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Structure of a lipid-bound extended synaptotagmin indicates a role in lipid transfer

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Abstract

Growing evidence suggests that close appositions between the endoplasmic reticulum (ER) and other membranes, including appositions with the plasma membrane (PM), mediate exchange of lipids between these bilayers. The mechanisms of such exchange, which allows lipid transfer independently of vesicular transport, remain poorly understood. The presence of a synaptotagmin-like mitochondrial-lipid-binding protein (SMP) domain, a proposed lipid-binding module, in several proteins localized at membrane contact sites has raised the possibility that such domains may be implicated in lipid transport1,2. SMP-containing proteins include components of the ERMES complex, an ER–mitochondrial tether3, and the extended synaptotagmins (known as tricalbins in yeast), which are ER–PM tethers4,5,6. Here we present at 2.44 Å resolution the crystal structure of a fragment of human extended synaptotagmin 2 (E-SYT2), including an SMP domain and two adjacent C2 domains. The SMP domain has a β-barrel structure like protein modules in the tubular-lipid-binding (TULIP) superfamily. It dimerizes to form an approximately 90-Å-long cylinder traversed by a channel lined entirely with hydrophobic residues, with the two C2A–C2B fragments forming arched structures flexibly linked to the SMP domain. Importantly, structural analysis complemented by mass spectrometry revealed the presence of glycerophospholipids in the E-SYT2 SMP channel, indicating a direct role for E-SYTs in lipid transport. These findings provide strong evidence for a role of SMP-domain-containing proteins in the control of lipid transfer at membrane contact sites and have broad implications beyond the field of ER-to-PM appositions.

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Figure 1: E-SYT2 structure.
Figure 2: E-SYT2 binds lipid-like molecules in the SMP hydrophobic channel.
Figure 3: ER–PM contact site and a model of E-SYT2.

Accession codes

Primary accessions

Protein Data Bank

Data deposits

Coordinates and structure factors have been deposited in the Protein Data Bank under accession number 4P42.

Change history

  • 25 June 2014

    The contrast in Fig. 2d has been improved; the figure was replaced on 25 June 2014.

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Acknowledgements

We are grateful to the staff at NE-CAT beamline 24ID-C at the Advanced Photon Source for assistance with data collection. We thank H. Shen and N. Kumar for discussion. This work was supported by the National Institutes of Health (GM080616 to K.M.R., R37NS36251 and DK082700 to P.D.C.). M.R.W. and F.T. are funded by grants from the National University of Singapore via the Life Sciences Institute (LSI) and the Biochemical Research Council (BMRC-SERC Diag-034). C.M.S. is recipient of an National Science Foundation Graduate Research Fellowship (DGE-1122492). Y.S. was supported by the Uehara Memorial Foundation and the Japanese Society for Promotion of Science.

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Authors and Affiliations

Authors

Contributions

All authors participated in the design of the experiments, data analysis, and manuscript editing. C.M.S., X.W., P.N., F.T. and Y.S. carried out the experiments. F.T., M.R.W., P.D.C. and K.M.R. wrote this manuscript.

Corresponding authors

Correspondence to Pietro De Camilli or Karin M. Reinisch.

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The authors declare no competing financial interests.

Extended data figures and tables

Extended Data Figure 1 E-SYT2 structure analysis.

a, E-SYT2 dimer coloured according to sequence conservation in E-SYT and tricalbin proteins30. C2A–C2B is pulled away from the SMP dimer (as in Fig. 1b, right). Residues at the C2A–C2B–SMP interface are outlined in green or yellow. The interfaces correspond to crystal contacts but are probably not physiologically important as these residues are not well conserved. Note the band of conserved residues at the SMP dimerization interface. b, A cross-section of the SMP barrel, exposing residues at the dimerization interface. The surface is coloured according to sequence conservation. c, E-SYT2 sequence. Secondary structure elements are coloured as in Fig. 1, right, and annotated, as is sequence conservation. Residues at the dimerization interface are underlined bold. Yellow stars and green squares indicate residues at the C2A–C2B–SMP interfaces and match their respective outline colour as in a.

Extended Data Figure 2 Binding of phospholipids by E-SYT2 expressed in bacteria.

ac, Positive-mode non-denaturing MS was carried out on E-SYT2 protein expressed in Escherichia coli. a, b, Spectra are shown before (a) and after (b) deconvolution with the apo (A; MW = 54,414.2 ± 1.5 Da) and the bound species (B and C, with one and two lipid molecules, respectively) represented. The average molecular weight difference between the different forms is shown and could correspond to the molecular mass of a phosphatidylglycerol (PG) (see c). c, When exposed to denaturing conditions but, partly, also in non-denaturing ones, E-SYT2 releases lipids that can be monitored in negative-mode MS. Inset, negative-mode product ion scan on m/z 745.48 released from E-SYT2 protein confirmed the identity of one ligand as PG 16:1/18:1.

Extended Data Figure 3 MS/MS fragmentation spectra in negative mode of two representative abundant glycerophospholipids released from bacterially expressed E-SYT2 under denaturing MS conditions.

a, Fragmentation of PG 16:0/16:1 (719.4648 m/z). b, Fragmentation of PE 16:0/16:1 (688.3992 m/z). All the other possible lipid ligands released from the protein and detected by MS were fragmented in the same way and the identity confirmed.

Extended Data Figure 4 Negative-mode MS spectra of glycerophospholipids released from E-SYT2, expressed in mammalian cells, under denaturing MS conditions.

The four most abundant glycerophospholipid classes (PE, PC, PS, PI) are shown. The acyl chain compositions of the most represented molecular species in each class are also shown.

Extended Data Figure 5 Positive- and negative-mode MS/MS fragmentation spectra of representative abundant glycerophospholipids released from E-SYT2 expressed in mammalian cells, under denaturing MS conditions.

a, Fragmentation of PC 16:0/18:1 (760.45 m/z). b, Fragmentation of PE 18:1/18:1 (742.53 m/z). c, Fragmentation of PI 18:1/18:1 (861.58 m/z). d, Fragmentation of PS 18:1/18:1 (786.53 m/z).

Extended Data Figure 6 Deconvoluted spectra of E-SYT2.

a, b, Deconvoluted spectra of the monomeric E-SYT2 form indicated as P (a) and of the lipid-bound dimeric form indicated as P2L4 (b) in Fig. 2c. c, Positive-mode ESI-MS spectrum in denaturing conditions (5% acetic acid in 20 mM ammonium acetate) of E-SYT2 expressed in mammalian cells. To exclude the possibility that the dimer-associated MS peaks in Fig. 2c and ED6b could be an artefact generated during ionization, the sample was denatured in acidic conditions. Only a charge distribution around +33 was detected (indicated as P), probably representing a more open and denatured monomeric apo protein conformation, confirmed by the deconvoluted spectrum (not shown). In these experimental conditions, no peaks corresponding to dimeric structures could be detected.

Extended Data Figure 7 Lipid competition assay.

NBD–PE-preloaded SMP–C2A–C2B mixed with methanol (control), phosphatidylcholine (PC), sphingomyelin (SM), ceramide (CER) or cholesterol (CH). Quantification of the five trials is shown. Standard deviation (STDEV) is indicated.

Extended Data Table 1 Data collection and refinement statistics
Extended Data Table 2 Phospholipids present as ligands in E-SYT2 purified from E. coli and observed after denaturing MS in negative-ionization mode
Extended Data Table 3 A targeted quantitative lipidomic approach based on LC-MS/MS was used to quantify the different lipid species released by E-SYT2 after organic solvent extraction

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Schauder, C., Wu, X., Saheki, Y. et al. Structure of a lipid-bound extended synaptotagmin indicates a role in lipid transfer. Nature 510, 552–555 (2014). https://doi.org/10.1038/nature13269

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