From the Max-Planck-Institut für Biochemie,
Abteilung Strukturforschung, Am Klopferspitz 18a, 82152 Planegg-Martinsried, Germany,
Max-Planck-Institut für Biochemie,
Abteilung Molekulare Strukturbiologie, Am Klopferspitz 18a, 82152 Planegg-Martinsried, Germany, and ¶ Macromolecular Structure
Laboratory, NCI-Frederick Cancer Research and Development Center,
Frederick, Maryland 21702
Received for publication, May 26, 2000, and in revised form, October 24, 2000
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() |
---|
cDNA of Capsicum annuum Yolo
Wonder (paprika) has been prepared from total cellular RNA, and
the complete gene encoding paprika ferredoxin-NADP+
reductase (pFNR) precursor was sequenced and cloned from this cDNA.
Fusion to a T7 promoter allowed expression in Escherichia coli. Both native and recombinant pFNR were purified to
homogeneity and crystallized. The crystal structure of pFNR has been
solved by Patterson search techniques using the structure of spinach ferredoxin-NADP+ reductase as search model. The structure
was refined at 2.5-Å resolution to a crystallographic
R-factor of 19.8% (Rfree = 26.5%). The overall structure of pFNR is similar to other members of
the ferredoxin-NADP+ reductase family, the major
differences concern a long loop (residues 167-177) that forms part of
the FAD binding site and some of the variable loops in surface regions.
The different orientation of the FAD binding loop leads to a tighter
interaction between pFNR and the adenine moiety of FAD. The
physiological redox partners [2Fe-2S]-ferredoxin I and
NADP+ were modeled into the native structure of pFNR. The
complexes reveal a protein-protein interaction site that is consistent
with existing biochemical data and imply possible orientations for the
side chain of tyrosine 362, which has to be displaced by the nicotinamide moiety of NADP+ upon binding. A reasonable
electron transfer pathway could be deduced from the modeled structures
of the complexes.
Ferredoxin-NADP+ reductase
(FNR)1 belongs to a family of
flavoproteins found in higher plants, eukaryotic algae, and
photosynthetic bacteria (1). This flavoprotein is the last enzyme in
the electron transport chain of linear photosynthesis, where electrons
are transferred through a series of electron carriers and finally produce NADPH. FNRs have also been identified in various tissues and
organisms not capable of photosynthesis, where FNR is involved in
nitrogen fixation and steroid hydroxylation (2). In photosynthesis, FNR
catalyzes the reduction of NADP+ to NADPH according to the
reaction,
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS AND DISCUSSION
REFERENCES
which can be divided into two half-reactions. For catalyzing this
reaction, FNR utilizes the noncovalently but strongly bound prosthetic
group FAD as the only redox center. In the first step, FNR catalyzes
the successive transfer of two electrons from each molecule of
the reduced one-electron carrier [2Fe-2S]-ferredoxin I (Fd) to a
single molecule of FAD. In the second step, FNR utilizes these two
electrons to convert NADP+ into NADPH via hydride
(H
) transfer from N5 of FAD (1).
The three-dimensional structures of the oxidized and reduced form of
native spinach FNR and of the complexes with the competitive inhibitor
2'-phospho-5'-AMP have been determined by Bruns and Karplus (3, 4) as
first structures for this family of flavoproteins. In 1996, Serre
et al. (5) reported the structure of Anabaena PCC
7119 FNR and that of a complex with NADP+. None of these
studies showed a complex characteristic for productive electron
transfer between FNR and NADP+. More recently, crystal
structures of the complexes between two mutants of pea FNR (Y308S and
Y308W) and NADP+ have been solved that reveal a single
productive NADP+ binding mode (6). FNR consists of two
distinct domains, one responsible for binding the prosthetic group FAD
and the other for NADP+ binding. The FAD-binding domain is
a six-stranded antiparallel -barrel, which is capped at one side by
an
-helix, while the NADP+ binding domain consists of a
five-stranded parallel
-sheet flanked by
-helices. This unique
structural two-domain motif has been proposed to be a prototype for a
large family of flavoproteins (7). Structurally characterized members
of this family involve phthalate dioxygenase reductase (PDR) (8), the
FAD-containing fragment of NADPH-dependent nitrate
reductase (9), cytochrome b5 reductase (10),
NADP-cytochrome P450 reductase (11), and flavodoxin reductase (12).
While all members show the characteristic two-domain FNR motif, some
members like PDR also contain extra domain(s) to extend their catalytic capability.
Electron transfer reactions involving protein-protein interactions require the formation of a transient complex that brings together the two redox centers exchanging electrons. Extensive biochemical studies revealed the involvement of electrostatic interactions in complex formation between FNR and its redox partner Fd (13) and implied a model for complex formation (14). In the proposed complex, FNR is predominantly positively and Fd is predominantly negatively charged. It was suggested that FNR and Fd are initially steered toward each other via complementary charge interactions of the molecular dipoles. Subsequently, interaction of basic residues on the surface of FNR and acidic residues on the surface of Fd are thought to help to attain an optimal orientation. Finally, short range forces, such as hydrophobic packing, van der Waals contacts, and hydrogen bonding should also contribute to fine structural rearrangements of the two redox partners, which optimize the interprotein electron transfer. Attempts to solve the three-dimensional structure of an efficient electron transfer complex between FNR and Fd (and NADP+) remained unsuccessful, despite extensive efforts and different crystallographic approaches (4).
To gain detailed insight into the catalytic reaction mechanism of FNR,
we have undertaken to sequence the gene of Capsicum annuum
FNR precursor and determined the crystal structure of mature pFNR at
2.5-Å resolution. Moreover, comparison of mature FNRs from different
species on the amino acid level as well as on the structural level
reveals structurally and functionally conserved features. The crystal
structure of pFNR enabled us to model complexes between FNR and its
physiological redox partner Fd, resulting in a complex consistent with
known biochemical data. This model suggests a mechanism for complex
formation between FNR and Fd. Furthermore, we were able to derive a
potential electron transfer pathway for successive electron transfer
between FNR and Fd. This should help us to gain further understanding
regarding the mechanisms of intramolecular and intermolecular electron
transfer processes, which are still debated.
![]() |
EXPERIMENTAL PROCEDURES |
---|
![]() ![]() ![]() ![]() ![]() ![]() |
---|
Extraction of RNA, cDNA Synthesis, and DNA Sequence
Analysis--
Fruit leaves of C. annuum Yolo Wonder were
harvested, frozen in liquid nitrogen, and stored at 70 °C until
use. With a mortar and a pestle, 3.5 g of frozen plant material
was ground to a fine powder under liquid nitrogen. The powdered leaves
were directly added to 10 ml of a buffer composed of 4 M
guanidine hydrochloride, 25 mM sodium citrate, 10 mM EDTA (pH 8.0), 0.1 M mercaptoethanol, and
0.5% (v/v) N-lauroylsarcosine and homogenized with an
Ultraturrax, followed by the addition of 1 ml of 2 M sodium
acetate buffer (pH 4.2). Nucleic acids were then isolated by
phenol-chloroform and chloroform extractions. Total RNA thus extracted
was prepared by precipitation with 0.1 volumes of 3 M
sodium acetate (pH 6.0) and 3 volumes of ethyl alcohol. The
precipitated total RNA was dissolved in 100 µl of water and stored at
20 °C. The RNA concentration was determined by UV absorption at
260 nm, while the intactness was analyzed by electrophoresis using
formaldehyde-agarose gels and ethidium bromide staining.
Poly(A)+ mRNA was isolated from total RNA by
chromatography on oligo(dT)-cellulose (Quiagen). 2 µg of the obtained
mRNA was used to synthesize cDNA with avian myeloblastosis
virus reverse transcriptase (Roche Molecular Biochemicals) according to
the manufacturer's instructions (15). Oligo(dT) primer and random
primer (hexanucleotides of random sequence) were used in this reaction
to prime synthesis of the first strand of cDNA. cDNA thus
synthesized was isolated by phenol-chloroform and chloroform
extractions, followed by precipitation with 10 µg of glycogen, 0.1 volumes of 3 M sodium acetate (pH 9.0), and 3 volumes of
ethyl alcohol. The precipitated cDNA was dissolved in 20 µl of
water and stored at
20 °C.
DNA sequencing was performed by PCR using degenerate oligonucleotides designed against the 3'-end and the 5'-end of the coding region of the FNR precursor gene. Sequence comparisons of known nucleic acid sequences from other plant species were used to design the degenerate oligonucleotides. PCR was performed with Deep VentR DNA Polymerase (New England BioLabs Inc.), screening the different degenerate oligonucleotides against each other at different annealing temperatures. PCR fragments of appropriate size were sequenced by automated Sanger dideoxynucleotide sequencing.
Sequence-- The complete coding sequence of C. annuum Yolo Wonder ferredoxin-NADP+ reductase precursor has been deposited with the EMBL Nucleotide Sequence Data base (accession code AJ250378).
Sequence Comparison-- Best fit alignments of the amino acid sequence of pFNR with known amino acid sequences of other plant FNRs as well as alignments of nucleic acid sequences were calculated with the PileUp program of the GCG package (Genetics Computer Group, Madison, WI).
Cloning and Expression of the fnr Gene--
To obtain the mature
form of pFNR, the proposed processing site of the chloroplast transit
peptide was deduced by sequence alignment. PCR using Deep
VentR DNA Polymerase was used to add flanking
BamHI (3' site) and NdeI (5' site) restriction
sites. To prevent translation of different pFNR polypeptides in
Escherichia coli, as reported for recombinant spinach FNR
(16), we converted the codons for residues Val75 and
Val77 from GTG to the synonymous GTT by site-directed
mutagenesis. The resulting construct was cloned into the
NdeI and BamHI sites of expression vector
pET22b(+) (Novagen) to yield pET22b(+)-fnr. E. coli strain
BL21(DE3) was transformed with pET22b(+)-fnr. The bacteria were grown
in 12 liters of LB medium containing 100 µg/ml ampicillin to an
A600 of 0.8, and expression of mature pFNR was induced with 1 mM isopropyl--thiogalactopyranosid for
5 h.
Purification of Native and Recombinant FNR--
For preparation
of the crude extract, the washed and precooled paprika leaves (~1 kg)
were homogenized in an appropriate amount of buffer A (25 mM Tris-HCl (pH 8.0), 150 mM NaCl, 10 mM EDTA, 2 mM -mercaptoethanol, 1 mM phenylmethanesulfonyl fluoride, 5 mM
benzamidine hydrochloride) using a Waring Blender with glass pearls.
The homogenate was mixed with DNase and RNase, filtered through a nylon
net and one layer of glass wool, and centrifuged at 10,000 × g (60 min, 4 °C) to remove the remaining solid
components. The supernatant was applied to a Reactive Red 120 affinity
column (50 ml; Sigma) equilibrated with buffer B (25 mM
Tris-HCl (pH 8.0), 50 mM NaCl). After washing off the
unabsorbed material with an excess of the same buffer, bound proteins
were eluted using a linear NaCl gradient from 0.05 to 1.0 M
in buffer B. The fractions containing pFNR were identified by
SDS-polyacrylamide gel electrophoresis, dialyzed against buffer B, and
applied to a DEAE-Sepharose column (100 ml; Sigma) equilibrated in
buffer B. After extensive washing with buffer B, a gradient from 0.05 to 0.5 M NaCl in the same buffer was started. The
pFNR-containing fractions were pooled and dialyzed against buffer C (25 mM Tris-HCl (pH 8.0), 1.5 M (NH4)2SO4) and then applied to a
phenyl-Sepharose column (50 ml; Sigma) and eluted in a linear
decreasing gradient from 1.5 to 0 M
(NH4)2SO4 in 25 mM
TRIS-HCl (pH 8.0). The resulting pFNR was concentrated and desalted by
dialysis against buffer B. Polyacrylamide gel electrophoresis and
UV-visible absorption spectra were used as a criterion for purity of
the protein. Protein concentrations were measured by the method of
Bradford (17) with a bovine serum albumin standard curve. For
purification of the recombinant pFNR, the E. coli cells that
overexpressed pFNR were harvested and resuspended in buffer A. Cells
were broken by sonification, and cell debris was removed by
centrifugation at 40,000 × g (60 min, 4 °C). The purification procedure for the recombinant protein was as described before.
Crystallization and X-ray Data Collection--
Crystals were
grown by sitting drop vapor diffusion against a reservoir containing
0.1 M Tris-HCl (pH 8.5) and 1.9 M
(NH4)2SO4. The droplets consisted
of 4.5 µl of protein solution (15.0 mg/ml, buffered with 20 mM Tris-HCl (pH 8.0)) and 4.5 µl of reservoir solution
and were equilibrated against 5 ml of reservoir solution at 20 °C.
Under these conditions, rectangular crystals grew to an approximate
size of 0.3 × 0.3 × 0.25 mm3 within 5 days. The
pFNR crystals belong to space group P21 with unit cell
parameters a = 44.72 Å, b = 108.98 Å,
c = 90.36 Å, /
= 90.0°, and
= 95.57°. X-ray diffraction data to 2.5-Å resolution were collected on
a MAR-Research image plate (Hamburg, Germany) equipped with a Rigaku
(Tokyo, Japan) RU 200 rotating anode x-ray generator. Data were
collected with a single pFNR crystal at room temperature in 1° frames
with exposure times of 1000 s/frame. The data were indexed with MOSFLM
(18) and scaled and merged using programs of the CCP4 suite (19).
Statistics for the data are shown in Table I.
Molecular Replacement and Structure Refinement--
Due to the
77% identity between the amino acid sequences of spinach FNR and pFNR,
it was expected that their overall three-dimensional structures would
be similar. Therefore, the structure was solved by Patterson search
techniques using the program package AMoRe (20). A poly(A)-model of the
1.7-Å resolution crystal structure of spinach FNR (4) was used as
search model. The FAD prosthetic group was omitted. A native Patterson
map was calculated using programs of the CCP4 suite (19). A
pseudo-origin peak could be detected in the Harker section at
y = 0.5. Self-rotation functions were calculated with
the program GLRF (21) to determine the noncrystallographic symmetry. In
the 180°, 120°, and 90° section, no peaks could be detected.
These calculations indicate that the asymmetric unit contains two
molecules of pFNR related by a translation. Therefore, Patterson search
techniques were applied for two pFNR molecules. Patterson rotation
searches were calculated with resolution limits of 15.0 to 3.5 Å and
an angular step size of 2.5°. Translation functions corresponding to
the highest rotation peaks resulted in a promising solution with two
monomers in the asymmetric unit. After a rigid body refinement,
correlation coefficient and Rcryst factor were
56.4 and 37.3%, respectively.
Structure refinement of the model was done with X-PLOR (22) using the
parameters of Engh and Huber (23). Model building into
2Fo Fc and
Fo
Fc maps calculated with
XPLOR was done using the program FRODO (24). The electron density maps
could not be improved by 2-fold real space averaging using MAIN (25)
with noncrystallographic symmetry operators determined with LSQMAN
(26). The electron density maps showed well defined density into which
we fitted the FAD molecule, using the FAD coordinates of spinach FNR as
starting coordinates. By contrast, the N-terminal residues 55-66
(first 12 N-terminal residues of mature pFNR) and the surface loop
comprising residues 171-175 remained poorly defined in the electron
density map. In further rounds of refinement, an overall anisotropic
B-factor was included, and in later rounds individual
isotropic B-factors were refined. Refinement parameters for
FAD were calculated with the program XPLO2D (27). To check the validity
of the different stages of the refinement process free R
values (28) were calculated from 5% of the diffraction data. When the
refinement nearly converged, one phosphate ion and 219 water molecules
per pFNR molecule were progressively added to the molecular model.
Finally, rebuilding and refinement cycles converged at
Rcryst and Rfree values
of 19.8 and 26.5%, respectively. The polypeptide chain could be traced from residue 66 to 362. The final model includes 592 residues, the two
FAD cofactors, two phosphate ions located at the
NADP+-binding sites, and 438 solvent molecules in the
asymmetric unit. The stereochemistry of the model was analyzed using
X-PLOR and PROCHECK (29). Details of the refinement are given in Table I.
Molecular Model Building of Substrate Complexes-- Molecular modeling to dock two macromolecules implementing the geometric surface recognition algorithm of Katchalski-Katzir (30) was done using the program FTDOCK (31). The refined structure of pFNR and the structure of the mutant E92K of [2Fe-2S]-ferredoxin I from Spinacia oleracea (32) were chosen for calculating the structure of an efficient electron transfer complex. During docking calculations, pFNR represented the fixed molecule and Fd the mobile molecule, respectively. The electrostatic function of FTDOCK was used to improve the final rank of the docking results. The 15 solutions with the highest scores for surface complementarity and favorable electrostatic interactions were used for refinement and energy minimization with MULTIDOCK (33). One final solution showing a large decrease in total energy during refinement resulted in a promising complex of the two molecules with favorable electrostatic interactions and high surface complementarity. Moreover, NADP+, which was generated by superimposing the structure of the NADP+-FNR(Y308S) complex (6) on the structure of pFNR, was positioned into the active site of the modeled pFNR-Fd complex, following the binding mode described by Deng et al. (6). Using the model building program O (34), new possible orientations were built for the C-terminal pFNR tyrosine residue 362, which has to be displaced to permit stacking interactions between the nicotinamide moiety of NADP+ and the isoalloxazine ring of FAD. Energy minimization of the pFNR-Fd-NADP+ complexes with different orientations of the C-terminal tyrosine 362 was then carried out using the program CNS (35). pFNR and both ligands were minimized simultaneously.
Coordinates--
The refined model of pFNR has been deposited
with the Research Collaboratory for Structural Bioinformatics Protein
Data Bank (accession code 1FB3).
![]() |
RESULTS AND DISCUSSION |
---|
![]() ![]() ![]() ![]() ![]() ![]() |
---|
Sequencing and Sequence Analysis of pFNR--
We identified the
gene encoding the FNR precursor of C. annuum Yolo Wonder by
PCR using a set of degenerate oligonucleotides targeted to the 3'-end
and the 5'-end of the coding region. A PCR fragment of 1089 base pairs
was obtained, which showed high sequence identities when compared with
FNR nucleic acid sequences of other plant species. The complete
nucleotide sequence of the gene was determined from the isolated
cDNA. The fnr gene of pFNR precursor codes for a
polypeptide of 362 amino acids with a calculated molecular mass of
40,407 Da and a pI of 8.57. pFNR precursor bears an amino-terminal
transit peptide consisting of residues 1-54, which allows the
protein to pass the chloroplast membrane, where it is proteolytically
processed to the mature form, which contains residues 55-362. Mature
pFNR has a molecular mass of 33,177 Da and a pI of 6.33. pFNR precursor
can be aligned significantly over its entire length with proteins of
the ferredoxin-NADP+ reductase family (Fig.
1). It is about 75% identical to spinach FNR precursor, ice plant FNR precursor, pea FNR precursor, bean FNR
precursor, and tobacco FNR precursor. The sequence similarity for the
FAD-binding domains and the NADP+-binding domains of
different members of the ferredoxin-NADP+ reductase family
is extremely high, but it is even considerable for the different
transit peptides containing many serine, threonine, and small
hydrophobic residues but few acidic residues. Regarding the degree of
similarity of the mature proteins, the first N-terminal amino acids
share only a few common features in contrast to the rest of the
protein. In the molecular structures of pFNR and spinach FNR, this
N-terminal part of the protein is even invisible in the electron
density map, indicating high flexibility and lower importance for
functional and structural integrity. The high degree of similarity
between the different mature FNRs on the amino acid level as well as on
a structural level (see below) indicate that both the overall fold and
the common amino acid features are important factors for electron
transfer.
|
Cloning, Recombinant Expression, and Purification of pFNR-- To have at hand large amounts of native pFNR for detailed mechanistic and structural studies, the NdeI-BamHI mature pFNR fragment was isolated by PCR from paprika cDNA and cloned in the expression vector pET22b(+). E. coli BL21(DE3) transformed with pET22b(+)-fnr, where fnr is under the control of a T7 promoter, produced the mature flavoprotein in a completely active form and yielded higher levels of soluble protein when E. coli cells were induced at temperatures lower than 37 °C. Furthermore, high levels of pFNR production were still obtained when cultures were induced in the late logarithmic rather than in earlier phases of growth, thus allowing a higher biomass yield.
We developed a purification procedure for the isolation of pFNR from paprika fruit leaves and from bacterial lysates, respectively. The main purification step of this procedure was the affinity chromatography to Reactive Red 120 Sepharose followed by ion exchange chromatography and hydrophobic interaction chromatography. Purified pFNR was more than 90% pure, as judged from Coomassie-stained SDS-polyacrylamide gels (not shown) and showed a major band at an apparent molecular mass of 33 kDa. The integrity of the purified samples was tested by UV-visible spectra showing the expected maxima at 280 nm (protein) and 385 and 458 nm (protein-bound FAD) (1).
Crystallization and Quality of the Final Model--
pFNR crystals
were grown by sitting drop vapor diffusion to an approximate size of
0.3 × 0.3 × 0.25 mm3, and x-ray diffraction
data were collected to 2.5-Å resolution. The structure of pFNR has
been solved by Patterson search techniques using spinach poly(A)-FNR as
search model (4). The final model includes two pFNR monomers in the
asymmetric unit, but the noncrystallographic symmetry between them was
restrained only in the earlier stages of refinement. Real space 2-fold
averaging did not improve the electron density maps. Unambiguous parts
and side chains could be added during refinement. In the final electron
density map, all residues are well defined except the N-terminal
segment, including residues 55-66. Moreover, residues 171-175 located
in an external loop on the FAD-binding site of pFNR are poorly defined
in the electron density map. This is comparable with the structures of spinach FNR (3, 4) and Anabaena FNR (5) and presumably reflects the flexibility of this part of the protein. For the reflection data between 2.5- and 8.0-Å resolution, the
crystallographic R-factor for the final model is 19.8%, and
the Rfree is 26.5%. The mean error of model
coordinates as estimated by a Luzzatti plot (37) as well as by the A
method (38) is 0.27 and 0.28 Å, respectively. The stereochemistry of
the model is well defined, since deviations from ideal values are 0.007 Å for bond length and 1.6° for bond angles (Table
I). All but three nonglycine main-chain
dihedral angles are within energetically favorable regions of the
Ramachandran plot (39), with 86.7% of the angles in the most favorable
area. For the three residues in disallowed regions, Lys69,
Lys70, and Asn170, the electron density is well
defined. Asn170 is located at the FAD-binding site and is
hydrogen-bonded to AN1 of the adenine moiety of FAD. Its unfavorable
main chain conformation may be required for tight binding of the
adenine portion of FAD to pFNR, whereas Lys69 and
Lys70 are in the N-terminal region located at the protein
surface with apparently higher flexibility.
|
Overall Structure of pFNR--
In the crystal structure of pFNR,
the molecule exists as a monomer like other members of the protein
family (3-5). pFNR is divided into two distinct domains, the
FAD-binding domain comprising residues 67-201 and the
NADP+-binding domain containing residues 202-362, which
are connected by a long loop (Fig. 2,
a and b). The secondary structure elements assigned to the two domains of FNR and their nomenclature are shown in
Fig. 2a.
|
The FAD-binding domain is made up of a barrel built of six antiparallel
-strands, which are arranged in two perpendicular
-sheets
(residues 86-95 (
1), 105-111 (
2), 123-127 (
3), 141-144 (
4), 158-164 (
5), and 191-199 (
6)). The bottom of
the barrel is capped by a short
-helix (residues 179-186 (
1))
and a long loop (residues 169-175) surrounded by two short
-strands
(residues 167 and 168 and residues 176 and 177, not numbered in the
nomenclature), both lying on the FAD-binding site of pFNR. The
FAD-binding domain contains the conserved cis-proline of the molecule,
Pro198, which is located at the end of sheet
6.
The core of the NADP+-binding domain contains a
five-stranded parallel -sheet (residues 212-218 (
1), 245-252
(
2), 275-281 (
3), 315-321 (
4) and 357-361 (
5))
surrounded by six
-helices (residues 223-234 (
1), 261-270
(
2), 295-299 (
3), 303-311 (
4), 324-340 (
5), and 344-354
(
6)). The NADP+-binding domain is similar to the
classical dinucleotide binding fold (42). The number of strands are,
however, different. The secondary structure of the FAD-binding domain
as well as the NADP+-binding domain is nearly identical to
other members of the FNR superfamily, but it is mainly the loops
between them that vary (Fig. 2c).
Bruns and Karplus (4) suggested that the C-terminal
NADP+-binding domain may consist of two subdomains. The
first contains all residues responsible for NADP+-binding,
and the second contains an invariant hydrophobic pocket, which may form
the attachment site of FNR to the thylakoid membrane or to a
corresponding membrane-docking protein. pFNR shows a similar architecture. The hydrophobic pocket is located on the far side of the
parallel -sheet and is lined with hydrophobic residues of the
-sheet and the opposite
-helices (
3,
4,
5, and
6; Fig. 2a). The cavity is close enough to the protein surface
to allow a hydrophobic attachment site to poke in. Besides the possible membrane attachment site, there exist two other large clusters of
buried hydrophobic side chains in pFNR important in determining its
three-dimensional fold. Cluster 2 includes the interior of the
six-stranded
-barrel, some side chains of the interdomain bridge,
and the extended N-terminal segment and thus establishes the structure
necessary for FAD binding. Cluster 3 comprises the interdomain region
and is responsible for the tight association of the two domains.
FAD Binding and Conformation--
The -barrel of the
FAD-binding domain exhibits a small gap between stands
4 and
5,
providing space for the isoalloxazine and ribityl moieties of the FAD
cofactor. The ribose and adenine moieties extend along the long
FAD-binding loop between
5 and
1, which is surrounded on either
side by two small
-strands (residues 167-177), and are well exposed
to solvent. pFNR binds its prosthetic group FAD through direct and
often water-mediated hydrogen bonds, van der Waals contacts, and
stacking interactions between the bases and aromatic amino acids (Fig.
3). Many of these interactions involve
main-chain atoms located in repetitive secondary structure
elements.
|
The binding mode of the isoalloxazine ring is similar in all members of
the superfamily. The isoalloxazine ring is tightly bound to the protein
between two tyrosine residues, Tyr143 on the
pro-si face and Tyr362 on the pro-re
face (Fig. 4a). Main-chain
atoms of residues Cys162 and Lys164 and the
side chain of Ser144 form direct hydrogen bonds to the
isoalloxazine ring, whereas Ser144 and Tyr362
interact through water-mediated hydrogen bonds. While a large part of
the isoalloxazine ring is deeply nestled into the opening of the
-barrel, the edge of the isoalloxazine ring (with its methyl groups
C7a and C8a) is exposed to solvent. This is presumably the site where
the iron-sulfur cluster passes its electrons to the FNR-bound FAD (see
below). In the structure of PDR, which exhibits a ferredoxin-like
iron-sulfur domain covalently attached to a FNR-like pair of domains,
the iron-sulfur cluster is indeed located very close to the C7a and C8a
methyl groups of the isoalloxazine ring (8).
|
The ribityl moiety and the phosphate group are also tightly bound to the protein mostly through direct hydrogen bonds from protein side chains and main chains. The hydrogen bonding pattern resembles that described for spinach FNR (4). Main-chain atoms of Leu142 and the side chain of Tyr143 participate in direct hydrogen bonds to the hydroxyl groups of the ribityl moiety, while main-chain atoms of Val179 and Ser181 and side-chain atoms of Ser181 and Arg141 form hydrogen bonds to the phosphate groups. Furthermore, there exist four hydrogen-bonded water molecules.
While the binding of the isoalloxazine ring, the ribityl moiety, and
the phosphate groups are well preserved in the reductase superfamily,
ribose and adenine binding show variations. The binding mode among
members of the superfamily is different. The FAD-binding loop (residues
167-177) between 5 and
1 is usually responsible for adenine
binding. In contrast to spinach FNR (4), where the adenosine moiety is
only loosely bound by a van der Waals contact with the side chain of
Tyr120, the adenine part in pFNR is well fixed. When
comparing the structures of pFNR, spinach FNR (4), and
Anabaena FNR (5), the unique feature of the pFNR FAD-binding
site is the bent conformation of the FAD-binding loop (residues
167-177 for pFNR) toward the adenine portion of FAD (Figs.
2c and 4b). This conformation brings the side
chain of Tyr168 in optimal orientation for stacking
interactions to the adenine ring. Besides that, Asn170 is
in a position in which it can form a direct hydrogen bond to AN1 of the
adenosine moiety, and Leu166 forms van der Waals contacts
to AC5 and AN7 (Fig. 4, a and b).
The conformation of the ribityl portion of the FAD molecule bound to pFNR is different from that reported for spinach FNR (4), since its hydroxyl groups at position 3 and 4 point in different directions. However, the electron density is unambiguous in this region. The tighter interaction between the protein and the adenine moiety in pFNR may possibly force the switched conformation of the ribityl part in paprika FAD.
Putative Ferredoxin Binding Site-- The redox partner of FNR, Fd, is a small acidic protein with an apparent molecular mass of 11 kDa, which contains a single [2Fe-2S] cluster coordinated by four cysteine residues as prosthetic group. Fd is involved in a number of different metabolic pathways (44) and in the photosynthetic electron transport chain, where Fd transfers its electron to FNR. The stability of the electron transfer complex between FNR and Fd depends on ionic strength, suggesting the involvement of intermolecular electrostatic interactions (13). Chemical modification experiments (45-48), chemical cross-linking experiments (49), and mutagenesis studies (50-52) on FNR and Fd of different species revealed a number of acidic residues in Fd and of basic residues in FNR that probably participate in complex formation. These experiments suggested that Fd binds within a large shallow cleft between the two domains of FNR. The FAD molecule is located in the center of this cavity, with its dimethylbenzene ring directed toward the putative Fd binding site.
The structure of pFNR exhibits this large concave cleft around the site of electron transfer. This cleft is surrounded by four major patches of positively charged residues grouped around the FAD cofactor. The first positively charged patch includes residues Lys136, His138, Lys139, and Arg141. A second patch of positively charged residues is located in the neighborhood of Lys201. The third patch is very large, including residues Lys348, Lys349, Lys352, and Lys353, while the fourth patch is located around the positively charged residue Lys323. Sequence comparison with spinach and Anabaena FNR show that most of these positively charged residues have been implicated in ferredoxin binding in the mentioned biochemical modification studies (45-52).
Modeling of Ferredoxin Binding Modes-- Attempts to crystallize FNR in complex with Fd have not been successful in our hands. Therefore, the putative binding of the redox partner [2Fe-2S]-ferredoxin I from S. oleracea (32) to pFNR has been modeled assuming that larger conformational changes and domain movements do not occur. The structure of PDR (8), which exhibits a ferredoxin-like iron-sulfur domain covalently attached to an FNR-like pair of domains, showed good correlation regarding the orientation of all secondary structure elements when superimposed with the structure of pFNR.
Models for the complex of spinach FNR with Spirulina
platensis (3) and Aphanothece sacrum Fds (53) and a
model (14) in which the spinach Fd sequence was fit to the structure of
the Fd from A. sacrum have been obtained previously.
Nevertheless, these models were obtained by manually placing the
iron-sulfur cluster of Fd to the proposed binding cleft of FNR near the
exposed portion of the flavin to account for the results of the
biochemical studies (45-52). Here we present an FNR-Fd docking model
generated by computational molecular modeling. We used the structure of [2Fe-2S]-ferredoxin I from S. oleracea (32) to dock onto
pFNR. The relative starting orientation of the two proteins used for molecular modeling was not set in advance. The complex was calculated using a geometric surface recognition algorithm and an electrostatic correlation function. The resulting preliminary pFNR-Fd complex was
then energy-minimized, allowing for side-chain conformational changes
and rigid body movements of the interacting proteins. The final complex
had minimized in total energy from 200 to
1047 kcal/mol including
the internal and intermolecular interaction energy.
A ribbon drawing of the modeled pFNR-Fd complex is shown in Fig.
5. Fd fills the large shallow cleft
between the two domains of FNR as predicted for ferredoxin binding and
is located close to the FAD and NADP+ binding sites. The
center of the iron-sulfur cluster adjusts on the geometric plane, which
is defined by the isoalloxazine ring of FAD, about 7.6 Å apart from
the dimethylbenzene ring of the isoalloxazine moiety, the part of the
cofactor through which the electrons are exchanged. In comparison with
the structure of PDR (8), where the iron-sulfur cluster is 7.4 Å apart
from the dimethylbenzene ring of the prosthetic group FMN, this
distance seems to be suitable for effective electron transfer.
Tyr37* of Fd,2
which is located close to the iron-sulfur cluster, faces directly into
the active site of FNR. Its phenol ring is oriented nearly parallel to
the plane defined by the isoalloxazine ring of FAD and faces toward the
C-terminal Tyr362 of pFNR (Fig.
6a). In this position,
Tyr37* is able to form a weak hydrogen bond (OH-OE1, 3.8 Å) to Glu360 and is in close van der Waals contact to the
dimethylbenzene ring of FAD. Tyr37* represents the only
residue of Fd that directly interacts with active site residues of
pFNR. In the protein-protein interface of the modeled complex, however,
numerous interactions, mainly of an electrostatic nature, can be
identified. Most of the interprotein interactions are formed between
residues of the NADP+-binding domain of pFNR and of Fd,
whereas residues of the FAD-binding domain are less involved in
interprotein interactions. Herein, we focus on some selected
interprotein interactions. Lys139 of the FAD-binding domain
of pFNR interacts with Glu93*, which is located at the C
terminus of Fd, while five important interactions are present between
the NADP+-binding domain of pFNR and Fd as follows:
Lys201 interacts with Asp62* and
Ser43*, Lys348 with Glu30*,
Lys352 with Asp26*, and Lys353 with
Asp60*. The pFNR residues involved in these interprotein
interactions correspond to the four major patches of positively charged
residues on the surface of pFNR described above. The Fd residues can be grouped into three major regions of negative charge on the surface of
Fd. Two distinct negative patches are located on both sides of the
iron-sulfur cluster, and a third patch is present near the C terminus
of Fd.
|
|
The structure of the pFNR-Fd complex is absolutely consistent with the results of the biochemical studies on electron transfer between FNR and Fd (45-52), while these were not able to give evidence for detailed interprotein interactions, and seems to meet all requirements for efficient electron transfer. Superposition of the structure of the pFNR-Fd complex and the structure of PDR (8) reveals no major structural differences between the two domains of pFNR and the two FNR-like domains of PDR, as expected. The orientations of the complexed Fd and the Fd-like domain of PDR are not identical, while the positions of both iron-sulfur clusters are nearly the same. The iron-sulfur cluster of PDR is moved a little bit out of the plane that is defined by the isoalloxazine ring of the prosthetic group. In comparison with the Fd-like domain of PDR, the complexed Fd seems to be rotated nearly 90° around a vertical axis running through its iron-sulfur cluster. However, a PDR-like rotation of the complexed Fd would no longer be consistent with the observed cross-linking results (45-52).
The modeling studies strengthen the postulated mode of FNR-Fd complex formation (14). Both protein surfaces possess several patches of oppositely charged residues that help in properly orientating the partner molecules. Interaction of the above mentioned basic residues on pFNR and acidic residues on Fd may then help in improving the orientation, as do the many short range forces, such as intermolecular van der Waals contacts, hydrophobic packing interactions, and hydrogen bonding. All of these interactions may contribute to fine structural rearrangements of both redox partners to optimize the interprotein electron transfer.
Electron Transfer Pathway--
Our modeling studies reveal a
potential orientation of the redox partners in the electron transfer
complex and also suggest a pathway for electron transfer between the
iron-sulfur cluster of Fd and the isoalloxazine ring of FNR-bound FAD.
The rate of electron transfer and the pathway of electrons within a
protein complex is controlled by distance, location, and environment of the redox components (54). It is generally accepted that the main
pathway of long distance electron transfer in proteins involves a chain
of covalently bonded atoms. Aromatic residues placed in appropriate
positions to the two redox centers may enhance electron transfer
through proteins by a more effective coupling through their extended
*-orbitals, since the energy gap between the tunneling electron and
the
-system is significantly smaller than between the tunneling
electron and
-orbitals (54). We suggest that the position of
Tyr37* in the modeled electron transfer complexes may
enhance the interaction between the iron-sulfur cluster and the
isoalloxazine ring, since the two ring systems are in van der Waals
contact, providing an electronic overlap and enabling electron
tunneling through the aromatic rings. Our FNR-Fd complex therefore
suggests the following electron transfer pathway. Starting at the
iron-sulfur cluster, Fe3+ transfers one electron through
the iron-ligating sulfur atom of Cys39* along the peptide
chain to the aromatic
-system of Tyr37*.
Tyr37* is in close van der Waals contact (~2.4-3.1 Å)
to the aromatic isoalloxazine ring, and by their electronic overlap
tunneling to FAD is enabled. In other systems that have been selected
by evolution for efficient electron transfer, aromatic residues have been found in similar positions, probably enhancing electron transfer. Examples are the tryptophan-mediated reduction of quinone in the photosynthetic reaction center (55, 56) and in the MADH/amicyanin system, where a tryptophan residue is placed at the interface of the
two proteins (57).
NADP+ Binding and Catalytic Mechanism-- The structures of two mutants of pea FNR (Y308S and Y308W, in which the C-terminal tyrosine residue is mutated) in complex with NADP+ and NADPH revealed a productive NADP+-binding mode for members of the FNR family, in which the nicotinamide ring is stacked against the flavin isoalloxazine ring at an angle of ~30° and occupies the same place as the side chain of the C-terminal tyrosine in native FNR (6). Due to the necessary displacement of this C-terminal tyrosine side chain, none of the previous structural studies on native FNR yielded productive NADP+ binding. NADP+-binding to wild type FNR occurs through the 2'-phospho-5'-AMP part of NADP+, whereas the nicotinamide part of NADP+ faces into the solvent (4, 5). Three mechanisms are possible for productive NADP+ binding. In the first binding mechanism, Fd binding to FNR provides a favorable environment for tyrosine displacement, followed by productive NADP+ binding (ordered sequential mechanism-Fd binding increases the affinity of FNR for NADP+). In the second mechanism, the tilted geometry of the isoalloxazine ring in reduced FAD forces tyrosine displacement and productive NADP+ binding (ping-pong mechanism). The third possibility is that both complex formation between FNR and Fd and the geometry of the reduced FAD force productive NADP+ binding (ordered sequential mechanism-Fd binding and the redox state of FAD both increase the affinity of FNR for NADP+). Here we present a model for productive NADP+-binding upon complex formation between pFNR and Fd following an ordered sequential reaction mechanism. Minor structural rearrangements upon pFNR-Fd complex formation may favor displacement of the C-terminal tyrosine, and the complex interface provides space in which the displaced tyrosine side chain is energetically stabilized. Tyr37*, which faces into the catalytic pocket of the complex, can also interact with the bound NADP+.
In the modeled pFNR-Fd complex (Fig. 6a), four positions are sterically accessible for the side chain of Tyr362. It could face into the gap between the adenine moiety of FAD and Fd exposed to solvent (position 1). On the other hand, Tyr362 could rotate toward the bound Fd pointing into the predominantly hydrophobic pocket, which is formed by residues Leu35*, Ile33*, Gly32*, Ala28*, and Val74* and is located on the left-hand side of Tyr37* (position 2). It could also move toward Tyr37*, where it would be stacked between the phenol ring of Tyr37* and the isoalloxazine ring of FAD (position 3). Finally, Tyr362 could face into the gap between the ribityl moiety of FAD and the bound Fd (position 4).
For modeling ternary complexes between Fd, FAD, and NADP+ according to an ordered sequential reaction, we positioned NADP+ according to the binding mode reported by Deng and coworkers (6) into the active site of the pFNR-Fd complex and changed the orientation of Tyr362 to the four positions described above (resulting in pFNR-Fd-NADP+ complexes 1-4). All four complexes minimized equally in total energy, accompanied by some smaller structural rearrangements. While ternary complexes 2 (Fig. 6b) and 4 (Fig. 6c) seem to reveal potential orientations for the tyrosine residue after displacement by NADP+, the position of this tyrosine residue seems to be energetically unfavorable in ternary complexes 1 and 3. In complex 3, Tyr362 moves during energy minimization toward the gap between the ribityl moiety of FAD and Fd and finally adopts the same orientation as in complex 4, while Tyr362 of complex 1 faces into the solvent, where the mainly hydrophobic side chain of Tyr362 is not stabilized. In complex 2 (Fig. 6b), in contrast, Tyr362 is deeply nestled into a hydrophobic pocket of Fd (described above). Its carboxyl group faces toward the ribityl part of NADP+, forming a hydrogen bond to one of the hydroxyl groups (O-OH9, 3.1 Å). Furthermore, the productive conformation of NADP+ is stabilized by the side chain of Tyr37*, which forms two strong interactions to the nicotinamide group (OH-O, 2.6 Å; OH-NH2, 3.5 Å) of NADP+. Similar stabilization conditions are present in complex 4 (Fig. 6c), and the side chain of Tyr362 faces into the gap between the ribityl moiety of FAD and Fd, forming a hydrogen bond to Asp34* (OH-OD1, 3.8 Å), while the carboxyl group forms a weak hydrogen bond to one of the hydroxyl groups of NADP+ (O-OH9, 3.5 Å).
These ternary complexes suggest a reasonable ordered sequential
mechanism for productive NADP+ binding involving complex
formation between FNR and its leading substrate Fd. Primarily,
NADP+ binds through its 2'-P-AMP portion to the uncomplexed
FNR, whereas the nicotinamide part still faces into the solvent as
reported for Anabaena FNR (5). In the leading step of the
ordered sequential reaction, Fd binds to FNR according to the binding
mode described above, producing a favorable environment for
Tyr362 displacement, which increases the affinity of the
enzyme for NADP+. In a second step, the nicotinamide
portion of NADP+ finally binds to FNR. The energetic cost
of displacing the tyrosine can presumably easier be compensated by
creation of an energetically favorable environment for the tyrosine
side chain, by the energetic gain due to nicotinamide binding and due
to its interaction with Tyr37*. Therefore, we favor this
electron transfer mechanism, including the formation of a ternary complex.
![]() |
ACKNOWLEDGEMENTS |
---|
We thank Gärtnerei Kamlah (München, Germany) for providing C. annuum Yolo Wonder fruit leaves. Silvia Gaertner and Irmingard Sures are gratefully acknowledged for giving advice on RNA isolation and cDNA synthesis.
![]() |
FOOTNOTES |
---|
* The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
The nucleotide sequence(s) reported in this paper has been submitted to the GenBankTM/EMBL Data Bank with accession number(s) AJ250378.
The atomic coordinates and the structure factors (code 1FB3) have been deposited in the Protein Data Bank, Research Collaboratory for Structural Bioinformatics, Rutgers University, New Brunswick, NJ (http://www.rcsb.org/).
§ To whom correspondence should be addressed. Tel.: 49-089-45563336; E-mail: anja.dorowski@gmx.de.
Supported by the Fonds der chemischen Industrie.
** Supported by a scholarship of Boehringer Ingelheim Fonds.
Published, JBC Papers in Press, October 26, 2000, DOI 10.1074/jbc.M004576200
2 Amino acids denoted with an asterisk are residues of Fd from S. oleracea within the modeled FNR-Fd complex.
![]() |
ABBREVIATIONS |
---|
The abbreviations used are: FNR, ferredoxin-NADP+ reductase; pFNR, paprika ferredoxin-NADP+ reductase; Fd, [2Fe-2S]-ferredoxin I; 2'-phospho-5'-AMP, adenosine 2',5'-diphosphate; PDR, phthalate dioxygenase reductase; PCR, polymerase chain reaction.
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() |
---|
1. | Carrillo, N., and Vallejos, R. H. (1987) Topics Photosynth. 8, 527-560 |
2. |
Arakaki, A. K.,
Ceccarelli, E. A.,
and Carrillo, N.
(1997)
FASEB J.
11,
133-140 |
3. | Karplus, P. A., Daniels, M. J., and Herriott, J. R. (1991) Science 251, 60-66[Medline] [Order article via Infotrieve] |
4. | Bruns, C. M., and Karplus, P. A. (1995) J. Mol. Biol. 247, 125-145[CrossRef][Medline] [Order article via Infotrieve] |
5. | Serre, L., Vellieux, M. D., Medina, M., Gómez-Moreno, C., Fontecilla-Camps, J. C., and Frey, M. (1996) J. Mol. Biol. 263, 20-39[CrossRef][Medline] [Order article via Infotrieve] |
6. | Deng, Z., Aliverti, A., Zanetti, G., Arakaki, A. K., Ottado, J., Orellano, E. G., Calcaterra, N. B., Ceccarelli, E. A., Carrillo, N., and Karplus, P. A. (1999) Nat. Struct. Biol. 6, 847-853[CrossRef][Medline] [Order article via Infotrieve] |
7. |
Correll, C. C.,
Ludwig, M. L.,
Bruns, C. M.,
and Karplus, P. A.
(1993)
Protein Sci.
2,
2112-2133 |
8. | Correll, C. C., Batie, C. J., Ballou, D. P., and Ludwig, M. L. (1992) Science 258, 1604-1610[Medline] [Order article via Infotrieve] |
9. | Lu, G., Campbell, W. H., Schneider, G., and Lindqvist, Y. (1994) Structure 2, 809-821[Medline] [Order article via Infotrieve] |
10. | Nishida, H., Inaka, K., Yamanaka, M., Kaida, S., Kobayashi, K., and Miki, K. (1995) Biochemistry 34, 2763-2767[Medline] [Order article via Infotrieve] |
11. |
Wang, M.,
Roberts, D. L.,
Paschke, R.,
Shea, T. M.,
Masters, B. S. S.,
and Kim, J. J. P.
(1997)
Proc. Natl. Acad. Sci. U. S. A.
94,
8411-8416 |
12. | Ingelman, M., Bianchi, V., and Eklund, H. (1997) J. Mol. Biol. 268, 147-157[CrossRef][Medline] [Order article via Infotrieve] |
13. | Foust, G. P., Mayhew, S. G., and Massey, V. (1969) J. Biol. Chem. 244, 964-979[Medline] [Order article via Infotrieve] |
14. |
De Pascalis, A. R.,
Jelesarow, I.,
Ackermann, F.,
Koppenol, W. H.,
Hirasawa, M.,
Knaff, D. B.,
and Bosshard, H. R.
(1993)
Protein Sci.
2,
1126-1135 |
15. | Sambrook, J., Fritsch, E. F., and Maniatis, T. (1989) Molecular Cloning: A Laboratory Manual , 2nd Ed. , Cold Spring Harbor Laboratory, Cold Spring Harbor, NY |
16. | Aliverti, A., Jansen, T., Zanetti, G., Ronchi, S., Herrmann, R. G., and Curti, B. (1990) Eur. J. Biochem. 191, 551-555[Abstract] |
17. | Bradford, M. M. (1976) Anal. Biochem. 72, 248-254[CrossRef][Medline] [Order article via Infotrieve] |
18. | Leslie, A. G. W. (1991) Recent Changes to the MOSFLM Package for Processing Film and Image Plate Data , SERC Laboratory, Daresbury, Warrington, UK |
19. | Collaborative Computational Project 4. (1994) Acta Crystallogr. Sec. D 50, 760-763[CrossRef][Medline] [Order article via Infotrieve] |
20. | Navaza, J. (1994) Acta Crystallogr. Sec. A 50, 157-163[CrossRef] |
21. | Tong, L. A., and Rossmann, M. G. (1990) Acta Crystallogr. Sec. A 46, 783-792[CrossRef][Medline] [Order article via Infotrieve] |
22. | Brünger, A. T. (1992) X-PLOR, a System for Crystallography and NMR, version 3.1 , Yale University Press, New Haven, CT |
23. | Engh, R. A., and Huber, R. (1991) Acta Crystallogr. Sec. A 47, 392-400[CrossRef] |
24. | Jones, T. A. (1985) Methods Enzymol. 115, 157-171[Medline] [Order article via Infotrieve] |
25. | Turk, D. (1992) Weiterentwicklung eines Programms für Molekülgrahik und Elektronendichte-Manipulation und seine Anwendung auf verschiedene Protein-Strukturaufklärungen, Ph.D. thesis , Technical University Munich |
26. | Kleywegt, G. J., and Jones, T. A. (1994) ESF/CCP 4 Newsletter 31, 9-14 |
27. | Kleywegt, G. J., and Jones, T. A. (1998) Acta Crystallogr. Sec. D 54, 1119-1131[CrossRef][Medline] [Order article via Infotrieve] |
28. | Brünger, A. T. (1992) Nature 355, 472-474[CrossRef] |
29. | Laskowski, R. A., MacArthur, M. W., Moss, D. S., and Thornton, J. M. (1993) J. Appl. Crystallogr. 26, 283-291[CrossRef] |
30. | Katchalski-Katzir, E., Shariv, I., Eisenstein, M., Friesem, A. A., Aflalo, C., and Vakser, I. A. (1992) Proc. Natl. Acad. Sci. U. S. A. 89, 2195-2199[Abstract] |
31. | Gabb, H. A., Jackson, R. M., and Sternberg, M. J. E. (1997) J. Mol. Biol. 272, 106-120[CrossRef][Medline] [Order article via Infotrieve] |
32. | Binda, C., Coda, A., Aliverti, A., Zanetti, G., and Mattevi, A. (1998) Acta Crystallogr. Sec. D 54, 1353-1358[CrossRef][Medline] [Order article via Infotrieve] |
33. | Jackson, R. M., Gabb, H. A., and Sternberg, M. J. E. (1998) J. Mol. Biol. 276, 265-285[CrossRef][Medline] [Order article via Infotrieve] |
34. | Jones, T. A., and Kjelgaard, M. (1991) O: The Manual , University of Uppsala, Uppsala, Sweden |
35. | Brünger, A. T., Adams, P. D., Clore, G. M., DeLano, W. L., Gros, P., Grosse-Kunstleve, R. W., Jiang, J-S., Kuszewski, J., Nilges, M., Pannu, N. S., Read, R. J., Rice, L. M., Simonson, T., and Warren, G. L. (1998) Acta Crystallogr. Sec. D 54, 905-921[CrossRef][Medline] [Order article via Infotrieve] |
36. | Barton, G. J. (1993) Protein Eng. 6, 37-40[Medline] [Order article via Infotrieve] |
37. | Luzzatti, V. (1952) Acta Crystallogr. Sec. A 5, 802-810[CrossRef] |
38. | Read, R. J. (1986) Acta Crystallogr. Sec. A 42, 140-149[CrossRef] |
39. | Ramachandran, G. N., and Sasisekharan, V. (1968) Adv. Protein Chem. 23, 283-437[Medline] [Order article via Infotrieve] |
40. | Kraulis, J. (1991) J. Appl. Crystallogr. 24, 946-950[CrossRef] |
41. | Merritt, E. A., and Bacon, D. J. (1997) Methods Enzymol. 277, 505-524 |
42. | Rossmann, M. G., Liljas, A., Brändén, C.-I., and Banaszak, L. J. (1975) The Enzymes 11, 61-102 |
43. | Evans, S. V. (1993) J. Mol. Graphics 11, 134-138[CrossRef][Medline] [Order article via Infotrieve] |
44. | Knaff, D. B., and Hirasawa, M. (1991) Biochim. Biophys. Acta 1056, 93-125[Medline] [Order article via Infotrieve] |
45. | Jelesarow, W., DePascalis, A. R., Koppenol, W. H., Hirasawa, M., Knaff, D. B., and Bosshard, H. R. (1993) Eur. J. Biochem. 216, 57-66[Abstract] |
46. | Medina, M., Méndez, E., and Gómez-Moreno, C. (1992) FEBS Lett. 298, 25-28[CrossRef][Medline] [Order article via Infotrieve] |
47. | Medina, M., Méndez, E., and Gómez-Moreno, C. (1992) Arch. Biochem. Biophys. 299, 281-286[Medline] [Order article via Infotrieve] |
48. |
Zanetti, G.,
Aliverti, A.,
and Curti, B.
(1984)
J. Biol. Chem.
259,
6153-6157 |
49. | Zanetti, G., Morelli, D., Ronchi, S., Negri, A., Aliverti, A., and Curti, B. (1988) Biochemistry 27, 3753-3759 |
50. | Hurley, J. K., Hazzard, J. T., Martínez-Júlvez, M., Medina, M., Gómez-Moreno, C., and Tollin, G. (1999) Protein Sci. 8, 1614-1622[Abstract] |
51. | Aliverti, A., Corrado, M. E., and Zanetti, J. (1994) FEBS Lett. 343, 247-250[CrossRef][Medline] [Order article via Infotrieve] |
52. | Schmitz, S., Martínez-Júlvez, M., Gómez-Moreno, C., and Böhme, H. (1998) Biochim. Biophys. Acta 1363, 85-93[Medline] [Order article via Infotrieve] |
53. | Karplus, P. A., and Bruns, C. M. (1994) J. Bioenerg. Biomembr. 26, 89-99[Medline] [Order article via Infotrieve] |
54. | Marcus, R. A., and Sutin, N. (1985) Biochim. Biophys. Acta 811, 265-322 |
55. | Deisenhofer, J., Epp, O., Miki, K., Huber, R., and Michel, H. (1984) J. Mol. Biol. 180, 385-398[Medline] [Order article via Infotrieve] |
56. | Plato, M., Michel-Beyerle, M. E., Bixon, M., and Jortner, J. (1989) FEBS Lett. 249, 70-74[CrossRef] |
57. | Chen, L., Durley, R. C. E., Mathews, F. S., and Davidson, V. L. (1994) Science 264, 86-90[Medline] [Order article via Infotrieve] |