From the Department of Biochemistry, Biophysics, and Molecular Biology, Iowa State University, Ames, Iowa 50011
Received for publication, October 30, 2002, and in revised form, January 12, 2003
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
In the neuron, soluble
N-ethylmaleimide-sensitive factor attachment protein
receptor (SNARE) proteins assemble into an Neurotransmitter release at synapses requires the fusion of
neurotransmitter-containing vesicles to the presynaptic plasma membrane. Membrane fusion is, however, an exceedingly difficult process
to go through without the assistant of specific proteins, because of
the protective nature of the biological membranes. In the neuron,
soluble N-ethylmaleimide-sensitive factor attachment protein
receptor (SNARE)1 proteins
play an essential role in promoting membrane fusion (1). It is proposed
that assembly of the SNARE complex initially bridges two membranes,
induces lipid mixing, and leads to the hemifusion state and the fusion
pore, of which the detailed mechanism is largely unknown (2-5).
Progress has been made in understanding the biophysical principles of
SNARE assembly. SNARE assembly starts with the interaction of
vesicle-associated membrane protein 2 (VAMP2 or synaptobrevin) with
target plasma membrane SNAREs Syntaxin 1A and SNAP-25. Interactions between SNARE proteins are mediated by "SNARE motifs" that are essentially coiled coil sequences and are present in all SNARE proteins
(3). For the SNARE complex, one SNARE motif each from Syntaxin 1A and
VAMP2 and two from SNAP-25 assemble into a 110-Å-long four-stranded
coiled coil (6, 7). It is worthwhile to note that target plasma
membrane SNAREs Syntaxin 1A and SNAP-25 also spontaneously assemble
into a similar but less stable four-stranded coiled coil (8-10).
How does coiled coil formation lead to membrane fusion? There are two
features of the SNARE coiled coil that might be important. First, the
helices are all aligned parallel, suggesting the co-location of two
membrane attachment points, which sets up a favorable geometry for
membrane fusion (6, 7, 11-13). Second, the coiled coil is highly
stable (14, 15). Therefore, coiled coil formation might have the
capacity to overcome the repulsive force between two apposing
membranes. Although this mechanistic model appears to be structurally
and energetically attractive, there are caveats that require careful
consideration. For example, if the SNARE core were tethered with
flexible linkers to membrane domains, coiled coil formation might not
be able to bring about membrane apposition no matter how strong the
pulling force it generates because the energy would be dissipated.
To validate this model, a direct coupling between the coiled coil and
membranes appears to be necessary. Previously, Brunger and co-workers
proposed a hypothetical model for the coiled coil-to-membrane coupling
(7). In this model, the coiled coil is linked to transmembrane domains
(TMD) as continuous helices. This model arbitrarily assumes some
bending flexibility of helices in short amino acid stretches at the
membrane-proximal region. Furthermore, helix-disrupting mutations or
amino acid insertions in the linker region have little or only moderate
effect on the SNARE fusion activity, inconsistent with this model (16,
17). How then is the coiled coil energetically coupled to membranes?
The answer to this fundamental question hinges on structural
information of the connection of the coiled coils to the membranes.
Recent EPR investigations of intact SNAREs using site-directed spin
labeling EPR have yielded new results that not only confirm the
existence of coupling between the coiled coil and the membrane but also
suggest a tentative mechanism of the SNARE core-membrane coupling. EPR
analysis indicated that the linker region of Syntaxin 1A, enriched with
basic amino acid residues, is unstructured but laterally inserted into
the membrane, tightly coupling the coiled coil to the membrane (18,
19). Importantly, clusters of basic amino acid residues are found in
the linker regions of all transmembrane SNAREs (20), raising the
possibility that SNARE linker regions generally insert into the
membrane. Further, this tentative model offers a plausible explanation
as to why this region is tolerant to helix-disrupting mutations.
However, structure and membrane topology of the VAMP2 linker region is
not experimentally confirmed yet.
In this work, we report the EPR and fluorescence investigations of the
membrane topology of the recombinant SNARE complex. Fluorescence
quenching analysis revealed that the native Trp residues at positions
89 and 90 in VAMP2 are inserted into the acyl chain region of the
bilayer. Further, the EPR results reveal that the core domain maintains
the coiled coil structure up to residue 92, suggesting that the SNARE
coiled coil is partially inserted into the head group region of the
bilayer. The EPR data also suggest that the coiled coil penetrates into
the membrane with an oblique angle. Taken together, the new results
further establish the concept of the tight SNARE core-membrane
coupling, providing structural basis for the force transmission from
the core region to the membrane during SNARE assembly.
Materials--
1-palmitoyl-2-oleoyl phosphatidylcholine (POPC)
and 1,2-dioleoyl phosphatidylserine (DOPS),
1-palmitoyl-2-stearoyl-(6,7)-dibromo-sn-glycero-3-phosphocholine (6,7-Br2-PC), and
1-palmitoyl-2-stearoyl-(11,12)-dibromo-sn-glycero-3-phosphocholine (11,12-Br2-PC) were purchased from Avanti Polar Lipids
(Birmingham, AL). (1-Oxyl-2,2,5,5-tetramethylpyrrolinyl-3-methyl)
methanethiosulfonate spin label (MTSSL) was obtained from Toronto
Research Chemicals (North York, Canada). The paramagnetic reagent,
nickel (II)-ethylenediamine-N,N'-diacetic acid
(NiEDDA) was synthesized following the procedure described elsewhere
(21). Pfu Turbo DNA polymerase and Escherichia
coli BL21-CodonPlus RIL were purchased from Stratagene (La Jolla,
CA). n-Octylglucoside (OG) was from Roche Molecular
Biochemicals. Bio-beads SM2 was obtained from Bio-Rad. Oligonucleotides
for site-directed mutagenesis were obtained from Qiagen Operon
Technologies (Alameda, CA). Ultrafree Centrifugal Devices Biomax-5K for
proteins was obtained from Millipore (Bedford, MA).
Nickel-nitrilotriacetic acid-agarose was purchased from Qiagen.
Glutathione-agarose, human thrombin, 4-(2-aminoethyl)benzenesulfonyl
fluoride, leupeptin, L-methionine, n-lauroyl
sarcosine, Triton X-100, Tween 20, glycerol, L-methionine,
isopropyl- Plasmid Constructs and Site-directed
Mutagenesis--
Full-length VAMP2 (amino acids 1-116), the soluble
SNARE motif of Syntaxin 1A (or Syntaxin H3) (amino acids 191-266), and the C-terminal SNARE motif of SNAP-25 (SNAP-25(C)) (amino acids 125-206) are inserted in the pGEX-KG vector as glutathione
S-transferase fusion proteins (22). On the other hand, the
N-terminal SNARE motif of SNAP-25 (SNAP-25(N)) (amino acids 1-82) is
in the pQE-30 vector (Qiagen) as a N-terminal His6-tagged
protein. To introduce a unique cysteine site for the specific nitroxide
attachment, native cysteine 103 of VAMP2 was changed to alanine. All of
the mutants were generated by QuikChange site-directed mutagenesis (Stratagene) and confirmed by DNA sequencing (Iowa State University DNA
Sequencing Facility).
Protein Expression, Purification, and Spin
Labeling--
Recombinant glutathione S-transferase fusion
proteins were expressed in E. coli BL21-CodonPlus RIL and
purified using glutathione-agarose chromatography. Briefly, the cells
were grown at 37 °C in LB medium with glucose (2 g/liter),
ampicillin (100 µg/ml), and chloramphenicol (50 µg/ml) until the
A600 reached 0.6-0.8. After the addition of isopropyl-
To purify the protein, the frozen cell pellet was resuspended in
PBST-Met buffer (phosphate-buffered saline, pH 7.4, with Tween 20 (0.05%) (percentages are v/v unless otherwise mentioned), 10 mM L-methionine, 2 mM
4-(2-aminoethyl)benzenesulfonyl fluoride, 1 µM leupeptin,
and 2 mM dithiothreitol. The cells were broken by
sonication on the ice bath. For full-length VAMP2, Triton X-100 (0.5%
(v/v) and n-lauroyl sarcosine (0.5%) were added to the
solution before sonication. After nutation for 30 min, the cell lysate was centrifuged at 15,000 × g for 15 min at 4 °C.
The supernatant was then mixed with glutathione-agarose beads in
PBST-Met buffer, and the mixture was left at 4 °C for 40 min. The
protein bound-beads were washed with an excess volume of PBST-Met
buffer for SNAP-25(C) and Syntaxin H3 or with PBST-Met-Triton buffer
(PBST-Met with 0.5% Triton X-100) for full-length VAMP2.
The cysteine mutants were spin-labeled, while the protein was bound to
the beads. After washing the beads with PBST-Met-Triton buffer without
dithiothreitol, a 10-fold molar excess of MTSSL was added. The sample
was initially reacted with MTSSL at room temperature for 1 h and
left to stand at 4 °C overnight. Free MTSSL was removed by washing
with PBST-Met-Triton buffer, and the spin-labeled protein was cleaved
off from the resin with thrombin in the cleavage buffer (50 mM Tris-HCl, pH 8.0, 150 mM NaCl, 2.5 mM CaCl2, 0.1% Triton X-100).
The His6-tagged protein SNAP-25(N) was expressed in
E. coli M15[pREP4] (Qiagen) and purified using
nickel-nitrilotriacetic acid-agarose affinity chromatography. Briefly,
the cells were grown at 37 °C in LB medium with glucose (2 g/liter),
ampicillin (100 µg/ml), and kanamycin (25 µg/ml) until the
A600 reached 0.6-0.8. After the addition of
isopropyl
For protein purification, the frozen cell pellet was resuspended in
buffer A (50 mM NaH2PO4, pH 8.0, 300 mM NaCl, 20 mM imidazole, 2 mM
4-(2-aminoethyl)benzenesulfonyl fluoride, 1 µM
leupeptin). After sonication on ice, the cell lysate was centrifuged at
15,000 × g for 15 min at 4 °C. The supernatant was
mixed with nickel-nitrilotriacetic acid-agarose beads in buffer A. The
mixture was left equilibrated at 4 °C for 40 min. After
equilibration the beads were washed with an excess volume of buffer A.
The protein concentration was estimated by Bio-Rad protein assay kit
using bovine serum albumin as a standard. The spin labeling efficiency
was estimated by comparing the double integration with the standard
Tempo sample at 100 µM. Spin labeling was nearly quantitative for all VAMP2 mutants.
Preparation of the Recombinant SNARE Complex--
Two separate
SNARE motifs of SNAP-25 lacking the long interhelical loop were used to
avoid the formation of domain-swapped complex (23). While
His6-tagged SNAP-25(N) was bound to nickel-nitrilotriacetic acid-agarose resin, excess amounts of purified Syntaxin 1A H3, SNAP-25(C), and full-length VAMP2 (molar ratio of 1:1:2) were added to
the solution. The solution contained 0.2% Triton X-100. The mixture
was left to stand at 4 °C for 60 h. After extensive washing
with buffer A containing 0.6% (w/v) OG, the recombinant SNARE complex
was eluted with buffer A with 250 mM imidazole and 0.6%
(w/v) OG. The complex was concentrated using a Centricon (5 K
cut-off). During the concentration process buffer was changed to buffer
B (25 mM HEPES, pH 7.7, 100 mM KCl, 10%
glycerol) with 0.6% (w/v) OG. The final protein concentration was in
the range of 80-100 µM.
Membrane Reconstitution of Recombinant SNARE Complex--
Large
unilamella vesicles with a 100-nm diameter (100 mM total
lipids) were prepared in buffer B without OG using an extruder. Vesicles of POPC containing 15 mol % of DOPS were first mixed with two
volumes of the SNARE complex. OG was added to the mixture to a final
concentration of 0.6%. After dilution with an equal volume of buffer
B, the samples were dialyzed against the buffer B containing Bio-beads
SM2 adsorbent at 4 °C for 40 h. During the dialysis, the buffer
was changed three times. The samples were centrifuged at 100,000 × g for 5 min to remove both the protein precipitates and
the fraction of large vesicles.
EPR Data Collection and Accessibility Measurements--
EPR
spectra were obtained using a Bruker ESP 300 spectrometer (Bruker,
Germany) equipped with a low noise microwave amplifier (Miteq,
Hauppauge, NY) and a loop-gap resonator (Medical Advances, Milwaukee, WI). The modulation amplitude was set at no greater than
one-fourth of the line width. The spectra were collected at room
temperature in first derivative mode. The gas exchange to the protein
sample was achieved with the TPX EPR tube for the loop-gap resonator.
For individual mutants, the power saturation curves were obtained from
the peak-to-peak amplitude of the central line (M = 0)
of the first derivative EPR spectrum as a function of incident
microwave power in the range 0.1-40 mW. Three power saturation curves
were obtained after equilibration: (i) with N2, (ii) with
air (O2), and (iii) with N2 in the presence of
200 mM NiEDDA. From saturation curves, the microwave power
P1/2, where the first derivative amplitude is
reduced to one-half of its unsaturated value, was calculated. The
quantity Fluorescence Quenching Experiment--
For fluorescence
measurements, membrane samples were prepared from the recombinant SNARE
complex with the wild-type sequences. Total lipid concentration was
~2.5 mM, whereas the concentration of the SNARE complex
was 5 µM. For acrylamide quenching, appropriate amounts
of the acrylamide stock solution (2 M) were added to the membrane sample to make the final concentration in the range of 0-160
mM, whereas lipids and protein concentrations remain
constant among samples. The fluorescence measurements were carried out with PerkinElmer fluorescence spectrophotometer. The samples were excited at 285 nm, and the emission spectra were collected in the range
of 300-400 nm. The total fluorescence intensity F was obtained by integrating the intensity in this spectral range. The
degree of quenching was analyzed according to the following Stern-Volmer equation.
To measure the membrane immersion depth of Trp residues,
membrane samples containing two types of lipid quenchers were prepared. Lipid quencher, 6,7-Br2-PC or 11,12-Br2-PC, was
added in replacement of part of POPC while maintaining the DOPC mole
fraction at 15%. The reconstitution of SNARE complex was carried out
by the same procedure described above. The degree of quenching was
determined as a function of the mole fraction of added brominated PC.
The averaged immersion depth of two Trp residues was calculated
according to the parallax analysis (25, 26). The distance of the Trp
residue from the bilayer center ZCF is given by
the following.
Fluorescence Quenching Experiments--
For fluorescence
measurement the recombinant complex was assembled from full-length
VAMP2, soluble Syntaxin 1A, and two separate SNARE motifs from SNAP-25
(Fig. 1). The TMD of Syntaxin 1A was not
included this time to avoid the coexistence of two TMDs in one membrane
and to best mimic the "hypothetical" trans-SNARE complex
in which two TMDs are separately anchored to two apposing membranes. In
the recombinant SNARE complex, there are total two native Trp residues.
Both of them reside at the membrane-proximal region of VAMP2
(Trp89 and Trp90) and belong to the coiled coil
in the core structure (Fig. 1). This is ideal for the investigation of
the possible coiled coil-membrane coupling using fluorescence.
First, to examine whether these residues are exposed to the solvent or
not, we monitored Trp fluorescence in the presence of an added quencher
acrylamide that is hydrophilic and partitions heavily into the solution
phase. For the detergent-solubilized SNARE complex, the
fluorescence intensity (F) decreases sharply as the
acrylamide concentration increases, suggesting that Trp residues are
solvent-exposed (Fig. 2a,
open circles). In contrast, for the membrane-reconstituted
complex, F decreases little in the presence of added
acrylamide (Fig. 2a, closed circles). This result
strongly implies that Trp residues in the SNARE complex are sequestered
from the water phase, suggesting the possibility of insertion into the
membrane.
Next, the insertion of Trp residues into the membrane is probed
utilizing brominated lipids in which bromines are attached to the acyl
chain of the lipid. In the presence of a brominated lipid, Trp
fluorescence is effectively quenched only when Trp is in contact with
the acyl chain region of the membrane. Otherwise, we would expect a
negligible effect. As the mole fraction of the brominated lipid
increases, a significant decrease in F was observed (Fig.
2b), suggesting that Trp residues are inserted into the membrane. The immersion depth of Trp residues was calculated based on
quenching efficiencies by the shallow lipid quencher
6,7-Br2-PC (the lipid with bromines at the sixth and
seventh carbon positions) and the deep lipid quencher
11,12-Br2-PC. Trp residues are ~8.8 Å below the
phosphate groups of lipids. Because there are two Trp residues, the
immersion depth determined here must be an average depth of two
residues. In conclusion, Trp89 and Trp90
that belong to the coiled coil are inserted in the membrane in the
SNARE complex.
Site-directed Spin Labeling EPR--
To investigate the membrane
topology of the SNARE complex further using site-directed spin labeling
EPR, residues of full-length VAMP2 near the membrane-water interface
were replaced with cysteines, to which a nitroxide spin label was
attached. We prepared 13 spin-labeled mutants (K83C-M95C) to
explore the interfacial region with EPR (Fig. 1). For EPR measurements,
the recombinant complex was assembled from spin-labeled VAMP2, soluble
Syntaxin 1A, and two separate SNARE motifs from SNAP-25. All
spin-labeled recombinant complexes were capable of forming the
SDS-resistant complex, which is one characteristic feature of
the core complex, as confirmed with SDS-PAGE (data not shown).
After reconstitution of the SNARE complex into POPC vesicles containing
15 mol % of DOPS, the EPR spectra were collected for spin-labeled
mutants at room temperature. The EPR spectrum is sensitive to the
tumbling rate of the nitroxide. EPR spectra shown in Fig.
3 are all relatively broad, indicating
slow motion. Slow motional spectra represent motionally restricted
nitroxides. There are three structural factors that might have
contributed to the immobilization of the nitroxide (28-30): (i) The
motional restriction of the peptide backbone, because of the
Additionally, to examine the possibility of intermolecular interactions
between SNARE complexes, we measured low temperature (130 K) EPR
spectra in which the spectral broadening caused by the spin-spin
interaction is readily identified (31). Comparison of the low
temperature EPR spectra with the standard confirmed that SNARE
complexes are separated from each other beyond the detectable distance
range (less than 25 Å) (data not shown), eliminating the possibility
of self-aggregation of the SNARE complex.
It is interesting to note that EPR spectra for positions 93-95 are
less broad than others. In fact, EPR spectra for these three positions
closely resemble those observed for the nitroxide attached to the
membrane-inserted linker region of Syntaxin 1A (18, 19).
EPR Accessibility Measurements--
The EPR line shape is a useful
parameter for the tumbling rate of the nitroxide that, in many cases,
provides a qualitative assessment of the local environment surrounding
the nitroxide (29). However, the line shape alone is often not
sufficient to yield information pinpointing the secondary and the
tertiary structures. Furthermore, the partial insertion of the protein into the bilayer makes the matters complicated for the SNARE complex. Here, we utilized the EPR saturation method to assess the local structure and the membrane topology of the linker region. For the
nitroxide, the EPR saturation method measures the accessibility to a
water-soluble paramagnetic reagent such as NiEDDA
(WNiEDDA) to estimate the solvent exposure of the
spin-labeled site, or the accessibility to a nonpolar paramagnetic
reagent such as molecular oxygen (WO2) to
probe, for example, the insertion into the membrane (32-34).
In Fig. 4 WNiEDDA and
WO2 for the SNARE complex are plotted against
the residue number, respectively. We observe an overall decrease of
WNiEDDA, whereas we detect an overall increase of WO2. Interestingly, however, there are
quite significant increases and decreases for WNiEDDA along
the sequence, which might imply a secondary structure such as
Quantitatively, the ratio of WNiEDDA to
WO2 has been shown to be a useful
parameter to characterize the secondary structure. For a
For lipid-exposed nitroxides, the ratio of WNiEDDA to
WO2 has been shown to hold a quantitative
relationship to the membrane immersion depth. However, EPR spectra for
positions 84, 87, 90, and 92 contain highly immobilized spectral
components with large outer hyperfine splitting. Such high
immobilization might be due to some tertiary interactions of the
nitroxide in the complex. This certainly limits the application of the
membrane depth analysis in the region of residues 83-92. To our
advantage, it is already shown that Trp89 and
Trp90 are inserted into the membrane 8.8 Å deep from the
lipid phosphate group.
As mentioned before EPR spectra for positions 93-95 exhibit
intermediate motional rates, characteristic of lipid-exposed
nitroxides. It is also highly likely that these three residues are
disordered when judged from the disagreement with the continued
Combining the EPR data and the fluorescence data, we conclude that the
C-terminal part of the coiled coil is inserted into the membrane at an
oblique angle. Further, we propose that residues 93-95 might be
unstructured although inserted into the membrane.
The location of VAMP2 residues Trp89 and
Trp90 in the SNARE complex inside the membrane is not
surprising taking into account the fact that Trp residues are commonly
found in membrane proteins near the membrane-water interface (37). It
is further supported by the fact that the membrane-proximal region of
VAMP2 has phospholipid binding affinity (38). The high affinity of Trp
to the membrane (approximately The EPR results suggest that a small C-terminal portion of the SNARE
coiled coil penetrates into the membrane with an oblique angle. Such a
tilted topology provides a favorable geometry for six nearby basic
residues (Lys83, Lys85, Arg86,
Lys87, Lys91, and Lys94) to be able
to effectively interact with the membrane surface charge
electrostatically. According to the previous estimation, each basic
residue contributes approximately We expect that the TMD of VAMP2 is nearly perpendicular to the membrane
surface (41). However, the EPR results suggest that the coiled coil
domain of the complex is tilted significantly with respect to
the membrane, requiring bending or disorder at the linker region. The
EPR results suggest that positions 93-95 are strong candidates for the
disordered connection between the TMD and the core. The saturation EPR
analysis suggested that those three positions are immersed in the acyl
chain region. In parallel, previous EPR studies demonstrated the
adhesion of the polybasic linker region (Arg262,
Arg263, Lys264, and Lys265) of
Syntaxin 1A onto the membrane (18, 19). It was found that this region
is laterally inserted into the membrane, similarly pulling the SNARE
core toward the membrane.
However, one must be very careful in interpreting the EPR immersion
depth data. Because EPR measures the position of the substituted nitroxide side chain, the immersion depths do not necessarily report
the actual location of the native amino acids when the native residues
are charged amino acids. For instance, EPR reported the depth of 12-13
Å for K94C (Fig. 5). However, the actual location of the lysine side
chain may be much shallower. The positive charge on Lys94
could snorkel out to seek the negative changes on phosphate, opposite
to what is expected for the relatively hydrophobic nitroxide side chain.
A model for the trans-SNARE complex that sums up
experimental results is depicted in Fig.
6. In this model, the tight coupling between the SNARE core and two apposing membranes is achieved by (i)
the electrostatic interactions between the basic residues and the
negatively charged lipids in the membrane and (ii) the insertion of
interfacial Trp residues into the membrane. We estimate that the total
free energy of the stabilization from these two sources amounts
approximately to -helical coiled coil that
bridges the synaptic vesicle to the plasma membrane and drives membrane
fusion, a required process for neurotransmitter release at the nerve
terminal. How does coiled coil formation drive membrane fusion? To
investigate the structural and energetic coupling between the coiled
coil and membrane, the recombinant SNARE complex in the phospholipid
bilayer was studied using fluorescence quenching and site-directed spin
labeling EPR. Fluorescence analysis revealed that two native Trp
residues at the membrane-proximal region of the coiled coil are
inserted into the membrane, tightly coupling the coiled coil to the
membrane. The EPR results indicate that the coiled coil penetrates into
the membrane with an oblique angle, providing a favorable geometry for
the basic residues to interact with negatively charged lipids. The
result supports the proposition that core complex formation directly
leads to the apposition of two membranes, which could facilitate lipid
mixing. Trp residues and basic residues are abundant at the
membrane-proximal region of transmembrane SNARE proteins, suggesting
the generality of the proposed mechanism for the SNARE complex-membrane coupling.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
-D-thiogalactopyranoside, dithiothreitol, ampicillin sodium salt, chloramphenicol, kanamycin, and the low molecular weight size marker for SDS-PAGE were all purchased from Sigma.
-D-thiogalactopyranoside (0.2 mM), the cells were grown further for 5 h more at
30 °C for SNAP-25(C), at 22 °C for full-length VAMP2 but at
16 °C for the Syntaxin 1A H3 domain.
-D-thiogalactopyranoside (0.2 mM), the cells were grown further for 5 h more at
30 °C.
P1/2 is the difference in
P1/2 values in the presence and absence of a
paramagnetic reagent. The
P1/2 value is
proportional to the diffusion co-efficient times the collision
frequency of the nitroxide to the freely diffusing reagents such as
oxygen and NiEDDA. Thus,
P1/2 is considered to be
equivalent to the accessibility W. The immersion depth is calculated based on the reference curves determined from a set
of lipid molecules spin-labeled at different acyl chain positions.
where Fo and F are the
fluorescence intensities in the absence and presence of acrylamide,
respectively, KSV is the Stern-Volmer constant
for collisional quenching, and [Q] is the concentration of the
quencher. The equation predicts a linear behavior of
Fo/F versus [Q] for a homogeneous
solution (24). After fluorescence measurements, the membrane samples
were treated with Triton X-100 (final concentration, 1%) to
subsequently measure the acrylamide quenching in a
detergent-solubilized state.
(Eq. 1)
where LC1 represents the distance from
the bilayer center to the shallow quencher (11 Å for
6,7-Br2-PC), C is the mole fraction of the
quencher divided by the area of the lipid molecule (70 Å2), F1 and
F2 are the relative fluorescence intensities of
the shallow (6,7-Br2-PC) and deep quenchers
(11,12-Br2-PC), respectively, and L is the
difference in the depth of the two quenchers (0.9 Å/CH2 or
CBr2 group). The thickness of the hydrophobic region was
approximated to be ~29 Å (27). For the calculation of immersion depth, the data collected for the 0.4 molar fraction quencher were used.
(Eq. 2)
RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
View larger version (9K):
[in a new window]
Fig. 1.
Structural elements of the recombinant SNARE
complex used in this study. Helical domains are shown as
bars. The soluble H3 domain of Syntaxin 1A (amino acids
191-266), the N-terminal SNARE motif of SNAP-25 (SNAP-25(N), amino
acids 1-82), and the C-terminal SNARE motif of SNAP-25 (SNAP-25(C),
amino acids 125-206) are used as target plasma membrane SNAREs,
whereas full-length VAMP2 (amino acids 1-116) including the TMD was
used as VAMP1 (or vesicle-associated SNARE). The N-terminal
His6 tag of SNAP-25(N) was not removed. The amino acids of
the membrane-proximal region of VAMP2 are shown. Two Trp residues are
indicated with bold letters.
View larger version (11K):
[in a new window]
Fig. 2.
Two native Trp residues of the SNARE complex
are inserted into the membrane. In this study the SNAP-25
interhelical loop and the Syntaxin N-terminal helical domain
were not included. Therefore, there are only two Trp residues
(Trp89 and Trp90 of VAMP2) left in the
recombinant complex. a, quenching of Trp fluorescence by
water-soluble acrylamide. The ratio of the fluorescence intensity in
the absence of acrylamide F0 to that in the
presence of acrylamide F,
F0/F is plotted as a function of
acrylamide concentration (closed circles). For the same
samples, the fluorescence measurements were repeated after disrupting
the membrane with Triton X-100 (v/v 1%). The ratios
F0/F for Triton X-100-solubilized
samples are also plotted for comparison (open circles).
b, quenching of Trp fluorescence by lipid quenchers
6,7-Br2-PC (filled circles) and
11,12-Br2-PC (open circles). The average
immersion depth of two Trp residues determined by the difference in
fluorescence quenching efficiency by the deeper and shallower quenchers
as described in experimental procedure is 8.8 Å from the lipid
phosphate group.
-helical secondary structure, could have reduced the tumbling rate
of the nitroxide. (ii) Tertiary interactions with other parts of the
protein would slow down the motion of the nitroxide significantly. On
the basis of the crystal structure (7), we expect that the
four-stranded coiled coil structure extends up to residue 92, which
gives rise to many potential tertiary contacts between helices. (iii)
From the fluorescence measurement it is clearly shown that
Trp89 and Trp90 are inserted into the acyl
chain region of the bilayer. Therefore, we expect that a significant
part of the region is immersed in the membrane, which exposes
nitroxides into the viscous membrane environment and the high density
head group region. It is likely that the combination of all three
factors contributed to the EPR spectral broadening.
View larger version (19K):
[in a new window]
Fig. 3.
First-derivative room temperature EPR spectra
of the spin-labeled SNARE complex. The SNARE complex was
reconstituted to the POPC vesicles containing 15 mol % of DOPS.
-helix. Further, the WO2 values show
variations that appear to be in opposite directions to those of
WNiEDDA, although the trend is much less clear. Such an
out-of-phase oscillatory behavior of WNiEDDA and
WO2 has been previously found for
-helical
peptides residing at membrane-water interface, which includes the
fusion peptide of influenza hemagglutinin and a synthetic amphiphilic
peptide (32, 35, 36).
View larger version (14K):
[in a new window]
Fig. 4.
Accessibility parameters WO2
(open circles) and WNiEDDA (filled
circles) are plotted as functions of the residue
number.
-helix we
expect a periodical behavior of this parameter along the sequence with
a periodicity of 3.5. In Fig. 5, the
value, which is defined as the logarithm of the ratio of
WO2 to WNiEDDA, is plotted as a
function of residue number. We observe a significant variation of the
value along the sequence. In particular, it appears that there is a
periodic oscillation of the
values in the region of residues
83-92. To better represent this oscillatory behavior, we fit the data
with a sine function of the 3.5 residue repeat, which represents the
-helical geometry. In this fit, we also take into account the
overall decreasing trend of the
values along the sequence as a
linear term added to the sine function. The EPR data are shown
overlapped with the fit in Fig. 5. Despite the qualitative nature of
the
value, the EPR results clearly fall into the pattern of the
-helical geometry, consistent with the structure inferred from the
crystal structure in which the last ordered residue of VAMP2 is 92. The fit also indicates that the helix is tilted with respect to the membrane surface, although the exact magnitude of the tilt could not be
estimated from the fit.
View larger version (12K):
[in a new window]
Fig. 5.
The value, defined as the logarithm of
the ratio of WNiEDDA to WO2, is plotted as
a function of the residue number. The solid line is the
fit with a modified sine function; a linear term is added to the sine
function to take into account the tilt. The EPR line shape analysis
allows the conversion of the
values to the membrane immersion
depths only for positions 93-95. These three positions are inserted
into the membrane as depicted to the right of the
dotted line.
-helical geometry. For these three positions, we compared the
values with those obtained from the spin-labeled lipids in the membrane of the same lipid composition containing the similar concentration of
the unlabeled SNARE complex. The immersion depth analysis revealed that
these three positions are all inserted into the membrane (Fig. 5,
panel to the right of the dashed
line).
DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
2.5 kcal/mol/residue) helps the
stabilization of the structure and topology of membrane proteins (39).
Trp residues are also found in multiple numbers in the linker region of
many transmembrane SNARE proteins (20), implying potential functional
roles of those Trp residues in stabilizing specific topological
structures of individual SNAREs and their complexes, which is perhaps
necessary to achieve membrane fusion.
1 kcal/mol to the free energy of
the membrane-peptide interactions (40). Therefore, we expect that the
total electrostatic contribution would be close to
6 kcal/mol.
Combining the free energy contributions from Trp residues and basic
residues, we speculate that the membrane-dipped SNARE complex is highly
stable with the free energy of as much as
11 kcal/mol.
16 kcal/mol.
View larger version (35K):
[in a new window]
Fig. 6.
Model for the trans-SNARE
complex. The highlight of this model is the insertion of the
C-terminal ends of coiled coil helices into two apposing membranes. The
insertion of interfacial Trp residues (yellow side chains)
and the electrostatic interactions between the basic residues
(green spheres) and the membrane surface charge contribute
to the stability. Coiled coil formation tightly pins two membranes to
allow the juxtaposition. SNARE proteins are color-coded:
red, Syntaxin 1A; blue, VAMP2; and
gray, SNAP-25.
The theoretical estimation of the free energy barrier for membrane
fusion is as high as 25 kcal/mol (42), a significant fraction of which
arises from the deformation of bilayers, a necessary step toward the
lipid mixing and the hemifusion intermediate. Evidence suggests that
coiled coil formation generates sufficient force to overcome this
energy barrier. The important question is, then; how is the force
delivered from the coiled coil to the membrane domains? To accomplish
this, the coiled coil must be structurally coupled to the membrane
domains. Otherwise, the force generated by the coiled coil formation
would be dissipated. The EPR analysis suggested that there is indeed
structural coupling between the coiled coil and the membrane.
Furthermore, we estimated that this coupling amounts to 16 kcal/mol.
However, it appears that the coupling force (
16 kcal/mol) is somewhat
smaller than the theoretically predicted membrane distortion force
during fusion (25 kcal/mol). Therefore, one SNARE complex might not
have the capacity to hold the membrane deformation force during fusion. This energetic insufficiency must be overcome by the coordinated effort
of more than one SNARE complex (23, 43-45). In fact, it has been
recently shown that three SNARE complexes cooperate for the successful
fusion (46).
Oblique dipping of the SNARE coiled coil to the membrane appears to have important implications regarding the trans-SNARE complex. In the trans state, the four-helix bundle, the diameter of which is as large as 20 Å (7), is sandwiched between two membranes. If the coiled coil were not inserted into the membrane, the juxtaposition of two membranes would be inherently inhibited by the existence of the coiled coil in the middle. In fact, it has been previously thought that the SNARE complex might hinder membrane apposition closer than 20 Å (47). At this distance, two membranes would remain well hydrated on the surface and would not proceed to fusion (48). The oblique insertion of the C-terminal end of the coiled coil would provide a solution to this geometric problem.
The functional significance of Trp and the basic residues discussed above is strongly supported by the recent experiment using PC12 cells. Using the tetanus toxin-resistant VAMP2 mutant, it has been shown that double mutation W89A/W90A reduced the secretion of human growth hormone significantly. Further, similar reduction of human growth hormone secretion has been observed for the double mutation K83A/K87V (49).
SNARE proteins are central to the membrane fusion machinery in the
neuron. However, membrane fusion is regulated by the Ca2+
influx. There is evidence that the vesicle protein synaptotagmin is a
Ca2+ sensor (50, 51). Although its function remains
elusive, recent studies have shown that synaptotagmin not only binds to
the membrane in a Ca2+-dependent manner but
also interacts with SNARE complexes. Coincidentally, loops 1 and 3 of
synaptotagmin inserts into the interfacial region of the bilayer,
similar to the linker regions of SNARE complex (52). We speculate that
the interaction between SNARE and synaptotagmin might occur in the
membrane environment, warranting further investigation.
![]() |
FOOTNOTES |
---|
* This work was supported by National Institute of Health Grant GM51290.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.
These authors contributed equally to this paper.
§ To whom correspondence should be addressed. Tel.: 515-294-2530; Fax: 515-294-0453; E-mail: colishin@iastate.edu.
Published, JBC Papers in Press, January 15, 2003, DOI 10.1074/jbc.M211123200
![]() |
ABBREVIATIONS |
---|
The abbreviations used are: SNARE, soluble N-ethylmaleimide-sensitive factor attachment protein receptor; 6, 7-Br2-PC, 1-palmitoyl-2-stearoyl-(6,7)-dibromo-sn-glycero-3-phosphocholine; 11, 12-Br2-PC, 1-palmitoyl-2-stearoyl-(11,12)-dibromo-sn-glycero-3-phosphocholine; DOPS, 1,2-dioleoyl phosphatidylserine; EPR, electron paramagnetic resonance; NiEDDA, nickel (II)-ethylenediamine-N,N'-diacetic acid; OG, n-octylglucoside; PBST-Met, phosphate-buffered saline pH 7.4, 0.05% (v/v) Tween 20, 10 mM L-methionine; POPC, 1-palmitoyl-2-oleoyl phosphatidylcholine; SNAP-25, 25-kDa, soluble N-ethylmaleimide-sensitive factor attachment protein; TMD, transmembrane domain(s); VAMP2, vesicle-associated membrane protein 2; MTSSL, (1-oxyl-2,2,5,5-tetramethylpyrrolinyl-3-methyl) methanethiosulfonate spin label.
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1. | Weber, T., Zemelman, B. V., McNew, J. A., Westermann, B., Gmachl, M., Parlati, F., Sollner, T. H., and Rothman, J. E. (1998) Cell 92, 759-772[Medline] [Order article via Infotrieve] |
2. | Rothman, J. E. (1994) Nature 372, 55-63[CrossRef][Medline] [Order article via Infotrieve] |
3. | Jahn, R., and Südhof, T. C. (1999) Annu. Rev. Biochem. 68, 863-911[CrossRef][Medline] [Order article via Infotrieve] |
4. | Lin, R. C., and Scheller, R. H. (2000) Annu. Rev. Cell Dev. Biol. 16, 19-49[CrossRef][Medline] [Order article via Infotrieve] |
5. | Brunger, A. T. (2001) Curr. Opin. Struct. Biol. 11, 163-173[CrossRef][Medline] [Order article via Infotrieve] |
6. | Poirier, M. A., Xiao, W., Macosko, J. C., Chan, C., Shin, Y.-K., and Bennett, M. K. (1998) Nat. Struct. Biol. 5, 765-769[CrossRef][Medline] [Order article via Infotrieve] |
7. | Sutton, R. B., Fasshauer, D., Jahn, R., and Brunger, A. T. (1998) Nature 395, 347-353[CrossRef][Medline] [Order article via Infotrieve] |
8. | Xiao, W., Poirier, M. A., Bennett, M. K., and Shin, Y.-K. (2001) Nat. Struct. Biol. 8, 308-311[CrossRef][Medline] [Order article via Infotrieve] |
9. |
Margittai, M.,
Fasshauer, D.,
Pabst, S.,
Jahn, R.,
and Langen, R.
(2001)
J. Biol. Chem.
276,
13169-13177 |
10. |
Zhang, F.,
Chen, Y.,
Kweon, D.-H.,
Kim, C. S.,
and Shin, Y.-K.
(2002)
J. Biol. Chem.
277,
24294-24298 |
11. | Hanson, P. I., Roth, R., Morisaki, H., Jahn, R., and Heuser, J. E. (1997) Cell 90, 523-535[Medline] [Order article via Infotrieve] |
12. | Lin, R. C., and Scheller, R. H. (1997) Neuron 19, 1087-1094[Medline] [Order article via Infotrieve] |
13. |
Katz, L.,
Hanson, P. I.,
Heuser, J. E.,
and Brennwald, P.
(1998)
EMBO J.
17,
6200-6209 |
14. |
Fasshauer, D.,
Bruns, D.,
Shen, B.,
Jahn, R.,
and Brunger, A. T.
(1997)
J. Biol. Chem.
272,
4582-4590 |
15. |
Fasshauer, D.,
Otto, H.,
Eliason, W. K.,
Jahn, R.,
and Brunger, A. T.
(1997)
J. Biol. Chem.
272,
28036-28041 |
16. | McNew, J. A., Weber, T., Engelman, D. M., Sollner, T. H., and Rothman, J. E. (1999) Mol. Cell. 4, 415-421[Medline] [Order article via Infotrieve] |
17. |
Wang, Y.,
Dulubova, I.,
Rizo, J.,
and Südhof, T. C.
(2001)
J. Biol. Chem.
276,
28598-28605 |
18. | Kweon, D.-H., Kim, C. S., and Shin, Y.-K. (2002) Biochemistry 41, 9264-9268[CrossRef][Medline] [Order article via Infotrieve] |
19. | Kim, C. S., Kweon, D.-H., and Shin, Y.-K. (2002) Biochemistry 41, 10928-10933[CrossRef][Medline] [Order article via Infotrieve] |
20. | Weimbs, T., Mostov, K., Low, S. H., and Hofmann, K. (1998) Trends Cell Biol. 8, 260-262[CrossRef][Medline] [Order article via Infotrieve] |
21. | Altenbach, C., Greenhalgh, D. A., Khorana, H. G., and Hubbell, W. L. (1994) Proc. Natl. Acad. Sci. U. S. A. 91, 1667-1671[Abstract] |
22. | Guan, K. L., and Dixon, J. E. (1991) Anal. Biochem. 192, 262-267[Medline] [Order article via Infotrieve] |
23. | Kweon, D.-H., Chen, Y., Zhang, F., Poirier, M., Kim, C. S., and Shin, Y.-K. (2002) Biochemistry 41, 5449-5452[CrossRef][Medline] [Order article via Infotrieve] |
24. | Birks, J. B. (1970) Photophysics of Aromatic Molecules , Wiley-Interscience, New York |
25. | Abrams, F. S., and London, E. (1992) Biochemistry 31, 5312-5322[Medline] [Order article via Infotrieve] |
26. | Chattopadhyay, A., and London, E. (1987) Biochemistry 26, 39-45[Medline] [Order article via Infotrieve] |
27. | McIntosh, T. J., and Holloway, P. W. (1987) Biochemistry 26, 1783-1788[Medline] [Order article via Infotrieve] |
28. | Hubbell, W. L., Gross, A., Langen, R., and Lietzow, M. A. (1998) Curr. Opin. Struct. Biol. 8, 649-656[CrossRef][Medline] [Order article via Infotrieve] |
29. | Mchaourab, H. S., Lietzow, M. A., Hideg, K., and Hubbell, W. L. (1996) Biochemistry 35, 7692-7704[CrossRef][Medline] [Order article via Infotrieve] |
30. | Columbus, L., and Hubbell, W. L. (2002) Trends Biochem. Sci. 27, 288-295[CrossRef][Medline] [Order article via Infotrieve] |
31. | Rabenstein, M. D., and Shin, Y.-K. (1995) Proc. Natl. Acad. Sci. U. S. A. 92, 8239-8243[Abstract] |
32. | Macosko, J. C., Kim, C.-H., and Shin, Y.-K. (1997) J. Mol. Biol. 267, 1139-1148[CrossRef][Medline] [Order article via Infotrieve] |
33. | Shin, Y.-K., Levinthal, C., Levinthal, F., and Hubbell, W. L. (1993) Science 259, 960-963[Medline] [Order article via Infotrieve] |
34. | Altenbach, C., Marti, T., Khorana, H. G., and Hubbell, W. L. (1990) Science 248, 1088-1092[Medline] [Order article via Infotrieve] |
35. | Han, X, Bushwell, J. H., Cafiso, D. S., and Tamm, L. K. (2001) Nat. Struct. Biol. 8, 715-720[CrossRef][Medline] [Order article via Infotrieve] |
36. | Russell, C. J., Thorgeirsson, T. E., and Shin, Y.-K. (1999) Biochemistry 38, 337-346[CrossRef][Medline] [Order article via Infotrieve] |
37. | Wimley, W. C., and White, S. H. (1996) Nat. Struct. Biol. 3, 842-848[Medline] [Order article via Infotrieve] |
38. |
Quetglas, S.,
Leveque, C.,
Miquelis, R.,
Sato, K.,
and Seagar, M.
(2000)
Proc. Natl. Acad. Sci. U. S. A.
97,
9695-9700 |
39. | Thorgeirsson, T. E., Russell, C. J., King, D. S., and Shin, Y.-K. (1996) Biochemistry 35, 1803-1809[CrossRef][Medline] [Order article via Infotrieve] |
40. | Kim, J., Mosior, M., Chung, L. A., Wu, H., and McLaughlin, S. (1991) Biophys. J. 60, 135-148[Abstract] |
41. | Fleming, K. G., and Engelman, D. M. (2001) Proteins 45, 313-317[CrossRef][Medline] [Order article via Infotrieve] |
42. |
Kuzmin, P. I.,
Zimmerberg, J.,
Chizmadzhev, Y. A.,
and Cohen, F. S.
(2001)
Proc. Natl. Acad. Sci. U. S. A.
98,
7235-7240 |
43. | Margittai, M., Otto, H., and Jahn, R. (1999) FEBS Lett. 446, 40-44[CrossRef][Medline] [Order article via Infotrieve] |
44. |
Laage, R.,
Rohde, J.,
Brosig, B.,
and Langosch, D.
(2000)
J. Biol. Chem.
275,
17481-17487 |
45. | Tokumaru, H., Umayahara, K., Pellegrini, L. L., Ishizuka, T., Saisu, H., Betz, H., Augustine, G. J., and Abe, T. (2001) Cell 104, 421-432[CrossRef][Medline] [Order article via Infotrieve] |
46. |
Hua, Y,
and Scheller, R. H.
(2001)
Proc. Natl. Acad. Sci. U. S. A.
98,
8065-8070 |
47. | Mayer, A. (2001) Trends Biochem. Sci. 26, 717-723[CrossRef][Medline] [Order article via Infotrieve] |
48. | Rand, R. P., and Parsegian, V. A. (1986) Annu. Rev. Phyiol. 48, 201-212[CrossRef] |
49. |
Quetglas, S.,
Iborra, C.,
Sasakawa, N.,
Haro, L.,
Kumakura, K.,
Sato, K.,
Leveque, C.,
and Seagar, M.
(2002)
EMBO J
21,
3970-3979 |
50. | Chapman, E. R. (2002) Nat. Mol. Cell. Biol. 3, 1-11[CrossRef] |
51. | Südhof, T. C. (2000) J. Biol. Chem. 277, 7629-7632 |
52. |
Bai, J.,
Earles, C. A.,
Lewis, J. L.,
and Chapman, E. R.
(2000)
J. Biol. Chem.
275,
25427-25435 |