Structural Analysis of the Neuronal SNARE Protein Syntaxin-1A†,‡
Jeffrey C. Lerman,§James Robblee,|Robert Fairman,|and Frederick M. Hughson*,§
Department of Molecular Biology, Princeton UniVersity, Princeton, New Jersey 08544, and Department of Cell, Molecular, and
DeVelopmental Biology, HaVerford College, HaVerford, PennsylVania 19041
ReceiVed February 21, 2000; ReVised Manuscript ReceiVed May 11, 2000
ABSTRACT: Intracellular trafficking depends on the docking and fusion of transport vesicles with cellular
membranes. Central to docking and fusion is the pairing of SNARE proteins (soluble NSF attachment
protein receptors) associated with the vesicle and target membranes (v- and t-SNAREs, respectively).
Here, the X-ray structure of an N-terminal conserved domain of the neuronal t-SNARE syntaxin-1A was
determined to a resolution of 1.9 Å using multiwavelength anomalous diffraction. This X-ray structure,
which is in general agreement with an NMR structure of a similar fragment, provides new insight into the
interaction surface between the N-terminal domain and the remainder of the protein. In vitro characterization
of the intact cytoplasmic domain of syntaxin revealed that it forms dimers, and probably tetramers, at low
micromolar concentrations, with concomitant structural changes that can be detected by limited proteolysis.
These observations suggest that the promiscuity characteristic of pairing between v-SNAREs and t-SNAREs
extends to the formation of homo-oligomeric t-SNARE complexes as well. They also suggest a potential
role for the neuronal Sec1 protein (nSec1) in preventing the formation of syntaxin multimers.
Neurotransmitter release requires three members of the
SNARE1family of proteins:
synaptobrevin (also called VAMP). The complex formed by
these three proteins is likely to provide an intimate physical
link between synaptic vesicles and the plasma membrane at
nerve terminals and thereby to facilitate membrane fusion
(1-3). These SNAREs, and especially syntaxin, also interact
directly with a number of additional proteins important for
synaptic transmission (4).
Syntaxin-1A is a 288-residue type II integral membrane
protein; residues 1-265 constitute the cytoplasmic domain,
while residues 266-288 form the carboxy-terminal trans-
membrane anchor (5). Among the three SNARE proteins
required for neurotransmitter release, only syntaxin appears
to adopt a well-folded tertiary structure in the absence of
the other two SNAREs (6, 7). By contrast, SNAP-25 and
the cytoplasmic domain of synaptobrevin are largely disor-
syntaxin, SNAP-25, and
dered under physiological conditions, folding only upon
SNARE complex formation.
Like all other SNARE proteins, the juxtamembrane region
of syntaxin contains heptad repeats of hydrophobic residues
(spanning residues 195-254) consistent with formation of
a coiled coil structure (8). Indeed, syntaxin forms an SDS-
resistant complex with SNAP-25 and synaptobrevin in which
this juxtamembrane region contributes one helix to a bundle
of four parallel R-helices (3). This configuration, which
would draw the transmembrane domains of synaptobrevin
(anchored in the synaptic vesicle) and syntaxin (anchored
in the presynaptic plasma membrane) into close proximity,
is likely to facilitate membrane fusion and consequent
In addition to the C-terminal transmembrane anchor and
the juxtamembrane region that binds the other SNARE
proteins, syntaxin has an approximately 15 kDa N-terminal
domain that appears as a flexible arm in electron micrographs
of SNARE complexes (1). This domain survives limited
proteolysis of SNARE complexes (9). Despite limited
sequence conservation, protease-resistant N-terminal domains
are a very common feature among widely diverse syntaxin
family members (10; L. Cavanaugh and F. M. Hughson,
unpublished). Two potential roles, which are not mutually
exclusive, have been ascribed to these N-terminal domains.
Studies with syntaxin and with its yeast homologue Sso1p
have suggested that the N-terminal domain inhibits SNARE
assembly, presumably by binding intramolecularly to the
juxtamembrane domain to produce a “closed” conformation
(7, 11-13). A second potential role is in binding to other
proteins with important roles in neurotransmitter release; the
binding site of Munc13 has been localized within the
N-terminal domain of syntaxin (14), and binding of the
†This work was supported by NIH Grant NS-38046, the Searle
Scholars Program, and the Beckman Young Investigators Program
‡X-ray coordinates and structure factors have been deposited in the
Research Collaboratory for Structural Bioinformatics, Rutgers Univer-
sity, New Brunswick, NJ (1EZ3).
* To whom correspondence should be addressed. Telephone: (609)
258-4982. Fax: (609) 258-6730. E-mail: email@example.com.
1Abbreviations: CD, circular dichroism; DTT, dithiothreitol; HEPES,
4-(2-hydroxyethyl)piperazine-1-ethanesulfonic acid; MAD, multiwave-
length anomalous diffraction; nSec1, neuronal Sec1 protein; NSF,
N-ethylmaleimide-sensitive fusion protein; PAGE, polyacrylamide gel
electrophoresis; PEG, polyethylene glycol; SDS, sodium dodecyl
sulfate; SeMet, selenomethionine; SNAP-25, synapse-associated protein
(25 kDa); SNARE, soluble NSF attachment protein receptor; t-SNARE,
target membrane SNARE; v-SNARE, vesicle SNARE; SRV, square
root of variance; VAMP, vesicle-associated membrane protein.
Biochemistry 2000, 39, 8470-8479
10.1021/bi0003994 CCC: $19.00© 2000 American Chemical Society
Published on Web 06/23/2000
neuronal Sec1 protein (nSec1; also called Munc18) is
dependent on this domain as well as on the juxtamembrane
region (15). nSec1 binds to a closed syntaxin conformation
(16), and mutations that apparently destabilize this closed
conformation compromise binding (7).
Fernandez et al. (17) have used solution NMR to charac-
terize the structure of the N-terminal domain of syntaxin-
1A. They found that residues 1-180, expressed as a
recombinant protein in Escherichia coli, contained disordered
ends, but that residues 27-146 formed an independently
folded domain. Structural analysis, using residues 27-146
with Cys 146 replaced with Ser, revealed a three-helix bundle
with a left-handed twist (17). The groove between the second
and third helices is formed by residues that are highly
conserved among plasma membrane syntaxins (17).
Here, we describe biophysical studies that provide further
insight into the overall structural organization of syntaxin-
1A. In particular, the X-ray structure of residues 24-150
has been determined and refined to 1.9 Å resolution. While
this structure is similar to that previously determined by
NMR spectroscopy (17), there are a number of differences,
particularly for residues that define the interactive surface
of the molecule. The X-ray structure reveals that the groove
between the second and third helices is not only conserved
but also quite hydrophobic, strengthening the hypothesis that,
in the closed form of intact syntaxin, it is involved in
interactions with the juxtamembrane region. Furthermore,
we have characterized the oligomeric state of the intact
cytoplasmic domain (residues 1-265) of syntaxin and find
that dimeric and probably tetrameric forms are significantly
populated at low micromolar protein concentrations. Syntaxin
dimers and tetramers may result from the parallel alignment
of juxtamembrane regions into a helical bundle structure
resembling that of the SNARE complex. These results add
a new level of complexity to previous findings suggesting
that heteromeric SNARE complex formation can be promis-
cuous (10, 18, 19), demonstrating that SNAREs can self-
partner as well.
Materials. All chemicals used for crystallization were
purchased from Fluka. Other materials were purchased from
Sigma (subtilisin, amino acids, and trace metal salts),
Boehringer Mannheim (proteinase K), and Novagen (BL21
& B834 E. coli strains).
Protein Expression. Protein expression plasmids were
constructed in pLM1 (20), a T7 promoter plasmid, using
PCR. The correctness of the coding sequence was confirmed
by DNA sequencing. Protein was expressed in E. coli
BL21(DE3) cells, except that BL21(DE3)(pLysS) cells were
used for production of syntaxin 1-265 and B834(DE3) cells
were used for the selenomethionine (SeMet) derivative of
SeMet DeriVatiVe Expression. SeMet-substituted syntaxin
24-150 was expressed essentially as described previously
(21) in modified M9 medium except that the starter culture
was supplemented with 5% LB medium and several amino
acids (alanine, aspartate, glutamate, glycine, histidine, as-
paragine, proline, glutamine, serine, tryptophan, and tyrosine)
were omitted from the modified M9 medium. SeMet
incorporation was confirmed by mass spectrometry.
Protein Purification. Syntaxin 24-150 (both the native
form and the SeMet derivative) was purified using Q-
Sepharose and phenyl-Sepharose columns and exchanged
into 10 mM dithiothreitol (DTT) by repeated concentration
and dilution in a centrifuge concentrator (Millipore). Syntaxin
1-253 and syntaxin 1-265 were purified by recovery from
inclusion bodies as described previously (22), subsequent
refolding by rapid dilution, and concentration by binding to
a Q-Sepharose column. The identity of the purified proteins
was confirmed by mass spectrometry.
Crystallization. Native syntaxin 24-150 was crystallized
by the hanging drop method. One microliter of 4 mg/mL
protein in 10 mM DTT was added to 1 µL of reservoir
solution [16% polyethylene glycol (PEG) 1500, 20 mM
sodium acetate (pH 5.0), and 10 mM DTT] on a coverslide
which was inverted over 1 mL of reservoir solution. Crystals
grew over a period of 1-4 days at 23 °C. SeMet syntaxin
24-150 derivative crystals grew under the same conditions,
except that the protein stock contained 8 mg/mL protein, 20
mM DTT, and 30 mM sodium acetate (pH 5.0).
X-ray Data Collection. Crystals were equilibrated in
several steps over 10 h into reservoir solution supplemented
with 18% glycerol and flash-frozen in a 100 K nitrogen cold
stream (Oxford). Data were collected at beamline X12C at
the National Synchrotron Light Source at Brookhaven
X-ray Data Processing. Data were indexed and scaled
using Denzo and Scalepack from the HKL software package
(version 1.11.0) (23). Initial phases were calculated from a
multiwavelength anomalous diffraction (MAD) data set using
MLPHARE and DM (24) and were later extended with
higher-resolution native data using X-PLOR (25) and CNS
version 0.5 (26). Cycles of model building and refinement
were carried out using X-PLOR, CNS, and O (27).
Protease Digestion. Syntaxin was exchanged into digestion
buffer [150 mM NaCl, 5 mM HEPES (pH 7.4), and 1 mM
DTT] using NAP-5 columns (Pharmacia), diluted to the
working concentration using additional digestion buffer, and
incubated on ice for 2 h to allow equilibration among
oligomeric states. Subtilisin was added to a substrate:protease
weight ratio of 90:1 to start the reaction. At the indicated
time points, samples were taken from the reaction and the
reactions stopped with 1 mM phenylmethanesulfonyl fluoride
followed by freezing in liquid nitrogen. After samples at all
time points had been collected, they were thawed, concen-
trated by trichloroacetic acid precipitation (15%), and an-
alyzed by SDS-PAGE. In separate experiments, several of
the digestion products were partially purified and character-
ized by amino-terminal sequencing and mass spectrometry.
Circular Dichroism. Circular dichroism (CD) data were
collected using an AVIV 62DS CD spectrometer and a
sample containing 10 µM syntaxin 24-150 in 1 mM
?-mercaptoethanol, 100 mM KF, and 1 mM sodium phos-
phate (pH 7.5). Spectra from three consecutive scans (250-
190 nm, 1 s averaging time, and 0.25 nm steps) were
averaged. For thermal unfolding experiments, data at 222
nm were recorded (1 min temperature equilibration, 0.5 min
averaging time, and 1 °C steps).
Analytical Ultracentrifugation. Protein was exchanged into
digest buffer containing 0.5 mM DTT. Sedimentation equi-
librium experiments were performed at 4 °C with a Beckman
Optima XL-A analytical ultracentrifuge at rotor speeds of
Structural Analysis of Syntaxin-1A
Biochemistry, Vol. 39, No. 29, 2000 8471
8000, 12 000, 16 000, and 20 000 rpm using an An-60 Ti
rotor. Six-channel, 12 mm path length, charcoal-filled Epon
cells with quartz windows were used. Twenty scans were
averaged. Temperature-corrected partial specific volumes of
both syntaxin-1A fragments were calculated from the
weighted average of the partial specific volumes of the
individual amino acids (28). Temperature-corrected solution
densities were calculated using standard tables listing coef-
ficients for the power series approximation of density (28).
The HID program from the Analytical Ultracentrifugation
Facility at the University of Connecticut (Storrs, CT) was
used for data analysis. Equilibrium sedimentation analysis
software implemented under IGOR Pro version 3.16 (Wave-
Metrics Inc., Lake Oswego, OR) using the Marquardt-
Levenberg algorithm for curve fitting was used for the
preparation of Figure 4.
Syntaxin 24-150. The structure of the cytoplasmic domain
of syntaxin-1A (residues 1-265) was probed by limited
digestion with nonspecific proteases. Subtilisin (Figure 1A)
and proteinase K reactions (data not shown) yielded very
similar patterns. In both cases, syntaxin is cleaved first
between residues 179 and 180 to yield two fragments,
residues 1-179 and 180-265 (Figure 1B). Additional
digestion yields a stable product comprising syntaxin residues
24-150. These findings are in good agreement with NMR
studies of syntaxin 1-180 indicating that residues 27-146
form a folded domain (17). Furthermore, in ternary syn-
taxin-SNAP-25-synaptobrevin complexes, a similar or
identical syntaxin domain is protected from proteolysis by a
variety of proteases (9).
Syntaxin 24-150, as expected, is highly R-helical (Figure
1C). In addition, it is thermostable, unfolding in a single
reversible cooperative transition with a midpoint of ap-
proximately 85 °C (Figure 1D). We have determined the
X-ray structure of this domain using selenomethionine-
substituted syntaxin 24-150. Phases estimated by MAD
yielded an electron density map that was, in general, readily
interpretable (Figure 2A). The structure was subsequently
refined to 1.9 Å resolution (Figure 2B) using data collected
from a native crystal (Table 1).
The X-ray structure shows a bundle of three R-helices
connected by short loops (Figure 2C,D), as expected on the
basis of the NMR structure of syntaxin 27-146 with the
point mutation Cys 146 f Ser (17). In keeping with the
notation introduced there (17), the three helices are termed
Ha (residues 28-64), Hb (residues 68-105), and Hc
(residues 110-146), although each helix is several residues
shorter in the NMR structure. The root-mean-square devia-
tion between the NMR and X-ray structures is relatively large
(2.1 Å over 124 equivalent CRatoms). Comparative analysis
of the X-ray and NMR structures reveals four principal
differences. First, the course of the backbone is well-defined
in the X-ray structure for most of the regions that are poorly
defined by the NMR structure. Second, many surface side
chains are also well-defined in the X-ray, but not the NMR,
structure. Third, a few well-defined side chains in the NMR
structure occupy significantly different rotamer positions in
the X-ray structure (see below). Fourth, the pronounced
curvature of the helix bundle observed in the NMR structure
FIGURE 1: Subtilisin digestion of the syntaxin cytoplasmic domain yields a protected N-terminal structural domain. (A) Recombinant
syntaxin 1-265 (2 µM) was mixed with subtilisin (90:1 w/w) on ice. Samples were withdrawn at the indicated times after protease addition,
and phenylmethanesulfonyl fluoride was added to a concentration of 1 mM to quench the reaction. Samples were analyzed on Coomassie-
stained SDS-PAGE gels. Digestion products as indicated were identified by amino-terminal sequencing and mass spectroscopy. (B) Schematic
interpretation of panel A. The initial products of digestion are fragments 1-179 and 180-265. Fragment 1-179 is subsequently trimmed
to 24-179 and ultimately to 24-150. (C) The CD spectrum of syntaxin 24-150 at 25 °C (black trace) indicates >90% R-helix content.
Almost complete reversibility is observed after heating to 95 °C and subsequent cooling back to 25 °C (gray trace). (D) Syntaxin 24-150
unfolds cooperatively with temperature as judged by its mean residue ellipticity at 222 nm.
8472 Biochemistry, Vol. 39, No. 29, 2000
Lerman et al.
is not seen in the X-ray structure, in which the bundle is
essentially straight. This difference may be attributable to
crystal lattice effects or to the lack of long-range distance
constraints in the determination of the NMR structure (29).
As noted previously (17), one side of syntaxin 24-150 is
decidedly acidic (Figure 3B). Two patches at either end of
the molecule are especially notable. One patch is formed by
Asp 68, Glu 69, Glu 73, Glu 74, Glu 76, Glu 77, and Asp
81. Many of these residues participate directly in an
interaction between syntaxin and the Ca2+-binding protein
syntaptotagmin, thought to be a principal Ca2+sensor in
regulating neurotransmitter release (17). A second, smaller
patch is formed by Glu 32, Glu 35, and Glu 38 (Figure 3B).
A ConserVed Hydrophobic GrooVe. Between helices Hb
and Hc is a wide groove (Figure 3A). This groove has a
pronounced hydrophobic character resulting from fully (Val
128) or partially (Ile 61, Leu 75, Leu 93, Phe 127, and Met
131) solvent-accessible nonpolar side chains. In the X-ray
(but not the NMR) structure, Gln 138 is tucked into the core
of the three-helix bundle, hydrogen bonded with the buried
side chain of His 58, further increasing the hydrophobic
character of the groove (Figure 2A,B). All of these residues
are highly conserved among syntaxins implicated in exo-
cytosis. Indeed, the residues lining the Hb-Hc groove are
significantly more conserved than those in either of the other
two helix-helix interfaces (17). The other two interhelical
grooves (Figure 3) are less hydrophobic, narrower, and
partially occluded by interhelical salt bridges: Lys 57 forms
an ion pair with Glu 74, bridging Ha and Hb, while nearby
Lys 55 forms a salt bridge network with Glu 52 and Glu
133, bridging Ha and Hc.
Although the exact role of the N-terminal domain in
syntaxin function is not known, interactions between the
N-terminal domain and the juxtamembrane C-terminal region
likely play important roles in regulating SNARE function.
Both syntaxin and its yeast homologue Sso1p exist in an
equilibrium between closed and open forms, distinguished
by whether the N-terminal domain is folded together with
the juxtamembrane region (closed) or not (open) (7, 12). In
the absence of direct high-resolution information character-
izing the closed conformation, two structural models have
been proposed in which the three-helix bundle is extended
by residues of the juxtamembrane domain to form either a
four-helix (13) or five-helix (7) bundle. In either case, these
juxtamembrane regions likely form one or more additional
R-helices that pack into the conserved hydrophobic groove
observed in the X-ray structure of syntaxin 24-150.
Our initial analysis of syntaxin 1-265, however, provided
little evidence for such a closed conformation. The earliest
protected proteolytic fragments comprised residues 1-179
and residues 180-265 (Figure 1A,B), and we first suspected
that this protection might be afforded by interactions between
FIGURE 2: Syntaxin 24-150 X-ray structure. (A) A portion of the model corresponding to the box in panel C is shown together with the
corresponding regions of the experimental 2.4 Å MAD electron density map contoured at 1.2 σ. Most of helix Ha (residues 27-57) and
residues 146-150 are omitted for clarity. The hydrogen-bonded side chains of His 58 and Gln 138 are shown in magenta. (B) σA-weighted
2Fo- Fcmap at 1.9 Å is shown together with the same portion of the model as in panel A. (C) Schematic overview of the structure.
Residues 27-150 are shown; residues 24-26 did not appear in the electron density and were not modeled. The structure is built primarily
of three R-helices: Ha (residues 28-63; blue), Hb (residues 69-106; green), and Hc (residues 111-145; red). Loop regions are shown in
gray. (D) Longitudinal view, where the model is rotated 90° from the orientation shown in panel C, with the N-terminus now toward the
viewer. These figures were created using Bobscript (46), Molscript (47), and Raster3D (48).
Structural Analysis of Syntaxin-1A
Biochemistry, Vol. 39, No. 29, 2000 8473
these fragments. However, they did not comigrate on gel
filtration columns, as might have been expected were they
associated (data not shown). Furthermore, syntaxin 180-
265 exhibited a strong tendency to aggregate, and precipitated
from digestion reaction mixtures in which this fragment was
present at concentrations of 40-100 µM (data not shown).
Thus, the protection of syntaxin 180-265 against proteolytic
digestion is not due to association with the N-terminal
domain but may instead be attributable to self-association.
The finding that syntaxin 180-265 is prone to intermolecular
association prompted us to investigate the oligomeric state
of syntaxin and whether oligomerization might affect its
Syntaxin Multimerization. Preliminary gel filtration experi-
ments (not shown) suggested that syntaxin 1-265 formed
several self-associated species. To characterize further the
nature and energetics of oligomeric syntaxin states, we used
analytical equilibrium ultracentrifugation. In addition to
syntaxin 1-265, we also analyzed syntaxin 1-253, a product
of botulinum neurotoxin C cleavage (30) that previous work
had suggested might be less susceptible to aggregation (7,
17). Data were collected at four rotor speeds (8000-20000
rpm) and four protein loading concentrations (2-100 µM)
to evaluate various self-association models.
We focus first on a discussion of the results for syntaxin
1-253 as this fragment exhibited more limited self-associa-
tion behavior than did syntaxin 1-265. For syntaxin 1-253,
self-association is evident from single-species analysis in
which the molecular mass is treated as a fitting parameter.
A diagnostic feature for multiple states in solution is that
the apparent molecular mass decreases as a function of
increasing rotor speed (Table 2). The fact that the additional
states are a consequence of self-association follows from the
observation that the apparent molecular mass increases from
40 800 ( 3000 Da to 60 900 ( 3400 Da over the range of
2-100 µM (data not shown); the monomer molecular mass
is 29 307 Da. Similar results are obtained for syntaxin
1-265, with the apparent molecular mass increasing from
64 300 ( 4900 to 83 400 ( 8300 over this same con-
centration range (monomer molecular mass of 30 751 Da).
These data, in agreement with gel filtration results (not
shown) and previous NMR studies (7), indicate that syntaxin
1-265 is more prone to oligomerization than syntaxin
The single-species analyses prompted us to evaluate a
series of models for the oligomeric state(s) of syntaxin
1-253. The square root of variance (SRV) was monitored
as a probe of the quality of the fits of various models to the
data (Table 2), with a lower SRV indicating a better model.
At high rotor speeds, we find that the minimal model which
fits the data is a monomer-dimer (1 T 2) model (Table 2).
The equilibrium dissociation constant, K21, for this reaction
is 9.9 ( 4.6 µM, predicting the formation of a stable dimer.
However, it is obvious that this model is insufficient to
describe the data at lower rotor speeds. The SRVs for this
model are poorer at lower speeds and seem to favor a higher-
order, 1 T 3 model. While this observation may, at first,
seem inconsistent, it is possible to reconcile the failure of a
two-state model analysis by considering a more complex
three-state model which includes the 1 T 2 model with the
addition of a higher-order oligomeric state. The trend, a
decreasing order of the single self-associated state as a
function of increasing speed, helps to support this conclusion.
Among the three-state models that were considered, a 1
T 3 T 6 model can be ruled out as this model is no better
than the simpler 1 T 3 model and is significantly worse
than the two-state 1 T 2 model, as judged by the SRVs
reported at 20 000 rpm. In contrast, the SRVs for either a 1
T 2 T 3 or a 1 T 2 T 4 model, in general, are lower than
that of any two-state model. While we could not rule out
the 1 T 2 T 3 model on the basis of the SRVs alone, this
model is less likely because it predicts no significant amount
of dimer and, thus, is in disagreement with the amount of
dimer that is predicted from the reasonable fit of the 1 T 2
model to the higher-speed data. In contrast, the 1 T 2 T 4
model predicts amounts of dimer similar to that with the 1
T 2 model. Therefore, we favor the 1 T 2 T 4 model.
An identical model analysis of syntaxin 1-265 yields
similar results, although in this case it is not possible to
distinguish with confidence among the three-state models
that are considered. Since this fragment self-associates to a
higher degree, we suspect that even more complex models
Table 1: Data Collection, Phasing, and Refinement Statistics
no. of unique
no. of total
I/σ (last shell)
2975619358 19472 18895
99.9 (99.7) 98.0 (95.1) 98.0 (94.7) 98.2 (96.4)
4.3 (16.3) 8.0 (34.9) 8.9 (35.3) 7.9 (38.9)
45.5 (9.4)18.6 (3.0) 20.0 (2.9) 19.5 (2.8)
no. of sites
no. of reflections
(no. of reflections)
0.81 (18603) 0.75 (18582) 0.80 (18162)
overall figure of merit0.75
no. of atoms (protein/water)
mean B factors (Å2) (protein/water)
bond lengths (Å)
bond angles (deg)
aLast shell of 10 shells; 1.97-1.90 Å for the native data set, and
2.28-2.20 Å for the SeMet data set.bRsymm) Σ|(Ih- 〈Ih〉)|/ΣIhover
all h, where Ihis the intensity of reflection h.cRcullis) Σ||Fph( Fp| -
|Fh||/Σ|Fph ( Fp|, where Fph and Fp are the derivative and native
structure factors, respectively, and Fh is the calculated heavy-atom
structure factor.dPhasing power ) 〈Fh〉/E, where 〈Fh〉 is the root-mean-
square heavy-atom structure factor and E is the residual lack of closure
error.eRcrystand Rfree) Σ||Fo| - |Fc||/Σ|Fo|. Rfreewas calculated using
5% of the data excluded from refinement.
8474 Biochemistry, Vol. 39, No. 29, 2000
Lerman et al.
are required, including states larger than a tetramer. Clear
evidence for higher-order aggregation is apparent in an
analysis of syntaxin 1-265 at the highest loading concentra-
tion (100 µM; data not shown). However, because it differs
from syntaxin 1-253 only in the presence of 12 additional
C-terminal residues, we hypothesize that syntaxin 1-265
forms stable dimers and tetramers in addition to higher-order
We can use these simple models to make predictions about
the stabilities of dimers and higher-order oligomeric states.
As an example, we assume a 1 T 2 T 4 model based on
our analysis of syntaxin 1-253. Equilibrium dissociation
constants derived for this model as fit simultaneously to all
the data (four rotor speeds and at least three protein loading
concentrations) indicate that syntaxin 1-265 dimers (K21)
1.4 ( 0.5 µM) are more stable than syntaxin 1-253 dimers
(K21) 5.8 ( 1.8 µM). Strikingly, syntaxin 1-265 tetramers
(K42 ) 12 ( 6 µM) are much more stable than syntaxin
1-253 tetramers (K42) 1.2 ( 0.5 mM). In Figure 4, we
show the quality of the fits using this model to the 2 µM
FIGURE 3: Syntaxin 24-150 surface properties. Each panel is organized with a ribbon diagram on the left, a molecular surface coded by
hydrophobicity in the center (green is hydrophobic and white is hydrophilic), and a molecular surface coded by electrostatic surface potential
on the right (red is acidic and blue is basic). Syntaxin 24-150 has been rotated 120° about its long axis between each successive panel. The
groove between helices Hb and Hc, proposed to bind the juxtamembrane helix of syntaxin, is outlined in magenta in panel A. The acidic
patch on helices Hb and Hc implicated in binding to synaptotagmin (see the text) is circled in yellow in panel B. These figures were created
using Molscript (47), Raster3D (48), and GRASP (49).
Table 2: Sedimentation Equilibrium Ultracentrifugation Data Analyzed as a Function of Speeda
square root of variance (×103)
syntaxin 1-253syntaxin 1-265
82 500 ( 10 400
12 000 rpm
70 000 ( 4800
16 000 rpm
57 300 ( 3300
20 000 rpm
46 300 ( 2800
112 500 ( 14 400
12 000 rpm
91 700 ( 7100
16 000 rpm
78 600 ( 4500
20 000 rpm
64 600 ( 4800
1 T 2
1 T 3
1 T 4
1 T 5
1 T 2 T 3
1 T 2 T 4
1 T 3 T 6
aThe 2, 8, 32, and 100 µM data were analyzed simultaneously at each rotor speed.bMolecular mass in daltons, Mr, is calculated from a single-
species analysis.cThe apparent molecular mass was greater than the dimer molecular mass; thus, the model could not be fit to the data.
Structural Analysis of Syntaxin-1A
Biochemistry, Vol. 39, No. 29, 2000 8475
ultracentrifugation data for both fragments (Figure 4A,B).
Although the equilibrium constants reported here are model-
dependent, it is clear that at low concentrations both syntaxin
fragments dimerize and that syntaxin 1-265 is significantly
more prone to self-association.
Self-Association Causes a Conformational Change. The
evidence presented so far shows that (i) syntaxin 180-265
aggregates, (ii) syntaxin 1-253 forms dimers and also larger
multimers, probably trimers and/or tetramers, and (iii)
syntaxin 1-265 self-associates to an even greater extent than
syntaxin 1-253. It therefore appears that, in addition to
mediating heteromeric interactions with SNAP-25 and syn-
aptobrevin, juxtamembrane regions of syntaxin mediate
homomeric interactions. Accordingly, we asked whether
conformational changes were associated with homomeric
interactions as they are with heteromeric ones (6, 12, 13,
31, 32), using limited proteolysis as a conformational probe.
Detailed studies were carried out comparing the limited
protease digestion patterns of syntaxin 1-265 and syntaxin
1-253 (Figure 5). A number of the resulting protein
fragments were identified by N-terminal sequencing and mass
Initial experiments comparing syntaxin 1-253 and syn-
taxin 1-265 were carried out at a substrate concentration
of 2 µM (Figure 5, central panels). At this concentration,
the majority of syntaxin 1-253 is monomeric, whereas the
majority of syntaxin 1-265 is dimeric. Surprisingly, syntaxin
1-253 is less susceptible to subtilisin cleavage than syntaxin
1-265, as judged by the disappearance of the full-length
protein (A1 or A2 in the central panels of Figure 5). If the
native conformation were simply destabilized by truncation,
an increase in protease sensitivity for syntaxin 1-253 would
have been anticipated. The greater protease sensitivity of
syntaxin 1-265 is due to an increased level of cleavage
between residues 179 and 180, since the products of this
cleavage reaction (C and F2) accumulate rapidly. The
accumulation of C is somewhat attenuated by its further
digestion to D and E. Taken together, however, the ac-
cumulation of C plus D and E is substantially more rapid
for syntaxin 1-265 than for syntaxin 1-253. Thus, syntaxin
1-265 is more sensitive to cleavage between residues 179
and 180 than syntaxin 1-253. Furthermore, whereas syntaxin
1-265 is cleaved preferentially between residues 179 and
180, the initial cleavage site for subtilisin on syntaxin 1-253
is between residues 220 and 221. Cleavage at this latter site
yields fragment B (residues 1-220) in Figure 5; the
corresponding C-terminal fragment is unlikely to be stable,
and in any case is too small to visualize using this gel system.
This fragment probably corresponds to the structured region
of the closed conformation (see the Discussion).
On the basis of these results, we hypothesized that the
increased protease sensitivity of syntaxin 1-265 between
residues 179 and 180 was coupled to its greater tendency to
self-associate (Figure 4). Furthermore, we reasoned that the
decreased sensitivity of syntaxin 1-253 at the same site
might be attributable to the closed conformation of the
monomeric protein. This model is depicted schematically in
Figure 6 (see also the Discussion), in which parentheses
indicate that a site is inaccessible (or less accessible) to
proteolytic cleavage. To further examine the potential
FIGURE 4: Syntaxin forms dimers and higher-order multimers at low micromolar concentrations. (A and B) Sedimentation equilibrium data
and fits derived from a 1 T 2 T 4 model. The 2 µM data for syntaxin 1-253 (A) or 1-265 (B) were fit (50) using dissociation constants
reported in the text. For comparison, we show fits assuming pure monomer (dashed lines). Nonrandom deviations in the fits are evident in
this model. If these data alone are fit to a 1 T 2 equilibrium, the fits are indistinguishable from the more complex 1 T 2 T 4 equilibrium
8476 Biochemistry, Vol. 39, No. 29, 2000
Lerman et al.
connection between syntaxin conformation and self-associa-
tion, we conducted additional proteolysis experiments at a
protein concentration favoring self-association (40 µM) and
at a protein concentration where both syntaxin constructs
are primarily monomeric (0.1 µM). For both syntaxin 1-253
and syntaxin 1-265, the site of initial proteolytic cleavage
was found to vary with protein concentration, providing
direct evidence that self-association is influencing protein
conformation in this system. In each case, high concentrations
(left panels in Figure 5) promoted cleavage between residues
179 and 180, resulting in the early accumulation of fragment
C. For example, whereas the initial cleavage of syntaxin
1-253 at lower concentrations strongly favors fragment B,
at high concentrations B and C appear with a similar time
course. Conversely, at low concentrations (right panels in
Figure 5), initial cleavage for both proteins is at the site
between residues 220 and 221, and fragment B is produced
preferentially. Furthermore, the accelerated disappearance of
syntaxin 1-265 compared to that of syntaxin 1-253 is not
seen at this low concentration, confirming that the relatively
greater protease sensitivity of syntaxin 1-265 is caused by
self-association. Thus, two lines of evidence from proteolysis
experiments indicate that conformational changes accompany
self-association: at each protein concentration, syntaxin
1-253 and syntaxin 1-265 display different protease
sensitivity, and for each protein construct, protease sensitivity
varies with protein concentration.
The structure of an N-terminal domain of syntaxin-1A was
determined at 1.9 Å resolution by MAD phasing. The
structure comprises a three-helix bundle with a conserved
hydrophobic groove. This groove in the N-terminal domain
likely interacts with juxtamembrane residues nearer the
C-terminus of the intact protein to create a “closed”
conformation. Further evidence for this closed protein
conformation comes from proteolytic data: significant ac-
cumulation of syntaxin 1-220 (B in Figure 5) is seen in all
protease protection experiments conducted under “monomer”
conditions (0.1 and 2 µM for syntaxin 1-253, and 0.1 µM
for syntaxin 1-265). This fragment corresponds to a
hypothetical closed conformation including the N-terminal
domain and a large portion of the juxtamembrane region
(Figure 6). This work therefore adds to previous evidence
suggesting that syntaxin can adopt a closed conformation
that might regulate its interactions with other proteins (7,
Previous experiments had suggested that the histidine-
tagged cytoplasmic domain of syntaxin is monomeric at
relatively high salt concentrations (300 mM) (6), while intact
syntaxin (including the transmembrane domain) forms SDS-
stable dimers (33). Here, we use equilibrium analytical
ultracentrifugation over a broad range of protein concentra-
tions to demonstrate that even the cytoplasmic domain of
untagged syntaxin self-associates under conditions that
approximate physiological ionic strength.
Formation of dimers or larger homomultimers has also
been observed for other t-SNAREs and may be biologically
significant. The cytoplasmic domains of yeast SNAREs,
including Sed5 (L. Cavanaugh and F. M. Hughson, unpub-
lished results) and Pep12 (34), have been observed to
dimerize in vitro. In addition, Drosophila Sed5 dimers are
formed in transfected COS cell membranes, as judged by
chemical cross-linking, and this dimerization requires the
FIGURE 5: Multimerization causes a conformational change in syntaxin. Recombinant syntaxin 1-253 (top) or syntaxin 1-265 (bottom)
was subjected to limited subtilisin digestion and analyzed as described in the legend of Figure 1. The initial syntaxin:subtilisin ratio was
held constant in all experiments, but three different initial substrate concentrations were tested: 40 µM (left), 2.0 µM (center), and 0.1 µM
(right). Molecular masses in kilodaltons are indicated at the far right. Syntaxin digestion products as labeled on the far left were identified
by amino-terminal sequencing and mass spectrometry as follows: (A1) 1-253, (A2) 1-265, (B) 1-220, (C) 1-179, (D) 24-179, (E)
24-150, (F1) 180-253, and (F2) 180-265.
juxtamembrane region. Closed monomers, represented schematically
as four-helix bundles, are preferentially cleaved by subtilisin
between residues 220 and 221, while open dimers are preferentially
cleaved between residues 179 and 180. Cleavage between residues
179 and 180 is suppressed in monomers by unknown structural
features. Little if any cleavage between residues 220 and 221 is
observed in multimers; instead, syntaxin 180-265 resulting from
initial cleavage is strongly protected from further digestion, as it is
in syntaxin-SNAP-25-synaptobrevin complexes (9, 40).
Model for syntaxin dimerization mediated by the
Structural Analysis of Syntaxin-1A
Biochemistry, Vol. 39, No. 29, 2000 8477
juxtamembrane domain (35). Most models for SNARE
function have suggested that the hetero-oligomerization of
v- and t-SNAREs on different membranes plays a central
role in the docking and fusion of vesicles with organelles or
the plasma membrane (36, 37). Nonetheless, several observa-
tions have suggested that t-SNAREs alone can suffice,
presumably by forming homomeric complexes that link
membranes. For example, homotypic vacuole fusion is only
about 2-fold less efficient in vitro when both partners carry
only t-SNAREs than it is when one partner carries only
v-SNAREs and the other carries only t-SNAREs (38).
Similarly, the t-SNARE Ufe1 has been shown to oligomerize
in vitro and has been proposed to mediate homotypic fusion
among endoplasmic reticulum membranes via t-SNARE-
t-SNARE interactions (39). Though no role for syntaxin
oligomers in neurotransmitter release has yet been docu-
mented, our results represent the most quantitative analysis
to date of t-SNARE self-association. Furthermore, the
growing evidence for t-SNARE homo-oligomerization points
toward an additional level of promiscuity in SNARE part-
nering that may be subject to regulatory control in vivo.
Sytaxin undergoes a conformational change upon self-
association (Figure 6). The proteolytic sensitivity of the
cleavage site between residues 179 and 180 appears to serve
as a readout for this change. Strikingly, the same site
becomes sensitive when heteromeric SNARE complexes are
formed (9). Thus, self-association seems to be accompanied
by a conformational opening of the closed syntaxin mono-
Guided by our analysis of protease sensitivitiy as well as
the known structural features of heteromeric SNARE com-
plexes, we propose a model for open syntaxin dimers (Figure
6). It seems plausible that the juxtamembrane heptad repeats
assemble to form parallel bundles in syntaxin multimers, as
seen in heteromeric complexes. Syntaxin 180-265 is pro-
tease-resistant under conditions where syntaxin is primarily
self-associated (Figure 5), and very similar syntaxin frag-
ments are protease-resistant in heteromeric complexes (3,
9, 40). Association between transmembrane domains, not
present in our study, would further stabilize the parallel
helical bundles in homomeric SNARE complexes (see ref
33). Peptides as short as 32 residues derived from the
juxtamembrane region of syntaxin display significant R-helix
content by CD at concentrations of 30-60 µM (41),
presumably due to multimeric associations such as those
proposed here. Flexibly attached N-terminal domains, as
observed for heteromeric complexes (1, 12), would account
for the increased proteolytic susceptibility of the bond
between residues 179 and 180. While this model is consistent
with the available data, an alternative self-association model
involving domain swapping (see ref 42 for review) cannot
be ruled out at present. Nonetheless, the parallel bundle
model seems more consistent with what is known about
heteromeric SNARE complexes, with the protection against
further proteolysis of residues 180-265, and with the dimer-
destabilizing effect of deleting residues 254-265.
Are there structural constraints that argue against the
formation of t-SNARE complexes in which the helical bundle
resembles that of heteromeric SNARE complexes? In the
only published SNARE complex structure (3), four ap-
proximately 85-residue protein fragments derived from
syntaxin (residues 180-262), SNAP-25, and syntaptobrevin
form an elongated bundle of parallel R-helices with a
generally hydrophobic core. The bundle contains a single
“layer” of polar core residues that serve to determine the
register of the four helices (3, 8). Synaptobrevin contributes
an Arg residue to this layer, the guanidino group of which
is buried within the core of the four-helix bundle. This buried
positive charge is neutralized by partial negative charges
carried on the side chain oxygens of glutamine side chains
contributed by each of the other three helices (3). It has been
suggested that most SNARE complexes would contain one
arginine-containing SNARE and three glutamine-containing
SNAREs (43). In this model, homomeric SNARE complexes
containing four glutamine-containing SNAREs would be
disallowed. However, all-glutamine layers are not uncommon
in coiled coil cores (unpublished observations), presumably
because the partial positive and partial negative characters
of each glutamine side chain can compensate one another
cyclically around the bundle. Thus, parallel helical bundles
with two or more syntaxin molecules are not implausible
on first principles.
The stoichiometry of syntaxin-SNAP-25 complexes in
vitro is 2:1 (31). Therefore, it might be anticipated that
syntaxin dimers would be on the kinetic pathway to syn-
taxin-SNAP-25 complexes. Furthermore, conformational
opening of the yeast syntaxin Sso1 accelerates SNARE
complex assembly 2000-fold (12), further suggesting that
the open syntaxin dimers might form complexes with SNAP-
25 more readily than the closed monomers. Nonetheless, the
rate of assembly with SNAP-25 is only subtly influenced
by the conformational opening mediated by syntaxin dimer-
ization (F. M. Hughson, unpublished results). Therefore,
contrary to expectation, it appears that the rate-limiting step
for assembly in vitro is not strongly sensitive to the
oligomeric state of syntaxin. It will be worthwhile in future
work to examine whether 1:1 and 2:1 syntaxin-SNAP-25
complexes bind the v-SNARE syntaptobrevin with equal
efficiency, or whether the need to displace a syntaxin
molecule from the 2:1 complexes slows the formation of the
ternary core complex.
Many intracellular trafficking steps appear to require a
member of the Sec1 protein family (see ref 44 and references
therein). Sec1 family members interact with syntaxin ho-
mologues and/or ternary SNARE complexes (7, 16, 44, 45),
but their molecular roles in vesicle targeting and fusion have
been controversial. In vitro, nSec1 binds to a closed form of
syntaxin and prevents it from forming heteromeric SNARE
complexes (7, 15, 16, 45). Our experiments suggest strongly
that the closed conformation prevents syntaxin multimer-
ization. Thus, an important function of nSec1 binding may
be to block the formation of homomeric syntaxin complexes.
nSec1 is well-suited to this role since it binds directly to the
juxtamembrane region of syntaxin that mediates multimer-
ization and since it holds syntaxin in a closed conformation
incompatible with multimer formation. nSec1 may even serve
to chaperone the assembly of the proper SNARE complexes.
We thank Larisa Sereda for early contributions to this
project, Vibha Rao and the staff of the National Synchrotron
Light Source beamline X12C for advice and assistance in
X-ray data collection, Saw Kyin for DNA sequencing and
8478 Biochemistry, Vol. 39, No. 29, 2000
Lerman et al.
protein microchemistry, Jim Lear for curve fitting procedures, Download full-text
Phyllis Hanson for reagents, and Bill Weis for sharing a
manuscript prior to publication. We gratefully acknowledge
helpful input from Yigong Shi, Gerry Waters, Jannette Carey,
Jeff Gerst, Phyllis Hanson, Graham Warren, and members
of the Hughson laboratory.
1. Hanson, P. I., Roth, R., Morisaki, H., Jahn, R., and Heuser, J.
E. (1997) Cell 90, 523-535.
2. Weber, T., Zemelman, B. V., McNew, J. A., Westermann, B.,
Gmachl, M., Parlati, F., So ¨llner, T. H., and Rothman, J. E.
(1998) Cell 92, 759-772.
3. Sutton, R. B., Fasshauer, D., Jahn, R., and Brunger, A. T.
(1998) Nature 395, 347-353.
4. Jahn, R., and Su ¨dhof, T. C. (1999) Annu. ReV. Biochem. 68,
5. Bennett, M. K., Calakos, N., and Scheller, R. H. (1992) Science
6. Fasshauer, D., Otto, H., Eliason, W. K., Jahn, R., and Bru ¨nger,
A. T. (1997) J. Biol. Chem. 272, 28036-28041.
7. Dulubova, I., Sugita, S., Hill, S., Hosaka, M., Fernandez, I.,
Sudhof, T. C., and Rizo, J. (1999) EMBO J. 18, 4372-4382.
8. Weimbs, T., Low, S. H., Chapin, S. J., Mostov, K. E., Bucher,
P., and Hofmann, K. (1997) Proc. Natl. Acad. Sci. U.S.A. 94,
9. Fasshauer, D., Eliason, W. K., Brunger, A. T., and Jahn, R.
(1998) Biochemistry 37, 10354-10362.
10. Fasshauer, D., Antonin, W., Margittai, M., Pabst, S., and Jahn,
R. (1999) J. Biol. Chem. 274, 15440-15446.
11. Calakos, N., Bennett, M. K., Peterson, K. E., and Scheller, R.
H. (1994) Science 263, 1146-1149.
12. Nicholson, K. L., Munson, M., Miller, R. B., Filip, T. J.,
Fairman, R., and Hughson, F. M. (1998) Nat. Struct. Biol. 5,
13. Fiebig, K. M., Rice, L. M., Pollock, E., and Brunger, A. T.
(1999) Nat. Struct. Biol. 6, 117-123.
14. Betz, A., Okamoto, M., Benseler, F., and Brose, N. (1997) J.
Biol. Chem. 272, 2520-2526.
15. Pevsner, J., Hsu, S.-C., Braun, J. E. A., Calakos, N., Ting, A.
E., Bennett, M. K., and Scheller, R. H. (1994) Neuron 13,
16. Yang, B., Steegmaier, M., Gonzalez, L. C., and Scheller, R.
H. (2000) J. Cell Biol. 148, 247-252.
17. Fernandez, I., Ubach, J., Dulubova, I., Zhang, X., Su ¨dhof, T.
C., and Rizo, J. (1998) Cell 94, 841-849.
18. Yang, B., Gonzalez, L., Prekeris, R., Steegmaier, M., Advani,
R. J., and Scheller, R. H. (1999) J. Biol. Chem. 274, 5649-
19. Tsui, M. M., and Banfield, D. K. (2000) J. Cell Sci. 113, 145-
20. MacFerrin, K. D., Chen, L., Terranova, M. P., Schreiber, S.
L., and Verdine, G. L. (1993) Methods Enzymol. 217, 79-
21. Leahy, D. J., Erickson, H. P., Aukhil, I., Joshi, P., and
Hendrickson, W. A. (1994) Proteins 19, 48-54.
22. Thogersen, H. C., and Nagai, K. (1987) Methods Enzymol.
23. Otwinowski, Z., and Minor, W. (1998) Methods Enzymol. 276,
24. Collaborative Computational Project Number 4 (1994) Acta
Crystallogr. D50, 760-763.
25. Bru ¨nger, A. T. (1992) X-PLOR (Version 3.1): A System for
X-ray Crystallography and NMR, Yale University Press, New
26. Brunger, A. T., Adams, P. D., Clore, G. M., Delano, W. L.,
Gros, P., Grosse-Kunstleve, R. W., Jiang, J.-S., Kuszewski,
J., Nilges, N., Pannu, N. S., Read, R. J., Rice, L. M., Simonson,
T., and Warren, G. L. (1998) Acta Crystallogr. D54, 905-
27. Jones, T. A., Zou, J.-Y., Cowan, S. W., and Kjeldgaard, M.
(1991) Acta Crystallogr. A47, 110-119.
28. Laue, T. M., Shah, B. D., Ridgeway, T. M., and Pelletier, S.
L. (1992) in Analytical Ultracentrifugation in Biochemistry
and Polymer Science (Harding, S. E., Rowe, A. J., and Horton,
J. C., Eds.) pp 90-125, The Royal Society of Chemistry,
29. Tjandra, N., and Bax, A. (1997) Science 278, 1111-1114.
30. Schiavo, G., Shone, C. C., Bennett, M. K., Scheller, R. H.,
and Montecucco, C. (1995) J. Biol. Chem. 270, 10566-10570.
31. Fasshauer, D., Bruns, D., Shen, B., Jahn, R., and Bru ¨nger, A.
T. (1997) J. Biol. Chem. 272, 4582-4590.
32. Rice, L. M., Brennwald, P., and Bru ¨nger, A. T. (1997) FEBS
Lett. 415, 49-55.
33. Margittai, M., Otto, H., and Jahn, R. (1999) FEBS Lett. 446,
34. Tishgarten, T., Yin, F. F., Faucher, K. M., Dluhy, R. A., Grant,
T. R., Fischer von Mollard, G., Stevens, T. H., and Lipscomb,
L. A. (1999) Protein Sci. 8, 2465-2473.
35. Banfield, D. K., Lewis, M. J., Rabouille, C., Warren, G., and
Pelham, H. R. B. (1994) J. Cell Biol. 127, 357-371.
36. Hughson, F. M. (1999) Curr. Biol. 9, R49-R52.
37. Mayer, A. (1999) Curr. Opin. Cell Biol. 11, 447-452.
38. Nichols, B. J., Ungermann, C., Pelham, H. R. B., Wickner,
W. T., and Haas, A. (1997) Nature 387, 199-202.
39. Patel, S. K., Indig, F. E., Olivieri, N., Levine, N. D., and
Latterich, M. (1998) Cell 92, 611-620.
40. Poirer, M. A., Hao, J. C., Malkus, P. N., Chan, C., Moore, M.
F., King, D. S., and Bennett, M. K. (1998) J. Biol. Chem.
41. Zhong, P., Chen, Y. A., Tam, D., Chung, D., Scheller, R. H.,
and Miljanich, G. P. (1997) Biochemistry 36, 4317-4326.
42. Schlunegger, M. P., Bennett, M. J., and Eisenberg, D. (1997)
AdV. Protein Chem. 50, 61-122.
43. Fasshauer, D., Sutton, R. B., Brunger, A. T., and Jahn, R.
(1998) Proc. Natl. Acad. Sci. U.S.A. 95, 15781-15786.
44. Carr, C. M., Grote, E., Munson, M., Hughson, F. M., and
Novick, P. J. (1999) J. Cell Biol. 146, 333-344.
45. Misura, K. M. S., Scheller, R. H., and Weis, W. I. (2000)
Nature 404, 355-362.
46. Esnouf, R. M. (1997) J. Mol. Graphics Modell. 15, 132-134.
47. Kraulis, P. (1991) J. Appl. Crystallogr. 24, 924-950.
48. Merritt, E. A., and Bacon, D. J. (1997) Methods Enzymol. 277,
49. Nicholls, A., Sharp, K. A., and Honig, B. (1991) Proteins 11,
50. Johnson, M. L., Correia, J. J., Yphantis, D. A., and Halverson,
H. R. (1981) Biophys. J. 36, 575-588.
Structural Analysis of Syntaxin-1A
Biochemistry, Vol. 39, No. 29, 2000 8479