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Dynamic binding mode of a Synaptotagmin-1–SNARE complex in solution

Nature Structural & Molecular Biology volume 22, pages 555–564 (2015)Cite this article

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Abstract

Rapid neurotransmitter release depends on the Ca2+ sensor Synaptotagmin-1 (Syt1) and the SNARE complex formed by synaptobrevin, syntaxin-1 and SNAP-25. How Syt1 triggers release has been unclear, partly because elucidating high-resolution structures of Syt1–SNARE complexes has been challenging. An NMR approach based on lanthanide-induced pseudocontact shifts now reveals a dynamic binding mode in which basic residues in the concave side of the Syt1 C2B-domain β-sandwich interact with a polyacidic region of the SNARE complex formed by syntaxin-1 and SNAP-25. The physiological relevance of this dynamic structural model is supported by mutations in basic residues of Syt1 that markedly impair SNARE-complex binding in vitro and Syt1 function in neurons. Mutations with milder effects on binding have correspondingly milder effects on Syt1 function. Our results support a model whereby dynamic interaction facilitates cooperation between Syt1 and the SNAREs in inducing membrane fusion.

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Figure 1: A polybasic region of the Syt1 C2B domain binds to the SNARE complex.
Figure 2: Δχ tensors defined by the PCSs induced in the SNARE complex by Dy3+-C2 labels on residue 166 or 41 of SNAP-25.
Figure 3: PCSs induced by SC166Dy and SC41Dy on the Syt1 C2B domain.
Figure 4: Analysis of the C2B–SNARE complex by MD simulations.
Figure 5: The Syt1 C2B–SNARE-complex binding mode.
Figure 6: Mutagenesis verifies the C2B–SNARE-complex binding mode.
Figure 7: Disruption of Syt1 function correlates with impairment of Syt1–SNARE-complex binding.
Figure 8: Disruption of Syt1–SNARE-complex binding correlates with impairment of the Ca2+-triggered step of release.

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Acknowledgements

We thank L. Kay for fruitful discussions and F. Peters (Max Planck Institute for Biophysical Chemistry) for generously providing a sample for Cys-Ph-TAHA–labeling. The DD2 console of one of the Agilent 600-MHz NMR spectrometers used for the research presented here was purchased with shared instrumentation grant S10RR026461 from the US National Institutes of Health (to M.K. Rosen; University of Texas Southwestern Medical Center). The authors acknowledge the Texas Advanced Computing Center at the University of Texas at Austin for providing high-performance computer resources that have contributed to the research results reported within this paper. This work was supported by Welch Foundation grant I-1304 (to J.R.), Australian Research Council (ARC) Discovery Grant DP150100383 (to B.G.), ARC Future Fellowship FT130100838 (to B.G.), Swiss National Science Foundation grant 200021_130263 (to D.H.) and US National Institutes of Health grants K99NS087086 (to T.B.) and NS040944 (to J.R.).

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Authors and Affiliations

  1. Department of Biophysics, University of Texas Southwestern Medical Center, Dallas, Texas, USA

    Kyle D Brewer, Jin Liu, Amy Zhou, Junjie Xu, Alpay B Seven, Eric A Prinslow, Rashmi Voleti, Diana R Tomchick & Josep Rizo

  2. Department of Biochemistry, University of Texas Southwestern Medical Center, Dallas, Texas, USA

    Kyle D Brewer, Jin Liu, Amy Zhou, Junjie Xu, Alpay B Seven, Eric A Prinslow, Rashmi Voleti, Diana R Tomchick & Josep Rizo

  3. Department of Pharmacology, University of Texas Southwestern Medical Center, Dallas, Texas, USA

    Kyle D Brewer, Jin Liu, Amy Zhou, Junjie Xu, Alpay B Seven, Eric A Prinslow, Rashmi Voleti & Josep Rizo

  4. Department of Molecular and Cellular Physiology, Stanford University Medical School, Stanford, California, USA

    Taulant Bacaj, Peng Zhou & Thomas C Südhof

  5. Howard Hughes Medical Institute, Stanford University Medical School, Stanford, California, USA

    Taulant Bacaj, Peng Zhou & Thomas C Südhof

  6. Institute for Research in Biomedicine, Bellinzona, Switzerland

    Andrea Cavalli

  7. Department of Chemistry, University of Cambridge, Cambridge, UK

    Andrea Cavalli, Carlo Camilloni & Michele Vendruscolo

  8. Monash Institute of Pharmaceutical Sciences, Monash University, Parkville, Victoria, Australia

    James D Swarbrick, Nicholas Barlow & Bim Graham

  9. Department of Chemistry, Southern Methodist University, Dallas, Texas, USA

    Jin Liu

  10. Department of Chemistry, University of Basel, Basel, Switzerland

    Daniel Häussinger

  11. Bijvoet Center for Biomolecular Research, Faculty of Science, Utrecht University, Utrecht, the Netherlands

    Alexandre M J J Bonvin

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  1. Kyle D Brewer
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Contributions

K.D.B., T.B., J.D.S., A.Z., P.Z., N.B., J.X., A.B.S. and E.A.P. performed experiments and analyzed data. A.C., C.C., J.L. and R.V. performed computational analyses. D.H., A.M.J.J.B., D.R.T., M.V., B.G. and T.C.S. designed experiments and analyzed data. J.R. analyzed data and wrote the manuscript with input from all coauthors.

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Correspondence to Josep Rizo.

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Integrated supplementary information

Supplementary Figure 1 Effects of Syt1 C2AB on the 1H-15N HSQC spectra of the SNARE complex.

(a) Diagram depicting the proteins involved in neurotransmitter release discussed in this study. The SNARE complex is depicted as a four-helix bundle with Synaptobrevin in red, Syntaxin-1 in yellow and SNAP-25 in green and blue. The N-terminal region of Syntaxin-1 is not shown for simplicity. The SNARE complex is shown partially formed and full zippering of the Synaptobrevin SNARE motif to complete the four-helix bundle is believed to be key for membrane fusion. Syt1 is shown in orange with the C2A and C2B domains labeled as A and B, respectively, while CpxI is shown in gray (central helix) and pink (accessory helix). The location of CpxI is based on the crystal structure of the CpxI(26-83)-SNARE complex1. The position of Syt1 is arbitrary and is the subject of this study. Only one molecule each of Syt1 and of CpxI are shown for simplicity but each SNARE complex is expected to have one molecule of Syt1 and one of CpxI bound. (b-d) 1H-15N TROSY HSQC spectra of 40 μM SNARE complex samples 2H,15N-labeled at the syntaxin-1 (b), synaptobrevin (c), SNAP-25 N-terminal (SNN) (d) and SNAP-25 C-terminal (SNC) (e) SNARE motifs in the absence (black contours) and presence (red contours) of 20 μM C2AB.

Supplementary Figure 2 Evaluations of PCSs induced by SC166Dy and SC41Dy.

(a-e) Leu,Val region of 1H-13C HMQC spectra of 30 μM 15N,2H-ILV-13CH3-labeled C2AB (a,d) or C2B (b,c,e) in the presence of SC166Dy (a-c) or SC41Dy (d,e) before (red contours) or after (black contours) removal of the tag by reduction. The concentration of SC166Dy or SC41Dy was 20 μM in (a,d) and 30 μM in (b,c,e). In (c), blue contours show a spectrum of 15N,2H-ILV-13CH3-C2B bound to SC166Dy in the presence of CpxI. Note that in panels (a,c) the contour levels were chosen to allow visualization of some of the PCSs and at the same time avoid the overcrowding observed at lower contour levels. Some of the cross-peaks from the red spectra are not observable at these contour levels due to the broadening caused by the lanthanide tag, but most cross-peaks are observable at lower contour levels and reveal PCSs that are parallel to those observed for C2B in panels (b,e). (f,g) Ile region of 1H-13C HMQC spectra of 30 μM SC166Dy (f) and SC41Dy (g) containing 15N,2H-ILV-13CH3-syntaxin-1 acquired after removal of the tag (black contours) or before removal of the tag in the absence (red contours) or presence (blue contours) of 30 μM C2B domain bearing the R398Q R399Q mutation (C2BRR).

Supplementary Figure 3 Comparing the SC166 and C2B166 tensors.

(a) Ribbon diagram of the C2B domain and isosurfaces representing regions with positive (blue) and negative (red) PCSs, contoured at ± 0.8 ppm with the C2B166 tensor. The values for Δχax and Δχrh (10−32 m3) are 32.0 and 18.4, respectively. (b) Ribbon diagram of the C2B domain and the SNARE complex with isosurfaces representing regions with positive (blue) and negative (red) PCSs, contoured at ± 0.8 ppm with the SC166 tensor. This is the same tensor illustrated in Fig. 2c but rotated to allow comparison with the C2B166 tensor shown in panel A. The position of C2B was derived by superimposing the centers of the SC166 and C2B166 tensors. (c) Correlation between experimental C2B PCSs caused by SC166Dy and PCSs calculated with the C2B166 tensor illustrated in panel (A). (d) Correlation between experimental C2B PCSs caused by SC166Dy and PCSs calculated with the SC166 tensor in the model resulting after superimposing the C2B166 and SC166 tensors (illustrated in panel b). (e) Correlation between experimental C2B PCSs caused by SC166Dy and PCSs calculated with the SC166 tensor in the model with C2B rotated, illustrated with C2B in orange in panel (f). (f) Ribbon diagram of the SNARE complex with isosurfaces representing regions with positive (blue) and negative (red) PCSs, contoured at ± 0.8 ppm with the SC166 tensor, showing the positions of C2B after superimposing the centers of the SC166 and C2B166 tensors (cyan ribbon) and after rotating C2B around the vertical axis (defined here by the negative lobes of the SC166 tensor) to make contact with the SNARE complex (orange ribbon). CpxI(26-83) is also shown (in pink) based on superimposing the models with the crystal structure of the CpxI(26-83)-SNARE complex (PDB code 1KIL) to illustrate that C2B would have steric clashes with CpxI(26-83) in the position of the orange ribbon. (g) Ribbon diagram of the SNARE complex with isosurfaces representing regions with positive (blue) and negative (red) PCSs, contoured at ± 0.8 ppm with the SC166 tensor, showing in orange the rotated position of C2B from panel (f) and in gray the position of C2B from the 166 manual model of Fig. 3e. In (a,b,f,g), the tensor center is indicated with a black sphere. Note that in (f) the rotated C2B (orange) is at the same distance from the tensor center as in the original model obtained by superimposing the centers of the C2B166 and SC166 tensors (cyan C2B), and that in (g) C2B from the 166 manual model (gray) is considerably closer to the tensor center than the rotated C2B (orange).

Supplementary Figure 4 Evaluation of selected models of the C2B–SNARE complex.

(a,c) Correlations between experimental C2B PCSs induced by SC166Dy and PCSs calculated with the SC166 tensor and the 166 HADDOCK model (a) or the 166 MD model (c). Correlation coefficients (r) and slopes (m) are indicated. (b,d) Representations of the 166 HADDOCK model (b) and 166 MD model (d) with the SNAREs and C2B shown as semi-transparent ribbons and the positive-negative lobes of the SC166 tensor represented by isosurfaces as in Fig. 2c. The C2B PCSs induced by SC166Dy are illustrated to visualize how well the positive-negative patterns match with the positive-negative lobes of the SC166 tensor. Amide hydrogens and methyl carbons are shown as spheres and color-coded according to the measured PCSs (dark blue, > 0.06 ppm; blue, 0.04 to 0.06 ppm; cyan, 0.02 to 0.04 ppm; pale cyan, 0.008 to 0.02 ppm; red, −0.04 to −0.06 ppm; salmon, −0.02 to −0.04 ppm; light pink, −0.008 to −0.02 ppm). Ca2+ ions are represented by yellow spheres.

Supplementary Figure 5 The 41 MD model.

Ribbon diagram of the 41 MD model with C2B shown in orange and Ca2+ ions represented by cyan spheres. Stick models show the side chains of basic (blue) and acidic (red) residues. Basic side chains from the polybasic strand and the concave side of C2B are labeled. N and C represent the N- and C-termini of the SNARE complex, respectively.

Supplementary Figure 6 Mutagenesis verifies the binding mode of the C2B–SNARE complex.

(a,b) Plots of normalized intensities of the SMRs in 1D 13C-edited 1H-NMR spectra of 3 μM WT or mutant 13C-C2AB as a function of SNARE complex concentration. C2AB mutants contained single substitutions in basic residues as indicated and color-coded. The data were acquired in 25 mM Tris (pH 7.4), 125 mM NaCl and 1 mM CaCl2, and were fitted to a single-site binding model2. (c,d) Bar diagrams illustrating the Kds derived from fitting the data of Figs. 5b,c to single-site binding models. The Kd values should be interpreted with caution because we did not use SNARE complex concentrations beyond 20 μM to minimize contributions from weaker binding modes and, as a consequence, the titrations were far from reaching saturation for the mutants with stronger effects on binding. Because of the uncertainty in the limiting intensities at infinite SNARE complex concentrations for the mutants that bind to the SNARE complex more weakly, this limiting value was forced to be 0.555 times the intensity at 0 SNARE complex concentration. The 0.555 factor was derived from averaging the ratios between intensities at 0 and infinite SNARE complex concentration in the fits obtained for WT C2AB. Bars show average Kds calculated from two independent experiments, and error bars show standard deviations.

Supplementary Figure 7 Overexpression levels of WT and double-mutant Syt1 used in rescue experiments.

(a) Sample Western blots illustrating the overexpression levels of the WT and double mutant Syt1 in the rescue experiments. (b) Quantification of the protein overexpression levels in three different independent experiments.

Supplementary Figure 8 Model of how Syt1 and the SNAREs trigger neurotransmitter release in an interplay with CpxI.

(a) Model of a primed state with a partially assembled SNARE complex where the C-terminus of the synaptobrevin SNARE motif remains flexible. CpxI(26-83) is shown according to the crystal structure of the CpxI(26-83)-SNARE complex (PDB accession code 1KIL). The position of Ca2+-free C2B corresponds to that of the 166 MD model. R398 and R399 at the bottom of C2B are shown as dark blue spheres and bound to the plasma membrane. (b) Model of a fused state with a fully assembled SNARE complex after Ca2+ influx, with C2B bound to the SNARE complex and to both membranes. Ca2+ ions are represented by yellow spheres. Key aspects of this model are: i) before Ca2+ influx, Syt1 binds to a partially assembled SNARE complex through the concave, basic side of the C2B domain and, in this primed state, the CpxI accessory helix (pink) repels the vesicle membrane, hindering membrane fusion; ii) Ca2+ binding to C2B induces binding to the vesicle membrane, forcing the inhibitory accessory helix to melt away; iii) simultaneous binding of R398-R399 of C2B to the plasma membrane and of the Ca2+-binding loops to the vesicle membrane forces the two membranes together which, together with full zippering of the SNARE complex, induces membrane fusion; iv) these actions of C2B may require re-orientation with respect to the SNARE complex, which would be facilitated by the dynamic nature of the C2B-SNARE complex interactions. This mechanism is consistent with the 166 MD model and 41 manual model, which we take as representatives of the preferred orientations defined by the two PCS datasets, as well as with many of the structures visited during our MD simulations. Hence, the proposed mechanism relies on the overall dynamic binding mode rather than on a biased choice of a particular model.

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Brewer, K., Bacaj, T., Cavalli, A. et al. Dynamic binding mode of a Synaptotagmin-1–SNARE complex in solution. Nat Struct Mol Biol 22, 555–564 (2015). https://doi.org/10.1038/nsmb.3035

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