High Resolution Structure of a Truncated Form of Cytochrome *f*

from *Chlamydomonas reinhardtii*^†⊥^

by

Young-In Chi^‡§^, Li-Shar Huang¶, Zhaolei Zhang¶^@^,

Javier G. Fernández-Velasco^\#^\* and Edward A. Berry¶\*

¶E. O. Lawrence Berkeley National Laboratory, Berkeley, CA 94720, USA

\@Graduate Group of Biophysics,

‡Department of Chemistry and

#Department of Plant and Microbial Biology

University of California, Berkeley, CA 94720, USA

(†)This work was supported in part by a grant from the National
Institutes of Health (GM-20571) to Prof. R. Malkin (Department of Plant
and Microbial Biology, UC-Berkeley ) and by the Office of Health and
Environmental Research, US Department of Energy, under contract
DE-AC03-76SF00098 to LBNL.

(\*) To whom correspondence should be addressed or reprints requested

Javier G. Fernández-Velasco Edward A. Berry

Tel:(510)642-9425 Tel: (510) 486-4336 Fax:(510)642-4995 Fax: (510)
486-5272

e-mail:jfernand@nature.berkeley.edu e-mail:eaberry@LBL.gov

(§)Present address: Joslin Diabetes Center and the Department of
Medicine, Harvard Medical School, Boston, MA 02215

(⊥)The coordinates of the structure presented here have been deposited
in the Protein Data Bank with the accession number 1CFM.

**Running Title :** Structure of *C. reinhardtii* cytochrome *f.*

**Abbreviations and Textual Footnotes**

**Abbreviations:**

F~o~, observed structure factor amplitudes; F~c~, amplitudes of
calculated structure factors; rms, root-mean-square; ncs, non
crystallographic symmetry; W, water molecule; Φc, phases of calculated
structure factors; σ, rms deviation from the mean value of an electron
density map.

**Textual Footnotes:**

^1^ A preliminary release of the program CNS was made available to the
Sung-Hou Kim lab for testing purposes by Axel BrŸnger.

^2^ All residue numbers in this work refer to the sequence of the mature
protein. For comparing with the sequence of the precursor available in
the databases, the length of the leader sequence (31 residues) must be
added.

^3^ The individual cytochrome monomers pack together in the crystal
forming 7 different inter-monomer interfaces: (1) The interface between
monomers A and B in the asymmetric unit involves H-bonds between E34,
N37, and K145 of monomer A and S186, A21, A260, and K165 of monomer B,
and close approach of A250 and Y143 of monomer A to P117 and F4 of
monomer B. (2) Monomer B has an interface with monomer C in a different
asymmetric unit which is nearly the same as the above interface of A and
B, respectively, with the addition of two H-bonds: one between R251 of B
and P117 of C and one between E34 of B and K189 of C. (3) The interface
between monomers A and C in the chosen asymmetric unit involves H-bonds
between Y1, Q7, N63, and K188 in A and N16, N23, and E241 in C. (4) K178
in monomer A forms H-bonds with T43 and D42 in monomer C of another
asymmetric unit. (5) Monomer B is related by a distorted 2-fold rotation
of 176° to monomer A of another asymmetric unit. This brings the
side-chain N of N231 in the two monomers into H-bonding distance. Also
K217 of each monomer ion-pairs with E14 of the other, and N231 N of each
H-bonds with L26 O of the other. The guanidino group of R18 in monomer A
H-bonds with the carbonyl of L214 in B, but the complementary
interaction is not made. (6) K58 of monomer B H-bonds with V203 of
monomer B in another asymmetric unit. (7) N99 of monomer C H-bonds with
D87 of monomer A in another asymmetric unit.

^4^ The ranges of temperature factors of Lys Nζ in the cluster detected
in the three monomers are 36-40, 30-40, 29-40, 45-78, and 37-78 Å^2^ for
K58, K65, K66, K188, and K189, respectively*.* Monomer C shows the
highest temperature values for ζN in K58, K65 and K66, whereas monomer A
shows the highest ones for ζN in K188 and K189. Apart from the cases of
ζN in K188 and K189 in monomer A (which show also the highest
temperature values for ζN in the entire structure, possibly indicating
multiple conformations), those temperature factors are not different
from the corresponding ones to all other ζN's outside the cluster. For
the three monomers, they are close to the median of the broad
distributions shown by the total of 21 lysines per monomer. The lowest
ζN temperature factor is 25 Å^2^ .

^5^ The range of values detected (2.4 to 75.4 Å^2^) was divided in
intervals of 1.25 Å^2^. The resulting distribution is broad and,
probably, multimodal. The global mean, median, and standard deviation of
the whole distribution are 36.6, 35.7, and 13.8 Å^2^, respectively. In
comparison, and for all the three monomers, temperature factors of W1-W5
lie before the first 13 % of the whole distribution, whereas W6-W7 can
have values extending before the median.

**ABSTRACT**

A truncated form of cytochrome *f* from *Chlamydomonas reinhardtii* (the
preferred eukaryotic model organism for photosynthetic electron transfer
studies) has been crystallized (space group P2~1~2~1~2~1~; 3 molecules/
asymmetric unit) and its structure determined to 2.0 Å by molecular
replacement using the coordinates of a truncated turnip cytochrome *f*
as a model. The structure displays the same folding and detailed
features as turnip cytochrome *f* including: a) an unusual heme Fe
ligation by α-amino group of tyrosine 1, b) a cluster of lysine residues
(proposed docking site of plastocyanin), and c) the presence of a chain
of 7 water molecules bound to conserved residues and extending between
the heme pocket and K58 and K66 at the lysine cluster. For this array of
waters we propose a structural role. Two non-crystallographically
related molecules are related by a distorted 2-fold rotation and may
represent the dimeric relation of the monomers in situ, however the heme
orientation in this putative physiological dimer is not consistent with
previous epr measurements on oriented membranes.

Biological systems couple redox energy to processes, such as
biosynthesis and transport, through electron transport chains and proton
electrochemical gradients across membranes (1-4). The electron transport
chains include a series of membrane-bound protein complexes. Each
complex functions as a redox enzyme in part of the overall transport of
electrons from low-potential donors to high-potential acceptors,
coupling the translocation of protons across the membrane and generating
a high-energy proton gradient. Two of those enzymes share an analogous
catalytic activity: a) the ubiquinol:cytochrome c oxidoreductase
(cytochrome ***bc~1~*** complex) in mitochondria (1, 2), prokaryotic
respiration (4), and prokaryotic anoxygenic photosynthesis (4); and b)
the plastoquinol:plastocyanin oxidoreductase (cytochrome ***b~6~ f***
complex) in eukaryotic and prokaryotic oxygenic photosynthesis (5, 6).

In order to understand the detailed mechanism of electron transfer and
proton translocation, high resolution structural information is needed.
While the high resolution structure of the multisubunit mitochondrial
cytochrome ***bc~1~*** complex (7, 8) have shed light in understanding
its mechanism, the structure of the cytochrome ***b~6~ f*** complex is
not available due to the difficulties involved in its crystallization.
At present, only the general dimeric nature of the ***b~6~ f*** complex
has been shown at 8Å resolution (9-14). Crystals of a cyanobacterial
***b~6~ f*** complex have been obtained (15), but considerable
improvement in the order will be required before an atomic resolution
structure can be obtained.

A useful strategy to gain structural information of multisubunit
proteins is to divide the large complexes into smaller components that
are more amenable to structure determination. Thus far, elucidation of
the structure of individual subunits has been successful for a truncated
form of cytochrome ***f*** (a component of the chloroplast cytochrome
***b~6~ f*** complex) from turnip (16, 17), as well as for truncated
forms of the Rieske iron-sulfur subunit of the cytochrome ***bc~1~***
(18) and ***b~6~ f*** (19, 20) complexes.

Cytochrome ***f*** is one of the four redox centers in the cytochrome
***b~6~ f*** complex of the thylakoid membrane in oxygenic
photosynthetic organisms and is analogous in function to the
mitochondrial/bacterial cytochrome ***c~1~*** of the cytochrome
***bc~1~*** complex. Cytochrome ***f*** is the electron donor to the
Cu-containing protein plastocyanin (21) (or to cytochrome *c*~6~
(*c*~552~) in some eukaryotic algae and cyanobacteria, when growing in
Cu deficiency (22-24) and is thought to be reduced directly by the
Rieske iron-sulfur protein (3). The protein has a predicted molecular
weight of ca. 32 kDa, contains one covalently bound heme c group per
molecule, and has a midpoint redox potential of ca. +350 mV (5, 25).
Hydropathy profiles have predicted that the protein has one
transmembrane spanning region, near its C-terminus, and that most of the
protein is localized in the thylakoid lumen (25). When a lumen-side
water-soluble fragment of cytochrome ***f*** was isolated from turnip,
it lacked the C-terminus transmembrane spanning region (33 residues)
(16, 26), but was still redox-active and possessed the same midpoint
redox potential as the intact cytochrome ***f*** (17). The structure of
this truncated form has been reported (16, 17).

In recent years, the eukaryotic unicellular green alga *Chlamydomonas
reinhardtii* has become the preferred model organism for studies of
eukaryotic photosynthesis. This alga has a photosynthetic electron
transfer chain similar to the one found in higher plants but with the
advantage over them that the chloroplast genome of *C. reinhardtii* is
easily transformable (27, 28). Thus, it allows the preparation of
site-directed mutants of chloroplast encoded proteins. Among these
proteins, it is of particular interest cytochrome ***f*** (chloroplast
petA gene) because in *C. reinhardtii* it is 72% identical to the higher
plant cytochrome (25) and because the electron transfer reactions of
cytochrome ***f*** can be studied by kinetic spectrophotometry *in situ*
in whole algal cells (29-32). These advantages for mutational analysis
make the algal cytochrome ***f*** a very good model of the higher plants
cytochrome ***f***. For these reasons, and in order to complement
functional descriptions with specific structural data, it is of great
interest to know the actual structure of the *C. reinhardtii* cytochrome
***f**.*

For this work, the non-photosynthetic strain F283ST of *C. reinhardtii*
(30) was used to purify a truncated soluble form of cytochrome ***f***
that is naturally overexpressed. In the petA gene of this strain a stop
codon replaces the codon for I252 (I283 numbered from the M initiation
codon in the preprocessed protein). Thus, the truncated cytochrome lacks
the last 35 residues that would involve the membrane anchoring α-helix.
Consequently, this modified protein is normally processed and
translocated into the thylakoid lumen where it remains soluble, being
redox active *in vivo* and having very similar spectral characteristics
with the wild type form. The transformation of this intrinsic membrane
protein into a soluble one, that represents the peripheral portion of
the native cytochrome, facilitates its purification and, in combination
with the conserved physicochemical properties, provides an attractive
system for structural studies. Here we present its crystal structure at
a 2.0 Å resolution and a comparison with the previously described one
from turnip. We found that both structures are very similar, although
there are some structural differences.

In this paper we present the crystal structure at 2.0 Å resolution of a
truncated form of cytochrome ***f*** from *C. reinhardtii* and compare
this structure with that previously described from turnip. We find that
the structures are similar but also contain some significant
differences. An abstract of this work has been previously communicated
(33).

**MATERIALS AND METHODS**

***Protein preparation and crystallization.** C. reinhardtii* strain
F283ST was kindly provided by Dr. Francis-Andrè Wollman (Institut de
Biologie Physico-Chimique, Paris). The microalga was grown in carboys
containing 12 L of TAP medium (34) at 28°C with strong aeration (2 L
min-1 per carboy) and moderate magnetic stirring. Dim incandescent light
(10 μEinstein m-2 s-1) was provided. Typically, the culture grew with a
duplication time of 16-18 hr and was harvested with a continuous flow
centrifuge at 29,000g when a density equivalent to A750nm= 0.6-0.8 was
reached. The cell paste was stored at -20 or -80°C.

For cytochrome purification (all steps in ice or 4°C), the cells (80g
wet weight) were thawed and suspended in 800 mL of 50 mM MOPS/KOH pH
7.2, 100 mM KCl, 3 mM NaN3, 0.5 mM Na-EDTA, and 0.1 mM
phenylmethylsulfonyl fluoride; sonicated (5 min, Heat Systems Ultra
Sonics Inc., Model W185, setting 9 (ca. 110W), 1 cm diameter probe); and
then centrifuged 2 hr at 11,300g. The supernatant was diluted to 6 L
with 0.5 mM Na-EDTA and loaded onto a 300 mL column of DEAE-Sepharose
CL6B (Pharmacia) equilibrated with 20 mM KPi (pH 7.5) and 0.5 mM EDTA.
The column was washed with 200 mL of the equilibration buffer and then
eluted with an 800 mL linear gradient 20-75 mM KPi (pH 7.5) and 0.5 mM
EDTA. The pink cytochrome ***f*** containing fractions were pooled and
diluted with an equal volume of distilled water and the ion exchange
chromatography was repeated in the absence of EDTA. Pooled fractions
from the second DEAE column were applied to a 75 mL column of Ceramic
Hydroxyapatite type II, 0.08 mm (Biorad). The column was washed with 200
mL of 50 mM KPi (pH 7.5) and then eluted with an 800 mL linear gradient
50-120 mM KPi. The pooled fractions were concentrated to
[\<]{.underline}5 mL by ultrafiltration (Amicon YM-10 membrane) and
loaded onto a 600 mL column of Sephacryl S-200 HR (Pharmacia)
equilibrated with 50 mM KPi (pH 7.5), 0.5 mM EDTA, to finally elute
cytochrome ***f*** with the same buffer. The protein is obtained in its
reduced state.

The crystals were initially screened by the sparse-matrix method (35)
and one promising set of conditions was optimized. For crystallization,
the pooled fractions from the Sephacryl column were concentrated by
ultrafiltration (Centricon P-10, Amicon) to about 2 mM based on an
extinction coefficient of 34 mM^-1^ cm^-1^ at 554 nm. Portions of 5‑10
μL were mixed with equal volume of precipitant containing 180 g/L
PEG-8000, 0.1 M Na-cacodylate (pH 6.5) and 0.2 M Zn(OAc)~2~; and allowed
to equilibrate at 4 °C by vapor diffusion against the same precipitant
until crystal growth stopped.

***Data collection*** . The native data set from a single crystal was
collected at 100°K on the R-AXISIIC imaging plate detector coupled with
Rigaku Rotaflex X-ray generator operating at 50kV and 100mA. The
crystals were transferred stepwise to a mixture of the mother liquor
containing 30%(v/v) glycerol, acting as a cryo-protectant, before being
flash frozen by liquid nitrogen stream at 100°K for data collection. The
data set was processed with DENZO (36) and scaled with CCP4 program
suite (37). In all, 158,195 observations of 47,961 unique structure
amplitudes to 2.0Å resolution were obtained. The data statistics are
shown in Table 1.

***Structure determination.*** The structure was determined by the
molecular replacement method using the AMORE program (38). The
coordinates of the truncated cytochrome ***f*** from turnip were
obtained from the Protein Data Bank (entry 1CTM) and used as the search
model. Solutions were easily obtained by using the entire model
(residues 1 to 250 except heme) without any side chain substitution or
deletion/insertion. A search carried out with the data between 10.0 and
4.0Å, and center of mass cutoff as 20Å, produced the best solution with
a correlation coefficient of 46.3% and an R-factor of 41.0%, which stood
out well above the next possible solution (18.8% higher in correlation
coefficient). In the program X-PLOR (39) this initial solution had an
R-factor of 41.8% for data between 10.0 and 2.3Å, which after rigid body
refinement was reduced to 39.4%. An Fo-Fc map calculated with phases
from this rigid-body-refined model (still lacking the heme) clearly
showed electron density for the heme group (Figure 1)*.* The heme was
then added, the sequence was changed to that of *C. reinhardtii*, and
those residues that differ between the two species or that did not fit
the electron density were rebuilt.

***Model refinement.*** After solving the molecular replacement, a new
higher resolution (2.0Å) data set was collected, and this was used for
further refinement. An anisotropic *B*-factor correction as well as
resolution-dependent weighting scheme implemented in X-PLOR 3.1 was
applied to the diffraction data. Initial positional refinement of the
rigid body-refined model using 10.0-2.0Å data followed by group
*B*-factor refinement dropped the Rfree to 38.6%. The refinement was
then continued with the simulated-annealing protocols implemented in
X-PLOR 3.1. Restraints were placed on bond lengths, bond angles,
non-bonded contacts and temperature factors of neighboring atoms; ncs
restriction was imposed; no correction for solvent continuum was made.
2Fo-Fc maps as well as omit maps were calculated at regular intervals to
allow manual rebuilding of side chains with correct sequence or a
different rotamer from the search model for the identical sequence. The
rebuilding of the model was done on the "O" graphic program (40) and
regularly checked against a structural database with pep_flip option.
Solvent molecules (all regarded as water) were added conservatively with
due regard for their environment including potential interactions with
hydrogen-bond partners. The solvent model was comprehensively checked
several times during refinement by omitting all water molecules that had
high *B* values (\>60Å2), or made either too-close contacts with each
other or with protein atoms, or made no potential hydrogen-bonded
contacts at all. The simulated annealing protocol did not improve the
model any further at the later stage of the refinement and positional
refinement was continued. Inclusion of individual atomic temperature
factors and removal of the ncs restriction during the final stages was
validated by a substantial decrease in the value of Rfree. The program
PROCHECK (41) was used to check the quality of the structure. At the end
of the X-PLOR refinement the crystallographic R factor was 21.5% and
Rfree value was 29.3%. Bulk solvent correction and anisotropic B-factor
refinement were then carried out in CNS 0.2^1^ using all data \>2σ
between 2.0 and 25 Å, which further reduced R and R~free~ to 21.4 and
27.1%. The model was then subjected to another cycle of Powell
minimization and individual atomic B-factor refinement in CNS with a
maximum likelihood target function, which did not significantly affect
the structure or R-factor. Compared with the final model from Xplor the
rms difference in Cα position was 0.2 Å, with no change greater than 0.3
excluding the N- and C-terminal regions. Rms deviations from ideal
values were 0.006 Å for bond lengths and 1.5° for angles. The standard
deviation in coordinate values estimated from cross-validated Sigma-A
treatment was 0.23 Å.

Representations of the structure were prepared using the progams "O"
(40), Molscript (42), Raster 3D (43), GRASP (44) and Rasmol (45).

***Other methods.*** Absorption spectra were taken with a computerized
Aminco DW**-**2 spectrophotometer in dual beam mode with a 1 nm spectral
band pass, 0.8 nm/sec scanning rate and 50 ms response time. The
wavelength scale was calibrated using the 656.1 nm peak of the deuterium
lamp. SDS-PAGE (46) was performed after denaturing samples in the
presence of 5% w/v SDS, with or without 50 mM dithiothreitol at 80 °C
for 5 min; samples were not centrifuged before electrophoresis.
Non-denaturing electrophoresis was performed with Tris-glycine pH 8.8
gels. In all cases, pre-cast gels 14% or 4-20% acrylamide (with
equivalent results), all buffers and MW standards (MW 200-2.5 kDa) as
well as running conditions were from NOVEX (San Diego, CA). Protein
determination, using BSA as standard, and hemechrome
extraction/measurements were performed as in (47). Analytical size
exclusion chromatography through a Pharmacia LKB Superdex 75 HR 10/30
was performed with a Pharmacia FPLC at 22 °C and 1 mL/min. The column
was calibrated with 12 standards (MW 66-3.5 kDa).

**RESULTS**

***Description of the purified protein.*** The truncated form of *C.
reinhardtii* cytochrome ***f**,* that is purified here for the first
time, has almost identical spectral characteristics as cytochrome
***f*** from a wild-type strain (30) (see results below). It is also
redox active; it can be rapidly photooxidized in intact cells (30); and
*in vitro* it can be reduced and oxidized by ascorbate and ferricyanide,
respectively. In addition, it can bind to and react with homologous
plastocyanin (48, 49).

After the last step of purification of cytochrome ***f*** (absorbance
ratio 554.5/280 nm is 0.72, for the fully reduced form), an overloaded
SDS-PAGE (30 µg protein/lane) stained with Coomasie blue shows only one
band and a mobility equivalent to a MW of 31[+]{.underline}1 kDa, both
with the sample prepared in reducing conditions (dithiothreitol) or not.
The same mobility is also observed with a load of 1 µg protein. FPLC
size exclusion chromatography of the cytochrome in non-denaturing
conditions (4 mM NaPi pH 6.0, 250 mM KCl), yields a slightly
asymmetrical tailing peak and a molecular weight of 32[+]{.underline}2
kDa. Non-denaturing electrophoresis shows (Coomasie) a main band, with
little tailing, and a minor component (10%) with a slightly higher
mobility.

The spectrum of the purified cytochrome in the reduced state (in 50 mM
KPi pH 7.4) shows peaks at 421.4, 523.9, 531.1 and 554.2 nm; whereas in
the oxidized state the maxima are at 410.8 and \~530 nm. The maxima of
the reduced-minus-oxidized spectrum are at 421.8, 524.1, 531.1 and 554.2
nm (this last value is 0.5 nm higher than the one found through the
light-induced differential spectrum *in situ* (30)). The isosbestic
points are at 414.9, 432.3 and about 509, 533, 543 and 560 nm.

Using the pyridine hemechrome quantification, the extinction
coefficients

(mM-1cm-1) were: for the reduced protein 34 at 554 nm, whereas in the
reduced-minus-oxidized spectrum, 26 at 554 nm and 28 at 554-538 nm.
These values are similar to the ones for purified spinach and turnip
cytochromes ***f*** (50).

The properties of this purified protein indicate that the nature of the
truncated cytochrome ***f*** is very similar to that of the native form
and that, therefore, it is a good model for studying
structural/functional relationships.

***Structural determination.*** Crystals of the type used for this study
have maximal and minimal dimensions of 1mm and 0.3 mm. The cytochrome is
crystallized in the reduced form. The crystals belong to the space group
P212121 (a=75.62 Å, b=94.95 Å, and c=120.20 Å). The asymmetric unit
contains three monomers of the cytochrome.

Diffraction data were phased by molecular replacement using the
coordinates of Brookhaven Protein Database entry 1CTM for a truncated
form of turnip cytochrome ***f*** (16) as a search model. A model of the
three cytochrome molecules was built and refined against 2.0 Å
diffraction data to a crystallographic R factor of 21.4% and R-free
value of 27.1%. The detailed refinement statistics are listed in Table
1.

Figure 1 shows an "omit" map of the heme region, phased from a model in
which the heme and residues Y1 and H25, that provides the 5th and 6th
ligands to the heme iron in turnip (16) (see below), were omitted. This
eliminates phase bias with respect to these residues. The electron
density clearly shows heme axial ligation by the N-terminal amino group
of Y1 and by Nδ of H25, as indicated in the model.

***Description of the structure.*** The final model (PDB entry 1CFM)
contains in the asymmetric unit 3 hemes, all residues of 3 monomers, and
1001 water molecules. As we expect from sequence homology (72% sequence
identity), the overall folding is very similar to that from *Brassica
campestris* (turnip) (16). The protein is made of two distinct domains
in an elongated form where the heme group sits within the large domain
near the interface between two domains (Figure 2). The large domain has
an immunoglobulin-like fold flanked by small α-helices, and the small
domain has a jelly-roll fold made up of 4 major and 3 minor β-strands. A
schematic drawing of the secondary structure arrangement is shown in
Figure 3.

The *C. reinhardtii* cytochrome ***f*** contains 1 more residue than the
turnip protein, 286 aminoacids rather than 285. Gray (25) aligned the
sequences (Figure 4) using 2 insertions in *C. reinhardtii* and one in
turnip, all in the region of relatively low homology between residues
169 and 204 (turnip).

Using this scheme, the backbone atoms of the large domain of the algal
cytochrome ***f*** monomers and that of the turnip cytochrome ***f***
can be superimposed accurately (with rms deviations [\<]{.underline} 0.5
Å), with the exception of the truncated C-terminus (R251) and the loop
after helix 1 (around residue 15). When this superposition is done
(Figure 5a), it is seen that the small domain has a slightly different
orientation in the four structures. This implies some flexibility of the
hinge between the two domains. The difference in orientation of the
small domains in the three independent molecules can be described as a
rotation of 2.3-3.2°*.* The rotations do not share a common axis, so the
movement is not a simple hinge motion. When these angles between the two
species are compared, larger rotation angles of 6.0-7.5° are observed. A
similar small change in relative orientation of the two folding domains
was also observed when two turnip cytochrome ***f*** structures at
different temperatures were compared (17).

In Gray\'s alignment ^2^, residues 1-177 of *C. reinhardtii* align with
1-177 of turnip, *C. reinhardtii* 179-187 align with turnip 178-186, *C.
reinhardtii* 189-199 with turnip 187-197, and *C. reinhardtii* 201-251
with turnip 200-250. However, when the backbone atoms of the small
domain are superimposed using Gray\'s alignment, residues 179-183 of the
algal protein do not superimpose on residues 178-182 of the turnip
protein. A much better fit is obtained superimposing residues *C.
reinhardtii* 178-182 onto turnip 178-182 (Figure 5b). This implies that
the β-strand 13 of the small domain aligns with same-numbered residues
of turnip and that both insertions in *C. reinhardtii* should be located
in the loop between the β-strands 13 and 14 (loop 1 in Figure 5b). This
loop is relatively disordered, as indicated by high temperature factors
(specially in monomer A) and large conformational differences between
these residues in the three different *C. reinhardtii* structures
(Figure 5b). Besides the conserved E186 (turnip) (E187 in *C.
reinhardtii*), this loop contains K187 (turnip) (K188 and K189 in *C.
reinhardtii* ). These lysines are believed to be involved in
plastocyanin binding (also see "Lysine cluster" section). The turnip
insertion is in the loop between the β-strands 14 and 15 in the small
domain (loop 2 in Figure 5b). This loop also appears to be flexible and
disordered in *C. reinhardtii*, specially in monomer B. The correct
alignment based on the structure is compared with Gray\'s alignment in
Figure 4.

Although the asymmetric unit contains three molecules of cytochrome
***f**, in vivo* the cytochrome would be part of a dimeric ***b~6~ f***
complex (13). The functionally homologous cytochrome ***c~1~*** forms
part of the dimer interface in the ***bc~1~*** complex (8). The three
monomers in the asymmetric unit are related by improper symmetry. The
choice of asymmetric unit is arbitrary, and was made to give a
relatively compact asymmetric unit. Because the crystal packing contacts
affecting the three monomers are different, those features that are the
same in all three monomers can be considered intrinsic features of the
cytochrome and not artifacts of packing forces. Those parts of the
structure that are different in the three monomers can be assumed to be
flexible, and the differences probably are due to packing forces. To
understand the crystal packing, and account for the different effects of
crystal contacts on the 3 monomers, a list of all inter-monomer contacts
closer than 3.5 Å was made ^3^. The analysis indicates that residues in
the front face of the heme binding pocket (Figure 6) Y1 of monomer A and
F4 of monomer B and C have contacts with other monomers inside or
outside the asymmetric unit. For residues of the Lys patch (see below),
only K58 of monomer B and K189 of monomer C have intermolecular
contacts. Monomer A shows more extensive involvement of the lysine
patch, but K58, K65 and K66 of monomer A bear no disturbances. It is
concluded that the model for the Lys patch is not much affected by
crystallographic contacts, whereas the one for the heme binding pocket
could be somewhat affected. Structural differences between the three
monomers are observed in the loop 1 region of the small domain (residues
185-190), see Figure 5b. Here monomers B and C are nearly the same
(Figure 5B), as expected from the involvment of this region in the
similar interactions (1) and (2) which involve S186 but not the charged
residues 187-189, while monomer A is somewhat different due to
involvement of K188 in interaction (3). The turnip structure is
different in this region due to insertion of residues L185 and K188 in
*C. reinhardtii.*

***Dimeric relation between two cytochrome f monomers.*** Although the
three monomers in the chosen asymmetric unit are related by improper
symmetry, the relation between monomer B in one asymmetric unit and
monomer A in another is a distorted (176°) proper 2-fold rotation
(described as contact 5 in footnote 3). The possibility that this
represents a physiological dimeric state will be approached in the
discussion. The putative dimer is depicted in Figure 7, with the likely
position of the membrane shown assuming (for lack of other information)
that the cytochrome ***f*** dimer 2-fold is perpendicular to the plane
of the membrane.

***Heme ligation.*** As seen in the cytochrome ***f*** structure from
turnip (16), the heme is covalently bound to the protein by thioether
bonds through strictly conserved cysteine residues (Cys21 and Cys24, in
fingerprint peptide Cys-X-Y-Cys-His sequence in all c-type cytochromes).
The heme Fe is ligated by the 4 tetrapyrrole N, by a proximal histidine
group of H25 and by the α-amino group of the N-terminus residue Y1
(Figure 1). This type of coordination was shown for the first time in
turnip cytochrome ***f**.* The distances between Fe atom and Nδ of H25
or N of Y1 are 2.6Å, in both cases. In all three monomers, the aromatic
ring of Y1 lies approximately parallel to the plane of the heme ring
(Figures 1 and 6). The heme is almost completely buried in a hydrophobic
pocket (established by highly conserved residues such as Y1, P2, F4,
F237, G72, V74, I19, V20, A5, and P161) that shields the redox center
from the solvent (Figure 6). The only portions of the heme that are
exposed to the surface are the two propionic acid side chains (whose
negative charges are partially counterbalanced by the fully conserved
residue R156, see Figure 6) and one of its saturated vinyl groups,
barely seen at the left of Figure 6.

***Lysine cluster.*** Figure 8 compares the electrostatic potentials at
the molecular surfaces of cytochrome ***f*** from *C. reinhardtii* (A)
and from turnip (B) in their reduced form. Both basic and acidic patches
are seen. These charge clusters are natural candidates for sites
involved in the stabilization of the quaternary structure of the ***b~6~
f*** complex and in the (transient) association with the intermediary
electron transfer carriers (e.g. plastocyanin or cytochrome *c*~6~).

In the case of turnip cytochrome ***f**,* a prominent positively charged
region at the interface between the large and small domains that
contains the basic residues K58, K65, K66, K187, and R209 (16) has been
proposed by many *in vitro* studies, including: kinetic, biochemical,
and cross-linking studies (references in (31, 51); electrostatic
analysis (52-54); as well as NMR studies (51), to be the site of
electrostatic interaction with plastocyanin. The first four of these
residues are highly conserved in eukaryotic cytochromes ***f*** (25). On
the other hand, most of the acidic residues in two
structurally-conserved, closely associated and negatively-charged
patches on the surface of eukaryotic plastocyanins are also conserved.
These last patches are proposed to be the counterpart binding site for
cytochrome ***f*** (21, 55, 56).

In *C. reinhardtii* cytochrome ***f**,* the equivalent cluster of basic
residues, also contributed by both the large and small domains, would be
composed of K58, K65, K66, K188 and K189 (Figure 6). Notice that the Lys
patch and the heme binding pocket are separated from each other through
both a translation and a ca. 60° rotation along the molecule main axis.
The temperature factors for the side chains of those residues in all
three monomers are relatively high, indicating conformational
flexibility ^4^.

The largest difference in the electrostatic landscape around the heme
face and lysine patch is due to the presence of aspartate 68 in *C.
reinhardtii* where the turnip protein has alanine and most other
cytochromes ***f*** have a neutral residue. This results in the negative
(red) patch to the right of the negative patch around the heme
propionates in Figure 8A. It also results in the complete masking (at
the surface) of the positive charge due to arginine 156, seen as a blue
spot above the heme propionates in the turnip (Figure 8B) which is
absent in the *C. reinhardtii* electrostatic surface. Recalculation of
the *C. reinhardtii* surface charge after removing the charge from the
carboxylic oxygens of D68 (not shown) resulted in essentially the same
charge distribution as seen for turnip in this region.

The lysine cluster, as defined originally for turnip, could be extended
to K121, K122, and K217 in the microalga (Figure 6). The ζN of K58, K65,
K188, K189 and K121 are approximately in the same plane, whereas that of
K66 is not, as its lateral chain points out of the cluster (Figures 5b
and 6). This characteristic of K66 is also seen in the turnip structure
(entry 1CTM), see Figure 5b.

***Water chain.*** An internal chain of buried water molecules extending
from the heme ligand H25 to a position close to K66 has been found in
the cytochrome ***f*** structure from turnip (17). The authors proposed
that it could constitute a "proton wire" (57, 58) and that, as such, it
could be involved in a pathway of proton transfer from the Rieske
iron-sulfur protein, through cytochrome ***f**,* and into the
chloroplast lumen, following the oxidation of plastoquinol at the quinol
oxidation site.

In our structure for *C. reinhardtii*, the chain of five buried water
molecules previously reported in the turnip cytochrome ***f*** structure
(17) is seen in all three independent monomers. In addition two more
water molecules (W6 and W7 in Table 2) can be identified in each
monomer, which extend the water chain all the way up to ζN of K66, a
surface residue that, *in situ*, is believed to be in contact with bulk
water and interacting with plastocyanin (Figure 9a). The locations of
the five buried water molecules are identical and their hydrogen bonding
network is almost identical to the respective information reported for
the turnip cytochrome ***f*** structure (17). The two additional waters
correspond to water residues 107 and 135 of the turnip structure. The
hydrogen bonding network of the water chain is shown in Figure 9b and
the refined parameters and residue numbers in the submitted coordinates
file 1cfm are listed in Table 2. In comparison with the turnip structure
1HCZ, the OD1 and ND2 atoms of water ligands N153 and N168 are
interchanged in the present structure based on H-bond analysis by the
verification program \"what-check\", however as these atoms cannot be
distinguished by the electron density, the difference may not be real.

The exposure of these molecules to the solvent is partial for W5 and
complete for W6 and W7; all three of them being less ordered than W1-W4,
which are completely buried. There are no other completely buried waters
in the cytochrome. Water molecules W5-W7 are visible on the surface in a
space filling model, however they are not seen in the orientation of
Figure 6. The inter-water hydrogen bonding chains are broken between W5
and W6 (distances are 4.0, 3.8 and 4.1 Å in monomers A, B and C,
respectively); however, the hydrogen bond chain is complete through
hydrogen bondings with the P232 carbonyl oxygen.

These water sites have relatively low temperature factors (a parameter
expressing the dynamic vibrations of an atom as well as its static
disorder in the crystal) and each water\'s position is within 0.2 Å of
the average position for that water in the three independent monomers.
In order to compare the degree of order of W1-W7 with the rest of the
water molecules in the asymmetric unit, we analyzed the distribution of
temperature factors of the whole population (1001 waters in identified
positions) ^5^. This analysis indicates that the degree of localization
of all these seven waters, although showing levels lower than the mean
and median, is also shared by many other water molecules in the crystal.

**DISCUSSION**

Cytochrome ***f*** is a subunit of the cytochrome ***b~6~ f*** complex,
a fundamental redox enzyme in oxygenic photosynthetic electron
transport. The folding and structural features described for turnip
cytochrome ***f*** (16, 17) were confirmed here for 3
non-crystallographic-related monomers of cytochrome ***f*** purified
from the microalga *C. reinhardtii* that, due to its genetic
amenability, is the preferred model organism for eukaryotic
photosynthesis. While significant differences are observed between the
turnip and the algal cytochromes in loops and at points of insertion or
deletion in the sequence, they are not much greater than the ones found
between the structures of the three monomers for the algal cytochrome
which, being from the same crystal, were obtained under identical
conditions of ionic strength, pH, and buffer composition. The two
insertions and one deletion in the amino acid sequence of the algal
protein occur at turns between β-strands and do not affect the overall
folding. The insertion of one Lys does affect the composition of the Lys
cluster (see below), which might be of functional significance to modify
specificity for the different electron acceptor partners.

The main structural characteristiscs of cytochrome ***f*** that are
currently discussed in relationship with function are: a) the conserved
residues of the heme pocket and the unusual heme Fe ligation by α-amino
group of tyrosine 1, b) the cluster of lysine residues that is proposed
to be the docking site of plastocyanin, and c) the chain of 7 water
molecules bound to conserved residues that extends between the heme
pocket and K58 and K66 at the lysine cluster.

***The front face of the heme binding pocket*** . The significance of
the conserved residues in the N-terminal end of the protein (25) has
been tested with site-directed mutagenesis in *C. reinhardtii* (32).
Mutants of Y1 or P2, showed normal cytochrome ***f*** photooxidation but
with its reduction inhibited in various degrees. This indicated that
residues 1 and 2 in the front face of the heme pocket play a direct or
indirect role in the redox equilibration with the FeS cluster in the
Rieske subunit, but not with plastocyanin (32). These results are in
contradiction with simulations (52, 53), and the NMR resolution (51) of
the plastocyanin/cytochrome ***f*** complex configuration. All these
works show Y1 as involved in the electron transfer path between both
molecules. The paradox will be solved with the structural resolution of
the cytochrome ***b~6~ f*** complex.

***Flexibility in the orientation of the small and large domains and of
the lateral chain at the conserved lysine patch.*** The lysine patch in
*C. reinhardtii* involving K58, K65, K66, K188 and K189 is very similar
to the one in turnip. Nevertheless, the following differences are found
between both proteins*.* Considering that in turnip the span 202 to 221
shows a single basic residue (R209) while in *C. reinhardtii* in the
span 190 to 216 there is a single basic residue (K207), it can be
proposed that K207 is the charged residue counterpart of R209.
Nevertheless, K207 in *C. reinhardtii* is not associated with the Lys
cluster described above (Figure 5b and 6), in contrast with the case of
R209 for turnip (16). Besides, K121 and K217 which could be proposed as
participants in the Lys cluster in *C. reinhardtii* (see Figure 6), are
replaced by Q121 and S216 in turnip, respectively. These subtle
differences in the Lys cluster in terms of charges and positions due to
mutations, as well as insertions and deletions (Figures 4 and 5b), could
be responsible for some differences in the electron partner specificity
of the two species. For example, the *C. reinhardtii* cytochrome ***f***
can interact with both plastocyanin and cytochrome c~6~ as mobile
electron acceptors (22, 31), whereas higher plant cytochrome ***f***
interacts *in situ* only with plastocyanin (22).

The role of these conserved residues in the redox reaction has been
studied in *C. reinhardtii* by site directed mutagenesis. With a series
of mutants, where K58, K65, K66, K188 and K189 have been replaced mainly
by polar residues or Glu, it has been shown that, in contrast with what
was expected, the complete neutralization of the charges in the Lys
cluster does not affect the kinetics of photooxidation of cytochrome
***f*** at pH 7 in uncoupled intact cells (31, 49) although reactions in
permeabilized cells (49, 59) and *in vitro* (60) are inhibited. The *in
vivo* results are also in disagreement with theoretically predicted
(52-54) and NMR resolved (51) structures of the reactive complex. The
reason for the discrepancy is unsettled.

The flexibility shown by these Lys lateral chains in the crystal is
still consistent with this cluster being involved in the plastocyanin
binding site, because a conformationally mobile area may be instrumental
in allowing "induced fit" to different electron acceptors (plastocyanin
or cytochrome *c*6) or for guiding the approach of them from different
orientations, see below. Decreased mobility of these residues upon
complexation could be expected, as seen in other protein complexes (61).

The 3 monomers show flexibility in the relative position of the
constituting large and small domains. The interface between these
domains involves a patch of Lys (putative binding site for plastocyanin)
with relatively mobile side chains. This flexibility might be needed to
accommodate different electron transfer partners, such as plastocyanin
or cytochrome c~6~, and to achieve a conformational rearrangement after
the initial docking (62). It can also be interpreted in terms of the
"Velcro" model of a binding site (63-65) According to this model there
is not one unique specific lock-and-key interaction, but rather the two
reacting molecules can interact through multiple different
configurations of similar energy. Such a dynamic configuration of the
binding between plastocyanin and cytochrome ***f*** has been deduced by
NMR (51). The interdomain flexibility could be important in the
mechanism of interaction of cytochrome ***f*** not only with its oxidant
partner(s) but also, in alternation, with the Rieske protein, involving
an oscillatory catalytic mechanism analogous to the one proposed in (8).
Finally, the interdomain flexibility could have a role in the correct
association of cytochrome ***f*** with other interfacing subunit(s) of
the cytochrome ***b~6~ f*** complex.

***An alternative structural role for the water array.*** As in turnip,
an array of buried waters is also present in *C. reinhardtii* cytochrome
***f***. A proton wire function has been proposed for this chain (17).
The residues participating in side-chain hydrogen bonds such as Q59,
N153, Q158, N168, and N233 (N232 in turnip) are strictly conserved in
all cytochromes ***f*** whose sequences are known (25). Furthermore,
those residues participating in backbone hydrogen bonds are also
strictly conserved in all cytochromes ***f*** with only one exception,
K58 which is replaced by Q in cyanobacteria (25).

Similar chains of hydrogen-bonded water molecules have been previously
observed in bacteriorhodopsin (66) and in a bacterial photosynthetic
reaction center (67, 68) and have been proposed to function as proton
wires. Nevertheless, the proposed proton translocation function of this
conserved water array in cytochrome ***f*** has not been yet
demonstrated nor, in principle, expected, as protons generated at the
quinol oxidizing site should have shorter pathways to reach the lumen
than one involving, in series, the Rieske and cytochrome ***f***
proteins. Mutational analysis of the residues associated with the
cytochrome ***f*** water array has not yield a unique answer yet (69).

In addition to the proton wire function, another purely structural
(hydrogen bonding) role can also be postulated. The analysis of the
structural data shown in Figures 2 and 8b is consistent with the
possibility of a structural role in the large domain. Most
significantly, due to their complete or partial enclosure in the protein
body and low temperature factors, W1, W2, W3, W4 and W5 make hydrogen
bonds between some of themselves and between stretches of residues that
are far away in sequence. In one case, they relate at the same time with
residues placed 80 positions apart, i.e. 153-156, 158, and 168 with
232-236. Consequently, one end of the β-strands S9 and S19, the loop
inter S18-S19, the loop between S9-S10 and the loop between S11 and S12
are connected by an abundant network of water-dependent hydrogen bonds.
In the other case, W1 and W2 establish bonds with the loop inter S1-H2
that is close to the N-terminal portion, more than 100 residues apart
from S9. We can consider this structure as a "core" contributed by
different secondary structure portions of the protein that is
stabilized, at least in part, by hydrogen bonds mediated by waters.
Probably less significant, due to their higher temperature factors and
exposure, W6 and W7, with W4 and W5, connect to the core domains S4 and
S5, which bear the conserved K58, K65 and K66. Other cases of buried
water molecules, either single or in arrays, are known (70-73). Even
water molecules bound to the protein surface can have a role in protein
folding, stability, recognition and activity (74).

Another explanation for the presence of the internal water array would
consider these waters as resulting from their entrapment during the
folding process. Those waters, without any functional role, would be
stabilized by circumstantial hydrogen bondings to the indicated
residues, which would have been conserved for other reasons.

***Putative in situ dimeric configuration of cytochrome f*** . The
physiological state of the membrane-embedded cytochrome ***b~6~ f***
complex is a dimer (9-12, 14), and the functionally homologous
cytochrome ***c~1~*** is involved in the dimer contact in the
mitochondrial cytochrome ***bc~1~*** complex (8). If cytochromes ***f***
of the dimeric ***b~6~ f*** complex also contribute to the dimer
interface, cytochrome ***f*** may have an affinity for itself which
could be revealed in the crystal packing. Of the seven different types
of inter-monomeric contacts observed in the crystal, the only one which
approximates 2-fold symmetry is that described as contact 5 in footnote
3. This association is the most intimate of the inter-monomer contacts,
with 3033 Å^2^ buried in dimer formation (about 1500 Å^2^, or more than
10% of the monomer surface area, is buried on each monomer). It is thus
tempting to speculate that this association between two monomers in the
crystal represents the arrangement of monomers in the dimeric cytochrome
***b~6~ f*** complex, as suggested by Figure 7.

This contact involves both large and small domains, in fact the
\"necks\" between the domains are interlocked in a way that provides
considerable rigidity to the putative dimer. The lysine patch and the
hydrophobic heme face (likely to be involved in interaction with
plastocyanin) are exposed. A solution structure for the complex of
poplar plastocyanin with turnip cytochrome ***f*** is available (51)
(PDB entry 2PCF). As illustrated in Figure 7, it is possible for
plastocyanin to bind either monomer, in the manner revealed by this NMR
structure, without interference from the other monomer. We assume that
the dimer 2-fold axis would be perpendicular to the membrane as
indicated in Figure 7. Both C-termini of the truncated proteins are
close to the membrane surface in this model, as expected if the
truncated portion is a transmembrane anchor.

In this model the normal to the heme plane is 79° from the dimer 2-fold
axis, therefore the heme plane is 79° from the plane of the membrane.
This is similar to the value (72°) calculated for cytochrome ***c~1~***
from structure 1BCC. However this is inconsistant with values around
25-30° estimated by epr spectroscopy of oriented membranes (75-77)**.**
Thus unless there is some artifact in the epr measurements, our dimer
from the crystal is probably not a good model for the cytochromes
***f*** in the dimeric cytochrome ***b~6~ f*** complex.

***Perspectives.*** In spite of functional similarities, the structure
of cytochrome ***f*** is different from that of mitochondrial cytochrome
***c~1~*** which is mainly made of α-helices and has different heme
ligation (8). Because they share a low degree of sequence identity and
have different cellular origin (cytochrome ***f*** is organelle encoded
and cytochrome ***c~1~*** is nuclear encoded in eukaryotes), they might
represent a convergent functional evolution (78).

As cytochrome ***f*** sequences are highly conserved across eukaryotes
and cyanobacteria (25), the structures presented here together with
those from turnip (17) and cyanobacteria (79) should serve as a
prototype for the main structural features of all cytochromes ***f**.*
Small differences could be attributed to functional specificities or
evolutionary drift.

Free subunits of macromolecular complexes, although easier to
crystallize than the entire complex, may have different structural and
physicochemical characteristics than *in situ*, where a variety of
interactions with other domains exists. Nevertheless, these results will
allow structure/function studies with solid structural foundation, and
will help in the resolution of the full cytochrome ***b~6~ f*** complex
structure, once suitable crystals are obtained.

**ACKNOWLEDGMENT**

We would like to thank Dr. F-A. Wollman (Institut de Biologie
Physico-Chimique, Paris) for providing strain F283ST; to Prof. S-H. Kim
(Calvin Lab, UC-Berkeley) for the use of his crystallographic facilities
and for financial support to Y-I. C; to Ms. A. Reyes and Mr. R. Solem
for assistance in growing the microalga; to Mr. T. Linden for assistance
in cytochrome ***f*** purification and crystallization, and to Mr. L. R.
Comolli for helpful discussions. J.G.F-V thanks Prof. R. Malkin
(Department of Plant and Microbial Biology, UC-Berkeley) for generously
providing laboratory facilities and financial support, and for
critically reading the manuscript.

**REFERENCES**

1\. Degli Esposti, M., DeVries, S., Crimi, M., Ghelli, A., Patarnello,
T., and Meyer, A. (1993) *Biochim. Biophys. Acta* *1143*, 243-271.

2\. Trumpower, B. L., and Gennis, R. B. (1994) *Annual Review of
Biochemistry* *63*, 675-716.

3\. Hope, A. B. (1993) *Biochim. Biophys. Acta* *1143*, 1-22.

4\. Gray, K. A., and Daldal, F. (1995) in *Anoxygenic Photosynthetic
Bacteria* (Blankenship, R. E., Madigan, M. T., & Bauer, C. E., Eds.) pp
747-774, Kluwer Academic Publishers, Dordrecht.

5\. Malkin, R. (1992) *Photosynthesis Research* *33*, 121-136.

6\. Hauska, G., Schutz, M., and Buttner, M. (1996) in *Oxygenic
Photosynthesis: The Light Reactions.* (Ort, D. R., & Yocum, C. F., Eds.)
pp 377-398, Kluwer Academic Publishers, Dordrecht.

7\. Xia, D., Yu, C. A., Kim, H., Xia, J. Z., Kachurin, A. M., Zhang, L.,
Yu, L., and Deisenhofer, J. (1997) *Science (Washington D C)* *277*,
60-66.

8\. Zhang, Z., Huang, L., Shulmeister, V. M., Chi, Y.-I., Kim, K. K.,
Hung, L.-W., Crofts, A. R., Berry, E. A., and Kim, S.-H. (1998) *Nature*
*392*, 677-684.

9\. Chain, R. K., and Malkin, R. (1991) *Photosynthesis Research* *28*,
59-68.

10\. Huang, D., Everly, R. M., Cheng, R. H., Heymann, J. B., Schagger,
H., Sled, V., Ohnishi, T., Baker, T. S., and Cramer, W. A. (1994)
*Biochemistry* *33*, 4401-4409.

11\. Chain, R. K., and Malkin, R. (1995) *Photosynthesis Research* *46*,
419-426.

12\. Breyton, C., Tribet, C., Olive, J., Dubacq, J. P., and Popot, J. L.
(1997) *J. Biol. Chem.* *272*, 21892-21900.

13\. Mosser, G., Breyton, C., Olofsson, A., Popot, J. L., and Rigaud, J.
L. (1997) *Journal of Biological Chemistry* *272*, 20263-20268.

14\. Bron, P., Lacapere, J. J., Breyton, C., and Mosser, G. (1999) *J.
Mol. Biol.* *287*, 117-126.

15\. Huang, D., Zhang, H., Soriano, G. M., Dahms, T. E. S., Krahn, J.
M., Smith, J. L., and Cramer, W. A. (1998) in *Photosynthesis:
Mechanisms and Effects* (Garab, G., Ed.) pp 1577-1581, Kluwer Academic
Publishers, Netherlands.

16\. Martinez, S. E., Huang, D., Szczepaniak, A., Cramer, W. A., and
Smith, J. L. (1994) *Structure (London)* *2*, 95-105.

17\. Martinez, S. E., Huang, D., Ponomarev, M., Cramer, W. A., and
Smith, J. L. (1996) *Protein Science* *5*, 1081-1092.

18\. Iwata, S., Saynovits, M., Link, T. A., and Michel, H. (1996)
*Structure (London)* *4*, 567-579.

19\. Zhang, H., Carrel, C. J., Huang, D., Sled, V., Ohnishi, T., Smith,
J. L., and Cramer, W. A. (1996) *Journal of Biological Chemistry* *271*,
31360-31366.

20\. Carrell, C. J., Zhang, H., Cramer, W. A., and Smith, J. L. (1997)
*Structure (London)* *5*, 1613-1625.

21\. Gross, E. L. (1996) in *Oxygenic Photosynthesis: The Light
Reactions.* (Ort, D. R., & Yocum, C. F., Eds.) pp 413-429, Kluwer
Academic Publishers, Dordrecht.

22\. Wood, P. M. (1978) *European Journal of Biochemistry* *87*, 9-19.

23\. Merchant, S., and Bogorad, L. (1986) *Molecular and Cellular
Biology* *6*, 462-469.

24\. Kerfeld, C. A. (1997) *Photosynthesis Research* *54*, 81-98.

25\. Gray, J. C. (1992) *Photosynthesis Research.* *34*, 359-374.

26\. Gray, J. C., Rochford, R. J., and Packman, L. C. (1994) *European
Journal of Biochemistry* *223*, 481-488.

27\. Rochaix, J.-D. (1995) *Annual Review of Genetics* *29*, 209-230.

28\. Davies, J. P., and Grossman, A. R. (1998) *J. Phycol.* *34*,
907-917.

29\. Delosme, R. (1991) *Photosynthesis Research* *29*, 45-54.

30\. Kuras, R., Wollman, F. A., and Joliot, P. (1995) *Biochemistry*
*34*, 7468-7475.

31\. Soriano, G. M., Ponamarev, M. V., Tae, G. S., and Cramer, W. A.
(1996) *Biochemistry* *35*, 14590-14598.

32\. Zhou, J., Fernandez-Velasco, J. G., and Malkin, R. (1996) *Journal
of Biological Chemistry* *271*, 6225-6232.

33\. Berry, E. A., Huang, L. S., Chi, Y., Zhang, Z., Malkin, R., and
Fernandez-Velasco, J. G. (1997) *Biophysical Journal* *72*, A125.

34\. Harris, E. H. (1989) *The Chlamydomonas Source Book. A
comprehensive Guide to Biology and Laboratory Use.*, Academic Press, San
Diego.

35\. Jancarik, J., and Kim, S.-H. (1991) *J. Appl. Cryst.* *24*,
409-411.

36\. Otwinowski, Z. (1993) in *Proceedings of the CCP4 Study Weekend:
\"Data Collection and Processing\"* (Sawyer, L., Isaacs, N., & Bailey,
S., Eds.) pp 56-62, SERC Daresbury Laboratory, Warrington, UK.

37\. CCP4:The SERC (UK) Collaborative computational project No.4 (1995)
*Acta Cryst.* *D50*, 760-763.

38\. Navaza, J. (1994) *Acta Cryst.* *A50,*, 157-163.

39\. BrŸnger, A. T. (1992) *X-PLOR manual version 3.1.*, Yale
University, New Haven, CT.

40\. Jones, T. A., Zhou, J.-Y., Cowan, S. W., and Kjeldgaard, M. (1991)
*Acta cryst.* *A47,*, 110-119.

41\. Laskowski, R. A., Macarthur, M. W., Moss, D. S., and Thornton, J.
M. (1993) *J. appl. crystallo.* *26*, 283-291.

42\. Kraulis, P. J. (1991) *Journal of Applied Crystallography.* *24*,
946-950.

43\. Merrit, E. A., and Murphy, M. E. P. (1994) *Acta Cryst.* *D50*,
869-873.

44\. Nicholls, A., Sharp, K. A., and Honig, B. (1991) *Proteins Struc.
Funct. Genet.* *11*, 281-296.

45\. Sayle, R. A., and Milner-White, E. J. (1995) *Trends in Biochemical
Sciences* *20*, 374-376.

46\. Laemli, U. K. (1970) *Nature* *227*, 680-685.

47\. Berry, E. A., and Trumpower, B. L. (1987) *Analytical Biochemistry*
*161*, 1-15.

48\. Fernandez-Velasco, J. G., Linden, T., Hung, T., Flores, J., Huang,
L.-s., and Malkin, R. (1998) *Biophysical Journal* *74(2)*, A78.

49\. Comolli, L. R., Zhou, J., Linden, T., Breitling, R., Flores, J.,
Hung, T., Jamshidi, A., Huang, L.-s., and Fernandez-Velasco, J. G.
(1988) in *Photosynthesis: Mechanisms and Effects* (Garab, G., Ed.) pp
1589-1592, Kluwer Academic Publishers, Netherlands.

50\. Metzger, S. U., Cramer, W. A., and Whitmarsh, J. (1997) *Biochimica
et Biophysica Acta* *1319*, 233-241.

51\. Ubbink, M., Ejdeback, M., Karlsson, B. G., and Bendall, D. S.
(1998) *Structure* *6*, 323-335.

52\. Pearson, D. C., Jr., Gross, E. L., and David, E. S. (1996)
*Biophysical Journal* *71*, 64-76.

53\. Soriano, G. M., Cramer, W. A., and Krishtalik, L. I. (1997)
*Biophysical Journal* *73*, 3265-3276.

54\. Ullmann, G. M., Knapp, E.-W., and Kostic, N. M. (1997) *J. Am.
Chem. Soc.* *119*, 42-52.

55\. Gross, E. L. (1993) *Photosynthesis Research* *37*, 103-116.

56\. Redinbo, M. R., Yeates, T. O., and Merchant, S. (1994) *Journal of
Bioenergetics and Biomembranes* *26*, 49-66.

57\. Nagle, J. F., and Morowitz, H. (1978) *Proc. Natl. Acad. Sci. USA.*
*75*, 298-302.

58\. Pomes, R., and Roux, B. (1996) *Biophysical Journal* *71*, 19-39.

59\. Fernandez-Velasco, J. G., Zhou, J., and Malkin, R. (1997)
*Biophysical Journal* *72*, A126.

60\. Soriano, G. M., Ponamarev, M. V., Piskorowski, R. A., Krishtalik,
L. I., and Cramer, W. A. (1998) *Biophys. J.* *74*, A41.

61\. Edwards, K. J., Ollis, D. L., and Dixon, N. E. (1997) *Journal of
Molecular Biology* *271*, 258-265.

62\. Qin, L., and Kostic, N. M. (1993) *Biochemistry* *32*, 6073-6080.

63\. McLendon, G. (1991) in *Metal Ions in Biological Systems.* (Sigel,
H., & Sigel, A., Eds.) pp 183-198, M Dekker, New York.

64\. McLendon, G. (1991) *Struct. Bond.* *75*, 159-174.

65\. McLendon, G., and Hake, R. (1992) *Chem. Rev.* *92*, 481-490.

66\. Cao, Y., Chang, V. M., Ni, B., Needleman, R., and Lanyi, J. K.
(1991) *Biochemistry* *30*, 10972-10979.

67\. Ermler, U., Fritzsch, G., Buchanan, S. K., and Michel, H. (1994)
*Structure (London)* *2*, 925-936.

68\. Fritzch, G., Kampmann, L., Kapaun, G., and Michel, H. (1998)
*Photosynthesis Research* *55*, 127-132.

69\. Ponamarev, M. V., and Cramer, W. A. (1998) *Biophys. J.* *74*, A41.

70\. James, M. N. G., and Sielecki, A. R. (1983) *J. Mol. Biol.* *163*,
299-361.

71\. Oefner, C., and Suck, D. (1986) *J. Mol. Biol.* *192*, 605-632.

72\. Jeffrey, G. A. (1991) *Hydrogen Bonding in Biological Structure*,
Springer-Verlag, Berlin, Germany.

73\. Meyer, E. (1992) *Protein Science* *1*, 1543-1562.

74\. Schoenborn, B. P., Garcia, A., and Knott, R. (1995) *Progress in
Biophysics and Molecular Biology* *64(2-3)*, 105-19.

75\. Bergstršm, J., and VŠnngard, T. (1982) *Biochim. Biophys. Acta*
*682*, 452-456.

76\. Crowder, M. S., Prince, R. C., and Bearden, A. (1982) *FEBS Lett.*
*144*, 204-208.

77\. Riedel, A., Rutherford, A. W., Hauska, G., MŸller, A., and
Nitschke, W. (1991) *J. Biol. Chem.* *266*, 17838-17844.

78\. Prince, R. C., and George, G. N. (1995) *Trends Biochem. Sci.*
*20*, 217-218.

79\. Carrell, C. J., Schlarb, B. G., Bendall, D. S., Howe, C. J.,
Cramer, W. A., and Smith, J. L. (1999) *Biochemistry* *38*, 9590-9599.

**[Table 1. Data processing and refinement statistics]{.underline}**

[Data statistics:]{.underline}

Resolution Reflections Redundancy Data coverage(%) I/σ Rsym(%)

2.0Å 47,961 3.3 (2.8) 91.6 (68.7) 14.8 (3.5) 4.2 (17.6)

[Refinement Statistics:]{.underline}

Resolution range 25.0 - 2.0 Å

Number of non-H Protein Atoms 5,877

Number of solvent molecules 1,001

R factor for 44561 reflections (F\>2σ, 95%) 21.4 % (24.1 %)

Rfree value for 2346 reflections (F\>2σ, 5%) 27.1 % (31.7 %)

rms deviation from ideal geometry

Bond lengths 0.006 Å

Bond Angles 1.5°

Dihedral angles 24.3°

Average temperature factors

Main chain atoms only 23.8 Å2

All protein atoms (excluding water) 24.8 Å2

Water molecules only 36.6 Å^2^

\_\_\_\_\_\_\_\_\_\_\_\_\_\_\_\_\_\_\_\_\_\_\_\_\_\_\_\_\_\_\_\_\_\_\_\_\_\_\_\_\_\_\_\_\_\_\_\_\_\_\_\_\_\_\_\_\_\_\_\_\_\_\_\_\_\_\_\_

Values in parentheses correspond to the highest-resolution shell
(2.13-2.0Å)

**Table 2.** Distance and temperature factor (*B*) information of each
atom participating in the direct hydrogen bonding network involving
water molecules W1-W7 in each of the three monomers residing in the
asymmetric unit. For each monomer the residue numbers that identify
water molecules W1-W7 in the submitted coordinates file 1CFM are shown
in parenthesis.

[Water O Atom]{.underline} [H-bonding Atom]{.underline}

*B* factor (Å2) Distance (Å) *B* factor (Å2)

(residue number)

\\Monomer Monomer Monomer

Water A B C Partner A B C A B C

**W1** 14.1 12.6 17.8 W2 2.9 2.8 2.6 11.6 13.9 20.2

(1041 1201 1249) Q158 OE1 2.7 2.9 3.1 10.9 12.6 16.7

N233 ND2 2.9 3.0 3.0 12.8 12.8 20.2

A27 N 3.2 3.1 3.0 16.0 12.7 26.5

L26 N 3.6 3.5 3.7 17.3 12.6 23.9

**W2** 11.6 13.9 20.2 W1 2.9 2.8 2.6 14.1 12.6 17.8

(1040 1333 1435) W3 2.8 2.9 2.7 10.6 9.5 12.1

H25 ND1 2.5 2.4 2.4 23.7 14.4 23.5

N153 OD1 2.5 2.5 2.6 11.2 15.6 20.4

G155 O 3.1 3.5 3.5 11.7 10.6 15.7

R154 O 3.1 3.2 3.3 16.8 15.1 19.4

G236 N 3.3 3.3 3.4 10.8 14.0 19.4

**W3** 10.6 9.5 12.1 W2 2.8 2.9 2.7 11.6 13.9 20.2

(1155 1030 1224) W4 2.8 2.9 2.8 14.2 9.5 17.7

R156 O 2.7 2.9 2.8 14.7 13.2 16.2

R156 N 3.4 3.4 3.4 13.8 13.5 16.6

G155 O 3.4 3.5 3.5 11.7 10.6 15.7

N233 OD1 2.5 2.5 2.7 12.5 13.0 20.3

G235 N 3.5 3.5 3.6 13.6 12.0 20.2

**W4** 14.2 9.5 17.7 W3 2.8 2.9 2.8 10.6 9.5 12.1

(1015 1054 1018) W5 3.4 3.4 3.5 13.6 13.0 19.0

Q59 OE1 3.0 2.9 2.9 14.4 14.0 19.5

N233 OD1 2.9 3.0 2.9 12.5 13.0 20.3

N168 ND2 3.1 3.1 3.1 15.7 16.8 19.7

V234 N 3.4 3.4 3.4 13.8 12.1 20.3

G235 N 3.3 3.2 3.0 13.6 12.0 20.2

**W5** 13.6 13.0 19.0 W4 3.4 3.4 3.5 14.2 9.5 17.7

(1039 1059 1274) P232 O 2.7 2.5 2.7 13.0 12.9 22.4

N168 OD1 2.6 2.5 2.5 16.8 16.1 23.6

V60 N 2.8 2.7 2.7 15.7 15.9 21.0

**W6** 14.8 28.9 30.4 W7 3.0 2.5 2.8 26.5 33.9 29.3

(1377 1749 1517) P232 O 2.7 2.7 2.6 13.0 12.9 22.4

**W7** 26.5 33.9 29.3 W6 3.0 2.5 2.8 14.8 28.9 30.4

(1378 1730 1314) K58 O 2.5 2.6 2.6 18.0 21.0 26.0

[K66 NZ 3.0 2.9 2.9 35.6 29.2 40.4]{.underline}

**FIGURE LEGENDS**

**Figure 1.** Omit map of the heme region. The electron density map is
made with coefficients of Fo-Fc and Φc, where Fc and Φc are amplitudes
and phases calculated from the final refined model after deleting the
heme and residues Y1 and H25 to avoid model bias in the heme region. The
map is contoured at the 3 σ level.

**Figure 2.** Stereo ribbon diagram of cytochrome ***f*** from *C.
reinhardtii* (Raster 3D (43) rendered). In the wild type the C-terminus
would be continuous forming a membrane spanning α-helix and a short
cytoplasmic segment.

**Figure 3.** Schematic drawing of the secondary structure arrangement
in the *C. reinhardtii* truncated cytochrome ***f**.*

**Figure 4.** Sequence alignment of *C. reinhardtii* (Chlamy) with
turnip cytochrome ***f*** based on Gray\'s alignment (25), compared with
alignment based on the structure (this work). In the latter, symbols
between the lines indicate the closeness of superposition of the aligned
residues, using the *C. reinhardtii* monomer B structure, turnip
structure (entry 1CTM), and *Phormidium laminosum* structure (entry
1CI3) superimposed as described in the legend of Figure 5b: (\|) Cα
positions differ by less than 1.0 Å after alignment. (×) Cα positions
differ by 1.0 Å or more. Insertions are placed to minimize the distance
between Cα\'s of aligned residues. The asterisk indicates the totally
conserved carboxylate E186(turnip), E187(*C. reinhardtii*), and D187
(25).

**Figure 5.** Comparison of the backbone representations of the three
ncs-related monomers of *C. reinhardtii* (A, B, and C) and of turnip
cytochrome ***f**.* The molecules were superimposed so as to minimize
the rms distance between corresponding Cα. The "loop 1" and "loop 2"
regions are discussed in the text.

a\. The backbone of the full protein- Residues of the large domain 1-13,
18-169, and 231-248 of *C. reinhardtii* were matched with residues 1-13,
18-169, and 230-247 of turnip, respectively. For these 183 residues, the
rms deviation in Cα position between the turnip cytochrome and the
monomer B of *C. reinhardtii* cytochrome was 0.5 Å. The corresponding
deviation of monomer A and monomer C with monomer B in *C. reinhardtii*
were 0.3 Å, in both cases.

b\. Detail of the small domain- Residues of the small domain 176-182,
191-197, and 204-222 of *C. reinhardtii* were matched with residues
176-182, 189-195, and 203-221 of turnip, respectively. For these 33
residues, the rms deviation in Cα position between the turnip cytochrome
and the monomer B of *C. reinhardtii* cytochrome was 0.4 Å. The
corresponding deviation of monomer A and monomer C with monomer B in *C.
reinhardtii* were 0.2 Å, in both cases. Side chains for residues in the
basic patch (putative plastocyanin binding site) are shown (with CPK
coloring code) for the turnip structure and for monomer B of *C.
reinhardtii* identified by T and C, respectively*.*

**Figure 6.** Space-filling Rasmol (45) representation of the central
part of the *C. reinhardtii* truncated form of cytochrome ***f***
(monomer B). The hydrophobic outer face of the heme binding pocket
(involving the N-terminus region) and the Lys cluster (putative
plastocyanin binding site) are shown, see text. The top and base of the
figure are areas distal and proximal to the membrane, respectively.
Color code: blue, Lys; gray-blue, Arg; pink, Glu; magenta, Asp; yellow,
12 aliphatic, 4 Gly and 4 Pro residues; green, aromatic; white, polar;
red, heme (only the two propionate chains are clearly seen in this
view). Due to the molecule orientation, exposed water molecules W5-W7
are not seen. The Lys ζN groups are identified with a star. Notice that
K66 ζN is not in the same plane with the other ζN's in the cluster and
is almost not seen in this view. This figure encompasses approximately
2/3 of the large domain (bottom) and 1/2 of the small domain (top).

**Figure 7.** Dimer-like noncrystallographic symmetry observed in the
crystal of *C. reinhardtii* cytochrome *f*. Above is a \"top\" view,
looking down the 2-fold axis of symmetry. Below is a side view, looking
parallel to a hypothetical membrane plane constructed perpendicular to
the 2-fold (orange). The blue and yellow space-filling models are
monomers A and B from the crystal of *C. reinhardtii* cytochrome
***f***. The hemes are shown as red space-filling models. The rectangles
below the C termini in the lower panel represent the lumenal ends of the
transmembrane helices. The magenta backbone drawings are plastocyanin
from the NMR structure of the plastocyanin-cytochrome ***f*** complex
(2PCF), oriented with the operators that best superimpose cytochrome
***f*** of that structure on each monomer of the *C. reinhardtii* dimer.
The copper atom of plastocyanin is shown as a green sphere. The
interactions betwen these two monomers in the crystal may be similar to
those involved in stabilizing a physiological dimer in vivo (see text).
In this figure monomer B is taken directly from the structure 1CFM;
monomer A is transformed by crystallographic symmetry operator \[1/2+x,
1/2-y, 1-z\] because the chosen asymmetric unit does not include this
dimer.

**Figure 8.** Electrostatic potentials at the molecular surface of the
reduced forms of cytochromes ***f*** from *C. reinhardtii* (monomer B)
(A) and from turnip (coordinates from 1CTM) (B). Two views rotated 180°
around the longest dimension of the molecules are shown. The orientation
of the molecules at the left panel is similar to the one shown in Figure
6. The surface electrostatic potentials are calculated by the program
GRASP (44) and displayed as a color gradient from red (electronegative,
[\<]{.underline} -5 kT/charge) to blue (electropositive,
[\>]{.underline}+5 kT/charge). The uncharged hydrophobic environment
around the heme pocket and the overall positive potentials of the lysine
cluster, with some differences between cytochromes ***f*** from two
species, are seen. The dark stick figures indicate the location and
orientation of the heme moiety, mainly buried in the protein (see Figure
6) with only the propionate groups exposed, if observed in the left view
(see Figure 6), and one vinyl group exposed, if observed in the right
view. In calculating electrostatic potentials, hydrogen atoms were
omitted and their charge included in the charge of the atom to which
they are bound (the default for GRASP). A charge of +1 was used for the
Nζ atoms of Lys, +0.5 for Nη1 and Nη2 of Arg, -0.5 for both carboxylate
oxygens of Asp, Glu and the heme propionates. All other atoms of the
heme were considered neutral, which is appropriate for the reduced heme.

**Figure 9.** The "water chain" A) Locations of the seven water
molecules forming the water chain (W1-W7, numbered green spheres; see
text) are shown relative to the whole molecule. Residues H25 and K66, at
the ends of the water chain, as well as the heme moieties are indicated
in a ball-and-stick representation. B) Schematic representation of the
hydrogen bonding network directly involving the buried (W1-W4) and
exposed (W5-W7) water molecules of cytochrome ***f*** from *C.
reinhardtii*. Side chains are not shown for those residues whose
backbone atom is forming a hydrogen bond. All the residues shown in this
network are strictly conserved in cytochromes ***f*** from *C.
reinhardtii* and higher plants (25).