1
Introduction
Redox
reactions play important roles in almost all biological processes,
including photosynthesis and respiration, which are two essential
energy processes that sustain all life on earth. It is thus not surprising
that biology employs redox-active metal ions in these processes. It
is largely the redox activity that makes metal ions uniquely qualified
as biological cofactors and makes bioinorganic enzymology both fun
to explore and challenging to study.
Even though most metal
ions are redox active, biology employs a
surprisingly limited number of them for electron transfer (ET) processes.
Prominent members of redox centers involved in ET processes include
cytochromes, iron–sulfur clusters, and cupredoxins. Together
these centers cover the whole range of reduction potentials in biology
(Figure 1). Because of their importance, general
reviews about redox centers
1−77
and specific reviews about cytochromes,
8,24,78−90
iron–sulfur proteins,
91−93
and cupredoxins
94−104
have appeared in the literature. In this review, we provide both
classification and description of each member of the above redox centers,
including both native and designed proteins, as well as those proteins
that contain a combination of these redox centers. Through this review,
we examine structural features responsible for their redox properties,
including knowledge gained from recent progress in fine-tuning the
redox centers. Computational studies such as DFT calculations become
more and more important in understanding the structure–function
relationship and facilitating the fine-tuning of the ET properties
and reduction potentials of metallocofactors in proteins. Since this
aspect has been reviewed extensively before,
105−110
and by other reviews in this thematic issue,
2000,2001,2002
it will not be covered here.
Figure 1
Reduction
potential range of redox centers in electron transfer
processes.
2
Cytochromes in Electron Transfer
Processes
2.1
Introduction to Cytochromes
Cytochromes
are a major class of heme-containing ET proteins found ubiquitously
in biology. They were first described in 1884 as respiratory pigments
(called myohematin or histohematin) to explain colored substances
in cells.
81,111
These colored substances were
later rediscovered in 1920 and named “cytochromes”,
or cellular pigments.
112
The intense red
color combined with relatively high thermodynamic stability makes
cytochromes easy to observe and to purify. As of today, more than
70 000 cytochromes have been discovered.
78
In addition, due to their small size, high solubility,
and well-folded helical structure and the presence of the heme chromophore,
cytochromes are one of the most extensively studied classes of proteins
spanning several decades.
79
Cytochromes
are present mostly in the inner mitochondrial membrane of eukaryotic
organisms and are also found in a wide variety of both Gram-positive
and Gram-negative bacteria.
113,114
Cytochromes play crucial
roles in a number of biological ET processes associated with many
different energy metabolisms. Additionally, cytochromes are involved
in apoptosis in mammalian cells.
115
Further
description of the latter role of cytochromes is beyond the scope
of this review, which is solely focused on the role of cytochromes
in ET. For a similar reason, another family of cytochromes, the cyts
P450 (CYP), which catalyze the oxidation of various organic substrates
such as metabolites (lipids, hormones, etc.) and xenobiotic substances
(drugs, toxic chemicals, etc.), will not be discussed in this review
either.
A number of books and reviews have appeared in the literature
describing
the role of cytochromes as ET proteins.
8,24,78−90
Here we summarize studies on both native and designed cytochromes
and their roles in biological ET processes.
2.2
Classification
of Cytochromes
Cytochromes
are classified on the basis of the electronic absorption maxima of
the heme macrocycle, such as a, b, c, d, f, and o types of heme. More specifically,
these letter names represent
characteristic absorbance maxima in the UV–vis electronic absorption
spectrum when the heme iron is coordinated with pyridine in its reduced
(ferrous) state, designated as the “pyridine hemochrome”
spectrum (Figure 2).
Figure 2
Representative pyridine
hemochromogen spectra of hemin cofactors:
(A) heme b, (B) heme a, and (C)
heme d
1. The spectrum of pyridine ferrohemochrome c is similar to that of heme b. Reprinted
with permission from ref (116). Copyright 1992 Springer-Verlag.
Table 1 shows the maximum peak positions
and their corresponding extinction coefficients of the pyridine hemochrome
spectra of various classes of cytochromes. These differences arise
from different substituents at the β-pyrrole positions on the
periphery of the heme.
Table 1
UV–Vis Spectral
Parameters
of Pyridine Hemochrome Spectra of Various Types of Cytochromesa
pyridine hemochromogen
heme
position
of α peak (nm)
εmM (at α peak)
α peak
(nm) of reduced protein
example
ref
protoheme IX (b)
557
34.4
557–563
cyt b
6
f complex
(117)
heme c
550
29.1
549–561
cyt c
(118)
heme a
587
26
587–611
cyt aa
3 oxidase
(117)
heme d
613
630–635
cyt bd oxidase
(116)
heme d
1
620
24
625
cyt cd
1 nitrite reductase
(116)
heme o
553
560
cyt bo
3 oxidase
(119)
a
Adapted with permission from ref (116). Copyright 1992 Springer-Verlag.
The word “heme”
specifically describes the ferrous
complex of the tetrapyrrole macrocyclic ligand called protoporphyrin
IX (Figure 3).
81
It is the precursor to various types of cytochromes through different
peripheral substitutions. Figure 3 shows a
schematic of these various types of hemes.
Figure 3
Different types of heme
found in cytochromes.
The b-type cytochromes have four methyl
substitutions
at positions 1, 3, 5, and 8, two vinyl groups in positions 2 and 4,
and two propionate groups at positions 6 and 7, resulting in a 22-π-electron
porphyrin. Hemes a and c are biosynthesized
as derivatives of heme b. In heme a, the vinyl group at position 2 of the porphyrin
ring of heme b is replaced by a hydroxyethylfarnesyl side chain while
the methyl group at position 8 is oxidized to a formyl group. These
substituents make heme a more hydrophobic as well
as more electron-withdrawing than heme b due to the
presence of farnesyl and formyl groups, respectively. Covalent cross-linking
of the vinyl groups at β-pyrrole positions 2 and 4 of heme b with Cys residues from
the protein yields heme c, where the vinyl groups of heme b are
replaced by thioether bonds.
The covalent cross-linking of the
two Cys residues from the protein
to the porphyrin ring occurs at the highly conserved -Cys-Xxx-Xxx-Cys-His-
sequences (Xxx=any amino acid). This cross-linking covalently attaches
heme c to the protein. The histidine residue in the
conserved sequence serves as an axial ligand to the heme iron. In
heme d, two cis-hydroxyl groups
are inserted at positions 5 and 6 on the β-pyrrole, which renders
heme d as a 20-π-electron chlorin. Heme d
1 contains two ketone groups in place of the
vinyl groups at positions 2 and 4, while two acetate groups are added
to positions 1 and 3 of the tetrapyrrole macrocycle, resulting in
18-π-electron isobacteriochlorins. The hemes f is similar to heme c, with the difference
in the
ligands that coordinate to the heme iron at the axial position (called
axial ligands) make hemes c and f spectroscopically distinct.
Common axial ligands found in
cytochromes are shown in Figure 4. With the
exception of cytochromes c′ (cyts c′),
all cytochromes with ET function contain 6-coordinate
low-spin (6cLS) hemes axially ligated to amino acids such as His or
an N-terminal amine group. Two axial His residues act as ligands to
the heme iron in b-type cytochromes. The only example
of bis-Met axial coordination to heme b is observed
in the iron storage protein bacterioferritin.
120,121
A common axial His ligand is found in all cyts c, where the axial His is a part of
the conserved -Cys-Xxx-Xxx-Cys-His-
sequence, through which the heme is covalently attached to the protein.
The most commonly encountered second axial ligand in c-type cytochromes is Met with
the exception of multiheme c-type cytochromes, which generally display bis-His axial
ligation of the heme iron (section 2.3.6).
80
In most cases, the His ligands are coordinated
to the heme iron by their Nε atom. However, an example
of Nδ coordination has been reported.
122
The f-type cytochromes contain
the same type of heme with one axial His ligand, as in cyts c; the only exception
is in the nature of the second axial
ligation in that the second axial ligand is the NH2 group
of an N-terminal tyrosine instead of the most commonly found Met or
His as the second axial ligand.
123
Not
surprisingly, the variation in the axial ligation makes each heme
type electronically unique, resulting in different out-of-plane distortions
of the heme iron from the heme plane (Figure 4) as well as different spectroscopic
features (Table 1).
Figure 4
Commonly found heme axial ligands in various cytochromes. (A) Class
I cyts c (PDB ID 3CYT) uses His/Met axial ligation. (B) Cyts b and multiheme cyts
c contain bis-His
ligation (bovine liver cyt b
5, PDB ID 1CYO). (C) An unusual
His/amine ligation is found only in cyt f (PDB ID 1HCZ). (D) Bis-Met ligation
is encountered in bacterioferritin (PDB ID 1BCF). For c-type cytochromes
the conserved -Cys-Xxx-Xxx-Cys-His- ligation and its covalent linkage
to the heme via Cys residues are shown.
2.3
Native Cytochromes c
2.3.1
Functions of Cytochromes c
Cytochromes c are involved in biological
ET processes in both aerobic and anaerobic respiratory chains. In
aerobic respiration, they are involved in the mitochondrial respiratory
chain to produce the energy currency ATP by transferring electrons
from the transmembrane bc
1 complex to
cyt c oxidase.
85,86
In addition, cyts c have also been recently discovered to play a crucial role
in programmed cell death (apoptosis), where they activate the protease
involved in cell death, caspase 3.
124−126
Other examples where c-type cytochromes are involved in ET include the reduction
of sulfate to hydrogen sulfide, conversion of nitrogen to ammonia
in nitrogen fixation, reduction of nitrate to dinitrogen in denitrification,
in phototrophs that use light energy to carry out various cellular
processes, and in methylotrophs that use methane or methanol as the
carbon source for their growth. Detailed descriptions of the roles
of cyts c in these cases will be discussed in the
following sections.
As cyts c are involved
in numerous crucial biological processes, they have been used extensively
as a hallmark system to study biological ET by site-directed mutagenesis,
which have elucidated the regions of the protein that are critical
for their ET properties as well as fine-tuning the reduction potentials.
87,127−131
In addition, various inorganic redox couples have been covalently
appended to surface sites of cyts c to study intraprotein
ET pathways.
24,132,133
Various complexes of cyts c with other protein
partners have also been prepared to study interprotein ET pathways.
134−149
2.3.2
Classifications of Cytochromes c
Cytochromes c generally contain ∼100–120
amino acids. Biosynthesis of cyts c involves the
formation of two thioether bonds between two Cys residues and the
two vinyl groups of heme b by post-translational
modification.
150,151
Primary amino acid sequence
alignment shows that the residue identity of cyts c is 45–100% among eukaryotes. The
electronic spectra of cyts c are dominated by the allowed porphyrin π →
π* transitions that are mixed together with interelectronic
repulsions that give rise to an intense band at ∼410 nm (called
the Soret or γ band) and two weaker signals in the 500–600
nm range (the α and β bands). The reduced form of the
protein shows a Soret band at 413 nm and sharp α and β
bands at 550 nm (ε = 29.1 mM–1 cm–1) and 521 nm (ε = 15.5 mM–1 cm–1), respectively,
with a ratio of α to β bands of 1.87
(Table 1). The electronic spectra of cyts c from other sources are very similar to
that of horse heart
cyt c. Originally classified by Ambler,
89,152
cyts c have been divided into four major classes
on the basis of the number of hemes, position and identity of the
axial iron ligands, and reduction potentials (Table 2).
Table 2
Axial Ligand Types and Reduction Potentials
of Various Cytochromesa
cytochrome
axial ligand
heme type
E (mV)b
mutant
E (mV)
ref
Nitrosomonas
europaea diheme cyt c peroxidase
His/Met
class I
450
(153, 154)
Rhodocyclus
tenuis THRC cyt c
class IV
420
(155)
HP1
His/Met
420
HP2
His/Met
110
LP1
bis-His
60
LP2
His/Met
Rhodopseudomonas
viridis THRC cyt c
class IV
380
(156,157)
H1 (c
559)
His/Met
330
H3 (c
556)
His/Met
20
H2 (c
552)
bis-His
–60
H4 (c
554)
His/Met
Rhodobacter
capsulatas cyt c
2
His/Met
class I
373
Gly29Ser
330
(158−160)
Pro30Ala
258
Tyr67Cys
348
Tyr67Phe
308
Chlamydomonas
reinhardtii cyt f
His/Ntr-Tyr
cyt f
370
Tyr1Phe
369
(161)
Tyr1Ser
313
Val3Phe
373
Phe4Leu
348
Phe4Trp
336
Tyr1Phe/Phe4Tyr
370
Tyr1Ser/Phe4Leu
289
Val3Phe/Phe4Trp
342
Rhodospirillum
rubrum cyt c
2
His/Met
class I
324
(156)
Pseudomonas
aeruginosa cyt c nitric oxide reductase
His/Met
class I
310
(162)
bis-His
cyt b
345
Pseudomonas
aeruginosa cyt c peroxidase
His/Met
class I
320
(163)
Arthrospira
maxima cyt c
6
His/Met
class I
314
(164)
Saccharomyces
cerevisiae iso-2-cyt c
His/Met
class I
288
Asn52Ile
243
(130)
Saccharomyces
cerevisiae iso-1-cyt c
His/Met
class I
272
Arg38Lys
249
(131, 165−173)
285
Arg38His
245
290
Arg38Gln
242
Arg38Asn
238
Arg38Leu
231
Arg38Ala
225
Asn52Ala
257
Asn52Ile
231
Tyr67Phe
234
Phe82Leu
286
Phe82Tyr
280
Phe82Ile
273
Phe82Trp
266
Phe82Ala
260
Phe82Ser
255
Phe82Gly
247
Pseudomonas
aeruginosa cyt c
551
His/Met
class I
276
(156)
horse cyt c
His/Met
class I
262
Met80Ala
82
(158, 174)
Met80His
41
Met80Leu
–42
Met80Cys
–390
rat cyt c
His/Met
class I
260
Pro30Ala
258
Pro30Val
261
Tyr67Phe
224
Rhodopseudomonas
palustris cyt c
556
His/Met
class II
230
(80)
Escherichia
coli cyt b
562
His/Met
cyt b (class II)
168
Phe61Gly
90
(175, 176)
Phe65Val
173
Phe61Ile/Phe65Tyr
68
His102Met
240
Arg98Cys/His102Met
440
Alicycliphilus
denitrificans cyt c′
His/Met
class II
132
(80)
Rhodopseudomonas
palustris cyt c′
His/Met
class II
102
(80)
cytochrome b
5
bis-His
cyt b
form A
80
(177)
form B
–26
Desulfovibrio
vulgaris cyt c
553
His/Met
class I
37
Met23Cys
29
(156, 178)
20 ± 5
Gly51Cys
28
Met23Cys/Met23Cys
88
Gly51Cys/Gly51Cys
105
bovine liver microsomal cyt b
5
bis-His
cyt b
3
protoheme IX
dimethyl ester
70
(179)
Saccharomyces
cerevisiae cyt b
2
bis-His
cyt b
–3
(156)
Chromatium
vinosum cyt c′
His
class II
–5
(80)
rat liver microsomal cyt b
5
bis-His
cyt b
–7 ± 1
(129, 180)
Rhodospirillum
rubrum cyt c′
His/Met
class I
–8
(80)
tryptic bovine hepatic cyt b
5
His/Met
class I
–10 ± 3
Val61Lys
17
(181)
Val61His
11
Val61Glu
–25
Val61Tyr
–33
Allochromatium
vinosum triheme cyt c
bis-His
class III
–20
(182)
His/Met
–200
His-Cys/Met
–220
Rhodobacter
sphaeroides cyt c′
His/Asn
cyt c
–22
(183)
cyt b
6
f complex
bis-His
cyt b
–45
(184)
–150
Thermosynechococcus
elongates PS cyt c
550
His/Met
class I
–80
in the absence of mediators
200
(185)
MamP magnetochrome
His/Met
class I
–76
(186)
rat liver OM cyt b
5
bis-His
cyt b
–102
His63Met
110
(187, 188)
Val45Leu/Val61Leu
–148
protoheme IX dimethyl ester
–36
Desulfovibrio
desulfuricans Norway cyt c
3
bis-His
class III
–132
(78)
bis-His
–255
bis-His
–320
bis-His
–360
Chlorella nitrate reductase cyt b
557
bis-His
cyt b
–164
(189, 190)
Ectothiorhodospira
shaposhnikovii cyt b
558
bis-His
cyt b
–210
(191)
Azotobacter
vinelandii bacterioferritin
bis-His
cyt b
–225
(192)
(in the presence of a nonheme
iron core)
–475
Desulfovibrio
vulgaris Hildenborough cyt c
3
bis-His
class III
–280
(192, 193)
bis-His
–320
bis-His
–350
bis-His
–380
Synechocystis sp. cyt c
549
bis-His
–250
(78)
Arthrospira
maxima cyt c
549
His/Met
–260
(164)
a
Adapted with permission from ref (78). Copyright 2004 Elsevier.
b
All reduction potentials
listed
in this review are versus standard hydrogen electrode (SHE) or normal
hydrogen electrode (NHE).
The class I cyts c include small (8–120
kDa) soluble proteins containing a single 6cLS heme moiety and display
a range of reduction potentials from −390 to +450 mV versus
standard hydrogen electrode (SHE) (Table 2).
78
On the basis of sequence and structural alignments,
class I cyts c have further been partitioned into
16 different subclasses.
88
The majority
of the subclasses include mitochondrial cyts c and
purple bacterial cyts c. Examples of other subclasses
represent a wide variety of different sources, including cyts c
551, cyts c
4, cyts c
5, and cyts c
6 from Pseudomonas, Chlorobium cyt c
555, Desulfovibrio (Dv.) cyts c
553, c
550 from cyanobacteria and algae, Ectothiorhodospira cyts c
551, flavocytochromes c, methanol dehydrogenase-associated
cyt c
550 or c
L, cyt cd
1 nitrite reductase, the cyt
subunit associated with alcohol dehydrogenase, nitrite reductase-associated
cyt c from Pseudomonas, and cyt c oxidase subunit II from Bacillus.
78
Class
I cyt c domains are characterized by their
signature cyt c fold and the presence of an N-terminal
conserved -Cys-Xxx-Xxx-Cys-His- sequence containing cysteines for
covalent cross-linking of the heme to the protein and the His, which
acts as the axial ligand to the heme iron. The class I cyt c fold is recognized as
having a total of five α-helices
arranged in a unique tertiary structure. There are two helices, one
each at the N- and C-termini, represented as α1 and α5,
respectively. In between, there is a small helix, α3 (also called
the 50s helix in mitochondrial cyts c), followed
by two other helices, α4 and α5, which are known as the
60s helix and 70s helix, respectively, in mitochondrial cyts c. The 70s helix precedes
a loop toward the C-terminus that
contains the second axial ligand, Met, to the heme iron. There are
examples where the second axial ligand is a residue other than Met,
e.g., Asn or His, or is even absent.
79
In
many cases, this core cyt c domain can be found fused
to other membrane proteins. General features of the class I cyt c fold are shown in
Figure 5.
Figure 5
Schematic representations
of various classes of cyts c. (A) Class I cyt c fold with His/Met heme axial
ligands (PDB ID 3CYT). Mitochondrial designation of the helices is also shown. (B)
Four-helix
bundle cyt c′ belongs to class II cyt c having a 5c heme with His120 as the sole axial
ligand
(PDB ID 1E83). (C) Tetraheme cyt c
3 belongs to class
III cyt c with bis-His ligation to all four hemes
(PDB ID 1UP9). Hemes I and III are attached to the protein via the highly conserved
-Cys-Xxx-Xxx-Cys-His- sequence, whereas hemes II and IV are covalently
bound to the protein by a -Cys-Xxx-Xxx-Xxx-Xxx-Cys-His- motif. In
(A)–(C) the covalent attachment of the heme to the protein
via Cys residues is shown. (D) Tetraheme cyt c from
the photosynthetic reaction center (RC) belongs to class IV cyt c. Hemes I, II, and
III have His/Met axial ligands, while
heme IV has bis-His axial ligation to the heme iron (PDB ID 2JBL). (E) Cyt c
554 from Nitrosomonas europaea belongs to a class of its own. Hemes I, III, and IV
have bis-His-ligated
heme iron, whereas heme II is 5c with His as the only axial ligand
(PDB ID 1BVB). Heme numbering in (C)–(E) is according to their attachment
occurring along the protein’s primary sequence. (F) Cyt f from chloroplast is unique
from all other classes of cytochromes
in that it mostly contains β-sheets and the heme is 6c with
a His and N-terminal backbone NH2 group of a Tyr residue
(PDB ID 1HCZ). It has been included as a subclass of cyt c because
the heme is covalently bound to the protein via the highly conserved
-Cys-Xxx-Xxx-Cys-His- signature motif for heme attachment ubiquitously
found in c-type cytochromes.
The class II cyts c consist of a c-type heme covalently attached to the highly conserved
C-terminal
-Cys-Xxx-Xxx-Cys-His- sequence, as in class I cyts c, with the Cys residues and the
His as one of the axial ligands.
80
Four α-helices and a left-handed twisted
overall structure represent this subclass of cyts c (Figure 5). The second axial ligand
to the
heme iron is variable.
194,195
The subclass cyt c′ is axially coordinated to a single His imidazole
ligand, lacks the second axial ligand, and has a relatively small
range of reduction potentials ranging from approximately −200
to +200 mV.
8,84,90
Members from this subclass represent a wide range of sources that
include photosynthetic, denitrifying, nitrogen-fixing, methanotrophic,
and sulfur oxidizing bacteria. This class has two subclasses based
on the distinct spin states displayed by the heme. Subclass IIa of
cyt c′ displays high-spin (HS) ferrous [Fe(II), S = 2] electronic configurations,
while the ferric form
shows either a HS S = 5/2 state or S = 3/2, S = 5/2 mixture of spin states.
196−202
The subclass IIa proteins, isolated from Rhodopseudomonas
palustris, Rhodobacter (Rb.) capsulatus, and Chromatium (Ch.) vinosum, display
a large amount of the S = 3/2 ground state in the
spin-state admixture, ranging from 40% to 57% as determined from electron
paramagnetic resonance (EPR) simulations.
196,201,203
The second subclass, IIb, includes
cyt c
556 from Rp. palustris,
204
Rb. sulfidophilus,
205
and Agrobacterium
tumefaciens(80) and cyt c
554 from Rb. sphaeroides,
206
which contain heme in the low-spin
(LS) configuration. This subclass of proteins has a second axial ligand
to the heme iron which is a Met residue located close to the N-terminus.
Class II cyts display reduction potentials ranging from −5
to +230 mV (Table 2).
Class III cyts c include proteins containing multiple
hemes with bis-His ligation and display reduction potentials in the
range of −20 to −380 mV (Table 2).
80,88,152,207−212
In some cases this class of cytochromes have up to 16 heme cofactors
and display no structural similarity with other classes of cyts c. They are found
as terminal electron donors in bacteria
involved in sulfur metabolism.
213
These
bacteria utilize sulfur or oxidized sulfur compounds as terminal electron
acceptors in their respiratory chain. One of the best studied proteins
in this class is cyt c
3 (∼13 kDa)
(Figure 5) from Desulfovibrio, which acts as a natural electron acceptor and donor
in hydrogenases
and ferredoxins.
214
The overall protein
fold containing two β-sheets and three to five α-helices
is conserved among the known structures of cyts c
3 as well as the orientation of the four hemes which are
located in close proximity to each other, with each of the heme planes
being nearly perpendicular to the others.
88
Each heme displays a distinct reduction potential spanning a range
from −200 to −400 mV.
215−219
Cyt c
555.1,
also known as cyt c
7 (∼9 kDa, 70
amino acids), from Desulfuromonas acetoxidans is another class III cyt c that contains
three
hemes.
220
These proteins have been proposed
to be involved in ET to elemental sulfur as well as in the coupled
oxidation of acetate and dissimilatory reduction of Fe(III) and Mn(IV)
as an energy source in these bacteria.
221
In cyt c
7, two of the hemes have a reduction
potential of −177 mV and the third heme has a reduction potential
of −102 mV.
222
Class IV cyts c fall into the category of large
molar mass (∼35–40 kDa) cytochromes that contain other
prosthetic groups in addition to c-type hemes such
as flavocytochromes c and cyts cd.
152
One example of class IV cyts c is revealed by the X-ray structure of the photosynthetic
reaction center (RC) from Rhodpseudomonas viridis, where light energy is harvested
and converted to chemically useful
energy. The cyt c in the RC consists of four c-type heme moieties covalently bound
to subunit C of the
RC. Three of the hemes have His/Met axial ligation, while the fourth
heme is bis-His-ligated. The four hemes are oriented in two types
of pairs. The porphyrin planes of hemes I/III and II/IV are orientated
parallel to each other, while the porphyrin planes of each pair of
hemes are mutually perpendicular to each pair’s porphyrin planes
(Figure 5).
223
Cyt c
554 is another tetraheme cytochrome
that is involved in the ET pathway of the biological nitrogen cycle
in the oxidation of ammonia in Nitrosomonas europaea.
122,224
This family of cytochromes does not fall
into either class III or class IV cytochromes and has been proposed
to belong to a class of its own. A pair of electrons are passed from
hydroxylamine oxidoreductase (HAO) to two molecules of cyt c
554 upon oxidation of hydroxylamine to nitrite.
One of the hemes is HS, and the other three are 6cLS with reduction
potentials of +47, +47, −147, and −276 mV, respectively.
Porphyrin planes of hemes III and IV are oriented almost perpendicular
to each other, while the heme pairs I/III and II/IV have parallel
orientation (Figure 5). The sets of parallel
hemes overlap at an edge, and such heme orientation has been observed
in HAO and cyt c nitrite reductase.
Cyt f is a high-potential (Table 2) electron
acceptor of the chloroplast cyt b
6
f complex involved in oxygenic photosynthesis
by passing electrons from photosystem II to photosystem I of the RC.
123,225
Cyt f accepts electrons from a Rieske-type iron–sulfur
cluster and passes electrons to the copper protein plastocyanin. Cyt f consists of
two domains primarily of β-sheets and
is anchored to the membrane by a transmembrane segment, while most
of the protein is located on the lumen side of the thylakoid membrane.
The heme is also located on the lumen side at the interface of the
two domains and is covalently attached to the protein via the signature
sequence of cyts c, -Cys-Xxx-Xxx-Cys-His-. The β-sheet
fold has not been observed in any other families of cytochromes and
is thus unique to cyts f. Intriguingly, this family
of cytochromes also contains an unusual second axial ligation to the
heme iron, an N-terminal −NH2 group of a Tyr residue
(Figure 5).
Quite uniquely, the only
exception to the bis-Cys covalent attachment
of the c-type hemes via the conserved -Cys-Xxx-Xxx-Cys-His-
motif in cyt c is found in eukaryotes from the phylum
Euglenozoa, including trypanosome and Leishmania parasites. In the mitochondrial cyt
c of these
organisms, the heme is attached to the protein via a single Cys residue
from the heme binding motif -Ala (Ala/Gly)-Gln-Cys-His-.
226−228
2.3.3
Conformational Changes in Class I Cytochromes c Induced by Changes in the Heme Oxidation
State
Many structural studies have been undertaken to determine whether
there is any effect on the protein structure associated with different
oxidation states of the heme iron. These studies include X-ray and
NMR structures of oxidized and reduced cyts c from
various sources,
229−235
which indicate that the oxidation state of the heme iron has a minimal
effect on the tertiary structures of the proteins (Figure 6). The major changes are
observed in the conformation
of some amino acid residues located close to the heme pocket. Among
these residues, Asn52, Tyr67, Thr78, and a conserved water (wat166)
molecule show maximal changes in conformations depending on the oxidation
state of the heme iron. These conserved residues,
236
along with the conserved water molecule, the axial ligand
Met80, and heme propionate 7, form a hydrogen-bonding network around
the heme site. The high-resolution X-ray structure of yeast iso-1-cytochrome c shows
that in the reduced state the heme is significantly
distorted from planarity into a saddle shape. The degree of heme distortion
in the oxidized state is even more pronounced compared to that of
the reduced state, suggesting that the planarity of the heme group
is dependent on the oxidation state of the iron. The major change
in the bond length of the heme iron ligands is observed in the case
of axial Met80, which increases from 2.35 to 2.43 Å in going
from the reduced to the oxidized state. On the contrary, the other
axial ligand, His18, shows a minute change of 0.02 Å, from 1.99
to 2.01 Å.
230
Figure 6
Overall structural overlay
of the reduced (cyan, PDB ID 1YCC) and oxidized (orange,
PDB ID 2YCC)
iso-1-cyt c (left). A close look at the heme site
and the nearby residues is shown on the right along with some hydrogen
bond interactions.
In the reduced state
of iso-1-cytochrome c, the
conserved water molecule is hydrogen bonded to Asn52, Tyr67, and Thr78
(Figure 6). Upon oxidation wat166 undergoes
a 1.7 Å displacement toward the heme, which results in the loss
of the hydrogen bond to Asn52, but interactions with Tyr67 and Thr78
are retained. Figure 6 shows an overlay of
the residues near the heme pocket between the reduced and oxidized
states of iso-1-cytochrome c.
87
Further analysis suggested that wat166 plays a key
role in stabilizing
both oxidation states of the heme iron by reorienting the dipole moment,
by changing the heme iron–wat166 distance, and by variations
in the nearby H-bonding network. Another noticeable change is observed
in the H-bonding between a conserved water, wat121, and heme propionate
7. In the reduced state, wat121 and Trp59 are hydrogen-bonded to O1A
and the O2A oxygen of propionate 7, respectively. In the oxidized
state, interaction between Trp59 and O2A of the heme propionate weakens,
while that of O2A and the conserved Gly41 increases. Additionally,
wat121 moves by 0.5 Å and causes a bifurcated hydrogen bond between
both O1A and O2A of the propionate.
230
Thus,
it appears that there are three major regions that show significant
changes in conformation between the two oxidation states: heme propionate
7, wat166, and Met80. A conserved region that does not show mobility
between oxidation states is the region encompassing residues 73–80
in iso-1-cytochrome c, which is linked to the three
major regions of conformation change through Thr78. On the basis of
this observation, it has been suggested that region 73–80 acts
as a contact point with redox partners and triggers the necessary
conformational changes in other parts of the protein that are required
to stabilize both oxidation states of cyt c.
230
A contrasting observation from NMR studies
is that wat166 moves 3.7 Å away from the heme iron when going
from the reduced to the oxidized state, rather than moving toward
the heme iron.
237,238
Similar to the changes
of heme propionate observed in eukaryotes,
cyts c
2(160,239−242) and c
6(220,243,244) from some prokaryotes also display conformational
changes in the heme propionate between the reduced and oxidized states
of the protein. In the cases of cyt c
H (reduces methanol oxidase in methylotropic bacteria) from Methylobacterium extorquens
and cyt c
552(245−247) (electron donor to a ba
3–cytochrome c oxidase) from T. thermophilus, there is no conserved water molecule
in the heme pocket, suggesting that the water-mediated H-bonding network
is not a critical requirement for ET.
248−250
2.3.4
Cytochromes c as Redox
Partners to Other Enzymes
In the following sections we summarize
some specific examples of native enzymes that use cyts c as the native electron donor
for performing various biochemical
processes.
2.3.4.1
Cytochrome c as a Redox
Partner to Cytochrome c Peroxidases
Cytochrome c peroxidases (CcPs) are a family of enzymes
that catalyze the conversion of H2O2 to water
and are found in both eukaryotes and prokaryotes. Eukaryotic CcPs are located in the
inner mitochondrial membrane and
contain a single heme cofactor, heme b, while prokaryotic
CcPs are located in the periplasmic space and contain
two covalently bound c-type hemes,
251,252
one of which is a low-potential (lp) heme and the other is a high-potential
(hp) heme. In general, the physiological electron donors to bacterial
CcPs are monoheme cyts c, although
other donors such as azurin (Az) or pseudoazurin have also been found
in some bacteria.
253
The hp heme is located
at the C-terminal domain and has a more positive reduction potential
than cyt c as it accepts electrons from cyt c. The reduction potential for the hp
heme varies depending
on the organism; e.g., the Ps. aeruginosa CcP hp site has a reduction potential of
+320 mV,
163
the Rb. capsulatus CcP hp site a reduction potential of +270 mV,
254
and the N. europaea CcP hp site a reduction potential of +130 mV.
154
The electrons are then transferred from the
hp heme to the lp heme of CcP. In some organisms,
e.g., Ps. aeruginosa and Rb. capsulatus, the hp heme should be in the ferrous
state for the enzyme to be active,
254,255
whereas in
other cases the enzyme is fully functional even with the ferric state
of the hp heme, e.g., in N. europaea.
154
The axial ligands for the hp heme
are a His and a Met, similar to most c-type cytochromes.
The lp heme is the site for H2O2 reduction.
It is located at the N-terminal domain and has two His residues as
axial ligands. The lp heme also displays a wide range of reduction
potentials from as low as −330 mV in Ps. aeruginosa(163) to as high as +70 mV in N.
europaea CcP.
154
Electron transfer between the hp and lp hemes, which are
10 Å apart, is thought to occur through tunneling.
255
Cyts c interact with
CcP at a small surface patch of the enzyme which
has a hydrophobic center and a charged periphery.
256
The small size of the surface patch suggests that the interaction
of the enzyme with the electron donor is transient, but at the same
time is highly specific, which ensures complex formation due to desolvation
of the surface waters and binding of cyt c. The charged
periphery has been shown to be important to guide the donor toward
the surface site, but it does not increase the specificity of the
interactions or the ET rate.
257
Mutagenesis
studies in Rb. capsulatus CcP have shown that the interface at which the enzyme interacts
with its electron donor cyt c
2 involves
nonspecific salt bridge interactions, as the extent of the interaction
is dependent on the ionic strength of the solution.
258
In contrast, in Ps. nautica CcP, the interaction surface between the enzyme
and the electron donor cyt c is highly hydrophobic
on the basis of studies which showed that the enzyme was active across
a wide range of ionic strength of the solution.
259
Studies from Pa. denitrificans CcP have shown that two molecules of horse heart
cyt c are able to bind to the enzyme surface.
260
Binding of an “active” and “waiting”
cyt c in a ternary complex with the enzyme has been
proposed to improve the ET rate. Structural studies of Pa. denitrificans CcP with
the monoheme
cyt c has shown that the heme of the donor binds
above the hp heme of CcP, while the two molecules
of horse heart cyts c bind between the two hemes
of the enzyme.
261
2.3.4.2
Cytochrome c as a Redox
Partner to Denitrifying Enzymes: Nitrite, Nitric Oxide, and Nitrous
Oxide Reductases
Denitrification is a stepwise process in
the biological nitrogen cycle where nitrogen oxides act as electron
acceptors and are sequentially reduced from nitrate to nitrite, nitrite
to nitric oxide, nitric oxide to nitrous oxide, and finally nitrous
oxide to nitrogen. These four steps of the nitrogen cycle are catalyzed
by a diverse family of enzymes, viz., nitrate reductase, nitrite reductase,
nitric oxide reductase, and nitrous oxide reductase, all of which
are induced under anoxic conditions.
262−264
Various cyt c domains act as electron donors in the denitrification
process. Reduction of nitrite to nitric oxide is catalyzed by one
of the two structurally diverse enzymes that also have different catalytic
sites: (a) cytochrome cd
1 nitrite reductase
(cyt cd
1 NiR)
265,266
and (b) multicopper nitrite reductase (CuNiR).
267,268
Cyt cd
1 NiRs are periplasmic, soluble
heterodimeric enzymes containing an ET cyt c domain
and a catalytic cyt d
1 domain in each
subunit, while multicopper nitrite reductases are homotrimeric enzymes
containing T1Cu as an ET site and T2Cu as a catalytic site. Cyts c
552 are the putative electron donors of cyt cd
1.
269
Multicopper
nitrite reductases have cupredoxin-like folds and use azurins and
pseudoazurins as their biological redox partner, and as such are not
expected to have cyt c domains. Contrary to this
expectation, two instances have been found where a fusion of multicopper
nitrite reductase and cyt c domains was discovered
in the genomes of Chromobacterium violaceum and Bdellovibrio bacteriovorus, where
in both cases the cytochrome c domain is present
at the end of a ∼500-residue-long sequence.
79
These cyt c sequences are similar to those
of the caa
3 oxidase sequences.
Nitric
oxide reductases (NORs) are integral membrane proteins that catalyze
the two-electron reduction of nitric oxide to nitrous reductase.
270,271
A recent X-ray structure of the Gram-negative bacterium Ps. aeruginosa cyt c-dependent
NOR
(cNOR) (Figure 7) shows that the enzyme consists
of two subunits.
272
The NorB subunit is
the transmembrane subunit and contains the binuclear active site consisting
of an HS heme b
3 and a nonheme iron (FeB) site. It also houses an LS ET cofactor heme b. NorC is
a membrane-anchored cyt c and contains
a c-type heme. Electrons are received from cyt c
552 or azurin to the heme c, which then passes the electrons to LS heme b and
then to HS heme b
3 of the catalytic binuclear
active site. The reduction potentials are +310, +345, +60, and +320
mV for heme c, heme b, heme b
3, and the FeB sites, respectively.
162
Figure 7
X-ray structure of cyt c-dependent NOR
(cNOR)
(PDB ID 3O0R) from Ps. aeruginosa.
2.3.4.3
Cytochromes c as Redox
Partners to Molybdenum-Containing Enzymes
Mononuclear molybdenum-containing
enzymes constitute a group of enzymes that catalyze a diverse set
of reactions and are found in both eukaryotes and prokaryotes.
273,274
The general function of these groups of enzymes is to catalytically
transfer an oxygen atom to and from a biological donor or acceptor
molecule, and these enzymes are thus referred to as molybdenum oxotransferases.
These enzymes possess a Mo=O unit at their active site and
an unusual pterin cofactor which coordinates to the metal via its
dithiolene ligand moiety. These Mo-containing enzymes are generally
classified into three families depending on their structures and the
reactions that they catalyze. The first one is xanthine oxidase from
cow’s milk, which has an LMoVIOS(OH) (L = pterin)
catalytic core and generally catalyzes the hydroxylation of carbon
centers. The second family includes sulfite oxidase from avian or
mammalian liver with a core coordination consisting of an LMoVIO2(S–Cys) moiety that
catalyzes the transfer
of an oxygen atom to or from the substrate’s lone pair of electrons.
The third family of oxotransferases shows diversity in both structure
and function and uses two pterin ligands instead of only one used
by the first two classes. The reaction occurs at the active site core
containing L2MoVIO(X), where X could be Ser
as in DMSO reductase or Cys as in assimilatory nitrate reductase.
Xanthine oxidases have been reported to be coexpressed with three
cyt c domains in Bradyrhizobium japonicum, Bordetella bronchiseptica, Ps. aeruginosa,
and Ps. putida; however, the exact cause of this association is not well understood
as these enzymes use flavins as their redox partners.
79
Sulfite oxidase catalyzes the oxidation of sulfite to sulfate
using 2 equiv of oxidized cyt c as physiological
oxidizing substrates (Scheme 1).
273
The molybdenum is reduced from the VI to the
IV oxidation state, and the reducing equivalents are then transferred
sequentially to the cyt c in the oxidative half-reaction.
The assimilatory nitrate reductases (NRs) are found in algae, bacteria,
and higher plants which uptake and utilize nitrate.
273
These enzymes contain a cyt b
557 and flavin adenine dinucleotide (FAD) in addition to the Mo center.
Electrons flow from FAD to cyt b
557 to
the Mo center under physiological conditions. The midpoint reduction
potentials for FAD and cyt b
557 from Chlorella NR have been determined to be −288
and −164 mV, respectively.
189,190,275
The Mo center displays reduction potentials of +15
mV for the MoVI/V couple and −25 mV for the MoV/IV couple. These reduction potentials
indicate that the physiological
direction of electron flow is thermodynamically favorable. The cyt b
557 domain of NR is homologous to the mammalian
cyt b
5, yeast flavo-cyt b
2, and cyt b domain of sulfite oxidase.
276
Scheme 1
Scheme Showing the Oxidation of Sulfite
to Sulfate by Cyt c in Sulfite Oxidase
Reprinted from ref (273). Copyright 1996 American
Chemical Society.
The DMSO reductase family
consists of a number of enzymes from
bacterial and archaeal sources that display remarkable sequence similarity.
Respiratory DMSO reductases are periplasmic and use membrane-anchored
multiheme cyts c as electron donors that transfer
electrons from the quinine pool to the periplasmic space. These cytochromes
are about 400 amino acids long and are encoded in the same operon
as the enzyme. In some γ-proteobacteria, the tetraheme cyts c occur as a fusion to
the C-terminal cyt c binding domain of the enzyme. On the other hand, in some ε-proteobacteria
single-domain cyts c have been coexpressed with the
DMSO reductase and act as electron donors to the enzyme. Nonetheless,
the cyt c sequences from both types of proteobacteria
are clustered together, suggesting that even though the mechanism
of ET is different, they are functionally similar.
79
Even though these ET proteins in DMSO reductases are referred
to as cyts c because they contain c-type hemes, their structural folds do not fall
into the uniquely
defined category of cyt c folds as mentioned in section 2.3.2.
2.3.4.4
Cytochrome c as a Redox
Partner to Alcohol Dehydrogenase
The type II quinohemoprotein
alcohol dehydrogenases are periplasmic enzymes that catalyze the oxidation
of alcohols to aldehydes and transfer electrons from substrate alcohols
first to the pyrroloquinoline quinone (PQQ) cofactor, which subsequently
transfers electrons to an internal heme group that is found in a cyt c domain.
277
This cyt c domain of about 100 residues contains three α-helices
in the core cytochrome domain and is similar to the cyt c domain in p-cresol methylhydroxylase
(PCMH) from Ps. putida(278) and the
cyt c551i from Pa. denitrificans.
279
2.3.4.5
Involvement
of Cytochromes c in Photosynthetic Systems
Photosynthesis involves the conversion
of light energy to useful chemical forms of energy, which is accomplished
by two large membrane protein complexes, photosystem I (PSI) and photosystem
II (PSII).
280
The catalytic cores of the
two PSs are referred to as the reaction centers, which have [4Fe−4S]
clusters and quinines as terminal electron acceptors for PSI and PSII,
respectively. Like algae and higher plants, cyanobacteria also use
PSI and PSII to convert light energy to chemical forms by producing
oxygen from water oxidation. Even though cyanobacteria have a bis-His-coordinated
PS-C550 cyt subunit in their PSII, apparently there is
no redox role of this cytochrome.
281,282
Being located
at the lumenal surface of the enzyme, PS-C550 cytochrome
acts as an insulator of the catalytic core from reductive attack and
contributes to structural stabilization of the complex.
283,284
The low midpoint reduction potentials of the soluble protein from
−250 to −314 mV exclude any redox role of this class
of cytochromes.
285−288
When complexed with PSII, more positive values of reduction potentials
have been determined.
288,289
A reduction potential of +200
mV in PS-C550 cytochrome from Thermosynechococcus
elongates has recently been reported,
185
which suggests a possible role of this cytochrome
in ET in PSII, despite a long distance (∼22 Å) between
the PS-C550 cytochrome and its nearest redox center, the
Mn4Ca cluster.
290
In cyanobacteria,
cyt c
6 is known to act interchangeably
with the copper protein plastocyanin as an electron donor to PSI,
depending on the availability of copper,
291−293
while in higher plants plastocyanin is the exclusive electron donor.
On the basis of this observation, it has been proposed that cyt c
6 is the older ancestor, which has been replaced
by plastocyanin during evolution due to the shortage of iron in the
environment.
294
Another cytochrome,
cyt c
M, is found
exclusively in cyanobacteria, but its role is ambiguous. It has been
shown to be expressed under stress-induced conditions such as intense
light or cold temperatures where the expression of both cyt c
6 and plastocyanin is suppressed.
295
Thus, it would be tempting to believe that
cyt c
M is a third electron donor to PSI
in cyanobacteria under stress conditions, but experimental evidence
goes against this hypothesis.
296
2.3.4.6
Cytochrome c as a Single-Domain
Oxygen Binding Protein
Sphaeroides heme protein (SHP) is
an unusual c-type cytochrome which was discovered
in Rb. sphaeroides.
183
SHP (∼12 kDa) has a single HS heme with a reduction
potential of −22 mV and an unusual His/Asn axial heme coordination
in the oxidized form. SHP is spectroscopically distinct from cyts c′, which also have
a HS heme. SHP was shown to bind
oxygen transiently during slow auto-oxidation of the heme. The Asn
axial ligand was shown to swing away upon reduction of the heme or
binding of small molecules such as cyanide or nitric oxide. The distal
pocket of SHP shows marked resemblance to other heme proteins that
bind gaseous molecules.
297
It has been
suggested that SHP could be involved as a terminal electron acceptor
in an ET pathway to reduce small ligands such as peroxide or hydroxylamine.
297
2.3.5
Cytochrome c Domains in
Magnetotactic Bacteria
Magnetotactic bacteria consist of
a group of taxonomically and physiologically diverse bacteria that
can align themselves with the geomagnetic field.
298
The unique property of these bacteria is due to the presence
of iron-rich crystals inside their lipid vesicles forming an organelle,
referred to as the magnetosome. From sequence analysis, three proteins,
MamE, MamP, and MamT, in the Gram-negative bacterium Magnetospirillum magneticum AMB-1
that contribute
to the formation of the magnetosome have been discovered to contain
a double -Cys-Xxx-Xxx-Cys-His- motif, characteristic of cyts c.
186
All three proteins were
expressed and purified in E. coli.
Subsequent characterization of these proteins confirmed that MamE,
MamP, and MamT indeed belong to c-type cytochromes,
and they have been designated as “magnetochromes”. Midpoint
reduction potentials were determined to be −76 and −32
mV for MamP and MamE, respectively. The presence of cyts c proteins in magnetotactic
bacteria is intriguing and suggests that
these proteins take part in ET, although the exact nature of their
ET partners is not known. It has been hypothesized that the magnetochromes
can either donate electrons to Fe(III) and participate in magnetite
[mixture of Fe(III) and Fe(II)] formation or accept electrons from
magnetite to maintain a redox balance, or they can act as redox buffers
to maintain a proper ratio of maghemeite (all ferric irons) and magnetite.
2.3.6
Multiheme Cytochromes c
Multiheme cyts c occur as both soluble
and membrane-anchored ET proteins in many enzymes across diverse functionalities.
79,299
Triheme cyts c
7 from Geobacter sulfurreducens and Dm. acetoxidans are involved in ET for Fe(III)
respiration,
207,300−303
although their exact roles are not known. These proteins have conserved
secondary structural elements consisting of double-stranded β-sheet
at the N-terminus followed by several α-helices. The protein
displays a miniaturized version of the cyt c
3 fold where heme II and the surrounding protein environment
are missing (Figure 8). The arrangement of
hemes is conserved in cyts c
7 in terms
of the distances between heme iron atoms and the angles between heme
planes. Hemes I and IV are almost parallel to each other and are mutually
perpendicular to heme III, which is in close contact with hemes I
and IV. NMR and docking experiments suggest that heme IV is the region
of interaction with similar physiological partners, while the other
interacting partner would most likely interact through the region
near hemes I and III. Such differences in interaction surfaces might
play a role in choosing the right redox partners to perform different
physiological functions.
Figure 8
(A) X-ray structure of triheme cyt c
7 (PDB ID 1HH5). All the hemes are bis-His-ligated. Cyt c
7 is a minimized version of cyt c
3 where heme II is missing. (B) Spatial arrangement of the
four hemes
in flavocytochrome c
3 fumarate reductase
(PDB ID IQO8). The heme irons of the heme pair II and III are in close proximity
at 9 Å from each other, and the heme edges are 4 Å away.
An unusual triheme cyt c is DsrJ from the purple
sulfur bacterium Allochromatium vinosum that is a part of a complex involved in sulfur
metabolism.
182,304
Sequence analysis suggested the presence of three distinct c-type hemes containing
bis-His, His/Met, and a very unusual
His/Cys axial ligation, respectively. Subsequent cloning and expression
of DsrJ in E. coli indeed confirmed
the presence of three hemes, and EPR data showed the presence of partial
His/Cys coordination to one of the hemes (His/Met is another possibility).
From redox titrations, reduction potentials of the hemes were determined
to be −20, −200, and −220 mV, respectively. Although
the exact role of DsrJ is still unknown, its involvement in catalytic
functions rather than in ET has been hypothesized.
182
Other examples of multiheme cyts c include a tetraheme
cyt c (NapC) involved in nitrate reductase from Pa. denitrificans,
305
an
Fe(III)-induced tetraheme flavocytochrome c
3 (Ifc3)
306
in fumarate reductase
(Fcc3) from Sh. frigidimarina, an HAO containing eight heme groups for hydroxylamine
oxidation
in N. europaea,
307
and a pentaheme nitrite reductase (NrfA) for nitrite reduction
in Sulfurospirillum deleyianum.
308,309
A periplasmic flavocytochrome c
3 which
is an isozyme of the soluble Fcc3 is also induced by Fe(III).
310−312
The X-ray structure of this protein shows that the tetraheme arrangement
in Fcc3 includes an intriguing heme pair where the two
irons are only 9 Å from one another and the closest heme edges
are within 4 Å (Figure 8).
The four
hemes from Ifc3 and Fcc3 can be
superimposed on four of the eight hemes in HAO.
307
All four hemes of Ifc3 overlay on four of the
hemes from the pentaheme NrfA,
308
and all
five hemes from NrfA overlay on five of the HAO hemes. Lastly, two
hemes from Ifc3 overlay on two of the four hemes of cyt c
554(122) from N. europaea, all four hemes of which overlay on four
hemes from HAO. Despite such similarities in heme arrangement, there
is no resemblance in the primary sequence of these enzymes. Nevertheless,
such similar heme arrangements in these proteins suggest that they
share a common ancestor, but have evolved divergently to perform four
different reactions, viz., Fe(III) reduction, fumarate reduction,
hydroxylamine reduction, and nitrite reduction.
313
Some membrane-bound multiheme cytochromes, belonging to
the NapC/NirT family, contain four heme binding sequences that have
evolved due to gene duplication of diheme domains.
314
In NapC and CymA all four hemes are 6cLS with bis-His axial
ligation and display reduction potentials of +10 and −235 mV,
respectively.
305,313
Sh. oneidensis MR-1 is a facultative
anaerobe that is capable of using many terminal electron acceptors
such as DMSO or metal oxides such as ferrihydrite and manganese dioxide
outside the outer cell membrane, accepting electrons from the quinol
pool and the tetraheme protein CymA.
317−325
Electron transfer in Sh. oneidensis MR-1 is facilitated by two periplasmic decaheme
cyts c, DmsE, which supplies electrons to DMSO, and MtrA, which is involved
in ET to metal oxides (Figure 9). Both of these
decaheme proteins have been proposed to be involved in a long-range
ET across a ∼300 Å “gap”
326
(∼230 Å periplasmic gap and ∼40–70
Å thick outer membrane). Using protein film voltammetry, a potential
window between −90 and −360 mV and an ET rate of ∼122
mV s–1 were measured for DmsE at pH 6.
315
The measured reduction potential window for
DmsE is shifted ∼100 mV lower than what was observed in MtrA,
327−329
although the rate of ET is similar in both proteins. Although the
MtrA and DmsE families of decaheme proteins facilitate long-range
ET in Sh. oneidensis, it is not clear
how ET is feasible across a 300 Å gap, especially given the fact
that MtrA spans only 105 Å in length.
330
Clearly, the arrangement of hemes must play a crucial role; however,
the exact mechanism of this ET process is yet to be determined. A
recent NMR study proposes the presence of two independent redox pathways
by which the ET occurs from the cytoplasm to electron acceptors on
the cell surface across the periplasmic gap in MtrA,
331
one involving small tetraheme cyt c (STC)
and the other involving FccA (flavocytochrome c).
Both of these proteins interact with their redox partners CymA (donor)
and MtrA (acceptor) through a single heme and show a large dissociation
constant for protein–protein complex formation. Together, these
facts suggest that a stable multiprotein redox complex spanning the
periplasmic space does not exist. Instead, ET across the periplasmic
gap is facilitated through the formation of transient protein–protein
redox complexes.
Figure 9
(A) Schematic model for DMSO reduction by DmsEFAB and
iron reduction
by MtrABC(DEF). Flows of electrons are shown with arrows. DmsE and
MtrA(D) are proposed to accept electrons from the menaquinone pool
via CymA. Multiheme groups in CymA, MtrACDF, and DmsE are shown. IM
= inner membrane, and OM = outer membrane. (B) “Staggered-cross”
orientation of the hemes in outer membrane decaheme MtrF (PDB ID 3PMQ). Heme numbering
is shown as Roman numerals, heme–iron distances are shown in
orange, and distances between heme edges are shown in blue. (A) Reprinted
with permission from ref (315). Copyright 2012 Biochemical Society. (B) Adapted from
ref (316) Copyright 2011 National
Academy of Sciences.
MtrF is a decaheme c-type cytochrome found
in
the outer membrane of Sh. oneidensis MR-1 (Figure 9) which has been proposed to
transfer electrons to solid substrates through the outer membrane,
like its homologue MtrC, with the help of periplasmic MtrA and a membrane
barrel protein, MtrE, that facilitates ET by forming contact between
MtrA and MtrF.
332,333
A recent crystal structure of
MtrF shows that the protein consists of four domains, domains I and
III containing β-sheets and domains II and IV being α-helices.
316
The arrangement of the 10 bis-His-ligated hemes
is like a “staggered cross” where four hemes (I, II,
VI, VII) are almost coplanar with each other and are almost perpendicular
to a group of three hemes (III, IV, V and VIII, IX, X) that are parallel
to each other (Figure 9).
The reduction
potentials of the hemes in MtrF lie in the range
of 0 to −312 mV as determined by both solvated and protein
film voltammetry. Unfortunately, reduction potentials of individual
hemes have not been possible to assign due to their similar chemical
nature. Molecular dynamics simulations show an almost symmetrical
free energy profile for ET. Additionally, the computed reorganization
energy range from 0.75 to 1.1 eV is consistent for partially solvent
exposed heme cofactors capable of overcoming the energy barrier for
ET.
334,335
Further molecular details of ET in MtrF
are unknown.
Multiheme cyts c also act as ET
agents in the
Fe(III)-respiring genus Shewanella.
299
However, due to the fact that Fe(III) is soluble
only at pH < 2, these organisms face the problem of moving electrons
from the cytoplasm across two cell membranes to the extracellular
space to reduce the insoluble extracellular species. It has been proposed
that these organisms circumvent this problem by employing a number
of tetraheme and decaheme cyts c which act as “wires”
to transfer electrons between the inner and outer membranes.
313,336
For tetraheme cyts c
3, hemes I
and
III are covalently attached to the protein segment by a conserved
-Cys-Xxx-Xxx-Cys-His- sequence, while hemes II and IV are linked to
the protein with the two Cys residues occurring in the sequence -Cys-Xxx-Xxx-Xxx-Xxx-Cys-His-.
337,338
Although the overall orientation of hemes is conserved, the order
of heme oxidation varies from source to source.
217,339,340
The hemes in cyts c
3 display redox cooperativity, such that the reduction
potential of one heme is dependent on the oxidation state of the other
hemes. The reduction potentials of the hemes in cyts c
3 are also dependent on the pH, called the redox-Bohr
effect,
340−342
due to the interactions of the heme propionates
in the H-bonding network and/or electrostatic interactions with the
residues in the vicinity.
341,343−345
Type I cyts c
3 are soluble, periplasmic
proteins and contain a patch of positively charged residues close
to heme IV which have been proposed to interact with its partners.
346
This class of cyts c
3 mediate ET between periplasmic hydrogenases and transmembrane ET
complexes where the electron acceptor is thought to be type II cyts c
3. Type II cyts c
3 are structurally similar to those of type I, but lack the lysine
patch.
347
It was proposed that type I cyts c
3 receive electrons from hydrogenase and deliver
them to type II cyts c
3. Recent experimental
evidence shows that these two types of cyts c
3 form a complex with each other and are indeed physiological
partners, but type I cyts c
3 transfer
only one electron to type II cyts c
3 in
solution.
348,349
2.3.7
Cytochromes b
5
Cyts b
5 are ET hemoproteins
containing bis-His-ligated b-type hemes and are found
ubiquitously in bacteria, fungi, plants, and animals. Cyts b
5 display reduction potentials that span a range
of ∼400 mV.
350−353
Mitochondrial and microsomal cyts b
5 are membrane-bound, while those from bacteria and erythrocytes are
soluble. In addition, there are various cyt b
5-like proteins that act as redox partners in various enzymes
such as flavocytochrome b
2 (l-lactate dehydrogenase), sulfite oxidase, assimilatory nitrate reductase,
and cyt b
5/acyl lipid desaturase fusion
proteins. The structures of cyts b
5 from
various sources reveal that there are two hydrophobic cores on each
side of a β-sheet that belong to the α + β class
(Figure 10).
350
The
larger hydrophobic core contains the heme binding crevice, while the
smaller hydrophobic core is proposed to have only a structural role.
About 3% of deoxyhemoglobin in adults is oxidized to inactive methemoglobin.
354
Soluble cyts b
5 in erythrocytes reduce methemoglobin to the functionally reduced
deoxy form that binds oxygen. For this reaction electrons are transferred
from NADH to methemoglobin via NADH cyt b
5 reductase and cyt b
5.
355
Microsomal cyts b
5 are found
in the membranes of the endoplasmic reticulum anchored to the membrane
by a stretch of 22 hydrophobic residues.
353
Microsomal cyts b
5 are known to function
by transferring electrons in fatty acid desaturation, cholesterol
biosynthesis, and hydroxylation reactions involving cyts P450.
356
Figure 10
Schematic representation of the X-ray structure
of bovine cyt b
5 that belongs to the α
+ β class
(PDB ID 1CYO). Two hydrophobic core domains, six α-helices, five β-strands,
and 6c bis-His-ligated heme are shown. Adapted from ref (357). Copyright 2011 American
Chemical Society.
Two different forms
of cyt b
5 have
been detected in rat hepatocyte; one is associated with the membrane
of the endoplasmic reticulum (microsomal, or Mc, cyt b
5), while the other is anchored to the outer membrane
of liver mitochondria (OM cyt b
5).
358−362
These two types of cyt b
5 display a
reduction potential difference of 100 mV (−107 mV for OM cyt b
5,
187,363
−7 mV for Mc
cyt b
5).
180
The rat OM cyt b
5 is involved in the
reduction of cytosolic ascorbate radical using NADH as the electron
source.
364,365
The mammalian OM cyt b
5 and Mc cyt b
5 have three different
domains, an N-terminal hydrophilic domain that binds the heme, an
intermediate hydrophobic domain, and a C-terminal hydrophilic domain.
The N-terminal heme binding domains for both types of cyts b
5 have very similar structural folds consisting
of six α-helices and four β-strands. The heme is bound
in a pocket formed by four α-helices and a β-sheet formed
by two of the β-strands.
141,366
Studies relating to
the complex formation and ET rates between cyts b
5 and its redox partners suggest that the nature of interactions
between two proteins is primarily electrostatic and the heme edges
of cyts b
5 make contacts with electron
donors and acceptors.
350
Within this general
area, there are multiple overlapping sites with which cyts b
5 interact with its various partners.
A gene encoding a cyt b
5-type heme
from the protozoan intestinal parasite Giardia lamblia was recently cloned into E.
coli as
a soluble protein.
367
The spectroscopic
properties of this cloned cyt b
5 are similar
to those of the microsomal cyts b
5, and
homology modeling suggests the presence of a bis-His-ligated heme.
Residues near the heme binding core from Giardia cyt b
5 are comprised of charged amino
acids and differ from those of other families of cyt b
5. The reduction potential of the heme was determined
to be −165 mV.
2.3.7.1
Heme Orientation Isomers
in Cytochromes b
5
Solution NMR
studies of the soluble
fragment of cyt b
5 suggested the coexistence
of two different species that contained two orientation isomers (forms
A and B, Figure 11) of heme that are related
by a 180° rotation about an axis through the heme α,γ-meso-carbon atoms.
368−372
Figure 11
Two orientation isomers (A and B forms) of heme observed in solution
studies of the soluble fragment of cyt b
5. The two isomers are related by a 180° rotation around the
α,γ-meso-carbon atoms.
The relative population of the two isoforms A and
B varies from
species to species. In bovine and rabbit, the A/B ratio is ∼10/1,
177,368,370,373
20/1 in chicken cyt b
5,
374
6/4 in rat Mc cyt b
5,
374
and 1/1 in the OM cyt b
5.
375
Even though reconstitution
of apo cyt b
5 with heme resulted in the
initial formation of a 1/1 ratio of species A and B, they converted
back to the proportion found in the thermodynamically stable native
state after some time.
370,373
Reduction potentials
of +0.8 and −26.2 mV were calculated for isoforms A and B,
respectively, from spectroelectrochemical titrations.
177
Interaction between the 2-vinyl group and side
chains of residues 23 and 25 was initially thought to be the driving
factor that dictated the heme orientation isomers.
368,374,376
This theory was disputed in
later studies.
375
It is now generally accepted
that the heme itself can adapt to the surrounding environment by a
rotation of the porphyrin plane around an axis perpendicular to the
iron, which is proposed to be the determining factor that caused the
different heme orientation in species A and B.
376−378
Several studies have indicated that residue His39 is the major determining
factor of the electronic state that orients the molecular orbitals
for easy ET through the exposed pyrrole ring III and meso-carbon heme edge.
370,379,380
2.3.8
Cytochrome b
562
Cyt b
562 is a 106-residue monomeric
heme protein of unknown function found in the periplasm of E. coli. It is a four-helix
bundle protein where
the helices are oriented antiparallel to each other (Figure 12).
381,382
Figure 12
NMR structure of the
antiparallel four-helix bundle cyt b
562 (PDB ID 1QPU). His/Met axial coordination to the heme
iron is shown.
The protein has a noncovalently
bound 6cLS heme with His102 and
Met7 axial ligands, even though this protein is structurally homologous
to cyt c′ that contains a covalently bound
5cHS c-type heme. In the oxidized unfolded state,
the heme of cyt b
562 is converted to 5cHS
with His102 as the only axial ligand.
383
The folding properties of this protein are highly dependent on the
pH. At pH 7 the reduction potential of the heme in the folded state
is 189 mV, while that of the unfolded state is −150 mV, suggesting
that the reduced state has a greater driving force for folding than
the oxidized state.
176,384−387
Unfolding of the oxidized state of the protein occurs reversibly
with a midpoint GuHCl concentration of 1.8 M, while the reduced state
shows irreversible unfolding at >5 M GuHCl due to heme dissociation.
Folding of the reduced state has been shown to be triggered by photoinduced
ET to the oxidized form of the protein under 2–3 M GuHCl concentrations.
A folding rate of 5 μs was extrapolated in the absence of denaturant,
which is similar to the intrachain diffusion time scale of the polypeptide.
388
2.4
Designed Cytochromes
In addition
to studying native systems by a top-down approach, in recent decades,
many groups have adopted a bottom-up approach of building minimal
functional proteins that mimic natural ones. The theoretical simplicity
and ubiquity of cytochromes has made them appealing targets for design,
and a number of artificial cytochrome-mimicking proteins have been
engineered, with varying levels of sophistication. In this issue of Chemical Reviews,
Pecoraro and co-workers give a thorough
review of protein design strategies and successes, including designed
heme ET proteins.
3000
Here, we give a brief
account focusing on the redox properties of designed 6-coordinate
heme proteins mimicking ET cytochromes.
2.4.1
Designed
Cytochromes in de Novo Designed
Protein Scaffolds
Two de novo heme proteins called VAVH25(S–S) and retro(S–S)
389
were designed to bind heme in a bis-His coordination, by strategically
engineering His residues into the de novo cystine-cross-linked, homodimeric
four-helix bundle called α2.
390−392
Both sequences yielded artificial cytochromes with dissociation
constants for heme in the submicromolar range, and spectroscopic properties
of these proteins were consistent with low-spin bisimidazole-ligated
heme, with reduction potentials of −170 and −220 mV
for each of the proteins. Although these potentials are nearly unchanged
from the potentials of bisimidazole heme in aqueous solution, the
success of incorporation demonstrated the power of rational de novo
design and set the stage for rapid development of more complex and
nativelike structures. Using an alternative tetrameric protein scaffold,
consisting of two pairs of disulfide linked α-helices, a series
of proteins mimicking the heme b domain of cytochrome bc
1 were also designed by strategic placement
of histidine residues. The designed proteins incorporated either two
or four hemes per bundle,
393
with potentials
of the individual sites reported to range from −230 to −80
mV in the tetraheme construct. More impressively, the sites showed
cooperative redox properties, with the presence of a second ferric
heme site proposed to raise the potential of the first by ∼115
mV through electrostatic interactions (vide infra).
393,394
In a systematic study of the electronic properties of this scaffold,
varying the heme, pH, and local charge could achieve a potential range
of 435 mV (−265 to +170 mV),
395
over
half the 800 mV range covered by native cytochromes. Interestingly,
investigation of the more natural mutation of one of the His ligands
with a Met resulted in only a 30 mV increase in reduction potential,
and substitution of heme b with heme c gave no significant change.
396
Rational
mutagenesis of several core residues, as well as incorporation of
helix–turn–helix and asymmetric disulfide bonds, further
improved the structural rigidity and uniqueness of the designed scaffolds.
397,398
Subsequently, this maquette system was extended in a variety of
ways to achieve coupling to electrode surfaces,
399
incorporation of non-natural amino acid ligands,
400
and binding of two different hemes—which
mimics the structure of ba
3 oxidases.
401
Particularly exciting is the demonstration
of coupling of ET and protonation of carboxylate residues on the protein,
402−404
which is relevant for understanding and engineering proton pumping.
On the basis of recent developments in structural understanding
of cytochrome bc
1 and improvements in
computational modeling, Ghirlanda et al. investigated designing a
more structurally unique mimic of the bc
1 complex. The structure of the heme b binding portion
of bc
1 was modeled as a coiled coil, and
secondary coordination sphere interactions to the coordinating histidines,
such as conserved Gly, Thr, and Ala residues, were added to stabilize
the orientation of the His ligand and tune its electronic properties
(Figure 13A).
405
The potentials were measured by cyclic voltammetry (CV) as −76
and −124 mV in the oxidative and reductive directions, respectively,
at pH 8, significantly higher than the potential of aqueous bisimidazole
heme and earlier bis-His-ligated designed proteins. The hysteresis
in the potentials is attributed to conformational reorganization of
the ligating His residues between the oxidized and reduced forms.
The model was further improved by linking and expression as a single
chain for more efficient structure determination studies,
408
as well as incorporation into a membrane.
409
Figure 13
Structural models of designed cytochrome models
in de novo scaffolds.
(A) A design model for a homodimeric four-helix tetraheme binding
protein inspired by cyt bc
1. Remade from
coordinates courtesy of G. Ghirlanda and W. F. DeGrado.
405
(B) Schematic representation of monomeric four-α-helix
maquettes used to mimic ET cytochromes. Reprinted with permission
from ref (406). Copyright
2013 Macmillan Publishers Ltd. (C) Crystal structure of Co(II) mimichrome
IV (PDB 1PYZ).
407
Most recently, Dutton and co-workers have reported the design
and
thorough characterization of a monomeric, single-chain four-α-helix
bundle maquette protein, which can bind up to two hemes (Figure 13B). It is particularly
noteworthy for the subject
of this review that the redox properties of this scaffold as a function
of charge distribution were systematically analyzed. By raising the
total charge uniformly from −16 to +11, the reduction potential
of both hemes changed from −290 to −150 mV, as expected.
Furthermore, the potentials of the hemes could be changed individually
by only increasing the charge at one end of the protein; the potentials
of the individual hemes were −240 and −150 mV. Finally,
it was demonstrated that the reduced negatively charged protein could
transfer an electron to native cytochrome c with
rate constants approaching those of native photosynthetic and respiratory
electron transport chains. Such a single-chain four-helix bundle was
also used to build an artificial oxygen binding cytochrome c with an intramolecular
B-type ET heme with a 60 mV lower
reduction potential, mimicking a natural ET chain.
410
More rational computational protein design algorithms
have also
been brought to bear on the de novo design of artificial cytochromes.
Xu and Farid used the algorithm named CORE
411
to design a nativelike four (27 amino acid)-helix bundle that binds
two to four hemes in a bis-His fashion.
412
The α-helical character was confirmed by circular dichroism
(CD), and the binding affinity for the first 2 equiv was determined
to be in the micromolar range, while, due to negative cooperativity,
the remaining sites had K
d > 3 mM.
The
measured potentials for the diheme and tetraheme protein were −133
to −91 and −190 to −0110 mV, respectively.
While the rationally guided design strategies described above have
been very successful, the lack of a priori knowledge about the necessary
structural features for design of functional metalloproteins limits
the scope of sequence and structure space that is probed by the strategy.
As a complementary approach, Hecht and co-workers have utilized a
semirational “binary code” library generation method
to produce 15 74-residue sequences that formed helical bundles and
bound heme,
413
one with submicromolar affinity.
Extending this scaffold further produced five 102-residue sequences
with higher stabilities and more “nativelike” structures.
414
Analysis of a handful of these proteins revealed
spectroscopic features typical of low-spin heme proteins and reduction
potentials ranging from −112 to −176 mV.
415
Furthermore, it was demonstrated that at least
one construct was electrically competent on an electrode.
416
A similar semirational combinatorial approach
was utilized by Haehnel and co-workers, who combined it with template-assisted
synthetic protein (TASP) methods, in which two sets of antiparallel
helices are templated onto a polypeptide ring, to design and screen
an impressive library of 399 cytochrome b mimicking
four-helix bundles.
417,418
Using a colorimetric screen,
the potentials were estimated to range from −170 to −90
mV. It was also demonstrated that the proteins could be incorporated
onto electrodes
419,420
and achieved estimated ET rate
constants comparable to those of native cytochromes.
A number
of smaller, water-soluble peptide-based cytochrome mimics
have also been developed, utilizing one or two short α-helical
peptides. Two groups independently developed heme compounds with covalently
attached, short α-helix-forming peptides, with His ligands.
In one case, peptide-sandwiched mesoheme (PSM) compounds were prepared
by covalently attaching a 12-mer peptide to each of the two propionate
groups of the heme via amide bonds with lysine groups on the peptide.
421
Although the helicity of the free peptide was
low, upon ligatation of the heme, the helicity was seen by CD to increase
to ∼50%, and the electronic spectra were consistent with bis-His
heme ligation, similar to b-type cytochromes.
421,422
Further work suggested that aromatic side chain interaction with
the heme, such as Phe and Trp, improves helix stability and heme binding,
423
and covalent linkage of the peptide termini
via disulfide bonds resulted in further stabilization.
424
Studies of the redox properties of a PSM and
a mutant with an Ala to Trp mutation, (called PSMW), highlight
the importance of stability in determining reduction potential, with
more stable helix binding in PSMW lowering the reduction
potential by 56 mV (−281 to −337 mV), due to the increased
ability of the His ligands to stabilize the Fe(III) state.
425
The authors propose that this effect may also
explain the difference in potential between mitochondrial and microsomal
cyts b
5.
Similarly, short α-helical
peptides, based on the heme binding
peptide fragment of myoglobin, have been covalently attached to deuterohem
by a similar amide-bond attachment strategy, yielding compounds known
as mimochromes.
426
It is noteworthy that
the peptides retained their α-helical character even in the
absence of heme binding.
426,427
The stability of the
model was further improved in later revisions by enhancing the intramolecular
interpeptide interactions through extending the peptide (mimochrome
II)
428
or rational mutagenesis (mimochrome
IV).
429
A crystal structure of the Co(II)
derivative of mimochrome IV has been obtained and substantiates the
designed structure (Figure 13C).
407
The reduction potential of Fe mimochrome (IV)
at pH 7 is −80 mV, though it exhibits strong pH dependence
over the range of pH from 2 to 10 (∼+30 to −170 mV).
429
The low-pH dependence is attributed to the
His ligands unbinding from the heme, while the high-pH transition
is proposed to be caused by deprotonation of a nearby arginine; however,
this is surprising due to the 4 orders of magnitude higher apparent
acidity and requires further investigation to be proven. Still, it
is exciting that this simple mimic is well folded enough to be crystallized
and has a potential in the range of those of native cytochromes.
Intermediate between these covalently attached heme–peptide
models and full polyhelical bundles described above, heme protein
complexes consisting of heme ligated by designed short peptides that
are not covalently attached have also been developed.
430−434
Studies on the binding of a variety 15-mer peptides showed a strong
correlation between peptide–heme affinity and reduction potential
(−304 to −218 mV), with lower potentials for more stable
complexes, consistent with the results of studies on PSMs.
425,431
The overall low potential was attributed to the inability of the
small peptides to reduce the strong dielectric constant of the solvent,
as native proteins do (vide infra). To further improve the stability,
two peptides were covalently linked at both ends by disulfide ligands,
resulting in a series of cyclic dipeptide heme binding motifs, with
reduction potentials ranging from −215 to −252 mV.
433
Interestingly, in a step away from the
helix bundle paradigm, Isogai
and co-workers were able to rationally design a series of de novo
proteins that would fold into a globin fold, but with only ∼25%
sequence identity to sperm whale myoglobin.
435,436
Although the proteins were designed for a 5-coordinate myoglobin-like
heme binding site, the resulting proteins were consistent with 6-coordinate
bis-His-ligated heme. In these scaffolds, the reduction potential
was in the range of −170 to −200 mV, similar to that
of aqueous bis-Im heme, which was attributed to higher solvent access
to the heme due to the molten-globular state of the proteins. This
was further supported by the re-engineering of a nonheme globin protein,
phycocyanin, into a heme binding protein (vide infra), which had a
more unique, hydrophobic, and nativelike core structure and 50 mV
higher reduction potential.
437
2.4.2
Designed Cytochromes in Natural Scaffolds
In addition
to designing scaffolds for cytochromes de novo, an
appealing alternative strategy is to make use of the diversity of
natural proteins as scaffolds. One of the most straightforward approaches
is to convert a non-cytochrome heme protein into a cytochrome by site-directed
mutagenesis. Along these lines, various myoglobins have also been
redesigned into bis-His cytochrome-like proteins, similar to b
5, by mutating the valine near the heme at position
E11 to histidine (Figure 14A).
438−440
The spectroscopic features of reduced and oxidized forms of these
mutants are consistent with low-spin bis-His-ligated heme, and the
crystal structure confirms the ligation.
440
The mutations result in a 170 mV decrease in the reduction potential
of myoglobin, from ∼60 to ∼−110 mV.
Figure 14
Structural
models of designed cytochrome models in native scaffolds.
(A) X-ray crystallographic model of a pig myoglobin designed to have
cytochrome-like bis-His ligation (PDB ID 1MNI).
440
(B) Molecular
dynamics model of a histidine mutant of the membrane protein, glycophorin
A, designed to bind heme in a cytochrome-like manner.
441
Coordinates provided by courtesy of G. Ghirlanda.
Similarly, natural nonheme proteins
can also be designed to bind
heme in a manner consistent with the cytochrome binding motif. As
briefly mentioned above, Isogai and co-workers introduced two histidines
into the natural nonheme plant globin phycocyanin
437
to generate a heme binding site. Although the protein was
designed as a myoglobin mimic, the spectral features were consistent
with low-spin bis-His coordination, similar to that of cytochromes b, with a one-electron
reduction potential of −120
mV.
Heme binding sites have also similarly been designed into
native
α-helical bundle proteins that do not have native heme binding
sites. Starting with the DNA binding protein rop, a specific bis-His
heme binding protein was designed by removing surface histidines and
introducing two internal histidine residues.
442
An alternative His/Met binding mode was also investigated.
443
Both proteins displayed electronic spectra
characteristic of low-spin heme, with reduction potentials of −155
and −88 mV, respectively. A cytochrome-like heme binding site
was also designed into the transmembrane protein glycophorin A (Figure 14B).
441,444
Each of the proteins
bound heme with submicromolar affinity, and the presence of aromatic
phenylalanine residues near the heme lowered the reduction potential
from −128 to −172 mV.
2.4.3
Conversion
of One Cytochrome Type to Another
In addition to designing
cytochrome sites in non-cytochrome proteins,
several groups have investigated the conversion of one type of cytochrome
into another.
445−449
Conversion of c-type to b-type
cytochrome has been achieved in cytochrome c
552 by removing the Cys residues in the -Cys-Xxx-Xxx-Cys-His-
heme binding motif with the Cys11Ala/Cys14Ala double mutation.
447
CD and NMR spectra confirmed that the structure
of the protein and heme site was maintained.
447,450
However, it was found that the removal of the c-type heme binding motif destabilized
the protein toward chemical
and thermal denaturation. While the electron-withdrawing potential
of the vinyl groups of heme b relative to the thioether
groups of heme c would be expected to raise the potential,
80
the resulting protein had a reduction potential
of 170 mV, 75 mV lower than that of the wild type, suggesting that
the electronic structure of the porphyrin is not the major determinant
of the reduction potential difference between cytochromes c and b (discussed in section
2.5).
Conversion from cyt b
562 to c-type heme has been achieved
by introducing the conserved -Cys-Xxx-Xxx-Cys-His- motif into the
wild-type protein by means of two mutations (Arg98Cys and Tyr101Cys).
449,451
The resulting c-type cytochrome displayed enhanced
stability toward chemical denaturants, maintaining the same protein
fold and axial His ligation. c-type heme attachment
has also been achieved in cytochrome b
5 by introducing a surface cysteine residue with the Asn57Cys mutation.
448
The resulting holoprotein was isolated in four
forms, with distinct forms of heme, one of which contained covalently
attached heme and a hemochrome α-band at 553 nm, intermediate
between those of b-type (556 nm) and c-type (551 nm) heme, suggesting the presence
of a single c-type thioether linkage. NMR further confirmed the stereochemical
nature of this linkage, and the protein displayed a reduction potential
of −19 mV, 23 mV lower than that of the wild-type b
5.
2.5
Structural Features Controlling
the Redox
Chemistry of Cytochromes
Being involved in distinct ET pathways,
each cytochrome has evolved its ET properties to match those of its
redox partners. Therefore, reduction potentials of cytochromes span
a range of almost 1 V, from −475 mV in bacterioferritin from Azotobacter vinelandii(192,452)
to +450 mV in the heme c of diheme cytochrome c peroxidase of N. europaea(153,154)
vs the SHE.
453
Through a variety of studies, many properties have been found to
be important in determining the redox properties of heme proteins.
As expected, the molecules in the first coordination sphere of the
iron, namely, the four pyrrole groups of the porphyrin and the axially
coordinating residues, are important in determining the baseline reduction
potential, as they interact directly with the iron center. These interactions
are also fine-tuned by the secondary coordination sphere—chemical
moieties that interact with the primary coordination sphere ligands
and adjust their properties. Secondary coordination sphere interactions,
such as H-bonding, can cause strengthening or weakening of ligand–metal
interactions. The overall charge as well as the electrostatic environment
of the metal center, which is determined by the surrounding charge,
dipole distribution, and solvent accessibility, also critically modulates
the redox properties.
2.5.1
Role of the Heme Type
It is known
that c-type hemes tend to be found in cytochromes
with more extreme potentials (much lower or much higher) relative
to b-type hemes; however, it is unclear whether a
direct causative relationship exists. One way to probe the role of
the heme type in a way that is less dependent on other factors is
to replace the heme in one protein with another. In studies of the
de novo designed four-helix bundles, the strongest effect on reduction
potential was attributed to the nature of the heme,
395
though unnatural hemes were used in the study. In the more
natural protein cases, several groups have interconverted b- and c-type hemes.
445−449
It has been found, however, that this interconversion shows little
inherent effect on the reduction potential
447,448
with no clear trend. For instance, it was found that converting
the c-type heme in cyt c
552 into a b-type heme by mutating away the conserved
Cys residues lowered the reduction potential by 75 mV.
447
On the other hand, introducing a thioether
bond between heme in cytochrome b
5 and
the protein, and therefore converting the b-type
heme into a c-type heme, also lowered the potential
by 23 mV.
448
It is clear that the choice
of heme c over heme b has little
effect on the reduction potential, and other effects, such as structural
changes or solvent accessibility, may play a bigger role.
If
the choice of heme c or heme b does
not play a significant role in determining the reduction potentials
of cytochromes, one may wonder why organisms invest in the energetically
expensive process of synthesizing c-type linkages.
Though the exact reason that Nature has chosen c-type
hemes in certain proteins remains to be fully understood, several
hypotheses have been proposed.
454−456
It is suggested that multiheme
cytochromes, such as c
3, with largely
exposed hemes in close proximity may utilize heme anchoring as a strategy
to ensure stable heme binding in the absence of well-defined hydrophobic
interactions.
457
Similarly, the high-potential
cyts c, with His/Met coordination, may use covalent
anchoring as a strategy to prevent heme dissociation due to the relatively
weaker binding of methionine to ferric heme.
457
Alternatively, it is proposed that covalent heme attachment may
help in protein folding and stability
454,456
or may strengthen
the Fe–His bond and help maintain a low-spin state.
456
Regardless, the choice of heme c over heme b likely does not itself directly tune
the reduction potential in a significant or consistent way, but may
allow the protein greater flexibility in achieving other functionality
and tuning the potential by other means, such as solvent accessibility.
In addition to hemes b and c,
heme a is a unique heme used for ET in enzymes such
as heme copper oxidases (HCOs). The heme incorporates two unique peripheral
structural features, namely, a hydroxyethylfarnesyl group and a formyl
group, and these functional groups have been suggested to play a role
in tuning the reduction potential of the heme. While heme a has been replaced with
other hemes in a native system,
458
detailed studies of how this substitution affects
the redox chemistry of the protein have not been reported. Using their
de novo designed scaffold (vide supra), Gibney and co-workers
459
have studied the redox properties of hemes a and b, as well as diacetyl heme, and
found that the electron-withdrawing acyl groups increased the potential
by ∼160 mV. This effect can be fully accounted for by the 200-fold
lower affinity of the ligands for the oxidized form over the reduced
form of the heme, and it is proposed that the hydrophobic farnesyl
group serves to anchor the heme stably in the protein
460
to compensate for the lower affinity of the
ferric state.
2.5.2
Role of Ligands
In addition to
the heme type, the identity of the axial ligands sets the baseline
for the reduction potentials of cytochromes.
457
Between the two most common ligands (His and Met), it has been found
that the Met ligation generally raises the potential of the heme by
∼100–150 mV, relative to the His ligation.
461−463
However, contrary to this theory, early work by Sligar and co-workers
found that redesigning bis-His cyt b
5 into
a His/Met cyt lowered the reduction potential by
∼240 mV. This opposite change in the reduction potential was
attributed to the change in the spin state of the heme, from low-spin
bis-His to high-spin His/Met cyt.
464
More
consistent with the theory, it was demonstrated that conversion of
bis-His to His/Met ligation in cyts c
3 results in a reduction potential increase of 160–180 mV.
192
Similarly, using a proteolytic fragment of
cyt c, it was found that methionine ligation in cyts c contributes 130 mV to the energy.
386
Conversely, a 105 mV drop in the reduction potential was
observed when the methionine in cytochrome c
551 was replaced with a histidine.
463
Interestingly, Hay and Wydrzynski
462
observed
a 260 mV decrease in reduction potential when they substituted the
native Met ligand in cyt b
562 with His,
yielding a typical bis-His cyt. This decrease is greater than ∼150
mV, and the authors attribute it to destabilization of the fold and
increased solvent exposure, which is known to significantly lower
the potential (vide infra). In contrast, an Arg98Cys and His102Met
double mutant of the same protein, cyt b
562, shows 6cLS bis-Met axial ligation at low pH, with a reduction potential
of +440 mV, ∼180 mV higher than that of native His/Met cyt b
562.
465
The authors
note that the effect of bis-Met ligation is likely to be slightly
higher at ∼200 mV, as they expect the c-type
thioether heme linkage to lower the potential. The stereochemical
alignment of the axial methionine ligands results in an almost axial
symmetry of the heme, caused by a 110° change in the torsion
angle between the sulfur lone pairs.
466
The reduction potential of this protein is 665 mV higher than that
of the only other known bis-Met axially ligated heme system in bacterioferritin
(−225 mV)
176
in which the ground
state of the oxidized form of the heme is highly rhombic in nature.
120,121,467
Therefore, factors other than
the differences in the ligand coordination are most likely to be involved
to account for the reduction potential difference.
78
In general, all else being equal, the preference of soft
methionine thioether for the softer ferrous heme over the harder ferric
heme contributes to a ∼100–200 mV increase in reduction
potential over His ligation.
2.5.3
Role
of the Protein Environment
2.5.3.1
Solvent Exposure
Consistently,
one of the most important factors in raising the reduction potentials
of the cytochromes is the extent of heme burial in the protein or,
alternatively, the extent of solvent exposure of the heme.
178,187,386,457,468−473
The basis for this effect lies in the lower dielectric constant
of proteins relative to aqueous solution, which significantly destabilizes
the charged ferric site over the neutral ferrous state of the heme.
For instance, Tezkan et al. estimated that solvent exclusion accounts
for ∼240 mV of the potential increase in cyt c.
386
Similarly, in a thorough computational
study of heme proteins spanning an 800 mV range of potentials, Zheng
and Gunner identified that heme solvent exclusion accounts for ∼20%
of the reduction potential difference between proteins.
457
Interestingly, the same study found less correlation
between the reduction potentials and the remaining individual factors
or energy terms, yet the computation was able to faithfully reproduce
and account for heme protein potentials over an 800 mV range. This
study elegantly demonstrates that the reduction potential is determined
by an intricate balance of numerous factors of comparable energy.
2.5.3.2
Secondary Coordination Sphere of the Ligand
Although the nature of the ligand itself determines primary interaction
energies with the heme, and therefore is the primary determinant of
the reduction potential, the electronic character of the ligand can
be further modulated by secondary noncovalent interactions, such as
hydrogen bonds. These so-called secondary coordination sphere effects
have been shown to be influential in determining the potentials of
a number of heme proteins, including cytochromes.
230,472,474−477
For instance, in cyt c in particular, Bowman et
al. demonstrated that strengthening the hydrogen bond between the
proximal His ligand and a backbone carbonyl through peripheral mutations
resulted in an almost 100 mV decrease in the reduction potential,
attributable to increased imidazolate character.
474
Similarly, Berguis et al. show in three different mutants
of yeast iso-1-cyt c that a disruption of the hydrogen
bond from tyrosine 67 to the methionine ligand consistently decreases
the potential by 56 mV, due to an increase in electron density on
the Met sulfur, stabilizing the ferric form of the heme,
230,476
and Ye et al., found that the presence of hydrogen bonds between
Gln64 and the axial Met ligand in Ps. aeruginosa and Hydrogenobacter thermophilus
cyt c lowered the potential by 15–30 mV.
477
In addition, aromatic interactions with the
axial ligand have also been implicated in tuning the heme reduction
potentials. For instance, it was shown that Tyr43, which interacts
with the π system of His 34, contributed a ∼35–45
mV decrease in reduction potential.
478
Therefore,
although the identity of the ligand is a primary determinant of the
reduction potential of the heme, the secondary coordination sphere
interactions with it also play a role of similar magnitude in determining
the reduction potential.
2.5.3.3
Local Charges and Electrostatics
Another important means by which cytochromes have been found to
modulate
their reduction potentials is through the judicious use of charge
and electrostatic interactions. For instance, by comparison and selective
mutagenesis of the structurally homologous cyts c
6 and c
6A, it was demonstrated
that the interaction of the positive dipole of the amide group of
a carefully positioned glutamine (residues 52 in c
6 and 51 in c
6A) with the
heme is a strategy used by Nature to raise the reduction potential
by ∼100 mV.
479
Similarly, Lett et
al. observed an increase in the reduction potential of cytochrome c by 117 mV through
the Tyr48Lys mutation.
480
Tyr48 is involved in a H-bonding interaction with a heme
propionate, and it is likely that introduction of lysine at this position
stabilizes the propionate negative charge and destabilizes the ferric
heme state. It has also been shown that replacement of a neutral residue
in contact with the heme in myoglobin with a polar or negatively charged
residue can reduce the potential by up to 200 mV.
481
Furthermore, a library screen of cytochrome b
562 mutants at four residues near the heme binding site
identified mutations that could gradually tune the potential over
a 160 mV range.
482
Even relatively distant
surface electrostatic interactions have been shown to control the
redox function of cytochromes.
483
These
reports demonstrate the critical role of local charge in determining
the reduction potential of the heme. In general, negative local charges
stabilize the ferric state and lower the reduction potential, and
the magnitude of this effect can be comparable to that of ligand substitution
or ligand secondary coordination sphere effects.
In addition
to charge interactions, more subtle effects such as electrostatic
interactions can also play an important role in determining redox
properties. As discussed in section 5.2.2 below,
a conserved aromatic residue in cyt b
6
f is found to be in contact with the heme f at position 4, and the identity of the
aromatic residue
differs between cyanobacteria and algae. Interconversion between Phe
and Trp at this position accounts for about half of the 70 mV difference
between these proteins.
161
The origin of
this effect is attributed to differential interaction of the side
chain electrostatic potentials with the porphyrin π system and
the Fe orbitals. A similar effect has also been reported in cyt c
3, where a phenylalanine in contact with heme
I is proposed to maintain its low potential by a π–π
interaction with the porphyrin π system.
484
Since many charged residues around the heme, such
as Glu, Asp,
Lys, and Arg, as well as the heme propionate group itself, can be
protonated or deprotonated depending on the pK
a values of the residues and pH of the solution, protonation
states of these groups will affect the reduction potential of the
heme by preferentially stabilizing one redox state over the other.
Therefore, the pH of the solution can have significant effects on
the reduction potentials in various cytochromes.
342,485−490
For example, protonation of a heme propionate in cyt c contributed an increase of
65 mV to the reduction potential.
485
Similar effects of 60 and 75 mV have been reported
in cyt c
551(491,492) and in cyt b
559,
490
respectively. In cyt c
2, pH-dependent
reduction potentials covered a range of ∼150 mV, between pH
4 and pH 10.
493
In their de novo designed
maquette, Dutton and co-workers observed a 210 mV range of reduction
potentials over a pH range of 3.5–10, and such a change was
attributed to the involvement of Glu residues near the heme site.
494
Furthermore, the role of the propionate charge
has been investigated specifically by studies in which the carboxylate
groups have been neutralized to their ester form. An increase of reduction
potential by ∼60 mV was reported,
495,496
consistent with those obtained from the studies described above.
A special case of the effect of local charges on reduction potential
is the cooperativity between nearby hemes in multiheme cytochromes.
497
It is known that the presence of multiple hemes
in various oxidation states greatly affects the macroscopic or observable
reduction potentials of the hemes. For instance, it has been demonstrated
in multiheme cyt c
3 that the interaction
energy between hemes can shift the reduction potential by 50–60
mV.
498−500
It is suggested that this effect may be
mediated by electrostatic interactions also involving local aromatic
groups.
484
The cooperativity between hemes
in multiheme cytochromes is proposed to be a major factor in their
reduction potential regulation.
In cyt c
3, the redox-Bohr effect can
result in pK
a differences of up to 2.8
pH units, and the coupling between protonation has been linked to
cooperativity between the hemes, resulting in concerted two-ET steps.
340,501,502
On the other hand, the pH-dependent
reduction potential difference, over a range of 10 pH units, can be
∼200 mV.
503
Such property is crucial
for proper charge separation to generate a promotive force that drives
ATP synthesis.
343,504
Similarly, this coupling of
proton and ET plays a key role in the proton pumping mechanism of
cytochrome c oxidase. Although there are several
proposed mechanisms, they share the common theme that proton uptake
to the heme sites and release into the P-side of the membrane are
driven by charge compensation during ET events from the low-spin to
high-spin heme.
505−507
It is clear that local electrostatic interactions
at heme redox centers are of immense physiological importance.
2.5.3.4
Heme Distortion/Ruffling
Another
significant contributor to heme redox properties is the plasticity
of the heme. It is now well-known that heme distortion or ruffling
plays an important role in the electronic sturcture of the porphyrins,
508,509
due to decreased delocalization of the π electrons.
510−516
While the phenomenon has been described in many heme proteins, including
cytochromes,
512,513,515,517,518
thorough investigation of how it affects redox properties is limited.
Recently, Marletta and co-workers demonstrated that protein-induced
heme distortion can account for up to a 170 mV increase in potential
in the heme nitric oxide/oxygen binding protein.
513
Furthermore, a basic computational model was implemented
by Senge and co-workers, and it was estimated that porphyrin distortion
can account for 54 mV of the difference between hemes in a bacterial
tetraheme cytochrome.
519
Further investigation
is needed to gain a more detailed understanding of the role of heme
distortion in the redox properties of typical cytochromes.
3
Fe–S Redox Centers in Electron Transfer
Processes
3.1
Introduction to Fe–S Redox Centers
Fe–S proteins are among the oldest metalloproteins on earth.
The early atmosphere, under which both sulfur and iron were abundant,
enabled the spontaneous assembly of these two elements into clusters,
mainly containing four iron and four sulfur atoms.
91,520
Early life took advantage of the redox properties of these clusters
and used them as redox centers. Despite the later shift to a more
oxidizing environment on earth, the established Fe–S proteins
continued to be used as electron carriers. Thus, these proteins are
found ubiquitously throughout all kingdoms of life and play roles
in crucial processes such as photosynthesis and respiration. The wide
range of reduction potentials these proteins can accommodate and their
diverse structural motifs allow them to interact with different redox
partners, acting as electron carriers in a variety of biological processes.
91−93
The Fe–S proteins were first discovered in the 1960s
on the basis of their unique g = 1.9 EPR signal that
appears upon reduction and was not observed before for any metalloproteins.
9000−9002
This discovery was aided by the abundance of these proteins and
their unique spectral features and often highly charged nature, which
made them easier to purify and analyze. Studies of these proteins
were further facilitated by advances in molecular biology and recombinant
protein expression, allowing the use of site-directed mutagenesis
to unravel important features of these proteins and their function.
While the Fe–S centers are well-known for their function
as electron carriers, they are also known to be involved in the active
sites of many enzymes, performing several functions
521
such as reduction of disulfide bonds and initiation or
stabilization of radical chain reactions,
523,525,529
or serving as Lewis acids.
524,527,543
In addition, the Fe–S
centers can simply function as structural elements that stabilize
the protein or another active site in the protein.
523,525,527,529
Furthermore, the sensitivity of the Fe–S centers to an oxidative
environment and their range of redox states make them good candidates
for sensing oxidative and metal stress and balancing the oxidative
homeostasis of the cells.
93,525,526,527,530−533,543
Functions in DNA repair have also been reported for several Fe–S
proteins.
532,534
Recently, a function for Fe-S
proteins has been proposed in formation of FemoCo cluster.
522
Finally it has been shown that the Fe–S
proteins can be used as a storage for sulfur or iron.
529,532
This review focuses exclusively on the ET function of the Fe–S
proteins.
3.2
Classification of Fe–S Redox Centers
and Their General Features
The Fe–S clusters are often
classified on the basis of the number of iron and sulfur atoms in
the cluster, as suggested by the Nomenclature Committee of the International
Union of Biochemistry (IUB) in 1989.
535
In this convention, the elements of the core cluster (iron and inorganic
sulfur atoms) are placed in brackets with the oxidized level of the
core cluster shown as a superscript outside the brackets (e.g., [2Fe–2S]2+). A comma
or a slash in the superscript can show multiple
possible oxidation states. A more expanded notation can be used to
show the ligands and the overall charge of the whole cluster, including
those ligands. Another common classification of Fe–S clusters,
which is used in this review, is based on the protein type. This scheme
classifies the Fe–S centers on the basis of not only the number
of iron and sulfur atoms but also certain structural motifs and spectroscopic
and electrochemical properties. In this classification, the Fe–S
proteins are divided into major groups as follows: rubredoxins (Rd’s;
[1Fe–4S]), ferredoxins (low-potential [2Fe–2S], [4Fe–4S],
[3Fe–4S], [3Fe–4S][4Fe–4S], and [4Fe–4S][4Fe–4S]),
Rieske proteins (which are high-potential [2Fe–2S] proteins),
and high-potential iron–sulfur proteins (HiPIPs, which are
high-potential [4Fe–4S] proteins) (Table 3). In addition, we will also describe more
complex Fe–S proteins
that contain multiple Fe–S cofactors or Fe–S cofactors
coupled with other cofactors, such as heme.
92,93,523,526,529,536−540
Table 3
Classification of Fe–S Proteins
Though certain structural elements may differ
between them, members
of each class of Fe–S proteins usually consist of a common
structural motif. Between classes the overall structure is distinct.
Despite these overall structural differences, however, the geometries
of the Fe–S clusters are quite similar, especially within each
cluster class. The iron cofactor has a distorted tetrahedral geometry
in almost all the Fe–S proteins. In the case of proteins with
more than one iron, the S–S distance is usually 1.3 times longer
than the Fe–Fe distance.
523
Each
iron atom is coordinated by a total of four ligands, typically cysteine
or inorganic sulfurs, although other ligands have been observed. For
instance, in Rieske proteins, two cysteine ligands have been replaced
with histidines. In some [3Fe–4S] clusters, an aspartate serves
as a ligand to iron. In certain enzymes such as aconitase, a hydroxyl
group from the solvent is shown to be one of the ligands.
541
While the geometry of Fe and its coordinating
cysteine/sulfur ligands
is very similar in all Fe–S proteins, the amino acid sequences
and peptide motifs that accommodate these clusters differ significantly
even in a given class, resulting in further categorization of each
group. Interestingly, the ligands of the Fe–S proteins usually
reside within loop regions. This structural flexibility is important
in accommodating the geometric requirement of the Fe–S clusters
and thus minimizing the reorganization energy required for rapid ET.
The iron site has large spin-polarization effects, strong Fe–S
covalency, and spin coupling through inorganic sulfurs.
542
The strong covalency and the delocalization
features of Fe–S proteins result in a low reorganization energy,
mostly by lowering the inner sphere effects. Gas-phase DFT calculations
give the following reorganization energies for different Fe–S
proteins in vacuum: 0.41 eV (1Fe, Rd) < 0.45 eV (4Fe, HiPIP) <
0.64 eV (4Fe, Fd) < 0.83 eV (2Fe, Fd).
542
The sulfur atom has several advantages over other ligands
for coordinating
Fe: it can occupy 3d orbitals of the iron, while the effects of its
nuclear charge are not significant, and as a weak ligand, it can keep
iron in a high-spin state.
543
However,
it imparts an intrinsic instability to the cluster, as sulfur is vulnerable
to oxidation. Moreover, due to having a weak ligand, Fe in Fe-S clusters
is in a high spin state.
523
As a result,
the Fe–S clusters are usually very sensitive to oxidation,
hydroxylation, and other chemical modifications.
523
In fact, one of the characteristic features of Fe–S
clusters is their being “acid-labile”.
1,538
The protein provides a protective, hydrophobic environment around
the Fe–S clusters, excluding solvent and improving stability.
523
The Fe–S proteins have long been
the focus of bioinorganic
studies due to their rich electronic structure and magnetism. The
presence of iron as the core redox-active center provides researchers
with a wealth of techniques to investigate this site which are not
easily applicable to most other redox-active metals. A very intriguing
feature of the Fe–S proteins is the presence of mixed-valence
species, which have been the subjects of extensive investigations.
All common bioinorganic methods have been applied to study Fe–S
proteins, including EPR, electron–nuclear double resonance
(ENDOR)/electron spin echo envelope modulation (ESEEM), 1D and 2D
NMR, X-ray absorption spectroscopy (XAS) analysis, X-ray crystallography,
Mössbauer, and CD/magnetic circular dichroism (MCD). Information
can be deduced even with simple electronic absorption spectroscopy
techniques.
537,538,540
3.3
Biosynthesis of Fe–S Proteins
In
vitro studies have shown that the Fe–S proteins can be
reconstituted by addition of FeCl3 and Na2S
in a reductive environment.
539,544,545
The presence of iron and sulfur in the solution is sufficient for
formation of a [4Fe–4S] cluster.
543
Despite the straightforward in vitro assembly, the assembly of the
Fe–S clusters in vivo is a more precise and complex process.
Multiple experiments have been performed with the aim of elucidating
the exact mechanism of assembly of different Fe–S clusters,
and every year, new discoveries are made in this field. Nif, Isc,
and Suf cluster-binding systems are the most common systems involved
in in vivo assembly of Fe–S proteins.
520
These systems are abundant in different organisms, and many organisms
have more than one of them. Briefly, all of these systems require
a cysteine desulfurase to produce sulfur from l-cysteine,
a scaffold that plays the role of a carrier for the formation of the
cluster, and a carrier to transfer the cluster to the final protein.
The source of iron remains to be definitively elucidated. The Nif
system is dedicated to maturation of nitrogenase and was first found
in Azotobacter vinelandii. Isc and
Suf systems, in contrast, are more general, and homologues of these
systems are found in mitochondria and chloroplasts, respectively.
The two systems are conserved between bacteria and eukaryotes. The
Isc system utilizes five proteins: IscU that acts as a scaffold, IscS
that generates sulfur from cysteine, HscA/B that facilitates the transfer
of the cluster to the protein, and the ferredoxin. The Suf system
is composed of two subcomplexes: One is SufBCD that can bind to and
transfer the [4Fe–4S] cluster to proteins. In this subcomplex,
SufB acts as a scaffold, SufD is important for iron entry, and SufC
is an essential ATPase. The other is the SufSE subcomplex that acts
as a heterodimer and donates sulfur to the cluster. SufS is the major
component with cysteine desulfurase activity, and SufE enhances its
activity. Several classes of proteins are important in transferring
the cluster to the apoprotein, but the so-called A-type proteins are
the most common. Recently, members of cytosolic iron-sulfur cluster
assembly machinery have been found as main components of the Fe–S
biogenesis in cytosol. The Fe–S biogenesis is tightly regulated
and correlated to oxidative and metal stresses.
520,547−554
3.4
Native Fe–S Proteins
3.4.1
Rubredoxin
3.4.1.1
Structural Aspects
Rd is the
simplest among Fe–S proteins. It is a robust small protein
usually composed of 45–54 amino acids with a molar mass of
6–7 kDa mainly found in bacteria, archaea, and anaerobes. It
contains a monoiron center, coordinated by four cysteines from two
Cys-(Xxx)2-Cys-Gly segments, with a distorted tetrahedral
geometry (Figure 15).
555,556
Sequence alignment reveals that the four cysteine residues are conserved
in rubredoxins from different sources. Moreover, nearby glycine and
proline residues, several aromatic residues such as tyrosine, tryptophan,
and phenylalanine, and two charged lysine residues are conserved as
well. However, a novel rubredoxin has been identified in several members
of the Desulfovibrio genus, possessing
an N-terminal Cys-(Xxx)4-Cys segment.
557
Figure 15
Crystal structure of CpRd (PDB ID 1IRO) at 1.1 Å resolution.
(a) Overall fold of chain A of CpRd. The Fe(Cys)4 center is displayed as a ball-and-stick
representation. (b)
NH···S H-bond interactions around the Fe(Cys)4 center of CpRd. The side chains of
C6, C39, V8,
Y11, L41, and V44 are omitted for clarity. Color code: Fe, green;
C, cyan; S, yellow, O, red; N, blue.
Rubredoxin from mesophilic Cl. pasteurianum (CpRd) is among the most well studied
members of
the family,
556
and rubredoxin from hyperthermophilic
archaeon Pyrococcus furiosus (PfRd) is one of the most thermally stable proteins with
a melting temperature of 200 °C.
558
The overall fold of rubredoxin is composed of a three-strand antiparallel
β-sheet with a hydrophobic core and two loops containing the
coordinating cysteines with pseudo-2-fold symmetry (Figure 15). The loop carrying
ligands Cys6 and Cys39 (numbering
of CpRd), buried inside the protein, is more constrained
by the rigid aromatic core of the protein. In combination with a bulky
aliphatic residue (Ile/Leu/Val33), these conserved aromatic residues
contribute to the stabilization of the overall three-dimensional structure
as well as exclusion of water from the metal center.
559,560
Charged residues, mainly glutamate and aspartate, are distributed
over the surface and result in high solubility and a very acidic isoelectric
point of about 4. The metal binding site is close to the protein surface,
between the two binding loops, and metal incorporation contributes
to stabilization of the protein as well.
The two coordinating
loops exhibit a pseudo-2-fold symmetry about
the [Fe(Cys)4] center with six NH···S H-bonds
in a range of 3.5–3.9 Å. The Fe–S bond distances
to the buried Cys6 and Cys39 ligands are slightly longer (2.28–2.30
Å on the basis of three different rubredoxins) than those of
Cys9 and Cys42, which are close to the surface (2.25–2.26 Å).
This is possibly because Cys6 and Cys39 are involved in two H-bonds
with the backbone amides of Thr7/Val8 and Pro40/Leu41, respectively,
while Cys9 and Cys42 have only one H-bond donor each, from the backbone
amides of Tyr11 and Val44, respectively (numbering of CpRd, Figure 15b).
561,562
Nine sp3-hybridized C–H···S weak
hypervalent interactions are identified by 13C NMR in CpRd, which contribute to stabilization
of the protein as
well.
563,564
X-ray absorption near-edge spectral (XANES)
fitting of the oxidized forms of recombinant CpRd
at pH 8.0 gave a bond length of 2.27(1) Å for Fe(III)–S,
562
comparable to the average bond length of 2.26(3)
Å from crystal structures.
556
3.4.1.2
Function
The electron-rich iron
center of rubredoxin is redox-active, and its Fe(II)/Fe(III) couple
is involved in a variety of biological ET processes.
565
No significant structural changes are observed by NMR and
crystallographic studies when the ferric center is reduced. Slight
lengthening of the Fe–S bonds by an average of 0.096 Å
(CpRd),
566
0.033 Å
(PfRd),
555
or 0.012 Å
(Leu41Ala CpRd),
567
as
well as shortening of the cysteine involved in H-bonds has been observed,
consistent with the valence change of the metal center. DFT calculations
reveal that the Fe–S center of rubredoxin from Dv. vulgaris has a low reorganization
energy during
oxidation due to high Fe–S bond covalency and large electronic
relaxation, which makes it well suited for fast ET.
568
Rubredoxin from Ps. oleovorans (PoRd) forms an ET complex with rubredoxin reductase
in its physiological environment and provides a good system for studies
of interprotein ET. PoRd transfers electrons from
rubredoxin reductase to a membrane-bound ω-hydroxylase for aliphatic
and aromatic hydrocarbon oxidation. The ET from NADH to Rd is gated
by a rate-limiting adiabatic step preceding the ET step.
569−572
Similarly, rubredoxin from Ps. aeruginosa is involved in alkane oxidation by transferring
electrons from NAD(P)H
via NAD(P)H:rubredoxin reductase to the terminal electron acceptor.
573
FAD-dependent NAD(P)H:rubredoxin reductase
has been cocrystallized with RubA2(PA5350), an AlkG2-type rubredoxin
from Ps. aeruginosa closely related
to PfRd,
574
and diffracted
to 2.45 Å. The shortest distance between redox centers has been
determined to be 6.2 Å, which leads to an estimated maximum ET
rate in the nanosecond range.
575,576
Rubredoxin from Dv. gigas is important
in the oxidative stress defense system in anaerobic organisms by functioning
as the redox partner of NADH:rubredoxin oxidoreductase and rubredoxin:dioxygen
oxidoreductase
561,577−579
and transferring electrons from ferredoxin:NADP+ oxidoreductase
to superoxide reductase (SOR) to reduce O2 or reactive
oxygen species (ROS).
580−582
It also donates electrons to rubrerythrin
or diiron SORs (i.e., rubredoxin oxidoreductase or desulfoferrodoxin;
see section 3.4.2.4) to reduce hydrogen peroxide
or superoxide, respectively, in Dv. vulgaris.
583
Rubredoxin is an electron acceptor
of carbon monoxide dehydrogenase
and pyruvate ferredoxin oxidoreductase in Chlorobium
tepidum(584) and intracellular
lactase dehydrogenase in Dv. vulgaris Miyazaki F.
585
Furthermore, nucleomorph-encoded
rubredoxin has been discovered to associate with PSII and proposed
to branch electrons from PSII to plastid membrane-located pathways
or replace some of the ET proteins in photosynthesis machinery under
certain circumstances.
586
Rubredoxin
also exhibits high electron self-exchange rates (k
ese). For example, the k
ese of CpRd has been determined to be 3 ×
105 M–1 s–1 at 30 °C
in 50 mM potassium phosphate at pH 7.
587
DFT calculations reveal that pathways through the two surface cysteines
dominate in the electron self-exchange process and surface-accessible
amides H-bonded to the cysteines play an important role as well.
568
3.4.1.3
Important Structural
Features
The reduction potential of the metal cofactor in
a protein is generally
determined by its ionization energy, electronic structure, reorganization
energy, and solvent accessibility during the redox process.
588
Specifically in the case of rubredoxin, the
NH···S H-bonding interactions and water solvation of
the active site are proposed to have a significant influence on the
reduction potential of the iron center. The reduction potentials of
rubredoxins vary in the range of −100 to +50 mV vs SHE (those
of the model complexes are around −1 V vs SHE)
92,588−590
and can be divided into two categories by
the residue at position 44 (Table 4).
590
Rubredoxins such as mesophilic CpRd with lower reduction potentials have a Val residue
at position
44 followed by Gly 45, while those such as hyperthermophilic PfRd with higher reduction
potentials (∼50 mV difference)
have an Ala residue at position 44 followed by Pro 45. Mutating Ala44
of CpRd to Val increases the reduction potential,
and changing Val44 of PfRd to Ala decreases the reduction
potential (Table 4). The short peptide Ala44Pro45
has higher backbone stability, and consequently a higher probability
of orienting the backbone dipole toward the redox center.
591−595
No correlation between reduction potential and Fe–S bond
covalency of CpRd and PfRd has been
observed by sulfur K-edge XAS studies.
596
Table 4
Reduction Potentials for Simple Rubredoxinsa
class
source
E
mν
b (mV)
I (V44)
Clostridium
pasteurianum
–77, −53
Chlorobium limicola
c
–61
Butyribacterium
methyltrophicum
–40
Heliobacillus
mobiliz
–46
Pyrococcus
furiosus A44V
–58
Cp Pf chimerasd
–46 to −67
II (A44)
Clostridium
pasteurianum V44A
–24, +31
Pyrococcus
furiosus
0 to +31
Desulfovibrio vulgaris
e H
0
Desulfovibrio vulgaris
f M
+5
Desulfovibrio
gigas
+6
Megasphaera
elsdenii
+23
Cp Pf chimerasd
+63 to +69
a
Reprinted with permission from ref (590). Copyright 2002 The Royal
Society of Chemistry.
b
Versus
SHE.
c
f. sp. thiosulfatophylum.
d
Constructions of fused domains from Clostridium
pasteurianum and Pyrococcus
furiosus.
e
Strain Hildenborough.
f
Strain
Mivazaki.
The reduction
potential of rubredoxin is pH-independent in the
pH range of 5–10, but pressure- and temperature-dependent.
The reduction potentials of CpRd and PfRd have been reported to linearly decrease
with an increase of temperature
(−1.6 and −1.8 mV/°C, respectively) and decrease
of pressure (0.028 and 0.033 mV/atm, respectively).
597
The phenomena could be rationalized by the dielectric constant
change of a solvent such as water, which is lower at higher temperature
and lower pressure, and consequently less efficient in protein solvation.
Since the stability of a protein oxidation state is dependent on the
solvent–solute interactions to neutralize the excess charge,
the oxidized state of Rd with less net charge is more stable at high
temperatures and low pressures.
598
Replacement of one of the surface cysteines with serine in CpRd resulted in a significant
decrease of the reduction
potential by up to 200 mV, while for internal cysteines only a 100
mV decrease was observed (Table 5). Sulfur
K-edge XAS studies of wild-type CpRd and the four
Ser mutants revealed an increase in the pre-edge energy of the Cys
for all four mutants compared to the wild type, indicating higher
d orbital energy for the mutants, arising from the more electronegative
olate serine ligand, which will lower the reduction potential as observed
experimentally. Consistent with the pre-edge data, extended X-ray
absorption fine structure (EXAFS) fitting shows longer average Fe–S
bonds for the four mutants. DFT calculations also indicate that an
alkoxide ligand stabilizes Fe(III) better than a thiolate ligand.
Changes of solvent accessibility, H-bonding, and the electrostatic
field around the site are other factors possibly involved.
599,600
The Ser mutants display strong pH dependence, possibly arising from
the protonation of coordinating oxygen of Ser following reduction
at neutral or low pH.
601−603
Table 5
Reduction Potentials
for CpRdsa
protein
E°, mV
protein
E°, mV
protein
E°, mV
native
–76
G43A
–93
V44G
0
recombinant
–77
G43V
–123
V44A
–24
C6S
–170
G10V/G43A
–134
V44I
–53
C39S
∼−190
G10V/G43V
–163
V8G/V44G
+39
C9S
–284
V8G
–7
V8I/V44I
–13
C42S
–273
V8A
–44
V44I/V44I
–55
G10A
–104
V8L
–82
V44L
–87
G10V
–119
V8I
–81
a
Square wave voltammetry data, vs
SHE.
Mutations of the secondary
sphere residues have been conducted
mainly on the conserved residues, and potential changes of 100 mV
in both directions have been achieved (Table 5).
604,605
In recombinant CpRd, Gly43Ala
eliminates the Val44 NH···S Cys42 H-bonding interactions,
and a Gly10Val mutation significantly perturbs the overall structure
of Cys9 containing loop by increasing steric hindrance. Replacement
by Val decreases the reduction potential more than Ala, and the mutations
lower reduction potentials up to −86 mV.
604,606,607
Side chain variation of surface
residue 44 of CpRd also could influence the reduction
potential of the metal center. Three mutants, Val44Ile, Val44Ala,
and Val44Gly, increase the reduction potential to −53, −24,
and 0 mV, respectively, from −77 mV of the wild type. The increase
of E° is well correlated with a decrease of
the NH···S H-bond distance determined by 15N NMR. A possible explanation of the trend
is that the shortening
of H-bonds might lead to increased capacity for electron delocalization
or decreased electron donation from the sulfur ligands and finally
to a higher reduction potential of the metal center.
608,609
Similarly, quantum mechanical calculations reveal that shortening
of H-bonds would decrease the energy of the reduced state faster than
that of the oxidized state and result in increased reduction potential.
610
Electrostatic effects of the charged
residues make important contributions
to the reduction potential of the iron center as well. Two neutral
surface residues, Val8 and Leu41, of CpRd close to
the iron center were replaced by positively charged Arg, and the resulting
mutants displayed increased reduction potentials as expected. However,
mutants Val8Asp and Leu41Asp, in which two negatively charged residues
were incorporated, also displayed higher reduction potentials. The
mutations might have also changed the solvent accessibility, and consequently
the dielectric constant around the metal center, leading to complicated
effects difficult to predict and explain simply by Coulomb’s
law.
611,612
A series of unnatural analogues of
tyrosine have been incorporated
into the Tyr10 position of PfRd close to the sulfur
of Cys38 (3.95 Å at the closest point) by native chemical ligation
methods, and the reduction potentials of the resulting proteins are
linearly correlated with the Hammett σp of the para
substituent of the phenyl ring. Electron-donating groups shift E° to more negative
values (Tyr10 PfRd, −78.0 mV; Phe10 PfRd, −69.5 mV;
4-F-Phe10 PfRd, −61.5 mV vs SHE), and electron-withdrawing
groups shift E° to more positive values (4-NO2-F10 PfRd, −49.5 mV; 4-CN-F10 PfRd,
−43.5 mV vs SHE).
613
The trend is not well correlated with the dipole movement of the
side chain
614
and is proposed to arise
from either electrostatic interaction
615,616
or modulation
of the H-bond strength between the sulfur of Cys38 and residue 10.
617−619
3.4.1.4
Spectroscopic Features
Ferrous
rubredoxin is colorless, with weak absorptions centered at 311 and
331 nm. On the other hand, ferric rubredoxin displays strong absorption
peaks at 350, 380, 490, and 570 nm from ligand to metal charge transfer
(LMCT) of the σ orbital and a weak peak at 750 nm from the π
orbital of the cysteinyl sulfur to the metal center (Figure 16a). Mutating one of
the Cys residues to Ser still
gives LMCT bands in ferric form, but with the peaks shifted to higher
energy together with some changes of intensity, consistent with a
decreased S to Fe(III) LMCT contribution.
562
CD spectra of rubredoxins display minima at 202 and 226 nm from
β-sheet structures in the protein.
620−622
Figure 16
Representative spectra of rubredoxins. (a) UV–vis spectra
of ferric and ascorbate reduced ferrous (inset) CpRd. (b) Mössbauer spectra of dithionite
reduced ferrous CpRd measured at 4.2 K under a magnetic field applied parallel
to the γ-rays. Reprinted from ref (623). Copyright 2002 American Chemical Society.
(c) EPR spectra of CpRd. Reprinted with permission
from ref (611). Copyright
1996 Elsevier.
Mössbauer spectra
of ferrous rubredoxin as purified give
parameters of an S = 2 Hamiltonian with D = 5.7(3) cm–1, E/D = 0.25(2), δ = 0.70(3)
mm/s, and ΔE
Q = −3.25(2) mm/s (Figure 16b).
623
Consistent with the Mössbauer
studies, experiments using broad-band quasi-optical HF-EPR reveal
a D value of 4.8 ± 0.2 cm–1 and E/D of 0.25 ± 0.01.
624
The ferric form is high-spin as well, as determined
by EPR spectroscopy, with a set of signals arising from an S = 5/2 spin state, including
g = 4.3 from
the middle Kramers doublet and g = 9.5 from the lowest
Kramers doublet (Figure 16c). The Mössbauer
spectrum of the oxidized form of CpRd shows δ
= 0.24 ± 0.01 mm/s at 4.2 K.
603,625
The
Fe–S covalency has also been probed using single-molecule
AFM by measuring the mechanical stabilities of Fe(III)–thiolate
bonds. The rupture forces of interior Fe–S bonds of PfRd are greater than those of
surface Fe–S bonds,
consistent with other experimental observations.
626
The mechanical stability of Fe–S bonds also shows
good correlation with the NH···S H-bond strength reflected
by the reduction potential.
627
The
dynamic properties of the redox iron center are important for
the redox properties of a protein. 57Fe nuclear resonance
vibrational spectroscopy (NRVS) of the oxidized form of PfRd, which is sensitive to
all normal modes involving the Fe center,
shows bands around 70, 150, and 364 cm–1. The 70
cm–1 signal is from collective motion of some or
all of the coordinating cysteines with respect to the iron center.
The ∼150 cm–1 signal mostly involves S–Fe–S
bending motion composed of a doubly degenerate E mode (ν2) and a mixed T2 ν4 mode
of Td
symmetry. The strong signal between 355 and 375
cm–1 is mainly from an asymmetric Fe–S stretch
mode, ν3, of Td
symmetry,
consistent with an average value of 362 cm–1 from
Raman spectra of Dv. gigas (Dg) Rd. In the case of the reduced form, the asymmetric
Fe–S stretching modes shift to 300–320 cm–1, bending modes shift slightly lower, and
collective motion modes
at ∼70 cm–1 do not change substantially.
Derived force constants of both stretching and bending modes are higher
in the oxidized form than in the reduced form.
614,628
The resonance Raman spectra of oxidized Rd display the strongest
band at ∼315 cm–1 from totally symmetric
Fe–S4 breathing modes.
614
The force constant of the ν3 frequency is lower
than in synthetic models, probably because of the H-bonding to the
S of the cysteines in the protein scaffold.
589
1H NMR has been utilized to study the magnetic
properties
of ferrous rubredoxin. Broadening and shifting of signals are observed
due to the presence of iron. To avoid the strong paramagnetism of
iron, other metals such as Zn, Cd, and Hg were used as a surrogate
of Fe(II) for structural studies. Paramagnetic contact shifts in 1H, 2H, 13C, and
15N nuclei
of oxidized CpRd have been measured experimentally,
and the data are consistent with high-level all-electron density functional
calculations based on high-resolution crystal structures. Computational
studies reveal that the experimental hyperfine shifts are mainly from
Fermi contact interactions.
629,630
NMR has also been
applied in measuring the magnetic susceptibility anisotropies of both
oxidized and reduced CpRd, demonstrating that pseudocontact
has negligible contributions to hyperfine shifts.
631
3.4.2
Rubredoxin-like Proteins
3.4.2.1
Flavorubredoxin
Flavorubredoxin
is a type of protein containing a rubredoxin-like domain coupled to
a flavodiiron protein and a flavodoxin domain binding one flavin mononucleotide.
632,633
It has been isolated from E. coli and Moorella (M.) thermoacetica and
discovered to be involved in ET pathways in reduction of nitric oxide
and conversion of CO2 to acetate.
634−637
The reduction potentials of flavorubredoxins from E. coli have been determined to
be −140 ±
20 mV at pH 7.6
635
and −120 ±
20 mV at pH 7.5.
636
The reduction potential
of flavorubredoxin from M. thermoacetica is −30 ± 10 mV at pH 7.0.
638,639
3.4.2.2
Diiron Rubredoxins
Diiron rubredoxin
is composed of two [FeCys4] domains connected by a 70–80
amino acid linker.
570,640
It can be readily prepared from
corresponding monoiron rubredoxin by precipitation and resolubilization
and is proposed to be the physiological form of rubredoxin. Though
less stable, it can transfer electrons from reduced spinach ferredoxin
reductase to cytochrome c just as the monoiron form.
The midpoint reduction potential of both of the two-electron reduction
processes is −10 mV vs SHE at pH 7.0, similar to that of monoiron
rubredoxins.
641
3.4.2.3
Desulforedoxin
Desulforedoxin
(Dx), isolated from sulfate reducing bacterium Dv.
gigas, is an α2 dimer with 36 amino
acids for each subunit. Each dimer contains a four-stranded antiparallel
β-sheet and several turns and interchain short β-sheets.
Each monomer has a high-spin rubredoxin-like [Fe(Cys)4]
center. The iron center is near the protein surface, coordinated by
four cysteine residues, Cys9-Xxx-Xxx-Cys12 and Cys28-Cys29. Unlike
rubredoxin, two of the four coordinating cysteines are consecutive,
making the tetrahedral coordination geometry distorted (Figure 17).
642,643
In addition, Dx only has one
aromatic residue, while Rd has up to six. The Fe–S bond lengths
of Dx range from 2.25 to 2.36 Å, and the S–Fe–S
angles vary from 102° to 119°.
Figure 17
Crystal structure of
desulforedoxin from Dv. gigas (PDB
ID 1DXG). The
[FeCys4] centers are displayed in ball-and-stick
mode and denoted. The backbones of coordinating cysteines are omitted
for clarity. Color code for the ball-and-stick mode: cyan, carbon;
green, iron; yellow, sulfur.
Oxidized Dx displays three major UV–vis absorptions
centered
at 278, 370, and 507 nm. The 370 and 507 nm absorptions arise from
the sulfur to iron charge transfer, and the extinction coefficient
of the 507 nm absorption is 4580 M–1 cm–1 per monomer, falling in the normal range
of Fe–S proteins.
Unlike the nearly rhombic EPR features of oxidized Rd (E/D = 0.28),
644
the EPR spectrum of oxidized Dx displays an S =
5/2 site with near axial symmetry, with g = 4.1,
7.7, and 1.8 from the ground Kramers doublet and g = 5.7 from the middle Kramers doublet.
645
This difference reflects different geometric and electronic structures
of the two iron sites. D = 2.2 ± 0.3 cm–1, ΔE
Q = −0.75
mm/s, and δ = 0.25 mm/s are obtained by Mössbauer studies
of oxidized Dx. The parameters of reduced Dx from Mössbauer
studies are D = −6 cm–1, E/D = 0.19, ΔE
Q = 3.55 mm/s, and δ = 0.70 mm/s. The positive ΔE
Q value of reduced Dx indicates that the ground-state
orbital is mainly d
x
2
–y
2, while the ΔE
Q value of reduced Rd is correlated to pure d
z
2 as the ground-state orbital.
642
Insertion of a Gly residue or Pro-Val
residues between Cys28 and
Cys29 makes the ferric center of Dx nearly spectroscopically identical
to that of Rd. However, both mutations are detrimental to the protein
stability.
646
Similar to Rd, Dx associates
with other metal centers in biological
systems. For example, desulfoferredoxin (Dfx) possesses a binding
motif for the Dx-type [FeCys4] center associated with another
nonheme monoiron center with N/O ligands
647
(see section 3.4.2.4). Moreover, Dx in Dv. gigas is reported to transfer electrons
to SOR
more efficiently than Rd.
648
3.4.2.4
Desulfoferrodoxin
Dfx is an α2 dimer
with a molar mass of ∼28 kDa belonging to the
diiron superoxide reductase family.
649,650
Each monomer
contains an [FeCys4] center (center I) and a nonheme iron
center coordinated by a four-His–one-Cys motif (center II).
651
The 1.9 Å resolution crystal structure
reveals that center I is structurally similar to the metal center
of Dx.
652
The midpoint reduction potential
of center I is around 0 mV, falling in the range of [FeCys4] centers in Dx and Rd.
647,653−656
Replacement of Cys13 of Dfx from Dv. vulgaris Hildenborough with serine results in
a [1Fe–3Cys–1Ser]
center instead of the Rd/Dx-like center. Redox titration reveals no
influence on the reduction potential of center II by such a mutation,
indicating the independence of the two cofactors.
657
On the other hand, reduction potentials of Dfx from hyperthermophilic
archaeon Archaeoglobus fulgidus are
+60 mV for center I and +370 mV for center II,
649
while E° is +230 mV for monoiron
SOR containing only the center II cofactor from the same genome.
658
The difference in E°
implies possible involvement of center I of Dfx in facilitating the
reduction of center II.
654
3.4.2.5
Rubrerythrins
Rubrerythrin (Rr),
an α2 dimer, is a nonheme iron protein with peroxidase
and in vitro ferroxidase activity.
583,659
Each monomer
contains a diiron–oxo site in the middle of a four-helix bundle,
and an [FeCys4] center at the C-terminus.
660,661
The [FeCys4] center is structurally very similar to that
of Rd, yet the midpoint reduction potentials are estimated to be +230
mV at pH 8.6 and +281 mV at pH 7.0, much higher than the normal value
of around 0 mV for Rd centers.
662,663
The crystal structure
reveals the dramatic potential increase and pH-dependent behavior
might be due to the polar and solvent-exposed environment around the
iron center created by nearby residues, including Asn160, His179,
and Ala176, which are not conserved in Rd.
660,664
Replacement of the iron in the Rd-like domain with zinc inhibits
the peroxidase activity of the protein, indicating the essential role
of the Rd domain in the ET process.
665
Desulforubrerythrin, a unique member of the rubrerythrin family,
has been isolated recently from Campylobacter jejuni. It is an α4 protein, and each
24 kDa monomer is
composed of three domains: a Dx-like N-terminal domain, a four-helix
bundle domain containing a μ-oxo-bridged diiron site, and an
Rd-like C-terminal domain. The reduction potentials of the [FeCys4] centers in the
N-terminal and C-terminal domains are +240
± 30 and +185 ± 30 mV, respectively, at pH.7.0 vs SHE.
666
Nigerythrin is an α2 dimer containing one diiron–oxo
center and an [FeCys4] center, very similar to rubrerythrin.
The reduction potential of the Rd-like center in nigerythrin from Dv. vulgaris is
+280 mV vs SHE at pH 7.5, comparable
to that of Rr as well.
663,667,668
3.4.3
Ferredoxins
3.4.3.1
Introduction
The term ferredoxin
refers to a wide range of small, low molar mass Fe–S proteins
that function solely as electron carriers in different biological
pathways including photosynthesis and respiration.
669
Ferredoxins first were observed on the basis of their distinct
rhombic EPR feature with g = 1.9. EPR studies with 57Fe later confirmed that the signal
is from a nonheme iron.
670
Evolution of H2S gas upon acid treatment
was an indicator of the presence of inorganic sulfur in this protein.
1,539,671
All ferredoxins share some common
features: They are all low molar mass, highly acidic proteins that
contain iron and inorganic or acid-labile sulfurs.
1,669
The Fe–S cluster resides in a hydrophobic patch within the
protein and gives the proteins a distinctive dark-brown color.
672,673
All ferredoxins go through a partial decrease in absorbance upon
reduction. Reduction can be achieved through chemical treatment by
sodium hydrosulfite or enzymatic treatment with H2 gas
and hydrogenase. The pattern of reduction is dependent on the method
and extent of reduction. After reduction, a rhombic EPR signal appears
with g < 2 (exact value depending on the cluster
type).
539,672
Ferredoxins usually have low reduction potentials
with an average of −400 mV and spanning a range of 800 mV depending
on the cluster type, protein structure, H-bonding network, water solubility
of the cluster, and ligands to the iron. This wide range enables ferredoxins
to serve as redox partners to a variety of molecules in a number of
important biological reactions. Due to the high acidity, these proteins
usually have high affinity for (diethylamino)ethanol (DEAE)-sepharose
and can be easily purified by acetone precipitation and DEAE-facilitated
separation. The purity can be monitored by the ratio of A390/A280.
539,672,673
It has been shown that the proteins can usually be reconstituted
by treatment with iron and Na2S under reducing conditions
(in the presence of β-mercaptoethanol).
539,672−674
All of the low reduction potential
ferredoxins seem to have evolved from a common ancestral polypeptide.
91
Despite different types, CD and optical rotatory
dispersion (ORD) studies show that all ferredoxins have a very similar
polar active site environment around the cluster in which the iron
assumes tetrahedral coordination geometry. The similarity of extinction
coefficients of their electronic absorption bands, mainly due to metal
to ligand charge transfer, also indicates a similar electronic structure
of the iron center.
539
Despite somewhat
surface-exposed iron, the reaction of the proteins with iron chelators
is usually slow, unless denaturing conditions are applied.
539
The ferredoxins are further divided into subcategories
on the basis of the number of iron molecules present in the cluster.
3.4.3.2
[2Fe–2S] Clusters
3.4.3.2.1
Structural Aspects
As their
name suggests, [2Fe–2S] clusters are a class of one-electron
transport ferredoxins containing two iron atoms that are coordinated
in a distorted tetrahedral geometry by two inorganic sulfurs and four
cysteine thiolates from the protein. The [2Fe–2S] cluster is
not completely planar, having a small tilt in the plane of the first
and second irons. Three of the four cysteines come from one loop in
the structure of the protein, with the other one being at the tip
of a β-strand in a different loop (3 + 1 arrangement). The cluster
is positioned close to the surface of the protein, surrounded by hydrophobic
residues. Except for the vicinity of the cluster, the surface of [2Fe–2S]
ferredoxins is highly acidic, covered with a large number of Asp and
Glu residues. This acidic patch is used to interact with the basic
surface of redox partners. After initial alignment through these electrostatic
interactions, hydrophobic interactions between the two surfaces and
water exclusion further facilitate the ET between the proteins.
540,677
A role for the orientation of redox partners with regards to each
other has been proposed in ET rates.
678
Lack of complete complementarity between the two surfaces ensures
the separation of oxidized ferredoxin and initiation of a new cycle.
540
There are several NH···S H-bonds
from backbone amides to the sulfurs of the cluster, with sulfur ligands
of Fe1 (the iron closer to the surface) being involved in more H-bonds
than those of Fe2.
678,683
It appears that the Fe–Fe
and Fe–Sγ bonds lengthen upon reduction while
the H-bonds strengthen and shorten, consistent with increased negative
charge on S.
683,685
Despite these
similar features, [2Fe–2S] ferredoxins can be further divided
into three subcategories on the basis of differences in sequence and
structural alignments and in the ligand Cys motifs (Figure 18). The details about
each category are briefly
explained below.
677
Figure 18
Structures of three
classes of [2Fe–2S] ferredoxins. Notice
that, in their physiological form, thioredoxin-like ferredoxins function
as a dimer.
3.4.3.2.1.1
Plant-Type Clusters
The archetype
of plant-type ferredoxins is chloroplast ferredoxin I. The members
of this family share a common β-grasp structural motif, which
consists of three to five β-strands, with one to three adjacent
α-helices and some additional secondary structures and loops.
91
Three of four coordinating Cys residues are
in a loop with a conserved Cys-(Xxx)4-Cys-(Xxx)2-Cys motif, and the fourth Cys is
29 amino acids away. The cluster
is usually buried at one end of the protein in a hydrophobic environment.
677,682,683
Although plant-type ferredoxins
have high sequence homology, there are multiple isoforms of them in
each organism, which suggests different roles of the isoforms in different
evolutionary and physiological conditions. Acidic residues are usually
distributed in an asymmetric fashion, resulting in a dipole with its
negative end near the Fe–S cluster. This dipole is shown to
be important in docking of the ferredoxin into its redox partner.
679−681
Several H-bonds anchor the cluster to the protein and are known
to be important in fine-tuning the reduction potential of the protein.
A water channel with five water molecules connects the solvent to
the proximity of the cluster in the C-terminal region of
the protein.
677,683−686
3.4.3.2.1.2
Mammalian/Mitochondrial Cluster
Mostly known for their hydroxylating role, these clusters include
mammalian [2Fe–2S] proteins as well as some bacterial [2Fe–2S]
proteins. The archetypes of this class are adrenodoxin and bacterial
putiredoxin. The overall fold and structure of this class are very
similar to those of plant-type clusters with the exception that they
have an additional interaction loop,
91
a
large hydrophobic domain that is used as an interacting domain with
the redox partner. The conserved ligating motif of this class is Cys-(Xxx)5-Cys-(Xxx)2-Cys,
with the fourth cysteine 35–37
residues away from the third ligand, further away than in plant-type
structures. This group has a very flexible C-terminal which is very
difficult to crystallize, but can be captured in the presence of its
redox partner. It also has a compact α + β structure,
characteristic of ferredoxins. Interestingly, the same fold has been
observed in enzymes containing Fe–S clusters as well as some
unrelated proteins that are void of Fe–S clusters. There has
been evidence of structural changes upon reduction in some loops as
well as the C-terminus. The solvent channel is shorter in mammalian-type
ferredoxins compared to plant-type ferredoxins.
677,681,682
3.4.3.2.1.3
Thioredoxin-like Clusters
These proteins are only reported
in bacteria, mostly in proteobacteria
and cyanobacteria.
689
They were first discovered
in Cl. pasteurianum(687) and Azotobacter vinelandii(670) due to their spectroscopic
features,
which are distinct from those of common [2Fe–2S] ferredoxins.
Their sequence and positioning of the cysteine ligands differ significantly
from those of other ferredoxins. These differences were further confirmed
by analyzing vibrational bands in resonance Raman studies.
688
Two features in the structure of this class
are known to cause these differences: a distortion of the loop containing
the Cys ligands and a H-bond between two cysteine residues. Proteins
of this class function as a dimer, each monomer having a thioredoxin-like
fold, despite low sequence homology (∼7%). Two regions are
notably distinct between these proteins and thioredoxins: a protruding
surface loop that has been shown to have no significant function and
an α-helix in one subunit and a short helix in the other subunit
that are important in interaction
689
between
the two subunits. The cluster lies within two loop regions in the
periphery of subunits in a conserved motif of Cys-(Xxx)10−12-Cys-(Xxx)29−34-Cys-(Xxx)3-Cys.
682,692
The fourth cysteine is placed in a protruding loop, which is absent
in other ferredoxins. Several studies showed that the position of
this Cys is flexible and that it can be moved to other positions in
the loop.
690,691
Some members of this class contain
five cysteines instead of four. ESEEM studies and mutational analyses
showed that loss of one of these cysteine residues can be compensated
by the other four.
9003
There are a small
number of conserved residues in the family, including the four cysteine
ligands and some cysteines in the dimer interface. The overall common
structure has five β-strands, two long α-helices, and
an additional short helix. The Cys ligands of the more buried iron
are provided by the loop that is longer. The cluster itself shows
some deviation from other ferredoxins with two irons. One difference
is a more compressed angle between two sulfurs of Cys ligands and
Fe2 (the more buried iron atom), and the other is a longer distance
between one of the Cys residues and Fe2 than other Fe–S distances.
The cluster is more surface-exposed in this class than the other two
classes of [2Fe–2S] ferredoxins.
91,689,692,693
3.4.3.2.2
Function
3.4.3.2.2.1
Plant-Type Ferredoxins
Plant-type
ferredoxins can usually be found in the stroma of chloroplasts of
higher plants and algae as well as the cytoplasm of cyanobacteria.
Ferredoxins play a role as the first electron acceptor in the stromal
side of chloroplasts and function mainly as electron distributors
in photosynthesis. They are also involved in a variety of other functions
such as sulfur and nitrogen assimilation, biosynthesis of several
compounds such as chlorophyll, and redox homeostasis of the cell.
538
The most important and well-studied function
of these proteins is the transfer of two electrons in two consecutive
steps from photoreduced PSI to ferredoxin:NADP reductase (FNR), which
will result in final CO2 assimilation.
538,683
The FNR forms a 1:1 complex with reduced ferredoxin and uses NADP+ to oxidize the
ferredoxin. The NADP+ and ferredoxin
have separate binding sites in FNR. It has been shown that binding
of one of these substrates (ferredoxin or NADP+) weakens
the binding of the other. Once oxidized, the ferredoxin has a lower
binding affinity to FNR and dissociates from the complex, while a
second reduced ferredoxin will replace it to complete the cycle.
2003
In organs that produce NADPH by the pentose
phosphate cycle, FNR acts in the reverse direction and reduces ferredoxin.
677
Ferredoxin also distributes electrons
from photoreduced PSI to
ferredoxin-dependent enzymes such as nitrite reductase, glutamate
synthase, and ferredoxin:thioredoxin reductase (FTR) for nitrogen
and sulfur assimilation. Cyanobacteria have a vegetative ferredoxin
that functions in photosynthesis and a heterocyst ferredoxin that
transfers electrons to nitrogenase. Ferredoxin from halobacteria can
function as an electron carrier in α-keto acid decarboxylation
or in nitrite reduction.
694
One of
the most studied realms in the field of ferredoxins is their
interaction patterns with their redox partners. These complexes have
been studied using several techniques such as cross-linking, NMR,
isothermal titration calorimetry (ITC), and site-directed mutagenesis;
however, it is not completely understood whether ferredoxin uses the
same surface, partially overlapping surfaces, or totally different
surfaces for interacting with different redox partners. The most likely
hypothesis is that ferredoxin acts as a mobile electron carrier between
PSI and other redox partners.
677
3.4.3.2.2.2
Interactions with Other Proteins
3.4.3.2.2.2.1
Interaction with Ferredoxin:NADP+ Reductase (FNR)
The most well-known partner of plant-type
ferredoxins is FNR. It has been shown that ferredoxin and FNR have
very tight binding with K
d in the range
of 10–7–10–8 M.
677
As discussed previously, several surface amino
acid residues are conserved in ferredoxins, and mutation of these
amino acids revealed important factors in interaction between these
redox partners. Binding of ferredoxin to FNR cause a negative shift
in Em of ferredoxin, which is suggested to be important in more efficient
ET between the two proteins. Laser flash photolysis is one of the
techniques that has been used to analyze the reactivity of several
ferredoxin mutants from Anabaena. Among
the conserved residues, Phe65 was the only one essential for tight
binding between ferredoxin and FNR.
684,697
Ser47, Glu94,
and Phe65 were also shown to be important in the rapid ET between
the two partners, though conservative mutations to other similar residues
were tolerated.
677,695
Interestingly, mutating residues adjacent to the above three residues
had a much less effect on the activity.
684
Mutational studies of Glu92 in spinach ferredoxin, which is analogous
to Glu94 in Anabaena, resulted in decreased
activity, but much less significant than that of the former. More
interestingly, this mutation resulted in an increase in reduction
potential and stimulation of NADPH–cytochrome c reductase activity catalyzed by FNR.
These mutants were more efficient
in transferring electrons in the direction opposite that of the physiological
ET pathway.
2004
Although several studies
have shown significant correlation between ET and reduction potential,
ET changes are thought to be more likely a result of changes in protein
orientation and transient changes in configuration rather than a consequence
of reduction potential changes. A thorough study of the mutants with
laser flash photolysis showed very similar effects of Glu92/94 mutation
in both spinach and Anabaena variants,
hence suggesting a difference between these results and previous NAD+ photoreduction
results.
677
ITC
studies suggested entropy as the main driving force of complex formation,
meaning that hydrophobic interactions are the major forces governing
the efficient interaction between the two partners. The proposed binding
surfaces of many ferredoxins are covered with water, so the binding
of the partners will release water molecules and favor the reaction
entropically.
694,698
Several models of complexes
between ferredoxins and FNRs have been made on the basis of experimental
evidence coming from chemical modification, cross-linking, partial
proteolysis, and mutational studies, as well as homology models. These
models predicted the binding site between ferredoxin and FNR to be
a large hollow surface near the dimethylbenzyl ring edge of the flavin
in FNR. The binding will bring the Fe–S cluster and the flavin
close, so that they can transfer electrons. While ferredoxin has an excess of positive
charge on the binding
surface, FNR has a net negative charge on its binding surface. The
specific orientations of dipoles in the two proteins have been shown
to be important in recognition between the two partners. Another model
proposes that electrostatic potential complementation plays an important
role. The two models differ in the orientation of the ferredoxin molecule
about the axis perpendicular to the protein–protein surface.
677,679,680
Cross-linking experiments have
been done to study the complex between ferredoxin and FNR (Figure 19). The cross-linked
molecule showed oligomer states
in the crystal structure that might be relevant to in vivo interactions.
699
Figure 19
Structure of ferredoxin (right) cross-linked
to FNR (left), PDB
ID 3W5U. As
shown, red acidic patches of ferredoxin are positioned in contact
with blue basic residues of FNR. A zoomed-in figure of the region
containing the cofactors (Fe–S and FAD) is shown at the bottom.
3.4.3.2.2.2.2
Interaction with Ferredoxin:Thioredoxin
Reductase, Nitrate, Nitrite and Sulfite Reductase, and Glutamate Synthase
Reduced ferredoxin donates electrons to FTR to reduce thioredoxin,
which is involved in multiple steps of the Krebs carbon cycle. FTR
is found only in oxygenic photosynthetic organisms. Chemical modification
of acidic residues on the surface showed that the Glu92–94
acidic patch is important for the interaction between the two partners.
A model has been proposed on the basis of the crystal structures of
the two partners. In this model, ferredoxin docks into the opposite
site of the flat, disklike structure of FTR in such a way as to position
itself close to the [4Fe–4S] cluster and the redox-active disulfide
bond in FTR.
700
In this ternary complex,
two successive one-ET reactions take place. The complex between ferredoxin
and FTR has very high affinity, with both electrostatic and hydrophobic
interactions being involved.
Site-directed mutagenesis and chemical
modification studies suggest that the same site of ferredoxin is responsible
for interacting with nitrite reductase, sulfite reductase, and glutamate
synthase.
677,701,702
The surface is formed in low ionic strength, indicating a role for
electrostatic interactions in formation of the complex.
694
Another site has also been proposed for sulfite
reductase (SiR).
694,703
While less is known for SiR,
NMR analyses of the contact shifts between the presumed complex confirmed
the important role of acidic surface residues on complex formation.
694
Nitrate reductase is found in cyanobacteria
and performs two-electron
reduction of nitrate to nitrite. It has been shown that there is only
one ferredoxin binding site in nitrate reductase, so the reduction
proceeds in two separate consecutive steps.
694
Nitrite reductase performs six-electron reduction of nitrite
to
ammonia. As with nitrate reductase, only one binding site exists for
ferredoxin. A conserved Trp residue has been shown to play an important
role in ET between the two partners.
694
A loop close to the [3Fe–4S] cluster of glutamate synthase
is responsible for binding of ferredoxin. CD analyses showed that
neither of the two proteins underwent significant conformational changes
upon binding.
694
3.4.3.2.2.2.3
Interaction with Photosystem I
Photosystem I (PSI) is an essential part of the photosynthetic
ET pathway in cyanobacteria and plants. This multisubunit complex
is a membrane-bound system that harvests light and helps convert it
into a chemical potential. The complex consists of multiple chlorophylls,
carotenoids, phylloquinones, bound lipids, and [4Fe–4S] clusters.
Three subunits at the stromal site of PSI are involved in docking
and reducing ferredoxin I: PsaC (with [4Fe–4S] clusters FA and FB), PsaD, and PsaE.
FA, FB, and FX are three low-potential [4Fe–4S]
clusters that lie in the stromal side of the PSI complex. FA and FB are bound to PsaC,
and FB functions
as a terminal electron acceptor (Figure 20).
FX is an interpolypeptide cluster, positioned between PsaA
and PsaB, and has the most negative reduction potential reported so
far for a [4Fe–4S] cluster (−705 mV).
705
Figure 20
Structure of PSI (PDB ID 1JB0). The top left figure shows the overall
structure,
and the bottom figure shows all the cofactors in the system. The top
right figure shows the PsaC, PsaD, and PsaE sites with FA and FB. Ferredoxin binds
in the interface between PsaC,
PsaD, and PsaE.
In vitro studies and
cross-linking experiments revealed PsaD as
the main docking site for ferredoxin I. A binding site for PsaC has
been also proposed on the basis of mutational studies. It has been
shown that PsaD and FNR compete with each other in binding to ferredoxin,
yet no ternary complex has been observed.
705
3.4.3.2.2.3
Mammalian-Type and Thioredoxin-like
Ferredoxins
The main function of mammalian-type ferredoxins
is ET in the mitochondrial ET chain, ET to P450’s, and Fe–S
biosynthesis. It has been shown that adrenodoxin has very tight binding
to both adrenodoxin reductase and cytochrome P450 on the order of
10–7–10–8 M.
677
As
with ferredoxin, adrenodoxin interacts with its redox partners through
an acidic surface,
706
with Asp76 and Asp79
being essential for the binding. The overlapping interaction surface
supports a mobile carrier hypothesis for the adrenodoxin. A model
based on the crystal structures of the partners suggests that adrenodoxin
binds in the cleft between two domains of adrenodoxin reductase, resulting
in a distance of 16 Å between the Fe–S cluster and the
isoalloxazine ring of the FAD in the reductase.
707,708
A specific ET path between the two has also been proposed.
708
Several studies on putiredoxin have shown the
same overlapping surface for reductase and P450 interaction. The crystal
structure of the complex between adrenodoxin and adrenodoxin reductase
further confirmed the importance of charged Asp and Glu residues on
the surface of ferredoxin in the formation of the complex (Figure 21).
709
Figure 21
Structure of adrenodoxin
(right) in complex with adrenodoxin reductase
(left) (PDB ID 1E6E). As shown, red acidic patches of adrenodoxin are positioned against
blue basic residues of adrenodoxin reductase. A zoom-in region of
the cofactors (Fe–S and FAD) is shown at the bottom.
No certain function has been determined
for thioredoxin-like ferredoxins
yet. However, their abundance in nitrogen-fixing bacteria suggests
a role in nitrogen metabolism. Some molecular dynamics and docking
studies have shown an interaction surface with this class of proteins
and the MoFe protein of nitrogenase, suggesting a role as electron
carriers to this complex.
692,689,710
To analyze the ET activity of [2Fe–2S] ferredoxins,
a simple
spectroscopic assay can be performed using cytochrome c as the final electron acceptor.
677
A
wealth of mutational studies showed the importance of entropy as the
main driving force in this interaction. While positive surface charges
are important in bringing the two proteins into proximity, hydrophobic
interactions are the major players in stabilizing the complex.
694
3.4.3.2.3
Important Structural
Features
The reduction potentials of ferredoxins from plants
and mammals are
between −460 and −300 mV.
683,694
On average, mammalian ferredoxins have higher reduction potentials
than plant-type ferredoxins,
683
due to
different patterns of electron delocalization, as observed by NMR.
711
Interestingly, mammalian ferredoxins show an
ionic strength- and pH-dependent redox behavior.
712
The average reduction potential for the thioredoxin-like
class is around −300 mV.
689
Multiple
methods have been used to measure reduction potentials of ferredoxins,
including direct protein film voltammetry,
714
and spectrochemical redox titration.
714
Several factors have been reported to be important in fine-tuning
the reduction potentials of ferredoxins. The overall protein fold
and solvent accessibility of the cluster are known to be important
in giving a low reduction potential range to ferredoxins compared
to Rieske centers that also have a [2Fe–2S] cluster core. These
factors are discussed in more detail in the section on Rieske centers
(section 3.4.4).
Models of [2Fe–2S]
proteins have been used to analyze the
reduction potential properties. These analyses have shown the nature
of the peptide to be important in reduction potential determination
and behavior.
717
Other factors such as
the H-bonding network from backbone amides to sulfurs and overall
charge of the protein are reported to play a role in determining the
reduction potential value within [2Fe–2S] ferredoxin classes.
In all the classes, there is a conserved H-bonding network, with sulfurs
ligating the higher potential iron being involved in more H-bonds
(Figure 22).
677,683
Figure 22
H-bonding
network in plant-type ferredoxins.
It has been suggested that the charge and H-bonding pattern
differences
between Thioredoxin-like ferredoxins and plant-type ferredoxins is
the cause of differences in their reduction potential. Indeed, point
mutations near the active site that change the charge of thioredoxin-like
ferredoxin resulted in a 100 mV change in reduction potential.
689
Three kinds of mutations were found to influence
the reduction potential in thioredoxin-like ferredoxins the most:
replacing Cys ligands, swapping ligands or changing the loop containing
them, and changing the charge in the vicinity of the cluster.
689
Interestingly,
changing the loop (either insertion or deletion) resulted in a reduction
potential correlated with the sum of the charged residues left in
the loop. Cys → Ser mutations caused a decrease in reduction
potential.
690,721
A 100 mV change in reduction
potential was observed upon mutating one of the Cys residues in thioredoxin-like
ferredoxins that have five Cys residues.
690
Cys to ser mutants of Anabaena [2Fe−2S]
cluster showed that the changes in reduction potential is dependent
on the position of ligating Cys.
678
Mutations
of Glu94 and Ser47 of Anabaena ferredoxin
showed a significant increase in the reduction potential of this protein
mostly due to rearrangement of the H-bonding network as well as removal
of a negative charge close to the cluster.
9004
3.4.3.2.4
Spectroscopic Features
All
[2Fe–2S] ferredoxins share very similar UV–vis spectra
with a protein peak at 280, a near ultraviolate peak at 330 nm, and
visible absorptions at 420 and 463 nm, and a shoulder at 560 nm in
the oxidized form (Figure 23). The relative
intensities of the 420 and 460 bands are inverted in thioredoxin-like
ferredoxins compared with the other two groups. Depending on the hydrophobicity
and H-bonding pattern around iron atoms, one of them, usually the
one closest to the surface, is reduced more easily. After reduction,
the spectral intensity decreases to about 50% of that of the oxidized
form and the band positions are altered to a maximum at 540, with
small peaks at 460, 390, 350, and 312 nm.
538,539
These proteins show similar CD and ORD spectra. A red shift was
observed in the spectra after selenium substitution. Strong positive
bands between 420 and 460 nm in the oxidized form dominate the CD
spectra. The reduced state has negative bands at 440 and 510 nm. From
these CD analyses, bands from d
z
2
→ d
xz
and d
z
2
→ d
yz
have been assigned.
539
Figure 23
Representative spectra
of [2Fe–2S] ferredoxins:
728
(a)
UV–vis spectra of reduced (thin
line) and oxidized (thick line) forms of ferredoxin from Aquifex aeolicus; (b) X-band
EPR of [2Fe–2S]+ ferredoxin from Aq. aeolicus at 20 K; (c) Mössbauer of the [2Fe–2S]2+
state of ferredoxin from Aq. aeolicus at 4.2 K in zero field (upper) and an 8.0 T
applied field parallel
to the observed γ radiation (lower). Reprinted from ref (728). Copyright 2002 American
Chemical Society.
Ferredoxins were first
identified through their unique EPR signal
in the reduced state (Figure 23). The two irons
in the Fe(III) state each have a spin of S = 5/2
and are antiferromagentically coupled, resulting in a final diamagnetic
EPR-silent species. Upon reduction of one of the iron ions to Fe(II),
the net spin changes to 1/2 and a rhombic EPR signal at g = 1.94 is observable at
temperatures below 100 K. When the iron
in the protein is replaced with 57Fe, the sample shows
a broader or split EPR signature, proving that the signal is from
iron.
670
Studies with S33 showed
that hyperfine splitting from S contributes to broadening of the signal
at g = 1.94.
539,722,723
ENDOR experiments were performed and provided information complementary
to that of EPR that is required for computer simulation of Mössbauer
data. These studies showed two nonequivalent iron sites in the reduced
form, consistent with the Mössbauer results. The same studies
also revealed some protons that are coupled to irons in the cluster.
539
While all studies are consistent with a localized
electronic structure of the irons in the reduced state, a Cys →
Ser mutant of a thioredoxin-like ferredoxin showed a valence-delocalized S = 9/2 feature
in EPR, which was further analyzed by variable
temperature magnetic circular dichroism.
724
Due to the centrosymmetric core of [2Fe–2S] ferredoxins
(D
2d
, oxidized; C
2v
, reduced), the ungerade
vibrations are Raman-inactive and the protein has fewer features than
its counterpart Rieske centers. They show a characteristic Bt3u peak at around the
283–291 cm–1 region, which shifts to 263–273 in the reduced form. Other
features are an Agt peak at 329–338 cm–1, a B1ut peak at 350–357 cm–1 (mostly Fe–St
stretching mode), and an Agb peak at 387–400 cm–1 in the oxidized form.
These peaks appear at 307–314, 319–328, and 370–385
cm–1 in the reduced form, respectively. Resonance
Raman spectra of thioredoxin-like ferredoxins are substantially different
from those of the other two categories due to different cluster environments.
The main peaks are observed at 208, 290, 313, 335, 353, 366, 387,
and 404 cm–1 in the oxidized form and at 267, 280,
310, 328, 370, and 390 cm–1 in the reduced form.
725
It was first shown by Mössbauer
that upon reduction one
of the irons changes to Fe2+ (Figure 23). Mössbauer of the oxidized state shows a
narrow quadruple
doublet with δ = 0.27 mm/s relative to iron and a splitting
of 0.6 mm/s. The doublet position is temperature-independent, and
the splitting shows a slight decrease at temperatures higher than
200 K. The spectrum in the reduced form is temperature-dependent and
more complex, primarily because of magnetic hyperfine interactions
and quadruple interactions. The reduced state shows δ = 0.55–0.59
mm/s at 200 K. The A tensor of these proteins is more
symmetric along the z axis. In the reduced state,
Mössbauer of ferredoxins reveals two quadruple doublets, one
at δ = 0.30 mm/s and the other at δ = 0.72 mm/s, indicating
two localized irons.
529,539,726
NMR studies show that, in the reduced state, the protein has
a
mixed-valence Fe2+/Fe3+ state, with the iron
closer to the surface being in the Fe2+ form. Solvent exchange
studies by NMR suggested that reduction of the cluster might increase
accessibility of protons to the cluster. NMR studies were used to
analyze the interaction of ferredoxins with their redox partners to
find their contact points. Chemical shift changes upon reduction have
been assigned. NMR has also been extensively used for structure assignment.
NMR studies showed differences between plant-type and mammalian-type
ferredoxins. While plant-type proteins show a downfield shift of Cys
ligands in the reduced state, with the ligands of Fe3+ showing
Curie-type behavior and Fe2+ ligands showing anti-Curie
behavior, vertebrate-type proteins have both upfield and downfield
signals of cysteine ligands in their reduced state, and all show Curie-type
behavior.
539,727
3.4.3.3
[3Fe–4S]
and [4Fe–4S] Clusters
3.4.3.3.1
Structural Aspects
These clusters
are mainly found in bacteria and usually consist of either one or
two [3/4Fe–4S] clusters. [4Fe–4S] clusters are known
to be the first clusters formed in the early earth environment and
function as ubiquitous ET members in most anaerobic bacteria. The
cluster takes the form of a distorted cube, with iron and sulfur atoms
positioned alternatively in the apexes. Three inorganic sulfurs and
one thiol from a cysteine in the protein coordinate each iron. The
cysteine ligands are arranged in a C-(Xxx)2-C-(Xxx)2-C motif, the so-called classic
[4Fe–4S] motif. The
cluster resides in a common ferredoxin motif (βαββαβ)
with four β-strands, two linking helices, and cluster binding
loops. This fold is the most ancient ferredoxin fold and is very versatile,
with lots of insertions and deletions observed in different proteins
of the family.
91,92,539
The 2[4Fe–4S] or eight iron clusters are hypothesized
to emerge from a gene duplication of the ancestral [4Fe–4S]
cluster.
91
A clostridial 2[4Fe–4S]
protein was the first ferredoxin discovered. Due to its high iron
content, a large portion of the protein consists of inorganic materials
in these proteins.
91
The positions of cysteines
in all [4Fe–4S] or 2[4Fe–4S] proteins are very similar.
The proteins with two clusters can be divided into five subcategories
on the basis of their sequence and evolutionary relationship, including
the clostridial type, chromatium type from green and purple bacteria,
azotobacter [3Fe–4S][4Fe–4S] type, archaebacteria type,
and single [4Fe–4S] type.
729
The
essence of this characterization is sequence homology of 27 ferredoxins
and their deviation from basal architecture, which is a two-subunit
structure resulting from gene duplication with a three-linker connector
and an (Xxx)7-CysI-(Xxx)2-CysII-(Xxx)2-CysIII-(Xxx)8-CysIV motif in each subunit (Figure
24).
729
Figure 24
Structures of the five
classes of two-subunit ferredoxins.
Clostridial-type ferredoxins follow the basal architecture
and
have a conserved motif of Cys-(Xxx)2-Cys-Gly-(Xxx)-Cys-(Xxx)3-Cys-Pro. This motif
usually contains no other cysteine except
in the case of a small number of proteins, including PaFd, which contains a ninth
cysteine in its 22 position. The proteins
consist of two homologous halves that arrange in a pseudo 2-fold symmetry,
with three of the cysteine ligands coming from one half and the fourth
cysteine being provided by the second half, adjacent to a proline.
In 2[4Fe–4S] clusters, the [4Fe–4S] clusters are surrounded
by two antiparallel β-strands and two α-helices. In the
final arrangement of the protein, two sets of antiparallel β-sheets
with two strands lie beneath the clusters and two short helices are
positioned on top of the cluster. An array of water molecules facilitates
H-bonding between two halves of the protein. In clostridial ferredoxins,
there is a conserved Pro after the last coordinating Cys. Although
mutations of this Pro has shown that it is not necessary for the cluster
arrangement, it provides an optimal environment for the next cluster
by both providing hydrophobicity and supporting a specific turn mode
for binding.
91,674,730
In contrast, chromatium-type ferredoxins in most cases contain
a ninth cysteine in positions 2–8, between the second and third
cysteines in the clostridial core. They also have a C-terminal extension
relative to the clostridial sequences. Further classifications within
this class are possible on the basis of the position of their ninth
cysteine and the length and arrangement of their extension, including
photosynthetic ferredoxins, chromatium-type ferredoxins, and dimeric
2[4Fe–4S] ferredoxins. Chromatium-type ferredoxins have their
ninth cysteine close to cluster I. In addition, they have an extended
loop and a short α-helix next to cluster II. The presence of
this loop results in a positive Fe–S–Cα–Cβ torsion angle, compared to the negative
angle in clostridial-type ferredoxins. Moreover, the backbone orientation
around this loop is changed so that this cluster I has one less NH···S
H-bond.
731
Lack of this H-bond results
in a slightly shorter Fe–S bond. These clusters are unstable
at room temperature, at pH values below 6.5, and in the presence of
oxygen.
674
The azotobacter-type ferredoxins
have two residues inserted after
CysII in their subunit 1, and the CysII is mutated to Ala. Their subunit
2 is intact, apart from a 48- or 49-residue extension of the C-terminus.
While this extension is similar within members of the group, it differs
substantially from that of other groups.
729
The archaebacteria-type ferredoxins have a conserved central
domain
in each subunit, but further modifications are observed in regions
before or after this domain, such as an extension of the N-terminus,
or an insertion before the linker. CysII in this class is mutated
to an Asp, resulting in a [3Fe–4S] cluster that can become
a [4Fe–4S] cluster under certain conditions.
729
The single [4Fe-4S] group has both domains, but the
conserved motif
in subunit II is disrupted due to replacement of two to four of the
cysteines with other nonligating residues. Members of this group cannot
be grouped further due to differences in their sequence and structure.
729
Chemical modification studies showed
that neither the N- nor C-terminal
Fe–S binding motif can form a stable cluster in 2[4Fe–4S]
proteins, but their combination will result in formation of a stable
cluster.
674
Using a protein maquette of
[4Fe–4S] ferredoxins and step-by-step replacement and truncation
of amino acids, several minimal essential features have been derived
for formation of a [4Fe–4S] cluster, including the spacing
between Cys residues, the importance of noncoordinating amino acids
in assembling and stabilizing the cluster, preferable use of Cys ligands,
the requirement of only three Cys ligands for formation of a single
cluster, and the requirement of only a consensus core motif of CysIleAlaCysGlyAlaCys.
732
Figure 25 shows consensus
motifs in [3/4Fe−4S] ferredoxins.
Figure 25
Consensus sequences
in ferredoxins. Reprinted with permission from
ref (740). Copyright
2007 University Science Books.
The [3Fe–4S] cluster can be thought of as a cubane
[4Fe–4S]
cluster missing one of the irons. This class is found exclusively
in bacteria, mainly anaerobic bacteria, and is involved in anaerobic
metabolism. The [3Fe–4S] clusters can emerge from oxidative
damage of [4Fe–4S] clusters, as in the case of aconitase, or
treatment of 4Fe clusters with potassium ferricyanide or can be found
as intrinsic constituents of natural proteins, such as mitochondrial
complex II and nitrate reductase. In all cases, the true reason for
the presence of such clusters is not yet completely understood. It
has been shown that [3Fe–4S] and [4Fe–4S] clusters can
be interconverted under certain physiological conditions and the exchange
between 3Fe and 4Fe can be used as a regulatory mechanism. The [3Fe-4S]
clusters have the Cys-(Xxx)2-Cys-(Xxx)2-Cys
motif similar to the [4Fe-4S] clusters but the middle Cys is replaced
by an Asp in most of them.
734
It has been
shown that replacement of the Asp with Cys can change the cluster
into a complete [4Fe–4S] type.
734,735
Addition
of two extra amino acids between the second and third cysteines can
also change a [4Fe–4S] cluster into a [3Fe–4S] cluster.
733
Another common motif, Cys-(Xxx)7-Cys, is found in [3Fe-4S]
cluster of 7Fe-containing proteins, some of which are thermostable
and air-stable. Another Cys following this motif serves as the third
ligand to the cluster. The presence of seven irons in [3Fe–4S][4Fe–4S]
clusters has been confirmed by a combination of techniques such as
EPR, Mössbauer, and X-ray crystallography. There are examples
of Asp residues and hydroxyl groups from the solvent as ligands. As
with 2[4Fe–4S] clusters, the [3Fe–4S][4Fe–4S]
clusters are capable of two-ET. The [3Fe–4S] cluster can be
found in two states: [3Fe–4S]1+ and [3Fe–4S]0, with overall spins of 1/2 and 2, respectively.
H-bonds play
an important role in stabilizing the reduced state. The number of
these bonds is related to the extent of solvent accessibility of iron,
but there are on average six such interactions that direct protons
to the site.
737
The N-terminal structure
of the 7Fe proteins is similar to that of 8Fe proteins, consisting
of a central part with four β-strands that have the Fe–S
cluster in the middle. Two short α-helices connect the loops
in β-sheets. The structure has a partial 2-fold symmetry that
is disrupted at the N-terminus by differences in Cys ligands to the
[3Fe–4S] cluster. There are two nonligand Cys residues next
to each cluster. Although the clusters are positioned close to the
surface, the presence of hydrophobic and aromatic residues protects
them from the solvent. The [3Fe–4S] cluster is very similar
to the [4Fe–4S] cluster, with Fe–Fe distances shorter
than S–S distances, and very similar Fe–S distances.
However, the protein matrix distorts the [3Fe–4S] cluster,
while the [4Fe–4S] cluster is more symmetric.
737
Conserved hydrophobic residues are shown to be important
for the
stability of the protein but not for ET.
738,1085
The thermostable ferredoxins have been shown to have extra salt
bridges in their C-terminus as well as an extra flexible hydrophobic
loop.
739
3.4.3.3.2
Function
The [4Fe–4S]
clusters are important in hydrogen evolution in anaerobic bacteria,
in which the reduced form of ferredoxin transfers electrons to H+ as the final acceptor.
In Clostridium, reduction of ferredoxin is coupled to pyruvate oxidation. The hydrogenase
complex further oxidizes the reduced ferredoxin. Ferredoxins have
been shown to be important in reactions that couple oxidation of the
substrate with reduction of NAD(P)+, flavin mononucleotide
(FMN), FAD, riboflavin, sulfite, and N2. They can bridge
excitation of chlorophyll by light to reduction of NAD. Conversion
of formate to CO2 is often ferredoxin-coupled.
9005
The role of [3Fe–4S] clusters
is less well-known. It has been reported that they can act in sulfite
reduction. A role as iron storage has also been proposed. The [3Fe–4S]
clusters have been observed in the monooxygenase system of Streptomyces griseolus.
741
The 2[4Fe–4S] clusters are mainly found in anaerobic
bacteria
and Clostridium species. However, there
are multiple reports of their occurrence in other organisms such as Micrococcus lactolyticus,
Peptostreptococcus
esldenii, Methanobacillus omelianski, certain photosynthetic bacteria such as Ch.
vinosum, Chlorobium limicola, and Rb. capsulatus, and several extremophiles.
674
There are several ways to test the activity
of [3/4Fe–4S]
ferredoxins. Clostridial-type ferredoxins are usually assayed using
their ability to reduce NADP either in an NADP:ferredoxin reductase
system or in a phosphoroclastic system. Coupling H2 oxidation
to the reduction of an organic dye is another assay used to monitor
the concentration and activity of ferredoxins.
674
3.4.3.3.3
Important Structural
Elements
The [3/4Fe–4S] clusters, like other Fe–S
clusters,
display very low reduction potentials. The reduction potential of
[4Fe–4S] clusters usually ranges from −250 to −650
mV, with an average of −400.
540,737
The common
reduction potential for [3Fe–4S] clusters ranges from −50
to −450 mV, with an average of −100 to −150.
540,737,887
Several methods have been used
to monitor the reduction potential of the clusters, such as potentiometric
CD titration, direct CV, and spectroscopic potentiometry.
737,742
In the case of 7/8Fe proteins, the reduction potentials of the two
sites can be similar (isopotential) or differ by values as high as
192 mV.
743
The same factors that control
the reduction potential of clusters affect the reduction potential
of each cluster within a multiple-cluster protein. Usually the greater
the difference between the reduction potentials of two clusters, the
lower the ET rate between the two. Mutational analyses of conserved
residues that are thought to be important in the intramolecular ET
showed no significant decrease, but less stability. It was postulated
that the geometry and relative orientation of the two clusters are
the factors important in determining this rate. A role for amide dipoles
has also been suggested.
737
It has been
shown that the number of these bonds and more importantly the overall
dipole around the cluster play essential roles in the reduction potential.
718,719
A major part of reduction potential analyses of these types
of ferredoxins deal with roots of differences between them and HiPIPs.
These types of studies are discussed in detail in the section on HiPIPs
(section 3.4.5).
Peptide models of [4Fe–4S]
proteins showed that the reduction
potential of the center is dependent on the number of Cys residues
in the oligomer and will stabilize higher oxidation states, hence
decreasing the reduction potential, with increasing cysteines. These
studies also showed the importance of NH···S in determining
the reduction potential of 4Fe ferredoxins and their difference from
HiPIPs.
717
The reduction potential
of the [3Fe–4S] cluster is pH-dependent.
The pH dependence is related to proton transfer via the conserved
Asp next to the cluster.
745
Mutation of
this Asp to Asn lowers the proton transfer and gates oxidation. Other
studies show a less significant role for the conserved Asp, suggesting
protonation of the cluster itself as the main cause of the pH-dependent
behavior.
746
Also, it has been shown that,
in a protein film electrochemical setup, a hyper-reduced [3Fe–4S]2– cluster can be
formed.
716,747
The presence of a fifth Cys residue close to the cluster can
lead
to formation of a SH···S H-bond and tune the activity
by lowering the reduction potential.
9006
This effect is important in fine-tuning the reduction potential
of proteins with two clusters. Moreover, there are around 15 partial
positive charges in ferredoxins that result in an overall positive
environment of the cluster, which is suggested to be a reason for
the lower reduction potential of these ferredoxins compared to their
higher reduction potential counterparts, HiPIPs.
674
Introduction of a His near the cluster of a 7Fe protein
caused
a 100–200 mV increase in the reduction potential. The reduction
potential of this variant was pH-dependent. At pH values where the
His was protonated, this large increase in reduction potential was
attributed to placement of a positive charge next to the cluster.
A dipole moment directed toward the cluster was proposed as the main
cause of increased reduction potential when the His was neutral.
277
Mutations of conserved Pro in CpFd resulted in
changes of the reduction potentials of the two clusters. NMR studies
of these mutants showed that signals from the β-proton to cysteine
sulfur were changed by these mutations.
730
Mutational analysis of conserved Asp and Glu residues in the CpFd show negligible
changes in the redox properties.
748
Replacement of AvFdI amino
acids with their counterparts in PaFd showed no change
except for small changes in the case of a Phe → Ile mutation,
casting doubt on the role of single amino acids in the reduction potential
differences.
749
A Cys→ Ala mutation
resulted in a 100 mV lower reduction potential of the cluster, mainly
due to changes in coordination geometry and accommodating a new Cys
ligand.
750
Resonance Raman studies
on the cluster showed a very similar environment
of the cluster in different proteins and suggested a role for the
Fe–S–Cα–Cβ torsional
angle in fine-tuning the reduction potential of the site.
618
Solvent accessibility and cluster solvation
also play important
roles in determining the reduction potential of these clusters. More
buried clusters have higher reduction potentials.
92,749,752
The protein dipole Langevine
dipole (PDLD) model was used to analyze
the important features of the reduction potential. On the basis of
these calculations, the number and orientation of amide dipoles, and
not necessarily their involvement in H-bonding, are the most important
factor sin defining the reduction potential. Addition of more amide
dipoles by site-directed mutagenesis indeed resulted in a more positive
reduction potential in cases where the backbone conformation did not
change drastically.
737
Another study suggested
that not the absolute number of H-bonds, but the net dipole moment
on the cluster is the determining factor in the reduction potential
of the cluster.
752
While factors
important in determining reduction potentials of
[3/4Fe–4S] clusters have been found, their effects are not
conclusive. It seems that different factors have different degrees
of importance in different proteins. While surface charges seem not
to be important in CpFd, they showed significant
effects on the reduction potential in other proteins.
753
Studies on CvFd showed that
the two clusters have different reduction potentials, with one being
extremely low, ∼−600 mV. Although it seems that the
cluster with classical geometry should be the one with a normal reduction
potential, thorough mutational and electrochemical studies on this
protein proved it to be the other way.
753
3.4.3.3.4
Spectroscopic Features
Proteins
with more than one cluster are usually brown in color, with a broad
absorption in the 380–400 nm region. Usually an R(390 nm)/Z(280 nm) ratio of more
than 0.7 is observed
for these proteins.
9005
CD and MCD analyses
showed that the [3Fe–4S] cluster of 7Fe proteins is protonated
at acidic pH.
539,746
The [4Fe–4S] clusters
go from a 2Fe3+–2Fe2+ EPR-silent state
(S = 0) to an Fe3+–3Fe2+ (S = 1/2) state with an EPR signal of around 1.96,
while [3Fe–4S] clusters have an EPR signal with a feature at
2.01, going from [3Fe–4S]1+ to [3Fe–4S]0 (Figures 26 and 27). Although the EPR signals
are similar between this class
of ferredoxins and [2Fe–2S] ferredoxins, the relaxation time
of the [2Fe–2S] clusters differs from that of the [4Fe–4S]
clusters, with a common trend of [2Fe–2S] < [3Fe–4S]
< [4Fe–4S]3+ < ferredoxin-type [4Fe–4S]1+. Therefore, the temperature dependence of
the EPR signal
can be used as a guide to the cluster type. However, care should be
taken in analysis of the signals, because spin–spin interactions
between clusters can lead to an enhanced relaxation time.
754
Figure 26
Representative spectra of [4Fe–4S] proteins.
(a, left) UV–vis
of the oxidized form. Reprinted with permission from ref (760) Copyright 2005 Springer-Verlag.
(b, middle) EPR of the [4Fe–4S]1+ state. Reprinted
with permission from ref (761). Copyright 1999 Elsevier. (c, right) Mössbauer of
the [4Fe−4S]2+ cluster of the E.
coli FNR protein, T = 4.2 K (top),
and the [4Fe−4S]1+ cluster of E.
coli sulfite reductase, T = 110 K
(bottom). Reprinted with permission from ref (529). Copyright 1997 American
Association for the Advancement of Science.
Figure 27
Representative spectra of the [3Fe–4S] cluster. (a, left
top) UV–vis of the oxidized form and (b, right) temperature-dependent
EPR of the [3Fe–4S]1+ cluster. Reprinted with permission
from ref (762). Copyright
2002 Elsevier. (c, left bottom) Mössbauer of the [3Fe–4S]1+ (top) and [3Fe–4S]0 (bottom)
clusters
. Reprinted with permission from ref (529). Copyright 1997 American Association for
the
Advancement of Science.
The [3Fe–4S] clusters have a Mössbauer spectrum
with
one quadruple doublet at δ = 0.27 mm/s, showing three equivalent
Fe3+ sites in the oxidized state (Figure 27). The reduced form shows two doublets
with a 1/2 ratio in
intensity. The minor doublet at δ = 0.32 mm/s is assigned to
Fe3+, while the major doublet at δ = 0.46 mm/s is
attributed to a delocalized mixed-valence Fe2.5+ state.
529
The Mössbauer features of the [4Fe–4S]2+ cluster have been discussed in detail in
the section dealing
with the spectroscopic features of HiPIPs (section 3.4.5.5).
NMR is one of the tools that has
been extensively used to analyze
[3/4Fe–4S] clusters. A higher number of total hyperfine shifted
resonances in NMR can indicate the presence of more than one cluster
in a given protein. Nine or twelve contact shifts are usually observed
for [3Fe–4S] or [4Fe–4S] clusters, respectively. The
[4Fe–4S] clusters are identified by the presence of peaks with
anti-Curie temperature dependence, while Curie-type behavior is indicative
of a [3Fe–4S] cluster. Typical 7Fe ferredoxins show five downfield
peaks, two with Curie-temperature-dependent behavior. There are, however,
7Fe proteins with quite different NMR spectra and more downfield peaks.
These 7Fe proteins usually have a short symmetric motif. A peak at
30.0 ppm is characteristic of mononuclear 3Fe clusters.
736
In NMR studies of the [3Fe–4S] clusters,
it has been shown that the contact shifts of His close to the conserved
Asp are pH-dependent and correlate with the pK
a of the Asp residue. Also, the effects of disulfide bonds
in the shifts were studied.
755
NMR of [4Fe–4S]
clusters showed very similar shifts for all Cys residues in the oxidized
form. Upon reduction, a similar pattern is observed for all [4Fe–4S]
proteins, with two Cys residues showing Curie-like behavior (Fe2.5+) and two showing
anti-Curie behavior (Fe2+).
This also suggests that there are two isoforms with an Fe2.5+ pair on the Cys I/III
or Cys II/IV pair. The former is more preferred,
and this preference is stronger when a disulfide bond is present,
as shown by NMR studies.
755
The effects
of other ligating residues were also analyzed in terms of NMR contact
shift. NMR was also used to analyze the self-exchange rate and hence
reorganization energy in ferredoxins.
756
NMR studies provided structures of several ferredoxins such as [4Fe–4S]
ferredoxin from Tt. maritima.
757
The resonance Raman spectra of [4Fe–4S]
ferredoxins can
be explained without considering coupling between Fe–S and
δ(S–C–C) modes. For these proteins at least seven
ν(Fe–Sβ) bands and three ν(Fe–St) bands are observable, with a band at 340 cm–1 being
the most prominent due to total symmetry of the cubane structure.
Resonance Raman was also used to study Se complexes of ferredoxins
as well as the presence of [3Fe–3S] clusters. Resonance Raman
studies revealed the solvent accessibility of H-bonds to the cluster,
the distorted D
2d
symmetry
of the cluster, and Fe–S–Cα–Cβ torsion angles.
618,758
NRVS was also used
to study the dynamics and the oxidized and reduced states of the [4Fe–4S]
cluster.
759
3.4.3.4
Ferredoxin-like
Proteins
A class
of so-called plant ferredoxin-like proteins (PLFPs) has been discovered
in the past few years. These proteins are known to play a role in
several cellular processes. The first PFLP was discovered in sweet
pepper. The protein consists of three domains: a N-terminal signal
peptide, a [2Fe–2S] domain, and a casein kinase II phosphorylation
(CK2P) site at the C-terminus. Phosphorylation of this domain is postulated
to be important in resistance to pathogens in Arabidopsis
thaliana,
763
and PLFPs
are evolved in plant defense mechanism pathways.
[4Fe–4S]
ferredoxin-like proteins are also common and are found in some bacteria
with a modified Cys-(Xxx)2-Cys-(Xxx)2-Cys-(Xxx)3-Cys motif at the N-terminus or Cys-(Xxx)2-Cys-(Xxx)8-Cys-(Xxx)3-Cys-(Xxx)5-Cys
at the C-terminus.
The ferredoxin-like protein in Rhizobium meliloti is shown to be important in nitrogen
fixation. The protein is located
in an operon with nif genes. Mutational analyses and molecular modeling
showed the importance of extra amino acids in positioning the loop
in a way that it could incorporate the cluster efficiently.
764,765
A PLFP has been discovered in Erwinia carotovora that is regulated by quorum sensing.
This ferredoxin has similarity
to plant ferredoxins with no significant similarity to bacterial ferredoxins.
766,767
PFLP genes in Helicobacter pylori and its corresponding ferredoxin reductase have
been shown to be
important in imparting metronidazole resistance to the bacteria.
768
PFLPs are known to be important in enhancing
plant resistance to bacterial pathogens. Transgenic expression of
PFLP from sweet pepper in calla lily resulted in more resistance to
soft rot bacterial diseases.
769
The same
transformation in tobacco, orchid, and rice plants enhanced their
resistance to Xanthomonas oryzae pv. oryzae.
767
3.4.4
Rieske Centers
3.4.4.1
Introduction/History
Rieske proteins
are [2Fe–2S] iron–sulfur proteins that are distinguished
by their unique His2-Cys2 ligation motif. The
first example of these proteins was discovered by Rieske in 1964,
who observed an EPR signal with g = 1.90 in the cytochrome bc
1 complex (complex III of the mitochondrial
electron transport chain
770
). Similar EPR
signals were later observed in the b
6
f complex of the photosynthetic chain, the membrane of bacteria
with a hydroquinone oxidizing ET chain, and soluble bacterial dioxygenases.
The coordination environment was first established by ENDOR and ESEEM
spectroscopy and further proved by the crystal structure.
773
There have been multiple reports of the presence
of several isoforms of Rieske proteins in the genome of prokaryotes.
The presence of these isoforms most likely aids the organism to adapt
better to environmental changes.
771
3.4.4.2
Structural Aspects
3.4.4.2.1
Primary Structure/Amino Acid Sequence
The first Rieske protein
to be sequenced was the Rieske protein
from the bc
1 complex of Neurospora crassa.
772
Subsequently,
other gene sequences of multiple Rieske proteins from a wide range
of organisms have been obtained. Sequence alignment and analysis revealed
a Cys-Xxx-His-(Xxx)15–47-Cys-Xxx-Xxx-His motif as
the conserved motif for [2Fe–2S] ligands.
773
On the basis of this sequence analysis, the proteins can
be divided into Rieske and Rieske-type subcategories.
Rieske
proteins can be found in bc complexes such as the bc
1 complex of mitochondria and bacteria, the b
6
f complex of chloroplast,
and corresponding subunits in menaquinone oxidizing bacteria. Three
residues other than Fe–S ligands are also conserved in this
class of Rieske proteins, two of which are cysteine residues that
form a disulfide bond important in the stability of the protein,
774
and the other is a Gly in a conserved Cys-Xxx-His-Xxx-Gly-Cys-(Xxx)12–44-Cys-Xxx-Cys-His
motif. Mutational analysis of
this class confirmed the presence of two histidines and four cysteines
essential for cluster formation.
775,776
Rieske proteins
that are not part of the bc complex also belong to
this class. Some of these proteins are within complexes that are not
well identified, and some belong to organisms that are devoid of the bc complex, such
as TRP from T. aquaticus and SoxF and SoxL from Sl. acidocaldarius.
773,777
Rieske-type proteins are typically
part of water-soluble dioxygenases.
This class of proteins can be further divided into four separate groups. Bacterial
Rieske-type ferredoxins are water-soluble ET proteins
with a [2Fe–2S] cluster that show no similarity to common ferredoxins
and share a conserved Cys-Xxx-His-(Xxx)16–17-Cys-Xxx-Xxx-His
motif. They have diverse sequences, but their three-dimensional structures
are very similar to those of other Rieske proteins. Bacterial
Rieske-type oxygenases have a Rieske center and a mononuclear
nonheme iron in their active site. In addition to four Rieske ligands,
four other residues are conserved in these proteins, including two
glycine residues, one tryptophan, and one arginine. Naphthalene dioxygenase
(NDO) is the archetype of this class. Eukaryotic homologues
of bacterial Rieske-type oxygenases also have a ligand set
for Rieske coordination and a site for mononuclear nonheme iron. Choline
monooxygenase and CMP-N-acetylneuraminic acid hydroxylase
are examples of this class. Lastly, there are proteins that have a putative Rieske
binding site, with a common motif of Cys-Pro-His-(Xxx)16-Cys-Pro-Xxx-His, but the
presence of a Rieske cluster has
not been confirmed in them yet.
773
3.4.4.2.2
Three-Dimensional Structure/Crystallographic
Analysis
Crystal structures of several Rieske proteins from
different categories have been solved. All Rieske proteins share the
so-called “Rieske fold”. This fold consists of three
antiparallel β-sheets that form a double β-sandwich (Figure 28). Sheet 1 consists
of three conserved strands,
1, 10, and 9. Strands 2, 3, and 4 form sheet 2, and strands 5–8
are in sheet 3. Sheet 2 is longer and interacts with both sheets 1
and 3. The interactions between sheets 2 and 1 are mostly of hydrophobic
nature. Most conserved residues are found in the loop regions connecting
the β-strands, especially loops β1−β2, β2−β3,
and β8−β9 (the so-called “Pro loop”).
91,773
Figure 28
Minimal Rieske fold with three β-sheets and loops coordinating
the [2Fe–2S] cluster with two His ligands and two Cys ligands
(from PDB ID 1NDO).
The cluster binding subdomain
is mainly located in sheet three
and two of its adjacent loops (β4−β5 and β6−β7).
Each loop provides one of the cysteine and histidine ligands, so the
pattern is 2 + 2, in contrast to the 3 + 1 pattern observed in most
ferredoxins. In mitochondrial and chloroplast Rieske proteins, there
is a disulfide bridge that connects the loops in Rieske proteins.
This disulfide bond is of prominent importance in maintaining structural
integrity in these proteins because their loops are exposed to solvent.
Rieske-type proteins do not have this conserved disulfide bridge.
It has been argued that this difference is due to the fact that buried
Rieske complexes are stable without the need to disulfide bond.
773
Rieske proteins from bc
1 or b
6
f complexes
have an additional
“Pro loop” with a highly conserved sequence of Gly-Pro-Ala-Gly
that covers the cluster and has been shown to be critical for the
stability.
773,779
In most cases the Fe2+ is the one that is more surface-exposed, and it is this iron
atom
that has two exposed His ligands. In buried Rieske complexes such
as NDO, the histidines are not solvent-exposed and usually form H-bonds
with acidic side chains in the active site.
780
The geometry of the Fe–S cluster is the same among all Rieske
proteins, forming a distorted tetrahedral conformation. In contrast
to the Cys ligands which impart a tetrahedral geometry, the His ligands
accommodate a geometry that is closer to octahedral (Figure 29).
773
Figure 29
Structure of the bc
1 complex from chicken
(PDB ID 3H1J) and its Rieske protein and Rieske center (left) and structure of
the b
6
f complex from Mastigocladus laminosus (PDB ID 1VF5) and its Rieske
protein and Rieske center (right).
Multiple H-bonds constrain and stabilize the cysteine ligands,
which are conserved between most bc
1 and b
6
f Rieske proteins. They are
three bonds with sulfur S1, two with sulfur S2, two with S
y
of cysteine in loop 1, and 1 with S
y
of loop 2. Usually there are H-bonds between the
sulfurs of coordinating cysteines and the main chain nitrogen of residue i + 2. These
H-bonds are known to stabilize type I turns.
Two of these H-bonds are of the OH···S type, one from
a conserved Ser to the bridging S1 and one from a conserved Tyr to
the Cys in loop 1. Rieske proteins from menaquinol oxidizing organisms
lack this Ser···Cys H-bond. Rieske-type proteins lack
three of these conserved H-bonds due to a lack of the conserved Ser
and Tyr. Multiple site-directed mutagenesis studies confirmed the
importance of these two H-bonds in maintaining the high reduction
potential of Rieske proteins.
773,781
Despite the
high degree of structural similarity between different
Rieske and Rieske-type proteins, each category has its unique features.
It seems that although the cluster-binding site and the minimal Rieske
fold are highly conserved among all classes of Rieske and Rieske-type
proteins, there are multiple insertions between elements of this minimal
fold, mainly in loop regions. These significant differences make sequence
alignments of Rieske proteins controversial, compared to their ribosomal
RNA alignments.
782
Rieske-type ferredoxins
have the closest structure to the minimal fold. Rieske proteins from
the b
6
f complex usually
have a C-terminal extension that is known to be important in stabilizing
the open conformation required for the activity. The same role was
proposed for helix–loop insertion in mitochondrial Rieske proteins.
Chloroplast Rieske proteins also show a distortion in the β-sheets,
forming a β-barrel rather than a β-sandwich.
773
Novel disulfide bonds have been reported at
the C-terminus of a thermophilic Rieske protein from Acidianus ambivalence that are
reported to be important
in higher stability of the protein.
783
A
disulfide bond and extended C-terminal region insertion have been
observed in archaeal Rieske proteins.
784
Some acidophilic proteins have extended β-strands in their
cluster binding domain. The peptide bond orientation differs in the
Pro loop of bc
1 and b
6
f complexes in regard to the cis or trans configuration.
773
Some Rieske proteins have a very long loop
in place of the Pro loop that is important for interacting with redox
partners.
785
Although the pattern of H-bonding
and salt bridges is similar, it is not identical, and the residues
that are involved are not conserved.
773
Another difference between Rieske proteins lies in their surface
charge distribution. These differences are required for interactions
with different redox partners. Different charge distribution also
reflects the variation of pH in which the proteins work, as exemplified
by a net negative charge on the surface of some acidophilic proteins.
786
The Rieske fold and the geometry of the
cluster are unique to Rieske
and Rieske-type proteins and differ significantly from those of the
other class of [2Fe–2S] ferredoxins. The most similar geometries
are those of rubredoxins and the zinc-ribbon domain, suggesting that
the Rieske fold may have arisen from a mononuclear ancestral fold.
91
3.4.4.3
Function
3.4.4.3.1
Rieske Clusters: Cytochrome bc Complexes
Mitochondrial bc
1 complexes and chloroplast b
6
f complexes are multisubunit proteins with four redox centers
organized in three subunits: two heme b centers in
a transmembrane domain of cytochrome b, cytochrome c
1/f, and the Rieske iron–sulfur protein.
All of them oxidize hydroquinone (ubihydroquinone/plastohydroquinone)
and transfer electrons to either cytochrome c or
plastocyanin, generating a proton gradient across the membrane through
the Q-cycle. For proper function of this cycle, the hydroquinone oxidation
reaction is strictly coupled. The Rieske protein is responsible for
hydroquinone oxidation and acts as the first electron acceptor. Electron
transfer is accomplished by direct interaction between the exposed
His ligand and the quinone substrate.
787
Since the function of the Fe–S cluster in these protein complexes
is tied to hemes, a more detailed explanation will be presented in
section 5.
3.4.4.3.2
Rieske-Type Clusters: Dioxygenases
Rieske-type clusters
are part of aromatic ring hydroxylating dioxygenase
enzymes that catalyze the conversion of aromatic compounds to cis-arenediols, a key
step in aerobic degradation of aromatic
compounds.
788
Dioxygenases contain a reductase,
a terminal oxygenase, and often a [2Fe–2S] ferredoxin. The
reductase part can be of two types: ferredoxin–NADP or glutathione.
The oxygenase part contains a Rieske center and a mononuclear nonheme
iron center (Figure 30). The Rieske center
transfers an electron from ferredoxin or reductase to the iron center.
788
Although these two centers are in different
domains that are far apart in a single subunit (45 Å), the quaternary
structure with 3-fold symmetry will bring them to a close distance
within 12 Å. In most cases the His ligand of the Rieske center
and one of the His ligands of iron are bridged by an Asp residue,
ensuring the rapid ET between the two centers (Figure 31). The removal of this conserved
Asp abolishes the activity
without changing the metalation.
789−791
In the case of 2-oxoquinoline
monooxygenase, the Asp changes its position after reduction of the
Rieske center to H-bond with a His ligand that is protonated upon
reduction. This repositioning will cause a conformational change that
results in generation of a 6-coordinated iron geometry which is more
active.
792
It has also been suggested that
the H-bonds provided by this Asp can help the Rieske center and catalytic
center to sense the redox state and ligand state of each other. Mutational
studies have been implemented to discover sites that are important
in specific interactions between these Rieske centers and their redox
partners.
793
Figure 30
Structure of naphthalene
1,2-dioxygenase (PDB ID 1NDO), the archetype
of Rieske-type proteins from two different views, and a close-up of
the active site Rieske center.
Figure 31
Interface between two monomers of naphthalene dioxygenase, NDO.
Asp205 from the polypeptide chain on the left bridges two His residues
that are ligands to the Fe–S cluster and catalytic nonheme
iron center (PDB ID 1NDO).
3.4.4.4
Important
Structural Elements
As with any other ET centers, the reduction
potential of Rieske centers
is one of the most important factors in determining its ET rate and
conveying its activity.
794
Any changes
in the reduction potential of Rieske and Rieske-type proteins have
been shown to affect their activity and the kinetics of the ET between
these centers and their redox partners. Reduction potentials of Rieske
centers vary in a wide range of −100 to +490 mV, which is significantly
higher than the average reduction potentials of ferredoxins. In general,
any factor that selectively stabilizes either the reduced or oxidized
state of a Rieske center will influence its reduction potential.
773
The difference between the overall charge of
the cluster (0/–1 in the case of Rieske proteins vs −2/–3
in the case of ferredoxins) and electronegativity of the ligands (histidine
vs cysteine) is the main reason for the higher reduction potential
of Rieske proteins. Different H-bonds to bridging or terminal sulfurs
and solvent exposure of the clusters are the main determinants of
different reduction potentials within the Rieske family. The reduction
potential range differs depending on the type of Rieske complex: 265–310
mV in the bc
1 complex and around 320 mV
in the b
6
f complex. The
reduction potentials of menahydroquinone oxidizing complexes are 150
mV lower than that of the ubihydroquinone bc
1 complex (the same difference that is observed between the
two types of quinones).
773
This lower reduction
potential has been attributed to a lack of a H-bond donated from a
conserved Ser, which is absent in the former class of Rieske proteins.
Different methods of reduction potential measurement have been applied
to Rieske proteins, such as chemical redox titration monitored by
EPR
795
or CD
796
and direct cyclic voltammetry,
797,798
which enables
measurement of thermodynamic parameters.
780
CV experiments also showed for the first time the second reduction
step to a 2Fe2+ state at very low reduction potentials
(∼−840 mV).
797
Computational
studies showed that the cluster distortions caused by the protein
environment play a prominent role in tuning the reduction potential
of the center. Accordingly, using active site structures determined
from x-ray crystallography will result in calculations that agree
much better with experimental values than idealized structures.
800
An interesting feature of Rieske proteins
is their pH-dependent
reduction potential, which decreases with increasing pH and is attributed
to deprotonation of a group in contact with the Rieske complex.
773,801
This phenomenon can be observed in the oxidized state where the
pK
a values of one of the His ligands are
near physiological pH (two pK
a values
of 7.8 and 9.6 vs one pK
a of around 12.5
in the reduced state
802
). This pH dependence
can be important in interactions and binding of Rieske proteins to
their redox partners. Moreover, this redox-dependent ionization may
be very important for their physiological function, as these proteins
are part of proton-coupled ET systems. The biomimetic models of Rieske
clusters prove the dependence of the reduction potential of the center
on the protonation state of its His ligands.
803
Shifts in the UV–vis absorption peaks and CD features upon
pH titration are consistent with the two protonation states of the
oxidized form.
804
Several studies have
shown that multiple inhibitors can bind to the sites close to the
cluster and affect the reduction potential of the site.
787,805,806
In a related study, diethyl
pyrocarbonate (DEPC) was used to react
with and trap deprotonated His. Addition of this ligand caused reduction
of the cluster as well as an increase in the overall reduction potential,
a phenomenon that was observed in the case of inhibitors such as stigmatellin,
immobilizing it in the b conformation. Moreover, if the protein was
reduced first, no addition would be observed, due to a lack of available
deprotonated His.
806,807
Analysis of some pH-independent
low reduction potential Rieske proteins suggests that the coupling
between the cluster oxidation state and the His protonation state
also has a role in determining the reduction potential of the cluster.
808
The reduction potentials of Rieske-type
clusters are lower than
those of Rieske clusters, with values around −150 to −100
mV.
773,780,788
One reason
for this difference is a lack of three out of eight conserved H-bonds
of Rieske proteins in Rieske-type proteins (Figure 32).
781
Reduction potential of Rieske-type
proteins is pH-independent due to less solvent accessibility in comparison
to Rieske proteins.
809,810
There are examples of Rieske-type
proteins that have very similar active site structure to Rieske centers,
but different loop orientations cause disruption of the H-bonding
network, resulting in proteins with reduction potentials around −150
mV.
810
A Rieske-type ferredoxin has been
found with a reduction potential around 170 mV. The higher reduction
potential in this Rieske-type protein has been attributed to the presence
of amino acid substitutions in positions around the metal center.
795
Figure 32
Differences in the H-bond pattern between the
Rieske fragment of
naphthalene dioxygenase, NDO (PDB ID 2NDO), the water-soluble Rieske fragment of
the bc
1 complex, ISF (PDB ID 1RIE), and the Rieske
fragment from the b
6
f complex, RFS (PDB ID 1RFS). Reprinted with permission from ref (773). Copyright
1999 Elsevier.
The most important residues involved
in the H-bonding network in
Rieske proteins are a conserved serine and a conserved tyrosine. It
has been suggested that this H-bond network stabilizes the reduced
state by charge delocalization, thereby increasing the reduction potential.
773,811
The electrostatic environment of the protein is another feature
that can influence the reduction potential, meaning that the presence
of charged residues on their own can increase the reduction potential
of the center. In one study, removal of negatively charged residues
in the vicinity of the Rieske center in Rieske ferredoxin from biphenyl
dioxygenase of Burkholderia sp. resulted
in a pK
a of the His ligands similar to
that of mitochondrial Rieske proteins.
812
Mutational analyses have been extensively used to reveal features
that are important in tuning the reduction potential. Gly143Asp, Pro146Leu,
and Pro159Leu mutations in the Pro loop resulted in a shift of about
50–100 mV toward more negative reduction potentials, mostly
due to distortion in the Fe–S environment and changes in the
H-bond network around it.
773
The cluster
content was decreased to 32–70% in these mutants. Another study
showed that mutations in the loop containing Fe-S ligands are the
ones that alter reduction potential.
779
Several site-directed mutations were made with the goal of
understanding
the role of H-bonds from conserved Ser and Tyr in different organisms.
781,813,814,817
Mutations of Ser to Ala and Tyr to Phe both decreased the reduction
potential.
781,815
When both mutations were made,
the effects on the reduction potential were observed to be additive.
It was shown that these mutations do not influence the stability of
the cluster or its interaction with quinone. However, the activity
was decreased, demonstrating the importance of the reduction potential
in hydroquinone oxidation activity.
9007
These mutations also increased the pK
a values of the His ligands. Different effects were observed when
these two residues were mutated into other amino acids. Mutations
of Tyr to nonphenolic amino acids targeted the Rieske protein to cytosolic
proteolytic cleavage machinery. A Ser to Cys mutation resulted in
expression of proteins that could no longer incorporate a Rieske cluster,
and in cases where it could, a slight increase in the reduction potential
was observed. A Ser to Thr mutation resulted in a protein with moderate
changes in the midpoint potential.
781
Mutations of a conserved Thr that packs tightly against the Pro
loop resulted in a lower reduction potential and a significant decrease
in the activity.
774
Mutations of a conserved
Leu residue that is supposed to protect the cluster from solvent were
analyzed as well.
816,817
Leu136Gly/Asp/Arg/His mutants
were analyzed and showed low activity and altered reduction potential.
Replacement of Leu with a neutral residue such as Ala caused a similar
change in both reduction potential and pK
a values of the His ligands, suggesting a causative effect of a change
in water accessibility.
816
Mutation to
a negative residue such as Asp has marginal effects on the reduction
potential, probably due to movement of the Asp side chain from His
and its solvation. However, placing a positive charge here resulted
in a significant increase in the reduction potential.
817
Several mutations in a flexible linker
distant from the cluster
binding site have been shown to increase the reduction potential.
773
Mutations in a hinge region were shown to increase
the E
m of the Rieske center of Rb. capsulatus. These mutations affect the reduction
potential in two ways: by altering the interaction mode with quinone,
which is known to affect the reduction potential, and by altering
the positioning of the [2Fe−2S]-containing domain of the Rieske
protein, which can impart changes in both the reduction potential
and the EPR signal shape.
818
Mutations
in the residues involved in disulfide bridge formation also showed
decreased reduction potential values. This lower reduction potential
is mainly due to removal of polarizable Cys groups and disturbance
of the loop conformation and pattern of H-bonds.
817,819
Analyses of a protein with a reduced disulfide also showed a small
decrease in the reduction potential that was attributed mainly to
changes in the H-bonding pattern and enthalpic effects.
820
Similar mutational studies of conserved
residues close to the cluster
binding domain of Rieske-type proteins have also been performed, showing
different effects depending on the mutation type. Mutations of a conserved
Asp residue in Rieske oxygenase resulted in a lower reduction potential
mainly due to deprotonation of a His ligand caused by loss of a H-bond
from Asp.
822
Another important factor
in determining the reduction potential
is the condition in which the protein performs its function. Studies
on extremophilic organisms revealed that Rieske centers from acidophilic
organisms have more positive midpoint potentials than neutral centers
whereas the potentials of acidophilic Rieske centers are significantly
lower than the expected value. Interestingly, the pK
a of the His ligand also shifted correspondingly in these
extremophilic organisms.
786,823
It should be
noted that there are exceptions to these general statements.
There are high reduction potential Rieske proteins, such as sulredoxin,
which lacks the hydroxyl group responsible for redox modulation and
shows a different pH-dependent redox response compared to other high
reduction potential Rieske proteins.
824
3.4.4.5
Spectroscopic Features of Rieske and Rieske-Type
Proteins
As with other Fe–S proteins, Rieske proteins
have broad absorption spectra resulting from overlapping bands from
S → Fe3+ charge transfer (Figure 33). CD and MCD spectroscopic techniques were used
to deconvolute
some of these spectra. In their oxidized form, Rieske proteins have
absorptions at 325 and 458 nm and a shoulder around 560–580
nm. Upon reduction, the position of the bands shifts to 380–383,
425–432, and 505–550 nm and the intensity of the bands
drops by 50%. The CD spectrum of Rieske proteins has features that
are unique among Fe–S proteins, showing two positive bands
between 310 and 350 nm, a negative band at 375–380 nm, and
a set of positive bands between 400 and 500 nm in the oxidized form.
In the reduced form, the CD spectrum shows a positive band at 314
nm, a negative band at 384–390 nm, a negative band at 500 nm,
and a band at 760 nm.
773
These bands are
attributed to d–d transitions of Fe2+ from the lowest
lying d orbital into t2g sets. The strong negative band
at 500 nm in the reduced state is an indicator of the redox state
even in the presence of other cofactors such as heme.
809
The CD spectrum of oxidized Rieske proteins
is pH-dependent in near-UV and visible regions due to the presence
of some deprotonation events.
804
Rieske
proteins show temperature-dependent MCD spectra with multiple positive
and negative bands in the reduced state, but the intense negative
band at 300–350 nm and positive band at 275 nm, which is observed
in rubredoxins and [2Fe–2S] ferredoxins, is not visible in
them due to a blue shift of the bands to higher energies because of
the nitrogen ligation from the His ligand.
773
Figure 33
Representative spectra of Rieske centers. (a) UV–vis of
the reduced (lower spectrum) and oxidized (upper spectrum) forms.
Reprinted with permission from ref (866). Copyright 2004 National Academy of Sciences.
(b) EPR of the reduced form. Reprinted with permission from ref (867). Copyright 2007
National
Academy of Sciences. (c) Mössbauer of the [2Fe−2S]+ cluster of the Rieske protein
from Ps. mendocina at T = 200 K. Reprinted with permission from ref (529). Copyright
1997 American
Association for the Advancement of Science.
Mössbauer studies of Rieske proteins show a temperature-independent
four-line spectrum resulting from two quadruple doublets of the same
intensity (Figure 33). The spectrum of the
reduced form is very similar to that of ferredoxins with a more positively
shifted δ (0.68 mm/s at 200 K), which is due to the less electron-donating
nature of the His ligands.
529,778,847
While the Fe3+ state shows quite isotropic features,
the Fe2+ state has an anisotropic A tensor.
The electric field gradient tensor is symmetric around x axis of the A tensor for
Fe2+, with the largest component
being positive.
773
Resonance Raman
studies of Rieske proteins using laser excitation
at different wavelengths showed features very similar to those of
ferredoxins in both the reduced and oxidized states, with some shifts
in the bands and additional vibrations due to the presence of the
His ligands.
848
The higher number of bands
in the 250–450 cm–1 region is an indicator
of a lower symmetry of the Rieske proteins than those of all cysteinate
[2Fe–2S] ferredoxins (C
2v
vs D
2h
or C
2h
symmetry). Rieske proteins
feature a weak peak at 266–270 cm–1 that
is assigned to the Fe(III)–N(His) stretching mode, which is
thought to have some Fe–Fe mixing character. The peak is shifted
8 cm–1 up in more basic pH, consistent with deprotonation
of the His ligand. The peaks at 260–261 cm–1 are assigned to Fe–His bending modes
and are also very sensitive
to 15N substitution. A peak at 357–360 cm–1 corresponds mainly to Fe(III)–Sterminal
stretching
(B2t) mode.
848
This peak is
very similar to that of ferredoxins, only upshifted due to either
a different H-bond pattern or Fe–Sγ–Cα–Cβ dihedral angles, which is
a sign of similar Fe3+ environments in the two classes
of proteins. This peak can be observed at 319–328 cm–1 after reduction.
725
pH-dependent studies
in the 250–450 cm–1 region show that there
are no resonance Raman-detectable changes at the pK
a of the first His ligand and changes are only observed
above the pK
a of the second His ligand.
These changes arise, however, from additional factors such as protonation
of some amide backbones and not solely in regions related to the Fe–Nimid vibrational
frequency. A lack of changes at physiological
pH can ensure rapid proton-coupled ET.
849
No significant change was observed for Rieske-type proteins. Most
resonance Raman features are due to the Fe–S stretch. The kinematic
coupling observed by resonance Raman and rigidity of the H-bond network
around the cluster help minimize the reorganization energy and hence
facilitate ET.
850
resonance Raman studies
were also performed to analyze the role of the H-bonding network in
Rieske proteins. It has been shown that the presence or removal of
the S···Tyr H-bond shows significant changes in resonance
Raman bands at 320–400 cm–1, whereas removal
of the S···Ser H-bond does not show a detectable resonance
Raman change.
849
XAS analysis showed
very similar geometries of clusters in Rieske
proteins and ferredoxins and also indicated the contraction of the
site upon oxidation. Early XAS analyses were hampered by the fact
that the presence of His ligands was not known. XAS studies of Rieske
oxygenases showed a small but significant change in the Fe–S
bond length upon reduction. A larger increase in the Fe–Nimid bond distance (0.1 A)
was observed through reduction,
which can facilitate ET between the Rieske center and its redox partner.
The edge feature has a shift toward lower energies upon reduction.
847,851
EPR spectroscopy is one of the first techniques used to identify
Fe–S proteins. The g values of Rieske proteins
are significantly lower than those of ferredoxins (1.9–1.91
vs 1.945–1.975) due to the presence of nitrogen ligands (Figure 33). This EPR signal
is mainly due to Fe3+ and its His ligands and environment.
773
EPR signals vary significantly among different groups of Rieske
proteins, with g
z
= 2.008–2.042, g
y
= 1.888–1.92, and g
x
= 1.72–1.834. The
rhombicity changes between 51% in the z axis and
100–59% in the x axis.
773
In Rieske proteins all g values correlate
with rhombicity, indicating that EPR properties are influenced mainly
by the protein environment. Changes in the EPR signal upon binding
to quinone or inhibitors will change the shape of the EPR signal and g values. These
effects can also be correlated to rhombicity
parameters.
773
An EPR study of a Rieske
protein at pH 14 showed increased g values with broadened
features. The appearance of these new features can be assigned to
a decrease in the energy difference between reductions of the Fe with
two His ligands and the one with two Cys ligands due to deprotonation
of both His ligands.
852
ENDOR and
ESEEM studies support the presence of two nitrogen ligands
in both Rieske and Rieske-type proteins.
853
Studies with 15N-labeled protein further support the
presence of nitrogen ligands.
845,854−858
X-band 14N hyperfine sub-level correlation (HYSCORE)
spectroscopy of reduced Rieske and Rieske-type proteins is dominated
by two histidine Nd ligands with hyperfine couplings of
∼4–5 MHz. A combination of site-specific 14/15N labeling together with orientation-selective
HYSCORE studies was
used to gain more insight into the nature of the H-bonding network
around the cluster and through-bond electrostatic effects.
814
ESEEM studies coupled with isotope exchange
with H2O were used to understand the proton environment
around Rieske proteins from Rb. sphaeroides.
859
The magnetic and structural features
of the Cys and His ligand protons and the protons involved in the
H-bonding network were analyzed.
859
1H ENDOR analysis of the Rieske proteins from the bovine mitochondrial bc
1 complex showed three peaks from orientation
behavior: two from β protons of Cys ligands and one from the
β proton of the His141 ligand. The direction of g
max lies in the FeS plane with the largest proton coupling
along g
int.
2006
NMR studies have been applied to different Rieske and Rieske-type
proteins.
861,862
Cysteines coordinated to Fe3+ show four strongly downshifted signals between 50 and
110
ppm. Temperature-dependent studies of Hβ protons
of the cysteines show that they follow Curie law. Hε1 of one of the histidine ligands
shows a sharp resonance at 25 ppm,
showing a weak Curie-temperature-dependent behavior. There are still
complications in assigning all the resonances in NMR spectra due to
the unique features of Rieske NMR. NMR studies were used to monitor
the H-bonding patterns
863
and solvent accessibility.
864
NMR studies on Rieske proteins from T. thermophilus revealed slight conformational
changes
that are dependent on both the oxidation state and ligand binding. 1H, 15N, and 13C
NMR analyses showed
that two of the observable prolyl backbones change from the trans to the cis mode
upon reduction.
865
3.4.5
HiPIPs
3.4.5.1
Introduction/History
HiPIPs are
a well-defined superfamily of Fe–S proteins found mainly in
photosynthetic anaerobic bacteria, although proteins from aerobic
bacteria have also been reported. HiPIPs were expressed in both aerobic
and anaerobic conditions.
868
HiPIPs contain
a [4Fe–4S] cluster as with ferredoxins. However, the higher
reduction potential of HiPIPs results in one less electron in both
the reduced and oxidized states of these proteins compared to ferredoxins,
meaning a [4Fe–4S]2+/3+ state.
872,873
3.4.5.2
Structural Aspects
HiPIPs are
usually small proteins (6–11 kDa). The [4Fe–4S] cluster
is embedded within a characteristic fold of HiPIPs. HiPIPs are highly
charged, either acidic or basic depending on the organism from which
they have been purified. Despite low sequence homology, the structures
of all HiPIPs share similar features, especially in loop regions.
HiPIPs were the first iron–sulfur proteins for which a crystal
structure in both the oxidized and reduced forms was obtained. The
small size of the protein requires that the [4Fe–4S] cluster
occupies a large portion of the total volume of the protein. Their
structures mainly consist of loops with two small α-helices
and five β-strands. The cluster is positioned in the C-terminal
domain of the protein (Figure 34). A conserved
Tyr in most HiPIPs is located in a small helix in N-terminal packs
against the cluster and interacts with one of the inorganic sulfurs,
S3. Two of the Cys ligands are in two β-strands in a twisted
β-sheet, and two hairpins provide the other two. Three of the
four cysteines form H-bonds with the backbone amides of residues i + 2. Aromatic side
chains from a C-terminal loop together
with the conserved Tyr from the N-terminal form a hydrophobic pocket
that further shields the cluster from solvent. HiPIPs share the consensus
motif of Cys-(Xxx)2-Cys-(Xxx)8–16-Cys-(Xxx)10–13-Gly-Trp/Tyr-Cys to coordinate the
[4Fe–4S]
cluster. Several loops around the protein make a hydrophobic pocket
for the protein to accommodate the cluster. In some cases conserved
water ligands have been shown to be important for stabilizing the
structure.
870
Figure 34
Structure of reduced
(PDB ID 1HRR) and oxidized (PDB ID 1NER) HiPIP from Ch. vinosum (top left and top
right, respectively).
The overlay of the structures and zoom-in of the Fe–S cluster
are shown at the bottom. As shown, only slight structural changes
occurred upon reduction.
The [4Fe–4S] cluster, as with ferredoxins, has a cubane
structure in which each iron is coordinated with three inorganic sulfurs
and one thiolate from cysteine. All the irons have tetrahedral geometry.
Fe–Fe distances are significantly shorter than S–S distances
(2.72 vs 3.58 Å), resulting in lower accessibility to the iron
atoms. The spin coupling between pairs of irons leads to Jahn–Teller
distortion and a D2d state rather
than a Td point group symmetry. There is also a conserved Gly close
to the conserved Tyr in most HiPIPs, which is believed to mainly play
a role in steric control.
9008
Mutational analysis of conserved
aromatic residues in HiPIPs confirmed
a protective role for these residues against hydrolysis of the cluster
by decreasing solvent accessibility.
715
Removal of this protection resulted in degradation of the cluster
through a [3Fe–4S] intermediate as evidenced by heteronuclear
multiple quantum coherence (HMQC) NMR.
875
Some HiPIPs form higher quaternary structures; HiPIP from Tb. ferrooxidans, for example,
was isolated in a
tetrameric state.
874
There are several
aromatic residues in close proximity to the Fe–S cluster in
HiPIPs. These residues have been hypothesized to play a role in ET,
reduction potential determination, and cluster stability. Several
mutational studies suggest that these residues play a major supportive
role against degradation.
873,875,877
3.4.5.3
Function
The HiPIPs appear to
be unique to the bacterial kingdom, and higher organisms replaced
them by other more sophisticated ET proteins. Despite thorough characterization
of these proteins, their function is not yet fully understood. HiPIPs
act as soluble periplasmic electron carriers in photosynthetic bacteria
between the photosynthetic reaction center and the cytochrome bc
1 complex. Other functions have been reported,
such as an iron oxidizing enzyme in Acidithiobacillus
ferrooxidans,
878
an electron
donor to cytochrome cd-type nitrate reductase in Paracoccus(868) species
or to terminal oxidases in Rhodothermus marinus,
879
or a role in thiosulfate oxidation.
880
The relative distribution of HiPIPs and their
redox behavior suggest an overlapping role of these proteins with
cytochrome c
2 as a final electron acceptor
in the photocycle.
872
However, other studies
have shown a role for HiPIPs distinct from that of cytochrome c.
881
HiPIPs are also found in
the membrane of some thermophilic organisms.
879
HiPIPs are mainly found in organisms with a photosynthetic reaction
center having a tetraheme cytochrome (THC) subunit. Multiple studies
have shown that HiPIPs could be the preferred electron carrier in
purple sulfur bacteria. Crystal structure analysis, molecular docking
studies, and computational modeling have suggested that the hydrophobic
patch of HiPIPs can interact with a hydrophobic patch in THC so that
it plays a role as a redox partner to this protein.
873,882,883
3.4.5.4
Important
Structural Elements
HiPIPs have three ferric ions and one
ferrous ion that occur as a
pair of two Fe3+ ions and a pair of two Fe2.5+ ions. In the reduced state, the cluster
has two ferric and two ferrous
ions, mainly existing as a set of mixed-valence Fe2.5+.
543,884
The reduction potentials of HiPIPs are very high, occupying a range
of 100–500 mV. Several methods have been applied to measure
the reduction potential of HiPIPs, including redox titration monitored
by EPR,
879,881
chemical redox titration,
876
and direct electrochemistry.
715,799
Some studies have suggested further delineation of HiPIPs into two
categories: the first with a narrow reduction potential range of around
330 mV and the second with a broader range that depends on protein
charges. However, only a few studies currently support this classification.
872,885
Two classes of factors should be considered while studying
the reduction potential of HiPIPs. The first class includes factors
that differentiate the reduction potentials of HiPIPs from those of
ferredoxins. The main explanation for the difference in reduction
potential between the HiPIPs and ferredoxins has been well established
now as the different redox states employed by the two proteins. While
the ferredoxins go through a [4Fe–4S]1+/2+ transition,
the HiPIPs have a [4Fe–4S]2+/3+ state. This oxidation
state has an intrinsically higher reduction potential.
719
It has been reported, however, that HiPIPs
can form a super-reduced state of [4Fe–4S]1+ if
unfolded in 80% Me2SO or by pulse radiolysis. The reduction
potential of this [4Fe–4S]2+/1+ state was calculated
to be 400–600 mV lower than that of the same pair in ferredoxins.
886
There are studies in support of the importance
of the overall structural and backbone conformation in determining
the overall potential range of the protein.
887
Also, these studies demonstrated the role of the protein environment
in ET not only by manipulating the driving force and reduction potential
but also through changing the activation energy via environmental
reorganization.
887
Resonance Raman, X-ray
crystal structure analysis, computational analysis, and spin echo
studies have all revealed an important role of solvent accessibility
in the higher reduction potential of HiPIPs vs ferredoxins.
872,873
Moreover, crystal structure analyses of HiPIPs have revealed conserved
NHamide···S H-bonds to the coordinating
sulfurs.
719,872,873
These H-bonds stabilize the reduced form of the protein by decreasing
the electron density on sulfurs, thereby increasing the reduction
potential. This effect was demonstrated by using chemically synthesized
peptides in which the peptide amide bond was replaced with an ester
linkage, thus removing the H-bond between the amide and Cys sulfu.
888
Ferredoxins have more of these amide H-bonds,
resulting in the alternate oxidation state of the [4Fe–4S]
cluster (Table 6).
93,617,618,719,887
When elongated or compressed,
the [4Fe–4S] cubanes have different spin topologies; however,
sulfur K-edge XAS, 2D NMR, and DFT calculations have shown that the
orientation of [Fe2S2]+ subclusters
is very similar in both ferredoxins and HiPIPs, suggesting a localized
oxidation or reduction in only one of the two subclusters
889
and making cluster spin topology an unlikely
source of redox-state differentiation.
Table 6
Reduction
Potential of Different Rieske
and Rieske-Type Proteinsa
protein
organism
E
m (mV)
ref
Rieske Proteins
bc
1 complex
pigeon
heart
285
(825)
bc
1 complex
beef heart
290
(806)
bc
1 complex
beef heart
304
(826)
bc
1 complex
beef heart
312
(798)
bc
1 complex
beef heart
306
(827)
bc
1 complex
beef heart
315
(828)
bc
1 complex
yeast
262
(779)
bc
1 complex
yeast
286
(829)
bc
1 complex
yeast
285
(781)
bc
1 complex
Paracoccus
denitrificans
298
(815)
bc
1 complex
Paracoccus
denitrificans
280
(830)
bc
1 complex
Rhodobacter
capsulatus
310
(831)
bc
1 complex
Rhodobacter
capsulatus
321
(832)
bc
1 complex
Rhodobacter
capsulatus
294
(832)
bc
1 complex
Rhodobacter
sphaeroides
285
(831)
bc
1 complex
Rhodobacter
sphaeroides
300
(796)
bc
1 complex
Rhodobacter
sphaeroides
300
(796)
bc
1 complex
Chromatium
vinosum
285
(833)
b
6
f complex
spinach
320
(834)
b
6
f complex
spinach
375
(835)
b
6
f complex
spinach
320
(835)
bc
1 complex
Nostoc
321
(836)
bc complex
Chlorobium
limicola
160
(837)
bc complex
Bacillus
alcalophilus
150
(838)
bc complex
Heliobacterium
chlorum
120
(839)
bc complex
Bacillus PS3
165
(837)
bc complex
Bacillus
firmus
105
(840)
Rieske protein
Thermus thermophilus
140
(841)
SoxFII
Sulfolobus
acidocaldarius
375
(842)
Rieske-Type Proteins
FdBED
Pseudomonas
putida
–155
(843)
FdBED
Pseudomonas
putida
–156
(844)
FdBED
Pseudomonas
putida
–155
(809)
benzene dioxygenase
Pseudomonas
putida
–112
(843)
2-halobenzoate 1,2-dioxygenase
Burkholderia
cepacia
–125
(845)
2-oxo-1,2-dihydoquinoline
8-monooxygenase
Burkholderia
cepacia
–100
(846)
a
Reprinted with permission from ref (773). Copyright 1999 Elsevier.
Specific interactions between hydrophobic
residues are also considered
a source of variation in reduction potential between HiPIPs and ferredoxins.
While in HiPIPs aromatic···S interactions are through
face of the aromatic ring, leading to interactions between the highest
occupied orbital of the cluster and the lowest unoccupied Tyr orbital,
ferredoxins have an interaction via the edge of Tyr with the highest
occupied Tyr orbital interacting with the lowest unoccupied cluster
orbital.
872
Some studies have suggested
that the main role of the conserved Tyr is to stabilize the cluster
through these aromatic and H-bond interactions and not to have any
profound effect on the reduction potential;
877
however, because the Tyr in different proteins tends to take a different
alignment, this hypothesis cannot be generalized to all HiPIPs.
540
The second class of factors of important
influence on the reduction
potential of HiPIPs includes interactions that fine-tune the reduction
potential. This class has not yet been fully elucidated; however,
solvation and net charges on the protein are postulated to play a
role in this class of proteins.
220,885,890,891
No correlation was
found between the orientation of aromatic residues in the protein
and its reduction potential.
892
Different
factors including the net surface charge of the protein, partial charges
of certain residues, atomic polarizability of protein atoms, and solvent
dipoles have been thoroughly studied in a number of HiPIPs, and the
only factor determined to correlate with the reduction potential was
the net charge on the protein surface (Table 7).
873,890
Table 7
Effect of the Net
Charge on the Reduction
Potential of Some HiPIPsa
protein source
E
m (mV)
net charge
ref
Chromatium
purpuratum
390
(902)
Chromatium
tepidum
323
–4
(903)
Thiocapsa
roseopersicina
346 or 325
–6
(904)
(905)
(906)
(907)
Chromatium
warmingii Bart
355
–4
(908)
Chromatium
uinosum
356
–5
(909)
Chromatium
gracile
350
–7
(906)
(910)
Thiocapsa
pfennigii
350
–9
(911)
Ectothiorhodospira
halophile
120 (iso I)
–12
(896)
(912)
(913)
Ectothiorhodospira
uacuolata
260 (iso I), 150 (iso II)
–5 (iso I), −8 (iso II)
(914)
(896)
Ectothiorhodospira
shaposhnikouii
270 (iso I), 155 (iso II)
–6 (iso I), −8 (iso II)
(914)
Rhodoferar
fermentans
351
(915)
(882)
(916)
Rhodopila
globiformis
450
–3
(917)
(896)
Rhodospirillum
salinarum
265 (iso I)
–5 (iso I), −1 (iso II)
(914)
(918)
Rhodopseudomonas
marina
345
5
(918)
Rhodocyclus
tenuis
300
2
(914)
(917)
(919)
Rhodocyclus
gelatinosus
332
3
(896)
(920)
(884)
Paracoccus
halodenitricans
282
–13
(921)
Thiobacillus
ferrooxidans
380
1
(878)
(878)
(917)
(874)
a
Reprinted with permission from ref (873). Copyright 1998 Elsevier.
The roles of different parameters
involved in determining the reduction
potential of HiPIPs have been explored through mutational studies.
In one such study, mutation of the Cys77 ligand of Ch. vinosum to Ser was analyzed
by NMR, which found
negligible conformational changes in this mutant.
894
The role of the conserved Phe66 in the same protein was
likewise investigated, finding that mutation to polar residues had
minimal effects (<25 mV) on the reduction potential.
799,876
Mutations in buried polar groups have indicated a role for these
groups in the reduction potential as well. Mutation of Ser79Pro in Ch. vinosum HiPIP
resulted in a 104 mV decrease in
reduction potential. It has been suggested that the different electrostatic
properties of the amide group between Ser and Phe and hence the ability
to H-bond are the main reasons for the observed effect.
9009
Mutations of conserved hydrophobic residues
around the Fe–S cluster (making the site more solvent-accessible)
resulted in minimal changes in the midpoint potential as well as entropy
and enthalpy of reduction.
875
Mutation
of a conserved Phe to Lys showed similar marginal changes in the reduction
potential. However, a 15-fold decrease in the self-exchange rate was
observed upon addition of positive charge to the protein surface.
The same protective roles have also been reported by mutation of conserved
Tyr19 from Ch. vinosum.
873
A CD analysis of different HiPIPs has
shown that the pH dependence
of the reduction potential in HiPIPs is very dependent on the proximity
of a His residue to the cluster. In HiPIPs from Thiocapsa
roseopersicina, which has His49, strong pH dependence
was observed, while in HiPIPs from Ch. vinosum and Rhodopseudomonas gelatinosa,
which have His42, show smaller pH dependence. In cases with no His,
the reduction potential was independent of the pH.
896
Recently, computational studies have been used to locate
residues that cause the pH dependence of a Ch. vinosum HiPIP and identified His42
as a candidate, which is consistent with
previous observations.
897
Studies
have shown a more prominent role of enthalpy in determining
the reduction potential of HiPIPs, noting a favorable change in bonding
upon reduction. These proteins also show a negative entropy change.
Increased loss of both entropy and enthalpy results from increasing
temperature, mainly due to elongation and breakage of H-bonds in the
oxidized state.
873
The covalency of the
Fe–S bond and geometry of the ligands in the structure have
been shown to play a role in different redox states and the reduction
potential between HiPIPs and ferredoxins (Table 8).
898
DFT and potential energy surface
(PES) studies have further shown that this difference in covalency
is mainly due to different arrangements of the ligands of the cluster.
899
Ligand K-edge XAS studies have also shown large
differences in Fe–S covalency between HiPIPs and ferredoxins.
The primary transition of the K-edge is 1s → 4p; however, the
covalent mixing from ligand 3p orbitals into unoccupied metal 3d orbitals
results in an additional observable 1s → 3p transition. XAS
studies demonstrated that the redox-active molecular orbital (RAMO)
in HiPIPs is the HOMO of the [4Fe–4S]2+ resting
state and has 50% sulfur ligand character. This results in a better
superexchange rate from cluster to surface, which is necessary for
the buried cluster in HiPIPs to transfer electrons.
900
Another XAS study found that the difference in charge donation
is due to different H-bonds to sulfur ligands between HiPIPs and ferredoxins.
A more recent XAS study suggested hydration of the clusters as the
main reason for the difference. This study showed that removal of
water from ferredoxins results in higher covalency. In a similar way,
exposure of the HiPIP cluster by unfolding decreases the covalency.
901
Table 8
Redox Potential of
Some HiPIPs and
Some Ferredoxins with the Number of Their NH···S H-Bond
Contactsa
protein
E
m (mV)
no. of H-bond
contacts
ref
Ectothiorhodospira
halophila I HiPIP
120
5
(922)
Ectothiorhodspira
vacuolata I1 HiPIP
150
5
(892)
Chromatium
vinosum HiPIP
360
5
(923)
Rhodocyclus
tenuis HiPIP
303
5
(919)
Bacillus
thermoproteolyticus Fd′
–280
8
(924)
Peptococcus
aerogenes Fdf
–430
8
(923)
Azotobacter
vinelandii Fd Ib
–650
8
(925)
a
Reprinted with permission from ref (873). Copyright 1998 Elsevier.
3.4.5.5
Spectroscopic
Features
The HiPIPs
have a brown-green color with a prominent band at 388 nm, with an R/Z ratio of ∼0.5,
which is bleached
after oxidation.
872,926
The oxidized form has a very
broad band with shoulders at 450, 735, and 350 nm. Both forms have
280 nm absorptions that are much higher than what is expected from
aromatic contents, indicating that the cluster has some absorption
in that region as well.
926
CD spectroscopy
in both visible and far/near-UV region has been used to probe the
effect of the protein environment on the properties of HiPIPs. It
has been shown that visible CD spectra of reduced HiPIPs are very
similar, implying strong homology in their cluster environment. Most
of the spectra show a positive feature at 450 nm and two distinct
negative features in the 350 and 390 nm regions, with some of them
showing a positive ellipticity at 330 nm. A group of HiPIPs show completely
different features, having two positive bands between 350 and 440
nm and a negative feature at around 460 nm. CD studies indicate that
the maximum band observable in absorption spectroscopy consists of
several transitions, mainly a S → Fe charge transition. Visible
CD of oxidized HiPIPs is usually featureless with broad maxima at
350, 400, and 450 nm. Near-UV CD spectra are very dependent on the
position of aromatic residues in the protein. Far-UV CD spectra showed
∼12–20% α-helical content in the protein structure
and slight changes upon oxidation and reduction.
926
HiPIPs were the first class of paramagnetic proteins
for which a thorough solution NMR study was able to determine the
structure in both the reduced and oxidized forms.
927
1H NMR studies confirmed the mixed-valence state
in HiPIPs
884
and provided additional structural
insights for these proteins.
928,929
NMR was also used
to find Fe–S–Cα–Cβ dihedral angles on the basis of hyperfine shifts of β protons
and α carbons.
930
Differences in
the electronic features of iron pairs in the oxidized and reduced
forms cause a significant hyperfine shift of 1H and 13C of the cysteine ligands of
the cluster. Similar shifts
of β carbons in the reduced state confirm the notion that they
all have similar electronic features. Most HiPIPs show at least two
isomeric electronic states apparent by room temperature NMR studies.
The best explanation for this phenomenon is that the mixed-valence
pair can switch from an iron(II/III) pair to an iron(III/IV) pair.
The reduction potential of irons in the cluster usually follows this
trend: Fe(III) > Fe(IV) ≈ Fe(II) > Fe(I), so only two
states
are observable in the oxidized state of HiPIPs, which explains the
presence of two electronic isomers observed in NMR and EPR.
884
NMR of the oxidized pair shows two downfield
signals arising from the mixed-valence pair and two upfield signals
(or extrapolated upfield, which is two downfield signals with anti-Curie
temperature dependence) assigned to the ferric pair with inverted
electron polarization.
895,931
1H 2D exchange
spectroscopy (EXSY) NMR studies have analyzed self-exchange rates
for HiPIP from Ch. vinosum and its
aromatic mutants. An exchange rate of 2.3 × 104 M–1 S–1 was observed for the native
protein at 298
K, with rates within 2-fold for the mutants. This study ruled out
the role of aromatic residues in ET.
876
β protons from cysteine ligands of the cluster experience
large contact shifts. Eight signals from +110 to −40 ppm can
be assigned to eight protons from four β-CH2 Cys
ligands. The assignment of protons that are involved in amide–S
H-bonds is more difficult due to their broad features that overlap
with other protons.
929,932
NMR experiments have also been
used to assess water accessibility of the cluster and its mutants
through analyzing the H2O/D2O exchange rates. 1H–13C heteronuclear correlation (HETCOR)
NMR was used to show that the oxidized cluster has an overall shorter
relaxation time than the reduced state.
933
EPR of HiPIPs shows a nearly axial signal with g values at 2.13 and 2.03 that result
from an S =
1/2 ground state in the oxidized form.
934
In contrast to ferredoxins, HiPIPs are EPR-silent in their reduced
state. Some HiPIPs show heterogeneous signals, probably due to sample
preparation or dimerization of the cluster.
799
ENDOR studies confirmed the presence of two pairs of irons in the
oxidized form of the protein.
935,936
EPR of most HiPIPs
has shown at least two populations. Four species can be observed by
EPR of HiPIPs with g
⊥ = 2.15–2.13,
2.13–2.11, 2.06–2.08, and maybe 2.09–2.11, with
the first two often being the most dominant species.
872
Assignment of these two species can be performed by correlating
the EPR data with room temperature 1H NMR.
Zero-field
Mössbauer studies of HiPIPs at temperatures above
100 K show a broad quadruple splitting, indicative of fast electronic
relaxation, with δ = 0.29–0.33 mm/s and quadruple splitting
values of 0.74–0.80 mm/s. At lower temperature (4.2 K) the
spectra show two nonequivalent iron pairs, one of which increases
quadruple splitting with increased applied field, whereas the other
decreases quadruple splitting. The subsets are assigned to a ferric
pair (δ = 0.27 mm/s, with a −0.87 mm/s splitting) and
a ferric–ferrous pair (δ = 0.37 mm/s with a splitting
value of −0.94 mm/s).
895
Mössbauer
shows nondistinguishable iron atoms in reduced HiPIPs. Mössbauer
studies of mutated Cys → Ser HiPIP have shown loss of covalent
iron features due to replacement of S with O and a different spectrum
of the Ser-bound iron in the reduced form, suggesting the importance
of Cys residues in maintaining the mixed-valence state of the cluster.
937
Mössbauer analyses of partially unfolded
HiPIPs have found a slight increase in Fe–S bond distances
without significant changes in the core cluster, indicating that the
cluster is not denatured in early steps of unfolding.
529,938
EXAFS analysis of the structure of the core cluster of HiPIPs
and
Fe–S distances has found a small temperature dependence. Analyses
of Cys → Ser mutants reveal slight changes to the core structure
and the Fe–S distances of intact cysteines, while the Fe–O
bond is shortened, suggesting that the entire cluster is shifted toward
the Ser ligand.
937
Ligand K-edge XAS studies
have also elucidated some of the differences between HiPIPs and ferredoxins.
900
3.4.6
Complex
Fe–S Centers
3.4.6.1
Hydrogenases
3.4.6.1.1
[NiFe] Hydrogenase Cluster
[NiFe]
hydrogenases catalyze interconversion of H2 and
H+ in microorganisms and ultimately provide electrons for
ATP synthesis. [NiFe] hydrogenases from different sources have a conserved
large domain of ∼60 kDa, containing the binuclear Ni–Fe
active site and a small Fe–S cluster domain for ET. [NiFe]
hydrogenase from Dv. gigas contains
two [4Fe–4S] clusters and one [3Fe–4S] cluster, supported
by EPR, Mössbauer,
939
and crystallographic
studies.
940,941
The reduction potentials are
−70 mV for the [3Fe–4S]+,0 cluster and −290
and −340 mV for the two flanking [4Fe–4S]2+,1+ clusters. The fully oxidized state
of the two clusters ([4Fe–4S]2+) gives an isomer shift of 0.35 mm/s and quadruple
splitting
of 1.10 mm/s. Upon reduction, the two clusters are separated. Cluster
I gives an isomer shift of 0.525 mm/s and quadruple splitting of 1.15
mm/s, and cluster II gives 0.47 and 1.35 mm/s, respectively. The parameters
of [3Fe–4S]1+ are δ = 0.47 mm/s and ΔE
Q = 1.67 mm/s, and those of [3Fe–4S]0 are δ = 0.39 mm/s and ΔE
Q = 0.38 mm/s. The three Fe–S clusters are arranged
linearly in the 3-D structure, with one [4Fe–4S] cluster proximal
to the Ni–Fe–S catalytic center, the other [4Fe–4S]
cluster at the surface, and the [3Fe–4S] cluster in the middle
of them (Figure 35),
940,941
suggesting the existence of an ET pathway.
Figure 35
Proposed ET pathway
in Dv. gigas [NiFe] hydrogenase. Selected
distances are given in angstroms. PDB
ID 1FRV. Color
code: Fe, green; Ni, gray blue; C, cyan; S, yellow, O, red; N, blue.
Reprinted with permission from ref (940). Copyright 1995 Macmillan Publishers Ltd.
[NiSeFe] hydrogenase, a subclass
of [NiFe] hydrogenases, contains
three [4Fe–4S] clusters.
942,943
The crystal
structure reveals that a cysteine residue near the middle cluster,
as opposed to proline usually observed in [NiFe] hydrogenases, serves
as an extra ligand and results in a [4Fe–4S] cluster instead
of a [3Fe–4S] cluster .
[NiFe] hydrogenase from Dv. fructosovorans is structurally similar to that
from Dv. gigas.
944
On the basis of observations made
with respect to [NiSeFe] hydrogenases, a Pro238Cys mutation has been
made. The [3Fe–4S]1+,0 cluster was successfully
converted to a [4Fe–4S]2+,1+ cluster and resulted
in a 300 mV decrease of the reduction potential with little influence
on activity, indicating that the [3Fe–4S]1+,0 cluster
is not essential in the ET pathway of [NiFe] hydrogenase.
Recently,
a new type of [NiFe] hydrogenase was discovered. Unlike
the usually air-sensitive members of the family, [NiFe] hydrogenases
from the bacteria Ralstonia eutropha, Ralstonia metallidurans, Hydrogenovibrio marinus,
and Aquifex
aeolicus could tolerate O2 to a limited
extent.
947
The oxygen tolerance arises
from neither modification of the [Ni–Fe] active site nor limited
access to O2. Crystal structures of the proteins have revealed
a novel Fe–S cluster proximal to the Ni–Fe center (Figure 36a).
948,949
Instead of the normal
proximal [4Fe–4S] cluster coordinated by four cysteines from
the protein, this cluster is a plastic [4Fe–3S] cluster bound
by six cysteines with a flexible glutamic acid residue nearby. Upon
oxidation, the backbone amide of the coordinating Cys26 is deprotonated
by the nearby glutamic carboxylate and replaces the bridging Cys25
(Figure 36b,c), analogous to the P cluster
in nitrogenases. The negative charge of amide will help to stabilize
the oxidized state. As a result, the [4Fe–3S] cluster could
transfer two electrons in a window of 200 mV and remain stable in
three oxidation states.
950
DFT calculations
have revealed that the supernumerary coordination frame provided by
the six cysteines and the flexible coordination sphere of the Cys26-bound
Fe lead to plasticity of the unique proximal [4Fe–3S] cluster
and, consequently, low reorganization energy in the reduced state.
945
Hence, the proximal cluster could not only
transfer electrons efficiently from the active site during H2 oxidation, but also
rapidly supply two electrons to the active sites
upon O2 binding, which in combination with one electron
from the middle [3Fe–4S] cluster would efficiently reduce O2 to H2O and prevent formation
of an inactive [Ni3+– –OOH–Fe2+] cluster,
the so-called Ni-A state, and overoxidation by O2.
951−953
Figure 36
(a) Crystal structure of O2-tolerant membrane-bound
hydrogenase from Ralstonia eutropha (PDB ID 3RGW). Reprinted from ref (945). Copyright
2013 American Chemical Society. (b) Reduced
[4Fe–3S] cluster from MBH (PDB ID 3AYX) (Reprinted with permission from ref (946).
Copyright 2012 Wiley-VCH)
and (c) oxidized [4Fe–3S] cluster from MBH (PDB ID 3AYZ). Reprinted with
permission from ref (946). Copyright 2012 Wiley-VCH. Color code: Fe, green; C, cyan;
S, yellow;
N, blue; Ni, orange.
3.4.6.1.2
[FeFe] Hydrogenases
[FeFe]
hydrogenases share a conserved catalytic subunit binding metal cluster,
called the H-cluster, as the catalytic site and have various Fe–S
subunits harboring different Fe–S clusters for ET to and from
the H-cluster. The Fe–S domains are usually located at the
N-terminus of the catalytic domain and contain [4Fe–4S] or
[2Fe–2S] binding motifs similar to those of ferredoxins.
954−956
For example, [FeFe] hydrogenase from Dv. desulfuricans ATCC 7757 possesses two [4Fe–4S]
clusters for ET,
957
and the protein from Cl. pasteurianum contains one [2Fe–2S] cluster and three [4Fe–4S]
clusters.
958
The Fe–S clusters in Cl. pasteurianum [FeFe] hydrogenase are separated
by 8–11 Å, indicating potential ET pathways through covalent
bonds or a H-bonding network (Figure 37). The
FS4C and FS2 near the protein surface possibly function as the initial
electron acceptors of external electron donors and transfer electrons
to the FS4B at the junction position. The FS4A is 10 Å from cluster
FS4B and 9 Å from the H-cluster and could mediate sequential
ET to and from the catalytic site.
Figure 37
(a) Location of Fe–S clusters
in [FeFe] hydrogenase (PDB
ID 1FEH). (b)
Proposed ET pathways for [FeFe] hydrogenase. Reprinted with permission
from ref (958). Copyright
1998 American Association for the Advancement of Science.
3.4.6.2
Molybdonum-Containing
Enzymes
273
3.4.6.2.1
[4Fe–4S] Cluster and P-Cluster
in Nitrogenase
Four types of nitrogenases have been discovered:
two containing Mo and Fe, one containing V and Fe, and one containing
only Fe in the catalytic site in a large domain with a molar mass
of 220–250 kDa. Among them, [FeMo] nitrogenase has been the
most extensively studied (Figure 38a). Besides
the active site, all nitrogenases contain an iron protein as α2 dimers with a molar
mass of 60–70 kDa. It contains
a single [4Fe–4S] cluster between the two monomers, which is
coordinated by one conserved cysteine from each monomer and is exposed
to water.
959
The cluster transfers electrons
efficiently via a MgATP hydrolysis reaction at the larger domain containing
a catalytic site, along with other functions, including involvement
in biosynthesis and insertion of FeMoco into [FeMo] nitrogenase and
regulation of biosynthesis in other nitrogenases.
960
Figure 38
(a) Overall structure of nitrogenase (PDB ID 1N2C). Cofactors are
shown as spheres and denoted. Reprinted with permission from ref (965). Copyright
1997 Macmillan
Publishers Ltd. (b) Reduced P cluster from nitrogenase (PDB ID 3U7Q) (Reprinted with
permission from ref (946). Copyright 2012 Wiley-VCH.) and (c) oxidized P cluster from
nitrogenase
(PDB ID 2MIN). Reprinted with permission from ref (946). Copyright 2012 Wiley-VCH.
Three oxidation states, +2, +1, and 0, have been observed
for the
[4Fe–4S] cluster, indicating that the cluster could transfer
one or two electrons to the catalytic domain. The reduction potential
to achieve an all-ferrous [4Fe–4S]0 cluster is −460
mV, and this is the first example of this oxidation state for [4Fe–4S]
clusters, both in proteins and in model complexes.
961−963
EXAFS studies show that changes of the Fe–S and Fe–Fe
distances are less than 0.02 Å from the [4Fe–4S]2+ cluster to the [4Fe–4S]1+ cluster.
964
The Fe protein can bind 2 equiv of MgATP
or MgADP, each in a Walker
A motif on one monomer. The Walker A binding site is 15–20
Å away from the [4Fe–4S] cluster with a series of salt
bridges and H-bonds in between. However, the reduction potential of
the [4Fe–4S] cluster decreases ∼100 mV upon binding
of either nucleotide, possibly arising from protein conformational
changes induced by binding and hydrolysis reactions.
965−970
The reduction potential change is proposed to be the driving force
for ET.
968
UV–vis, resonance Raman,
and EPR spectroscopic studies indicate that the [4Fe–4S] cluster
could reversibly cycle between a regular [4Fe–4S] cluster in
the reduced state and two [2Fe–2S] clusters in the oxidized
state.
971
The [FeMo] domain contains
the FeMoco cluster and a P-cluster.
The FeMoco cluster is the catalytic center and will not be discussed
here. The P-cluster is situated at the interface of the α and
β subunits of the [FeMo] domain. It is an [8Fe–7S] cluster,
with a 6-coordinate sulfur at the center. The structure of the P-cluster
changes with the oxidation state. The dithionite reduced P cluster
(PN) is bound by six cysteines from the protein, four of which coordinate
a single iron, and the remaining two function as bridging ligands
(Figure 38b).
972
After two-electron oxidation of PN, a form called Pox is obtained.
In the Pox cluster, the coordination between the center 6-coordinate
sulfur and two irons associated with the β subunit is replaced
by the amide N of Cys88 of the α subunit and side chain hydroxyl
of Ser188 of the β subunit (Figure 38c), similar to the changes of oxygen-tolerant
[NiFe] hydrogenases
mentioned above (see Figure 36). The changes
are proposed to be related to the proton-coupled electron transfer
process in nitrogenases.
972−974
3.4.6.2.2
Aldehyde Oxidoreductases
Aldehyde
oxidoreductase belongs to the molybdoflavoenzymes. It is a homodimer
and usually requires Fe–S clusters, a molybdopterin or tungstopterin
site, and sometimes an FAD cofactor for substrate oxidation. Aldehyde
oxidoreductase from Dv. gigas is composed
of four domains, including two small N-terminal domains binding two
types of [2Fe–2S] clusters and two large domains containing
the molybdopterin cofactors.
975,976
The first Fe–S
domain (residue 1–76) is similar to that of spinach ferredoxins,
and the [2Fe–2S] cluster is coordinated by Cys40, Cys45, Cys47,
and Cys60. The second Fe–S domain (residues 84–156)
is a four-helix bundle, and the [2Fe–2S] cluster is coordinated
by Cys100, Cys103, Cys137, and Cys139. The molybdopterin is 15 Å
from the surface and 14.9 Å from the Fe–S cluster of the
second domain. Recently, the crystal structure of aldehyde oxidase
of mouse liver has been reported. The overall fold is very similar
to that from Dv. gigas, but that of
the mammalian protein has an additional FAD domain.
977
EPR studies revealed two types of [2Fe–2S]
clusters, named Fe–SI and Fe–SII.
978−981
Fe–SI is observable at 77 K with g values
of 2.021, 1.938, and 1.919, while Fe–SII is only observable
below 40 K with g values of 2.057, 1.970, and 1.900.
The reduction potentials of Fe–SI and Fe–SII are −260
and −280 mV, respectively.
In the presence of the substrate
benzaldehyde, partial reduction
of the Fe–S clusters has been detected in Mössbauer
studies, indicating participation of the Fe–S clusters in the
catalytic reaction and fast ET from the molybdopterin center.
982
3.4.6.3
Ni-Containing CO Dehydrogenase
and Hybrid
Cluster Protein
3.4.6.3.1
Ni-Containing CO
Dehydrogenase
CO dehydrogenases (CODHs) catalyze oxidation
of CO to CO2 along with dehydrogenation of water and release
of protons and electrons.
It is important in the oxygen-based respiratory process in hydrogenogenic
bacteria. There are two types of CODHs. One is Mo-based CODHs with
a mono-Mo cofactor coordinated by dithiolene sulfurs of a pterin ligand
found in aerobic organisms, which is beyond the scope of this review
but has been reviewed extensively in other papers.
983,984
The other is Ni-containing CODHs with a Ni–Fe–S cluster
as well as multiple Fe–S clusters found in anaerobic organisms
985−987
and will be discussed briefly below.
Ni CODHs are β2 homodimers.
988,989
Each monomer contains a Ni–Fe–S
cluster (cluster C) as the catalytic site and a [4Fe–4S] cluster
(cluster B). In addition, another [4Fe–4S] cluster (cluster
D) is situated at the interface of the two monomers and coordinated
by residues from both monomers (Figure 39a).
Clusters B and D transfer electrons between cluster C and external
redox regents. They also bind acetyl-CoA synthases to form α2β2 bifunctional enzymes
acetyl-CoA synthases/carbon
monoxide dehydrogenases (ACSs/CODHs).
990
Two additional [4Fe–4S] clusters, E and F, have been found
in an extra subunit of the ACS/CODH complex.
991
The crystal structure of Ni CODH from Carboxydothermus
hydrogenoformans reveals that cluster C is a [Ni–4Fe–5S]
cluster (Figure 39b). The geometries of the
irons are approximately tetrahedral, and that of Ni is close to square
planar. It is associated with the protein through four cysteines and
one histidine.
988
On the other hand, the
structures of Rhodospirillum rubrum Ni CODHs
989
and the M.
thermoacetica ACS/CODH complex
991
show cluster C as [Ni–4Fe–4S], coordinated
similarly by five cysteines and one histidine from the protein (Figure 39c). The Ni
is also coordinated by an external nonprotein
ligand.
Figure 39
(a) Crystal structure of Rs. rubrum Ni CODH. Clusters are shown as spheres. PDB ID
1JQK. (b) [4Fe–5S–Ni]
cluster C of Ca. hydrogenoformans Ni
CODH. PDB ID 1SU8. (c) [4Fe–4S–Ni] cluster C of M. thermoacetica Ni CODH. PDB ID 1MJG.
Reprinted with permission from ref (990). Copyright 2011 Elsevier.
3.4.6.3.2
Hybrid Cluster Proteins
Hybrid
cluster proteins (HCPs) are a type of Fe–S proteins with unknown
functions. However, they have been detected in more than 15 bacteria
and archaea. There are three categories of HCPs. The first is found
in anaerobic bacteria such as Dv. vulgaris and Dv. desulfuricans or methanogen
archeon Methanococcus jannaschii, with
coordinating cysteines arranged in the sequence Cys-(Xxx)2-Cys-(Xxx)7–8-Cys-(Xxx)5-Cys.
The second
is found in facultative anaerobic Gram-negative bacteria such as E. coli, Morganella
morganii, or Tb. ferrooxidans, with the sequence
Cys-(Xxx)2-Cys-(Xxx)11-Cys-(Xxx)6-Cys. The third is found in (hyper)thermophilic bacteria
or archaea,
including Methanobacterium thermoautotrophicum, Pyrococcus abyssi, or Tt. maritima,
with the same sequence arrangement
as the first category but with smaller size due to residue deletion
downstream of the N-terminal cysteine region.
HCP from Dv. vulgaris contains three domains (Figure 40a).
992,993
A [4Fe–4S]
cluster is bound to domain 1 by Cys3, Cys6, Cys15, and Cys21 from
the N-terminal region, similar to the cubane cluster in ferredoxins
except that no cysteine is from the C-terminal region. This Cys-(Xxx)2-Cys-(Xxx)8-Cys-(Xxx)5-Cys
motif is
conserved in all HCPs, and HCPs from both categories 1 and 3 contain
a [4Fe–4S] cluster linked by this motif. HCPs from category
2, on the other hand, might instead have two [2Fe–2S] clusters
at this position.
994
Figure 40
Hybrid clusters in HCP.
(a) Overall structure of as-isolated Dv. vulgaris HCP. Metal clusters are shown as
spheres.
PDB ID 1W9M.
(b) Superposition of Dv. vulgaris HCP
(cyan) and NiCODH (red, PDB code 1SU7). (c) Hybrid cluster in the as-isolated
oxidized form of Dv. vulgaris HCP prepared
anaerobically. PDB ID 1W9M. (d) Hybrid cluster in the reduced form of Dv. vulgaris
HCP. PDB ID 1OA1. Residue backbones are omitted for clarity.
Bonds inside the cluster are shown as dotted lines, and bonds between
residues and the cluster are shown as solid lines. Color code: Fe,
green; C, cyan; S, yellow; O, red; N, blue. Reprinted with permission
from ref (995). Copyright
2008 International Union of Crystallography.
HCPs also contain a unique hybrid cluster, [4Fe–2S–3O],
which was isolated in the oxidized form from Dv. vulgaris HCP (Figure 40c),
995
and [4Fe–3S] with a water molecule between Glu494 and His244
in the reduced form (Figure 40d).
996
In the former state, the cluster is linked
to the protein by Cys12, Cys434, Cys459, thio-Cys406 (Cys with an
additional S on the S(Cys), called Css406), His244, Glu268, and Glu494,
and in the latter case Css406 is reduced to cysteine. The EPR signal
of HCP is similar to that of the prismane model complex (Et4N)3[Fe6S6(SC6H4-p-Me)6]3+.
997
Therefore, the four oxidation states of the hybrid cluster
are named analogously to those of the prismane complex as “3+”,
“4+”, “5+”, and “6+”. The
midpoint reduction potentials of the Dv. vulgaris HCP hybrid cluster range from −200
to +300 mV at pH 7.5.
998
It is noteworthy that HCPs demonstrate
a high degree of similarity
to Ni CODHs.
992,993,999
They not only share similar overall folding, but also exhibit similar
cluster positions and structures inside the monomer (Figure 40b). The closest distance
between the [4Fe–4S]
cluster and hybrid cluster is 10.9 Å, with Tyr493, Thr71, Asn72,
and Glu494 in between. In addition, two tryptophan residues, Trp292
and Trp293, are located between the hybrid cluster and the protein
surface. The arrangements indicate possible ET pathways, yet no involvement
in such processes has been detected so far. The protein can be reduced
by NAD(P)H oxidoreductase,
994
but there
is no genomic evidence for the existence of a similar redox partner
in the sources from which HCP has been detected or isolated.
3.4.6.4
Siroheme Fe–S Proteins
Siroheme is an iron-containing
reduced tetrahydroporphyrin of the
isobacteriochlorin class (Figure 41a). Siroheme
proteins are a type of iron–sulfur protein containing a siroheme
conjugated to a [4Fe–4S] cluster through a thiolate bridge.
1000
Siroheme is the catalytic center, and the
[4Fe–4S] cluster serves as an electron trapping and storage
site. Siroheme proteins includes sulfite reductases and nitrite reductases,
and they are important in assimilation and dissimilation of sulfite
and nitrite.
1001,1002
Figure 41
(a) Structure of siroheme.
(b) Siroheme and the [4Fe–4S]
cluster of spinach nitrite reductase. PDB ID 2AKJ. Color code: Fe,
green; C, cyan; S, yellow; O, red; N, blue.
3.4.6.4.1
Nitrite Reductase
NiR catalyzes
the six-electron reduction of nitrite to ammonia. It exists in both
eukaryotes and prokaryotes. There are two types of NiR categorized
by the physiological electron donor: ferredoxin-dependent NiR in photosynthetic
organisms and NAD(P)H-dependent NiR in most heterotrophic organisms.
276,1003−1005
Ferredoxin-dependent NiR contains a siroheme
and a [4Fe–4S] cluster, while NAD(P)H-dependent NiR contains
an additional FAD cofactor bound at an extended N-terminal region.
276
Spinach nitrite reductase is a type of
ferredoxin-dependent NiR isolated from higher plants. It is composed
of 594 amino acids divided into three α/β domains. The
siroheme cofactor is situated at the interface of the three domains
and bridged to the [4Fe–4S] cluster via Cys486 (Figure 41b). The [4Fe–4S] cluster
is also coordinated
by Cys441, Cys447, and Cys482. The midpoint reduction potentials are
−290 mV for the siroheme and −365 mV for the [4Fe–4S]
cluster. Although the two cofactors are magnetically coupled with
a distance of 4.2 Å, they are independent in redox titration
processes.
1006,1007
Spinach NiR can form a 1:1
complex with ferredoxin with electrostatic interactions between acidic
residues from NiR and basic residues from ferredoxin. The interprotein
ET chain has been established as from photoexcited photosystem I via
the [2Fe–2S] cluster of ferredoxin to the [4Fe–4S] cluster
of NiR followed by intraprotein transfer to the siroheme.
1006−1008
3.4.6.4.2
Sulfite Reductase
Sulfite reductase
catalyzes the six-electron reduction of sulfite to sulfide in biological
systems and can be categorized as assimilatory sulfite reductase (aSiR)
or dissimilatory sulfite reductase (dSiR). aSiR reduces sulfite directly
to sulfide, while dSiR provides a mixture of sulfide, trithionate,
and thiosulfate in in vitro experiments.
1009
The aSiRs are found in archaebacteria, bacteria, fungi, and
plants.
1010,1011
Assimilatory ferredoxin-dependent
sulfite reductases from plant chloroplasts and cyanobacteria are soluble
monomeric proteins with molar masses of ∼65 kDa. They contain
a siroheme linked to a [4Fe–4S] cluster structurally similar
to those in nitrite reductase, and they undergo reduction by ferredoxin
from photoreduced photosystem I as well.
1002
They can also catalyze the reduction of nitrite to ammonia, the
reaction catalyzed by NiR, but with a higher K
M for nitrite than sulfite, further demonstrating the significant
similarity of the two types of enzymes.
1002,1012,1013
For maize sulfite reductase,
the midpoint potentials of siroheme and the [4Fe–4S] cluster
have been determined to be −285 ± 5 and −400 ±
5 mV, respectively, at pH 7.5 in Tris buffer by redox titrations.
Although the E° of the [4Fe–4S] cluster
is more negative than that of spinach nitrite reductase (E° = −375 ± 10 mV at pH 7.5
in Tris buffer), reduction
by ferredoxin (E° = −430 mV) is still
a thermodynamically favorable process. In the presence of cyanide,
the E° of siroheme shifts positively to −155
± 5 mV, while that of the [4Fe–4S] cluster shifts negatively
to −455 ± 10 mV, possibly due to decreased affinity of
the enzyme for cyanide upon reduction of the [4Fe–4S] cluster.
Similar trends are observed in spinach nitrite reductase as well.
1014
The aSiR from E. coli is a 780 kDa hemeoflavoprotein with an α8β4 arrangement. The
α subunit, known as sulfite reductase
flavoprotein, contains FAD and FMN, while the β unit, named
sulfite reductase hemoprotein, harbors the associated [4Fe–4S]
cluster and siroheme. The ET pathway is in the FAD–FMN–[4Fe–4S]–siroheme
sequence, with NADPH as the initial donor and sulfite as the terminal
acceptor.
1015
dSiRs exist in sulfate
reducing microorganisms.
1010,1011
dSiR is composed
of two types of subunits, DsrA and DsrB, generally
in a heterotetrametric α2β2 arrangement
with similar overall folds for all dSiRs from different sources.
1016,1017
Some dSiRs form a complex with two additional subunits of DsrC and
result in an α2β2γ2 arrangement. The dSiR contains eight [4Fe–4S] clusters together
with four sirohemes or two sirohemes and two sirohydrochlorins (the
metal-free form of siroheme) (Figure 42a,b),
and only two of the four sites are catalytically active. In Dv. gigas, desulfoviridin,
a subcategory of dSiR,
a [3Fe–4S] cluster is associated with the siroheme instead
of a [4Fe–4S] cluster in one active form, DsrII (Figure 42c). The relative position
of siroheme and the [4Fe–4S]
cluster is similar to that in aSiRs, and both the [4Fe–4S]
clusters proximal to and remote from the siroheme are coordinated
by four cysteines from the protein.
1018−1020
Figure 42
(a) Siroheme group and
[4Fe–4S] cluster of DsrI. PDB ID 3OR1. (b) Sirohydrochlorin
group and [4Fe–4S] cluster of DsrII. PDB ID 3OR2. (c) Siroheme group
and [3Fe–4S] cluster of DsrII. PDB ID 3OR2. Color code: Fe,
green; C, cyan; S, yellow; O, red; N, blue.
3.4.6.5
Respiratory Complex Chain
The
mitochondrial respiratory system is the main energy producer in eukaryotic
cells.
1021,1022
It consists of five membrane
complexes, complex I,
1023
complex II (succinate
dehydrogenase),
1024,1025
complex III (cytochrome bc
1 complex),
1026−1029
complex IV (cytochrome c oxidase complex),
1030,1031
and complex V (ATPase).
1032
The first four complexes are located on the
inner membrane and function by transferring electrons from electron
donors, NADH and succinate, to the final electron acceptor, oxygen,
and meanwhile pump protons across the membrane. This proton gradient
is utilized by ATPase to generate ATP.
3.4.6.5.1
Respiratory Complex I
Respiratory
complex I (CI), also known as NADH:ubiquinone oxidoreductase or NADH
dehydrogenase, is involved in one of the ET pathways of the respiratory
chain. It is composed of the following steps: (1) NADH donates electrons
through CI to reduce ubiquinone to ubiquinol. (2) Ubiquinol transfers
electrons through complex III to cytochrome c. (3)
Cytochrome c is oxidized by complex IV and transfers
electrons to O2 to produce water. In this process, each
electron transferred is associated with five protons pumped from the
matrix to the inner membrane space.
Although CI is the most
complicated complex in the mitochondrial respiratory chain, important
breakthroughs have been achieved, and multiple structures have been
reported recently.
1023,1033−1036
Mammalian CI (∼980 kDa) is composed of up to 45 different
subunits, including 7 subunits in hydrophilic parts harboring one
FMN and eight Fe–S clusters, 7 subunits in transmembrane parts,
and ∼30 accessory subunits.
1022,1037
Bacterial
NADH dehydrogenase (∼550 kDa) only contains 13–16 subunits,
which is sufficient for complete CI function as well.
1023,1038−1040
The crystal structure of the hydrophilic
part of complex I from T. thermophilus(1023) reveals for the first time the main
ET pathway of the protein as shown in Figure 43: electrons from NADH are transferred
through FMN to N3, followed
by N1b, N4, N5, N6a, and N6b sequentially, and finally through N2
to ubiquinone coupled with proton translocation.
1022
Figure 43
Crystal structure of mitochondrial respiratory complex
I from T. thermophilus. PDB ID 4HEA. Cofactors involved
in the ET pathway
are shown on the right side with distances and directions denoted.
Reprinted with permission from ref (1022). Copyright 2013 Elsevier.
3.4.6.5.2
Respiratory Complex II (Succinate Dehydrogenase)
and Fumarate Reducatse
Complex II in the respiratory chain
(CII), also known as succinate dehydrogenase (SDH) or succinate:quinone
reductase, is a membrane-bound protein involved in the citric acid
cycle and the second ET pathway in the mitochondrial respiratory chain.
In the mitochondrial respiratory chain, electrons are transferred
from succinate to ubiquinone through complex II, then to cytochrome c through complex
III, and finally to O2 through
complex IV. This process is less efficient than the process associated
with complex I, and each electron transferred will pump only three
protons across the membrane.
CII catalyzes oxidation of succinate
to fumarate by a hydrophilic catalytic domain composed of a large
flavoprotein (Fp; 65–79 kDa) with a covalently bound FAD cofactor
and an iron–sulfur protein (Ip; 25–37 kDa) containing
[2Fe–2S] (center S1), [4Fe–4S] (center S2), and [3Fe–4S]
(center S3) clusters.
1024,1025,1041
The catalytic domain is anchored to the membrane by one or two hydrophobic
domains (CybL, CybS) harboring usually b-type cytochromes
(Figure 44). The [2Fe–2S] center is
coordinated by four cysteines close to the N-terminus, and the [4Fe–4S]
and [3Fe–4S] clusters are coordinated near the C-terminus by
two cysteine-containing sequences: Cys-(Xxx)2-Cys-(Xxx)2-Cys-(Xxx)3-Pro and Cys-(Xxx)2-Xxx-(Xxx)2-Cys-(Xxx)3–Cys-Pro
(Xxx = Ile, Val, Leu, or Ala), similar to 7Fe
ferredoxins. The [4Fe–4S] cluster usually has a low reduction
potential and functions as the energy barrier of the ET process to
direct the electron flow and, consequently, the reaction pathway.
1042
The [3Fe–4S] cluster is involved in
a direct ET process from the initial electron donor quinones.
1043−1045
The midpoint reduction potential of the [3Fe–4S]1+,0 cluster is in the range of +60
to +90 mV, and the potential of the
initial electron donor ubiquinone is +65 mV.
1046
SDH from Sl. acidocaldarius contains a [4Fe–4S] center instead of a [3Fe–4S] center
for cluster S2 and displays poor reactivity toward caldariella quinone.
1047
Figure 44
Crystal structure of mitochondrial respiratory
complex II. FAD
binding protein (Fp) is shown in blue, iron–sulfur protein
(Ip) is shown in cream, hydrophobic domains are shown in pink and
orange, and the putative membrane is shown in gray shading. PDB ID 1ZOY. Cofactors
involved
in the ET pathway are shown on the right side, with distances, reduction
potential, and directions denoted. Reprinted with permission from
ref (1024). Copyright
2005 Elsevier.
It is noteworthy that
heme b (E° = +35 mV) in the
hydrophobic domain of SDH is not involved
in the ET pathway mentioned above. It is proposed that heme b in SDH of E. coli functions
as an electron sink and reduces ROS to protect FAD and Fe–S
clusters.
1025
However, the reduction potential
of heme b in SDH of porcine is −185 mV,
1048
much lower than that of E.
coli. Therefore, the electron sink mechanism is less
effective in this case and needs further investigation.
Fumarate
reductase is a member of the succinate–ubiquinone
oxidoreductase superfamily as well. It catalyzes the reduction of
fumarate to succinate, the reverse reaction of SDH. It is very similar
to SDH in subunit composition and cofactors.
1049,1050
Its three iron–sulfur clusters are linked to the protein
by cysteine residues in E. coli, which
are conserved in other fumarate reductases too. The midpoint reduction
potential is between −70 and −20 mV, and that of the
initial electron donor menaquinol is −74 mV.
1046
3.5
Engineered Fe–S
Proteins
3.5.1
Artificial Rubredoxins
A rubredoxin-like
[FeCys4] center has been constructed into thioredoxin by
computational design. The first coordination sphere is composed of
two cysteines, Cys32 and Cys35, which form a disulfide bond in wild-type
thioredoxin, as well as two cysteines introduced by mutation, Trp28Cys
and Ile75Cys. The resulting monoiron center resembles Rd in UV–vis
and EPR spectra, and the mimic protein is able to undergo three cycles
of air oxidation and β-mercaptoethanol reduction.
1051
The redox process of rubredoxin is not
fully reversible due to the instability of the reduced form. Nanda
et al. have constructed a minimal rubredoxin mimic, RM1, on the basis
of computational design for a more restrained tertiary structure derived
from PfRd. RM1 is a domain-swapped dimer fused with
a highly stable hairpin motif tryptophan zipper and displays spectroscopic
properties very similar to those of native Rd’s. Moreover,
it shows a reduction potential of 55 mV vs SHE and maintains redox
activity for up to 16 cycles under aerobic conditions.
1051
3.5.2
Artificial [4Fe–4S]
Clusters
There have been numerous studies focusing on making
model compounds
of ferredoxins
1052−1054
and using those models to elucidate features
of natural Fe–S clusters using several methods.
803,1055,1056,1057
In addition to synthetic models of ferredoxins that are discussed
in a review in this journal,
2007
protein
and peptide models of ferredoxins have also been made. These models
have been discussed in detail in another review in this thematic issue,
3000
and we will discuss them here only briefly.
Almost all of these mimics are modeled after [4Fe–4S] clusters,
usually made by placing the conserved motif within a scaffold. These
model systems have been used for unraveling the minimal structures
required for binding of Fe–S clusters.
732,1058,1059,1061
A 16 amino acid peptide has been modeled to incorporate a
low-potential
[4Fe–4S] cluster. More detailed sequence alignments resulted
in design of peptides with better cluster binding features that mimic
FA and FB of photosystem I.
705
Other peptide models have also been made to analyze reduction
potential properties of different Fe–S clusters, including
[4Fe–4S] clusters, [2Fe–2S] clusters, and rubredoxins.
717
Four-helix bundle models of [4Fe–4S]
clusters are among
the most common systems to build and study these clusters. Both a
single [4Fe–4S] cluster and a [4Fe–4S] cluster together
with a heme cofactor have been designed in such four-helix bundles.
1061,1062
Recently, a “metal first” approach has been taken
to introduce a [4Fe–4S] cluster into a non-natural α-helical
coiled coil structure. The design then went through further optimization
and addition of secondary sphere interactions to stabilize the reduced
form and prevent aggregation. Such designs that are independent of
structural motifs can be used as a platform for the future design
of multiclusters to be used as biological “wires” that
transfer electrons through a chain of proteins.
1063
3.6
Cluster Interconversion
Although
the Fe–S clusters are mostly classified on the basis of the
number of iron atoms in the center, there are several cases in which
changing one cluster to another type has been observed. These cluster
interconversions can happen through three types of processes: natural
changes in the environment of the cluster, chemical treatments of
the cluster, or amino acid replacements.
One of the most common
types of cluster interconversion is the change from a [4Fe–4S]
cluster to a [2Fe–2S] cluster. This kind of conversion has
been observed in hydrogenases and nitrogenases. While CD and MCD analyses
show that MgATP/ADP binding to the [4Fe–4S] cluster of Fe hydrogenase
does not result in conversion to a [2Fe–2S] cluster,
1064
addition of α,α′-dipyridyl
to the [4Fe–4S] cluster of nitrogenase resulted in formation
of a [2Fe–2S] cluster in the presence of MgATP.
1065,1066
The [4Fe–4S] to [2Fe–2S] cluster conversion has been
observed in enzymes such as ribonucleotide reductase
1067
and pyruvate formate activating enzyme
1068
as well, usually upon oxidation in air or
chemical treatment.
A very well studied case of the role of
[4Fe–4S] to [2Fe–2S]
cluster conversion in regulating cellular responses is that of fumarate
nitrate reduction transcription factor. It has been shown that this
protein undergoes the conversion upon O2 stress. The excess
oxygen will oxidize S ligands and generate disulfide cysteines. The
formation of a disulfide Cys-ligated [2Fe–2S] cluster will
result in a monomerization of the fumarate nitrite reduction transcription
factor dimer, hence unbinding from DNA.
1069,1070
The conversion is composed of two steps: first, the [4Fe–4S]
cluster undergoes a one-electron oxidation to form a [3Fe–4S]1+ intermediate after
releasing an Fe2+. Second,
the [3Fe–4S]1+ cluster converts to a [2Fe–2S]
cluster and releases an Fe3+ and two sulfide ions.
1071,1072
Mutating Ser24 into Phe and shielding Cys23 could inhibit step 1.
1073
Chelators of both Fe2+ and Fe3+ could accelerate step 2 significantly.
1074
Another very common interconversion is [4Fe–4S]
to [3Fe–4S]
interconversion. The [4Fe–4S] clusters are very sensitive to
air, and oxidation in air can remove one of the irons, resulting in
a 3Fe cluster.
1075
The most well studied
case of this interconversion is the enzyme aconitase. Aconitase has
a [4Fe–4S] cluster in its active form, which is very sensitive
to air. Aerobic purification of the protein causes formation of an
inactive enzyme with a 3Fe cluster. Addition of extra Fe, however,
can reverse the conversion and reactivate the enzyme.
1076
Exposure of the [3Fe–4S] aconitase
to high pH (>9.0) will result in the formation of a purple species
that has been attributed to a linear [3Fe–4S] cluster. This
purple protein can be activated again through reduction in the presence
of Fe.
1077
While more often clusters
of higher iron number convert into clusters
with fewer iron atoms, the reverse case has also been observed. In
biotin synthase, there are two [2Fe–2S] clusters that can convert
to a [4Fe–4S] cluster after reduction. UV–vis and EPR
studies reveal that the conversion process occurs through dissociation
of Fe from the protein followed by slow reassociation.
1078
Ferredoxin II of Dv. gigas has a [3Fe–3S] cluster that can convert into a [4Fe–4S]
cluster through incubation with excess Fe, presumably through a non-Cys
ligand.
1079
The [3Fe–4S]1+ and [2Fe–2S]2+ clusters in isolated pyruvate formate–lyase
can both be converted to [4Fe–4S] clusters with mixed valences
of +1 and +2 upon dithionite reduction.
1080
Interconversion between [4Fe–4S] and [3Fe–4S]
clusters
has been investigated through mutational studies. Removal of Cys ligands
in [4Fe–4S] clusters results in the formation of [3Fe–4S]
clusters. Replacement of the conserved Asp in [3Fe–4S] clusters
with a ligating residue such as His or Cys causes formation of [4Fe–4S]
clusters.
735,944,1081,1082
In [NiFe] hydrogenase, mutating
a conserved Pro residue into Cys near the [3Fe–4S] cluster
has successfully converted it to a [4Fe–4S] cluster accompanied
by a 300 mV decrease in the reduction potential,
944
while in F420 reducing hydrogenase of Methanococcus voltae the [4Fe–4S] to [3Fe–4S]
conversion has been achieved by replacing a Cys residue, producing
a ∼400 mV increase in the reduction potential.
1081
Addition of other metal ions in place
of the fourth iron into a
[3Fe–4S] cluster is sometimes also called interconversion.
There are multiple reports of the formation of such hybrid clusters
with Zn, Tl, and other metal ions.
1083,1084
3.7
Structural Features Controlling the Redox
Chemistry of Fe–S Proteins
The Fe–S proteins
cover a wide range of reduction potentials, mostly in the lower or
negative end of the range. Several parameters are known to be important
in the ability of Fe–S proteins to accommodate such a wide
range of reduction potentials. Unique electronic structures of iron
in different clusters and different protein environments are among
the most important factors. The ability of each iron to go through
2+ to 3+ oxidation states will allow multiple states for the core
cluster, each of which having a different reduction potential range.
This factor is more evident in the case of HiPIPs vs ferredoxins.
Solvent accessibility, H-bonding patterns around the cluster, the
net charge of the protein, partial charges around the cluster, and
the identity of the ligands are among the other features that contribute
to fine-tuning the reduction potential. Detailed examples of the role
of each feature are discussed in section 3.4.3.3.3, “Important Structural Elements”.
Below is
a summary of these features and their effects in different Fe–S
proteins.
3.7.1
Roles of the Geometry and Redox State of
the Cluster
As with other redox-active metal centers, the
primary coordination sphere of a metal ion plays an important role
in its redox properties. The iron center(s) has the same distorted
tetrahedral structure in almost all Fe–S proteins; however,
it has been shown that slight changes in this structure will result
in changes in the reduction potentials. Differences in the Fe–S–Cα–Cβ torsion angle
618,731,1085
and distortion of the cuboidal
structure in some [3Fe–4S] clusters
1086
are examples of this distortion. Different geometries can lead to
slight differences in electronic structures that will affect the redox
properties of the protein.
Another important feature that influences
the reduction potential is the number of redox centers in the cluster
and the redox state of the cluster. While rubredoxin has only one
iron that simply switches between Fe2+ and Fe3+ states, the same transition differs
significantly in a [4Fe–4S]
cluster in an environment with three more irons and a mixed-valence
state (e.g., 2Fe3+–2Fe2.5+ and Fe2.5+). Even the same cluster can undergo different
redox transitions,
as has been observed in the case of HiPIPs and ferredoxins.
719
3.7.2
Role of Ligands
While sulfurs are
the most dominant ligands in Fe–S proteins, it has been shown
that other ligands can replace sulfurs in some cases and that these
ligands play a prominent role in fine-tuning the reduction potential
of the proteins.
541
Generally speaking,
ligands that are less electron-donating than sulfur will increase
the reduction potentials by selectively destabilizing the oxidized
state. A well-established example of this principle is the increased
reduction potential of [2Fe–2S] clusters in Rieske proteins
compared to ferredoxins due to replacement of two of the Cys ligands
with His residues. Mutational studies on Cys ligands, mostly replacement
with Ser, have shown an increased reduction potential compared to
that of the wild-type (WT) proteins.
721,750,773,1087
3.7.3
Role of the Cellular Environment
As mentioned earlier
in this review, some Fe–S proteins such
as vertebrate ferredoxins and certain [3Fe–4S] clusters and
Rieske proteins show pH-dependent redox behavior. This behavior can
be due to the presence of a protonable residue such as Asp or His
residue as a ligand or near the active site.
712,746,801
Therefore, proteins in the presence
of different pH values in different cellular compartments should demonstrate
different reduction potentials. Another effect of the environment
is indirect through evolution: as shown in the case of ferredoxins,
organisms subjected to extreme environments will undergo changes in
the overall charges of proteins, which will affect the reduction potentials.
823
Peptide models of different Fe–S clusters
have demonstrated the impact of solvent composition in ET features
of the cluster.
717
3.7.4
Role of the Protein Environment
Several studies have
shown the importance of the protein environment
in fine-tuning the reduction potentials of metal centers. The protein
environment is one of the, if not the, most important factors determining
the reduction potential in Fe–S proteins because the general
geometry and primary coordination of iron are very similar in this
family of proteins. The protein environment conveys its effect via
several routes.
3.7.4.1
Solvent Accessibility/Cluster
Burial
Solvent accessibility has been shown to be a very
important factor
in the reduction potential for different metal centers, including
Cu centers, hemes, and Fe–S clusters. As a general rule of
thumb, the more buried a cluster, the higher or more positive the
reduction potential will be. This is mainly due to the electrostatic
destabilization of more positive charges in the clusters. Being more
buried is proposed to be one of the most important reasons behind
the difference between the reduction potentials of the [4Fe–4S]
clusters in HiPIPs vs ferredoxins.
618,749,752
Hydration of the cluster can influence the covalency
of Fe–S bonds, hence affecting the reduction potential.
901
Cluster burial can be accomplished through
physical positioning of the cluster by covering it with more secondary
structure elements or partially via more hydrophobic residues around
the cluster. As discussed earlier, there are exceptions to this trend,
and there are clusters that are significantly more solvent-exposed,
but little reduction potential change is observed for them.
875
It should be noted that cluster burial is dependent
on the size of the protein, the location of the cluster, and the extent
of solvent interaction, so it is difficult to make a fair comparison
of the effect of cluster burial among different proteins.
92
3.7.4.2
Secondary Coordination
Sphere
While ligands in the primary coordination sphere are
very important
in tuning the reduction potentials of the Fe–S centers, the
role of secondary coordination sphere interactions cannot be ignored.
A mounting number of studies support the essential roles of these
interactions in fine-tuning the reduction potentials.
1088
In the case of Fe–S proteins, secondary
coordination interactions are the major cause of differences in the
reduction potentials within a class of proteins.
887
The number of backbone to amide H-bonds has been shown
to be important in redox potential differences between HiPIPs and
ferredoxins.
617,618
As described in each section,
a conserved H-bonding pattern is observed in each subclass of ferredoxins,
and this pattern differs from one subclass to another.
718,719
Removal of some conserved H-bonds from this pattern is shown to
be one of the main causes of different reduction potentials between
different types of ferredoxins.
718,719
Removal of
conserved H-bonds in several cases resulted in a decrease in the reduction
potential.
773,780
It is important to mention that
although H-bonds are important, they are not the sole cause of differences
in the reduction potentials. Moreover, their analyses are complicated
in some cases due to ambiguity in their assignment and variation in
their number based on the environmental condition.
92
3.7.4.3
Electrostatics and
Local Charges
Local charges can selectively stabilize either
the reduced or oxidized
form of the cluster and influence the reduction potential. Many studies
of the Fe–S proteins showed that although these proteins usually
have conserved charged residues (such as positive charges in ferredoxins),
these charges are mainly important for interaction with the redox
partner, and usually their mutations do not cause significant changes
in the reduction potential.
749
In cases
where these residues are very close to the cluster, unpredictable
effects have been observed.
611
However,
the total charge of the cluster has been suggested to be an important
factor influencing the higher reduction potential of Rieske proteins
compared to ferredoxins.
773
Mutational
analysis on rubredoxins and thioredoxin-like ferredoxins confirmed
an important role for the charges around the cluster in the reduction
potential of the protein. There is convincing evidence for the role
of backbone amides and partial positive charges in the reduction potential
of Fe–S centers.
887
It has been
proposed that the diploes induced by the these backbone amides can
influence the reduction potential of different clusters, such as HiPIPs
and ferredoxins. The net protein charge and the dipole induced from
backbone amides have been shown to be important in determining the
reduction potential of HiPIPs.
752,873,890
While all these features are important, it should be noted
that none of them are the sole determinants of the reduction potential
in Fe–S proteins, and it has been found that different features
act as the major contributors to differences in the reduction potential
between different classes of the Fe–S proteins. Even among
members of a class, the same factor might not play the same role.
3.7.5
Computational Analysis of the Reduction
Potentials of Fe–S Proteins
To further understand
factors influencing the reduction potentials, computational methods
have been developed for calculating the reduction potential of Fe–S
proteins on the basis of their structures.
591,887
One of these methods uses Gunner’s multiconformational continuum
electrostatics method and has been calibrated using proteins with
known structure and reduction potential.
780
In another method a combined quantum-chemical and electrostatic
calculation was used to generate predictions for reduction potentials.
Poisson–Boltzmann electrostatic methods in combination with
QM/MM studies have also been used to analyze the reduction potentials
of Fe–S proteins.
93
The PDLP method
was applied to HiPIPs to analyze the effects of solvent accessibility
on the reduction potentials of these proteins.
92,719
B3LYP density functional methods have been used in combination with
broken symmetry to analyze factors that are important in tuning the
reduction potential of Rieske proteins.
800
Broken symmetry in combination with hybrid density functional theory
has also been used to characterize Rieske proteins.
1089
4
Copper Redox Centers in Electron
Transfer Processes
4.1
Introduction to Copper
Redox Centers
Copper is the second most abundant transition
metal in biological
systems, next to iron.
1090
In addition
to their critical role in electron transfer process, copper-containing
proteins catalyze a variety of reactions. In this section, we focus
on copper proteins that merely function as ET mediators, which include
blue or type 1 (T1) copper and CuA centers. A number of
reviews on these two centers have appeared in the literature.
94−104
Despite the lack of modern structural and computational methods,
initial attempts to understand the structure and function of copper
redox centers were very successful. This success was in part due to
the strong colors and interesting magnetic properties displayed by
these redox centers that allowed various spectroscopic studies. The
blue copper proteins were so-named because they display an intense
blue color, due to a strong absorption around 600 nm, first observed
in the 1960s.
1091,1092
It was found that this T1 copper
protein also displayed an unusual EPR spectrum with narrow hyperfine
splittings, suggesting the presence of Cu in a different ground state
compared to the normal copper complexes.
1093
The electronic structure of the blue copper center was further elucidated
with low-temperature absorption, CD, MCD, single-crystal EPR, XAS,
and computational studies.
96,99,1094,1095
The results of all these studies
demonstrated that the 600 nm band is associated with a S →
Cu charge transfer transition and that the highly covalent nature
of the Cu–S bond is responsible for the narrow hyperfine splitting
in the EPR spectra. The crystal structure of poplar plastocyanin later
confirmed that T1 copper proteins contain a copper site with an unusual
geometry.
1096
Although the existence
of copper in cytochrome c oxidases (CcOs) has been known since the 1930s,
the nature of the CuA centers was not established until
much later due to the presence of heme cofactors that complicated
interpretation of the spectroscopic results.
1097
EPR and elemental analyses have revealed that two copper-binding
sites exist in CcOs.
1098−1100
MCD studies by Thomson
and co-workers showed features at 475, 525, and 830 nm corresponding
to a CuA center.
1101,1102
Kinetic measurement
of reoxidation of reduced CcO, performed by a flow-flash
technique, indicated that the CuA is the ET center in CcO.
1103,1104
From 1987 to 1993, Buse and
co-workers performed chemical analysis of CcO with
inductively coupled plasma atomic emission spectroscopy, leading to
the conclusion that three copper atoms exist in one protein along
with two hemes.
1105,1106
Later, resonance Raman,
1107
EXAFS,
1108
and
finally crystal structures
1030,1109
revealed an unusual
dinuclear copper structure for the CuA center, which will
be discussed in detail in section 4.5.
4.2
Classification of Copper Proteins
As a diverse family
of proteins, copper proteins could be divided
into several types according to ligand sets, spectroscopic features,
and functions (Table 9).
1110,1111
Mononuclear T1 copper centers and dinuclear CuA centers
are the two types which act only as ET mediators. T1 copper centers
and CuA centers share several common features. First, both
centers contain Cu–thiolate bond(s), which are highly covalent
and display rich spectroscopic signatures.
99,1095,1112−1115
Second, both centers are located in a cupredoxin fold.
94,100,103
Finally, they are highly optimized
for ET, showing low reorganization energies and high ET rate constants.
These two types of copper proteins are collectively called cupredoxins,
analogous to ferredoxin for Fe–S-based ET centers.
1116
Other types of copper proteins may also involve
ET as part of their enzymatic reactions, including peptidylglycine
α-hydroxylating monooxygenase and dopamine β-monooxygenase,
1117
but will not be discussed here.
Table 9
Different Types of Copper Proteinsa
mononuclear
dinuclear
tetranuclear
type 1
type 2
type 3
CuA
CuZ
UV–vis spectrum
strong absorption, ∼600 nm and (in some proteins) 450 nm
weak absorption, ∼700 nm
300–400 nm
strong absorption, ∼480
and 530 nm
strong absorption, ∼640 nm
EPR
spectrum
four-line (A
|| < 80 × 10–4 cm–1)
four-line (A
|| ≈ (130–180) × 10–4 cm–1)
nondetectable
seven-line (A
|| ≈ 30–40 × 10–4 cm–1)
2 × four-line (A
|| ≈ 61 × 10–4 cm–1 and A
|| ≈ 24 × 10–4 cm–1)
common ligands
His, Cys (Met)
His, Asp (Tyr)
His (Tyr)
His, Cys (Met)
His, S2–
active site geometry
trigonal pyramidal or distorted
tetrahedral
distorted
tetragonal
tetragonal
trigonal planar
m
4-S2– tetracopper cluster
examples
azurin
superoxide
hemocyanin
cyt c oxidase
N2O reductase
plastocyanin
dismutase
tyrosinase
N2O reductase
stellacyanin
galactose
oxidase
catechol
menaquinol NO reductase
nitrite reductase
amine oxidase
oxidase
nitrite reductase
laccase
laccase
laccase
a
Reprinted with permission from ref (98). Copyright 2004 Elsevier.
4.3
Native Type 1 Copper Proteins
Exclusively
serving as ET centers, T1 copper proteins are distinct from other
copper proteins because of their unique geometry and ligand sets.
The copper ion is normally coordinated to two histidines and one cysteine
in a trigonal plane with the axial position often occupied by a methionine
at a relatively longer distance. They contain a highly covalent copper–thiolate
bond that imparts an intense blue color to the T1 centers, due to
absorption at ∼600 nm, and narrow four-line hyperfine splitting
in the EPR spectra.
99,1118
The T1 copper centers
reside in either single- or multiple-domain proteins.
1119
The former includes the most common T1 copper
proteins, such as plastocyanin, azurin, and amicyanin, while the latter
includes stellacyanin, uclacyanin, and dicyanin. The T1 copper centers
are also found in multicopper centers involving other types of copper
centers, such as in nitrite reductases, laccases, and ascorbate oxidases.
We will discuss the T1 copper centers in single- and multiple-domain
proteins in this section, while the T1 copper centers in multicopper
proteins will be discussed in section 4.3.4.
The T1 copper proteins are found in archaea, bacteria, and
plants.
In addition to the cupredoxin fold, genes containing the T1 copper
proteins may contain other components (Figure 45). All T1 copper proteins have an
N-terminal signal peptide or transit
peptide. With the signal peptide, the T1 copper proteins from bacteria
or archaea are directed into the periplasmic space. Their counterparts
in plants, on the other hand, are transported to the extracellular
milieu and anchored to the cell surface through an additional C-terminal
hydrophobic sequence.
1119
Plastocyanin
is guided to the chloroplast in plant cells by a transit peptide sequence
that is cleaved in the mature protein.
1120
Figure 45
Domain arrangement of type 1 copper protein. Reprinted with permission
from ref (1119). Copyright
2006 Wiley-VCH.
4.3.1
Structures
of the Type 1 Copper Proteins
The first crystal structure
of the T1 copper protein, plastocyanin
from poplar leaves (Populus nigra var. italica), was reported in 1978.
1096
Since then, crystal structures of many other T1 copper
proteins have been reported, as listed in Table 10. Despite the fact that sequence
identity between the T1 copper
proteins is less than 20%,
1154
the overall
structural folds of different T1 copper proteins are highly conserved.
This common fold is called cupredoxin fold, which consists of eight
β-strands arranged into a Greek key β-barrel as shown
in Figures 46 and 47.
94
There are also one to two α-helices
in different locations outside the core fold of the protein. This
fold is present not only in T1 copper proteins and the CuA domain,
1155
but also in other copper
proteins, such as Cu–Zn SOD,
94,1156
and in proteins
without metal cofactors, such as immunoglobins.
94,1157
Table 10
Properties of T1 Copper Proteins
name
organism
isolated from
first reported
PDB code
for first structure
ligand set
E
m (mV)
redox partner
Single
Domain
azurin
bacteria
1962
1121
1AZU
1Cys, 2His, 1Met, 1 carbonyl
oxygen
310
1122
amicyanin
methylotrophic bacteria
1981
1123
1MDA
1Cys, 2His, 1Met
260
1124
methylamine dehydrogenase,
cytochrome c
551
plastocyanin
plant/algae/cyanobacteria
1960
1125
1PLC
1Cys, 2His, 1Met
370
1126
cytochrome f, P700+
pseudoazurin
denitrifying bacteria and
methylotrophs
1973
1127
1PAZ
1Cys, 2His, 1Met
280
1128
nitrite reductase
rusticyanin
acidophilic bacteria
1975
1129
1RCY
1Cys, 2His, 1Met
670
1130
cytochrome c, cytochrome c
4
auracyanin
photosynthetic bacteria
1992
1131
1QHQ
1Cys, 2His, 1Met
240
1131
plantacyanin
plants
1974
1132
2CBP
1Cys, 2His, 1Met
310
1133
halocyanin
haloalkaliphilic archaea Natronobacterium
pharaonis
1993
1134
1Cys, 2His, 1Met
183
1134
sulfocyanin
acidophilic archaea Sulfolobus acidocaldarius
2001
1135
1Cys, 2His, 1Met
300
1135
nitrosocyanin
autotrophic bacteria
2001
1136
1IBY
1Cys, 2His, 1Glu,
1H2O
85
1137
Multidomain Protein with T1 Center
stellacyanin
plants
1967
1138
1JER
1Cys, 2His, 1Gln
190
1133
uclacyanin
plants
1998
1139
1Cys, 2His, 1Met
320
1139
dicyanin
plants
2000
1140
1Cys, 2His, 1Gln
Multidomain Protein with T1 Center and Other Copper
Center
laccase
fungi
1A65
1Cys, 2His (1Leu/Phe)
465–778
1141−1143
Pplants
1Cys, 2His, 1Met
434
1144,1145
ascorbate oxidase
plants
1AOZ
1Cys, 2His, 1Met
350
1146
ceruloplasmin
animals
1948
1147
1KCW
1Cys, 2His (1Leu)
>1000
1148
(redox-inactive)
ceruloplasmin
1Cys, 2His, 1Met
448
1149
(redox-active)
hephaestin
mammals
1999
1150
Fet3p
yeast
1994
1151
1ZPU
1Cys, 2His
427
1152
nitrite
reductase
plants,
bacteria
1NIA
1Cys, 2His, 1Met, 1 carbonyl
oxygen
260
1153
Figure 46
Crystal structures of the T1 copper proteins. The secondary
structure
(α-helix and β-sheet) is shown in cartoon format, copper
is shown as a purple ball, and ligands are shown in stick format.
The name of the protein and its PDB ID are given below each structure.
Figure 47
Topology diagram showing the scheme of
the secondary structure
of azurin. β-Strands are shown as arrows, and the α-helix
is shown as a cylinder. Copper ligands between β-strands 3 and
4 and between β-strands 7 and 8 are shown as blue polygons,
while copper is shown as a purple circle.
Most of the ligands to the T1 copper center resides at the
C-terminal
end of the cupredoxin fold. As shown in Figure 47, one of the His ligands is the first
residue of the fourth β-strand
and is referred to as N-terminal His. Carbonyl oxygen, the fifth ligand
of azurin, is located in the loop between the third and fourth β-strands.
Other ligands, including Cys, the second His on the trigonal plane,
and the axial ligand, are located in or adjacent to the loop between
the seventh and eighth β-strands, close to the C-terminus of
the protein. Cys is the last residue of the seventh β-strand,
while the second His is in the middle of the loop and is referred
as the C-terminal His. Met is the first residue of the eighth β-strand.
The three ligands are arranged in Cys-(Xxx)
n
-His-(Xxx)
m
-Met fashion, where n and m could vary between 2 and 4 in different
T1 copper proteins. This variation in length and amino acid composition
is important for the function of T1 copper proteins. In section 4.4.5 we discuss the
implications of the variations
based on loop-directed mutagenesis results.
While X-ray crystallography
could give a fairly good description
of the overall structure, EXAFS is more accurate in determining the
metal–ligand distance because it is sensitive to oxidation
state of the metal ion.
1158
The short Cu–S
distance was first revealed by EXAFS.
99,1159
By comparing
data from oxidized and reduced plastocyanin and azurin, it was found
that an average increase of ∼0.06 and ∼0.08 Å for
Cu–N(His) and Cu–S(Cys), respectively, happens upon
reduction.
99
These small changes upon reduction
are consistent with data from crystallography and suggest a small
reorganization energy for the redox process.
4.3.1.1
Copper
Ligands
Even though the
amino acid sequences and overall structures vary among different T1
copper proteins, the ligand composition, ligand–metal distance,
and geometry of the T1 copper centers are almost identical (Figure 48).
94,95,99
As the most conserved structural feature, T1 copper centers invariably
contain two His residues and one Cys residue as equatorial copper
ligands. In T1 copper proteins, the His coordinates with copper through
Nδ, in contrast to Nε used by T2 and most other copper
proteins. The Cu–His bond length is about 2.0 Å in T1
copper proteins, which is normal for such types of bonds. On the other
hand, the Cu–Cys bond lengths range from 2.07 to 2.26 Å,
which is short compared to those of normal copper complexes and other
copper proteins (Table 11). The short Cu–S
distance is key to the unique spectroscopic properties of T1 copper
and is maintained through extensive H-bonding within the protein scaffold,
as will be discussed later in this section. The 2N and 1S from His
and Cys, respectively, form a pseudotrigonal plane, with average
bond angles in the Cu(II) state being 101°, 117°, and 134°
with RMS deviations of 2.5°, 4.1°, and 2.8°, calculated
from crystal structures with resolution of 2.0 Å or higher.
1119
The Cu–Sγ–Cβ–Cα and Sγ–Cβ–Cα–N dihedral angles
are also consistently close to 180°, making the Cu–Sγ bond coplanar with the Cys side
chain and backbone.
Figure 48
T1 copper
centers in plastocyanin, azurin, plantacyanin, and amicyanin.
Reprinted with permission from ref (1119). Copyright 2006 Wiley-VCH.
Table 11
Distances (Å) between Cu or
Other Substituted Metals and Ligands in T1 Copper Proteinsa
P. aeruginosa azurin
pH
Cu–Nδ (His46)b
Cu–S (Cys112)b
Cu–Nδ (His117)b
Cu–S (Met121)b
Cu–O (Gly45)b
resolution
(Å)
PDB ID
ref
Cu(II)
5.5
2.08(6)
2.24(5)
2.01(7)
3.15(7)
2.97(10)
1.9
4AZU
(1160)
Cu(I)
5.5
2.14(9)
2.29(2)
2.10(9)
3.25(7)
3.02(8)
2.0
1E5Y
Cu(II)
9.0
2.06(6)
2.26(4)
2.03(4)
3.12(7)
2.94(11)
1.9
5AZU
(1160)
Cu(I)
9.0
2.20(11)
2.30(23)
2.21(12)
3.16(9)
3.11(11)
2.0
1E5Z
T. ferrooxidans rusticyanin
pH
Cu–Nδ (His85)
Cu–S (Cys138)
Cu–Nδ (His143)
Cu–S (Met148)
–
resolution (Å)
PDB ID
ref
Cu(II)
4.6
2.04
2.26
1.89
2.88
–
1.9
1RCY
(1161)
Cu(I)
4.6
2.22
2.25
1.96
2.75
–
2.0
1A3Z
P. nigra plastocyanin
pH
Cu–Nδ (His37)
Cu–S (Cys84)
Cu–Nδ (His87)
Cu–S (Met92)
–
resolution (Å)
PDB ID
ref
Cu(II)
6.0
1.91
2.07
2.06
2.82
–
1.33
1PLC
(1162)
Cu(I)
7.0
2.13
2.17
2.39
2.87
–
1.80
5PCY
(1163)
P. denitrificans amicyanin
pH
Cu–Nδ (His53)
Cu–S (Cys92)
Cu–Nδ (His95)
Cu–S (Met98)
–
resolution (Å)
PDB ID
ref
Cu(II)
6.0
1.95
2.11
2.03
2.90
–
1.31
1AAC
(1164)
Cu(I)
7.7
1.95
2.12
unbound
2.91
–
1.30
2RAC
(1165)
C. sativus cucumber basic protein
pH
Cu–Nδ (His39)
Cu–S (Cys79)
Cu–Nδ (His84)
Cu–S (Met89)
–
resolution (Å)
PDB ID
ref
Cu(II)
6.0
1.93
2.16
1.95
2.61
–
1.80
2CBP
(1166)
C. sativus stellacyanin
pH
Cu–Nδ (His46)
Cu–S (Cys89)
Cu–Nδ (His94)
–
Cu–O (Gln89)
resolution
(Å)
PDB ID
ref
Cu(II)
7.0
1.96
2.18
2.04
–
2.21
1.60
1JER
(1167)
a
Adapted with permission from ref (104). Copyright 2012 Elsevier.
b
Average of distances for four
molecules
in the asymmetric unit. Errors are 1 standard deviation.
The axial ligand in the T1 copper
center is less conserved. A Met
is present at 2.6–3.2 Å in this axial position in most
proteins, while a Gln is found in stellacyanin and dicyanin. In the
T1 center of fungal laccase and ceruloplasmin, a noncoordinating ligand
such as Phe or Leu takes this axial position. In azurin, there is
an additional backbone carbonyl oxygen at the opposite end of the
axial position to Met, giving the T1 copper site a trigonal bipyrimidal
geometry.
4.3.1.2
Secondary Coordination
Sphere
While the above mentioned ligands exert significant
influence on
the properties of T1 copper centers, the protein scaffold should not
be viewed as a passive entity to hold the copper site. On the contrary,
it can play important roles. First, it can shield the copper site
from water, raising the reduction potential and lowering the reorganization
energy for ET. More importantly, the extensive H-bond network surrounding
it can fine-tune the properties of the T1 copper site.
94,98
As shown in Figure 49, the Cys112 in
azurin forms two hydrogen bonds with adjacent backbone amide groups
of Asn47 and Phe114 at ∼3.5 Å. Together with S–Cu
and S–Cβ covalent bonds, these H-bonds form
a tetrahedral geometry around Sγ of Cys (Figure 49A). Plastocyanin, pseudoazurin,
and amicyanin have
only one H-bond around the Cys as a Pro in the site eliminates the
other amide bond. Additionally, cucumber basic protein has a very
weak H-bond at 3.7–3.8 Å. These H-bonds modulate the electron
density of S on Cys, which is crucial for the highly covalent nature
of the Cu–S bond.
Figure 49
H-bonding around Cys112 (A) and other ligands
(B) of azurin. PDB
ID 4AZU.
In azurin, the N-terminal His
coordinates with Cu through Nδ,
whereas Nε is hydrogen-bonded to the carbonyl oxygen of Phe15.
The same His is hydrogen-bonded to the Gln49 side chain in amicyanin,
the side chain of Asn80 in rusticyanin, and a water molecule in phytocyanins.
The C-terminal His is in a hydrophobic patch of the protein packed
against other residues. The Nε of C-terminal His is hydrogen-bonded
to a water molecule. The axial Met/Gln usually packs against aromatic
side chains such as Phe15 in azurin (Figure 49). In azurin, the carbonyl oxygen is
held in place by the secondary
structure of the loop and packs with Phe114.
There are more
H-bonding interactions beyond the copper center.
For example, an Asn close to the N-terminal His in the first ligand
loop is hydrogen-bonded to residues from the other ligand loop. This
interaction, acting like a zipper, further holds the copper site together.
Extensive H-bonding around the copper site in T1 copper proteins
has important functional implications, as we will address in section 4.4.2.
4.3.1.3
Comparison of Structures
in Different
States
As suggested by the “rack mechanism”
1168,1169
or entatic state,
1170
the active site
structure is predetermined by the protein scaffold. Thus, there is
little change in the structures of T1 copper proteins at different
oxidation states, with different metals, or even in the absence of
metal ions.
As shown in Table 11, compared
to the same protein with Cu(II), the metal to ligand bonds elongated
by 0.1 Å or less in protein containing Cu(I). Similar results
were obtained by EXAFS, which provides a more accurate determination
of the bond length.
99
The small change
in bond length is crucial for the low reorganization energy of the
T1 copper site and, thus, fast ET for its function. However, bond
lengths in X-ray crystal structures should be interpreted with caution,
as it has been shown that Cu(II) ions in protein undergo photoreduction
during X-ray exposure.
1171,1172
It will be useful
to conduct single-crystal microspectrophotometry concurrent with X-ray
diffraction to make sure that the oxidized protein is not reduced
during diffraction.
1173
On the other hand,
the oxidation state of the Cu ion can be easily monitored at the edge
and XANES regions of its X-ray absorption spectrum. Bond lengths derived
from carefully designed and conducted EXAFS should reflect the actual
bond lengths at the corresponding oxidation states.
Besides
structures with copper in oxidized or reduced states, crystal
structures of apo and metal-substituted T1 copper proteins also shed
light on how proteins interact with copper. Structures of apo forms
of azurin,
1174,1175
plastocyanin,
1176
pseudoazurin,
1177
and amicyanin
1178
show little difference
(0.1–0.3 Å) from that of the copper-bound form, confirming
the entatic state hypothesis.
Metal substitution is useful in
spectroscopic studies, such as
electronic absorption
1118,1179
and NMR.
1180
Due to the different sizes and ligand affinities
of different metals, the bond length and overall geometry are changed
upon substitution, but only to a small extent due to confinement of
the protein scaffold.
1181−1183
4.3.2
Spectroscopy
and Electronic Structure
Intense (∼5000 M–1 cm–1) electronic absorption at ∼600 nm
is the hallmark of T1 copper
proteins (Figure 50). Solomon and co-workers
attributed the origin of the ∼600 nm absorption to the S(Cys)pπ
→ Cu
x
2–y
2
LMCT transition.
1094,1184,1185
Another feature at ∼450 nm is not prominent
in plastocyanin or azurin, but is more pronounced in a perturbed T1
copper sites such as that of cucumber basic protein. This absorption
is attributed to S(Cys)pπ → Cu
x
2–y
2
LMCT. The
geometry of the copper site is believed to be important for the ratio
between the two peaks at ∼600 and ∼450 nm.
1095,1186
A series of weak absorption peaks from 650 to 1050 nm are attributed
to a d → d transition or ligand field transition.
1184
Figure 50
Electronic absorption (A) and EPR (B) spectra
of azurin.
EPR provides a sensitive
way to determine the copper site geometry.
T1 copper proteins exhibit a distinctive small hyperfine splitting
(<100 × 10–4 cm–1) in
the EPR spectrum, as opposed to that of T2 copper and other complexes
(>150 × 10–4 cm–1).
1119
Through S K-edge XAS, Solomon and co-workers
showed that the small hyperfine splitting is due to high covalency
between Cu and S, which delocalizes unpaired electrons onto S, thus
decreasing the electron density on Cu.
1187
Other spectroscopic techniques, such as resonance Raman spectroscopy
and Cu L-edge and S K-edge XAS, have also been important in deciphering
the electronic structures of T1 copper proteins. They are beyond the
scope of this review, but there are excellent reviews elsewhere
1095,1119
and in this issue that cover more details about these techniques.
2000
4.3.3
Redox Chemistry of Type
1 Copper Protein
As a class of proteins dedicated to ET,
T1 copper proteins display
various features for facile redox chemistry.
4.3.3.1
Redox
Partner
T1 copper proteins
shuttle electrons between donor and acceptor proteins as redox partners.
So far five T1 copper proteins with known physiological redox partners
have been identified: plastocyanin, amicyanin, rusticyanin, pseudoazurin,
and azurin. As an electron carrier in chloroplasts in plants, plastocyanin
accepts electrons from cytochrome f of membrane-bound
cytochrome b
6
f complex
and transfers them to P700+ in photosystem I.
256,1188−1192
Amicyanin accepts electrons from methylamine dehydrogenase and transfers
them to cytochrome c oxidase via a c-type cytochrome.
279,1193−1200
Rusticyanin is suggested to shuttle electrons between cytochrome c and cytochrome
c
4.
1201,1202
Pseudoazurin reduces nitrite reductase, but its electron donor is
not yet known.
1203−1207
Azurin is likely to interact with aromatic amine dehydrogenase in
vivo, as suggested by coexpression, the kinetics of reduction, and
the crystal structure.
1208−1210
Interaction between a
T1 copper protein and its redox partner is generally weak and transient.
NMR and crystallographic studies have revealed a structural basis
for this interaction. Interactions between plastocyanin from various
organisms and cyt f have been extensively studied
by NMR spectroscopy (Figure 51). Chemical shift
analysis and rigid-body structure calculations have demonstrated that
the hydrophobic patch around His87, the C-terminal His ligand to copper,
mediates the interaction between plastocyanin and cyt f.
1211,1212
Besides that, two acidic patches around
Tyr83 have been shown to interact with positively charged residues
of cyt f.
1213
Mutation
of Tyr83 to Phe or Leu drastically decreases the ET rate between the
two proteins, indicating that Tyr83 is involved in binding to cyt f and ET.
1214
The absence of
acidic patches also demolishes ET activity at low ionic strength,
showing they are involved in the interaction with cyt f.
1215,1216
However, interaction between acidic patches
and cyt f is not very specific as small changes in
acidic patches have a minimal effect on the interaction between two
proteins.
1216,1217
Figure 51
Structures of plastocyanin
(left) and the complex of plastocyanin
and cyt f (right). Left: copper ion is represented
as a purple ball, His87 and Tyr 83 are represented in licorice format,
and residues in two acidic patches are represented as ball and stick
models. Right: plastocyanin is colored cyan, and cyt f is orange. The copper ion and
His87 from plastocyanin and heme from
cyt f are also shown.
Another demonstration of the interaction between the T1 copper
proteins and their redox partners comes from X-ray crystallography.
The structures of the amicyanin–methylamine dehydrogenase complex
and methylamine dehydrogenase–amicyanin–cytochrome c
551 ternary complex have been determined.
279,1196
These structures further confirmed that the hydrophobic patch surrounding
His95 (the C-terminal His ligand equivalent to His87 in plastocyanin
and His117 in azurin) interacts with a hydrophobic patch on methylamine
dehydrogenase. An ET pathway from Trp57 and Trp108 in methylamine
dehydrogenase to His95 in amicyanin and eventually to copper has been
proposed from these structures.
Recently, the crystal structure
of the azurin and aromatic amine
dehydrogenase complex from Alcaligenes faecalis has been solved.
1208
In this structure,
only one azurin molecule is present in complex with four molecules
of aromatic amine dehydrogenase. The B factor of
the azurin structure is high except for those residues in the interface.
This result is consistent with the transient nature of the interaction
between the T1 copper proteins and their redox partners. The interaction
is very similar to the one between amicyanin and methylamine dehydrogenase.
The T1 copper proteins show promiscuity in reacting with proteins
other than their physiological redox partners,
64,1218
including small inorganic complexes such as [Fe(CN)6]3– and [Co(phen)3]3+,
31,44,1219
small molecules such as flavins
and ascorbate, and the proteins themselves through electron self-exchange
reactions.
100
Gray and co-workers have
used Ru derivatives of T1 copper proteins as a model to study long-range
ET in biological systems.
24,31,44,2005
4.3.3.2
Electron
Transfer Rate
T1 copper
proteins are involved in long-range ET in vivo and in vitro. For a
more detailed review of long-range ET, please refer to the review
in this issue by Gray et al.
2005
The process
can be described by the semiclassical Marcus equation:
Marcus equation
1
In this equation,
ΔE° is the difference in reduction potential
between the donor and acceptor sites (also known as the driving force), H
AB is the donor–acceptor electron coupling
or electron matrix coupling element, and λ is the reorganization
energy required for ET. Under the same driving force, the rate is
maximized when H
AB is large and λ
is small. In long-range ET, there is little direct coupling between
the donor and the acceptor. The coupling is mediated by intervening
atoms via the superexchange mechanism. H
AB is determined by the distance between the donor and acceptor and
the covalency of the metal–ligand bond.
1220−1222
Electron transfer rates between T1 copper proteins and their
redox partners have been measured by kinetic UV–vis spectroscopy
or cyclic voltammetry.
1223−1226
The k
ET between
plastocyanin and cyt f has been determined to be
2.8–62 s–1,
1227−1229
while the constant
between plastocyanin and P700+ has been determined to be
38–58 s–1.
1191,1192,1230,1231
Davidson and co-workers
have used kinetic UV–vis spectroscopy to measure the k
ET between amicyanin and methylamine dehydrogenase,
which was determined to be ∼10 s–1.
1232,1233
Suzuki and co-workers have determined the k
ET between pseudoazurin and nitrite reductase to be (0.8–7)
× 105 M–1 s–1 by
kinetic UV–vis spectroscopy or cyclic voltammetry.
1204,1224,1234−1236
As several studies have pointed out, the rate constant measurement
for interprotein ET processes is complicated by other processes, such
as multiple binding sites of the two proteins, transient formation
of conformational intermediates, and protonation/deprotonation processes.
1225,1237
There are two methods to measure the ET rate in T1 copper proteins
without involvement of a redox partner: pulse radiolysis and NMR.
Pulse radiolysis
1238
uses a short pulse
(typically 0.1–1 μs) of high-energy (2–10 MeV)
electrons to excite and decompose solvent molecules. A typical reaction
generates the CO2
•– radical:
Radicals
generated in solvent molecules trigger downstream reactions.
In azurin, CO2
– can reduce either Cu(II)
or the disulfide bond between Cys3 and Cys26 at a nearly diffusion-controlled
rate. Molecules with a reduced disulfide bond (RSSR–) can further reduce Cu(II) in
the same protein via intramolecular
ET:
101
By monitoring absorbance changes at 410 nm (RSS•R–) and 625 nm (Cu(II)), a fast reduction
process
corresponding to reduction of Cu(II) or RSS•R– by CO2
•– and a
slower process of intramolecular ET between RSSR and Cu(II) can be
resolved. The ET rate and driving force (ΔG°) can be calculated from the kinetics
of intramolecular ET.
By running experiments at different temperatures, the activation enthalpy
and activation entropy of the ET process can be calculated.
Using this method, Farver and Pecht determined the rate constant
of intramolecular ET of WT azurin to be 44 ± 7 s–1 at pH 7.0 and 25 °C with a driving
force ΔG° = −68.9 kJ mol–1. The activation
enthalpy and activation entropy were calculated to be 47.5 ±
4.0 kJ mol–1 and −56.5 ± 7.0 J K–1 mol–1.
1239
ET rates for azurin of different origins and mutations have been
measured and reviewed by Farver and Pecht.
101
Electron self-exchange is an intrinsic property of all redox
systems.
1240
Exchange of electrons happens
to two molecules
of the same complex at different oxidation states. Only one redox
couple is involved, and there is no driving force for this reaction.
Measuring electron self-exchange rate constants by NMR provides a
more universal way to measure ET transfer activity as it is carried
out in T1 copper centers
1241−1249
(reviewed in ref (100)) as well as in other redox centers.
1250−1252
Electron self-exchange
rate constants (k
SES) of T1 copper proteins
range from 103 to 106 M–1 s–1 at moderate to low ionic strength. The electron self-exchange
is thought to happen through a hydrophobic patch as the rate constant
is affected by the presence of an acidic patch
1248
or basic residues
1253
close
to the hydrophobic patch.
4.3.3.3
Reduction
Potential
T1 copper
proteins have reduction potentials ranging from 183 to 800 mV (see
Table 10). Compared to the aqueous Cu(I)/Cu(II)
couple (which has a reduction potential of ∼150 mV), copper
complexes, and other copper proteins, T1 copper proteins have unusually
high reduction potentials. Their potentials also span a wide range
(>600 mV), nearly half the range of biologically relevant potentials
(Figure 1). Within the T1 copper proteins,
groups of proteins are apparent when sorted on the basis of the midpoint
reduction potential (E
m). Nitrite reductases,
1153
stellacyanins,
1133
amicyanins,
1124
and pseudoazurins
1128
natively have substantially lower (∼100
mV) E
m values as compared to azurin.
98
Azurin and umecyanins have moderate E
m values natively around 200–300 mV vs
SHE. On the other end of the scale, rusticyanins have E
m values ∼400 mV higher than that of azurin. Understanding
the origin of this variance and the structural features involved in
tuning the reduction potential are of great importance. By comparing
the native proteins with different axial ligands (Table 12), it is revealed that proteins
with Gln as an
axial ligand generally have lower reduction potentials (190–320
mV), proteins with Met axial ligands have higher potentials (183–670
mV), and proteins with a noncoordinating ligand in multicopper proteins
have the highest potentials (354–800 mV). This trend is further
confirmed by mutagenesis studies that are discussed in section 4.4.1.
Table 12
Dependence of E°
on the Axial Ligand in Blue Cu Proteinsa
E° (mV)
Phe/Leu/Thr
Met
Gln
ref
fungal laccase
770
680
(1254−1256)
azurin
412
310
285
(1122, 1257)
cuc. stellacyanin
500
420
260
(1139)
nitrite reductase
354
247
(1258)
rusticyanin
800
667
563
(1259)
mavicyanin
400
213
(1260)
amicyanin
250
165
(1261)
a
Reprinted from
ref (99). Copyright
2004 American
Chemical Society.
Variation
within proteins containing the same axial ligand indicates
that there are more factors affecting the reduction potentials of
the T1 copper center. These factors have been uncovered by mutagenesis
studies and engineering of copper proteins and are discussed in section 4.4.
4.3.4
T1
Copper Center in Multicopper Proteins
The T1 copper center
exists not only in single-domain proteins,
but also in multidomain proteins with multiple copper cofactors. These
proteins include multicopper oxidases and nitrite reductases (Table 9). The former
contain a T1 copper (blue copper),
a type II copper (abbreviated as T2), and a pair of type III copper
centers (Figure 53).
1262−1266
The latter contain T1 and T2 copper centers and are evolutionarily
related to the multicopper oxidases.
1265−1267
As shown in Figure 52, multicopper oxidases and
nitrite reductases are closely related and are composed of two, three,
or six domains.
1265
In multicopper oxidases,
the T1 copper center resides in the cupredoxin-like domain while the
T2 and T3 copper centers are located between domains.
Figure 52
Domain organization
and copper center distribution in multicopper
oxidases. Reprinted with permission from ref (1265). Copyright 2011 Wiley-VCH.
Figure 53
Active site of the multicopper oxidases.
Cu sites are shown as
green spheres. Figure generated from the crystal structure of ascorbate
oxidase (PDB ID 1AOZ). Reprinted from ref (1264). Copyright 2007 American Chemical
Society.
T1 copper centers in multicopper oxidases (MCOs)
are very similar
to those in single-domain T1 copper proteins. The copper ion is coordinated
by one Cys residue and two His residues at its equatorial positions.
In plant laccases, ascorbate oxidases, and nitrite reductases, axial
Met coordinates with copper and forms a trigonal pyramidal geometry.
In fungal laccase, ceruloplasmin, and Fet3p, the axial ligand is a
noncoordinating Leu or Phe, leaving equatorial ligands and copper
in a more trigonal geometry.
1262,1265,1266
One noticeable feature for T1 copper centers in MCOs is their high
reduction potential compared with that of single-domain T1 copper
proteins. Ceruloplasmin has the highest reduction potential
1148
(>1000 mV) reported in T1 centers, while
TvLac
has the second highest reduction potential
1141−1143
(778 mV). The high reduction potential is partially attributed to
the more hydrophobic axial ligand, while other factors such as hydrogen
bonding around the T1 Cu centers may contribute too.
1268
4.3.5
A Novel Red Copper Protein—Nitrosocyanin
Recently, a mononuclear red copper protein, nitrosocyanin from N. europaea, an ammonia
oxidizing bacterium, was
isolated and structurally characterized (Figure 54).
1137,1269−1271
The crystal structure shows that the copper ion is coordinated by
two His residues, one S(Cys), and a side chain O(Glu) and has an additional
fifth water ligand in the oxidized form, but not in the reduced form.
Nitrosocyanin shows a strong absorption band at 390 nm (ε =
7000 M–1 cm–1), a large hyperfine
splitting value (147 × 10–4 cm–1) in the EPR spectrum, and a very low reduction potential
of 85 mV
(compared with those of the T1 copper proteins, which are in the range
of 150–800 mV).
1137,1271
With an exogenous
water ligand, the reorganization energy of this protein is calculated
to be 2.2 eV, significantly higher than those of T1 copper proteins.
1271
Similar to T1 copper proteins, nitrosocyanin
has copper–thiolate coordination and strong UV–vis absorbance.
However, the water ligand in nitrosocyanin has not been observed in
T1 copper proteins before. Its copper site geometry and absorption
at ∼400 nm are also different from those of T1 copper proteins.
Its EPR spectrum, reorganization energy, and reduction potential more
closely resemble those of T2 copper proteins. Solomon and co-workers
attribute these properties to the relative orientation of the Cu-N-N-S
and Cu-S-Cβ planes, which in turn is due to “coupled
distortion” between the axial ligand and the whole copper center.
1095,1186,1271
Figure 54
Crystal structures of
(A) the oxidized red copper site in nitrosocyanin,
(B) the oxidized T1 copper site in plastocyanin, and (C) the reduced
red copper site in nitrosocyanin. Reprinted from ref (1271). Copyright 2005 American
Chemical Society.
The biological role
of this protein, however, has not yet been
identified. It has been proposed that it might be involved in ET or
serve some as-yet-unknown catalytic function due to the presence of
the open coordination site.
1269,1270
4.4
Structural Features Controlling the Redox
Chemistry of Type 1 Copper Proteins
Although the study of
native proteins provides valuable information about the structure,
spectroscopy, and function of T1 copper centers, it is difficult to
draw any conclusion only by comparing copper centers from different
scaffolds with low sequence homology. With the advancement of modern
molecular biology, powerful tools such as mutagenesis are available
to research groups, allowing the amino acid sequence to be modified
at will. Methods of unnatural amino acid mutagenesis have further
expanded the toolbox for bioinorganic chemists.
1272−1274
With these methods, not only amino acid residues directly coordinating
to copper, but also residues beyond the first coordination sphere
have been changed. Mutagenesis reveals how different components of
the protein contribute to the structure, spectroscopy, and function,
especially in reduction potential tuning.
4.4.1
Role
of Axial Met
The T1 copper
center has highly conserved equatorial ligands, two His residues and
one Cys residue. The axial position of the T1 copper center shows
more variation, as Met, Gln, and noncoordinating residues can all
be found in the native proteins. Mutagenesis of the axial ligand has
been carried out in azurin,
1122,1275−1278
nitrite reductase,
1235,1258,1279
amicyanin,
1261
rusticyanin,
1259
pseudoazurin,
1234
laccase,
1256
and stellacyanin.
1139,1280,1281
Mutation of the axial ligand
in different T1 copper proteins generally results in a protein that
retains copper-binding ability but with a different reduction potential
or altered spectroscopic properties. An early work replaced Met121
in azurin with all other 19 amino acids with minimal alteration of
the T1 character of the copper center.
1276
While changing the axial ligand to hydrophobic ligands such as Ala,
Val, Leu, or Ile increases the reduction potential by 40–160
mV,
1122
substitution with Glu or Gln decreases
the reduction potential by 100–260 mV.
1122,1257
As the axial ligand is changed from Gln to Met to more hydrophobic
residues, the reduction potential of the protein increases. Theoretical
studies have suggested that the axial ligand is involved in tuning
the potential.
1282,1283
To test the role of the axial
ligand in tuning the reduction potential of the T1 copper protein,
Lu and co-workers incorporated unnatural amino acid analogs of Met
with different hydrophobicities at the axial position in azurin.
1284,1285
The reduction potential varied from 222 to 449 mV at pH 4.0. Such
a replacement of Met with its iso-structural analogs allowed conclusive
identification of hydrophobicity of the axial ligand as the major
factor in tuning reduction potentials, because a linear correlation
was found between the reduction potential and hydrophobicity of the
axial ligand. Likewise, Dennison and co-worker mutated the axial Met
of cucumber basic protein to Gln and Val. As the axial ligand was
changed from Gln to Met to Val, the electron self-exchange rate increased
by 1 order of magnitude, and the reduction potential increased by
∼350 mV.
1286
These studies have
firmly established a correlation between hydrophobicity of the axial
ligand and reduction potential, providing a better understanding of
the role of the axial ligand in reduction potential tuning.
Within T1 copper proteins, there are two classes of proteins with
slightly different spectroscopic features. Typical T1 copper proteins,
such as plastocyanin and azurin, have absorption at ∼600 nm
and an axial EPR signal, whereas “perturbed” T1 copper
proteins or green copper proteins have an additional ∼400 nm
absorption peak in their UV–vis spectra, as well as rhombic
EPR signals. At the same time, the perturbed T1 copper proteins have
longer Cu–S(Cys) distances and shorter Cu–axial ligand
distances.
1283
A more extreme case comes
from the newly discovered protein nitrosocyanin, which has a cysteine
ligand and dominant ∼400 nm absorption in its UV–vis
spectrum, resulting in a red color.
1137,1271
Although
the strong absorption and 1Cys/2His/1Glu ligand set resembles those
of T1 copper proteins, nitrosocyanin has large hyperfine splittings
(A
|| ≈ 150 × 10–4cm–1) in its EPR spectrum and a low reduction potential
(85 mV), which falls into the range of T2 copper proteins.
1136,1137,1271
Solomon and co-workers proposed
coupled distortion theory on the basis of a suite of spectroscopic
studies in combination with theoretical calculations to explain the
variance in electronic absorption and concomitant color change from
blue to green to red in native proteins. This theory states that shorter
Cu–axial ligand distances result in distortion of the T1 copper
geometry toward tetragonal, which elongates the Cu–S(Cys) distance.
1283
This distortion renders the pσ(Cys)–Cu
CT more favorable than pπ(Cys)–Cu CT, which causes an
increase in the ∼400 nm absorption over the ∼600 nm
absorption in the UV–vis spectrum. Mutational studies on the
axial ligand of various T1 copper proteins have validated the coupled
distortion theory. By changing a weak Met to a stronger His
1277,1287,1288
or Glu ligand,
1289−1291
the blue copper protein azurin can be converted to a green copper
protein. By mutating Met to a weaker ligand such as Thr, the natively
green copper protein, nitrite reductase, has been converted to a blue
copper protein.
1292
Recently, the axial
Met was mutated to Cys, a strong ligand, and then to the unnatural
amino acid homocysteine (Hcy), a strong ligand with a longer side
chain. The resulting Met121Cys azurin has an additional ∼450
nm absorption, while in Met121Hcy the ∼410 nm peak dominates
over the ∼625 nm peak. Together with EPR evidence, it was shown
that, within the same scaffold, blue copper protein azurin was converted
to a green copper protein and then to a red copper protein.
1293
Interestingly, the engineered red copper protein,
Met121Hcy azurin, has a low reduction potential (113 mV) similar to
that of nitrosocyanin (85 mV).
4.4.2
Role
of His Ligands
Although equatorial
His residues are highly conserved in T1 copper proteins, their mutation
does not impair the copper binding ability of the protein. Canters
and co-workers mutated two His residues into Gly separately, and the
resulting protein still had T1 characteristics.
1294,1295
As His to Gly mutation creates extra space around copper, exogenous
ligands such as halides, azides, and imidazoles could diffuse into
His46Gly and His117Gly azurins and coordinate with copper. Depending
on the type of external ligand, the mutants will be either T1 or T2
copper proteins.
1294−1296
His117Gly and His46Gly mutations also changed
solvent exposure of the copper site. Without external ligands, His117Gly
azurin has a reduction potential of 670 mV, much higher than that
of WT azurin (310 mV). The high reduction potential is due to loss
of a water ligand during reduction. Addition of external ligands will
lower the reduction potential.
1297
The
open coordination site of His117Gly makes it possible to study ET
using imidazole-modified complexes.
1298,1299
The mutants
generally exhibit a lower ET rate. As the properties of exogenous
imidazoles affect the ET rate, it has been suggested that His is also
important in the WT protein.
1300
4.4.3
Role of Cys Ligands
As the Cu–S(Cys)
bond defines the properties of type I copper sites,
99
mutation of Cys to other natural amino acids will dramatically
alter the copper site in T1 copper proteins (Figure 55). Substitution of any other
amino acid for Cys will result
in loss of the intense LMCT bands, which is due to the interaction
of the Cys S with copper. As an isostructural analogue of Cys, selenocysteine
(SeC) can replace Cys without major structural perturbation. This
strategy has been employed as a spectroscopic probe for T1 copper
centers.
1301−1303
The protein Cys112SeC azurin showed a reduction
potential similar to that of WT azurin (328 mV vs 316 mV at pH 4)
and a red-shifted LMCT band at 677 nm.
1301
So far, only Cys112Asp mutation in azurin has been characterized.
Mutation of Cys to Asp makes azurin a T2 copper protein, as evidenced
by large hyperfine splitting (A
|| ≈
152 × 10–4 cm–1) in the EPR
spectrum and slow ET.
1304−1307
Addition of another mutation at the axial
position, Met121Leu (Phe/Ile), results in a novel copper center called
type 0 copper, which has the small parallel hyperfine splittings and
rapid ET characteristic of T1 copper centers but no longer fits the
classification of T1 copper due to the loss of the copper–thiolate
interaction.
1308−1311
Moreover, there is only a slight increase of the reorganization
energy to 0.9–1.1 eV compared with that of WT azurin, much
less than that of T2 copper proteins. The ET rate of type 0 copper
protein is 100-fold faster than that of the Cys112Asp mutant, a typical
T2 protein.
1308,1309,1311
Figure 55
Active sites of type 2, type 1, and the newly constructed type
0 copper. In the center, a plot shows (in the shaded ovals) the typical
values of two electron paramagnetic resonance spectroscopy parameters, A
∥ and g
∥, for type 1 (lower) and type 2 (upper) copper sites and the values
of type 0 copper (green, red, and black points, right center), showing
that type 0 copper does not fall into the typical ranges for these
other kinds of sites. Reprinted with permission from ref (1308). Copyright 2009 Macmillan
Publishers Ltd.
4.4.4
Role
of Structural Features in the Secondary
Coordination Sphere
Copper ligands exert a significant influence
on the spectroscopic features and reduction potentials of T1 copper
proteins. However, copper ligands cannot fully account for variation
in the reduction potentials of T1 copper proteins. Mutation of copper
ligands usually results in loss of T1 characteristics or reduction
of ET activity. For the limited mutations that maintain T1 characteristics
and ET activity, the reduction potential is tuned over a 227 mV range
by introducing Met analogues at the axial position, which is far less
than the 600 mV range reported in native proteins.
1285
As discussed in section 4.3.3, the
H-bonding network beyond the T1 copper center plays an important role
in maintaining the structure and function of T1 copper centers. Mutagenesis
studies focusing on changes of hydrogen bonds have revealed important
information about how the reduction potential and other properties
are tuned in T1 copper proteins.
Rusticyanin has a higher potential
relative to other T1 copper proteins. By sequence comparison, it is
identified that there is a Ser in rusticyanin at the position corresponding
to Asn that “zips” two ligand loops together. Asn has
been proposed to decrease the E
m by strengthening
the H-bonding interactions between two ligand-containing loops. Mutating
Ser86 in rusticyanin to Asn established such a hydrogen bond and lowered
the E
m by 77 mV.
1312
On the other hand, changing Asn in azurin to Ser eliminates
one hydrogen bond between two loops (Figure 56) and results in a protein with a 131
mV higher reduction potential.
1293
Figure 56
X-ray structures of Az and selected variants.
(a) Native azurin
(PDB ID 4AZU). (b) N47S/M121L azurin (PDB ID 3JT2). (c) N47S/F114N azurin (PDB ID
3JTB). (d) F114P/M121Q
azurin (PDB ID 3IN0). Copper is shown in green, carbon in cyan, nitrogen in blue,
oxygen
in red, and sulfur in yellow. Hydrogen-bonding interactions are shown
by dashed red lines. Reprinted with permission from ref (1088). Copyright 2009 Macmillan
Publishers Ltd.
By comparing certain
cupredoxins that natively have lower E
m than the rest of the family, it is observed
that they share a conserved Pro residue that is two residues after
the copper-ligating Cys.
114,1313
The backbone amide
in the equivalent residue in azurin hydrogen bonds to the thiolate
of Cys112.
1160
Placing a Pro in this position
converts this secondary amide to a tertiary amide, which is incapable
of donating a hydrogen bond. The Phe114Pro mutant has a lower reduction
potential.
114
It is proposed that deleting
the hydrogen bond to the thiolate gives Cys112 more conformational
freedom, and it allows for the electron density that was previously
tied up in a hydrogen bond to contribute to the Cu–SCys interaction.
114
Another examination
of cupredoxin crystal structures reveals the
presence of backbone carbonyl oxygen from Gly45 near the copper ion
in azurin, which is missing in other cupredoxins such as rusticyanin.
97,98,1314
This ionic interaction in azurin
is proposed to result in higher electron density near the copper,
preferentially stabilizing the Cu(II) form of the protein and, therefore,
lowering the E
m.
98,481,1315
Phe114Asn mutation was made
to hydrogen bond with Gly45 backbone carbonyl and decrease the effect
of carbonyl oxygen in Az. The mutant showed a 129 mV higher reduction
potential compared to that of the wild type.
1088
With all of these individual factors in mind, Lu
and co-workers
combined mutations on both the copper ligands and residues in the
secondary coordination sphere. These mutations showed an additive
effect on the reduction potential in azurin. With different combinations,
the reduction potential was tuned from 90 to 640 mV, which is beyond
the reported range of native T1 copper proteins and their mutants
(Figure 57).
1088
Figure 57
Reduction potentials for a number of Az mutants versus a measure
of the hydrophobicity (log P), revealing the linear
trend with respect to the axial position (residue 121). Reprinted
with permission from ref (1088). Copyright 2009 Macmillan Publishers Ltd.
Unlike mutations on the copper ligands, mutations
of residues in
the secondary coordination sphere are less likely to change the T1
characteristics according to UV–vis, EPR,
1293
and resonance Raman
1316
spectroscopy.
DFT studies were able to separate the effects of covalent interaction
and nonlocal electrostatic components; while the covalent and nonlocal
electrostatic contributions can be significant and additive for active
H-bonds, they can be additive or oppose one another for dipoles (Figure 58).
1317
Figure 58
Illustration of the
experimentally derived covalent and nonlocal
electrostatic contributions to E° for the variants
of Az relative to WT Az and their comparison to calculations. Reprinted
from ref (1316). Copyright
2012 American Chemical Society.
Lower reorganization energies in the ET process generally
increase
the ET rate constants and efficiency. However, rational design of
ET centers to lower the reorganization energy has so far not been
demonstrated. Such a task is particularly challenging for ET proteins
such as the blue copper protein azurin that have already been shown
to possess very low reorganization energies in comparison to the majority
of the other proteins. A study of intramolecular ET by pulse radiolytically
produced disulfide radicals to Cu(II) in the above rationally designed
azurin mutants showed that the reorganization energies of the designed
mutants are lower than that of WT azurin, increasing the intramolecular
ET rate constants almost 10-fold.
1317
More
interestingly, analysis of structural parameters of these mutants
suggested that this lowering in reorganization energy is correlated
with increased flexibility of the copper center.
4.4.5
Role of Ligand Loop
Besides directly
mutating individual ligands, loop-directed mutagenesis containing
the ligands to the copper center enables manipulation of copper center
by changing the protein structure on a broader scale. T1 copper proteins
and CuA domains in heme–copper oxidases share the
same cupredoxin fold, with three ligands of T1 copper and four ligands
of CuA residing in the so-called “ligand loop”
(Figure 59). By careful design, it is possible
to transplant the ligand loop of one protein into another, enabling
interconversion between T1 copper and CuA and between different
T1 copper proteins (section 4.5.3).
Figure 59
Ligand and
loop structure in different T1 copper proteins, CuA from T. thermophilus heme–copper
oxidase, and red copper protein nitrosocyanin: (A) amicyanin (PDB
ID 1AAC); (B)
pseudoazurin (PDB ID 1PAZ); (C) plastocyanin (PDB ID 1PLC); (D) azurin (PDB ID 2AZA);
(E) rusticyanin
(PDB ID 1RCY); (F) CuA from T. thermophilus heme–copper oxidase (PDB ID 1CUA); (G)
nitrosocyanin (PDB ID 1IBY).
An early example of loop-directed mutagenesis comes
from interconversion
between different copper centers, as two research groups independently
installed a ligand loop from the CuA domain of cytochrome c oxidases on amicyanin
and azurin, converting the T1 copper
proteins to a CuA protein,
1318,1319
discussed
in detail in section 4.5. Recently, Berry and
co-workers transplanted the ligand loop of nitrosocyanin, a newly
discovered red copper protein, to azurin.
1320
The resulting protein, NCAz, has UV–vis and EPR features
similar to those of nitrosocyanin despite having His instead of Glu
as the fourth ligand.
Although the T1 copper proteins have a
conserved ligand set (section 4.3.1.1), the
ligand loops from different proteins
show variation in length and sequence (Figure 59). Loop-directed mutagenesis has been
carried out between different
T1 copper proteins. Ligand loops from azurin, pseudoazurin, plastocyanin,
rusticyanin, and nitrite reductase were introduced into the amicyanin
scaffold to create loop elongation mutants.
1321−1324
Later, the ligand loop from amicyanin, which is the shortest among
T1 copper proteins, was introduced into azurin, pseudoazurin, and
plastocyanin scaffolds to create loop contraction mutants.
1325,1326
The ligand loop from plastocyanin was introduced into the azurin
scaffold as well.
1327
All of the loop-directed
mutants maintain T1 copper spectroscopic characteristics, indicating
a similar structure in the Cu(II) state. On the other hand, the loop
length has been shown to affect the pK
a of C-terminal His and the Cu(I)–N(His) distance.
1326,1327
It has been observed that introducing the short loop of amicyanin
into pseudoazurin and plastocyanin increases the pK
a of C-terminal His, probably due to an entropically favored
Cu(I)–N(His) interaction with a longer, more flexible loop.
1324−1326
As expected, the reduction potentials of loop-directed mutants
are between the reduction potentials of donors of the loops and scaffolds.
Amicyanin has the second lowest reduction potential in T1 copper proteins
(see Table 10). Introducing the amicyanin loop
into other copper protein scaffolds decreases their reduction potentials
by 30–60 mV.
1326
On the other hand,
introducing loops of other T1 copper proteins into amicyanin increases
its reduction potential.
1322−1324
The ET activity of loop-directed
mutants has been measured by the
electron self-exchange rate constant (k
SES). The loop elongation mutants generally have 10-fold lower k
SES values, while loop contraction has less
influence on k
SES.
1322,1323,1326
All the studies indicate that,
T1 copper proteins can accommodate changes in loops and assume the
same active site structure, consistent with the “rack”
or entatic state of the T1 copper center.
95,1168,1170
4.5
CuA Centers
4.5.1
Overview of the CuA Centers
The CuA is a binuclear copper
center bridged by two
cysteine ligands to form a Cu2S2 “diamond-core”
structure, which has been found naturally in CcOs,
1030,1109,1328
nitrous oxide reductases (N2ORs),
1329,1330
the oxidase from Sl. acidocaldarius (SoxH),
1331
and a nitric oxide reductase (qCuANOR)
1332,1333
to date (Figure 60). Interestingly, all of
these proteins are terminal electron acceptors of ET processes; e.g.,
CcO is the terminal electron acceptor in aerobic
respiration, and N2OR is the terminal electron acceptor
in anaerobic respiration. One of the most important features of the
CuA center is that the two copper ions form a direct metal–metal
bond. Therefore, the unpaired electron is delocalized between two
copper ions, and the resting state of the CuA center is
a Cu(+1.5)–Cu(+1.5) state rather than a Cu(+2)–Cu(+1)
state. This is the first example of a metal–metal bond found
in biology, which makes it unique compared to centers of other metalloproteins.
In addition to the bridging Cys ligands, the copper ions are coordinated
by a His from the equatorial position to form a trigonal NS2 coordination. There is
a weak distal axial ligand on each copper
ion. The axial ligands are a methionine at one copper and a backbone
carbonyl at the other. Considering only each copper ion, the CuA center is very similar
to the T1 blue copper center with
an overall distorted tetrahedral geometry. Hence, the CuA center can be treated as
two T1 copper centers joined together with
a Cu–Cu bond in between, suggesting an evolutionary relationship
between these two centers. Indeed, such a relationship has been proposed
on the basis of three-dimensional structure comparison and construction
of phylogenetic trees, indicating that T1 copper and CuA proteins share a common ancestor
and are developed in part by divergent
evolution.
1334,1335
Figure 60
Crystal structures of
cytochrome c oxidase (PDB
ID 3HB3) and
nitrous oxide reductase (PDB ID 1FWX). The CuA sites are highlighted
(copper is in green, sulfur is in yellow, nitrogen is in blue, and
carbon is in cyan).
The UV–vis absorption
spectrum of CuA shows two
intense absorbance bands at ∼480 and 530 nm that arise from
S(Cys) → Cu charge transfer in the visible region and also
a broad band at ∼760–800 nm that arises from Cu(+1.5)–Cu(+1.5)
intervalence charge transfer.
860,1113−1115
The reduced Cu(I)–Cu(I) form is colorless because of the
d
10
electronic configuration at each copper
center. The more oxidized Cu(II)–Cu(II) state has not been
observed in proteins to date.
1336,1337
Attempts to oxidize
the CuA site normally give an irreversible anodic current
at around 1 V, probably due to oxidation of the bridging dithiolate
to disulfide.
1337,1338
Therefore, the CuA site acts as a one-ET center under physiological condition.
72
The Cu–Cu bond in CuA sites has been the subject
of extensive debate.
1353
Later, the structure
of the CuA site was confirmed by different spectroscopic
methods. Blackburn et al. reported the extended EXAFS studies of the
CuA binding domain of B. subtilis CcO, which showed a strong Cu–Cu interaction
of ∼2.5 Å together with a short 2.2 Å Cu–S
interaction.
1108
The Cu–Cu bond
distance is nearly identical to the results from EXAFS studies of
native CcO from bovine heart mitochondria, which
is 2.46 Å.
1354
The dinuclear nature
and the unusually short Cu–Cu distance of ∼2.55 Å
were further established by X-ray crystal structures of CcO from Pa. denitrificans
and bovine
heart mitochondria,
1030,1109
as well as an engineered CuA center in CyoA.
1349
Similar structural
features were also observed in the crystal structure of N2OR from Ps. nautica.
1329,1330
The most intense bands at 339, 260, and 138 cm–1 observed in resonance Raman spectroscopy
of the Pa.
denitrificans CcO CuA domain
were assigned to symmetric stretches involving primarily the Cu–S(Cys),
Cu–N(His), and Cu–Cu bonds, respectively.
1107
The Cu–Cu bond in the CuA site causes a valence
delocalization between the two copper ions and produces a seven-line
hyperfine splitting pattern in the EPR spectra. This unique EPR pattern
can be explained by the delocalized unpaired electron coupled with
two equivalent copper ions with a nuclear spin I =
3/2.
1112,1355,1356
Compared
to centers in T1 blue copper proteins, CuA centers show
even smaller A
∥ on the basis of
EPR simulations,
1114,1339,1342,1345,1346,1357
reflecting greater covalent
interaction and unpaired electron delocalization between the copper
ions and the bridging Cys residues.
4.5.2
CuA Centers in Water-Soluble
Domains Truncated from Native Proteins
Historically, studying
the biochemical role and probing the unique structure of CuA centers have not been
easy due to complications arising by overlapping
spectroscopic features of other metal centers present in the native
proteins containing the CuA center. For instance, the CcO is a membrane protein containing
two heme groups (heme a and heme a
3), two copper centers
(CuA and CuB), a zinc ion, and a magnesium ion.
To overcome these inherent difficulties in studying native CuA centers, two strategies
are developed: producing truncates
of native proteins containing CuA sites
742,1331,1339,1341,1342,1345,1352,1358−1361
and designing CuA centers into small, soluble proteins.
1318,1362,1363
In the first strategy,
the sequence of the CuA subunit from CcO or SoxH was isolated and recombinantly expressed
without the helices
that normally anchor this domain to the membrane. This way, a water-soluble
protein containing only the CuA site was obtained. Such
truncates have been constructed for CcO from B. subtilis,
1345
Pa. dentrificans,
742,1339,1358,1361
Procambarus
versutus,
1360
Synechocystis PCC 6803,
1352
and T. thermophilus(1341,1342,1359,1361) and for SoxH from Sl. acidocaldarius.
1331
The UV–vis, EPR, and EXAFS
spectroscopic characterizations as well as the reduction potentials
measurments for these soluble truncates are consistent with each other
(Table 13).
742,1339,1358,1361
To date, only the
truncate from T. thermophilus has been
successfully crystallized.
1359
Table 13
Summary of the Spectroscopic Parameters
of CuA Sites in Different Proteins
CuA site containing
protein
organism
λmax (nm) (extinction coefficient, M–1cm–1)
reduction
potential vs SHE (mV)
ERP params (g
x
, g
y
, g
z
)
Cu–Cu distance (Å)
ref
subunit
II of cytochrome c oxidase
Paracoccus
denitrificans
363 (1200), 480 (3000),
530, 808 (1600) (pH 7)
240
g
x
= g
y
= 2.03, g
z
= 2.18, A
z
= 3.5 mT
2.6
(742, 1109, 1339)
subunit II of cytochrome ba
3
Thermus thermophilus
363 (1300), 480 (3100),
530 (3200), 790 (1900)
250 (pH 8.1), 240 (pH 8), 297 (pH 4.6)
g
x
= 1.99, g
y
= 2.00, g
z
= 2.17, A
z
= 3.1 mT
2.43
(1337, 1341−1344)
subunit II of caa
3-type
cytochrome c oxidase
Bacillus
subtilis
365, 480, 530, 775–800
g
x
= g
y
= 1.99–2.03, g
z
= 2.178, A
z
= 3.82 mT
2.44
(1344, 1345)
nitrous oxide reductase
Paracoccus
dentrificans
480, 540 (1700), 800
(1330)
nitrous oxide reductase
Pseudomonas
stutzeri
480, 540
g
x
= g
y
= 2.03, g
z
= 2.18, A
z
= 3.83 mT
2.44
(1346)
nitrous oxide reductase
Achromobacter
cycloclastes
350, 481 (5200), 534 (5300),
780 (2900)
g
x
= g
y
= 2.045
(1347)
biosynthetic model
in CyoA
protein
Escherichia
coli
360, 538 (2000)
g
x
= 2.03, g
y
= 2.03, g
z
= 2.18, A
z
= 6.8, 5.3 mT
2.48
(1114, 1348, 1349)
biosynthetic model
in amicyanin
360, 483, 532, 790
g
x
= g
y
= 1.99–2.02, g
z
= 2.18, A
z
= 3.24 mT
(1318)
biosynthetic model
in azurin
360 (550), 485 (3730), 530
(3370), 770 (1640)
g
x
= g
y
= 2.06, g
z
= 2.17, A
z
= 5.5 mT
2.39
(1319, 1350)
nitrous oxide reductase
Pseudomonas
nautica 617
480, 540, 800
260
g
x
= g
y
= 2.021, g
z
= 2.178, A
z
= 7 mT
(1351)
subunit II of SoxM
Sulfolobus
acidocaldrius
361 (2300), 478 (3200),
538 (3700), 789 (2400)
237
g
x
= g
y
= 2.01, g
z
= 2.20
(1331)
subunit II of cytochrome c oxidase
Synechocystis PCC
6803
359 (1580),
482 (2820),
535 (3080), 785 (1840)
216 (pH 7)
(1352)
4.5.3
Engineered CuA Centers
The second strategy
to study CuA sites is designing this
site into other proteins, first accomplished in a quinol oxidase.
1362
The authors aligned subunit II of cytochrome c and quinol oxidases and found that
the C-terminal of both
proteins contained a subdomain in a Greek key β-barrel scaffold.
This alignment suggested that both proteins contain a basic structural
motif characteristic of cupredoxins. The CyoA lacked the putative
ligands for the formation of the CuA in CcO. The CuA ligand set was thus introduced
by extensive
mutagenesis of the isolated cupredoxin domain. This engineered CyoA
bound copper and showed two strong peaks at 358 and 536 nm, a shoulder
at 475 nm, and a broad peak between 750 and 780 nm, as well as an
EPR pattern similar to that observed in native CuA from
CcO. Later, the crystal structure of CyoA was reported
with 2.3 Å resolution.
1349
The distance
between the two coppers is 2.5 Å. Shortly after the release of
the purple CyoA study, two other research groups independently developed
designed CuA centers in T1 copper proteins.
1318,1319
Dennison et al. replaced the C-terminal loop of the blue copper
protein amicyanin, which contained three of the four T1 Cu-binding
ligands, with a CuA binding loop. After copper binding,
a purple protein was produced with UV–vis absorbance at 360,
483, and 532 nm and a broad absorption at approximately 790 nm, almost
identical to that of the native CuA domain of CcO from B. subtilis.
1318
The EPR spectrum of the CuA amicyanin
contained signals from two Cu(II) species; a distinctive T2 copper
site, and a CuA center.
1364
Hay et al. constructed a CuA protein from a recombinant
T1 copper protein, Ps. aeruginosa azurin,
by replacing the loop containing the three ligands to the blue copper
center with the corresponding loop of the CuA site in CcO from Pa. dentrificans.
1319
The UV–vis and EPR spectra of this
protein (CuAAz) were remarkably similar to those of native
CuA sites in CcO from Pa.
dentrificans (Figure 61). The
UV–vis absorption spectrum of CuAAz features two
S(Cys) → Cu CT bands at 485 (ε ≈ 3700 M–1 cm–1) and 530 nm (ε ≈ 3400 M–1 cm–1),
1113,1350
compared to 480–485 and 530–540 nm for native CuA centers.
98
CuAAz also
showed a broad band centered at 760–800 nm (ε ≈
2000 M–1 cm–1), typical of the
Cu–Cu ψ → ψ* transition, suggesting that
CuAAz had reproduced the Cu–Cu bond. Additionally,
the EPR spectrum of CuAAz displayed a seven-line hyperfine
splitting pattern, demonstrating that this biosynthetic model duplicated
the mixed-valence ground state of native CuA centers.
1319,1350
EXAFS, CD, MCD, and resonance Raman analyses of the CuAAz also suggested a high level
of electronic and structural identity
with CuA centers from CcO.
1113,1319,1350,1364,1366
The X-ray crystal structure
of CuAAz showed a very similar arrangement of ligands around
the copper ions and a Cu–Cu distance that was even slightly
shorter than the native CuA center in CcO, confirming the presence of a Cu–Cu bond.
1367
Figure 61
(A) Crystal structure of the biosynthetic model of the
CuA site in azurin (PDB ID 1CC3). (B) Comparison of UV–vis spectra
between
the soluble CuA domain in cytochrome c oxidase (green line), wild-type azurin (blue
line), and the biosynthetic
CuA model in azurin (purple line). (C) Comparison of X-band CW EPR
between wild-type azurin (blue line) and the biosynthetic CuA model
in azurin (purple line), four-line splitting vs seven-line splitting.
Reprinted with permission from ref (1365). Copyright 2010 Springer-Verlag.
4.5.4
Mutations of the Axial
Met
The
weaker axial methionine ligand has been investigated through mutagenesis
in CcO from Pa. denitrificans and Rb. sphaeroides. The Met227Ile
mutation in CcO from Pa. denitrificans resulted in a protein with unchanged stoichiometry
of the metals.
However, the two copper ions in the CuA site were no longer
equivalent and converted from a delocalized Cu(+1.5)–Cu(+1.5)
system to a localized Cu(+1)–Cu(+2) system on the basis of
EPR and near-IR studies.
1368
The ET from
cytochrome c to CuA was not affected,
but the rate of ET to heme a was significantly diminished
in the mutant protein compared with the wild-type protein due to an
altered reduction potential of the CuA site. It was concluded
that the weak axial Met was not essential for copper binding, but
it was important for maintaining the mixed-valence electronic structure
of the CuA site. The Met263Leu mutation in CcO from Rb. sphaeroides also showed
the binding of two copper ions and proton pumping activity. Multifrequency
EPR studies showed that the two copper ions in the CuA site
were still electronically coupled. While all the other metals remained
unchanged on the basis of UV–vis, EPR, and FTIR spectroscopy,
the mutant only maintained 10% of the activity
1369
of the native enzyme. The kinetic analysis of ET showed
a decrease of ET rate from heme c to CuA to 16 000 s–1 in the mutant, compared to
40 000 s–1 in the wild type. The rate constant
for the reverse reaction was increased to 66 000 s–1, compared to 17 000 s–1 in the
wild type.
This result was attributed to an increased reduction potential of
120 mV relative to that of the native enzyme.
1370
The perturbation of the weak axial methionine ligand
was also tested in the soluble CuA-containing subunit of
cytochrome ba
3 from T.
thermophilus.
1357
The mutants,
Met160Gln and Met160Glu, affected the g
z
region of the EPR spectra and the Cu hyperfine became
more resolved and larger in both mutants. Notably, the A
z
values of both mutants were increased
from 3.1 to 4.2 mT, larger than most of characterized native CuA sites. The UV–vis
spectra showed enhanced intensity
and a blue shift relative to that of the wild type. The EPR and UV–vis
data suggested that the axial ligand to copper interaction became
stronger, moving from WT to Met160Gln and then to Met160Glu. The effects
of both mutations were further studied by pulsed EPR/ENDOR spectroscopy.
1371
The results from this study showed an increase
of A
∥, larger hyperfine coupling,
and reduction in the isotropic hyperfine interaction and the axial g tensor. All these
effects were associated with an increase
in the Cu–Cu distance and changes in the geometry of the Cu2S2 core structure. The
mutant Met160Gln was also
studied by paramagnetic 1H NMR spectroscopy.
1372
The fast nuclear relaxation in this mutant
suggested that a low-lying excited state had shifted to higher energies
compared to that of the wild-type protein.
Blackburn et al.
reported a selenomethionine-substituted T. thermophilus cytochrome ba
3 and characterized it
with Cu K-edge EXAFS.
1373
Interestingly,
the optical and EPR spectra of the selenomethionine-substituted
CuA site were essentially identical to those of the native
CuA site as was the reduction potential. These data suggested
that whatever role the S(Met) atom played in the electronic structure
of the CuA site was also carried out by the Se(Met) atom.
The axial Met in CuAAz was mutated to Asp, Glu, and
Leu, covering the entire range of the hydrophobicity among the natural
amino acids. The measured reduction potentials for these axial Met
variants showed very little change, only about 20 mV, from that of
the original CuAAz, despite some visible perturbation to
the UV–vis and EPR spectra of these mutants. The significantly
smaller effect of axial ligand in tuning reduction potential of CuAAz compared with
WT-Az may reflect the resilience of the diamond
core of CuA. In other words, the stability of the interactions
making up the diamond core—the bridging Cys thiolates and copper–copper
bond—may lead to greater resistance to perturbations arising
from the axial position.
1374
Recently,
a different set of axial Met mutants was generated in the truncated
water-soluble CuA domain from T. thermophilus.
1375
By introducing Gln, His, Ser, Tyr,
and Leu at the axial Met position, a change of about 200 mV in reduction
potential was observed. The difference between the results from the
truncated CuA domain and CuAAz was attributed
to the difference in Cu–S(Met) bond lengths in these two systems:
2.47 Å in the truncated CuA domain vs 3.07 Å
in CuAAz. Another explanation is that CuAAz
contains the shortest Cu–Cu bond length (∼2.4 Å),
hence enhances resistance of the diamond-core structure toward ligand
changes.
It is interesting to note that the reduction potentials
of the
native CuA site from the soluble fragment of subunit II
of T. thermophilus
ba
3 at different pH values showed no significant changes.
1376
However, the engineered CuA site
in azurin exhibited strong pH dependence of the redox properties.
This difference might be caused by protonation and dissociation of
one of the histidine ligands in the engineered CuA center.
4.5.5
Mutations of the Equatorial His Ligands
The equatorial His ligands bind to the copper ions with a bond
length of ∼2.0 Å. In principle, mutations at this His
position would result in a significant perturbation of the CuA site. This assumption
has been proven to be true in the native
system. The His260Asn mutant in cytochrome c oxidase
from Rb. sphaeroides only exhibited
1% of the wild-type activity.
1369
The 850
nm band was shifted, and the extinction coefficient was diminished
to around 1230 M–1 cm–1, compared
with 1900 M–1 cm–1 in the wild
type. No apparent hyperfine splitting pattern was observed in the
EPR spectrum. The kinetic analysis of ET rates showed that the rate
constant for ET from CuA to heme c was
decreased to 11 000 s–1, compared to 40 000
s–1 in the wild type. The ET rate from CuA to heme c was decreased to 45 s–1, compared
with 90 000 s–1 in the wild type.
An increase of 90 mV in the reduction potential was also observed.
1370
However, dramatic differences were observed
in the biosynthetic model of CuA in azurin. The mutation
of His120 to Ala yielded a UV–vis spectrum similar to that
of the original CuAAz, including the Cu–Cu ψ
→ ψ* band at ∼760 nm.
1377,1378
The EPR spectrum of His120Ala only showed a four-line hyperfine
splitting pattern, suggesting that the active site had undergone a
transformation to trapped valence, although a Q-band ENDOR study of
His120Ala CuAAz showed evidence for the CuA site
still being delocalized.
1379
Xie et al.
applied a series of spectroscopic techniques, including EPR, UV–vis,
MCD, resonance Raman, and XAS to both CuAAz and His120Ala
CuAAz and correlated the results with DFT calculations.
1380
The surprising conclusion of this work was
that a minute, 1% mixing of the 4s orbital of one copper ion into
the ground-state spin wave function caused the collapse to a four-line
hyperfine splitting pattern in the EPR spectrum of His120Ala, not
a change from valence-delocalized to trapped valence. The resonance
Raman and MCD spectra both demonstrated that the valence delocalization
of the CuA center was still intact, although slightly perturbed,
despite the loss of His120 as a ligand. The authors attributed the
ability of CuA in azurin to remain valence-delocalized,
even with the loss of such a strong ligand, to the large electronic
coupling matrix element, which arises from the strong and direct Cu–Cu
bond. Thus, the diamond core of CuA plays an immense role
in the robust nature of this center.
4.5.6
Mutations
of the Bridging Cys Ligands
Mutagenesis studies of the CuA binding ligands in native
CcO from Pa. denitrificans and N2OR from Ps. stutzeri have demonstrated that the cysteine
ligands play an important role
in the functions of the enzymes and the spectroscopic features of
CuA. Mutating one of the two bridging cysteines to serine,
Cys216Ser, in CcO from Pa. denitrificans resulted in a type 1 blue copper site with
four-line EPR hyperfine
splitting rather than the seven-line EPR signal observed in the CuA site, and only
retained below 1% of the wild-type activity.
The Cys216Ser mutant no longer exhibited the near-IR absorption in
the optical spectrum, indicating the loss of the Cu–Cu bond.
Mutation of the second cysteine, Cys220Ser, resulted in 5–10%
of the wild-type activity. The higher activity in Cys220Ser is suggested
to be due to the intact binuclear copper site on the basis of the
metal/protein ratio and copper/iron ratio.
1381
The Cys618Asp mutant in N2OR resulted in almost complete
loss of activity. The copper was bound only weakly and was hardly
detectable on the gel filtration column. In contrast to the Cys618Asp
mutant, the Cys622Asp mutant retained some copper binding ability
and activity, although the characteristic multiline feature of the
mixed-valence CuA was no longer resolved in EPR.
1382
Similar to the studies in the native
system, the bridging Cys ligands were also individually mutated to
Ser in the biosynthetic model of CuA in azurin.
1383
Although the resulting mutants still bound
to the copper ions, the features of the Cu–Cu bond were completely
lost in that the Cys112Ser mutant resulting in two T2 copper sites.
The Cys116Ser mutation resulted in a T1 copper site. To account for
the loss of symmetry in a single Cys to Ser mutant, a double Cys to
Ser construct was made.
1384
At high pH,
the double mutant indeed bound two coppers, but the EPR spectrum showed
that the two copper ions were in two distinct T2 copper sites rather
than a mixed-valence site with seven-line hyperfine splitting.
4.5.7
Tuning the CuA Center through
Noncovalent Interactions
The H-bonding and hydrophobic interactions
around the active site of copper proteins can significantly tune the
ET process.
1088
Two mutations, Asn47Ser
and Glu114Pro, were made in CuAAz.
1385
Both the Asn47Ser and Phe114Pro mutations alter H-bonding
interactions near the Cys112 ligated to a copper ion, but the Phe114Pro
mutation decreases the reduction potential by deleting the hydrogen
bond between Cys112 and the backbone NH group,
114
while the Asn47Ser mutation increases the reduction potential
by affecting the rigidity of the copper binding site and most likely
forming a direct hydrogen bonds between the protein backbone and Cys112
(Figure 62).
1088
Interestingly, by placing both CuA and T1 blue copper
centers in the same scaffold of azurin, Lu and coworkers were able
to demonstrate that the same mutations in the secondary coordination
sphere resulted in similar decease or increase of the reduction potentials
of the copper centers, but the magnitude of the effect is much smaller
in CuA center, probably because its “diamond core”
structure is more resistant to the perturbation (Figure 62).
1088
Figure 62
Tuning the reduction
potential at blue copper azurin and CuA azurin by redesigning
the second coordination sphere. The
effects of these mutants are in the same direction, but the magnitude
is smaller in the CuA site due to the electron delocalization
between the two copper ions. Adapted with permission from ref (1385). Copyright 2012
The
Royal Society of Chemistry.
4.5.8
Electron Transfer Properties of the CuA Center
The CuA site is the point of entry
of the electrons from cytochrome c. In CcO, the CuA receives electrons from cytochrome
c and transfers them to cytochrome a. However,
in N2OR, the CuA is believed to transfer electrons
between cytochrome c and the catalytic site where
nitrous oxide is reduced. The characterization of the ET between cytochrome c and
cytochrome c oxidase has been a difficult
problem. The stopped-flow method has been used to study the kinetics
of electron transfer but does not have sufficient time resolution
to monitor such a rapid ET process.
The electron transfers between
bovine cytochrome c oxidase and horse cytochrome c labeled with (dicarboxybipyridine)bis(bipyridine)ruthenium(II)
were studied by laser flash photolysis.
1386
The electron was transferred from Lys25 ruthenium-labeled cytochrome c to the CuA
site with a rate constant of 11 000
s–1. The CuA site then transferred an
electron to cytochrome a with a rate constant of
23 000 s–1. Lys7, Lys39, Lys55, and Lys60
ruthenium-labeled derivatives showed nearly the same kinetics.
The ET between the CuA site and heme a in bovine cytochrome c oxidase was measured
by
pulse radiolysis.
1387
The rate constant
of ET was 13 000 s–1 from the CuA site to heme a, and 3700 s–1 for the reverse process.
From this study a low activation barrier
was observed, suggesting a small reorganization energy during the
ET process. The method was also applied to study the electron transfer
between the CuA site and heme a in cytochrome c oxidase from Pa. denitrificans.
1340
The ET rates were found to be 20 400
and 10 030 s–1 for the forward and reverse
reactions, respectively.
The T1 blue copper sites and CuA sites are commonly
used as ET centers found in many biological systems. However, direct
comparison between the ET rates of these two centers is difficult
to achieve due to different protein scaffolds and redox partners.
The engineered CuA site in azurin provides a great opportunity
to eliminate the protein structure contribution to the ET process
since the ET rates are measured in the same azurin scaffold.
1388
The authors first radiolytically reduced the
disulfide bond within the azurin scaffold and then measured the long-range
ET rate from the reduced disulfide bond to the oxidized CuA center. The rate constant
of this intramolecular ET process in CuAAz is ∼650 s–1. Although CuAAz has a smaller
driving force (0.69 eV for CuAAz vs 0.76
eV for blue copper azurin), the ET rate of CuAAz is almost
3-fold faster than for the same process in the WT-Az (∼250
s–1). The calculated reorganization energy of the
CuA center is only ∼0.4 eV, which is 50% of that
found for the blue copper azurin. The low reorganization energy of
CuA was also observed in the truncated soluble CuA domain of CcO from T. thermophilus.
1337
Farver et al. studied the ET rates
and reorganization energies of the mixed-valence CuAAz
site and trapped-valence His120Ala CuAAz.
1389
They found that changing from the mixed-valence
to the trapped-valence state increased the reorganization energy by
0.18 eV, but lowering the pH from 8.0 to 4.0 resulted in a ∼0.4
eV decrease in the reorganization energy, suggesting that the mixed-valence
state only played a secondary role in controlling the ET property.
4.5.9
pH-Dependent Effects
As an electron
entry site for cytochrome c oxidase, the CuA center receives electrons from cytochrome
c and
transfers the electrons to the heme a site. The electrons
are finally transferred to the heme a
3–CuB site where dioxygen reduction takes place.
The reduction results in a proton gradient, which in turns drives
the synthesis of ATP. For the CcO to function well,
a regulator is needed for initiating and shutting down the whole ET
process and dioxygen reduction reaction. A pH-dependent study on engineered
CuAAz suggested that the CuA site may play such
a role.
1390
CuAAz displayed
a seven-line EPR hyperfine with a mixed-valence state. When the pH
was decreased from 7.0 to 4.0, the absorption at 760 nm shifted to
810 nm; at the same time, a four-line EPR hyperfine was observed.
The pH dependence was reversible, and the mixed-valence state was
restored when the pH was increased back to 7.0. A dramatic increase
in reduction potential was also observed from 160 to 340 mV when the
pH was decreased from 7.0 to 4.0. It was identified that the protonation
of C-terminal His120 caused such a pH-dependent transition, as the
His120Ala mutation completely abolished this observation. A feedback
mechanism was proposed to explain how the CuA site regulated
the function of cytochrome c oxidase. The pumped
proton may result in protonation of the C-terminal His and cause a
different valence state of the CuA site. The increased
reduction potential in the new state will stop the whole ET process
and proton pumping (Figure 63). This hypothesis
is further supported by ET studies in the His260Asn mutant in cytochrome c oxidase
from Rb. sphaeroides which showed that protonation of the C-terminal histidine resulted
in a change in the valence state and an increase of the reduction
potential by 90 mV.
1370
The ET rate from
the CuA site to heme a decreased by over
4 orders of magnitude. The His260 in cytochrome c oxidase corresponds to His120 in
CuAAz.
Figure 63
Schematic model of different
states of the CuA center
in cytochrome c oxidase: (A) mixed-valence form at
neutral pH and (B) trapped-valence form at low pH. Subunit I is in
light blue, and subunit II is in pink. Black arrows represent the
flow of electrons, and orange arrows represent the flow of protons.
Reprinted with permission from ref (1390). Copyright 2004 National Academy of Sciences.
4.5.10
Copper
Incorporation into the CuA Center
While numerous
studies have established the structural
features of CuA, the question of how copper ions are delivered
into the CuA sites in vivo is still poorly understood.
In the cytoplasm, copper levels are rigorously regulated, and free
copper levels are extremely low and estimated to be at the attomolar
level.
1391−1397
Although it has been proposed that a metallochaperone called Sco
is responsible for metalation of the CuA site, delivering
the copper ions to the CuA site in CcO
by Sco proteins has not been demonstrated.
1398
Besides the delivery of copper ions by Sco proteins, another
possibility is unmediated metalation. The CcOs from
eukaryotes are located in mitochondrial membranes.
1399
In Gram-negative bacteria, CuA in CcO is exposed to the periplasmic space. However,
in Gram-positive
bacteria, CuA in CcO is exposed to the
extracellular space.
1109,1393,1400,1401
In addition, the N2OR is a soluble protein also located in the periplasmic space.
1402
In periplasmic and extracellular spaces, copper
levels are not regulated as rigorously as inside the cell, and the
free copper ion concentration could be much higher. In fact, unmediated
CuA metalation has been considered as a possibility for
CuA metalation in N2OR.
1403−1405
From this view, the studies of free copper ion incorporation into
CuA sites in vitro may provide important insights into
this process, although they do not perfectly reflect the process in
vivo.
In an early study of CuAAz, the metalation
of apo-CuAAz by adding a 10-fold excess of CuSO4 was studied
by stopped-flow UV–vis spectroscopy.
1406
A single intermediate with intense absorbance at 385 nm was observed,
which is characteristic of the Cys S → Cu CT bands of tetragonal
T2 copper centers.
98,1095
This T2 copper intermediate
formed with k
obsd = 1.2 × 103 s–1 and subsequently decayed with k
obs = 3.1 s–1; meanwhile the
absorptions corresponding to the CuA site increased. An
isosbestic point between the ∼385 nm band and the ∼485
nm band of the CuA site was observed, indicating the T2
copper intermediate was converted to CuA. Because only
Cu(II) ion was added during metalation, a reducing agent must be supplied
by the system itself to form a Cu(+1.5)–Cu(+1.5) site, indicating
that the free thiols in apo-CuAAz were providing electrons
by forming disulfide bonds.
1407−1409
Adding ascorbate or Cu(I) salt
increased the yield of CuA center formation.
Figure 64
Proposed
mechanism of copper incorporation into the biosynthetic
CuA model in azurin. Reprinted with permission from ref (103). Copyright 2012 Elsevier.
A similar study was investigated
in N2OR from Pa. denitrificans.
1410
Different from the previous study,
two intermediates were observed
upon adding Cu(II) salt. These two intermediates formed within a similar
time scale and also decayed at the same time with simultaneous formation
of CuA sites. Two isosbestic points were present between
the absorption bands of both intermediates and the CuA absorption
bands, strongly suggesting conversion of these intermediates to CuA. One of these
two intermediates has spectral features typical
of T2 copper centers with thiolate ligation, and another shows the
characteristics of a T1 copper center. These observations suggested
that the purple CuA site contained the essential elements
of T1 and T2 copper centers and provided experimental evidence for
a previously proposed evolutionary link between the cupredoxin proteins.
1334,1335
Guided by the observation of both T1 copper and T2 copper
intermediates
in the metalation of the CuA site in N2OR, the
metalation of CuAAz was revisited by varying both the copper
concentration and pH.
1411
When the CuAAz concentration was greater than the CuSO4 concentration,
both T2 copper and T1 copper intermediates were observed, similar
to the results obtained for N2OR. Global fitting of the
UV–vis absorption kinetic data and time-dependent EPR together
with previously studied mutants of CuAAz provided valuable
information about the mechanism of copper incorporation where a new
intermediate, I
x
, was observed. When Cys112
was mutated to Ser, a T2 copper site formed, with UV–vis and
EPR spectra similar to those of the T2 copper intermediate. From this
study it was inferred that the T2 copper intermediate is a capture
complex with Cys116, which is also supported by the greater solution
accessibility of this residue, compared to Cys112. Conversely, when
Cys116 was changed to Ser, a T1 copper center formed, with UV–vis
and EPR spectra nearly identical to those of the T1 copper intermediate
(Figure 64).
1383
4.5.11
Synthetic Models of the CuA Center
Another approach to study the CuA center is to synthesize
small-molecule mimics of CuA.
1412
This has been proven to be a difficult task because of the formation
of disulfide bonds between free thiols mediated by copper ions.
1338
Also, the most important feature in the CuA site, the diamond-core structure with
Cu–Cu bond bridging
by thiolates, is difficult to achieve. Besides the first coordination
sphere, the second coordination sphere has also proven to be important
in tuning the properties of the CuA site, which is even
harder to mimic in small-molecule compounds.
1385
However, model compounds have met with varying degrees
of success.
369,1413−1428
Houser et al. reported a fully delocalized mixed-valence dicopper
complex with bis(thiolate) bridging which was the first closet small-molecule
CuA mimic. The crystal structure of this model complex
showed that the Cu2S2 core is planar with an
average Cu–Cu distance of 2.92 Å. However, it is still
longer than the Cu–Cu distance (2.46 Å by EXAFS
1354
and 2.55 Å by X-ray crystal structures
1030,1109
) in native CuA centers.
1416
The EPR spectrum recorded at 4.2 K clearly showed the seven-line
hyperfine splitting indicating the fully delocalized electronic structure.
More recently, Gennari et al. reported a new bis(μ-thiolato)dicopper
complex that mimicked most of the important spectroscopic features
of the CuA site.
1429
Notably,
this dicopper complex is the first CuA model with a Cu2S2 core that can be reversibly
oxidized or reduced
between the Cu(+1.5)–Cu(+1.5) state and the Cu(+1)–Cu(+1)
state. However, the short Cu(+1)–Cu(+1) distance (2.64 Å)
and long Cu(+1.5)–Cu(+1.5) distance (2.93 Å) significantly
increased the reorganization energy of ET, which was much higher compared
to the reorganization energy observed in the water-soluble CuA domain of T. thermophilus
cytochrome ba
3.
1337
4.6
Structural Features Controlling the Redox
Chemistry of the Cupredoxins
4.6.1
Role of the Ligands
As the immediate
residues that coordinate to the copper centers, the ligands exert
a huge influence on the redox properties of cupredoxins. The strong
Cu–thiolate bond(s) playd the dominant role in defining T1
Cu and CuA centers in both their electronic structures
and ET functions. Except for a few unnatural amino acids, mutation
of Cys will change the T1 copper character. The same happens in the
CuA center in that mutation of Cys to Ser will result in
either T1 or T2 center.
The His residues are important for shielding
the copper center from the solvent and for directing ET. C-terminal
His is on a hydrophobic patch of T1 copper proteins. The hydrophobic
patch directly interacts with redox partners of T1 copper proteins.
Mutation of either His to Gly creates an open binding site, where
external ligands could coordinate with copper and influence the properties
of T1 copper proteins. Due to the open binding site, the His to Gly
mutant exhibited a high reorganization energy and low ET rate.
The axial Met is less conserved in T1 copper proteins. Besides
Met, native T1 copper proteins could have the more hydrophilic Gln
or the more hydrophobic, noncoordinating Leu/Phe at the axial position.
There is a general trend that proteins with Gln as their axial ligand
have the lowest reduction potentials, proteins with Met have intermediate
reduction potentials, and proteins with Leu/Phe have the highest potentials.
The reduction potential tuning role of the axial ligand has been further
confirmed by mutagenesis studies. The correlation between the hydrophobicity
of the axial ligand and the reduction potential has been established
by incorporation of a series of Met analogues. The role of the highly
conserved axial methionine ligand was performed by glutamate, aspartate,
and leucine in the engineered CuAAz.
1374
In contrast to the same substitutions in the structurally
related blue copper azurin, much smaller changes (∼20 mV) in
reduction potential were observed, indicating that the diamond-core
structure of the CuA site is much more resistant to variation
in axial ligand interactions than the distorted tetrahedral structure
of the blue copper protein.
4.6.2
Role
of the Protein Environment
The first coordination sphere
directly affects the spectroscopic
properties and ET of the T1 copper proteins. Beyond the first coordination
sphere, the protein scaffold holds copper ligands together and forces
trigonal geometry regardless of the oxidation state of copper, as
suggested by the rack mechanism
1168
or
the entatic state.
1170
Furthermore, the
environment around the primary coordination sphere can fine-tune the
electronic structure and redox properties of the copper centers by
noncovalent interactions such as a H-bonding network to the copper
ligands.
94,1119,1430
Through manipulation of H-bonding networks in the secondary coordination
sphere, Marshall et al. managed to tune the reduction potential of
azurin over the natural range while maintaining T1 character in the
copper center.
1088
The same mutations that
affected the noncovalent interactions in azurin were introduced to
tune the reduction potentials of engineered CuAAz.
1385
The effects of these mutations were in the
same direction, but with smaller magnitude in the CuA site
due to dissipation of the effects by two copper ions rather than the
single copper ion in blue copper proteins.
All these findings
are important in understanding the different roles of the two cupredoxins.
Since the T1 blue copper proteins are used in a wide range of ET processes,
the reduction potentials of the blue copper proteins need to be tuned
to fit a wide range. Such a tuning is mainly achieved by changing
the axial ligands and H-bonding network in the secondary coordination
sphere.
95,1088
However, the CuA sites are only
found in terminal electron acceptors with very small potential differences
between redox partners where a wide range of reduction potentials
is not preferred. The diamond-core structure of CuA sites
decreases the reorganization energies and enables fast ET processes.
4.6.3
Blue Type I Copper Centers vs Purple Cu
A
Centers
The type I blue copper centers
are widely found as ET centers common in many biological systems.
However, the CuA centers are only found in CcOs, N2ORs, and the oxidase from Sl. acidocaldarius
(SoxH). Several key questions that have been raised regarding these
sites are concerned with how such a mixed-valence binuclear copper
site was selected, what the advantage of such a site compared to T1
blue copper sites is, and why the CuA sites are only found
in terminal electron acceptors. To answer these questions, a direct
comparison of the ET rates of these two centers is required. The engineered
CuA site in azurin provides a great opportunity to eliminate
the protein structure contribution to the ET process since the ET
rates are measured in the same azurin scaffold.
1388
The CuAAz demonstrated that CuA is
a more efficient ET site even with a smaller driving force between
the reduced disulfide and CuA site than between the reduced
disulfide and blue copper site. The calculated reorganization energy
of the CuA site is only half that of the blue copper site,
which is due to the rigid structure of the diamond core in the CuA site. Both CcOs
and N2ORs are
large enzymes that contain multiple ET sites. As the electrons are
transferred along the chain, the difference in reduction potentials
as the driving force must fall within a narrow range of values. In
this case, the ET sites with lower reorganization energy would be
preferred because the driving force might be small.
5
Enzymes Employing a Combination of Different
Types of Electron Transfer Centers
5.1
Enzymes
Using Both Heme and Cu as Electron
Transfer Centers
5.1.1
Cytochrome c and CuA as Redox Partners to Cytochrome c Oxidases
The CcO is a terminal protein complex in the respiratory
electron
transport chain located in the bacterial or mitochondrial membranes.
This large protein complex receives four electrons from cyt c that are used to efficiently
reduce molecular oxygen to
water with the help of four protons from the aqueous phase without
producing any reactive oxygen species such as superoxide and peroxide.
In addition, it translocates four protons across the membrane, which
establishes an electrochemical potential gradient used for ATP synthesis.
Out of many different types of CcOs from various
different organisms, the families involved in aerobic respiration
that generally use cyt c as their biological electron
donors are caa
3, aa
3, cbb
3, ba
3, co, bb
3, cao, and bd oxidases.
1431
Cyts caa
3 and cbb
3 oxidases contain a distinct cyt c domain
integrated into the cyt c oxidase enzyme complex.
Cyt aa
3 oxidase is the mitochondrial counterpart
of cyt caa
3 except that it does not contain
the cyt c domain at the C-terminal end of subunit
II (Cox2) of the enzyme complex. Subunit II also contains the binuclear
CuA center. Cyt cbb
3 oxidases
do not contain the CuA center, but they contain both a
monocytochrome c subunit (FixO or CcoO) and a dicytochrome c subunit (FixP or CcoP).
79,1432
Many facultative
anaerobes use bo and bo
3 oxidases which use quinol as the substrate instead of cyts c. Depending on the organism,
the cyts c are associated with the enzyme complex by either covalent or noncovalent
interactions.
1433
For example, in the bacterium
PS3, cyt c binds covalently to the protein
complex at the C-terminal end of subunit II.
1434−1438
In Pa. denitrificans, the cyt c subunit is tightly bound to the oxidase subunit by
covalent
interactions and can be removed by treatment of a high concentration
of detergent. In eukaryotes, cyts c bind to the cyt c oxidase loosely, which can be
removed at high salt concentrations.
Mammalian cyt c oxidases have been shown to bind
one molecule of cyt c at a high-affinity site, which
serves as the electron entry point.
1439−1441
There is evidence
of the presence of a second low-affinity site, but the role of such
secondary interactions between cyt c and the oxidase
is not well-known. It has been shown that cyts c use
a series of several (six or seven) positively charged lysines near
the heme edge which form complementary electrostatic interactions
with negatively charged carboxylates on the high-affinity site of
subunit II of the oxidase. Such electrostatic interactions are important
for placing the substrate in the correct orientation to bind to the
oxidase complex.
1442,1443
Available data suggest
that electrons are transferred from reduced
cyt c, one at a time, to the oxidized CuA.
1444,1445
Then internal ET takes place from the reduced
CuA to the LS heme a and to the binuclear
active site consisting of HS heme a
3 and
CuB where the dioxygen reduction takes place (Figure 65). It has been measured that
the ET rate constant
from CuA to heme a is 20 400 s–1 and the rate of the reverse process, from heme a
to CuA, is 10 030 s–1 in Pa. denitrificans cytochrome c oxidase by pulse radiolysis.
1340
A similar study was also applied to cytochrome ba
3 from T. thermophilus,
and the first-order rate constants are 11 200 and 770 s–1, respectively.
1340
Electron
transfer from cyt c to CuA and CuA to heme a is fast,
1445,1446
while the ET from heme a to the heme a
3/CuB site is slow and has been proven to be
the rate-limiting step of the reaction.
1447,1448
It has also been shown that the presence of CuA is not
required for the oxidase activity as the deletion of the CuA gene from beef heart
cyt c oxidase slows down the
ET rate, but still maintains some oxidase activity.
1449,1450
Figure 65
Cyt c oxidase from Pa. denitrificans (PDB ID 3HB3). The ET pathway is shown with arrows.
Binding of cyt c to the oxidase causes conformational
changes in both the protein partners.
1451,1452
The major
changes are observed upon reduction of the CuA and heme a centers. It has been proposed
that the reduction of these
two redox centers causes a conformational change of the binuclear
active site from a closed to an open state that facilitates the intramolecular
ET that couples the subsequent redox reaction and proton translocation.
1453−1456
NRVS on cyt c
552 from Hydrogenobacter thermophilus has indicated the presence
of strong vibrational dynamic coupling between the heme and the conserved
-Cys-Xxx-Xxx-Cys-His- motif of the polypeptide chain.
1457
Such vibrational coupling has been proposed
to lower the energy barrier for ET by either transferring the vibration
energy released upon protein–protein complex formation or by
modulating the heme vibrations.
A recent NMR study has shown
that the hydrophobic residues near
the heme of cyt c form hydrophobic interactions with
cyt c oxidase and are major contributors to the complex
formation, while the charged residues near the hydrophobic core dictate
the alignment and orientation of cyt c with the enzyme
to ensure efficient ET.
1458
The affinity
of oxidized cyt c for complex formation with CcO is significantly lower, suggesting
that ET is gated by
the dissociation of oxidized cyt c from CcO. The rate of dissociation of oxidized
cyt c is dictated by the affinity of oxidized cyt c for
CcO that provides facile ET.
5.1.2
CuA and Heme b as Redox Partners
to Nitric Oxide Reductases
Although the
NORs from Gram-negative bacteria use cyt c as the
biological electron donor to the heme c, one NOR
(qCuANOR) purified from the Gram-positive bacterium B. azotoformans shows the presence
of a quinol binding
site and uses the binuclear CuA site as an electron acceptor
instead of heme c.
1332,1333
This family
of NORs use melaquinol as the physiological electron donor to the
CuA site instead of cyt c. Electrons are
passed from melaquinol to the CuA site and are then transferred to
the LS heme b and onto the binuclear active site
consisting of a HS heme b
3 and a nonheme
FeB site.
5.1.3
Cytochrome c and CuA as Redox Partners to Nitrous Oxide Reductases
The
N2OR is the last enzyme in the denitrification pathway
which reduces nitric oxide to dinitrogen.
1329,1330,1459
N2ORs are homodimeric
periplasmic enzymes containing the binuclear ET site CuA which receives electrons
from cyt c and a tetranuclear
catalytic site, CuZ. A unique N2OR has been
reported from Wolinella succinogenes which has a C-terminal cytochrome c domain that
is suggested to be the biological electron donor to the CuA center.
1460
5.2
Enzymes
Using Both Heme and Iron–Sulfur
Clusters as Electron Transfer Centers
5.2.1
As
Redox Partners to the Cytochrome bc
1 Complex
The coenzyme Q–cytochrome c oxidoreductase,
also called the cytochrome bc
1 complex
or complex III, is the third complex in the
electron transport chain playing a crucial role in oxidative phosphorylation
or ATP generation. The bc
1 complex is
a multisubunit transmembrane protein complex located at the mitochondrial
and bacterial inner membrane that catalyzes the oxidation of ubihydroquinone
and the reduction of cyt c(1461) coupled to the proton translocation from the matrix
to the cytosol.
The catalytic core of the bc
1 complex
consists of three respiratory subunits: (1) subunit cyt b that contains two b-type
hemes, b
L and b
H, (2) subunit cyt c, containing a heme c
1, and
(3) iron–sulfur protein subunit containing a Rieske-type [2Fe–2S]
cluster (Figure 66). While in some α
proteobacteria such as Paracoccus, Rs. rubrum, and Rb. capsulatus, this enzymatic
core containing the three subunits is catalytically
active, several additional (seven or eight) subunits are present in
the mitochondrial cytochrome bc
1 complexes.
86,1462
Figure 66
Bovine cytochrome bc
1 complex (PDB
ID 1BE3). Different
ET domains and their cofactors are shown. bL = low-potential
heme, bH = high-potential heme, and Q = ubiquinol. Electron
transfer pathways are shown with arrows.
Structures of the bc
1 complex
from
various resources such as yeast, chicken,
1029
rabbit,
1029
and cow
1026,1029,1463
show that the cyt b subunit consists of eight transmembrane helices designated as
A–H.
Hemes b
L and b
H are contained in a four-helix bundle formed by helices A–D
and are separated by a distance of 8.2 Å. The axial ligands for
both hemes are all His and are located in helices B and D. His83 and
His182 are bound to heme b
L, while His97
and His196 are axial ligands for heme b
H. The cyt c subunit containing cyt c
1 is anchored to the membrane by a cytoplasmic domain
and belongs to the Ambler type 1 cyt c based on the
protein fold and the presence of the signature sequence -Cys-Xxx-Xxx-Cys-His-.
Electron transfer has been proposed to occur through the exposed “front”
face of the corner of the pyrrole II ring.
1029
One of the His residues that acts as a ligand to the [2Fe–2S]
cluster is 4.0 Å from an oxygen atom of heme propionate-6 and
8.2 Å from the C3D atom of the heme edge of cyt c
1. Such proximity of the heme group and the Rieske-type
cluster has been proposed to facilitate ET. Using this distance of
8.2 Å, a rough estimation of the ET rate from the iron–sulfur
protein to cyt c
1 has been calculated
to be 4.8–80 × 106 s–1.
On the basis of the relative orientations of the prosthetic groups
as discussed above, an ET pathway has been proposed where in round
I an electron is transferred from a bound ubiquinol to the Rieske-type
cluster into the cyt c
1 via its heme propionate-6
and out of cyt c
1 via its pyrrole II heme
edge to the cyt c (not the same as cyt c
1).
78,1029
At the same time the low-potential
heme (bL) pulls an electron from the ubiquinol and transfers
it to the high-potential heme (bH), which is ultimately
picked up by an oxidized ubiquinone. The same cycle is repeated in
round II.
Mitochondrial cyt c or bacterial
cyt c
2 connects the bc
1 complex
with the photosynthetic reaction center or cyt c oxidase.
80,1464
The mode of interaction between cyt c (or c
2) with its redox partners has been proposed
to involve docking of cyt c with its solvent-exposed
heme edge (called the “front” side). There are multiple
dynamic H-bonding and salt bridge interactions between the cyt c and cyt c
1 of the bc
1 complex.
1465
The
front side is composed of a ring of positively charged Lys residues
near the exposed heme edge. The opposite side, called the “back”
side, is composed of several negatively charged residues. This charge
separation creates a dipole moment in both bacterial cyts c
2 and mitochondrial cyt c.
1466,1467
The positively charged front side forms complementary interactions
with the negatively charged surface of its partner, which orients
the electron donor in proper alignment for facile ET. EPR experiments
with cyt c
2 from Rb. capsulatus have demonstrated that the dipolar nature of cyt c
2 influences its orientations, which facilitate ET to
its partner under physiological conditions.
1468−1470
Rieske protein can accommodate three conformations in the
complex:
The first is the c1 position in which the His ligand is H-bonded to
propionate of heme in cyt c, and fast ET (60 000
s–1)
1471
between the
two will occur.
1026
At this state the cluster
is far from the quinone binding site. The b position allows interaction
between the cluster and quinone. This position was stabilized by interaction
of H161 with the inhibitor stigmatellin that mimics the H-bond pattern
of semiquinone.
223,1029
The final conformation is an
intermediate state in which the Rieske protein cannot interact with
either cytochrome or quinone.
865
The cycle starts from an intermediate state (Figure 67). Upon binding of reduced hydroquinone,
the Rieske protein
will move to state b and an electron will be transferred to hydroquinone,
generating a semiquinone, which binds tightly to the Rieske protein.
This tight interaction will become loose by transfer of a second electron
from semiquinone to heme b
L and generation
of quinone. The thermodynamically disfavored reduction of heme b
L by semiquinone is coupled to favorable oxidation
of hydroquinone by the Rieske center. As a result the reduction potential
of the Rieske center is of significant importance to the rate of reduction
of heme b
L. Reduction of the Rieske center
and heme b
L happens within a half-life
of 250 μs as evident by freeze quench EPR. The semiquinone intermediate
has a very high affinity for the Rieske protein. This tight binding
will increase the reduction potential of the Rieske center by 250
mV. This binding mode and increased reduction potential will ensure
that the Rieske center will not reduce cyt c before
heme b
L is reduced and quinone is formed.
The reduced Rieske center will then move to its c
1 state and transfer an electron to cyt c. After complete transfer of both electrons,
the Rieske protein will
go back to its intermediate state for the second cycle.
773,787,865
The binding of quinone and Rieske
protein is redox-dependent. While the kinetics of ET to cyt c is pH-dependent due
to the pH dependence of the reduction
potential, it has been proposed that the rate-limiting step in this
reaction is mostly the transition from one state (e.g., state b) to
another state (e.g., state c
1) of the
Rieske center and not the ET, considering the same rate observed in
mutants with different reduction potentials.
1078
Figure 67
Schematic cycle of Rieske positions in the bc
1 complex. Reprinted from ref (865). Copyright 2013 American Chemical Society.
Although the mechanism of proton
transfer is not very well understood
in this system, evidence suggested that the two protons are bound
to the Rieske center, one to each His in the reduced state. The oxidized
state can have no protons, one proton, or two protons depending on
the pH. It has been shown that removal or mutation of the Rieske cluster
will result in a proton-permeable bc
1 complex,
suggesting a role as a proton gate for the Rieske protein.
1472
NMR was used to calculate the pK
a of His ligands in the T. thermophilus Rieske protein. In this study, residue-selective
labeling was used
to unambiguously assign the NMR shifts. The results were consistent
with other pH-dependent studies of Rieske proteins, showing that one
of the water-exposed His ligands that is close to quinone undergoes
large redox-dependent ionization changes. Their system also supports
proton-coupled ET in the Rieske–quinone system.
864
Analysis of driving forces using a Marcus–Bronsted
method in mutants that had distorted H-bonding due to mutation of
either conserved Ser or Tyr resulted in the proposal of a proton-first-then-electron
mechanism in which the ET follows the transfer of a proton between
hydroquinone and the imidazole ligand of the Rieske cluster.
814
5.2.2
As Redox Partners to
the Cytochrome b
6
f Complex
Cyt b
6
f (plastoquinol–plastocyanin
or cyt c
6 oxidoreductase) is a protein
complex belonging to a “Rieske–cytochrome b” family of energy-transducing protein
complexes found in
the thylakoid membrane in the chloroplasts of green algae, cyanobacteria,
and plants and catalyzes ET from plastoquinol to plastocyanin or cyt c
6 (PSII to PSI) coupled with the proton translocation
across the membrane for ATP generation.
282,1473−1476
It is located in between the PSII and PSI reaction centers in oxygenic
photosynthesis (Figure 68). The b
6
f complex is analogous to the bc
1 complex of the mitochondrial electron transport
chain. The b
6
f complex
comprises seven subunits: a cyt b
6 with
a low-potential (b
p) and a high-potential
(b
n) heme, a cyt f, a
Rieske iron–sulfur protein, subunit IV, and three low molar
mass (∼4 kDa) transmembrane subunits.
1473
There are a total of seven prosthetic groups that are found
in the b
6
f complex: cyt f, hemes b
n and b
p, a Rieske [Fe2–S2] cluster, chlorophyll a, β-carotene, and a c-type heme
designated as c
n, c
x
, or c
i
. This heme, located close to the quinone reductase site near
the electronegative side of the membrane, is linked to the protein
via a single thioether linkage, lacks any axial ligands, and has been
shown to be critical for function of the b
6
f complex.
225,1478−1481
The cyt b
6 subunit contains two bis-His-ligated
hemes, a high-potential heme (−45 mV) on the luminal side and
a low-potential heme (−150 mV) on the stromal side of the thylakoid
membrane. EPR and Mössbauer data reveal that both hemes are
6cLS and have His planes that are perpendicular. Cyt b
6 and subunit IV of the b
6
f complex are structurally similar to cyt c of the bc
1 complex,
184
while there is no structural similarity between
cyt f and cyt c
1 even
though they are functionally similar.
123,1029
The cyt b
6
f complex takes part in linear
electron flow between PSII and PSI where it links the plastoquinone
pool of PSII to plastocyanin or cyt c
6 to PSI as well as in cyclic electron flow within PSI (Figure 68). The linear electron
flow path involves oxidation
of quinol to quinone from PSII to PSI coupled to the generation of
ATP and reduced ferredoxin, which reduces NADP+ to NADPH
via an oxidoreductase FNR. Cyclic electron flow in PSI involves electron
flow via the b
6
f complex
back to the P700 reaction center of PSI. In both the cases two electrons
are passed from plastoquinol at the quinol oxidation site (QP) near the lumenal, electropositive
site of the membrane to the one-electron
acceptor plastocyanin, which is coupled to the “Q-cycle”
1482,1483
involving proton translocation across the membrane. One of the electrons
from plastoquinol is transferred to PSI via the high-potential chain,
while the second electron is passed onto the low-potential, transmembrane
chain on the electronegative side of the membrane where plastoquinone
reduction takes place.
Figure 68
Cyt b
6
f complex in
the photosynthetic electron transport chain. P680 = reaction center
chlorophylls of PSII, QA, QB = quinones of PSII, PQ/PQH2 pool = plastoquinone/plastoquinol
pool, Fe–S = Rieske cluster, f = cyt f of the high-potential chains (blue arrows),
Qp, Qn = plastoquinol
oxidation and plastoquinone reduction sites, b
p, b
n, c
n = hemes of the low-potential chain (red arrows), Fd = ferredoxin,
and P700 = reaction center chlorophylls of PSI. The domain movement
of the Rieske protein is shown by a two-sided arrow. The direction
of proton translocation across the membrane is shown by proton arrows.
The electronegative (cytoplasmic) (n) and electropositive (luminal)
(p) sides of the membrane are labeled, and ET pathways are shown by
arrows. A possible direct ET path from PSI to the cyt b
6
f complex is shown as the dashed line
from Fd to the Qn site. Reprinted with permission from ref (1477). Copyright 2012
Springer
Science+Business Media.
On the His ligation side of the heme, a chain of five conserved
water molecules oriented in an L-shaped manner have been identified
from the X-ray structure, which form hydrogen bonds with ten amino
acid residues from the protein, seven of which are conserved.
1473,1484,1485
These water molecules have
been proposed to act as “proton wires” in coupling of
the ET with proton transfer across the membrane.
1485,1486
The heme of cyt f is located in a hydrophobic environment
and is protected from the solvent by Tyr1, Pro2, Ile3, and Phe4 (or
Trp4 in cyanobacteria).
161
The side chain
of residue 4 is located close to the heme edge and oriented almost
perpendicular to the heme plane (Figure 69).
1485
This edge-to-face interaction of the Trp4
and the heme has been proposed to be responsible for tuning the reduction
potential of the heme by interaction with the porphyrin π molecular
orbitals. Such edge-to-face interactions have been observed in cyt b
5 (Phe58, Phe35),
141,366
cyt b
562 (Phe61),
382
and peptide-sandwich mesoheme model systems reported by
Benson and co-workers (Trp or Phe).
423,1487
In these
peptide mesoheme sandwich complexes the heme–Trp interaction
has been shown to be important to stabilize the α-helical scaffold
as well as the ferric state of the heme iron.
1488
Such interactions also stabilize the ferric state of the
heme iron in the cyanobacterium cyt f.
Figure 69
Environment
around the heme of cyt f (PDB ID 1HCZ). Hydrophobic residues
are shown as gray sticks. The “edge-to-face” interaction
at 4 Å between Phe4 and the heme that is proposed to be important
to tune the reduction potential of the heme iron is shown. The five
conserved molecules that have been proposed to act as “proton
wires” that couple ET with proton transfer are shown as red
spheres. Residue numbering of waters is arbitrary.
The chloroplast Rieske proteins work in the same
way. It has been
shown that the movement of these Rieske proteins will also function
as a redox-state sensor that can balance the light capacity of the
two photosystems. This state transition can also act as a switch between
cyclic and linear electron flow.
1489
5.2.3
As Redox Centers in Formate Dehydrogenases
Formate dehydrogenases (Fdh’s) catalyze decomposition of
formate to CO2. They exist in both prokaryotes and eukaryotes.
Fdh’s are mainly NAD+-dependent in aerobic organisms
and NAD+-independent in anaerobic prokaryotes, donating
electrons from formate to a terminal electron acceptor other than
O2.
1490
Structural studies reveal
that Fdh’s contain one to three subunits with either W or Mo
in the active site.
1491−1493
Fdh-N from E. coli is among the most well studied Fdh’s. It is important in
the nitrate respiratory pathway under anaerobic conditions. It is
a membrane-bound trimer (α3β3γ3) with a molar mass of 510 kDa. It harbors a Mo-bis-MGD
cofactor
and a [4Fe–4S] cluster in the catalytic α subunit, four
[4Fe–4S] clusters in the β subunit, and two heme b groups in the γ subunit (Figure
70).
1492
The β subunit transfers
electrons between the α and γ subunits, similar to other
membrane-bound oxidoreductases that bind four [4Fe–4S] clusters,
such as nitrate reductases, [NiFe] hydrogenases, DMSO reductase, and
thiosulfate reductase.
1494
Figure 70
Overall structure of
Fdh-N from E. coli. Cofactors are displayed
as spheres and denoted accordingly on the
right. The putative membrane is shown in gray shading. PDB ID 1KQF. Reprinted with
permission from ref (1492). Copyright 2002 American Association for the Advancement
of Science.
Fdh from Dv. desulfuricans is an
αβγ protein with a molar mass of ∼150 kDa.
It contains four different types of redox centers, including four
heme c centers, two [4Fe–4S] clusters, and
a molybdopterin.
1495
EPR studies showed
the existence of two types of Fe–S clusters after reduction,
i.e., center I with g values of 2.050, 1.947, and
1.896 and center II with g values of 2.071, 1.926,
and 1.865. Midpoint reduction potentials of the two Fe–S clusters
are −350 ± 5 mV for center I and −335 ± 5
mV for center II.
Fdh from Dv. gigas is an αβ
protein
1493
containing tungsten instead
of molybdenum. It also possesses two [4Fe–4S] clusters similar
to Fdh from Dv. desulfuricans.
981,1496
5.2.4
As Redox Centers in Nitrate Reductase
NARs reduce nitrate to nitrite, a vital component in the nitrogen
respiratory cycle. Most NARs isolated so far contain three subunits,
NarG (112–140 kDa), NarH (52–64 kDa), and NarI (19–25
kDa). NarG harbors a Mo-bis-MGD cofactor and a [4Fe–4S] cluster,
NarH contains one [3Fe–4S] cluster and three [4Fe–4S]
clusters, and NarI immersed in the membrane binds two b-type hemes (Figure 71).
1497−1502
The overall folding and cofactor positions are strongly homologous
to those of Fdh from E. coli.
1503
The eight redox centers are separated by 12–15
Å from each other and form an ET pathway about 90 Å long.
NAR from Cupriavidus necator does not
contain the NarH domain and harbors two c-type hemes
in the small subunit.
1504
Figure 71
Overall three-dimensional
structure of NarGHI from E. coli K12.
PDB ID 1Q16.
Subunit and cofactor names are denoted.
Reprinted with permission from ref (1505). Copyright 2006 Elsevier.
6
Summary and Outlook
This review summarizes three important classes of redox centers
involved in ET processes. Although each class spans a wide range of
reduction potentials, none of them can cover the whole range needed
for biological processes. Together, however, they can cover the whole
range, with cytochromes in the middle, Fe–S centers toward
the lower end, and the cupredoxins toward the higher end (Figure 1). All three redox
centers have structural features
that make them unique, and yet they also show many similarities that
make them excellent choices for ET processes.
Because the redox-active
iron is fixed into a rigid porphyrin that
accounts for four of the iron’s six coordination sites, most
of the electronic structure and redox properties remain similar between
different cytochromes. In completing the primary coordination sphere
of the iron, cytochromes typically use a combination of nitrogen and
sulfur ligations from histidine or methionine side chains, respectively;
terminal amine ligation has also been observed. In general, mutagenesis
studies reveal that methionine ligation raises the reduction potential
by 100–200 mV, relative to histidine ligation, primarily due
to the lower affinity of thioether to the higher oxidation state of
the heme, and that the effect is generally additive.
192,386,461−463,465
Heme puckering or flexing has
been demonstrated to tune the reduction potentials by up to 200 mV.
513
Changes in the heme type between b and c would be expected to change the electronic
properties of the heme; however, the effect on the reduction potential
is small and varies depending on the systems studied.
446,448
It is clear, on the other hand, that the electron-withdrawing formyl
group on heme a appears to be responsible for the
increase in the reduction potential by ∼160 mV.
459,460
For iron–sulfur proteins, the reduction potential ranges
are influenced to some extent by the number of irons because it affects
the redox states and transitions. In the case of clusters with the
same number of irons, the higher the redox pair, the higher the reduction
potentials (e.g., HiPIPs have a [4Fe–4S]2+/3+ pair,
while ferredoxins have a [4Fe–4S]1+/2+ pair).
719
In addition, the cluster geometry such as Fe–Sγ–Cα–Cβ torsional angles, the Fe–Fe
distance, and covalency of Fe–S
bonds also play important roles in some proteins.
618,901,1085,1506
Electron delocalization of the cluster and the net charge of the
cluster are also important. For example, it has been shown that the
net charge of the protein is the main factor determining the reduction
potential within HiPIPs. Electrostatic effects of the charged residues
in the secondary coordination sphere can influence the solvent accessibility
and consequently the dielectric constant around the metal center.
However, the effects are usually complicated and difficult to rationalize
by just Coulomb’s law. For example, in rubredoxin from Cl. pasteurianum, replacement
of a neutral surface
residue by a positively charged Arg or a negatively charged Asp has
led to an increase of reduction potentials in both cases.
611,612
Finally, the direct ligands to iron and H-bonding interactions with
the direct ligands make significant contributions to the reduction
potential.
541
When the common Cys thiolate
ligand was replaced with a His imidazole ligand, naturally in the
Rieske proteins, or with Ser by site-directed mutagenesis, the reduction
potentials changed accordingly.
721,773,1087
The multiple NH···S H-bonding interactions
in rubredoxin render the reduction potential of the [FeCys4] center to fall in the
range of −100 to +50 mV, while reduction
potential of the corresponding model complexes without the H-bonding
networks is around 1 V.
92,588−590
The NH···S H-bonds have also been shown to be important
in determining reduction potentials between different ferredoxins
as well as ferredoxins vs HiPIPs.
617,618,718,719
For cupredoxins,
the metal centers cannot be easily fixed like
in either porphyrin or thermodynamically stable iron–sulfur
clusters and proteins play a more prominent role in enforcing the
unique trigonal geometry and strong copper–thiolate bond to
maintain a low reorganization energy for the ET function. In this
class of proteins, both the geometry and the ligands, particularly
the strictly conserved Cys, play a dominant role in controlling the
redox properties. In T1 copper protein azurin, changing axial Met
to a stronger cysteine or homocysteine induced a geometry change and
weakened the Cu–S bond. These changes in turn resulted in a
>100 mV decrease in the reduction potential.
1293
Deleting the H-bonding to Cys, realized through the Phe114Pro
mutation in azurin, affected the covalency of the Cu–S bond
and lowered the reduction potential of azurin.
114,1088,1316
Despite the differences
in the primary coordination spheres, all
three redox centers employ noncovalent secondary coordination interactions
in fine-tuning the redox properties.
The first common feature
is the control of the degree of solvent
exposure; the deeper the redox centers are buried into the hydrophobic
center of the protein, the higher the reduction potential and the
smaller the changes in the reorganization energy due to influences
by the solvent. For example, redox center burial is considered to
be one of the main factors for differences in reduction potentials
between different HiPIPs and ferredoxins.
618,719,749,752
Furthermore, a computational study of heme proteins over an 800
mV range has attributed the greatest correlation with the reduction
potential to solvent exposure.
457
The second common feature is the electrostatic interactions. For
example, the net charge of protein is shown to be the only factor
that correlates with the reduction potentials of different HiPIPs.
715,752,890
The number of amide dipoles
and not necessarily H-bonding is shown to be important in reduction
potential determination in ferredoxins.
718,719
In myoglobin, Val68, which was in the van der Waals interaction
distance with the heme group, was replaced by Glu, Asp, and Asn. A
200 mV decrease in reduction potential was observed for the Glu and
Asp mutants compared to the wild type.
481
This study demonstrated that replacement of hydrophobic Val68 by
charged and polar residues led to substantial changes in the reduction
potential of the heme iron. In a number of different cytochromes,
electrostatic polar and charged groups near the heme were shown to
vary the potential by 100–200 mV.
169,479,481,482
For instance, in cyts c
6 and c
6A, the glutamine at positions 52 and 51, respectively,
were shown to raise the potential ∼100 mV,
479
and in cyt c, the Tyr48Lys mutation raised
the potential 117 mV;
480
all these effects
can be attributed to charge compensation in the heme pocket. Similarly,
replacing Met121 with Glu or Asp in T1 copper azurin resulted in 100
and 20 mV decreases in the reduction potentials, respectively.
1278,1289
Beyond copper ligands, mutating Met44 in azurin to Lys destabilizes
Cu(II), causing a 40 mV increase of the reduction potential.
1507
The final common feature is the presence
of a hydrogen-bonding
network around the ligands to the metal center, especially those to
the ligand that dominates the metal–ligand interactions. For
example, the NHamide···Scys H-bonds
are known to be important in different reduction potentials between
rubredoxins, HiPIPs, and ferredoxins.
617,618,718,719
They are also shown
to play a role in different reduction potentials of different ferredoxins.
Other than backbone amide H-bonds, H-bonds from side chains are also
important. A good example of such is H-bonds from conserved Ser and
Tyr in Rieske proteins and a lack of thereof in Rieske-type proteins,
hence differences in the reduction potential.
781
In cytochromes, H-bonding interactions with the axial ligands
can tune the potential by up to 100 mV.
474,476,477,1508
For instance, increasing the imidazolate character of the axial
His ligand in cyt c by strengthening H-bonding from
the H to the Nε increased the potential by nearly 100 mV,
474
and disrupting the hydrogen bond donation from
Tyr67 to the axial Met resulted in a 56 mV decrease in potential.
476,1508
Similarly, the H-bonding interactions to the Cys in cupredoxins
are known be responsible for their reduction potential differences.
114
A test of how much we understand these
structural features responsible
for the redox properties is to start with a native redox center and
use the above knowledge to fine-tune the redox properties. A pioneering
work in this area is the demonstration of a ∼200 mV decrease
in the reduction potential of myoglobin when a buried ionizable amino
acid (Glu) was introduced into the distal pocket of the protein, and
such a change has been attributed to electrostatic interactions.
481
Since then, not many examples have shown similar
magnitude changes of reduction potentials by electrostatic interactions,
perhaps due to the compensation effect by ions in the buffer or other
ionizable residues nearby. Instead, hydrophobicity and H-bonding network
have been shown to play increasing roles, and a combination of these
effects has been shown to fine-tune the reduction potentials of T1
copper azurins by more than 700 mV, beyond its natural range.
1088
These features were further shown to be additive,
making reduction potential tuning predictable. Such rational design
also allowed the lowering of the reorganization energy of azurin,
1317
which is already known to be very low in comparison
to those of other redox centers. With more such successful examples
in other systems, we will be able to achieve a deeper understanding
of ET reactivity in proteins and facilitate de novo design of ET centers
for applications such as advanced energy conversions.