction potentials of the various
players, we can draw a possible pathway for electrons from Fe(I1) to CO2 in R.palustris
(Figure 2.2).
22
Figure 2.2. Electron transfer in phototrophic iron oxidizers. (A)Potential path electrons
might take from Fe(II). The outer membrane, inner membrane and inner cytoplasmic
membrane (ICM; lamellar stacks) are shown; the photosynthetic machinery resides in
the ICM. Abbreviations: PioB, R.palustris strain TIE-i outer membrane protein which
might be involved in Fe transfer in and out of the cell; PioA, TIE-1 cytochrome; FoxE,
Rhodobacter sp. SW2 cytochrome; PioC, TIE-1 high potential iron protein; FoxY, SW-2
quinoprotein; reaction center, phototrophic reaction center; c2, cytochrome C2 ; bc 1 ,
cytochrome bci; Q, ubiquinones; NADHi1, NADH dehydrogenase. Dotted arrows denote
electron transfer. (B) Schematic of reduction potentials of the photosynthetic electron
transfer pathway. Red arrows indicate downhill reactions, blue arrows indicate uphill
reactions, and yellow brackets indicate the potential range. Abbreviations: P870,
bacteriochlorophyll; hv, light energy; Bph, bacteriopheophytin; UQA/UQB, ubiquinone A
and B; HiPIPs, known range for high potential iron proteins; Fe(llI)NTA, Fe(Ill)
nitrilotriacetic acid; Fe(llI)cit, Fe(Ill) citrate.
23
Open questions
The model in Figure 2.2 leaves many open questions related to the challenges
described earlier.
First, what form of Fe(II) does the cell oxidize? TIE-1 and SW2 can grow using
either Fe2+ or Fe(Il)-nitrilotriacetic acid (NTA) and Fe(ll)-citrate at pH 7. The reduction
potentials of these different forms range from -0.2 V for Fe2+/goethite to +0.385 V for
Fe-citrate, which is nearly as high or higher than the reduction potential of cytochrome
c2 from R.palustris (which increases with decreasing pH; [44]). Oxidation of bound Fe(II)
through the pathway shown in Figure 2.2 would therefore require PioA and PioC to have
reduction potentials between +0.385 and +0.45 V. It is also possible that the cell has
some way of removing Fe(II) from citrate or NTA and maintaining it in an environment
that lowers its reduction potential. If so, the mechanism awaits discovery.
Second, how do electrons get to NAD*? The model in Figure 2.2 shows electrons
flowing to the reaction center, a thermodynamically favorable reaction. However, it is
also possible that electrons are actually transferred to the bci complex, which would be
favorable in the case of Fe2+/ferrihydrite and is the pathway suggested for acidophilic
bacteria. It is also interesting to speculate what path the external electrons take once
they enter the transport chain. It is generally thought that they end up at the quinone
pool and are transferred to NAD* via the NADH1 complex. But how do they get to the
quinone pool? Do they flow in reverse, to bci, and finally to the quinone pool? Or do
they become excited at the reaction center and go through the forward cycle to the
quinone pool? Or is there another, separate pathway by which they get there?
Third, where is Fe(II) oxidized? This question is intimately tied to the problem of
Fe(Ill) precipitation. Logically, it seems that Fe(II) should be oxidized at the cell surface
bypassing the problem of periplasmic precipitation. Intriguingly, however, all the
proteins involved in phototrophic Fe(II) oxidation described so far (i.e. PioA, PioC, FoxE
and FoxY) are predicted to be periplasmic based on their sequence and some
preliminary biochemical evidence in the case of PioA [35]. If oxidation does occur in the
periplasm, there are several potential ways that the cell might avoid periplasmic Fe(IllI)
24
precipitation, such as producing ligands to bind Fe(IllI) or rapidly transporting Fe(Ill) out
of the cell. Miot et al. [45] have shown that in SW2, Fe precipitation occurs outside the
cell on organic fibers that are attached to the bacteria and that the precipitates start as
Fe(lll)/Fe(II) mixed valence minerals which are converted to Fe(Ill) minerals over time.
These results are consistent either with oxidation taking place outside the cell, or with
Fe(Ill) oxidized in the periplasm being rapidly pumped out of the cell and precipitating
on the surface. This second possibility might be facilitated by a close association of the
oxidase with an outer membrane protein (e.g. PioA and PioB in TIE-1); in this case, we
would predict Fe(II) oxidation to localize to the periplasmic face of the outer membrane
so that Fe(Ill) is transported out of the cell before it has a chance to precipitate
intracellularly. It is more difficult to imagine the results of Miot et al. being due to ligand
binding of Fe(Ill), unless the Fe(llI) has a stronger affinity for the carbon fibers than for
the ligand.
Fe(lII) reduction
In the absence of oxygen, many microbes can use Fe(Ill) as an electron acceptor,
reducing it to Fe(ll). Iron reduction has been observed under both acidophilic and
neutrophilic conditions.
As discussed above, the Em of Fe(lll)/Fe(II) at low pH is quite high: +0.77 V. The
coupling of organic carbon oxidation to Fe(Ill) reduction is therefore a quite favorable
reaction. While a wide range of acidophilic bacteria are capable of taking advantage of
this potential in the absence of oxygen [46], relatively little is known about how they do
this, thus we restrict the remainder of our discussion to what is known about the
molecular mechanisms of neutrophilic Fe(Ill) reducers.
Neutrophilic Fe(Ill) reduction poses two main challenges for microorganisms.
First, Fe(Ill) is often in solid form. Second, different forms of iron have widely varying
midpoint potentials, some of which are quite low, thus limiting the energy available
from organic carbon oxidation. Here, we will describe what is known about the
25
bioenergetics of Shewanella and Geobacter species, the two best studied Fe(lll)reducing bacteria.
Both Shewanella and Geobacter are able to grow heterotrophically by conserving
energy from the breakdown of organic carbon. In the general model of heterotrophic
growth (in both bacteria and mitochondria), electrons from organic carbon reduce a
redox active small molecule, such as NAD* to NADH. Electrons from NADH are then
passed to the quinone pool via the NADH1 complex, which translocates protons and
builds up the proton gradient. During aerobic respiration, electrons proceed through the
ubiquinone pool and several additional proton translocating complexes until they reach
02
(shown for Shewanella in Figure 2.4B). Protons translocated through this process
reenter the cytoplasm through the ATP synthase (generating ATP), or through other
channels, doing other work. When a less thermodynamically favorable acceptor than 02
is used, electrons are instead transferred from NADH1 to the menaquinone pool and
eventually to the terminal electron acceptor.
Pathways for electron transfer to Fe(Ill)
The pathway for Fe(Ill) reduction by Shewanella and Geobacter has been
described in several reviews [5-8] and isshown in Figures 2.3A and 2.4A. Electrons are
thought to travel from the inner membrane through the periplasm and the outer
membrane through a series of multiheme cytochromes with overlapping reduction
potentials (Figures 2.3 and 2.4) [47]. Three mechanisms-which are not mutually
exclusive-have been suggested for electron transfer from the outer membrane to
Fe(Ill) minerals: (i) directly from outer membrane c-type cytochromes, (ii) indirectly via
electron shuttles, or (iii) indirectly via the solubilization of Fe(Ill) by organic chelators
[48]. In support of the first mechanism, Shewanella and Geobacter outer membrane
cytochromes have been shown to reduce iron in vitro [49-52], and Geobacter
metallireducens requires direct contact with solid Fe(Ill) to grow [53]. However, in
theory, direct electron transfer would require that the donating cytochrome be within
20 angstroms of solid Fe(Ill) [54] which would severely limit electron flow within a
26
biofilm, and suggests that other mechanisms must also be at play [55]. In support of the
second mechanism, Shewanella has been shown to secrete riboflavins and flavin
mononucleotides when growing on iron, fumarate, and electrodes [56]. These flavins
act as electron shuttles to solid, extracellular oxidants [e.g. Fe(Ill) minerals or
electrodes], becoming reduced at the cell surface and oxidized extracellularly.
Geobacter species do not appear to produce endogenous electron shuttles but Fe(Ill)
reduction can be greatly stimulated by the addition of exogenous electron shuttles (i.e.
flavins, quinones, and humic substances) [53]. Another solution to transferring electrons
at a distance has been suggested to be through 'nanowires', long pili-like appendages
produced by both Shewanella and Geobacter that have been shown to conduct
electrons [57, 58]. Finally, in support of the third mechanism, soluble Fe(Ill) has been
detected in cultures of Fe(Ill) grown Shewanella alga [59]. It has also been reported that
Shewanella putrefaciens and oneidensis species produce an organic chelator [60], but
these claims are controversial. More work is needed to confirm and identify this
putative chelator.
27
B
chelated Fe(Ill)
Outer
Membrane
H*
Periplasm 2H+
Inner Membrane
Cytoplasm
-MQ
NADH
ATF as*
..
-
NAD* H+
ADP + P,
2H+
ATP
H+
A
v
-0.42-0
0
+0.2
+0.4
OmB
ferriydrite/Fe2.
OmcS
NADHI
i
NAD-/NADH
PdcA MCINH
2
F.(ftIWe(I) ci
Fe IINTAl I)NTA
+0.6
+0.8
+1
Figure 2.3. Model of Fe(lll) reduction in Geobacter sulfurreducens. (A)Potential
pathway for electrons through the membranes and periplasm. The pathway is still
uncertain, as there are many cytochromes expressed in Geobacter under Fe(Ill) reducing
conditions. Abbreviations: NADH1, NADH dehydrogenase; MQ, menaquinone; PpcAD
and OmcBES, cytochrome c type proteins; OmpB, multicopper protein. (B)Schematic of
reduction potentials of the Geobacter electron transfer pathway. Abbreviations: NTA,
nitrilotriacetic acid; cit, Citrate.
28
A
.
Fla ,in
Chelated Fe(Il).
Outer
Membrane
Periplasm
H+
--
Inner
Membrane
+
MQ-+
2eCytoplasm
NADH
NAD*
H+4
Acetate + ATP
Lactate
B
V
-0.4
-0.2
0
a
9gnqt1
C mA
2
MtrC OmcAMt
STC Cy+A--.-M
2
ferrihydrteFe
Fe(Ill)cft/Fe(lI) cit
+0.4
FelIQN7AlF(lI)NTA
AD
ADH
rUQ
+
+0.2
H
Cc
+0.6a
+0.8
02/H20
+1
Figure 2.4. Electron transfer to Fe(Ill) in Shewanella. (A) Path electrons take from the
cytosol to Fe. How electrons enter the electron transport chain is unclear, so we indicate
a general protein at the start of the chain (green box, labeled with '?'). MtrC and OmcA
are thought to be donors to Fe(Ill). Whether they interact directly with solid Fe(III),
chelated Fe(Ill), electron shuttles such as flavins, or all three is unclear. ATP generation
occurs via substrate level phosphorylation. (B) Schematic of reduction potentials of the
Shewanella electron transfer pathway. Arrow on scale bar denotes the energetically
favorable direction. Blue arrows denote the aerobic path electrons take to 02, red
arrows denote the path to Fe(Ill), and yellow bars denote potential ranges. The
potentials of NADH1, bc1 , and aai are simplified for clarity; in reality, each of these
protein complexes has multiple redox centers and a range of potentials. Abbreviations:
CymA, MtrA, STC, MtrC, and OmcA, c-type cytochromes; MtrB, outer membrane
protein; UQ ubiquinone; MQ4 menaquinone; bci, bci cytochrome complex; Cyt c,
cytochrome c; aai, cytochrome c oxidase; NTA, nitrilotriacetic acid; cit, Citrate.
29
Bioenergetics of Fe(IlI) reduction
The environmentally relevant potentials of various Fe(llI)/Fe(II) couples range
from +0.382 V to -0.3 V (Table 2.1), and the energy available from organic carbon
oxidation changes accordingly. Chelated Fe [such as Fe(Ill)-citrate and Fe(Ill)-NTA] is on
the favorable end of the spectrum; however, neither Geobacter nor Shewanela extracts
the maximum energy available from chelated Fe(Ill) as evidenced by their poor growth
yields.
In Geobacter, growth yields on acetate/Fe(Il)-citrate are far below the
predictions based on the free energy (AG) available from the reaction [61, 62]. There are
two reasons for this low efficiency. First, the extracellular location of iron reduction is
energetically costly. Organic carbon oxidation takes place inside the cell, and produces 1
H per e~. When a cytoplasmic acceptor such as 02 or fumarate is used, the proton
production is balanced by the consumption of 1 H* per electron accepted. In contrast,
an external electron acceptor such as iron consumes no cytoplasmic protons. Using any
external electron acceptor thus costs the cell one proton per electron transferred.
The second reason for low efficiency is that the electron transport chain is short.
In the current model of electron transfer to Fe(Ill), no energy is harvested after the
electrons reach the periplasm. The amount of energy extracted from the electron
transport chain therefore depends not on the potential difference (AE) between NADH
and the final acceptor, but on the AE between NADH and the periplasmic acceptor. In
Geobacter, the periplasmic cytochrome PpcA isthought to accept electrons from an
unknown membrane partner (Figure 2.3A). The AG for electron transfer from NADH to
PpcA is much closer to the energy extracted by the cell, based on growth yields. It
therefore seems that Geobacter has adapted to use low potential substrates [e.g. Fe(lll)
minerals] rather than maintaining an electron transport chain that would allow it to
extract greater energy from higher potential acceptors [e.g. Fe(Ill)-citrate]. This is born
out experimentally: Geobacter biofilms grow on electrodes with an Em as low as -0.15 V
[63, 64], and more positive electrodes do not yield more cells [65].
30
Unlike Geobacter, Shewanella oneidensis cannot anaerobically grow on acetate.
Furthermore, deletion of the ATP synthase does not significantly impair the growth of S.
oneidensis strain MR-1 on lactate with fumarate as the electron acceptor [66]. This
suggests that Shewanella relies on substrate level phosphorylation rather than the
electron transport chain for ATP synthesis and NAD* regeneration when growing on
fumarate. This is not surprising because in S. oneidensis fumarate is reduced (and uses
up protons) in the periplasm instead of the cytoplasm [67]. Substrate level
phosphorylation also has been suggested to be the primary mode of energy generation
during growth on Fe(llI) [66}. Interestingly, Shewanella grows on extremely low
potential acceptors such as magnetite [68]. If it uses substrate level phosphorylation to
gain energy when reducing Fe(Ill), this may explain why Shewanella can take advantage
of an even wider range of potential electron acceptors than Geobacter.
Box 1: General Iron Chemistry
In order to understand the nuances of biological transformation, we must first
have a basis in the general chemistry of iron. The state of iron in the environment
depends on many factors, including pH, Eh, and the presence of complexing agents. The
reduction potential (E)of Fe2 +/Fe3 , in the absence of precipitation (which only occurs at
pH < 3), is about 0.77 V. At pH > 3, the formation of a solid effectively removes Fe(Ill)
from solution, making iron oxidation more favorable (lowering E). The amount that the
reduction potential is lowered depends on the solubility of the mineral formed. Iron
hydroxide [Fe(OH) 3], for example, is poorly crystalline and more soluble than ordered
minerals such as hematite [Fe 20 3]. The potential of Fe(OH) 3/Fe(II) is therefore
significantly higher than that of, for example, Fe20 3/Fe(II) because hematite is a less
soluble mineral. Chelators can lower or raise the reduction potential, depending on
their properties. Citrate and nitrilotriacetic acid (NTA), for example, bind more tightly to
Fe(lli) and stabilize it, thus lowering the reduction potential [69]. Other chelators bind
more tightly to Fe(II) thus raising the reduction potential. Some of the potentials
relevant to this review are listed in Table 2.1.
31
It is important to note that while this review deals primarily with thermodynamic
constraints on microbial iron metabolisms, not everything can be explained from a
thermodynamic perspective. This is because metabolic reactions must not only be
thermodynamically favorable, but kinetically favorable as well. Abiotic oxidation must
proceed slowly enough that microorganisms can take advantage of the reaction; at the
same time, the iron must be in a form that isaccessible to the microorganisms. Several
studies have shown that rates of Fe(Ill) reduction by microorganisms depend on the
solubility of iron minerals (70] or the affinity of the ferric chelator [71]. Understanding
iron metabolisms in the environment thus requires a grasp of both thermodynamics and
kinetics. Finally, random quirks of evolution ultimately dictate what is possible or not:
even if a substrate is thermodynamically and kinetically favorable, an organism must
possess the machinery required to recognize it.
Conclusions
Iron has been used by bacteria to generate energy for billions of years, yet only
recently have we begun to understand how they accomplish this. While we have a
general idea of how a few select bacteria have organized their membranes to gain
energy from oxidizing Fe(ll) or reducing Fe(Ill), much remains to be learned about these
processes. Moreover, many other organisms that utilize iron for energy have not yet
been studied in mechanistic detail. Understanding the different ways bacteria have
evolved to grow on iron isfascinating at a basic level because of the interesting cell
biological and biochemical challenges iron metabolisms pose. Mechanistic insights into
these processes also have the potential to be useful in interpreting iron biosignatures on
the early Earth [41] and informing applications ranging from biomining to alternative
energy. Towards these ends, a few questions that only scratch the surface of the
opportunities that exist for future research are listed below in Box 2. It is our hope that
this review will encourage new investigators to enter this field, bringing with them tools
to answer these questions, and fresh perspectives to ask new ones.
32
Box 2: Outstanding questions
e
How do microbes couple Fe(II) oxidation to the reduction of nitrate and
other electron acceptors such as perchlorate? Can a mechanistic
understanding be exploited in the context of bioremediation?
*
How do microbes deal with insoluble byproducts of iron oxidation at
neutral pH?
e
How do the integral membrane components of the electron transport
chain in acidophiles, such as the ATP synthase, the terminal oxidase, the
bci and the NADH1 complexes deal with the pH difference between the
periplasm and the cytoplasm?
e
How do Fe(II)-oxidizing microbes balance the need for ATP with the need
for reducing equivalents? In other words, how is reverse electron
transport achieved and regulated?
e
What forms of iron do cells actually 'see' in the real world? Defining the
nature of Fe(II) and Fe(Ill) complexes iscrucial both for accurate
bioenergetic calculations and biochemical understanding. For that
matter, what isthe local environment like around any given step in the
electron transport pathway? Perhaps steps we currently perceive as
being thermodynamically 'uphill' [e.g. electron transfer from Fe(II) to
Cyc2 in At. ferrooxidans], might simply be an artifact of our incomplete
understanding of the local environment relevant for that step. For
example, the midpoint potential of CycAl in At. ferrooxidans has been
shown to increase by 0.60 V when it forms a complex with rusticyanin
[72] while that of rusticyanin decreased by > 0.1 V when it is associated
with Cyc1 [72, 73].
e
How do electrons enter the electron transport chain in Fe(Ill)-reducing
bacteria? We have shown the pathways involving NAD*/NADH; however,
it is possible that other electron carriers, such as formate from the
breakdown of pyruvate [66], may also play a role.
33
e
How do the proteins involved in these processes localize to their proper
sites in the cell?
e
What are the 'design principles' that underpin the cellular localization of
the proteins and small molecules required for these metabolisms? Are
there trade-offs between energetic efficiency, minimization of toxicity,
and/or metabolic range?
Acknowledge me nts
We thank Jeffrey Gralnick and Daniel Bond for helpful conversations, and
Sebastian Kopf for assistance in thermodynamic calculations. DKN and UB are supported
by the Howard Hughes Medical Institute.
34
Table 2.1. Reduction potentials and free energies of relevant compounds and proteins.
Reduction pair
Eenv* (volts)a
AG (kJ/mol)b
Refs
Components of photosynthetic electron transport chains relevant to Figure 2.2
P8 70
Ps70*
Bph
UQA
UQ
Cytochrome bc (b)
Cytochrome bci (c1)
Cytochrome bc2 (Rieske)
+0.45
-1.1
-43.4
106.1
[16]
[16]
-0.6
-0.2
+0.08
+0.05 and -0.09
+0.285
+0.28
57.9
38.6
-15.4
-4.8 /8.7
-27.5
-27
[16]
[16]
[16]
[79]
[80]
[81]
+n- PC
f~i1
35
Endogenous and exogenous
Riboflavin
Monoflavin nucleotide
Humic substances
-0.2 to +0.3
-77 to 19
(87)
indicates environmentally relevant midpoint potentials: pH 7 except where noted,
standard concentrations except for solid Fe minerals, for which Fe2+ is 100 P M.
bAG calculations assume standard conditions and pH 7, except in the case of iron
aEenv*
minerals where [Fe 2+]is assumed to be 100 M.
36
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44
Chapter 3: Iron and copper act synergistically to
delay anaerobic growth in bacteria
Lina J. Bird, Maureen L. Coleman, and Dianne K. Newman
This chapter was adapted from the following manuscript: Bird L. Coleman M, and
Newman, D. (2013) Iron and copper act synergistically to delay anaerobic growth in
bacteria. Applied and Environmental Microbiology in press.
Contribution:
Dr. Coleman collaborated to perform the microarray and analyze the resulting data. She
did some of the initial growth curves with iron. I wrote the text with input from Dr.
Coleman and Dr. Newman, and prepared the figures.
45
Abstract
Transition metals are known to cause toxic effects through their interaction with
oxygen, but toxicity in anoxic conditions is poorly understood. Here we investigated the
effects of iron (Fe) and copper (Cu) on anaerobic growth and gene expression in the
purple phototrophic bacterium Rhodopseudomonas palustris TIE-1. We found that Fe(Il)
and Cu(II) act synergistically to delay anaerobic growth at environmentally relevant
metal concentrations. Cu(l) and Cu(ll) had similar effects both alone and in the presence
of ascorbate, a Cu(II) reductant, indicating that reduction of Cu(II) to Cu(l) by Fe(II) is not
sufficient to explain the growth inhibition. Addition of Cu(II) increased the toxicity of
Co(II) and Ni(ll); by contrast Ni(II) toxicity was diminished in the presence of Fe(II).
Synergistic anaerobic toxicity of Fe(II) and Cu(II) was also observed in Escherichia coli
MG1655, Shewanella oneidensis MR1, and Rhodobacter capsulatus SB1003. Gene
expression analyses in R.palustris identified three regulatory genes that respond to
Cu(II) and not Fe(lI): homologs of cueR and cusR, two known proteobacterial copper
homeostasis regulators, and csoR, a copper regulator recently identified in
Mycobacterium tuberculosis. Two P-type ATPase efflux pumps, along with an F1 Fo ATP
synthase, were also upregulated by Cu(II) but not Fe(II). An E.coli mutant deficient in
copA, cus, and cueO showed a smaller synergistic effect, indicating that iron might
interfere with one or more of the copper homeostasis systems. Our results suggest that
interactive effects of transition metals on microbial physiology may be widespread
under anaerobic conditions, though the molecular mechanisms remain to be more fully
elucidated.
46
Introduction
Copper (Cu) and iron (Fe) are essential but potentially toxic metals. While much
is known about these metals individually, their combined biological effect(s) under
environmentally relevant conditions has not been studied. This gap in our knowledge is
partially due to the fact that although iron is abundant in the Earth's crust, in the
presence of oxygen it occurs mainly as poorly soluble Fe(Ill), and aerobic organisms
more often face iron deficiency than excess. However, that bacteria thrive under anoxic
conditions in diverse habitats has been appreciated since the work of Winogradsky in
the late
19 th
century (1). Today, it is well known that soluble Fe(II) can build up in anoxic
environments, and this is often due to microbial activity (2). Indeed, soluble iron in
groundwater has been measured at concentrations up to several hundred micromolar
(3). The effect of excess dissolved iron on microbial physiology, especially under
anaerobic conditions, deserves further attention.
Iron is known to be toxic through the Fenton reaction (H20 2 + Fe2 -+ Fe3 + HO- +
OH-) (4-6), but this mechanism depends on the presence of 02 and thus cannot account
for anaerobic toxicity. Indeed, iron has not traditionally been considered a toxic metal
under anoxic environmentally relevant conditions, although there are a few reports of
anaerobic iron toxicity in the literature (7, 8). A mechanism for anaerobic iron toxicity
has not been established, and the toxicity reported is not a universal trait: a number of
bacteria, including dissimilatory iron reducers such as Shewanella and Geobacter and
iron oxidizers such as Rhodobacterferooxidans SW2 and Rhodopseudomonas palustris
TIE-1, tolerate millimolar concentrations of Fe(II) with no obvious growth handicap.
Copper toxicity, in contrast, has been much better studied because it is a welldocumented pollutant. A survey of public testing data showed that copper levels in US
groundwater range from undetectable to approximately 1 mM, with most samples in
the nanomolar range (9). Micro- to millimolar concentrations that represent the high
end of the range, are found particularly in industrial and mining waste streams (10, 11).
There are several mechanisms by which copper is toxic (6, 12, 13). Under aerobic
conditions, copper, like iron, can react with hydrogen peroxide to form oxygen radicals
47
(H 20 2 + Cu() -
Cu(Il) + HO- + OH-) (14). Cu(I) also interferes with the iron-sulfur clusters
of some proteins in an oxygen-independent process (15). To combat these toxic effects,
bacteria have sophisticated copper homeostasis systems that can include several types
of efflux pumps, chaperones, ligands, and oxidases (6, 12, 13).
Copper toxicity has also been studied in conjunction with other metals (16). The
effects of copper combined with toxic metals such as zinc, cadmium, lead, silver, and
nickel have been extensively documented in a range of organisms (16, 17). In most of
these studies, the effects were found to be additive or even antagonistic (combinations
were less toxic than single metals), but in some cases synergistic toxicity was apparent.
To our knowledge, all of these studies were performed under oxic conditions.
The combined effect of copper and iron on microorganisms is worth considering
because these metals co-occur in a number of environments, particularly anthropogenic
environments: studies of groundwater and soil near landfills (18), soil in cemeteries
(19), and soil near metal recycling plants (20) have all documented the co-occurrence of
significant levels of copper and iron. Wetlands can also show increased concentrations
of dissolved iron and copper in their anoxic zones (21), and because wetlands are often
constructed to treat mine, industrial and municipal wastes that can contain very high
heavy metal levels (22), metal behavior and influence in wetlands is of particular
interest.
To explore the possible synergistic effects of Fe(II) and Cu(ll) on anaerobic
bacterial growth, we chose Rhodopseudomonas palustris TIE-1, a purple non-sulfur
phototroph, as our model organism. R. palustris strains have long been studied for their
diverse metabolisms, and their ability to break down aromatic compounds (23, 24) has
generated interest in the context of bioremediation. Because sites polluted with
aromatics may also contain metal contamination, R. palustris' response to trace metals
is relevant to bioremediation efforts.
48
Materials and Methods
Growth media and culturingconditions
Table 3.1 lists all strains tested in this work. R.palustris TIE-1 and R.capsulatus
SB1003 were grown anaerobically in minimal freshwater medium as previously
described (25) with a lowered phosphate concentration and an alternate trace element
mix: the basal media consisted of 100 pM KH2 PO4 , 5.61 mM NH4 CI, 0.68 mM CaCl 2, and
2 mM MgSO 4. After autoclaving, media was cooled under 20% CO2 80% N2 gas. After
cooling, 30 mM sterile NaHCO 3 , vitamin mix (0.29 pM 4-Aminobenzoic acid, 41 nM D (+)
biotin, 0.81 pM Nicotinic acid, 0.21 iM Ca-(+) pantothenate, 0.41 pM Pyridoxamine
dihydrochloride, 0.29 pM Thiamine dichloride, 1.32 pM Riboflavin), B12(0.1 mg/ml) and
trace element mix (7.2 pM FeCI 2e4H 20, 0.51pM ZnCl 2 , 0.48 pM MnCl 204H 20, .76 pM
CoCl 2 e6H 2 0, 11.1 nM CuCl 2 e2H 2 0, 95.8 nM NiC
2 e6H 2
0, 0.146 pM Na2 MoO 4 e2H 20, 97
nM H3 BO3 ) (26) were added. The cooled medium was adjusted to pH 7 with 1M Na2 CO3
and stored in an anaerobic glove box (Coy Laboratory Products) under a 5% H2 15% Co2
80% N2 atmosphere. Acetate was added before inoculation (10mM final concentration).
For routine aerobic growth YPS-MOPS medium (3g/L yeast extract, 3g/L peptone, 10
mM succinate, 20 mM MOPS, pH 7)was used. Cultures were grown in sealed balch
tubes at 300 Cin a light incubator for anaerobic cultures, and in a dark shaking incubator
for aerobic cultures.
For E.coli MG1655 and S. oneidensis MR-i growth curves, minimal MOPS media
was used containing 1 mM MgSO 4 , 3 mM KH2 PO4 , 7.5 mM (NH4 )2SO4, 30 mM NaCl, and
30 mM MOPS pH 7 (27). After autoclaving and cooling under N2, 5 mg/L thiamine and
0.5 mM histidine were added from filter sterilized stock, along with the vitamin and
mineral mixes described above. Media was stored in the anaerobic chamber until use.
Before inoculation 0.3% glucose was added to E. coli cultures, and fumarate and lactate
(final concentration 20 mM each) were added for S. oneidensis. Cultures were grown in
sealed balch tubes at 370 C. LB broth was used for routine aerobic growth.
For CFU counts, freshly inoculated cultures were serially diluted in anaerobic
YPS-MOPS media and plated on YPS-MOPS plates (YPS-MOPS media with 15% Bacto-
49
agar). Plates were grown at 300 Cunder light in a Coy anaerobic chamber for 5 days,
and resulting colonies were counted. Additional experiments were performed with E.
coli strains provided by J.Imlay (see "Analysis of E.coli growth curves" below).
Microarray
R.palustris cultures were grown to mid-exponential phase and transferred to an
anaerobic chamber (Coy). 7 ml of each culture was withdrawn and added to an equal
volume of RNALater (Ambion). These samples were then spun for 6 minutes at 5,000xg,
the supernatant was poured off, and the pellets flash frozen and stored at -800 C.
Cultures were then shocked with 5mM Fe(II) (contaminated with 260 nM copper).
Samples of shocked cultures were taken at 15 minutes to determine the rapid
transcriptional response. Many bacteria respond to stress within 15 minutes, and
preliminary tests indicated this held true for R.palustris.
RNA extraction was performed with the RNeasy minikit (Qiagen) using the
digestion, lysozyme, and mechanical disruption method described in the manufacturer's
protocol. Isolated RNA was treated with Turbo DNAse (Ambion) using the rigorous
treatment protocol to remove contaminating DNA and quantified by absorbance with a
Nanodrop spectrophotometer. RNA quality was confirmed on an Agilent Bioanalyzer,
and by checking 260/280 and 260/230 nm absorbance ratios on a Nanodrop
spectrophotometer. Total RNA was amplified using MessageAmp 11Bacterial RNA kit
(Ambion) according to manufacturer's directions, with aminoallyl UTPs incorporated at
the final step. Cy3 dyes (Amersham/GE) were incorporated using the standard protocol.
Labeled RNA was hybridized to a custom Agilent microarray, which contained 60-mer
oligonucleotide probes covering all protein-coding genes and non-coding RNA features
based on the R.palustris TIE-1 genome annotation available in Genbank (accession
number NC_011004). The microarray data was analyzed in Rusing the limma package
(28). Differentially expressed genes were defined as having a p-value less than 0.05 after
Bonferroni correction for multiple testing.
50
Metal shocks, RNA isolation and qRT-PCR
R. palustris cultures were grown anaerobically as described above and brought
into an anaerobic chamber for the shock. In order to obtain enough cell mass for RNA
isolation, we performed the shock on mid-log phase cultures. Shocks were performed
with 1 mM Fe(II) and/or 1 pM Cu(ll). These concentrations were chosen to avoid
significant copper contamination in the Fe(II) shocks, while maintaining metal
concentrations high enough to ensure a transcriptional response. After 15 minutes, 6
mls of culture were added to an equal volume of RNALater (Ambion), and centrifuged
for 6 minutes at 6,000xg at room temperature. The supernatant was poured off, and
the pellets frozen in liquid nitrogen and stored at -80 0 C until RNA extraction.
RNA was extracted using the method described above. 40 ng of RNA was used
to make cDNA using the iScript" cDNA Synthesis Kit (Bio-Rad) with random primers. 3
pl of the cDNA reaction was assayed in a 25 pal qRT-PCR reaction using the iTAQ SYBR
Green supermix with ROX (Bio-Rad). qRT-PCR was performed with the 7500 fast real
time PCR system (Applied Biosystems), and CT values were calculated using Sequence
Detection Software Version 1.4.0.25. Gene expression was normalized to recA using the
AACT method (29). clpX was used as an internal control, and samples that showed a
more than 2-fold change in the corrected c/pX value were re-run. Primers used for each
gene are listed in Table 3.2.
Growth curves and preparation of metal stocks
To better characterize the effects of metals on R. palustris, cultures were
inoculated in media containing metals, and growth was monitored. Measuring growth in
the presence of metals provided a simple and reproducible way to measure the
inhibitory effect of metals, and has been used in other toxicity studies (15, 27). Growth
curves were performed in 10 ml of the media listed above, using sealed 25 ml balch
tubes. For growth assays, metals were added to individual tubes before inoculation. R.
palustris and R. capsulatus growth curves were inoculated in a Coy anaerobic chamber
from stationary anaerobic cultures with a 1:200 inoculum. E. coli cultures were started
by diluting an overnight anaerobic minimal medium culture 1:100, and S. oneidensis was
51
diluted 1:50 from an anaerobic LB culture. Fe(II), Cu(II), Ni(II), and Co(II) stocks were
prepared by adding FeCl 2 (Sigma 98% pure or Alfa Aesar 99.9% pure) , CuCl 2, NiC 2 or
CoC12 (Fisher) to nanopurified water under anoxic conditions. Cu(l) was prepared under
anoxic conditions from CuCl (Alfa Aesar) as a 100 pM stock in 1 M NaCl, with heating to
370 Cto aid dissolution. Bacterial culture growth was monitored by measuring optical
density of balch tubes on a Spectronic 20D+ spectrophotometer at 660 nm for R.
palustris and R. capsulatus and 600 nm for E.coli and S. oneidensis.
ICP-MS
Levels of copper in the stocks of iron prepared from Sigma FeCl 2 (98% pure) and
Alfa Aesar FeCl 2 (99.9% pure) were measured at the Caltech Environmental Analysis
Center on a Hewlett-Packard 4500 ICP-MS System. Samples of 1 M stock solution were
diluted 1:450 in nitric acid, and sampled. The results were quantified using the standard
addition method, by adding pure CuCI 2 to each sample to a final concentration of 10, 20,
and 30 pg/L. The lines obtained were extrapolated to a Y of 0, and the absolute value of
the X intercept was taken as the amount of copper in the original sample.
Abiotic Cu(II) reduction
Abiotic Cu(ll) reduction was tested by mixing 40 pM Cu(II) with 100 pM Fe(II) or
ascorbate in fresh water medium in an anaerobic chamber. Cu(l) concentrations were
determined using the bathocuproine assay (30): 200 pL of sample was added to a 96well plate containing 50 pL bathocuproine (50% ammonium acetate, .03%
bathocuproine sulfonate, 2 mM EDTA). Samples were read at 484 nm on an anaerobic
Synergy 4 plate reader (BioTek). A standard curve was made with CuCl.
Analysis of E. coli growth curves
Growth curves were performed as described above for E. coli MG1655* and the
mutant copA::kan AcueO AcusCFBA::cm (Table 3.1). Lag phases were calculated in R
using the Gompertz model (31). Averages from triplicate cultures were used for the
52
comparison of lag times for different Cu(II) and Fe(Il) concentrations. The delay in
growth caused by addition of copper or iron was calculated by subtracting the lag phase
of the control culture from the lag phase of the metal treated culture. The synergistic
effect of Fe(ll) and Cu(II) was calculated by dividing the delay in growth of the Fe(ll)
Cu(II) condition by the delay in growth of the Cu(II) only condition.
53
Results
Microarray of Fe(II) shock
Our initial goal was to investigate the effect of high Fe(ll) levels on R.palustris
under anoxic conditions. R.palustris is able to grow using 5 mM Fe(ll) as the sole electron
donor, and would therefore be expected to be resistant to high Fe(ll) levels. To our
surprise, we found that 5 mM Fe(ll) arrested growth for many hours under anaerobic
conditions when added to log phase cultures growing on acetate (Figure 3.1A). To probe
the effect mechanistically, we measured genome-wide transcriptional responses to Fe(ll)
shock using a custom microarray. A number of genes were upregulated after a 15
minute, 5 mM Fe(ll) shock (Figure 3.1B, Table 3.2). Among the genes upregulated after 15
minutes were those predicted to encode three P-type ATPase pumps, six RND-type efflux
pumps, an F1FO ATP synthase, a multicopper oxidase, and a number of regulatory
proteins. Many of the highly upregulated genes in the microarray were annotated as
being involved in copper homeostasis, in addition to many genes of unknown function.
54
2.0 -
control
* 5mM Fe(ll)
0
0
1.5
A
A
A
A.
A
.~
-
AA
A
-'-a-
131.0
~
0
A
AA
*i~
£
A
~
0.5
A
A
0.0
0
20
40
60
hours
80
100
nknown (34)
other (39)
I
ATP synthase
(7, 1 operon)
ribosomal proteins (8)
outer membrane efflux
pumps (6)
sigma/transcription/
longation factors (10)
P-type ATPases (3)
B
Figure 3.1. Effect of Fe(II) and copper on growing R.palustris cells. A) A shock of 5 mM
Fe(II) (260 nM copper) temporarily arrested growth of log phase R.palustris cultures.
Lines show the median of three cultures, with individual data points shown. B)
Categorization of genes up-regulated more than 5-fold in the 15 minute shock with
Fe(ll)/copper microarray.
55
Synergistic effect of Cu(II) and Fe(II)
Our initial experiments and microarray were performed with a 98% pure Fe(II)
stock (Sigma). As a control against contamination, we tested growth of R.palustris with a
more pure stock of iron (99.99% purity, Alfa Aesar). Using this new Fe(II) source, the
effect of Fe(lI) at the concentrations previously tested disappeared. An ICP-MS analysis of
both Fe(II) stocks indicated that copper levels were higher in the 98% pure stock of iron
from Sigma: 5 mM of the 98% pure Fe(II) contained 260 nM copper as measured by ICPMS, while 5 mM of the 99.99% pure Fe(II) contained only 10 nM copper. Our analysis did
not allow us to determine the oxidation state of the copper contamination, although it
seems likely that it would be present as Cu(I), due to the high concentration of Fe(II), as
described below. We hypothesized that the results described above were due not simply
to Fe(II), but to the combination of Fe(ll) and copper. To test this possibility, we set up
experiments with pure Fe(II) and Cu(Il) in varying concentrations to explore the possible
effects of iron and copper combined.
We found iron and copper alone have very little effect even at 1 mM and 1 pM
respectively, but they act synergistically to delay bacterial growth at 100 pM Fe(II) and
100 nM Cu(ll) (Figure 3.2A). The effect decreased with the concentration of each metal,
with a slight effect seen at 50 nM Cu and 10 pM Fe. We tested other metals as well
(Figure 3.2B-C): 100 nM Cu(II) increased toxicity of 50 pM Ni(II) and 500 pM Co(ll), while
100 pM Fe(II) had no effect on Co(II) and decreased the toxic effect of Ni(II). We also
tested Mn(II) with Cu(Il): 500 pM Mn(II) had no effect on growth, either with or without
100 nM Cu(II) (Figure 3.2D).
To further determine the nature of the iron/copper synergistic growth inhibition,
we performed colony-forming-unit (cfu) counts after inoculating cultures to determine
whether cells were dying or simply arrested in their growth. We found that cells were not
killed by the metals: 30 minutes after inoculation into media containing 100 nM Cu(II)
plus 100 pM Fe(II), cfu counts were unchanged relative to control cultures: triplicate cfu
56
counts for metal treated cultures ranged from ~2 to 3 x 106/ml, while control cultures
had counts ranging from ~2 to 4 x 106/ml.
57
1.5
no metal
-1
pM cu(II)1000 pM Fe(ll)
-0.1 pMCui 100 pM Fe
1.0
no metal
50 pM Ni(Il)
50 pM Ni(II) 0.1 pM Cu(Il)
50 pM Ni(II) 100 pM Fe(ll)
0
-
*.
*
0.8
1.0
0.6
0
0
0.5
0.4
*
U'
-.
.
30
10 20
-
0.2
40
B
50
6
houm
A
0.0
0
+no
1.0
40
20
hours
60
0.0 *
0
80
1.5
metal
no metal
500 PM R
00MFe(II) 500pM Co(II)
0.1 pM Cu(Ii) 500 pM Co(IlI)
0.8
-.
1 iM r.
1.0
I,+
5+
0
C0.6
0
0.5
0.4
0.2
1*
D
C
0.0
0
20
40
hours
80
0
20
40
hours
60
80
Figure 3.2. Anaerobic growth curves of R.palustris with Cu(ll) in the presence of Fe(ll),
Co(II), or Ni(ll). All cultures were grown in fresh water-acetate media under light. Lines
show the median of triplicate cultures, shown as individual points. A) Comparison of
Fe(II) and Cu(II) compared to each metal alone. B) Comparison of Ni(II) alone and with
Fe(II) or Cu(lI). 50 pM Ni(II) alone had an inhibitory effect that was increased by Cu(II) and
decreased by Fe(II). C)Effect of Co(ll) alone and with Fe(ll) or Cu(II). Cu(II) increased the
toxic effect of 500 pM Co(ll), while Fe(ll) has no effect.
58
Effect of Cu(lI) and Fe(ii) on other bacterial taxa
To determine whether the synergistic effect of Cu(II) and Fe(II) was unique to R.
palustris, we tested the effect of Cu(II) and Fe(II) on E.coli MG1655, Shewanella
oneidensis MR-1, and Rhodobacter capsulatus SB1003. The physiology of these
organisms has been extensively studied, and their copper resistance systems are at least
partially characterized (12, 32-34). Intesting the sensitivity of these organisms to copper
and iron, we chose varying copper and iron concentrations to reflect the varying
sensitivities to copper of the different bacterial taxa. We found that Cu(ll) and Fe(II)
combined cause a longer growth lag than either metal alone in all three organisms (Figure
3.3). Given that this synergistic effect holds true in four different bacterial species, what
could lead to this synergistic effect?
59
no metal
no metal
+
-n 5
- 1 pM Cu(IlI)
. 1pM Cu, 200 pM Fe
Ci ll
-&0dPM Fe((1)
0.20 - 50 pM Cu(I) 300 pM Fe()
-e
0. 200 pM Fe
*
----
0.15
0.3
e
8
0 0.10
00.2
0.1
.'* *
0.05
0
5
10
'''--7-.B
A
4
-
0.0
15
hours
20
25
0.00
30
5
0
hours
10
1.2
40 pM Fe(II)
0.04 pM Cu(Il
pM
1.0 -0.04 pM Cu
j
j4
I
0.8
0.6
0
'I
I.
I;
~I
A
0.4
0.2
A
-- I
*
0.0
U
0
10
20
hours
30
40
Figure 3.3. Effect of Cu(Il) and Fe(ll) on other bacteria. A) E.coli MG1655, B)S.
oneidensis MR1, and C)R. capsulatus SB1003. Lines show the median of triplicate
cultures, shown as individual points.
60
15
Influence copper oxidation state on the synergistic effect
Fe(lI) is known to reduce Cu(II) to Cu(l) [Fe(ll)+Cu(II) -+ Fe(Ill) +Cu(l)] (35), and we
confirmed that this reaction takes place in our anaerobic freshwater medium (Figure 3.4).
Because Cu(l) is generally considered more toxic than Cu(ll) (15), it seemed likely that
Cu(ll) reduction might be responsible for the synergistic inhibition observed. To test this
hypothesis, we tested the effect of Cu(ll) with ascorbate, which also rapidly reduced Cu(ll)
(Figure 3.4). We found that ascorbate plus Cu(ll) did not inhibit growth as severely as
Fe(lI) plus Cu(lI) (Figure 3.5A). To more directly test the effect of the copper oxidation
state on toxicity, we compared the effect of Cu(l) and Cu(II). We found that Cu(l) and
Cu(II), either alone or in the presence of ascorbate, did not strongly affect growth,
whereas either Cu(l) or Cu(ll) in the presence of Fe(ll) caused a growth lag (Figure 3.5B).
Hence it appears that the synergistic effect cannot be attributed exclusively to the
reduction of Cu(ll) by Fe(ll).
61
20 -. Fe(ll
- ascorbate
0
15
...............................
010
5
0
0
5
10
15
seconds
20
25
30
Figure 3.4. Abiotic reduction of Cu(Il) by Fe(ll) and ascorbate. 30 PM Cu(II) was added to
fresh water medium under anoxic conditions. 100 pM of either Fe(ll) or ascorbate was
added to the mix, and samples were taken at appropriate intervals and tested for Cu(I)
accumulation with the bathocuproine assay as described in the methods. Lines depict the
averages of two experiments, shown as individual points.
62
1.5
-control
100 pM ascorbate 0.1 pM Cu(lI)
+ 100 pM Fe(II) 0.1 pM Cu(lI)
t
100 pM ascorbate
-
1.0
0p
0.5
0.0
A
0
040
-
1.0
60
hours
control
3 0.1 pM Cu(I)
-
+0.1 pM Cu(I) 100 pM ascorbate
-0.1 pM Cu(II) 100 pM ascorbateb.
+ 0.1 pM Cu(I) 100 pM Fe(il)
0.1 pM Cu(II) 100 pM Fe(II)
,
I
0.5
.b/, .
1
B
0.0
0
10
20
30
hours
40
50
Figure 3.5. Effect of Cu(I) vs. Cu(II) on growth of R. palustris.A)Ascorbate plus Cu(II)
causes a slight delay in growth; however, Fe(ll) plus Cu(II) causes a longer delay. B) Cu(l)
and Cu(II) have identical effects: at 100 nM, there is no effect alone, a very slight effect in
the presence of ascorbate, and a significant growth lag in the presence of Fe(II). Lines
show the median of triplicate cultures, shown as individual points.
63
Specificity of gene expression to Cu(II) or Fe(II)
Several genes of interest were upregulated in the microarray, including a number
of genes predicted to be involved in copper homeostasis and a secondary F1Fo ATP
synthase with an unknown function (Table 3.2). To test the metal specificity of these
transcriptional responses, we subjected R.palustris cells to a 15 minute shock using either
1mM Fe(II), 1 IM Cu(II), or 1 mM Fe(ll) + 1 pM Cu(II). We used higher metal
concentrations for the shock than for the growth assay because lower amounts did not
have an effect on mid-log phase cultures, and we wished to be certain we were triggering
a response. We then measured the transcriptional response of eight genes likely to be
involved in copper homeostasis (Figure 3.6): atpD from the upregulated ATP synthase
operon (Rpal_1057), the putative multicopper oxidase (Rpal_1091), the three P-type
ATPases (Rpal_1072, 1856, and 3679), a homolog of E.coli's copper responsive cusR
(Rpal_2141), and a homolog to csoR, a recently discovered copper regulator in
Mycobacterium tuberculosis and Bacillus subtilis (Rpal_1857) (36, 37). We also tested the
response of Rpal_3680 because although it was not highly upregulated in the microarray
(2.7 fold, with a p-value of .07), it belongs to the MerR family of metal-responsive
transcriptional regulators, as does E.coli's copper resistance regulator CueR. All the
genes tested responded to Cu(ll) and not to Fe(II) with the exception of the P-type ATPase
Rpal_3679, which did not respond to either metal. Intriguingly, the multicopper oxidase
and the cusR homolog were upregulated to a lesser degree in the presence of Cu(II) plus
Fe(II) than in the presence of Cu(II) alone, while the other genes tested were more
strongly upregulated in the presence of both metals.
64
ofitrol 1mM FeIII
M
1 pM CUIIII
&ooA
II
B
Rpal 1091
(Multicopper oxidasi
-
Rpal 1057
(ATP synthase)
C40C
10
'2200
C
D
Rpal 2141
c
Rpal 1072
(P-type ATPase)
(CusR homoloqi
S
4
40
20
6o
so
E
Rpal 1857
(CsoR homo9ogi
Rpa 3680
(CueR homolog)
0
530
Q2
920
8C
GRpal
1856
2
(P-type Atpasei
H
4C
EV
C
-
Apal 3679
(P-type Atpase)
L
V4
mm=PAE
-.
Figure 3.6. Q-RT-PCR showing fold change after 15 minute shock (T15/TO) by Cu(ll) and
Cu(Il) + Fe(lt). A) Rpal_1057, an AtpD homolog; B) Rpal_1091, a putative multicopper
oxidase; C)Rpal_2141, a CusR homolog; D) Rpal_1072, the P-type ATPase with the
greatest similarity to the copper efflux pumps, E) Rpal_1857, the CsoR homolog; F)
Rpal_3680, the CueR homolog; and G,H) Rpal_1856 and 3679, both P-type ATPases. Bars
show the averages of triplicate cultures, shown as individual points.
65
Synergistic effect in copper homeostasis deficient E. coli strain
Based on our expression results, we speculated that Fe(II) might interfere with
one or more proteins in the copper homeostasis systems. To test this prediction, we used
E.coli as a model organism because the copper detoxification system is well understood.
We therefore examined the synergistic effect in E. coli strain LEM59 (JImlay, unpublished
strain) which has the genotype copA::kan AcueO AcusCFBA::cm. Without CopA (a P-type
ATPase copper efflux pump), CueO (a multicopper oxidase), and Cus (an RND efflux
pump), this strain is much more sensitive to copper than its wild-type parent (strain
MG1655* from J.Imlay; note that this version of the strain has a different history than
that of MG1655 shown in Figure 3.3, so they are not directly comparable). We grew the
parent strain with 1 IM Cu(II) with and without 200 pM Fe(II), and the mutant strain with
300 nM Cu(ll) with and without 200 pM Fe(II). The different concentrations of Cu(II)
reflected the different copper sensitivities of the parent and mutant strains: we chose
Cu(II) concentrations for which each strain showed a reproducible response. We found
that when the parent and mutant strains were assayed side by side, the mutant was less
affected by Cu(II) with Fe(II) than the wild-type (Figure 3.7, Table 3.3): while Cu(II) caused
a longer growth lag in the mutant than in the wild-type, the addition of Fe(II) increased
the growth delay only 2.8-fold in the mutant strain but 23-fold in the parent strain.
66
0.6
wt control
+
LEM59
Swt 1pM Cu
0.5
+ LEM59 0.3pM Cu(ll)
-wt 1 pM Cu 200 pM Fe - LEM59 0.3 pM Cu(lI) 200 pM Fe(Il)
0
0.4
a
@5
0
8
C0.3
*
0
%
.
0.2
0.
0.1
10
0.0
0
30
hours
4
50
Figure 3.7. Growth curves of the parent and the mutant strain LEM59 in the presence of
metals. As expected LEM59 is more sensitive to Cu(II); however, the synergistic effect of
Fe(II) and Cu(II) is less in the LEM59 than in MG1655*. Although the absolute length of
the lag phase in the presence of Fe(ll) and Cu(II) varied widely between experiments, this
trend remained the same over 3 independent experiments.
67
Discussion
It iswell known that different toxic metals can exhibit additive or synergistic toxic
effects on various organisms under oxic conditions (16). Our results show that under
anoxic conditions, Cu(ll) and Fe(II) interact synergistically to delay the growth of diverse
bacteria, including R.palustris. To our knowledge, this isthe first report of an anoxic,
synergistic toxicity effect between Fe(II) and Cu(II) at environmentally relevant
concentrations.
Our comparative experiments with other metals reveal that while Cu(II) increases
the inhibitory effect of Co(ll) and Ni(II) on R.palustris, Fe(II) does not alter the effect of
Co(II) and actually decreases the inhibitory effect of Ni(II) (Figure 3.2). Copper's
enhancement of cobalt and nickel's negative impact is not surprising - such interactions
between toxic metals have been observed before (16, 17). Iron's mitigation of nickel
toxicity is also predictable: it supports previous research showing that nickel disrupts iron
containing enzymes and increases expression of the iron uptake machinery in E.coli (38,
39). Cu(ll), like Ni(II), is known to interfere with iron containing enzymes (15, 40), which
makes our finding that Fe(II) and Cu(II) interact synergistically to inhibit growth
particularly interesting.
We hypothesized that one mechanism underlying this inhibitory synergistic effect
might be reduction of Cu(ll) by Fe(II). Fe(II) has been shown to reduce Cu(II) under
environmentally relevant conditions (35). Our own experiments showed that in our
growth medium, Fe(II) could quickly reduce a significant amount of Cu(II) to Cu(I)
abiotically (Figure 3.4). We tested copper reduction as a possible cause of the inhibitory
effect by using ascorbate as an alternate reductant to Fe(ll): when added at the same
concentrations as Fe(Il), ascorbate also rapidly reduced Cu(ll), but resulted in only a slight
delay in growth compared to Fe(II). Moreover, Cu(I) had the same effect on R.palustris as
Cu(lI) either alone, with ascorbate, or with Fe(II). If the reduction of Cu(II) by Fe(Il) were
the sole cause of toxicity, then we would expect Cu(I) to have greater effect than Cu(II).
68
Together, these results suggest that Cu(lI) reduction to Cu(I) is not sufficient to explain
the observed growth inhibition.
What then might underpin the synergistic effect? Our microarray provides a
snapshot of the genes that are upregulated after a shock of Fe(ll) and Cu, which hint at
alternative mechanisms. A putative operon upregulated in the microarray was
Rpal_1089-Rpal_1093, which includes a gene predicted to encode a multicopper oxidase.
Multicopper oxidases are a common component of copper resistance systems in bacteria,
although they are generally considered to be useful only in the presence of oxygen (41).
Based on our qRT-PCR results the multicopper oxidase responds to Cu(II) and not Fe(II);
however, the upregulation by Fe(II) plus Cu(ll) was less than the upregulation by Cu(II)
alone (Figure 3.5B). This trend also held true for the CusR homolog, though not for the
other genes tested. While it is unclear whether this difference in RNA levels would
translate into phenotypic differences, it suggests that Fe(II) may interfere with the
regulation of some of the copper resistance genes, which could explain the inhibitory
effect we observe. It is also possible that iron may inhibit other portions of the copper
homeostasis system - for example, inhibition of a copper efflux pump or chaperone
would also explain the effect we observe.
To test whether the synergistic effect of iron might come from interference with
bacterial copper resistance mechanisms, we used LEM59, an E. coli strain deficient in
CopA, the copper efflux P-type ATPase, Cus, the RND-efflux pump, and CueO, the
multicopper oxidase. These experiments were complicated by the fact that LEM59 is
more sensitive to copper, and needed to be tested at lower Cu(II) concentrations than its
parent strain. Furthermore, the absolute length of the lag phase varied somewhat
between experiments. Despite these issues, a robust trend emerged when the parent and
mutant strain were compared side by side in the same experiment: although the
synergistic effect of Fe(ll) and Cu(ll) in was still evident in LEM59, it was much smaller
than in the parent strain (Table 3.3). This data supports our hypothesis that Fe(ll) might
interfere with one or more components of E.coli's copper homeostasis machinery,
69
although this is clearly not the only cause of the synergistic effect. More work is needed
to determine which proteins are affected, how, and whether other bacterial species
demonstrate the same effects.
Regardless of the mechanisms responsible for the synergistic inhibitory effect, our
microarray suggests that the regulation of genes involved in copper homeostasis may be
more complicated in R.palustris than other studied Proteobacteria. This contention
springs from our measurements of the transcriptional response of three genes likely to be
involved in copper homeostasis in the presence of different combinations of Fe(II) and
Cu(II): homologs of cueR, cusR and csoR. In E.coli, CueR regulates a P-type ATPase metal
efflux pump and a multicopper oxidase. CusR regulates the transcription of a protondriven RND efflux pump. Although not previously reported to regulate the copper
response in Proteobacteria, CsoR is a copper resistance regulator in M. tuberculosis and B.
subtilis (36, 37). Interestingly, in R.palustris, all three putative regulators are upregulated
by Cu(II) but not by Fe(II). This suggests that CueR, CusR and CsoR homologs may help
regulate copper homeostasis in R.palustris. The response of R.palustris's CueR homolog
to copper is surprising because it lacks several of the conserved motifs thought to be
important for determining copper specificity: proteins binding to the monocations Cu*,
Ag*, and Au* share a SXXV[K/R] signature and a CXGXXXXDCP metal-binding loop (42). In
R.palustris's CueR homolog, neither of these motifs is fully conserved. More work is
needed to test whether CueR, CusR and CsoR regulate copper homeostasis genes in R.
palustris.
Among the genes likely controlled by the regulators described above are the Ptype ATPases. Of the P-type ATPases upregulated in our microarray, Rpal_1072 and
Rpal_1856 both belong to the copper specific protein family TIGR01511 and share
conserved residues in transmembrane helices 5, 6, 7, and 8 with CopA proteins (43). It is
thus not surprising that they respond to copper and not to iron. Rpal_3679, on the other
hand, contains an WIY[R,K] motif thought to indicate Zn/Cd specificity in P-type ATPases.
Our qRT-PCR results indicate that it does not respond to copper or iron, and that its
70
upregulation in the microarray was spurious. The fact that it is located next to Rpal_3680
(the upregulated CueR homolog) suggested that it might have some connection to
copper; however, both bioinformatic analysis and qRT-PCR data indicate that this is not
the case, and that R. palustris' copper response system makes use of only two P-type
ATPases.
It is also of interest that a putative ATP synthase operon was upregulated in our
microarray. Quantitative RT-PCR indicates that it responds to copper and not to iron.
Bioinformatic investigation of the putative operon suggests that it does not encode the
primary ATP synthase of R. palustris, but a secondary complex present only in R. palustris
TIE-1, not related strains. The operon structure, as well as the presence of certain
subunits, indicates that it belongs to the family of Na-dependent ATP synthases (44). In
Archaea, these Na-ATP synthases function as the primary manufacturer of ATP. Their
function in Bacteria is not known, though the operon is present in a variety of bacteria.
The large positive transcriptional response of R. palustris'operon to copper raises the
possibility that bacterial Na-ATP synthase may be involved in metal resistance, but this
remains to be experimentally verified.
In conclusion, we have demonstrated that environmentally relevant
concentrations of Fe(II) and Cu(II) can significantly inhibit the growth of microorganisms.
The concentrations of Fe(II) and Cu(II) used in our experiments are similar to those found
in a number of natural and polluted environments. This underscores the importance of
paying attention to trace metals in the design of laboratory studies. Moreover, the
synergistic inhibitory phenomenon described here highlights the need to consider all
metals in an environment when predicting microbial behaviors or planning a
bioremediation system, not simply the conventionally toxic ones.
71
Acknowledgments
We thank Nathan F.Dalleska (Caltech Environmental Analysis Center) for
obtaining the ICP-MS data, Vijaya Kumar and Igor Antoshechkin (Caltech Millard and
Muriel Jacobs Genetics and Genomics Laboratory) for microarray preparation, and Jim
Imlay (UICU) for helpful discussion and for E.coli strains. The Howard Hughes Medical
Institute (HHMI) supported this work and D.K.N. is an HHMI Investigator.
72
Table 3.1. Strains used in this work.
genotype
Wild type
Figures with strain
1, 2, 4, 5
Source
This lab (45)
Wild type
3
Wild type
3
Wild type
3
Haselkorn, Robert
(46)
Myers, Charles R.
(47)
Gross, Carol A.
Wild type
6
Imlay, James A.
Escherichia coli
copA::kan AcueO
6
Imlay, James A.
LEM59
AcusCFBA::cm
Strain
Rhodopseudomonas
palustris TIE-1
Rhodobacter
capsulatus SB1003
Shewanella
oneidensis MR1
Escherichia coli
MG1655
Escherichia coli
MG1655*
unpublished
73
Table 3.2. Genes upregulated more than 5-fold and with corrected p-values <0.05,
following a 15-minute shock with 5 mM Fe(ll) and approximately 260 nM copper.
74
Gene Locus
Annotation
Fold-change
Rpal_1060
Rpal_1062
Rpal_1071
Rpal_1070
Rpal_1057
Rpal_1058
Rpal_1605
Rpal_1064
Rpal_1857
Rpal_1604
Rpal_1056
Rpal_1047
Rpal_1089
Rpal_2016
Rpal_1072
Rpal_2140
Rpal_1055
Rpal_2344
Rpal_1059
Rpal_2017
Rpal_4973
Rpal_0153
Rpal_1091
Rpal_1090
Rpal_1073
Rpal_4604
Rpal_2345
Rpal_1046
Rpal_4085
Rpal_0028
Rpal_1054
Rpal_1856
Rpal_0222
Rpal_3672
Rpal_1247
Rpal_2298
hypothetical protein
putative exported protein of unknown function
hypothetical protein
hypothetical protein
FOFI ATP synthase subunit beta
RNA polymerase sigma factor
putative exported protein of unknown function
putative membrane protein of unknown function
protein of unknown function DUF156
efflux transporter, RND family, MFP subunit
FOF1 ATP synthase subunit epsilon
outer membrane efflux protein
hypothetical protein
RNA polymerase sigma factor
heavy metal translocating P-type ATPase
protease Do
FOF1-ATPase subunit
outer membrane efflux protein
protein of unknown function DUF1109
protein of unknown function DUF1109
efflux transporter, RND family, MFP subunit
GCN5-related N-acetyltransferase
multicopper oxidase type 3
outer membrane efflux protein
hypothetical protein
methionine sulfoxide reductase A
efflux transporter, RND family, MFP subunit
efflux transporter, RND family, MFP subunit
hypothetical protein
phospho-2-dehydro-3-deoxyheptonate aldolase
hypothetical protein
heavy metal translocating P-type ATPase
hypothetical protein
30S ribosomal protein S12
quinolinate synthetase
outer membrane efflux protein
208.7
203.0
156.5
151.7
104.0
71.0
60.5
54.8
43.3
43.0
41.4
40.4
37.0
36.5
33.7
31.0
30.7
28.8
28.3
26.0
25.5
24.8
22.2
20.7
18.4
15.6
15.4
14.9
12.8
12.2
12.0
11.8
10.7
10.6
10.4
9.8
Rpal_2263
Rpal_2814
Rpal_3879
Rpal_1053
Rpal_1603
Rpal_1068
Rpal_2343
Rpal_0242
Rpal_0966
Rpal_3671
Rpal_1052
Rpal_1093
Rpal_1747
Rpal_4459
Rpal_0934
putative exported protein of unknown function
pentapeptide MXKDX repeat protein
hypothetical protein
FOF1 ATP synthase subunit A
heavy metal efflux pump, CzcA family
RNA polymerase sigma factor
hypothetical protein
50S ribosomal protein L19
dihydrodipicolinate synthase
30S ribosomal protein S7
FOF1 ATP synthase subunit C
hypothetical protein
ribulose bisophosphate carboxylase
methionine sulfoxide reductase B
GreA/GreB family elongation factor
9.3
9.0
8.8
8.6
8.4
8.1
7.9
7.1
6.7
6.7
6.5
6.3
6.3
6.3
6.3
two component transcriptional regulator, winged
6.2
Rpal_1051
Rpal_5052
Rpal_4790
Rpal_3619
Rpal_4711
Rpal 2057
Rpal_0612
Rpal_0439
Rpal_1207
Rpal_4666
efflux transporter, RND family, MFP subunit
hypothetical protein
hypothetical protein
hypothetical protein
Integrase catalytic region
heavy metal translocating P-type ATPase
hypothetical protein
hypothetical protein
hypothetical protein
blue (type 1) copper domain protein
oxidoreductase molybdopterin binding
H+transporting two-sector ATPase B/B' subunit
protease Do
phosphoserine aminotransferase
hypothetical protein
hypothetical protein
transcriptional regulator, MucR family
hypothetical protein
ribosome-binding factor A
transcriptional regulator, PadR-like family
Integral membrane protein TerC
6.2
6.0
5.8
5.8
5.8
5.7
5.7
5.5
5.5
5.5
5.4
5.4
5.1
5.1
5.0
5.0
4.8
4.7
4.7
4.6
4.6
Rpal_2769
FMN-binding negative transcriptional regulator
4.6
Rpal_1748
Ribulose-bisphosphate carboxylase
4.5
Rpal 2141
-_
Rpal_2299
Rpal_2992
Rpal_1662
Rpal_4684
Rpal_1039
Rpal_3679
Rpal_3083
Rpal_4799
Rpal_0936
Rpal_1092
Rpal_0223
_
helix family
75
Rpal_2286
Rpal_0038
Rpal_4606
Rpal_0938
Rpal_3880
Rpal_3240
Rpal_3487
Rpal_1069
Rpal_1574
Rpal_2294
Rpal_2242
76
cation diffusion facilitator family transporter
phenylalanyl-tRNA synthetase, alpha subunit
transcription elongation factor GreA
Ornithine decarboxylase
hypothetical protein
hypothetical protein
30S ribosomal protein S6
protein of unknown function DUF1109
sulfatase
flagellar basal body rod protein
acetolactate synthase 3 catalytic subunit
4.5
4.4
4.3
4.3
4.3
4.3
4.2
4.2
4.2
4.1
4.0
Table 3.3. Comparison of the growth delays of the metal treated cultures from Figure
3.7. The lag phase for each of the experimental conditions was subtracted to obtain the
extent of the delay in growth caused by Cu(II) alone or Cu(II) and Fe(ll) combined. The
synergistic effect was measured by taking the ratio of the delay of Cu(II) with Fe(ll) to
Cu(II) alone. Due to the difference in copper sensitivity, the parent strain was assayed
with 1 pM Cu(II), and LEM59 was assayed with 0.3 pM Cu(II). The delay times are
averages of 3 biological replicates, and are representative of 3 independent trials.
Parent strain
LEM59
Lag of Cu(II)
Lag of Cu(II) with Fe(ll)
Cu(ll) with Fe(II) delay
- Lag of
- Lag of control
/Cu(II) delay
control
1.4
9.7
33.2
27.6
23.4
2.8
77
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82
Chapter 4: Non-redundant roles for cytochrome c2
and two HiPIPs in the photoferrotroph
Rhodopseudomonas palustrisTIE-1
Lina J BirdE, Ivo H SaraivaE, Shannon Park, Eduardo Calgada, Carlos A. Salgueiro, Wolfgang
Nitschke, Ricardo 0 Louro§ and Dianne K Newman§
Eco-first authors
§co-corresponding authors
This chapter will be submitted to Journal of Bacteriology
Contributions:
I collected the data for and generated figures 4.1,4.2, and 4.8. I collected the data for
figure 4.7 in collaboration with Dr. Nitschke. The text was written in collaboration with Ivo
Saraiva.
83
Abstract
The purple bacterium Rhodopseudomonas palustris TIE-1 expresses multiple small
high potential redox proteins during photoautotrophic growth, including two HiPIPs (PioC
and Rpal_4085) and a cytochrome c2. We have evaluated the role of these proteins in TIE1through physiological and biochemical analysis. A deletion mutant of cytochrome c2 was
found to be unable to perform photosynthesis indicating that this protein cannot be
replaced by either HiPIP in cyclic electron flow. PioC was previously implicated in
photoferrotrophy, an unusual form of photosynthesis in which reducing power is
provided through ferrous iron oxidation. Using cyclic voltammetry (CV), electron
paramagnetic resonance (EPR), and flash induced spectrometry techniques we show that
PioC has a midpoint potential of 450 mV, contains all the typical features of a HiPIP, and
can reduce the reaction centers of an R.palustris TIE-1 membrane suspension in a light
dependent manner at a much lower rate than cytochrome c2. This data supports the
hypothesis that PioC's role is linear electron transfer from iron, while cytochrome c2 is
required for cyclic electron flow. Rpal_4085, despite having similar spectroscopic
characteristics and reduction potential to PioC, is unable to reduce the reaction center.
Rpal_4085 is upregulated by divalent metals, indicating that it might have a role in
mediation of metal toxicity.
Introduction
The small soluble cytochromes c and high potential iron proteins (HiPIPs) are both
small redox active proteins that have long been known to act as donors to the reaction
centers (RCs) of anoxygenic phototrophs (1). The HiPIPs contain a [4Fe-4S] cubane cluster
coordinated by four cysteine residues, and have reduction potentials that range from 50
to 500 mV (2). They occur in a wide variety of organisms, both phototrophic and nonphototrophic. The small soluble cytochromes used in photosynthesis (most often c2)
contain a single c-type heme and have reduction potentials ranging from 150 to 380 mV
(3).
84
A survey of photosynthetic bacteria (4) shows that many contain either a HiPIP or
a small cytochrome c, leading to the conclusion that both proteins can serve the same
role: that of an electron shuttle between the bc1 complex and the reaction center during
photosynthetic energy generation. Many bacteria contain both small cytochromes and
HiPIPs, sometimes multiple copies of each, a redundancy that has yet to be fully
explained. Biophysical studies have shown that, in at least some of the organisms
containing a HiPIP and a small cytochrome, both are capable of reducing the reaction
center (5-8). This observation has supported the idea that these proteins have
overlapping functions, and are at least partially redundant - although other studies have
shown that in whole cells under a given condition, a single protein is generally the
dominant electron donor (6).
In the phototrophs studied so far, those in which the dominant electron donor is a
HiPIP also contain a tetraheme cytochrome bound to the reaction center (4). The
tetraheme cytochrome reduces the bacterial chlorophyll and is in turn reduced by a
soluble donor. All biophysical studies on photosynthetic HiPIPs in wild type bacteria show
that the HiPIP interacts with the tetraheme cytochrome rather than the
bacteriochlorophyll (5-7, 9, 10). However, one study in a Rubrivivax gelatinosus mutant
lacking the tetraheme cytochrome showed that the HiPIP can still donate to the reaction
center, though at a reduced rate (11). In addition, there are a number of photosynthetic
bacteria that contain HiPIPs in their genome but lack the tetraheme cytochrome.
Precisely how these bacteria utilize their HiPIPs is an interesting question.
Rhodopseudomonas palustris is an example of this last category. It is a purple
non-sulfur phototroph that has both types of electron shuttles: its genome encodes two
HiPIPS and two cytochromes c2. Its genome has been fully sequenced, and does not
contain the tetraheme cytochrome. R. palustris is also genetically tractable, which makes
it an excellent model in which to study the roles of multiple small redox active proteins in
a single organism.
85
A metabolically versatile bacterium, R.palustris is capable of growing aerobically
using various sources of energy and anaerobically performing different forms of
anoxygenic photosynthesis. Some strains, in particular R.palustris TIE-1, can grow by
oxidizing ferrous iron photosynthetically (i.e. photoferrotrophy) (12). Previous studies
have shown that one of TIE-1's HiPIPs, PioC, is required for photoferrotrophic growth
(13). Interestingly, although the ApioC mutant could not grow on iron, it was still able to
oxidize Fe(II), though at a slower rate than the wild type strain. The cause of this residual
oxidation activity was hypothesized to be the second HiPIP, Rpal_4085. The ApioC
mutant was also not impaired for other forms of photosynthetic growth, indicating that it
plays little, if any, part in cyclic electron transfer.
Although genetic studies indicated that PioC has a specialized role in Fe(II)
oxidation, questions about the function and mechanism of the HiPIPs in R. palustris
remain. Do the HiPIPs donate to the reaction center, even without a tetraheme subunit?
Can they substitute for each other, or for the cytochrome c2? What is the primary role of
Rpal_4085, the second HiPIP? We undertook to answer these questions through a
combination of physiological and biochemical techniques.
Experimental procedures
Media and growth conditionsfor R. palustris
R.palustris TIE-1 was grown in minimal medium as previously described (14) with
the following modifications: the KH2 PO4 concentration was reduced to 100 mM and the
NaHCO 3 concentration was increased to 30 mM. For hydrogen growth, the cooled
medium was immediately aliquoted into sealed balch tubes and the headspace was
flushed with 80% hydrogen as the electron donor (balance CO2). For acetate growth, 10
mM acetate was added to anaerobic medium. For routine aerobic growth YPS-MOPS
medium (3 g/L yeast extract, 3 g/L peptone, 10 mM succinate, 50 mM MOPS, pH 7) was
used.
86
RNA isolationand q-rtPCR
Samples with an O.D.650nm of 0.3 when 0.2 mL were measured on a Synergy 4 plate
reader from Biotech were used for analysis. 3 mL of culture were added to 3 mL RNALater
(Ambion) in a Coy anaerobic chamber. Samples were centrifuged for 6 minutes at 6,000xg
at room temperature. The supernatant was poured off, and the pellets frozen at -80 *C
until RNA extraction.
Cells were lysed using the lysis, digestion and mechanical disruption protocol from
Qiagen, and RNA purification was performed with the Qiagen RNeasy kit according to the
manufacturer's instructions. Purified samples were treated with Ambion Turbo DNase
using the rigorous treatment to remove contaminating DNA and quantified with a
Nanodrop. 40 ng of RNA was used to make cDNA using the BioRad cDNA kit. 1 pl of the
cDNA reaction was assayed in a 25 pl qPCR reaction using the iTAQ SYBR green supermix
with ROX from BioRad. Standard curves were made using RNA synthesized with the
Ambion MEGAscript Kit. The resulting RNA was diluted to a known concentration and
treated the same way as experimental samples. qPCR was performed with the 7500 fast
real time PCR system from Applied Biosystems. Data from four independent cultures
were analyzed and standard curves generated using Sequence Detection Software
Version 1.4.0.25.
Constructionof deletion and substitutionstrain
Strains used in this study are listed in Table 4.1 and primers are listed in Table 4.2.
To delete Rpal_4085,
1 kb fragments of the upstream and downstream regions were
amplified using primer sets HipipUpF2/HipipUpR and HipipDownF/HipipDownR2. These
fragments were recombined and cloned into pMQ84 using the yeast cloning system (15).
The recombined fragment was cut out of pMQ84 and cloned into the suicide vector
pJQ200sk. This vector was transformed into E. coli S17-1 and mated into R. palustris TIE-1
as previously described (13). The substitution strain was made in a similar fashion: the
upstream, signal sequence, and downstream regions of PioC were amplified with the
87
primer sets PioCmchF1C/PiC-HfusionsigRlb and PioC-HfusionsigF2/PioCHfusionR2, and
the coding region (without the signal sequence) of Rpal 4085 was amplified with the
primer set PioCHfusionF3/PioCmchR3C.
Cell suspension assay
The cell suspension assay was performed as previously described (13). Briefly,
cells were harvested by centrifugation at an O.D. at 650 nm of ~0.3, rinsed anaerobically
in cell suspension buffer (50 mM HEPES, 20 mM NaCl, 20 mM NaHCO 3, pH 7), and
resuspended in cell suspension buffer plus ~600 pM FeCl 2 to an OD 65 0 of ~0.8. 100 pl of
each sample was aliquoted into a clear flat bottomed 96 well plate, and incubated under
a 60 watt light bulb. 50 pl of sample were taken from a new well at each time point and
combined with 50 pl ferrozine. Absorbance was measured at 570 nm after 10 a minute
incubation.
Cloning, expression, and purificationof the HiPIPs
Primers used in this study are shown in Table 4.2. Amplification of pioC was
achieved by PCR with forward primer PioC-N-NcoCap and reverse primer PioC-C-XhoCap
using genomic DNA extracted from aerobically grown TIE-1 as template. The PCR product
was digested with restriction enzymes Ncol and Xhol and ligated into pET32h vector to
generate plasmid pET32hPioC. PioC without its signal sequence was PCR-amplified from
pET32hPioC using forward and reverse primers PioCwithoutTAT and PioC-C-XhoCap,
respectively. The resulting PCR product was cloned into pET32h by using Ncol and Xhol
restriction enzymes.
The same expression protocol was used for the two proteins. PioC and Rpal_4085,
were expressed in E. coli BL21 with a pET32h plasmid. The signal sequences predicted
with TatP (16) were excluded from the resulting construct: thioredoxin - 6xHis - thrombin
cleavage site - HiPIP. Both strains were grown in LB medium at 37 "C.At an O.D. at 600
nm of 1 expression was induced with 0.5 mM of IPTG at 30 *C.After 4 hours cells were
harvested by centrifugation and resuspended in binding buffer (50 mM phosphate buffer,
88
300 mM NaCl, pH 8). Cells were lysed with a Thermo Scientific French-press and cell
debris was removed by centrifugation. The resulting supernatant was loaded into a
Qiagen Ni-NTA agarose column equilibrated with binding buffer. Unbound proteins were
discarded and bound proteins were eluted with elution buffer (50 mM phosphate buffer,
300 mM NaCl, 250 mM imidazole, pH 8). After elution, the bound fraction was incubated
with GE Healthcare thrombin overnight according to the manufacturer protocol. The
enzymatic digest was dialyzed against binding buffer and reloaded into the reequilibrated Ni-NTA column. Target proteins did not bind to the column and were
collected in the flow-through. Purity was evaluated by SDS-PAGE with coomassie blue
staining. N-terminal sequencing showed that the obtained proteins were correctly
digested by thrombin.
Membrane suspension EPR assay
To prepare a suspension of double mutant TIE-1 membranes, cells were grown
anaerobically on minimal acetate medium to early stationary phase. The cell pellets were
resuspended in 5 mL of 50 mM potassium phosphate buffer at pH 6 and passed three
times through a Thermo Scientific French-press. The cell lysate was ultra-centrifuged for
30 min in a Beckman-Coulter TLA-100.3 rotor at 60000 rpm. The supernatant was
discarded and the pelleted membranes were resuspended in 4 mL of the same buffer. 0.2
mL of 0.5 mM HiPIP solution was mixed with 0.1 mL of membrane suspension in a 3 mm
EPR tube. The tubes were incubated for 30 min ~10 cm from a 36 W white fluorescent
lamp or in the dark prior to freezing with liquid N2. HiPIP oxidation was detected by EPR
spectroscopy. The spectra were recorded as described below.
Cyclic voltammetry
Cyclic voltammograms were recorded using an Autolab PSTAT10 potentiostat. The
working electrode was a pyrolytic graphite-edge disc, a platinum wire was used as
counter electrode and a Ag/AgCI (1M KCI) electrode was used as reference. Reduction
89
potentials are reported versus standard hydrogen electrode (SHE) by adding +222 mV to
the experimental readings made at 252 C. Prior to each experiment the working electrode
was washed with Millipore water (18 MO.cm). For each voltammogram 20 pL of HiPIP at
5 pM in 50 mM TrisHCI, 300 mM NaCl, pH 9 buffer were used. The scan rate was 1 mV.s'.
An initial potential of 750 mV was applied to the sample for 10 min before recording of
each voltammogram. Data was processed with SOAS using a noise filter with automatic
threshold and subtraction of the background current (17).
EPR spectroscopy
EPR spectra of pure HiPIPs were recorded on a Bruker EMX spectrometer
equipped with a dual-mode cavity and an Oxford Instruments continuous flow cryostat.
The experimental conditions were: temperature 10 K; microwave frequency 9.66 GHz;
microwave power 6.346 mW; modulation amplitude 0.5 mT; receiver gain 1.00 x 10s.
HiPIP concentration was 200 IIM in 100 mM potassium phosphate buffer, 400 mM KCI, pH
8. Sample oxidation was achieved by addition of potassium ferricyanide.
Membrane fractions were prepared by cell lysis with BugBuster, according to the
manufacturer protocol, followed by ultracentrifugation at 200000 g. The pelleted
membranes were resuspended in 20 mM phosphate buffer, pH 7 and oxidized with
potassium hexachloroiridate(IV). The spectra of membrane fractions and of the
membrane suspension assay were recorded in a Bruker EMX spectrometer equipped with
an ESR-900 continuous-flow helium cryostat. The experimental conditions were:
temperature 13 K; microwave frequency 9.39 GHz; microwave power 2.012 mW;
modulation amplitude 1 mT; receiver gain 5.02 x 104.
Metal and non-metal shocks of TIE-1 cultures
To determine the transcriptional response of Rpal_4085, we grew 8 ml triplicate
cultures anaerobically on acetate to mid log phase and transferred them to an anoxic
chamber. A 4 ml aliquot was added to RNAlater as described above, and the remaining
90
cultures were shocked with either 50 pM NiCI 2, 1 mM CoCl 2 , 5 mM FeCl 2, 300 pM H2 0 2 , or
0.8 M NaCl. All stocks were dissolved in anaerobic water, with the exception of NaCl,
which was added as a solid to avoid large volume changes. The concentrations were
chosen to reflect the various levels of sensitivity TIE-1 growth displays to each compound.
Catalase was added, where indicated, to a concentration of 2900 U/ml
Cytochrome c2 purification
Cytochrome c2 was purified using a modified version of the native purification
protocol described by Bartsch (18). 6 liters of TIE-1 was grown anaerobically on YPSMOPS medium until stationary phase, harvested, washed in 1/10th volume of 100 mM
TRIS buffer pH 8, and resuspended in the same. Cells were lysed using a French press,
and ultracentrifuged for 1.5 hrs. at 100,000xg. The resulting lysate was fractionated by
ammonium sulfate precipitation: a 30% ammonium sulfate was added, and the lysate
stirred for 10 minutes at 40 C,then centrifuged for 20 minutes at 25,000 x g. Additional
ammonium sulfate was added to the supernatant to the final concentration of 90%. The
lysate was again stirred and centrifuged, and the resulting pellet was resuspended in 10
mM Tris pH 8. The sample was then desalted using a HiPrep 26/10 desalting column (GE
Healthcare) into 1 mM Tris pH 8. The desalted sample was then passed through DEAE
sephacel resin (GE Healthcare) using a low pressure BioRad pump. The flow through,
which contained cytochrome c2, was adjusted to pH 5.5 by slow addition of 1 M acetic
acid, then passed through CM sepharose FastFlow resin equilibrated with 1 mM
phosphate buffer pH 6. The column was rinsed with 10 mM phosphate buffer pH 6, and
the protein was eluted with 20 mM Na-phosphate, pH 6. The eluate was diluted to 10
mM phosphate, and loaded on a prepacked 1 ml CaptoS FastFlow column using an ACTA
purifier FPLC (GE). The column was rinsed with 10 mM Na-phosphate, and eluted with
100 mM phosphate buffer, pH 6. The eluate was concentrated to 500 pI using a 3 kD spin
column, and run on a gel filtration column as a final polishing step. The final protein was
checked for purity by UV-Vis spectra and coomasie gel.
91
Electron transfer of purifiedproteins in membrane fragments
Membrane fragments for reaction center reduction studies were prepared from
acetate grown cultures. Late log phase cells were lysed by French press, spun at low
speed to remove unbroken cells, and ultracentrifuged for 1 hr. at 150,000 g. Membranes
were rinsed once in 50 mM MOPS pH 7, and resuspended in the same.
Membrane suspensions were added to a quartz cuvette, and measured on a Joliot
type Jts-10 flash induced spectrometer (Bio Logic). Membrane protein interactions were
measured with 0.5 mM ascorbate and 9 pM myxothiazol to provide a reducing
environment and inhibit the bc 1 complex, respectively.
Results
Mutant analysis and qPCR
Cytochrome c2 is a common periplasmic electron shuttle between the cytochrome
bc2 complex and the reaction center, however, in some organisms HiPIPs also assume this
role (6). Given that TIE-1 has two HiPIPs, we wanted to determine whether cytochrome
c2 is required for photosynthetic growth. Accordingly, we made a clean deletion of cyc2
and tested the resulting mutant strain for phototrophic growth on acetate or hydrogen.
Acyc2 could not grow under these conditions, demonstrating that cytochrome c2 is
essential for photosynthesis in TIE-i and that it cannot be replaced by either HiPIP (Figure
4.1).
We next sought to determine whether the HiPIPs could substitute for each other.
Previous work has shown that although TIE-1 ApioC is unable to grow using Fe(II) as an
electron donor, it still has some Fe(II) oxidation activity (7). We hypothesized that the
residual Fe(II) oxidation activity was due to the presence of another HiPIP (Rpal_4085) in
ApioC. To test this hypothesis, we deleted Rpal 4085 in the ApioC background. To our
surprise, the ApioCARpal 4085 mutant was able to oxidize Fe(II) at the same rate as
ApioC (Figure 4.2), indicating that Rpal_4085 was not substituting in the pathway at all.
We next set out to determine why, given that both proteins are small redox active
92
proteins in the same family, Rpal_4085 could not at least partially substitute for PioC in
the iron oxidation pathway. In order to determine whether the lack of iron oxidation
activity attributable to Rpal_4085 was due to a lack of protein expression levels and/or a
difference in localization between the two HiPIPs, we replaced the coding region of pioC
with the coding region of Rpal 4085, leaving the TAT signal sequence of pioC intact. This
strain (pioC -> Rpal 4085) oxidized Fe(II) at the same rate as ApioC, as shown in Figure
4.2.
Next, we used qPCR to determine whether pioC -
Rpal_4085 was properly
expressing the Rpal 4085 gene. The results (Figure 4.3A) show that in the wild type
strain, Rpal 4085 expression is several orders of magnitude lower than pioC. In the
replacement strain pioC -> Rpal 4085, the expression of Rpal 4085 is increased to within
the range of pioC expression in TIE-1. Translation and proper folding of both HiPIPs were
confirmed by the presence of characteristic signals in the EPR spectra of membranes of
both strains (Figure 4.3B).
Given these results, we conclude that Rpal_4085 cannot substitute for PioC during
iron oxidation in wild type TIE-1. This raised the question: what is the biochemical basis of
Rpal_4085's inability to substitute for PioC?
As a step towards answering this question,
we measured the ability of the two HiPIPs to donate electrons to the reaction center and
characterized their electronic properties.
93
I
/
1.0
/
-
Acetate WT
C0
(0
-
Hydrogen WT
0
- -
Acetate Acyc 2
Hydrogen Acyc 2
-
0.5
/
/
/
/
/
0.0
__-
6
50
-U
-
-U
10
Hours
Figure 4.1. WT TIE-i and Acyc2 growing photoheterotrophically with acetate and
photoautotrophically with hydrogen. While WT TIE-1 was able to grow under both
conditions, Acyc2 was not. The lines indicate the median values of 3 independent
biological cultures, and the error bars represent the data range.
94
2.0
.I
1.5
I
.1
-TIE-1
--pioC
-- ploCrpal_4085
- -
pioCrpal_4085
.biotic
0.5
0
5
10
15
Hours
Figure 4.2. Fe(ll) oxidation rates of TIE-1 and mutant strains. Lines are the medians of 4
independent biological cultures, while the error bars show the range.
95
A
1e+07 -
-I--
I
E
-
0
0-L1e+030
le+01 -
strain
wild type
gene
rpal 4085
pioC- rpal_4085
rpal_4085
wild type
pioC
B
2.12
2.08
ARpal 4085
320
310
2.12
I
V30
340
0
2.08
I
pioC -+ Rpal_4085
310
320
3
0
0
Figure 4.3. Rpal_4085 and pioC transcription and localization in strains TIE-i and
pioC-+Rpal_4085. (A)QRT-PCR measurements of expression for the different strains.
Bars are averages of the same 4 biological replicates used to measure Fe(ll) oxidation
rates in Figure 4.2. Points show individual samples. (B)X-Band (9.39 GHz) EPR data
showing the presence of HiPIPs in membrane fragments of wild type and
pioC-+Rpal 4085. The peaks marked at g-values 2.12 and 2.0 are unique to HiPIP
proteins, and are only visible when the HiPIPs are in the oxidized form.
96
HiPiP purification
First, we expressed and purified the two HiPIPs from TIE-1 using a fusion with Histagged thioredoxin. Because thioredoxin is placed at the N-terminal, the periplasmic TAT
signal sequence of the HiPIPs was removed in the cloning so the fusion protein would
accumulate in the cytoplasm. This strategy, followed by affinity chromatography and
proteolytic cleavage of thioredoxin, resulted in pure HiPIPs with a yield of 3 mg per liter of
culture.
EPR spectroscopy
Reduced HiPIPs have a diamagnetic ground state. Because only the ground state is
occupied at the temperature of the EPR experiments, both proteins were EPR silent in the
reduced form (not show). In the oxidized form the ground state is paramagnetic and
HiPIPs display a characteristic EPR spectrum (19) (Figure 4.4). The spectra in Figure 4.4
show that the main features of these signals have g- values of 2.12 and 2.03 for PioC and
2.12 and 2.04 for Rpal_4085. Both HiPIPs show minor species with the gma-value at 2.08.
To test the ability of each protein to donate electrons to the reaction center, we
incubated a reduced sample of each protein with a suspension of TIE-1 membranes either
under light or in the dark. As observed through EPR spectroscopy (Figure 4.5), PioC, but
not Rpal_4085, is oxidized by the membrane suspension in a light-dependent manner.
Cyclic voltammetry
The reduction potentials of the two HiPIPs were determined at pH 9 by cyclic
voltammetry using a scan rate of 1 mVs 1 (Figure 4.6). Both proteins show reversible
reactions with midpoint potentials of 450 mV and 470 mV for PioC and Rpal_4085,
respectively. Faster scan rates resulted in irreversible reactions. At pH 7, scans were also
irreversible, possibly due to poor interaction between the HiPIPs and the electrodes used.
97
2.12
310
320
2.08
2.03
I(
350
330 mT
2.12
PioC
2.04
360
Rpal_4085
2.0
310
320
330
mT
4
350
360
Figure 4.4. X-Band (9.66 GHz) EPR spectra of oxidized purified PioC and Rpal_4085. The
g-values (which reflect the environment of the electron detected) of the main features of
the two HiPIPs are indicated. The features shown here are typical of correctly processed
HiPIP proteins. The y-axis is arbitrary, and hence is not shown.
98
dark
310
320
3
J
340
350
360
mT
2.12
310
light
2.03
320
3 0
r350
360
mT
Figure 4.5. X-Band (9.39 GHz) EPR spectra of PioC incubated with a suspension of TIE-1
membranes in the dark (top) or under light (bottom). Because the peaks that denote a
HiPIP are only visible when the protein is oxidized, these results show that membranes in
the presence of light are able to oxidize PioC. This result was not observed for Rpal_4085,
indicating that it does not interact with the reaction center. The y-axis is arbitrary, and
hence is not shown.
99
PioC
250
3
550
625
700
550
625
700
Rpal_4085
250
32!
E/mV (SHE)
Figure 4.6. Cyclic voltammograms of PioC and Rpal_4085. The experiments were
performed at 25 2C with a 1mVs 1 scan rate in 100 mM potassium phosphate buffer, 300
mM NaCl, pH 9. The midpoint potential for PioC was 450 mV, and the midpoint potential
for Rpal_4085 was 480 mV.
100
Flash induced absorbance changes of membranes and proteins
Both PioC and cytochrome c2 increased the rate of reaction center re-reduction in
membranes after an oxidizing flash (Figure 4.7), indicating that both can react with the
reaction center. However, the maximum rate and extent of reduction by cytochrome c2
was much faster then that of PioC. This observation is consistent with earlier findings
that the tetraheme cytochrome is required for rapid electron transfer from HiPIPs to the
reaction center. During these experiments, the purified Rpal_4085 proved to be
degraded and could not be used. Because the EPR experiments with membranes
indicated that it could not be reduced by the reaction center at all, this experiment was
not pursued.
The presence of a biphasic background reduction was noted in the control
experiments with membrane fragments and ascorbate. It is unclear what causes this
background. Addition of PioC also increased the initial Aabsorbance at 435 nm.
Rpal_4085 Transcriptionalresponse to divalent cations
Because Rpal_4085 cannot be oxidized by the reaction center, we looked for
insight into its function by investigating its transcriptional response to several
environmental stresses. In a previously performed microarray (Chapter 3), Rpal_4085 was
upregulated by an iron shock, while PioC was not. We therefore hypothesized that it
might be involved in mitigating metal toxicity. Using qPCR, we tested the response of
Rpal_4085 under anoxic conditions to the cations Mn(II), Fe(II), Co(II), Ni(II), Cu(II), Zn(II),
Ca(II), and the oxidative stress inducer H2 0 2. Co(II), Ni(II), Cu(lI) have inhibitory effects on
TIE-1, (Chapter 3). With the exception of Zinc and Calcium, these divalent metals are
known to be redox active with the ability to produce oxygen radicals within the cell (2024). In order to determine whether oxygen contamination was leading to production of
oxygen radicals by these metals in our experiment, we used H20 2 as an oxidative stress
inducing control. and Zn(II) and Ca(lI) were included as non-redox active controls to
determine whether the bacterium was responding to some other property of divalent
101
metals. We found that in TIE-1 cultures grown on acetate, Rpal_4085 was highly
upregulated in response to a shock with Mn(II), Fe(II), Co(II), and Ni(II), but not to Zn(II) or
Ca(II). Cu(II) and H2 0 2 produced a only relatively minor upregulation that was not fully
reproducible between experiments (Figure 4.8).
102
2500
2000
E
-0
nM c2
50nM c2
101500
200nM c2
-- 400 nM c2
--
---
4800 nMcQ
4400 nM c2
4800nMc2
-
500
0A
0
50
100
150
milliseconds
200
3000
M2000
-0 nM
----50 nM PioC
200 nM PioC
-- 300 nM PIoC
- 400 nM PioC
-500 nM PioC
--- ---
21000
B
0
0
50
100
milliseconds
150
200
Figure 4.7. re-reduction of the reaction center in membrane fragments. A) Titration with
purified cytochrome c2 of membrane fragments and 2 mM ascorbate. B) Titration with
purified PioC of membrane fragments and 0.5 mM ascorbate. Cytochrome c2 reduces the
photooxidized reaction center far more rapidly and to a greater extent than PioC does.
The absorbance is measured at 435 nm, in which a positive change indicates oxidation of
the reaction center and a return to zero indicates the reaction center's re-reduction.
Membrane fragments were exposed to 3 rapid excitation flashes to ensure complete
photooxidation of the reaction center. Each trace is the average of 9 technical replicates
to reduce machine noise.
103
0
0
so
25
E10
1 LZ
control
j
=
1 mM Ca(II)
1 mM Zn(Il)
1mM F9(11) 1mM MnI)
300 pM
hydrogen peroxide
50 pI NII)
1 mM Co(I1)
2 MMCu(lI)
Figure 4.8. Transcriptional reponse of Rpal_4085 to metals: non-redox active divalent
cations (Zn(II) and Ca(ll)), oxidative stress (hydrogen peroxide), and 7 potentially redox
active divalent cations (Mn(ll), Fe(II), Ni(II), Co(ll) and Cu(ll)). Rpal 4085 responds to
Mn(II), Fe(II), Ni(II), and Co(II), but not to Zn(II) or Ca(II). The response to both Cu(II) an
hydrogen peroxide was minimal. Bars show the median values of triplicate cultures,
shown as individual points, while the fold change is the ratio of RNA levels measured
before and after a 20 minute metal shock, measured by qPCR with an external standard.
104
Discussion and conclusion
R. palustris TIE-1 expresses two HiPIPs (PioC and Rpal_4085) and a cytochrome c2
during growth. Both protein types have been shown to be capable of supporting
phototrophic growth in other species. Despite the fact that all are small redox active
proteins with potentials in the correct range to reduce the reaction center, only one
(Rpal_1724, cytochrome c2) is required for photosynthetic growth in TIE-1, and only PioC
is specifically involved in photoferrotrophy.
Although the two HiPIPs from TIE-1 are different with respect to sequence (35%
identical and 65% similar over 54 amino acids, excluding the TAT sequences), the two
proteins are similar spectroscopically and show the typical EPR patterns of HiPIPs. The
reduction potentials of the two proteins are also very similar, with differences of less than
20 mV. Reduction potential values determined for the RCs of purple bacteria range
between 450 and 500 mV (25). The reduction potentials determined for both HiPIPs are
within the known range of values determined for RCs. Therefore, we tried to recover the
iron oxidation rate of the ApioC mutant by replacing pioC with Rpal 4085 in the TIE-1
genome, leaving the promoter and periplasmic signal sequence of pioC intact. This
ensured the same expression levels and localization as wt. PioC. Our results show that
Rpal_4085 is not responsible for the residual iron oxidation observed in ApioC.
Furthermore, whereas an illuminated suspension of TIE-1 membranes was capable of
oxidizing PioC, the same was not observed for Rpal_4085. These results indicate that
Rpal_4085 is not involved in photoferrotrophy, or any other metabolism that relies on the
reaction center for oxidation.
There are examples of HiPIPs in non-phototrophic organisms that interact with the
cytochrome bc2 complex, such as those in the respiratory chains of Rhodothermus
marinus (26, 27) and Acidithiobacillusferrooxidans (28, 29). It is possible that Rpal_4085
plays a similar role in some type of aerobic metabolism in TIE-1, although ARpal 4085 has
no defect when growing aerobically on acetate and succinate. Another possibility is
105
suggested by the fact that Rpal_4085 is up-regulated anaerobically by Mn(II), Fe(II), Co(Il),
and Ni(II) (Figure 4.8). Although all four of these metals are known to produce reactive
oxygen species in cells to varying degrees (20, 21, 23, 30), indicating that they are redox
active under physiological conditions. Rpal_4085 was not up-regulated in response to the
non-redox active metals Zn(II) and Ca(II), suggesting that reduction/oxidation activity of
the metals is involved in the regulation and/or function of Rpal_4085. In this context, the
low response of Cu(ll) is puzzling: however, the lack of response may be due to the fact
that the relevant redox couple of copper under physiological conditions is Cu(ll)/Cu(I).
Under physiological conditions, therefore, the divalent Cu(II) would not be oxidizable. In
this context, Rpal_4085 may act as a sensor for divalent, redox active metals in the
periplasm. Although HiPIPs have not been described in this role, iron sulfur proteins are
well known to have sensing and regulatory functions in a number of contexts (31),
including iron sensing. It thus seems feasible that Rpal_4085 might play be a sensor for
redox active metals. It is also possible that Rpal_4085 might mitigate metal toxicity by
oxidizing either the metal or some reactive product of intracellular metal activity.
Although this role has not before been shown for a HiPIP, other redox active proteins do
play such a role. One example isthe role that multicopper oxidases play in detoxification
of copper through their ability to oxidize Cu(l) to Cu(II). More research will be needed to
determine the precise role that Rpal_4085 plays in R.palustris TIE-1.
PioC, on the other hand, has a clear role in Fe(II) oxidation. This role was first
suggested by genetic work (13); here, we demonstrate for the first time that PioC is
capable of performing the role assigned to it in our model (Figure 4.9), although the slow
rate and extent of the reaction indicate that the situation may be more complicated in
vivo. Interestingly, with a reduction potential of 450 mV, PioC is nearly isopotential with
the reaction center. The reduction potential of PioC may be even higher than 450 mV
under physiological conditions: research on other HiPIPs suggests that the reduction
potential can increase with decreased pH (32). This result suggests that an intermediate
between PioC and the reaction center would be redundant, and makes direct transfer
106
more likely. Our results with washed membrane fragments indicate that PioC does not
require a soluble partner to be oxidized by the reaction center (Figure 4.5), and
experiments with flash induced absorbance spectroscopy indicate that, at least in vitro,
there is no cytochrome involved in electron transfer from PioC to the reaction center. A
comparison of the in vitro rate of PioC vs. cytochrome c2 reduction of the reaction center
in membrane fragments indicates that PioC is a relatively slow electron donor compared
to cytochrome c2. This is consistent with the fact that organisms utilizing HiPIPs as cyclic
electron donors require a tetraheme cytochrome, and with the observation that deleting
the tretraheme slows electron transfer (11). This slow rate may be the reason for PioC's
inability to substitute for cytochrome c2 in cyclic electron transfer. It also raises the
interesting possibility that slow reaction center reduction by PioC might allow the cell to
maintain a high rate of cyclic electron transfer (for energy generation) relative to electron
acquisition. However, it is important to remember that the experiments described here
reflect in vitro characteristics, and PioC's function in the cell may be entirely different.
The work presented here demonstrates that three soluble electron donors in R.
palustris TIE-1 have distinct and non-overlapping functions in TIE-1. The function of
cytochrome c2 is in cyclic electron transfer for photosynthetic energy generation,
shuttling electrons from the bc1 complex to the reaction center. The two HiPIPs appear
biochemically similar, but only one, PioC, is capable of donating to the reaction center,
and plays a role in photoferrotrophy. The second HiPIP, Rpal_4085, appears to have an
entirely different function, possibly related to metal homeostasis. More work is required
to more fully determine the role of Rpal_4085, and to elucidate the physical differences
between the two HiPIPs.
107
-300
-200 -100
-
0 mV
mv 100 -
Fe2'
PioA?
200 -
bc,
300 400
500
-7
C
4085
PioCRC
Figure 4.9. electron flow chart in R.palustrisTIE-1. The solid arrows indicate the
predicted electron flow. The dashed arrow indicates the ET through the quinone pool
from the light excited reaction center to the cytochrome bc2. The vertical lines indicate
the potential range of known forms. The potential of PioA is not yet determined.
108
Acknowledgeme nts
The N-terminal data was provided by the Analytical Laboratory, Analytical Services
Unit, Instituto de Tecnologia Quimica e Biol6gica, Universidade Nova de Lisboa. We thank
Pablo Gonzales at FCT-UNL for the EPR spectra of pure HiPIPs and Miguel Teixeira at ITQB
for obtaining the other EPR data and for helpful discussions. We thank the Howard
Hughes Medical Institute (HHMI) for supporting this work. DKN is an HHMI Investigator.
This research was supported by the MIT-Portugal Program (MIT-Pt/BS-BB/0014/2008).
This work was also supported by FCT through grant [PEst-OE/EQB/LA0004/2011], and
grant PTDC/BIA-PRO/098158/2008. Rede Nacional de RMN (REDE/1517/RMN/2005) was
supported by POCI 2010 and Fundagio para a Ciencia e a Tecnologia (FCT). I.H.S. is the
recipient of FCT grant SFRH/BD/36582/2007.
109
Table 4.1. Strains used in this work.
Name
Rhodopseudomonas
palustris TIE-1
ApioC
ARpal_4085
Description
Wild type strain
R. palustris TIE-1 ApioC
R. palustris TIE-1
Source
Jiao & Newman
2005
Jiao & Newman
2007
This study
ARpal_4085
ApioC ARpal_4085
R. palustris TIE-1 ApioC
This study
ARpal 4085
pioC -+ Rpal_4085
Replacement of PioC
with Rpal_4085 in a wt.
This study
background
Acyc2
R. palustris TIE-1 with
deletion of cytochrome
C2
110
This study
Table 4.2. Primers used in this work.
Name
PioC-NNcoCap
PioCwithout
TAT
Sequence (5 prime-3 prime)
CATGCCATGGGTATGAACGACA
AACGCAAC
CATGCCATGGGTAACGCCCAGG
TCACCAAGA
Source
Jiao thesis
CCGCTCGAGTTATGCCTTGCCG
GCGTAGA
GGACTAGTACGAACAGAGCACC
CACAG
Jiao thesis
CAGAGTGTTGCCGAAAATTCAG
AGTCCCCCCTTTCGT
This study
GAAAGGGGGGACTCTGAATTTT
CGGCAACACTCTGC
This study
GGACTAGTGATCACCGGCTTGA
CCTG
This study
Binds upstream of pioC,
with overhang for
ATTTAATCTGTATCAGGCTGAA
AATCTTCTCTACTAGTCGGCATC
This study
pMQ84
GACATCACCT
PiCHfusionsigR
lb
PioCHfusionsigF
2
Binds at pioC signal
sequence with overhang
for Rpal 4085
Binds Rpal 4085 with
overhang for pioC signal
sequence
CGGCGTGGCTGCGGCGCAGCC
GGCCGCCGGGTGTTCGGCGGT
GATCCCGCGA
GCGATGCTCGGCGCCGGCGTG
GCTGCGGCGCAGCCGGCCGCC
GGGTGTTC
This study
PioCHfusion
R2
Binds Rpal 4085 with
overhang for pioC
downstream region
Binds pioC downstream
region with overhang
for Rpal 4085
Binds down stream of
pioC with overhang for
pMQ84
GCGCCGCTCGTCCGTGCGCAGC
AGATGACGTTACAGCCGGATCG
GCCCGTC
GCTGCGAGCCCGACGGGCCGA
TCCGGCTGTAACGTCATCTGCT
GCGCACG
CTGATTCTGTGGATAACCGTATT
ACCGCCTTTACTAGTGGCGGTG
CTGATGCTC
This study
PioC-CXhoCap
HipipUpF2
HipipUpR
HipipDownF
Description
Forward primer in
cloning pioC
Forward primer in
cloning pioC without
signal sequence
Reverse primer for
cloning pioC
Upstream F primer for
making Rpal 4085
deletion
Upstream R primer for
making Rpal 4085
deletion
Downstream F primer
for making Rpal 4085
This study
This study
deletion
HipipDown
R2
Downstream R primer
for making Rpal 4085
deletion
PioCmchFl
C
PioCHfusion
F3
PioCmchR3
C
This study
This Study
This study
111
Cyc2upF
Cyc2upR
Cyc2downF
Cyc2downR
112
Forward upstream
primer for deleting cyc2
Reverse upstream
GGACTAGTGTCGCAATCTTCGT
TGGTC
CGACTGTTTGTCGTGTTTGGGA
primer for deleting cyc2
AGCTCTCTGC
Forward downstream
CTTCCCAAACACGACAAACAGT
primer for deleting cyc2
CGCATCCAAA
Reverse downstream
GGACTAGTGATGAACGAGGAC
primer for deleting cyc2
GCTGAG
This study
This study
This study
This study
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genetically tractable photoautotrophic Fe(Il)-oxidizing bacterium, Rhodopseudomonas
palustris strain TIE-1. Appl Environ Microbiol. 71(8):4487-4496.
13. Jiao Y, Newman DK. 2007. The pio operon is essential for phototrophic Fe(II)
oxidation in Rhodopseudomonas palustris TIE-1. JBacteriol. 189(5):1765-1773.
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116
Chapter 5: Visualizing photosynthesis in whole cells
Lina J Bird, Wolfgang Nitschke, Dianne K Newman
This chapter will be further developed and submitted to Biochimica et Biophysica Acta.
Contributions:
All the experiments in this work were done in collaboration with Dr. Nitschke in Marseille,
France.
I wrote the text and generated the figures therein.
117
Abstract
This chapter describes the visualization of whole TIE-1 cells with and without
ferrous iron using flash induced absorbance spectrometry. By recording the difference
spectra at specific wavelengths, we were able to follow the light induced oxidation and
re-reduction of the reaction center and the c-type cytochromes, as well changes in
electrochemical potential across the membrane. This allowed us to distinguish between
cultures in which the cell's quinone pool was mostly reduced and those in which it was
partially oxidized. The effect of Fe(II) on cells with an oxidized quinone pool was
surprising: cultures incubated with Fe(II) became reduced independently of the Pio
pathway, and the reduction was blocked by the bci inhibitor antimycin. Based on these
results, we propose a new model for electron transfer from Fe(II), in which electrons
travel through the bc 1 complex instead of the reaction center.
Introduction
The anoxygenic photosynthesis performed by purple bacteria is of interest both as
a stepping-stone to understanding the more complex oxygenic photosynthesis and as an
interesting metabolism in its own right. Under anaerobic conditions, purple
photosynthetic bacteria generate energy through cyclic electron flow: electrons energized
by light at the photosynthetic reaction center travel through the electron transport chain
to the bc1 complex and are returned to the reaction center via a soluble electron carrier,
such as cytochrome c2 or a High Potential Iron Protein (HiPIP) (1). Energy is harvested in
this process through proton pumping coupled to the electron transfer, yielding a proton
gradient that drives ATP synthesis through the membrane-bound ATP synthase (2).
The cyclic flow of electrons provides a ready source of energy to the bacteria as
long as light is present; however, to reduce CO2 for biosynthesis, an outside source of
electrons is needed. These electrons can come from a variety of donors, such as organic
compounds, reduced sulfur compounds, and iron. The flow of electrons from these
118
sources, particularly from iron, is poorly understood in comparison with our knowledge of
cyclic electron flow. In the case of iron, this is because relatively few organisms are
capable of phototrophic iron oxidation, and of those, only one thus far discovered is
genetically tractable - Rhodopseudomonas palustris TIE-1 (3).
R.palustris TIE-i is a purple, non-sulfur bacterium able to grow both aerobically
and anaerobically. In the presence of oxygen, it can grow heterotrophically, while under
anoxic conditions it grows both photoheterotrophically on a number of reduced carbon
sources and photoautotrophically on thiosulfate, H2, and Fe(II). Previous work has shown
that three genes, encoded in the Pio operon, are required for photoautotrophic growth
on Fe(II) (4). Genetic homology provided putative identities for these proteins. The first
gene, pioA, encodes a putative decaheme cytochrome c-type protein with homology to
MtrA, a decaheme cytochrome involved in iron reduction in Shewanella sp. The second
gene, pioB, encodes a putative outer membrane porin with some similarity (39% similar
at the amino acid level) to MtrB, the porin found in the iron reduction system of
Shewanella. Finally, pioC encodes a high potential iron protein (HiPIP), a type of protein
that is often found donating to the reaction center in photosynthetic systems. The model
developed based on the predicted identities of these proteins is described in Chapter 2 of
this work: PioA was proposed as the primary iron oxidase, with PioC shuttling electrons to
the reaction center. PioB was postulated to either assist in iron transport or to associate
with PioA at the outer membrane to allow the hemes to contact Fe(II). This latter
possibility is similar to the proposed model of iron reduction in Shewanella oneidensis, in
which decaheme cytochromes associate with an outer membrane porin to transfer
electrons across the outer membrane (5, 6).
Biochemical detail on the Pio mediated pathway was challenging to obtain:
purification of PioA and PioB in particular proved elusive. PioC alone was successfully
characterized and shown to be a typical HiPIP that reacts with TIE-1's reaction center at a
much slower rate than the primary electron shuttle cytochrome c2 (Chapter 4). Although
119
this in vitro result suggested that PioC shuttles electrons to the reaction center, we
sought to validate our model for PioA and PioC function in whole cells.
We chose to examine the function of the reaction center and its donor proteins in
whole cells using flash induced absorbance spectrometry, a technique that has been
successfully used to study the reaction center of a number of photosynthetic organisms
(7-9). This technique depends on the typical differences seen between the absorbance
spectra of several components in the photosynthetic pathway: the bacteriochlorophyll
special pair (Figure 5.1A (10)), the cytochromes c (Figure 5.1B, (11)), and the b hemes
(Figure 5.1C (12)). The technique cannot determine changes in the quinone pool or
HiPIPs, as the absorption changes of these molecules are too small to detect. However,
the activity of these portions of the electron transfer chain can be inferred, as described
below.
Flash induced absorbance spectroscopy utilizes these absorption differences by
photooxidizing the reaction centers with short pulses of bright light and then monitoring
absorbance changes on the timescale of microseconds through seconds using a weaker
detection light at specific wavelengths. By monitoring different wavelengths, we can
measure changes in the oxidation states of the reaction center, cytochromes, and light
harvesting pigments. The main components, along with their absorbance changes, are
summarized in Figure 5.2.
The first observable change isthat of the reaction center: a very rapid increase in
absorbance can be observed at 435 nm when the reaction center is photooxidized. The
absorbance at 435 nm immediately begins dropping again as the reaction center is
reduced by the cytochrome c2. At the same time, the oxidation of c2 as it donates to the
reaction center can be seen at 420 nm, where a large drop in absorbance indicates
cytochrome c oxidation. The re-reduction of cytochrome c2 by cytochrome ci is not
straightforwardly observable at a single wavelength because both proteins absorb in the
420 nm region. However, cytochrome ci reduction is observable as the 420 nm
absorbance return to zero over longer time scales. Cytochrome c1 is re-reduced through
120
the b heme, which manifests as a negative change in absorbance at 435 nm. Finally, the
bc1 complex is re-reduced by the quinone pool (which does not have a detectable
absorbance change), and the photosynthetic system (if left in the dark) returns to
equilibrium.
121
2000
0.2
1500
e
1000500
80.0J
0
-500
AOO
400
425
-01
1VI
I
1
450
475
500
525
Wavelength (nm)
550
575
600
400
450
500
550
600
650
700
Wavelength (nm)
0.010CU
0.005
E
*0
0.000
U
-0.010
400
430
T
-
T
460
490
520
550
580 600
Wavelength (nm)
Figure 5.1: Difference spectra of oxidized vs reduced photosynthetic components. A)
Spectra of photooxidized vs. reduced membrane fragments from R.palustris TIE-1.
Membranes were exposed to a brief pulse of saturating light and absorbance was
measured before the pulse and after 10 microseconds. B) Difference of oxidized and
reduced absorbance spectra for purified cytochrome c2 from R.palustris TIE-1.
Cytochrome c2 was purified in reduced form: full reduction was assured by addition of
ascorbate, and the protein was oxidized by addition of ferricyanide. Oxidation of c2 in
whole cells was followed at 420 nm. C)Dithionite reduced-ascorbate reduced membrane
fragments from TIE-1. Because ascorbate will not reduce the b hemes but dithionite will,
this spectra shows the reduced-oxidized b heme difference. Although the spectrum has a
high noise level caused by other components in the preparation, the b heme peak can be
clearly seen at 432 nm.
122
The energy from the downhill movement of electrons through the electron
transport chain is harvested through proton movement across the membrane to form an
electrical and osmotic gradient. The electrical component of this gradient can be
detected through absorption changes in specific pigments in the light-harvesting antenna
known as carotenoids. This phenomenon, known as electrochromism or the Starks effect,
occurs because light absorption by carotenoids causes a change in energy state from the
ground to the excited state. When the ground state and excited state have different
dipole moments (separation of charges), they can be altered to different extents by the
electric field in and across the membrane. This changes the difference between the
ground and excited states, changing the absorption pattern of the molecule. (13). The
electrochromic response in R.palustris TIE-1 can be detected as an absorbance increase
at 553 nm; first we see a rapid change due to the separation of negative charge by the
reaction, and then a further change isobserved as the membrane potential increases. As
the proton gradient is dissipated through ATP generation and other uses of membrane
potential (such as active membrane transport), this absorbance change returns to zero.
Taken together, these absorbance changes offer a fairly comprehensive view of
the events that occur during photosynthesis. It should also be theoretically possible to
observe electron transfer from Fe(lI) to the reaction center. Cyclic electron flow can be
blocked using bc1 complex inhibitors, and the oxidation and reduction of the reaction
center and of PioA should then be detectable. Because absorbance changes of HiPIPs are
too small to observe in these experiments, the contribution of PioC would be detectable
by reduction of the reaction center that was not matched by cytochrome oxidation i.e.,
electrons reducing the reaction center from a source other than cytochrome c2.
We set out to test the model for iron oxidation in TIE-1 that was described in
Chapter 2 by observing and analyzing the flash induced absorbance changes. Specifically,
we wanted to determine whether electrons flow to the reaction center in the order we
have suggested - from Fe(ll) to PioA to PioC and finally to the reaction center - and
whether there are other factors involved in Fe(II) oxidation.
123
Fe(Il): electrons
Light: energy
bch
I
2
QH 2
-QHQH
QH
H-E
AAbs: 420 nm,
A Abs: 435 nm,
AAbs: 435 nm,
AAbs: 553 nm,
- when oxidized
+ when oxidized
- when oxidized
+ with electric field
bph
caratenoids
RC
NAD-NADH
CO 2 : building
biomass
Figure 5.2: Model of the electron transfer system components and their light induced
absorbance shifts. Colors indicate the wavelength each component is measured at and
the direction of the change. The pigments, noted in blue, are carotenoids that shift
absorbance maxima in response to changes in electric potential across the membrane,
including either the movement of a negative charge from the reaction center or of
protons across the membrane. Abbreviations: bc1 : the bci complex; b: b hemes; c:
cytochromes c (1 or 2); Q: quinones; QH 2: reduced quinones (quinols); bch:
bacteriochlorophyll; bph: bacteriopheophytin; RC: reaction center. Solid arrows indicate
the flow of electrons through the system while dashed arrows indicate the movement of
protons.
124
Materials and methods
Media and growth conditions
Rhodopseudomonas palustris was grown aerobically on YPS-MOPS medium,
containing 3% yeast extract, 3% peptone, 30 mM sodium succinate, and 25 mM MOPS
pH7. For plates, 7.5% bacto-agar was added to the YPS-MOPS liquid medium. For
anaerobic growth, we used minimal freshwater medium, as described in Chapter 1 of this
work. Acetate cultures were supplemented with 10 mM sodium acetate. For growth on
H2 , cooled medium was immediately aliquoted, sealed, and flushed with 80% H2/20%
C0 2.
Cultures were grown under bright light at 370 C. For electron limited growth, H2
flushed tubes were placed upright to limit H2 diffusion.
Flash induced absorbance spectroscopy
Flash induced absorbance spectra were taken with a Joliot-type Jts-10
spectrometer (Bio Logic) with a custom-made light ring of actinic LEDs (800 nm) for
excitation and wavelength filters at 420, 435, and 553 nm for detection. Samples were
placed in a round quartz cuvette with a silicone stopper. The light ring functioned to
apply excitation light to the sample evenly from all sides, ensuring a saturating flash. In
order to preserve anaerobicity, the headspace of the cuvette was flushed with argon and
additions to the cuvette were made with a syringe. Fe(Il)-NTA and acetate were added
from 10 mM stocks dissolved in anaerobic fresh water medium to a final concentration of
100 pM. Antimycin was added from a 1 mM stock solution dissolved in ethanol to a final
concentration of 10 pM.
For each sample, 9 scans were averaged to reduce machine noise, with 10 seconds
between each scan to allow the sample to return to equilibrium. The baseline was
determined by machine software from 6 initial points before the initial flash.
125
Results
The quinone pools of cells growing slowly with H2 were oxidized compared to fast
growing cultures
When TIE-1 cultures were grown under bright light at 370 Cwith tubes placed
upright, they grew significantly more slowly (doubling time of a day or more) than
cultures in tubes placed on their sides at 300 C(doubling time 12 hours). Cultures with
acetate, in contrast, grew rapidly (with a doubling time of under 4 hours) both at 30 C
and at 370 C. Given that a higher temperature and a smaller surface area would reduce
the level and speed at which H2 would dissolve in the media, we assumed that the
difference in the growth rates was due to a limitation of reducing substrate (H2) available
to the bacteria. There was some variation between batches of media: in one batch of
media, hydrogen cultures grew relatively quickly even when placed upright at 370 C,
possibly due to a higher concentration of hydrogen dissolved in the media.
We could reasonable expect cultures with limited access to reductant to be in a
more oxidized state than cultures with a plentiful source of external reductant.
Specifically, if electrons for CO2 fixation are taken from the quinone pool (Figure 5.2),
electron limited cells should have a high ratio of quinone/quinol. Our experimental
results indicate that this is the case: the observations made at each wavelength (420 nm
to measure oxidation of cytochromes c, 435 nm to measure reaction center and b heme
oxidation, and 553 to measure electrochemical gradient) all suggested that slow growing
cells have a higher proportion of oxidized quinone in their quinone pool than those in
more rapidly growing cultures. The changes at each wavelength are shown in Figure 5.3
and the behavior of each component measured is described below.
A) Rate of cytochrome c re-reduction: the absorbance changes at 420 nm
showed that the cytochromes, while they were rapidly oxidized by the reaction center in
all cases, took significantly longer to become re-reduced in slow growing cultures (Figure
5.3A, blue trace) than in fast growing cultures (Figure 5.3A, red and green traces). This is
expected because the electrons used to re-reduce the cytochromes c ultimately come
126
from the quinone pool, and a low abundance of quinols would lower the rate of electron
flow to the cytochromes c.
B) Reaction center absorbance changes: when cells from all three cultures were
exposed to 3 oxidizing flashes in rapid succession and monitored at 435 nm, the third
flash in acetate and fast hydrogen grown cultures induced a significantly smaller
absorbance increase than the first two, while in the slow growing hydrogen cultures all
three flashes yielded roughly the same absorbance increase (Figure 5.3B). Because the
rapid changes at 435 nm are due to the reaction center, this suggests that in fast growing
cells, the third light induced oxidation of the reaction center could not be detected. The
explanation for this phenomenon lies in the oxidation state of the quinone/quinol pool.
During the first two flashes, the quinone bound to the reaction center (QB in Figure 5.2) is
reduced: the electron excited by the first flash reduces it to semiquinone, and the
electron from the second flash converts it to a quinol (14). In order for the reaction center
to become stably oxidized by the
3 rd
flash, the reduced quinol must be replaced by an
oxidized quinone. If the quinone pool contains a significant fraction of oxidized quinone,
this replacement happens rapidly. If the pool consists primarily of reduced quinol
molecules, this replacement happens more slowly than the arrival of the third flash, and
the electron mobilized by this third flash cannot leave the reaction center and rapidly
recombines with the remaining positive charge on the photooxidized bacteriochlorophyll.
This back-reaction is faster than the time resolution of our experiment and manifests
itself as a lack of reaction center oxidation. Because the oxidation state of the quinone
pool is difficult to measure directly, this experiment provided a good indirect indicator of
the quinone/quinol ratio - although it is not quantitative, and the precise ratios could not
be determined because the rate constant for quinone binding to the TIE-1 reaction center
is not known.
C) Lack of b-heme oxidation. The absorbance changes 435 nm over longer time
scales (milliseconds in Figure 5.3) indicate that in slow growing cells (Figure 5.3, blue
trace) there is very little oxidation of the b hemes. This is consistent with an oxidized
127
quinone pool in that if the quinone pool is highly oxidized, the b-hemes will already be
partially oxidized. As electrons flow from the reaction center, the b-hemes will be slowly
reduced and then immediately re-oxidized by cytochrome c.
D) Lack of rapid electrochemical membrane gradient changes: the slow
absorbance change at 553 nm, indicative of a buildup of the electrochemical gradient
across the membrane, was absent in the slow hydrogen grown cultures. In the acetate
grown cultures, this change was rapid, occurring during the second and third flashes, as
seen by the increase in total absorbance after each flash. In fast growing hydrogen
cultures, this phase lasted a longer time (Figure 5.3C). In the slow growing cultures, the
553 nm absorbance began dropping immediately, indicating that proton gradient buildup
was minimal. This is consistent with an oxidized quinone pool and particularly with the bheme changes described above: turnover of the bc 1 complex is happening too slowly to
cause large changes in the electrochemical field.
128
A
B
C
.
2000
1500
1000
500
0
-1000
-2000
-3000
I
3000
2000
1000
-5W0
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amm
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U
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-500
400
-1000
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200
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30
40
milliseconds
50
6(0)
0
10
20
30
40
milliseconds
50
60
0
20
40
60
milliseconds
80
Figure 5.3. Absorbance changes in acetate grown cultures (red) vs. cultures growing
rapidly (green) or slowly (blue) on hydrogen. The difference in absolute signal is due to
the different densities of the cultures. The three rapid flashes occur in the first 300
microseconds of the experiment. A)The absorbance changes at 420 primarily reflect the
changes in cytochrome c oxidation state. B) The absorbance changes at 435, which show
reaction center oxidation. The first two flashes excite the reaction center, leading to a
rapid increase in absorbance, seen as two peaks, with the first peak nearly at zero. The
third flash does the same, but in the case of the acetate and fast hydrogen cultures the
quinone pool is strongly reduced and therefore unable to rapidly replace the quinone
Q8H2 that has been reduced by the first two flashes and the response to the third flash is
only a small peak. Over a longer time scale, oxidation of the b hemes is also detectable in
the acetate and fast hydrogen cultures, seen as a negative change in absorbance. C)
Changes at 553 nm, reflecting the changes in electrical field.
129
Oxidized cells were not reducible with acetate, but were reducible with Fe(ll)-NTA
When cultures that had been grown on hydrogen under electron limiting
conditions were exposed to acetate, their quinone pools (as determined by the effect of
the third flash on the reaction center and the re-reduction of the cytochrome c2)
remained oxidized even after a 30 minute incubation (Figure 5.4A), indicating that they
could not immediately use reduced carbon as an electron source. Fe(Il)-NTA, however,
reduced the quinone pool of the cells after a 5-10 minute incubation (Figure 5.4B).
Surprisingly, this effect was evident even when cultures were incubated in the dark: after
Fe(II) was added, the cuvette was placed in the spectrometer and kept in darkness. An
initial measurement of the culture indicated that the quinone pool was still oxidized, but
after approximately 10 minutes (the exact time was somewhat variable), a second round
of measurements indicated the quinone pool of the culture was reduced.
130
A
B
600
200
400
100
E
E
U1 200
-
0
Bow
-ynVen
-
wdh acett
slow hoge
with FeII}.NTA
0
-100
-00
-200
4000
5
10
15
20
muisconde
25
30
0
5
10
15
20
25
30
meconds
Figure 5.4. The effect of acetate and Fe(ll) on oxidized cultures. A) Wild type oxidized
(i.e. slow hydrogen) cultures alone and with acetate after a 30 minute incubation. There
was no oxidation of the quinone pool (all three flashes were the same), indicating that
the cells cannot rapidly use organic material to reduce their quinone pools. B) Wild type
oxidized cultures before and after a 10 minute incubation with Fe(Il)-NTA. Cells show
clear signs of having highly reduced quinone pool, as indicated by the lack of response to
the third flash and the rapid oxidation of the b heme (indicated by the negative 435
absorbance). A fourth flash was used in this experiment and showed the same response
as the third flash, as expected.
131
Reduction by Fe(lI)-NTA was blocked by antimycin
In order to isolate electrons coming from Fe(II), we used the bci inhibitor
antimycin to block cyclic electron flow. The effect of antimycin could be seen through
changes in the b heme signal at 435 nm (Figure 5.5A): after the third flash, instead of
becoming negative as the b hemes are oxidized, the absorbance becomes positive as the
bc, complex becomes fully reduced. This is because antimycin blocks the Q site, which
blocks electrons from returning from the bci complex back into the quinone pool. The Qo-site
will therefore reduce both b hemes, which remain reduced until the system slowly
returns to redox equilibrium.
Contrary to our expectations, the reduction of the quinone pool by Fe(II) was
blocked when the cells were treated with the bc1 complex inhibitor antimycin (Figure
5.5B). The quinone pool in antimycin-inhibited cells remained oxidized for many minutes
after Fe(II) addition, even when the cells were exposed to light. This results strongly
suggests that the bci complex is required for transfer of electrons from Fe(ll) to the
quinone pool.
It is interesting to note that the effect of antimycin was somewhat unreliable:
acetate and fast growing hydrogen cells were either unaffected by antimycin or were
affected only for a short time. Antimycin was more stable in oxidized cells, but in some
cases, even these samples stopped being inhibited by antimycin partway through the
experiment (as indicated by the loss of the b heme reduction described above). When
antimycin stopped working, it did so rather spectacularly: the cells quinone pool became
more reduced then it had been even before the addition of antimycin, with a strong b
heme oxidation signal (Figure 5.5A, red trace). This quinone pool reduction indicated that
the cells were somehow obtaining electrons from the addition of antimycin, either from
the drug itself or from the ethanol it was dissolved in. The reason for TIE-1's resistance to
antimycin is unclear. The bc1 complex appears structurally similar to that found in other
organisms, containing the both Rieske protein and the bci subunit. Antimycin effectively
132
inhibited the bc 1 complex in washed membrane fragments, indicating the resistance was
due to cell function. It is possible that TIE-1 can either export antimycin or can oxidize it.
The second possibility was suggested by the quinone pool reduction in the antimycinresistant cells, though the quinone pool reduction may simply indicate that the cells could
metabolize ethanol, which was also added. Highly oxidized cells were more susceptible to
antimycin.
133
10w
A
50O
E
nunT~y0n
Utg
10 mkAg
1Wha
hytgen VA"N
-1000
-10
10
0
30
20
50
40
"WAseonds
80
B
300
E200
conditon
contro
vUlhanbrnyan andF(I)TA
100
.5
0
5
10
me
15
20
25
30
Figure 5.5. Effect of antimycin on cells. A) The effect and failure of antimycin. When
antimycin successfully inhibited the bc1 complex, the absorbance change at 435 nm was
positive over longer time periods due to the reduction of the b hemes. When it failed,
the cells became highly reduced, with a very large negative absorbance change on longer
time scales, indicating the oxidation of the b hemes. B) When oxidized cultures are
successfully inhibited with antimycin, the addition of Fe(Il)-NTA had no effect on the 3 rd
flash, indicating that it did not reduce the quinone pool.
134
Reduction of TIE-1 cells by Fe(lI)-NTA is independent of pioABC
Although the effect of antimycin on Fe(Il)-mediated reduction of the cells was
unexpected, the ability of Fe(II) to reduce wild type TIE-1 cells was not, and the model for
PioABC oxidation of Fe(ll) predicts such an effect. Subsequently, we tested the APIoA,
APioB, and APioC mutants under the same conditions.
To our complete surprise, we found that all three mutant strains were easily
reduced by Fe(Il)-NTA, including ApioA (Figure 5.6), which has no Fe(ll) oxidation activity
in bulk experiments. In single turnover experiments, no obvious differences were
apparent once the cells had been reduced: 435 nm traces showed that the quinone pool
was reduced, 420 nm traces showed that the cytochromes were rapidly re-reduced, and
553 nm showed that cells with Fe(II) added were able to produce a strong field. Figure 5.6
shows these changes for wild type TIE-1 and ApioA cultures.
135
B
A
4M UKt AA
0
0
-200
-500
.400
-1000-
400-
-1500 -1000
00
300p200
400-xidized
200
0
wt
w~h F9QI)
2-
0
-100~
-200_
-200~
300-
200
200-
100-0
100/K
-100-
0
-100-10
efized
A
-200
0
10
20
30
40
50 60
-10
0
10
20 30
40
50
60
ms
Figure 5.6. Response of A) ApioA, and B)wild type TIE-1 to the addition of Fe(ll). All
three wavelengths indicate that the quinone pool of both strains becomes reduced after
incubation Fe(ll)-NTA. This reduction generally occurred within 10 minutes, even if the
cells were placed in the dark immediately after adding Fe(II).
136
Discussion and conclusions
Previous work with R.palustris TIE-1 unambiguously showed that the organism's
ability to oxidize Fe(II) was dependent on the Pio proteins, particularly the decaheme
cytochrome PioA. Based on this data, a model was developed for electron flow in which
electrons are transferred to the photosynthetic reaction center through PioA and PioC.
The findings described in Chapter 4 of this work partially supported this model: PioC has a
reduction potential close to that of the reaction center, and in vitro experiments showed
that PioC is capable of reducing the reaction center, albeit slowly.
The model proposed made clear predictions regarding electron flow in the wild
type and mutant strain whole cells: first, that electrons should not be able to enter the
transport chain in the pioA mutant; second, that electron transfer from Fe(ll) to the
reaction center should not be hindered by blocking downstream steps, such as transfer to
the bc1 complex. Surprisingly, neither prediction was supported by our flash induced
spectrometry experiments: both the wild type and the mutant strains were reduced by
Fe(II), and antimycin (a bc1 complex inhibitor) blocked the reduction of cells by Fe(II).
Our results thus require a new model to explain them. But what would this model
include?
A missing part of the puzzle (shown as a black box in Figure 5.2) is the reverse
electron transfer from Fe(II) to NAD. Almost nothing is known about the mechanisms of
reverse electron transfer in phototrophs. We therefore looked for inspiration to the
studies done on the acidophilic oxidizing bacterium Acidithiobacillusferrooxidans. The
model for reverse electron transfer in this organism (described in Chapter 2 of this work)
is through the bc 1 complex. If a similar system is present in TIE-1, it could explain the data
presented in this chapter. This new possible model isvisualized in Figure 5.7.
If the route of electron transfer were through the bci complex, electrons would
feed into the quinone pool - which would explain why the quinone pool became reduced.
This reaction would require energy in order to push electrons uphill from the bci complex
137
to the quinone pool, but it would not require light directly. This is consistent with the fact
that once Fe(II) was added to the cells, they could then be incubated in the spectrometer,
in the dark, and still have their quinone pools reduced. The observation that antimycin
blocks reduction by Fe(II) is consistent with electrons moving through the bc1 complex the model would clearly predict this result.
The fact that electrons from Fe(II) still reduce the quinone pool in the pio mutants,
however, remains an enigma, especially in the case of ApioA. The complete lack of
detectable Fe(II) oxidation activity in the pioA mutant indicates that PioA serves an
essential function in iron oxidation. Given the nature of the protein (a decaheme
cytochrome), a function in transferring electrons from Fe(II) still seems the most likely
scenario. However, reduction of the quinone pool in the mutant indicates that PioA is not
the only route electrons can take from Fe(II). It is important to note, however, that the
amount of Fe(II) oxidized in these flash induced experiments is extremely low - likely
submicromolar amounts. It is possible that although enough electrons can take an
alternate route from Fe(ll) to the quinone pool to be visible in these highly sensitive
experiments, truly efficient catalytic oxidation requires PioA. It is also possible that PioA
serves some kind of detoxification function - if it associates with PioB at the outer
membrane as has been postulated, it may keep Fe(ll) out of the periplasm and hence
avoid any toxicity issues that may arise from excess Fe(II) in the periplasm.
138
Fe(II)
PioA
PioC???
HiZ
+ji
H+
bch
IC
membl
'2
Q
QH2
-I
QH
-
\bh
caratenoid
RC
f r
NAD-NADH
CO 2
Figure 5.7. A new possible model for Fe(ll) oxidation. Reverse electron transport is
shown in green, and this path for electrons explain the data we have collected. The role
of the Pio proteins in the new model is somewhat unclear, especially that of PioC. It is
possible that it donates to the bc1 complex, or that it has an unknown purpose in
reduction the reaction center.
139
Our new model would also explain the longstanding observation that TIE-1 prefers
more reduced electron donors to Fe(ll): cultures growing on acetate show little to no
Fe(II) oxidation activity, expression of the Pio proteins is down regulated on other
substrates (15), hydrogen grown cultures oxidize Fe(ll) more slowly than Fe(II) grown
cultures (3), and hydrogen inhibits Fe(lI) oxidation (16). In the old model, in which
electrons flow downstream to the reaction center and enter the quinone pool through
light excitation, the thermodynamic advantage of using more reduced electron donors
was not obvious - although the preference for hydrogen could partly be explained
through kinetic arguments (16). If reverse electron transfer is an active process through
the bc1 complex, however, this strategy would make perfect sense: electrons from more
reduced sources could enter the electron transport chain at an earlier point (such as
hydrogenase in the case of hydrogen) and could be used more efficiently.
The issue of efficiency is one reason reverse electron transport through the bc1
complex initially seemed less likely in phototrophs. Lithotrophic bacteria have no other
option, but given the high potential of the reaction center, phototrophs can
(theoretically) transfer electrons from sources such as Fe(II) to the reaction center and
then to the quinone pool via the light excited reaction center without expending
previously harvested energy (although the next step, transferring electrons from the
quinone pool to NAD*, would still be an uphill reaction). Why would they expend energy
to push electrons uphill into the quinone pool using the proton motive force when
reaction center excitation does it more efficiently? Seen from another angle, however, it
is logical for the bacterium to decouple electron acquisition from light. It is easy to
speculate that availability of electron donors might vary in the environment depending on
external inputs (through, for example, water currents) and on the other types of
organisms present. If electron donors were available sporadically, then it would be
advantageous for a phototroph to be able to use them whenever they are present and
not just in the presence of light - which, naturally, is only available during the day. Thus,
140
an electron-limited cell could build up energy stores from light, and use those stores to
obtain electrons as soon as they became available.
This hypothesis of reverse electron transfer through the bc 1 complex raises several
questions: first, what role do the Pio proteins play in iron oxidation? Second, what other
proteins are involved? PioC represents a particular puzzle; although we have shown that
it can reduce the reaction center in vitro, it is not clear how relevant this is in vivo if
electrons from Fe(II) are donated to the bc1 complex. One possibility is that PioC actually
donates to the bc 1 complex in whole cells, either directly or through another
intermediate, and that in vitro reduction of the reaction center is an off target effect which would be an alternate explanation for its relatively sluggish donation to the
reaction center. Donation to the bci complex is at odds with its measured reduction
potential - however, at least one other HiPIP is known to shift reduction potential
drastically when complexed with its physiological partner. Such shifts could occur for
PioC and would explain this discrepancy. It is also possible our in vitro experiments do
accurately reflect in vivo function, and that PioC-mediated reduction of the reaction
center is playing an as yet undetermined role in Fe(II) oxidation.
If the Pio proteins are not the only mechanism for oxidizing Fe(II), then what are
the other proteins involved? In At. ferrooxidans, Fe(II) electrons are thought to proceed
through an outer membrane cytochrome to a copper protein and through another
cytochrome before reaching the bci complex. The final donor to the bc 1 complex is
thought to be CycAl, a c4 cytochrome. TIE-1 has many predicted cytochrome c type
proteins in its genome, but it is intriguing to note that there is a gene (rpal 0810)
predicted to encode a c4 cytochrome in the TIE-1 genome in the same region as the pio
operon. Rpal_0810 is predicted to be a 21 kD protein with two CXXCH heme binding
motifs. Although CyclA is also a diheme cytochrome, the similarity between the two
proteins is low: 32% identical and 45% similar. Homologues are found in other R.
palustris strains, as well as in the related Bradyrhizobiumjaponicum. While its proximity
141
to the pio operon is merely suggestive of a possible function in reverse electron transfer,
this gene might be a good target for future study.
The use of flash induced absorbance spectroscopy has great potential to answer
some of the questions raised by the data presented in this chapter. Future work using
this and other techniques will help us develop a more complete model of Fe(II) oxidation
in TIE-1 and of reverse electron flow in photosynthetic organisms.
142
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Cusanovich MA, Bartsch RG, Fischer U, Van Beeumen JJ. 2003. Amino acid sequences
and distribution of high-potential iron-sulfur proteins that donate electrons to the
photosynthetic reaction center in phototropic proteobacteria. J Mol Evol. 57(2):181199.
2. White D. The physiology and biochemistry of prokaryotes. 3rd ed. New York: Oxford
University Press; 2007.
3. Jiao Y, Kappler A, Croal LR, Newman DK. 2005. Isolation and characterization of a
genetically tractable photoautotrophic Fe(Il)-oxidizing bacterium, Rhodopseudomonas
palustris strain TIE-1. Appl Environ Microbiol. 71(8):4487-4496.
4. Jiao Y, Newman DK. 2007. The pio operon is essential for phototrophic Fe(II)
oxidation in Rhodopseudomonas palustris TIE-1. J Bacteriol. 189(5):1765-1773.
5. Bewley KD, Ellis KE, Firer-Sherwood MA, Elliott SJ. 2013. Multi-heme proteins:
Nature's electronic multi-purpose tool. Biochim Biophys Acta.
6.
Fonseca BM, Paquete CM, Neto SE, Pacheco I, Soares CM, Louro RO. 2013. Mind the
gap: cytochrome interactions reveal electron pathways across the periplasm of
Shewanella oneidensis MR-1. Biochem J. 449:101-108.
7. Hochkoeppler A, Zannoni D, Ciurli S, Meyer TE, Cusanovich MA, Tollin G. 1996.
Kinetics of photo-induced electron transfer from high-potential iron-sulfur protein to
the photosynthetic reaction center of the purple phototroph Rhodoferax fermentans.
Proc Natl Acad Sci U S A. 93(14):6998-7002.
8. Meyer TE, Cusanovich MA. 2003. Discovery and characterization of electron transfer
proteins in the photosynthetic bacteria. Photosynthesis Research. 76(1-3):111-126.
9.
Menin L, Schoepp B, Parot P, Vermeglio A. 1997. Photoinduced cyclic electron
transfer in Rhodocyclus tenuis cells: participation of HiPIP or cyt c8 depending on the
ambient redox potential. Biochemistry. 36(40):12183-12188.
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10. Trosper TL, Benson DL, Thornber PJ. 1977. Isolation and spectral characteristics of the
photochemical reaction center of Rhodopseudomonas viridis. Biochim Biophys Acta.
460(2):318-330.
11. Orlando JA. 1967. Rhodopseudomonas spheroides cytochrome c2. Biochim Biophys
Acta. 143(3):634-636.
12. Gabellini N, Hauska G. 1983. Characterization of cytochrome b in the isolated
ubiquinol-cytochrome c2 oxidoreductase from Rhodopseudomonas sphaeroides GA.
FEBS letters. 153(1):146-150.
13. Bailleul B, Cardol P, Breyton C, Finazzi G. 2010. Electrochromism: a useful probe to
study algal photosynthesis. Photosynth Res. 106(1-2):179-189.
14. Lancaster CRD, Michel H. 2006. Photosynthetic Reaction Centers of Purple Bacteria.
Handbook of Metalloproteins: John Wiley & Sons, Ltd.
15. Bose A, Newman DK. 2011. Regulation of the phototrophic iron oxidation (pio) genes
in Rhodopseudomonas palustris TIE-i is mediated by the global regulator, FixK. Mol
Microbiol. 79(1):63-75.
16. Croal LR, Jiao Y, Kappler A, Newman DK. 2009. Phototrophic Fe(II) oxidation in an
atmosphere of H2: implications for Archean banded iron formations. Geobiology.
7(1):21-24.
144
Chapter 6: A larger perspective
145
The work for this thesis began with a single, well defined goal: to understand the
biochemical details of ferrous iron oxidation in R.palustris TIE-1. We began with a simple
and self contained model which included two redox active proteins; characterizing these
two electron transfer proteins would, we thought, validate and add detail to the model.
However, over the course of the project, we discovered that thinking about iron
oxidation as an isolated catabolic process was not sufficient to understand it. The first
challenge came when we realized that under some conditions, iron could have a negative
effect on TIE-1 (Chapter 3). This observation made it clear that TIE-1's interactions with
iron were not as simple as we had initially thought, and highlighted an important point:
bacteria can interact with their environment on multiple levels, and the same compound
can have multiple effects. The involvement of copper in the negative effects of Fe(II)
added another layer of complexity to the story, and reinforced the need to think about
bacterial environments as a whole, even when that environment is a (supposedly) well
controlled culture tube.
The second major shift in our thinking came when we probed iron oxidation in
whole cells. Although in vitro characterization of PioC supported our initial model for
Fe(II) oxidation (Chapter 4), the whole cell data did not (Chapter 5). This contradiction
reminds us of the importance of validating in vitro results in the whole organism although biochemical studies can tell us a great deal about a protein's function, when it is
removed from its cellular environment, the activities we observe may or may not reflect
its behavior in vivo.
The results we obtained in whole cells could only be explained with an entirely
new model. In order to develop it, we needed to stop thinking about iron oxidation as an
isolated unit, and consider it in the context of its larger physiological function - that is, in
the context of reverse electron transfer.
146
Although we had thought about reverse electron transfer through the bc 1
complex, in the context of lithotrophic iron oxidation (summarized in Chapter 2), we had
not seriously considered such a system for photoferrotrophy. Reverse electron transfer
through the bc 1 complex seemed unlikely in photoferrotrophs for two reasons: 1) PioC,
which we assumed was the final donor in the chain, has been well documented as
reducing the reaction center. 2) Pushing electrons uphill through the bci complex seemed
like an inefficient way to get electrons to the quinone pool compared to delivering them
to the reaction center.
This second point is very much a reflection of habits of thought formed by years of
thinking about heterotrophic and lithotrophic bacteria. In these cases (where a great deal
of basic biology has been done), the bacterium is defined by energy limitation - either by
a limitation in reduced compounds or of terminal electron donors. The phototrophic
case, however, is entirely different - as long as light is present, energy is not the limiting
factor. The evolutionary pressure to function in the most energy efficient way possible
will not be the same for phototrophs as for non-phototrophic organisms. Flexibility in
acquiring the limiting resource - electrons - could easily be more important than
efficiency.
Thinking about TIE-i in this broader way helped us to develop a new model for
Fe(ll) oxidation, and indeed for TIE-1's interactions with iron as a whole. It will be exciting
to test these new ideas and see whether they are correct, and to discover in greater
detail how TIE-1 accomplishes reverse electron flow mechanistically. To this end, several
experiments come to mind that will help us validate the model and better understand the
system:
e
Determine whether cytochrome c4, Rpal_0810, is involved in Fe(ll) oxidation. If
TIE-1's reverse electron transport chain is similar to the acidophilic Fe(Il) oxidation
system, then it likely contains at least one more cytochrome. The location of the
147
gene proximal to those encoding the Pio proteins on the chromosome, as well as
the class of cytochrome, makes Rpal_0810 a strong candidate for future study.
Use flash induced absorbance spectrometry to determine whether there are other
cytochromes involved in the pathway. The experiments described in Chapter 5
were all done at discrete wavelengths, and did not allow us to distinguish between
different cytochrome c reactions. However, it is also possible to take wavelength
scans, by taking flash induced absorbance measurements at 1 nm intervals. While
all cytochrome c type proteins show a shift in absorbance around 420 nm, the
precise peak varies with the specific protein. It is possible to de-convolute these
and determine whether all the cytochrome c reduction we see is attributable to
c2 , c1, or some other cytochrome.
*
Determine whether there are differences in NAD/NADH ratios in the wild type and
pio mutants when oxidized cells are exposed to Fe(II). If the Pio proteins are
involved in reverse electron transfer, then we would expect to see some
difference in the rate at which it proceeds when they are absent. The nature of
the flash induced experiments made it difficult to quantify the time it took for the
quinone pool to become reduced. The shift in NAD/NADH ratios should have a
more gradual shift over time, and thus give us more detailed information about
the change in cell redox state.
*
Determine whether there is any growth defect in ApioA in the presence of iron
when other electron donors are present. PioA may have another role besides
simple electron transfer, and mitigating iron toxicity seems the most likely.
Looking for differences in growth under electron limiting conditions is the first
step in testing this possibility.
e
On a related note, significant work remains to be done to determine the precise
function of Rpal_4085 in metal toxicity. Tests with the deletion mutant can help
determine which metals, if any, Rpal_4085 detoxifies, and further regulatory
studies will better define which metals it is upregulated by.
148
These experiments will help untangle TIE-1's complex interaction with iron, and
will no doubt suggest a number of further areas for future study. They will also offer a
glimpse into an area that is almost entirely unstudied - that of reverse electron transfer
in photosynthetic bacteria.
149
Appendix A: Attempted purification of PioA
150
Introduction
Although significant progress has been made in understanding the iron oxidation
system of TIE-1 the detailed function of PioA, the presumed iron oxidase, remains an
enigma. Several related proteins have been characterized in recent years, including
several decaheme iron reductases from Shewanella oneidensis (See Chapter 2) and the
decaheme iron oxidase MtoA from the neutrophilic microaerobic sideroxydans
lithotrophicus ES-1 (1). However, PioA remains the only photosynthetic decaheme iron
oxidase known to date. Although it issimilar to other decahemes in its heme-binding
region, PioA also has a unique N-terminal domain, which has an unknown function.
Many of the questions about PioA would be answered by the characterization of
the purified protein. However, the purification of PioA has proven a challenge. Because
PioA is natively expressed at significant levels only during growth on Fe(II), which
produces low cell mass, an over expression system is required. This appendix describes
several attempts to over express PioA, and makes suggestions for future efforts.
Materials and methods
Heterologous expression in E. coli
E. coli does not efficiently produce heme proteins under normal aerobic growth;
however, incorporation of the plasmid pEC86, which constitutively expresses the heme
maturation machinery, has been successfully used to express cytochromes. I attempted
PioA over expression using plasmids developed by Yuri Londer (2) specifically for
production of multi-heme proteins. The first, pMKL1, fuses the inserted protein with a
histidine tag and maltose binding protein, to increase solubility and allow for affinity
purification. The second, pFCM21, co expresses the chaperone protein FkpA which is
known to assist in protein solubility and folding.
151
PCR amplificationand cloning
Primers used are listed in Table A.1. PioA was amplified using the FailSafe system,
with 1 pM each primer, 35 ng DNA, 1.25 U FailSafe enzyme, and FailSafe Buffer D,
according to the manufacturer's directions. The amplified PCR product was cloned into
pMKL1 and pFCM21 using the lic system (2), transformed into electrocompetent E. coli
DH5c cells using Gene Pulser 2 (BioRad) at 2400 V with a resistance of 200 ohms and a
capacitance 25 pF. Transformed cells were recovered on SOB media, and plated on the
appropriate antibiotic plates.
Colonies that grew overnight were restruck and checked for the insert with colony
PCR, and the plasmids from positive colonies were isolated using the Qiagen miniprep kit.
Inserts were sequenced to ensure that the insert was correct, and the plasmids were then
transformed into E. coli BL21 with pEC86 or E.coli C43 with pEC86 for expression studies.
Expression media and procedures
Several media were tested for optimal induction conditions: LB broth was
purchased premixed, as was Plasmid Gro (ISC BioExpress, Utah). Autoinduction media
was made using the protocol developed by Studier (3).
Additional iron was added from
an Fe(Ill)-HCI stock. Samples spun down at 6,000g for 10 minutes, rinsed and
resuspended in buffer, and either lysed either by sonication or French Press. The lysate
was clarified by centrifugation at 20,000g for 30 minutes. Spectra and optical densities
were taken on a DU 800 spectrophotometer (Beckman-Coulter).
Protein gels and heme stain
Samples for were boiled for 5 minutes in Laemmli buffer without a reducing agent
to avoid removing the hemes. Gels were run using the BioRad SDS-PAGE system with
purchased polyacrylamide gels and SDS buffer (BioRad). Gels were run at 100-130 V. For
protein staining, gels were rinsed in diH 20 and stained with BioSafe commasie (BioRad).
For the heme stain, run gels were soaked in 12.5% trichloroacetic acid for 30 minutes and
152
washed in diH 2 0 for 30 minutes. The heme stain was prepared as follows: 200 mg odianosine (Sigma) was added to 180 ml diH 2 0, and stirred rapidly for 15-30 minutes. 20
ml of 0.5 M Na-citrate buffer, pH 4.4 was added, and .5 ml of 30% H202 was quickly
added. The mixture was swirled a few times and immediately poured over the gel. The
gel was incubated in a covered container and checked periodically. Bands appeared at
varying times ranging from 2 hrs. to overnight.
Heterologous expression in Shewanella oneidensis
PCR and cloningprotocols
Because PioA contains an internal sequence that is very similar to its 5' end,
certain primer sets did not amplify the full gene. In order to obtain the gene with the
appropriate restriction sites, the pioA gene was amplified in two sections and
recombined. The successful PCR reaction was adapted from (4), and contained 1 PM
each primer, 25-100 ng DNA, 10 mM Tris buffer pH 8.3, 50 mM KCI, 1.5 mM MgCI, .2 mM
dNTPs, 0.6 M Betaine, and 1.25 units FailSafe polymerase. Reaction conditions were: 3
min 950C,then (1 min 950C,30 sec 550 C, 1 min 700 C)for 40 cycles, with a final 70
extension for 3 minutes. Reactions were assembled on ice and placed in a preheated
cycler.
Yeast recombination
The two PCR fragments obtained were recombined with each other and the
plasmid pMQ87 by transforming the fragments into Saccharomyces cerevisiae (5). Briefly,
0.5 mIs of an overnight S.cerevisiae culture were pelleted at 8,000 rpm for 10 seconds
and washed with 0.5 ml sterile TE buffer. The pellet was re-suspended in Lazy Bones
Solution: (40% Polyethylene glycol (MW 3350), 01. M Lithium acetate, 10 mM Tris-HCI, 1
mM EDTA). 20 pl of 2 mg/ml denatured salmon sperm DNA, 5 pl linearized plasmid DNA,
and 45 pl of each PCR reaction were added. The tube was vortexed for 1 minute, and
incubated on the bench for 2-3 days. The tube was then shocked at 42 0Cfor 10 minutes,
153
washed and re-suspended in 200 VI TE buffer, plated onto SC-URA plates, and incubated
for 2 days at 300 C. Several colonies were selected and grown in liquid culture. Plasmid
was recovered from the yeast cultures using a Qiagen miniprep kit according to the userdeveloped yeast prep protocol.
Transformation of E. coli
Plasmid recovered from yeast was transformed into electro competent E. coli
DH5a using Gene Pulser 2 (BioRad) at 2400 V with a resistance of 200 ohms and a
capacitance 25 pF. Cells were recovered for 1 hr. in SOB media at 370 C, and plated on
selective media for overnight growth. Colonies were restruck and confirmed with PCR.
Plasmids from colonies containing the insert were purified using the Qiagen miniprep kit,
and sequenced to confirm the accuracy of the clone. Correct clones were then cut
sequentially with EcoRI and Hindlil (NEB), and ligated into pBAD18-kan (6), cut with the
same enzymes. Transformants were screened for the insert, and a correct PioA-pBAD18
plasmid purified using the Qiagen miniprep kit.
Transformation into Shewanella oneidensis
The PioA-pBAD18 plasmid was transformed into Shewanella oneidensis MR1
AmtrA &dmsE AmtrD Aso4360 (generously provided by Jeff Gralnick) using the following
protocol (7): the bacteria were grown to OD 0.5 in LB. 1 ml of the culture was pelleted,
washed, and re-suspended in 40 pl of sterile 1 M sorbitol. The cells were chilled on ice,
and 200 ng plasmid was added to the tube. The cells were electroporated with 550 V,
and re-suspended in 0.8 ml SOC medium. The cells were concentrated and plated on 100
pg/ml kanamycin plates. Colonies appeared after 2 days.
Homologous over expression in TIE-1
Cloning of the PioA constructs
Constructs were made using the broad range expression vector pBBR and its
derivative pSRK-kan (8), which is replicated in TIE-1 and contains a tightly regulated lac
154
operon. 2 constructs were cloned: the first, PioA-pSRK, contained pioA with its native
signal sequence under the control of the lac operon. The second, cycPio-pBBR, contained
the entire pio operon with pioA fused to the signal sequence and upstream region of TIE1's cytochrome c2. The second construct would, in theory, express the Pio genes under
anaerobic growth conditions, when cytochrome c2 is expressed at high levels.
The PioA-pSRK construct was created as described above for PioA-pBAD18k construct,
with the exception that the restriction enzymes used were Ndel and Nhel.
CycPio-pBBR was made by amplifying the pio operon fragments without pioA's
signal sequence and the upstream/signal sequence from cyc2 using the PCR protocol
described for PioA-pBAD, and the primers listed in Table A.1. The fragments were
recombined with pBBR using Gibson recombination (9), and transformed into E. coli
DH5c. Plasmid purifications were done on several transformants, and the inserts
sequenced. Correct clones were transformed into TIE-1 by first transforming the plasmids
into the E. coli mating strain S-17, and then mating them into TIE-1; 0.5 ml each of a 2-day
culture of TIE-1 and overnight culture of the donor E. coli strain were spun down, rinsed
in fresh YP-succinate media, and combined. The mixture was spun down and resuspended in 200 pl YPS medium, spotted onto a YPS plate, and incubated overnight at
30 0 C. Positive colonies selected for on YPS plates with 400 pg/ml kanamycin, which TIE-1
could survive, but E. coli could not.
Induction of PioA constructsin TIE-1
Cultures were grown anaerobically in fresh water medium under incandescent
light bulbs for all induction experiments. Cells were harvested by centrifugation at 7,000
g for 20 minutes at 40 C, French pressed, then spun at 30,000 for 15 minutes, and
ultracentrifuged at 100,000 g for 1 hr. to obtain the soluble fraction. Samples were run
on BioRad SDS page gels and heme stained to determine whether PioA was expressed.
155
Activityassay for PioA
In order to test the Fe(II) oxidation activity of lysates, we attempted to develop an
in vitro activity assay in which the oxidation of Fe(II) would be linked to the reduction of a
redox active compound, so that it could be followed kinetically over the course of the
reaction. A number of compounds were tested for their abiotic reactivity with Fe(II),
including the copper protein azurin, methylene blue, phenazine ethosulfate, and toluene
blue. Of these, methylene blue and toluene blue were the least reactive in the absence of
lysate. Both dyes were then tested with lysate and Fe(II). The initial experiments were
done in a Coy anaerobic chamber with an N2 H2 atmosphere. However, both compounds
were rapidly reduced by lysate even in the absence of Fe(II). Experiments in an Mbraun
chamber indicated that this was due to H2 acting as an electron donor, and further
experiments were done in the Mbraun. Dye and Fe(II) were tested at varying
concentrations for each component and at various pHs. Experiments were done by
mixing the components in a UV-vis disposable cuvette with a round top, sealing it with a
green rubber stopper, then moving it out of the chamber as quickly as possible and
measuring the absorbance changes over the course of 30 minutes. An alternate method
involved mixing all but one component in a quartz cuvette with a screw cap and rubber
septum, and injecting the final component just before measurement.
Results
Cloning PioA constructs
The constructs PioA-pFCM21, PioA-pMKL1, PioA-pBAD18k, PioA-pSRK, and cycPiopBBR were successfully constructed and sequenced. Diagrams of the composition of each
are shown in Figure A.1. Pio-pSLM2 was attempted twice, and contained sequence errors
each time. Expressions studies were only performed on the correct clones.
156
PioA-pMKL1
IacP]6xHS-MBP
PioA
PioA-pFCM21
IacP
PioA
PioA-pBAD__
araP ig
PioA
PioA-psrk
IacP
pjo
PioA-pBBR
cyc2P/sig
p oABC
Figure A.1: PioA expression constructs made in this study. lacP -lac promoter; sig- sec
signal sequence; 6xHis-MBP -histidine tag followed by maltose binding protein tag; araP arabinose promoter; cyc2 P/sig - cyc promoter and signal sequence.
157
Heterologous expression conditionsfor PioA
Various conditions were tested in an attempt to optimize PioA expression. Table
A.2 summarizes a number of these attempts. The final conditions selected as the most
promising were: either PioA-pSRK or PioA-pFCM21 in BL21 with pEC86 grown in
PlasmidGro media with 100 pM Fe(lll)-NTA aerobically to an OD of 0.6-1, then induction
with 30 pM IPTG at 30' Covernight. This condition resulted in a significant amount of
pink visible in pelleted cells. However, once the cells were lysed, much of the color often
remained in the pellet, indicating that expressed PioA was precipitating.
Solubilization ofheterologouslyexpressed PioA
Various lysis conditions were tested in order to maximize soluble PioA in E. coli
expression. These conditions were attempted with both the pFCM21 and pMKL1
constructs. The effects of each buffer were quantified by taking the ratio of the 410/280
nm absorbance (samples were diluted to obtain accurate readings). As shown in Figure
A.2, the expressed protein displayed a typical heme signal, with a large peak at 410 nm
and smaller peaks at ~520 nm and ~550 nm. The absorbance at 280 indicated the overall
protein recovered: thus, a higher 410/280 ratio indicated that a greater percentage of the
soluble protein was heme protein, presumably PioA. The conditions are summarized in
Table A.3, and those with the highest ratios were run on SDS-PAGE gels using the heme
stain. A representative gel isshown in Figure A.3: in all cases, the majority of the heme
proteins stayed in the well, ran at very high molecular weights, or smeared throughout
the lane. A native gel run without SDS had the same result. Furthermore, attempts to
enrich PioA from these lysates on a nickel column (PioA-pMKL1) or on a CaptoS column at
pH 5.5 (PioA-pFCM21) failed due to the heme protein not sticking to the column. These
results indicated that soluble PioA was not being correctly expressed at high levels in E.
coli.
158
3.5
2.5
1.5
1
0.5
0
260
310
360
410
460
510
560
Figure A.2: Spectra of E. coli BL21 cells expressing PioA. Due to the high concentration of
pure protein in clarified lysate, the absorbance below 300 nm is above the maximum of
the machine, and accurate readings were taken by diluting the lysate 1:10. However, the
peaks at 410, 520, and 550 are clearly visible and are characteristic of reduced
cytochromes.
159
Figure A.3: Gel of PioA-pFCM21 expression in E. coli. 1: empty vector, clarified lysate. 2:
empty vector resuspended pellet. 3: marker. 4: BICENE 8.5 resuspended pellet. 5: BICENE
8.5 clarified lysate. 4: PIPES 6.0, resuspended pellet. 7: PIPES 6.0, clarified lysate. 8: whole
cells boiled in SDS. Although a slight band is visible at ~55 kD, much of the heme protein
is stuck in the well or at high molecular weights, indicated that even the "soluble" fraction
was in fact not fully soluble.
160
Expression in Shewanella oneidensis
Expression in Shewanella was not observed at high levels; due to the large number
of native cytochromes expressed, Shewanella is naturally pink. This was true even for the
deletion strain used, which had a number of cytochromes removed from its genome.
Heterologous expression could therefore only be followed by heme stain, and no
additional bands were observed in induced cultures. However, the PioA-pBAD18
construct was not extensively tested.
Expression of PioA in TIE-1 under an IPTG induciblepromoter
TIE-1 strains containing the over expression constructs were grown anaerobically
in fresh water media containing 100 ptg/ml kanamycin. Induction conditions are listed in
Table A.4. Results based on heme stain were variable: the soluble fraction often
contained a band the size of PioA even in uninduced controls. The intensity of the heme
stained bands varied, likely reflecting differences in staining efficiencies rather than
different levels of expression. The heme stain is not quantitative, which made it difficult
to detect subtle differences in expression. None of the conditions tested resulted in a
thick band, which would a highly over expressed band, and no overexpressed band was
visible by coomassie stain.
Even after ultracentrifugation, samples contained photosynthetic pigments that
absorb at 415 nm. This background, along with the presence of other cytochromes, made
it difficult to use 415 nm absorbance as a measure of PioA in crude samples. Purification
was therefore attempted on H2 grown cultures supplemented with Fe(lI)-NTA that had
been induced for 2 days with 1 mM IPTG. This condition consistently had a band in the
heme stain at the correct size for PioA, and logically should have had the greatest
induction.
161
Several attempts were made to purify PioA using CaptoQ and CaptoS ion exchange
resins. All of these attempts failed: once the pigments and other cytochromes had been
fractionated, the amount of PioA present proved to be miniscule. While it might be
possible to scale up in order to obtain enough biomass for a successful purification, we
opted not to pursue this route.
Expression of PioA in the cycPio-pSRK construct
Although this construct did show some expression of PioA during anaerobic
photoheterotrophic growth, expression (as judged by the heme stain) was still low, and
relatively unreliable. Controlled testing of photoautrophic growth was not done but
holds promise, as the Pio pathway is generally more highly expressed during
photoautotrophic growth.
PioA activityassay
Of the compounds tested, methylene blue proved to be the best electron accepter
for the PioA activity assay. Methylene blue reduction occurred in the presence of TIE-1
lysate and not in lysate free controls. The reaction, and especially the lack of background
reaction, was dependent on pH, with a pH of 7 in Tris buffer giving the least background
while still showing activity in the presence of lysate. An increase in pH led to abiotic
reduction of methylene blue by Fe(ll). The final, most effective conditions settled on
were: 50 pM Methylene blue, 200 pM Fe(II), 50 mM Tris pH7. To this mixture was then
added the sample, and time points were taken as rapidly as possible.
Although cell lysate successfully catalyzed ethelyne blue reduction by Fe(II), no
reproducible difference was observed between lysates from wt. and ApioA cultures
grown on hydrogen and lysate grown on Fe(II), which suggested that significant non-PioA
specific activity was present. Furthermore, the necessity of setting up the experiments in
the Mbraun made this assay cumbersome to perform, and not a helpful diagnostic tool in
the purification of PioA.
162
Conclusions
Although over expression of PioA was ultimately unsuccessful, the results
presented here suggest directions for future attempts.
While some multiheme cytochromes are expressible in E. coli, PioA does not
appear to be one of them. There are many possible reasons for this, and while there may
be a way to overcome the difficulties here described, E. coli expression seems the least
likely to produce results
Expression in Shewanella oneidensis was not deeply explored in this work, and
holds promise. Shewanella already encodes and expresses a number of decaheme
cytochromes, which means it has the machinery to correctly process them. A successful
strategy might involve tweaking the current pBAD system, or creating a new lac inducible
construct, possibly with a signal sequence from a highly expressed Shewanella
cytochrome, such as a small tetraheme cytochrome (STC). This method has been
successfully employed for Cytochrome c nitrite reductase (10), and may well work for
PioA.
The other most likely option is homologous expression. When attempting PioA
expression in TIE-1, a balance needs to be struck between growth yield and PioA
concentration; under conditions which produce rapid growth and high cell yields (aka
photoheterotrophic growth), iron oxidation activity and pio expression is low. The pio
operon is expressed when Fe(ll) is the only available electron donor, but cell yields under
these conditions are too low to make a practical expression system It may be possible
however, to grow cells very slowly, for example with limited H2 present, in order to obtain
higher cell yields while maintaining conditions under which PioA can be expressed. If
enough volume could be grown under these conditions, it may be possible to obtain
enough biomass with significant PioA levels.
During purification, the presence of PioA would ideally be followed with a
combination of the heme stain, to confirm the correct size, and 410 nm absorbance, to
163
estimate concentrations. While TIE-1 does have a number of other cytochromes, the
most abundant ones should be removable early in the purification, allowing tracking of
PioA by absorbance spectra to continue with relative confidence.
164
Table A.1: Primers used in making constructs.
name
PiopMKL1F
sequence
TACTTCCAATCCAATGCG
construct
PioA-pMKL1
comments
CTGGCCTGGGGGACCTG
PioPMKL1R
TAATCCACTTCCAATGCTATC
PioA-pMKL1
GGTGCCAGCGCGATCC
PioA-pFCM21
PioApLBM2
CCGTTGCGTTTGCA
F
CTGGCCTGGGGGACCTG
PioApLBM2
R
PioApMQ87
F
GCTTGTCGTCGTGCA
CTATCGGTGCCAGCGCGATCC
ATTTAATCTGTATCAGGCTGA
AAATCTTCTCTGAATTCGGAA
TGGGAGGCTCTCGGG
CTGATTCTGTGGATAACCGTA
TTACCGCCTTTTCTAGATATCG
GTGCCAGCGCGATCCGGA
GGTCGGACACACCGCCTTGC
PioA-pFCM21
TGAAGTTCTGTTCGGTCTTGC
PioA-pBAD18,
PioA-pSRK
PioApMQ87
R
PioAinternal
F
PioAinternal
R
PioA-pBAD18
Forward primer,
contains EcoRi site
PioA-pBAD18
Reverse primer,
contains Xbal site
PioA-pBAD18,
PioA-pSRK
Forward
binds at
PioA
Reverse
binds at
primer,
site 137 of
Primer,
site 266 in
PioA
PioApMQ87
F3
ATTTAATCTGTATCAGGCTGA
AAATCTTCTCTCATATGGGAG
PioA-pSRK
restriction site
GCTCTCGGGGGGC
PioApMQ87
R
CTGATTCTGTGGATAACCGTA
TTACCGCCTTTCGATCGCTA
TCGGTGCCAGCGCGATCCGG
Forward primer,
contains Ndel
PioA-pSRK
Reverse primer
contains Nhel
restriction site
cycPio-pBBR
Forward primer for
cyc2 upstream
region, with pBBR
A
Cyc2ABCF1
TGTGAGTTAGCTCACTCATTA
GGCACCCCAGGCGGGAAAAT
TAGCCGAATTGTCTAAC
overlap
Cyc2ABCR1
GTTCGGCGCCGGCTGGCCAG
GTCCCGGCCGAGGCAGTGCC
cycPio-pBBR
with PioA overlap
GATCGAC
Cyc2ABCF2
GTCGATCGGCACTGCCTCGGC
CGGGACCTGGCCAGCCGGCG
CCGAAC
Reverse primer for
cyc2 signal sequence
cycPio-pBBR
Forward primer for
PioA with cyc2 signal
overlap
165
Cyc2ABCR2
ATTTTAACAAAATATTAACGC
TTACAATTTACTAGTTTATGCC
TTGCCGGCGTAG
CAACACCAGCGCGCTGAGCA
TCTGGAACGGCGTCGGCAC
cycPio-pBBR
Cyc2ABCRi
GTGCCGACGCCGTTCCAGATG
CTCAGCGCGCTGGTGTTG
cycPio-pBBR
Reverse internal
primer, binds PioB
pBBrcyc2
gttagacaattcggctaattttcCCGC
CTGGGGTGCCTAATGAG
cycPio-pBBR
Binds pBBR with
cyc2 overhang
pBBrpioCen
d
TACGCCGGCAAGGCATAAAC
TAGTAAATTGTAAGCGTTAAT
cycPio-pBBR
Binds pBBR with
PioC overhang
Cyc2ABCFi
ATTTTG
166
cycPio-pBBR
Reverse primer for
PioC with pBBR
overlap
Internal forward
primer, binds PioB
Table A.2: induction conditions for heterologous expression
Strain
Media
Induction
Pellet
induct.
time
appearance
Some pink
in all
pellets,
more in
aerobic
Pink
Ind
pM
OD
temp
02?
PiopFCM21
BL21
PlasmidGro
1
IPTG
0, 10, 30
30
Yes,
No
12, 14
hrs
PiopFCM21
LB
.5
IPTG
0, 30
30
yes
16 hrs
PlasmidGro,
0.5
IPTG 30
30
yes
16 hrs
pMKL1 not
pink, pFC21
pinkish,
blobby
yes
16 hrs
Not pink
yes
14 hrs
RT blobby,
others
pinkish.
BL21
PiopMKL1,
PiopFCM21
BL21
PiopMKL1,
PiopFCM21
autoinduction
medium
N/A
N/A
BL21
PiopFCM21
BL21
PlasmidGro
100 pM Fe
0.7
10, 30
22,
30,
37
300 the
PiopFCM21
PlasmidGro
0.52
30
30
yes
17 hrs
best.
dark pink,
pink in
lysate
BL21
PiopMKL1
BL21
PlasmidGro
.7
30
16,
20,
30
yes
17 hrs
PiopBAD18 in
S.
oneidensis
LB
.85
Arabinose
1000
On
bench
yes
16 hrs
30 dark
pink, other
temps less
so.
Naturally
pink, heme
stain
showed no
induction
PiopMKL1
PlasmidGro
0.8
IPTG
3, 10, 30
30
Yes
19 hrs
No color,
nothing on
167
C43
PiopFCM21
heme stain
PlasmidGro
+/-0.3%
glucose
168
.8
IPTG 25
30
Yes
15 hrs
Glucose
made no
difference
Table A.3: buffers tested for processing heterologous expression
strain
PioA-pMKL1
PioA-pMKL1
PioA-pMKL1
PioA-pMKL1
PioA-pMKL1
PioA-pMKL1
PioA-pMKL1
PioA-pMKL1
PioA-pMKL1
PioA-pFCM21
PioA-pFCM21
PioA-pFCM21
PioA-pFCM21
PioA-pFCM21
PioA-pFCM21
PioA-pFCM21
PioA-pFCM21
PioA-pFCM21
PioA-pFCM21
PioA-pFCM21
PioA-pFCM21
PioA-pFCM21
PioA-pMKL1
PioA-pMKL1
PioA-pMKL1
PioA-pMKL1
PioA-pMKL1
PioA-pMKL1
PioA-pMKL1
PioA-pMKL1
Buffer (50 mM)
HEPES
HEPES
BICENE
MES
MES
PIPES
PIPES+DMSO
MOPS
MOPS+DMSO
HEPES
HEPES
BICENE
BICENE
MES
MES
PIPES
PIPES
MOPS
MOPS
Phosphate
Phosphate + 10 mM
betaine
Tris
BICENE+ betaine
BICENE+100 mM NaCI
BICENE+200 mM NaCI
BICENE+500 mM NaCI
BICENE+.5%Triton X
BICENE+.5%Triton X
BICENE+.5% Tween20
BICENE+1.5%
Tween20
pH
7.5
8
8.5
6
5.5
6.5
6
7.5
7
7.5
8
8.5
8
5.5
6
6
6.5
7
7.5
6.6
6.6
410/280
0.13
0.08
0.18
.11
.1
.1
.07
.14
.1
.22
.25
.27
.22
.29
.18
.29
.2
.26
.17
.15
.15
7.5
8.5
8.5
8.5
8.5
8.5
8.5
8.5
8.5
.17
.18
.15
.16
.16
.01
.2
.21
.3
169
Table A.4: Conditions tested for Homologous expression
IPTG
60 pM
500 pM
1 mM
1mM
100 PM
500 pM
1 mM
1.5 mM
1 mM
1 mM
1mM
1 mM
600 pM
500 iM
1mM
5mM
10 mM
15 mM
170
Time of
induction
inoculation
inoculation
inoculation
1 day
inoculation
inoculation
inoculation
1 day
inoculation
inoculation
inoculation
inoculation
inoculation
1 day
1day
1 day
1 day
1 day
Induction
length
2 days
2 days
2 days
1 day
2 days
2 days
2 days
1 day
1 day
2 days
3 days
4 days
4 days
1 day
1day
1 day
1 day
1 day
Media
temp
H2
H2
H2
H2
H2+Fe-NTA
H2 +Fe-NTA
H2+Fe-NTA
H2+Fe-NTA
H2+Fe-NTA
H2+Fe-NTA
H2+Fe-NTA
H2+Fe-NTA
H2+Fe-NTA
H2
H2
H2
H2
H2
30
30
30
30
24
24
24
30
30
30
30
30
30
30
30
30
30
30
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