BIOCHEMICAL, SPECTROSCOPIC, AND STRUCTURAL
INVESTIGATIONS ON [FeFe]-HYDROGENASE MATURATION
AND COMPLEX METALLOCLUSTER ASSEMBLY
by
David Wayne Mulder
A dissertation submitted in partial fulfillment
of the requirements for the degree
of
Doctor of Philosophy
in
Biochemistry
MONTANA STATE UNIVERSITY
Bozeman, Montana
April 2010
©COPYRIGHT
by
David Wayne Mulder
2010
All Rights Reserved
ii
APPROVAL
of a dissertation submitted by
David Wayne Mulder
This dissertation has been read by each member of the dissertation committee and
has been found to be satisfactory regarding content, English usage, format, citation,
bibliographic style, and consistency and is ready for submission to the Division of
Graduate Education.
Prof. John W. Peters
Approved for the Department of Chemistry and Biochemistry
Prof. David J. Singel
Approved for the Division of Graduate Education
Dr. Carl A. Fox
iii
STATEMENT OF PERMISSION TO USE
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doctoral degree at Montana State University, I agree that the Library shall make it
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David Wayne Mulder
April 2010
iv
ACKNOWLEDGEMENTS
I would like to thank my advisor, Professor John Peters, for giving me the
opportunity to study and conduct research in a truly exciting project in his laboratory and
for his constant council and positive motivation throughout graduate school.
I am
grateful to John for his mentorship both in science and daily life and for making graduate
school a pleasant experience. I also would like to thank Professors Joan Broderick and
Robert Szilagyi for guidance and many helpful discussions while conducting research.
I am very grateful to all the members of the Peters lab for creating a positive, fun,
and healthy research environment and for much help over the past 5 years in solving
many different research problems. I express gratitude too toward Eric Shepard for many
helpful conversations and advice while doing research.
Lastly, I would like to express many thanks to my friends from Gallatin Valley
Presbyterian Church, my mom Nancy and dad Jim, brothers and sisters Doug and Robyn,
Sharon and Kevin, Beth and Ron, and fiancé Linnea Butler for never-ending
encouragement, support, and confidence in me to complete this degree. They were all an
enormous factor toward making this possible and I am forever grateful.
v
DEDICATION
In loving memory of Dan Russcher (Dec 19, 1982 – Feb 4, 2006)
vi
TABLE OF CONTENTS
1. INTRODUCTION ...........................................................................................................1
Biological Role, Diversity, and Structure of Hydrogenases ...........................................4
[NiFe]-Hydrogenase Functional Diversity .................................................................6
[NiFe]-Hydrogenase Structure and Function ..............................................................8
Active Site Structure ..............................................................................................9
[FeFe]-Hydrogenase Functional Diversity ...............................................................11
[FeFe]-Hydrogenase Structure and Function ............................................................13
Active Site Structure ............................................................................................15
Ligand Exchangeable Site and H2 Catalysis ........................................................17
Dithiolate Ligand ..................................................................................................19
Oxygen Sensitivity and Model Complexes ..........................................................21
Hydrogenase Maturation ...............................................................................................22
Other Maturation Systems ........................................................................................22
[NiFe]-Hydrogenase Maturation ...............................................................................25
[FeFe]-Hydrogenase Maturation ...............................................................................27
Radical SAM Proteins ..........................................................................................29
Radical SAM Based Hypothesis ..........................................................................33
In Vitro Activation ...............................................................................................35
HydF as a Scaffold Protein ...................................................................................36
Substrate for CO and CN- Ligands .......................................................................37
Substrate for Dithiolate Ligand ............................................................................39
Current Hypothesis ...............................................................................................41
Research Directions .......................................................................................................42
2. ACTIVATION OF HYDA∆EFG REQUIRES A PREFORMED [4Fe-4S] CLUSTER .....
.......................................................................................................................................45
Introduction ...................................................................................................................45
Experimental Procedures ...............................................................................................48
Cloning and Cell Growth Conditions .......................................................................48
HydA∆EFG Purification ..............................................................................................49
Assays .......................................................................................................................51
Electronic Absorption Spectroscopy.........................................................................51
EPR Spectroscopy.....................................................................................................52
Mössbauer Spectroscopy ..........................................................................................52
Fe K-edge X-ray Absorption Spectroscopy ..............................................................53
Reconstitution of Iron-Sulfur Clusters in HydA∆EFG ................................................54
Preparation of Apo-HydA∆EFG and Reconstitution ...................................................55
Results and Discussion ..................................................................................................55
HydA∆EFG Binds a [4Fe-4S] Cluster .........................................................................55
vii
TABLE OF CONTENTS CONTINUED
Mössbauer Spectroscopic Characterization of HydA∆EFG ........................................59
Fe K-edge X-ray Absorption Spectroscopic Characterization of HydA∆EFG ............63
Reconstitution of the [4Fe-4S] Cluster in HydA∆EFG................................................68
Metal Chelation and Reconstitution..........................................................................69
Relevance to H-cluster Biosynthesis ........................................................................71
Conclusions ...................................................................................................................74
3. STEPWISE [FeFe]-HYDROGENASE H-CLUSTER ASSEMBLY REVEALED
IN THE STRUCTURE OF HYDA∆EFG ........................................................................76
Introduction ...................................................................................................................76
Results and Discussion ..................................................................................................79
Overall Structure .......................................................................................................79
Active Site Structure .................................................................................................84
Difference Fourier Analysis .................................................................................85
Channel Formation and Cluster Insertion .................................................................87
Implications for H-cluster Biosynthesis....................................................................89
Complex Metallocluster Assembly ...........................................................................89
Evolutionary Discussion ...........................................................................................90
Conclusions ...................................................................................................................92
Experimental Procedures ...............................................................................................93
Structure Determination and Refinement .................................................................93
Phylogenetic Analysis ...............................................................................................95
4. CONCLUDING REMARKS ........................................................................................97
REFERENCES CITED ....................................................................................................101
APPENDIX A: Table of Accession Numbers for Known HydA, HydE, HydF,
and HydG Homologues Compiled for Chapter 3 ................................127
viii
LIST OF TABLES
Table
Page
2.1. Representative fitting parameters for the C. reinhardtii HydA∆EFG EXAFS
spectrum using Fe-S composition from Mössbauer measurements .......................65
2.2. Representative fitting parameters for the C. reinhardtii HydA∆EFG EXAFS
spectrum using Fe-S composition from Mössbauer measurements including
the free Fe(II) content ..............................................................................................68
3.1 Data collection and refinement statistics for the x-ray crystal structure
determination of C. reinhardtii HydA∆EFG ............................................................95
ix
LIST OF FIGURES
Figure
Page
1.1. Active sites of [NiFe]-, [FeFe]-, and [Fe]-hydrogenases .......................................2
1.2. Active site of complex metalloenzymes .................................................................4
1.3. Bidirectional NAD-linked [NiFe]-hydrogenase .....................................................7
1.4. X-ray crystal structure of the [NiFe]-hydrogenase from D. gigas .........................8
1.5. [NiFe]-hydrogenase active site .............................................................................10
1.6. H-cluster binding motif alignment .......................................................................12
1.7. X-ray crystal structure of [FeFe]-hydrogenases CpI and DdH ............................14
1.8. Homology model of Chlamydomonas reinhardtii ...............................................15
1.9. [FeFe]-hydrogenase active site .............................................................................16
1.10. H-cluster Hox, Hred, and HoxCO states...................................................................18
1.11. H-cluster with di(thiolmethyl)amine ligand .........................................................20
1.12. P-cluster and FeMo-co of nitrogenase..................................................................23
1.13. Scheme for Mo-nitrogenase biosynthesis.............................................................24
1.14. Scheme for [NiFe]-hydrogenase biosynthesis ......................................................26
1.15. Genomic arrangement of [FeFe]-hydrogenases biosynthetic genes.....................28
1.16. [4Fe-4S] cluster reaction center of radical SAM proteins ....................................30
1.17. Radical SAM protein sequence alignment and reactions .....................................32
1.18 Hypothetical scheme for [FeFe]-hydrogenase maturation and H-cluster
biosynthesis based on parallels to radical SAM chemistry ..................................34
1.19 X-ray crystal structure of HydE from T. maritima ...............................................40
x
LIST OF FIGURES CONTINUED
Figure
Page
1.20 Current hypothetical scheme for [FeFe]-hydrogenase maturation and
H-cluster biosynthesis ..........................................................................................41
2.1 SDS-PAGE gel and UV-visible spectra of C. reinhardtii HydA∆EFG ..................56
2.2 EPR spectrum of reduced C. reinhardtii HydA∆EFG ............................................58
2.3 Temperature and power dependence of the S = ½ signal for C. reinhardtii
HydA∆EFG .............................................................................................................59
2.4 Mössbauer spectra of C. reinhardtii HydA∆EFG ...................................................60
2.5 Mössbauer spectra of C. reinhardtii HydA∆EFG ...................................................62
2.6
FT-EXAFS plot and individual EXAFS contributions for C. reinhardtii
HydA∆EFG with a representative fit containing Fe…Fe, Fe-S(sulfide), and
Fe-S(thiolate) scattering paths ..............................................................................64
2.7 FT-EXAFS plot and individual EXAFS contributions for C. reinhardtii
HydA∆EFG with a representative fit containing Fe…Fe, Fe-S(sulfide),
Fe-S(thiolate), and Fe-low Z(O/N) scattering paths .............................................67
2.8 Flow scheme for different forms of C. reinhardtii HydA∆EFG and
corresponding iron content and hydrogenase activity upon in vitro
activations .............................................................................................................70
2.9 Hypothetical scheme for [FeFe]-hydrogenase maturation ...................................73
3.1 Ball and stick representation of the H-cluster ......................................................77
3.2 X-ray crystal structure of C. reinhardtii HydA∆EFG and comparison to the
x-ray crystal of CpI...............................................................................................80
3.3 Unrooted phylogram of representative HydA and HydA homologs ....................82
3.4 Phylogenetic reconstruction of HydA and HydA homologs ................................83
3.5 Active site comparison between C. reinhardtii HydA∆EFG and Clostridium
pasteurianum (CpI) ..............................................................................................84
xi
LIST OF FIGURES CONTINUED
Figure
Page
3.6 2Fo – Fc, Fo – Fc, and anomalous difference Fourier map for the active site
of C. reinhardtii HydA∆EFG ..................................................................................86
3.7. Channels for insertion into hydrogenase and nitrogenase during complex
Fe-S-cluster assembly...........................................................................................88
xii
ABSTRACT
Metals are present in nearly half of all enzymes, often at the active site, where
they modulate catalytic function. Some of these metalloenzymes exist with a single
bound metal ion while many others contain complex metal clusters. Complex FeS
assemblies are associated with the interconversion of the small molecules H2, CO, CO2,
N2, and NH3. One such complex metalloenzyme, [FeFe]-hydrogenase, catalyzes the
reversible oxidation of molecular H2. The active site of [FeFe]-hydrogenases, the Hcluster, exists as a [4Fe-4S]-subcluster bridged by a protein thiolate ligand to a 2Fesubcluster which contains biologically unique CO and CN- ligands and a dithiolate
ligand. The H-cluster is synthesized by the activities of the hydrogenase maturation
enzymes HydE, HydF, and HydG and until recently little was known concerning the
biosynthetic pathway for the H-cluster. The results presented here provide significant
insight into the stepwise mechanism of H-cluster biosynthesis. Biochemical and
spectroscopic characterization of the structural [FeFe]-hydrogenase enzyme expressed in
a genetic background devoid of maturation genes hydE, hydF, and hydG (HydA∆EFG)
indicates by the presence of a [4Fe-4S] cluster required for [FeFe]-hydrogenase
activation that the [4Fe-4S]-subcluster and 2Fe-subcluster of the H-cluster are
synthesized independently. The determination of the x-ray crystal structure of HydA∆EFG
confirms this by revealing the presence of the [4Fe-4S]-subcluster and an open binding
pocket for the 2Fe-subcluster, indicating that H-cluster synthesis is directed in a stepwise
manner with synthesis and insertion of the [4Fe-4S]-subcluster occurring first by
generalized host cell machinery followed by synthesis and insertion of the 2Fe-subcluster
by specialized hyd encoded maturation machinery. The structure also reveals that
insertion of the 2Fe-subcluster occurs through a positively charged channel that collapses
following incorporation, as a result of conformational changes in two conserved loop
regions. By utilizing complementary gene data base searching with these structural
studies, new insight is made known into the evolutionarily relationships between [FeFe]hydrogenases present in microorganisms and the eukaryotic Nar1 family of proteins
which function in iron-sulfur cluster biosynthesis. The work presented as a whole, by
establishing parallels to complex metal cofactor biosynthesis in nitrogenase, reveals
unifying themes in complex metal cluster assembly and fundamental features of
metalloenzyme evolution.
1
CHAPTER 1
INTRODUCTION
The [NiFe]- and [FeFe]-hydrogenases catalyze the activation of molecular H2
through the reversible reaction, H2 2H+ + 2e-, and function to either couple H2
oxidation to energy yielding processes or reduce protons as a mechanism to recycle
reduced electron carriers that accumulate during fermentation (1).
A third type of
hydrogenase, [Fe]-hydrogenase or Hmd-hydrogenase also exists and catalyzes the
dehydrogenation of methylene-tetrahydromethanopterin to form H2 and methenyltetrahydromethanopterin (2).
Hydrogenases are widely distributed in diverse
microorganisms and ongoing microbial genome sequencing efforts continue to
demonstrate the ubiquitous occurrence and diversity of these enzymes (3-8). The [NiFe][FeFe]- and [Fe]-hydrogenases are phylogenetically distinct, do not share sequence
similarity, and therefore are classified according to the metals present at their active site
(4). [NiFe]-hydrogenases have been identified in archaea, bacteria, and cyanobacteria,
[FeFe]- in bacteria and eukarya, and [Fe]-hydrogenases in methanogenenic archaea (7, 8).
There are only a subset of organisms where [NiFe]- and [FeFe]-hydrogenases occur
together (such as sulfate reducing bacteria) and to date there appears to be a strict
segregation with respect to the occurrence of [NiFe]- and [FeFe]-hydrogenases in wateroxidizing phototrophs; only the [FeFe]-hydrogenases are found in eukaryotic green algae,
while only [NiFe]-hydrogenases are found in cyanobacteria (6).
2
Despite different ancestries, unique sequences, different metals at their active
sites, and different protein folds, the enzyme classes are unified in that their active sites
contain the biologically novel ligands CO and CN- coordinated to Fe (Figure 1.1). These
biologically unusual ligands, often associated with inhibition and poisoning, separate
hydrogenase active sites from presumably all other organometallic cofactors in nature
and are responsible for the unique electronic properties of the hydrogenase active site
necessary to efficiently catalyze the reversible oxidation of H2 (9, 10).
Figure 1.1. Active sites from left to right of [NiFe]-, [FeFe]-, and [Fe]-hydrogenases.
Extensive research efforts are currently aimed at using [NiFe]- and [FeFe]hydrogenases to generate H2 for use as a renewable energy carrier, and at using these
enzymes as a platinum substitute in fuel cells (11-21). Because [FeFe]-hydrogenases
typically display higher catalytic rates of hydrogen production than [NiFe]-hydrogenases
(1), in general more attention is given to these enzymes for developing biomimetic
hydrogen production catalysts, genetically engineered hydrogenases for biological
hydrogen production, and ultimately for developing hydrogen as a renewable fuel.
Advances in the basic understanding of hydrogenase function and assembly will help
3
develop the effective use of hydrogenases in biotechnological applications and may shed
light on how these evolutionary unrelated enzymes evolved to develop a unique, complex
active site, capable of catalyzing the reversible oxidation of hydrogen.
Much interest also surrounds the hydrogenases because of their close links to prebiotic chemistry (22-24).
Hydrogenases fit into a broader class of enzymes which
catalyze interconversion reactions necessary for life including the interconversion of H2,
CO, CO2, N2, and NH3 (25). These enzymes, which include carbon monoxide
dehydrogenase, acetyl coenzyme A synthase, and nitrogenase, in addition to hydrogenase
as discussed here, all exist as complex modifications to simple FeS clusters (eg. [2Fe-2S]
and [4Fe-4S] clusters) (Figure 1.2). A great deal of research is directed at how these
complex FeS biological enzymes associated with essential interconversion reactions
necessary for life may have developed and transitioned from basic iron-sulfur mineral
catalysts (26-32). For the case of hydrogenases, the presence of a unique active site
Fe(CO)x(RS-) core, common only among three diverse but yet phylogentically distinct
enzymes, makes this likely an example of functional convergent evolution in nature. The
relevance of hydrogenases to prebiotic chemistry as well as their practical applications to
renewable energy paves the pathway for diverse and exciting research in a rapid
developing field.
Although the primary focus of this thesis lies within [FeFe]-
hydrogenases, the [NiFe]-hydrogenases will also be discussed initially for comparison
purposes.
Hmd hydrogenases will not be further discussed due to fundamental
differences of the active site (mononuclear) and the reaction which they catalyze.
4
Figure 1.2 Complex active sites from carbon monoxide dehydrogenase (top left), acetyl
coenzyme A synthase (top right), nitrogenase (bottom left), and [FeFe]-hyrogenase (bottom
right).
Biological Role, Diversity, and Structure of Hydrogenases
The initial discovery of hydrogenases came in 1931 when they were found by
Stephenson and Stickland in colon bacteria (33). Since then, hydrogenases have been
found in taxonomically diverse microorganisms bacteria, archaea, and lower eukaryotes
including protists (4, 8, 34). Hydrogen metabolism is an essential component to living
microorganisms, and [NiFe]- and [FeFe]-hydrogenases facilitate this by catalyzing the
interconversion of hydrogen and protons according to the reaction H2 2H+ + 2e-.
[NiFe]- and [FeFe]-hydrogenases either function to oxidize molecular H2 to provide
5
reducing equivalents for metabolic processes (eg. methanogenesis, sulfate reduction,
acetogenesis, denitrification, nitrogen fixation, and photosynthesis) or to produce H2 via
proton reduction (1, 35). The latter is coupled to the oxidation of reduced electron carriers
generated by fermentation and/or photosynthesis and the proton is the terminal electronacceptor. Generally, [NiFe]-hydrogenases are involved in hydrogen uptake and [FeFe]hydrogenases with hydrogen production (1). Hydrogenases are vital in balancing cellular
proton gradients and redox potentials as well as cycling H2 for the transfer of chemical
energy in microbial communities (36, 37). Hydrogen-producing microorganisms and
hydrogen-utilizing mircroganisms are often interdependent, and hydrogen-utilizing
microorganisms are highly efficient at sequestering H2 to recycle the reducing potential
toward energy yield processes. Accordingly, H2 does not usually accumulate in natural
systems making it necessary to generate creative ideas to utilize the enzymes for biohydrogen applications.
Several approaches can be taken to utilize the catalytic activities of the [NiFe]and [FeFe]- hydrogenases for energy production applications (11-21). These include
fermentative approaches, in which reduced electron carriers generated during
fermentation are reoxidized by hydrogenase and photobiological approaches that use low
potential reductants generated by photosynthesis for H2 production. A number of in vitro
and bio-inspired approaches using hydrogenase enzymes or biomimetics are also being
explored for use in H2 production and oxidation applications. One drawback is that the
rates and/or yields of applied biological and bioinspired H2 production typically are not
high (3, 11, 15).
Additional examination of hydrogenase structure, function, and
6
diversity are necessary to make possible the effective and efficient utilization of these
enzymes in applied technology.
[NiFe]-Hydrogenase Functional Diversity
The [NiFe]-hydrogenases are dimers that consist of a large subunit harboring the
binuclear active site and a small subunit containing at least one [4Fe-4S] cluster (4, 5,
38). [NiFeSe]-hydrogenases have also been characterized, in which a selenocysteine
replaces cysteine in coordinating the active site Ni (39, 40). The phylogenetic analysis of
the [NiFe]-hydrogenases as well as biochemical studies have been used to group these
enzymes into several classes and subclasses (4, 5, 41). These include the (1) membrane
associated H2 uptake, (2a) cyanobacterial uptake, (2b) H2 sensing, (3a) F420 reducing, (3b)
bifunctional hyperthermophilic, (3c) MV-reducing, (3d) bidirectional NAD-linked, and
(4) membrane bound H2 evolving [NiFe]-hydrogenases (4). Accordingly, the respective
enzymes are often involved in a variety of metabolic functions and exhibit a broad range
of biochemical characteristics.
The respiratory uptake [NiFe]-hydrogenases (Group 1 or membrane-bound H2
uptake) typically oxidize H2 to supply reducing equivalents for cellular growth and
metabolism. The group 1 [NiFe]-hydrogenases are the most common of the classes of
[NiFe]-hydrogenases, and function to link H2 oxidation to the reduction of a variety of
electron acceptors, which include CO2, fumarate, O2, NO3, SO4 or metal ions. These
hydrogenases are often membrane associated and can couple electron transfer with transmembrane proton translocation (42, 43).
7
The group 2 [NiFe]-hydrogenases are typically located in the cytoplasm and
include the H2 sensing enzymes that regulate hydrogenase transcription in response to H2
(44, 45), as well as the cyanobacterial uptake [NiFe]-hydrogenases, which can recycle H2
produced by nitrogenase during N2 fixation in nitrogen-fixing organisms (19).
The group 3 [NiFe]-hydrogenases are generally multimeric, reversible enzymes
that contain additional subunits.
Figure 1.3. Cartoon representation of bidirectional NAD-linked [NiFe]-hydrogenase.
These subunits, which vary among the different group 3 enzymes, can interact with
soluble cellular redox components such as NAD, NADP or F420. For the bidirectional
NAD-linked [NiFe]-hydrogenases (group 3d), biochemical characterization shows that
they are heteropentameric, consisting of a large and small subunit (HoxYH) as well as a
diaphorase component (HoxEFU) which functions to couple NAD(H) oxidation or
reduction with hydrogenase activity (Figure 1.3) (18, 19).
The group 4 H2 evolving [NiFe]-hydrogenases are most often involved with H2
production in vivo in contrast to the more common hydrogen uptake functionality.
Members of this enzyme class include the hydrogenase 3 in E. coli (38), which couples
8
formate oxidation to proton reduction and has served as a prototypical example for
understanding [NiFe]-maturation (38, 46). Interestingly, class 4 enzymes typically lack
the C-terminal extension in the large subunit that is proteolytically cleaved in most NiFe
hydrogenases after insertion of Ni into the active site.
[NiFe]-Hydrogenase Structure and Function
The structure of [NiFe]-hydrogenases has been studied in detail from numerous
organisms and the x-ray crystal structures of [NiFe]-hydrogenases from sulfate reducing
bacteria Desulfovibrio (D.) gigas (47, 48), D. vulgaris (49, 50), D. fructosovorans (5153) and D. desulfuricans (54) have been determined.
Figure 1.4. Ribbon representation of the x-ray crystal structure of the [NiFe]-hydrogenase from
sulfate reducing bacteria D. gigas. The large subunit is colored blue and small is colored violet.
9
In addition, structures of [NiFe(Se)]-hydrogenases from Desulfomicrobium baculatum
(55) and D. vulgaris Hildenborough (56) have been determined. The preliminary x-ray
analysis for the photosynthetic bacterium Allochromatium vinosum has also been reported
(57). In their simplest and most characterized form, membrane-bound H2 uptake [NiFe]hydrogenases are heterodimeric and composed of a large (~60 kDa) and small subunit
(~30 kDa) (4, 5, 8) (Figure 1.4). For other classes of [NiFe]-hydrogenases, additional
subunits are often present. Hydrogen catalysis takes place at the [NiFe] active site
present in the large subunit. The small subunit contains at least one [4Fe-4S] cluster
proximal to the large subunit domain in addition to other accessory FeS clusters which
presumably function to assist electron transport to and from the catalytic site that is
buried well within the protein. For the membrane-bound H2 uptake [NiFe]-hydrogenases,
the small subunit contains 3 FeS clusters spaced ~12 Å apart, making them ideal to
mediate electron transfer from the active site. The clusters include a proximal [4Fe-4S]
cluster, medial [3Fe-4S] cluster, and distal [4Fe-4S] cluster in relation to the large
subunit. They play a critical role for hydrogen oxidation and respiratory uptake and their
special arrangement across the enzyme makes possible electron transport from the active
site to the protein surface where electrons can be delivered to different electron acceptors
(eg. cytochrome c3).
Active Site Structure. The heterobimetallic active site is comprised of a Ni atom
bridged by two cysteine thiolate ligands to an Fe atom (Figure 1.5). The Ni atom is
coordinated to the protein by two cysteine thiolate ligands. The Fe atom is coordinated
10
by one terminal CO and two terminal CN- ligands, which were originally detected using
IR-spectroscopy (48, 58, 59).
Figure 1.5. Ball and stick representation of the [NiFe]-hydrogenase active site determined by xray crystallography from D. gigas.
Also present in the as-isolated oxidized form is a species believed to be either a peroxide
molecule or a sulfenate from the oxidation of the bridging cysteine thiolate ligands (53,
60-62). [NiFe]-hydrogenases can be isolated aerobically and activated under reducing
conditions and the reactivation cycle involves the loss of the bridging oxygen species (50,
55).
Hydrogen activation is believed to take place at the Ni atom by nucleophilic
addition and heterolytic bond cleavage (63-65). Also, hydrophobic channels allowing for
H2 diffusion to the active site have been identified (51, 66).
Molecular dynamic
simulations of H2 diffusion from solvent to the active site of D. gigas show that H2
approaches the active site in every simulation from the Ni side (67).
During hydrogen catalysis, the [NiFe] active site passes through multiple redox
and structural states. These states have been termed Ni-A, Ni-B, Ni-C, Ni-L, and Ni-R
11
and have been the targets of many spectroscopic and structural studies (reviewed in (6871). The Ni atom passes through three different oxidation states (Ni1+, Ni2+, Ni3+) while
the Fe atom remains as Fe2+ throughout the cycle (72-76). The variable redox state of the
Ni atom provides support that it is the site for H2 activation and heterolytic bond
cleavage. In the as-isolated oxidized state, the active site can exist in two forms: Ni-A
(“unready”) and Ni-B (“ready”) (66, 70). While, the Ni-B state can be activated by the
reduction of H2 in seconds, activation of the Ni-A state is much slower and is on the time
scale of hours (61, 77). Activation of the two oxidized states involves the loss of the
bridging ligand between Ni and Fe atoms (50, 55) and it has been determined that this
ligand is different for the two states and thus can logically be attributed to the difference
in time required for activation. For the Ni-B state it is likely a single oxygen species (i.e.
hydroxide) and for the Ni-A state likely a multiple oxygen species (i.e. peroxide) (53, 6062). Other redox detected states of the [NiFe]-active site are created by the reduction of
the Ni-A and Ni-B states. One electron reduction and loss of the bridging species yields
the EPR active Ni-C state, which is light sensitive and can give way to the photoproduct
Ni-L at cryogenic temperatures. The fully reduced state is termed Ni-R. Also, similar to
[FeFe]-hydrogenases, the [NiFe] active site is reversibly inhibited by CO and binding has
been shown to take place at the Ni atom (78-81).
[FeFe]-Hydrogenase Functional Diversity
Genes encoding for [FeFe]-hydrogenase (HydA) have been identified in species
of bacteria and eukarya, with the latter primarily detected in lower order eukaryotes (4,
7). The primary structure of [FeFe]-hydrogenases is diverse, varying from proteins
12
comprising only the catalytic domain to those which contain up to six additional FeS
cluster binding domains. A common feature among [FeFe]-hydrogenases is that they
contain a highly conserved residue core, termed the H-cluster binding domain, which is
responsible for creating the unique chemical active site environment for the efficient
catalysis of reversible hydrogen oxidation. Residues associated with H-cluster ligation
include three distinct binding motifs termed L1 (TSCCPxW), L2 (MPCxxKxxE) and L3
(ExMACxxGCxxGGGxP) and are typically observed in the [FeFe]-hydrogenase primary
sequence (Figure 1.6) (4). Moreover, in the majority of [FeFe]-hydrogenases that have
been biochemically analyzed, a conserved GGV sequence is found between the L2 and
L3 motifs. The conserved H-cluster binding motifs allows for simple identification of
true [FeFe]-hydrogenases and distinguishes related sequences of HydA homologs.
Cp1
Cr1
DdH
Tma
L1
TSCCPGW
TSCCPGW
TSCCPGW
TSCCPAW
L2
MPCTSKKFE
MPCTRKQSE
MPCIAKKYE
MPCTAKKFE
L3
EVMACHGGCVNGGGQP
EIMACPAGCVGGGGQP
EYMACPGGCVCGGGQP
EVMACNYGCVGGGGQP
Figure 1.6. Amino acid alignments of the conserved H-cluster binding motifs for representative
[FeFe]-hydrogenases (CpI, CrI, DdH, and Tma). Cysteines ligating the H-cluster are highlighted
in green and additional conserved residues are highlighted in yellow.
For example, yeast and some eukaryotes have been found to contain HydA homologues
(Narf or Nar1). These homologues lack the cysteine residue from the L1 sequence motif
and typically have only a single [4Fe-4S] cluster N-terminal to the domain with H-cluster
similarity, whereas bacterial [FeFe]-hydrogenases typically contain eight cysteines in this
domain, and coordinate two [4Fe-4S] clusters. Although the role(s) of the eukaryotic
13
[FeFe]-hydrogenase homologs have yet to be firmly established, Nar1 in yeast from
Saccharomyces cerevisiae is proposed to be required for cytosolic and nuclear FeS
protein maturation and it is clear that they do not catalyze the production of molecular H2
(82, 83).
[FeFe]-Hydrogenase Structure and Function
Two [FeFe]-hydrogenases, Clostridium pasteurianum (CpI) and Desulfovibrio
desulfuricans (DdH) have been structurally characterized by x-ray crystallography (84,
85) and are the benchmark when comparing other biochemically characterized [FeFe]hydrogenases (Figure 1.7). CpI, a 60 kDa monomeric enzyme localized in the cytoplasm,
and DdH, a 53 kDa dimeric enzyme localized in the periplasm, are associated with H2
production and H2 uptake, respectively. The CpI enzyme is used to recycle reduced
ferredoxin, which oxidizes reduced pyruvate ferredoxin oxidoreductase and NADH
during fermentation. CpI and DdH differ in their complement of accessory Fe-S domains.
CpI has an H-cluster binding domain, a two [4Fe-4S] cluster binding domain which is
similar to bacterial ferredoxins, a domain that binds a unique [4Fe-4S] cluster that is
ligated by three cysteines and a histidine, and an N-terminal [2Fe-2S] plant-type
ferredoxin binding domain. In contrast, the DdH enzyme consists of an H-cluster domain
and a domain containing two [4Fe-4S] clusters. Often, characteristic motifs of cysteine
residues facilitate the identification of ferredoxin homologous domains in deduced amino
acid sequences and the x-ray crystal structures of various ferredoxins containing [4Fe-4S]
and [2Fe-2S] clusters have been reported (86-98).
14
Figure 1.7. Ribbon and space-filling representation of the x-ray crystal structures of [FeFe]hydrogenases from CpI (left) (84) and DdH (right) (85). For both structures, the H-cluster
catalytic domain is colored blue, the homologous C-terminus domain is red, and the two [4Fe-4S]
cluster domain is green. For CpI, additional [4Fe-4S] cluster and [2Fe-2S] cluster ferredoxin-like
domains are color violet and magenta, respectively.
Of other biochemically characterized [FeFe]-hydrogenases, differences in the
complement of FeS cluster accessory domains which link the catalytic domain to various
electron acceptors and donors are often observed (7). These variations can be attributed to
different physiological roles and locations of [FeFe]-hydrogenases. Enzymes consisting
of only the H-cluster domain and those having the H-cluster domain in addition to two,
three, and four additional FeS cluster binding domains have been identified.
The
majority of [FeFe]-hydrogenases identified in the bacterial genome contain three or more
additional FeS cluster binding domains. The FeS clusters in these accessory domains are
15
presumed to mediate electron transfer to and from the active site.
The [FeFe]-
hydrogenases characterized from eukaryotic green algae (e.g., Chlamydomonas
reinhardtii, Chlorella fusca, Scenedesmus obliquus) consist of just the H-domain (13, 99103) (Figure 1.8). These proteins, representing the most simple [FeFe]-hydrogenase
form, are of biochemical and biotechnological interest as they lack the additional FeSclusters observed in most native [FeFe]-hydrogenases which can complicate the direct
biophysical examination of the H-cluster Fe atoms.
Figure 1.8. Homology model of the eukaryotic green algae [FeFe]-hydrogenase C. reinhardtii.
Active Site Structure. The unique catalytic site of [FeFe]-hydrogenases, the Hcluster, consists of a [4Fe-4S] cluster coordinated to four cysteines and connected via a
single bridging cysteine thiolate to a unique binuclear 2Fe center which exists without
further coordination to the protein (Figure 1.9). Terminal CO and CN- ligands are bound
to each Fe atom of the 2Fe center, and a third CO ligand bridges both of the Fe atoms in
16
CpI and is terminal bound in DdH. A unique non-protein dithiolate ligand, proposed to be
either di(thiomethyl)amine (104), di(thiomethyl)ether (105) or dithiolpropane (85), also
bridges the 2Fe atoms of the active site.
Figure 1.9. Ball and stick representation of the [FeFe]-hydrogenases H-cluster determined by xray crystallography from CpI (84). The unknown atom of the dithiolate ligand is colored magenta
and a water molecule is present at the distal Fe atom of the 2Fe-subcluster in the oxidized state of
the H-cluster.
This unusual ligand set with π-acceptor ligands CO and CN- presumably functions to tune
the unique chemistry of the H-cluster, stabilizing low spin and low valance Fe that
facilitate H2 binding and reversible H2-oxidation properties normally found in second and
third row transition metals (9, 10).
In the oxidized state, the H-cluster displays a
characteristic EPR rhombic signal that is replaced by an axial signal after addition of CO
(106). The H-cluster has been termed by Holm and others into a class known as complex
bridged metal assemblies (22, 107, 108), which includes the nitrogenase FeMo-co and Pcluster and the active sites of acetyl CoA synthase and sulfite oxidase.
17
Ligand Exchangeable Site and H2 Catalysis. Similar to [NiFe]-hydrogenases,
[FeFe]-hydrogenases are reversibly inhibited by CO and x-ray structural analysis of the
CO inhibited state of CpI clearly demonstrates that CO binding occurs at the Fe of the
2Fe-subcluster distal from the [4Fe-4S]-subcluster (Figure 1.10) (104, 106, 109-112).
This observation along with the structural differences observed for the enzyme in various
oxidation states indicate that H2 binding and H2 production may occur at the distal Fe
atom of the 2Fe-subcluster (104, 106, 109-112). The CpI and DdH active site structures
represent different oxidation states for the H-cluster (Hox and Hred, respectively) and
differ in coordination of the distal Fe atom of the 2Fe-subcluster in respect to the [4Fe4S]-subcluster. In the CpI structure, which represents the oxidized state of the enzyme,
the Fe atom is in an octahedral coordination environment with two sulfur atoms of the
dithiolate ligand, two CO ligands (terminal and bridging), a terminal CN- ligand, and a
terminal bound water molecule (Figure 1.10) (84). In the DdH structure, representing the
reduced state of the enzyme, the Fe atom has an open coordination site from the absence
of the terminal bound water and is in a square pyramidal geometry (Figure 1.10) (85,
104). However, given that DdH crystals were grown in the presence of H2, it is likely
that a hydride or hydrogen is bound at the open coordination site of the distal Fe of the
2Fe-subcluster (109). Therefore, the distal Fe of the 2Fe-subcluster contains a ligand
exchangeable site. The bridging CO ligand, as observed by a shift in the IR band from
1802 cm-1 to 1894 cm-1 when going from oxidized CpI to reduced DdH, is terminally
bound in DdH (104).
18
Figure 1.10. Ball and stick representation of the H-cluster in the Hox, Hred, and HoxCO states. For
each state, a summary of the Mössbauer, EPR, and IR spectroscopic characteristics is given.
Adapted from (113).
These insights can begin to provide mechanistic clues as to how reversible H2
oxidation takes place at the 2Fe-subcluster and a scheme can be proposed in which this
ligand exchangeable site is occupied by a water molecule in the oxidized state but when
the metal cluster is reduced or hydrogen is available as a coordinating group, water is
displaced by H2 and the bridging CO ligand shifts from bridging terminal. In this
hypothesis, heterolytic H2 bond cleavage would take place at the terminal position of the
distal Fe site (104, 114-116). Also, hypotheses involving bridging hydrides have been
put forth and involve too migration of the bridging CO ligand to terminal coordination
and water displacement (117-119). As for the [4Fe-4S] cluster, it may serve as an
electron reservoir to mediate electron transfer to the 2Fe-subcluster during H2 catalysis
(117). Interestingly, density functional theory calculations on the entire H-cluster show
19
delocalization between the molecular orbitals of the [4Fe-4S]-subcluster and 2Fesubcluster, supporting that the H-cluster is an electronically linked 6Fe cluster (120, 121).
Looking at the oxidation states of the Fe atoms of the H-cluster, for both the
oxidized and reduced states, the [4Fe-4S] cluster remains in the EPR silent [4Fe-4S]2+
form [15, 145].
For the 2Fe-subcluster there is some discrepancy. Mössbauer studies
suggested that the oxidations states were Fe3+Fe2+ (Hox), Fe2+Fe2+ (Hred) (122, 123),
however, it was noted that Fe2+Fe1+ (Hox), Fe1+Fe1+ (Hred) would also fit the data. It is
becoming clearer from synthetic models (124) and density functional studies (116) that
the 2Fe-subcluster exists in the later, lower oxidation states. Given the nature of the CO
and CN- ligands, this does not come as a surprise.
Dithiolate ligand. A key remaining question in regards to active site H-cluster of
[FeFe]-hydrogenases is the nature of the non-protein dithiolate ligand, specifically if its
identity is di(thiomethyl)amine, di(thiomethyl)ether, or dithiolpropane . The composition
of this non-protein dithiolate ligand has been of significant interest since the central or
sometimes termed bridgehead group of the ligand is located in close proximity to the
distal site. It has been suggested that if the bridgehead group were an amine, it could
cycle between different protonation states and serve as a proton donor/acceptor group
during catalysis and H2 heterolytic bond cleavage (Figure 1.11) (104, 114, 125).
20
Figure 1.11. Potential mechanism for H2 heterolytic bond cleavage if the bridging dithiolate
ligand is di(thiomethyl)amine.
The chemical composition of this group to date has not been determined but recently a
spectroscopic study supports that an amine group is located at this position (126).
Computational work, however, questions whether an amine group at this position could
act as a proton acceptor group in catalysis and supports the presence of an ether group
(105). Also, another recent computational study using the same crystal structure as the
previous report contrarily supports N as the bridgehead atom and di(thiomethyl)amine as
the dithiolate ligand (127). Given these differences, additional experimental studies are
needed to shed light on the nature of the ligand and not any of three possible identities
should be ruled out at this time.
The H-cluster active site is the target of many inorganic and organometallic
synthetic chemists and many numerous cluster mimics have been synthesized
incorporating the unique CO and CN- ligands in addition to potential dithiolate ligands
(128, 129). Besides desired applications to synthesizing model complexes mimicking the
[FeFe]-hydrogenase H-cluster which are capable of producing hydrogen, these studies
can be alternative approaches toward probing unknown structural and binding
21
characteristics of the H-cluster including the composition the dithiolate ligand and
mechanisms for H2 catalysis.
Oxygen Sensitivity and Model Complexes. [FeFe]- and [NiFe]-hydrogenases
differ in their sensitivity to oxygen and generally [NiFe]-hydrogenases display higher
levels of O2 tolerance than [FeFe]-hydrogenases (1, 130-134).
Unlike [FeFe]-
hydrogenases which are irreversibly inactivated by O2, [NiFe]-hydrogenases can be
reversibly activated under O2, which involves loss of an oxygen species. Three [NiFe]hydrogenases from Ralstonia eutropha have even been shown to catalyze hydrogenase
oxidation in the presence of O2, although at significantly slower rates (131, 135, 136).
For both [NiFe]- and [FeFe]-hydrogenases, O2 diffusion pathways have been identified
by analysis of the crystal structures and by molecular dynamics calculations giving way
to experiments designed to engineer hydrogenases that are O2 tolerant and/or have a
decreased level of oxygen sensitivity (51, 109, 130, 134). Experiments aimed at mutating
amino acid residues along identified hydrophobic channels to decrease O2 diffusion, O2
access to the active site, and ultimately lower the O2 sensitivity have been conducted and
this approach has been demonstrated to be successful for a [NiFe]-hydrogenase (137,
138). Despite [FeFe]-hydrogenases high sensitivity to oxygen, they still remain top
targets for biotechnological studies since they display one-to-two orders of magnitude
higher catalytic activities than the [NiFe]-hydrogenases (1).
22
Hydrogenase Maturation
Taking into consideration the unique properties of the active site of hydrogenases
including the presence of CO and CN- ligands not common in biology and normally
associated with inhibition and poisoning, the process for how these unprecedented active
sites are constructed is intriguing. For [NiFe]-hydrogenase, the biosynthesis of the active
site also has to involve incorporation of the toxic metal Ni which adds another step into
the already complicated process. Unlike [NiFe]-hydrogenases, very little was known
concerning the maturation of [FeFe]-hydrogenases and the assembly of the complex Hcluster site until very recently. In fact, until 2004 little to nothing was known about the
maturation of [FeFe]-hydrogenase and H-cluster biosynthesis. In contrast, the maturation
of [NiFe]-hydrogenases have been studied longer and at least six gene products have
been identified to be required for the biosynthesis of its active site (139). Interestingly,
for [FeFe]-hydrogenases, it was determined recently that only three gene products are
required for the biosynthesis of the H-cluster (140).
Other Maturation Systems
Like the hydrogenases, nitrogenases have complex FeS cofactors that exist as
modifications to basic FeS clusters.
Studies on nitrogenases maturation have been
ongoing for approximately thirty years and many insights have been made as to how the
complex cofactors, the P-cluster, an [8Fe-7S] cluster coordinated by six Cys ligands, and
FeMo-cofactor, a [Mo-7Fe-9S] cluster with a Cys and His ligand in addition to a non-
23
protein ligand homocitrate, are assembled (Figure 1.12) (141, 142). The latter is
responsible for biological nitrogen fixation.
Figure 1.12. Ball and stick representation of the P-cluster (left) and FeMo-co site (right) of
nitrogenase. The unknown interstitial atom of the FeMo-co is colored magenta.
FeMo-co biosynthesis is a complicated process requiring multiple enzymes, scaffolds,
and carriers (fifteen gene products are linked to this process) and in many respects FeMoco biosynthesis can serve as a standard paradigm for complex FeS cluster biosynthesis
(Figure 1.13).
It is an ideal model system for experimental design in probing
hydrogenase maturation and specially the many unstudied details of [FeFe]-hydrogenase
maturation and H-cluster biosynthesis.
In addition to the nitrogenase system (NIF), the ISC assembly and SUF systems
have been studied intensely for the assembly of bacterial Fe-S proteins and general
principles from all of these systems together is a particular valuable resource for
understanding FeS cluster biogenesis in the context of hydrogenases (143-146). The ISC
and SUF systems, often referred to as general house-keeping FeS cluster maturation
machinery, function to biosynthesize basic FeS clusters (eg. [2Fe-2S] and [4Fe-4S]
cluster) under normal and oxidative-stress conditions, respectively. Common to the NIF,
24
ISC and SUF systems for the overall assembly of FeS proteins is a two part overarching
process consisting of the assembly or construction of the FeS cluster on a scaffold protein
which is followed by subsequent transfer to a target apoprotein (147).
Figure 1.13. Flow scheme depicting overall process for nitrogenase maturation and FeMo-co
biosynthesis (from reference (142)).
This theme is very clear among all three of these systems and can be useful for
probing unknowns of other biosynthetic pathways such as [FeFe]-hydrogenases
maturation. Moreover, steps in this process all have common factors including the
presence of a cysteine desulfurase for release of sulfur from cysteine and reduction to
sulfide for FeS cluster formation, iron incorporation from free iron (Fe2+) which often
requires specific iron donors, the presence of scaffold proteins, and the presence of FeS
cluster transfer proteins.
25
[NiFe]-Hydrogenase Maturation
The maturation of the active site of [NiFe]-hydrogenases has been studied in
detail using a number of different organisms, and a model for the individual steps in the
process has been developed for hydrogenase-3 in E. coli (reviewed in (139)). At least six
maturation enzymes are required for the synthesis of the active site of [NiFe]hydrogenases (139). This is not too surprising given the characteristics of nickel and its
toxicity, and often additional proteins are involved in the biosynthesis process for
activation, transport, insertion and binding of nickel. Taking into account the diversity of
[NiFe]-hydrogenases, it is not unexpected that unique maturases are required for
organisms in certain cases (139, 148, 149).
Below, the function of six essential maturation enzymes HypA, B, C, D, E, and F
(encoded by hyp genes) in the maturation of hydrogenase-3 in E. coli is detailed. The
overall process of maturation requires the synthesis and insertion of the Fe atom bound to
non-protein ligands CO and CN- and the activation, transport and insertion of Ni for
incorporation into the active site. This process takes place stepwise with the synthesis
and insertion of the Fe(CN)2CO moiety occuring before the incorporation of Ni (139,
150, 151) (Figure 1.14). In the first step, HypE and HypF form a complex to allow
synthesis of thiocyanate from carbamoyl phosphate, the biological precursor for the CNligands (152-155). By hydrolysis of carbamoyl phosphate on HypF and subsequent
transfer of the carbomyl group to HypE followed by hydrolysis, a thiocyanate is attached
to the sulfur of the C-terminal residue on HypE. HypC and HypD also complex together
and HypE transfers the thiocyanate to the complex where it is able to bind Fe, although
26
the exact mechanism is not known (156-158). At this point CO must also be incorporated
into the precursor, however, unlike CN-, the substrate for CO and details for this step are
not yet clear (159-161).
Figure 1.14. Biosynthetic pathway of the active site of [NiFe]-hydrogenases based on
hydrogenase-3 in E. coli. Adapted from (166).
27
Ultimately, the HypCD complex transfers the Fe(CN)2CO moiety into the apohydrogenase large subunit before Ni insertion. HypA and HypB are involved in the
insertion of Ni (162, 163) and recent the crystal and solution structures of HypA have
been reported (164, 165). Although the exact mechanism for Ni insertion is not known,
HypA and HypB interact with each other to form a complex (163). HypB is a GTPase
and the insertion of Ni involves GTP hydrolysis by HypB (167). HypA may act as a
metallochaperone between HypB and the large hydrogenase subunit in the insertion
process (168, 169). Following complete maturation of the active site is proteolysis of an
extended C-terminus present in some [NiFe]-hydrogenases by an endopeptidase (170173).
[FeFe]-Hydrogenase Maturation
The proteins required for [FeFe]-hydrogenase maturation were initially
discovered in the eukaryotic green alga C. reinhardtii, when it was shown that two novel
radical S-adenosylmethionine (radical SAM or AdoMet) proteins, HydEF and HydG, are
necessary for C. reinhardtii [FeFe]-hydrogenase enzyme activity (140).
In the C.
reinhardtii mutant hydEF-1, the hydEF gene is disrupted. This was discovered by
screening random C. reinhardtii mutants unable to produce H2. Further studies
demonstrated that the hydEF-1 mutant is unable to assemble an active [FeFe]hydrogenase and that complementation of a wild-type copy of the hydEF gene to the
hydEF-1 mutant restores hydrogenase activity. A second gene required for [FeFe]hydrogenase assembly, hydG, is directly adjacent to the hydEF gene in the C. reinhardtii
genome. The direct involvement of three gene products in hydrogenase maturation was
28
confirmed by their requirement for the heterologous expression of active hydrogenase in
E. coli (140, 174, 175). The three genes, hydE, hydF, and hydG, along with the [FeFe]hydrogenase structural encoding gene hydA, appear to be the only hydrogenase-specific
genes common to all organisms possessing [FeFe]-hydrogenases, suggesting that these
are the only genes required for synthesis and insertion of the H-cluster (Figure 1.15).
Figure 1.15. Gene clusters for representative [FeFe]-hydrogenase showing the presence of
[FeFe]-hydrogenase maturation genes products HydE, HydF and HydG along with the [FeFe]hydrogenase structure encoding gene product HydA.
In some organisms, additional genes cluster with the hydE, hydF and hydG
[FeFe]-hydrogenase maturation genes such as ammonium aspartate lyase. These
additional genes are not observed in all organisms containing [FeFe]-hydrogenases and it
is currently unknown whether these gene products participate in [FeFe]-hydogenase
maturation or are perhaps associated with some other aspect of H2 metabolism.
29
Initial characterization of all three [FeFe]-hydrogenase maturation proteins has
been carried out and HydE, HydF, and HydG from the extremophile Thermatoga
maritima have been heterologously expressed, purified, and anaerobically reconstituted
(176, 177). Both the T. maritima HydE and HydG proteins contain two distinct FeS
clusters, and both have the expected SAM activity, cleaving SAM into 5’deoxyadenosine (AdoH) and methionine in the presence of DTT after reduction with
sodium dithionite (177).
Initial characterization of HydF demonstrates that is has
GTPase activity and coordinate an FeS cluster upon reconstitution (176).
Comparing to the maturation of [NiFe]-hydrogenases, there is no cross-over
between the maturation enzymes required for [NiFe]- and [FeFe]-hydrogenase
maturation. Each class of hydrogenase requires unique maturation enzymes (4, 8). In
contrast to only three maturation enzymes required for [FeFe]-hydrogenases, at least six
maturation enzymes are required for the synthesis of the active site of [NiFe]hydrogenases (139). This is not unexpected given the characteristics of nickel and its
toxicity and often additional proteins are involved in the biosynthesis process for
activation, transport, insertion and binding of nickel. As for the nitrogenase system,
fifteen proteins have been linked to the maturation process (142).
Radical SAM Proteins. Of the three strictly conserved proteins identified for
[FeFe]-hydrogenase maturation, HydE and HydG belong to the radical SAM superfamily
of proteins. HydF on the other hand contains a putative GTPase domain. HydE and
HydG both have the C-X3-C-X2-C radical SAM signature motif. Additionally, HydG
proteins have a second conserved cysteine motif, C-X2-C-X22-C, in the C-terminal
30
portion of the protein. HydF contains an NTPase domain comprised of Walker A P-loop
and Walker B Mg2+ binding motifs, as well as a putative iron-sulfur cluster binding motif,
C-X-H-X(44-53)-HC-X2-C. The three cysteines of the radical SAM motifs in both HydE
and HydG have been mutated to serine in C. acetobutylicum HydE and were shown to be
critical for [FeFe]-hydrogenase assembly in vivo, using an E. coli heterologous
expression system (175). Mutation of the conserved cysteines in the C-terminal motif of
HydG, as well as amino acids in the metal binding motif and NTPase domain of HydF,
were also shown to be critical for [FeFe]-hydrogenase maturation in this system (175).
Radical-SAM proteins were recognized as a protein superfamily when advanced
sequence profiling methods demonstrated that several hundred proteins involved in
diverse cellular processes share significant sequence similarity, primarily the C-X3-C-X2C motif (178-184). In these prtoeins, a [4Fe-4S] cluster is coordinated by the three
cysteines of the radical-SAM motif and the methionine carboxylate and amine of SAM
bind the [4Fe-4S] cluster at the open iron coordination site (184-189) (Figure 1.16).
Figure 1.16. Unique [4Fe-4S] cluster in radical SAM proteins which coordinates methionine
carboxylate and amine of SAM.
31
Radical SAM enzymes catalyze the reduction of SAM by the reaction
to generate methionine and the 5’-deoxyadenosyl (DOA) radical. The highly reactive
DOA radical is subsequently utilized to catalyze difficult biochemical transformations or
reactions with high activation barriers that would not proceed without radical chemistry.
The structures of several radical-SAM enzymes including biotin synthase (BioB) (190),
coproporphyrinogen III oxidase (HemN) (191), molybdopterin (MoaA) (192), pyruvate
formate-lyase-activating enzyme (PFL-AE) (193), and lysine 2,3-amino mutase (LAM)
(194) have been determined. Although the structures of the characterized Radical-SAM
proteins are similar, each protein contains a unique N- and/or C-terminal region(s) that is
proposed to modulate substrate access to the active site.
The precise mechanism of [FeFe]-hydrogenase assembly is currently unknown,
however, early hypotheses could be made from parallels to the radical-SAM superfamily
of proteins. First, three different Radical-SAM enzymes, LipA (lipoate synthase), BioB,
and MiaB are known to incorporate sulfur into substrates (195, 196). Interestingly, HydE
most closely aligns with these proteins suggesting that it may function to synthesize the
dithiolate ligand of the H-cluster (Figure 1.17). Second, iron and sulfur originating from
32
the NifB cofactor, where NifB is a radical SAM protein, ultimately become incorporated
into nitrogenase (197, 198).
Figure 1.17. Portion of sequence alignment including C-X3-C-X2-C motif of radical SAM
proteins HydE and HydG relative to known functioned radical-SAM BioB, LipA, PFL-AE, LAM
and ThiH (top). Also shown are the reactions catalyzed by the respected proteins, sulfur insertion
reactions and reactions resulting formation of amino acid radicals. Adapted from (113).
33
The involvement of this radical SAM protein in the assembly of an iron metalloenzyme
may have parallels to the biosynthesis of the [FeFe]-hydrogenase active site. Third, the
reactions catalyzed by PFL-AE (199) , LAM (180), and the thiH gene product (200, 201)
all occur via the formation of amino acid radicals (Figure 1.17). This makes it attractive
to think that an amino acid substrate and an amino acid radical intermediate could serve
as a source of the CO and CN- ligands in the H-cluster. Interestingly, HydG shows
significant homology to ThiH, which catalyzes tyrosine cleavage to dehydroglycine and
ultimately thiamine biosynthesis (202).
Lastly, members of the Radical-SAM
superfamily make possible a number of difficult synthetic reactions, including several
anaerobic oxidations. In addition to the bridging dithiolate ligand, the 2Fe catalytic
center contains CO and CN- ligands. As mentioned above, no homologs of the [NiFe]hydrogenase assembly proteins are found in the genomes of organisms containing only an
[FeFe]-hydrogenase (4, 8, 139); therefore, unique pathways for the synthesis of these
ligands must exist.
Radical SAM Based Hypothesis. An initial hypothesis regarding the roles of
HydE, HydF, and HydG, in the biosynthesis of the H-cluster of hydrogenase was made
by Peters et al. and based primarily on similarities to known radical SAM chemistry (84)
(Figure 1.18). In the first biosynthetic step of this hypothesis, HydE or HydG convert a
standard [2Fe-2S] cluster (bound to HydE or HydG or to the scaffold protein HydF) to a
dithiolate-bridged [2Fe-2S] cluster. The dithiolate ligand is generated by alkylation of the
two bridging sulfides of the cluster. Radical SAM enzymes LipA (lipoate synthase) and
34
BioB (biotin synthase) both catalyze this type of reaction involving the insertion of a
bridging cluster sulfide into an alkane C-H bond to generate a thiolate (203, 204).
Figure 1.18. Potential mechanism for [FeFe]-hydrogenase maturation which is based on radical
SAM chemistry. Adapted from (84).
35
In the case of both BioB and LipA, the sulfur has been proposed to originate from a
second conserved FeS cluster unique from the SAM binding [4Fe-4S] cluster (196, 205208).
For HydE/HydG, sulfur insertion would occur twice to yield a dithiolate
coordinated to the [2Fe-2S] cluster. Alkylation of the sulfides protects the sulfur atoms
from further modification and shifts reactivity to the Fe atoms in the cluster.
The
modified, dithiolate-bridged 2Fe cluster would then be transferred from the first radicalSAM enzyme (HydE or HydG) to the second (HydE or HydG), or would remain bound to
the scaffold HydF, for the next step in cluster assembly, the generation of the carbonyl
and cyanide ligands. The second radical-SAM enzyme, either HydE or HydG, is then
proposed to generate glycyl radicals that react with the Fe atoms of the dithiolate bridged
[2Fe-2S] cluster. Two successive cycles of glycyl radical decomposition at these Fe
atoms would generate CO and CN- at an equivalent stoichiometry at each iron atom. The
glycyl radical decomposition to CO and CN- is supported in the hypothesis by DFT
calculations, which demonstrate that the high-energy requirement for coordination of
glycine to a reduced iron would be overcome through the generation of the radical
intermediate. The scaffold protein HydF would serve as the final site of H-cluster
precursor assembly, and the site from which this precursor is transferred to the
hydrogenase structural protein (HydA) to effect activation. Incorporation of the [4Fe4S]-subcluster is presumed to occur by means of generalized host cell FeS cluster
machinery (143) and inserted prior to the 2Fe-center.
In Vitro Activation.
Experimental steps toward deciphering the stepwise
assembly of the H-cluster and [FeFe]-hydrogenase maturation were established by the
36
development of heterologous expression system in E. coli and subsequently an in vitro
system for HydA maturation. In this system, [FeFe]-hydrogenase maturation could be
achieved in less than five minutes by incubating E. coli extracts expressing HydE, HydF,
and HydG from C. acetobutilycum with extracts expressing only HydA from CpI (209).
The extremely rapid rate of [FeFe]-hydrogenase activation was indicative of the presence
of a H-cluster precursor that could be readily transferred to HydA and was able to readily
activate the structural enzyme. Activation proceeded without addition of small molecule
reagents, suggesting that the H-cluster precursor was already assembled in the
HydE/HydF/HydG extract and was readily transferred to the HydA apoenzyme to
generate active [FeFe]-hydrogenase.
It was also demonstrated that the activation
component was protein-associated and not a freely diffusing small molecule.
These
observations suggest a model for H-cluster biosynthesis in which one or more of the
accessory proteins, and not the hydrogenase structural protein, acts as the physical
scaffold for the assembly of an H-cluster precursor.
HydF as a Scaffold Protein. Demonstration that HydF acts as a scaffold during
H-cluster assembly was achieved by purification of HydF heterologously expressed in E.
coli cell extracts, in a genetic background of HydE and HydG (HydFEG). The purified
HydFEG could activate HydA expressed heterologously in E. coli lacking HydE, HydF,
and HydG (HydA∆EFG) (210). The identification of HydF as a scaffold is significant and
strongly suggests that radical SAM proteins HydE and HydG enact chemistry upon HydF
to produce an H-cluster intermediate, which is then transferred to HydA to accomplish
activation.
Also, concerning the GTPase activity of HydF, resent results by Shepard
37
indicate that this GTPase activity is not associated with transfer of the subcluster from
HydF to HydA (211). Therefore, an alternative hypothesis is that GTP hydrolysis may be
associated with the interactions between HydF and HydE and/or HydG during synthesis
of H-cluster 2Fe-subcluster precursors (210, 211).
The spectroscopic characterization of HydF provides supportive evidence that the
scaffold enzyme contains an H-cluster 2Fe-subcluster precursor (211). When HydF is
heterologously expressed in E. coli in the absence of HydE and HydG, it exhibits
characteristic EPR spectra consistent with having both [4Fe-4S] clusters and [2Fe-2S]
clusters. However, when HydF is expressed in a genetic background in which HydE and
HydG are co-expressed (HydFEG), the [2Fe-2S] cluster is not observed by EPR. Fourier
transform infrared (FTIR) spectroscopy on HydFEG shows sharp vibrational bands at
2046, 2027, 1940, and 1881 cm-1, which can be assigned to CO (1940 and 1881 cm-1)
and CN- (2046, 2027 cm-1) ligands. These vibrational bands have not been observed for
HydF∆EG samples. These results provide clear evidence for the presence of CO and CNligands on HydF and support a model in which a basic [2Fe-2S] cluster on HydF is
converted to an H-cluster precursor 2Fe-subcluster which contains CO and CN- ligands
(211).
These results also coincide with the recently reported spectroscopic
characterization of HydF from Clostridium acetobutylicum which suggests the presence
of a 2Fe cluster with CO and CN- ligands on HydF (212).
Substrate for CO and CN- Ligands. A major breakthrough for deciphering the
maturation process was made in the discovery that HydG catalyzes the degradation of
tyrosine to produce cresol in a similar way to that of the homologous radical SAM
38
enzyme ThiH, which cleaves tyrosine to cresol and dehydroglycine in thiamine
biosynthesis (see Figure 1.16 above) (202). Subsequently, it was also noted that addition
of tyrosine, cysteine, and SAM could enhance [FeFe]-hydrogenase maturation (213).
Although it was speculated that the formation of dehydroglycine from trysosine could
serve as a precursor for the synthesis of the dithiolate ligand (202), HydG catalyzed
tyrosine cleavage resulting in the production of CN- in equivalent stoichiometry to pcresol was soon after observed (214).
This result was quickly followed by the
observation that HydG also catalyzes tyrosine cleavage to produce CO (215). Both of
these reactions utilized chemically reconstituted HydG along with tyrosine, AdoMet, and
sodium dithionite. CN- was detected using a derivatization method utilizing fluorescence
and HPLC analysis (214) while CO was detected using deoxyhemoglobin assays taking
advantage of shifts in the UV-VIS soret band of deoxyhemoglobin upon binding of CO
(215).
These results together clearly indicate that the role of HydG in H-cluster
biosynthesis is to synthesize the CO and CN- ligands from the substrate tyrosine by the
following reaction:
Significantly, formation of CO and CN- through the amino acid radical on tyrosine
represents a new chemical reaction catalyzed by a radical SAM enzyme (HydG).
39
Because p-cresol is formed in 1:1 stoichiometry to CN- (214), it can be hypothesized that
the reaction takes place through the intermediate dehydroglycine:
This hypothesis is warranted since another reaction product of the homologous radical
SAM enzyme ThiH, which has the same substrates as HydG and three identical products
(methionine, DOA, and p-cresol), is dehydroglycine.
The hydrolysis product of
dehydroglycine, glyoxylate, has been detected in small amounts in the tyrosine cleavage
reaction catalyzed by HydG, supporting the notion that dehydroglycine is an intermediate
in the HydG-catalyzed tyrosine cleavage reaction resulting in diatomic ligands CO and
CN-. Decarbonlyation of dehydroglyince could be a potential mechanism for formation
of CO and CN- products in a single step:
NHCHCO2H → HCN + CO + H2O
This reaction mechanism has been reported previously in literature (216).
Substrate for Dithiolate Ligand. The discovery that HydG serves to synthesize
CO and CN- ligands from tyrosine suggests that HydE may function to synthesize the
dithiolate ligand of the H-cluster and studies are underway to identify possible substrates.
40
The crystal structure of the recombinant, reconstituted form of HydE from Thermotoga
maritima has been solved but its function still remains elusive (Figure 1.19) (217).
Figure 1.19. Ribbon representation of the x-ray crystal structure of recombinant, reconstituted
HydE from T. maritima colored according to secondary structure. The [4Fe-4S] cluster and [2Fe2S] cluster are represented as space-filling models and SAM is depicted in stick representation.
The structure, similar to that of biotin synthase, shows the presence of a [4Fe-4S]
cluster coordinated to SAM and also a [2Fe-2S] cluster which is most likely the result of
chemical reconstitution (217). The role of HydE and substrate for the dithiolate ligand
are among several of the many questions remaining to be determined for the maturation
process. Determination of the substrate for the dithiolate ligand should also shed light as
to the actual composition of the ligand and consequently give significant progress toward
understanding how H2 catalysis at the H-cluster takes place.
41
Current Hypothesis. Based on the above observations a refined hypothetical
scheme for H-cluster biosynthesis can be put together (Figure 1.20).
Figure 1.20. Refined hypothetical scheme for the roles of HydE, HydF, and HydG in [FeFe]hydrogenase maturation and H-cluster biosynthesis. Adapted from (211).
In this hypothetical scheme backed by much experimental evidence, HydF serves as a
chemical scaffold on which a basic [2Fe-2S] cluster is transformed to an H-cluster 2Fesubcluster precursor with unique non-protein ligands. HydE synthesizes the dithiolate
ligand from an unknown substrate, HydG synthesizes the diatomic CO and CN- ligands
from substrate tyrosine. The [4Fe-4S] cluster is synthesized by general FeS cluster
biosynthetic machinery on the structural protein (HydA), and in the final step of
maturation the 2Fe-subcluster complete with ligands is transferred to HydA which
already contains the [4Fe-4S] cluster, thus completing H-cluster biosynthesis and [FeFe]hydrogenase maturation. This scheme provides the groundwork for the next generation
of experiments probing the mechanism of H-cluster biosynthesis.
42
Research Directions
Similar to other maturation systems (eg NIF, ISC, and SUF), the hypothetical
scheme for [FeFe]-hydrogenase maturation (see above) can be interpreted as an
overarching two part progression consisting of the synthesis of a complex FeS cluster on
a scaffold protein followed by subsequent transfer to a target apo-protein which results in
a protein that is catalytically active (218). For [FeFe]-hydrogenases, the complex FeS
cluster is the 2Fe-subcluster of H-cluster which contains unique non-protein ligands CO
and CN-, the scaffold protein is HydF, the apo-protein in the hypothetical scheme is the
[FeFe]-hydrogenase structural protein (HydA), and the matured active protein catalyzes
reversible oxidation of molecular H2. A major premise of the hypothesis is that the [4Fe4S]-subcluster and 2Fe-subcluster of the H-cluster are synthesized independently. Many
of the insights made regarding the biosynthesis of the 2Fe-subcluster outline above were
a direct result of experiments probing the function of maturation enzymes HydE, HydF,
and HydG, however others were a result of an alternative, complementary approach
probing [FeFe]-hydrogenase maturation and the synthesis of its complex active site, the
H-cluster, by characterization of the structural protein, HydA. This approach is presented
herein and was taken simultaneously in parallel to the numerous experiments focusing on
the maturation enzymes.
In the early hypothetical scheme concerning the maturation of [FeFe]hydrogenase put forth by Peters et al., it was hypothesized that H-cluster biosynthesis
occurs on a site of an accessory protein scaffold and not at the actual structural [FeFe]hydrogenase (HydA) (218).
Furthermore, it was hypothesized that the [4Fe-4S]-
43
subcluster and 2Fe-subcluster are synthesized independently, with a preformed [4Fe-4S]
cluster already present at the active site of HydA prior to 2Fe-subcluster synthesis and
insertion by the maturation enzymes HydE, HydF, and HydG.
Theoretically, it is
possible that the maturation enzymes HydE, HydF, and HydG could serve to synthesize
the entire H-cluster itself or its individual components such as the [4Fe-4S]-subcluster,
2Fe-subcluster, and/or unique ligands. Connection to other maturation systems that have
been studied, however, can be used to rule out several of these options and warrants the
reasoning of the original hypothesis by Peters et al. For example, it is known that general
house-keeping FeS cluster maturation machinery can synthesize and insert basic [4Fe-4S]
clusters (143). Also, in analogy to nitrogenase maturation, the active site FeMo-cofactor
is synthesized on a protein scaffold and transferred the apo-nitrogenase protein in the
final step of maturation to generate active enzyme (142). These examples serve as a
sound foundation for the hypothesis that [4Fe-4S]-subcluster and 2Fe-subcluster
synthesis occurs separately, but yet experimental evidence is needed to support the claim.
Heterologous expression of [FeFe]-hydrogenase in E. coli (175) along with the in-vitro
activation of [FeFe]-hydrogenases (209) presents an ideal system to probe the nature of
the [FeFe]-hydrogenase structural protein which can be expressed in a background
lacking the maturation enzymes (HydA∆EFG).
This system was utilized and the
biochemical and spectroscopic characterization of HydA∆EFG from the green algae
Chlamydomonas reinhardtii, along with its relevance to [FeFe]-hydrogenase maturation,
is presented in Chapter 2. The results of this chapter provide initial, key evidence that
[FeFe]-hydrogenase maturation enzymes are directed at the synthesis of the 2Fe-
44
subcluster and that the [4Fe-4S]-subcluster is synthesized independently by general FeS
housekeeping maturation machinery.
Another intriguing aspect not only to [FeFe]-hydrogenase maturation but also to
other biological systems is how complex FeS clusters are inserted into apo-enzymes and
how complex cofactors are assembled in nature. For nitrogenase, the structure of the
cofactor deficient form of the structural enzyme revealed significant structural
rearrangement resulting in a positively charged funnel to the active site (219). Not only
did this discovery make known the mechanistic pathway for FeMo-co insertion into the
cofactor-less enzyme but is also provided clues as to how FeS cluster insertion in general
may take place. Maintaining the maturation of nitrogenase as a principle analogy to
serve as a foundation for probing [FeFe]-hydrogenase, it is intriguing to hypothesize if
these features also parallel the cluster insertion process for [FeFe]-hydrogenase
maturation. Chapter 3 presented herein probes by x-ray crystallography the structure of
HydA∆EFG Chlamydomonas reinhardtii and subsequently the 2Fe-subcluster insertion
process for [FeFe]-hydrogenase and the stepwise biosynthesis of the H-cluster. The
results of this chapter are of broad interest and provide exciting new insight into the
mechanism for complex metallocluster assembly, establish unifying themes in complex
metal cluster assembly, and reveal fundamental features of metalloenzyme evolution.
45
CHAPTER 2
ACTIVATION OF HYDA∆EFG REQUIRES A PREFORMED [4Fe-4S] CLUSTER
Introduction
The [NiFe]- and [FeFe]- hydrogenases are widely distributed in nature and
efficiently catalyze the reversible oxidation of molecular hydrogen (H2 ↔ 2 H++ 2 e-).
The [NiFe]-hydrogenases, present in archaea and bacteria, generally function to oxidize
molecular H2 and provide reducing equivalents for metabolic processes, while the [FeFe]hydrogenases, present in bacteria and eukarya, function more broadly to catalyze both
proton reduction and H2 oxidation (1, 8). Recently, there has been a growing interest in
these metalloenzymes due to their inherent applicability toward developing renewable
H2- based energy technology.
The active sites for both [NiFe]- and [FeFe]- hydrogenases have been solved by
x-ray crystallography and are united by the presence of π acceptor CO and CN- ligands,
which are not common in biology. These diatomic ligands stabilize low-spin and lowvalent oxidation states of the metal centers at the active sites.
For the [NiFe]-
hydrogenase, the active sites from a variety of different sulfate reducing bacterial sources
have been determined to consist of a Ni atom coordinated to an Fe atom via two thiolate
ligands and a bridging oxygen species (47). The Ni atom is further coordinated by two
cysteine ligands from the protein, while the Fe atom is coordinated to two terminal CNligands and one terminal CO ligand. In comparison, the [FeFe]-hydrogenase active site
This section has been coauthored by David W. Mulder, Danilo O. Ortillo, David J. Gardenghi,
Anatoli V. Naumov, Shane S. Ruebush, Robert K. Szilagyi, BoiHanh Huynh, Joan B. Broderick,
and John W. Peters (2009) Biochemistry 48, 6240-6248.
46
contains a 6Fe-containing complex cluster termed the H-cluster, as determined for
Clostridium pasteurianum (CpI) (84, 105) and Desulfovibrio gigas (85, 104). The Hcluster consists of a [4Fe-4S]-subcluster coordinated to a 2Fe-subcluster via a cysteine
thiolate ligand. The two Fe centers in the 2Fe-subcluster are bridged via a five atom
dithiolate ligand and a CO ligand. The chemical composition of the dithiolate ligand has
not yet been determined unambiguously and has been proposed to be dithiomethylether
(105), propane dithiolate (85), or dithiomethylamine (104). In addition, both Fe centers
contain terminal CO and CN- ligands. For the presumed oxidized state of CpI, a water
molecule is present at the distal Fe center in proximity to the [4Fe-4S]-subcluster.
Biosynthesis and maturation of the [NiFe]-hydrogenases has been thoroughly
studied including identification of at least six gene products involved in formation of
active [NiFe]-hydrogenases and the interactions between gene products during
maturation. The metabolic source of diatomic CN- ligands has been identified to be
carbamoyl phosphate (139) whereas the metabolic source for the CO ligand is still in
question.
In contrast, relatively little is known concerning the biosynthesis and
maturation of [FeFe]-hydrogenases.
By analysis of several mutant strains of
Chlamydomonas reinhardtii that are unable to produce hydrogen, the genes hydEF, and
hydG were discovered to be required for maturation of [FeFe]-hydrogenases (140).
Subsequent expression studies revealed that formation of an active [FeFe]-hydrogenase
was achieved only when HydA was heterologously expressed in a background of coexpressed gene products HydEF and HydG in Escherichia coli (140). In most organisms
hydEF exists as two separate genes, hydE and hydF (140), and it has been shown that the
47
coexpression in E. coli of HydE, HydF, and HydG from Clostridium acetobutylicum with
the [FeFe]-hydrogenase structural gene product from various algal and bacterial sources
is sufficient to effect expression of active [FeFe]-hydrogenase (175). Deduced amino
acid sequence analysis of HydE, HydF, and HydG gene products reveal HydE and HydG
to be radical-SAM Fe-S enzymes as they both have the C-X3-C-X2-C radical-SAM
signature motif (139, 140) and preliminary biochemical characterization has revealed
SAM cleavage activity (177).
In addition, it has been shown that upon reconstitution
HydF binds an Fe-S cluster and exhibits GTPase activity (176, 177). Recently, the x-ray
crystal structure of HydE from Thermatoga maritima was determined (217), although its
function in relation to the [FeFe]-hydrogenase maturation process is not yet well
understood.
The involvement of HydE, HydF, and HydG maturation enzymes in the
biosynthesis of the H-cluster was further elaborated when it was shown that HydA,
expressed in a genetic background devoid of HydE, HydF, and HydG (HydA∆EFG) is a
stable protein capable of being activated in vitro by the aforementioned proteins (209). It
was also determined that HydF behaves as a scaffold protein in which an H-cluster
precursor is assembled and can be subsequently transferred to HydA∆EFG resulting in the
formation of an active [FeFe]-hydrogenase (210). Both in vitro activation studies imply
that cluster biosynthesis does not take place on the structural protein (HydA∆EFG) and that
a chemical precursor to the H-cluster is synthesized in the absence of HydA∆EFG that
upon transfer to HydA∆EFG results in its activation. Although no chemical precursors or
intermediates to the H-cluster have yet been characterized or even identified, it can be
48
hypothesized that HydE, HydF, and HydG are directed toward the synthesis of (1) the
entire 6Fe-containing H-cluster, (2) the 2Fe-subcluster of the H-cluster, or (3) the
biologically unique ligands of the 2Fe-subcluster of the H-cluster. The characterization
of HydA∆EFG provides a critical link in our understanding of this fascinating process by
defining the substrate for the Hyd maturation proteins.
In this study, we present spectroscopic and biochemical characterizations of
HydA∆EFG from C. reinhardtii to provide insights into the [FeFe]-hydrogenase maturation
and H-cluster biosynthesis.
HydA from the eukaryotic green algae C. reinhardtii
contains only the H-cluster binding domains and represents the simplest [FeFe]hydrogenase known. Unlike [FeFe]-hydrogenases from C. pasteurianum and D. gigas,
the [FeFe]-hydrogenases from eukaryotic green algae do not contain additional accessory
Fe-S clusters with plant-type ferredoxin domains that would complicate spectroscopic
characterization of the Fe-S clusters present at the active site (99-101).
Our
characterization of HydA∆EFG from C. reinhardtii indicates that a [4Fe-4S] cluster is
present in HydA∆EFG and is required for in vitro activation by the HydE, HydF, and HydG
maturation enzymes.
Accordingly, it follows that the aforementioned maturation
enzymes are not directed at the synthesis of the entire 6Fe-containing H-cluster.
Experimental Procedures
Cloning and Cell Growth Conditions
HydA∆EFG from C. reinhardtii was cloned into a pET Duet vector as described
previously (175) and modified for the presence of an N-terminal 6x-histidine tag.
49
HydA∆EFG from C. reinhardtii was expressed in E. coli BL21(DE3) cells and cultivated in
either 2 L flasks with 1 L medium volume or in a 10 L bench top fermentor (New
Brunswick) containing modified MOPS minimal medium (220) supplemented with 5.5%
glucose and 150 µg/mL ampicillin.
Either 5 mL or 50 mL overnight cultures of
BL21(DE3) cells were used to inoculate 1L and 10L cultures, respectively. Later, cell
expressions were cultivated in LB media buffered with 50 mM phosphate pH 7.6,
supplemented with 5.5% glucose and 150 µg/mL ampicillin. The cells were grown at
37°C with vigorous shaking (flasks) and agitation/aeration (fermentor, 250 rpm, 3 L/min)
to an optical density of 0.5 (measured at 600 nm with a visible spectrophotometer from
Thermo Spectronic) and induced by addition of IPTG to 1 mM final concentration.
(NH4)2Fe(SO4)2·6H2O (191 µM) was also added at induction. Induction was allowed to
proceed aerobically for 2 hours at 37°C followed by 16 hours at 4°C under nitrogen purge
with additional supplemented (NH4)2Fe(SO4)2·6H2O (191 uM). The cells were harvested
anaerobically (8,000 rpm, 4°C), washed with buffer A (50 mM HEPES pH 7.6, 150 mM
NaCl, 1 mM DTT) in an anaerobic Coy chamber (Coy Laboratories), and stored at -80°C.
HydA∆EFG Purification
Cell-free extracts were prepared by resuspending the cells (described above) in
degassed anaerobic buffer B (5 mL of buffer per gram of cells, 50 mM HEPES pH 7.6,
50 mM NaCl, 5% (w/v) glycerol, 1% (w/v) Triton X-100, 10 mM MgCl2, 1 mM PMSF, 1
mM DTT, and trace quantities of lysozyme and DNAase), and placing the suspension in a
pressure cell bomb (1000 psi nitrogen pressure, 1 hour) followed by centrifugation of the
lysate (36,000 x g, 45 minutes). HydA∆EFG was purified from the cell lysate by anion
50
exchange chromatography on Q-sepharose resin (GE Healthcare) followed by His-tag
affinity chromatography to a Co2+ resin (Talon resin; Clontech); all purification steps
were done anaerobically using thoroughly degassed buffers under positive nitrogen
pressure.
Cell lysates were loaded onto a 100 mL Q-sepharose column previously
equilibrated in buffer C (50 mM HEPES, pH 7.6, 20% glycerol, 1 mM DT). The column
was washed with buffer C supplemented with 100 mM NaCl, and HydA∆EFG was eluted
with buffer C in arrangement with a NaCl gradient up to 1 M final concentration while
monitoring the absorbance at 405 nm.
HydA∆EFG protein containing fractions were
golden-brown in color, consistent with the presence of Fe-S clusters. These fractions
were loaded onto a 25 mL Co2+ affinity column. The column, equilibrated with buffer D
(50 mM HEPES, pH 7.0, 300 mM NaCl, 20% glycerol, 0.5 mM DT, 5 mM βME), was
washed with buffer D supplemented with 10 mM imidazole, and HydA∆EFG was eluted
using an imidazole gradient up to 100 mM in buffer D while monitoring the absorbance
at 405 nm. The golden-brown protein containing fractions were concentrated using an
Amicon concentration cell under positive argon pressure with a 30 kDa cutoff membrane
filter. The protein was desalted in a Coy anaerobic chamber using a Sephadex 2 mL G25 column (GE Healthcare) equilibrated with buffer C and stored under liquid nitrogen.
Overall, six different HydA∆EFG protein preparations were carried out.
For later
preparations, it was determined that the quality of the spectroscopic data collected could
be improved by selectively saving HydA∆EFG fractions with the highest 405 to 280 nm
absorbance ratios as measured after elution from the Q-sepharose and Co2+ His-tag
affinity columns.
51
Assays
Protein concentrations were determined using the Bradford assay (221) with
bovine serum albumin (Sigma) as the standard. Specific hydrogenase activity of the
purified HydA∆EFG was measured using gas chromatography upon maximum in vitro
activation with cell lysates containing maturation proteins HydE, HydF, and HydG from
C. acetobutylicum as described by McGlynn et al. (209). In the assays, dithionite (20
mM) was used as the reducing agent, methyl viologen (10 mM) was used as the electron
carrier, and HydE, HydF, and HydG cell extracts from C. acetobutylicum were used to
activate HydA∆EFG. The iron content of HydA∆EFG was determined using a procedure
described by Fish (222), which uses ferrozine under reductive conditions after digestion
of the protein in 4.5% (w/v) KMnO4/1.2 N HCl. Iron standards were prepared by dilution
of a commercial Fe AA standard (Ricca Chemical Co.). Sulfide assays were conducted
according to a procedure described by Beinert (223) and Broderick (224) and sulfide
standards were prepared from Na2S·9H2O.
Electronic Absorption Spectroscopy
For UV-visible spectroscopic experiments, samples were prepared inside an
anaerobic Coy chamber and transferred to anaerobic 1 mL cuvettes (NSG Precision Cells,
Inc). UV-visible spectra were recorded at room temperature with a Cary 300 (Varian)
spectrophotometer. Reduced samples were prepared by adding 2 mM DT.
52
EPR Spectroscopy
Low temperature X-band EPR spectra were recorded using a Bruker ESP300E
spectrometer equipped with a liquid helium cryostat and temperature controller from
Oxford Instruments. Typical EPR parameters were as follows: sample temperature 12 K;
microwave frequency 9.36 GHz, attenuation 20.4 dB; microwave power 1.85 mW.
Sample concentrations varied between 128 µM and 512 µM. Reduced samples were
prepared by adding 2 mM DT and oxidized samples were prepared by titrating in
increasing concentrations of ferricyanide (2-11 mM). Spin concentration was determined
by double integration of the sample spectra using CuSO4 (0.20 mM) / EDTA (2.0 mM) as
the standard measured under identical conditions. Basic analysis of the collected spectra
was done using the computer software program SpinCount (Michael Hendrich, Carnegie
Mellon University).
Mössbauer Spectroscopy
57
Fe was purchased from Cambridge Isotope Laboratories, Inc and dissolved in
hot concentrated hydrochloric acid. The pH was adjusted with NaOH. HydA∆EFG asisolated
57
Fe samples were prepared from 10 L cultures using the defined minimal
medium (220) described above with 57Fe substituted in at the same molar concentrations
for 56Fe.
57
Fe (191 µM) was also added at induction with IPTG. Protein samples (500-
800 µM) were loaded into 450 µL cups and stored under liquid nitrogen. Mössbauer
spectra were recorded on a Mössbauer spectrometer equipped with a Janis 8DT variable
temperature cryostat and operated at a constant acceleration mode in transmission
geometry. The zero velocity refers to the centroid of a room temperature spectrum of a
53
metallic iron foil. Analysis of the spectra was performed with the WMOSS program
(WEB Research).
Fe K-edge X-ray Absorption Spectroscopy
HydA∆EFG samples (1.9 mM) were prepared from HydA∆EFG protein isolated as
described above. The EXAFS cells (Delrin) sealed with thin Fe-free Kapton tape were
loaded with ~100 µL of sample. Fe K-edge X-ray absorption spectroscopic (XAS)
measurements were carried out at beamline 7-3 (BL7-3) of the Stanford Synchrotron
Radiation Lightsource (SSRL) under storage ring (SPEAR3) conditions of 3 GeV energy
and 100-80 mA current on two different occasions. BL7-3 is a 20-pole, 2 T Wiggler
beam line equipped with a Si(220) downward reflecting, double-crystal monochromator.
Data were collected in the energy range from 6785 eV to k=17 Å-1 above the Fe K-edge
using an unfocused beam. The frozen solution samples were mounted under liquid
nitrogen and measured in a liquid He cryostat at about 11 K temperature. The beam line
parameters were optimized at 8000 eV. The Fe Kα fluorescence signal was collected
using a 30-element Ge array detector and with a Soller slit and Z-1 (Mn) filter. The
energy windowing of the detector was carefully done to minimize fluorescence signal due
to scattering and other non-Fe Kα emission sources.
ATHENA (225), a graphical-user interface to IFEFFIT (225) was used for
averaging and background subtraction. The data were calibrated to the first rising-edge
inflection point of the XAS spectra that was assigned to 7111.2 eV of an iron foil. The
data are averages of at least 5 scans before normalization or background subtraction.
54
ATHENA and AUTOBK were used to spline the post-edge region and to obtain the EXAFS
with Rbkg of 1.
ARTEMIS (225), ATOMS (225), and FEFF (226) were used to model and fit the
data, and
calculate Fe…Fe and Fe-S scattering paths, respectively.
The structural
models used in scattering calculations were derived from combination of average Fe…Fe
and Fe-S distances of reduced and oxidized [4Fe-4S] model compounds (227). Due to
the presence of various Fe environment which lead to various reasonable fits [defined as
R(fit) < 10-3], we used the composition of Fe-S clusters in the EXAFS fits as constraints
from corresponding Mössbauer measurements.
Reconstitution of Iron-Sulfur Clusters in HydA∆EFG
HydA was subjected to reconstitution conditions following the general procedures
described for biotin synthase (196). The protein (10 µM) was incubated with FeCl3 (100
µM), Na2S (100 µM) and DTT (1 mM) in buffer E (50 mM HEPES pH 7.0, 300 mM
NaCl, 20% glycerol) for 2-3 hours with constant stirring in an anaerobic Coy chamber.
Visually, the color of diluted protein solution was observed to change to golden brown.
All reagents were added sequentially and following reconstitution excess ions were
removed using a G-25 Sephadex column. The resulting reconstituted HydA∆EFG was
assayed for hydrogenase activity by in vitro activation with HydE, HydF, and HydG from
C. acetobutylicum and iron content was also analyzed as described earlier.
55
Preparation of Apo-HydA∆EFG and Reconstitution
Apo-HydA∆EFG from C. reinhardtii was prepared by stripping out all Fe-S clusters
using EDTA as an Fe chelator. HydA∆EFG from C. reinhardtii was incubated aerobically
with EDTA (100 mM) for one hour in buffer E. Fe-S chelation was monitored visually
and within 1 hour the golden brown protein solution turned colorless. The resulting apoprotein was made anaerobic by degassing it under vacuum with sequential nitrogen purge
and incubation with 5 mM DTT. Excess EDTA was removed with a G-25 Sephadex
column and verification that all Fe was stripped from the protein was carried out by Fe
analysis. Apo-HydA∆EFG from C. reinhardtii was reconstituted by incubating the protein
(1 µM) with a large excess of FeCl3, Na2S and DTT (1 mM) in buffer E for 2-3 hours.
Residual FeCl3 and Na2S were removed with a G-25 Sephadex column and the protein
was concentrated with a 30 kDa Centricon® centrifugal filter device (Millipore). The
colorimetric iron assay was used to monitor addition of Fe. The hydrogenase activities of
the different HydA∆EFG forms (HydA∆EFG, apo- HydA∆EFG, and reconstituted apoHydA∆EFG) were determined following activation with the HydE, HydF, and HydG
maturation enzymes.
Results and Discussion
HydA∆EFG Binds a [4Fe-4S] Cluster
Anaerobic purification of heterologously expressed C. reinhardtii HydA∆EFG (49
kDa, Figure 2.1A) in E. coli gives a stable protein that is capable of being activated in
vitro by C. acetobutylicum extracts containing HydE, HydF and HydG.
56
Figure 2.1. (A) SDS-PAGE gel (10%) showing purification of HydA∆EFG from C. reinhardtii
following elution from the Co2+His-tag affinity column. The left lane is the protein marker
standards and the right lanes show the pure HydA∆EFG from C. reinhardtii at 49 kDa. (B) UVvisible spectra of HydAÄEFG as-isolated (blue) and HydA∆EFG reduced with 2 mM DT
(red/slashed). The feature centered at 400 nm is characteristic for the presence of Fe-S cluster
LMCT bonds.
Specific activities of the 6 separate HydA∆EFG protein purifications after in vitro
activation with HydE, HydF, and HydG enzymes ranged from 10 - 38 µmol H2 min-1mg-1
HydA∆EFG. For in vivo co-expression of HydA from C. reinhardtii with the maturation
genes from C. acetobutylicum in E. coli, H2 evolution activity was previously reported to
57
be 150 µmol H2 min-1mg-1 (175). The difference in activity between in vitro and in vivo
expression systems is likely attributed to low occupancy of H-cluster activating
precursors assembled by HydE, HydF, and HydG, as well as the heterogeneity of the
maturation system (210).It was evident that HydA∆EFG binds Fe-S clusters as observed by
the dark brown color of the protein solution following purification as well as Fe (3.1 ±
0.5 Fe atoms / HydA∆EFG) and S2- (1.8 ± 0.5 S2- atoms / HydA∆EFG) analyses. It is
important to note that the amount of Fe and S2- bound / HydA∆EFG varied slightly between
different purifications, suggesting that the Fe-S cluster present may be slightly labile
during purification.
The UV-visible absorbance spectrum shows a broad shoulder
centered near 405 nm that upon reduction with DT decreases in intensity (Figure 2.1B).
These absorbance features are consistent with the presence of Fe-S clusters in HydA∆EFG.
The presence of Fe-S clusters in HydA∆EFG was further confirmed by the low
temperature X-band EPR spectrum of reduced HydA∆EFG which revealed an axial S = ½
signal (g = 2.04, 1.91) characteristic of a reduced [4Fe-4S]1+ cluster (Figure 2.2) (228).
Also, a slight shoulder was observed at g = 2.04, however appearance of this feature
varied between different protein preparations and could be minimized when the protein
was further purified based on collecting protein fractions with maximal 405/280 nm
absorbance. The temperature and power dependence of the axial S = ½ signal showed a
maximum intensity around temperatures of 10 K and diminished significantly at
temperatures greater than 30 K at 2 mW power (Figure 2.3). The signal was no longer
observed above temperatures of 40 K and also did not saturate with increasing
microwave power. These temperature and power signal characteristics are typical for the
58
presence of a [4Fe-4S]1+ cluster (229). Spin quantification of the signal, using CuSO4
(0.20 mM) / EDTA (2.0 mM) as a standard, gave 0.17 spins/Fe atom. Upon oxidation of
HydA∆EFG with ferricyanide the [4Fe-4S]1+ cluster signal disappeared and no EPR signal
was observed (data not shown).
Figure 2.2 EPR spectrum of reduced HydA∆EFG. Parameters for EPR measurement were as follows:
sample temperature 12 K; microwave frequency 9.36 GHz; attenuation, 20.4 dB; microwave power 1.85
mW; receiver gain 2.0 x 104. The reduced sample was prepared by adding 2 mM DT to as-isolated
HydA∆EFG (235 uM). Upon oxidation of HydA∆EFG with ferricyanide, the [4Fe-4S] cluster signal is no
longer observed.
59
Figure 2.3. Temperature (top) and microwave power (bottom) dependency of the S = ½ signal (g
= 2.04, 1.91) from reduced HydA∆EFG. Changes in the signal amplitude (S) of the spectral feature
at g = 2.04 were monitored at various temperatures and powers (P). For temperature dependence,
samples were measured at 2.0 mW and for power dependence the temperature was constant at 10
K. Maximum signal was observed at 10K and diminished significantly in intensity above 30K.
For increasing powers, the signal showed no saturation. These characteristics are typical for a
[4Fe-4S] cluster signal. Other EPR settings are as follows: microwave frequency 9.24 GHz; gain
1.6 x 103; modulation amplitude 10.00 G; time constant 0.50 sec.
Mössbauer Spectroscopic Characterization of HydA∆EFG
Figure 2.4 shows the Mössbauer spectra of the 57Fe-enriched HydA in its aspurified (A) and dithionite reduced (B and C) forms. The spectra were recorded at 4.2 K
60
in a magnetic field of 50 mT applied parallel (A and B) and perpendicular (C) to the γ
radiation.
Figure 2.4. Mössbauer spectra of the as-purified (A) and dithionite-reduced (B and C) HydA∆EFG.
The spectra (hatched marks) were recorded at 4.2 K in a magnetic field of 50 mT applied parallel
(A and B) and perpendicular (C) to the γ radiation. The solid lines plotted above the data are
simulated spectra for the [4Fe-4S]2+ cluster (red), [4Fe-4S]1+ cluster (blue) and Fe(II) impurities
(green), normalized to the following percent absorptions: (A) 70% [4Fe-4S]2+, 15% [4Fe-4S]1+
and 15% Fe(II), (B and C): 6% [4Fe-4S]2+, 76% [4Fe-4S]1+ and 18% Fe(II). The black lines overlaid
with the experimental spectra are composite spectra. Parameters used for the simulations are
given in the text (for the [4Fe-4S]2+ and Fe(II) impurities) and in the supporting information (for
the [4Fe-4S]1+ cluster).
61
Analysis of the data indicates that all three spectra are composed of three sub-spectral
components: a central quadrupole doublet (solid red lines in Figure 2.4), a magnetically
split spectrum (blue lines), and an outer quadrupole doublet with broad and asymmetric
absorption lines (green lines). The central quadrupole doublet is attributed to a [4Fe4S]2+ cluster and can be simulated as a superposition of two unresolved equal-intensity
quadrupole doublets representing the two mixed-valence Fe(II)Fe(III) pairs within the
cluster. The parameters obtained for the two unresolved doublets (∆EQ(1) = 1.34 mm/s,
δ(1) = 0.45 mm/s, and Γ(1) (line-width) = 0.30 mm/s; ∆EQ(2) = 1.05 mm/s, δ(2) = 0.44
mm/s, and Γ(2) = 0.43 mm/s) are typical for [4Fe-4S]2+ clusters (123, 230).
The
magnetically split spectral component exhibits an intensity pattern that depends on the
direction of the applied field (blue lines in B and C), indicating that it originates from an
EPR active Fe center. As reported above, reduced HydA displays an S =½ EPR signal
that can be assigned to a [4Fe-4S]1+ cluster.
The magnetic Mössbauer spectral
component is therefore attributed to this S = ½ Fe cluster. Consistent with the [4Fe-4S]1+
assignment, initial analysis of this spectral component indicates that it may be simulated
as a superposition of two spectral components arising from the Fe(II)Fe(II) and Fe(II)Fe(III)
pairs of a [4Fe-4S]1+ cluster (Figure 2.5) (123, 230). On the basis of the parameters
obtained for the outer quadrupole doublet (∆EQ = 2.77 mm/s, δ = 1.30 mm/s, Γ(left) =
0.53 mm/s and Γ(right) = 0.80 mm/s), this component is attributed to non-cysteine
coordinated extraneous FeII impurities, which may be generated via cluster degradation.
62
Figure 2.5. 4.2-K Mössbauer spectra of the [4Fe-4S]1+ cluster in HydA∆EFG. The spectra (Hatched
marks) were prepared by removing the contributions of the [4Fe-4S]2+ cluster (6%) and Fe(II)
impurities (18%) from the raw data shown in Figure 2.4B and 2.4C. The purple and cyan lines
are, respectively, simulated spectra for the Fe(II)Fe(II) and Fe(II)Fe(III) pairs of the [4Fe-4S]1+ cluster.
The parameters used are ∆EQ = 1.7 mm/s, δ = 0.62 mm/s, η = 1, Axx/gnβ n = 18.3 T, Ayy/gnβ n = 2.8
T, Azz/gnβ n = 4.6 T and Γ = 0.47 mm/s for the Fe(II)Fe(II) pair and ∆EQ = 1.6 mm/s, δ = 0.49 mm/s,
η = 0, Axx/gnβ n = −21.2 T, Ayy/gnβ n = −14.5 T, Azz/gnβ n = −15.6 T and Γ = 0.50 mm/s for the
Fe(II)Fe(III) pair. The blue lines are the composite spectra.
Decomposition of the Mössbauer spectra into the above described spectral
components shows that in the as-purified HydA∆EFG sample (see Figure 2.4A) the
majority (70%) of the Fe are present in the oxidized [4Fe-4S]2+ form while a small
percentage (15%) are in the reduced [4Fe-4S]1+ form. The remaining Fe (15%) is present
63
as Fe(II) impurities. Upon reduction (see Figure 2.4B and 2.4C), a substantial amount of
the [4Fe-4S]2+ clusters are reduced to the [4Fe-4S]1+ clusters, resulting in a decrease in
the Mössbauer absorption of the [4Fe-4S]2+ cluster from 70% to 6% and an increase in
the [4Fe-4S]1+ absorption to 76%. The Fe(II) impurities also increase slightly to 18%,
suggesting that the [4Fe-4S] cluster in HydA∆EFG is slightly unstable under dithionite
reducing conditions. Thus, the Mössbauer data unambiguously show that the as-purified
HydA∆EFG contains predominantly [4Fe-4S] clusters. No other types of Fe-S clusters are
detected. The observation that the [4Fe-4S]2+ cluster in HydA∆EFG can be reduced by
dithionite to the [4Fe-4S]1+ state establishes further that HydA∆EFG contains a redox
active [4Fe-4S]2+/1+ cluster. Taking into consideration the Fe and protein content (1.9 Fe
atoms/ HydA∆EFG) determined for the as-purified
57
Fe-enriched HydA∆EFG and the total
percent absorption of the [4Fe-4S] cluster detected by the Mössbauer measurement
(85%), a stoichiometry of 0.4 [4Fe-4S] clusters/ HydA∆EFG for the as-purified HydA∆EFG
may be concluded.
Fe K-edge X-ray Absorption Spectroscopic Characterization of HydA∆EFG
The Fourier transform of EXAFS (FT-EXAFS) for the as-isolated HydA sample
(black trace) and with a representative fit (red trace) are shown in Figure 2.6.
64
Figure 2.6. FT-EXAFS plot (A) and individual EXAFS contributions (B) for as-isolated
HydA∆EFG sample with a representative fit containing Fe…Fe, Fe-S(sulfide), Fe-S(thiolate)
scattering paths.
The fitting parameters are summarized in Table 2.1. Among numerous reasonable fits
with R(fit) values of 10-4, Table 2.1 and Figure 2.6 reports the one which was obtained
using the Mössbauer results (see above) as constraints for the amount and distribution of
65
various Fe sites. The initial parameters of the fit were set to represent the 70% oxidized
and 15% reduced protein embedded [4Fe-4S] cluster content. The average Fe…Fe, FeSt, and Fe-Ss distances in synthetic [4Fe-4S] complexes are 2.74 ± 0.01 Å, 2.25 ± 0.01 Å,
and 2.27 ± 0.03 Å (227). The protein bound counterparts of Fe-S distances tend to be
about 0.02 Å longer due to dipole and hydrogen bonding interactions to the sulfides and
thiolate sulfurs, which reduces the nucleophilicity of the sulfur atoms and thus the
covalency of the sulfur-iron bonds.
Table 2.1. Representative fitting parameters for the as-isolated HydA∆EFG sample using Fe-S
composition from Mössbauer measurements
scatterers
Fe-Ss
Fe-St
Fe…Fe
a
parametersa
r, Å
N, Α, −
σ2, Å2
r, Å
N, Α, −
σ2, Å2
r, Å
N, Α, −
σ2, Å2
fitted value
2.27
3
0.80
0.003
2.27
1
0.46
0.003
2.72
3
0.44
0.003
R (fit)
E0, eV
7.65·10-4
0.790
r: scattering path, N: coordination number, A: scattering path amplitude, σ2 Debye-Waller factor
as a measure of thermal displacement and disorder;
k3-weighted data fit in the k-range of 1-14 Å-1; FT-EXAFS window range fitted 1.5-3.3 Å
These differences between the synthetic model and protein bound Fe-S clusters can be
observed by EXAFS as demonstrated by a series of FT-EXAFS analysis of Fe-S clusters
(231-233). Using the initial distances with standard deviations as initial Debye-Waller
66
factors of 2.74 ± 0.005 Å, 2.24 ± 0.003 Å, 2.28 ± 0.003 Å for Fe…Fe, Fe-St, and Fe-Ss
scattering paths, we obtained a reasonable fit that is shown in Figure 2.6. In order to
avoid negative Debye-Waller factors (σ2) and large deviations of the edge positions (E0)
the final fit was obtained by linking all the σ2 and E0 values, while allowing the
individual path lengths and their amplitudes vary. The parameters in Table 2.1 fit
acceptably well with the bulk of the experimental data (black trace), while showing poor
fit at the shorter distances. The residual FT-EXAFS intensity at these distances is
consistent with the presence of about 15% of free iron that is likely partially solvated
and/or coordinated with low Z atoms, such as O and N from protein residues as suggested
by the Mössbauer results. The peak at around 1.9±0.1 Å can be reasonably well fitted
with an Fe center surrounded by six low Z (O,N) scatterers in 15% abundance. The
relatively short Fe-O distance of 1.9 Å is indicative of the presence of negatively charged
O/N ligands for the Fe(II) impurities (see above). The fit components and parameters for
the latter are given in Figure 2.7 and Table 2.2. For both FT-EXAFS fits the resulting
Fe…Fe (2.71-2.72 Å) and Fe-S (2.27 Å) distances are highly similar to those observed
for the oxidized and reduced [4Fe-4S] cluster in Av2 (2.72 and 2.73 Å; 2.27 and 2.29 Å,
respectively) (233) and thus further support the presence of [4Fe-4S] cluster in
HydA∆EFG.
67
Figure 2.7. FT-EXAFS plot (A) and individual EXAFS contributions (B) for as-isolated
HydA∆EFG sample with a representative fit containing Fe…Fe, Fe-S(sulfide), Fe-S(thiolate), Felow Z(O/N) scattering paths.
68
Table 2.2 Representative fitting parameters for the as-isolated HydA∆EFG sample using Fe-S
composition from Mössbauer measurements including the free Fe(II) content
scatterers
Fe-Ss
Fe-St
Fe…Fe
Fe-O
a
parametersa
r, Å
N, Α, −
σ2, Å2
r, Å
N, Α, −
σ2, Å2
r, Å
N, Α, −
σ2, Å2
E0, eV
fitted value
2.27
3
0.88
0.004
2.27
1
0.48
0.004
2.71
3
0.48
0.004
-0.436
r, Å
N, Α, −
σ2, Å2
E0, eV
1.87
1b
0.55
0.002
4.609
R (fit)
E0, eV
3.89⋅10-4
-0.436
r: scattering path, N: coordination number, A: scattering path amplitude, σ2 Debye-Waller factor
as a measure of thermal displacement and disorder;
k3-weighted data fit in the k-range of 1-14 Å-1; FT-EXAFS window range fitted 1.5-3.3 Å
Reconstitution of the [4Fe-4S] Cluster in HydA∆EFG
It was previously reported that HydA∆EFG can be activated by cell extracts
containing the maturation enzymes HydE, HydF, and HydG (209).
This observation,
coupled with our evidence described above for a [4Fe-4S] cluster in HydA∆EFG, suggests
that the [4Fe-4S] form of HydA∆EFG is the substrate for assembly of the H-cluster by
HydE, HydF, and HydG, and further that the [4Fe-4S] cluster present in HydA∆EFG
69
becomes part of the H-cluster upon activation.
In order to further explore the
requirement for a preformed [4Fe-4S] cluster in HydA∆EFG during activation with HydE,
HydF, and HydG, HydA∆EFG was chemically reconstituted with FeCl3 and Na2S.
Fe
analysis of the reconstituted HydA∆EFG gave 4.0 ± 0.1 Fe / HydA∆EFG, and the EPR
spectrum of reconstituted reduced HydA∆EFG indicates the presence of a [4Fe-4S] cluster,
with an axial S = ½ EPR signal (g = 2.04 and 1.91) essentially identical to that of the asisolated enzyme. In vitro activation of the reconstituted HydA∆EFG with HydE, HydF,
and HydG gave hydrogenase activity (31.5 ± 0.5 µmol H2 min-1mg-1 HydA∆EFG)
approximately two-fold greater than that of the as-isolated HydA∆EFG activated with
HydE, HydF, and HydG (17.1 ± 2.4 µmol H2 min-1mg-1 HydA∆EFG). These observations
provide additional evidence that the [4Fe-4S] form of HydA is the substrate for H-cluster
assembly by HydE, HydF, and HydG, as increasing the [4Fe-4S] content increases the
ability to activate HydA∆EFG. In addition, the reconstitution results are consistent with
our previous suggestion that the [4Fe-4S] cluster in HydA∆EFG is labile during
purification and is subsequently repopulated or repaired during reconstitution.
In
contrast, if the [4Fe-4S] cluster present in HydA∆EFG was not required for activation by
HydE, HydF, and HydG, it is likely that reconstitution would inhibit in vitro activation by
generating a cluster at the H-cluster site that would prevent activation by the maturation
enzymes.
Metal Chelation and Reconstitution
To further explore the hypothesis that the [4Fe-4S] cluster present in HydA∆EFG is
required for in vitro activation with HydE, HydF, and HydG, apo- HydA∆EFG was
70
prepared and then subsequently reconstituted with FeCl3 and Na2S; both apo and
reconstituted samples were subjected to in vitro activation by HydE, HydF, and HydG
(Figure 2.8).
Figure 2.8. Flow scheme outlining Fe content and hydrogenase activity during preparation of
apo- HydA∆EFG by stripping out all existing Fe-S cluster in HydA∆EFG and ensuing reconstitution
of apo- HydA∆EFG. Apo-HydA∆EFG shows no hydrogenase activity following in vitro activation.
Reconstituted apo-HydA∆EFG displayed 90% of initial hydrogenase activity (compared to asisolated HydA∆EFG) upon in vitro activation with maturases. Also, Fe analysis of the
reconstituted apo- HydA∆EFG gave 4.0 ± 0.1 Fe atoms / protein.
The apo-sample was found to contain 0 Fe atoms / HydA∆EFG, and upon in vitro
activation with HydE, HydF, and HydG maturation enzymes, the apo- HydA∆EFG protein
showed no hydrogenase activity. Following chemical reconstitution of apo- HydA∆EFG
with FeCl3 and Na2S the protein contained 4.0 ± 0.1 Fe atoms / HydA∆EFG and had an
EPR spectrum essentially identical to that of the as-isolated enzyme. Subsequent in vitro
activation of the reconstituted enzyme with HydE, HydF, and HydG produced
hydrogenase activity at a level of 90% that of as-isolated HydA∆EFG. The slightly lower
activity in combination with increased Fe content relative to that obtained for the as-
71
isolated enzyme is likely a reflection that Fe may bind advantageously during
reconstitution.
Nevertheless, these results clearly demonstrate that activation of
HydA∆EFG by HydE, HydF, and HydG requires the presence of a preformed [4Fe-4S]
cluster on HydA.
Relevance to H-cluster Biosynthesis
In relation to the overall scheme of [FeFe]-hydrogenase maturation and H-cluster
biosynthesis, these results provide significant insights into the role of HydE, HydF, and
HydG maturation machinery. Because our results show that HydA∆EFG contains a redox
active [4Fe-4S]2+/+ cluster that is required for in vitro activation, logically it follows that
HydE, HydF, and HydG maturation machinery does not transfer a complete 6Fecontaining H-cluster to HydA∆EFG.
Moreover, HydE, HydF, and HydG maturation
enzymes must be directed toward the synthesis of the 2Fe-subcluster of the H-cluster or
perhaps only the unique dithiolate and diatomic ligands of the 2Fe-subcluster. The later
hypothesis, however, seems more unlikely as CO and CN- ligands would presumably
need a chemical platform of some sort (such as a 2Fe-cluster) to be introduced into the
maturation scheme. In light of previous work demonstrating that HydF serves as a
scaffold or carrier protein which harbors an H-cluster precursor that is able to activate
HydA∆EFG (210), the current results suggest this precursor may be some form of the 2Fesubcluster of the H-cluster.
We hypothesize that HydE and HydG function to synthesize the unique ligands on
the 2Fe-subcluster (CO, CN- , and dithiolate ligands) using HydF as a scaffold (Figure 6).
72
Once ligand modified, the oxidation state of the 2Fe-subcluster in its reduced active state
is believed to be Fe(I)Fe(I) as indicated by Mössbauer and theoretical studies (116, 122,
123, 234). We have previously proposed that HydE and/or HydG initially modify a basic
[2Fe-2S] cluster with a dithiolate ligand in chemistry analogous to sulfur-carbon bond
insertion reactions by radical-SAM enzymes BioB (235) and LipA (236) in the synthesis
of biotin and lipoic acid, respectively (237).
This ligand modification should shift
reactivity to the Fe atoms setting up for further modification by addition of CO and CNligands. HydE and/or HydG could also be responsible for the generation CO and CNligands by glycine radical decomposition in a reaction analogous to radical-SAM
enzymes (pyruvate formate lyase activating enzyme (199), lysine amino mutase (180),
and ThiH (200, 201)) that catalyze the formation of amino acid radicals. Alternative
related proposals have been offered by Fontecave, Fontecilla-Camps, and coworkers
invoking precursors such as thiocyanate and glycine as substrates for the non-protein
ligands of the H-cluster (202, 217).
Regardless these proposals, the unifying point
between hypotheses is that in the final step of [FeFe]-hydrogenase maturation, HydF
transfers the ligand containing 2Fe-subcluster to HydA∆EFG, which already contains the
[4Fe-4S] cubane of the H-cluster, and is poised to accept the insertion of the 2Fesubcluster completing [FeFe]-hydrogenase activation (Figure 6).
73
Figure 2.9. Hypothetical scheme for [FeFe]-hydrogenase maturation. In two steps, radical-SAM
enzymes HydE and HydG use HydF as a scaffold to modify a basic [2Fe-2S] cluster with the
addition of (1) a dithiolate ligand and (2) CO and CN- ligands. The first step is proposed to take
place via a sulfur-carbon bond insertion reaction and the second step from an amino acid
precursor by radical formation (237). After assembly of the ligand modified 2Fe-subcluster,
HydF transfers it to HydA∆EFG, which already houses the [4Fe-4S]-subcluster of the H-cluster,
completing activation of [FeFe]-hydrogenase and H-cluster biosynthesis. An atomic model of the
H-cluster in activated HydA∆EFG is depicted showing the coupling of the [4Fe-4S] cubane to the
2Fe-subcluster with CO, CN- and dithiolate ligands via a bridging cysteine ligand (from PDB
code 3C8Y (105)); the coloring scheme is as follows: Fe: dark red, S: orange, O: red, N: blue, C:
dark grey, unknown atom of dithiolate ligand: magenta. Also, a water molecule is present at the
distal Fe of the 2Fe-subcluster in the presumed oxidized state of the H-cluster. The homology
models of HydA∆EFG and HydA of C. reinhardtii were constructed using the homology server
Phyre (238) and HydA from C. reinhardtii was threaded on HydA from C. pasteurianum (CpI)
during sequence alignment. Ribbon representations of the structures were made in PyMOL
(239) with the c-terminus domain colored in red.
74
It is reasonable that HydA∆EFG can contain a [4Fe-4S] cluster without the hydrogenase
maturation proteins since E. coli has endogenous machinery (eg. ISC, iron sulfur cluster
assembly system) that can easily assemble basic [4Fe-4S] clusters (240). Transfer of the
2Fe-subcluster to HydA∆EFG, already containing the [4Fe-4S]-subcluster of the H-cluster,
was originally proposed in a hypothesis paper regarding [FeFe]-hydogenase maturation
based solely upon enzyme function and energetic (237).
Conclusions
We have demonstrated that HydA∆EFG from C. reinhardtii contains a [4Fe-4S]
cluster at partial occupancy and further that the presence of this [4Fe-4S] cluster is
required for in vitro activation. The presence of the iron sulfur cluster was evidenced by
chemical analysis for iron and acid-labile sulfide, as well as by spectroscopic techniques
that together pointed to a [4Fe-4S]2+/+ cluster, with an occupancy of ~40%, in the asisolated enzyme. This [4Fe-4S] cluster-containing enzyme is capable of undergoing
activation by HydE, HydF, and HydG to produce an active hydrogenase.
Apo-
HydA∆EFG, in contrast, does not produce an active hydrogenase upon incubation with
HydE, HydF, and HydG under activation conditions. These results suggested that the
[4Fe-4S] cluster present in the as-isolated enzyme is required to be present on HydA∆EFG
prior to activation by HydE, HydF, and HydG. Further support for this conclusion was
provided by the observation that apo-HydA∆EFG reconstituted to contain a [4Fe-4S]
cluster could subsequently be converted to an active hydrogenase by HydE, HydF, and
HydG. Together, our results provide important new insights into the process by which
75
HydA is matured to an active enzyme in vivo.
First, a [4Fe-4S] cluster must be
assembled on HydA, in a process that is presumably dependent on the housekeeping ironsulfur cluster assembly proteins. In the second step of maturation, HydE, HydF, and
HydG likely serve to synthesize, assemble, and insert the 2Fe-subcluster to generate the
H-cluster on HydA. Our previous results provide evidence that HydF serves as a scaffold
in this process (210), and the radical AdoMet enzymes HydE and HydG have been
proposed to catalyze the synthesis of the CO, CN-, and dithiolate ligands to the 2Fesubcluster (237).
76
CHAPTER 3
STEPWISE [FeFe]-HYDROGENASE H-CLUSTER ASSEMBLY REVEALED IN THE
STRUCTURE OF HYDA∆EFG
Introduction
Complex Fe-S cluster-containing enzymes are ubiquitous in nature where they are
involved in a number of fundamental processes including carbon dioxide fixation,
nitrogen fixation, and hydrogen metabolism. The biosynthesis and assembly of complex
metalloclusters requires multiple enzymes, scaffolds, and carriers, although the detailed
stepwise process is not well understood in any case (147, 241, 242). One of the more
complex and unusual biological clusters is found in the [FeFe]-hydrogenase; the activesite H-cluster in these enzymes has a [4Fe-4S]-subcluster bridged via a cysteine thiolate
to a 2Fe-subcluster, which in turn is coordinated by terminal CO and CN- ligands, a
bridging CO ligand, and a bridging dithiolate (84, 85) (Figure 3.1). While the exact
composition of the bridging dithiolate ligand is unknown, it is hypothesized to be either
propane dithiolate (85), dithiomethylamine (104), or dithiomethylether (105), with recent
spectroscopic results pointing to dithiomethylamine (243).
Interestingly, both the
[NiFe]- and the Hmd or [Fe]-hydrogenases also contain unusual non-protein ligands to
the active site metals, however these enzymes are evolutionarily and structurally
unrelated to each other and to the [FeFe]-hydrogenase. [FeFe]-hydrogenases exhibit
higher catalytic rates than [NiFe]-hydrogenases and are therefore being targeted for the
This section has been coauthored by David W. Mulder, Eric S. Boyd, Ranjana Sarma, Rachel K.
Lange, James A. Endrizzi, Joan B. Broderick, and John W. Peters. An abbreviated form is
accepted for publication in Nature (2010, DOI 10.1038/nature08993).
77
clean production of H2 and as an alternative biological catalyst to precious metalcontaining catalysts such as platinum in hydrogen fuel cell technology.
Figure 3.1. Ball and stick representation of the H-cluster from CpI (PDB code: 3C8Y). The Hcluster is bound to the protein (tube representation) by 4 cysteine ligands of the [4Fe-4S]subcluster which is further linked by a cysteine thiolate ligand to a 2Fe-subcluster with unique
non protein ligands including CO, CN-, and a dithiolate ligand. A water molecule is coordinated
to the distal Fe of the 2Fe-subcluster in the presumed active, oxidized state. Coloring scheme: Fe
(rust), S (orange), C (gray), N (blue), O (red), central atom of the dithiolate ligand (magenta).
Three gene products, HydE, HydF, and HydG, are required for the maturation of
the active site H-cluster present in the [FeFe]-hydrogenase structural protein HydA (140).
HydE and HydG are members of the radical-SAM superfamily of enzymes and HydF is a
GTPase (140, 176, 177). HydG has been shown to catalyze tyrosine cleavage in a
manner similar to that of the radical-SAM enzyme ThiH, leading to the hypothesis that
78
tyrosine may be a substrate for synthesis of the non-protein ligands of the H-cluster
(202). Evidence to date suggests that HydE, HydF, and HydG function to couple radical
SAM chemistry and nucleotide binding and hydrolysis to ligand synthesis, cluster
assembly and insertion, and finally [FeFe]-hydrogenase maturation (202, 210, 217, 244).
Although several plausible scenarios have been proposed for the generation of the carbon
monoxide, cyanide, and dithiolate ligands at the Fe site, including radical SAM mediated
sulfur insertion coupled to the decomposition or condensation of amino acids (217, 244,
245), the precise mechanism by which the various enzymes, scaffolds, and carriers
coordinate H-cluster maturation is unknown. Advancements in understanding how the
H-cluster is synthesized by HydE, HydF and HydG could contribute significantly to both
the genetic engineering of hydrogen producing microorganisms and the synthesis of
biomimetic hydrogen production catalysts.
HydF has been shown to transfer a cluster precursor to HydA in the final stage of
[FeFe]-hydrogenase maturation, suggesting that the 2Fe-subcluster of the H-cluster may
be assembled on HydF (210). In addition we have shown that the H-cluster maturation
machinery (the activities of HydE, HydG, and HydF) is directed at the synthesis of only
the 2Fe unit of the 6Fe cluster, and that the [4Fe-4S]-subcluster can be synthesized
independently on HydA expressed in a genetic background devoid of hydE, hydF, and
hydG (HydA∆EFG) (246). In support of this work, the recent EPR and IR spectroscopic
characterization of HydF shows HydF to possess a CO and CN- ligated Fe-containing
cofactor when expressed in the presence of HydE and HydG but in the absence of HydA
(212). Together, these observations indicate that the 2Fe-subcluster of the H-cluster must
79
be synthesized elsewhere from HydA∆EFG and later inserted into the structural protein to
achieve maturation. To better understand the synthesis and insertion of the individual
[4Fe-4S]-subcluster and 2Fe-subcluster components of the H-cluster during [FeFe]hydrogenase maturation, we determined the x-ray crystal structure of HydA∆EFG.
Results and Discussion
The structure of HydA∆EFG from the phototrophic alga Chlamydomonas reinhardtii
(Cr), heterologously expressed in Escherichia coli, was determined by molecular
replacement using the structure of Clostridium pasteurianum (CpI) (84) as a search
model and refined to 1.97 Å resolution (Figure 3.2A). We focused the study on the
[FeFe]-hydrogenase from Cr because our complementary biochemical and spectroscopic
analysis examining maturation were conducted using this enzyme (246), and it is a
particularly useful model since it does not contain accessory FeS cluster domains which
are present in bacterial HydA (7, 99).
In addition, HydA from Cr is of broad
biotechnological interest and no structural information yet exists on this enzyme.
Overall Structure
The overall structure of Cr HydA∆EFG (Figure 3.2A) is similar to that observed for
the active site domain of the previously characterized [FeFe]-hydrogenases purified from
the native hosts from the anaerobic soil bacterium CpI (84) (Figure 3.2C) and the sulfate
reducing bacterium Desulfovibrio desulfuricans DdH (104). The active site domain of
Cr HydA∆EFG consists of two twisted β-sheet regions surrounded by a number of α
helices.
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Figure 3.2. X-ray crystal structure of C. reinhardtii HydA∆EFG determined to 1.97 Å compared to HydA
from CpI. (A) Ribbon diagram of the overall HydA∆EFG structure with space-filling representation of the
associated [4Fe-4S] cluster. Two conserved loop regions implicated in undergoing major conformational
rearrangement are colored green. (B) HydA∆EFG active site area where a 2Fe-subcluster recipient cavity is
adjacent to the [4Fe-4S] cluster. (C) Ribbon diagram of the overall CpI structure in the same orientation as
HydA∆EFG with space-filling model of the intact H-cluster. The regions of CpI corresponding to the loop
regions of HydA∆EFG shown in (A) are colored green. (D) CpI active site region with ball-and-stick
representation of the H-cluster. (E) Root mean square deviation (RMSD) in the position of Cα atoms
between HydA∆EFG and the catalytic domain of CpI after superimposition of Cα atoms of the two respected
structures using the program LSQKAB (247). The x-axis is numbered according the sequence number of
HydA∆EFG from Cr. Below the x-axis, the amino acid sequences and secondary structures of HydA∆EFG and
CpI are aligned from ClustalW2 (248) and PROCHECK (249), respectively. Loops, α-helices, and β-sheets
are represented as lines, coils, and block arrows, respectively. The coloring scheme of the graph and
alignments corresponds to that of the protein secondary structure above. The atomic coloring in the figure
is the same as described for Figure 3.1.
81
The C-terminus region (a short, separate β-subunit in DdH) runs perpendicular between
the two β sheet regions, dividing the overall structure into two halves with the active site
binding domain occurring at the interface of these two β sheet regions.
Covalent
coordination of the H-cluster is provided by four cysteinyl sulfurs with two provided
from loops from each of the β-sheet regions.
Compared to the structure of HydA purified from its native host, the primary
difference in HydA expressed in a background devoid of the maturation machinery is the
absence of the 2Fe-subcluster (Figure 3.2B, D) leaving an open cavity adjacent to the
[4Fe-4S] cluster that is linked to the protein surface through an open cationically-charged
channel. The channel is formed by significant structural rearrangement in two loop
regions (loop1: residues 240-255 and loop 2: 279-286) in HydA∆EFG as revealed by the
superimposition of HydA∆EFG to CpI (Figure 3.2E). These structural differences indicate
that the two loop regions adopt an alternate conformation upon insertion of the 2Fesubcluster, effectively closing the channel and shielding the active site from surface
exposure. Interestingly, a sequence alignment of HydA from a diversity of organisms
indicates that loop region 1 is highly conserved and loop region 2 is partially conserved
in HydA from bacteria and several lower eukaryotes including Cr but not in eukaryotic
Nar1 homologues (Figure 3.3 and Figure 3.4). Nar1 homologues contain only the [4Fe4S] cluster and are present in the genomes of nearly all eukaryotes where they function in
cytosolic and nuclear Fe-S cluster maturation (82).
82
Figure 3.3. Unrooted phylogram of representative HydA and HydA homologues. Unrooted radial
phylogram is inset. ME denotes mitochondriate eukaryotes, AE denotes amitochondriate
eukaryotes, and no designation denotes bacterial sequences. The designations and numerals
following designations correspond in both figures. Closed circles denote 90-100% bootstrap
support (BS); gray circles denote 80-89% BS; open circles denote 70-79% BS support; no symbol
indicates <69% BS.
83
Figure 3.4. Phylogenetic reconstruction of HydA and HydA homologs. The sequence names for
terminals correspond with those in Figure 3.3. Terminals have been right right-justified and
aligned with the corresponding loop regions to illustrate conservation in taxa that contain
homologs of hydE, hydF, and hydG (black lineages) and that lack homologues of hydE, hydF, and
hydG (red lineages). Closed circles denote 90-100% bootstrap support (BS); gray circles denote
80-89% BS; open circles denote 70-79% BS support; no symbol indicates <69% BS. The asterisk
denotes a region of the alignment that was deleted to conserve space. Residues that are conserved
in 20% of the sequences are indicated in color.
84
Active Site
The active site of HydA∆EFG, when compared to that of CpI, exists with an open
cavity at the position of the CpI 2Fe-subcluster which is linked to the protein surface via
the aforementioned channel which presumably facilitates the insertion of the 2Fesubcluster (Figure 3.5).
Figure 3.5. Active site comparison between HydA∆EFG and CpI. (A) Stereoview of the [4Fe-4S]
cluster active site environment in HydA∆EFG. Residues are labeled according to amino acid
abbreviations and sequence number. (B) Stereoview of the H-cluster active site environment in
CpI, shown in the identical orientation as HydA∆EFG. Corresponding residues in HydA∆EFG of CpI
residues 417 and 418 have migrated out of the region. The atomic coloring in the figure is the
same as described for Figure 3.1.
85
Two residues in the active site region, Phe250 and Gly251 (corresponding to CpI residues
Phe417 and Gly418) show major structural rearrangement in HydA∆EFG and belong to the
conserved loop region 1 mentioned above.
The structural rearrangement of these
residues in HydA∆EFG creates a connection between the open cavity and channel,
consequently creating a clear pathway for 2Fe-subcluster insertion. Other residues in the
immediate vicinity of the active site show little deviation from the corresponding residue
position in CpI. Even Cys381, the only protein ligand for the 2Fe-subcluster and site for
covalent attachment to the [4Fe-4S] cluster, resides at the same position as Cys503 in
CpI. The effect of the absence of the 2Fe-subcluster in the active site can be seen on
residue Cys129. While Cys129 is located at the same position of the corresponding
residue in CpI (Cys299), it can adopt different rotamers. In CpI, it has been pointed out
that the cysteine residue hydrogen bonds with the bridging dithiolate ligand of the 2Fesubcluster (84).
Because this interaction is not possible in HydA∆EFG, Cys129 has
increased conformational freedom and its side chain sulfur atom is observed in three
different states.
Given these observations, the active site environment of HydA∆EFG
exists as a [4Fe-4S] cluster adjacent to a well-defined binding cavity in a state ready for
insertion of the 2Fe-subcluster.
Difference Fourier Analysis. Although it is clear an intact 2Fe-subcluster is not
present in the active site, analysis of Fo – Fc electron density maps reveals some residual
density adjacent to the [4Fe-4S] cluster where the 2Fe-subcluster would be expected to
reside (Figure 3.6A).
86
Figure 3.6. (A) Stereoview of the 2Fo – Fc map (purple) calculated for the [4Fe-4S] cluster and
coordinating cysteine residues. The map is contoured at 1.5σ. Also, the stereoview of the Fo – Fc
map (cyan) is shown for the active site area and contoured at 3.5σ. (B) Stereoview of the
anomalous difference Fourier map (green). The map is contoured at 4.5σ.
To verify the absence of Fe in the residual density, we performed Fe edge
anomalous difference Fourier analysis that confirmed that the only anomalous scatterers
at the Fe absorption edge were within the [4Fe-4S] cluster (Figure 3.6A). A single
acetate molecule and chloride ion, both of which were present in the crystallization
87
buffer, could be modeled reasonably well where the unknown density resided. Thus, the
active site clearly exists as a [4Fe-4S] cluster adjoined to an open binding cavity for
insertion of the 2Fe-subcluster.
Channel Formation and Cluster Insertion
The overall surface representations of HydA∆EFG reveal a positively charged
channel leading to the active site area (Figure 3.7A-C), a feature that is absent in the
intact and active hydrogenase from CpI presumably due to conformational change
following insertion of the 2Fe-subcluster (Figure 3.7D). The channel for 2Fe-subcluster
insertion is solvent accessible, 8 to 15 Å in width, and ~25 Å deep with positively
charged residues (Arg275, Lys288, and Lys409) lining the channel entrance. Lys188,
which equivalent residue (Lys 358) hydrogen bonds to a cyanide ligand of the H-cluster
in the intact hydrogenase (84), lies at the end of the channel at the active site cavity and
may have a role in orienting the 2Fe-subcluster during insertion.
Importantly, the
bridging cysteine thiolate ligand (Cys381) between the [4Fe-4S] cluster and 2Fesubcluster is located ~19 Å into the channel. The Cys381 thiolate sulfur side chain is
exposed on the surface of the channel, providing the site for covalent attachment of the
2Fe-subcluster following insertion which is likely entropically-driven as the channel is
solvent accessible and hydrophilic.
88
Figure 3.7. Channels for insertion into hydrogenase and nitrogenase during complex Fe-S-cluster
assembly. (A) Surface representation of HydA∆EFG (grey) superimposed on native CpI (green ribbon
representation). (B) HydA∆EFG channel leading to the active site 2Fe-subcluster recipient cavity. (C,
D) Electrostatic surface representations of HydA∆EFG and the catalytic domain of CpI, respectively,
calculated with the PyMol plugin APBS. Red, (−10kBT/e); blue, (10kBT/e); kB, Boltzmann’s constant;
e, elementary charge. (E) Surface representation of FeMo-co deficient form of NifDK (Av1∆nifB) (grey,
PDB code: 1L5H) superimposed on native nitrogenase NifDK protein (Av1) (PDB code: 3MIN)
(green ribbon representation). (F) Av1∆nifB NifDK channel leading to the active-site FeMo-cofactor
recipient cavity. (G, H) Electrostatic surface representations of NifDK Av1∆nifB and NifDK,
respectively, calculated with identical APBS parameters as C and D. The atomic coloring scheme is
the same as described for Figure 3.1.
89
Implications for H-cluster Biosynthesis
The structure of HydA∆EFG presented here provides strong support for a stepwise
mechanism for H-cluster biosynthesis in which the [4Fe-4S]-subcluster insertion
precedes 2Fe-subcluster insertion. Previously we showed that the [4Fe-4S]-subcluster
synthesis and 2Fe-subcluster synthesis occur independently (246). We now show in the
current work that the [4Fe-4S]-subcluster makes up part of a binding cavity that is linked
to a channel leading to the surface of the protein. In the absence of the [4Fe-4S]subcluster, the binding cavity would not exist. Furthermore, since the [4Fe-4S]-subcluster
is at the base of the channel it would be impossible to insert the [4Fe-4S] cluster
subsequent to insertion of the 2Fe-subcluster since the pathway would be effectively
blocked. In addition, if the process is conserved through all organisms, alternative paths
for insertion of the [4Fe-4S] cluster would be precluded in the majority of organisms
which express [FeFe]-hydrogenases with accessory Fe-S cluster domains adjacent to the
H-cluster [4Fe-4S]-subcluster. Therefore, the structural observations from HydA∆EFG
indicate that the [4Fe-4S]-subcluster would need to be synthesized and inserted first by
Fe-S cluster generalized host cell machinery (143, 147) followed by the synthesis and
insertion of the 2Fe-subcluster by Hyd maturation (202, 210, 217, 246).
Complex Metallocluster Assembly
The implied structural rearrangement that occurs during 2Fe-subcluster insertion
reveals an interesting parallel to the maturation of the nitrogenase complex.
Molybdenum nitrogenase is comprised of an iron protein (NifH) and a MoFe protein
(NifDK) with the latter containing P-clusters (8Fe–7S) and FeMo-cofactors (Mo–7Fe–
90
9S–X–homocitrate, where X = C, O, N) (250, 251). Structural determination of a form of
Azotobacter vinelandii (Av1) NifDK deficient in FeMo-co (Av1∆nifB) reveals major
structural rearrangement in the domain involved in FeMo-co binding, resulting in channel
formation to the FeMo-co site in Av1∆nifB (Figure 3.7E-G) (219). Superimposition of
Av1 NifDK (containing FeMo-co) (250, 251) and Av1∆nifB NifDK (lacking FeMo-co)
(219) indicates that the FeMo-co site is at the end of a cationic channel (Figure 3.7E-G)
that is absent from the structure of Av1 NifDK (Figure 3.7H).
The structural
rearrangements resulting in the formation of a cationic channel in both HydA and NifDK
indicate that the process for complex Fe-S cluster insertion into apo-proteins may be
conserved and that the evolution of functional HydA may have occurred stepwise, a
feature which is consistent with the evolution of nitrogenise (252) and possibly Fe-S
enzymes in general. Moreover, these features suggest that these enzymes have evolved
more complex active site clusters, perhaps to direct substrate specificity or to enhance
catalytic activity.
Evolutionary Discussion
We investigated the evolutionary history of Hyd by screening all available genome
sequences for the presence of homologs of HydA, HydE, HydF, and HydG. With few
exceptions, bacterial genomes that encoded for HydA also encoded for HydE, HydF, and
HydG (see Appendix A). In contrast, of the 78 eukaryotic genomes which encoded for
HydA homologs, only Cr and Trichomonas vaginalis (Tv) contained clear homologues of
HydE, HydF, and HydG protein-encoding genes (see Appendix A) (7). The degree of
conservation in HydA loop regions 1 and 2 that are implicated in insertion of the 2Fe-
91
subcluster in organisms whose genome encoded for HydE, HydF, and HydG is high
(Figure 3.3). Conservation in loop regions was significantly lower in eukaryotic Nar1
homologues whose genomes did not encode for HydE, HydF, and HydG, when compared
to genomes that encoded for HydA, HydE, HydF, and HydG. These observations suggest
the presence of a common selective pressure for conserving loop regions and for
maintaining maturation genes and indicate that these gene products have co-evolved.
Maximum likelihood and Bayesian inference indicates that HydA and Nar1
homologs branch distinctly, forming two separate stem lineages (Figure 3.3). Although
the overall support for the HydA/Nar1 phylograms is not high, support in key nodes is
observed.
One stem lineage contains several well-supported Nar1 lineages from
eukaryotic genomes that lack hydE, hydF, and hydG. The other stem contains several
well-supported lineages of HydA from bacteria and several eukaryotic genomes that
generally contain hydE, hydF, and hydG, including HydA from both Cr and Tv. HydA
from Cr and Tv are nested with high statistical support among bacterial sequences,
suggesting that HydA in these organisms derive from lateral gene transfer (LGT) from a
bacterium and/or endosymbiosis of a bacterium (7).
It has been previously suggested the Nar1 was acquired in eukarya via
endosymbiosis of or LGT with a bacterium (7). The lack of conservation in loop regions
and the lack of hydE, hydF, and hydG in eukaryotic taxa with Nar1 suggest that the
acquisition of the Nar1 ancestor in Eukarya occurred prior to the recruitment of hydE,
hydF, and hydG and the ability to generate the 2Fe-subcluster.
This hypothesis is
bolstered by spectroscopic and structural studies of Nar1, which indicate that these
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proteins lack the 2Fe-subcluster (82, 83, 253).
Together, this set of observations
indicates that the active site of the HydA/Nar1 ancestor would have only contained the
[4Fe-4S] cluster and not the 2Fe-subcluster and thus most likely resembled that of extant
Nar1 sequences. While the function of Nar1 proteins is not understood, it is clear that
they do not catalyze hydrogen activation (83).
Conclusions
The structure of HydA∆EFG presented here reveals the stepwise assembly of the Hcluster in [FeFe]-hydrogenases and the structural pathway for 2Fe-subcluster insertion in
the final step of [FeFe]-hydrogenase maturation. By providing significant insights into Hcluster biosynthesis, the results provide a foundation for enhancing mechanisms of
biological hydrogen production in genetically engineered hydrogenases and for the
synthesis of biomimetic catalysts, and thus ultimately for developing hydrogen as a
renewable fuel. In addition, this structure reveals several novel and unifying themes for
Fe-S enzyme biosynthesis and evolution. Both the FeMo-cos of NifDK and the 2Fesubcluster of the H-cluster of HydA are synthesized on specialized scaffold proteins and
are inserted into the respective enzyme through a conserved cationically charged channel,
resulting in catalytically active proteins (142, 202, 210, 217, 242). Recent evolutionary
studies of the proteins involved in synthesizing the FeMo-cofactor of nitrogenase indicate
that, like the 2Fe-subcluster of HydA, the FeMo-cofactor in the active site of NifDK was
not present in the last universal common ancestor, indicating that it is a more recent
innovation (252, 254). Thus, the stepwise evolution of complex Fe-S metalloenzymes
93
may be pervasive in biology, resulting in protein complexes with improved specificity
and/or enhanced catalytic efficiency.
Experimental Procedures
Structure Determination and Refinement
HydA∆EFG from Cr was heterologously expressed in Escherichia coli and purified
under strict anaerobic conditions as described previously (246). Prior to crystallization,
purified HydA∆EFG was diluted over a SephadexTM G-25 column (GE Healthcare) in 50
mM Tris buffer (pH 7.8) with 300 mM NaCl, 20% glycerol, and 1 mM dithionite to a
final concentration of 28 mg/mL. Crystals were obtained at room temperature (298 K)
using the anaerobic microcapillary batch diffusion method (255) in a nitrogen atmosphere
glovebox (UniLAB, MBRAUN, NH) using 25.5% polyethylene glycol 8000 as
precipitate and 0.085 M sodium cacodylate (pH 6.5) with 0.17 M sodium acetate
trihydrate. The initial data were collected from a single flash-cooled crystal on beamline
9-1 at the Stanford Synchrotron Radiation laboratory (SSRL), with a continuous flow of
liquid nitrogen at 100K, and a single wavelength data set (λ = 0.954) was collected up to
resolution 1.97 Å (Table 3.1). An additional single anomalous wavelength data set at the
Fe-edge (λ=1.74) were collected at a later time on the identical crystal at beamline 9-2 at
SSRL, up to resolution 3.00 Å (Table 3.1). The 1.97 Å data were processed and scaled
using DENZO and SCALPACK of the HKL2000 software package (256) (version 1.98.7.
The structure was solved by molecular replacement using AutoMR of the CCP4
suite of programs (version 6.0) (257) with CpI (Protein Data Bank entry 1FEH (84)) as a
94
search model.
The solution was subjected to rigid body refinement in REFMAC5
(version 5.2.0019) (258) and further improved with ARP/wARP (version 7.0) (259). The
maps obtained were solvent flattened and histogram averaged using RESOLVE (260).
Model building was subsequently completed manually using COOT (version 0.3.3)
(261) with following refinement (REFMAC5) using NCS and B-factor restraints. This
resulted in the determined structure. The final model was refined on the 1.97 Å data to
Rfactor = 17.2% and Rfree = 21.6% (Table 3.1). The positions of residues 1 to 24, 116 to
120, 201 to 203, 312 to 322, and 451 to 457 were not able to be determined due to
disorder. Also, the side chain sulfur atom of Cys129 could be modeled in three different
states. In the final model, 98.4% of all residues were in favoured regions and 100% of all
residues were in allowed regions of the Ramachandran plot (calculated with
MOLPROBITY (version 3.17).
Because of undetermined density adjacent to the [4Fe-4S] cluster site, an
additional single anomalous wavelength data set at the Fe-edge (λ=1.74), at resolution
3.00 Å, was used to confirm the Fe positions in the determined structure. The data were
processed as describe above and Fe positions determined using SOLVE (version 2.13)
(262). The native and anomalous reflection data was merged using CAD in CCP4 and a
difference Fourier synthesis between the native model phases and anomalous scattering
was calculated using FFT in CCP4 (263).
95
Table 3.1. Data collection and refinement statistics
HydA∆EFG from Cra
Data collection
Space group
Cell dimensions
a, b, c (Å)
α, β, γ (°)
Wavelength
Resolution (Å)
Rsym (%)
I/σI
Completeness (%)
Redundancy
P32
P32
70.9, 70.9, 155.4
90.0, 90.0, 120.0
0.95369
50.00 - 1.97 (2.04-1.97)b
8.0 (40.9)
14.0 (2.4)
97.9 (97.2)
4.4 (4.2)
70.9, 70.9, 155.4
90.0, 90.0, 120.0
1.73950
50.00 - 3.00 (3.05-3.00)
6.6 (14.1)
29.2 (7.8)
81.7 (28.9)
6.2 (2.8)
Refinement
Resolution (Å)
20.00 - 1.97
No. reflections
57,708
Rwork / Rfree
17.0 / 21.6
No. Atoms
Protein
6204
2 (16)
[Fe4-S4] (No. Atoms)
Acetate (No. Atoms) / Cl
2 (8) / 2
Water
775
B-factors
25.8
Protein
21.1
[Fe4-S4]
39.6
Acetate / Cl
35.6
Water
R.m.s deviations
Bond lengths (Å)
0.006
Bond angles (º)
0.924
a
Native and anomalous (Fe edge) data sets were collected from the same crystal . The atomic
coordinates and structure factors have been deposited in the protein data bank (PDB Code:
3LX4) . bHighest resolution shell is shown in parenthesis.
Phylogenetic Analysis
BLASTp was used to compile all HydA-, HydE-, HydF-, and HydG-deduced and
HydA-homolog -deduced amino acid sequences from genomic sequences using the DOEIMG and the NCBI Genome Blast servers (See Appendix A). A total of 435 HydA and
HydA homolog sequences were compiled and these were aligned using ClustalX (version
2.0.8) with the Gonnet 250 protein matrix and default gap extension and opening
96
penalties (248).
The alignment was scrutinized and manually aligned using known
catalytic residues (7). A single maximum likelihood phylogenetic tree was computed
using PhyML (version 3.0) (264) employing the LG substitution matrix (265) (Figure
3.2). This tree was used to empirically select 90 HydA sequences that represented the
primary lineages. Sequences were trimmed to contain only the H-cluster domain present
in Cr (7). The trimmed sequences were realigned using ClustalX as described above.
ProtTest (version 2.0) (266) was used to select the WAG+I+G as the best-fit
protein evolutionary model. The phylogeny of each locus was evaluated using PhyML
with the WAG evolutionary model with a proportion of invariable sites and gammadistributed rate variation (I+G). A composite phylogram was constructed from 100
bootstrap replicate phylograms and the tree was projected using FigTree (version 1.2.2)
(http://tree.bio.ed.ac.uk/software/figtree/) (Figure 3.3). Similarly, the phylogeny of each
locus was evaluated using MrBayes (version 3.1.2) (267, 268). A composite phylogram
was constructed using the WAG evolutionary model with gamma-distributed rate
variation with a proportion of invariable sites (I+G). Tree topologies were sampled at
likelihood stationary during 2 separate runs every 500 generations by MrBayes over
3.7e6 generations with a parameter of burnin=2.0e6 (standard deviation of split trees was
<0.07).
97
CHAPTER 4
CONCLUDING REMARKS
[FeFe]-hydrogenase maturation and synthesis of its active site, the H-cluster, is a
complex process, requiring the activity of three specialized maturation enzymes, HydE,
HydF, and HydG in addition to the structural enzyme, HydA, to facilitate catalysis of
reversible oxidation of molecular H2 by the reaction H2 2H+ + 2e-. Overall, this work
provides key information for elucidating the biosynthetic pathway of the H-cluster during
[FeFe]-hydrogenase maturation and specifically makes clear the details of the last step of
[FeFe]-hydrogenase maturation in which the 2Fe-subcluster of the H-cluster is inserted
into the structural [FeFe]-hydrogenase to effect activation. The results presented in
Chapter 2 answer several initial questions of the biosynthetic assembly process and
establish which cofactor components are targeted by hydrogenase and general housekeeping FeS cluster maturation machinery.
By demonstrating through various
spectroscopic characterization techniques that HydA∆EFG contains a [4Fe-4S] cluster and
biochemically that the presence of the [4Fe-4S] cluster is required for [FeFe]hydrogenase activation, the conclusion can be made that the [4Fe-4S]-subcluster and 2Fesubcluster of the H-cluster are synthesized independently by different maturation
machineries. Significantly, it follows that that the 2Fe-subcluster, and not the entire Hcluster or simply just the ligands of the H-cluster, is the target of maturation enzymes
HydE, HydF, and HydG. The [4Fe-4S]-subcluster, conversely, can be synthesized
independently by general house-keeping FeS cluster maturation machinery.
98
The results presented in Chapter 3 not only definitively confirm this but also
further advance the understanding of the final step of [FeFe]-hydrogenase maturation by
revealing the precise stepwise synthesis of the H-cluster and structural pathway for
insertion of the 2Fe-subcluster in the final step of [FeFe]-hydrogenase activation. The xray crystal structure of the key intermediate HydA∆EFG determined to 1.97 Å resolution
from the green algae Chlamydomonas reinhardtii reveals the presence of the [4Fe-4S]subcluster linked to a positively-lined channel accessible to the protein surface. This
channel is the mechanistic pathway for insertion of the 2Fe-subcluster, making available
essential interactions to incorporate the cluster to complete biological assembly of the Hcluster. Following incorporation of the 2Fe subcluster the channel collapses as a result of
conformational changes in two conserved loop regions, giving way to active [FeFe]hydrogenase. The presence of the [4Fe-4S]-subcluster linked to a structural channel to the
surface of the protein also makes it clear that the 2Fe-subcluster must be synthesized by
HydE, HydF, and HydG maturation enzymes prior to the synthesis of the [4Fe-4S]subcluster by general house-keeping FeS cluster maturation machinery
Both the results of Chapter 2 and Chapter 3 support the early hypothesis for
[FeFe]-hydrogenase biosynthesis by Peters et al. Furthermore, the results, in light of ongoing experiments probing the function of the maturation enzymes themselves, begin to
make the picture for the biosynthetic pathway of the H-cluster more clear.
From
experiments aimed at the maturation enzymes, it is evident that HydE, HydF, and HydG
serve to modify a basic [2Fe-2S] cluster with unique CO and CN- ligands and also a
dithiolate ligand. Beginning the biosynthetic process, radical SAM enzyme HydE likely
99
modifies a basic [2Fe-2S] cluster with a dithiolate ligand, using HydF as a scaffold. This
is presumably followed by the radical SAM enzyme HydG modifying the 2Fe cluster
with CO and CN- ligands by means of unprecendented radical Sam chemistry, also using
HydF as a scaffold. Once the synthesis of the 2Fe-subcluster is complete, the results
presented in Chapter 2 and Chapter 3 indicate that it is inserted into the structural [FeFe]hydrogenase by means of a positively lined channel ending at a binding cavity linked to
the [4Fe-4S]-subcluster. Interesting, this picture for the biosynthetic pathway of the Hcluster as described here follows the general scheme for the synthesis of other FeS
clusters and involves modification of a simple FeS cluster on a scaffold enzyme and
subsequent insertion of cluster to the structural enzyme, completing maturation.
The work presented herein is not only significant toward deducing the
biosynthetic pathway for H-cluster biosynthesis and [FeFe]-hydrogenase maturation but
also has several broad implications on complex metallocluster assembly and
metalloenzyme evolution, making this work of general interest.
For complex
metallocluster assembly, the close parallels to nitrogenase maturation revealed by the xray crystal structure of HydA∆EFG presented in Chapter 3 suggest that these conclusions
may be thematic for complex metalloenzymes in general. Like the [4Fe-4S]-subcluster
of the H-cluster, the P-cluster of nitrogenase is synthesized independently of the FeMoco. Similarly, as the 2Fe-subcluster of the H-cluster is inserted in the structural [FeFe]hydrogenase enzyme by means of a positively charged channel lined to the active site, the
FeMo-co also is inserted into the structural nitrogenase enzyme by means of a positively
charged channel lined to the active site.
This connection and similar structural
100
rearrangements between two phylogentically distinct proteins catalyzing two different
reactions is significant and strongly suggests that the process for insertion of complex
metallocofactors into apo-enzymes in general is a conserved process.
For metalloenzyme evolution, these results again establish an interesting parallel
to nitrogenase and present an additional theme for metalloenzymes in general. The
evolutionary studies presented here on [FeFe]-hydrogenase, made possible only because
of the determination of the x-ray crystal structure of HydA∆EFG, indicate that HydA
emerged within bacteria most likely from a Nar1-like ancestor lacking the 2Fe-subcluster,
followed by acquisition in several lower order eukaryotes. Likewise, for nitrogenase,
recent studies involving the nitrogenase maturation proteins indicate that the FeMocofactor in the active site of NifDK may also be a relatively recent evolutionary feature.
Together, these findings suggest the possibility that metalloenzymes in general may
evolve complex active sites to direct substrate specificity or to enhance catalytic activity.
Interestingly, the discoveries of this work demonstrate that the approach of utilizing
structural biology to set-up evolutionary studies is a powerful method for deciphering
how complex metallocofactors may have developed and evolved over time and provide
the groundwork for similar studies.
101
REFERENCES CITED
102
1.
Adams, M. W. W. (1990) The structure and mechanism of iron-hydrogenases,
Biochim Biophys Acta 1020, 115-145.
2.
Shima, S., and Thauer, R. K. (2007) A third type of hydrogenase catalyzing H2
activation, Chem Rec 7, 37-46.
3.
Boichenko, V. A., Greenbaum, E., and Seibert, M. (2004) Hydrogen production
by photosynthetic microorganisms, In Photoconversion of solar energy,
molecular to global photosynthesis (Archer, M. D., and Barber, J., Eds.), pp 397452, Imperial College Press, London.
4.
Vignais, P. M., Billoud, B., and Meyer, J. (2001) Classification and phylogeny of
hydrogenases, FEMS Microbiol Rev 25, 455-501.
5.
Vignais, P. M., and Colbeau, A. (2004) Molecular biology of microbial
hydrogenases, Curr Issues Mol Biol 6, 159-188.
6.
Ludwig, M., Schulz-Friedrich, R., and Appel, J. (2006) Occurrence of
hydrogenases in cyanobacteria and anoxygenic photosynthetic bacteria:
implications for the phylogenetic origin of cyanobacterial and algal hydrogenases,
J Mol Evol 63, 758-768.
7.
Meyer, J. (2007) [FeFe] hydrogenases and their evolution: a genomic perspective,
Cell Mol Life Sci 64, 1063-1084.
8.
Vignais, P. M., and Billoud, B. (2007) Occurrence, classification, and biological
function of hydrogenases: an overview, Chem Rev 107, 4206-4272.
9.
Armstrong, F. A., and Fontecilla-Camps, J. C. (2008) Biochemistry. A natural
choice for activating hydrogen, Science 321, 498-499.
10.
Kubas, G. J. (2007) Fundamentals of H2 binding and reactivity on transition
metals underlying hydrogenase function and H2 production and storage, Chem
Rev 107, 4152-4205.
11.
Ghirardi, M. L., King, P. W., Posewitz, M. C., Maness, P. C., Fedorov, A., Kim,
K., Cohen, J., Schulten, K., and Seibert, M. (2005) Approaches to developing
biological H(2)-photoproducing organisms and processes, Biochem Soc Trans 33,
70-72.
12.
Ghirardi, M. L., Zhang, L., Lee, J. W., Flynn, T., Seibert, M., Greenbaum, E., and
Melis, A. (2000) Microalgae: a green source of renewable H2, Trends Biotechnol
18, 506-511.
103
13.
Happe, T., Hemschemeier, A., Winkler, M., and Kaminski, A. (2002)
Hydrogenases in green algae: do they save the algae's life and solve our energy
problems?, Trends Plant Sci 7, 246-250.
14.
Kruse, O., Rupprecht, J., Bader, K. P., Thomas-Hall, S., Schenk, P. M., Finazzi,
G., and Hankamer, B. (2005) Improved photobiological H2 production in
engineered green algal cells, J Biol Chem 280, 34170-34177.
15.
Melis, A., and Happe, T. (2001) Hydrogen production. Green algae as a source of
energy, Plant Physiol 127, 740-748.
16.
Melis, A., and Happe, T. (2004) Trails of green alga hydrogen research - from
hans gaffron to new frontiers, Photosynth Res 80, 401-409.
17.
Melis, A., Zhang, L., Forestier, M., Ghirardi, M. L., and Seibert, M. (2000)
Sustained photobiological hydrogen gas production upon reversible inactivation
of oxygen evolution in the green alga Chlamydomonas reinhardtii, Plant Physiol
122, 127-136.
18.
Schutz, K., Happe, T., Troshina, O., Lindblad, P., Leitao, E., Oliveira, P., and
Tamagnini, P. (2004) Cyanobacterial H(2) production -- a comparative analysis,
Planta 218, 350-359.
19.
Tamagnini, P., Costa, J. L., Almeida, L., Oliveira, M. J., Salema, R., and
Lindblad, P. (2000) Diversity of cyanobacterial hydrogenases, a molecular
approach, Curr Microbiol 40, 356-361.
20.
Hallenbeck, P. C. (2009) Fermentative hydrogen production: Principles, progress,
and prognosis, Int J Hydrogen Energy 34, 7379-7389.
21.
Hawkes, F., Dinsdale, R., Hawkes, D., and Hussy, I. (2002) Sustainable
fermentative hydrogen production: challenges for process optimisation, Int J
Hydrogen Energy 27, 1339-1347.
22.
Holm, R. H., Kennepohl, P., and Solomon, E. I. (1996) Structural and functional
aspects of metal sites in biology, Chem Rev 96, 2239-2314.
23.
Wachtershauser, G. (1992) Groundworks for an evolutionary biochemistry: the
iron-sulphur world, Prog Biophys Mol Biol 58, 85-201.
24.
Cody, G., Boctor, N., Filley, T., Hazen, R., Scott, J., Sharma, A., and Yoder Jr, H.
2000, Primordial Carbonylated Iron-Sulfur Compounds and the Synthesis of
Pyruvate, Science 289, 1337–1340.
104
25.
Drennan, C. L., and Peters, J. W. (2003) Surprising cofactors in metalloenzymes,
Curr Opin Struct Biol 13, 220-226.
26.
Wachtershauser, G. (1988) An All-Purine Precursor of Nucleic-Acids, Proc Natl
Acad Sci USA 85, 1134-1135.
27.
Wachtershauser, G. (1990) The Case for the Chemoautotrophic Origin of Life in
an Iron-Sulfur World, Origins of Life and Evolution of the Biosphere 20, 173-176.
28.
Wachtershauser, G. (1993) The Cradle Chemistry of Life - on the Origin of
Natural-Products in a Pyrite-Pulled Chemo-Autotrophic Origin of Life, Pure and
Appl Chem 65, 1343-1348.
29.
Wachtershauser, G. (1998) Origin of life in an iron-sulfur world, In The
molecular origins of life: assembling pieces of the puzzle (Brack, A., Ed.), pp 206218, Cambridge University Press.
30.
Tian, F., Toon, O. B., Pavlov, A. A., and De Sterck, H. (2005) A hydrogen-rich
early Earth atmosphere, Science 308, 1014-1017.
31.
Martin, W., and Russell, M. (2003) On the origins of cells: a hypothesis for the
evolutionary transitions from abiotic geochemistry to chemoautotrophic
prokaryotes, and from prokaryotes to nucleated cells, Philosophical Transactions
of the Royal Society B: Biological Sciences 358, 59.
32.
Schoonen, M., Smirnov, A., and Cohn, C. (2004) A perspective on the role of
minerals in prebiotic synthesis, AMBIO: A Journal of the Human Environment 33,
539-551.
33.
Stephenson, M., and Stickland, L. H. (1931) Hydrogenase: a bacterial enzyme
activating molecular hydrogen: The properties of the enzyme., Biochem J 25, 205214.
34.
Posewitz, M. C., Mulder, D. W., and Peters, J. W. (2008) New frontiers in
hydrogenase structure and biosynthesis, Curr Chem Biol 2, 178-199.
35.
Peters, J. W. (1999) Structure and mechanism of iron-only hydrogenases, Curr
Opin Struct Biol 9, 670-676.
36.
Vignais, P. (2008) Hydrogenases and H ()-reduction in primary energy
conservation, Results Probl Cell Differ 45, 223-252.
105
37.
Spear, J. R., Walker, J. J., McCollom, T. M., and Pace, N. R. (2005) Hydrogen
and bioenergetics in the Yellowstone geothermal ecosystem, Proc Natl Acad Sci
USA 102, 2555-2560.
38.
Hedderich, R., and Forzi, L. (2005) Energy-converting [NiFe] hydrogenases:
more than just H2 activation, J Mol Microbiol Biotechnol 10, 92-104.
39.
He, S. H., Teixeira, M., LeGall, J., Patil, D. S., Moura, I., Moura, J. J.,
DerVartanian, D. V., Huynh, B. H., and Peck, H. D., Jr. (1989) EPR studies with
77
Se-enriched (NiFeSe) hydrogenase of Desulfovibrio baculatus. Evidence for a
selenium ligand to the active site nickel, J Biol Chem 264, 2678-2682.
40.
Sorgenfrei, O., Klein, A., and Albracht, S. P. (1993) Influence of illumination on
the electronic interaction between 77Se and nickel in active F420-non-reducing
hydrogenase from Methanococcus voltae, FEBS Lett 332, 291-297.
41.
Wu, L. F., and Mandrand, M. A. (1993) Microbial hydrogenases: primary
structure, classification, signatures and phylogeny, FEMS Microbiol Rev 10, 243269.
42.
Sargent, F., Bogsch, E. G., Stanley, N. R., Wexler, M., Robinson, C., Berks, B.
C., and Palmer, T. (1998) Overlapping functions of components of a bacterial
Sec-independent protein export pathway, Embo J 17, 3640-3650.
43.
Dubini, A., and Sargent, F. (2003) Assembly of Tat-dependent [NiFe]
hydrogenases: identification of precursor-binding accessory proteins, FEBS Lett
549, 141-146.
44.
Kleihues, L., Lenz, O., Bernhard, M., Buhrke, T., and Friedrich, B. (2000) The H2
sensor of Ralstonia eutropha is a member of the subclass of regulatory [NiFe]
hydrogenases, J Bacteriol 182, 2716-2724.
45.
Vignais, P. M., Dimon, B., Zorin, N. A., Tomiyama, M., and Colbeau, A. (2000)
Characterization of the hydrogen-deuterium exchange activities of the energytransducing HupSL hydrogenase and H-2-signaling HupUV hydrogenase in
Rhodobacter capsulatus, J Bacteriol 182, 5997-6004.
46.
Bohm, R., Sauter, M., and Bock, A. (1990) Nucleotide sequence and expression
of an operon in Escherichia coli coding for formate hydrogenlyase components,
Mol Microbiol 4, 231-243.
47.
Volbeda, A., Charon, M. H., Piras, C., Hatchikian, E. C., Frey, M., and FontecillaCamps, J. C. (1995) Crystal structure of the nickel-iron hydrogenase from
Desulfovibrio gigas, Nature 373, 580-587.
106
48.
Volbeda, A., Garcin, E., Piras, C., de Lacey, A. L., Fernandez, V. M., Hatchikian,
E. C., Frey, M., and Fontecilla-Camps, J. C. (1996) Structure of the [NiFe]
hydrogenase active site: Evidence for biologically uncommon Fe ligands, J Am
Chem Soc 118, 12989-12996.
49.
Higuchi, Y., Yagi, T., and Yasuoka, N. (1997) Unusual ligand structure in Ni-Fe
active center and an additional Mg site in hydrogenase revealed by high resolution
X-ray structure analysis, Structure 5, 1671-1680.
50.
Higuchi, Y., Ogata, H., Miki, K., Yasuoka, N., and Yagi, T. (1999) Removal of
the bridging ligand atom at the Ni-Fe active site of [NiFe] hydrogenase upon
reduction with H2, as revealed by X-ray structure analysis at 1.4 A resolution,
Structure 7, 549-556.
51.
Montet, Y., Amara, P., Volbeda, A., Vernede, X., Hatchikian, E. C., Field, M. J.,
Frey, M., and Fontecilla-Camps, J. C. (1997) Gas access to the active site of NiFe hydrogenases probed by X-ray crystallography and molecular dynamics, Nat
Struct Biol 4, 523-526.
52.
Volbeda, A., Montet, Y., Vernede, X., Hatchikian, E., and Fontecilla-Camps, J.
(2002) High-resolution crystallographic analysis of Desulfovibrio fructiosovorans
[NiFe] hydrogenase, Int J Hydrogen Energ 27, 1449-1461.
53.
Volbeda, A., Martin, L., Cavazza, C., Matho, M., Faber, B. W., Roseboom, W.,
Albracht, S. P., Garcin, E., Rousset, M., and Fontecilla-Camps, J. C. (2005)
Structural differences between the ready and unready oxidized states of [NiFe]
hydrogenases, J Biol Inorg Chem 10, 239-249.
54.
Matias, P. M., Soares, C. M., Saraiva, L. M., Coelho, R., Morais, J., Le Gall, J.,
and Carrondo, M. A. (2001) [NiFe] hydrogenase from Desulfovibrio
desulfuricans ATCC 27774: gene sequencing, three-dimensional structure
determination and refinement at 1.8 A and modelling studies of its interaction
with the tetrahaem cytochrome c3, J Biol Inorg Chem 6, 63-81.
55.
Garcin, E., Vernede, X., Hatchikian, E. C., Volbeda, A., Frey, M., and FontecillaCamps, J. C. (1999) The crystal structure of a reduced [NiFeSe] hydrogenase
provides an image of the activated catalytic center, Structure 7, 557-566.
56.
Marques, M. C., Coelho, R., De Lacey, A. L., Pereira, I. A., and Matias, P. M.
(2009) The Three-Dimensional Structure of [NiFeSe] Hydrogenase from
Desulfovibrio vulgaris Hildenborough: A Hydrogenase without a Bridging Ligand
in the Active Site in Its Oxidised, "as-Isolated" State, J Mol Biol 396, 893-907.
107
57.
Kellers, P., Ogata, H., and Lubitz, W. (2008) Purification, crystallization and
preliminary X-ray analysis of the membrane-bound [NiFe] hydrogenase from
Allochromatium vinosum, Acta Crystallogr Sect F Struct Biol Cryst Commun 64,
719-722.
58.
Happe, R. P., Roseboom, W., Pierik, A. J., Albracht, S. P., and Bagley, K. A.
(1997) Biological activation of hydrogen, Nature 385, 126.
59.
Pierik, A. J., Roseboom, W., Happe, R. P., Bagley, K. A., and Albracht, S. P. J.
(1999) Carbon monoxide and cyanide as intrinsic ligands to iron in the active site
of [NiFe]-Hydrogenases. NiFe(CN)2CO, biology's way to activate H2, J Biol
Chem 274, 3331-3337.
60.
Ogata, H., Hirota, S., Nakahara, A., Komori, H., Shibata, N., Kato, T., Kano, K.,
and Higuchi, Y. (2005) Activation process of [NiFe] hydrogenase elucidated by
high-resolution X-ray analyses: conversion of the ready to the unready state,
Structure 13, 1635-1642.
61.
Lamle, S. E., Albracht, S. P., and Armstrong, F. A. (2004) Electrochemical
potential-step investigations of the aerobic interconversions of [NiFe]hydrogenase from Allochromatium vinosum: insights into the puzzling difference
between unready and ready oxidized inactive states, J Am Chem Soc 126, 1489914909.
62.
van Gastel, M., Stein, M., Brecht, M., Schroder, O., Lendzian, F., Bittl, R., Ogata,
H., Higuchi, Y., and Lubitz, W. (2006) A single-crystal ENDOR and density
functional theory study of the oxidized states of the [NiFe] hydrogenase from
Desulfovibrio vulgaris Miyazaki F, J Biol Inorg Chem 11, 41-51.
63.
Fichtner, C., Laurich, C., Bothe, E., and Lubitz, W. (2006)
Spectroelectrochemical characterization of the [NiFe] hydrogenase of
Desulfovibrio vulgaris Miyazaki F, Biochemistry 45, 9706-9716.
64.
Pardo, A., De Lacey, A. L., Fernandez, V. M., Fan, H. J., Fan, Y., and Hall, M. B.
(2006) Density functional study of the catalytic cycle of nickel-iron [NiFe]
hydrogenases and the involvement of high-spin nickel(II), J Biol Inorg Chem 11,
286-306.
65.
Niu, S., and Hall, M. B. (2001) Modeling the active sites in metalloenzymes 5.
The heterolytic bond cleavage of H2 in the [NiFe] hydrogenase of Desulfovibrio
gigas by a nucleophilic addition mechanism, Inorg Chem 40, 6201-6203.
108
66.
Fontecilla-Camps, J. C., Volbeda, A., Cavazza, C., and Nicolet, Y. (2007)
Structure/function relationships of [NiFe]- and [FeFe]-hydrogenases, Chem Rev
107, 4273-4303.
67.
Teixeira, V. H., Baptista, A. M., and Soares, C. M. (2006) Pathways of H2 toward
the active site of [NiFe]-hydrogenase, Biophys J 91, 2035-2045.
68.
Armstrong, F. A. (2004) Hydrogenases: active site puzzles and progress, Curr
Opin Chem Biol 8, 133-140.
69.
de Lacey, A., Fernandez, V., and Rousset, M. (2005) Native and mutant nickeliron hydrogenases: Unravelling structure and function, Coord Chem Rev 249,
1596-1608.
70.
De Lacey, A. L., Fernandez, V. M., Rousset, M., and Cammack, R. (2007)
Activation and inactivation of hydrogenase function and the catalytic cycle:
spectroelectrochemical studies, Chem Rev 107, 4304-4330.
71.
Ogata, H., Lubitz, W., and Higuchi, Y. (2009) [NiFe] hydrogenases: structural
and spectroscopic studies of the reaction mechanism, Dalton Trans, 7577-7587.
72.
Foerster, S., Stein, M., Brecht, M., Ogata, H., Higuchi, Y., and Lubitz, W. (2003)
Single crystal EPR studies of the reduced active site of [NiFe] hydrogenase from
Desulfovibrio vulgaris Miyazaki F, J Am Chem Soc 125, 83-93.
73.
Trofanchuk, O., Stein, M., Gessner, C., Lendzian, F., Higuchi, Y., and Lubitz, W.
(2000) Single crystal EPR studies of the oxidized active site of [NiFe]
hydrogenase from Desulfovibrio vulgaris Miyazaki F, J Biol Inorg Chem 5, 3644.
74.
Fan, H. J., and Hall, M. B. (2002) High-spin Ni(II), a surprisingly good structural
model for [NiFe] hydrogenase, J Am Chem Soc 124, 394-395.
75.
Gu, W., Jacquamet, L., Patil, D. S., Wang, H. X., Evans, D. J., Smith, M. C.,
Millar, M., Koch, S., Eichhorn, D. M., Latimer, M., and Cramer, S. P. (2003)
Refinement of the nickel site structure in Desulfovibrio gigas hydrogenase using
range-extended EXAFS spectroscopy, J Inorg Biochem 93, 41-51.
76.
Huyett, J. E., Carepo, M., Pamplona, A., Franco, R., Moura, I., Moura, J. J. G.,
and Hoffman, B. M. (1997) 57Fe Q-band pulsed ENDOR of the hetero-dinuclear
site of nickel hydrogenase: Comparison of the NiA, NiB, and NiC states, J Am
Chem Soc 119, 9291-9292.
109
77.
Fernandez, V., Hatchikian, E., and Cammack, R. (1985) Properties and
reactivation of two different deactivated forms of Desulfovibrio gigas
hydrogenase, Biochim Biophys Acta 832, 69-79.
78.
van der Zwaan, J. W., Coremans, J. M., Bouwens, E. C., and Albracht, S. P.
(1990) Effect of 17O2 and 13CO on EPR spectra of nickel in hydrogenase from
Chromatium vinosum, Biochim Biophys Acta 1041, 101-110.
79.
Bagley, K. A., Van Garderen, C. J., Chen, M., Duin, E. C., Albracht, S. P., and
Woodruff, W. H. (1994) Infrared studies on the interaction of carbon monoxide
with divalent nickel in hydrogenase from Chromatium vinosum, Biochemistry 33,
9229-9236.
80.
Ogata, H., Mizoguchi, Y., Mizuno, N., Miki, K., Adachi, S., Yasuoka, N., Yagi,
T., Yamauchi, O., Hirota, S., and Higuchi, Y. (2002) Structural studies of the
carbon monoxide complex of [NiFe]hydrogenase from Desulfovibrio vulgaris
Miyazaki F: suggestion for the initial activation site for dihydrogen, J Am Chem
Soc 124, 11628-11635.
81.
Pandelia, M. E., Ogata, H., Currell, L. J., Flores, M., and Lubitz, W. (2009)
Inhibition of the [NiFe] hydrogenase from Desulfovibrio vulgaris Miyazaki F by
carbon monoxide: An FTIR and EPR spectroscopic study, Biochim Biophys Acta.
82.
Balk, J., Pierik, A. J., Netz, D. J., Muhlenhoff, U., and Lill, R. (2004) The
hydrogenase-like Nar1p is essential for maturation of cytosolic and nuclear ironsulphur proteins, EMBO J 23, 2105-2115.
83.
Balk, J., Netz, D. J. A., Tepper, K., Pierik, A. J., and Lill, R. (2005) The essential
WD40 protein Cia1 is involved in a late step of cytosolic and nuclear iron-sulfur
protein assembly, Molecular and Cellular Biology 25, 10833-10841.
84.
Peters, J. W., Lanzilotta, W. N., Lemon, B. J., and Seefeldt, L. C. (1998) X-ray
crystal structure of the Fe-only hydrogenase (CpI) from Clostridium pasteurianum
to 1.8 angstrom resolution, Science 282, 1853-1858.
85.
Nicolet, Y., Piras, C., Legrand, P., Hatchikian, C. E., and Fontecilla-Camps, J. C.
(1999) Desulfovibrio desulfuricans iron hydrogenase: the structure shows unusual
coordination to an active site Fe binuclear center, Structure Fold Des 7, 13-23.
86.
Moulis, J. M., Sieker, L. C., Wilson, K. S., and Dauter, Z. (1996) Crystal structure
of the 2[4Fe-4S] ferredoxin from Chromatium vinosum: evolutionary and
mechanistic inferences for [3/4Fe-4S] ferredoxins, Protein Sci 5, 1765-1775.
110
87.
Adman, E. T., Siefker, L. C., and Jensen, L. H. (1976) Structure of Peptococcus
aerogenes ferredoxin. Refinement at 2 A resolution, J Biol Chem 251, 3801-3806.
88.
Bertini, I., Donaire, A., Feinberg, B. A., Luchinat, C., Piccioli, M., and Yuan, H.
(1995) Solution structure of the oxidized 2[4Fe-4S] ferredoxin from Clostridium
pasteurianum, Eur J Biochem 232, 192-205.
89.
Dauter, Z., Wilson, K. S., Sieker, L. C., Meyer, J., and Moulis, J. M. (1997)
Atomic resolution (0.94 A) structure of Clostridium acidurici ferredoxin. Detailed
geometry of [4Fe-4S] clusters in a protein, Biochemistry 36, 16065-16073.
90.
Unciuleac, M., Boll, M., Warkentin, E., and Ermler, U. (2004) Crystallization of
4-hydroxybenzoyl-CoA reductase and the structure of its electron donor
ferredoxin, Acta Crystallogr D Biol Crystallogr 60, 388-391.
91.
Giastas, P., Pinotsis, N., Efthymiou, G., Wilmanns, M., Kyritsis, P., Moulis, J. M.,
and Mavridis, I. M. (2006) The structure of the 2[4Fe-4S] ferredoxin from
Pseudomonas aeruginosa at 1.32-A resolution: comparison with other highresolution structures of ferredoxins and contributing structural features to
reduction potential values, J Biol Inorg Chem 11, 445-458.
92.
Tsukihara, T., Fukuyama, K., Mizushima, M., Harioka, T., Kusunoki, M.,
Katsube, Y., Hase, T., and Matsubara, H. (1990) Structure of the [2Fe-2S]
ferredoxin I from the blue-green alga Aphanothece sacrum at 2.2 A resolution, J
Mol Biol 216, 399-410.
93.
Tsukihira, T., Fukuyama, K., Nakamura, M., Katsube, Y., Tanaka, N., Kakudo,
M., Wada, K., Hase, T., and Matsubara, H. (1981) X-ray analysis of a [2Fe-2S]
ferrodoxin from Spirulina platensis. Main chain fold and location of side chains at
2.5 A resolution, J Biochem 90, 1763-1773.
94.
Rypniewski, W. R., Breiter, D. R., Benning, M. M., Wesenberg, G., Oh, B. H.,
Markley, J. L., Rayment, I., and Holden, H. M. (1991) Crystallization and
structure determination to 2.5-A resolution of the oxidized [2Fe-2S] ferredoxin
isolated from Anabaena 7120, Biochemistry 30, 4126-4131.
95.
Jacobson, B. L., Chae, Y. K., Markley, J. L., Rayment, I., and Holden, H. M.
(1993) Molecular structure of the oxidized, recombinant, heterocyst [2Fe-2S]
ferredoxin from Anabaena 7120 determined to 1.7-A resolution, Biochemistry 32,
6788-6793.
96.
Ikemizu, S., Bando, M., Sato, T., Morimoto, Y., Tsukihara, T., and Fukuyama, K.
(1994) Structure of [2Fe-2S] ferredoxin I from Equisetum arvense at 1.8 A
resolution, Acta Crystallogr D Biol Crystallogr 50, 167-174.
111
97.
Fukuyama, K., Ueki, N., Nakamura, H., Tsukihara, T., and Matsubara, H. (1995)
Tertiary structure of [2Fe-2S] ferredoxin from Spirulina platensis refined at 2.5 A
resolution: structural comparisons of plant-type ferredoxins and an electrostatic
potential analysis, J Biochem 117, 1017-1023.
98.
Frolow, F., Harel, M., Sussman, J. L., Mevarech, M., and Shoham, M. (1996)
Insights into protein adaptation to a saturated salt environment from the crystal
structure of a halophilic 2Fe-2S ferredoxin, Nat Struct Biol 3, 452-458.
99.
Happe, T., and Kaminski, A. (2002) Differential regulation of the Fe-hydrogenase
during anaerobic adaptation in the green alga Chlamydomonas reinhardtii, Eur J
Biochem 269, 1022-1032.
100.
Winkler, M., Heil, B., and Happe, T. (2002) Isolation and molecular
characterization of the [Fe]-hydrogenase from the unicellular green alga Chlorella
fusca, Biochim Biophys Acta 1576, 330-334.
101.
Forestier, M., King, P., Zhang, L., Posewitz, M., Schwarzer, S., Happe, T.,
Ghirardi, M. L., and Seibert, M. (2003) Expression of two [Fe]-hydrogenases in
Chlamydomonas reinhardtii under anaerobic conditions, Eur J Biochem 270,
2750-2758.
102.
Florin, L., Tsokoglou, A., and Happe, T. (2001) A novel type of iron hydrogenase
in the green alga Scenedesmus obliquus is linked to the photosynthetic electron
transport chain, J Biol Chem 276, 6125-6132.
103.
Wunschiers, R., Stangier, K., Senger, H., and Schulz, R. (2001) Molecular
evidence for a Fe-hydrogenase in the green alga Scenedesmus obliquus, Curr
Microbiol 42, 353-360.
104.
Nicolet, Y., de Lacey, A. L., Vernede, X., Fernandez, V. M., Hatchikian, E. C.,
and Fontecilla-Camps, J. C. (2001) Crystallographic and FTIR spectroscopic
evidence of changes in Fe coordination upon reduction of the active site of the Feonly hydrogenase from Desulfovibrio desulfuricans, J Am Chem Soc 123, 15961601.
105.
Pandey, A. S., Harris, T. V., Giles, L. J., Peters, J. W., and Szilagyi, R. K. (2008)
Dithiomethylether as a ligand in the hydrogenase h-cluster, J Am Chem Soc 130,
4533-4540.
106.
Bennett, B., Lemon, B. J., and Peters, J. W. (2000) Reversible carbon monoxide
binding and inhibition at the active site of the Fe-only hydrogenase, Biochemistry
39, 7455-7460.
112
107.
Holm, R. H. (1995) Chemical approaches to bridged biological metal assemblies,
Pure Appl Chem 67, 217.
108.
Holm, R. H., and Solomon, E. I. (1996) Bioinorganic enzymology - Preface,
Chem Rev 96, 2237-2237.
109.
Nicolet, Y., Lemon, B. J., Fontecilla-Camps, J. C., and Peters, J. W. (2000) A
novel FeS cluster in Fe-only hydrogenases, Trends Biochem Sci 25, 138-143.
110.
Lemon, B. J., and Peters, J. W. (1999) Binding of exogenously added carbon
monoxide at the active site of the iron-only hydrogenase (CpI) from Clostridium
pasteurianum, Biochemistry 38, 12969-12973.
111.
Lemon, B. J., and Peters, J. W. (2000) Photochemistry at the Active Site of the
Carbon Monoxide Inhibited Form of the Iron-Only Hydrogenase (CpI), J Am
Chem Soc 122, 3793-3794.
112.
Chen, Z., Lemon, B. J., Huang, S., Swartz, D. J., Peters, J. W., and Bagley, K. A.
(2002) Infrared studies of the CO-inhibited form of the Fe-only hydrogenase from
Clostridium pasteurianum I: examination of its light sensitivity at cryogenic
temperatures, Biochemistry 41, 2036-2043.
113.
Peters, J. W. (2009) Carbon monoxide and cyanide lignads in the active site of
[FeFe]-hydrogenase, In Metal ions in life sciences (Sigel, A., Sigel, H., and Sigel,
R. K. O., Eds.), pp 179-218, RSC Pub.
114.
Fan, H. J., and Hall, M. B. (2001) A capable bridging ligand for Fe-only
hydrogenase: density functional calculations of a low-energy route for heterolytic
cleavage and formation of dihydrogen, J Am Chem Soc 123, 3828-3829.
115.
Liu, Z., and Hu, P. (2002) Mechanism of H2 metabolism on Fe-only
hydrogenases, J Chem Phys 117, 8177-8180.
116.
Cao, Z., and Hall, M. B. (2001) Modeling the active sites in metalloenzymes. 3.
Density functional calculations on models for [Fe]-hydrogenase: structures and
vibrational frequencies of the observed redox forms and the reaction mechanism
at the diiron active center, J Am Chem Soc 123, 3734-3742.
117.
Zhou, T., Mo, Y., Zhou, Z., and Tsai, K. (2005) Density functional study on
dihydrogen activation at the H cluster in Fe-only hydrogenases, Inorg Chem 44,
4941-4946.
113
118.
Bruschi, M., Fantucci, P., and De Gioia, L. (2003) Density functional theory
investigation of the active site of [Fe]-hydrogenases: effects of redox state and
ligand characteristics on structural, electronic, and reactivity properties of
complexes related to the [2Fe]H subcluster, Inorg Chem 42, 4773-4781.
119.
Zhou, T., Mo, Y., Liu, A., Zhou, Z., and Tsai, K. R. (2004) Enzymatic mechanism
of Fe-only hydrogenase: density functional study on H-H making/breaking at the
diiron cluster with concerted proton and electron transfers, Inorg Chem 43, 923930.
120.
Schwab, D. E., Tard, C., Brecht, E., Peters, J. W., Pickett, C. J., and Szilagyi, R.
K. (2006) On the electronic structure of the hydrogenase H-cluster, Chem
Commun, 3696-3698
121.
Fiedler, A. T., and Brunold, T. C. (2005) Computational studies of the H-cluster
of Fe-only hydrogenases: geometric, electronic, and magnetic properties and their
dependence on the [Fe4S4] cubane, Inorg Chem 44, 9322-9334.
122.
Popescu, C., and Munck, E. (1999) Electronic structure of the H cluster in [Fe]hydrogenases, J Am Chem Soc 121, 7877-7884.
123.
Pereira, A. S., Tavares, P., Moura, I., Moura, J. J., and Huynh, B. H. (2001)
Mössbauer characterization of the iron-sulfur clusters in Desulfovibrio vulgaris
hydrogenase, J Am Chem Soc 123, 2771-2782.
124.
Razavet, M., Borg, S., George, S., Best, S., Fairhurst, S., and Pickett, C. (2002)
Transient FTIR spectroelectrochemical and stopped-flow detection of a mixed
valence {Fe (i)–Fe (ii)} bridging carbonyl intermediate with structural elements
and spectroscopic characteristics of the di-iron sub-site of all-iron hydrogenase,
Chem Commun 2002, 700-701.
125.
Liu, X. M., Ibrahim, S. K., Tard, C., and Pickett, C. J. (2005) Iron-only
hydrogenase: Synthetic, structural and reactivity studies of model compounds,
Coordination Chemistry Reviews 249, 1641-1652.
126.
Silakov, A., Wenk, B., Reijerse, E., and Lubitz, W. (2009) 14N HYSCORE
investigation of the H-cluster of [FeFe] hydrogenase: evidence for a nitrogen in
the dithiol bridge, Phys Chem Chem Phys 11, 6592-6599.
127.
Ryde, U., Greco, C., and De Gioia, L. (2010) Quantum Refinement of [FeFe]
Hydrogenase Indicates a Dithiomethylamine Ligand, J Am Chem Soc.
114
128.
Tard, C., and Pickett, C. J. (2009) Structural and functional analogues of the
active sites of the [Fe]-, [NiFe]-, and [FeFe]-hydrogenases, Chem Rev 109, 22452274.
129.
Gloaguen, F., and Rauchfuss, T. B. (2009) Small molecule mimics of
hydrogenases: hydrides and redox, Chem Soc Rev 38, 100-108.
130.
Cohen, J., Kim, K., King, P., Seibert, M., and Schulten, K. (2005) Finding gas
diffusion pathways in proteins: application to O2 and H2 transport in CpI [FeFe]hydrogenase and the role of packing defects, Structure 13, 1321-1329.
131.
Burgdorf, T., Lenz, O., Buhrke, T., van der Linden, E., Jones, A. K., Albracht, S.
P., and Friedrich, B. (2005) [NiFe]-hydrogenases of Ralstonia eutropha H16:
modular enzymes for oxygen-tolerant biological hydrogen oxidation, J Mol
Microbiol Biotechnol 10, 181-196.
132.
Vincent, K. A., Parkin, A., Lenz, O., Albracht, S. P., Fontecilla-Camps, J. C.,
Cammack, R., Friedrich, B., and Armstrong, F. A. (2005) Electrochemical
definitions of O2 sensitivity and oxidative inactivation in hydrogenases, J Am
Chem Soc 127, 18179-18189.
133.
Cracknell, J. A., Wait, A. F., Lenz, O., Friedrich, B., and Armstrong, F. A. (2009)
A kinetic and thermodynamic understanding of O2 tolerance in [NiFe]hydrogenases, Proc Natl Acad Sci USA 106, 20681-20686.
134.
Liebgott, P. P., Leroux, F., Burlat, B., Dementin, S., Baffert, C., Lautier, T.,
Fourmond, V., Ceccaldi, P., Cavazza, C., Meynial-Salles, I., Soucaille, P.,
Fontecilla-Camps, J. C., Guigliarelli, B., Bertrand, P., Rousset, M., and Leger, C.
Relating diffusion along the substrate tunnel and oxygen sensitivity in
hydrogenase, Nat Chem Biol 6, 63-70.
135.
Vincent, K. A., Cracknell, J. A., Lenz, O., Zebger, I., Friedrich, B., and
Armstrong, F. A. (2005) Electrocatalytic hydrogen oxidation by an enzyme at
high carbon monoxide or oxygen levels, Proc Natl Acad Sci USA 102, 1695116954.
136.
Goldet, G., Wait, A. F., Cracknell, J. A., Vincent, K. A., Ludwig, M., Lenz, O.,
Friedrich, B., and Armstrong, F. A. (2008) Hydrogen production under aerobic
conditions by membrane-bound hydrogenases from Ralstonia species, J Am Chem
Soc 130, 11106-11113.
115
137.
Leroux, F., Dementin, S., Burlat, B., Cournac, L., Volbeda, A., Champ, S.,
Martin, L., Guigliarelli, B., Bertrand, P., Fontecilla-Camps, J., Rousset, M., and
Leger, C. (2008) Experimental approaches to kinetics of gas diffusion in
hydrogenase, Proc Natl Acad Sci USA 105, 11188-11193.
138.
Dementin, S., Leroux, F., Cournac, L., de Lacey, A. L., Volbeda, A., Leger, C.,
Burlat, B., Martinez, N., Champ, S., Martin, L., Sanganas, O., Haumann, M.,
Fernandez, V. M., Guigliarelli, B., Fontecilla-Camps, J. C., and Rousset, M.
(2009) Introduction of methionines in the gas channel makes [NiFe] hydrogenase
aero-tolerant, J Am Chem Soc 131, 10156-10164.
139.
Bock, A., King, P. W., Blokesch, M., and Posewitz, M. C. (2006) Maturation of
hydrogenases, Adv Microb Physiol 51, 1-71.
140.
Posewitz, M. C., King, P. W., Smolinski, S. L., Zhang, L., Seibert, M., and
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, 25711-25720.
141.
Hu, Y., Fay, A. W., Lee, C. C., Yoshizawa, J., and Ribbe, M. W. (2008)
Assembly of nitrogenase MoFe protein, Biochemistry 47, 3973-3981.
142.
Rubio, L. M., and Ludden, P. W. (2008) Biosynthesis of the iron-molybdenum
cofactor of nitrogenase, Annu Rev Microbiol 62, 93-111.
143.
Johnson, D. C., Dean, D. R., Smith, A. D., and Johnson, M. K. (2005) Structure,
Function, and Formation of Biological Iron-Sulfur Clusters, Ann Rev Biochem 74,
247-281.
144.
Ayala-Castro, C., Saini, A., and Outten, F. (2008) Fe-S cluster assembly pathways
in bacteria, Microbiol Mol Biol Rev 72, 110.
145.
Fontecave, M., and Ollagnier-de-Choudens, S. (2007) Iron–sulfur cluster
biosynthesis in bacteria: mechanisms of cluster assembly and transfer, Arch
Biochem Biophys 474, 226-237.
146.
Bandyopadhyay, S., Chandramouli, K., and Johnson, M. (2008) Iron-sulfur
cluster biosynthesis, Biochem Soc Trans 36, 1112-1119.
147.
Lill, R. (2009) Function and biogenesis of iron-sulphur proteins, Nature 460, 831838.
116
148.
Bleijlevens, B., Buhrke, T., van der Linden, E., Friedrich, B., and Albracht, S. P.
(2004) The auxiliary protein HypX provides oxygen tolerance to the soluble
[NiFe]-hydrogenase of Ralstonia eutropha H16 by way of a cyanide ligand to
nickel, J Biol Chem 279, 46686-46691.
149.
Lenz, O., Gleiche, A., Strack, A., and Friedrich, B. (2005) Requirements for
heterologous production of a complex metalloenzyme: the membrane-bound
[NiFe] hydrogenase, J Bacteriol 187, 6590-6595.
150.
Blokesch, M., Paschos, A., Theodoratou, E., Bauer, A., Hube, M., Huth, S., and
Bock, A. (2002) Metal insertion into NiFe-hydrogenases, Biochem Soc Trans 30,
674-680.
151.
Maier, T., and Bock, A. (1996) Generation of active [NiFe] hydrogenase in vitro
from a nickel-free precursor form, Biochemistry 35, 10089-10093.
152.
Paschos, A., Glass, R. S., and Bock, A. (2001) Carbamoylphosphate requirement
for synthesis of the active center of [NiFe]-hydrogenases, FEBS Lett 488, 9-12.
153.
Paschos, A., Bauer, A., Zimmermann, A., Zehelein, E., and Bock, A. (2002)
HypF, a carbamoyl phosphate-converting enzyme involved in [NiFe] hydrogenase
maturation, J Biol Chem 277, 49945-49951.
154.
Reissmann, S., Hochleitner, E., Wang, H., Paschos, A., Lottspeich, F., Glass, R.
S., and Bock, A. (2003) Taming of a poison: biosynthesis of the NiFehydrogenase cyanide ligands, Science 299, 1067-1070.
155.
Blokesch, M., Paschos, A., Bauer, A., Reissmann, S., Drapal, N., and Bock, A.
(2004) Analysis of the transcarbamoylation-dehydration reaction catalyzed by the
hydrogenase maturation proteins HypF and HypE, Eur J Biochem 271, 34283436.
156.
Blokesch, M., Albracht, S. P., Matzanke, B. F., Drapal, N. M., Jacobi, A., and
Bock, A. (2004) The complex between hydrogenase-maturation proteins HypC
and HypD is an intermediate in the supply of cyanide to the active site iron of
[NiFe]-hydrogenases, J Mol Biol 344, 155-167.
157.
Blokesch, M., and Bock, A. (2006) Properties of the [NiFe]-hydrogenase
maturation protein HypD, FEBS Lett 580, 4065-4068.
158.
Watanabe, S., Matsumi, R., Arai, T., Atomi, H., Imanaka, T., and Miki, K. (2007)
Crystal structures of [NiFe] hydrogenase maturation proteins HypC, HypD, and
HypE: insights into cyanation reaction by thiol redox signaling, Mol Cell 27, 2940.
117
159.
Roseboom, W., Blokesch, M., Bock, A., and Albracht, S. P. (2005) The
biosynthetic routes for carbon monoxide and cyanide in the Ni-Fe active site of
hydrogenases are different, FEBS Lett 579, 469-472.
160.
Forzi, L., Hellwig, P., Thauer, R. K., and Sawers, R. G. (2007) The CO and CNligands to the active site Fe in [NiFe]-hydrogenase of Escherichia coli have
different metabolic origins, FEBS Lett 581, 3317-3321.
161.
Lenz, O., Zebger, I., Hamann, J., Hildebrandt, P., and Friedrich, B. (2007)
Carbamoylphosphate serves as the source of CN-, but not of the intrinsic CO in
the active site of the regulatory [NiFe]-hydrogenase from Ralstonia eutropha,
FEBS Lett 581, 3322-3326.
162.
Waugh, R., and Boxer, D. H. (1986) Pleiotropic hydrogenase mutants of
Escherichia coli K12: growth in the presence of nickel can restore hydrogenase
activity, Biochimie 68, 157-166.
163.
Maier, T., Jacobi, A., Sauter, M., and Bock, A. (1993) The product of the hypB
gene, which is required for nickel incorporation into hydrogenases, is a novel
guanine nucleotide-binding protein, J Bacteriol 175, 630-635.
164.
Xia, W., Li, H., Sze, K. H., and Sun, H. (2009) Structure of a nickel chaperone,
HypA, from Helicobacter pylori reveals two distinct metal binding sites, J Am
Chem Soc 131, 10031-10040.
165.
Watanabe, S., Arai, T., Matsumi, R., Atomi, H., Imanaka, T., and Miki, K. (2009)
Crystal structure of HypA, a nickel-binding metallochaperone for [NiFe]
hydrogenase maturation, J Mol Biol 394, 448-459.
166.
Peters, J. W., Boyd, E. S., D’Adamo, S., Mulder, D. W., Therien, J., and
Posewitz, M. C. (submitted) Hydrogenases, nitrogenases, anoxia, and H2
production in water-oxidizing phototrophs, In Algae for Biofuels and Energy
(Borowitzka, M. A., Ed.), Springer/Humana Press.
167.
Maier, T., Lottspeich, F., and Bock, A. (1995) GTP hydrolysis by HypB is
essential for nickel insertion into hydrogenases of Escherichia coli, Eur J
Biochem 230, 133-138.
168.
Atanassova, A., and Zamble, D. B. (2005) Escherichia coli HypA is a zinc
metalloprotein with a weak affinity for nickel, J Bacteriol 187, 4689-4697.
169.
Leach, M. R., and Zamble, D. B. (2007) Metallocenter assembly of the
hydrogenase enzymes, Curr Opin Chem Biol 11, 159-165.
118
170.
Menon, A. L., and Robson, R. L. (1994) In vivo and in vitro nickel-dependent
processing of the [NiFe] hydrogenase in Azotobacter vinelandii, J Bacteriol 176,
291-295.
171.
Menon, N. K., Robbins, J., Der Vartanian, M., Patil, D., Peck, H. D., Jr., Menon,
A. L., Robson, R. L., and Przybyla, A. E. (1993) Carboxy-terminal processing of
the large subunit of [NiFe] hydrogenases, FEBS Lett 331, 91-95.
172.
Rossmann, R., Maier, T., Lottspeich, F., and Bock, A. (1995) Characterisation of
a protease from Escherichia coli involved in hydrogenase maturation, Eur J
Biochem 227, 545-550.
173.
Rossmann, R., Sauter, M., Lottspeich, F., and Bock, A. (1994) Maturation of the
large subunit (HYCE) of Escherichia coli hydrogenase 3 requires nickel
incorporation followed by C-terminal processing at Arg537, Eur J Biochem 220,
377-384.
174.
Posewitz, M. C., King, P. W., Smolinski, S. L., Smith, R. D., Ginley, A. R.,
Ghirardi, M. L., and Seibert, M. (2005) Identification of genes required for
hydrogenase activity in Chlamydomonas reinhardtii, Biochem Soc Trans 33, 102104.
175.
King, P. W., Posewitz, M. C., Ghirardi, M. L., and Seibert, M. (2006) Functional
studies of [FeFe] hydrogenase maturation in an Escherichia coli biosynthetic
system, J Bacteriol 188, 2163-2172.
176.
Brazzolotto, X., Rubach, J. K., Gaillard, J., Gambarelli, S., Atta, M., and
Fontecave, M. (2006) The [Fe-Fe]-hydrogenase maturation protein HydF from
Thermotoga maritima is a GTPase with an iron-sulfur cluster, J Biol Chem 281,
769-774.
177.
Rubach, J. K., Brazzolotto, X., Gaillard, J., and Fontecave, M. (2005)
Biochemical characterization of the HydE and HydG iron-only hydrogenase
maturation enzymes from Thermatoga maritima, FEBS Lett. 579, 5055-5060.
178.
Sofia, H. J., Chen, G., Hetzler, B. G., Reyes-Spindola, J. F., and Miller, N. E.
(2001) Radical SAM, a novel protein superfamily linking unresolved steps in
familiar biosynthetic pathways with radical mechanisms: functional
characterization using new analysis and information visualization methods,
Nucleic Acids Res 29, 1097-1106.
179.
Fontecave, M., Atta, M., and Mulliez, E. (2004) S-adenosylmethionine: nothing
goes to waste, Trends Biochem Sci 29, 243-249.
119
180.
Frey, P. A. (2001) Radical Mechanisms of Enzymatic Catalysis, Annu. Rev.
Biochem. 70, 121-148.
181.
Frey, P. A., and Magnusson, O. T. (2003) S-Adenosylmethionine: A wolf in
sheep's clothing, or a rich man's adenosylmethionine?, Chem. Rev. 103, 21292148.
182.
Layer, G., Kervio, E., Morlock, G., Heinz, D. W., Jahn, D., Retey, J., and
Schubert, W. D. (2005) Structural and functional comparison of HemN to other
radical SAM enzymes, Biol Chem 386, 971-980.
183.
Marsh, E. N. G., Patwardhan, A., and Huhta, M. S. (2004) S-Adenosylmethionine
radical enzymes, Bioorg Chem 32, 326-340.
184.
Walsby, C., Ortillo, D., Yang, J., Nnyepi, M., Broderick, W. E., Hoffman, B. M.,
and Broderick, J. B. (2005) Spectroscopic approaches to elucidating novel ironsulfur chemistry in the "Radical SAM" protein superfamily, Inorg Chem 44, 727741.
185.
Chen, D., Walsby, C., Hoffman, B. M., and Frey, P. A. (2003) Coordination and
mechanism of reversible cleavage of S-adenosylmethionine by the [4Fe-4S]
center in lysine 2,3-aminomutase, J Am Chem Soc 125, 11788-11789.
186.
Cosper, M. M., Jameson, G. N. L., Davydov, R., Eidsness, M. K., Hoffman, B.
M., Huynh, B. H., and Johnson, M. K. (2002) The [4Fe-4S]2+ cluster in
reconstituted biotin synthase binds S-adenosyl-L-methionine, J Am Chem Soc
124, 14006-14007.
187.
Walsby, C. J., Ortillo, D., Broderick, W. E., Broderick, J. B., and Hoffman, B. M.
(2002) An anchoring role for FeS clusters: Chelation of the amino acid moiety of
S-adenosylmethionine to the unique iron site of the 4Fe-4S cluster of pyruvate
formate-lyase activating enzyme, J Am Chem Soc 124, 11270-11271.
188.
Walsby, C. J., Hong, W., Broderick, W. E., Cheek, J., Ortillo, D., Broderick, J. B.,
and Hoffman, B. M. (2002) Electron-nuclear double resonance spectroscopic
evidence that S-adenosylmethionine binds in contact with the catalytically active
[4Fe-4S]+ cluster of pyruvate formate-lyase activating enzyme, J Am Chem Soc
124, 3143-3151.
189.
Krebs, C., Broderick, W. E., Henshaw, T. F., Broderick, J. B., and Huynh, B. H.
(2002) Coordination of adenosylmethionine to a unique iron site of the [4Fe-4S]
of pyruvate formate-lyase activating enzyme: a Mossbauer spectroscopic study, J
Am Chem Soc 124, 912-913.
120
190.
Berkovitch, F., Nicolet, Y., Wan, J. T., Jarrett, J. T., and Drennan, C. L. (2004)
Crystal structure of biotin synthase, an S-adenosylmethionine-dependent radical
enzyme, Science 303, 76-79.
191.
Layer, G., Moser, J., Heinz, D. W., Jahn, D., and Schubert, W. D. (2003) Crystal
structure of coproporphyrinogen III oxidase reveals cofactor geometry of Radical
SAM enzymes, Embo Journal 22, 6214-6224.
192.
Hanzelmann, P., and Schindelin, H. (2004) Crystal structure of the Sadenosylmethionine-dependent enzyme MoaA and its implications for
molybdenum cofactor deficiency in humans, Proc Natl Acad Sci USA 101, 1287012875.
193.
Vey, J. L., Yang, J., Li, M., Broderick, W. E., Broderick, J. B., and Drennan, C.
L. (2008) Structural basis for glycyl radical formation by pyruvate formate-lyase
activating enzyme, Proc Natl Acad Sci USA 105, 16137-16141.
194.
Lepore, B. W., Ruzicka, F. J., Frey, P. A., and Ringe, D. (2005) The x-ray crystal
structure of lysine-2,3-aminomutase from Clostridium subterminale, Proc Natl
Acad Sci USA 102, 13819-13824.
195.
Cicchillo, R. M., Lee, K. H., Baleanu-Gogonea, C., Nesbitt, N. M., Krebs, C., and
Booker, S. J. (2004) Escherichia coli lipoyl synthase binds two distinct 4Fe-4S
clusters per polypeptide, Biochemistry 43, 11770-11781.
196.
Ugulava, N. B., Gibney, B. R., and Jarrett, J. T. (2001) Biotin synthase contains
two distinct iron-sulfur cluster binding sites: Chemical and spectroelectrochemical
analysis of iron-sulfur cluster interconversions, Biochemistry 40, 8343-8351.
197.
Allen, R. M., Chatterjee, R., Ludden, P. W., and Shah, V. K. (1995) Incorporation
of iron and sulfur from NifB cofactor into the iron-molybdenum cofactor of
dinitrogenase, J Biol Chem 270, 26890-26896.
198.
Curatti, L., Ludden, P. W., and Rubio, L. M. (2006) NifB-dependent in vitro
synthesis of the iron-molybdenum cofactor of nitrogenase, Proc Natl Acad Sci
USA 103, 5297-5301.
199.
Kulzer, R., Pils, T., Kappl, R., Huttermann, J., and Knappe, J. (1998)
Reconstitution and characterization of the polynuclear iron-sulfur cluster in
pyruvate formate-lyase-activating enzyme - Molecular properties of the
holoenzyme form, J Biol Chem 273, 4897-4903.
121
200.
Begley, T., Downs, D., Ealick, S., McLafferty, F., Van Loon, A., Taylor, S.,
Campobasso, N., Chiu, H., Kinsland, C., and Reddick, J. (1999) Thiamin
biosynthesis in prokaryotes, Arch microbiol 171, 293-300.
201.
Leonardi, R., Fairhurst, S., Kriek, M., Lowe, D., and Roach, P. (2003) Thiamine
biosynthesis in Escherichia coli: isolation and initial characterisation of the
ThiGH complex, FEBS letters 539, 95-99.
202.
Pilet, E., Nicolet, Y., Mathevon, C., Douki, T., Fontecilla-Camps, J. C., and
Fontecave, M. (2009) The role of the maturase HydG in [FeFe]-hydrogenase
active site synthesis and assembly, FEBS Lett. 583, 506-511.
203.
Wang, S. C., and Frey, P. A. (2007) S-adenosylmethionine as an oxidant: the
radical SAM superfamily, Trends Biochem Sci 32, 101-110.
204.
Frey, P. A., Hegeman, A. D., and Ruzicka, F. J. (2008) The Radical SAM
Superfamily, Crit Rev Biochem Mol Biol 43, 63-88.
205.
Bui, B. T. S., Lotierzo, M., Escalettes, F., Florentin, D., and Marquet, A. (2004)
Further investigation on the turnover of Escherichia coli biotin synthase with
dethiobiotin and 9-mercaptodethiobiotin as substrates, Biochemistry 43, 1643216441.
206.
Cicchillo, R. M., and Booker, S. J. (2005) Mechanistic investigations of lipoic
acid biosynthesis in Escherichia coli: Both sulfur atoms in lipoic acid are
contributed by the same lipoyl synthase polypeptide, J Am Chem Soc 127, 28602861.
207.
Cosper, M. M., Jameson, G. N. L., Hernandez, H. L., Krebs, C., Huynh, B. H.,
and Johnson, M. K. (2004) Characterization of the cofactor composition of
Escherichia coli biotin synthase, Biochemistry 43, 2007-2021.
208.
Jarrett, J. T. (2005) Biotin synthase: enzyme or reactant?, Chem Biol 12, 409-410.
209.
McGlynn, S. E., Ruebush, S. S., Naumov, A., Nagy, L. E., Dubini, A., King, P.
W., Broderick, J. B., Posewitz, M. C., and Peters, J. W. (2007) In vitro activation
of [FeFe] hydrogenase: new insights into hydrogenase maturation, J Biol Inorg
Chem 12, 443-447.
210.
McGlynn, S. E., Shepard, E. M., Winslow, M. A., Naumov, A. V., Duschene, K.
S., Posewitz, M. C., Broderick, W. E., Broderick, J. B., and Peters, J. W. (2008)
HydF as a scaffold protein in [FeFe] hydrogenase H-cluster biosynthesis, FEBS
Lett 582, 2183-2187.
122
211.
Shepard, E. M., Mcglynn, S. E., Bueling, A., Grady-Smith, C., George, S.,
Winslow, M. A., Cramer, S. P., Peters, J. W., and Broderick, J. B. (submitted)
Synthesis of the 2Fe-subcluster of the [FeFe]-hydrogenase H-cluster on the HydF
scaffold, Proc Natl Acad Sci USA
212.
Czech, I., Silakov, A., Lubitz, W., and Happe, T. (2009) The [FeFe]-hydrogenase
maturase HydF from Clostridium acetobutylicum contains a CO and CN(-) ligated
iron cofactor, FEBS Lett 584, 638-642.
213.
Kuchenreuther, J., Stapleton, J., and Swartz, J. (2009) Tyrosine, Cysteine, and SAdenosyl Methionine Stimulate In Vitro [FeFe] Hydrogenase Activation PLoS
One 4, e7565.
214.
Driesener, R. C., Challand, M. R., McGlynn, S. E., Shepard, E. M., Boyd, E. S.,
Broderick, J. B., Peters, J. W., and Roach, P. L. (2010) [FeFe]-Hydrogenase
Cyanide Ligands Derived From S-Adenosylmethionine-Dependent Cleavage of
Tyrosine, Angew Chem Int Ed Engl 22, 1687-1690.
215.
Shepard, E. M., Duffs, B. R., Mcglynn, S. E., Challand, M. R., Swanson, K. D.,
Roach, P. L., Peters, J. W., and Broderick, J. B. (submitted) [FeFe]-Hydrogenase
Maturation: HydG-Catalyzed Synthesis of CO, J Am Chem Soc.
216.
Dean, R. T., Padgett, H. C., and Rapoport, H. (1976) A high yield regiospecific
preparation of iminium salts, J Am Chem Soc 98, 7448-7449.
217.
Nicolet, Y., Rubach, J. K., Posewitz, M. C., Amara, P., Mathevon, C., Atta, M.,
Fontecave, M., and Fontecilla-Camps, J. C. (2008) X-ray structure of the [FeFe]hydrogenase maturase HydE from Thermotoga maritima, J Biol Chem 283,
18861-18872.
218.
Peters, J. W., Szilagyi, R. K., Naumov, A., and Douglas, T. (2006) A radical
solution for the biosynthesis of the H-cluster of hydrogenase, FEBS Lett. 580,
363-367.
219.
Schmid, B., Ribbe, M. W., Einsle, O., Yoshida, M., Thomas, L. M., Dean, D. R.,
Rees, D. C., and Burgess, B. K. (2002) Structure of a cofactor-deficient
nitrogenase MoFe protein, Science 296, 352-356.
220.
Neidhardt, F. C., Bloch, P. L., and Smith, D. F. (1974) Culture medium for
enterobacteria, J Bacteriol 119, 736-747.
221.
Bradford, M. (1976) A rapid and sensitive method for the quantitation of
microgram quantities of protein utilizing the principle of protein-dye binding,
Anal Biochem 72, 248.
123
222.
Fish, M. W. (1988) Rapid colorimetric micromethod for the quantitation
of complexed iron in biological samples., Methods Enzymol 158, 357-364
223.
Beinert, H. (1983) Semi-Micro Methods for Analysis of Labile Sulfide and of
Labile Sulfide Plus Sulfane Sulfur in Unusually Stable Iron Sulfur Proteins, Anal
Biochem 131, 373-378.
224.
Broderick, J. B., Henshaw, T. F., Cheek, J., Wojtuszewski, K., Smith, S. R.,
Trojan, M. R., McGhan, R. M., Kopf, A., Kibbey, M., and Broderick, W. E.
(2000) Pyruvate formate-lyase-activating enzyme: Strictly anaerobic isolation
yields active enzyme containing a 3Fe-4S (+) cluster, Biochem Biophys Res
Commun 269, 451-456.
225.
Ravel, B., and Newville, M. (2005) ATHENA, ARTEMIS, HEPHAESTUS: data
analysis for X-ray absorption spectroscopy using IFEFFIT, J Synchrotron
Radiation 12, 537-541.
226.
Rehr, J., and Albers, R. (2000) Theoretical approaches to x-ray absorption fine
structure, Rev mod phys 72, 621-654.
227.
Rao, P. V., and Holm, R. H. (2004) Synthetic analogues of the active sites of ironsulfur proteins, Chem Rev 104, 527-559.
228.
Johnson, M. K. (1994) Iron-Sulfur Proteins, In Encyclopedia of Inorganic
Chemistry (King, R. B., Ed., Ed.), pp 1896-1915, John Wiley & Sons, Chichester.
229.
Rupp, H., Rao, K. K., Hall, D. O., and Cammack, R. (1978) Electron spin
relaxation of iron-sulphur proteins studied by microwave power saturation,
Biochim Biophys Acta 537, 255-260.
230.
Middleton, P., Dickson, D. P., Johnson, C. E., and Rush, J. D. (1978)
Interpretation of the Mossbauer spectra of the four-iron ferredoxin from Bacillus
stearothermophilus, Eur J Biochem 88, 135-141.
231.
Teo, B., Shulman, R., Brown, G., and Meixner, A. (1979) EXAFS studies of
proteins and model compounds containing dimeric and tetrameric iron-sulfur
clusters, J Am Chem Soc 101, 5624-5631.
232.
Beinert, H., Emptage, M., Dreyer, J., Scott, R., Hahn, J., Hodgson, K., and
Thomson, A. (1983) Iron-sulfur stoichiometry and structure of iron-sulfur clusters
in three-iron proteins: Evidence for [3Fe-4S] clusters, Proc Nat Acad Sci USA 80,
393.
124
233.
Musgrave, K., Angove, H., Burgess, B., Hedman, B., and Hodgson, K. (1998)
All-ferrous titanium (III) citrate reduced Fe protein of nitrogenase: An XAS study
of electronic and metrical structure, J Am Chem Soc 120, 5325-5326.
234.
Liu, Z. P., and Hu, P. (2002) A density functional theory study on the active
center of Fe-only hydrogenase: characterization and electronic structure of the
redox states, J Am Chem Soc 124, 5175-5182.
235.
Duin, E. C., Lafferty, M. E., Crouse, B. R., Allen, R. M., Sanyal, I., Flint, D. H.,
and Johnson, M. K. (1997) 2Fe-2S to 4Fe-4S cluster conversion in Escherichia
coli biotin synthase, Biochemistry 36, 11811-11820.
236.
Miller, J. R., Busby, R. W., Jordan, S. W., Cheek, J., Henshaw, T. F., Ashley, G.
W., Broderick, J. B., Cronan, J. E., and Marletta, M. A. (2000) Escherichia coli
LipA is a lipoyl synthase: In vitro biosynthesis of lipoylated pyruvate
dehydrogenase complex from octanoyl-acyl carrier protein, Biochemistry 39,
15166-15178.
237.
Peters, J. W., Szilagyi, R. K., Naumov, A., and Douglas, T. (2006) A radical
solution for the biosynthesis of the H-cluster of hydrogenase, FEBS Lett 580, 363367.
238.
Bennett-Lovsey, R. M., Herbert, A. D., Sternberg, M. J., and Kelley, L. A. (2008)
Exploring the extremes of sequence/structure space with ensemble fold
recognition in the program Phyre, Proteins 70, 611-625.
239.
DeLano, W. L. (2002)
http://www.pymol.org.
240.
Barras, F., Loiseau, L., and Py, B. (2005) How Escherichia coli and
Saccharomyces cerevisiae build Fe/S proteins, Adv Microb Phys, Volume 50, 41.
241.
Fontecilla-Camps, J. C., Amara, P., Cavazza, C., Nicolet, Y., and Volbeda, A.
(2009) Structure-function relationships of anaerobic gas-processing
metalloenzymes, Nature 460, 814-822.
242.
Schwarz, G., Mendel, R. R., and Ribbe, M. W. (2009) Molybdenum cofactors,
enzymes and pathways, Nature 460, 839-847.
243.
Silakov, A., Kamp, C., Reijerse, E., Happe, T., and Lubitz, W. (2009)
Spectroelectrochemical characterization of the active site of the [FeFe]
hydrogenase HydA1 from Chlamydomonas reinhardtii, Biochemistry 48, 77807786.
The
PyMol
Molecular
Graphics
System
at
125
244.
McGlynn, S. E., Mulder, D. W., Shepard, E. M., Broderick, J. B., and Peters, J.
W. (2009) Hydrogenase cluster biosynthesis: organometallic chemistry nature's
way, Dalton Trans 22, 4274-4285.
245.
Peters, J. W., and Szilagyi, R. K. (2006) Exploring new frontiers of nitrogenase
structure and mechanism, Curr Opin Chem Biol 10, 101-108.
246.
Mulder, D. W., Ortillo, D. O., Gardenghi, D. J., Naumov, A. V., Ruebush, S. S.,
Szilagyi, R. K., Huynh, B., Broderick, J. B., and Peters, J. W. (2009) Activation
of HydA∆EFG requires a preformed [4Fe-4S] cluster, Biochemistry 48, 6240-6248.
247.
Kabsch, W. (1976) A solution for the best rotation to relate two sets of vectors,
Acta Crystallogr A32, 922-923.
248.
Larkin, M. A., Blackshields, G., Brown, N. P., Chenna, R., McGettigan, P. A.,
McWilliam, H., Valentin, F., Wallace, I. M., Wilm, A., Lopez, R., Thompson, J.
D., Gibson, T. J., and Higgins, D. G. (2007) Clustal W and Clustal X version 2.0,
Bioinformatics 23, 2947-2948.
249.
Laskowski, R. A., MacArthur, M. W., Moss, D. S., and Thornton, J. M. (1993)
PROCHECK: a program to check the stereochemical quality of protein structures,
J Appl Crystallogr 26, 283-291.
250.
Peters, J. W., Stowell, M. H., Soltis, S. M., Finnegan, M. G., Johnson, M. K., and
Rees, D. C. (1997) Redox-dependent structural changes in the nitrogenase Pcluster, Biochemistry 36, 1181-1187.
251.
Einsle, O., Tezcan, F. A., Andrade, S. L., Schmid, B., Yoshida, M., Howard, J. B.,
and Rees, D. C. (2002) Nitrogenase MoFe-protein at 1.16 A resolution: a central
ligand in the FeMo-cofactor, Science 297, 1696-1700.
252.
Fani, R., Gallo, R., and Lio, P. (2000) Molecular evolution of nitrogen fixation:
the evolutionary history of the nifD, nifK, nifE, and nifN genes, J Mol Evol 51, 111.
253.
Nicolet, Y., Cavazza, C., and Fontecilla-Camps, J. C. (2002) Fe-only
hydrogenases: structure, function and evolution, J Inorg Biochem 91, 1-8.
254.
Soboh, B., Boyd, E. S., Zhao, D., Peters, J. W., and Rubio, L. M. (2010) Substrate
specificity and evolutionary implications of a NifDK enzyme carrying NifB-co at
its active site, FEBS Lett.
126
255.
Georgiadis, M. M., Komiya, H., Chakrabarti, P., Woo, D., Kornuc, J. J., and Rees,
D. C. (1992) Crystallographic structure of the nitrogenase iron protein from
Azotobacter vinelandii, Science 257, 1653-1659.
256.
Otwinowski, Z., Minor, W., and Charles W. Carter, Jr. (1997) Processing of Xray diffraction data collected in oscillation mode, Methods Enzymol 276, 307-326.
257.
Collaborative. (1994) The CCP4 suite: programs for protein crystallography, Acta
Crystallogr D50, 760-763.
258.
Murshudov, G. N., Vagin, A. A., and Dodson, E. J. (1997) Refinement of
macromolecular structures by the maximum-likelihood method, Acta Crystallogr
D53, 240-255.
259.
Perrakis, A., Morris, R., and Lamzin, V. S. (1999) Automated protein model
building combined with iterative structure refinement, Nature Struct Biol 6, 458463.
260.
Terwilliger, T. C. (2003) SOLVE and RESOLVE: automated structure solution
and density modification, Methods Enzymol. 374, 22-37.
261.
Emsley, P., and Cowtan, K. (2004) Coot: model-building tools for molecular
graphics, Acta Crystallogr D60, 2126-2132.
262.
Terwilliger, T. C., and Berendzen, J. (1999) Automated MAD and MIR structure
solution, Acta Crystallogr D55, 849-861.
263.
Ten, L. F. (1973) Crystallographic fast Fourier transforms, Acta Crystallogr A29,
183-191.
264.
Guindon, S., and Gascuel, O. (2003) A simple, fast, and accurate algorithm to
estimate large phylogenies by maximum likelihood, Syst Biol 52, 696-704.
265.
Le, S. Q., and Gascuel, O. (2008) An improved general amino acid replacement
matrix, Mol Biol Evol 25, 1307-1320.
266.
Abascal, F., Zardoya, R., and Posada, D. (2005) ProtTest: selection of best-fit
models of protein evolution, Bioinform 21, 2104-2105.
267.
Ronquist, F., and Huelsenbeck, J. P. (2003) MrBayes 3: Bayesian phylogenetic
inference under mixed models, Bioinform 19, 1572-1574.
268.
Huelsenbeck, J. P., and Ronquist, F. (2001) MRBAYES: Bayesian inference of
phylogenetic trees, Bioinform 17, 754-755.
127
APPENDIX A
TABLE OF ACCESSION NUMBERS FOR HYDA, HYDE, HYDF, AND HYDG
HOMOLOGS COMPILED FOR CHAPTER 3. SEQUENCES WERE COMPILED
FROM THE DOE-IMG AND THE NCBI DATABASE IN SEPTEMBER 2009
128
Taxon
HydA
HydE
HydF
HydG
Abiotrophia defectiva
ATCC 49176
ZP_04452658
ZP_04451558
ZP_04451560
ZP_04451559
Acidaminococcus sp. D21
ZP_03929102
ZP_03929126
ZP_03928631
ZP_03989861
ZP_03989427
Acyrthosiphon pisum
XP_001943301
Aedes aegypti
XP_001655974
Alistipes putredinis DSM
17216
ZP_02424371
ZP_02425117
ZP_02425119
ZP_02425118
Alkaliphilus
metalliredigens QYMF
YP_001318525
YP_001321844
YP_001321843
YP_001321845
YP_001512534
YP_001512536
YP_001512535
YP_001321847
Alkaliphilus oremlandii
OhILAs
YP_001512473
YP_001512533
Anaerocellum
thermophilum DSM 6725
YP_002573172
YP_002574485
YP_002572378
YP_002572094
Anaerococcus
hydrogenalis DSM 7454
ZP_03305079
ZP_03304005
ZP_03304701
ZP_03304702
Anaerococcus prevotii
DSM 20548
ZP_04049650
ZP_04049876
ZP_04049877
ZP_04049878
ZP_03930686
ZP_03930687
YP_003152043
Anaerococcus tetradius
ATCC 35098
ZP_03929493
ZP_03930685
129
Anaerofustis
stercorihominis DSM
17244
ZP_02861877
ZP_02860849
ZP_02860851
ZP_02860850
ZP_02417503
ZP_02421058
ZP_02418081
ZP_02441588
ZP_02441586
ZP_02441587
ZP_01962185
ZP_01962187
ZP_01962186
ZP_02862336
Anaerostipes caccae
DSM 14662
ZP_02417588
ZP_02417946
ZP_02419016
Anaerotruncus
colihominis DSM 17241
ZP_02444642
Anopheles gambiae str.
PEST
XP_319772
Arabidopsis thaliana:
Chromosome: 4
NP_567496
Aspergillus fumigatus
Af293
XP_751635
Aspergillus nidulans
FGSC A4: Chromosome
2
XP_661236
Aspergillus niger CBS
513.88 clone An01
XP_001389148
Aspergillus oryzae
RIB40: genomic DNA
BAE58127
Aspergillus terreus
NIH2624
XP_001213045
Bacteroides caccae
ATCC 43185
ZP_01960540
Bacteroides capillosus
ATCC 29799
ZP_02035215
130
ZP_02037133
Bacteroides
cellulosilyticus DSM
14838
ZP_03634729
ZP_03676190
ZP_03676192
ZP_03676191
ZP_03676194
Bacteroides intestinalis
DSM 17393
ZP_03015926
ZP_03015922
ZP_03015924
ZP_03015923
Bacteroides ovatus ATCC
8483
ZP_02068583
ZP_02067573
ZP_02067575
ZP_02067574
Bacteroides pectinophilus
ATCC 43243
ZP_03462801
Bacteroides
thetaiotaomicron VPI5482
NP_809037
Blastocystis sp. NandII
ACD10930.1
Blautia hansenii DSM
20583
ZP_03548879
Blautia
hydrogenotrophica DSM
10507
ZP_03780903
NP_810750
NP_810748
ZP_03781584
ZP_03783789
ZP_03781703
ZP_03780910
ZP_03781171
Bos taurus
A4FV58.2
Brachyspira
hyodysenteriae WA1
YP_002722659
YP_002720232
YP_002720231
YP_002720230
Brachyspira murdochii
DSM 12563
ZP_04048507
ZP_04046659
ZP_04046658
ZP_04046657
131
Branchiostoma floridae
XP_002241183
Bryantella formatexigens
DSM 14469
ZP_03686653
ZP_05347601
ZP_05346246
ZP_05346250
ZP_03687910
Caenorhabditis elegans
AAF60782.5
Caenorhabditis elegans:
Chromosome 3
NP_498092
Caldicellulosiruptor
saccharolyticus DSM
8903
YP_001180640
YP_001179319
YP_001179924
YP_001181192
Campylobacter rectus
RM3267
ZP_03610805
ZP_03610813
ZP_03610840
ZP_03610801
Candida albicans
SC5314: Chromosome 1
XP_723443
XP_723254
Candida glabrata str.
CBS138
XP_448037
Candidatus Cloacamonas
acidaminovorans
YP_001741248
YP_001740557
YP_001741048
YP_001741000
Candidatus Desulforudis
audaxviator MP104C
YP_001716352
YP_001716362
YP_001716361
YP_001716359
YP_001716354
YP_001717478
YP_001717682
132
Canis familiaris
XP_547207
Chlamydomonas
reinhardtii
XP_001693376
XP_001691465
XP_001691465
ZP_03652138
ZP_03653464
XP_001691319
XP_001694503
Chlorella' fusca
CAC83291
Ciona intestinalis
XP_002131190
Clostridiales bacterium
1_7_47FAA
ZP_03651101
ZP_03651102
ZP_04669283
ZP_03651544
Clostridium
acetobutylicum
AAB03723
Clostridium
acetobutylicum ATCC
824
NP_346675
NP_348258
NP_348277
NP_347984
Clostridium
asparagiforme DSM
15981
ZP_03760839
ZP_03759890
ZP_03759888
ZP_03759889
ZP_02210998
ZP_02211233
ZP_02211340
ZP_03760851
ZP_03761376
Clostridium bartlettii
DSM 16795
ZP_02210685
ZP_02210702
133
ZP_02210839
ZP_02212112
Clostridium beijerinckii
NCIMB 8052
YP_001308901
YP_001307445
YP_001310328
YP_001311504
ZP_02087032
ZP_02087803
ZP_02087802
YP_001311066
YP_001311176
Clostridium bolteae
ATCC BAA-613
ZP_02083481
ZP_02083500
ZP_02089645
ZP_02089736
Clostridium botulinum A
str. ATCC 19397
YP_001384105
YP_001385894
YP_001385896
YP_001385895
Clostridium botulinum A
str. ATCC 3502
YP_001254348
YP_001256061
YP_001256063
YP_001256062
Clostridium botulinum A
str. Hall
YP_001387645
YP_001389301
YP_001389303
YP_001389302
Clostridium botulinum A2
str. Kyoto
YP_002804182
YP_002806126
YP_002806128
YP_002806127
Clostridium botulinum A3
str. Loch Maree
YP_001787172
YP_001788925
YP_001788927
YP_001788926
Clostridium botulinum B
str. Eklund 17B
YP_001884306
YP_001884593
YP_001887658
YP_001886192
134
YP_001886193
YP_001886496
Clostridium botulinum B1
str. Okra
YP_001781395
YP_001783218
YP_001783220
YP_001783219
Clostridium botulinum
Ba4 str. 657
YP_002862803
YP_002864592
YP_002864594
YP_002864593
Clostridium botulinum Bf
ZP_02619183
ZP_02618324
ZP_02618326
ZP_02618325
Clostridium botulinum C
str. Eklund
ZP_02621445
ZP_02620058
ZP_02620056
ZP_02620057
Clostridium botulinum E3
str. Alaska E43
YP_001919488
YP_001919779
YP_001922641
YP_001921205
YP_001920992
YP_001921030
YP_001921431
Clostridium botulinum F
str. Langeland
YP_001391107
YP_001392937
YP_001392939
YP_001392938
Clostridium botulinum
NCTC 2916
ZP_02615089
ZP_02615713
ZP_02615715
ZP_02615714
Clostridium butyricum
5521
ZP_02948973
ZP_02950586
ZP_02948272
ZP_02950303
YP_002507459
YP_002507375
YP_002507376
ZP_02951233
Clostridium
cellulolyticum H10
YP_002506554
135
YP_002506620
YP_002506776
Clostridium difficile 630
YP_001089830
YP_001088668
YP_001088667
YP_001088669
ZP_05351338
ZP_05351337
ZP_05351339
YP_001089927
Clostridium difficile
ATCC 43255
ZP_05352557
ZP_05352607
Clostridium difficile CIP
107932
ZP_03120112
ZP_03121291
ZP_03121290
ZP_03121292
Clostridium difficile
QCD-23m63
ZP_03125532
ZP_03125948
ZP_03125947
ZP_03125949
ZP_01803193
ZP_01803192
ZP_01803195
ZP_05397567
ZP_05397566
ZP_05397568
ZP_02748020
ZP_02748019
ZP_02748021
ZP_03125632
Clostridium difficile
QCD-32g58
ZP_01802842
ZP_01801566
Clostridium difficile
QCD-37x79
ZP_05398734
ZP_05398850
Clostridium difficile
QCD-63q42
ZP_02740355
ZP_02747361
136
Clostridium difficile
QCD-76w55
ZP_02756550
ZP_02757889
ZP_02757890
ZP_02757888
ZP_02756608
ZP_02763890
Clostridium difficile
QCD-97b34
ZP_02763950
ZP_02762976
ZP_02762975
ZP_02762977
Clostridium hiranonis
DSM 13275
ZP_03292845
ZP_03292564
ZP_03292566
ZP_03292565
Clostridium hylemonae
DSM 15053
ZP_03479501
ZP_03778460
ZP_03480976
ZP_03778461
ZP_03480570
ZP_03480974
ZP_03778462
YP_001391107
EDK32659
YP_001395713
YP_001397177
YP_001395640
YP_001394007
YP_002471219
YP_002470999
YP_002472515
YP_002473846
ZP_03481026
Clostridium kluyveri
DSM 555
Clostridium kluyveri
NBRC 12016
YP_002472451
Clostridium leptum DSM
753
ZP_02082403
ZP_02081236
ZP_02081234
ZP_02081235
Clostridium
methylpentosum DSM
5476
ZP_03705348
ZP_03708102
ZP_03708100
ZP_03708101
Clostridium nexile DSM
1787
ZP_03288284
ZP_03290619
ZP_03290617
ZP_03290618
137
ZP_03290965
Clostridium novyi NT
YP_877778
YP_877565
YP_877567
YP_877566
YP_697148
YP_694567
YP_694570
ZP_02636843
ZP_02637493
ZP_02637489
ZP_02864195
ZP_02863313
ZP_02863304
ZP_02640777
ZP_02639022
ZP_02639020
ZP_02953761
ZP_02953748
ZP_02953735
YP_878332
Clostridium pasteurianum
AAA23248
Clostridium perfringens
BAA74726
BAA74738
Clostridium perfringens
ATCC 13124
YP_695522
YP_697035
Clostridium perfringens B
str. ATCC 3626
ZP_02635975
ZP_02637414
Clostridium perfringens C
str. JGS1495
ZP_02864829
ZP_02865968
Clostridium perfringens
CPE str. F4969
ZP_02637950
ZP_02638882
Clostridium perfringens D
str. JGS1721
ZP_02862336
138
ZP_02953701
Clostridium perfringens E
str. JGS1987
ZP_02631057
ZP_02632860
ZP_02631742
ZP_02631740
ZP_02643845
ZP_02643500
ZP_02643504
YP_699717
YP_697443
YP_697445
ZP_02631180
Clostridium perfringens
NCTC 8239
ZP_02641711
ZP_02642314
Clostridium perfringens
SM101
YP_698262
YP_699606
Clostridium perfringens
str. 13
NP_563262
NP_563376
NP_561024
NP_561026
Clostridium
phytofermentans ISDg
YP_001557216
YP_001559190
YP_001559188
YP_001559189
ZP_02428409
ZP_02428411
ZP_02428410
ZP_02432053
ZP_02432055
ZP_02432054
YP_001559163
YP_001560891
Clostridium ramosum
DSM 1402
ZP_02427918
Clostridium
saccharobutylicum
AAA85785
Clostridium scindens
ATCC 35704
ZP_02431953
ZP_02432096
139
ZP_02433354
Clostridium sp. L2-50
ZP_02074529
ZP_02074795
ZP_02074771
ZP_02074773
Clostridium sp. M62
ZP_03733976
ZP_03730736
ZP_03730734
ZP_03730735
Clostridium sp. SS2/1
ZP_02440568
ZP_02437903
ZP_02438347
ZP_02440252
Clostridium spiroforme
DSM 1552
ZP_02868187
ZP_02867474
ZP_02867472
ZP_02867473
Clostridium sporogenes
ATCC 15579
ZP_02996025
ZP_02993906
ZP_02993904
ZP_02993905
Clostridium thermocellum
ATCC 27405
YP_001036773
YP_001038253
YP_001036476
YP_001037082
ZP_03150788
ZP_03149990
ZP_03150639
ZP_03798186
ZP_03797942
YP_002246510
YP_002246509
YP_001036861
YP_001039392
Clostridium thermocellum
DSM 4150
ZP_03150969
Collinsella aerofaciens
ATCC 25986
ZP_01772411
Coprococcus comes
ATCC 27758
ZP_03798014
ZP_03799143
Coprothermobacter
proteolyticus DSM 5265
YP_002246551
Cryptococcus neoformans
var. neoformans B3501A: Chromosome 8
XP_773874
YP_002246511
140
Cryptococcus neoformans
var. neoformans JEC21
XP_572892
Danio rerio
NP_001074460
Danio rerio:
Chromosome 12
NP_001002342
XP_684304
XP_687073
Danio rerio:
Chromosome 1
XP_683711
Debaryomyces hansenii
CBS767
XP_458135
Dechloromonas
aromatica RCB
YP_284205
Dehalococcoides
ethenogenes 195
YP_180897
Dehalococcoides sp.
BAV1
YP_001213691
Dehalococcoides sp.
CBDB1
YP_307338
Dehalococcoides sp. VS
ZP_02203712
Desulfitobacterium
hafniense DCB-2
YP_002457890
YP_002458510
YP_002461058
YP_002461288
141
Desulfitobacterium
hafniense Y51
YP_517169
YP_521191
YP_520202
YP_520945
Desulfobacterium
autotrophicum HRM2
Desulfotalea psychrophila
LSv54
YP_002601445
YP_002601439
YP_002603916
YP_002602797
YP_064215
YP_064211
YP_002601437
YP_066114
YP_066113
ZP_04351988
YP_003189760
YP_001113494
YP_001113492
YP_066112
Desulfotomaculum
acetoxidans DSM 771
ZP_04351973
ZP_04351986
ZP_04351977
ZP_04352660
ZP_04353069
ZP_04353070
ZP_04354102
Desulfotomaculum
reducens MI-1
YP_001111834
YP_001113004
YP_001113007
YP_001113493
142
YP_001114615
Desulfovibrio
desulfuricans
AAK11625
Desulfovibrio
desulfuricans G20
YP_386577
YP_388769
YP_388768
YP_388770
YP_002480084
YP_002480081
YP_002480085
YP_386971
YP_388773
Desulfovibrio
desulfuricans
subsp. desulfuricans str.
ATCC 27774
YP_002480083
Desulfovibrio
fructosovorans
CAA72423
YP_002480520
AAA87057
Desulfovibrio salexigens
DSM 2638
ZP_03330130
ZP_03464477
ZP_03464478
YP_002989973
ZP_03464471
ZP_03467901
ZP_03466104
YP_002990936
YP_002990940
ZP_03467900
ZP_03467896
ZP_03464474
Desulfovibrio vulgaris
AAA23373
CAA40970
143
CAA26266
Desulfovibrio vulgaris
subsp. vulgaris DP4
YP_966829
YP_966833
YP_966832
YP_966835
YP_010985
YP_010986
YP_010983
ZP_03729906
ZP_03729908
ZP_03729909
ZP_02233324
ZP_02233323
ZP_01994144
ZP_01994143
YP_966831
Desulfovibrio vulgaris
subsp. vulgaris str.
Hildenborough
YP_010987
YP_010989
Dethiobacter alkaliphilus
AHT 1
ZP_03729918
Dictyoglomus
thermophilum H-6-12
YP_002250290
YP_002250294
Dictyoglomus turgidum
DSM 6724
YP_002352460
YP_002352464
Dictyostelium discoideum
AX4: Chromosome 5
XP_635360
Dorea formicigenerans
ATCC 27755
ZP_02234450
ZP_02234883
Dorea longicatena DSM
13814
ZP_01996214
144
Drosophila grimshawi
XP_001988084
Drosophila melanogaster
AAL48960
AAF45404
Drosophila melanogaster:
Chromosome 2R
NP_001036428
Drosophila mojavensis
XP_002002631
Drosophila persimilis
XP_002023111
Drosophila virilis
XP_002051294
Drosophila willistoni
XP_002065237
Elusimicrobium minutum
Pei191
YP_001875425
Entamoeba histolytica
AAG09783
Entamoeba histolytica
HM-1
XP_652137
XP_652137
XP_656685
XP_652839
XP_656685
YP_001876216
YP_001875165
YP_001876220
145
Equus caballus
XP_001496296
Eremothecium gossypii
ATCC 10895
NP_982742
Eubacterium hallii DSM
3353
ZP_03715049
ZP_03717668
ZP_03716281
ZP_03715773
ZP_03716960
ZP_03718050
Eubacterium siraeum
DSM 15702
ZP_02422892
ZP_02421770
ZP_02421772
ZP_02421771
Fervidobacterium
nodosum Rt17-B1
YP_001409749
YP_001409764
YP_001409769
YP_001409762
YP_001409760
YP_001409840
Gallus gallus
XP_414836
Giardia lamblia ATCC
50803
EAA39802
Giardia lamblia ATCC
50803 strain WB C6
XP_769974
Gibberella zeae PH-1:
Chromosome 1
XP_380651
Hahella chejuensis KCTC
2396
YP_431454
146
Halothermothrix orenii
H168
YP_002507938
YP_002508054
YP_002508385
Heliobacterium
modesticaldum Ice1
YP_001679556
Histomonas meleagridis
ACI16483
Holomastigotoides
mirabile
BAF82037
Homo sapiens
BAB15199.
BAB15261
NP_071938
AAD51446
NP_036468
Homo sapiens:
Chromosome 16
NP_071938
Homo sapiens:
Chromosome 17
NP_036468
Hydra magnipapillata
XP_002157697
Kluyveromyces lactis
CAA49833
YP_002509658
YP_002509673
YP_002509674
147
Kluyveromyces lactis
NRRL Y-1140
XP_453943
Laribacter hongkongensis
HLHK9
YP_002796320
Macaca mulatta
XP_001118535
Magnaporthe grisea 70
XP_001402943
Malassezia globosa CBS
7966
EDP43317
Mariprofundus
ferrooxydans PV-1
ZP_01452314
Megasphaera elsdenii
AAF22114
Methylocella silvestris
BL2
YP_002363034
Mitsuokella multacida
DSM 20544
ZP_03471817
Monodelphis domestica
XP_001365922
Moorella thermoacetica
ATCC 39073
YP_430562
YP_430726
YP_430731
Mus musculus
NP_080514
Mus musculus:
Chromosome 11
NP_080548
YP_002796322
YP_002796319
ZP_03473484
ZP_03473489
YP_430579
YP_430134
YP_002796319
YP_430578
148
Mus musculus:
Chromosome 17
NP_080514
Nasonia vitripennis
XP_001605725
Natranaerobius
thermophilus JW/NMWN-LF
YP_001916287
Nematostella vectensis
XP_001630338
Neocallimastix frontalis
AAK60409
AAN39277
Neurospora crassa
OR74A: Chromosome 1
XP_965304
XP_965304
Nyctotherus ovalis
CAA76342
CAA76373
AAU14235
Opitutus terrae PB90-1
YP_001818428
Oribacterium sinus F0268
ZP_03990780
Ornithorhynchus anatinus
XP_001513248
Oryza sativa
XP_469746
ZP_03991992
ZP_03991990
ZP_03991991
149
Ostreococcus lucimarinus
CCE9901 chromosome 3
XP_001416706
Pan troglodytes
XP_510719
Parabacteroides
distasonis ATCC 8503
YP_001302432
Parabacteroides
johnsonii DSM 18315
ZP_03475733
Parabacteroides merdae
ATCC 43184
ZP_02034360
Pediculus humanus
corporis
XP_002429397
Pelobacter carbinolicus
DSM 2380
YP_357019
YP_357135
YP_357047
YP_357021
YP_355770
YP_357911
YP_357046
YP_357045
Pelobacter propionicus
DSM 2379
YP_901164
YP_901165
YP_901168
YP_901166
Pelotomaculum
thermopropionicum SI
YP_001211218
YP_001210799
YP_001211532
YP_001211534
YP_001211927
YP_001211533
YP_001568874
YP_001568876
YP_357043
YP_356037
YP_001212560
YP_001213196
Petrotoga mobilis SJ95
YP_001567770
YP_001568875
150
YP_001568190
Pichia stipitis CBS 6054:
Chromosome 7
XP_001386245
Piromyces sp. E2
AAL90459
Pongo abelii
NP_001124725
Populus trichocarpa
XP_002323132
Pseudotrichonympha
grassii
BAF82035
BAF82036
Psychromonas ingrahamii
37
YP_942648
Ralstonia metallidurans
CH34: Chromosome 1
YP_583674
Rattus norvegicus
NP_001013201
Rattus norvegicus:
Chromosome 10
NP_001013201
NP_001034296
Rhodoferax ferrireducens
T118
YP_525088
Rhodopseudomonas
palustris
YP_782767
Rhodopseudomonas
palustris CGA009
NP_945487
151
Rhodopseudomonas
palustris TIE-1
YP_001989175
Rhodospirillum rubrum
ATCC 11170
YP_425402
Ruminococcus gnavus
ATCC 29149
ZP_02042261
ZP_02041949
ZP_02041947
YP_001568876
Ruminococcus lactaris
ATCC 29176
ZP_03166398
ZP_03168009
ZP_03168845
ZP_03168010
Ruminococcus obeum
ATCC 29174
ZP_01965473
ZP_01965040
ZP_01962531
Ruminococcus torques
ATCC 27756
ZP_01966799
ZP_01969065
ZP_01968929
ZP_01969064
Saccharomyces cerevisiae
AAT93011
NP_014159
Saccharomyces cerevisiae
AWRI1631: Chromosome
14
EDZ69818
Saccharomyces cerevisiae
YJM789: Chromosome
14
EDN62585
Scenedesmus obliquus
CAC34419
Schizosaccharomyces
pombe
CAB40177
Schizosaccharomyces
pombe 972h
NP_588309
Shewanella halifaxensis
HAW-EB4
YP_001673633
YP_001673638
YP_001673639
YP_001673636
Shewanella oneidensis
MR-1
NP_719451
NP_719456
NP_719457
NP_719454
152
Shewanella sp. ANA-3:
Chromosome 1
YP_868355
YP_868350
YP_868349
YP_868352
Shewanella sp. MR-4
YP_735375
YP_735380
YP_735381
YP_735378
Spironucleus barkhanus
AAG31038
Strongylocentrotus
purpuratus
XP_001191432
Subdoligranulum
variabile DSM 15176
ZP_03775084
Symbiobacterium
thermophilum IAM 14863
YP_077035
YP_074182
YP_074181
YP_074183
YP_844976
YP_844973
YP_845963
YP_845961
YP_844980
YP_845964
YP_753702
YP_753705
YP_753032
YP_753707
YP_461123
YP_461140
YP_461122
YP_077118
Syntrophobacter
fumaroxidans MPOB
Syntrophomonas wolfei
subsp. wolfei str.
Goettingen
YP_844974
YP_753709
YP_754497
YP_754593
YP_755096
Syntrophus aciditrophicus
SB
YP_461142
153
Taeniopygia guttata
XP_002188515
XP_002193820
Tetraodon nigroviridis
CAG11445
Thalassiosira pseudonana
CCMP1335
XP_002295160
Thermoanaerobacter
pseudethanolicus ATCC
33223
YP_001665442
YP_001664645
Thermoanaerobacter sp.
X514
YP_001663746
YP_001662207
YP_001664250
YP_001665676
YP_001662174
YP_001662764
Thermoanaerobacter
tengcongensis MB4
NP_622546
NP_622716
NP_622467
NP_623168
YP_002249509
YP_002249510
YP_002249512
NP_623465
Thermodesulfovibrio
yellowstonii DSM 11347
YP_002249517
YP_002249518
Thermosinus
carboxydivorans Nor1
ZP_01665950
ZP_01666842
ZP_01667192
ZP_01666986
Thermosipho africanus
TCF52B
YP_002334586
YP_002335576
YP_002335288
YP_002335578
YP_002334590
YP_002335580
154
Thermosipho
melanesiensis BI429
YP_001305742
YP_001306826
YP_001306593
YP_001306828
YP_001470582
YP_001469919
YP_001470656
YP_001471366
YP_001470586
YP_001471364
NP_229074
NP_228255
NP_229072
YP_002534027
YP_002533831
YP_002533769
YP_002534848
YP_002534609
YP_002534846
YP_001305746
YP_001306830
Thermotoga lettingae
TMO
YP_001470971
Thermotoga maritima
MSB8
AAD35293
AAD36491
AAD36496
NP_228016
NP_229221
NP_229226
Thermotoga neapolitana
DSM 4359
YP_002534613
155
Thermotoga petrophila
RKU
YP_001244319
YP_001244156
YP_001244957
YP_001245086
YP_001244075
YP_001245088
YP_001738529
YP_001739572
XP_001309182.1
XP_001313153
YP_001244961
Thermotoga sp. RQ2
YP_001738782
YP_001739570
YP_001739342
YP_001739667
YP_001739346
Tolumonas auensis DSM
9187
YP_002892704
Treponema denticola
ATCC 35405
NP_972199
Tribolium castaneum
XP_001808932
Trichomonas vaginalis
AAG31037
AAC47159
AAC47160
Trichomonas vaginalis
G3
XP_001305709
XP_001310180
XP_001322682
XP_001326754
XP_001310059
156
XP_001328981
XP_001330775
XP_001580286
XP_001584012
Trichoplax adhaerens
XP_002117790
Trypanosoma brucei
TREU927: Chromosome
10
XP_823285
Ustilago maydis 521:
Chromosome 6
XP_758949
Veillonella dispar ATCC
17748
ZP_04598684
ZP_04600260
ZP_04598665
ZP_04599378
ZP_03855979
ZP_03855126
ZP_03855724
ZP_01924295
ZP_01925163
ZP_04599185
Veillonella parvula DSM
2008
ZP_03855144
ZP_03855575
Victivallis vadensis
ATCC BAA-548
ZP_01923911
ZP_01924420
Xenopus (Silurana)
tropicalis
NP_001106557
Yarrowia lipolytica
CLIB122
XP_500499