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The Journal of Neuroscience, March 1, 2001, 21(5):1421-1433
synaptotagmin Mutants Reveal Essential Functions
for the C2B Domain in Ca2+-Triggered Fusion and
Recycling of Synaptic Vesicles In Vivo
J. Troy
Littleton4,
Jihong
Bai1,
Bimal
Vyas1,
Radhika
Desai1,
Andrew E.
Baltus4,
Martin B.
Garment2,
Stanley D.
Carlson2,
Barry
Ganetzky3, and
Edwin R.
Chapman1
Departments of 1 Physiology and
2 Entomology, and 3 Laboratory of Genetics,
University of Wisconsin, Madison, Wisconsin 53706, and
4 Center for Learning and Memory and Department of Biology,
Massachusetts Institute of Technology, Cambridge, Massachusetts 02139
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ABSTRACT |
Synaptotagmin has been proposed to function as a
Ca2+ sensor that regulates synaptic vesicle
exocytosis, whereas the soluble N-ethylmaleimide-sensitive factor attachment protein
receptor (SNARE) complex is thought to form the core of a
conserved membrane fusion machine. Little is known concerning the
functional relationships between synaptotagmin and SNAREs. Here we
report that synaptotagmin can facilitate SNARE complex formation
in vitro and that synaptotagmin mutations
disrupt SNARE complex formation in vivo. Synaptotagmin oligomers efficiently bind SNARE complexes, whereas
Ca2+ acting via synaptotagmin triggers cross-linking
of SNARE complexes into dimers. Mutations in Drosophila
that delete the C2B domain of synaptotagmin disrupt clathrin
AP-2 binding and endocytosis. In contrast, a mutation that blocks
Ca2+-triggered conformational changes in C2B and
diminishes Ca2+-triggered synaptotagmin
oligomerization results in a postdocking defect in neurotransmitter
release and a decrease in SNARE assembly in vivo. These
data suggest that Ca2+-driven oligomerization via
the C2B domain of synaptotagmin may trigger synaptic vesicle fusion via
the assembly and clustering of SNARE complexes.
Key words:
exocytosis; synaptotagmin; SNARE; Ca2+; synaptic vesicle; membrane fusion; C2 domain; Drosophila
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INTRODUCTION |
Neuronal exocytosis is precisely
controlled by Ca2+ ions (Katz, 1969 ) and
is extremely rapid (Llinas et al., 1981). The speed of exocytosis
dictates that a small number of molecular rearrangements couple
Ca2+ influx to the catalysis of bilayer
fusion. Recent studies have established that cycles of
Ca2+-triggered exocytosis require the
assembly and disassembly of the soluble
N-ethylmaleimide-sensitive factor attachment protein receptor (SNARE) complex (Söllner et al., 1993; Littleton
et al., 1998) (for review, see Rothman, 1994; Scheller, 1995; Jahn and
Südhof, 1999). In synapses, this complex is composed of the target membrane SNAREs (t-SNAREs) syntaxin (Bennett et al., 1992) and
synaptosomal associated protein of 25 kDa (SNAP-25) (Oyler et
al., 1989), and the vesicle membrane SNARE (v-SNARE)
synaptobrevin/vesicle-associated membrane protein (VAMP)
(Trimble et al., 1988). The core of the ternary complex (Fasshauer et
al., 1998; Poirier et al., 1998a) is a parallel four-helix bundle
(Sutton et al., 1998) that, upon assembly, brings the vesicle and
target membranes together, potentially driving bilayer fusion (Poirier
et al., 1998b; Hanson et al., 1997; Sutton et al., 1998).
Consistent with this model, SNAREs reconstituted into proteoliposomes
can assemble and catalyze membrane fusion in vitro (Weber et
al., 1998).
Although there is evidence that the SNARE complex serves as the core of
the fusion machinery, it is unclear how SNARE-mediated fusion is
regulated by Ca2+. The synaptic vesicle
protein synaptotagmin I (Matthew et al., 1981; Perin et al., 1990)
binds Ca2+ (Brose et al., 1992) and has
been shown, via genetic studies, to be essential for efficient and
rapid excitation-secretion coupling in vivo (Littleton et
al., 1993, 1994; Nonet et al., 1993; DiAntonio and Schwarz,
1994; Geppert et al., 1994). Whether this effect is attributable to a
loss of Ca2+ sensing (Geppert et al.,
1994; Littleton et al., 1994), failure of vesicles to be recycled
(Jorgensen et al., 1995), failure to dock efficiently at release sites
(Reist et al., 1998), failure of the release machinery to be
sequestered near Ca2+ channels (Sheng et
al., 1997), or combinations of these defects in knock-out animals,
remains the subject of debate. However, mutations in
synaptotagmin can alter the
[Ca2+]-response curve for secretion
(Littleton et al., 1994), and disruption of the synaptotagmin
I gene in mice selectively inhibits the fast synchronous component
of exocytosis (Geppert et al., 1994). Furthermore, the equilibrium and
kinetic Ca2+-binding properties of
synaptotagmin are consistent with the Ca2+
requirement and speed of secretion (Davis et al., 1999). These data
support a model in which synaptotagmin functions as a
Ca2+ sensor for secretion, albeit via an
unknown mechanism.
Synaptotagmin spans the vesicle membrane once and binds
Ca2+ via two C2 domains designated C2A and
C2B (Südhof and Rizo, 1996; Desai et al., 2000). One way to
better define the function of synaptotagmin would be to generate
animals that are selectively defective in the
Ca2+-sensing ability of each C2 domain.
Here, we use a genetic approach to determine whether the
Ca2+-sensing ability of the C2B domain of
synaptotagmin functions in synaptic transmission. Furthermore, we
investigate the biochemical relationship between synaptotagmin and
SNARE dynamics and propose a molecular model by which synaptotagmin may
regulate SNARE-catalyzed membrane fusion.
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MATERIALS AND METHODS |
Individual recombinant proteins. cDNA encoding rat
synaptotagmin I (Perin et al., 1990; Osborne et al., 1999) and
human SNAP-25B (Bark and Wilson, 1994) were kindly provided by
T. C. Südhof (Dallas, TX), G. Schiavo (London, UK), and M. Wilson (Albuquerque, NM), respectively. cDNA encoding rat syntaxin 1A
(Bennett et al., 1992) and synaptobrevin II/VAMP II (Elferink et al.,
1989) were kindly provided by R. Scheller (Stanford, CA). Soluble forms
of syntaxin, SNAP-25B, and synaptobrevin were prepared by subcloning into pTrcHisA (Invitrogen, San Diego, CA), resulting in fusion proteins
with T7 and His6 tags at their N termini. His-tagged proteins were
expressed and purified as described (Chapman et al., 1995, 1996).
Wild-type, AD1, and AD3 mutant rat and
Drosophila synaptotagmins were generated by PCR, subcloned
into pGEX-2T, expressed, purified, and cleaved from the GST-fusion
moiety with thrombin, as described (Chapman et al., 1996). All
constructs were confirmed by DNA sequencing.
There are two reported rat synaptotagmin I sequences: one with a
aspartate at position 374 (D374; Perin et al., 1990) and another with a
glycine at this position (G374; Osborne et al., 1999). We have
confirmed that both forms of synaptotagmin are expressed in rats and
have observed that the G374, but not the D374, form clusters in
response to Ca2+ (Davis et al., 1999;
Desai et al., 2000). The basis for this sequence variability is under
investigation. To simplify interpretation of synaptotagmin-SNARE
coimmunoprecipitation experiments, the D374 form was used. In the
assembly experiments shown in Figure 2, the D374 form was used, but
similar results were observed with the G374 form as well as with the
AD3 mutant (data not shown). The G374 form was used in the
oligomerization assays shown in Figure 3.
Midi SNARE complexes. Midi SNARE complexes were composed of
residues 180-262 of syntaxin 1A, full-length SNAP-25A, and residues 1-96 of synaptobrevin II. The components used to assemble midi complexes differ from those used in all other experiments. To obtain
high level expression of SNAP-25, rat SNAP-25A was subcloned into the
vector pET28a (Novagen, Madison, WI) via NheI and
XhoI restriction sites resulting in an N-terminal His6-tag.
In addition, four cysteines (cys 84, 85, 90, and 92) were replaced with
serines using the overlapping primer method (Chapman and Jahn, 1994). These mutations facilitated expression and purification and had no
apparent effects on the structure or binding properties of SNAP-25
(Fasshauer et al., 1999). The cytoplasmic domain of synaptobrevin II
was generated as described (Fasshauer et al., 1997), and
syntaxin fragment 180-262 was expressed using pET15b. SDS-resistant
midi-SNARE complexes were assembled and purified to homogeneity as
described (Fasshauer et al., 1997, 1998).
Immunoprecipitation and antibodies. Mouse monoclonal
antibodies directed against rat synaptotagmin I (41.1), syntaxin
(HPC-1), SNAP-25 (71.2), and synaptobrevin (69.1) were kindly provided by S. Engers and R. Jahn (Göttingen, Germany), and the anti-T7 tag antibody was from Novagen. Monoclonal antisera against
Drosophila syntaxin (8C3) was used at 1:2000, polyclonal
DSYT2 against Drosophila synaptotagmin I at 1:2000
(Littleton et al., 1993), and polyclonal anti-Drosophila
 -adaptin at 1:2000 (Gonzalez-Gaitan et al., 1996).
All immunoprecipitation and bead-binding experiments were performed at
4°C. Immunoprecipitation of recombinant SNAREs and SNARE complexes
was performed as described (Chapman et al., 1995). Briefly, recombinant
individual SNAREs or SNARE complexes were incubated with recombinant
synaptotagmin in Tris-buffered saline (TBS; 20 mM Tris, pH
7.4, 150 mM NaCl) plus 0.5% Triton X-100 in the presence
of 2 mM EGTA or 1 mM
Ca2+ for 2 hr. Syntaxin, synaptobrevin, or
SNAP-25 was immunoprecipitated by incubating the samples with HPC-1 (5 µl), 69.1 (1.5 µl), or 71.1 (5 µl) ascites, respectively, for 2 hr and 12 µl of Protein G Sepharose Fast-flow (Amersham Pharmacia
Biotech) for 1 hr. The immunoprecipitates were washed three times and
analyzed by SDS-PAGE and either immunoblotting or staining with
Coomassie blue. As a control for nonspecific precipitation of
synaptotagmin, samples were also prepared lacking SNAREs. In each case,
the immunoprecipitating antibodies did not bind synaptotagmin, and,
under the conditions of the binding assays, synaptotagmin did not
precipitate in the absence of SNAREs. Thus, for experiments shown in
the Figures ( SNARE) samples also lacked immunoprecipitating
antibodies. Coimmunoprecipitation of synaptotagmin was quantified using
a Bio-Rad (Hercules, CA) GS-670 Imaging Densitometer. Generation of
Drosophila head homogenates for AP-2 binding assays was as
previously described (Littleton et al., 1998).
[Ca2+] determination.
For Ca2+ titration experiments,
[Ca2+]free was
determined using a Microelectrode MI-600
Ca2+ electrode, MI-402 microreference
electrode (Bedford, NH), and World Precision Instruments (Sarasota, FL)
Ca2+ standards
(pCa2+ range of 1-8).
Ca2+ concentrations <100
µM were buffered using 2 mM EGTA.
In vitro assembly of SDS-resistant 7S SNARE complexes.
His6-tagged synaptobrevin II (1-96), SNAP-25B (1-206), and syntaxin 1A (1-265) were incubated at 0.5 µM with
constant mixing for 0, 5, 15, or 120 min at 25°C with or without 2 µM recombinant cytoplasmic domain of
synaptotagmin Ia. All assembly reactions were performed using freshly
purified proteins in 150 µl of TBS supplemented with either 2 mM EGTA, 1 mM
Mg2+, or 1 mM
Ca2+. In some experiments, assembly was
performed in the presence of 1 mM DTT.
Assembly reactions were stopped by adding 15 µl of 3× SDS sample
buffer, containing 10%  -mercaptoethanol, to 30 µl of the reaction
mixture. Samples were loaded onto discontinuous 9-15% SDS-PAGE
mini-gels (Bio-Rad) without boiling (except where indicated) and
separated at 15 mA per gel. The gels were immunoblotted using
monoclonal antibodies directed against syntaxin, synaptobrevin, or
synaptotagmin; immunoreactive bands were visualized using enhanced chemiluminescence. Synaptotagmin consistently enhanced 7S assembly, but
this effect was variable, ranging from 1.5- to 4-fold. A representative experiment showing a threefold enhancement at 5 min is shown in Figure
2A.
Interaction of synaptotagmin oligomers with midi-SNARE
complexes. Fifteen micrograms of GST-tagged synaptotagmin
(amino acids 96-421; G374-version) immobilized on beads was incubated
with 10 µM soluble synaptotagmin (amino acids
96-421; G374-version) for 1.5 hr in 150 µl of HEPES-buffered saline
(HBS; 50 mM HEPES, pH 7.4, 100 mM NaCl)
plus 0.5% Triton X-100 in either 2 mM EGTA or 1 mM Ca2+. Beads were
washed three times in binding buffer plus 2 mM
EGTA or 1 mM
Ca2+, and then incubated with 2 µM midi-SNARE complex for 1.5 hr. Beads were
washed three times as described above, and 25% of the samples were
subjected to SDS-PAGE; gels were stained with Coomassie blue.
To determine whether SNARE complexes can inhibit the binding of soluble
synaptotagmin to immobilized synaptotagmin, oligomerization assays were
performed as described above. However, free soluble synaptotagmin was
not removed by washing, and the concentration of SNARE complex was
titrated (1, 3, and 6 µM). For analysis, samples were
boiled, separated by SDS-PAGE, bound synaptotagmin was visualized by
staining with Coomassie blue, and SNAREs were detected by immunoblot analysis.
Isolation of Drosophila SNARE complexes. Flies of
the indicated genotype were frozen in liquid nitrogen, vortexed, and 10 heads for each genotype were homogenized in 50 µl of SDS sample buffer on ice. The samples were briefly centrifuged to pellet cuticle,
and 20 µl of the supernatant was resuspended in 30 µl of SDS sample
buffer. Samples were loaded onto discontinuous 9 and 15% SDS-PAGE gels
without boiling and separated at 15 mA per gel. The gels were
immunoblotted with anti-syntaxin monoclonal antibody 8C3 at 1:2000
dilution. Immunoreactive bands were visualized using ECL.
EM. Transmission electron microscopy quantification
at photoreceptor synapses was done as previously described (Littleton et al., 1998). The number of vesicles per T-bar was determined by
counting vesicles that were under the arms of an active zone T-bar and
within 40 nM of the presynaptic membrane. Error
measurements are reported in SD.
Drosophila genetics. Flies were cultured on standard medium
at 23°C.
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RESULTS |
Synaptotagmin drives SNARE complex assembly, and
Ca2+-synaptotagmin drives the cross-linking
of SNARE complexes into dimers
To define the biochemical relationship between synaptotagmin
activity and SNARE complex assembly, we undertook a detailed analysis
of the interaction dynamics between these two essential elements of the
vesicle fusion machinery. Direct interactions between synaptotagmin and
t-SNAREs have been previously demonstrated; synaptotagmin binds, in a
stoichiometric and Ca2+-promoted manner,
to both syntaxin and SNAP-25 (Chapman et al., 1995; Schiavo et al.,
1997; Davis et al., 1999; Gerona et al., 2000). To further explore the
interaction of synaptotagmin with SNAREs, we assembled syntaxin and
SNAP-25 with synaptobrevin to form SDS-resistant ternary SNARE
complexes. Because synaptotagmin binds solely to the H3-domain of
syntaxin (Chapman et al., 1995; Kee and Scheller, 1996; Davis et al.,
1999), we used "midi" SNARE complexes composed of residues 180-263
of syntaxin 1A, full-length SNAP-25A, and residues 1-96 of
synaptobrevin II. Assembly of the midi complex is confirmed in Figure
1A (left
panel), where the complex runs at ~67 kDa on SDS
polyacrylamide gels and, after boiling, dissociates into the three
individual SNARE proteins. Midi complexes were incubated with
increasing concentrations of recombinant rat synaptotagmin in either
EGTA or Ca2+ and then immunoprecipitated
with anti-synaptobrevin antibodies. Immunoprecipitates were boiled and
analyzed by SDS-PAGE and Coomassie staining. As shown in Figure
1A (middle and right panels),
synaptotagmin bound to midi complexes in both EGTA and in
Ca2+. Under these conditions,
Ca2+ increased the affinity of
synaptotagmin for the midi complex by approximately an order of
magnitude. In the presence of Ca2+, the
EC50 for synaptotagmin binding to SNARE complexes
was 1-2 µM. At saturation, the stoichiometry
was 0.5 moles of synaptotagmin per mole of complex, suggesting that one
copy of synaptotagmin binds two copies of the SNARE complex.
Furthermore, the
[Ca2+]1/2 for the
interaction of synaptotagmin with the SNARE complex was ~100
µM Ca2+ (Fig.
1B), consistent with the
Ca2+ dependence for secretion in retinal
bipolar neurons (194 µM
Ca2+; Heidelberger et al., 1994) and
within a factor of 10 of the Ca2+
dependence for secretion at the calyx of Held (10-20
µM Ca2+; Bollman
et al., 2000; Schneggenburger and Neher, 2000). Binding was selectively
promoted by Ca2+ versus other divalent
cations (Fig. 1C).
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Figure 1.
Interaction of synaptotagmin with
assembled SNARE complexes. A, Left panel,
The "midi" SNARE complex is SDS-resistant. Midi SNARE complex (1.5 µg) was dissociated into its component parts (residues 1-96 of
synaptobrevin, 1-206 of SNAP-25, and 180-262 of syntaxin) by boiling
in SDS sample buffer. Middle panels, Increasing
concentrations of synaptotagmin were incubated with midi complexes (2 µM) in the presence of EGTA or Ca2+ in
a 75 µl reaction volume. Synaptotagmin binding was assayed by
coimmunoprecipitation using anti-synaptobrevin antibodies. Proteins
were separated by SDS-PAGE and visualized with Coomassie blue.
Forty percent of the bound material was loaded onto the gel.
Right panel, Coimmunoprecipitated synaptotagmin was
quantified by densitometry. The level of binding in EGTA (open
circles) and Ca2+ (closed
circles) was normalized to the maximum level of binding and
plotted versus [synaptotagmin]. In the presence of
Ca2+, the EC50 was 1.7 µM;
at saturation the stoichiometry was 0.5 mol of synaptotagmin per
mole of midi complex. B, Left
panel, Synaptotagmin (3 µM) was mixed with
midi-SNARE complex (2 µM) in 75 µl of HBS-0.5%
Triton X-100 plus EGTA (2 mM) or the indicated
concentration of Ca2+ for 2 hr at 4°C. SNARE
complexes were immunoprecipitated with an anti-synaptobrevin antibody.
Proteins were separated by SDS-PAGE and stained with Coomassie blue.
Forty percent of the bound material was loaded onto the gel;
total corresponds to 10% of the binding reaction.
Right panel, Coimmunoprecipitated synaptotagmin was
quantified by densitometry, normalized, and plotted versus the
free Ca2+ concentration. The
[Ca2+]1/2 was ~100 µM.
C, Synaptotagmin-midi-SNARE complex formation was
monitored as described in B in the presence of the
indicated divalent cations (1 mM Mg2+;
200 µM Ca2+, Ba2+,
Sr2+). The synaptotagmin and midi-SNARE complex
concentrations were 2 µM. Synaptotagmin binding was
normalized (binding in 2 mM EGTA and 200 µM
Ca2+ were set at 0 and 100% binding, respectively),
and the means from triplicate determinations are plotted. Error bars
represent the SD from triplicate determinations.
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Isolated t-SNAREs have a distinct and less ordered conformation than
t-SNAREs that are assembled into the four helix bundle that constitutes
the core of the SNARE complex (Fasshauer et al., 1997; Sutton et al.,
1998; Fiebig et al., 1999). The observation that synaptotagmin binds to
isolated t-SNAREs, as well as to assembled SNARE complexes, efficiently
and in a Ca2+-regulated manner, indicates
that isolated t-SNAREs are ordered into their ternary "SNARE-complex
conformations" after complex formation with synaptotagmin. This model
predicts that synaptotagmin, via its "ordering" of t-SNAREs, would
facilitate assembly of SNARE complexes. To test this prediction, we
incubated purified SNAREs (syntaxin, SNAP-25, and synaptobrevin) with
and without rat synaptotagmin in the presence and absence of
Ca2+, for increasing periods of time.
SDS-resistant SNARE complexes were detected using antibodies directed
against syntaxin (Fig. 2A), synaptobrevin
(Fig. 2C), or SNAP-25 (data not shown). These complexes were
disassembled into monomeric SNAREs after boiling in SDS (Fig.
2A,C). Consistent with previous reports,
SDS-resistant SNARE complexes formed in the absence of synaptotagmin
and Ca2+ (Fig. 2A;
Hayashi et al., 1994). Under these conditions,
Ca2+ had no apparent effect on the rate or
extent of SDS-resistant SNARE complex assembly. However, addition of
synaptotagmin to mixtures of isolated SNAREs accelerated SNARE complex
assembly (Fig. 2A,B). This effect is marked at early
time points; at 5 min, synaptotagmin drove a threefold enhancement of
SNARE complex assembly, and by 120 min equal amounts of SNARE complex
accumulated in the presence and absence of synaptotagmin (Fig.
2C). Surprisingly, the ability of synaptotagmin to drive
complex assembly was Ca2+-independent,
despite the fact that Ca2+ promotes
binding of synaptotagmin to t-SNAREs. Therefore, in addition to the
kinetics experiments shown in Figure 2A, we also conducted synaptotagmin-titration experiments. In all kinetic and
titration experiments, the ability of synaptotagmin to enhance the rate
of SNARE complex assembly was independent of
Ca2+ (data not shown). The reason for this
lack of a Ca2+-effect is unclear. One
possibility is that the difference in affinity of synaptotagmin for
SNAREs in the presence and absence of Ca2+
is not great enough to yield differences under the conditions of our
assembly experiments where we measured assembly on minute rather than
millisecond time scales. However, we observed that Ca2+-synaptotagmin can trigger the
formation of SDS-resistant SNARE complex dimers (Fig. 2C).
These dimers result from intermolecular disulfide bonds that can be
disrupted by DTT (Fig. 2C). Thus, Ca2+ can act via synaptotagmin to drive
the "cross-linking" of two SNARE complexes together. These findings
are congruent with the estimated stoichiometry of one synaptotagmin
bound to two SNARE complexes as described in Figure
1A. We note that in some experiments, Ca2+ alone was able to trigger low levels
of cross-linked dimer formation, perhaps via direct effects on the
SNARE complex (Sutton et al., 1998). However, in all experiments, this
effect was markedly enhanced by the addition of synaptotagmin.
Furthermore, cross-linking is specifically driven by
Ca2+-synaptotagmin, 1 mM Mg2+ does not
trigger cross-linking, nor does it inhibit
Ca2+-synaptotagmin-driven cross-linking
(Fig. 2C). Control experiments demonstrated that the dimers
were composed of all three SNAREs; omission of any of the SNAREs
precluded dimer formation (Fig. 2C), and dimers were
recognized by antibodies directed against each component of the SNARE
complex (Fig. 2A,C; data not shown). In summary,
these data indicate that Ca2+, acting via
synaptotagmin, can drive conformational changes in SNARE complexes that
result in the formation of cross-linked SNARE complex dimers.
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Figure 2.
Synaptotagmin facilitates SNARE complex assembly
in vitro. A, Recombinant his6-syntaxin,
his6-SNAP-25B, and his6-synaptobrevin were incubated in the presence
and absence of recombinant synaptotagmin in 2 mM EGTA
(Ca2+) or 1 mM Ca2+
(+Ca2+) for 0, 5, or 15 min at room
temperature. SDS-resistant 7S SNARE-complex formation was
assayed by subjecting the samples to SDS-PAGE, without previous boiling
(except where indicated), and immunoblotting with anti-syntaxin
antibodies. Immunoreactive bands were visualized using enhanced
chemiluminescence. 7S denotes an SDS-resistant complex
consisting of syntaxin, SNAP-25, and synaptobrevin.
dimer denotes the trace formation of disulfide-bonded
SNARE complex dimers that form under these conditions.
B, The optical densities of 7S complexes from the
+Ca2+ lanes in A are plotted versus
time of incubation. C, Assembly experiments were
performed as described in A for 2 hr but in the absence
of DTT, except where indicated. Assembly reactions were conducted with
(+) or without () synaptotagmin in either 2 mM EGTA, 1 mM Mg2+, 1 mM
Ca2+, or 1 mM Mg2+
plus 1 mM Ca2+. As controls, samples
were prepared that lacked either synaptobrevin, syntaxin, or SNAP-25.
As a further control, samples were boiled before analysis. Samples were
analyzed by immunoblotting with a mixture of anti-synaptobrevin and
anti-synaptotagmin antibodies. Ca2+ and
synaptotagmin enhanced the formation of SDS-resistant dimers. These
dimers are disulfide-linked and are dissociated by DTT. SDS-resistant
7S complex formation only occurs in the presence of all three SNAREs,
and complexes are dissociated by boiling.
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Assembly of synaptotagmin oligomers with SNARE complexes
One prominent feature of the synaptotagmin family is the ability
of synaptotagmins to undergo both homo- and hetero-oligomerization in a
Ca2+-dependent manner (Sugita et al.,
1996; Chapman et al., 1998; Osborne et al., 1999; Desai et al., 2000).
This property is conserved from invertebrates to mammals (Littleton et
al., 1999; Desai et al., 2000), suggesting it may be essential
for the function of synaptotagmin in neurotransmitter release. Given
that Ca2+-triggered synaptotagmin
self-association occurs on rapid time scales (Davis et al., 1999), one
hypothesis is that oligomerization may play an important role in late
stages of SNARE assembly and clustering of SNARE complexes into a
collar-like fusion pore. If this model is correct, synaptotagmin should
be able to oligomerize and bind to SNARE complexes at the same time. To
examine this possibility, synaptotagmin was immobilized on beads and
assayed for SNARE complex binding activity. We made use of the
observation that synaptotagmin, fused to GST, cannot efficiently bind
SNARE complexes. After removal of the GST moiety, high-affinity
complexes between synaptotagmin and SNARE complexes are efficiently
assembled (Chapman et al., 1996). This result is shown in Figure
3A, where the immobilized
GST-synaptotagmin fusion protein bound SNARE complexes only weakly in
both the absence and presence of Ca2+
(Fig. 3A). We then assembled soluble synaptotagmin onto
immobilized GST-synaptotagmin by virtue of the
Ca2+-triggered clustering activity of the
protein. Unbound synaptotagmin was removed by washing, and the ability
of the synaptotagmin-GST-synaptotagmin oligomer was assayed for SNARE
complex binding activity. As shown in Figure 3A,
oligomerized synaptotagmin efficiently captured assembled SNARE
complexes.
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Figure 3.
Oligomerized synaptotagmin binds to assembled
SNARE complexes. A, GST and GST-synaptotagmin were
immobilized on beads (15 µg per data point) and assayed for binding
to midi-SNARE complexes (2 µM) in 2 mM EGTA
(Ca2+) or 1 mM Ca2+
(+Ca2+) in 150 µl of HBS using a cosedimentation
assay, as described in Materials and Methods. To leave SNARE complexes
intact, samples were subjected to SDS-PAGE without previous boiling.
Coomassie staining revealed only low levels of SNARE binding to
immobilized synaptotagmin in either condition. Immobilized
synaptotagmin was then preincubated with soluble synaptotagmin (10 µM) in EGTA or Ca2+. Beads were washed
three times to remove unbound soluble synaptotagmin, and the soluble-
immobilized synaptotagmin oligomers were assayed for binding to
midi-SNARE complexes. Twenty-five percent of the bound material was
loaded onto the gel; the left two lanes correspond to
0.3 and 0.5 µg of soluble synaptotagmin and midi-SNARE complex,
respectively. Coomassie staining revealed efficient binding of soluble
synaptotagmin to immobilized synaptotagmin. Furthermore, midi-SNARE
complexes efficiently bound to the soluble-immobilized synaptotagmin
oligomers. These results demonstrate that synaptotagmin, which has
oligomerized, is capable of binding SNARE complexes. *Denotes
proteolytic fragments from GST-synaptotagmin. Note,
Ca2+ induces a shift in the mobility of
synaptotagmin that has not been boiled. Therefore, soluble and
GST-synaptotagmin are indicated with double arrows.
B, SNARE complexes do not inhibit synaptotagmin
oligomerization. GST (12 µg per data point) and GST-synaptotagmin
(8 µg per data point) were immobilized on beads. Soluble
synaptotagmin (1.5 µM; +) and midi-SNARE complex (6 µM; +) or the indicated [SNARE complex] were incubated
with the beads in 2 mM EGTA () or 1 mM
Ca2+ (+) for 1.5 hr. Samples were also prepared that
lacked SNARE complexes () or soluble synaptotagmin (). Bound
material was boiled in SDS sample buffer and subjected to SDS-PAGE.
Twenty-five percent of the bound material was loaded onto the gel;
total corresponds to the mixture of 0.3 and 0.7 µg of soluble
synaptotagmin and midi-SNARE complex. Gels were stained with Coomassie
blue to visualize bound synaptotagmin. Staining of disassembled SNARE
complexes was poor, therefore SNARE binding was detected by
immunoblotting with anti-SNAP-25 and anti-syntaxin antibodies.
Immunoreactive bands were visualized using enhanced
chemiluminescence.
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As a further test of the model in which synaptotagmin oligomerizes and
binds to SNAREs at the same time, we determined whether SNARE complexes
act as competitive inhibitors of synaptotagmin oligomerization. For
these experiments we monitored the
Ca2+-dependent binding of soluble
synaptotagmin to immobilized synaptotagmin in the presence of
increasing concentrations of SNARE complexes. As shown in Figure
3B, addition of SNAREs did not impede oligomerization. Our
biochemical observations are consistent with a role for synaptotagmin oligomerization and SNARE binding in triggering vesicle fusion. To directly test this model, we characterized mutations in
Drosophila synaptotagmin I that block
Ca2+-dependent oligomerization.
The C2B domain of synaptotagmin is required for both
exocytosis and endocytosis of synaptic vesicles in
vivo
A collection of 20 different alleles of synaptotagmin I
(syt) have been generated in Drosophila
(Littleton et al., 1993, 1994; DiAntonio and Schwarz, 1994), providing
useful experimental material to determine the mechanism by which
synaptotagmin functions in synaptic vesicle cycling. Many of these
mutations in syt, including P-element insertions,
enhancer/promoter deletions, and early stop codons in the open reading
frame (DiAntonio and Schwarz, 1994; Littleton et al., 1994),
disrupt synaptic function by decreasing the levels of wild-type
synaptotagmin at synapses. However, several syt alleles
display intragenic complementation (Littleton et al., 1994). This form
of complementation is often observed for genes that encode proteins
that are part of multimeric complexes and that contain multiple
distinct functional domains. Thus, intragenic complementation for
syt mutations suggests the presence of several independent
domains within synaptotagmin that mediate distinct steps in
neurotransmitter release. Two of the syt alleles involved in
intragenic complementation are AD1 and AD3. Flies
containing various heteroallelic combinations with AD1 and
AD3 display defects including a severe lack of coordination
and dramatically decreased viability. Previous electrophysiological
analysis of heteroallelic combinations involving AD1 and
AD3 (Littleton et al., 1994) demonstrated a profound
decrease in synaptic exocytosis. At low
Ca2+ concentrations, evoked release is
virtually abolished in AD3 mutants. By raising extracellular
Ca2+ to 6 mM,
exocytosis in AD3 mutants can be partially rescued (Fig. 4C). In contrast,
AD1 mutants have severe defects in synaptic transmission
that cannot be rescued by higher levels of extracellular Ca2+ (Fig. 4C). In addition,
recordings from synaptotagmin alleles that are viable with
AD1 (T7 and T41) and AD3
(T7, T41, T11, D2, D3, D37, D45) show either the
AD1 or AD3 phenotype regardless of the other
allele with which AD1 or AD3 are paired. Indeed, the same synaptotagmin mutants (T41, T7)
behave dramatically different when paired with AD1 or
AD3. Thus, the AD1 and AD3 alleles
confer the dominant phenotype to any synaptotagmin allele
with which they are paired (even when paired with the same
alleles T7, T41), leading us to focus on the molecular
defects in the AD1 and AD3 mutants.
Immunolocalization studies reveal that the mutant synaptotagmins are
targeted to synapses in AD1 and AD3 mutants (data
not shown). The amount of synaptotagmin that is present at mutant
synapses is difficult to quantify precisely because we do not know how these mutations affect the ability of our anti-synaptotagmin I antibody
to detect the mutated protein in vivo. However,
AD1 and AD3 mutants over a deletion that
completely removes synaptotagmin are far less severe phenotypically
than null mutants such as T77 and AD4 over
deletion (Littleton et al., 1994) and survive much longer as larva than
do null mutants. These observations directly demonstrate that the
AD1 and AD3 mutant synaptotagmin proteins are
made and have partial function at synapses, allowing these mutants to
survive and function more efficiently than mutants that completely
remove synaptotagmin and die as embryos. Thus, an altered
func- tion of the mutant synaptotagmins, rather than a loss of
the protein at synapses, is likely the cause of the electrophysiological defects. We cannot completely rule out some contribution to the phenotype from altered protein levels that are
beyond our detection. Sequence analysis of AD1 and
AD3 revealed that the AD1 phenotype is caused by
a premature stop codon that deletes the C2B domain. AD3
results from a Y to N substitution in C2B (DiAntonio and Schwarz, 1994)
at a residue (364) that is highly conserved in all synaptotagmin
isoforms from Caenorhabditis elegans to humans (Fig.
4A). The crystal structure of the cytoplasmic domain
of rat synaptotagmin III (Sutton et al., 1999) indicates that the
AD3 mutation lies near two conserved aspartate residues that
may function as Ca2+ ligands (Fig.
4B). These two mutants allow us to investigate the
in vivo roles of the C2B domain of synaptotagmin I in
synaptic function.
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Figure 4.
Mutations in the C2B domain of
Drosophila synaptotagmin I. A, Alignment
of the C2B domain sequence surrounding the Y364N change found in the
AD3 mutant (DiAntonio and Schwartz, 1994). The five
putative Ca2+ ligands are highlighted in
gray, whereas the AD3 change is indicated
in black. Y364 is conserved among all synaptotagmin
isoforms from C. elegans to humans. B,
Predicted structure of the AD1 and AD3 mutant proteins based on the
crystal structure of synaptotagmin III (see Fig. 9 for details). The
location of the Y to N change in AD3 is indicated by the
arrow. The AD1 mutations result in
a premature stop codon deleting the C2B domain. C, The
electrophysiological defects observed in AD3 and
AD1 heteroallelic combinations (Littleton et al., 1994)
are plotted against the responses of the control cn bw
sp line. Recordings were made in 0.4 or 6.0 mM
Ca2+ in Jan's Ringer's solution.
Excitatory junctional potential (EJP) amplitude at muscle fiber
6 in segments A3-A5 is plotted vs the extracellular
Ca2+ concentration. At low Ca2+,
both AD1 and AD3 exhibit a profound block in evoked secretion. At
higher Ca2+ levels, the defects in
AD3 mutants can be partially rescued, whereas
AD1 mutants continue to have dramatically abnormal
synaptic responses. These EJP responses have not been corrected
for nonlinear summation. Thus, both synaptotagmin
mutants still have significant defects compared with control responses
even in high calcium, where the control responses already saturated at
these calcium levels. Dominant defects from the AD1 and
AD3 alleles when paired with a wild-type allele of
synaptotagmin have not been observed (Littleton et al., 1994).
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We first examined morphological defects in AD1 and
AD3 mutants to determine where in the synaptic vesicle cycle
each mutant is blocked. For this analysis we examined the first optic
neuropil containing 800 highly stereotypic optic cartridges with
defined synaptic contacts between photoreceptor axons and laminar
neurons that can be readily identified by the presence of presynaptic T-bars. We focused specifically on the histaminergic synapses between
photoreceptors (R1-R6) and postsynaptic laminar neurons (L1 and L2).
Electroretinogram (ERG) recordings, in which synaptic transmission
between photoreceptors and second order neurons in the lamina is
indicated by the on and off transients in response to a light flash,
revealed that both AD1 and AD3 heteroallelic mutants lack these transients (Fig. 5).
Thus, these mutations disrupt synaptic transmission at photoreceptor
synapses as well as at neuromuscular junctions.
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Figure 5.
Ultrastructural analysis of stimulated synapses in
C2B mutants. Ultrastructural defects in control cn
(A), AD3 cn/T11 cn
(B), and AD1 cn/T41 cn
(C) photoreceptor synapses were examined by
driving photoreceptors with constant light stimulation for 10 min,
followed by rapid fixation. Both AD1 and
AD3 mutants lack the on-off transients measured during
ERG recordings in the retina (shown on the right),
demonstrating that synaptic transmission is disrupted at these
photoreceptor synapses. AD1 mutants show a decrease in
the overall number of synaptic vesicles, whereas AD3
synapses do not show a depletion of synaptic vesicles, but rather a
defect in the ability of docked synaptic vesicles to fuse.
Quantification of vesicles per photoreceptor synapse for each of the
genotypes was: AD1 cn/T41 cn, 25 ± 14 SD;
AD1 cn/T7 cn, 27 ± 20 SD; AD3 cn/T11
cn, 88 ± 28 SD; cn controls, 96 ± 37 SD. Quantification of vesicles per T-bar for each of the genotypes was:
AD1 cn/T41 cn, 1.4 ± 0.9 SD; AD1 cn/T7
cn, 1.9 ± 0.9 SD; AD3 cn/T11 cn, 2.6 ± 1.4 SD; cn controls, 2.3 ± 0.9 SD.
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To examine the morphological correlates of the block in synaptic
transmission in AD1 and AD3, electron microscopy
was performed on the photoreceptor synapses after 10 min of constant
light stimulation before fixation to drive continuous vesicle cycling.
A total of 168 micrographs were examined from cn,
AD1/T41, AD1/T7, and AD3/T11 flies
(n = 3-10 flies for each genotype). The overall
architecture of the lamina was normal in syt mutants. The
most dramatic difference was a decrease (p < 0.05, unpaired Student's t test) in the number of synaptic
vesicles in photoreceptor terminals of AD1/T7 and AD1/T41 mutants compared with controls (AD1 cn/T41
cn, 25 + 14 SD; AD1 cn/T7 cn, 27 ± 20 SD; AD3 cn/T11 cn, 88 ± 28 SD; cn controls, 96 ± 37 SD (Fig. 5, compare A, B),
suggesting a defect in endocytosis in AD1 heteroallelic
mutants. Although we cannot rule out that the loss of synaptic vesicles
in AD1 mutants is caused by a defect in vesicle biogenesis
from an internal compartment as opposed to a direct defect in
endocytosis, the lack of any vesicle biogenesis defect in
synaptotagmin null mutants (Reist et al., 1998) argues against this alternative interpretation. In contrast, AD3
mutants did not have depleted nerve terminals compared with controls, indicating that endocytosis in not disrupted in this mutant. Indeed, synaptic vesicles could be clearly visualized in contact with the
presynaptic membrane under T-bars (Fig. 5C), indicating that AD3 mutant vesicles can undergo docking but are defective at
a later step in exocytosis (2.6 ± 1.4 SD docked vesicles in
AD3 cn/T11 cn compared with 2.3 ± 0.9 in cn controls).
The morphological and electrophysiological analysis of AD1
and AD3 suggest that fundamentally different processes are
affected in the two mutants. AD1 terminals are relatively
depleted of synaptic vesicles compared with controls (Fig.
5B), and synaptic transmission cannot be rescued by high
extracellular Ca2+ (Fig. 4C).
AD3 mutants have a defect in exocytosis, not endocytosis, (Fig. 5C; see Fig. 7A), and release can be
partially rescued by high extracellular
Ca2+ (Fig. 4C). Thus, the
morphologically docked vesicles in AD3 mutants are also
physiologically competent for release but require significantly higher
Ca2+ concentrations. We therefore examined
the biochemical defects caused by AD1 and AD3 to
determine which activities of the C2B domain are required for
endocytosis and exocytosis, respectively. For this analysis we
generated GST fusion proteins containing the cytoplasmic domains from
wild-type Drosophila synaptotagmin, AD1 (C2A domain alone
lacking C2B), and AD3. To determine whether AD1 and AD3 are able to
penetrate membranes in the presence of Ca2+, we tested the immobilized
recombinant cytoplasmic domains for their ability to bind liposomes
(25% phosphatidyl serine, 75% phosphatidyl choline) with or without
Ca2+ (Fig.
6A). Wild-type, AD1,
and AD3 fusion proteins all showed robust
Ca2+-dependent phospholipid binding, an
activity previously shown to be mediated by the C2A domain of
synaptotagmin Ia (Bai et al., 2000; Desai et al., 2000). We next
examined binding to the t-SNARE, syntaxin 1A, whose
Ca2+ dependence is also mediated by
Ca2+ ligands in the 2A domain of
synaptotagmin Ia (Bai et al., 2000; Desai et al., 2000). Recombinant
Drosophila syntaxin 1 was able to interact with wild-type,
AD1, and AD3 fusion proteins (Fig. 6B). We also
examined the interaction of synaptotagmin with the clathrin adapter
AP-2. Binding of the AP-2 complex from Drosophila head
extracts was detected with an antibody generated against -adaptin
(Gonzalez-Gaitan et al., 1996). Whereas wild-type and AD3 synaptotagmin
bound AP-2 in the absence or presence of
Ca2+, AD1 synaptotagmin did not bind AP-2
under either condition (Fig. 6B). Thus, as reported
for mammalian synaptotagmin I, AP-2 binding to Drosophila
synaptotagmin is also mediated through the C2B domain (Zhang et al.,
1994). We conclude that AP-2 binding to synaptotagmin and subsequent
clathrin recruitment is altered in AD1 mutants, leading to
defective endocytosis and a relative depletion of synaptic vesicles in
stimulated synapses. Although complete removal of the C2B domain would
also be expected to disrupt the exocytotic activities mediated by C2B,
the loss of vesicles in the AD1 mutant dominates the
morphological and electrophysiological phenotype. A similar
morphological depletion of synaptic vesicles has been observed in a
C. elegans synaptotagmin mutant that also deletes the C2B
domain (Jorgensen et al., 1995). These endocytotic defects preclude the
investigation of the role of the C2B domain in exocytosis in AD1
mutants. The lack of any defect in AP-2 binding by the AD3 mutant protein and the corresponding lack of an
endocytotic defect by morphological or electrophysiological criteria in
AD3 mutants defines a second function for the C2B domain of
synaptotagmin in synaptic vesicle exocytosis that is disrupted in
AD3 mutants.
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Figure 6.
Synaptotagmin AD1 mutants fail to bind AP-2.
A, Ca2+-dependent phospholipid
binding of immobilized recombinant wild-type
(WT), AD3, or AD1 synaptotagmin I proteins. Both
AD1 and AD3 recombinant proteins showed robust
Ca2+-stimulated phospholipid binding. Phospholipid
binding assays were conducted as previously described (Littleton et
al., 1999). B, Binding of recombinant syntaxin (5 µM) and native AP-2 -adaptin (0.2 mg of
Drosophila head membranes) to 30 µg of recombinant WT,
AD3, or AD1 Drosophila synaptotagmins in 2 mM EGTA or 1 mM Ca2+ for 2 hr at 4°C. For detection of recombinant syntaxin binding to
synaptotagmins, Western analysis with the monoclonal anti-syntaxin
antisera 8C3 was performed. For analysis of AP-2 binding, fly head
membranes were prepared as previously described (Littleton et al.,
1998), and AP-2 binding was detected with a polyclonal antibody
generated against -adaptin (Gonzalez-Gaitan and Jackle, 1996).
Immunoreactive bands were visualized by enhanced chemiluminescence.
Both AD1 and AD3 mutant proteins showed
Ca2+-dependent binding to syntaxin. However, only
AD3 showed an interaction with AP-2.
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The AD3 mutation selectively impairs
Ca2+-driven conformational changes and
Ca2+-triggered oligomerization of the C2B domain of
synaptotagmin
Synaptic vesicles in AD3 mutants are capable of
translocating and docking at active zones during synaptic stimulation,
as shown in Figure 7A. Thus,
it is likely that the defect in AD3 mutants lies somewhere
after docking. This interval encompasses both priming and fusion,
although it is unknown what molecular events occur during this period.
One possibility is that individual SNAREs assemble into various stages
of "loose" and "tight" states of SNARE complexes (Xu et al.,
1999), generating a potential fusion pore that can be triggered to open
by Ca2+. We were thus interested in
determining whether there were defects in a specific stage of SNARE
assembly in the AD3 mutant. One possibility is that
synaptotagmin is required to trigger SNARE complex assembly. Another
possibility is that synaptotagmin binds preassembled SNARE complexes
and prevents them from mediating full fusion until arrival of a
Ca2+ signal. To explore these
possibilities, we examined 7S complexes in head extracts of
AD3 mutants. SNARE complexes do not form in SDS (Littleton
et al., 1998), but once the complex is formed, they are resistant to
dissociation by SDS unless the sample is boiled (Hayashi et al., 1994).
To assay complex formation in flies, supernatants from SDS-solubilized
control and AD3 mutant fly heads were separated on SDS-PAGE
gels and probed with an anti-syntaxin antiserum. Monomeric 35 kDa
syntaxin can be detected, as well as the 73 kDa SNARE complex
containing syntaxin, synaptobrevin, and SNAP-25. Because 7S SNARE
complexes do not form in SDS, the complex we detect corresponds to that
present in vivo. AD3 mutants show a dramatic decrease in the
amount of 7S complex (Fig. 7B), consistent with a defect in
the ability of synaptotagmin to trigger SNARE complex formation rather
than a defect in triggering fusion after SDS-resistant SNARE complex
assembly. The block in SNARE complex assembly in AD3 mutants
suggests an activity mediated by the C2B domain of synaptotagmin that
also acts late in the exocytotic pathway to trigger SNARE assembly and
consequent fusion. We therefore undertook an investigation of the
defects resulting from the Y364N change in C2B in the AD3
mutant to uncover a biochemical link between synaptotagmin, assembly of
the SNARE complex, and vesicle fusion.
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Figure 7.
Synaptotagmin AD3 mutants
decrease SNARE complex assembly in vivo.
A, Enlarged image of an active zone in
AD3/T11 mutant photoreceptor terminals demonstrating
docked vesicles (arrowheads) under a T-bar that have not
fused. B, 7S complexes from 10 control
(CS) or synaptotagmin AD3/T11 mutants
were isolated. Syntaxin is present in a 35 kDa monomeric form and in a
73 kDa complex with SNAP-25 and synaptobrevin in wild-type flies. A
severe reduction in the amount of 7S complex was found in
AD3/T11 synaptotagmin mutants. C, Both
wild-type and AD3 recombinant synaptotagmins are able to bind SNARE
complexes in a Ca2+-stimulated manner. Either 3 µM wild-type (sytWT) or AD3 mutant
synaptotagmin (sytAD3) was incubated with 3 µM midi-SNARE complex for 1.5 hr in either 2 mM EGTA (E) or 1 mM
Ca2+. Midi-SNARE complex was immunoprecipitated,
and samples were separated by SDS-PAGE and stained with Coomassie
blue. As a control, samples were prepared that lacked midi-SNARE
complex and immunoprecipitating antibodies. Thirty percent of the
immunoprecipitated material was loaded onto the gel; total corresponds
to 6% of the binding reaction. Note: the asterisk
indicates a proteolytic fragment present in preparations of soluble AD3
mutant rat synaptotagmin. D, Both wild-type and AD3
synaptotagmins are able to bind the mammalian synprint peptide. Ten
micrograms of GST or GST fused to the cytoplasmic domain of WT or
AD3 mutant synaptotagmin were immobilized on beads and incubated with 1 µM T7-tagged synprint for 2 hr in 2 mM EGTA
(E) or 1 mM Ca2+.
Samples were washed, and bound material was subjected to SDS-PAGE and
immunoblot analysis using an anti-T7 tag antibody and enhanced
chemiluminescence. Twelve percent of the bound material was loaded onto
the gel; total corresponds to 3.5% of the binding reaction.
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For this analysis, we engineered the AD3 mutation into rat
synaptotagmin I (corresponding to amino acid 311 in the rat sequence) to be able to use the biochemical reagents available for this organism.
One possibility to explain the AD3 phenotype is that the
mutant protein fails to interact with the SNARE complex to trigger
Ca2+-induced conformational changes
required for fusion. However, both wild-type and AD3 mutant
synaptotagmin showed comparable binding to preassembled SNARE complexes
(Fig. 7C). Another possibility is that the interaction of
synaptotagmin with the synprint domain of the presynaptic
Ca2+ channel is affected, resulting in an
alteration in Ca2+ entry that could block
the interaction of a second Ca2+ sensor
with the SNARE components. Although the C2B domain is required for the
interaction with synprint, the AD3 mutation does not affect
this interaction (Fig. 7D).
The AD3 mutation lies in close proximity to a set of five
conserved amino acid residues that coordinate divalent cations in a
number of C2 domains (Fig.
8A). These structural
data suggest that the AD3 mutation may impair the putative
Ca2+-binding properties of C2B. To test
this hypothesis, the C2B domain of wild-type and AD3 mutant rat
synaptotagmin were immobilized as GST fusion proteins (the isolated C2B
domain of Drosophila is insoluble and could not be produced
in sufficient quantities to perform this analysis). Fusion proteins
were incubated with increasing concentrations of chymotrypsin in the
presence of EGTA or Ca2+, and the
proteolysis patterns were analyzed by SDS-PAGE. As shown in Figure
8B, the degradation patterns in EGTA versus
Ca2+ were distinct, demonstrating that C2B
undergoes a conformational change after binding
Ca2+. A protease-resistant fragment
accumulated in the presence of Ca2+,
suggesting that Ca2+ binds to and
stabilizes this domain. This result is analogous to the data from
limited proteolysis of the C2A domain (Davletov and Sudhof,
1994). In contrast, a different proteolysis pattern was observed with
the AD3 mutant C2B domain and the presence of Ca2+ has no effect on this pattern. These
results suggest that the AD3 mutation impairs
Ca2+-driven conformational changes in C2B.
We note that the ability of the AD3 mutant to bind SNARE complexes,
AP-2, and the synprint peptide, as described above, demonstrates that
the C2B domain is not misfolded, but rather, exhibits a selective loss
of function.
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Figure 8.
The AD3 mutation
blocks Ca2+-driven conformational changes within the
C2B domain of synaptotagmin and disrupts
Ca2+-triggered oligomerization activity.
A, Crystal structure of the C2B domain of synaptotagmin
III. This image was modified from Sutton et al. (1999); the structure
of the C2B domain of synaptotagmin I has not been reported,
however, all known C2B domains share similar structures. The tyrosine
that is mutated to an asparagine in the AD3 mutant
allele of Drosophila synaptotagmin is indicated, as are
five putative Ca2+ ligands and a single bound
Mg2+ ion. B, The C2B domain of WT
(GST-C2BWT) and AD3 (GST-C2BAD3)
rat synaptotagmin Ib were immobilized as a GST fusion proteins (20 µg/data point) and subjected to limited proteolysis in the presence
of 2 mM EGTA () or 1 mM
Ca2+ (+) at the indicated [chymotrypsin] for 60 min at rt. Samples were boiled in SDS sample buffer, analyzed by SDS
PAGE, and stained with Coomassie blue. C,
Ca2+-triggered synaptotagmin oligomerization is
impaired by the AD3 mutation. Eight micrograms of GST or GST fused to
the cytoplasmic domain of wild-type (sytWT) or AD3
mutant (sytAD3) Drosophila
synaptotagmin was immobilized on beads. Beads were incubated with
1.5 µM soluble WT or AD3 mutant Drosophila
synaptotagmin for 1.5 hr in 150 µl of TBS plus 0.5% Triton X-100 and
either 2 mM EGTA (), 1 mM
Ca2+ (+), or the indicated concentration of
Ca2+. Beads were washed three times with binding
buffer and boiled in SDS sample buffer. Three percent of the soluble
synaptotagmin from the binding assay (left two lanes)
and 25% of the bound material (remaining lanes) were
subjected to SDS-PAGE and visualized by staining with Coomassie blue.
Syt, Cytoplasmic domain of synaptotagmin I. Note: the
asterisk indicates a proteolytic fragment present in
preparations of GST-fused AD3 mutant synaptotagmin. D,
Data from two oligomerization assays (as described in C)
were quantified by densitometry, normalized to the pixel intensity in
the "total" lanes, and plotted versus the free
[Ca2+]. Closed circles, Wild-type
synaptotagmin; open circles, AD3 synaptotagmin.
E, Soluble Drosophila synaptotagmin I (5 µM) was incubated with 30 µg of either
wild-type synaptotagmin IV, AD3 synaptotagmin IV, or synaptotagmin IV
containing a KK to AA substitution (Chapman et al., 1998) at amino
acids 385 and 386. Binding of synaptotagmin I was visualized by Western
analysis with anti-synaptotagmin I DSYT2 antisera (Littleton et al.,
1993).  denotes binding reactions lacking soluble synaptotagmin.
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The C2B domain of synaptotagmin mediates
Ca2+-triggered oligomerization as
previously described (Chapman et al., 1996; Sugita et al., 1996; Desai
et al., 2000). To determine whether this oligomerization activity is
affected by the observed alteration in
Ca2+-dependent conformational changes in
the AD3 C2B domain, oligomerization of the cytoplasmic domains of
wild-type and AD3 Drosophila synaptotagmin were assayed at
increasing concentrations of Ca2+. As
shown in Figure 8, C and D, wild-type
synaptotagmin efficiently bound to immobilized synaptotagmin in
response to Ca2+. In contrast,
Ca2+-triggered oligomerization of the AD3
mutant was reduced to ~40% of the wild-type clustering activity.
These data suggest that the AD3 phenotype results from a
defect in the Ca2+-sensing ability of the
C2B domain, causing a loss of Ca2+-induced
synaptotagmin oligomerization and subsequent SNARE assembly in
vivo. To obtain additional evidence for this model, we tested whether the AD3 Y to N change also alters
Ca2+-dependent oligomerization when
introduced into other isoforms of synaptotagmin. As shown in Figure
8E, when the AD3 change is engineered into
Drosophila synaptotagmin IV,
Ca2+-dependent hetero-oligomerization with
wild-type synaptotagmin I is decreased by >80%. The AD3 mutation
disrupted oligomerization as effectively as the well characterized
K326,327A substitution within the C2B domain of rat synaptotagmin I
(Chapman et al., 1998; Desai et al., 2000; note: this mutation
corresponds to K385, K386 in Drosophila synaptotagmin IV).
Under our assay conditions, oligomerization of wild-type synaptotagmin
was half-maximal at ~6 µM
Ca2+. This value is considerably lower
than the Ca2+ dependence for exocytosis in
retinal bipolar neurons (Heidelberger et al., 1994) but is consistent
with the Ca2+ dependence for exocytosis at
the axosomatic synapse formed by the calyx of Held (Bollman et al.,
2000; Schneggenburger and Neher, 2000).
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DISCUSSION |
Mutations in synaptotagmin I result in profound defects
in neurotransmitter release. In Drosophila, these defects
include a severe reduction in evoked release, an increase in
spontaneous fusion, and delays in the onset of vesicle fusion
(Littleton et al., 1993, 1994; DiAntonio and Schwarz, 1994). In mice,
disruption of the synaptotagmin I gene results in the selective loss of
the rapid synchronous component of exocytosis without affecting the frequency of spontaneous fusion events (Geppert et al., 1994). In
contrast, removal of the t-SNARE syntaxin eliminates both evoked and
spontaneous fusion (Schulze et al., 1995), and temperature-sensitive mutations in syntaxin that block SNARE assembly result in paralysis and
an accumulation of unfused vesicles at release sites (Littleton et al.,
1998). These results are consistent with mounting functional data
indicating that assembly of the SNARE complex is essential for vesicle
fusion (Weber et al., 1998; Chen et al., 1999; Xu et al., 1999). The
more variable effects on secretion in synaptotagmin mutants
are consistent with the possibility that synaptotagmin plays a
regulatory role in promoting evoked release through direct interactions
with the fusion machinery, without being absolutely necessary for
vesicle fusion. Here, we provide evidence for two independent functions
for the C2B domain of synaptotagmin I in synaptic vesicle cycling.
AD1 mutations, which lack the C2B domain, disrupt
synaptotagminAP-2 interactions (Zhang et al., 1994) and lead to a
fourfold reduction in the total number of synaptic vesicles at mutant
terminals during nerve terminal stimulation. These findings are similar
to studies of synaptotagmin mutants in C. elegans (Jorgensen et al., 1995). AD1 terminals do harbor some
synaptic vesicles surrounding active zones and have an elevated
frequency of spontaneous fusions when examined electrophysiologically.
Thus, it is likely that additional endocytotic pathways are capable of
maintaining the smaller pool of vesicles that are recycled in the
immediate vicinity of the active zone. Our observations are consistent
with the distribution of Drosophila AP-2, which is absent
near active zones, but concentrated in the synaptic periphery
(Gonzalez-Gaitan et al., 1996). The lack of a complete loss of
endocytosis in Drosophila synaptotagmin null mutants (Reist et al., 1998) also indicates that in unstimulated synapses, other endocytotic pathways that bypass synaptotagmin I can refill nerve terminals given enough time. One possible candidate for mediating this
endocytotic trafficking in the absence of synaptotagmin I is
synaptotagmin IV, which is also present on synaptotagmin I-containing vesicles and binds to AP-2 as effectively as synaptotagmin I (Li et
al., 1995; Littleton et al., 1999).
In contrast with the endocytotic defect manifested in AD1
mutants, the AD3 mutant phenotype results from a postdocking
defect in synaptic vesicle exocytosis and a failure to assemble SNARE complexes in vivo. Our biochemical analysis revealed that
this mutation disrupts Ca2+-induced
conformational changes in the C2B domain and inhibits Ca2+-induced oligomerization. Whether the
failure to assemble SNARE complexes results from a direct defect in the
acceleration of SNARE formation by synaptotagmin or an alteration in
additional downstream SNARE interactions after synaptotagmin clustering
requires further analysis. Nonetheless, these results indicate that the C2B domain of synaptotagmin must be able to bind
Ca2+, change conformation, and cluster to
trigger coordinated vesicle fusion at nerve terminals. These results
provide biochemically supported genetic evidence that
synaptotagmin is indeed a Ca2+ sensor for
fast exocytosis, as proposed in previous studies (Brose et al., 1992;
DiAntonio and Schwarz, 1994; Geppert et al., 1994; Littleton et al.,
1994). How oligomerization of synaptotagmin leads to vesicle fusion is
unknown, but it is likely to involve direct effects on the SNARE
complex. We suggest that clustering of SNARE complexes by
Ca2+-synaptotagmin leads to rapid
triggering of fusion via formation of SNARE-dependent fusion pores. The
Ca2+-independent interaction of
synaptotagmin with SNAREs would allow these vesicles to remain in a
fusion-ready state and may contribute to the suppression of spontaneous
fusion events (DiAntonio and Schwartz, 1994, 1999; Littleton et al.,
1994). The loss of SNARE clustering activity in synaptotagmin mutants
would prevent rapid, Ca2+-dependent
vesicle fusion as has been observed in syt mutants. The
partial rescue of release in syt mutants at very high
Ca2+ levels could reflect the ability of
other Ca2+ sensors to trigger fusion under
these conditions. These other Ca2+ sensors
might be other members of the synaptotagmin family, which contains
seven members in Drosophila. Biochemical characterization of
the Ca2+-dependent oligomerization
properties of these synaptotagmins might uncover other candidates for
mediating fusion at high Ca2+ concentrations.
Scale models reveal precisely how close complete assembly of the SNARE
complex would bring the vesicle and target membranes (Fig.
9A; Sutton et al., 1998). A
number of experiments indicate that SNARE complexes do not fully
assemble before fusion (Chen et al., 1999; Xu et al., 1999). Thus,
final "zippering" of the complex may not occur until arrival of the
Ca2+ signal that triggers fusion (Chen et
al., 1999). In this case, Ca2+-triggered
oligomerization of synaptotagmin and its association with partially
assembled SNARE complexes could drive final assembly of the base of the
complex (Chapman et al., 1995; Kee and Scheller, 1996; Davis et al.,
1999; Gerona et al., 2000) to accelerate SNARE-mediated membrane
fusion and SDS-resistant SNARE complex assembly (Weber et al., 1998).
The facilitation of SNARE complex assembly, in vitro and
in vivo, reported here supports this model. Furthermore, Ca2+-synaptotagmin triggers the formation
of disulfide-bonded SNARE complex dimers. This finding, in conjunction
with the 1:2 stoichiometry of saturated synaptotagmin-SNARE complexes,
argues that Ca2+-synaptotagmin can bring
at least two SNARE complexes into close proximity. The ability of
synaptotagmin to oligomerize and simultaneously bind SNAREs
suggests that synaptotagmin can cluster multiple SNARE complexes into a
higher ordered assembly that might correspond to the exocytotic fusion
pore. In this light we point out that SNARE complexes alone appear to
cluster only weakly (Fasshauer et al., 1997; Hohl et al., 1998; but see
also Poirier et al., 1998) and that viral fusion proteins are
homotrimers that must assemble into oligomers to form functional fusion
pores (Blumenthal et al., 1996; Danieli et al., 1996). Similarly, a
protein designated EEA1 has recently been reported to cluster
syntaxin-13 into oligomeric structures that are required for endosomal
fusion (McBride et al., 1999). An analogous model for synaptotagmin in
neuronal exocytosis is shown in Figure 9. Synaptotagmin binds to the
membrane proximal region of the SNARE complex while simultaneously
penetrating into membranes (Davis et al., 1999; Gerona et al.,
2000; Fig. 9A). We speculate that these interactions, in
conjunction with the ability of synaptotagmin to oligomerize and to
facilitate SNARE complex assembly (Fig. 9B), leads to
formation of an open fusion pore. In support of this model, we have
recently shown that disrupting C2B-mediated oligomerization of
synaptotagmin I in vitro can lead to postdocking vesicle
fusion defects in cracked PC12 cells (Desai et al., 2000).
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Figure 9.
Model depicting the synaptotagmin-SNARE
complex. A, The core of the SNARE complex, the Habc
domain of syntaxin, the cytoplasmic domain of synaptotagmin, and a
simulated lipid bilayer were modified from Sutton et al. (1998),
Fernandez et al. (1998), Sutton et al. (1999), and Heller et al.
(1993), respectively, and rendered using MOLSCRIPT (Kraulis, 1991). The
regions that interact are indicated with brackets; both
C2 domains of synaptotagmin are required for high affinity binding to
the base of the SNARE complex (Chapman et al., 1995; Davis et al.,
1999; Gerona et al., 2000). The transmembrane anchors of syntaxin,
synaptobrevin, and synaptotagmin were generated by molecular modeling.
B, Model for synaptotagmin-mediated assembly and
clustering of SNARE complexes. One synaptotagmin can interact with two
SNARE complexes; this interaction is depicted as two grooves within
synaptotagmin that bind and assemble SNAREs (which are shown in an
"end view" in which each strand of the four-helix bundle is
depicted as a quarter of a circle).
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Finally, we point out that the N terminus of synaptotagmin contains a
novel Ca2+-independent clustering domain
that may play an important role in both endocytosis (von Poser et al.,
2000) and exocytosis (Bai et al., 2000). We speculate that N-terminal
clustering activity, in conjunction with the weak
Ca2+-independent components of C2B-C2B and
synaptotagmin-SNARE interactions, may serve to poise
synaptotagmin-SNARE complexes for rapid conformational changes,
including the formation of a stable ring-li |
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ABSTRACT
INTRODUCTION
