From the Department of Cell Biology and Physiology, Washington University School of Medicine, St. Louis, Missouri 63110
Received for publication, February 25, 2003 , and in revised form, April 24, 2003.
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
![]() |
INTRODUCTION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
-SNAP (a ubiquitous SNAP isoform) serves as the requisite link
between NSF and SNARE complexes. It binds to SNARE complexes, and together
they recruit NSF (4).
SNARE-bound
-SNAP stimulates ATP hydrolysis by NSF, leading to
conformational changes and concomitant disassembly of the SNARE complex
(2,
4). Scanning transmission
electron microscopy has been used to define the probable composition of the
complex containing
-SNAP, NSF, and SNARE complex, also referred to as
20 S complex (5). Each 20 S
complex consists of one SNARE complex, three
-SNAPs, and one NSF
hexamer. Interestingly,
-SNAP only binds NSF in the absence of SNARE
complex when forced into an oligomeric form by fusion to an intrinsically
trimeric protein (5) or when it
is immobilized on plastic (6).
Electron micrographs of
-SNAP bound to SNARE complex and an antibody
specific for the N terminus of
-SNAP show that
-SNAP binds SNARE
in an antiparallel orientation. This positions its N terminus near the
membrane and its C terminus away from the membrane, where it can interact with
NSF (2,
7). Consistent with this, the
-SNAP C terminus has been shown to play a critical role in stimulating
NSF ATPase activity and promoting SNARE complex disassembly
(8).
Despite limited overall sequence similarity among SNAREs on different
membranes, all SNARE complexes so far examined bind -SNAP and can be
disassembled by NSF
(914).
SNARE complexes are stable
-helical structures containing specific
combinations of four SNARE helices (for review, see Ref.
15). Crystal structures of the
synaptic and late endosomal SNARE complexes show that these two complexes are
rod-shaped and held together by interactions among conserved, mostly
hydrophobic residues within the core of the complex
(1618).
Although both complexes have largely acidic surface potentials, there are few
specifically conserved residues on the outer, solvent-exposed, surfaces
(19,
20).
Little is known about how -SNAP recognizes the variety of SNARE
complexes involved in different membrane fusion reactions. Based on deletion
studies (8,
2124)
and electron micrographs in which
-SNAP molecules appear to ensheathe
the SNARE complex (7), a
significant portion of
-SNAP is thought to participate in its
interaction with the SNARE complex. The structure of the Saccharomyces
cerevisiae
-SNAP orthologue, Sec17p
(25), provides some clues to
where the interfaces between SNAPs and SNARE complexes might be. Sec17p is a
14-helix
/
protein with two principal domains, an N-terminal
twisted sheet (nine
-helices arranged in antiparallel to form a
sheet-like structure) and a C-terminal globular bundle
(25). Each helix in the
twisted sheet is slightly askew from its neighbors, giving the domain a
concave face whose curvature complements that of the convex surface of the
SNARE complex. One edge of the twisted sheet is longer than the other and
curved in a way that could allow it to fit into the shallow grooves that lie
between individual helices of the SNARE complex
(25). Both the concave face
and the longer edge of the twisted sheet contain residues that are conserved
among SNAP orthologues (25).
Shape complementarity, sequence conservation, and overall surface charge
distribution led to the proposal that either the concave face or longer edge
of the Sec17p (or the
-SNAP) twisted sheet domain might be responsible
for SNARE complex binding
(25). Whether this is the case
and, if so, how
-SNAP actually contacts SNARE complexes has not been
experimentally examined.
In this report, we present evidence showing that the SNARE complex binding
surface -SNAP is the concave face of its twisted sheet domain. In a
coimmunoprecipitation-based binding assay, point mutations in charged residues
on this surface strongly affect SNARE complex binding, whereas mutations
elsewhere do not. Most of these mutations similarly affect the ability of
-SNAP to promote SNARE complex disassembly by NSF. Based on these
results we present a model of how
-SNAP interacts with SNARE complexes
and discuss its implications for SNARE complex disassembly.
![]() |
EXPERIMENTAL PROCEDURES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Cloning and Site-directed MutagenesisBovine -SNAP
DNA (from pQE-9
-SNAP) was cloned into
NdeI/HindIII-digested pET28 (Novagen) to generate a
His6-
-SNAP expression construct. The resulting protein had
the sequence MGSSHHHHHHSSGLVPRGSH at its N terminus. During sequencing we
found that the sequence of pQE-9
-SNAP differed from the published
bovine
-SNAP sequence (PubMed 228 accession number AAB25812
[GenBank]
; Ref
26) at residues 7071
(HV
QL) and 243244 (RI
LM). With these changes, the
bovine
-SNAP sequence matches human
-SNAP at all four residues
and bovine
-SNAP at two residues. All sequencing was performed using a
cycle-sequencing kit (Big Dye version 2; ABI Systems) and processed by the
Protein and Nucleic Acid Laboratory (Washington University).
Eleven -SNAP mutants (F32G, E39A, E43A, K56A, D87A, K93A, R116A,
K140A, E155A, K163A, and Y200K) were generated by PCR using an overlap
extension method with pQE-9
-SNAP as template and subcloned into pET28.
The remaining mutants (E39K/E40K, R47A, K53A, K53E, Q74A, D80A, K94A, E99A,
E109A, K122A, E129A, E132A, E134A, D150A, E156A, K167A, K203A, and D273R) were
generated using the QuikChange mutagenesis kit (Stratagene) with wild-type
pET28
-SNAP as template.
Because of a PCR error, some of the SNAP mutants (E39K/E40K,
Q74A, K94A, E99A, E109A, E129A, D150A, K203A, and D273R) contained a C84S
mutation. However, wild-type and C84S
-SNAP were functionally
equivalent in our assays (data not shown), and the mutation was, therefore,
left in place.
Protein Expression and Purification-SNAP was
expressed in Escherichia coli BL21 (DE3) cells. 12 liters of
Terrific Broth cultures were grown 1014 h at 30 °C to
A600 =
2 and induced for 24 h with 0.4
mM isopropyl-1-thio-
-D-galactopyranoside.
Bacterial pellets were resuspended in lysis buffer (20 mM Tris, 250
mM NaCl, 1 mM phenylmethylsulfonyl fluoride, 5%
glycerol, pH 7.4) and stored at 20 °C. After thawing, pellets were
sonicated (2 x 30 s), Triton X-100 was added to a final concentration of
0.1%, and lysates were centrifuged for 20 min at 18,500 x g.
-SNAP was purified from the supernatant by binding to nickel
nitrilotriacetic acid-agarose (Qiagen) and eluting with imidazole. After
dialysis into Mono Q buffer (20 mM Tris, 100 mM NaCl, 1
mM dithiothreitol, pH 7.4)
-SNAP was further purified on a
Mono Q anion exchange column (Amersham Biosciences). Fractions containing
-SNAP were pooled and quantitated by measuring absorbance at 280 nm
(
= 40,200
M1cm1 in 20
mM Tris, 100 mM NaCl, pH 7.4; determined by quantitative
amino acid analysis). Typical preparations had a concentration of 50100
µM and were snap-frozen and stored at 70 °C.
Full-length His6-tagged CHO NSF was expressed and purified as previously described (2). SNARE complex was assembled by mixing three purified SNAREs (rat His6-synaptobrevin II (196) (2), mouse SNAP-25b-His6-Myc (2), and rat syntaxin1a (184265)-His6) together at equimolar ratios in Mono Q buffer. After overnight assembly, SNARE complex was separated from unincorporated individual SNAREs by chromatography on a Mono Q column. Column fractions were analyzed by SDS-PAGE either with or without boiling; fractions containing SDS-resistant complex were pooled, quantitated using a Bradford assay with BSA as standard, and snap-frozen for storage.
Circular Dichroism SpectroscopyCircular dichroism (CD)
spectra were collected on a Jasco J600 spectropolarimeter; each spectrum was
the average of five scans from 190 to 250 nm in 0.4-nm increments at 50
nm/min. Wild-type and mutant -SNAP proteins were diluted to 2.9
µM in 20 mM Tris, 100 mM NaCl, pH 7.4.
Measurements were taken at room temperature using a quartz cell with a 1-mm
path length. Mean residue ellipticity (degree x cm2/dmol) was
calculated after subtracting the signal from buffer alone.
-Helical
content was calculated using the mean residue ellipticity at 222 nm
(
222 nm) and the assumption that for 100%
-helix,
222 nm = 39,500 (12.57/n), where
n is the average number of residues per helix
(27). Based on homology with
the Sec17p structure (25),
n for
-SNAP is 15 residues.
Immunoprecipitation-based SNARE Complex Binding
Assay-SNAP (concentrations are noted in Figs.
1 and
3), Myc-tagged SNARE complex (1
µM), and anti-Myc IgG (0.04 mg/ml final concentration
(28)) were combined in binding
buffer (20 mM Hepes, 100 mM NaCl, 5 mM
imidazole, 0.2% Triton X-100, pH 7.6) and mixed with protein G-Sepharose
(Amersham Biosciences) for a total volume of 100 µl in silane-treated
polypropylene tubes. After 40 min of shaking at room temperature, samples were
washed twice in 1 ml of binding buffer for a total washing time of
30 s.
Samples lacking SNARE complex were used to measure nonspecific binding. 10
µl of gel loading buffer (100 mM Tris, pH 7.6, 4% SDS, 0.2%
bromphenol blue, 25% glycerol, 10% 2-mercaptoethanol) was added to the beads
in each tube, tubes were boiled for 3 min, and each reaction was loaded onto
an 11% Tris-Tricine polyacrylamide gel, modified from Schagger and Von Jagow
(29) (Bio-Rad acrylamide
solution 37.5:1 acrylamide:bisacrylamide or 30% total monomer, 2.6%
cross-linker). Gels were run in standard Tricine buffers (anode 0.2
M Tris, pH 8.9; cathode 0.1 M Tris, 0.1 M
Tricine (Serva), 0.1% SDS), placed in 10% methanol, 7.5% acetic acid to strip
SDS from the gel, incubated 30 min in 0.05% SDS, and finally stained with
SYPRO Red (Molecular Probes) according to the manufacturer's instructions.
Images were collected using a STORM 860 scanner (Molecular Dynamics);
fluorescence was detected using a 635-nm diode laser and a 650-nm long-pass
filter. Staining was quantitated using ImageQuant software (Molecular
Dynamics).
|
|
Fluorescence Resonance Energy Transfer (FRET)-based SNARE Complex Disassembly AssayFluorescently tagged SNARE proteins were used to build a SNARE complex in which FRET could be used to monitor SNARE complex assembly and disassembly.2 Briefly, SNARE complex consisting of syntaxin 1A (1265)-cyan fluorescent protein (CFP), synaptobrevin II (196)-yellow fluorescent protein (YFP), and SNAP-25b was assembled and purified using a Mono Q column as for the SNARE complex above. SNARE complex fluorescence was monitored using a SPEX Fluorolog 3 with excitation at 433 nm and emission at 477 nm (CFP emission) and 527 nm (YFP emission). Slits were 3 nm. The ratio of YFP to CFP emission (YFP527/CFP477) was monitored as a function of time. Fully assembled SNARE complex had a YFP/CFP ratio of 2.2, whereas dissociated SNAREs had a YFP/CFP ratio of 0.6.
To examine the requirements for -SNAP in SNARE disassembly, SNARE
complex (50 nM) was incubated with
-SNAP (250 nM)
for 10 min at 30 °C in 500 µl of fluorimetry buffer (30 mM
Hepes, 100 mM glutamic acid, 10 mM MgCl2, 2
mM ATP, 0.1 mg/ml bovine trypsin inhibitor, pH 7.6). A kinetic scan
was then started. After obtaining a base line, disassembly was initiated by
adding NSF (0.5 µg = 2nM). The ratio of YFP/CFP emission was
measured every 3 s. The initial rate of SNARE complex disassembly was derived
by fitting a straight line to the first 10 data points collected.
-SNAP Structure ModelA structural model of bovine
-SNAP was constructed with the SWISS-MODEL Protein Homology Modeling
Server (30)
(www.expasy.org/swissmod/SWISS-MODEL.html)
using the coordinates of S. cerevisiae Sec17p (Protein Data Bank
(PDB) code 1QQE
[PDB]
) and a sequence of bovine
-SNAP adjusted to compensate
for a shorter N-terminal sequence in Sec17p, gaps and insertions in loops
between helices, and a loop between helices 11 and 12 that is disordered in
the Sec17p structure.
-SNAP-SNARE Complex Binding ModelProtein Data Bank
files for the
-SNAP comparative homology model and the synaptic SNARE
complex (one of the three in the unit cell, PDB code 1SFC
[PDB]
) were combined using
the Swiss Protein Data Bank Viewer
(30). Basic residues in
-SNAP whose mutations reduced SNARE complex binding were manually
aligned with three sets of conserved acidic residues on the SNARE complex
surface (Fig. 7). Side chain
torsion angles were modified to maximize the number of interprotein hydrogen
bonds without producing steric clashes. Modeling was done both in the presence
and absence of
-SNAP C-terminal globular domain; a substantially better
fit was obtained when the C-terminal domain was allowed to move and left out
of the model. Features of the model are described elsewhere (see
"Discussion").
|
|
![]() |
RESULTS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
At high -SNAP concentrations, approximately three
-SNAPs were
bound per SNARE complex (Fig.
1). SYPRO dyes stain different proteins differently, rendering
measurements of stoichiometry only semiquantitative
(31,
32). It is, however, clear
that more than one, and most likely three,
-SNAPs bind to a single
SNARE complex. This is consistent with early estimates of the stoichiometry
(22) and with calibrated
scanning transmission electron microscopy analysis of the 20 S complex, which
suggests that it contains one SNARE complex, one NSF, and three molecules of
-SNAP (5). Differences
between the three
-SNAP binding sites on the SNARE complex are not
apparent in our binding data.
Interestingly, binding of -SNAP to SNARE complex was sharply reduced
when the concentration of NaCl in the assay was increased from 100 to 500
mM (Fig. 1). This
demonstrates that electrostatic interactions are important for
-SNAP
binding to the SNARE complex. It also confirms the specificity of binding
detected in our assay, even at the highest concentrations of
-SNAP.
Design and Production of -SNAP MutantsTo
define functionally important contacts between
-SNAP and the synaptic
SNARE complex, we generated a series of
-SNAP mutants based on a
homology model of
-SNAP. We built this model using the crystal
structure of the S. cerevisiae
-SNAP orthologue, Sec17p
(25). Sec17p is 35% identical
and 15% similar to
-SNAP and can functionally replace
-SNAP in
in vitro Golgi transport assays
(6). To provide evidence that
the homology model reasonably predicts the actual
-SNAP structure, we
used circular dichroism (CD) spectroscopy to estimate
-SNAP helical
content (Fig. 2). Based on its
mean residue ellipticity at 222 nm,
-SNAP is
70% helical. The
helical content of Sec17p predicted from its crystal structure
(25) is 73%, and that of
-SNAP from the homology model is 74%. The close correlation between the
observed and predicted helical content of these proteins both substantiates
the
-SNAP homology model and provides independent evidence that the
Sec17p crystal structure reflects its structure in solution.
|
As discussed in the introduction, the nine N-terminal -helices in
Sec17p form a twisted sheet domain which has two surfaces that are good
candidates for SNARE complex binding. These two surfaces, the concave face and
longer edge, are also found on the
-SNAP homology model. The concave
face, which is part of the front face of the protein, is complementary in
shape and charge to the convex, mostly acidic surface of the SNARE complex,
and many of its exposed residues are strongly conserved among SNAP orthologues
(25). The longer edge of the
twisted sheet also has several conserved residues, and its curvature
complements that of the shallow grooves on the surface of the SNARE complex
that run between the SNARE helices.
The SNAP-SNARE complex interface is, therefore, likely to involve either
the front face or the longer edge of the -SNAP twisted sheet domain.
Because of this and because the salt sensitivity of binding
(Fig. 1) indicated that
electrostatic interactions are important for
-SNAP binding to the SNARE
complex, we introduced point mutations at all conserved charged residues on
these two surfaces (Fig. 3). In
addition, we generated mutations at three conserved residues (Tyr-200,
Lys-203, and Asp-273) on the edge of the globular bundle near the twisted
sheet and at one conserved hydrophobic residue (Phe-32) on the longer edge of
the twisted sheet. Conservation on the back face is less extensive than on the
front face, but several charged and polar residues are moderately conserved.
To examine the role of the back face in interaction with the SNARE complex, we
mutated five of these. Most mutations were to alanine to remove the charge
from the side chain without imposing new structural constraints.
Because we introduced mutations only at positions predicted to be
solvent-exposed, we did not expect the mutations to cause significant
structural disturbances. Indeed, as with wild-type -SNAP, mutant
proteins could be expressed at high levels in E. coli and were not
proteolyzed during growth or purification. All were soluble at high
concentrations (50100 µM). The one exception was
-SNAP(D273R), which was largely insoluble in E. coli and was,
therefore, not included in the rest of this study. To further confirm that the
mutations did not change the
-SNAP structure, we compared CD spectra of
wild-type
-SNAP and each of the mutants used in this study
(Fig. 2). All of the proteins
showed similar helical content, with an average mean residue ellipticity at
222 nm of 24160 x 1280 degree x cm2/dmol. In
addition, we treated a subset of the mutants with limited concentrations of
trypsin. There were no detectable differences in the proteolytic patterns of
wild-type, F32G, R116A, and E155A
-SNAP (data not shown).
SNARE Complex Binding by -SNAP Surface
MutantsTo compare binding of wild-type and mutant
-SNAP to
SNARE complex, we carried out binding reactions as described in
Fig. 1. A representative
experiment is shown in Fig. 4.
-SNAP(K122A) did not bind appreciably to SNARE complex at
concentrations of up to 15 µM. In contrast,
-SNAP(D80A)
bound with higher affinity than wild-type
-SNAP to the SNARE complex,
with an EC50 of
2.5 µM. Both decreased and
increased binding were readily apparent when 7.5 µM
-SNAP
was added to the complex in this and similar experiments. We, therefore, used
this concentration to compare SNARE complex binding by all
-SNAP
mutants to that of wild-type
-SNAP
(Fig. 5).
|
The effects of the mutations fell into three categories: depressed binding
(<50% of wild-type -SNAP binding; colored blue in
Fig. 5), enhanced binding
(>130% that of wild-type
-SNAP; colored red in
Fig. 5), and binding that was
not very different from that of wild-type
-SNAP (colored
yellow in Fig. 5).
Nine of the 11 mutations that reduced binding were of basic residues on the
front face of
-SNAP. In contrast, mutations of seven acidic residues on
the front face enhanced SNARE complex binding. Mutations that did not affect
SNARE complex binding included changes in all residues on the back face and
four of the five residues on the longer curved edge of
-SNAP. Together
these data indicate that much of the
-SNAP front face and, in
particular, the concave face of the twisted sheet is involved in SNARE complex
binding. The extensive binding surface defined here is consistent with the
large minimal binding domain predicted by previous studies of truncation
mutants (8,
2124).
Although most mutations were alanine substitutions, three mutations were
different. Two of these included mutations to lysine that had opposite effects
on binding. Changing Glu-39 and Glu-40 to lysine (E39K/E40K) enhanced binding
as had alanine mutations at acidic residues on the front face. In contrast, a
Y200K mutation caused -SNAP to bind poorly to the SNARE complex,
suggesting that nonionic interactions involving this residue are important for
SNARE complex binding. Finally, a glycine mutation at Phe-32 led to only a
small decrease in SNARE complex binding. This was unexpected because Phe-32 is
the only conserved residue not on the back face among the first 35 amino acids
of
-SNAP, a region reported to be critical for SNARE complex binding
(24).
SNARE Complex Disassembly Directed by -SNAP
Mutants
-SNAP recruits NSF to disassemble SNARE complexes.
It is, therefore, likely that mutations that affect
-SNAP binding to
SNARE complex will also affect its ability to mediate SNARE complex
disassembly. However, because how NSF actually promotes disassembly is
unclear, it is possible that individual mutations in
-SNAP could change
SNARE complex disassembly differently from how they change binding.
Identifying such mutations in
-SNAP could yield insight into the role
of
-SNAP in SNARE complex disassembly.
We monitored SNARE complex disassembly by measuring FRET between CFP and
YFP tags that were attached to the C termini of the SNAREs syntaxin 1a and
synaptobrevin II in place of their transmembrane domains.2 FRET
between the CFP and YFP moieties is lost when SNAREs in the SNARE complex are
separated by NSF and -SNAP. To determine how mutations in
-SNAP
affected its ability to promote disassembly, we measured the initial velocity
of the reaction using an
-SNAP concentration determined to be
approximately the EC50 for disassembly by wild-type
-SNAP
(Table I). In most cases,
mutations affected disassembly similarly to how they affected binding, thereby
confirming the results of the binding assays
(Fig. 6). On the front face of
-SNAP, where all mutations that changed binding are located, mutations
that reduced binding also reduced disassembly. Among the mutations that
enhanced binding, most also enhanced disassembly, although less so than
binding. Interestingly, several mutations that did not increase binding did
enhance disassembly (E99A, E109A, and K140A;
Fig. 6). These residues are on
the back face of the
-SNAP twisted sheet and may contribute to the
SNAP-NSF interface.
|
|
![]() |
DISCUSSION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Mutating basic residues on the front face of -SNAP reduced SNARE
complex binding by up to approximately 20-fold
(Fig. 5). The effects of these
mutations were specific since similar mutations on the back face of
-SNAP did not affect SNARE complex binding. Similar effects of alanine
mutations have been observed at other protein-protein interfaces where the
interactions are primarily electrostatic, including the interface between
tubulin and kinesin (33) and
that between barnase and barstar
(34,
35). Interestingly, in
vitro phosphorylation of
-SNAP by protein kinase A reduces SNARE
complex binding by 10-fold
(36), demonstrating in another
way that positive charge on the surface of
-SNAP is important for its
binding to SNARE complex.
The EC50 of wild-type -SNAP binding to SNARE complex
(
5 µM, Fig.
1) is consistent with data in previous studies that examine
binding between
-SNAP and SNARE complexes using affinity matrix-based
binding assays (22,
36,
37). This EC50 is
nevertheless surprisingly low given that the EC50 of
-SNAP
for mediating SNARE complex disassembly is
0.2 µM
(Table I). 0.2 µM
-SNAP is closer to what is needed for functional reconstitution of NSF-
and SNAP-dependent processes in vitro
(24,
38,
39). That
co-precipitation-based assays inherently underestimate affinity may in part
explain this discrepancy. In addition, NSF may enhance SNAP-SNARE complex
binding (4).
The fact that binding to the synaptic SNARE complex was readily enhanced by
mutagenesis (Fig. 5) is
probably not surprising given the diversity among SNARE complexes throughout
the cell to which -SNAP must bind; optimized binding to one SNARE
complex could result in impaired binding to another. Because dissociation of
-SNAP from the SNARE complex is a step in disassembly, one might
imagine that enhancing binding would eventually impair disassembly. In fact,
excess
-SNAP (38) and
Sec17p (40) have been found to
impair SNARE complex-dependent processes in vitro. Although they were
not inhibitory, most mutations that enhanced binding only slightly enhanced
disassembly, and two did not enhance disassembly at all
(Fig. 6).
A group of mutations that did not enhance SNARE complex binding but did
enhance its disassembly included residues on the back face of -SNAP
(E99A, E109A, and K140A). These residues are likely to remain accessible to
NSF when
-SNAP is bound to the SNARE complex and may, therefore, play a
role in coupling conformational changes in NSF to SNARE complex
disassembly.
SNAP-SNARE Complex Binding ModelTo develop a model for how
-SNAP binds SNARE complexes we examined shape and charge
complementarity between the
-SNAP surface shown in this study to be
critical for binding and the surface of the SNARE complex. Six of the 10
residues whose mutation reduced SNARE complex binding (Lys-56, Lys-93, Lys-94,
Lys-122, Lys-163, and Lys-167; Fig.
5) are basic and form a diagonal band across the front face of the
-SNAP twisted sheet domain (Fig.
7). Diagonal bands of acidic residues on the SNARE complex surface
define three possible binding sites; similar groups of residues are also
present in other well characterized SNARE complexes
(17,
20,
41). Pairing basic and acidic
residues allowed us to manually align three
-SNAP twisted sheet domains
with a single SNARE complex (Fig.
7). In this arrangement, shape complementarity between
-SNAP and the SNARE complex is maximized and is substantially greater
than when SNAPs are placed directly parallel to individual SNARE helices
within the complex (not shown). Each
-SNAP interacts with three SNAREs.
This may explain why
-SNAP binding to individual SNAREs is weaker than
to the SNARE complex (21,
42) and why
-SNAP
dissociates after SNARE complex disassembly.
Each putative -SNAP binding site contains 4 or 5 acidic residues
close enough to
-SNAP residues Lys-56, Lys-93, Lys-94, Lys-163, and/or
Lys-167 to form salt bridges (Fig.
7). Mutation of any one of these
-SNAP residues
significantly reduced SNARE complex binding
(Fig. 5). Whether
-SNAP
mutations directly impair binding to one, two, or all three of the sites or
whether the mutations affect one critical site and thereby disrupt cooperative
interactions with other sites cannot be distinguished in our binding assay.
-SNAP binding to the site designated Site 1 positions it over the
groove between syntaxin and synaptobrevin in which complexin binds
(43,
44). This would be expected to
interfere with complexin binding and is, therefore, consistent with the
reported competition between
-SNAP and complexin for binding to the
SNARE complex (45). The
largest gap between
-SNAP molecules is between sites 3 and 1. The
interhelical linker that connects the two helices of SNAP-25 could fit between
these two
-SNAPs.
The model shown in Fig. 7
accounts for most of the experimental data described in this report. There
are, however, a few observations it does not presently explain that raise
questions for further study. Four basic residues on the twisted sheet of
-SNAP (Arg-47, Lys-53, Arg-116, and Lys-122) appear to be important for
SNARE complex binding (Fig. 5)
but are not predicted to directly interact with acidic residues on the SNARE
complex. These residues may play other roles; for example, Lys-53 is likely to
be involved in an interhelical salt bridge within
-SNAP, and Arg-47 may
counteract the charge of a nearby acidic residue. Second, to dock the
-SNAP twisted sheet domains closely against the SNARE complex, the
N-terminal globular bundle domains have been left out of the model shown in
Fig. 7. If kept in the model
these domains would collide with the SNARE complex. The Sec17p crystal
structure is of isolated protein
(25) and does not necessarily
reflect the conformation of SNARE complex-bound
-SNAP. The globular
bundle domain may, therefore, move in order for the twisted sheet domain to
effectively bind SNARE complex. Experiments to examine this possibility are in
progress.
Implications for SNARE Complex DisassemblyOur model of the
SNAP-SNARE complex interaction is compatible with a variety of mechanisms for
SNARE complex disassembly. Importantly, it implies that -SNAP molecules
are the primary connection between NSF and the SNARE complex. The simultaneous
binding of three
-SNAP molecules to the SNARE complex suggests that NSF
may use its nucleotide-driven conformational changes to promote disassembly by
affecting some property of all three SNAP-SNARE interfaces. An attractive
target for NSF action is found at the level of the SNARE complex ionic layer,
a conserved point of interaction among the four SNARE helices
(19). This layer is probably
the least stable region of the SNARE complex
(18,
44) and has been proposed to
be essential for efficient SNARE complex disassembly
(47). A number of
-SNAP
mutations that affected binding and disassembly lie near this layer. These
mutations may alter critical connections between the region of the SNARE
complex surrounding the ionic layer,
-SNAP, and NSF, which in turn
impair the ability of NSF to effect complex disassembly. Further studies to
explore these possibilities are in progress.
![]() |
FOOTNOTES |
---|
A W. M. Keck Foundation Distinguished Young Scholar, a McKnight Endowment Fund
for Neuroscience Scholar, a Searle Scholar, and an Alfred P. Sloan Foundation
Research Fellow. To whom correspondence should be addressed: Dept. of Cell
Biology and Physiology, Washington University School of Medicine, 660 S.
Euclid Ave, Box 8228, St. Louis, MO 63110. Tel.: 314-747-4233; Fax:
314-362-7463; E-mail:
phanson{at}cellbiology.wustl.edu.
1 The abbreviations used are: NSF, N-ethylmaleimide-sensitive
factor; SNAP, soluble NSF attachment protein; SNAP-25, synaptosome-associated
protein of 25 kDa; SNARE, SNAP receptor; YFP, yellow fluorescent protein; CFP,
cyan fluorescent protein; EC50, 50% effective concentration;
Tricine, N-[2-hydroxy-1,1-bis(hydroxymethyl)ethyl]glycine; PDB,
Protein Data Bank; FRET, fluorescence resonance energy transfer.
2 J. M. Lauer and P. I. Hanson, manuscript in preparation.
![]() |
ACKNOWLEDGMENTS |
---|
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|