[FEFE]-HYDROGENASE SUBSTRATE TRANSPORT MECHANISMS AND INVESTIGATION
OF ALGAL HYDROGEN METABOLISM
By
Adam Cornish

A DISSERTATION
Submitted to
Michigan State University
In partial fulfillment of the requirements
For the degree of
DOCTOR OF PHILOSOPHY
Biochemistry and Molecular Biology
2012

ABSTRACT
[FEFE]-HYDROGENASE SUBSTRATE TRANSPORT MECHANISMS AND
INVESTIGATION OF ALGAL HYDROGEN METABOLISM
By
Adam Cornish
The global population has recently exceeded 7 billion people and the demand for
energy continues to expand as the number of industrialized countries grows. Currently,
the energy economy is dominated by the utilization of polluting and non-renewable fossil
fuels. Both the collection and use of petroleum-based fuels is destructive to the
environment and is not sustainable over a long time-scale, which justifies the
investigation into the development of renewable, alternative fuels. Of the various fuels
that have been proposed, molecular hydrogen (H2) in particular holds great promise as
a clean-burning fuel capable of supplementing the current energy economy, especially
as the combustion of H2 generates only water vapor as by-product. H2 can be
generated via a number of chemical processes, but current H2 technologies either
require fossil fuels as inputs or are energy-inefficient. The biological production of H2,
however, has garnered a great deal of interest because microorganisms are able to
drive H2 synthesis using energy derived from both light and dark fermentative
metabolisms. This manner of production does not depend on mining non-renewable
resources and microbes can be cultured at the industrial scale without competing with
arable land needed for agriculture.
H2 evolution in these microorganisms is dependent on nitrogenases and/or
hydrogenases, enzymes which utilize unique metal centers for catalysis. Hydrogenases

have been of particular interest for industrial-scale H2 production because these
enzymes are found in a diverse array of organisms and require only protons and
electrons as substrates. In particular, [FeFe]-hydrogenases have very high turnover
numbers and catalysis can be coupled to photosynthesis. Unfortunately, these enzymes
are inactivated by molecular oxygen (O2), and a number of studies have therefore
attempted to engineer O2-tolerant hydrogenases. However, engineering enzymes to
introduce optimal qualities has been impeded by an incomplete understanding of the
overall reaction mechanism.
Substrate (protons, electrons, and H2) transport is essential to hydrogenase
activity, yet relatively little information is available regarding the intraprotein transport of
substrate in [FeFe]-hydrogenase. I focused my investigation on identifying and testing
pathways important for substrate transport between the enzyme surface and the active
site in the Clostridium pasteurianum [FeFe]-hydrogenase. I have elucidated a key
pathway for proton transport and confirmed that two iron-sulfur clusters are essential in
an electron transfer relay, contributing to the overall characterization of [FeFe]hydrogenase activity.
Green algae utilize [FeFe]-hydrogenases to catalyze H2 production using
reducing equivalents derived from photosynthesis and these enzymes are an integral
component of anaerobic metabolism in these microalgae. I explored the H2 production
capabilities of a multicellular green alga, Volvox carteri, and characterized two
hydrogenases likely responsible for this activity. In addition, a unique hydrogenase gene
cluster discovered within the Volvox carteri genome provided interesting hints into the
origin of [FeFe]-hydrogenase in green algae.

To my mother, father, and sister, for supporting me through
both the good and the bad times.
To Michael, for being my partner in life,
and for loving duckies.

iv

ACKNOWLEDGEMENTS

First and foremost, I would like to thank my excellent mentor, Dr. Eric Hegg, for
his guidance and encouragement throughout the course of my PhD studies. It has been
his great ideas, his willingness to let me fail when I needed to, and his ability to set me
back on the right path that I have arrived to this point in my career. Thank you, Eric.
I want to thank all past and current members of the hydrogenase team. Many
thanks to Katrin for her enthusiasm and technical skills. Thanks to Saundra for helping
me grow as mentor and for all of the coffee, and to Robin for being so passionate about
science. Thanks to Travis, who is far more responsible for his scientific development
than I am, and a big thank you to Adam T., for learning what a 1:1000 dilution is.
I also have to express my appreciation for the entire Hegg lab and their
persistent ability to ignore what I say and understand what I mean. Emily, Hui, Vaidy
and others, you’ve made this lab a great place to work.
I’d like to give a very special thank you to Dr. Changcheng Xu, who first
encouraged me in the world of scientific research, and Dr. Christoph Benning, who
continues to put out papers with my name on them.
Thank you to all of my science friends (Dorothy Tappenden, Chris Buehl, and
Eric Carter) and my “real-life” friends (Kristi Tilford, Tiffany Leusby, and everyone at
Animosity). You made my time outside of lab so much more sane.
I give sincere thanks to the members of my guidance committee: Drs.Robert
Hausinger, Charles Hoogstraten, Gemma Reguera, and Shelagh Ferguson-Miller. Their
advice and direction were integral to my success through my PhD.

v

Most of all, I want to thank the Department of Biochmemistry and Molecular
Biology, as well as Michigan State University, for giving me the opportunity to study and
grow in a fantastic learning environment.

vi

TABLE OF CONTENTS

LIST OF TABLES…………………………………………………………………………...x
LIST OF FIGURES………………………………………………………………………....xi
LIST OF ABBREVIATIONS……………………………………………………………....xii
CHAPTER 1
INTRODUCTION…………………………………………………………………………. 1
A Sustainable Fuel Economy…………………………………………………… 2
Hydrogen in Biology……………………………………………………………… 4
Industrial Hydrogen Production………………………………………………… 7
Hydrogen-Forming Enzymes…………………………………………………… 9
The [FeFe]-hydrogenase Active Site…………………………………………... 14
Substrate Transport……………………………………………………………… 18
[FeFe]-hydrogenase Gene Evolution and Horizontal Gene Transfer ……... 23
Thesis Goals……………………………………………………………………… 26
References………………………………………………………………………... 28
CHAPTER 2
INVESTIGATING THE MECHANISM OF PROTON TRANSFER IN
[FEFE]-HYDROGENASE FROM CLOSTRIDIUM PASTEURIANUM……………… 43
Introduction……………………………………………………………………….. 44
Methods…………………………………………………………………………… 48
Multiple Sequence Alignment…………………………………………… 46
Plasmid Construction……………………………………………………. 46
Site-Directed Mutagenesis……………………………………………… 46
Cell Growth and Induction………………………………………………. 50
Protein Purification……………………………………………………….. 50
Hydrogenase Activity Assays…………………………………………… 51
Azide Rescue Assay…………………………………………………….. 51
Zn2+ Inhibition Assay…………………………………………………….. 52
Activation Energy Assays……………………………………………….. 52
Results…………………………………………………………………………….. 53
Proton Pathway Identification…………………………………………… 53
Amino Acid-Substituted Enzyme Studies……………………………… 55
Chemical Rescue of Variant Enzyme Activity by Sodium Azide……. 59
Zn2+-based Inhibition…………………………………………………….. 61
Discussion………………………………………………………………………… 65
References………………………………………………………………………... 71

vii

CHAPTER 3
AN ELECTRON TRANSFER RELAY IN THE CLOSTRIDIUM PASTEURIANUM
[FEFE]-HYDROGENASE……………………………………………………………….. 77
Introduction……………………………………………………………………….. 78
Methods…………………………………………………………………………… 82
Cell Growth, Induction, and Purification……………………………… 83
Circular Dichroism……………………………………………………….. 84
Hydrogen Uptake Activity Assays……………………………………… 84
Hydrogen Evolution Activity Assays……………………………………. 85
Results…………………………………………………………………………….. 86
Enzyme Purification and Circular Dichroism Spectroscopy…………. 86
H2 Uptake Assays………………………………………………………... 89
H2 Evolution Assays………………………………………………………93
Discussion………………………………………………………………………… 95
References………………………………………………………………………... 98
CHAPTER 4
CHARACTERIZATION OF HYDROGEN METABOLISM IN THE
MULTICELLULAR GREEN ALGA VOLVOX CARTERI……………………………... 101
Introduction……………………………………………………………………….. 102
Methods…………………………………………………………………………… 105
Growth Conditions……………………………………………………….. 105
Anaerobic Acclimation ………………………………………………….. 105
Hydrogen Production Measurements………………………………….. 105
RNA Isolation……………………………………………………………... 106
RT-PCR…………………………………………………………………… 107
Cloning of HYDA1 and HYDA2…………………………………………. 107
Overexpression, Protein Purification, and Hydrogen
Evolution Assay…………………………………………………………... 107
Phylogenetic Analysis…………………………………………………… 109
Results…………………………………………………………………………….. 110
Hydrogen Evolution Studies…………………………………………….. 110
Fermentative H2 Production…………………………………………….. 113
Transcript Analysis………………………………………………………. 113
Hydrogenase Cloning and Characterization…………………………... 115
Phylogenetic Analysis…………………………………………………… 115
Discussion………………………………………………………………………… 121
References………………………………………………………………………... 125
SUPPLEMENTAL CHAPTER 1
DISCOVERY AND IDENTIFICATION OF NOVEL [FEFE]-HYDROGENASE
GENES …………………………………………………………………………………… 131
Introduction……………………………………………………………………….. 132
Methods…………………………………………………………………………… 134
Phylogenetic Tree Construction………………………………………… 134
Multiple Sequence Analysis…………………………………………….. 135
viii

Degenerate Primer-Based PCR Amplification………………………… 135
Cloning of Gene Fragments…………………………………………….. 135
Gene Fragment Identifaction…………………………………………… 136
Whole Genome Amplification…………………………………………… 136
Sequence Extension…………………………………………………….. 137
Results…………………………………………………………………………….. 138
Targeted Degenerate Primer Design…………………………………... 138
General Degenerate Primer Design and Testing…………………….. 142
Investigation of G. ahangari 234 Genomic DNA……………………… 142
Investigation of Environmental Genomic DNA Samples from YNP… 144
Full-Length Gene Sequence Determination…………………………... 149
Phylogenetic Analysis of Gene Fragments……………………………. 151
Discussion………………………………………………………………………… 154
References………………………………………………………………………... 159
SUPPLEMENTAL CHAPTER 2
GENOME ANNOTATION AND INVESTIGATION OF H2 METABOLISM OF
NANNOCHLOROPSIS OCEANICA CCMP1779……………………………………... 162
Introduction……………………………………………………………………….. 163
Methods…………………………………………………………………………… 165
Manual Annotation of Genome…………………………………………. 165
Growth Conditions……………………………………………………….. 165
Hydrogen Evolution Assay……………………………………………… 165
Results…………………………………………………………………………….. 167
Manual Gene Annotation………………………………………………... 167
Hydrogen Evolution……………………………………………………… 174
Discussion………………………………………………………………………… 176
References………………………………………………………………………... 179

ix

LIST OF TABLES

Table 2.1 Primer pairs used for site-directed mutagenesis PCR of the
C. pasteurianum HydA gene in the pAC-BAD vector………………………………… 49
Table 2.2. Measured properties of native and variant enzymes……………………. 60
Table 3.1. Primer pairs used for site-directed mutagenesis PCR of the
C. pasteurianum HydA gene in the pAC-BAD vector………………………………… 82
Table. 3.2. Secondary structure analysis of native and variant enzymes………….. 87
Table 3.3. Native and variant enzyme kobs

rel

values……………………….……….. 90

Table 3.4. Kinetic values of native and variant enzymes……………………………. ..92
Table 4.1. List of primers used for cloning and cDNA amplification………………... 108
Table S1.1. Table of Clade-Specific [FeFe]-hydrogenase Degenerate Primers….. 139
Table S1.2. Table of General [FeFe]-hydrogenase Degenerate Primers………….. 143
Table S1.3. Table of Gene Fragment DNA Sequences………………………………145
Table S2.1 Manually annotated genes in N. oceanica CCMP1779………………… 168

x

LIST OF FIGURES

Figure 1.1. Structures of the active sites found in the three different classes of
hydrogenases…………………………………………………………………………….. ..11
Figure 1.2: Proposed mechanism for H-cluster assembly by the maturation
proteins HydE, HydF, and HydG……………………………………………………….. ..16
Figure 1.3. Putative proton transport pathway………………………………………... ..19
Figure 1.4. Electron transfer relay of the C. pasteurianum [FeFe]-hydrogenase….. 22
Figure 2.1. [FeFe]-hydrogenase structure and putative proton transfer pathway.... ..46
Figure 2.2. Multiple sequence alignment……………………………………………......54
Figure 2.3. Relative reduction in hydrogenase activity of proton pathway
variants……………………………………………………………………………………. ..56
Figure 2.4. Two potential proton transfer relays identified via analysis of the
C. pasteurianum [FeFe]-hydrogenase crystal structure……………………………… ..58
Figure 2.5. Inhibition plots of the effect of ZnCl2 on the [FeFe]-hydrogenase
from C. pasteurianum as a function of the concentration of methyl viologen……… ..62
Figure 2.6. Inhibitory effects of ZnCl2 on activity of native and variant enzymes…... 64
Figure 3.1. Iron-sulfur clusters and cluster domains in [FeFe]-hydrogenases…….. ..80
Figure 3.2. CD spectra of native and variant enzymes………………………………. ..88
Figure 3.3. Relative reduction in hydrogenase activity of singly- and
doubly-substituted variants……………………………………………………………… ..91
Figure 4.1. Partial multiple sequence alignment of V. carteri and C. reinhardtii
HYDA amino acid sequences…………………………………………………………... 111
Figure 4.2. H2 evolution rate of aerobically- or anaerobically-acclimated
V. carteri cells…………………………………………………………………………….. 112
Figure 4.3. Gene transcript accumulation observed by RT-PCR…………………… 114
Figure 4.4. [FeFe]-hydrogenase operon-like gene cluster in V. carteri.................... 116

xi

Figure 4.5. Phylogenetic analysis of representative HYDA homologs……………... 118
Figure 4.6. [FeFe]-hydrogenase gene clusters observed in V. carteri,
F. nodosum, F. pennivorans, and T. thermarum……………………………………… 120
Figure S1.1. Multiple sequence analysis of both clade-specific and
general hydrogenase proteins………………………………………………………….. 140
Figure S1.2. Multiple sequence analysis of gene fragment and
[FeFe]-hydrogenase sequences………………………………………………………... 148
Figure S1.3. Gene sequence extension methodology……………………………….. 150
Figure S1.4. Evolutionary relationship of translated gene fragments and
[FeFe]-hydrogenases……………………………………………………………………. 152
Figure S2.1. [FeFe]-hydrogenase operon-like gene cluster in
N. oceanica CCMP1779.......................................................................................... 173
Figure S2.2. H2 evolution rate of aerobically- or anaerobically-acclimated
N. oceanica CCMP1779 cells…………………………………………………………... 175

xii

LIST OF ABBREVIATIONS

AA

amino acid

BLAST

basic local alignment search tool

BSA

bovine serum albumin

BV

benzyl viologen

CD

circular dichroism

CpI

Clostridium pasteurianum [FeFe]-hydrogenase

DdH

Desulfovibrio desulfuricans [FeFe]-hydrogenase

Fed

distal iron of [FeFe]-hydrogenase H-cluster

DTMA

dithiomethylamine

DTME

dithiomethylether

Ea

activation energy

EDTA

ethylenediaminetetraacetic acid

EPR

electron paramagnetic resonance

ESEEM

electron spin echo envelope modulation

Hmd

H2-forming methylene-H4MPT dehydrogenases

Hox

oxidized form of the [FeFe]-hydrogenase

Hred

reduced form of the [FeFe]-hydrogenase

HYSCORE

hyperfine sub-level correlation

kcat

catalytic rate constant

kDa

kilodalton

Ki

inhibition constant
xiii

rel

kobs

normalized rate of activity

LB

lysogeny broth

NCBI

National Center for Biotechnology Information

NTA

nitriloacetic acid

methenyl-H4MPT

+

methenyltetrahydromethanopterin

methylene-H4MPT

methylenetetrahydro-methanopterin

MV

methyl viologen

PCR

polymerase chain reaction

PETF

plastidic ferredoxin

PSI

photosystem I

PSII

photosystem II

QM/MM

quantum mechanical and molecular mechanics calculations

ROS

reactive oxygen species

RT-PCR

reverse transcription polymerase chain reaction

SAM

S-adenosyl methionine

SVM

standard Volvox medium

YNP

Yellowstone National Park

xiv

CHAPTER 1

Introduction

1

A Sustainable Fuel Economy

In June of 2012, the Arctic lost ~1 million square miles of ice (four times the area
of Texas), which is the largest loss of ice in June in the satellite record (1). That month
also marked the June with the highest globally-averaged land surface temperature on
record (1.93 ˚F above the average) , and these are only two of many climate-related
records being broken in 2012 (2). A strong correlation can be deduced between the
temperature reports, the disappearance of arctic ice mass, and the fact that the United
States this year alone has released ~7 million metric tons of carbon dioxide (CO2) into
the atmosphere (3). Climate change is a reality and, recognizing that the rate of
greenhouse gas emissions is increasing with each passing year (3), alternative energy
options need to be investigated and pursued.
Currently, primary global fuel sources are petroleum-derived products (gasoline,
diesel, jet fuel, etc), and the processes of extraction, preparation, and combustion of
these products have negative impacts upon the global environment (4, 5). Even if fuels
made from petroleum were environmentally safe to process and utilize, petroleum is still
a finite resource—every gallon of gasoline burned is a gallon that has been converted
into CO2 and is contributes an additional burden onto the carbon cycle. Petroleum
resources must be mined as current stores are consumed, and while isolating crude oil
is a process that can reap large rewards (high yields of oil), it also comes at great risk
(high yields of oil into the ocean), as exemplified by the Deepwater Horizon oil spill of
2010. Unfortunately, the economic costs of oil spills do not always directly affect the
2

price of gasoline at the pump, and this price is often misleading, because it does not
take into account public health or environmental risks. If these hidden costs were
included in the price of gas, the cost per gallon could be far greater than it is currently
(6, 7, 8).
Natural gas, coal, and ethanol are a few of the available alternative energy
carriers that supplement petroleum-based fuels (9, 10). Unfortunately, these fuels share
similar problems with crude oil and petroleum, especially as natural gas and coal
generate CO2, a potent greenhouse gas, upon combustion. Natural gas and coal must
still be mined and extracted from the earth, often through hydraulic fracking (natural
gas) and mountaintop removal mining (coal), all of which cause intense geological
stress, damaging the environment, and often contaminating groundwater (11, 12, 13).
The production of ethanol has a less immediate impact upon the environment, but does
compete with local agriculture, as the majority of ethanol produced is derived from
staple crops such as potatoes, sugar cane, and corn (14). Although “cleaner” in some
ways than petroleum, these fuels also struggle with environmental impacts and longterm sustainability.
Another alternative fuel is diatomic hydrogen (H2), which has many ideal
properties and has the potential to supplement currently existing transportation fuels. H2
is the most abundant molecule in the solar system, has a high energy density per unit
mass, and generates only water as a by-product of combustion (Reaction 1).
Unfortunately, the light weight of H2 allows it to escape Earth’s gravity, resulting in
atmospheric concentrations of roughly only 1 ppm H2, yet atomic hydrogen is extremely
abundant on the planet’s surface in the form of hydrocarbons and water. Steam
3

reforming of either natural gas or coal is currently the most common method for
collecting H2 but requires extremely high temperatures, generates carbon monoxide
(CO) as a by-product, and has the other aforementioned problems associated with
natural gas (15). Several methods of direct electrolysis can be used to generate
gaseous O2 and H2 from liquid water without the formation of hazardous by-products,
but implementation of this technology is impeded by high costs and expensive material
requirements (16, 17). Microorganisms also have the capability to generate H2 without
the use of fossil fuels and many of these organisms are able to couple the fermentation
of sugars or energy derived from sunlight to H2 synthesis (18, 19). Significantly, this
biological production of H2 does not require arable land, thus avoiding competition with
modern agriculture (20) and providing rationale for the use of biological H2 production.

H2 + 1/2 O2 → H2O(g)

(Reaction 1)

Hydrogen in Biology

The role of H2 in organismal metabolism must be understood before utilizing
these microorganisms for large-scale H2 production. Although H2 exists at ~1 ppm, it is
often an important component of the microbial lifecycle (21, 22). On early Earth,
relatively high concentrations of H2 were likely present and this small molecule could
have played an integral part in many beginning biological functions (23). Today, H2
serves an essential function in the metabolism of many microbes, acting as both an
energy carrier and an endpoint of proton reduction, with recent evidence suggesting that
4

H2 may also have a role in pathogen survival during infection (22, 24).
A variety of bacteria and cyanobacteria produce H2 as an obligate by-product of
nitrogen (N2) fixation (21, 25, 26). Catalyzed by nitrogenase, the theoretical
stoichiometry of the reaction involves consumption of 16 ATP molecules for every two
molecules of ammonia and one molecule of H2 produced (Reaction 2), but reactions
utilizing more ATP or generating greater than stoichiometric amounts of H2 have been
reported. (27, 28, 29, 30). Due to the oxygen-sensitivity of nitrogenase, some
filamentous cyanobacteria form heterocysts—microoxic differentiated cells—allowing for
both nitrogen fixation and H2 formation under aerobic conditions (21, 31).

+

-

N2 + 8 H + 8 e + 16 ATP → 2 NH3 + H2 + 16 ADP + 16 Pi

(Reaction 2)

Under anaerobiosis, H2 formation can be coupled to sugar fermentation, to
anoxygenic photosynthesis, or to the oxidation of metals (22, 32, 33). To sustain ATP
and NADH production under microoxic conditions, microbes degrade organic molecules
via anaerobic respiration, generating electrons which are usually funneled into the
respiratory electron chain (34). Lacking the typical terminal electron acceptor (O2),
excess electrons derived during anaerobiosis can be channeled into proton reduction
(Reaction 3) to generate H2 (35). This process eliminates buildup of largely negative
reduction potentials, effectively decreasing formation of damaging radical species.

+

-

2 H + 2 e → H2

5

(Reaction 3)

Anaerobic protists and pathogenic fungi are also able to evolve H2, coupling
proton reduction to ATP synthesis utilizing mitochondria-like organelles called
hydrogenosomes (36, 37, 38). Under anaerobiosis, these organelles produce H2 during
the conversion of pyruvate to CO2 and acetyl-CoA, ultimately leading to ATP formation
via succinyl coenzyme A synthetase (39). Hydrogenosome-containing organisms
release H2 and acetate as by-products, which can then be utilized by other microbes in
the community.
H2 molecules diffuse easily through cell membranes, representing a loss or gain
of electrons as H2 passes between microbes and carrying electrons with them. A
number of organisms utilize H2 uptake enzymes under anaerobiosis to gain electrons to
drive anabolic processes. Cyanobacteria are able to take up H2 to regain “wasted”
electrons that were channeled into H2-production during N2 fixation, effectively recycling
electrons for further ammonia production (40, 41). Bacteria and algae which evolve H2
during fermentation often grow syntrophically with a variety of organisms (e.g., Knallgas
bacteria) which are then able to take up exogenous H2, utilizing the electrons from H2 to
power CO2 fixation and basic anabolic metabolism (42, 43, 44).
H2 uptake can also be important in the conversion of CO2 to methane in
methanogenic archaea, where the H2 often acts as an integral component in multiple
steps of the process. The decomposition of H2 to protons is fundamental in driving a
+

+

+

Na /H antiporter, which then regenerates the Na gradient essential to methyl-group
transfer (45). Electrons derived from H2 are used to reduce ferredoxin, and these
electrons are then donated to generate the first compound in methane synthesis,
formyl-methanofuran (46). Essential to intermediary steps, hydrides from H2 are added
6

to

both

methenyltetrahydromethanopterin

+

(methenyl-H4MPT )

and

methylenetetrahydro-methanopterin (methylene-H4MPT) (45, 47), further demonstrating
the important role of H2 in methanogen metabolism.
Pathogenic bacteria have been shown to utilize H2 to increase survival during
infection. To colonize the intestine, pathogens must first endure the low pH levels (pH 22.5) of acid in the stomach. Recently, Escherichia coli has been shown to deacidify
highly acidic media via proton uptake and subsequent reduction to H2, effectively
increasing the local pH and enhancing acid resistance (48). H2 metabolism may also be
important in the response of pathogenic bacteria to oxidative stress, antibiotic
treatments, and macrophage attack. Microbial and cellular H2 production has been
linked with resistance to oxidative stress induced by antibiotics and human immune
cells (24, 49, 50), especially in regards to OH•, a highly cytotoxic ROS (51).

Industrial Hydrogen Production

Biological H2 production has the potential to supplement and bolster current
technologies using renewable and/or non-polluting raw materials. A broad range of
microbes (e.g., green algae, cyanobacteria, sugar-reducing thermophilic bacteria) are
able to couple anaerobic photosynthetic or fermentative processes to H2 production
(22). For industrial biological H2 production, ideal microorganisms would need to be
amenable to large-scale culturing using nutrient-rich waste water (52, 53). At the end of
their lifecycles, H2-producing organisms can be harvested for agricultural feedstocks or
used in the manufacture of other biofuels, such as bioethanol or biodiesel (54, 55).
7

Finally, a biological system selected for industrial H2 production needs to require few
material inputs, utilize genetically tractable organisms, and be able to produce large
quantities of H2.
One popular approach for large-scale H2 production involves supplying
anaerobically-cultured bacteria with inexpensive fermentable carbon sources. A variety
of sugars can be utilized to drive H2 formation during fermentation, principally
depending on the identity of the microbe (22, 56, 57). Glycerol is one candidate to feed
into fermentative H2-production, as this 3-carbon product is an obligate waste material
generated during biodiesel refinement, which has increased the availability of glycerol
and sharply decreased its price on the global market (58, 59). Another candidate is
lignocellulosic biomass (e.g., corn stover), which is a popular substrate within the
industrial agricultural complex, providing another source of sugars which could be used
for fermentative biohydrogen production (60, 61), although bioethanol remains the
principal focus of cellulosic biofuel research (62).
Another approach to biological H2 production is the use of microalgae—
organisms which utilize photosynthesis to either directly or indirectly drive H2 synthesis.
A standard method for this synthesis involves alternating between light and dark cycles,
allowing these organisms to undergo oxygenic photosynthesis—fixing CO2 and storing
reducing equivalents as starch—followed by anaerobic respiration and concomitant
production of H2 (19, 63). Unfortunately, all H2-synthesizing enzymes are extremely O2sensitive and require anaerobic conditions for maximal activity, which presents
difficulties during oxygenic photosynthesis (35, 64). To circumvent this O2 sensitivity,
sulfur deprivation has been utilized in the green alga Chlamydomonas reinhardtii,
8

eliminating photosystem II activity and allowing non-oxygenic photosynthesis to drive H2
production under microoxic conditions (65, 66).

Hydrogen-Forming Enzymes

As already discussed, organisms have developed a large toolset for the
production of H2 from metabolic, organic, and photosynthetic sources. Key enzymes in
these reactions are nitrogenase and hydrogenase, enzymes which utilize unique
organometallic active sites to catalyze the production of H2 (19). A detailed understanding of the enzymes, metabolic processes, and organisms involved in generating
molecular hydrogen is essential to build and sustain efficient biological H2-synthesis
technologies.
Nitrogenases are complex metalloenzymes which form H2 as a catalytic byproduct during N2 fixation and, like all H2-forming enzymes, nitrogenases are
inactivated by O2. There are three primary classes of nitrogenase based on the active
site composition, but the molybdenum (Mo)-nitrogenase is the best characterized of
these enzymes. The iron (Fe) protein of Mo-nitrogenase is a dimer that is bridged by a
[4Fe4S] cluster, and each subunit contains one MgATP-binding site. The Monitrogenase is composed of two Fe proteins and the tetrameric α2β2 molybdenum iron
(MoFe) protein, which contains two P-clusters ([Fe8S7]) and two FeMoco ([MoFe7S9C])
cofactors per nitrogenase (27, 28). Recent targeted substitution experiments have
generated nitrogenase variants which still evolve H2 but lack N2-fixation activity,
effectively separating these two processes from one another (67).
9

Hydrogenases are enzymes which catalyze both proton reduction and H2
oxidation at an organometallic active site (22, 32, 68). Protons for this reaction are
supplied by water, whereas the electrons needed for reduction are derived from
fermentation or photosynthesis and are shuttled to the hydrogenase via a variety of
physiological carriers (e.g., ferredoxin, NAD(P)H, cytochrome c3) (22, 69, 70, 71, 72).
As noted previously, these enzymes are rapidly inactivated by O2 and are therefore
primarily active in organisms during anaerobic metabolism (73, 74, 75). The catalytic
turnover of H2 in hydrogenase can be ~1000-fold greater than that of nitrogenase and,
importantly, hydrogenases are ATP-independent and can be directly linked to
photosynthesis, providing a rationale for using these enzymes in industrial-scale H2
production (18). Like nitrogenases, hydrogenases are classified according to the metal
composition of their respective active sites: [FeFe], [NiFe], and [Fe] (22, 76, 77, 78, 79).
[FeFe]-hydrogenases are bidirectional enzymes, although preferential H2
evolution has been observed for some organisms in vivo (80). The [FeFe]-hydrogenase
active site (H-cluster) is composed of a cysteine-coordinated [4Fe4S] cubane cluster
connected by a proteinaceous thiolate ligand to a unique diiron sub-cluster ([2Fe]H)
-

(Figure 1.1A). This [2Fe]H is further ligated by two cyanide (CN ) and three carbon
monoxide (CO) molecules, as well as a non-proteinaceous dithiolate ligand, and proper
assembly of the cluster requires three accessory proteins (76, 77). Encoded by the
hydA gene, this class of enzyme is primarily encoded in the genomes of sulfate- or
metal-reducing bacteria, hyperthermophilic bacteria, and parasitic anaerobes. Genes
encoding hydrogenase are also found in eukaryotic genomes, such as green algae,
anaerobic protozoans, and fungi (22, 81).
10

Figure 1.1. Structures of the active sites found in the three different classes of
hydrogenases. A) Structural representation of the [FeFe]-hydrogenase active site,
emphasizing the diiron cluster and associated ligands. The identity of X is proposed to
be a nitrogen atom, although oxygen and carbon atoms remain possibilities. B)
Structural representation of the [NiFe]-active site coordinated by four proteinaceous
cysteine thiolates. C) Structural representation of the [Fe]-hydrogenase iron guanylyl
pyridone cofactor; a water molecule is modeled in the “open” coordination site of the
cofactor.

A)

B)

C)

11

[NiFe]-hydrogenases compose the largest group of known H2-reactive enzymes
and contain the greatest diversity of enzymes, many of which participate in H2
evolution, H2 uptake, and/or regulatory functions. The [NiFe]-hydrogenase active site is
best described as a nickel-iron center coordinated by four proteinaceous cysteine thiols,
-

with one CO and two CN molecules ligating the iron atom (Figure 1.1B) (78, 82, 83).
[NiFe]-hydrogenases are multimers, but the essential core structure is a heterodimer
composed of a large catalytic subunit and a smaller subunit containing three iron-sulfur
clusters involved in electron transfer to the active site (78, 84). [NiFe]-hydrogenases can
be further classified into five distinct subclasses according to protein sequence: I)
Membrane-localized H2-uptake enzymes involved in oxidizing H2 to feed electrons into
anaerobic respiration, IIa) Soluble uptake enzymes which are often associated with H2
recycling during nitrogen-fixation, IIb) H2-sensing enzymes which regulate the gene
expression of H2-uptake enzymes, III) Cytoplasmic bidirectional enzymes which are
+

NAD(P)H/NAD(P) -dependent,

and

IV)

Membrane-bound

H2-evolving

enzymes

composed of six or more subunits (22). Examples of each of these subclasses can be
found in a great variety of bacteria, whereas only subclasses IIa and III are utilized by
cyanobacteria, and archaeal hydrogenases only fit within subclasses I, III, and IV (22).
This great diversity of [NiFe]-hydrogenases is associated with increased complexity in
maturation, requiring a minimum of seven assembly proteins for proper active site
assembly and insertion (85).
[Fe]-hydrogenases are formally known as H2-forming methylene-H4MPT
dehydrogenases (Hmd) and catalyze reversible hydride transfer onto methenyl+

H4MPT . In nature, these hydrogenases are utilized by methanogenic archaea to
12

catalyze a step in the conversion of CO2 to methane, specifically the conversion of
+

methenyl-H4MPT to methylene-H4MPT (86). A three-dimensional crystal structure of
Hmd reveals a single catalytic iron atom that is at the center of an iron guanylyl pyridone
cofactor (FeGP cofactor). The iron atom is octahedral and is ligated by two CO
molecules, the nitrogen and an acyl carbon of a pyridinol ring, a cysteinyl thiolate, and
an “open” site (modeled with a water molecule) (Figure 1.1C) (79, 87). Two putative
maturases may be important for Hmd active site assembly, but the assembly
mechanism remains uncharacterized (88). Although of great interest for its role in CO2
reduction to methane, Hmd is not an ideal catalyst for large-scale H2 production, as the
H2 evolution activity would be dependent on methane oxidation (47, 86).
Phylogenetic analyses of the hydrogenase classes, in conjunction with active site
composition and coordination environment, strongly suggest distinct evolutionary origins
for all three enzymes (81). Additionally, the maturation machinery is specific to each
class, requiring 3, 7-13, or 2 active-site assembly proteins for [FeFe]-, [NiFe-], and [Fe]hydrogenase, respectively (22, 85, 88, 89). This finding lends further support to the
hypothesis that these enzymes are the result of convergent evolution, explaining the
existence of unique properties observed between the enzyme classes (81).
A great deal of research has focused on the [FeFe]- and [NiFe]-hydrogenases for
large-scale production of H2, as the only direct substrates for this reaction are protons
and electrons. In comparison to [FeFe]-hydrogenase, a greater diversity of [NiFe]hydrogenases has been discovered, including relatively O2-insensitive enzymes like the
[NiFeSe]-hydrogenase, which replaces an active site cysteinyl ligation with a selenium
atom (90). Still, most of these [NiFe]-hydrogenases are not ideal for H2 formation due to
13

a catalytic bias for H2 oxidation over proton reduction under physiological conditions
(19, 91). [NiFe]-hydrogenase active site assembly is also very complex and many of
these hydrogenases are membrane-bound (22, 85). Conversely, [FeFe]-hydrogenases
have some properties which are ideal for H2 production. For instance, the maximal
catalytic turnover of several characterized HydA proteins is 2-3 orders of magnitude
greater than the highest-producing [NiFe]-hydrogenases (91, 92), and the maturation of
[FeFe]-hydrogenase is comparatively simple (93). Additionally, [FeFe]-hydrogenases are
soluble proteins that can be readily expressed, characterized, and used industrially
without the concerns that accompany membrane-bound enzymes (94, 95, 96).

The [FeFe]-hydrogenase Active Site

The active sites of [FeFe]-hydrogenases from Clostridium pasteurianum (CpI)
and Desulfovibrio desulfuricans (DdH) are remarkably similar when observed by x-ray
crystallography, but differ in coordination of the distal iron (Fed) (76, 77). The Fed is
proposed to be the site of proton reduction as the H-cluster cycles between oxidized
and reduced forms (Hox and Hred) (97, 98). In the CpI structure (presumed Hox state),
Fed is in an octahedral coordination environment with two carbon monoxide ligands
(terminal and bridging), a terminal cyanide ligand, a terminally-bound water molecule,
and two thiolate sulfurs from the dithiolate ligand (76). The DdH structure (Hred state)
lacks the terminally-bound water molecule observed in the CpI Fed (octahedral
geometry), leaving open an a coordination site where H2 is proposed to bind (77, 99).
Due to limitations in x-ray crystallography, the identity of the bridging dithiolate ligand
14

was initially proposed to be propanedithiolate, but subsequent electron paramagnetic
resonance (EPR) experiments and density functional theory calculations indicate that
dithiomethylamine (DTMA) or dithiomethylether (DTME) are more likely (100, 101, 102).
Utilizing either a bridging nitrogen or oxygen atom (DTMA and DTME, respectively), this
dithiolate ligand could be involved in proton transfer to the distal iron site (102, 103), as
proposed by quantum mechanical and molecular mechanics (QM/MM) simulations (104,
105). These observations indicate that the unique [FeFe]-hydrogenase active site is
ideally poised to participate in the reversible reduction of protons to H2.
The [2Fe]H of the H-cluster is an organometallic complex that does not form
spontaneously and requires assembly before insertion into the apo-enzyme (89). A suite
of maturation proteins (HydE, HydF, and HydG) are essential in the synthesis of [2Fe]H,
although the mechanism of this assembly and insertion remains to be fully elucidated
(Figure 1.2). Homologous to GTPases, HydF both hydrolyzes GTP and acts as a
scaffold for [2Fe]H biosynthesis, although this activity is not dependent on GTPhydrolysis (106, 107). HydG, a radical S-adenosyl methionine (SAM) enzyme, catalyzes
-

the conversion of L-tyrosine to CO and CN ligands for attachment to the diiron cluster
(108, 109, 110). Similar to HydG, HydE also has radical SAM enzyme activity and,
although the role of this enzyme in [2Fe]H assembly remains to be elucidated, HydE is
hypothesized to be involved in synthesis of the bridging dithiolate ligand (111, 112). A
recently solved crystal structure reveals a surface-exposed channel for insertion of
[2Fe]H and shows that the active site [4Fe4S] cluster is present before insertion (93,
106, 112). [FeFe]-hydrogenase activity is dependent on proper insertion and assembly
of the [2Fe]H, as no appreciable activity is observed without a complete H-cluster (113,
15

114).

16

Figure 1.2. Proposed mechanism for H-cluster assembly by the maturation proteins HydE, HydF, and HydG. The
HydF protein is proposed to act as a scaffold, ligating a [2Fe2S] cluster which will form the core of the [2Fe]H. HydE is
proposed to synthesize and attach the dithiolate ligand, while HydG has been shown to catalyze the synthesis of the CO
and CN ligands. Both HydE and HydG can associate with HydF and attach the ligands to [2Fe]H in any order, although
only one assembly factor binds to HydF at a time. Once the H-cluster is fully synthesized, the active site can be
transferred to the apo-hydrogenase by HydF.

17

The H-cluster of HydA is irreversibly inactivated in the presence of O2, effectively
eliminating hydrogenase activity under aerobiosis, although some reports indicate O2insensitivity when in the Hox state, possibly due to full-occupancy of the distal iron
coordination sphere (115, 116). The primary hypothesis for this inactivation is
attachment of O2 to the distal iron, leading to the formation of ROS, which then can
directly attack the cubane cluster, culminating in complete degradation of the active site
(64, 73, 117). DFT calculations have modeled several gas channels within HydA for H2
and O2 diffusion, which could be targets for engineering to decrease O2-based
inactivation (118, 119). Supporting this rationale, previous investigation of a [NiFe]hydrogenase indicate that narrow gas channels may contribute to O2-insensitivity, while
still allowing for diffusion of the much smaller H2 molecule (120). Unfortunately,
subsequent mutagenesis studies have not succeeded in significantly improving
insensitivity to O2 for either [NiFe]- or [FeFe]-hydrogenases (75).
In addition to O2-inactivation, [FeFe]-hydrogenase can be reversibly inhibited by
several other small molecules, providing more methods to probe the mechanisms of
hydrogenase activity. A potent inhibitor of H2 uptake activity, CO tightly binds to the
distal iron of the H-cluster, occluding both H2- and O2-binding and rendering CO-bound
HydA insensitive to O2-based inactivation (64, 121, 122). CO is often used to model O2
diffusion in the enzyme, as it is a small diatomic molecule that can be labeled (123). In
contrast to CO, formaldehyde specifically inhibits H2 evolution activity more strongly
than H2 uptake activity and is thought to bind the partially reduced H-cluster at either
the distal iron or dithiolate bridge, preventing proton-transfer (124, 125). In addition to
diatomic gaseous molecules, divalent metal cations (e.g., Cd
18

2+

,Cd

2+

2+

, Hg , Zn

2+

)

have also been reported to impair hydrogenase activity (126, 127). Sequestered ~20 Å
within the enzyme, the active site metal atoms are unlikely to be exchanged with these
cations, and, while early reports suggested that this inhibition was specific to electron
transfer (126).

Substrate Transport

[FeFe]-hydrogenase activity is dependent on both the intermolecular and
intramolecular transport of protons, electrons, and H2. While transport of these
substrates is essential to enzymatic activity, the majority of [FeFe]-hydrogenase
research has focused on either optimizing catalytic turnover or understanding and
decreasing O2-based inactivation (118, 128, 129, 130). Research focused on
engineering improved hydrogenases is of great importance, but this work has been
hindered by an incomplete understanding of substrate transport mechanisms within the
enzyme.
When the first two crystal structures of [FeFe]-hydrogenases were solved, two
potential proton transfer pathways were hypothesized based on amino acid sequence
conservation, hydrogen-bonding capabilities, and relative distances within the structure
(131). Intramolecular protein transfer is typically achieved via hydrogen-bonding
networks between protonatable amino acid side-chains and water residues (132, 133).
Recently, QM/MM studies on the CpI and DdH hydrogenases provided computational
evidence that residues of a putative proton pathway (Figure 1.3) could accept protons
from solvent and may form a hydrogen-bonding network extending from the enzyme
19

surface to the active site (104). To determine the absolute role of these residues in
proton transfer, however, direct experimental evidence is required.

20

Figure 1.3. Putative proton transport pathway. A theorized proton transfer relay
composed of four conserved amino acids and a modeled water molecule that connect
the enzyme surface to the H-cluster. For interpretation of the references to color in this
and all other figures, the reader is referred to the electronic version of this dissertation.

21

Intermolecular

transport

of

electrons

to/from

[FeFe]-hydrogenase

is

accomplished through interactions with physiological electron carriers, and these
interactions are dependent on the nature of the hydrogenase. [FeFe]-hydrogenases can
differ in the number of both subunits and iron-sulfur cluster-binding domains, as
described in detail by Meyer (81). Sulfate- and metal-reducing bacteria donate electrons
from cytochrome c3 to dimeric hydrogenases to mediate formation of a chemiosmotic
gradient in the periplasm (134, 135, 136). The Acetobacterium woodii bifurcating
tetrameric hydrogenase couples H2 oxidation to the reduction of NADH and ferredoxin
in a 1:1 ratio (overcoming the endergonic H2-based ferredoxin reduction) and effectively
feeding energy into CO2-fixation (72). In green algae, photosynthetically-derived
electrons are transferred to the monomeric hydrogenase from a plastidic ferredoxin
(PETF), directly linking photosynthesis to H2 production (71). The interactions between
hydrogenase and PETF have been investigated computationally, and the results
suggest that the association is mediated largely by electrostatic interactions (137).
Similar to algal hydrogenases, monomeric bacterial hydrogenases also accept electrons
from ferredoxin to drive H2 production during dark fermentation (138). The interplay
between physiological electron donors and H2-reactive enzymes appears to be
dependent on the structural organization of the hydrogenase, and may imply a positive
correlation between subunit composition, number of iron-sulfur clusters, and reactivity
with electron carriers.
Intramolecular electron transfer in [FeFe]-hydrogenases is presumed to be
accomplished via electron transfer relays composed of iron-sulfur clusters and the relay
present in M3 HydA architecture is the most prevalent among sequenced
22

hydrogenases. The basic M3 architecture is hypothesized to constitute a functional
electron transfer relay (76) and is composed of four iron-sulfur cluster-binding domains
(F-domains) that are observed in ~75% of sequenced [FeFe]-hydrogenases. A [2Fe2S]
cluster and two [4Fe4S] clusters are ligated by a typical four cysteine thiolatecoordination in three F-domains (designated FS2, FS4A, and FS4B, respectively). The
fourth domain (FS4C) uses a histidine and triple cysteine linkage to coordinate a third
cubane cluster, which is similar to the distal iron-sulfur cluster in the [NiFe]-hydrogenase
electron transfer relay (139). Intriguingly, the proposed electron transfer relay in the C.
pasteurianum enzyme branches past FS4B, potentially terminating at either the FS4C
or the FS2 cluster (Figure 1.4). Earlier reports have hypothesized that these two
clusters could serve to modulate the reduction potential of the enzyme for interaction
with electron carriers, provide specific binding-sites for electron donors, or act as
different electrons “routes” for proton reduction and H2 oxidation (131). Due to the
prevalence of this putative relay in [FeFe]-hydrogenases, the characterization of its role
in hydrogenase activity is essential to understanding the complete catalytic mechanism.
Molecular H2 transfer to and from the H-cluster is a passive action and does not
require direct mediators. Molecular dynamics studies have modeled H2 diffusion within
the enzyme and have indicated multiple potential routes which often overlap with
putative O2 diffusion pathways (140). A single permanent H2 channel is unlikely to exist,
as transient cavities are formed on the nanosecond timescale while the protein
undergoes dynamic motion, allowing H2 molecules to diffuse through the enzyme (119).
Recently, mutagenesis studies were utilized to investigate putative channels in the
enzyme, but no definitive routes of H2 diffusion were confirmed (141).
23

Figure 1.4. Electron transfer relay of the C. pasteurianum [FeFe]-hydrogenase. The
electron transfer pathway is composed of three [4Fe4S] clusters and a [2Fe2S] cluster.

24

Recent investigations have targeted individual [FeFe]-hydrogenase amino acid
residues via substitution to identify roles in O2 sensitivity, but have had limited success.
In particular, Lautier et al. (118) substituted residues within putative gas channels to
occlude O2 from the active site. These substitutions either had no effect on oxygensensitivity or sharply impaired hydrogenase activity. Although intriguing, these results
are inconclusive in regards to hydrogenase activity without a greater understanding of
substrate transport, especially as both studies substituted key residues previously
implicated in proton transfer. The elucidation of the proton and electron transport
mechanisms is essential in building a comprehensive model for hydrogenase activity,
which can then be used for future enzyme engineering and optimization.

[FeFe]-hydrogenase Gene Evolution and Horizontal Gene Transfer

[FeFe]-hydrogenase genes are observed in a broad range of bacteria, green
algae, protists, and fungi (22, 81). Prokaryotic in origin, genes encoding the catalytic
hydrogenase and assembly proteins (hyd genes) are hypothesized to have been
acquired by eukaryotes via horizontal gene transfer (35, 81, 95, 142, 143). As gene
acquisition via multiple specific gene transfer events into a single genome is
improbable, the suite of hydrogenase and maturase genes in eukaryotes was likely
acquired as an operon or operon-like cluster from a prokaryotic ancestor. Multiple hyd
gene clusters can be observed in the genomes of modern-day bacteria (81), providing
further evidence for this hypothesis.
Among eukaryotes with known [FeFe]-hydrogenases, green algae contain HYD
25

proteins with the greatest deviation in protein structure from bacterial homologs. First
observed in the C. reinhardtii genome, the hydE and hydF genes (bacterial
nomenclature) are absent and are replaced by a fusion gene, HYDEF (algal
nomenclature) (143). Heterologous expression studies indicate that HYDEF synthesized
in a bacterial system is suboptimal for hydrogenase maturation (144), whereas C.
reinhardtii HYDA can be fully activated using the bacterial HydE, HydF, and HydG suite
of proteins (145, 146). This may suggest that, although the mechanism of [2Fe]H
assembly and insertion could differ subtly between organisms, there is probably little
discrimination at the level of [FeFe]-hydrogenase identity in relation to maturation.
Most known green algal HYDA proteins lack the F-domains which ligate ironsulfur clusters proposed to participate in intramolecular electron transfer (147, 148). The
physiological significance of this truncation is unknown, although green algal
hydrogenase catalytic turnover is among the highest recorded (149). In addition, HYDA
is localized to the chloroplast in green algae and accepts electrons from PETF, a
ferredoxin associated with photosynthesis (71), suggesting that hydrogenases lacking
the N-terminal F-clusters may be optimally poised for accepting photosyntheticallyderived electrons. Among green algae studied for hydrogenase activity, only the
Chlorella variabilis HYDA proteins contain F-domains (150), which may indicate either
recent acquisition of the genes or minimal gene restructuring. This question can be
further explored once the C. variabilis genome has been sequenced, as the presence of
the HYDEF fusion gene and/or clustering of hyd genes within the genome could indicate
evolutionary relationships.
Within most green algae studied for H2 metabolism at the protein level, two
26

separate [FeFe]-hydrogenases are present, HYDA1 and HYDA2 (151). A recent report
provides evidence of gene duplication, which could explain the presence of these two
highly homologous genes (150), although single acquisition of two hydrogenase genes
from a gene cluster is still possible. Intriguingly, a unique amino acid domain is
conserved among most green algal HYDA sequences, but is absent in C. variabilis
HYDA sequences, further emphasizing the disparity between these proteins and other
green algal hydrogenases.
[FeFe]-hydrogenases have high amino acid sequence homology to nuclear
prelamin A recognition factor (NARF) proteins, which are essential for cytoplasmic ironsulfur cluster biosynthesis (in contrast to mitochondrial synthesis) (36, 152, 153). Known
as IOP1/NARFL/NARF in Homo sapiens and Arabidopsis thaliana and Nar1p in
Saccharomyces cerevisiae, no functional homologs have been observed in prokaryotic
systems (152). Phylogenetic analysis of NARF proteins and [FeFe]-hydrogenases
indicates a strong evolutionary relationship between these two enzymes, although
NARF lacks a catalytic H-cluster and has not been reported to demonstrate H2production or -uptake activity (152). Based on sequence similarity, NARF essentially
mimics the catalytic domain of HydA and utilizes four cysteine residues to ligate a
[4Fe4S] cluster that is analogous to the cubane of the H-cluster. A second [4Fe4S]
cluster is ligated by a FS4A-like domain, and both of these clusters in NARF have been
shown to be essential for protein function and cellular viability (154). Although several
HydA-specific motifs are altered in NARF, the extent of amino acid substitution is not
constant between species, providing further evidence that these proteins evolved from a
common HydA ancestor. As a gene that is divergent from hydA, NARF is an excellent
27

outgroup during evolutionary analysis of hydrogenase genes and can be used as a root
in phylogenetic trees.
The origin of NARF proteins has been hypothesized to be linked to the evolution
of hydrogenosomes—organelles that couple H2 production to ATP synthesis (36).
Recent comparison of hydrogenosomal and mitochondrial DNAs suggests that these
organelles are the product of a single endosymbiotic event, with rapid differentiation
occurring before the roles of the respective organelles were fully established (38, 153).
A unifying metabolic process between hydrogenosomes and mitochondria is the
organellar synthesis of iron-sulfur clusters (38, 155, 156). An especially interesting
feature observed in hydrogenosome-containing organisms is that, while hydA genes are
present, no NARF proteins are encoded within either the nuclear or the
hydrogenosomal genomes. These observations may indicate that the original
endosymbiont—hypothesized to be an α-proteobacteria (37)—contained a hydA gene,
which either changed in function during the organellar transition to a mitochondrion or
retained H2-evolving functionality within the hydrogenosome (91).

Thesis Goals

As the availability of fossil fuels diminishes and demand for energy continues to
rise, an alternative to unsustainable 20

th

century energy technologies is required.

The biological production of H2 holds great promise as a source of renewable and clean
fuel. Industrial-scale levels of H2 are needed to supplement both petroleum-based and
alternative fuels in the global economy, and many important questions remain regarding
28

H2-catalyzing enzymes and the organisms which utilize these proteins.
[FeFe]-hydrogenases have tremendous potential to contribute to the large-scale
synthesis of H2. A vital component of the enzymatic activity is intramolecular substrate
transport, yet a detailed mechanism of this transport remains to be described. To
elucidate and confirm pathways essential for proton (Chapter 2) and electron transport
(Chapter 3) within the C. pasteurianum [FeFe]-hydrogenase, I have generated aminoacid substituted variant enzymes and provided strong evidence for the roles of
individual residues in substrate transport activities.
Green algae are intriguing photosynthetic organisms which have the potential to
be utilized for industrial-scale biofuel production, especially in the generation of
biohydrogen. The multicellular green alga Volvox carteri, a close relative of C.
reinhardtii, has been intensely studied at the level of cell differentiation and
sexual/asexual reproduction, yet there remains a knowledge gap regarding anaerobic
metabolism in V. carteri. To provide insight into this metabolism (Chapter 4), I have
investigated H2-evolution activity in the organism, characterized the enzymes
responsible for this activity, and identified a unique gene cluster that may provide hints
into the evolutionary origins of hydrogenase within modern-day green algae.

29

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Cohen J., Kim K., Posewitz M., Ghirardi M. L., Schulten K., Seibert M., King P. (2005).
Molecular dynamics and experimental investigation of H2 and O2 diffusion in [Fe]hydrogenase. Biochem. Soc. Trans. 33(Pt 1), 80-82.

141.

Lautier T., Ezanno P., Baffert C., Fourmond V., Cournac L., Fontecilla-Camps J. C.,
Soucaille P., Bertrand P., Meynial-Salles I., Leger C. (2011). The quest for a functional
substrate access tunnel in FeFe hydrogenase. Faraday Discuss 148, 385-407; discussion
421-341.

142.

Horner D. S., Heil B., Happe T., Embley T. M. (2002). Iron hydrogenases--ancient
enzymes in modern eukaryotes. Trends Biochem. Sci. 27(3), 148-153.

143.

Posewitz M. C., King P. W., Smolinski S. L., Smith R. D., Ginley A. R., Ghirardi M. L.,
Seibert M. (2005). Identification of genes required for hydrogenase activity in
Chlamydomonas reinhardtii. Biochem. Soc. Trans. 33(Pt 1), 102-104.

144.

Posewitz M. C., King P. W., Smolinski S. L., Zhang L., Seibert M., Ghirardi M. L.
(2004). Discovery of two novel radical S-adenosylmethionine proteins required for the
assembly of an active [Fe] hydrogenase. J. Biol. Chem. 279(24), 25711-25720.

145.

Kuchenreuther J. M., Grady-Smith C. S., Bingham A. S., George S. J., Cramer S. P.,
Swartz J. R. (2010). High-yield expression of heterologous [FeFe] hydrogenases in
Escherichia coli. PLoS One 5(11), e15491.

146.

Sybirna K., Antoine T., Lindberg P., Fourmond V., Rousset M., Mejean V., Bottin H.
(2008). Shewanella oneidensis: a new and efficient system for expression and maturation
of heterologous [Fe-Fe] hydrogenase from Chlamydomonas reinhardtii. BMC Biotechnol
42

8, 73.
147.

Happe T., Kaminski A. (2002). Differential regulation of the Fe-hydrogenase during
anaerobic adaptation in the green alga Chlamydomonas reinhardtii. Eur. J. Biochem.
269(3), 1022-1032.

148.

Winkler M., Heil B., Happe T. (2002). Isolation and molecular characterization of the
[Fe]-hydrogenase from the unicellular green alga Chlorella fusca. Biochim. Biophys. Acta
1576(3), 330-334.

149.

Kamp C., Silakov A., Winkler M., Reijerse E. J., Lubitz W., Happe T. (2008). Isolation
and first EPR characterization of the [FeFe]-hydrogenases from green algae. Biochim.
Biophys. Acta 1777(5), 410-416.

150.

Meuser J. E., Boyd E. S., Ananyev G., Karns D., Radakovits R., Narayana Murthy U. M.,
Ghirardi M. L., Dismukes G. C., Peters J. W., Posewitz M. C. (2011). Evolutionary
significance of an algal gene encoding an [FeFe]-hydrogenase with F-domain homology
and hydrogenase activity in Chlorella variabilis NC64A. Planta 234(4), 829-843.

151.

Forestier M., King P., Zhang L., Posewitz M., Schwarzer S., Happe T., Ghirardi M. L.,
Seibert M. (2003). Expression of two [Fe]-hydrogenases in Chlamydomonas reinhardtii
under anaerobic conditions. Eur. J. Biochem. 270(13), 2750-2758.

152.

Hackstein J. H. (2005). Eukaryotic Fe-hydrogenases -- old eukaryotic heritage or
adaptive acquisitions? Biochem. Soc. Trans. 33(Pt 1), 47-50.

153.

Shiflett A. M., Johnson P. J. (2010). Mitochondrion-related organelles in eukaryotic
protists. Annu. Rev. Microbiol. 64, 409-429.

154.

Urzica E., Pierik A. J., Muhlenhoff U., Lill R. (2009). Crucial role of conserved cysteine
residues in the assembly of two iron-sulfur clusters on the CIA protein Nar1.
Biochemistry 48(22), 4946-4958.

155.

Lill R., Kispal G. (2000). Maturation of cellular Fe-S proteins: an essential function of
mitochondria. Trends. Biochem. Sci. 25(8), 352-356.

156.

Tachezy J., Sánchez L. B., Müller M. (2001). Mitochondrial type iron-sulfur cluster
assembly in the amitochondriate eukaryotes Trichomonas vaginalis and Giardia
intestinalis, as indicated by the phylogeny of IscS. Mol. Biol. Evol. 18(10), 1919-1928.

43

CHAPTER 2

Investigating the mechanism of Proton Transfer in [FeFe]-hydrogenase from
1
Clostridium pasteurianum

1

This chapter is adapted from Cornish A. J., Gärtner K., Yang H., Peters J. W., Hegg E.
L. (2011). Mechanism of proton transfer in [FeFe]-hydrogenase from Clostridium
pasteurianum. J. Biol. Chem. 286(44), 38341-38347.
43

INTRODUCTION

Hydrogenases are found in a diverse array of microorganisms where they mainly
function either to a) reduce protons utilizing electrons accumulated during fermentation
or b) couple the oxidation of H2 to energy-yielding reactions (1). Hydrogenases are
separated into three subclasses based on the composition of their active sites: [FeFe]-,
[NiFe]-, and [Fe]-hydrogenase (2). [FeFe]-hydrogenases demonstrate the highest in
vitro catalytic turnover among hydrogen-producing enzymes, and there is great interest
in elucidating the enzymatic mechanism of these hydrogenases (3, 4). Current research
has primarily focused on examining the catalytic activity (5, 6), active site assembly (7,
8, 9, 10), and irreversible oxygen inactivation of these enzymes (11, 12, 13), but
relatively few data are available concerning intramolecular transport of substrates
between the active site and the enzyme surface (14, 15, 16). Proton transfer is an
essential component of reversible hydrogen production by [FeFe]-hydrogenases, and
biochemical investigation is required for a complete understanding of the enzymatic
mechanism.
Two structures of [FeFe]-hydrogenases have been solved by crystallography
(Clostridium pasteurianum (17, 18) (Figure 2.1A) and Desulfovibrio desulfuricans (19)),
and there is considerable structural homology near the active site. This active site (Hcluster) is best described as a [4Fe4S] cubane cluster attached via a cysteinyl sulfur
linkage to a diiron moiety. Each iron is coordinated by a terminal cyanide and a carbon
monoxide ligand, and an additional CO bridges the two irons (17, 20). A five-atom
dithiolate ligand bridges the diiron moiety (Figure 2.1B). The exact identity of this ligand

44

has not been determined through direct chemical or biochemical analysis, but this
ligand has been proposed to be dithiomethylamine (DTMA) because of the close
proximity of the amine group to the presumed H2 oxidation site, and the ability of amines
to cycle between protonation states at physiological pH (21). The assignment of an
amine group at this position is consistent with the results of a recent electron spin echo
envelope modulation (ESEEM) and hyperfine sub-level correlation (HYSCORE)
spectroscopic study (22). Direct experimental results addressing whether an amine`
group within the dithiolate ligand could cycle between protonation states in the enzyme,
however, is still lacking.

45

Figure 2.1. [FeFe]-hydrogenase structure and putative proton transfer pathway. A, Crystal structure of the [FeFe]hydrogenase from C. pasteurianum (3C8Y www.pdb.org) resolved to 1.39 Å. The protein is a monomer separated into a
catalytic domain containing the [6Fe6S] H-cluster and three N-terminal domains containing three [4Fe4S] clusters and a
[2Fe2S] cluster. B, Chemical structure of the H-cluster emphasizing the diiron active site. X is generally postulated to be
nitrogen, although oxygen and carbon remain possibilities. C, Putative proton transfer relay inferred by multiple sequence
alignment analysis and examination of the crystal structure of the [FeFe]-hydrogenase from C. pasteurianum. Hcluster→C299→H2O612→E279→S319→E282.

46

Intramolecular transfer of substrates is necessary for efficient catalysis at the
active site, which is sequestered ~20 Å within the interior of the enzyme. Accessory
iron-sulfur clusters provide pathways to facilitate the transport of electrons between the
protein surface and the H-cluster (23, 24). Rationalizing pathways for bidirectional
proton transfer, however, is less intuitive. In previous work, it has been observed that
proton exchange within proteins can be facilitated via negatively charged side-chains
and water molecules (25, 26, 27). Understanding this key mechanism in [FeFe]hydrogenases is critical for targeted protein engineering, as perturbing proton transfer
can cause unintended deleterious effects on the catalytic rate (28). To date, detailed
experiments investigating proton transport within [FeFe]-hydrogenases have not been
reported.
In this chapter, we provide biochemical evidence that four residues (Cys299,
Glu279, Ser319, and Glu282) in the [FeFe]-hydrogenase from C. pasteurianum are
important for hydrogenase activity and are likely to be involved in proton transfer
between the active site and the enzyme surface. The strict conservation of these
residues among this class of hydrogenases suggests a ubiquitous proton transport
pathway. In addition, we have observed that Zn
hydrogenase specifically targets this proton pathway.

47

2+

-based inhibition of [FeFe]-

METHODS

Multiple Sequence Alignment—Using the C. pasteurianum [FeFe]-hydrogenase as the
base sequence, we performed Position-Specific Iterated Basic Local Alignment Search
Tool (BLAST) (29) to identify an additional 62 unique amino acid sequences. These 63
[FeFe]-hydrogenase sequences (acquired from National Center for Biotechnology
Information [NCBI]) were examined for side-chain conservation using ClustalX (30).

Plasmid Construction—The coding sequence of the [FeFe]-hydrogenase from C.
pasteurianum was amplified by Polymerase Chain Reaction (PCR) and ligated into the
NcoI/SacI sites of pAC-BAD (a modified pBAD/D-TOPO vector from Invitrogen lacking
the N-terminal thioredoxin tag) containing a kanamycin resistance cassette and an Larabinose inducible promoter. Constructs were transformed into DH5α Escherichia coli
competent cells and selected for resistance to 50 μg/mL kanamycin.

Site-directed Mutagenesis—Specific amino acid codons in the [FeFe]-hydrogenase
gene from C. pasteurianum were mutated by site-directed mutagenesis PCR (pfu turbo,
Stratagene) using appropriately designed primers (Table 2.1) (31). Segregation of
mutated DNA from wild-type DNA was achieved by DpnI digestion of methylated DNA.
After isolation of mutated DNA from E. coli, the sequences were confirmed and the
constructs transformed into S. oneidensis MR-1 ΔhydA/ΔhyaB electrocompetent cells
as described by Ozawa (32) and selected for resistance to 50 μg/mL kanamycin.

48

Table 2.1. Primer pairs used for site-directed mutagenesis PCR of the C.
pasteurianum HydA gene in the pAC-BAD vector

49

Cell Growth and Induction—A 0.5 mL inoculum of overnight transformant S. oneidensis
MR-1 ΔhydA/ΔhyaB culture was transferred to 50 mL of 50 μg/mL kanamycinsupplemented lysogeny broth (LB) in a 250-mL Erlenmeyer flask and was shaken at
200 rpm at 30 °C until reaching an OD600 of 0.4 (Eppendorf BioPhotometer). The entire
culture was transferred into 500 mL of 50 μg/mL kanamycin-supplemented LB in a 2-L
Erlenmeyer flask and shaken at 30 °C until reaching an OD600 of 0.4. The culture was
then transferred to a 2-L round-bottom flask, supplemented with ammonium iron citrate
to a final concentration of 100 μM, and sparged with Ar to remove ambient O2. After 30
min, 0.5 mL of 1.3 M L-arabinose (final concentration 1.3 mM) was added and the
culture was sparged for an additional 30 min. The flask was then sealed with a rubber
septum and shaken at 200 rpm at 30 °C for an additional 16 h.

Protein

Purification—All

steps

for

hydrogenase

purification

were

performed

anaerobically. Cells were collected by spinning the culture in airtight 250-mL Nalgene
tubes at 2,800× g for 20 min. Inside an anaerobic Coy Chamber, the cell pellet was
resuspended in 7.0 mL of Ni-NTA wash buffer (100 mM Tris-HCl [pH 8.0], 200 mM
NaCl, 5.0 % glycerol) supplemented with 10 mM sodium dithionite. The resuspended
cell pellet was transferred to 1.7 mL Eppendorf tubes in 1.0 mL aliquots and sonicated 8
times for 4 s each at a power setting of 4 (Fischer Scientific Sonic Dismembrator Model
100). The sonicated cell suspension was cleared of cell debris by centrifugation at
20,000× g in airtight 40-mL tubes. A column containing 1.0 mL Ni-nitriloacetic acid
(NTA) agarose resin (Qiagen) was first equilibrated with 20 mL of wash buffer
supplemented with 10 mM sodium dithionite to remove residual O2, and then the
50

supernatant was passed over the column. The column was washed with 15 mL of wash
buffer supplemented with 20 mM imidazole. The protein was eluted from the column in
1.0 mL fractions by increasing the concentration of imidazole to 100 mM.

Hydrogenase Activity Assays—Hydrogen evolution and uptake assays were modified
from King et al. (33). Briefly, H2 evolution was measured by incubating 0.1 mL of protein
sample in 1.9 mL of H2 evolution assay buffer (50 mM HEPES [pH 7.0], 500 mM NaCl,
100 mM sodium dithionite, 10 mM methyl viologen (MV)) in a 13 mL serum vial at 25 °C
with continuous shaking. A 100 µL syringe was used to inject 50 μL of headspace gas
into a TRACE GC Ultra Gas Chromatograph (Thermo Scientific) and H2 accumulation
was measured over time by plotting the peak area against a standard curve. To
measure H2 consumption, 0.1 mL of protein sample was incubated in 1.9 mL of H2
uptake assay buffer (50 mM Tris-HCl [pH 8.0], 10 mM benzyl viologen (BV), 0.2% triton
x-100) in a 13 mL serum vial at 25 °C with 2.5% H2 in the headspace and shaken
continuously. Uptake of H2 was measured over time by injecting 50 μL of the
headspace into the gas chromatograph and plotting the peak area against a standard
curve.

Azide Rescue Assay—To determine the effect of sodium azide on hydrogenase activity,
H2 evolution activity of native and variant enzymes was measured (see Hydrogenase
Activity Assays) in the presence of different amounts of sodium azide (0, 50, 100, 250
mM).

51

Zn

2+

Inhibition Assay—Zn

2+

inhibition assays were performed measuring H2 evolution

activity (see Hydrogenase Activity Assays) in a 2.0 mL total volume. Because ZnCl2
only dissolves in the H2 evolution assay buffer below pH 6.3, 0.02 mL of the buffer (pH
2+

6.3), with or without dissolved ZnCl2, was added to each assay. The effect of Zn

on

the native enzyme was assayed by measuring H2 evolution activity over several ZnCl2
concentrations (0, 0.05, 0.125, 0.25. 0.50 mM) as the concentration of methyl viologen
was varied (3.0, 5.0, 8.0, 10.0 mM) and the data were graphed on both Dixon and
Cornish-Bowden plots. The effect of ZnCl2 on the activity of amino acid-substituted
[FeFe]-hydrogenases was assayed at either 0 or 0.50 mM ZnCl2.

Activation Energy Assays—H2 evolution and uptake assays were performed as
previously noted over a range of temperatures (0˚C, 8 ˚C, 25 ˚C, 30 ˚C, 35 ˚C, and 40
˚C). The natural log of kobs was plotted against the reciprocal of the temperature. The
resulting slope was multiplied by the negative value of the gas constant R (8.314 J K-1
mol-1) to calculate the activation energy (Ea).

52

RESULTS

Proton Pathway Identification—Based on their crystallographic analysis, a putative
proton transfer relay composed of Cys299, a modeled water, Glu279, Ser319, and
Glu282 that connect the enzyme surface to the H-cluster of the C. pasteurianum [FeFe]hydrogenase was initially proposed by Peters et al. (17). Reasoning that residues
involved in proton transport should be highly conserved, we performed multiple
sequence alignment on 63 unique [FeFe]-hydrogenases (Figure 2.2). Our results
revealed that these four residues are strictly conserved among both algal and bacterial
[FeFe]-hydrogenase genes. In comparison, these residues are not conserved among
Nar1 proteins, a class of protein that share a number of sequence motifs with [FeFe]hydrogenases (8, 34). Together, these data strongly implicate the importance of these
four residues to hydrogenase function.

53

Figure 2.2. Multiple sequence alignment. Partial multiple sequence alignment (ClustalX) of [FeFe]-hydrogenases from
C. pasteurianum, D. desulfuricans, Chlamydomonas reinhardtii, Thermotoga maritima, Trichomonas vaginalis,
Shewanella oneidensis, and Neocallimastix frontalis. Amino acids predicted to participate in proton transfer are denoted
by asterisks. Strictly conserved residues are in a black box while highly conserved residues are in bold. Residues with low
conservation but chemically similar side-chains are boxed. Every tenth amino acid of the C. pasteurianum sequence is
denoted by a black dot.

54

Amino Acid-Substituted Enzyme Studies—The coding sequence of the putative proton
transport residues was mutated by site-directed mutagenesis PCR in the [FeFe]hydrogenase gene from C. pasteurianum. N-terminal His-tagged wild-type and mutated
genes were overexpressed anaerobically in a ΔhydA/ΔhyaB strain of S. oneidensis
MR-1 and the resulting enzymes purified by Ni-NTA affinity chromatography. The kcat of
native C. pasteurianum hydrogenase H2 evolution activity was calculated to be ~1,000
-1

s . When compared to native enzyme, all amino acid-substituted [variant] enzymes
demonstrated sharp decreases in hydrogenase activity (Figure 2.3). Interestingly, in all
variant enzymes, the hydrogen evolution activity is depressed to a greater extent than
the hydrogen uptake activity.
In close proximity to the H-cluster, Cys299 is an ideal candidate for donating
protons to the dithiolate ligand at the active site. Substitution of this residue by alanine
rel

or serine strongly decreased kobs

[specific activity of variant enzyme relative to native

enzyme] for both evolution and uptake activities. Substitution by serine retained slightly
more hydrogenase activity, which may be due to the fact that the serine side-chain
preserves a hydrogen-bonding hydroxyl group, while the alanine side-chain is aliphatic
and unable to participate in proton transfer.
The residues adjacent to Cys299 in the putative pathway, Glu279 and Ser319,
are also important for enzyme activity. When these residues were substituted, H2
uptake activity of the resulting enzymes was reduced to ~85% of kobs
activity was decreased by >99%.

55

rel

while evolution

Figure 2.3. Relative reduction in hydrogenase activity of proton pathway variants.
rel
kobs
(specific activity of variant enzyme relative to the specific activity of native
enzyme) values for H2 evolution and uptake activities (black and gray bars,
respectively). To determine the effect of substitution, the specific activity of native
rel
enzyme was normalized to 1.0. Variant enzyme kobs
values were calculated from
individual experiments and averaged. Error bars denote one unit of standard deviation.
N=3-8.
Evolution
Uptake

56

At the enzyme surface, the Glu282 residue is solvent-exposed and ideally
positioned to exchange protons between Ser319 and bulk solvent. The residual activity
of Glu282 variant enzymes was dependent on the specific substitution. A greater
reduction in activity was observed in the E282L variant compared to the E282D variant,
especially with regard to H2 uptake. These results are consistent with the structural
differences of the side chains. Whereas aspartic acid and glutamic acid differ only in
side-chain length, leucine lacks a protonatable side chain.
To probe the apparent discrepancy noted between the percent reduction in H2
uptake vs. evolution activities in the amino acid-substituted hydrogenases, we
calculated the Ea for each direction for the native enzyme. Our results revealed
activation energies of 24 ± 3 kJ/mol (n = 5) and 45 ± 2 kJ/mol (n = 3) for H2 evolution
and uptake, respectively. The difference in the activation energies could partially explain
the apparent discrepancy in the uptake vs. evolution assays to amino acid substitutions.
A second putative proton transfer pathway can be imagined from the C.
pasteurianum hydrogenase crystal structure, starting at Cys299, passing through
several modeled water molecules and Ser298, and ending at the non-conserved Lys571
residue at the enzyme surface (Figure 2.4). However, when we substituted Ser298 to
alanine, the resulting enzyme had activity similar to the native enzyme (Figure 2.3),
indicating that this residue is not critical for activity.

57

Figure 2.4. Two potential proton transfer relays identified via analysis of the C.
pasteurianum [FeFe]-hydrogenase crystal structure. A. Residues composing
pathway examined in this paper (DTMA → Cys299 → H 2O612 → Glu279 → Ser319 →
Glu282). B. Residues composing alternate pathway (Cys299 → H 2O612 → H 2O605 →
Ser298 → H 2O594 → H 2O675 → H 2O668 → Lys571). However, Lys298 is not strictly
conserved, and our data reveal that Ser298 is not required for activity (Figure 2.3).

A

B

58

Chemical Rescue of Variant Enzyme Activity by Sodium Azide—To examine further the
role of the residues identified as being potentially involved in proton transfer, hydrogen
evolution activity was stimulated by the addition of a chemical recovery agent. Sodium
azide has been previously demonstrated to enhance specifically in vitro proton transfer
activity of variant enzymes deficient in proton transport (35, 36, 37). Importantly, the H2
evolution activity of the native enzyme was unaffected by the addition of sodium azide;
the chemical neither significantly promotes nor represses the hydrogenase activity. The
activities of C299 and E279 variants were not stimulated by addition of sodium azide,
possibly as a result of azide’s inability to penetrate into the interior of the protein. In
contrast, the H2 evolution activities of the S319A, E282D, and E282L variants were all
stimulated as the concentration of sodium azide was increased, providing additional
evidence that the S319 and E282 residues participate in proton transfer (Table 2.2).

59

rel

Table 2.2. Measured properties of native and variant enzymes. kobs values of
2+
evolution activities calculated as described in Figure 2.3. Zn -binding constants, Ki,
were determined for native and variant enzymes at 0.50 mM ZnCl2. Fold-stimulation
values are the ratio of enzyme activity with and without addition of 250 mM sodium
azide. N=3-8.

60

Zn

2+

-based Inhibition—Previous studies have revealed that certain divalent metal

cations inhibit [FeFe]-hydrogenase, but the specific mechanism for this inhibition was
not elucidated (38, 39). Importantly, divalent cations such as Zn

2+

have been previously

shown to inhibit intramolecular proton transfer in a variety of biomolecular reactions (27,
2+

40). To determine the inhibitory effect of Zn

on C. pasteurianum [FeFe]-hydrogenase

activity, we measured the H2 evolution activity over a range of ZnCl2 and MV
2+

concentrations. H2 evolution activity decreased as the Zn

concentration increased.

Kinetic analyses from Dixon and Cornish-Bowden plots indicated non-competitive mixed
inhibition (Figure 2.5), with nearly equal values of Ki and Ki´, 0.171 and 0.170 mM,
2+

respectively. These results indicate that Zn

likely binds with similar affinity to the

enzyme and substrate-bound enzyme and is not competing for binding with MV.
Addition of 0.50 mM CaCl2 or NaCl has only a negligible effect on hydrogenase activity
(data not shown), indicating that the inhibition is not due to chloride ions or a change in
ionic strength. When the Zn

2+

chelator ethylenediaminetetraacetic acid (EDTA), was

titrated into a reaction containing Zn

2+

, the enzyme activity was restored in proportion to

amount of chelator added. Together, these results clarify that Zn

2+

does not occlude

MV-binding to the C. pasteurianum [FeFe]-hydrogenase, and that Zn
enzyme activity by inhibiting proton transport.

61

2+

may depress

Figure 2.5. Inhibition plots of the effect of ZnCl2 on the [FeFe]-hydrogenase from
C. pasteurianum as a function of the concentration of methyl viologen. A. Dixon
plot: Concentration of ZnCl2 versus the reciprocal of velocity. B. Cornish-Bowden plot:
Concentration of ZnCl2 versus the concentration of substrate over velocity. The lines
from each data set intersect, indicating mixed inhibition. The Ki and Ki´ calculated from
each plot is 0.171 µM and 0.170 µM, respectively, indicative of noncompetitive mixed
inhibition.

A

B

62

Recognizing that Zn
2+

hypothesized that Zn

2+

has high affinity for negatively-charged residues, we

might inhibit [FeFe]-hydrogenase activity through coordination

to the surface-exposed Glu282 residue, thereby blocking proton transfer. To examine
more fully the effect of ZnCl2, native and variant enzymes were assayed for H2 evolution
activity in both the presence and absence of 0.50 mM ZnCl2. Addition of 0.50 mM ZnCl2
to the assay mixture results in a 70-80% reduction in relative H2 evolution activity for
native and most variant enzymes, while the activity of the E282D variant is depressed
by only ~40% (Figure 2.6). The binding constants (Ki) of Zn

2+

to the native and most

variant enzymes range between 0.1 mM and 0.4 mM (Table 2.2), with the exception of
E282L. This variant enzyme maintains the same level of activity independent of Zn

2+

concentration, with a binding constant an order of magnitude greater than that observed
in the native enzyme. These data indicate that the E282L substitution renders the
enzyme insensitive to Zn

2+

, providing experimental evidence that Zn

hydrogenases by binding to Glu282.

63

2+

inhibits [FeFe]-

Figure 2.6. Inhibitory effects of ZnCl2 on activity of native and variant enzymes.
Relative H2 evolution activity after addition of 0.05, 0.10, 0.25, or 0.50 mM ZnCl2.
Residual activity is relative to enzyme activity with 0 mM ZnCl2 (1.0). Error bars denote
one unit of standard deviation. N=3-8.

64

DISCUSSION

The activity of [FeFe]-hydrogenase is dependent on the transfer of protons
between the enzyme surface and the H-cluster. Active intramolecular proton transport is
an essential function for a variety of enzymes, and this process occurs via a protonhopping method (Grothuss mechanism) between water molecules and the side chains
of amino acid residues (26, 41, 42). Although hypothesized, this essential transfer
activity has not been characterized experimentally for [FeFe]-hydrogenases.
The C. pasteurianum [FeFe]-hydrogenase is a soluble 63 kilodalton (kDa)
periplasmic protein. Four iron-sulfur clusters, termed F clusters, are sequestered in the
three N-terminal domains, while the H-cluster is buried within the catalytic domain
(Figure 2.1A). From analysis of the crystal structure, the cysteine residue at position 299
has been proposed to act as a proton donor/acceptor for the H-cluster (17, 43).
When the structure of the C. pasteurianum hydrogenase was first determined, a
putative proton transfer pathway was envisioned involving residues within hydrogen
bonding-distance that link the H-cluster to the enzyme surface (17). This pathway
consists of Cys299, a water molecule, Glu279, Ser319, and Glu282 (24) (Figure 2.1C).
Importantly, we determined that these residues are strictly conserved, supporting the
viability of this pathway (Figure 2.2).
Recent theoretical work has examined this proton pathway via quantum
mechanical and molecular mechanics calculations (QM/MM) (44). As part of this study,
direct proton transport between the active site dithiolate ligand and a nearby cysteine
residue (analogous to Cys299) in the D. desulfuricans [FeFe]-hydrogenase was
65

calculated to be energetically favorable. Parallel simulations on the C. pasteurianum
hydrogenase demonstrate low energy barriers for proton transfer from Glu282 to
Glu279 and from Glu279 to Cys299 (44). This work therefore provides computational
evidence for a putative proton transfer relay in C. pasteurianum [FeFe]-hydrogenase,
and it establishes a basis for direct biochemical experimentation.
To investigate experimentally the role of Cys299, Glu279, Ser319, and Glu282 in
the C. pasteurianum hydrogenase, we generated amino acid substitutions of these
residues to limit hydrogen-bonding and, thus, proton transfer. Substitution of these
residues causes a dramatic reduction in both H2 uptake and evolution, indicating their
importance for hydrogenase activity. Based on the crystal structure, these residues are
unlikely to participate in electron transfer or significantly alter the redox potential of any
of the N-terminal [FeS] clusters (Figure 2.1A) (45). Substitution of a residue responsible
for coordinating either the FS2 or the FS4C [FeS] cluster (C34S and C98S,
respectively) has only a moderate effect on hydrogenase activity when MV is the
electron donor (data not shown). Additionally, the similarity between Ki values of the
native and proton pathway variant enzymes indicates that the reduction in kobs

rel

is not

due to structural changes. Finally, to examine the proton transfer activity of these
residues, sodium azide was used as a chemical rescue agent. Addition of sodium azide
stimulates the H2 evolution activity of the E282 and S319 variants, whereas the native
enzyme is not enhanced by azide (Table 2.2). Based on this stimulation of variant
enzyme activity by azide, the noted Zn

2+

insensitivity of the E282L variant, and

computational calculations (44), the most parsimonious explanation for the observed
reduction in activity of the variants is that proton transfer has been impaired.
66

Within hydrogen-bonding distance of the H-cluster’s dithiolate ligand, Cys299 is
an ideal candidate as the direct proton donor to the active site. Substitution of this
residue demonstrates a dramatic reduction in hydrogenase activity and matches
QM/MM predictions previously described (44). The C299S variant retains low level
activity and can be further inhibited by the addition of Zn

2+

, with a Ki similar in value to

the native enzyme. Together with its close proximity to the dithiolate ligand, these
results indicate that Cys299 could participate in proton transfer to the H-cluster.
In recent work examining H2 diffusion channels, Lautier et al. measured the
effect of changing the residue analogous to C299A in the [FeFe]-hydrogenase from
Clostridium acetobutylicum (C298A) (16). The reported H2 oxidation kcat value of the
C298A variant was 6-fold less than native enzyme, indicating that, under their assay
conditions, a significant decrease in activity could be noted, albeit to a lesser extent
than noted in our experiments (~20-fold decrease for H2 uptake). This difference in the
magnitude of reduced activity for the C299/C298 variant enzyme might be explained by
the very different reaction conditions used by Lautier et al. (16). Nevertheless, both
studies clearly indicate the importance of this residue for hydrogenase activity.
The solvent-exposed Glu282 residue is expected to act as a proton acceptor at
the protein surface. Substitution of this residue by either leucine or aspartate causes a
rel

decrease in kobs . Interestingly, whereas the E282D variant is susceptible to Zn

2+

-

based inhibition, the E282L variant is unaffected even at 0.50 mM ZnCl2. These results
are consistent with previous work which established that divalent metal cations can
specifically inhibit proton transfer via interactions with negatively-charged residue sidechains (27, 40). Together, our work suggests a mechanism for preferential binding of
67

Zn

2+

to the surface-exposed Glu282 residue, presumably occluding proton binding and

transfer.
rel

An apparent discrepancy was noted between the kobs (uptake) and
rel

kobs (evolution) of the variant enzymes, with H2 evolution activity more strongly
impaired by substitution relative to uptake activity (Figure 2.3). To investigate this
apparent discrepancy, we measured the activities of native and variant enzymes
(C299S, E279D, and E282D) with a variety of buffers, salt concentrations, and varying
rel

pH. No set of tested conditions resulted in kobs (uptake) being equal to
rel

kobs (evolution). Next, we determined the Ea of the native enzyme for both evolution
and uptake (24 ± 3 kJ/mol and 45 ± 2 kJ/mol, respectively). The fact that the two values
are different is not surprising because the forward and reverse directions of a reaction
will always have different Eas unless the free energies of the products and reactants are
identical. In this particular case, however, this difference is also impacted by the fact
that the H2 uptake and evolution reactions are performed under distinct assay
conditions (pH and redox partner).
rel

The noted discrepancy (kobs

of evolution versus uptake) is likely a result of this

disparity between the calculated Ea values. The effect of mutating a residue important
for hydrogenase activity is expected to increase the energy of the transition state by the
same magnitude for both directions of the reaction. Because H2 uptake has a higher
activation energy under our reaction conditions, however, the percent change in the Ea
between native and variant enzyme will be less for H2 uptake. Thus, one would predict
that mutations would have a smaller observed effect on the rate constant for H2 uptake,
which is exactly what we observe experimentally. Ultimately, because of the different
68

reaction conditions, these systems are not directly comparable, and the apparent
discrepancies therefore do not imply different proton pathways for H2 evolution and H2
uptake.
Hong et al. identified a second potential proton transfer pathway, composed of
Cys299, Ser298, five modeled water molecules, and Lys571 (44) (Figure 2.4). The
surface-exposed lysine, however, is not well-conserved among [FeFe]-hydrogenase
genes. In addition, our results indicate that the activity of the S298A variant enzyme is
not impaired under our reaction conditions relative to the native enzyme. Thus, this
potential pathway does not appear to be essential for proton transfer activity, although
its potential role as a substitute pathway under certain conditions cannot be dismissed.
The five crystallographically modeled water molecules described above are in
close proximity to the putative proton transfer pathway examined in this study. Thus, it
could be argued that the residues we have examined play only an indirect role in proton
transfer via interactions with this water channel. While theoretically possible, this
scenario seems unlikely given the insensitivity of hydrogenase activity to amino acid
substitutions of Ser298 and the importance of Glu282 to proton transport as determined
by Zn

2+

inhibition. Importantly, there does not appear to be any other chain of

crystallographically-observed water molecules in the vicinity of our putative proton
channel.
In conclusion, we demonstrated the importance of Cys299, Glu279, Ser319, and
Glu282 in the [FeFe]-hydrogenase from C. pasteurianum for activity. Together with a
crystallographically characterized water molecule, these residues form a potential
hydrogen bonding chain from the active site to the enzyme surface. Amino acid
69

substitution of any of these residues causes a severe reduction in hydrogenase activity.
In many cases, this activity could be partially restored by incubation with sodium azide,
providing additional evidence that these residues (Ser319 and Glu282) are important for
proton transfer. Substitution of the solvent-exposed Glu282 by leucine renders the
enzyme insensitive to ZnCl2, indicating that Zn

2+

inhibits enzyme activity by binding to

this residue and occluding the proton transfer pathway. Because Cys299, Glu279,
Ser319, and Glu282 are strictly conserved, these residues may constitute a key proton
transport relay in all [FeFe]-hydrogenases.

70

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71

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76

CHAPTER 3

An Electron Transfer Relay in the Clostridium pasteurianum [FeFe]-hydrogenase

77

INTRODUCTION

[FeFe]-hydrogenases catalyze the reversible reduction of protons to hydrogen
gas (H2) utilizing a unique catalytic [6Fe6S]-cluster (1, 2). Electrons for this reduction
are donated by intracellular carriers (e.g., ferredoxin) and are typically derived from
fermentation or photosynthesis (3, 4). Intermolecular electron transfer between various
carriers and [FeFe]-hydrogenases has been investigated (5, 6, 7, 8), but there remains
a paucity of information regarding intramolecular electron transfer. In [NiFe]hydrogenases, a series of iron-sulfur clusters has been reported to act cooperatively as
a relay to shuttle electrons between the active site and enzyme surface for H2
production/oxidation (9). Likewise, electron transfer in [FeFe]-hydrogenases also
presumably occurs through coordinated relays of iron-sulfur clusters, and understanding
this pathway is integral to enhancing [FeFe]-hydrogenase activity.
[FeFe]-hydrogenases can be classified based on the number of iron-sulfur
cluster-binding domains (F-domains) present in the N-terminal region of the protein.
Clusters ligated by these F-domains in the hydrogenase resemble ferredoxin clusters
and are either [2Fe2S] or [4Fe4S]-clusters (10, 11). Individual F-domains are
characterized by their binding to either [4Fe4S] or [2Fe2S] clusters, as well as the
coordination environment. The number of these domains can vary between different
[FeFe]-hydrogenases (12), except for green algal hydrogenases, which lack F-domains
and instead permit direct transfer between an electron carrier and the active site (1, 6).
The [FeFe]-hydrogenase from Clostridium pasteurianum has four F-domains, three of
which utilize a typical four-cysteine motif to ligate one [2Fe2S]-cluster and two [4Fe4S]78

clusters (FS2, FS4A, and FS4B, respectively). A fourth F-domain coordinates an
additional [4Fe4S]-cluster, which is ligated by three cysteines and a histidine (FS4C).
This overall domain structure is conserved among approximately 75% of known and
putative [FeFe]-hydrogenases (Figure 3.1A).
By examining the C. pasteurianum hydrogenase three-dimensional structure
(3C8Y, www.pdb.org), a putative electron transfer relay can be inferred between the
active site and medial FS4B From this cluster, the relay branches into two distal Fdomains (FS4C and FS2), preventing a definitive role for these two clusters in electron
transfer activity to be assigned (Figure 3.1B) (13). An identical electron transfer relay in
Clostridium acetobutylicum HydA has been shown to be functional during light-induced
H2 production experiments directly connecting the hydrogenase to photosystem I (14).
In addition, Brownian dynamics studies have indicated that ferredoxin could potentially
bind to positively-charged residues on the enzyme surface and pass electrons to the
FS4C (15). Without direct experimental evidence, however, the role of the FS4C and
FS2 clusters in electron transfer can only be surmised, and the prevalence of this
domain structure among [FeFe]-hydrogenases provides a rationale for investigating the
importance of these clusters.

79

Figure 3.1 Iron-sulfur clusters and cluster domains in [FeFe]-hydrogenases. A) Graphic illustration of iron-sulfur
cluster-binding domains and the amino acid motifs associated with each domain. B) Putative electron transfer of
Clostridium pasteurianum [FeFe]-hydrogenase (3C8Y (16)) composed of four iron-sulfur clusters between the catalytic
active site and the enzyme surface. Based on sequence homology, ~75% of sequence [FeFe]-hydrogenases are
predicted to have similar cluster organization.

A

80

To test the role of this relay in hydrogenase activity, we exchanged two cysteine
residues with serines (C98S and C34S) in the C. pasteurianum [FeFe]-hydrogenase to
alter the reduction potential of the two distal iron-sulfur clusters (FS4C and FS2,
respectively). Single substitutions of these residues caused a decrease in both in vitro
and in vivo activities when compared to the native enzyme, and this effect was
compounded in the doubly-substituted variants. Protein-folding analyses demonstrated
that the variant and native enzymes share similar folding characteristics, suggesting that
these amino-acid substitutions alter [FeFe]-hydrogenase activity by disrupting the FS4C
and FS2 clusters.

81

METHODS

Overexpression, Plasmid Construction, and Site-directed Mutagenesis—The C.
pasteurianum hydA gene was PCR amplified and inserted into the pAC-BAD vector as
previously described by Cornish et al. (17). Selected amino acid codons in the hydA
gene were mutated by site-directed mutagenesis PCR using the primers (Pfu turbo,
Stratagene) listed in Table 3.1 (18). Mutated PCR product was isolated by DpnI
digestion (New England Bioloabs) of the methylated template DNA at 37˚C for 2 h ,
followed by transformation into DH5α E. coli cells and selection on LB plates
supplemented with 50 µg/mL kanamycin sulfate. DNA was isolated from appropriate
transformant colonies, and the desired mutations were confirmed by DNA sequencing.
The constructs were transformed into electrocompetent Shewanella oneidensis MR-1
ΔhydAΔhyaB cells and selected for resistance to 50 μg/mL kanamycin (19).

Table 3.1. Primer pairs used for site-directed mutagenesis PCR of the C.
pasteurianum HydA gene in the pAC-BAD vector
Identity
C34S Sense
C34S Antisense
C98S Sense
C98S Antisense

Sequence
5'-ATATCTGCACTGTCTTTTTTAAATAAT-3'
5'-ATTATTTAAAAAAGACAGTGCAGATAT-3'
5'-CATGAATTCAAATCTGGTCCTTGCAAT-3'
5'-ATTGCAAGGACCAGATTTGAATTCATG-3'

82

Cell Growth, Induction, and Purification—S. oneidensis MR-1 ΔhydAΔhyaB cells
harboring the wild-type and mutated pAC-BAD:hydA constructs were cultured and gene
expression was induced as described by Cornish et. al (17), except that all cultures
were grown in a final volume of 1 L of antibiotic- and L-arabinose-supplemented LB
media. All purification steps were performed anaerobically, using either a glove box
(Coy Laboratory Products) or airtight Nalgene tubes. 16 h after addition of L-arabinose,
the cells were pelleted by centrifugation at 2,800× g and the supernatant was discarded.
The pellet was resuspended in 7 mL of nickel-nitrilotriacetic acid (Ni-NTA) wash buffer
(100 mM Tris-HCl [pH 8.0], 200 mM NaCl, and 5.0% glycerol) that was supplemented
with 10 mM Na2S2O4. The resuspension was aliquoted into seven 1.7 mL disposable
tubes (Eppendorf) and was lysed by sonicating each sample 8 times at 4 s intervals and
a power setting of 4 using the Fisher Scientific Model 100 Sonic Dismembrator at 4 sec
intervals. Lysed cells were centrifuged at 20,000× g to clear the lysate. Columns for
protein purification were filled with 1 ml Ni-NTA agarose (Qiagen) and washed with 15
mL of 10 mM Na2S2O4-supplemented Ni-NTA buffer to remove residual O2. The
supernatant was applied to the column and allowed to completely flow through. The
column was sequentially washed with 10 mL of Na2S2O4-supplemented Ni-NTA buffer,
10 mL of unsupplemented Ni-NTA buffer, and 5 mL of 20 mM imidazole-supplemented
Ni-NTA buffer. Bound protein was eluted from the column in 1 mL-fractions by applying
100 mM imidazole-supplemented Ni-NTA buffer. Qualitative hydrogenase uptake
activity was measured by incubating 20 μL of elution fraction with 20 μL of 10 mM
benzyl viologen dichloride (BV) for 5 min and noting the appearance of a purple color.
Consistently, the second elution fraction from each purification showed the greatest
83

change in color. The protein concentration of the elution fractions was determined using
the Pierce BCA Assay Kit and comparison to a bovine serum albumin (BSA) standard
curve.

Circular Dichroism—Protein samples were desalted and concentrated prior to
measuring the circular dichroism (CD) spectra. To desalt/concentrate the sample, a 0.5
mL aliquot of an elution fraction containing the 100 mM imidazole and purified enzyme
was applied to an Amplicon Ultra-0.5 centrifugal filter device (Millipore). The sample
was by centrifuged at 14,000× g for 10 min and the flow through was discarded. The
remaining elution fraction (~0.4 mL) was applied to the filter and the procedure
repeated. The samples were then resuspended in 0.4 ml of Ni-NTA buffer and
centrifuged again for 10 min at 14,000× g for 10 min to desalt the solution. This step
was repeated once more.
The concentrated sample was transferred to a quartz cuvette with a 1.0 mm
pathlength, and CD spectra were recorded using a Chirascan CD spectrometer (Applied
Photophysics). The scan speed was 6 nm/min, and the spectra were recorded using a 1
nm bandwidth over the range of 180 - 260 nm. Analysis was carried out utilizing the
DichroWeb Interface (http://dichroweb.cryst.bbk.ac.ukl), using the CDSSTR program
(20) and comparison to the Set 7 reference.

Hydrogen Uptake Activity Assays—Hydrogen uptake activity was measured by
preparing 1.9 mL of H2 uptake buffer [100 mM potassium phosphate buffer (pH 6.8), 2
mM dithiothreitol, 10 mM methyl viologen dichloride (MV) or BV] in a Pyrex Schlenk
84

cuvette. After sealing the cuvette with a rubber septum, the headspace gas was
exchanged with 100% H2 gas at 1 atm. The cuvette was returned to the glove box and
0.1 mL of the protein sample was added to the reaction. The Abs604 of the samples was
measured every 20 sec using a UV-Visible Spectrophotometer (Hewlett-Packard) using
the “kinetics” mode on the Agilent UV-Vis ChemStation software. The reaction time was
3 min and the cuvette was briefly shaken between measurements.

Hydrogen Evolution Activity Assays—H2 evolution activity with an abiotic electron donor
was measured by first preparing 1.9 mL of H2 evolution buffer (100 mM potassium
phosphate buffer [pH 6.8], 100 mM Na2S2O4, 10 mM MV) in 10 mL serum vials
(Wheaton). The vials were sealed with rubber septa (Bellco) and the headspace gas
exchanged with Ar at 1 atm. After returning the vials to the glove box, 0.1 mL of protein
sample was added to each and the vials were shaken at ~100 rpm at 25 A 100 μL
˚C.
syringe (Hamilton) was used to inject 20 μL of headspace gas into a TRACE GC Ultra
gas chromatograph (Thermo Scientific) and the resulting peak area was converted to
μmol H2 by comparison to a standard curve. The rate of H2 evolution was measured as
the change in μmol of H2 in the headspace over time per mg of protein. For
determination of Km and Vmax, the concentration of MV was varied from 0.5 to 10 mM.
H2 evolution activity with a biotic electron donor was measured as described for
activity with an abiotic donor, except that the 10 mM MV was replaced by 20 µM C.
pasteurianum ferredoxin. In addition, the vials were incubated at For
˚C. 35
determination of Km and Vmax the concentration of ferredoxin was varied from 5 to 20
µM.
85

RESULTS

Enzyme Purification and Circular Dichroism Spectroscopy—Specific cysteine residues
ligating the FS4C and FS2 clusters (Cys98 and Cys34, respectively) were substituted
with serine residues, generating both singly- and doubly-substituted forms of the
enzyme. N-terminally His-tagged forms of both the native and amino acid-substituted
proteins [variants] were purified from S. oneidensis MR-1 ΔhydAΔhyaB cells using NiNTA affinity chromatography.
Altering the native ligand coordination of an iron-sulfur cluster has the potential to
disrupt binding and cause loss of the cluster, thereby destabilizing protein folding. To
investigate folding properties of the native and variant enzymes, protein samples were
washed (to remove imidazole) and concentrated, and the overall folding of each
enzyme samples was analyzed by CD spectroscopy. Analyses of the spectra indicate
that the length and number of α-helices and β-strands are consistent between the
enzymes, suggesting that the variants have similar folding characteristics to the native
enzyme (Table 3.2, Figure 3.2). In addition, these spectra are in agreement with
secondary structure predictions of the C. pasteurianum hydrogenase, as well as solved
crystal structures of the enzyme. The overall structures of the variants are similar to the
native enzyme and changes in folding are unlikely to disrupt hydrogenase activity.

86

Table. 3.2. Secondary structure analysis of native and variant enzymes.
“Observed” denotes expected results based on the solved crystal structure of the C.
pasteurianum [FeFe]-hydrogenase.

α-helices per 100 residues
Average length of α-helices
β-strands per 100 residues
Average length of β-strands

Observed Variant C34S C98S C34S/C98S
3.6
3.5
4.2
3.5
3.5
13.3
16.3
14.0 16.1
14.7
3.6
4.0
3.8
3.5
4.3
4.6
4.6
4.3
4.7
4.3

87

Circular Dichroism (millidegrees)

Figure 3.2. CD spectra of native and variant enzymes. Circular dichroism spectra of
native and variant enzymes recorded in millidegrees, ranging from 200-260 nm. Spectra
<200 nm are unordered and were not used for analysis. Spectra have not been
normalized to protein concentration.

50
30
10
-10
-30
-50
-70
-90
-110
-130
-150

CpHydA
C34S
C98S
C34S/C98S

200

210

220
230
240
Wavelength (nm)

88

250

260

H2 Uptake Assays—Purified native and variant proteins were tested for H2 uptake
activity in the presence of an abiotic electron acceptor (BV) under 1 atm H2. A sharp
rel

decrease in the kobs

[specific activity of variant enzyme relative to that of the native

enzyme] for both the C34S and C98S variants was noted (Table 3.3, Figure 3.3). The
doubly-substituted variant (C34S/C98S) demonstrated uptake activity that was barely
above detection limit of the assay (0.00075 AU/s) and was only 0.3% as active as the
native enzyme. These results indicate that proper ligation of both clusters is required for
maximal H2 uptake activity.
To learn more about the interaction between BV and the enzymes during H2
uptake, Km and Vmax values of the native and variant proteins for H2 uptake were
calculated as a function of BV concentration (Table 3.4). The Vmax values calculated for
the singly- and doubly-substituted variants were 5- and >100-fold less, respectively,
than the native enzyme Vmax, closely resembling the kobs

rel

data. Conversely, there

was little difference between the calculated BV Km values, suggesting that, although
hydrogenase activity is impaired in the variants, the relationship between enzyme and
substrate may remain unaltered.

89

rel

rel

Table 3.3. Native and variant enzyme kobs
values. Comparison of kobs
H2
uptake activities using benzyl viologen (BV) and evolution activities using methyl
viologen (MV) and ferredoxin. Values are calculated as described in Figure 3.3.

90

Figure 3.3. Relative reduction in hydrogenase activity of singly- and doublyrel
substituted variants. kobs
(rate of variant enzyme relative to the rate of native
enzyme) values for H2 uptake using benzyl viologen (BV) and evolution activities using
methyl viologen (MV) or ferredoxin. To determine the effect of substitution, the specific
rel
activity of native enzyme was normalized to 1.0. Variant enzyme kobs values were
calculated from individual experiments and averaged.

91

Table 3.4. Kinetic values of native and variant enzymes. Km and Vmax values
calculated for H2 uptake using benzyl viologen (BV) and H2 evolution using methyl
viologen (MV) and ferredoxin.

92

H2 Evolution Assays—To examine the effect of the serine substitutions on H2 evolution,
the native and variant enzymes were tested for H2 evolution activity using an abiotic
electron donor system (10 mM MV and 100 mM Na2S2O4). All three variant enzymes
were deficient in evolution activity relative to the native enzyme, retaining only 5-12% of
rel

kobs

(Table 3.3, Figure 3.3), similar to the H2 uptake data.
To calculate Km and Vmax values for H2 evolution, the primary electron donor

(Na2S2O4) concentration was kept constant as the concentration of the direct electron
donor (MV) was varied between 1 and 10 mM. As observed with the kobs

rel

data, the

singly-substituted enzyme Vmax values were greatly reduced in comparison to the
native enzyme, while the calculated Vmax for C34S/C98S variant was <1% (Table 3.4),
again closely matching the H2 uptake results, indicating that the FS2 and FS4C clusters
must be fully ligated for optimal activity with abiotic donors.
The variant enzymes are decreased in activity relative to the native enzyme
when supplied with an artificial electron carrier, but this provides no direct information
on putative interactions with biotic electron donors. To explore these interactions, native
and variant enzymes were tested for H2 evolution activity using 20 µM C. pasteurianum
ferredoxin (and 100 mM Na2S2O4 as a reductant) as a biotic electron donor system.
rel

The kobs

values for all of the variants were less than the native enzyme, generating

values which closely matched the pattern of the abiotic donor H2 evolution activities
(Table 3.3, Figure 3.3), and suggesting that disrupting the clusters has a similar impact
on accepting electrons from both abiotic and biotic donors. When kinetic parameters
were calculated using ferredoxin, all enzymes were found to have similar Km values,

93

whereas the variant Vmax values were at least 5-fold reduced in comparison to the
native enzyme (Table 3.4), again resembling the MV data.

94

DISCUSSION

The C. pasteurianum [FeFe]-hydrogenase ligates four iron-sulfur clusters that are
hypothesized to compose an electron transfer relay between the active site and enzyme
surface. The iron-sulfur cluster-binding domains (FS4A, FS4B, FS4C, and FS2) which
coordinate the clusters in this relay are conserved among 75% of sequenced [FeFe]hydrogenases. This relay branches near the enzyme surface to the FS4C and FS2
clusters, but the role of this branch point has yet to be directly investigated.
During H2 oxidation, electrons are passed from the H-cluster to FS4A and then
FS4B, which is the branchpoint for the relay. From FS4B, electrons can be transferred
to an electron acceptor at the enzyme surface via either the FS4C or FS2 clusters (Fig
3.1). Peters et al. hypothesized either that these clusters could modulate the redox
potential for optimal electron transfer, that specific clusters were associated with only H2
evolution or H2 uptake activities, or that different physiological electron carriers might
interact separately with these clusters (13) and we set out to test these hypotheses.
To investigate the roles of the FS4C and FS2 clusters in both electron transfer and
hydrogenase activity, residues ligating these clusters (Cys98 and Cys34, respectively)
were substituted with serine residues to alter the reduction potential of the clusters (21).
Serine can act as a ligand to iron-sulfur clusters, and this type of substitution has been
used previously to alter the reduction potential of a cluster while still maintaining a
stable coordination environment (9). To test if the single or double substitutions in the C.
pasteurianum hydrogenase disrupted protein folding, the secondary structure of the
native and variant enzymes was investigated using CD spectroscopy and subsequently
95

compared to predicted structural models. As seen in Figure 3.2, the spectra are similar
in shape, although differ in amplitude, which may be due to differences in protein
concentration between the samples. Importantly, spectral analysis indicates that all of
the enzymes share similar secondary structures to one another. In addition, the values
of the native and variant proteins closely matched predicted structural models, further
suggesting that protein folding is not disrupted in the variant enzymes.
When serine was substituted for cysteine, the activity of the singly-and doublysubstituted variants were reduced in comparison to the native enzyme for both H2
evolution and H2 uptake activities (kobs
rel

reductions in H2 evolution kobs

rel

and Vmax). Additionally, extremely similar

and Vmax values were calculated when the enzymes

were supplied with either biotic or abiotic donors. Together, these results provide
evidence that both H2 evolution and uptake are impaired by these mutations, that
incorrect ligation of either of the distal clusters can cause this impairment, and that the
decrease in H2 evolution activity of the variants is independent of the electron donor
identity.
Comparison of the H2 uptake and H2 evolution data reveals that, for the C34S and
C98S variants in relation to native enzyme, there was a greater reduction in H2
evolution activity than there was in H2 uptake activity. This effect was observed
previously by Cornish et al. while investigating hydrogenase proton transfer (17). A
possible explanation for this disparity may lie in the difference between the driving
forces and that each reaction was far outside of equilibrium (excess substrate
conditions). In addition, the reduction potentials for MV and BV differ (-450 mV and -350

96

mV, respectively), which could also cause a disparity between the measured H2 uptake
rel

and H2 evolution kobs

values (22).

A strange pattern was observed in examining the Km values calculated using abiotic
donors. These data would suggest that disruption of one or both distal clusters leads to
an increase in reactivity with BV during H2 uptake and a decrease in reactivity with MV
during H2 evolution, and this does not seem to match the values calculated with
ferredoxin. These experiments need to be repeated using a broader range of substrate
concentrations to determine the cause of these conflicting data.
In conclusion, we have explored the importance of two distal iron-sulfur clusters that
constitute a branching point in the C. pasteurianum [FeFe]-hydrogenase. Substitution of
residues affecting either cluster causes a reduction in both H2 evolution and H2 uptake
activity relative to native enzyme, and activity of the doubly-substituted enzyme is
reduced by >90%. CD spectral analysis provide strong evidence that impairment of
activity does not correlate to protein misfolding. Together, these results demonstrate
that this relatively common electron transfer relay is dependent on both of the distal
clusters for hydrogenase activity, as these clusters are essential in either directly
accepting electrons or modulating the reduction potential of the cluster involved in
electron transfer. Importantly, further characterization of this relay may provide
important insight into the mechanisms of [FeFe]-hydrogenase activity.

97

REFERENCES

98

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1.

Vignais P. M., Billoud B. (2007). Occurrence, classification, and biological function of
hydrogenases: an overview. Chem. Rev. 107(10), 4206-4272.

2.

Adams M. W. W. (1990). The structure and mechanism of iron-hydrogenases. Biochim.
Biophys. Acta 1020(2), 115-145.

3.

Ghirardi M. L., Dubini A., Yu J., Maness P. C. (2009). Photobiological hydrogenproducing systems. Chem. Soc. Rev. 38(1), 52-61.

4.

Kim D. H., Kim M. S. (2011). Hydrogenases for biological hydrogen production.
Bioresour. Technol. 102(18), 8423-8431.

5.

Schuchmann K., Müller V. (2012). A bacterial electron-bifurcating hydrogenase. J. Biol.
Chem. 287(37), 31165-31171.

6.

Winkler M., Hemschemeier A., Jacobs J., Stripp S., Happe T. (2010). Multiple ferredoxin
isoforms in Chlamydomonas reinhardtii - their role under stress conditions and
biotechnological implications. Eur. J. Cell. Biol. 89(12), 998-1004.

7.

Jacobs J., Pudollek S., Hemschemeier A., Happe T. (2009). A novel, anaerobically
induced ferredoxin in Chlamydomonas reinhardtii. FEBS Lett. 583(2), 325-329.

8.

ElAntak L., Morelli X., Bornet O., Hatchikian C., Czjzek M., Dolla A., Guerlesquin F.
(2003). The cytochrome c3-[Fe]-hydrogenase electron-transfer complex: structural model
by NMR restrained docking. FEBS Lett 548(1-3), 1-4.

9.

Dementin S., Belle V., Bertrand P., Guigliarelli B., Adryanczyk-Perrier G., De Lacey A.
L., Fernandez V. M., Rousset M., Leger C. (2006). Changing the ligation of the distal
[4Fe4S] cluster in NiFe hydrogenase impairs inter- and intramolecular electron transfers.
J. Am. Chem. Soc. 128(15), 5209-5218.

10.

Vignais P. M., Billoud B., Meyer J. (2001). Classification and phylogeny of
hydrogenases. FEMS Microbiol. Rev. 25(4), 455-501.

11.

Mulder D. W., Shepard E. M., Meuser J. E., Joshi N., King P. W., Posewitz M. C.,
Broderick J. B., Peters J. W. (2011). Insights into [FeFe]-hydrogenase structure,
mechanism, and maturation. Structure 19(8), 1038-1052.

12.

Meyer J. (2007). [FeFe] hydrogenases and their evolution: a genomic perspective. Cell.
Mol. Life Sci. 64(9), 1063-1084.

13.

Peters J. W. (1999). Structure and mechanism of iron-only hydrogenases. Curr. Opin.
Struct. Biol. 9(6), 670-676.
99

14.

Lubner C. E., Knorzer P., Silva P. J., Vincent K. A., Happe T., Bryant D. A., Golbeck J.
H. (2010). Wiring an [FeFe]-hydrogenase with photosystem I for light-induced hydrogen
production. Biochemistry 49(48), 10264-10266.

15.

Long H., Chang C. H., King P. W., Ghirardi M. L., Kim K. (2008). Brownian dynamics
and molecular dynamics study of the association between hydrogenase and ferredoxin
from Chlamydomonas reinhardtii. Biophys. J. 95(8), 3753-3766.

16.

Pandey A. S., Harris T. V., Giles L. J., Peters J. W., Szilagyi R. K. (2008).
Dithiomethylether as a ligand in the hydrogenase h-cluster. J. Am. Chem. Soc. 130(13),
4533-4540.

17.

Cornish A. J., Gärtner K., Yang H., Peters J. W., Hegg E. L. (2011). Mechanism of
proton transfer in [FeFe]-hydrogenase from Clostridium pasteurianum. J. Biol. Chem.
286(44), 38341-38347.

18.

Zheng L., Baumann U., Reymond J. L. (2004). An efficient one-step site-directed and
site-saturation mutagenesis protocol. Nucleic Acids Res. 32(14), e115.

19.

Ozawa K., Yasukawa F., Fujiwara Y., Akutsu H. (2001). A simple, rapid, and highly
efficient gene expression system for multiheme cytochromes c. Biosci. Biotechnol.
Biochem. 65(1), 185-189.

20.

Sreerama N., Woody R. W. (2000). Estimation of protein secondary structure from
circular dichroism spectra: comparison of CONTIN, SELCON, and CDSSTR methods
with an expanded reference set. Anal. Biochem. 287(2), 252-260.

21.

Babini E., Bertini I., Borsari M., Capozzi F., Dikiy A., Eltis L. D., Luchinat C. (1996). A
serine → cysteine ligand mutation in the high potential iron−sulfur protein from
Chromatium vinosum provides insight into the electronic structure of the−4S]
[4Fe
cluster. J. Am. Chem. Soc. 118(1), 75-80.

22.

Wardman P. (1991). The reduction potential of benzyl viologen: an important reference
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100

Chapter 4

Characterization of hydrogen metabolism in the multicellular green alga Volvox
1
carteri

1

This chapter is adapted from Cornish A. J., Gärtner K., Green, R., Mason, S., Hegg E.
L. (2012). Characterization of hydrogen metabolism in the multicellular green alga
Volvox carteri. Appl. Environ. Microbiol. Submitted
Saundra Mason and Robin contributed significantly to H2 evolution assays, RNA
isolation, and plasmid construction.
101

INTRODUCTION

Hydrogen gas is a potent energy carrier and has the potential to supplement
existing transportation fuels (1, 2). A clean-burning gas, H2 can be synthesized from
renewable sources (3) and releases only water vapor during energy conversion via
combustion. Certain autotrophic microorganisms, such as green algae, are able to
channel photosynthetically-derived reducing equivalents to H2 production (4, 5, 6, 7).
Many green algae can be cultured at an industrial scale utilizing non-arable land,
thereby avoiding competition with agricultural feedstocks (8). Extensive studies of H2
metabolism in several algal species have established a basis for investigation of algal
biohydrogen production, and these studies can be used to inform future efforts to
generate renewable alternative fuels (9, 10).
Green

algae

evolve

H2

using

[FeFe]-hydrogenases

(EC#1.12.7.2),

metalloproteins which catalyze both the reduction of protons and the oxidation of H2
(11, 12). Electrons for H2 production can either be channeled from photosynthetic
water-splitting or be obtained by the fermentation of carbon sources (11, 13). In a
variety of green algae, hydrogenases are encoded by two gene paralogs (HYDA1 and
HYDA2) that have high sequence similarity, although the respective physiological
functions of each enzyme are still unclear (14, 15). Maturation proteins are required to
assemble the catalytic active site, and the corresponding genes (HydE, HydF, and
HydG in bacteria - HYDEF and HYDG in green algae) are ubiquitous in the genomes of
organisms which utilize [FeFe]-hydrogenase (16). Although these genes in green algae
were likely acquired by lateral gene transfer (17), hydrogenase gene clustering has not
102

been noted in sequenced green algal genomes. Despite this lack of clustering,
hydrogenase and maturation factor gene expression is tightly co-regulated. In addition,
the enzyme is irreversibly-inactivated by the presence of O2, and the HYD genes are
expressed only under anaerobiosis (12, 18).
The O2-sensitivity of [FeFe]-hydrogenase renders H2 synthesis dependent on
micro-aerobic conditions and thus limits production during light-driven oxygenic
photosynthesis (5, 19, 20). In Chlamydomonas reinhardtii, a model unicellular green
alga, this inhibition may be overcome by sulfur deprivation, which limits O2 production
by photosystem II (PSII) while still allowing electrons gained from photosystem I (PSI) to
be coupled to H2 production (21). In addition, anaerobic conditions are quickly
established in the dark as respiration depletes O2, thereby allowing carbon stores
generated during photosynthesis to be utilized for H2 production (5, 6). Both of these
methods allow for channeling of electrons to H2 synthesis.
Volvox carteri is a multicellular green alga separated from C. reinhardtii by
approximately 220 million years of evolution (22). V. carteri is composed of two cell
types, gonidia and somatic, which are embedded within an extracellular matrix (23, 24).
Recently, the V. carteri genome was sequenced and, as expected, it shares a great
deal of genome sequence similarity and metabolic processes (22, 25) with C.
reinhardtii. Two putative [FeFe]-hydrogenase genes were annotated in the genome of
V. carteri (22). However, the ability of this organism to evolve H2 was not established.
In this manuscript we demonstrate that the green alga V. carteri is able to utilize
an abiotic electron donor system to drive H2 production. To our knowledge, this is the
first report of H2 metabolism in a multicellular eukaryote with differentiated cells. Genes
103

responsible for hydrogenase assembly and maturation are up-regulated under
anaerobic conditions, and both HYDA genes encode functional [FeFe]-hydrogenases. In
relation to other green algae, these genes in V. carteri are uniquely arranged in an
operon-like structure within the genome, providing evolutionary evidence for the
potential origins of horizontal gene transfer. Together, these data support a role for H2
in the metabolism of V. carteri and provide a basis for further investigation of the
ancestral acquisition of [FeFe]-hydrogenase genes in green algae.

104

METHODS

Growth Conditions—Unless otherwise noted, V. carteri f. nagariensis EVE (22) cells
were grown with continuous light (90 µE m

-2

-1

s ) and shaking (100 rpm) in standard

Volvox medium (SVM) at 22 ˚C as previously described (26).

Anaerobic Acclimation—To acclimate V. carteri cells to anaerobiosis, a culture of lightadapted cells was centrifuged at 1,000× g for 5 min. The resulting cell pellet was
washed three times in an anaerobic chamber (Coy Laboratory Products) with degassed
SVM supplemented with 0.16 mM MgCl2 rather than 0.16 mM MgSO4 [SVM (-SO4)] to
exclude sulfate from the medium. Sulfate-depleted medium was utilized to decrease
photosystem II activity and thus limit oxygenic photosynthesis. The cells were
resuspended in degassed SVM (-SO4) to a final concentration of 75 μg chlorophyll/mL,
and 2 mL of this mixture were sealed in a 10 mL air-tight serum vial (Wheaton). The
resuspension was shaken for 4 h. To prevent any remaining photosynthetic activity from
photosystem II, the vials were wrapped with aluminum foil.

Hydrogen Production Measurements—H2 evolution using an artificial electron donor
was measured by incubating 0.1 mL of either aerobically- or anaerobically-acclimated
cells (75 μg chlorophyll/mL) with 1.9 mL of H2 evolution assay solution [SVM (-SO4),
100 mM sodium dithionite, 10 mM methyl viologen dichloride (MV)] in a sealed 10 mL
serum vial at 22 ˚C in the dark with continuous shaking. At various time points, 20 μL of
headspace gas were injected into a TRACE GC Ultra Gas Chromatograph (Thermo
105

Scientific) using a 100 µL syringe. The peak at 1.4 min corresponded to H2, and the
absolute value of H2 gas was determined by comparison to a standard curve. The
resulting values were plotted versus time to monitor the accumulation of H2 in the
headspace, and the rates were determined per μg of chlorophyll.
H2 evolution measurements using exogenous carbon sources were performed as
described above, except that the cell resuspension was incubated in 1.9 mL of SVM
(-SO4) supplemented with either 0.174 mM sodium acetate, D-glucose, or glycerol.

RNA Isolation—To isolate total RNA from V. carteri, cell cultures were centrifuged at
10,000× g for 10 min. The resulting pellets were resuspended in 0.5 mL of TRIzol
(Invitrogen) and frozen in liquid nitrogen. The frozen TRIzol/cell mass mixture was
crushed using a mortar and pestle, thawed, and resuspended with an additional 0.5 mL
of TRIzol. To this suspension, 0.2 mL of chloroform was added, and the mixture was
vortexed at top speed at room temperature for 15 min. To pellet cell debris, the
suspension was centrifuged at 10,000× g for 15 min at 10 ˚C. The aqueous phase was
transferred to a 1.7 mL tube (Denville) containing 0.25 mL of 0.8 M sodium citrate and
1.0 M sodium chloride, and the suspension was gently mixed by inversion. Following
the addition of 0.25 mL of isopropanol, the mixture was centrifuged at 10,000× g for 25
min at 10 ˚C. The supernatant was discarded and the RNA pellet washed twice with icecold 75% ethanol. The pellet was allowed to dry and then resuspended in 20 µL
diethylpyrocarbonate-treated water.

106

RT-PCR—Reverse transcription of mRNA was performed using M-MLV Reverse
Transcriptase (Invitrogen), and the cDNA was amplified via PCR using GoTaq Green
Master Mix (Promega) and the primer sets listed in Table 4.1. Genomic and cDNA
sequences of HYDA1, HYDA2, HYDEF, and HYDG can be accessed from the
DDBJ/EMBL/GenBank database

with

the

accession numbers

XM_002948441,

XM_002948437, XM_002948568, and XM_002948439, respectively.

Cloning of HYDA1 and HYDA2—The HYDA1 and HYDA2 genes were PCR amplified
with PfuTurbo DNA polymerase (Stratagene) from reverse-transcribed cDNA using the
primers listed in Table 4.1. The amplified products were ligated into the SacI/HindIII site
of pAC-BAD, a pBAD/D-TOPO (Invitrogen) expression vector (contains a kanamycinresistant cassette and an L-arabinose inducible promoter) that was modified to remove
the N-terminal thioredoxin-tag (27). The constructs were transformed into S. oneidensis
MR-1 ΔhydA/ΔhyaB (28) electrocompetent cells as described by Ozawa (29) and
selected for antibiotic resistance on 50 µg/mL kanamycin LB plates.

Overexpression, Protein Purification, and Hydrogen Evolution Assay—Cells harboring
the pAC-BAD_HYDA1 and pAC-BAD_HYDA2 vectors were induced for gene
overexpression and protein synthesis as previously described (27). Enzymes were
purified in an anaerobic chamber and assayed for H2 evolution activity as described by
Cornish et al. (27).

107

Table 4.1. List of primers used for cloning and cDNA amplification. Restriction digest sites are italicized. 6xHis-tag
sequences are underlined. Genomic sequences amplified by the cloning primers are in bold.

108

Phylogenetic Analysis—Using BLAST, 86 unique [FeFe]-hydrogenase amino acid
sequences were collected from the National Center for Biotechnical Information and
aligned by MUSCLE (30) using a gap extension of 13 and opening penalty of 0.05.
Pairwise distances were calculated from the alignment and sequences with a distance
<0.1 were eliminated. The remaining 52 sequences were aligned as stated previously,
and the alignment was trimmed to isolate only the representative H-cluster-binding
domain from HYDA. Evolutionary history was inferred using the Neighbor-Joining
method (31) and evolutionary distances computed using the p-distance method (32).
2000 bootstrap replicates were performed. All analyses were conducted in MEGA5 (33).

109

RESULTS

Hydrogen Evolution Studies—V. carteri is closely related to C. reinhardtii, a green alga
with well-described hydrogen metabolism (19, 20). Recent sequencing and annotation
of the V. carteri genome (22) uncovered two genes with sequence similarity to algal
[FeFe]-hydrogenase genes (HYDA1, HYDA2) (Figure 4.1) as well as two genes
predicted to be necessary for [FeFe]-hydrogenase maturation (HYDEF, HYDG).
V. carteri was tested for in vivo H2 production in SVM (-SO4) after a 4 h
anaerobic

acclimation

period.

Aliquots of

anaerobically-acclimated

cells

were

transferred to anaerobic assay vials containing an abiotic electron donor system (MV
and Na2S2O4), and the headspace was tested for accumulation of H2. H2 production
could be measured within the first 45 min of incubation with MV/Na2S2O4, eventually
achieving a rate of 380 nmol H2/mg chlorophyll/s after 3 h (Figure 2). When oxygenexposed cells (i.e. aerobic cells) were assayed under similar conditions, no H2
accumulation was observed over the course of the assay (Figure 4.2). This indicates
that V. carteri requires anaerobic acclimation for H2 evolution. Cells that were not
supplemented with MV/Na2S2O4 did not accumulate a significant amount of H2 in the
headspace even 48 h after initiating the assay.

110

Figure 4.1. Partial multiple sequence alignment of V. carteri and C. reinhardtii HYDA amino acid sequences. Black
boxes denote strictly conserved residues, while regions with high conservation are boxed and are in boldface. The
HYDA1 proteins from V. carteri and C. reinhardtii share 74.1% sequence identity, while the HYDA2 proteins share 75.1%
sequence identity. Accession numbers for C. reinhardtii HYDA1 and HYDA2 are XP_001693376 and XP_001694503,
respectively.

111

Figure 4.2. H2 evolution rate of aerobically- or anaerobically-acclimated V. carteri
cells. Following a 4 h acclimation period, cells were incubated in SVM (-SO4) with and
without an abiotic electron donor system. H2 production rates were measured over a 3 h
period and normalized on a per mg chlorophyll basis. Error bars denote standard
deviation (n ≥ 3).

112

Fermentative H2 Production—Having established H2 metabolism in V. carteri using an
artificial electron system, we sought to drive H2 production utilizing electrons generated
during fermentation. A recent report suggests that V. carteri may be able to import
glycerol (34), which could then be fed into fermentation. To test for fermentative H2
evolution in V. carteri, anaerobically-induced cultures were supplemented with either
0.174 mM sodium acetate,

D-glucose,

or glycerol and compared to the H2 evolution

from cells in unsupplemented medium. No quantifiable increase in H2 could be noted
even 48 h after supplementation, indicating that addition of these exogenous carbon
sources does not drive H2 production under these conditions.

Transcript Analysis—Based on H2 accumulation under anaerobiosis, we predicted that
the putative HYDA genes played a role in H2 production. To test for HYDA expression
during anaerobiosis, RNA was collected from both aerobically- and anaerobically
acclimated cells. The RNA was reverse-transcribed and PCR amplified with primers
specific to the putative [FeFe]-hydrogenase genes, HYDA1 and HYDA2. Strong
amplification of mRNA for both genes was observed in anaerobic samples, while very
little transcript could be observed in the aerobic samples when compared to a
housekeeping gene, ACTA (accession number XP_002955536) (Figure 4.3). These
results indicate that the anaerobic expression of HYDA1 and HYDA2 correlate with the
H2 production observed in V. carteri.

113

Figure 4.3. Gene transcript accumulation observed by RT-PCR. HYDA1 and HYDA2
transcripts were strongly amplified using RNA isolated from anaerobically-acclimated V.
carteri cells. The ACTA gene was constitutively expressed under both aerobic and
anaerobic conditions and was used as a control.

114

Hydrogenase Cloning and Characterization—The two putative HYDA genes have high
sequence similarity to known green algal hydrogenase genes (Figure 4.1) and likely
code for active H2-producing enzymes. To test for this, the two genes were amplified
from cDNA, cloned into pAC-BAD, and transformed into S. oneidensis ΔhydAΔhyaB
cells. Gene expression was induced by the addition of L-arabinose, and the N-terminally
His-tagged proteins were purified by nickel-affinity chromatography. The HYDA1 and
HYDA2 proteins were active when supplied with MV/Na2S2O4, and both had similar Km
values when measured for MV (2.76 mM and 2.82 mM, respectively). The specific
activity of HYDA1 was ~1,000-fold greater than HYDA2 (4,000 nmol H2/mg/s and 2.2
nmol H2/mg/s, respectively). This differs from previously reported values for
recombinant C. reinhardtii HYDA1 and HYDA2 specific activities, which were nmol
H2/mg/s and nmol H2/mg/s, respectively (35).

Phylogenetic Analysis—The two hydrogenase genes and the two assembly factor
genes in the V. carteri genome (HYDA1, HYDA2, HYDEF, HYDG) were found to be in
relatively close proximity to one another. The distance between the furthest genes
(HYDA1 and HYDA2) is ~57 kb, and the main cluster of genes is within a ~23 kb span
(Figure 4.4). This level of [FeFe]-hydrogenase gene clustering has not been previously
observed in the sequenced genomes of green algae (C. reinhardtii and Chlorella
variabilis). Green algae are theorized to have obtained their hydrogenase genes via
horizontal gene transfer, likely from an operon or operon-like gene structure.

115

Figure 4.4. [FeFe]-hydrogenase operon-like gene cluster in V. carteri. Four genes with sequence similarity to HYDA1,
HYDA2, HYDEF, and HYDG are arranged within 60 kb of one another. Three additional putative genes are within this
region; ACKA (XP_002948613), a phosphate acetyltransferase (XP_002948484), and a predicted gene, Q85
(XP_002948486). The predicted gene products are not hypothesized to catalyze H2 production or assist in the assembly
of the [FeFe]-hydrogenase active site, and these genes are not found within investigated bacterial [FeFe]-hydrogenase
gene clusters.

116

To investigate potential origins of this operon-like cluster in V. carteri, a
phylogenetic analysis was performed utilizing [FeFe]-hydrogenase protein sequences.
As previously observed, green algal hydrogenase sequences group closely together,
with the sequences from C. variabilis showing greater divergence (Figure 4.5) (17). A
grouping of thermophilic anaerobic bacterial sequences suggests a relationship to the
green algal clade, indicating that these genes may have arisen from a common
ancestor. Within the clade, the bacterial genomes each contain an operon-like cluster
composed of hydrogenase and hydrogenase maturation genes. In particular, the
genomes

of

Fervidobacterium

nodosum,

Fervidobacterium

pennivorans,

and

Thermotoga thermarum contain operon-like clusters composed of five hydrogenase
genes that cover a <22 kb span (Figure 4.6). These findings provide hints for a common
ancestor among these bacteria and the possible origin of hydrogenase genes in green
algae.

117

Figure 4.5. Phylogenetic analysis of representative HYDA homologs. Evolutionary
history of [FeFe]-hydrogenase amino acid sequences was inferred by the NeighborJoining method and the bootstrap consensus tree was inferred from 2,000 replicates.
Partitions reproduced in less than 50% bootstrap replicates are collapsed and bootstrap
values are represented at individual nodes. The tree is drawn to scale, with evolutionary
distances calculated by the p-distance method and the units are described as the
number of amino acid differences per site.

118

Figure 4.5 (cont’d)

119

Figure 4.6. [FeFe]-hydrogenase gene clusters observed in V. carteri, F. nodosum, F. pennivorans, and T.
thermarum. Clusters are all on the same scale and the putative HYD genes are denoted by color (HYDA sequences,
orange and pink; HYDE, green; HYDF, blue; HYDG, purple; HYDEF fusion gene, green-to-blue). The coding sequence of
the F. nodosum and T. thermarum clusters has been reversed (reading 3’ to 5’) to more closely match the orientation of
the V. carteri cluster.

120

DISCUSSION

A variety of green algae have been characterized for their ability to couple energy
derived from photosynthesis to the production of H2, especially the model organism C.
reinhardtii (4, 5, 6). V. carteri is separated from C. reinhardtii by ~220 million years of
evolution, yet the two organisms share many traits at the physiological and genetic
levels (22). This forms a logical basis for investigating the unique multicellular alga V.
carteri for H2 metabolism.
To test the H2 metabolism of V. carteri, anaerobic algal cell cultures were
provided with an electron donor system and H2 accumulation was measured. Under the
conditions tested, V. carteri cultures evolved appreciable amounts of H2 when supplied
with an exogenous electron source (MV/Na2S2O4). The addition of fermentable carbon
sources (acetate, glycerol, and D-glucose) did not drive H2 production in anaerobicallyacclimated cells. Based on previous reports, V. carteri cannot take up acetate under
dark anaerobiosis (36) and lacks glucose importers (37), and therefore the inability of
acetate and glucose to drive H2 production is to be expected. Intriguingly, although
Anderberg et al. hypothesized that V. carteri could take up glycerol via putative
aquaporins (34), no H2 production was noted upon glycerol supplementation. This result
could be explained by the inability of glycerol to enter the cells under our assay
conditions or because electrons derived from glycerol-breakdown are not channeled
into H2 metabolism.
Transcript for the hydrogenase homologs HYDA1 and HYDA2 accumulated
under anaerobic conditions, and the heterologously-expressed gene products
121

demonstrated quantifiable in vitro H2 evolution activity when supplied with an artificial
electron donor. In addition, HYDA1 and HYDA2 have similar Km values of ~2.8 mM for
MV. Interestingly, the specific activities measured for each enzyme were dissimilar, with
HYDA1 being approximately 1000-fold more active than HYDA2. This may be due to
incomplete maturation of HYDA2 in the S. oneidensis MR-1 expression system.
In addition to similar kinetic values, HYDA1 and HYDA2 also share common
structural features with other characterized green algal [FeFe]-hydrogenases. For
example, both proteins contain two short sequences not observed in non-algal
hydrogenases and also lack the canonical iron-sulfur cluster-containing F-domains (18).
When analyzed with several localization prediction servers (ChloroP 1.1 (38), PCLR
(39), WoLF PSORT (40), and CELLO (41)), a chloroplast-targeting peptide was
identified at the N-terminus of HYDA2, which is a typical feature of green algal
hydrogenases. Due the ubiquitous nature of this peptide among algal hydrogenases, it
was intriguing to find that these programs do not predict that HYDA1 is chloroplastlocalized. If the localization prediction is accurate, targeting to different subcellular
compartments could imply the utilization of different physiological donors.
While examining the hydrogenase genes within the V. carteri genome, it was
noted that the HYDA1 and HYDA2 genes are within proximity to genes encoding the
maturation proteins HYDEF and HYDG. This clustering is unique to V. carteri among
currently sequenced green algal genomes, but an [FeFe]-hydrogenase gene cluster
was also recently reported in the genomes of the photosynthetic heterokonts
Nannochloropsis oceanica CCMP1779 (42) and Nannochloropsis gaditana CCMP526
(43). Currently, hydrogenase genes within eukaryotic genomes have been assumed to
122

be acquired from bacteria via horizontal gene transfer, although the exact origin of the
HYD genes is not known (44, 45).
We investigated possible evolutionary relationships between HYDA genes via
phylogenetic analysis, particularly concentrating on known green algal protein
sequences, and including representative bacterial and protist homologs. Our phylogram
matches well with analyses conducted by Meuser et al. (17), showing a distinct
grouping of green algal genes (Figure 4.5). In addition, a clade of thermophilic bacterial
anaerobes demonstrates a relationship to this group of green algal genes, and three
genomes of bacteria within this clade contain operon-like clusters (>22 kb span) with
genes encoding two hydrogenases and all three maturation proteins. However, distinct
differences exist between the V. carteri and bacterial gene clusters, such as the
orientation of the genes and the fusion of the HYDE and HYDF genes in V. carteri. It is
therefore intriguing that the relative positions of the genes are identical between V.
carteri and bacterial clusters, although the bacterial genes, lacking introns, are shorter
in length (Figure 4.6). Together, this information suggests that HYD genes in modern
green algae may have been attained via lateral gene transfer from an ancestor of the
thermophilic bacteria clade. In addition, sequences from several protists, including N.
oceanica CCMP1779, also have a close relationship to these algal sequences,
indicating that these organisms may have acquired HYDA genes from a similar bacterial
ancestor.
In conclusion, we have provided the first direct evidence for H2 metabolism in a
multicellular eukaryote with differentiated cells. The green alga V. carteri is able to drive
H2 production in the dark utilizing at least one of two [FeFe]-hydrogenases, both of
123

which were show to be active in vitro and have similar Km values for MV. A unique gene
cluster encodes the hydrogenases and maturation machinery in the V. carteri genome,
and this cluster provides new hints to the origin of these genes in green algae.

124

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125

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130

SUPPLEMENTAL CHAPTER 1

Discovery and Identification of Novel [FeFe]-hydrogenase Genes

1

1

Initial degenerate primer PCR conditions were determined, in part, by Saundra Mason.
131

INTRODUCTION

The use of hydrogenases to produce H2 has great potential to supplement existing
energy technologies and expand the availability of renewable, clean fuels (1, 2). In
particular, [FeFe]-hydrogenases have been investigated for industrial-scale H2
production, as the catalytic turnover of these proteins is the highest measured among
H2-evolving enzymes (3, 4). For industrial use of these hydrogenases, additional
properties would be required for an ideal enzyme, such as: oxygen-insensitivity, a broad
pH range, salt tolerance, and thermostability (5, 6, 7). A relatively small number of
[FeFe]-hydrogenases have been characterized that possess these properties, and there
remains potential for discovery of enzymes with unique qualities.
Degenerate primer-based PCR is one method which could be used to discover new
[FeFe]-hydrogenase genes. This technique involves designing primers that utilize
nucleotide degeneracy of regions within a gene which are conserved at the amino acid
level. Using this approach, homologous protein-encoding genes can be identified, even
when large amounts of variability exist elsewhere in the gene. In addition, the efficacy of
this method was recently illustrated as degenerate primer-based PCR was used to
identify new [FeFe]-hydrogenase genes from a variety of genomic samples (8, 9, 10).
Two primary genomic DNA sources were of interest during our search for new
hydrogenase genes. Geoglobus ahangari 234 is a hyperthermophilic archaeon that
grows anaerobically and is able to oxidize H2 as an electron source (11). To date, no
archaeal genome has been found to contain a gene homolog encoding a functional
[FeFe]-hydrogenase. However, the availability of G. ahangari 234 genomic DNA, the
132

greater thermostability of proteins isolated from similar hyperthermophiles, and the
noted H2-oxidizing activity of the organism piqued our interest. Genomic samples from
Yellowstone National Park (YNP) were also of interest, as a number of hot springs in
the park contain great microbial biodiversity (12, 13). Colonizing and associating with
microbial mats in these springs are a variety of auto-, hetero-, and mixotrophic
organisms. In addition to high temperatures, these microorganisms often have to deal
with both high salinity and extremes in pH, and many are predicted to participate in H2
metabolism, providing excellent rationale for investigating environmental genomic
samples from YNP.
After performing a phylogenetic analysis of known [FeFe]-hydrogenase genes, we
identified key regions that were highly conserved between homologs. Using designed
primers, ideal PCR conditions were established. No gene fragments homologous to
[FeFe]-hydrogenase were identified from G. ahangari 234, but 9 gene fragments were
found from environmental samples isolated at YNP. Although we extended our
knowledge of the fragments, we did not establish the full-length sequence of any one
gene fragment and continuing work will focus on this effort.

133

METHODS

Phylogenetic Tree Construction—The phylogenetic tree used to identify specific clades
of organisms was generated using the PHYlongeny Inference Package (PHYLIP) (14).
The NCBI database was used to collect 148 annotated and putative [FeFe]hydrogenase amino acid sequences and these were aligned using ClustalX (15). A
protein distance matrix based on the sequences was calculated with the “protdist”
algorithm and the tree assembly was carried out using the “Fitch-Margoliash” method.
No outgroup was selected and 1,000 bootstrap replicates were performed.
To compare the translated sequences of the identified gene fragments, the
alignment software MEGA5 (16) was used to generate a phylogenetic tree comparing
98 [FeFe]-hydrogenase homolog amino acid sequences and the translated gene
fragment sequences. [FeFe]-hydrogenase sequences were collected as described
previously and the sequences were aligned using MUSCLE (17) with opening gap and
gap extension penalties of 13 and 0.05, respectively. Pairwise distances were
calculated from the alignment and sequences with a distance <0.1 were eliminated. The
remaining 64 sequences were aligned as stated previously, and the alignment was
trimmed to isolate only the representative H-cluster-binding domain from HydA. The
phylogram was generated by the Neighbor-Joining method (18) and the evolutionary
distances calculated by the p-distance method (19). The Homo sapiens NARFL protein
and Saccharomyces cerevisiae Nar1p amino acid sequences were used as outgroups
and 2,000 bootstrap replicates performed.

134

Mulitple Sequence Analysis—Sequences were aligned using the TCOFFEE software
tool available at http://igs-server.cnrs-mrs.fr/Tcoffee/tcoffee_cgi/index.cgi.

Degenerate Primer-Based PCR Amplification—Basic PCR amplification was performed
as described in section 8.18 of Molecular Cloning (20), using GoTaq Green Master Mix
(Promega) and 40 pmol of each primer in a total of 25.0 µL reaction buffer. No BSA was
added to the Fe4A reactions, while 200 ng of BSA was added to the Fe9 reactions.
Using an Eppendorf Mastercycler, the initial denaturation step was at 95 for 5 min,
˚C
followed by 40 cycles composed of (a) 30 s of denaturation at 95
˚C,

(b) 30 s of

annealing at 49 ˚C for Fe9 and 57 ˚C for Fe4A, and (c) 1 min of elongation at 72 ˚C. The
PCR reaction was finished with an elongation step of 72 ˚C for 5 min.

Cloning of Gene Fragments—Gene fragments generated during degenerate primerbased PCR were run on a 0.8 % agarose TAE gel (section 5.4 of Molecular Cloning)
(20) at 80 V for 30 min. Bands were observed by ethidium bromide-staining and UV
illumination. Amplified products were cut from the gel using a clean razor blade and
isolated using the QIAGEN Gel Extraction Kit. The isolated amplicons were ligated into
the pGEM-T Easy vector (Promega) as described by the kit protocol. The recombinant
plasmid was transformed into DH5α Escherichia coli chemically-competent cells as
described in section 1.116 of Molecular Cloning (20), and selected for resistance to 50
µg/mL ampicillin. Further selection was based on appearance of white transformant
colonies in the presence of 20 µg/mL 5-bromo-4-chloro-indolyl-β-D-galactopyranoside
(X-gal), denoting β-galactosidase gene interruption by insertion of a gene fragment.

135

Gene Fragment Identification—White transformant colonies were individually picked
with a toothpick and first dipped into a PCR tube containing 11.5 µL sterile water before
being streaked onto an LB plate containing 50 µg/mL ampicillin and allowed to grow
overnight at 37˚C. Reactions for colony PCR were prepared by adding 12.5 µL of
GoTaq Green Master Mix (Promega) and 0.5 µL each of 25 µM forward and reverse
M13 primers (5’-GTAAAACGACGGCCAG-3’ and 5’-CAGGAAACAGCTATGAC-3’,
respectively) to each PCR tube. Using a thermocycler, the initial denaturation step was
at 94 ˚C for 5 min followed by 30 cycles composed of (a) 30 s of denaturation at 94 ˚C,
(b) 30 s of annealing at 48˚C, and (c) 2 min of elongation at 72 ˚C. The PCR reaction
was finished with an elongation step of ˚C for 5 min. The PCR products were
72
analyzed on a 0.8% agarose TAE gel and clones that generated DNA fragments
>200bp were flagged for DNA isolation and sequencing. Bacterial cultures were started
using flagged transformants in 5 mL of 50 µg/mL ampicillin-supplemented LB and
shaken overnight at 37˚C. DNA was isolated from these cultures using the QIAGEN
Miniprep DNA Kit, and DNA samples were sent for sequencing to the Michigan State
University Research Technology Support facility (http://rtsf.msu.edu/). Resulting
sequences were translated, BLASTed against the NCBI database, and flagged for
further investigation if they matched an [FeFe]-hydrogenase gene with >55% query
coverage and had a <0.001 e-value.

Whole Genome Amplification—To amplify the entirety of the environmental genomic
DNA samples, the REPLI-g Mini Kit was used (QIAGEN). The reaction was carried out
as recommended by the kit protocol, aliquoting 2.5 µL of template DNA to 50 µL of

136

reaction and incubating at 30 for 16 h
˚C

. After amplification, the enzyme was

inactivated at 65 ˚C for 3 min.

Sequence Extension—The known sequence of each gene fragment was extended in a
sequential process. A 23.5 µL aliquot of REPLI-g amplified genomic sample was mixed
with 2.5 µL of 10X restriction digest buffer, 2.5 µL of 10X BSA, and 1.0 µL of desired
restriction enzyme (New England Biolabs) and incubated at 37 two h. The
˚C for
digestion reaction was inactivated at 80 ˚C for 15 min. The digested sample was ligated
using T4 DNA ligase (New England Biolabs) overnight at room temperature. The ligated
samples were amplified via PCR using GoTaq Green Master Mix with a primer pair
specific to the gene fragment of interest. Resulting PCR products were cloned into the
pGEM-T Easy vector and sequenced as described above. The new sequence was then
compared to the gene fragment of interest. If significant overlap existed, the gene
fragment sequence was extended by adding the newly sequenced fragment to either
the 5’- or 3’-terminus.

137

RESULTS

Targeted Degenerate Primer Design—In an effort to design targeted degenerate
primers, a phylogenetic tree of 148 [FeFe]-hydrogenases was constructed and
individual clades were classified (data not shown). To identify conserved sequences
unique to each clade, multiple sequence alignments were generated (TCOFFEE) using
clade-specific amino acid sequences. Four clades contained regions of conserved
residues unique to those individual clades and degenerate primers of 15-20 bp in length
were designed using these regions (Table S1.1, Figure S1.1abcd). Regions of strictly
conserved residues among all [FeFe]-hydrogenases were not considered in this
analysis.

138

Table S1.1. Table of Clade-Specific [FeFe]-hydrogenase Degenerate Primers.
Strictly conserved amino acid (AA) residues are in bold. Moderately conserved AA
residues are denoted as X/Y, where “X” and “Y” are the residues most commonly
observed.
Primer
EH295
EH296
EH297
EH298
EH299
EH300

Clade
Green Algae
Green Algae
Cellulolytes
Fungi
Fungi
Thermophiles

Strand
Sense
Anti
Sense
Sense
Anti
Anti

Sequence
H/QLEQ/AHP
NAKKLI
LCG/RD/RC
VENPI
PKSAD
IYT/S/EIDE

139

Degenerate Primer Sequence
CAYYTNGARSVNCAYCCNMA
NATNAGYTTYTTNGCRTG
CTNTGYGGNGAYTGYG
GTNGARAAYCCNATHCC
RTCNGCNGAYTTNGG
YTCRTCDATNKHRTADAT

Figure S1.1. Multiple sequence analysis of both clade-specific and general
hydrogenase proteins. Using TCOFFEE, amino acid sequences from either cladespecific proteins or from the Clostridium pasteurianum HydA protein were aligned.
Degenerate primer design was based on regions of high conservation, which are
highlighted in the each of the alignments. A) Green algal clades sequences. B) Fungi
clade sequences. C) Hyperthermophilic bacteria clade sequences. D) Cellulolytic
bacterial clade sequences. E) Various sequences from each mentioned clade and the
C. pasteurianum HydA sequence.

140

Figure S1.1 (cont’d)

141

General Degenerate Primer Pair Design and Testing—The degenerate primers
described above were designed to amplify specific regions that are not ubiquitous to all
[FeFe]-hydrogenase genes. To discover new genes from a wide variety of organisms,
however, general primers which apply to all hydrogenase genes were required.
Previously, Zac Stephens and Pamela Smith from the Hegg lab designed three sets of
degenerate primer pairs which target several regions of high conservation among
[FeFe]-hydrogenase genes (Table S1.2, Figure S1.1e). To optimize PCR conditions
with the general degenerate primers, S. oneidensis MR-1 genomic DNA, which encodes
an [FeFe]-hydrogenase, was used to ascertain the best conditions and degenerate
primer pair combinations for amplification of a valid gene fragment from the DNA. Ideal
PCR conditions were established and are described in the Methods section. The primer
pairs best suited to isolate the S. oneidensis [FeFe]-hydrogenase gene were pair Fe4A
(Fe_4F, Fe_6R) and Fe9 (Fe_9F, Fe_9R) and were used for subsequent degenerate
primer-based PCR amplification reactions.

Investigation

of

G.

ahangari

234

Genomic

DNA—Genomic

DNA

from

the

hyperthermophilic archeon G. ahangari 234 was supplied by Dr. Kazem Kashsefi from
the Department of Microbiology and Molecular Genetics at Michigan State University.
This DNA was amplified by PCR with both the Fe4A and Fe9 primer pairs under a
variety of conditions (i.e. varying [Mg2+] and annealing temperatures). Out of 20 PCR
products initially identified and sequence, no amplified products demonstrated
significant homology to known [FeFe]-hydrogenase sequences.

142

Table S1.2. Table of General [FeFe]-hydrogenase Degenerate Primers. Strictly
conserved amino acid (AA) residues are in bold. Moderately conserved AA residues
are denoted as X/Y, where “X” and “Y” are the residues most commonly observed.
Primer
Fe_6F
Fe_1F
Fe_4F
Fe_9F
Fe_4R
Fe_9R
Fe_1R
Fe_6R

Strand
Sense
Sense
Sense
Sense
Anti
Anti
Anti
Anti

AA Sequence
ADL/MTI/VM/LEE
PM/LFTSCCP
PM/LFTSCCP
PM/LFTSCCP
M/TEAV/AL/IR
M/TEAV/AL/IR
MM/ACP/HG/AGC
MM/ACP/HG/AGC

Degenerate Primer Sequence
GAYYTNACNATHATGGARGAR
CANTCNTGYTGYCCNG
CCNATGTTYACNAGYTGYTGY
CCICARTTYACNWSNTGYTGY
CKNTCNGCYTCNHMYTTYTT
KWNCKNARNGCNGCYTCCAT
RTNNCCNGGRCANGCCAT
RCANCCNCCNGGRCANGCCAT

143

Investigation of Environmental Genomic DNA Samples from YNP—A relatively small
amount of environmental genomic DNA samples from YNP was available for
experimentation. To generate more DNA sample, each environmental DNA sample was
amplified using REPLI-g, a whole genome amplification kit. The products of the REPLI-g
reaction were then used in further experiments.
Environmental genomic DNA samples isolated from 10 distinct sites in the YNP
were kindly donated by the Lucigen Corporation (http://www.lucigen.com/). DNA from
these samples was amplified by PCR using both the Fe4A and Fe9 primer pairs and
performed under the idealized conditions established previously. Nine gene fragments,
out of a total of 60 that were isolated and sequenced, were found to have similarity to
[FeFe]-hydrogenase genes. Of these 9 fragments, 5 were determined to be distinct from
one another and were used for further experiments (Table S1.3). A multiple sequence
alignment of these 5 fragments can be seen in Figure S1.2.

144

Table S1.3. Table of Gene Fragment DNA Sequences
Gene
Fragment
Length Sequence
5sisN3.1
453 bp ATTCCGCAGTTTACGTGGTGTTGTCCAGCCTGG
GTTCGGTAAAAACCTATGGTGCAAAAGAGGTTTG
GAAAGTAAACCCTGAAGACATGTATATGGTTGGC
GTTATGCCCTGTACTGCCAAGAAGTTTGAAGCTT
CGCGTCCAGAGTTCCAAAGTGCTGCCCATTATTG
GCAGACCCAGGGTAGATCAGGCAGCTATCCAGA
TATTGATGTAGTTCTTACTACCAGAGATTTAGCCC
GTCTGTTCAAGAAACTAAACATCGACATCAAAAC
AGTCGCTGAATTTAGCGACAAAGATAATCCGCTA
GCCCAATACAGTGGGGCAGGTACTATTTTTGCCA
ACACCGGCGGTGTAATGGAAGCCGCCTTCCGCT
CA
AZTEC.1
777 bp GCGCAAACAGCTCCATCTGTCAGGGTTGCAATA
GGTGAAGAATTTGGCATGGAACCCGGAAGCATAA
GCACAGGGAAGATGGTTGCTGCACTTAGAAGACT
TGGTTTCGACTATGTATTTGACACAAACTTCGCAG
CTGACCTTACAATTATGGAAGAGGGTTACGAACT
GATTGGAAGGCTCCAAAACGGCGGTAAGTTCCC
AATGTTTACGAGGTGTTGTCCTGGTTGGGTAAACG
AAATGGAGAAAGAATGGCCCGAACTCAGAGAAC
ACCTTTCAACGGCAAAGTCCCCACAACAGATGAT
GAGTAGTCTTGTCAAAACATACTTTGCACAAAAGA
TAGGTGTGAAACCTGAAGATATAGTTATGGTATCA
ATTATGCCATGTACAGCTAAGAAAGACGAAATAA
CAAGACCACAACAATTGGTCGACGGTATCAAAGT
AACGGATTACGTCATTACAACAAGAGAACTCGGA
AAGCTCATTAAACTCAAAGGTATACCATTCGTAAA
TCTTCCAGAAGAACAGTATGATAGCCCACTTGGA
ACATCAACAGGTGCTGCTGCGCTCTTTGGTGTGA
CTGGCGGTGTTATGGAAGCAGCTCTCAGAACAGC
TTACGAAGTTTTGACAGGCGAGAAATTACCGAAA
CTTGTGTTCGAAAGCGTACGAGGCCTGGATGGTG
TAAGAGAAGCAGAAATTGATATCAATGGAAGATG
CTAAACAGCAAGGAACCCTAATTCTTCACCTATT
GCAACGGACAAGACTACTCATCATCTA-3'
AZTEC.2
786 bp CATGGCGGCCGCGGGAATTCGATTCCAATGTTTA
CGAGCTGCTGTCCGGGCTGGATTGCGATGCTGG
AAAAAAGCTATCCGGATCTGATTCCGTATGTGAG
CAGCTGCAAAAGCCCGCAGATGATGCTGGCGGC
GATGGTGAAAAGCTATCTGGCAGAAAAAAAAGG
145

Table S1.3 (cont’d)

MNDWOOD.1

796 bp

CATTGCGCCGAAAGATATGGTGATGGTGAGCATT
ATGCCGTGCACCCGTAAACAGAGCGAAGCGGAT
CGTGATTGGTTCTGCGTGGATGCGGATCCGACCC
TGCGTCAGCTGGATCATGTGATTACCACCGTGGA
ACTGGGCAACATTTTTAAAGAACGTGGCATTAAC
CTGGCGGAACTGCCGGAAGGCGAATGGGATAAC
CCGATGGGCGTGGGCAGCGGCGCGGGCGTGCT
GTTCGGCACCACCGGCGGCGTGATGGAAGCGG
CGCTGCGTACCGCGTATGAACTGTTTACCGGCAC
CCCGCTGCCGCGTCTGAGCCTGAGCGAGGTGCG
TGGCATGGATGGCATTAAAGAGACCAACATTACC
ATGGTGCCGGCGCCGGGCAGCAAATTTGAAGAA
CTGCTGAAACATCGTGCGGCGGCGCGTGCGGAA
GCGGCGGCGCATGGCACCCCGGGCCCGCTGG
CGTGGGATGGCGGCGCGGGCTTTACCAGCGAAG
ATGGCCGTGGCGGCATTACCCTGCGTGTGGGCG
TGGCGAACGGCCTGGGGCACGCGAAAAACTGAT
TACCAAATGCAGGCGGGCGAAGCGAATATGATTT
GTGGGAATATGCATGTCCCGCG
TCCGATGTTCACGAGTTGCTGTCCCAGCTGGGTG
AAGTTTGCCGAACAGTATTACCCCGAGCTGTTGC
CCAATTTGTCCACCTGCAAGTCGCCCCAGCAGA
TGTTTGGCAGCGTGGCCCGCGAGGTTCTTCCTAA
ATTGCTCAATATCAAGCCCGAAAACCTTGTCATTG
TGTCAATTATGCCTTGCACGGCCAAAAAGTATGA
GGCCAAACGCGATGAGTTTATCCATGATGGCCTT
GCCGAGGTTGACCATGTGCTCACTACGGAAGGT
CTTGGCCGCATGATACACGAAACCGGATTACAGT
TCAATAAGCTGAAGCCCGAGTCGTTCGATATGCC
ATTGGGCTTTAAGACCGGAGCAGGGGTAATATTC
GGTAACACTGGCGGAGTGAGCGAAGCAGTATTG
CGTTTCGCCTACGAAAAGATTACCGGTGAAACCC
TGATCGACACCGACATCCGGCAGACCCGTGGTA
TGGAAGGTATTCGCACAGTGGAGATGCAACTCGG
CGAAACCAGAATTAAGCTCGGGATTGCGCATACC
CTCTCGAATGCCCGCAAACTGTGCGATGCCATAG
TGAAAGGCGAAGCCGATTACGACCTTGTGGAAGT
GATGGCCTGCCCGGGCGGCTGTATTGCAGGTGG
CGGACAACCAGTGAGCTTCGACCCCGAATTCAG
ACAAAAGCGAATTCAAGGGATCTACAATGCCGAT
AAACAGCTCGAACTGCACAAATCGCAGGACAATC
146

Table S1.3 (cont’d)
MNDWOOD.2

910bp

CTTATGTTAAGGAGTTGTATCAGAATATTTTAGGCG
GAAATAGGCGGACACAG
CATTGCCGGAGGAGGACAGCCTTATCACCATGG
CAATGATGAGATTGTTAAAAAACGCAGGGAAGCC
ATATTTGAAGAGGATAGGAACAAAAAGATAAGAA
AATCCCATGAGAACAAAGAAATATTGGAACTATA
GCAAAAACCTACTATGCTGAAAAGATGGGAATTG
ACCCCGACAATATTGTTATGGTATCTGTCATGCC
CTGCATAGCAAAGAAAGGCTGAAGCCAAACGTC
CCGAGCTGACAAAGGACGAACACAACAATGTAG
ACATATCAATAACCACCTCCGGGAGCTTGGTGCC
ATGATACATGAAGCGGGAATAGAATTTGCAAAGC
TGCCCGACAGTGAATTTGACAGTCCGCTTGGTGA
ATCCTCGGGGGCAGCCGTAATGTTCGGTACTGCC
GGAGGCGTTATTGAGGCGGCTCTGCGTACAGCC
TCAGAATGGCTGACAGGTGAACCTCTTAAGAAAA
TAGAATTTGAAGATTTAAGGGGTATGGAGGGAGTT
CGCAGAGCTGCTGTCAAAATAGGAGACAAGGAA
CTCAAAATAGGTATTACAAACGGTCTGGGCAATG
CACGGCATATATTGGAGGATATCCGCGACGGAA
AGGCGGACTACCATGCTATTGAGATAATGGCTTG
TCCCGGAGGCTGCATTGCCGGAGGAGGACAGC
CTTATCACCATGGCAATGATGAGATTGTTAAAAA
ACGCAGGGAAGCCATATTTGAAGAGGATAGGAA
CATGAAGATAAGAAAATCCCATGACAACAAAGAA
ATATTGGAATTATATAAAACTTATCTGGGTGAACC
TTTTAACAAAATAGAATTTGAAGATTTAAAGGGTAT
GGAGGGAGTTCTCAGAGCTGCTGTCATACTATGA
TACCAGGAACTCAAAATCACTAATGAATTCT

147

Figure S1.2. Multiple sequence analysis of gene fragment and [FeFe]-hydrogenase
sequences. Using TCOFFEE, amino acid sequences of the translated gene fragments
were compared with known [FeFe]-hydrogenase protein sequences by sequence
alignment, as well as the homolog Nar1p from S. cerevisiae.

148

Full-Length Gene Sequence Determination—To determine the full-length gene
sequence from the identified fragments, a common restriction site was found within
each gene fragment and genomic samples were digested with the desired restriction
enzyme. After inactivating the restriction enzyme, ligase was added to each sample to
ligate digested fragments within the reaction, essentially generating a variety of circular
DNA molecules, some containing the gene fragment of interest. Next, primer pairs were
designed at the 5’- and 3’-ends of the fragment, as well as near the restriction cut-site,
to amplify away from the fragment and into “unknown” gene sequence. The ligation
products were then amplified by PCR using the newly-designed primers (Figure S1.3,
Sequence Extension Method). The resulting PCR products were sequenced, thereby
extending the sequence coverage of the unknown gene.
This method was applied to each of the 5 gene fragments, extending the
fragment gene sequence of each except 5sisN3.1 and AZTEC.2 (Table S1.3). At the
time of this report, none of the full-length genes had been completely sequenced.

149

Figure S1.3. Gene sequence extension methodology. Graphic representation of the
Sequence Extension Method. The gene fragment of interest is light gray, while the
“unknown” sequence is dark gray. A common four-base restriction site (HhaI, GTAC)
which occurs once in the gene fragment is marked by└┐”. Arrows represent primers
“
designed to amplify away from the gene fragment and into unknown sequence.

150

Phylogenetic Analysis of Gene Fragments—To determine a relationship between the
identified fragments and known [FeFe]-hydrogenases, a phylogenetic analysis was
performed (Figure S1.4). The 5sisN3.1 fragment grouped mostly closely with the S.
oneidensis MR-1 hydA gene, but also groups with genes from Desulfovibrio species.
The AZTEC.2 fragment groups well with green algal genes and mostly likely comes
from a a member of this species, whereas the AZTEC.1 fragment demonstrates a
strong relationship to both Fervidobacterium and Thermosipho species’ hydrogenase
genes. The MndWood.1 gene demonstrates a relationship only with the Ruminococcus
gnavus hydA gene. The MndWood.2 fragment was found to associate with
Desulfotomaculum species and grouped loosely with Alkaliphilus metalliredigens and
Clostridium ljungdahlii, species which are obligate anaerobes.

151

Figure S1.4. Evolutionary relationship of translated gene fragments and [FeFe]hydrogenases. Phylogenetic analysis of 64 [FeFe]-hydrogenase amino acid sequences
and translated gene fragments using the MEGA5 software program. Evolutionary
relationships are inferred by the Neighbor-Joining method and 2,000 bootstrap
replicates were performed. Evolutionary distances were calculated by the p-distance
method and the branch lengths are equal to the number of amino acid substitutions per
site. The tree was rooted using Nar1 sequences from H. sapiens and S. cerevisiae.

152

Figure S1.4. (cont’d)

153

DISCUSSION

Current Results and Interpretation—In an effort to discover new and potentially novel
[FeFe]-hydrogenases, degenerate primers were used to amplify genomic DNA samples.
In particular, primer pairs that are specific to [FeFe]-hydrogenase-conserved amino acid
sequences were used for this amplification. Uniquely conserved regions can be
observed within several clades identified via phylogenetic analysis, and these regions
were used to design clade-specific degenerate primers. Currently, no experiments have
been performed utilizing these clade-specific primers, principally due to the samples
available to us. G. ahangari 234 is an archeon and does not match any clade identified
on the phylogram, thereby greatly decreasing the utility of clade-specific primer pairs for
this organism. Additionally, the YNP environmental samples were composed of
unknown genomic DNA and likely contain DNA from many organisms that are not
targets of the clade-specific primers. Thus, our studies utilized primer pairs specific to
strictly conserved amino acid regions (Fe4A and Fe9) in order to isolate the maximum
number of gene fragments available. In future experiments, the clade-specific primers
could be used to target hydrogenase genes in genomic DNA from individual organisms.
G. ahangari 234 genomic DNA was amplified using optimized PCR conditions with
both the Fe4A and Fe9 degenerate primer pairs, and the resulting PCR fragments were
sequenced. None of the resulting PCR fragments were homologous to known [FeFe]hydrogenases. This result is not surprising because all characterized H2-producing
enzymes found in archaea are [NiFe]-hydrogenases. Although an unidentified [FeFe]hydrogenase could be responsible for the noted H2 metabolism of G. ahangari 234,

154

based on the data collected here, a different enzyme is likely responsible for this
metabolism.
Genomic DNA samples from YNP were amplified utilizing the Fe4A and Fe9
degenerate primer pairs and 9 gene fragments were identified. Utilizing multiple
sequence analysis, 4 of the fragments were found to be highly homologous to the
AZTEC.2 gene fragment and, as the purpose of this work was to find novel genes, fulllength determination of these fragments was not pursued. Taking advantage of
restriction sites commonly found in DNA (specifically, short recognition sequences), the
environmental genome samples were digested with restriction enzymes, re-ligated to
form small DNA molecules, and amplified using primers specific to the gene fragments.
The amplified products were sequenced, examined for overlap with the gene fragments,
and added to the fragment sequences when positive overlap was determined (Figure
S1.3). The AZTEC.1, MndWood.1, and MndWood.2 sequences were all extended by
200-700 bp using this method, but unfortunately no new sequence was added to the
5sisN3.1 or AZTEC.2 sequences.
A number of interesting aspects were noted when examining each of the gene
fragments. First, all of the fragments contained canonical [FeFe]-hydrogenase motifs:
“VXXMPTC” and “GGVXMEAAXR”. Second, the well-conserved motif “TSCCP”
matched exactly with the MndWood.1 and AZTEC.2 gene fragment sequences, but was
not observed in the MndWood.2 sequence. This may be due to inaccuracies in
amplification during extension of the gene fragment or to the presence of “filler”
sequence not normally observed in this region of HydA protein sequence. Intriguingly, in
the amino acid sequence of the 5sisN3.1 and AZTEC.1 fragments the serine residue of

155

the “TSCCP” motif is replaced by tryptophan and arginine, respectively. A role for this
serine in hydrogenase activity has yet to be determined, but this residue is strictly
conserved among all known HydA proteins. Although this may be a novel finding, it is
more likely due to errors in amplification and sequencing, as a small change in the
triplet codon for serine (TCN and AGY) can easily lead to tryptophan (TGG) or arginine
(AGG).
A phylogenetic analysis was performed to provide insight into the evolutionary
relationship between each of the gene fragments and known hydA genes. Little
information could be gleaned from the MndWood.1 fragment, which only showed a
relationship to the HydA protein from R. gnavus, a gut microbe that is unlikely to be
closely-related to YNP hot springs microbes. The MndWood.2 fragment showed
similarity to several strict anaerobes, but more information on the complete gene
sequence is needed for further identification. The 5sisN3.1 fragment clustered with
genes from bacteria commonly involved in sulfate-reduction, indicating that this
fragment may originate from similar organisms. Relationships between the AZTEC
fragments were among the most interesting, clustering well with green algal (AZTEC.2)
and hyperthermophilic bacterial proteins (AZTEC.1). As seen in Figure S1.4 and
discussed previously in relation to Volvox carteri (Chapter 4), green algal [FeFe]hydrogenase genes were presumably acquired via horizontal gene transfer, and the two
clades discussed previously are hypothesized to share a common ancestor gene.
Because these gene fragments were isolated from the same environmental genomic
DNA sample, this strongly suggests that organisms with homologous genes to these
fragments currently coexist in YNP hot springs. Assuming that AZTEC.2 and AZTEC.1

156

are green algal and hyperthermophilic bacterial in origin, respectively, this further
suggests that green algae acquired [FeFe]-hydrogenases via horizontal gene transfer,
and this event may have occurred in a YNP hot spring-like environment. Additional
evidence, such as determining the full-length sequence of the AZTEC fragments would
be required to lend more support to this claim.
Due to a change in focus of the thesis project, no additional experiments or analyses
were undertaken.

Future

Directions—Several

gene

fragments

have

been

isolated

from

YNP

environmental genomic samples and the translated sequences show strong similarity to
known HydA proteins. The full-length sequence of these fragments remains to be
determined and this can be accomplished using the Sequence Extension method
described previously. Additionally, using knowledge gained from the phylogenetic
analysis, clade-specific degenerate primers could be used to extend sequence
knowledge of the AZTEC fragments, which demonstrated clustering with both the green
algal and the hyperthermophilic bacterial clades. Once the full-length sequences are
determined, the genes can be amplified, cloned into the pAC-BAD expression vector,
and transformed into the S. oneidensis MR-1 ΔhydAΔhyaB overexpression system. The
expressed and purified gene products can then be characterized on the basis of H2
evolution activity, thermostability, salt tolerance, pH stability, and resistance to O2induced inactivation. In addition, the full-length sequences of the putative genes can be
directly compared to known hydA sequences, and a more definitive evolutionary
analysis conducted by phlyogram.
157

Conclusions—Using a phylogenetic tree of HydA homologs, we designed several cladespecific degenerate primers, concentrating on regions of residues with high
conservation between HydA proteins within that clade. Degenerate primer pairs
designed to target highly-conserved regions of all HydA genes were utilized to optimize
conditions for PCR amplification of S. oneidensis MR-1 genomic DNA. These conditions
were used to PCR amplify putative hydA gene fragments from G. ahangari 234 genomic
DNA and YNP environmental genomic DNA. Under the conditions tested, 0 and 9
amplicons with homology to hydA genes were identified from G. ahangari 234 and the
YNP samples, respectively. Extension of the YNP gene fragment sequences was
attempted for 5 of the 9 YNP fragments, meeting with moderate success. Future work
will focus on determining the full-length sequence of these fragments and characterizing
the gene products.

158

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161

SUPPLEMENTAL CHAPTER 2

Genome Annotation and Investigation of H2 Metabolism of Nannochloropsis
1
oceanica CCMP1779

1

This chapter is adapted and contains data from Vieler A., Wu G., Tsai C.-H., Bullard
B., Cornish A. J., Harvey C., Reca I.-B., Thornburg C., Achawanantakun R., Buehl C. J.,
Campbell M. S., Cavalier D., Childs K. L., Clark T. J., Deshpande R., Erickson E.,
Ferguson A. A., Handee W., Kong Q., Li X., Liu B., Lundback S., Peng C., Roston R. L.,
Sanjaya, Simpson J. P., TerBush A., Warakanont J., Zäuner S., Farre E. M., Hegg E. L.,
Jiang N., Kuo M.-H., Lu Y., Niyogi K. K., Ohlrogge J., Osteryoung K. W., Shachar-Hill
Y., Sears B. B., Sun Y., Takahashi H., Yandell M., Shiu S.-H., Benning C. (2012).
Genome, functional gene annotation, and nuclear transformation of the heterokont
oleaginous alga Nannochloropsis oceanica CCMP1779. PLoS Genet. Published
Online Ahead of Print.

162

INTRODUCTION

In recent years, microalgae have shown promise as sustainable sources of
renewable fuels, such as biodiesel, bioethanol, and biohydrogen (1, 2, 3, 4). To date,
only a handful of microalgal species have been characterized with appropriate depth for
biofuel production, especially at the industrial scale (5). Among well-characterized
microalgae, few of these organisms have properties ideal for the targeted engineering
required in high-volume fuel production (e.g., efficient knock-out/knock-down of specific
genes and optimized expression of transgenes).
Nannochloropsis oceanica CCMP1779 is a photoautotrophic microalga that lies
within the class Eustigmatophyceae of Heterokontophyta (6). Organisms in the genus
Nannochloropsis have been targeted for biofuel production due to lipid accumulation
under stress and ease of both gene deletion and expression (7, 8, 9). In addition, N.
oceanica CCMP1779 can utilize sugars from the media, is sensitive to a range of
antibiotics, and can integrate selectable markers into its nuclear genome (10).
Recently, the N. oceanica CCMP1779 genome was sequenced by Drs. Christoph
Benning and Shin-Han Shiu (Michigan State University) in an effort to characterize the
putative metabolism further and investigate unique properties at the genomic level (10).
The genome was found to be 28.7 Mb in size and a total of 11,973 genes were
identified from RNA sequencing data. Although automated annotation programs were
utilized, specific sets of genes were manually annotated to improve identification
accuracy and to incorporate the genes into a larger biochemical perspective (10).

163

Due to our knowledge in respiration, intracellular electron transfer, and hydrogen
metabolism, our research group manually annotated genes in the N. oceanica
CCMP1779 genome. We identified several suites of genes required for the described
processes and, based on the presence of hydrogenase genes, we also tested for and
measured H2 production from N. oceanica CCMP1779.

164

METHODS

Manual Annotation of Genome—Nucleic acid sequences for genes of interest were
collected from the Chlamydomonas reinhardtii, Ectocarpus siliculosus, and Phytophthora infestans genomes using the NCBI database (http://www.ncbi.nlm.nih.gov/).
Homologous sequences in the N. oceanica CCMP1779 genome were discovered using
TBLASTX with a threshold of <0.001 and
≥55% query coverage.

Putative genes were

confirmed by BLASTX against the NCBI database, again with a threshold of <0.001 and
≥55% query coverage required for confirmation. The genes were then assigned a
putative identity, and the designation and location in the genome noted.

Growth Conditions—N. oceanica CCMP1779 strains were grown in 250 mL Erlenmeyer
flasks containing 50 mL F/2 Nutrient Media (17 g/L Tropic Marine Salt enriched with f/2
nutrients, trace elements and vitamins (11), and 40 mM bicarbonate and 15 mM Tris
buffer [pH 7.6] to prevent carbon limitation). The cultures were shaken at 105 rpm in
continuous light (85 μmol photons m

-2 -1

s ) and kept at 22 °C (12).

Hydrogen Evolution Assay—To acclimate the cells to anaerobiosis, a culture was
concentrated to 75 μg chlorophyll/mL in 2 mL of anaerobic F/2 nutrient media, sealed in
a 10 mL serum vial (Wheaton), and shaken at 100 rpm at 22 in the dark
˚C

for 4 h.

“Aerobic” cells were treated similarly, except that they were exposed to the atmosphere
during the 4 h acclimation. To measure H2 evolution, 0.1 mL of aerobically- or
anaerobically-acclimated cells were added to 1.9 mL of H2 evolution assay buffer (F/2
165

nutrient media, 100 mM sodium dithionite, 10 mM methyl viologen) in a 10 mL serum
vial and incubated with shaking in the dark. H2 accumulation was measured over time
by injecting 10 µL of gas from the headspace into a TRACE GC Ultra Gas
Chromatograph (Thermo Scientific) using a 100 µL syringe. Absolute amounts of H2
were determined by comparison of the peak area against a standard curve.

166

RESULTS

Manual Gene Annotation—The N. oceanica CCMP1779 nuclear genome was
sequenced by 454 and Illumina technology. RNA-sequencing was utilized to generate a
de novo assembly of 65,321 gene transcripts, of which 35,756 were then mapped onto
the final genome assembly (10).
Oxidative phosphorylation is ubiquitous among aerobically-grown organisms.
Genes were found within the N. oceanica CCMP1779 genome encoding key subunits of
NADH dehydrogenase (complex I), succinate dehydrogenase (complex II), cytochrome
c reductase (complex III), cytochrome c oxidase (complex IV), and FoF1 ATP synthase
(complex V), as well as numerous accessory proteins required for the proper folding
and the insertion of cofactors into these protein complexes (Table S2.1). In addition,
three genes were identified with strong homology to alternative oxidases, proteins which
allow for the reduction of O2 without the concomitant synthesis of ATP (13). Together,
the presence of these genes strongly implies a typical respiratory electron transfer chain
used to couple O2 reduction to ATP synthesis.
Reactive oxygen species (ROS) are unavoidable byproducts of cellular
respiration and photosynthesis and are able to cause cellular damage (14). Examining
the genome, 4 genes with homology to ROS scavenging genes which are typically
targeted to both the cytosol and mitochondria were identified (Table S2.1), suggesting a
functional ROS scavenging system is present in N. oceanica CCMP1779.

167

Table S2.1. Manually annotated genes in N. oceanica CCMP1779
Description
Name ID
HYDROGEN
[FeFe]-hydrogenase
[FeFe]-hydrogenase maturation
factor
[FeFe]-hydrogenase maturation
factor

HydA1 CCMP1779_5967-mRNA-1
augustus_maskedHydG nanno_856abinit-gene-0.2-mRNA-1*
HydF

CCMP1779_5970-mRNA-1

HydE

augustus_maskednanno_856abinit-gene-0.1-mRNA-1*

Sdh2

CCMP1779_10056-mRNA-1

Sdh1

CCMP1779_10769-mRNA-1

Sdh3

CCMP1779_9122-mRNA-1

succinate dehydrogenase subunit D

Sdh4

heme O synthase

Cox10

cytochrome c oxidase subunit 6b

Cox12

heme A synthase
cytochrome c oxidase assembly
cytochrome c oxidase assembly
cytochrome c oxidase assembly
cytochrome c oxidase Cu chaperone
cytochrome c oxidase chaperone
cytochrome c oxidase subunit II
cytochrome c oxidase subunit I
cytochrome c oxidase subunit III

Cox15
Cox11
Shy1
Sco1
Cox17
Ssc1
Cox2
Cox1
Cox3

alternative oxidase 2

Aox2

CCMP1779_3454-mRNA-1
augustus_masked-nanno1077
-abinit-gene-1.10-mRNA-1*
augustus_masked-nanno604abinit-gene-0.21-mRNA-1*
CCMP1779_925-mRNA-1
CCMP1779_4145-mRNA-1
CCMP1779_22-mRNA-1
CCMP1779_6751-mRNA-1
CCMP1779_9640-mRNA-1
CCMP1779_8242-mRNA-1
CUFF.84573.1+**
CUFF.95393.1+**
CUFF.95359.1+**
augustus_masked-nanno2785
-abinit-gene-0.2-mRNA-1*

[FeFe]-hydrogenase maturation
factor
RESPIRATION
succinate dehydrogenase
flavoprotein
succinate dehydrogenase subunit A
succinate dehydrogenase
cytochrome b subunit

*This gene model is from augustus or snap gene annotation and was found
superior to the final maker annotation after manual examination.
Transcript from expressed sequence tag data, as no final gene model was
available.
168

Table S2.1 (cont’d)
Description
alternative oxidase 2
alternative oxidase 3
NADH:ubiquinone oxidoreductase
subunit 10
NADH:ubiquinone oxidoreductase
subunit 17
NADH:ubiquinone oxidoreductase
subunit E
NADH:ubiquinone oxidoreductase
subunit F
NADH:ubiquinone oxidoreductase
subunit I
NADH:ubiquinone oxidoreductase
subunit 9
NADH:ubiquinone oxidoreductase
18 kDa subunit
NADH:ubiquinone oxidoreductase
subunit B14
NADH:ubiquinone oxidoreductase
subunit B16
NADH:ubiquinone oxidoreductase
subunit B18
NADH:ubiquinone oxidoreductase
subunit 11
NADH:ubiquinone oxidoreductase
subunit S4
NADH:ubiquinone oxidoreductase
subunit S5
NADH dehydrogenase subunit 4

Name
Aox1

ID
augustus_masked-nanno1051
-abinit-gene-0.9-mRNA-1*
Aox3
augustus_masked-nanno3659
-abinit-gene-0.0-mRNA-1*
Nuo10 CCMP1779_10235-mRNA-1
Nuo17 CCMP1779_11866-mRNA-1
NuoE

CCMP1779_9179-mRNA-1

NuoF

CCMP1779_4574-mRNA-1

NuoI

CCMP1779_7640-mRNA-1

Nuo9

CCMP1779_6889-mRNA-1

Nuo18k CCMP1779_11055-mRNA-1
NuoB14 CCMP1779_6453-mRNA-1
NuoB16 CCMP1779_9016-mRNA-1
NuoB18 CCMP1779_211-mRNA-1
Nuo11 augustus_masked-nanno_224abinit-gene-1.7-mRNA-1
NuoS4 CCMP1779_2896-mRNA-1
NuoS5 CCMP1779_10934-mRNA-1
Nuo4

CUFF.95343.1+**

*This gene model is from augustus or snap gene annotation and was found
superior to the final maker annotation after manual examination.
**Transcript from expressed sequence tag data, as no final gene model was
available.

169

Table S2.1 (cont’d)
Description
NADH dehydrogenase subunit 5
NADH dehydrogenase subunit 1
NADH:ubiquinone oxidoreductase
subunit 7
NADH dehydrogenase subunit 2
NADH dehydrogenase subunit 3
F1F0 ATP synthase gamma subunit

Name
Nuo5
Nuo1

ID
CUFF.84531.1+**
CUFF.95403.1+**

Nuo7

CUFF.95391.1+**

CUFF.95399.1+**
CUFF.95361.1+**
augustus_masked-nanno_54
42-abinit-gene0.2-mRNA-1*
Atp5C1 CCMP1779_10955-mRNA-1
Atp5A1 CCMP1779_10718-mRNA-1
Atp5B
CCMP1779_9984-mRNA-1
Atp6
CUFF.95401.1+**

Nuo2
Nuo3
Atp3

ATP synthase O subunit
F0F1 ATP synthase subunit alpha
F-type H-ATPase beta subunit
ATP synthase F0 subunit 6
ATP synthase mitochondrial F1
AtpAF2 CCMP1779_10134-mRNA-1
assembly factor 2
ATP synthase mitochondrial F1
AtpAF1 CCMP1779_6914-mRNA-1
assembly factor 2
ubiquinol cytochrome c
CytC1 CCMP1779_9301-mRNA-1
cytochrome c1
ubiquinol cytochrome c reductase
Qcr7
CCMP1779_9021-mRNA-1
14kDa subunit
cytochrome b-c1 complex subunit
RipI
CCMP1779_8327-mRNA-1
Rieske
cytochrome b-c1 complex 50 kDa
Qcr1
CCMP1779_635-mRNA-1
subunit 1
ubiquinol-cytochrome c reductase
Risp1
CCMP1779_8327-mRNA-1
iron-sulfur subunit 1
**
cytochrome b
CytB
CUFF.95357.1+
*This gene model is from augustus or snap gene annotation
and was found superior to the final maker annotation after
manual examination.
**Transcript from expressed sequence tag data, as no final
gene model was available.

170

Table S2.1 (cont’d)
Description
FERREDOXINS
[2Fe2S] ferredoxin
ferredoxin, adrenodoxin-like protein
putative chloroplast ferredoxin 1
putative chloroplast ferredoxin 6
Rieske [2Fe2S] ferredoxin 1
Rieske [2Fe2S] ferredoxin 2
putative chloroplast ferredoxin 2
putative chloroplast ferredoxin 5
putative chloroplast ferredoxin 3

Name

ID

Fdx
Mfdx1
Fdx1
Fdx6
rFDX1
rFDX2
Fdx2
Fdx5
Fdx3

CCMP1779_688-mRNA-1
CCMP1779_10471-mRNA-1
CCMP1779_7894-mRNA-1
CCMP1779_3054-mRNA-1
CCMP1779_7488-mRNA-1
CCMP1779_4457-mRNA-1
CCMP1779_2103-mRNA-1
CCMP1779_7881-mRNA-1
CCMP1779_1904-mRNA-1

IRON SULFUL CLUSTER BIOSYNTHESIS
iron-sulfur cluster assembly protein
IscA1
iron-sulfur cluster assembly protein
IscA2
iron-sulfur cluster biosynthesis
Nar1
protein
ROS SCAVENGING
glutathione synthase
microsomal glutathione Stransferase
superoxide dismutase
glutathione peroxidase

CCMP1779_11149-mRNA-1
CCMP1779_6581-mRNA-1
CCMP1779_4004-mRNA-1

GST

CCMP1779_1871-mRNA-1

mGST

CCMP1779_9028-mRNA-1

SOD
GPX

CCMP1779_6610-mRNA-1
CCMP1779_10092-mRNA-1

171

A number of photosynthetic microorganisms are capable of H2 production or
oxidation under anaerobic conditions. The genome was investigated for genes involved
in H2 metabolism and a single gene was identified to encode a putative [FeFe]hydrogenase (hydA), as well as three genes coding for proteins required in
hydrogenase maturation (hydE, hydF, hydG) (Table S2.1). In addition, these genes are
organized in an operon-like gene cluster (Figure S2.1) similar to the cluster seen in V.
carteri (Chapter 4).
Ferredoxins are proteins integral to much of intracellular electron transport within
an organism, especially for delivery of electrons during photosynthesis, H2 production,
+

and NAD(P) reduction (15, 16, 17). Ferredoxins require iron-sulfur clusters for electron
transfer activity, and iron-sulfur cluster biosynthesis enzymes are responsible for the
assembly and insertion of these clusters in a variety of proteins (18). Within the
genome, 9 genes were found with homology to ferredoxins, while 3 putative iron-sulfur
cluster biosynthesis genes were identified (Table S2.1).

172

Figure S2.1. [FeFe]-hydrogenase operon-like gene cluster in N. oceanica CCMP1779. Four genes with sequence
similarity to hydA, hydE, hydF, and hydG are arranged within 14 kb of one another.

173

Hydrogen Evolution—To test for its ability to evolve H2, anaerobically-acclimated N.
oceanica CCMP1779 was incubated both with and without an abiotic electron donor
system and accumulation of H2 in the headspace was then measured. Headspace H2
accumulation could be observed when an electron donor was supplied, whereas
unsupplemented culture did not accumulate H2 even 48 s after the beginning of the
assay (Figure S2.2). Cells that were acclimated to aerobiosis produced considerably
less H2 in the headspace over the same 48 hour-time period, which was likely due to an
initial lack of proteins required for H2 production until anaerobiosis was established
during the assay.

174

Figure S2.2. H2 evolution rate of aerobically- or anaerobically-acclimated N.
oceanica CCMP1779 cells. Following a 4 h acclimation period, cells were incubated in
F/2 media with and without an abiotic electron donor system. H2 accumulation in the
headspace was measured at 3, 24, and 48 h time-points and normalized on a per mg
chlorophyll basis. Error bars denote S.D. (n ≥ 3).

175

DISCUSSION

N. oceanica CCMP1779 is a photoautotrophic microalga that accumulates high
concentrations of lipids and fatty acids under a variety of conditions. To gain a greater
perspective on this organism, the genome was sequenced as a joint venture between
Drs. Benning and Shiu at Michigan State University. Manual annotation of the genome
was accomplished through collaboration with various experts in metabolic processes,
including the Hegg lab.
Before the results detailed in this report were published, a manuscript detailing
the genomic sequence of Nannochloropsis gaditana CCMP526, a close relative of N.
oceanica CCMP1779, was released (19). When compared, the two genomes were
found to be similar in sequence length, although the N. oceanica CCMP1779 genome
contained more predicted protein models. The N. oceanica CCMP1779 genome was
also found to contain ~6,500 species-specific genes in comparison to N. gaditana
CCMP526, using defined orthologous group comparisons (10). These results further
affirm the differences that exist between the species and provide a rationale for studying
both organisms.
To help confirm gene identity during annotation, putative genes were BLASTed
against the NCBI database. The majority of highly homologous genes were encoded by
the genomes of two organisms, Ectocarpus siliculosus and Phytophthora infestans.
Both of these organisms are classified as heterokonts. E. siliculosus is a filamentous
brown alga that forms large colonies on water plants, whereas P. infestans is an
infectious oocyte that colonizes potatoes and tomatoes. Because of the similarity
176

between the noted homologs, genes from these organisms were utilized to target genes
in N. oceanica CCMP1779 that were difficult to identify using typical gene sequences.
The N. oceanica CCMP1779 genome was examined for genes important in
cellular respiration, ROS scavenging, H2 metabolism, electron transport, and iron-sulfur
cluster biosynthesis. A majority of the genes examined were positively identified,
implying that these metabolic processes and pathways are present in the organism.
Several mitochondrial genes required for respiration (Cox1, Cox2 and others) could only
be identified from expressed transcripts, as the mitochondrial genome was not
sequenced. This suggests that the annotation process may have missed genes
encoded by the chloroplast and mitochondrial genomes that were not expressed under
the tested conditions.
The N. oceanica CCMP1779 hydrogenase genes are within a 12 kb sequence of
the genome, organized in an operon-like gene cluster (Figure S2.1), similar to a cluster
observed in the V. carteri genome. Unlike V. carteri, the maturation proteins hydE and
hydF have not fused to form the atypical green algal HYDEF (green algal
nomenclature). Also unlike green algae, hydA more closely resembles bacterial [FeFe]hydrogenase genes, containing F-domains that ligate additional iron-sulfur clusters.
When BLASTed against the NCBI database, three of the genes had the most similarity
to green algal sequences, in contrast to hydF, which shared the most sequence identity
to a gene in Nagleria gruberi, a non-photosynthetic amoeba. Together, these results
may suggest that this hydrogenase cluster in N. oceanica CCMP1179 shares ancestry
with the genes acquired in green algae and that comparatively less genetic drift has
occurred since this acquisition. Finally, measured rates of H2 production and the
177

presence of putative [FeFe]-hydrogenase genes within the genome suggest that N.
oceanica CCMP1779 has functional hydrogenase genes that allow for H2 metabolism
under anaerobiosis.
In summary, the N. oceanica CCMP1779 genome contains putative genes
involved in a variety of basic metabolic processes. Of particular interest were several
genes required for H2 production. A putative role for these genes in H2 metabolism was
strengthened by experiments performed under dark anaerobiosis with an electron donor
system, as N. oceanica CCMP1779 generated appreciable amounts of H2 under these
conditions.

178

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