From the Departments of Structural Biology and
§ Biochemistry, St Jude Children's Research Hospital,
Memphis, Tennessee 38105 and the ¶ Department of Molecular
Sciences, University of Tennessee Health Science Center, Memphis,
Tennessee 38163
Received for publication, January 9, 2001
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ABSTRACT |
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In Escherichia coli, the expression
of fatty acid metabolic genes is controlled by the transcription
factor, FadR. The affinity of FadR for DNA is controlled by long
chain acyl-CoA molecules, which bind to the protein and modulate gene
expression. The crystal structure of FadR reveals a two domain dimeric
molecule where the N-terminal domains bind DNA, and the C-terminal
domains bind acyl-CoA. The DNA binding domain has a winged-helix motif,
and the C-terminal domain resembles the sensor domain of the Tet
repressor. The FadR·DNA complex reveals how the protein interacts
with DNA and specifically recognizes a palindromic sequence. Structural and functional similarities to the Tet repressor and the BmrR transcription factors suggest how the binding of the acyl-CoA effector
molecule to the C-terminal domain may affect the DNA binding affinity
of the N-terminal domain. We suggest that the binding of
acyl-CoA disrupts a buried network of charged and polar residues
in the C-terminal domain, and the resulting conformational change is
transmitted to the N-terminal domain via a domain-spanning Fatty acid synthesis and degradation are important facets of
bacterial physiology, and the regulation of these pathways has been
extensively studied in the model prokaryote, Escherichia coli (1). Fatty acids are vital constituents of the cell
membranes, but they also represent a source of energy. Thus, fatty acid
degradative and biosynthetic metabolic pathways must be switched on and
off based on the availability of extracellular fatty acids. In E. coli, the transcription factor, FadR, functions as a switch that coordinately regulates the machinery required for fatty acid
It has been known for some time that the enzymes responsible for fatty
acid degradation in E. coli are inducible, and the isolation
of regulatory mutants, fadR, suggested the existence of a
single repressor that controlled the entire set of degradative (fad) genes (4-6). Mutagenesis studies identified the
fadR gene and verified that this single repressor controls
the transcription of the whole fad regulon (7, 8). Following
the cloning (9) and sequencing (10) of the fadR gene, it was
predicted that FadR contains a
HTH1 motif, which is
consistent with its proposed function as a DNA-binding protein. The
fad genes are only induced in the presence of long-chain fatty acids, suggesting that FadR is a classical bacterial repressor. Thus, in the absence of fatty acids, the protein binds at a site downstream of the promoters of the fad genes and represses
transcription. When long chain fatty acids become available, they are
converted to CoA thioesters, bind to FadR, and elicit a conformational
change that releases the protein from DNA, thereby removing the
repression. Subsequent studies on the fadB gene (11)
confirmed this general outline and revealed that the FadR DNA binding
site in fadB is close to the +1 region relative to the start
site of transcription.
The subsequent observation that fadR mutants were also
defective in unsaturated fatty acid production (12) led to the
important discovery that FadR is also a transcriptional activator (13, 14). Specifically, it controls the expression of the fabA
gene that encodes the enzyme FabA, which introduces double bonds into the growing acyl chain (13, 14). Analysis of the fabA gene revealed a canonical FadR binding site in the To understand the mechanism of FadR at the molecular level, we have
performed a crystallographic analysis of the protein. Here we report
the structure of the FadR dimer at 1.5 Å in the absence of
bound DNA and long chain acyl-CoA thioesters, and also the structure of
the FadR·DNA complex at 3.25 Å. The crystal structure of FadR in the
absence of DNA has recently been determined independently at 2.0 Å resolution (16), and therefore the emphasis of this report is how FadR
interacts with DNA. Structural homologies to other transcription
factors provide important clues as to how ligand binding controls the
DNA affinity.
Cloning the fadR Gene--
The fadR open reading
frame (GenBankTM/EBI accession X08087) was amplified from
E. coli strain UB1005 in a polymerase chain reaction using
the primers fadr-Nco (5'-CCATGGTCATTAAGGCGCAAAGC) and fadr-Bam
(5'-GGATCCTTATCGCCCCTGAATGGC), and was cloned into pCR2.1 (Invitrogen).
Following transformation into INV Purification of FadR--
Native FadR was purified by tandem
anion and cation ion-exchange chromatography essentially as described
previously (11). [SeMet]FadR did not bind to cation-exchange resins,
and the following alternative purification method was developed. Cells
from 1 liter of culture were resuspended in buffer A (20 mM
Tris, pH 8.0, 1 mM EDTA, 1 mM dithiothreitol)
and lysed in a French pressure cell at 16,000 psi. Cell debris was
removed by centrifugation, and the supernatant was directly applied to
a 30 ml DE52 column pre-equilibrated with 20 mM Tris, pH
8.0. The protein was eluted with a linear gradient of KCl in 20 mM Tris, pH 8.0. The fractions containing FadR were
identified by SDS-polyacrylamide gel electrophoresis and pooled. The
pooled fractions were concentrated with an Amicon stirred cell and
applied to a Sephacryl S200 gel filtration column equilibrated in
buffer A containing 100 mM NaCl. FadR eluted as a single
peak at a position consistent with a dimer and was essentially pure as
judged by SDS-polyacrylamide gels.
Crystallization--
FadR in the absence of bound DNA was
crystallized using the hanging drop procedure at 18 °C by mixing
equal volumes of the protein solution at 10 mg/ml with a reservoir
solution containing 0.1 M sodium citrate, pH 5.6, 0.7 M ammonium sulfate, 0.2 M lithium sulfate, and
5 mM zinc chloride. Although crystals grew from ammonium sulfate or lithium sulfate alone, a mixture of the two gave the best
crystals. [SeMet]FadR crystallized in the same condition, but without
zinc chloride. Crystals typically grew within 2 weeks and were in space
group P21 with unit cell dimensions a = 59.4 Å, b = 87.0 Å, c = 59.1 Å, and
Data Collection and Processing--
All diffraction data used in
these studies were collected at the Structural Biology Center (SBC)
beamline 19ID at the Advanced Photon Source in Argonne National
Laboratory, Chicago. Data were recorded on a 3 × 3 CCD detector
from crystals flash-frozen in liquid nitrogen at 100 K. MAD data were
collected from Se-Met-labeled crystals that were cryoprotected with
30% sucrose. The peak, inflection point and high energy remote
wavelengths were determined from an x-ray fluorescence spectrum
collected from the mounted crystal, and 360° of data were collected
at each wavelength in two 180° sweeps. The 1.5 Å native dataset was
collected from an unlabeled frozen crystal that was also cryoprotected
in 30% sucrose. Data were collected from a frozen FadR·DNA complex
crystal that was cryoprotected in 20% MPD. Integration, scaling, and
merging of data were performed with the program HKL2000 (18).
Structure Determination and Refinement--
The FadR structure
in the absence of DNA was determined directly from the MAD data. All
Patterson search, MAD phasing, electron density map calculations, and
density modification procedures were carried out using the CNS program
suite (19). Four selenium atoms were identified at equivalent locations
in the unit cell from each of the three MAD datasets. These initial
positions were refined and then used in a MAD phasing calculation at
1.7 Å resolution. The experimental electron density map was of high
quality, but density modification produced a greatly improved and
easily interpretable map at 1.7 Å resolution. A model was built into
the density using the O program (20), and refined against the MAD
remote data using simulated-annealing, positional, and individual
B-factor refinement. The partially refined model was then further
refined in the same manner against the superior quality 1.5 Å resolution native data. The final model includes residues 5-230 in
each monomer, 416 water molecules, 6 sulfate ions, and 1 zinc ion
located on the local molecular 2-fold axis. 95.5% of residues are in
the core regions of a Ramachandran plot performed in PROCHECK (21), and
only the two glutamine residues at position 6 in the dimer are in
forbidden regions.
The structure of the FadR·DNA complex was solved with
molecular replacement methods using AMoRe (22). The 1.5 Å resolution native FadR dimer was used as a search model. After rigid body refinement performed in AMoRe, the R-factor was 45.5% with a
correlation coefficient of 0.505 calculated with data between 10.0 and
3.2 Å resolution. A preliminary model of the bound DNA molecule was built into the visible density, and the structure was improved iteratively with successive rounds of model building using the O
program (20) and refinement using CNS (19). Only grouped B-factor
refinement was applied. Non-crystallographic symmetry restraints were
applied during the early stages of refinement and later removed. The
final model contains one FadR dimer (residues 7 to 228), one DNA
duplex, and one magnesium ion.
The FadR Crystal Structure--
The MAD technique, using
metabolically incorporated selenomethionine, was used to solve
the FadR structure. The selenium positions were identified using
automated Patterson search methods, and phasing was carried out to the
resolution limit of the MAD data (1.7 Å). The data collection and
phasing statistics are shown in Table I.
The electron density map after density modification was of superb
quality, and the complete FadR molecule was visible apart from four
residues at the N terminus and nine residues at the C terminus.
Refinement by simulated annealing was performed against a native
dataset that extended to 1.5 Å resolution. The refinement statistics
and model quality parameters are listed in Table
II.
The structure of the FadR monomer is shown in Fig.
1, and it is essentially identical to
that reported previously (16). Briefly, it can be divided into three
regions, an Homologous Structures to FadR--
Known protein structures with
homology to FadR were identified using the Dali search engine (25). The
coordinates of the N- and C-terminal domains were submitted separately.
As noted earlier, the N-terminal domain shows clear homology to the
winged-helix motif (23), and the Dali search identified CAP (26) as the most similar winged-helix transcription factor. A number of proteins were shown to have similar helical arrangements to the FadR C-terminal domain within their structures, for example ATP synthase, myosin, and
serum albumin, but the most intriguing was the C-terminal domain of the
tetracycline (Tet) repressor (27). Like FadR, the Tet repressor
contains a regulatory C-terminal domain that recognizes a specific
ligand (tetracycline), which, in turn, mediates its ability to bind
cognate DNA (28). Fig. 1 shows the FadR, CAP, and Tet repressor domains
in equivalent orientations to highlight their similarities.
Based on sequence alignment, it was previously suggested that FadR
might be homologous to the GntR family of transcription factors (16,
29). However, the sequence homology is marginal, and there are no
structures of any of the GntR family members to confirm this suggested
relationship. We consider that the homologies to CAP and the Tet
repressor based on structure are more relevant. Therefore, in terms of
structural genomics, FadR can be categorized as a chimera of two motifs
that have previously been characterized, the winged-helix motif and the
C-terminal domain of the Tet repressor.
The Crystal Structure of the FadR·DNA Complex--
The structure
of the FadR·DNA complex was determined by molecular replacement using
the dimer as the search model. Phases from the molecular replacement
solution produced an electron density map in which the sugar-phosphate
backbone of the bound DNA was clearly visible. Iterative rounds of
model building and refinement generated the final structure. Because
the FadR·DNA complex is the asymmetric unit in the crystal,
non-crystallographic symmetry constraints were incorporated into the
early rounds of refinement, but these were relaxed in the final rounds.
Also, to guard against model bias, a number of simulated annealing omit
maps were calculated during refinement in which regions at the
protein-DNA interface were removed from the model. A representative
region of the final electron density map is shown in Fig.
2A, and the pertinent
statistics of the final model are listed in Table II.
The complete structure of the FadR·DNA complex is shown in Fig.
2B. The conformation of the FadR dimer in the complex is
virtually identical with that in the absence of bound DNA. The RMSD on
alpha carbons is 0.717 Å. All of the duplex DNA is visible in the
electron density map apart from the overhanging ends. In the crystal,
the ends of the DNA interact with the C-terminal domains of neighboring FadR molecules, and the 5'-cytosine of one chain is flipped out of the
duplex to form one of these interactions (Fig. 2B). Apart from the central G(11)-C(11') base pair, the DNA in the complex is
palindromic, and the pseudo 2-fold axis is coincident with the local
2-fold axis of FadR, with the major groove facing the protein and the
minor groove facing away from the protein. The result is that each
N-terminal domain interacts in an identical fashion with its DNA
half-site. The DNA has a B-form conformation with a curvature of 20°
toward the protein, and this results in a contraction of the central
major groove and an expansion of the opposite minor groove. The gross
features of the B-form helix are recognized by the two N termini of the
paired
A detailed view of the FadR-DNA interface is shown in Fig.
3A, and only one monomer is
presented, because both half-sites are identical. The interactions that
determine the specificity are shown diagrammatically in Fig.
3B. Although there are a number of nonspecific interactions
with the DNA sugar-phosphate backbone, only three adjacent base pairs
are formally recognized within the complex, T/A (6/16'), G/C (7/15')
and G/C (8/14'). These interact with His-65, and arginines 35 and 45 respectively, which are invariant in the three known sequences of FadR
(16). His-65 is at the very tip of the wing and is buried deep in the
minor groove where the N The Architecture and Specificity of the FadR-DNA
Interaction--
The structure of the FadR·DNA complex provides a
firm basis for understanding how this transcription factor controls the
expression of bacterial fatty acid metabolic genes. A key finding is
that the N-terminal winged-helix motif does not bind DNA in the
classical manner. The structures of several protein-DNA complexes
involving winged-helix motifs have now been determined (23) and,
typically, the second recognition helix in the HTH region binds along
the DNA major groove, and the
The oligonucleotide that was used to produce crystals of the FadR·DNA
complex was based on the highest affinity cognate sequence that is
found within the fadB promoter (11). The structure reveals that only three residues (Arg-35, Arg-45, and His-65) formally recognize this sequence, and FadR appears to have specificity for the
palindromic consensus sequence 5'-TGGNNNNNCCA-3'. It is clear why the
two G/C base pairs in each half-site are important specificity elements
in the complex (Fig. 3). Both are involved in specific and identical
Arg/G/C triple interactions, and the adjacent triples are able to form
a stable stacked configuration within the half-site. The specificity
for the flanking T/A base pairs in the minor groove is probably related
to the close approach of the wing and the sidechain of His-65, which is
facilitated by the lack of a sidechain on the adjacent and conserved
Gly-66. A G/C base pair in this position would introduce an amine group into the minor groove (N2 from the guanine), and the close approach would be compromised. Finally, the five central spacer elements are
necessary to match the architecture of the dimeric N-terminal domains
and the resulting distances between the 2-fold related specificity
elements. The consensus sequence based on our structure is supported by
binding data on other cognate sequences where deviations generally
reduce the affinity (15). However, the conservation within these
sequences also suggests that a better consensus is 5'-TGGTNNNACCA-3',
and this is consistent with independent in vitro selection
studies on the FadR binding site within the iclR promoter
(31). The inner T/A base pair is present in the oligonucleotide used in
this study, and a specific interaction is feasible with Arg-49 that is
adjacent in the complex. However, this is not supported by our
structure in which Arg-49 forms a salt bridge with Glu-50'.
The FadR Switching Mechanism--
Studies both in vitro
and in vivo have revealed that long-chain acyl-CoA
thioesters are the actual ligands that control the DNA binding affinity
of FadR, and that they interact directly and reversibly with a specific
region of the C-terminal domain (14, 24, 32-34). Based on the
functionally and structurally related Tet repressor that has been fully
characterized by crystallographic studies (27), we can predict that the
interaction results in a conformational change that affects the
structure, and hence the DNA binding affinity, of the N-terminal
domain. In the Tet repressor, ligand binding within the C-terminal
domain causes the movement of an
Mutagenesis experiments have identified Gly-216, Glu-218, Ser-219,
Trp-223, and Lys-228 within helix
As regards the interdomain communication mechanism, helix -helix.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
-oxidation and the expression of a key enzyme in fatty acid
biosynthesis (2, 3).
40 region of the promoter where positive activators of the
70 factor
typically bind. Thus, the role of FadR as a repressor or an activator
depends on the location of its binding site within the promoter. When
bound downstream of the RNA polymerase binding site of the
fad promoters, it blocks the polymerase and acts as a
repressor. However, when bound close to the
40 region, it promotes the binding of the RNA polymerase. A number of FadR binding sites have
now been characterized, and all conform to this straightforward model
(15).
EXPERIMENTAL PROCEDURES
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ABSTRACT
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DISCUSSION
REFERENCES
F' (Invitrogen), a plasmid with
the correct sequence was isolated, and the gene was released by
digestion with NcoI and BamHI. This fragment was ligated into similarly digested pET-15b (Novagen) to create pPJ139 for
expression of full-length FadR. Plasmids were transformed into
B834(DE3) (Novagen) for expression. To produce FadR containing selenomethionine ([SeMet]FadR), the cells were resuspended in minimal
medium containing selenomethionine (0.05 g/liter; Sigma) immediately
prior to induction. This method is fully described elsewhere (17).
Generally, the level of FadR expression was greater than 20% of the
soluble cell protein.
= 120.2°. There is one dimer in the asymmetric unit.
To prepare the FadR·DNA complex, the oligonucleotides, 5'-CGATCTGGTCCGACCAGATGCT-3' and 5'-GCATCTGGTCGGACCAGATCGA-3', were
dissolved in buffer A, annealed, and mixed with FadR at a protein/DNA
molar ratio of 1:3. Crystals of the complex were also obtained by the
hanging drop method at 18 °C by mixing equal volumes of 8 mg/ml
complex solution with a reservoir solution containing 10% isopropyl
alcohol, 50 mM MES, pH 6.0, and 10 mM
MgCl2. Crystals grew in 1 month and were in space group
P212121 with the cell dimensions
a = 83.7 Å, b = 110.1 Å, and
c = 130.2 Å. There is one dimer complex in the
asymmetric unit.
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ABSTRACT
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DISCUSSION
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Crystallographic data and phasing statistics for the FadR structure
analysis
Refinement Statistics
/
N-terminal domain (
1-
1-
2-
3-
2-
3),
an
-helical C-terminal domain
(
6-
7-
8-
9-
10-
11-
12), and a linker comprising two
short
-helices (
4-
5). The two major domains only interact via
the linker region, and the overall shape of the monomer therefore
resembles a dumbbell. The structure of the N-terminal domain conforms
to the so-called winged-helix motif (23), consistent with its role in
binding DNA. The larger C-terminal domain is essentially an
antiparallel array of six
-helices that form a barrel-like
structure, with a seventh
-helix (
10) forming a lid at the end
closest to the N-terminal domain. FadR forms a dimeric structure in the
crystal, which is consistent with independent studies showing a dimer
to be the functional state of the protein in solution (24). The
monomers are packed together in a parallel fashion, and the two domains
and the linker regions make reciprocal interactions across the
interface. The monomers are tightly wrapped, and ~1600
Å2 of surface area are buried in forming the dimer. A
region of tetrahedral electron density on the local 2-fold axis was
interpreted as a zinc atom coordinated to two water molecules and the
O
1 atoms of asparagines 81 and 81' (the prime refers to the second monomer in the dimer structure). Although zinc ions were not required to grow crystals of FadR, they were essential to obtain the highest resolution crystals from which the 1.5 Å native dataset was
collected.
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Fig. 1.
The structure of the FadR monomer
(left), and its homology to the winged-helix domain
of CAP and the ligand binding domain of the Tet repressor
(right). The monomer is color-coded to
emphasize the proposed homologies: magenta, the winged-helix
motif; green, the Tet repressor motif; red, the
two additional helical regions in the C-terminal domain of FadR
compared with the Tet repressor. The figure was produced using
MOLSCRIPT (36) and rendered with Raster3D (37).
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Fig. 2.
The structure of the FadR·DNA
complex. A, the 2Fo-Fc
calculated phased map from the final refined coordinates of the
FadR·DNA complex at 3.25 Å. The area shown is the recognition of G7,
G8, and A16' by Arg-35, Arg-45, and His-65, respectively. The map is
contoured at the 1.5 level and displayed using the O program
(20). B, stereoview of the FadR·DNA complex. The
central paired
3 helices are in the major groove, and the two wings
are within the flanking minor grooves. Note that the DNA is bent toward
the FadR molecule, and that the major and minor grooves are contracted
and expanded respectively as a result. The 5'-cytosine at one end of
the DNA is flipped out of the duplex and makes crystal contacts with a
neighboring FadR molecule. The picture was produced using the Ribbons
program (38).
3 helices that project orthogonally into the central major
groove and the two
-ribbon wings that dock into the flanking minor grooves.
2 nitrogen forms a hydrogen bond with N3 of
A16. The guanidinium groups of arginines 35 and 45 are hydrogen-bonded
to the O6 and N7 atoms of G7 and G8 in the major groove. Other
conserved residues in the complex include Arg-49, that forms a salt
bridge interaction with the phosphate group of G8, and threonines 44, 46, and 47 at the N terminus of helix
3 that interact with the
sugar-phosphate backbone. Finally, glutamic acids 34 and 50' appear to
be key residues, because they form electrostatic interactions with, and presumably stabilize the positions of, arginines 35, 45, and 49.
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Fig. 3.
Details of the FadR·DNA complex showing
specific protein-DNA interactions. A, stereoview
showing a close up of the FadR-DNA interface within one monomer. See
text for details. The picture was produced using the Ribbons program
(38). B, schematic of the FadR-DNA interface showing the
important residues and how they interact with the DNA. Amino acids from
one monomer are shown in the same colored text. Note that
the interactions are identical across the local 2-fold axis of the
DNA.
DISCUSSION
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ABSTRACT
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EXPERIMENTAL PROCEDURES
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DISCUSSION
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-ribbon binds within the adjacent
minor groove. However, the precise details of the interaction can vary (23, 30), and FadR is unusual because only the N terminus of the
3
recognition helix is within the major groove. The most similar example
to the FadR variation is found in CAP (26), although, unlike CAP, FadR
does not induce a sharp 90° bend in the DNA. It is therefore not
surprising that the FadR and CAP winged-helix domains are structurally
very similar. The way in which FadR binds DNA is determined by the
location of the
3 recognition helices that are paired together at
the dimer interface. The FadR dimer cannot bind DNA with the
3
helices along the major groove unless the N-terminal domains move apart
to expose the surfaces of these helices. This was implied in a model of
the complex based on the apo-FadR structure (16), but we can now
confirm that the N-terminal domains continue to interact via their
3
helices in the FadR·DNA complex.
-helix that connects to, and also
forms part of, the N-terminal domain (27). In the paired N-terminal
domains, the result of this movement is that the two DNA binding
helices are no longer able to bind in the major groove. Therefore, the connecting
-helix essentially represents a mechanism for
communicating structural changes from the regulatory domain to the DNA
binding domain. By analogy, to establish how the FadR switching
mechanism operates, it is necessary to understand the nature of the
ligand-induced conformational change, the interdomain communication
mechanism, and the resulting effects on the N-terminal domain.
12 as components of the acyl-CoA
binding site. Also, affinity labeling experiments with a palmitoyl-CoA
analog tagged the peptide 187-195 in the adjacent helix
11 (32).
The interior of the C-terminal domain contains an unusual cluster of
aromatic, charged, and polar residues, which is directly adjacent to
these putative acyl-CoA binding regions. The cluster is composed of
tyrosines 172, 193, and 215, Arg-105, Thr-106, Asp-145, and Ser-219,
and it is knitted together by a lattice of hydrogen-bonding
interactions that involves five buried water molecules (Fig.
4A). This lattice links
together five of the seven surrounding
-helices including
11 and
12, and we suggest the binding of acyl-CoA disrupts this lattice and produces a large conformational change within the C-terminal domain. This suggestion is based on structural studies of the BmrR
transcription factor, which controls the expression of the Bmr
multidrug-efflux transporter in Bacillus subtilis (35). The
C-terminal sensor domain of BmrR contains a similar buried cluster
composed of three tyrosines and a glutamic acid, which forms part of
the binding site for aromatic/hydrophobic cationic drugs. The structure
of the drug-bound complex showed that the aromatic ligand accesses this
buried array and induces a large conformational change in the sensor
domain.
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Fig. 4.
The FadR switch mechanism. A,
stereoview of the putative switch region in the C-terminal domain of
FadR. This constellation of buried aromatic, polar, and charged
residues is extensively hydrogen-bonded into a tight lattice and
adjacent to the putative acyl-CoA binding site involving helices 11
and
12. Note that the lattice includes a number of water molecules,
and it directly links five of the seven
-helices that comprise the
C-terminal domain. For clarity, the two remaining helices,
7 and
9, are not shown in the figure. B, structure of the FadR
dimer color-coded to emphasize the important interfaces that we suggest
are important to the switching mechanism. Note the polar residues on
helix
3 at the interface of the N-terminal domains and the aromatic
and hydrophobic residues at the boundaries of the N- and C-terminal
domains. The green sphere is the putative zinc ion at the
local 2-fold axis. The orange circle indicates the location
in the C-terminal domain of the constellation shown in A.
The figure was produced using MOLSCRIPT (36) and rendered with Raster3D
(37).
5 in the
linker region appears to be a key component, because it has extensive
and conserved hydrophobic interfaces with the two flanking domains
(Fig. 4B). Specifically, leucines 86 and 89, and Ile-82
interact with the N-terminal domain, and leucines 80 and 83, and Ala-87
interact with the C-terminal domain. The interface with the N-terminal
domain is also characterized by a conserved cluster of aromatic
residues comprising tryptophans 21, 60, and 75, and Phe-74. Therefore,
helix
5 may have a similar role to the equivalent domain-spanning
helix of the Tet repressor. Unfortunately, the BmrR structural studies
do not provide clues as to how the conformational change in the sensor
domain modulates DNA affinity because the DNA binding domain is missing
(35). One scenario for FadR is that the interface between the
3
helices is broken by movements in the C-terminal domain, and this leads
to a disruption of the precise DNA binding architecture of the
N-terminal domain dimer. This is plausible because the interface
between the N-terminal domains is only mediated by the paired
3
helices, and the interactions are exclusively ionic and polar (Fig.
4B). The 2-fold related arrays of interacting amino acids
comprise arginines 49, 54', and 57, Glu-50', Gln-53, and Asp-58'.
Glu-13 from helix
1 is the only other residue within FadR that
contributes to this interface. Thus, a separation of the N-terminal
domains would not be structurally unfavorable, because it would only
result in the exposure of two hydrophilic surfaces.
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ACKNOWLEDGEMENTS |
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We are particularly grateful to Rongguang Zhang and the rest of the SBC staff at the APS for help in collecting and processing the diffraction data. We also thank Hee-Won Park for invaluable help, and Suzanne Jackowski and Allen Price for insights and discussions. Pam Jackson, Amy Sullivan, Xiaoping He, and Charles Ross II provided excellent technical assistance.
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FOOTNOTES |
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* This work was supported by National Institutes of Health Grant GM34496 (to C. O. R.), Cancer Center (CORE) Support Grant CA 21765, and the American Lebanese Syrian Associated Charities (ALSAC).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 atomic coordinates and the structure factors (code 1HW1 and 1HW2) 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: Dept. of
Structural Biology, St Jude Children's Research Hospital, 332 N. Lauderdale, Memphis, TN 38105-2794. Tel.: 901-495-3040; Fax:
901-495-3032; E-mail: stephen.white@stjude.org.
Published, JBC Papers in Press, February 13, 2001, DOI 10.1074/jbc.M100195200
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ABBREVIATIONS |
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The abbreviations used are:
HTH, helix-turn-helix;
CAP, catabolite gene activator protein;
CoA, coenzyme
A;
FabA, -hydroxydecanoyl-ACP dehydratase;
MAD, multiwavelength
anomalous dispersion;
MPD, 2-methyl-2,4-pentanediol;
RMSD, root
mean-squared deviation;
MES, 4-morpholineethanesulfonic acid;
Tet, tetracycline.
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REFERENCES |
---|
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---|
1. | Cronan, J. E., Jr., and Rock, C. O. (1996) Escherichia coli and Salmonella typhimurium: Cellular and Molecular Biology , pp. 612-636, American Society for Microbiology, Washington, D. C. |
2. | Cronan, J. E., Jr., and Subrahmanyam, S. (1998) Mol. Microbiol. 29, 937-943[CrossRef][Medline] [Order article via Infotrieve] |
3. | DiRusso, C. C., and Nyström, T. (1998) Mol. Microbiol. 27, 1-8[CrossRef][Medline] [Order article via Infotrieve] |
4. | Overath, P., and Raufuss, E. M. (1967) Biochem. Biophys. Res. Commun. 29, 28-33[Medline] [Order article via Infotrieve] |
5. | Overath, P., Pauli, G., and Schairer, H. U. (1969) Eur. J. Biochem. 7, 559-574[Medline] [Order article via Infotrieve] |
6. | Klein, K., Steinberg, R., Fiethen, B., and Overath, P. (1971) Eur. J. Biochem. 19, 442-450[Medline] [Order article via Infotrieve] |
7. | Simons, R. W., Egan, P. A., Chute, H. T., and Nunn, W. D. (1980) J. Bacteriol. 142, 621-632[Medline] [Order article via Infotrieve] |
8. | Simons, R. W., Hughes, K. T., and Nunn, W. D. (1980) J. Bacteriol. 143, 726-730[Medline] [Order article via Infotrieve] |
9. | DiRusso, C. C., and Nunn, W. D. (1985) J. Bacteriol. 161, 583-588[Medline] [Order article via Infotrieve] |
10. | DiRusso, C. C. (1988) Nucleic Acids Res. 16, 7995-8009[Abstract] |
11. |
DiRusso, C. C.,
Heimert, T. L.,
and Metzger, A. K.
(1992)
J. Biol. Chem.
267,
8685-8691 |
12. | Nunn, W. D., Giffin, K., Clark, D., and Cronan, J. E., Jr. (1983) J. Bacteriol. 154, 554-560[Medline] [Order article via Infotrieve] |
13. | Henry, M. F., and Cronan, J. E., Jr. (1991) J. Mol. Biol. 222, 843-849[Medline] [Order article via Infotrieve] |
14. | Henry, M. F., and Cronan, J. E., Jr. (1992) Cell 70, 671-679[CrossRef][Medline] [Order article via Infotrieve] |
15. | DiRusso, C. C., Black, P. N., and Weimar, J. D. (1999) Prog. Lipid Res. 38, 129-197[CrossRef][Medline] [Order article via Infotrieve] |
16. |
van Aalten, D. M. F.,
DiRusso, C. C.,
Knudsen, J.,
and Wierenga, R. K.
(2000)
EMBO J.
19,
5167-5177 |
17. | Davies, C., Heath, R. J., White, S. W., and Rock, C. O. (2000) Structure 8, 185-195[CrossRef][Medline] [Order article via Infotrieve] |
18. | Otwinowski, Z., and Minor, W. (1996) Methods Enzymol. 276, 307-326 |
19. | Brünger, A. T., Admas, P. D., Clore, G. M., Delano, W. L., Gros, P., Grosse-Kunstleve, R. W., Jiang, J. S., Kuszewski, J., Nilges, M., and Pannu, N. S. (1998) Acta Crystallogr. Sect. D 54, 905-921[CrossRef][Medline] [Order article via Infotrieve] |
20. | Jones, T. A., Zou, J. Y., Cowan, S. W., and Kjeldgaard, M. (1991) Acta Crystallogr. Sect. A 47, 110-119[CrossRef][Medline] [Order article via Infotrieve] |
21. | Laskowski, R. A., Marcarthur, M. W., Moss, D. S., and Thornton, J. M. (1993) J. Appl. Crystallogr. 26, 283-291[CrossRef] |
22. | Navaza, J. (1994) Acta Crystallogr. Sect. A 50, 157-163[CrossRef] |
23. | Gajiwala, K. S., and Burley, S. K. (2000) Curr. Opin. Struct. Biol. 10, 110-116[CrossRef][Medline] [Order article via Infotrieve] |
24. |
Raman, N.,
Black, P. N.,
and DiRusso, C. C.
(1997)
J. Biol. Chem.
272,
30645-30650 |
25. | Holm, L., and Sander, C. (1995) Trends Biochem. Sci. 2, 478-480[CrossRef] |
26. | Schultz, S. C., Shields, G. C., and Steitz, T. A. (1991) Science 253, 1001-1007[Medline] [Order article via Infotrieve] |
27. | Orth, P., Schnappinger, D., Hillen, W., Saenger, W., and Hinrichs, W. (2000) Nat. Struct. Biol. 7, 215-219[CrossRef][Medline] [Order article via Infotrieve] |
28. | Betrand, K. P., Postle, K., Wray, L. V., Jr., and Reznikoff, W. S. (1983) Gene (Amst.) 23, 149-156[Medline] [Order article via Infotrieve] |
29. | Haydon, D. J., and Guest, J. R. (1991) FEMS Microbiol. Lett. 79, 291-296[CrossRef] |
30. | Gajiwala, K. S., Chen, H., Cornille, F., Roques, B. P., Reith, W., Mach, B., and Burley, S. K. (2000) Nature 403, 916-921[CrossRef][Medline] [Order article via Infotrieve] |
31. | Gui, L., Sunnarborg, A., and LaPorte, D. C. (1996) J. Bacteriol. 178, 4704-4709[Abstract] |
32. |
Raman, N.,
and DiRusso, C. C.
(1995)
J. Biol. Chem.
270,
1092-1097 |
33. | Cronan, J. E., Jr. (1997) J. Bacteriol. 179, 1819-1823[Abstract] |
34. |
DiRusso, C. C.,
Tsvetnitsky, V.,
Hojrup, P.,
and Knudsen, J.
(1998)
J. Biol. Chem.
273,
33652-33659 |
35. | Zheleznova, E. E., Markham, P. N., Neyfakh, A. A, and Brennan, R. G. (1999) Cell 96, 353-362[Medline] [Order article via Infotrieve] |
36. | Kraulis, P. J. (1991) J. Appl. Crystallogr. 24, 946-950[CrossRef] |
37. | Merritt, E. A., and Bacon, D. J. (1997) Methods Enzymol. 277, 505-524 |
38. | Carson, M. (1997) Methods Enzymol. 277, 493-505 |