Structural and Functional Study of SNARE Machinery in Neurotransmitter Release Shuang Song Iowa State University

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2013 Structural and functional study of SNARE machinery in neurotransmitter release Shuang Song Iowa State University

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Structural and functional study of SNARE machinery in neurotransmitter release

Shuang Song

A dissertation submitted to the graduate faculty

in partial fulfillment of the requirements for the degree of

DOCTOR OF PHILOSOPHY

Major: Molecular, Cellular, and Developmental Biology

Program of Study Committee:

Yeon-Kyun Shin, Major Professor

Linda Ambrosio

Edward Yu

Alan DiSpirito

Diane Bassham

Iowa State University

Ames, Iowa

2013

TABLE OF CONTENTS

ABSTRACT ...... iv

CHAPTER 1: GENERAL INTRODUCTION ...... 1 1.1 Membrane fusion and neurotransmitter release ...... 1 1.2 SNARE proteins ...... 2 1.3 Structure of SNARE proteins and SNARE complex ...... 3 1.4 SNARE assembly ...... 5 1.5 Mechanics of SNARE-mediated membrane fusion ...... 7 1.6 Regulators of SNARE-mediated membrane fusion ...... 8 1.7 Study methods ...... 12 1.8 References ...... 16 1.8 Figures and Captions ...... 24

CHAPTER 2: ACCESSORY α-HELIX OF COMPLEXIN I CAN DISPLACE VAMP2 LOCALLY IN THE COMPLEXIN-SNARE QUATERNARY COMPLEX * ...... 38 2.1 Abstract ...... 38 2.2 Introduction ...... 39 2.3 Results ...... 41 2.4 Discussion ...... 46 2.5 Materials and Methods ...... 49 2.6 References ...... 52 2.7 Figures and Captions ...... 56 2.8 Supplementary Data ...... 61

CHAPTER 3: LINKER REGION OF NEURONAL SNARE COMPLEX FORMS INDEPENDENT α-HELICES WHEN SNARE CORE ASSEMBLY IS COMPLETED* ...... 62 3.1 Abstract ...... 62 3.2 Introduction ...... 63 3.3 Results ...... 65 3.4 Discussion ...... 72 3.5 Materials and Methods ...... 75 iii

2.8 Reference ...... 78 2.7 Figures and Captions ...... 84

CHAPTER 4: GENERAL CONCLUSIONS ...... 95 5.1 Summary ...... 95 5.2 Future plan ...... 97 5.3 Reference ...... 99

ACKNOWLEDGEMENTS………………………………………………………………………95

ABSTRACT

Synaptic neurotransmitter release is the most critical communication process for the connections between neurons and between neurons and target cells. SNAREs (soluble N- ethylmaleimide sensitive factor attachment protein receptors) are believed to be highly involved in docking and fusion of synaptic vesicles to the pre-synaptic plasma membrane.

In vivo, synaptic vesicle exocytosis is a regulated and extremely rapid process. Numerous regulatory proteins are also required to achieve the fast speed and Ca 2+ dependency of synaptic neurotransmitter release.

Our research mainly focuses on investigating the mechanism and protein structural basis of SNARE mediated membrane fusion. Site-directed spin labeling (SDSL) and electron paramagnetic resonance (EPR) spectroscopy are powerful tools to study the structure and membrane topology of membrane proteins in lipid bilayer. We use these advanced techniques to probe the structure of SNARE proteins and the important regulators in neurotransmitter release.

Neurotransmitter release takes place on a much shorter time scale compared to other kind of exocytosis and this fast release is accurately coupled with Ca 2+ signaling. In this dissertation, we have revealed some structure details that contribute to understand the underlying mechanism of this fine-tuning process. Firstly, the structural analysis of complexin/SNARE complex discloses a balance between different interaction patterns of complexin and SNARE. The exchange of these interaction patterns might switch the v complexin function between stimulation and inhibition during different fusion steps.

Then we further investigate the linker region structure of SNARE complex in different zippering stages and obtain detailed information about the conformational changes of this linker region during SNARE zippering process. The results we have got from these studies may shed light on the molecular basis of the efficient and precise control of SNARE machinery on neurontransmitter release. 1

CHAPTER 1: GENERAL INTRODUCTION

1.1 Membrane fusion and neurotransmitter release

The eukaryotic cytosol is partitioned into distinct compartments in order to implement the diverse biochemical processes under their favorable conditions. The exchange of material between different organelles is achieved by the trafficking of vesicles 1.

Transport vesicles containing proteins and lipids bud from a donor organelle and then target, dock and fuse with an acceptor organelle to accomplish the transportation of substance 2. Exocytosis is defined as the fusion of a transport vesicle with the plasma membrane. Some proteins are secreted constantly and no stimulation is required which is referred to as constitutive exocytosis. Others are secreted only under certain circumstances and are triggered by some second messengers such as Ca 2+3. In the nervous system, neurotransmitter release of the pre-synaptic membrane is the first step of chemical synaptic transmission and is also a regulated exocytosis process.

The chemical synaptic transmission is a process that neurons use to communicate with each other and with their target cells through transportation of chemical messengers called neurotransmitters 4. Firstly, synaptic vesicles carrying neurotransmitters are secreted by the endosome of neurons. The filled vesicles then accumulate to form the vesicle cluster and dock at the active zone of the pre-synaptic membrane. These vesicles need to undergo a priming stage to be prepared for a fast fusion to happen within ~2ms

2 once the action potential arrives and Ca 2+ flows in ( Figure 1 ). The membrane fusion between the vesicle and plasma membrane requires proteins specialized to act as a fusion machinery to conquer the repulsive ionic forces and dissipate the hydration between the two opposite bilayers 5. The SNARE family of proteins have been identified as the conserved core protein apparatus for all intracellular membrane fusion events from yeast to human organism 2; 6; 7; 8; 9 .

1.2 SNARE proteins

SNARE (soluble N-ethyl-maleimide-sensitive factor attachment protein receptors) has been proposed as the minimal fusion machinery in almost all intracellular membrane fusion. In mammalian cells, SNAREs from different cellular compartments can be categorized based on sequence homology and domain structure as members of syntaxin

(STX1) 10 , SNAP-25(25 kDa synaptosome-associated protein) 11 and VAMP (vesicle- associated membrane protein, also named as synaptobrevin) 1; 12 . They are specifically involved in different vesicular trafficking steps. Because syntaxin and SNAP-25 reside on the target membrane (plasma membrane), they are also called target membrane (t-)

SNARE. Both syntaxin and VAMP are anchored to the membrane by a carboxy-terminal transmembrane domain, whereas SNAP-25 is attached to the membrane surface by palmitoylation 8.

Our research mainly focuses on SNAREs particularly involved in synaptic vesicle exocytosis. These SNAREs were established to be critical for synaptic neurotransmission by studies revealing that they are the targets of CNT proteases 6; 13; 14 . More experiments have shown the importance of SNAREs in gene knockout animal models such as knockout mice lacking VAMP show decreased synaptic vesicle fusion rate 15 and knockout mice lacking SNAP-25 is particularly deficient in Ca 2+ -triggered neurotransmitter release 16 .

1.3 Structure of SNARE proteins and SNARE complex

SNARE proteins are structurally characterized by its evolutionarily conserved domain named as the SNARE motif, which is a homologous ~60-70 amino acids sequence containing about 8 heptad repeats suggesting a potential α-helical structure. Most of the

SNAREs contain a variable N-terminal sequence as well as a C-terminal transmembrane domain. Syntaxin has an independently folded domain at the N-terminus known as the

Habc domain. This 120-residue N-terminal domain is conserved in plasma membrane syntaxins only which may indicate a specific role in exocytosis. It has a three-helix bundle structure and potentially regulates the SNARE complex assembly 17; 18 (Figure 2 ).

SNAP-25 contains two SNARE motifs linked by a loop structure of 37 residuals with 4 conservative cysteines, which is probably palmitoylated in vivo and used as a membrane anchor 19 . Both of the transmembrane proteins syntaxin and VAMP have a ~10 amino

4 acids linker region preceding the transmembrane domain. Solution NMR studies and circular dichroism (CD) experiments have shown that the cytoplasmic domains of isolated VAMP and SNAP-25 are unfolded 20; 21; 22 , whereas those of syntaxin are partially folded 21 . In the following part we will specifically discuss the neuronal SNARE proteins which have been best characterized and are our major research objects. Therefore I refer to the neuronal SNARE isoforms syntaxin 1A as simply syntaxin, VAMP2 as VAMP, and SNAP-25A as SNAP-25 without further specification. A scheme chart of domain structure of these isoforms is shown in Figure 3 .

During membrane fusion, vesicle-associated (v-) SNARE and target membrane (t-)

SNARE form a SNARE complex which brings the two opposite membranes together. A parallel four-stranded coiled-coil model for the structure of the core SNARE complex omitting its N-terminal and C-terminal regions was first revealed by spin labeling electron paramagnetic resonance (EPR) 23 and determined by crystal structure in 1998 24 .

Since the transmembrane domains are not necessary for the SNARE core assembly, many early structural studies were carried out with only the soluble parts of SNARE proteins and this core complex has similar biophysical properties as the full-length complex 25; 26; 27 .

Three SNARE motifs from t-SNAREs (two from SNAP-25 and one from syntaxin) and one from v-SNARE can spontaneously assemble into a four-helix bundle which is composed of 16 hydrophobic layers ( Figure 4 ). This synaptic core complex is highly stable and is even resistant to SDS and heat up to 90ºC 26; 28; 29 . An ionic “0” layer is present in the middle of the core complex which is made up of one Arg residue

(contributed by VAMP) and three Gln residues (contributed by syntaxin and SNAP-25)

(Figure 4c). These residues are very conservative throughout the entire SNARE super family. The surface of the bundle is highly grooved and is rich of distinct hydrophilic, hydrophobic and charged regions which might facilitate the binding of potential regulatory factors 24 .

1.4 SNARE assembly

As mentioned before, the individual SNARE core domains are mostly unstructured 20; 21;

22 . Therefore upon complex formation, dramatic structural changes occur to accomplish a disorder-to-order transition. Studies of the yeast SNARE complex has also shown similar phenomenon 21 . Firstly, the N-terminal regulatory Habc domain of syntaxin is detached from its SNARE core probably with the assistance of SM protein, Munc18-130 and switches the SNARE protein from a closed-form to an open-form 31; 32 (Figure 5).

Secondly, binary complex can be formed by syntaxin and SNAP-25 in a mixture of 1:1 and 2:1 complexes 20 . Studies by spin labeling EPR spectroscopy suggested that a 2:1 binary SNARE complex is highly coiled 33 which is somewhat contradictory to a previous study which proposes a disordered midsection 34 and an uncoiled C-terminal region 35 .

However in both of these cases, the 2:1 form of the syntaxin·SNAP-25 complex adopts an identical parallel four-helix bundle structure as the ternary SNARE complex (Figure

6) . This 2:1 syntaxin·SNAP-25 complex might suggest a dead-end status in which a ternary complex is formed and the excess of syntaxin prevents VAMP to bind. So this

6 complex is not supposed to be functional for membrane fusion. Unfortunately, the 1:1 syntaxin·SNARE complex structure is not accessible due to its transition characteristic.

Thirdly, once the v-SNARE is involved, formation of ternary SNARE complex between binary complex and VAMP starts along with induction of more α-helix structures in previously unstructured regions of the SNARE motifs. Before the membrane fusion is completed, the intermediate state of the SNARE complex is referred to as the

“trans -SNARE complex” (“SNAREpin”)9. Again direct structure evidence is missing because of the difficulty of catching a trans -SNARE complex in the middle of fusion.

The SNARE complex formation is widely believed to initiate from the N-terminal region and zipper towards the C-terminal membrane-proximal region 36 . However, we are still lack of hard evidence to support that hypothesis. When fusion is accomplished, the fully zippered SNARE complex ended up residing on the same bilayer to form the “cis -

SNARE complex” which represents a post-fusion state ( Figure 7 ). The core SNARE complex whose structure was described before is a representation of the post-fusion cis -

SNARE complex. This cis-SNARE complex will be disassembled to join another cycle which requires the involvement of ATPases: NSF ( N-ethylmaleimide sensitive factor) and α-SNAP (soluble NSF attachment protein) 37; 38

The parallel arrangement of SNARE motifs spatially brings the two transmembrane domains of syntaxin and VAMP together along with the corresponding juxtaposed membranes into close proximity. Kinetically, the high stability of the cis -SNARE complex indicates that the assembly of SNARE complex might be responsible for generating the energy that is required to overcome the energy barrier for membrane

7 fusion 28; 39 . Single SNARE complex might not be enough for providing the energy required in fusion pore opening. Several SNARE complexes may be involved in a single fusion event, by forming a ring structure around the site of fusion, the bulk energy formed by these complexes together will easily exceed the energy barrier for membrane fusion 40 . People believe that a limited number of assembling SNARE complexes may remain the intermediate trans-SNARE state without inducing fusion (Figure 8, right) until the accumulation of many SNARE complexes is accomplished which will cause sufficient strain on the membranes to force the merger 41 . However, there is also literature showing that one SNARE complex is sufficient for membrane fusion 42 .

1.5 Mechanics of SNARE-mediated membrane fusion

When the SNARE complex manages to bring the two opposite membranes into a close distance, the two membranes begin to merge, resulting in the opening of a “fusion pore”.

As the fusion pore expands, the two membranes gradually form a uniform bilayer and leads to full content mixing. The pore was once hypothesized to be proteinaceous, composed of proteins that span the two bilayers 43; 44; 45 . However a scenario which has become more commonly accepted is that the active pore harbors predominantly lipidic intermediates such as the hemifusion state, while the outer membranes are merged and the inner leaflet remains intact 46; 47 . This hemifusion state has been suggested by either the observation of lipid mixing without content mixing 48; 49 or by the difference in kinetics between outer- and inner-leaflet mixing 50; 51 . And direct evidence for the

8 existence of this hemifusion state is also provided by single-vesicle fusion assay 52 . The formation of the lipid fusion pore requires a protein scaffold to generate the force needed to conquer the high membrane tension and curvature. The assembly of SNARE complex is supposed to act like this protein scaffold by initiating inward and lateral movement in both membranes and exerting the force from the core complex formation to the membrane anchors through a linker region in between53 . A similar mechanism of membrane fusion has been discovered in viral system54 . Although SNAREs alone are believed to be the minimal fusion machinery, fusion of phospholipid bilayer vesicles reconstituted with only SNARE proteins has been found to be very slow. It may require hours before reaching the saturation state. In the nervous system, synaptic exocytosis is an extremely fast (~2ms) and tightly regulated process as mentioned in Chapter 1.1.

Other than the residual spontaneous neurotransmitter release which can happen even in the absence of VAMP or SNAP-25, for the most part, neurotransmitter release is Ca 2+ - triggered. It can be further classified into fast Ca 2+ -triggered synchronous release and slower Ca 2+ -triggered asynchronous release 4; 55 which both depend on SNARE proteins.

To achieve the exquisite Ca 2+ sensitivity and fast speed, neurons need a more complex protein apparatus.

1.6 Regulators of SNARE-mediated membrane fusion

There are many other SNARE-interacting proteins involved in the Ca 2+ -triggered neurotransmitter release. They are not only responsible for expediting and synchronizing

9 the membrane fusion during the fusion process, but also have been shown to possibly promote a 1:1 binary t-SNARE complex formation over its 2:1 dead-end counterpart 56 .

Some of them bind with the SNARE proteins directly and others function indirectly as an upstream or downstream regulator. In this thesis study, we just focus on two of these regulatory protein families, complexin (Cpx) and synaptotagmin (Stg).

1.6.1 Complexin

Complexins are a family of small, highly charged neuronal proteins in mammals.

There are four isoforms of complexin. Complexin I and complexin II are 86% identical in rat. They also function redundantly, whereas complexin I is specifically localized in the central nervous system while complexin II is also found in non-neuronal tissues 57 .

Complexin III and complexin IV show only limited homology with complexin I or complexin II, they have been found to function in retinal neurotransmitter release 58 .

Complexin I will be referred to as simply complexin in the following part.

Complexin is a soluble protein of molecular weight ~15kDa which can bind with

7 -1 -1 SNARE complex rapidly (5×10 M S ) and with high affinity (Km=10nM).

Investigations about complexin have shown controversial results these years.

Physiological data reveals both stimulatory and inhibitory functions in different animal models 59; 60; 61 . Deletion of complexin I and complexin II in mice abolishes synchronous release while asynchronous release remains intact, which is similar to, but less severe

10 than the synaptotagmin knock-out phenotype 60 . This observation suggests an essential positive role of complexin in Ca 2+ -triggered neurotransmitter release as well as potential coupled function of complexin and synaptotagmin 62 . On the other hand, the knock-out experiments in drosophila melanogaster reveal a highly inhibitory effect of complexin 63 .

A dual function model of complexin has also been proposed on the basis of both physiological experiments 64 and biophysical data 59 . Application of single-vesicle fluorescence fusion assay enables direct observation of the two-faceted functions of complexin in a reconstituted bilayer system. Yoon and colleagues have been able to observe an inhibitory function of complexin in preventing SNARE complex formation at the vesicle docking stage and a stimulatory function in stabilizing the assembled SNARE complex in response to Ca 2+ at the fusion step with different requirements of concentration. A hypothetical model of the complexin dual function during the entire fusion procedure is shown in Figure 10.

Structurally, complexin binds ternary SNARE complex through a central α-helical domain (residues 48-70). It also possesses an accessory α-helix (residues 26-47) contiguous to the central helix and additional N- and C-terminal sequences with unknown structure and functions. Quantitative structure-function analysis of complexin I in mouse suggests that distinct domains may act differently in regulating synaptic exocytosis 64 . It shows that the accessory α-helix may have an inhibitory function on neurotransmitter release and the N-terminal domain may perform a stimulatory function. But the exact molecular mechanism remains elusive. Diverse interaction modes have been discovered between complexin and SNARE complex 65 . Crystal structures have been resolved for

11 both an additive binding mode 66 which represents a post-fusion state ( Figure 11 ) and is considered to be in favor of the stimulatory role and a cross-linked zigzag model 66 which may mimic a intermediate state and is believed to perform the inhibitory role.

1.6.2 Synaptotagmin

Syntaptotagmin is a transmembrane protein with two C2 domains (C2A and C2B), which have been specialized as the Ca 2+ binding sites. It has been believed to be the Ca 2+ sensor in the neuron. Unlike complexin, synaptotagmin has been found in several isoforms in both neurons and non-neuronal cells 67 . Synaptotagmin I and II are specifically localized on synaptic vesicles 68 . Next I will refer to the best characterized isoform synaptotagmin I as simply synaptotagmin. Knock-out of synaptotagmin I gene is lethal and cultured hippocampal neurons from the knock-out mice exhibite a specific defect in Ca 2+ -triggered neurotransmitter release while the spontaneous release remains normal 55 . As mentioned previously, synaptotagmin and complexin have similar knockout phenotype in mice, both of them impact the fast synchronous Ca 2+ -triggered neurotransmitter release, but slow asynchronous Ca 2+ -triggered release is not altered by deletion of neither complexin nor synaptotagmin. So they may act cooperatively to fine- tune the subtle Ca 2+ -triggered neurotransmitter release process. In the “complexin-clamp model”69 , it has ever been claimed that Ca 2+ -bound synaptotagmin will release the clamping effect of complexin on the readily primed synaptic vesicles. People have also raised the hypothesis that primed metastable vesicles form the substrate for

12 synaptotagmin to trigger the fast synchronous release upon Ca 2+ influx by displacing complexin, and coupling the SNARE complex to phospholipids (Figure 12). However, not enough evidence has been provided yet for an explicit model for the coordinating mechanism of synaptotagmin and complexin.

1.7 Study methods

1.7.1 Site-directed spin labeling (SDSL) and EPR method

Site-directed spin labeling (SDSL) combined with electron paramagnetic resonance

(EPR) spectroscopy is a well-established technique for investigations in structure and topology of membrane proteins. Site-directed spin labeling is a powerful tool for monitoring the structure and dynamics of proteins 70; 71 . A nitroxide side chain is introduced via cysteine substitution mutagenesis followed by attachment of a methanethoisulfonate spin label (MTSSL) to the cysteine residue through a disulfide bond (Figure 13a).

EPR is a powerful and sensitive technique for the study of macromolecular structure and has been widely used in detecting protein structure as well. The EPR signal is generated by the unpaired electron on the nitroxide side chain of MTSSL, which is rare in biological system. Compared to other methods of structure study, crystallization and

NMR, EPR has neither requirement of crystal form nor strict constraints in molecular size

13 and sample concentration 72 . The EPR line shape is a function of the nitroxide side chain tumbling rate. We can get information about the structural environment of the spin label, such as its solvent accessibility, local geometry, flexibility and structural changes upon chemical or physical event. For example, an isotropic spectrum shown in Figure 13b represents a freely moving spin label side chain in solution without any restrictions. In contrast, a restricted spin label side chain may exhibit broad and less isotropic peaks as shown in Figure 13c . However, partially broadened spectra may also be observed, representing an immobilized subpopulation. Therefore by scanning the EPR signal of a sequence of residues in our interested region, we can map the protein structure in that area.

Since the EPR line shape yields information about the mobility and structural environment of a spin label, which also reflects the structural environment of the protein backbone, changes in the spectrum may also be indicative of conformational changes. For example, EPR spectroscopy has been successfully applied to investigating the protein function and dynamics in T4 lysozyme 73; 74 . A switch from the “open” conformation to a

“closed” active state has been detected by introducing double spin labels into pairs of cysteine residues simultaneously. Some of the doubly spin-labeled mutants exhibited

EPR spectra which were considerably broadened or sharpened in the presence of the substrate indicating a basic conformational change. While the sum of these EPR spectra between the corresponding singly spin-labeled mutants remained the same. Taken together, followed by some quantitative analysis of the spin-spin interactions,

McHaourab and co-workers successfully reveal a substrate-induced conformational change in T4 lysozyme based on the EPR method.

1.7.2 EPR accessibility measurement

Sometimes, the EPR line shape alone is not sufficient to yield information about the entire secondary and ternary structures. And the membrane associated characteristic of

SNARE complex makes the matters even more complicated. In this case, the EPR saturation method is a powerful tool to assess the local structure and membrane topology of the membrane-proximal regions of proteins 75; 76; 77 . EPR saturation method measures the accessibility of the nitroxide to a polar paramagnetic reagent such as NiEDDA

(W NiEDDA ) to estimate the solvent exposure of the spin-labeled side chain, or the accessibility to a non-polar paramagnetic reagent such as O 2 (Wo 2) to determine the membrane insertion. A relatively low value of W NiEDDA may suggest that the protein side chain could be either inserted into the membrane or buried in the protein environment.

Correspondingly, a high value of Wo 2 indicates a membrane-inserted nitroxide, because oxygen heavily prefers the membrane phase to the protein interior.

By plotting the value of W NiEDDA and Wo 2 of each spin-labeled residue against the residue number, we can observe the general tendency of accessibility to the membrane and water phase of the scanned region. A set of example plots of these power saturation

77 curves is shown in Figure 14 . The ratio of W NiEDDA and Wo 2 is also a useful parameter

15 for the secondary structure prediction. For example, a periodical behavior of this parameter along the sequence may indicate an α-helical structure. In addition to investigating the secondary structure and conformational change of proteins, EPR method has also been applied to determining the membrane immersion depth of protein side- chains in lipid bilayers. The immersion depth measurement of a nitroxide on a lipid chain in a pure bilayer has been reported first 78; 79 . Then in 1994, EPR method has been further developed to measure the immersion depth of a nitroxide on transmembrane domains of proteins 80 . The primary principle this approach bases on is that the collision rates of the nitroxides with paramagnetic reagents are associated with position. Small molecules in a membrane/water system are partitioned between the hydrophilic and the hydrophobic phase according to their polarity. Obviously, polar molecules prefer the water phase and nonpolar molecules prefer the membrane phase. However, the interior of bilayer has nonuniform gradients of fluidity and polarity along the direction of the bilayer normal.

Therefore, nitroxides located on the protein side chains are supposed to have a depth- dependent collision rate with a paramagnetic reagent corresponding to the phases. The two paramagnetic reagents mentioned in the accessibility measurement experiment are both fast-relaxing reagents along the gradients. So the values we got in the accessibility experiment can also be used to calculate the membrane immersion depth of the nitroxide side chain with certain formulas. However, we need to be very careful in interpreting the

EPR immersion depth data. The immersion depth reports the location of the substituted nitroxide side chain other than the actual location of the native amino acid when the native residue is charged, because the charged residues may tend to approximate the charged phosphate head groups while the nitroxide prefers the relatively hydrophobic

16 interior of the membrane. In summary, these EPR methods provide useful strategies for investigating the membrane protein interface, but further thoughts are still necessary due to the complicated circumstance.

1.8 References

1. Rothman, J. E. (1994). Mechanisms of intracellular protein transport. Nature 372 , 55-63.

2. Chen, Y. A. & Scheller, R. H. (2001). SNARE-mediated membrane fusion. Nat Rev Mol Cell Biol 2, 98-106.

3. Burgess, T. L. & Kelly, R. B. (1987). Constitutive and regulated secretion of proteins. Annu Rev Cell Biol 3, 243-93.

4. Sudhof, T. C. (2004). The synaptic vesicle cycle. Annu Rev Neurosci 27 , 509-47.

5. Meyer B. Jackson, E. R. C. (2006). Fusion Pores and Fusion machines in Ca 2+ - Triggered Exocytosis. Annu. Rev. Biohphys. Biomol. Struct. 35 .

6. Blasi, J., Chapman, E. R., Link, E., Binz, T., Yamasaki, S., De Camilli, P., Sudhof, T. C., Niemann, H. & Jahn, R. (1993). Botulinum neurotoxin A selectively cleaves the synaptic protein SNAP-25. Nature 365 , 160-3.

7. Chen, Y. A., Scales, S. J., Patel, S. M., Doung, Y. C. & Scheller, R. H. (1999). SNARE complex formation is triggered by Ca2+ and drives membrane fusion. Cell 97 , 165-74.

8. Sollner, T., Bennett, M. K., Whiteheart, S. W., Scheller, R. H. & Rothman, J. E. (1993). A protein assembly-disassembly pathway in vitro that may correspond to sequential steps of synaptic vesicle docking, activation, and fusion. Cell 75 , 409- 18.

9. Weber, T., Zemelman, B. V., McNew, J. A., Westermann, B., Gmachl, M., Parlati, F., Sollner, T. H. & Rothman, J. E. (1998). SNAREpins: minimal machinery for membrane fusion. Cell 92 , 759-72.

10. Bennett, M. K., Calakos, N. & Scheller, R. H. (1992). Syntaxin: a synaptic protein implicated in docking of synaptic vesicles at presynaptic active zones. Science 257 , 255-9.

11. Oyler, G. A., Higgins, G. A., Hart, R. A., Battenberg, E., Billingsley, M., Bloom, F. E. & Wilson, M. C. (1989). The identification of a novel synaptosomal- associated protein, SNAP-25, differentially expressed by neuronal subpopulations. J Cell Biol 109 , 3039-52.

12. Trimble, W. S., Cowan, D. M. & Scheller, R. H. (1988). VAMP-1: a synaptic vesicle-associated integral membrane protein. Proc Natl Acad Sci U S A 85 , 4538- 42.

13. Schiavo, G., Benfenati, F., Poulain, B., Rossetto, O., Polverino de Laureto, P., DasGupta, B. R. & Montecucco, C. (1992). Tetanus and botulinum-B neurotoxins block neurotransmitter release by proteolytic cleavage of synaptobrevin. Nature 359 , 832-5.

14. Yamasaki, S., Baumeister, A., Binz, T., Blasi, J., Link, E., Cornille, F., Roques, B., Fykse, E. M., Sudhof, T. C., Jahn, R. & et al. (1994). Cleavage of members of the synaptobrevin/VAMP family by types D and F botulinal neurotoxins and tetanus toxin. J Biol Chem 269 , 12764-72.

15. Schoch, S., Deak, F., Konigstorfer, A., Mozhayeva, M., Sara, Y., Sudhof, T. C. & Kavalali, E. T. (2001). SNARE function analyzed in synaptobrevin/VAMP knockout mice. Science 294 , 1117-22.

16. Washbourne, P., Thompson, P. M., Carta, M., Costa, E. T., Mathews, J. R., Lopez-Bendito, G., Molnar, Z., Becher, M. W., Valenzuela, C. F., Partridge, L. D. & Wilson, M. C. (2002). Genetic ablation of the t-SNARE SNAP-25 distinguishes mechanisms of neuroexocytosis. Nat Neurosci 5, 19-26.

17. Margittai, M., Fasshauer, D., Jahn, R. & Langen, R. (2003). The Habc domain and the SNARE core complex are connected by a highly flexible linker. Biochemistry 42 , 4009-14.

18. Fernandez, I., Ubach, J., Dulubova, I., Zhang, X., Sudhof, T. C. & Rizo, J. (1998). Three-dimensional structure of an evolutionarily conserved N-terminal domain of syntaxin 1A. Cell 94 , 841-9.

19. Hess, D. T., Slater, T. M., Wilson, M. C. & Skene, J. H. (1992). The 25 kDa synaptosomal-associated protein SNAP-25 is the major methionine-rich polypeptide in rapid axonal transport and a major substrate for palmitoylation in adult CNS. J Neurosci 12 , 4634-41.

20. Fasshauer, D., Otto, H., Eliason, W. K., Jahn, R. & Brunger, A. T. (1997). Structural changes are associated with soluble N-ethylmaleimide-sensitive fusion protein attachment protein receptor complex formation. J Biol Chem 272 , 28036- 41.

21. Fiebig, K. M., Rice, L. M., Pollock, E. & Brunger, A. T. (1999). Folding intermediates of SNARE complex assembly. Nat Struct Biol 6, 117-23.

22. Hazzard, J., Sudhof, T. C. & Rizo, J. (1999). NMR analysis of the structure of synaptobrevin and of its interaction with syntaxin. J Biomol NMR 14 , 203-7.

23. Poirier, M. A., Xiao, W., Macosko, J. C., Chan, C., Shin, Y. K. & Bennett, M. K. (1998). The synaptic SNARE complex is a parallel four-stranded helical bundle. Nat Struct Biol 5, 765-9.

24. Sutton, R. B., Fasshauer, D., Jahn, R. & Brunger, A. T. (1998). Crystal structure of a SNARE complex involved in synaptic exocytosis at 2.4 A resolution. Nature 395 , 347-53.

25. Fasshauer, D., Bruns, D., Shen, B., Jahn, R. & Brunger, A. T. (1997). A structural change occurs upon binding of syntaxin to SNAP-25. J Biol Chem 272 , 4582-90.

26. Fasshauer, D., Eliason, W. K., Brunger, A. T. & Jahn, R. (1998). Identification of a minimal core of the synaptic SNARE complex sufficient for reversible assembly and disassembly. Biochemistry 37 , 10354-62.

27. Rice, L. M., Brennwald, P. & Brunger, A. T. (1997). Formation of a yeast SNARE complex is accompanied by significant structural changes. FEBS Lett 415 , 49-55.

28. Poirier, M. A., Hao, J. C., Malkus, P. N., Chan, C., Moore, M. F., King, D. S. & Bennett, M. K. (1998). Protease resistance of syntaxin.SNAP-25.VAMP complexes. Implications for assembly and structure. J Biol Chem 273 , 11370-7.

29. Hayashi, T., McMahon, H., Yamasaki, S., Binz, T., Hata, Y., Sudhof, T. C. & Niemann, H. (1994). Synaptic vesicle membrane fusion complex: action of clostridial neurotoxins on assembly. Embo J 13 , 5051-61.

30. Misura, K. M., Scheller, R. H. & Weis, W. I. (2000). Three-dimensional structure of the neuronal-Sec1-syntaxin 1a complex. Nature 404 , 355-62.

31. Christie, M. P., Whitten, A. E., King, G. J., Hu, S. H., Jarrott, R. J., Chen, K. E., Duff, A. P., Callow, P., Collins, B. M., James, D. E. & Martin, J. L. Low- resolution solution structures of Munc18:Syntaxin protein complexes indicate an

open binding mode driven by the Syntaxin N-peptide. Proc Natl Acad Sci U S A 109 , 9816-21.

32. Dulubova, I., Sugita, S., Hill, S., Hosaka, M., Fernandez, I., Sudhof, T. C. & Rizo, J. (1999). A conformational switch in syntaxin during exocytosis: role of munc18. Embo J 18 , 4372-82.

33. Zhang, F., Chen, Y., Kweon, D. H., Kim, C. S. & Shin, Y. K. (2002). The four- helix bundle of the neuronal target membrane SNARE complex is neither disordered in the middle nor uncoiled at the C-terminal region. J Biol Chem 277 , 24294-8.

34. Xiao, W., Poirier, M. A., Bennett, M. K. & Shin, Y. K. (2001). The neuronal t- SNARE complex is a parallel four-helix bundle. Nat Struct Biol 8, 308-11.

35. Margittai, M., Fasshauer, D., Pabst, S., Jahn, R. & Langen, R. (2001). Homo- and heterooligomeric SNARE complexes studied by site-directed spin labeling. J Biol Chem 276 , 13169-77.

36. Hua, S. Y. & Charlton, M. P. (1999). Activity-dependent changes in partial VAMP complexes during neurotransmitter release. Nat Neurosci 2, 1078-83.

37. Rice, L. M. & Brunger, A. T. (1999). Crystal structure of the vesicular transport protein Sec17: implications for SNAP function in SNARE complex disassembly. Mol Cell 4, 85-95.

38. Lenzen, C. U., Steinmann, D., Whiteheart, S. W. & Weis, W. I. (1998). Crystal structure of the hexamerization domain of N-ethylmaleimide-sensitive fusion protein. Cell 94 , 525-36.

39. Antonin, W., Fasshauer, D., Becker, S., Jahn, R. & Schneider, T. R. (2002). Crystal structure of the endosomal SNARE complex reveals common structural principles of all SNAREs. Nat Struct Biol 9, 107-11.

40. Cohen, F. S. & Melikyan, G. B. (2004). The energetics of membrane fusion from binding, through hemifusion, pore formation, and pore enlargement. J Membr Biol 199 , 1-14.

41. Chen, X., Arac, D., Wang, T. M., Gilpin, C. J., Zimmerberg, J. & Rizo, J. (2006). SNARE-mediated lipid mixing depends on the physical state of the vesicles. Biophys J 90 , 2062-74.

42. van den Bogaart, G., Holt, M. G., Bunt, G., Riedel, D., Wouters, F. S. & Jahn, R. One SNARE complex is sufficient for membrane fusion. Nat Struct Mol Biol 17 , 358-64.

43. Almers, W. & Tse, F. W. (1990). Transmitter release from synapses: does a preassembled fusion pore initiate exocytosis? Neuron 4, 813-8.

44. Han, X. & Jackson, M. B. (2005). Electrostatic interactions between the syntaxin membrane anchor and neurotransmitter passing through the fusion pore. Biophys J 88 , L20-2.

45. Han, X., Wang, C. T., Bai, J., Chapman, E. R. & Jackson, M. B. (2004). Transmembrane segments of syntaxin line the fusion pore of Ca2+-triggered exocytosis. Science 304 , 289-92.

46. Monck, J. R. & Fernandez, J. M. (1994). The exocytotic fusion pore and neurotransmitter release. Neuron 12 , 707-16.

47. Chernomordik, L. V. & Kozlov, M. M. (2005). Membrane hemifusion: crossing a chasm in two leaps. Cell 123 , 375-82.

48. Reese, C., Heise, F. & Mayer, A. (2005). Trans-SNARE pairing can precede a hemifusion intermediate in intracellular membrane fusion. Nature 436 , 410-4.

49. Giraudo, C. G., Hu, C., You, D., Slovic, A. M., Mosharov, E. V., Sulzer, D., Melia, T. J. & Rothman, J. E. (2005). SNAREs can promote complete fusion and hemifusion as alternative outcomes. J Cell Biol 170 , 249-60.

50. Lu, X., Zhang, F., McNew, J. A. & Shin, Y. K. (2005). Membrane fusion induced by neuronal SNAREs transits through hemifusion. J Biol Chem 280 , 30538-41.

51. Xu, Y., Zhang, F., Su, Z., McNew, J. A. & Shin, Y. K. (2005). Hemifusion in SNARE-mediated membrane fusion. Nat Struct Mol Biol 12 , 417-22.

52. Yoon, T. Y., Okumus, B., Zhang, F., Shin, Y. K. & Ha, T. (2006). Multiple intermediates in SNARE-induced membrane fusion. Proc Natl Acad Sci U S A 103 , 19731-6.

53. Stein, A., Weber, G., Wahl, M. C. & Jahn, R. (2009). Helical extension of the neuronal SNARE complex into the membrane. Nature 460 , 525-8.

54. Skehel, J. J. & Wiley, D. C. (1998). Coiled coils in both intracellular vesicle and viral membrane fusion. Cell 95 , 871-4.

55. Geppert, M., Goda, Y., Hammer, R. E., Li, C., Rosahl, T. W., Stevens, C. F. & Sudhof, T. C. (1994). Synaptotagmin I: a major Ca2+ sensor for transmitter release at a central synapse. Cell 79 , 717-27.

56. Weninger, K., Bowen, M. E., Choi, U. B., Chu, S. & Brunger, A. T. (2008). Accessory proteins stabilize the acceptor complex for synaptobrevin, the 1:1 syntaxin/SNAP-25 complex. Structure 16 , 308-20.

57. McMahon, H. T., Missler, M., Li, C. & Sudhof, T. C. (1995). Complexins: cytosolic proteins that regulate SNAP receptor function. Cell 83 , 111-9.

58. Reim, K., Wegmeyer, H., Brandstatter, J. H., Xue, M., Rosenmund, C., Dresbach, T., Hofmann, K. & Brose, N. (2005). Structurally and functionally unique complexins at retinal ribbon synapses. J Cell Biol 169 , 669-80.

59. Yoon, T. Y., Lu, X., Diao, J., Lee, S. M., Ha, T. & Shin, Y. K. (2008). Complexin and Ca2+ stimulate SNARE-mediated membrane fusion. Nat Struct Mol Biol 15 , 707-13.

60. Reim, K., Mansour, M., Varoqueaux, F., McMahon, H. T., Sudhof, T. C., Brose, N. & Rosenmund, C. (2001). Complexins regulate a late step in Ca2+-dependent neurotransmitter release. Cell 104 , 71-81.

61. Takahashi, S., Ujihara, H., Huang, G. Z., Yagyu, K. I., Sanbo, M., Kaba, H. & Yagi, T. (1999). Reduced hippocampal LTP in mice lacking a presynaptic protein: complexin II. Eur J Neurosci 11 , 2359-66.

62. Tang, J., Maximov, A., Shin, O. H., Dai, H., Rizo, J. & Sudhof, T. C. (2006). A complexin/synaptotagmin 1 switch controls fast synaptic vesicle exocytosis. Cell 126 , 1175-87.

63. Huntwork, S. & Littleton, J. T. (2007). A complexin fusion clamp regulates spontaneous neurotransmitter release and synaptic growth. Nat Neurosci 10 , 1235-7.

64. Xue, M., Reim, K., Chen, X., Chao, H. T., Deng, H., Rizo, J., Brose, N. & Rosenmund, C. (2007). Distinct domains of complexin I differentially regulate neurotransmitter release. Nat Struct Mol Biol 14 , 949-58.

65. Lu, B., Song, S. & Shin, Y. K. Accessory alpha-helix of complexin I can displace VAMP2 locally in the complexin-SNARE quaternary complex. J Mol Biol 396 , 602-9.

66. Chen, X., Tomchick, D. R., Kovrigin, E., Arac, D., Machius, M., Sudhof, T. C. & Rizo, J. (2002). Three-dimensional structure of the complexin/SNARE complex. Neuron 33 , 397-409.

67. Li, C., Davletov, B. A. & Sudhof, T. C. (1995). Distinct Ca2+ and Sr2+ binding properties of synaptotagmins. Definition of candidate Ca2+ sensors for the fast and slow components of neurotransmitter release. J Biol Chem 270 , 24898-902.

68. Perin, M. S., Fried, V. A., Mignery, G. A., Jahn, R. & Sudhof, T. C. (1990). Phospholipid binding by a synaptic vesicle protein homologous to the regulatory region of protein kinase C. Nature 345 , 260-3.

69. Giraudo, C. G., Eng, W. S., Melia, T. J. & Rothman, J. E. (2006). A clamping mechanism involved in SNARE-dependent exocytosis. Science 313 , 676-80.

70. Cornish, V. W., Benson, D. R., Altenbach, C. A., Hideg, K., Hubbell, W. L. & Schultz, P. G. (1994). Site-specific incorporation of biophysical probes into proteins. Proc Natl Acad Sci U S A 91 , 2910-4.

71. Hubbell, W. L., McHaourab, H. S., Altenbach, C. & Lietzow, M. A. (1996). Watching proteins move using site-directed spin labeling. Structure 4, 779-83.

72. Biswas, R., Kuhne, H., Brudvig, G. W. & Gopalan, V. (2001). Use of EPR spectroscopy to study macromolecular structure and function. Sci Prog 84 , 45-67.

73. McHaourab, H. S., Oh, K. J., Fang, C. J. & Hubbell, W. L. (1997). Conformation of T4 lysozyme in solution. Hinge-bending motion and the substrate-induced conformational transition studied by site-directed spin labeling. Biochemistry 36 , 307-16.

74. McHaourab, H. S., Lietzow, M. A., Hideg, K. & Hubbell, W. L. (1996). Motion of spin-labeled side chains in T4 lysozyme. Correlation with protein structure and dynamics. Biochemistry 35 , 7692-704.

75. Kweon, D. H., Kim, C. S. & Shin, Y. K. (2003). Insertion of the membrane- proximal region of the neuronal SNARE coiled coil into the membrane. J Biol Chem 278 , 12367-73.

76. Kweon, D. H., Kim, C. S. & Shin, Y. K. (2002). The membrane-dipped neuronal SNARE complex: a site-directed spin labeling electron paramagnetic resonance study. Biochemistry 41 , 9264-8.

77. Yu, Y. G., Thorgeirsson, T. E. & Shin, Y. K. (1994). Topology of an amphiphilic mitochondrial signal sequence in the membrane-inserted state: a spin labeling study. Biochemistry 33 , 14221-6.

78. Dalton, L. A., McIntyre, J. O. & Fleischer, S. (1987). Distance estimate of the active center of D-beta-hydroxybutyrate dehydrogenase from the membrane surface. Biochemistry 26 , 2117-30.

79. Pali, T., Bartucci, R., Horvath, L. I. & Marsh, D. (1992). Distance measurements using paramagnetic ion-induced relaxation in the saturation transfer electron spin resonance of spin-labeled biomolecules: Application to phospholipid bilayers and interdigitated gel phases. Biophys J 61 , 1595-602.

80. Altenbach, C., Greenhalgh, D. A., Khorana, H. G. & Hubbell, W. L. (1994). A collision gradient method to determine the immersion depth of nitroxides in lipid bilayers: application to spin-labeled mutants of bacteriorhodopsin. Proc Natl Acad Sci U S A 91 , 1667-71.

1.8 Figures and Captions

Südhof TC, Annu. Rev. Neurosci., 2004

Figure 1 Secretory pathways in neurons. The center part of this drawing schematically illustrates the pathway of neurotransmitter release by synaptic vesicle exocytosis (step 1- 9). Synaptic vesicles are filled with neurotransmitters (step 1), and form the vesicle cluster (step 2). Vesicles dock at the active zone (step 3) to undergo a priming stage (step 4) and get ready for Ca 2+ -triggered fusion pore opening (step 5). After fusion pore opening, synaptic vesicles can go through three alternative pathways shown by step 6, 7, 8 and 9.

Misura KM, Nature, 2000

Figure 2 Structure of syntaxin. a, Topology diagram of syntaxin. b, Ribbon representation of syntaxin. Habc domain, red; Habc/H3 linker, orange; H3 region, purple. The approximate dimensions are also shown in b.

Brunger AT, Q. Rev. Biophys., 2005

Figure 3 Primary domain structure of neuronal SNAREs. Primary structure diagram for syntaxin (red), SNAP-25 (green), and synaptobrevin (blue). TM, transmembrane domain is located at the C-terminal of syntaxin and VAMP. The SNARE motifs are defined through the 16 layers as found in the crystal structure (Sutton et al . 1998). Green lines indicate the palmitoylation sites in SNAP-25. The two SNARE motifs are named as SN1 and SN2 in SNAP-25. The N-terminal Habc domain in syntaxin is not shown in this figure.

Sutton RB et al., Nature, 1998

Figure 4 Crystal structure of neuronal SNARE complex. a, Backbone ribbon drawing of the SNARE core complex. VAMP, blue; syntaxin, red; SNAP-25 (SN1 and SN2), green. b, Organization of the SNARE core complex. b, The 16 hydrophobic layers of the

SNARE core complex are shown by virtual bonds between corresponding C α positions. c, Ionic ‘0’ layer of the synaptic fusion complex.

Christie MP et al. , Proc. Natl. Acad. Sci. USA, 2012

Figure 5 Closed and open conformation of Munc18: Syntaxin complexes. a, Crystal structure of closed syntaxin1A (red) bound to Munc18 -1. b, Model of Munc18 -1 (blue) in complex with syntaxin1A. c, Model of Munc18c (blue) in complex with syntaxin 4 (red). Ref ined against solution scattering data and distance restraints indicates that syntaxin1A and syntaxin 4 both adopt an open conformation when bound to Munc18 -1. For both b and c, the position of the H3 helix is just predicted. d, Schematic model showing a mo del switch between the closed and open conformation of syntaxin1A, involving an N -peptide regulation.

Xiao WZ et al., Nature Structural Biology, 2001

Figure 6 Schematic representations of binary and ternary SNARE complexes. a, t- SNARE complex. b, ternary SNARE complex. Syntaxin, green; SN1, red; SN2, blue; VAMP2, yellow. The dashed line indicates that the interspin distances is larger than 25Å.Interspin distances differing by 3Å or more between t-SNARE and SNARE complexes are shown in white boxes.

Südhof TC et al., Science, 2009

Figure 7 Model for trans -SNARE and cis -SNARE conformation. a, Three helices anchored in the plasma membrane (t-SNARE) assemble with the fourth helix from the vesicle membrane (v-SNARE) to form a trans-SNARE intermediate called the trans - SNARE complex. b, The SNARE complex ends up with the four-helix bundle residing on the same membrane called the cis -SNARE complex.

Rizo J et al., Nat. Struct Mol Biol., 2008

Figure 8 SNARE complex assembly pathway. a, Partially assembled SNARE complexes (center) could either induce membrane fusion as they fully assemble (left) or could remain fully assembled between the two membranes (right) if the linker regions between the SNARE motifs and the transmembrane domains are flexible. VAMP, orange; SNAP-25, green; syntaxin, yellow. b, Munc18-1 bound to the closed conformation of syntaxin (left) and partially assembled SNARE complexes (center) then induce fusion as the SNARE complexes fully assembled (right). Munc-18, purple. Munc13s and RIMs are potential regulators of the transition between the two complexes.

Yoon TY and Shin YK, Cell. Mol. Life Sci., 2008

Figure 9 Two models of membrane fusion pathway . a, Pre-docking stage: t-SNARE proteins in the plasma membrane exist in a clustered form. b, Docking and gap junction- like structure: the SNARE complexes bridge two membrane-spanning structures which is similar to the structure of gap junction. c, Hemifusion state: Some of the lipid molecules form membrane connectivity in the form of a lipid stalk. D, Fusion pore opening: inner leaflets merge and the transmembrane domains completely dissociate from each other to open a fusion pore.

Xue MS et al., Nat. Struct. Biol.,2007

Figure 10 Model for complexin dual function in Ca 2+ -triggered neurotransmitter release. a, Before the fusion process, t-SNARE and v-SNARE resides on opposite membranes with the t-SNARE partially structured. b, The SNARE complex forms but zippering is not complete, the C terminus of the SNARE complex is still partially unstructured. c, Complexin binds the partly assembled SNARE and helps to stabilize the SNARE complex. The accessory α-helix occupies the position of the C-terminal VAMP and thereby hinders full assembly of the SNARE complex before Ca 2+ influx. d, After Ca 2+ influx, Ca 2+ -bound synaptotagmin interacts with phospholipid membrane and the SNARE complex, and fully assembles the SNARE complex. e, Fusion-pore opens, and complexin is displaced from the SNARE complex during this process.

Xue MS et al., Nat. Struct. Biol., 2002

Giraudo CG et al., J. Biol. Chem. 2008

Figure 11 Domain structure of complexin I and crystal structure of the SNARE/complexin quaternary complex. a, Schematic diagram of domain organization of Complexin I. b, Three-dimensional structure of the SNARE/complexin complex. Complexin, magenta; VAMP, red; syntaxin1, green; SN1, blue; SN2, yellow. The residues in complexin facing the SNARE complex are labeled in orange, and those that are conserved among different complexins are highlighted in white. c, Three-dimensional structure of SNARE/complexin complex showing residues in complexin facing out from the SNARE complex in light blue.

Tang J. et al., Cell, 2006

Figure 12 Model of complexin and synaptotagmin coordination in Ca 2+ -triggered release. Docked vesicles containing unassembled SNARE complexes are primed by partial SNARE-complex assembly (step 1). The resulting primed vesicles can undergo two release pathways: asynchronous release (step 2 and 3), and synchronous release (step 4, 5 and 6). Synaptotagmin binding involves the phospholipids can displace complexin and result in fusion pore opening (step 5). Opened fusion pores can then dilate to complete fusion (step 6).

Biswas R. et al., Sci. Prog., 2001

Figure 13 A schematic description of site-directed spin labeling (SDSL) strategy . a, The site-directed spin labeling strategy. b, Example EPR spectra of a nitroxide spin label in aqueous solution at room temperature. c, Example EPR spectra of a nitroxide spin label in glycerol at room temperature. d, Example EPR spectra of a nitroxide spin label frozen in aqueous solution.

Yu YG. et al., Biochemistry, 1994

Figure 14 Schemetic power saturation curves in different polar paramagnetic solvent. These example curves are from the PCOX peptide in POPC vesicles containing 10 mol % POPG. The data were obtained by measuring the peak-to-peak amplitude of the central peak of the first-derivative EPR spectrum. The EPR spectra were collected at 0.1, 0.25, 0.4, 1.0, 4.0, 10, 16, 25, and 40mW incident microwave powers under three different conditions: under nitrogen flow in the absence of paramagnetic reagents; under nitrogen in the presence of 10mM NiEDDA; and under the flow of air. The curves are the least-squares fits to certain standard equations.

CHAPTER 2: ACCESSORY α-HELIX OF COMPLEXIN I CAN DISPLACE VAMP2 LOCALLY IN THE COMPLEXIN-SNARE QUATERNARY COMPLEX *

Bin Lu †, Shuang Song † and Yeon-Kyun Shin

2.1 Abstract

The calcium-triggered neurotransmitter release requires three SNARE (soluble N- ethylmaleimide-sensitive factor attachment protein receptor) proteins: synaptobrevin 2

(or VAMP2) on the synaptic vesicle and syntaxin I and SNAP-25 at the presynaptic plasmas membrane. This minimal fusion machinery is believed to drive fusion of the vesicle to the presynaptic membrane. Complexin, also known as synaphin, is a neuronal cytosolic protein that acts as a major regulator of synaptic vesicle exocytosis. Stimulatory and inhibitory effects of complexin have both been reported, suggesting the duality of its function. To shed light on the molecular basis of the complexin’s dual function, we have performed an EPR investigation of the complexin-SNARE quaternary complex. We found that the accessory α-helix (amino acids 27-48) by itself has the capacity to replace the C-terminus of the SNARE motif of vesicle-associated membrane protein 2 in the four-helix bundle and makes the SNARE complex weaker when the N-terminal region of complexin I (amino acids 1-26) is removed. However, the accessory α-helix remains detached from the SNARE core when the N-terminal region of complexin I is present.

*This paper is published in J. Mol. Biol . (2010) 396, 602-609. Necessary modifications are made to fit the format of this dissertation. †Contributed equally to this work. Author contributions: B.L., and S.S. performed research and analyzed data; Y.K.S. designed research and wrote the paper.

Thus, our data show the possibility that the balance between the activities of the accessory α-helix and the N-terminal domain might determine the final outcome of the complexin function, either stimulatory or inhibitory.

2.2 Introduction

Synaptic vesicle fusion is driven by assembly of trans-SNARE complex between the vesicle and the plasma membrane, which facilitates merging of the opposite bilayers 1; 2; 3;

4; 5; 6; 7 . The physiological trigger for this process is the spike in the local concentration of calcium. As Ca 2+ enters the presynaptic cytoplasm, the Ca 2+ -sensor synaptotagmin I8; 9; 10;

11 , which is localized on synaptic vesicles, interacts with the SNARE complex and phospholipids simultaneously 12; 13; 14 , leading to the fast opening of the fusion pore 15; 16; 17;

18 .

In addition to synaptotagmin I, complexins also participate in the fast Ca 2+ -dependent neuronal exocytosis 19; 20; 21; 22 . Complexins are small soluble cytoplasmic protein (15 to

18kDa) that are mainly found in the presynaptic part of neuronal cells 23; 24; 25 and the central region (amino acis 48-70) binds to the SNARE core as an antiparallel α-helix, which attaches complexin to the SNARE complex 26; 27 .

Complexin is believed to have a dual function 20; 28; 29 in synaptic membrane fusion- both stimulation 19; 26; 27 and inhibition 21; 30; 31; 32; 33; 34; 35 . Although the exact molecular mechanism is unknown, the N-terminal region of complexin is shown to play an

40 important role in promoting the neurotransmitter release28; 29; 36 . It is proposed that the N- terminal region interacts with the membrane-proximal region of VAMP2 (vesicle- associated membrane protein 2) and functions as a force transducer 29 . However, under different conditions, complexin exhibits the inhibitory function 28 , which is the basis for the recent “complexin-clamp model”34 . Two possible molecular-level mechanistic models have been proposed to explain the inhibition by complexin. Firstly, complexin is found to have the micromolar affinity to the t-SNARE (target membrane SNARE) complex in addition to its submicromolar affinity to the SNARE complex 20; 26 . Thus, under favorable conditions, complexin could compete with the VAMP2 binding to the t-

SNARE complex, which would competitively inhibit SNARE assembly and membrane fusion. Secondly, when bound to the SNARE core, the accessory α-helix of complexin, which franks the central SNARE-anchor domain and the stimulatory N-terminal region, might replace the C-terminal portion of the VAMP2 helix to form an alternative four- helix bundle with the helices from t-SNAREs 28; 34; 37 . Such structural replacement would prevent formation of the complete SNARE four-helix bundle, thereby inhibiting membrane fusion.

In this work, we performed the EPR investigation of the conformational changes in the vesicle-associated SNARE VAMP2 after complexin binding to the SNARE complex.

The results show that there is no major conformational change, such as the replacement of VAMP2 by complexin, occurring at the membrane-proximal region of VAMP2 upon binding of the full-length complexin to the SNARE complex. However, when the stimulatory N-terminal region of complexin is removed, the accessory α-helix replaces

41 the C-terminal region of VAMP2 partially. Thus, our results show that the balance between the stimulatory and inhibitory activities of the accessory α-helix and the adjacent

N-terminal region, respectively, might determine the exact function of the particular complexin.

2.3 Results

2.3.1 Complexin-SNARE quaternary complexin formation

The recent study by Rothman et al. suggests that the accessory α-helix of complexin replaces VAMP2 and forms an alternative four-helix bundle with t-SNAREs near the membrane, thereby inhibiting complete SNARE complex formation and membrane fusion 34 . According to this proposed model, the displaced sequence of VAMP2 includes both the cleavage site (between residues 76 and 77) and recognition site (residues 62 to

71) for BoNT/B toxin. Therefore, our EPR investigation was focused on the structure of the C-terminal region of VAMP2. We prepared five single cysteine mutants of VAMP2

(Figure 1a ), including A69C within the recognition site of BoNT/B and Q76C on its cleavage site and K83C, R86C, and W89C in the membrane-proximal region. The cysteine mutants were derivatized with methanethiosulfonate spin label [(1-oxyl-2,2,5,5- tetramethyl-D-pyrroline-3-methyl) methanethiosulfonate] for EPR measurements. Next, we used the pull-down method ( Figure 1b ) to prepare the complexin-SNARE quaternary

42 complex that includes the spin labeled soluble VAMP2, full-length syntaxin 1A, SNAP-

25 (synaptosome-associated protein of 25 kDa), and complexin. As the bait, His 6-tagged

SNAP-25 was attached to the nickel-nitrilotriacetic acid (Ni-NTA) beads and the SNARE complex was first formed by flowing syntaxin 1A and soluble VAMP2 into the bead solution. Complexin was next added to the column to form the complexin-SNARE quaternary complex. The transmembrane domain (TMD) of VAMP2 was not included in order to avoid the interaction between two TMDs in one membrane and to best mimic the trans -SNARE complex. For individual mutants, we analyzed formation of the complexin-

SNARE quaternary complex by the SDS-PAGE gel after heating. We found that the immobilized His 6-tagged SNAP-25 co-purified with stoichiometric amounts of syntaxin

1A, soluble VAMP2, and complexin ( Figure 1c ).

2.3.2 Full-length complexin does not replace VAMP2

The EPR line shape is a sensitive function of the motional rate. The slower the motion of the nitroxide side chain, the broader the EPR spectrum. Therefore, if the proposed structure of the quaternary complex in Figure 1a were to be true, we expect the narrowing of the EPR spectra for the spin-labeled positions when compared with the EPR spectra from the SNARE complex only. EPR spectra of the SNARE complex before and after binding to full-length complexin I were shown in Figure 2 . The spectra for position

60 and 76 of the complexin-SNARE complex became much broader than those from the

SNARE complex only. This tendency was more obvious at position 69, indicating that

43 there was strong tertiary interaction at this site. Our data were also consistent with the deuterium exchange results, which showed that position 60 was highly protected after complexin binding 27 . Thus, the EPR results for position 69 and 76 were not consistent with the proposed structure in Figure 1a . Instead, the results suggest that complexin bind on the surface of the SNARE four-helix bundle, as shown in the crystal structure 27 .

For positions 83, 86, and 89, which are located in the membrane-proximal region, there was no obvious narrowing due to the replacement of VAMP2 by complexin either.

Unlike the cases with positions 69 and 76, we did not observe extra line broadening due to the complexin binding, suggesting that complexin might not even interact with the

SNARE complex locally in the region. Further addition of complexin to the complexin-

SNARE quaternary complex did not change the EPR line shape either ( Figure S1 ), suggesting that the reason we did not observe complexin-induced line broadening was neither due to the insufficient complexin binding nor due to the intermolecular complexin binding equilibrium. Therefore, the EPR analysis of the quaternary complex suggested that full-length complexin I does not replace VAMP2 when bound to the SNARE complex.

2.3.3 The accessory α-helix of complexin has no obvious interaction with the SNARE core

We now examined the conformation of complexin in the complexin-SNARE quaternary complex using spin-labeled complexin. We aligned the sequences of complexin with VAMP2 in an antiparallel orientation as they were shown in the crystal structure 27; 34 and selected five positions R42, E35, P28, E25, and G22 in the accessory α- helix as the spin labeling sites. EPR spectra for these sites in the complexin-SNARE complex were shown in Figure 3 . The line shapes for spin-labeled complexin alone were all very sharp, reflecting the freely diffusing random coil. When the spin-labeled mutants were bound to the SNARE complex, the spectra became broader, reflecting the slower motion of the nitroxide, most likely due to formation of α-helix. However, the spectra were still not so broad with any indication of any tertiary contact. In fact, the spectra are sharper than those for the solvent exposed positions on the surface helix 38; 39 , indicative of substantial motion of the accessory α-helix. Therefore, our EPR analysis shows that the accessory α-helix most likely has a little or no tertiary interaction with the SNARE core.

2.3.4 The accessory α-helix of complexin can partially displace VAMP2 locally when

N-terminal region is absent

When complexins are deleted from the neuron, the Ca 2+ -triggered fast release is severely reduced. In contrast, the frequency of the Ca 2+ -independent spontaneous release is dramatically enhanced. However, when the 26 N-terminal residues are removed from complexin, both the evoked and the spontaneous releases are reduced when compared

45 with those of complexin-null mutant 29 . Thus, these results show that the shortened complexin mutant lacking the stimulatory N-terminal sequence has an inhibitory function for the spontaneous release. In this case, it is possible that the accessory α-helix displaces

VAMP2, thereby inhibiting membrane fusion. To test this possibility, we prepared the shortened construct of complexin lacking the 26 N-terminal residues (amino acids 27-134) and investigated the structure of the complexin ∆26-SNARE quaternary complex employing the spin-labeled VAMP2 mutants (Figure 4a and b). For nitroxides attached to positions 69 and 76, the EPR spectra were very similar to those obtained from the full- length complexin-SNARE quaternary complexin (Figure 2). However, for three C- terminal spin labeled positions 83,86, and 89, the EPR spectra showed a small but distinctly sharp spectral component (Figure 4b, see arrows, 7-17%), which is most likely from the frayed VAMP2 chain that is freely diffusing and fast moving in solution. Thus, the EPR analysis shows that the accessory α-helix of complexin has the capacity to replace VAMP2 and makes the SNARE complex somewhat weaker in the absence of the

N-terminal region. It appears though that the sharp spectral component for N-terminal position 83(17%) is larger than those for C-terminal positions 86 and 89 (9% and 7%, respectively). One, however, must be cautious in interpreting these numbers when taking into account the perturbation by nitroxides and the uncertainty of the spectral subtraction analysis. Therefore, given the experimental uncertainty, we would not want to put much weight on the order of the extent of displacement among the spin labeled positions.

Our EPR analysis shows that the displacement of VAMP2 by complexin ∆26 happens locally in the membrane-proximal region. If true, we expect that the accessory α-helical

46 region of complexin ∆26 would interact with the t-SNARE helices in replacement of

VAMP2. Indeed, for spin labeled complexin ∆26 mutants R37C and L41C, we observe small but distinct line broadening (Figure 4c ) most likely due to the tertiary interaction between the accessory α-helix and the t-SNARE core. Although positions 37 and 41 are not in register with VAMP2 residues 83-89, the results show that the tertiary interaction between complexin I ∆26 and the t-SNARE core might occur in the part preceding this region.

2.4 Discussion

The EPR analysis of the complexin-SNARE quaternary complex revealed that the accessory α-helix has small but distinct capacity to displace the C-terminal part of

VAMP2 from the SNARE four-helix bundle. However, in full-length complexin I, such an inhibitory function of the accessory α-helix against SNARE complex formation is nearly completely suppressed by the presence of the N-terminal sequence. It is unclear how the N-terminal sequence might override the inhibition by the accessory α-helix. One possible scenario is that the N-terminal region directly interacts with the accessory α- helix to pull it out from its interaction with the SNARE complex. An alternative scenario is that the N-terminal region itself binds to the SNARE complex in a yet unknown site in such a way that it destabilizes the binding of accessory α-helix to the three-helix bundle made of t-SNAREs syntaxin 1A and SNAP-25 non-competitively.

In the previous NMR study of complexin-SNARE quaternary complex using an N- terminal truncated fragment 27 , such displacement of the membrane proximal region of

VAMP2 was not observed. A fundamental difference between EPR and NMR is in the time scales of measurement. The EPR time scale is at least 2 orders of magnitude faster than that of NMR and it can pick up fast-exchanging events. We speculate that the equilibrium exchange between the accessory α-helix and VAMP2 is sufficiently fast for

NMR not to be able to detect. Further, only ~10% insertion exacerbates the situation for the detection by NMR.

Although complexin’s ability to displace VAMP2 in the quaternary complex appears to be small, it could actually be much bigger for the trans-SNARE complex in which the interaction between membrane-anchored VAMP2 and t-SNARE on the opposite membrane is expected to be much weaker than that in our model system. In our system, the TMD of VAMP2 was deleted to avoid the situation of the cis -SNARE complex in which the TMDs of vesicle-associated SNAREs and t-SNAREs are in the same membrane, imposing no repulsion of the bilayer-bilayer interaction to the SNARE core.

The modes of interaction between complexin and the SNARE complex appear to be quite diverse. So far, three different interaction modes are discovered. Firstly, complexin has the capacity to interact with the SNARE complex additively without changing the basic coiled-coil structure of the SNARE core 27 . Such an interaction is expected to work favorably for the stabilization of the SNARE complex and believed to play a stimulatory role in SNARE-dependent membrane fusion. Secondly, the accessory α-helix displaces the C-terminal part of VAMP2 to form an alternative four-helix bundle locally while the

48 four plus one helix bundle structure is maintained in the middle 28; 34 . Our EPR results, however, show that only a small fraction of the quaternary complex has such a structure.

It is most likely that the first and the second modes are interconvertible and in equilibrium. Thirdly, complexin interacts with the t-SNARE complex, which acts as the competitive inhibitor for VAMP2 binding to the t-SNARE complex 20; 40 . The binding constant for the complexin/t-SNARE complex is reported to be somewhat higher than its affinity to the ternary SNARE complex 20 . It is highly likely that all three interaction modes exist in neuronal cells and the delicate balance between these three modes of interaction produces the inhibitory-stimulatory control of the neurotransmitter release.

For example, the knockout experiments reveal that complexin from Drosophila melanogater is highly inhibitory 22; 32 , while complexin from rat is stimulatory 19 . We speculate that such species-specific function of complexin stems from the balance between the functional strengths between the inhibitory accessory α-helix and the stimulatory N-terminal sequence.

The interaction mode in which the accessory α-helix replaces the VAMP2 while bound to the SNARE core constitutes the structural basis for the fusion clamp model 34 . Indeed, in the absence of the N-terminal sequence, the accessory α-helix has the capacity of replacing VAMP2 partially and locally from the SNARE core while the other part of the complex is fully engaged. Such local structural inhibition may be advantageous over the fully competitive inhibition relying on the t-SNARE/complexin binding because the system could react more quickly in responding to the Ca 2+ signal. Paradoxically, however, for full-length complexin, where the N-terminal sequence is present, such replacement

49 was not seen. In pure speculation, if another protein factor such as synaptotagmin I could sequester the N-terminal sequence, the inhibition of SNARE assembly might be possible, and the inhibition could be removed by the influx of Ca 2+ , which might produce synchronized membrane fusion. The results from the present analysis show that such a mechanistic model is structurally possible.

2.5 Materials and Methods

2.5.1 Plasmid construction and site-directed mutagenesis

DNA sequences encoding full-length syntaxin 1A (amino acids 4-288), the soluble

VAMP2 (amino acids 1-94), full-length complexin I (amino acids 1-134 with one native cysteine C105 replaced by alanine), and the truncated complexin I ∆26 (amino acids 27-

134) were cloned into the pGEX-KG vector and expressed as the N-terminal glutathione

S-transferase (GST) fusion protein. SNAP-25 (amino acids 1-206 with four native cysteines C85, C88, C90, and C92 replaced by alanines) was cloned into pET-28b vector and expressed as N-terminal His 6-tagged protein. All cysteine mutants were generated by

QuikChange Site-Directed Mutagenesis Kit (Stratagene), and they were confirmed by

DNA sequencing (Iowa State University DNA Sequencing Facility).

2.5.2 Protein expression, purification, and spin labeling

GST fusion proteins were expressed in Escherichia coli Rosetta (DE3) pLysS

(Novagene) and purified using glutathione-agarose beads (Sigma) 41 . Briefly, the cells were grown at 37ºC in LB with glucose (2g/l), ampicillin (100 µg/ml), and chloramphenicol (50 µg/ml) until A600 reached 0.6-0.8. After adding 0.3 mM IPTG, the cells were further grown for 6 h at 22ºC for GST-soluble VAMP2, GST-complexin I, and

GST-complexin I ∆26 but at 16ºC for GST-syntaxin 1A. Finally, the protein was cleaved to remove the GST tag by thrombin in the cleavage buffer (50mM Tris-HCl and 150 mM

NaCl, pH8.0). We add 1% n-octyl-glucoside (OG) in the cleavage buffer for syntaxin 1A.

His 6-tagged SNAP-25 was expressed in E.coli BL21 (DE3) Codon Plus RIL (Stratagene) and purified using Ni-NTA resin (Qiagen) 41 . The cells were grown at 37ºC in LB with glucose (2 g/l), kanamycin (34 µg/ml), and chloramphenicol (50 µg/ml) until A600 reached 0.6-0.8. Protein expression was induced by 0.5 mM IPTG, and the cells were grown for an additional 6h at 30ºC. Finally, the protein was eluted from the Ni-NTA resin by elution buffer (25 mM Hepes and 100mM KCl with 250 mM imidazole, pH 7.4).

The cysteine mutants of soluble VAMP2 and complexin I were reacted with (1-oxyl-2,

2, 5, 5-tetramethyl-D-pyrroline-3-methyl) methanethiosulfonate spin label at 4ºC overnight while the proteins were bound to the GST-agarose beads. To remove free spin label, the beads with bound proteins were extensively washed with the cleavage buffer and then cleaved by thrombin (Sigma). The spin-labeling efficiency was determined by

51 comparing the spin concentrations with the 50-µM 2, 2, 6, 6-tetramethyl-4-piperidine N- oxide standard. For all samples, the efficiency was ~80%.

2.5.3 Preparation of the complexin-SNARE quaternary complex

Purified His 6-tagged SNAP-25 was first added to the Ni-NTA resin solution and nutated for 1 h at room temperature. After washing out the free proteins, we mixed purified syntaxin 1A and soluble VAMP2 with His 6-SNAP-25 to form the SNARE complex with the molar ratio of 2:5:1. The mixture was incubated at 4ºC overnight. After washing one time, 2-fold excess of purified complexin I or complexin I ∆26 was added to the solution and incubated at 4ºC overnight again. After extensive washing to remove the unbound proteins, the complexin-SNARE quaternary complex was eluted with a buffer containing 250 mM imidazole and 1% OG. The formation of the complex was confirmed with SDS-PAGE gel. We obtained identical results when the quaternary complex was formed by adding all four components simultaneously.

2.5.4 EPR data collection

EPR spectra were obtained using a Bruker ESP 300 spectrometer (Bruker, Germany) equipped with a low noise microwave amplifier and a loop-gap resonator. The modulation amplitude was set at no greater than one-fourth of the line width. Spectra

52 were collected at room temperature in the first-derivative mode with 1 mW microwave power.

As for the complexin-SNARE quaternary complex in the detergent (1% OG), the samples were concentrated to the final concentration of 50-100 µM using a 5-kDa cutoff centrifugal filter (Millipore) before EPR spectra collection. For the purified complex in the membrane, the samples were reconstituted into vesicles at about 1:200 protein-to- lipid molar ratio using the published method 42 . The large unilamellar vesicles (~100nm in diameter) of 1-palmitoyl-2-oleoyl-sn -glycero-3-phosphatidylserine were prepared in a detergent-free buffer using an extruder (Avanti). The detergent was removed by treating the sample with the dialysis buffer (25 mM Hepes and 100 mM KCL, pH 7.4) at 4ºC overnight. After dialysis, the sample solution was concentrated using a 100-kDa cutoff centrifugal filter (Millipore) before taking an EPR spectrum.

2.6 References

1. Sollner, T., Bennett, M. K., Whiteheart, S. W., Scheller, R. H. & Rothman, J. E. (1993). A protein assembly-disassembly pathway in vitro that may correspond to sequential steps of synaptic vesicle docking, activation, and fusion. Cell 75 , 409- 18.

2. Weber, T., Zemelman, B. V., McNew, J. A., Westermann, B., Gmachl, M., Parlati, F., Sollner, T. H. & Rothman, J. E. (1998). SNAREpins: minimal machinery for membrane fusion. Cell 92 , 759-72.

3. Sutton, R. B., Fasshauer, D., Jahn, R. & Brunger, A. T. (1998). Crystal structure of a SNARE complex involved in synaptic exocytosis at 2.4 A resolution. Nature 395 , 347-53.

4. Poirier, M. A., Xiao, W., Macosko, J. C., Chan, C., Shin, Y. K. & Bennett, M. K. (1998). The synaptic SNARE complex is a parallel four-stranded helical bundle. Nat Struct Biol 5, 765-9.

5. Jahn, R. & Scheller, R. H. (2006). SNAREs--engines for membrane fusion. Nat Rev Mol Cell Biol 7, 631-43.

6. Jahn, R., Lang, T. & Sudhof, T. C. (2003). Membrane fusion. Cell 112 , 519-33.

7. Yoon, T. Y. & Shin, Y. K. (2009). Progress in understanding the neuronal SNARE function and its regulation. Cell Mol Life Sci 66 , 460-9.

8. Fernandez, I., Arac, D., Ubach, J., Gerber, S. H., Shin, O., Gao, Y., Anderson, R. G., Sudhof, T. C. & Rizo, J. (2001). Three-dimensional structure of the synaptotagmin 1 C2B-domain: synaptotagmin 1 as a phospholipid binding machine. Neuron 32 , 1057-69.

9. Chapman, E. R. (2002). Synaptotagmin: a Ca(2+) sensor that triggers exocytosis? Nat Rev Mol Cell Biol 3, 498-508.

10. Koh, T. W. & Bellen, H. J. (2003). Synaptotagmin I, a Ca2+ sensor for neurotransmitter release. Trends Neurosci 26 , 413-22.

11. Saraswati, S., Adolfsen, B. & Littleton, J. T. (2007). Characterization of the role of the Synaptotagmin family as calcium sensors in facilitation and asynchronous neurotransmitter release. Proc Natl Acad Sci U S A 104 , 14122-7.

12. Perin, M. S., Fried, V. A., Mignery, G. A., Jahn, R. & Sudhof, T. C. (1990). Phospholipid binding by a synaptic vesicle protein homologous to the regulatory region of protein kinase C. Nature 345 , 260-3.

13. Petrenko, A. G., Perin, M. S., Davletov, B. A., Ushkaryov, Y. A., Geppert, M. & Sudhof, T. C. (1991). Binding of synaptotagmin to the alpha-latrotoxin receptor implicates both in synaptic vesicle exocytosis. Nature 353 , 65-8.

14. Davletov, B. A. & Sudhof, T. C. (1993). A single C2 domain from synaptotagmin I is sufficient for high affinity Ca2+/phospholipid binding. J Biol Chem 268 , 26386-90.

15. Chen, Y. A., Scales, S. J., Patel, S. M., Doung, Y. C. & Scheller, R. H. (1999). SNARE complex formation is triggered by Ca2+ and drives membrane fusion. Cell 97 , 165-74.

16. Dai, H., Shen, N., Arac, D. & Rizo, J. (2007). A quaternary SNARE- synaptotagmin-Ca2+-phospholipid complex in neurotransmitter release. J Mol Biol 367 , 848-63.

17. Tucker, W. C., Weber, T. & Chapman, E. R. (2004). Reconstitution of Ca2+- regulated membrane fusion by synaptotagmin and SNAREs. Science 304 , 435-8.

18. Wang, C. T., Bai, J., Chang, P. Y., Chapman, E. R. & Jackson, M. B. (2006). Synaptotagmin-Ca2+ triggers two sequential steps in regulated exocytosis in rat PC12 cells: fusion pore opening and fusion pore dilation. J Physiol 570 , 295-307.

19. Reim, K., Mansour, M., Varoqueaux, F., McMahon, H. T., Sudhof, T. C., Brose, N. & Rosenmund, C. (2001). Complexins regulate a late step in Ca2+-dependent neurotransmitter release. Cell 104 , 71-81.

20. Yoon, T. Y., Lu, X., Diao, J., Lee, S. M., Ha, T. & Shin, Y. K. (2008). Complexin and Ca2+ stimulate SNARE-mediated membrane fusion. Nat Struct Mol Biol 15 , 707-13. 21. Tang, J., Maximov, A., Shin, O. H., Dai, H., Rizo, J. & Sudhof, T. C. (2006). A complexin/synaptotagmin 1 switch controls fast synaptic vesicle exocytosis. Cell 126 , 1175-87.

22. Huntwork, S. & Littleton, J. T. (2007). A complexin fusion clamp regulates spontaneous neurotransmitter release and synaptic growth. Nat Neurosci 10 , 1235-7.

23. McMahon, H. T., Missler, M., Li, C. & Sudhof, T. C. (1995). Complexins: cytosolic proteins that regulate SNAP receptor function. Cell 83 , 111-9.

24. Ishizuka, T., Saisu, H., Odani, S. & Abe, T. (1995). Synaphin: a protein associated with the docking/fusion complex in presynaptic terminals. Biochem Biophys Res Commun 213 , 1107-14.

25. Reim, K., Wegmeyer, H., Brandstatter, J. H., Xue, M., Rosenmund, C., Dresbach, T., Hofmann, K. & Brose, N. (2005). Structurally and functionally unique complexins at retinal ribbon synapses. J Cell Biol 169 , 669-80.

26. Pabst, S., Margittai, M., Vainius, D., Langen, R., Jahn, R. & Fasshauer, D. (2002). Rapid and selective binding to the synaptic SNARE complex suggests a modulatory role of complexins in neuroexocytosis. J Biol Chem 277 , 7838-48.

27. Chen, X., Tomchick, D. R., Kovrigin, E., Arac, D., Machius, M., Sudhof, T. C. & Rizo, J. (2002). Three-dimensional structure of the complexin/SNARE complex. Neuron 33 , 397-409.

28. Xue, M., Reim, K., Chen, X., Chao, H. T., Deng, H., Rizo, J., Brose, N. & Rosenmund, C. (2007). Distinct domains of complexin I differentially regulate neurotransmitter release. Nat Struct Mol Biol 14 , 949-58.

29. Maximov, A., Tang, J., Yang, X., Pang, Z. P. & Sudhof, T. C. (2009). Complexin controls the force transfer from SNARE complexes to membranes in fusion. Science 323 , 516-21.

30. Archer, D. A., Graham, M. E. & Burgoyne, R. D. (2002). Complexin regulates the closure of the fusion pore during regulated vesicle exocytosis. J Biol Chem 277 , 18249-52.

31. Itakura, M., Misawa, H., Sekiguchi, M., Takahashi, S. & Takahashi, M. (1999). Transfection analysis of functional roles of complexin I and II in the exocytosis of two different types of secretory vesicles. Biochem Biophys Res Commun 265 , 691-6.

32. Schaub, J. R., Lu, X., Doneske, B., Shin, Y. K. & McNew, J. A. (2006). Hemifusion arrest by complexin is relieved by Ca2+-synaptotagmin I. Nat Struct Mol Biol 13 , 748-50.

33. Giraudo, C. G., Eng, W. S., Melia, T. J. & Rothman, J. E. (2006). A clamping mechanism involved in SNARE-dependent exocytosis. Science 313 , 676-80.

34. Giraudo, C. G., Garcia-Diaz, A., Eng, W. S., Chen, Y., Hendrickson, W. A., Melia, T. J. & Rothman, J. E. (2009). Alternative zippering as an on-off switch for SNARE-mediated fusion. Science 323 , 512-6.

35. Melia, T. J., Jr. (2007). Putting the clamps on membrane fusion: how complexin sets the stage for calcium-mediated exocytosis. FEBS Lett 581 , 2131-9.

36. Xue, M., Stradomska, A., Chen, H., Brose, N., Zhang, W., Rosenmund, C. & Reim, K. (2008). Complexins facilitate neurotransmitter release at excitatory and inhibitory synapses in mammalian central nervous system. Proc Natl Acad Sci U S A 105 , 7875-80.

37. Giraudo, C. G., Garcia-Diaz, A., Eng, W. S., Yamamoto, A., Melia, T. J. & Rothman, J. E. (2008). Distinct domains of complexins bind SNARE complexes and clamp fusion in vitro. J Biol Chem 283 , 21211-9.

38. McHaourab, H. S., Lietzow, M. A., Hideg, K. & Hubbell, W. L. (1996). Motion of spin-labeled side chains in T4 lysozyme. Correlation with protein structure and dynamics. Biochemistry 35 , 7692-704.

39. Kweon, D. H., Kim, C. S. & Shin, Y. K. (2003). Regulation of neuronal SNARE assembly by the membrane. Nat Struct Biol 10 , 440-7.

40. Weninger, K., Bowen, M. E., Choi, U. B., Chu, S. & Brunger, A. T. (2008). Accessory proteins stabilize the acceptor complex for synaptobrevin, the 1:1 syntaxin/SNAP-25 complex. Structure 16 , 308-20.

41. Zhang, Y., Su, Z., Zhang, F., Chen, Y. & Shin, Y. K. (2005). A partially zipped SNARE complex stabilized by the membrane. J Biol Chem 280 , 15595-600.

42. Chen, Y., Xu, Y., Zhang, F. & Shin, Y. K. (2004). Constitutive versus regulated SNARE assembly: a structural basis. Embo J 23 , 681-9.

2.7 Figures and Captions

Figure 1 Characterization of the complexin-SNARE quaternary complex. (a) The fusion clamp model proposes that complexin can replace the C-terminus of VAMP2 upon binding to the trans -SNARE complex (adapted from Ref. 34). Positions 69, 76, 83, 86, and 89 are the spin-labeled sites of the soluble VAMP2. Positions 35 and 28 of complexin have also been shown in the model. Positions 42, 25, and 22, which are located at the N-terminus of complexin, have not been shown. (b) The flow diagram of purification of the complexin-SNARE quaternary complex. (c) SDS-PAGE analysis of the purified complexin-SNARE quaternary complex.

Figure 2 EPR spectra of spin-labeled soluble VAMP2 in the full-length complexin- SNARE quaternary complex at room temperature. First-derivative mode EPR spectra for the complex in the detergent (a) and in the membrane (b) are shown.

Figure 3 EPR spectra of the spin-labeled complexin in the full-length complexin- SNARE quaternary complex at room temperature . First-derivative mode EPR spectra for the complex in the detergent and in the membrane are shown. Positions 22, 25, 28, 35, and 42 of complexin correspond to positions 89, 86, 83, 76, and 69 of VAMP2, respectively.

Figure 4 (a and b) EPR spectra of the spin-labeled soluble VAMP2 in the truncated complexin ∆26-SNARE quaternary complex at room temperature. First-derivative mode EPR spectra for the complex in the detergent (a) and in the membrane (b) are shown. The arrowheads indicate the sharp component from the locally displaced VAMP2. The percentage of the sharp component was calculated using the spectral subtraction method (ref. 39). (c), EPR spectra of the spin-labeled full-length and truncated complexin upon binding with the ternary SNARE complex at room temperature. The inset figures show the difference between full-length and truncated complexin involved in the quaternary complex much more clearly.

2.8 Supplementary Data

Figure S1 . EPR spectra of the spin-labeled soluble VAMP2 in the complexin- SNARE quaternary complex at room temperature when adding more complexin . First-derivative mode EPR spectra for the complex in the membrane are shown. Two-fold molar excess of complexin are added into the purified complexin-SNARE quaternary complex. There are still broad spectra at positions 83,86, and 89 of VAMP2.

CHAPTER 3: LINKER REGION OF NEURONAL SNARE COMPLEX FORMS INDEPENDENT α-HELICES WHEN SNARE CORE ASSEMBLY IS COMPLETED*

Shuang Song†, Xiaochu Lou†, Jaeil Shin and Yeon-Kyun Shin

3.1 Abstract

Neurotransmitter release relies on the membrane fusion between synaptic vesicles and the membrane of nerve cells, which is a process requiring the assembly of several SNARE (soluble N-ethylmaleimide-sensitive factor attachment protein receptor) proteins: VAMP2 (vesicle associated membrane protein 2) on the synaptic vesicle and syntaxin 1A and SNAP-25 on the plasma membrane of the nerve cell. These proteins are believed to form in trans between the two membranes as a four-helix bundle which can finally drive the opposite membranes together. However, it is still unclear where the energy needed to overcome the repulsive forces is generated and transferred to the fusion pore and how the conformational change of SNARE complex is linked to fusion initiation. Here, we report the results of a thorough EPR investigation of the SNARE complex linker-region structure. The results show that the linker region of syntaxin and VAMP2 in a SNARE complex can independently form helical extension beyond the core SNARE complex only if the SNARE core four-helix bundle is completely assembled. These results provide further evidence for the hypothesis that the energy generation and transduction is operated stepwise and coupled with corresponding conformational change of SNARE complex. The final step of SNARE assembly will directly lead to membrane merger.

*This paper is being prepared for submission. Necessary modifications are made to fit the format of this dissertation. †Contributed equally to this work. Author contributions: X.C.L., and S.S. performed research and analyzed data; Y.K.S. designed research and wrote the paper.

3.2 Introduction

Membrane fusion is a critical step which is required in a large variety of cellular processes-including neurotransmitter release, protein transportation and fertilization 1; 2 . And the biological system has specialized proteins to fulfill this task 3; 4; 5; 6; 7 . SNARE proteins form a superfamily characterized by conserved stretches of 60-70 amino acids arranged in heptad repeats called SNARE motifs 1; 8; 9; 10 . In most SNAREs, there is a single transmembrane domain C-terminally connected to this SNARE motif by a short linker region 11; 12 . Different intracellular transportations are considered to be operated by different members of the SNARE family 13; 14; 15 . While in the process of neurotransmitter release, syntaxin 1A, VAMP2 and SNAP-25 can form the neuronal SNARE complex which acts as the minimal fusion apparatus 16; 17; 18 . Both of syntaxin 1A and VAMP 2 have a linker region between their SNARE motifs and a C-terminal transmembrane domain. SNAP-25 is attached to the surface of plasma membrane by palmitoylation through the cysteine residues in the loop region between its two SNARE motifs. Individual SNARE proteins are largely unstructured, but spontaneously assemble into a stable SDS-resistant ternary complex which has a melting temperature as high as 90ºC19; 20; 21; 22 . X-ray structure of this SNARE core complex has been revealed as a four parallel α-helical bundle with each helix contributed by one SNARE motif 23 . The assembly of this synaptic fusion complex hypothetically drives the vesicles and target membranes together and provides the energy required to conquer the repulsion between the opposite membranes during membrane fusion 22; 24 .

Assembly of SNARE complexes is believed to initiate from the N-terminus and proceed to the C terminus progressively as stated in the zipper model 25; 26 . However, there is no hard proof of this model but indirect evidence 25; 26; 27; 28; 29; 30; 31 . Interestingly, EPR study on yeast SNAREs proposes the possibility that the coiled-coil structure of the C- terminal part of SNARE motif can be interfered by the membrane and becomes partially unstructured which makes the SNARE motif less likely to serve as the energy source for the fusion 32 and this is different from what has been known in neuronal SNAREs.

If the zippering of SNARE four-helix bundle is responsible for providing and transducing energy to overcome the repulsive force between two opposing membranes, it is necessary that the helical structure remains intact all the way through the fusion pore to avoid dissipating the energy 23; 33 . Crystal structure of SNARE complex formed with full length syntaxin 1A and VAMP2 has also been resolved which reported helical extension throughout the linkers and transmembrane domains in cis -SNARE complex 33 . But there remains a question that whether the helical extension is formed along zippering proceeding or it has already been formed beforehand. And how is the coiled coil formation energetically coupled to membrane merger. Previous EPR analysis has reported that the linker region of syntaxin 1A is unstructured but laterally inserted into the membrane in a mimic trans-SNARE complex 34; 35 . This insertion results in a firm grip on to the membrane and drags the two membranes into contact with the force produced by SNARE core assembly and also leads to the deformation of the membrane.

To further investigate the structure and membrane topology of the SNARE complex using site-directed spin labeling EPR 36; 37 and power saturated accessibility measurements 38 . We conducted consecutive cysteine mutants at the linker regions of syntaxin 1A and VAMP2. EPR spectra revealed a general α-helical structure of both syntaxin 1A and VAMP2 linker regions in solution. Interestingly, even when one of the linker regions was removed, it didn’t affect the general structure of the other one. However, when VAMP2 was further truncated in the middle of its SNARE motif, the structure of syntaxin 1A linker region was significantly affected. The power saturated accessibility measurement generated similar structural information for single linker region in absence of the other linker as our previous experiment did which involved both of the linker regions 34; 39 . Thus, our results showed that for a trans -SNARE state in both solution and membrane, the linker regions of syntaxin1A and VAMP2 can form independent helical structure once the SNARE core motif is fully assembled which indicated a direct coupling of the four-helix bundle formation and energy transduction to the membrane interface.

3.3 Results

3.3.1 SNARE core complex formation

To investigate the linker region structure of SNARE complex using site-directed spin labeling EPR, residues of full-length syntaxin 1A and VAMP2 in the corresponding linker regions were replaced with cysteines to facilitate a further attachment of nitroxide spin label through disulfide bond. We prepared 10 cysteine mutants (K256C-K265C) of syntaxin1A and 7 cysteine mutants (W89C-M95C) in their linker regions. The cysteine mutants were purified and derivatized with methanethiosulfonate spin label [(1-oxyl- 2,2,5,5-tetramethyl-D-pyrroline-3-methyl) methanethiosulfonate for EPR measurements. Two truncated versions of synatxin 1A and three of VAMP2 were generated as shown in Figure 1 which would be used in our following study about the independency of linker region structure. To investigate the linker region structure of syntaxin 1A, the recombinant complex was firstly assembled from full length syntaxin1A, soluble VAMP2 with linker region residues Met 1-Met 96 (VpS) or soluble VAMP2 without linker region residues Met 1- Leu 84 (VpDL) and SNAP-25 (synaptosome-associated protein of 25 kDa). His 6-tagged SNAP-25 was attached to the nickel-nitrilotriacetic acid (Ni-NTA) beads as the bait and the SNARE complex was formed by flowing syntaxin 1A and soluble VAMP2 into the bead solution and incubating at 4ºC overnight. Unbound portion of syntaxin1A and soluble VAMP2 was eluted from the column to avoid extra free signals generated by unbound syntaxin 1A. The SNARE complex formed was eluted by buffer containing 0.8% n-nonyl β-D-glucopyranoside (OG). For each mutant, the formation of SNARE complex was confirmed by the SDS-page gel after heating. It turned out that the immobilized His 6-tagged SNAP-25 co-purified with stoichiometric amounts of syntaxin 1A and soluble VAMP2.

3.3.2 The absence of VAMP2 linker region does not change structure of syntaxin 1A linker region

The EPR line shape is a sensitive function of the nitroxide side chain tumbling rate. A broaden EPR spectrum indicates a slower tumbling rate which might due to the restriction of interaction and local environment. EPR spectra taken at room temperature of the SNARE complex with spin labeling on syntaxin 1A linker region with either soluble VpS or soluble VpDL were shown in Figure 2 . Therefore, if the linker region structure formation of syntaxin 1A is related to the linker region of VAMP2, we should expect observing differences of the EPR spectra for the spin-labeled positions between SNARE complexes formed by VpS and VpDL, probably narrower spectra for the VpDL SNARE complexes due to the lack of VAMP2 linker region. This VpDL version of SNARE complex is supposed to represent an earlier stage of SNARE zippering compared to the VpS version. However, the spectra for positions 256-265 of syntaxin 1A with VpS and VpDL are barely different from each other. Both of them showed similar degree of line broadening which indicated slower motion of the nitroxide, most likely due to a secondary α-helical structure and potential tertiary interactions between side chains. But we cannot conclude that the linker region of syntaxin 1A has an α-helix conformation without further evidence. We can just infer that the linker region structure of syntaxin 1A remains the same if the VAMP2 linker region is removed.

3.3.3 The linker region of syntaxin 1A has no obvious structure change in trans- SNARE and cis-SNARE complexes.

To examine if this linker region structure of syntaxin 1A will remain the same till the post-fusion stage in a cis -SNARE complex, we also performed experiments with the same set of syntaxin 1A linker region mutants and full-length version of VAMP2 (VpFL). We aligned these EPR spectra with the corresponding ones with VpDL and the results are shown in Figure 3 . Interestingly, the spectra for those labeling positions of syntaxin1A in

SNARE complexes formed with VpDL and VpFL coincided with each other very well, indicating that the syntaxin 1A linker region structure didn’t change significantly during the two transmembrane domains aligning and membrane merging. In another word, the structure formation of syntaxin 1A linker region has already completed as early as the SNARE motif zippering is completed at a trans -SNARE stage. Crystal structure of SNARE complex including both linker regions and transmembrane regions of syntaxin 1A and VAMP2 has already revealed a continuous helical extension structure beyond the SNARE core and all the way through the transmembrane domains 33 . Therefore, we propose that in our trans -SNARE complex, syntaxin 1A has already formed similar linker region α-helical extension.

3.3.4 The helical-extension of linker region depends on fully zippering of the SNARE motifs.

We now examine an earlier stage of SNARE zippering by using a further truncated version of VAMP2, VpDS. VpDS is truncated near the center of its SNARE motif, which is supposed to be incapable of forming a complete SNARE core complex with full length syntaxin 1A and SNAP25, and therefore representing an N-terminal partially zippered SNARE complex. EPR spectra for the same set of spin-labeled syntaxin 1A in this partially zippered SNARE complex formed by VpDS were aligned with those spectra with VpS and shown in Figure 4 . The EPR spectra line shape is a reflection of the motion for the nitroxide side chain and is related to three kinds of movements: the rotation of the entire protein; the rotational isomerization about the bonds that link the nitroxide to the backbone; the segmental motion of the backbone relative to the average protein structure. EPR spectra for different spin-labeled positions representing distinct topographical sites have been well-established in T4 lysozyme whose three-dimensional structure is well-know (Mchaourab et al., 1996). In fact, our EPR spectra for the spin- labeled syntaxin 1A in SNARE complex with VpDS were very similar to those for the solvent exposed positions on the surface helix revealed in this study which were supposed to be indicative of substantial motion of an accessory α-helix. That implied that

68 even at a relative early stage in which a complete SNARE core complex has not been formed yet, the linker region of syntaxin 1A has already accomplished its secondary structure formation as a hypothetical α-helix. And this result is consistent with a published crystallization data which also claimed that the syntaxin 1A α-helices are almost fully formed even in the absence of the VAMP2 C-terminus 40 . However, we also observed obvious differences between the EPR lines for syntaxin 1A linker region in SNARE complex formed by VpS and VpDS. In the cases of VpDS, the EPR spectra tended to lose their broad component starting from position 260, which might reflect a loss of potential tertiary interaction compared with the corresponding VpS cases. These results suggested that the linker region of syntaxin 1A had a relatively free movement pattern with less restriction in this N-terminal partially zippered SNARE complex. But the remaining question is which part of the rest of the SNARE complex syntaxin 1A linker region interacts with when SNARE core complex is fully zippered. For nitroxides attached to positions 256-259, the EPR spectra were very similar to those obtained from the SNARE complex formed by VpS. According to our previous study 41 , a coiled-coil structure of syntaxin1A was maintained up to residue 259 in a ternary complex. This means that the upstream part preceding the progressively more mobile region toward the C-terminal end of syntaxin 1A remains its hypothetical coiled-coil structure in an incompletely zippered SNARE complex. Meanwhile, the completion of SNARE core zippering will bring an extra ternary interaction to the C-terminal part of syntaxin 1A linker region.

3.3.5 The linker region of VAMP2 has no obvious structure change in SNARE complex with and without syntaxin 1A linker region.

Besides the syntaxin 1A side, we are also interested in what happens on VAMP2 side when syntaxin 1A linker region is removed. To investigate the linker region structure of VAMP2, the recombinant complex was assembled from spin-labeled full length VAMP2, soluble syntaxin1A with linker region residues Ala 191- Ile 266 (SynSH3) or soluble syntaxin 1A without linker region residues Ala 191- Lys 256 (SynDL) and SNAP-25

(synaptosome-associated protein of 25 kDa). The two sets of EPR spectra were aligned and plotted in Figure 5. Similarly, if the linker region conformation of VAMP2 is related to the linker region of syntaxin1A, we should expect observing differences of the EPR spectra for the spin-labeled positions between SNARE complexes formed by SynSH3 and SynDL, probably narrower spectra for the SynDL SNARE complexes due to the failure of helical structure formation of the VAMP linker region. Same as what we observed on the syntaxin 1A side, the spectra for position 86-95 of VAMP2 turned out to be very similar in SNARE complexes with SynSH3 and SynDL in general shape. This result was not as surprising as the previous ones because it has already been shown that the membrane-proximal linker region of VAMP2 alone can form a helical segment of two-turn, which is considered as an important regulatory module for SNARE assembly. This study also suggested a possibility of SNARE complex formation initiated at the C- terminal region which was contradictory to what we aimed to illustrate here. Interestingly, for position 86, 87, 88, 92 and 95, we observe varying degrees of an extra broaden component in the EPR spectra for the SNARE complex formed by full length VAMP2. Notice that the crystallization study 33 of cis -SNARE complex showed that four VAMP2 transmembrane domains built the core of a four SNARE complexes association surrounded by four syntaxin 1A transmembrane domains and there were multiple intermolecular side chain contacts between the linker regions. Position 86, 88, 92 and 95 are supposed to be just the sites of forming those interactions with syntaxin 1A linker regions. Obviously, removal of the syntaxin 1A linker region may result in losing those tertiary interactions which leads to a narrowed EPR spectra. So at this point, we can just conclude that the full length VAMP2 linker region shows similar secondary structural characteristics in either VMAP2 monomer, or in SNARE complexes with and without syntaxin 1A linker region.

3.3.6 The absence of syntaxin 1A linker region does not change structure of VAMP2 linker region in a soluble SNARE complex without both transmembrane domains.

Since the previous circular dichrosim (CD) study suggested that the existence of VAMP2 transmembrane domain might interfere the packing of the four helix bundle further upstream 33 , we wanted to further investigate that whether the involvement of VAMP2 transmembrane domain contributed to stabilize the linker region structure or hinder the coiled-coil formation. We collected another set of EPR spectra for spin labeled soluble VAMP2 (VpS) in SNARE complexes formed by SynH3 or SynDL. However, spectra for position 89 and 95 are missing here. As the results shown in Figure 6, EPR spectra for VAMP2 linker region in a soluble SNARE complex still exhibited a highly restricted movement pattern which was very similar to what we observed in the full length VAMP2 case. And the absence of syntaxin 1A linker region also didn’t show significant effect on VAMP2 linker region structure formation in this context. Although this experiment didn’t provide further support of what has been observed in the CD experiment, it did contribute to confirm an N- to C-terminal direction progressively zippering model of SNARE complex.

3.3.7 EPR accessibility measurements study further confirms the α-helical structure of syntaxin1A and VAMP2 linker region.

All of the evidence we have at hand up to now only pointed to the fact that syntaxin 1A and VAMP2 linker regions have secondary structure or a potential tertiary interaction with some other unknown part of the SNARE complex. A hypothesis of α-helical structure seems to be legitimate based on our result and previous studies. Although the EPR line shape itself is a useful parameter for the tumbling rate of the nitroxide that can provide a qualitative view of the local environment for the labeling position, we still need a more powerful tool to validate our α-helical model and the EPR power saturation method is a natural choice. It will not only generate more detailed information about the secondary and tertiary structures, but also about the membrane topology.

For the nitroxide on the labeling reagent, the EPR saturation method measures the accessibility to the water-phase paramagnetic reagent such as NiEDDA (W NIEDDA ) to

71 estimate the solvent exposure of the spin-labeled site. On the other hand, we also measure the accessibility to membrane-phase nonpolar paramagnetic reagent such as molecular oxygen (W O2) to estimate, for example, the insertion into the membrane of the side chain 42; 43; 44 .

SNARE complex samples were prepared using the same procedure as stated previously. The SNARE complex obtained in solution with 0.8% detergent was then reconstituted into vesicles composed by 45 mol% 1-palmitoyl-2-oleoylphosphatidylcholine (POPC), 15 mol% dioleoylphosphatidylserion (DOPS) and 40 mol% cholesterol, a lipid composition used to mimic the native cellular membrane 45 . EPR saturation experiments were performed for all spin-labeled variants under nitrogen, under air, and in the presence of NiEDDA under nitrogen. The standard EPR spectra are also collected in room temperature, under air and at 1mW incident microwave power, same condition as all other EPR spectra shown previously to make sure that the complexes are successfully formed. The EPR spectra for spin-labeled full length syntaxin 1A in SNARE complex with VpS or VpDL are shown in Figure 7 . Those for spin-labeled full length VAMP2 in SNARE complex with SynFL or SynDL are shown in Figure 8 . All of the EPR spectra are broad and even broader than those we observed for the same composition of SNARE complex but in solution state. Some of the sharp components in solution which might represent a freely moving subpopulation just disappeared when the SNARE complex is reconstituted into vesicles. Moreover, we observe increased overall breadth of the spectrum along the magnetic field axis and even a hyperfine line in high magnetic field, which all indicated highly immobilized nitroxides 46 . Similar slow motional spectra are commonly observed for spin-labeled proteins bound to the membrane 47; 48; 49 . Therefore, we have evidence to propose that the membrane contributed to further stabilize the SNARE complex linker region. In fact, our previous study has already shown that both syntaxin 1A and VAMP2 linker regions were inserted into the membrane.

A set of example EPR saturation curves for one of the spin-labeled sites under various power conditions are shown in Figure 9. The W NiEDDA and W O2 values are the coefficients obtained by fitting those curves with a specific exponential function. Then,

72 the accessibility values were normalized by the corresponding central line width of the spectra taken under air at 1mW. The normalized accessibility parameters for the labeled syntaxin 1A in SNARE complexes formed by VpS or VpDL were plotted against the corresponding residue number, respectively in Figure 10. Same kind of plot for spin- labeled VAMP2 with SynH3 or SynDL is shown in Figure 11. The observed accessibility of the nitroxide to NiEDDA was much lower than typical values for fully solvent-exposed nitroxides which was expected to be as high as 40 under the same conditions 42 . But the accessibility parameters before normalization were around 15~25 in our experiments. This suggested that the nitroxides attached to the SNARE complex were not exposed to the water phase. Again, this indicated that the side chains could be either buried in the protein environment or inserted into the membrane, which was consistent with what we inferred from the broad EPR line shapes. In addition, in both Figure 10 and Figure 11, the accessibility parameters were not significantly different in the left panel and the right panel either in absolute values or general pattern. That further confirmed our previous conclusion that the linker regions for syntaxin 1A and VAMP2 were structured independently without the involvement of each other. We were also aware of that there are significant increases and decreases for W NiEDDA along both the syntaxin 1A and VAMP2 linker region sequence, which might imply an α-helix structure. But at this point, we don’t want to draw a conclusion about the α-helical structure.

3.4 Discussion

Both the EPR spectra and the power saturation accessibility measurement experiments revealed a capacity of syntaxin 1A and VAMP2 linker regions to independently form a robust secondary structure, perhaps α-helix. And this structured linker region is stable in either solution state or in the membrane. Although our previous study has suggested a possibility that the linker region of syntaxin 1A might be unstructured and bound laterally to the membrane and the broad EPR spectra were contributed by interaction with the membrane 34 , our conclusion might be slightly different because we were also able to

73 observe slow motion EPR spectra of SNARE complex in solution state which could merely due to interaction from the protein environment. However, both of these results address to rule out the possibility that a flexible tether existents between the SNARE core and the membrane, and therefore confirm the function of SNARE complex linker region as the critical energy transducer.

Our discovery of independently forming α-helices is well related with the crystal structure of a post-fusion state cis -SNARE complex which revealed the continuous helices beyond the core SNARE complex throughout the linker regions and transmembrane domains. These α-helices in a trans -SNARE state may continue to zip up with the assistance of side-chain interactions in the linker region, then directly bring the membrane merger. Previous EPR distance measurement also revealed that in some non- functional SNARE complexes like t-SNARE complex and the syntaxin 1A dimer, the interfacial linker region is disordered 35 . Although, homotypic membrane fusion has also been found in some studies, the rate and efficiency is much lower 50; 51; 52 . That also reinforces the significance of a readily structured linker region in optimum fusion machinery. So our results further verified the N- to C-terminal direction zippering model and provided powerful evidence to explain the mechanism of energy generation and transduction for membrane fusion.

Interestingly, we were also able to observe a ternary interaction between syntaxin 1A linker region and some other part inside the SNARE complex which happened only if the SNARE core complex was completely zippered. But the remaining question is which part of the SNARE complex this syntaxin 1A linker region interacts with. Previous studies have shown that the cleavage of SNAP25 SN2 C-terminus by neurotoxins (BoNT/A and BoNT/E) will significantly block neurotransmitter release in vivo 18; 53; 54 . So it has been proposed that the C-terminal truncated SNAP25 mutants will disrupt the assembly of SNARE core complex and then inhibit the initial fusion pore opening. Therefore it is also likely that the engagement of SNAP25 C-terminus promotes fusion by interacting with the linker region of syntaxin 1A and contributes to stabilize and tighten up the coiled coil structure up to the linker region. Our previous study has pointed out that in a binary

SNARE complex, the C-terminus of SNAP25 SN1 domain was fully assembled but the C-terminus of SN2 domain was not completely involved (unpublished data). The complete formation of ternary SNARE complex included the recruitment of SNAP25 C- terminus by VAMP2 first. In our experiment, the SNARE complex formed with VpDS which was truncated in the central layer of SNARE motif might partially or completely lose its ability to recruit the SN2 C-terminus, and therefore SN2 C-terminus failed to establish its connection with syntaxin 1A linker region. The interaction with SN2 C- terminus might contribute to strengthen the secondary structure of SNARE complex linker region and tightly control the energy transfer. So removal of the SN2 C-terminus highly affected the fusion activity.

Additionally, the extra ternary interaction with the membrane is very important in the coupling of energy transfer and membrane juxtaposition. Only if the linker region α- helices are laterally bound to the membrane, it is possible for them to physically bring the membranes into close proximity and finally accomplish the task of transferring the energy to the site of membrane decomposition initiated. This kind of membrane bound pattern has been found by EPR immersion depth measurement in both syntaxin and VAMP2 linker regions from either binary or ternary SNARE complex.

Based on our EPR spectra analysis, it seems that the potential α-helical structure of the linker regions have been readily formed even before the core complex is completely zippered. If that is true, it will highly improve our understanding about how could the Ca 2+ -triggered fusion happen within millisecond scale. The structured linker region might provide a scaffold for the binding of late regulatory proteins beforehand such as synaptotagmin and complexin to guarantee that all the accessories are ready to operate whenever the Ca 2+ signaling comes into play. Actually, we also hypothesized that complexin binding to the SNARE machinery might hinder the continuity of syntaxin 1A linker region, therefore acted as a fusion clamp to inhibit at the docking stage. But the highly stable nature of SNARE complex in solution state will override this disruption effect of complexin. It is highly necessary for us to build up a more flexible system which

75 can better mimic the situation of a trans -SNARE complex in order to catch up this kind of intermediate states.

The one step ahead feature of the SNARE complex linker region structure formation may largely improve the efficiency of SNARE complex fully assembly and membrane fusion. It also enables early involvement of regulatory factors. Our study still has several weaknesses that may need to be addressed by further experimental data. Firstly, the results to support an α-helical structure is not conclusive, we still need other powerful evidence to confirm this hypothesis. Secondly, the ternary interaction mode of the linker region with C-terminal SNAP25 SN2 domain is purely hypothetical. There still exists other possible interaction modes like association of multiple SNARE complexes linker regions 55; 56 . To further investigate when this secondary linker region structure was initiated, we may still need to trace back further to even earlier steps in SNARE core complex zippering. To summarize, SNARE complex acts not only like a fusion operator but also a distributing center of all fusion related factors. Regulatory proteins come on- and-off recursively to guarantee the continuity of membrane fusion activity.

3.5 Materials and Methods

3.5.1 Plasmid construction and site-directed mutagenesis

DNA sequences encoding full-length syntaxin 1A (amino acids 4-288), the soluble SynSH3 (amino acids 191-266), soluble SynDL (amino acids 191-256), the full-length VAMP2 (amino acids 1-116), soluble VpS (amino acids 1-96), soluble VpDL (amino acids 1-84), soluble VpDS (amino acids 1-60) were cloned into the pGEX-KG vector and expressed as the N-terminal glutathione S-transferase (GST) fusion protein. SNAP-25 (amino acids 1-206 with four native cysteines C85, C88, C90, and C92 replaced by alanines) was cloned into pET-28b vector and expressed as N-terminal His 6-tagged protein. All cysteine mutants were generated by QuikChange Site-Directed Mutagenesis

Kit (Stratagene), and they were confirmed by DNA sequencing (Iowa State University DNA Sequencing Facility).

3.5.2 Protein expression, purification, and spin labeling

GST fusion proteins were expressed in Escherichia coli Rosetta (DE3) pLysS (Novagene) and purified using glutathione-agarose beads (Sigma) 32 . Briefly, the cells were grown at 37ºC in LB with glucose (2g/l), ampicillin (100 µg/ml), and chloramphenicol (50 µg/ml) until A600 reached 0.6-0.8. After adding 0.3 mM IPTG, the cells were further grown for 6 h at 22ºC for GST-VpS, GST-VpDL, GST-VpDS, GST- SynSH3 and GST-SynDL but at 16ºC for GST-syntaxin 1A and GST-VAMP2. Finally, the protein was cleaved to remove the GST tag by thrombin in the cleavage buffer (50mM Tris-HCl and 150 mM NaCl, pH8.0). We add 0.8% n-octyl-glucoside (OG) in the cleavage buffer for full length syntaxin 1A and VAMP2 with transmembrane domain.

His 6-tagged SNAP-25 was expressed in E.coli BL21 (DE3) Codon Plus RIL (Stratagene) and purified using Ni-NTA resin (Qiagen) 32 . The cells were grown at 37ºC in LB with glucose (2 g/l), kanamycin (34 µg/ml), and chloramphenicol (50 µg/ml) until

A600 reached 0.6-0.8. Protein expression was induced by 0.5 mM IPTG, and the cells were grown for an additional 6h at 30ºC. Finally, the protein was eluted from the Ni-NTA resin by elution buffer (25 mM Hepes and 100mM KCl with 250 mM imidazole, pH 7.4).

The cysteine mutants of full length syntaxin 1A and VAMP2 were reacted with (1- oxyl-2, 2, 5, 5-tetramethyl-D-pyrroline-3-methyl) methanethiosulfonate spin label at 4ºC overnight while the proteins were bound to the GST-agarose beads. To remove free spin label, the beads with bound proteins were extensively washed with the cleavage buffer and then cleaved by thrombin (Sigma). The spin-labeling efficiency was determined by comparing the spin concentrations with the 50-µM 2, 2, 6, 6-tetramethyl-4-piperidine N- oxide standard. For all samples, the efficiency was ~80%.

3.5.3 Preparation of the SNARE ternary complex

Purified His 6-tagged SNAP-25 was first added to the Ni-NTA resin solution and nutated for 1 h at room temperature. After washing out the free proteins, we mixed corresponding versions of purified syntaxin 1A and VAMP2 for each separate experiment with His 6-SNAP-25 to form the SNARE complex with the molar ratio of 2:5:1. The mixture was incubated at 4ºC overnight. After extensive washing to remove the unbound proteins, the SNARE ternary complex was eluted with a buffer containing 250 mM imidazole and 0.8% OG. The formation of the complex was confirmed with SDS-PAGE gel. We obtained identical results when the ternary complex was formed by adding all three components simultaneously.

3.5.4 EPR data collection

As for the ternary SNARE complex in the detergent (0.8% OG), the samples were concentrated to the final concentration of 50-100 µM using a 10-kDa cutoff centrifugal filter (Millipore) before EPR spectra collection. For the purified complex in the membrane, the samples were reconstituted into vesicles at about 1:200 protein-to-lipid molar ratio using the published method 57 . The large unilamellar vesicles (~100nm in diameter) of 1-palmitoyl-2-oleoyl-sn -glycero-3-phosphatidylserine were prepared in a detergent-free buffer using an extruder (Avanti). The detergent was removed by treating the sample with the dialysis buffer (25 mM Hepes and 100 mM KCL, pH 7.4) at 4ºC overnight. After dialysis, the sample solution was concentrated using a 100-kDa cutoff centrifugal filter (Millipore) before taking an EPR spectrum.

3.5.5 EPR accessibility measurements

The gas exchange to the protein sample was achieved with the TPX EPR1 tube for the loop-gap resonator. For individual mutants, 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 of 0.1-40mW. Three power saturation curves were obtained after equilibration (1) with N 2, (2) with air (O 2), and (3) with N 2 in the presence of 200mM NiEDDA (nickel ethylenediaminediacetic acid). 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 ∆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 coefficient times the frequency of collision of the nitroxide with the freely diffusion reagents such as oxygen and NiEDDA. Thus, ∆P1/2 is considered to be equivalent to the accessibility W.

2.8 Reference

1. Jahn, R. & Sudhof, T. C. (1999). Membrane fusion and exocytosis. Annu Rev Biochem 68 , 863-911.

2. Rothman, J. E. (1994). Mechanisms of intracellular protein transport. Nature 372 , 55-63.

3. Bennett, M. K., Calakos, N. & Scheller, R. H. (1992). Syntaxin: a synaptic protein implicated in docking of synaptic vesicles at presynaptic active zones. Science 257 , 255-9.

4. Oyler, G. A., Higgins, G. A., Hart, R. A., Battenberg, E., Billingsley, M., Bloom, F. E. & Wilson, M. C. (1989). The identification of a novel synaptosomal- associated protein, SNAP-25, differentially expressed by neuronal subpopulations. J Cell Biol 109 , 3039-52.

5. Trimble, W. S., Cowan, D. M. & Scheller, R. H. (1988). VAMP-1: a synaptic vesicle-associated integral membrane protein. Proc Natl Acad Sci U S A 85 , 4538- 42.

6. Trimble, W. S. & Scheller, R. H. (1988). Molecular biology of synaptic vesicle- associated proteins. Trends Neurosci 11 , 241-2.

7. Baumert, M., Maycox, P. R., Navone, F., De Camilli, P. & Jahn, R. (1989). Synaptobrevin: an integral membrane protein of 18,000 daltons present in small synaptic vesicles of rat brain. Embo J 8, 379-84.

8. Jahn, R. & Scheller, R. H. (2006). SNAREs--engines for membrane fusion. Nat Rev Mol Cell Biol 7, 631-43.

9. Brunger, A. T. (2005). Structure and function of SNARE and SNARE-interacting proteins. Q Rev Biophys 38 , 1-47.

10. Kloepper, T. H., Kienle, C. N. & Fasshauer, D. (2007). An elaborate classification of SNARE proteins sheds light on the conservation of the eukaryotic endomembrane system. Mol Biol Cell 18 , 3463-71.

11. Hong, W. (2005). SNAREs and traffic. Biochim Biophys Acta 1744 , 493-517.

12. Fasshauer, D. (2003). Structural insights into the SNARE mechanism. Biochim Biophys Acta 1641 , 87-97.

13. Chen, Y. A. & Scheller, R. H. (2001). SNARE-mediated membrane fusion. Nat Rev Mol Cell Biol 2, 98-106.

14. Mayer, A. (2002). Membrane fusion in eukaryotic cells. Annu Rev Cell Dev Biol 18 , 289-314.

15. Rizo, J. & Sudhof, T. C. (2002). Snares and Munc18 in synaptic vesicle fusion. Nat Rev Neurosci 3, 641-53.

16. Schiavo, G., Benfenati, F., Poulain, B., Rossetto, O., Polverino de Laureto, P., DasGupta, B. R. & Montecucco, C. (1992). Tetanus and botulinum-B neurotoxins block neurotransmitter release by proteolytic cleavage of synaptobrevin. Nature 359 , 832-5.

17. Schiavo, G., Shone, C. C., Bennett, M. K., Scheller, R. H. & Montecucco, C. (1995). Botulinum neurotoxin type C cleaves a single Lys-Ala bond within the carboxyl-terminal region of syntaxins. J Biol Chem 270 , 10566-70.

18. Blasi, J., Chapman, E. R., Link, E., Binz, T., Yamasaki, S., De Camilli, P., Sudhof, T. C., Niemann, H. & Jahn, R. (1993). Botulinum neurotoxin A selectively cleaves the synaptic protein SNAP-25. Nature 365 , 160-3.

19. Sollner, T., Bennett, M. K., Whiteheart, S. W., Scheller, R. H. & Rothman, J. E. (1993). A protein assembly-disassembly pathway in vitro that may correspond to sequential steps of synaptic vesicle docking, activation, and fusion. Cell 75 , 409- 18.

20. Hayashi, T., Yamasaki, S., Nauenburg, S., Binz, T. & Niemann, H. (1995). Disassembly of the reconstituted synaptic vesicle membrane fusion complex in vitro. Embo J 14 , 2317-25.

21. Fasshauer, D., Otto, H., Eliason, W. K., Jahn, R. & Brunger, A. T. (1997). Structural changes are associated with soluble N-ethylmaleimide-sensitive fusion protein attachment protein receptor complex formation. J Biol Chem 272 , 28036- 41.

22. Poirier, M. A., Hao, J. C., Malkus, P. N., Chan, C., Moore, M. F., King, D. S. & Bennett, M. K. (1998). Protease resistance of syntaxin.SNAP-25.VAMP complexes. Implications for assembly and structure. J Biol Chem 273 , 11370-7.

23. Sutton, R. B., Fasshauer, D., Jahn, R. & Brunger, A. T. (1998). Crystal structure of a SNARE complex involved in synaptic exocytosis at 2.4 A resolution. Nature 395 , 347-53.

24. Antonin, W., Fasshauer, D., Becker, S., Jahn, R. & Schneider, T. R. (2002). Crystal structure of the endosomal SNARE complex reveals common structural principles of all SNAREs. Nat Struct Biol 9, 107-11.

25. Sorensen, J. B., Wiederhold, K., Muller, E. M., Milosevic, I., Nagy, G., de Groot, B. L., Grubmuller, H. & Fasshauer, D. (2006). Sequential N- to C-terminal SNARE complex assembly drives priming and fusion of secretory vesicles. Embo J 25 , 955-66.

26. Pobbati, A. V., Stein, A. & Fasshauer, D. (2006). N- to C-terminal SNARE complex assembly promotes rapid membrane fusion. Science 313 , 673-6.

27. Chen, Y. A., Scales, S. J. & Scheller, R. H. (2001). Sequential SNARE assembly underlies priming and triggering of exocytosis. Neuron 30 , 161-70.

28. Fiebig, K. M., Rice, L. M., Pollock, E. & Brunger, A. T. (1999). Folding intermediates of SNARE complex assembly. Nat Struct Biol 6, 117-23.

29. Hua, S. Y. & Charlton, M. P. (1999). Activity-dependent changes in partial VAMP complexes during neurotransmitter release. Nat Neurosci 2, 1078-83.

30. Melia, T. J., Weber, T., McNew, J. A., Fisher, L. E., Johnston, R. J., Parlati, F., Mahal, L. K., Sollner, T. H. & Rothman, J. E. (2002). Regulation of membrane fusion by the membrane-proximal coil of the t-SNARE during zippering of SNAREpins. J Cell Biol 158 , 929-40.

31. Xu, T., Rammner, B., Margittai, M., Artalejo, A. R., Neher, E. & Jahn, R. (1999). Inhibition of SNARE complex assembly differentially affects kinetic components of exocytosis. Cell 99 , 713-22.

32. Zhang, Y., Su, Z., Zhang, F., Chen, Y. & Shin, Y. K. (2005). A partially zipped SNARE complex stabilized by the membrane. J Biol Chem 280 , 15595-600.

33. Stein, A., Weber, G., Wahl, M. C. & Jahn, R. (2009). Helical extension of the neuronal SNARE complex into the membrane. Nature 460 , 525-8.

34. Kweon, D. H., Kim, C. S. & Shin, Y. K. (2002). The membrane-dipped neuronal SNARE complex: a site-directed spin labeling electron paramagnetic resonance study. Biochemistry 41 , 9264-8.

35. Kim, C. S., Kweon, D. H. & Shin, Y. K. (2002). Membrane topologies of neuronal SNARE folding intermediates. Biochemistry 41 , 10928-33.

36. Hubbell, W. L., McHaourab, H. S., Altenbach, C. & Lietzow, M. A. (1996). Watching proteins move using site-directed spin labeling. Structure 4, 779-83.

37. Hubbell, W. L., Gross, A., Langen, R. & Lietzow, M. A. (1998). Recent advances in site-directed spin labeling of proteins. Curr Opin Struct Biol 8, 649-56.

38. Altenbach, C., Greenhalgh, D. A., Khorana, H. G. & Hubbell, W. L. (1994). A collision gradient method to determine the immersion depth of nitroxides in lipid bilayers: application to spin-labeled mutants of bacteriorhodopsin. Proc Natl Acad Sci U S A 91 , 1667-71.

39. Kweon, D. H., Kim, C. S. & Shin, Y. K. (2003). Insertion of the membrane- proximal region of the neuronal SNARE coiled coil into the membrane. J Biol Chem 278 , 12367-73.

40. Kummel, D., Krishnakumar, S. S., Radoff, D. T., Li, F., Giraudo, C. G., Pincet, F., Rothman, J. E. & Reinisch, K. M. Complexin cross-links prefusion SNAREs into a zigzag array. Nat Struct Mol Biol 18 , 927-33.

41. Zhang, F., Chen, Y., Kweon, D. H., Kim, C. S. & Shin, Y. K. (2002). The four- helix bundle of the neuronal target membrane SNARE complex is neither disordered in the middle nor uncoiled at the C-terminal region. J Biol Chem 277 , 24294-8.

42. Macosko, J. C., Kim, C. H. & Shin, Y. K. (1997). The membrane topology of the fusion peptide region of influenza hemagglutinin determined by spin-labeling EPR. J Mol Biol 267 , 1139-48.

43. Shin, Y. K., Levinthal, C., Levinthal, F. & Hubbell, W. L. (1993). Colicin E1 binding to membranes: time-resolved studies of spin-labeled mutants. Science 259 , 960-3.

44. Altenbach, C., Marti, T., Khorana, H. G. & Hubbell, W. L. (1990). Transmembrane protein structure: spin labeling of bacteriorhodopsin mutants. Science 248 , 1088-92.

45. Weber, T., Zemelman, B. V., McNew, J. A., Westermann, B., Gmachl, M., Parlati, F., Sollner, T. H. & Rothman, J. E. (1998). SNAREpins: minimal machinery for membrane fusion. Cell 92 , 759-72.

46. McHaourab, H. S., Lietzow, M. A., Hideg, K. & Hubbell, W. L. (1996). Motion of spin-labeled side chains in T4 lysozyme. Correlation with protein structure and dynamics. Biochemistry 35 , 7692-704.

47. Kim, C. H., Macosko, J. C. & Shin, Y. K. (1998). The mechanism for low-pH- induced clustering of phospholipid vesicles carrying the HA2 ectodomain of influenza hemagglutinin. Biochemistry 37 , 137-44.

48. Perozo, E., Cortes, D. M. & Cuello, L. G. (1998). Three-dimensional architecture and gating mechanism of a K+ channel studied by EPR spectroscopy. Nat Struct Biol 5, 459-69.

49. Gross, A. & Hubbell, W. L. (2002). Identification of protein side chains near the membrane-aqueous interface: a site-directed spin labeling study of KcsA. Biochemistry 41 , 1123-8.

50. Nichols, B. J., Ungermann, C., Pelham, H. R., Wickner, W. T. & Haas, A. (1997). Homotypic vacuolar fusion mediated by t- and v-SNAREs. Nature 387 , 199-202.

51. Marash, M. & Gerst, J. E. (2001). t-SNARE dephosphorylation promotes SNARE assembly and exocytosis in yeast. Embo J 20 , 411-21.

52. Schoch, S., Deak, F., Konigstorfer, A., Mozhayeva, M., Sara, Y., Sudhof, T. C. & Kavalali, E. T. (2001). SNARE function analyzed in synaptobrevin/VAMP knockout mice. Science 294 , 1117-22.

53. Schiavo, G., Shone, C. C., Rossetto, O., Alexander, F. C. & Montecucco, C. (1993). Botulinum neurotoxin serotype F is a zinc endopeptidase specific for VAMP/synaptobrevin. J Biol Chem 268 , 11516-9.

54. Binz, T., Blasi, J., Yamasaki, S., Baumeister, A., Link, E., Sudhof, T. C., Jahn, R. & Niemann, H. (1994). Proteolysis of SNAP-25 by types E and A botulinal neurotoxins. J Biol Chem 269 , 1617-20.

55. Tokumaru, H., Umayahara, K., Pellegrini, L. L., Ishizuka, T., Saisu, H., Betz, H., Augustine, G. J. & Abe, T. (2001). SNARE complex oligomerization by synaphin/complexin is essential for synaptic vesicle exocytosis. Cell 104 , 421-32.

56. Margittai, M., Otto, H. & Jahn, R. (1999). A stable interaction between syntaxin 1a and synaptobrevin 2 mediated by their transmembrane domains. FEBS Lett 446 , 40-4.

57. Chen, Y., Xu, Y., Zhang, F. & Shin, Y. K. (2004). Constitutive versus regulated SNARE assembly: a structural basis. Embo J 23 , 681-9.

2.7 Figures and Captions

Figure 1 Schematic domain structures of SNARE proteins and their truncation mutants used in this study. The SNARE motifs (grey), N-terminal Habc domain of syntaxin 1A (blue) and transmembrane regions of syntaxin 1A and VAMP2 (red) are shown. Full length syntaxin 1A, truncated form of syntaxin 1A with the SNARE motif and the linker region (residues 191-266) and truncated form of syntaxin 1A with only the SNARE motif (residues 191-256) are named as SynFL, SynSH3 and SynDL respectively. Full length VAMP2, truncated form of VAMP2 (residues 1-96) and truncated form of VAMP2 (residues 1-85) are named as VpFL, VpS and VpDL respectively. The amino acid sequence of the point mutation positions is displayed.

Figure 2 EPR spectra of spin-labeled full-length syntaxin 1A in SNARE complex formed with soluble VpS or soluble VpDL in solution. First-derivative mode EPR spectra taken at room temperature in detergent are shown. Ternary SNARE complex were formed by SNAP25, SynFL and VpS (black) or VpDL(red).

Figure 3 EPR spectra of spin-labeled full-length syntaxin 1A in SNARE complex formed with full-length VAMP2 or soluble VpDL in solution . First-derivative mode EPR spectra taken at room temperature in detergent are shown. Ternary SNARE complex were formed by SNAP25, SynFL and VpFL (black) or VpDL (red). The SNARE complex formed by SNAP25, SynFL and VpFL represents a cis -SNARE structure in detergent by including both of the TMRs (black lines) .

Figure 4 EPR spectra of spin-labeled full-length syntaxin 1A in SNARE complex formed with soluble VpDS or soluble VpS in solution. First-derivative mode EPR spectra taken at room temperature in detergent are shown. Ternary SNARE complex were formed by SNAP25, SynFL and VpS (black) or VpDS (red). The VpDS mutant is a truncated version of VAMP2 which lacks half of its SNARE motif.

Figure 5 EPR spectra of spin-labeled full-length VAMP2 in SNARE complex formed with soluble SynDL or soluble SynSH3 in solution . First-derivative mode EPR spectra taken at room temperature in detergent are shown. Ternary SNARE complex were formed by SNAP25, VpFL and SynSH3 (black) or SynDL (red). Both of SynSH3 and SynDL mutants don’t have the N-terminal Habc domain for sake of simplicity. Habc domain turns out to be not involved in the SNARE four-helix bundle formation.

Figure 6 EPR spectra of spin-labeled soluble VpS in SNARE complex formed with soluble SynDL or soluble SynSH3 in solution . First-derivative mode EPR spectra taken at room temperature in detergent are shown. Ternary SNARE complex were formed by SNAP25, VpS and SynSH3 (black) or SynDL (red). Both of SynSH3 and SynDL mutants don’t have the N-terminal Habc domain for sake of simplicity. Habc domain turns out to be not involved in the SNARE four-helix bundle formation.

Figure 7 EPR spectra of spin-labeled full-length syntaxin 1A in SNARE complex formed with soluble VpS or soluble VpDL in the membrane. First-derivative mode EPR spectra in the membrane taken at room temperature are shown. Ternary SNARE complex were formed by SNAP25, full length syntaxin 1A and VpS (left panel) or VpDL (right panel).The sample is used for the EPR power saturation experiment.

Figure 8 EPR spectra of spin-labeled full-length VAMP2 in SNARE complexformed with full-length syntaxin 1A or soluble SynDL in the membrane. First-derivative mode EPR spectra in the membrane taken at room temperature are shown. Ternary SNARE complex were formed by SNAP25, full length VAMP2, and full-length syntaxin1A (left panel) or SynDL (right panel).The sample is used for the EPR power saturation experiment.

Amplitude

0 2 4 6 (P (mW))1/2

Figure 9 Example plots of power saturation curves for the recombinant ternary

SNARE complex formed by spin-labeled VpFK91C, His 6 tagged SN25, and SynDL in vesicles containing 45% POPC, 15% POPS, 40% CHO. The data were obtained by measuring the peak-to-peak amplitude of the M I=0 line of the first-derivative EPR spectrum. The EPR spectra were collected at 0.1, 0.25, 0.4, 1.0, 4.0, 10, 16, 25, and 40mW incident microwave powers under three different conditions: under nitrogen flow in the absence of paramagnetic reagents (open circles); under nitrogen in the presence of 10mW NiEDDA (open squares); and under the flow of air (closed circles). The curves are the least-squares fits to eq 1.

Figure 10 Normalized accessibility to the paramagnetic reagents of syntaxin 1A. (a)

The normalized accessibility parameter to oxygen (WO2 ) (open circles) and that to NiEDDA (W NiEDDA ) (filled circles) for full length syntaxin 1A in SNARE complex formed with VpDL are plotted against the residue numbers. (b) The normalized accessibility parameter to oxygen

(W O2 ) (open circles) and that to NiEDDA (W NiEDDA ) (filled circles) for full length syntaxin 1A in SNARE complex formed with VpS are plotted against the residue numbers.

Figure 11 Normalized accessibility to the paramagnetic reagents of VAMP2. (a)

The normalized accessibility parameter to oxygen (WO2 ) (open circles) and that to NiEDDA (W NiEDDA ) (filled circles) for full length VAMP2 in SNARE complex formed with SynDL is plotted against the residue numbers. (b) The normalized accessibility parameter to oxygen

(W O2 ) (open circles) and that to NiEDDA (W NiEDDA ) (filled circles) for full length VAMP2 in SNARE complex formed with SynSH3 is plotted against the residue numbers.

CHAPTER 4: GENERAL CONCLUSIONS

5.1 Summary

The SNARE system for neurotransmitter release is advantageous over that for vesicle trafficking in yeast in term of its more sophisticated fine-tuning characteristic which can perfectly couple the Ca 2+ signaling and the fast fusion event initiation 1; 2; 3 . This more developmental task is accomplished by more complicated fusion machinery. The neuronal SNAREs is able to involve other regulatory factors in addition to the basic SNARE components 4. These various regulatory proteins control different fusion steps and finally complete an elaborate procedure together with the SNARE complex.

In this dissertation study, I not only focus on investigation of SNARE-complexin complex, but also on the structural basis of the SNARE complex functional domains. Various EPR techniques including the spectrum analysis and power saturation method have been used to develop the detailed structure information. Two main projects have been presented in this thesis to show the revealed mechanism of the interplay between SNAREs and regulatory proteins and how the SNARE complex modulates its structure to facilitate the regulatory function.

Complexin is one of these important regulatory proteins which have been studied extensively 5; 6; 7; 8 . And these studies have also raised intensive controversial issues about complexin function 9; 10 . Initially, a fusion “clamp” model has been proposed evidenced by inhibitory effects observed in different physiological systems 8; 11; 12 . However, a dual function model has become more and more extensively accepted recently 6; 9 . But the underlying mechanism of this dual function remains elusive. It has also been claimed that complexin can function like an energy transducer 10 . Complexin might also function to control the Ca 2+ -triggering step in membrane fusion and it might either have a cooperation with synaptotagmin or a redundant function 5; 7 . Rothman et al . suggests that complexin accessory α-helix can replace VAMP2 C-terminal, thereby inhibiting SNARE assembly and membrane fusion 11 . However, in our EPR analysis, this replacement effect

96 turns out to be very small and almost suppressed by the presence of the complexin N- terminal sequence. There are several possible scenarios about the balancing between the N-terminal stimulatory effect and the accessory α-helix inhibitory effect. And this delicate balance between those dual functions somewhat produces the on-and-off switch control of the complexin overall performance, and then contributes to the highly regulatory characteristic of neurotransmitter release.

The progress in resolving crystal structure of SNARE complex with full length syntaxin 1A and VAMP2 is a great breakthrough in understanding the mechanism of SNARE complex as a fusion catalyst 13 . And the N- to C- direction zippering model 14 of SNARE complex assembly has been further confirmed by this study. The continuity of α- helical structure leaves no flexible region between the membrane proximal region and the SNARE core which guarantees that the energy transfer pathway won’t be disrupted or dissipated. While this further raise the interest about the structural features of a trans - SNARE complex. Here in this study, we used different versions of truncated mutants of syntaxin 1A and VAMP2 to block the progressive zippering of SNARE complex in different intermediate stages. And hopefully, these intermediate states would partially represent different trans -SNARE stages and give us a primary idea about the SNARE structure during zippering. Based on both previous EPR studies and our new development, we found out that SNARE proteins underwent dramatic conformational changes during the transition from trans -SNARE to cis-SNARE. A relatively “loose” conformation of t-SNARE would be progressively tightened up upon the engagement of v-SNARE. But the secondary structure of individual syntaxin 1A and VAMP2 linker region had already been formed separately before the zippering was up to the linker region. Our EPR investigation has also successfully revealed a potential ternary interaction between the syntaxin 1A linker region and some other part of the SNARE complex, which is likely to be the C-terminus of SNAP25 SN2 domain by a natural speculation. We were also able to observe a membrane bound fashion of the syntaxin 1A and VAMP2 linker regions from both our EPR line shapes and power saturation experiment, which was consistent with previous results and physically meaningful.

Our findings in these studies all supported the zippering model of SNARE complex assembly. We focused on the underlying structural details about the efficient and exquisite fusion activity of SNARE proteins. And we successfully gained useful information to fill up the gaps of previous research. The results shown in this dissertation further enriched our understanding of the temporally and spatially regulated mechanisms of SNARE-mediated membrane fusion.

5.2 Future plan

5.2.1 New system to investigate structure of trans-SNARE complex

Although we have kept using the truncated versions of VAMP2 aiming to mimic a trans -SNARE state, the actual situation for the tran -SNARE complex in which VAMP2 and t-SNARE reside on the opposite membrane is expected to be much weaker than that in our model system. In our system, no matter which kind of truncated version we use, we just avoid the situation of the cis -SNARE complex in which the transmembrane domains of VAMP2 and syntaxin 1A are in the same membrane, but imposing no repulsion of bilayer-bilayer interaction to the SNARE core. And some of the activities of regulators in trans -SNARE state might be covered up in this experiment system. So a more suitable working system to investigate the trans -SNARE state is highly necessary to be built up. We are currently working on that using both lipid bilayer and nanodisc to stabilize transmembrane domains seperately. And one of our future directions would be optimize the conditions of this system and set up a more mature platform for experiments to validate all of our hypotheses under the trans -SNARE context.

5.2.2 Identify the interaction partner of syntaxin 1A linker region

Based on our linker region structure study, we identified an interaction between the syntaxin 1A linker region and some other part in the SNARE complex. We have proposed a hypothesis that this part might be the SN2 C-terminal domain of SNAP25. EPR experiments could be done to examine this proposal. We will screen spin-labeled cysteine mutants of SN2 C-terminal domain and detect which region will have a structure change upon VAMP2 binding. If we successfully locate a specific area that shows this kind of expected effect, we have reason to believe that this region is highly likely interacting with syntaxin 1A linker region, or VAMP2 linker region as well.

5.2.3 Crystalization of SNARE-complexin full length quaternary complex

Although several crystal structures characterizing the SNARE-complexin complex has been resolved representing different binding modes of complexin with SNARE complex 15; 16 , no complete structure of the whole quaternary complex with full length versions of each individual component has been studied. Structures of complexin bound to a cis -SNARE complex gave rise to an insight regarding the facilitatory mechanism. The N-terminal and C-terminal truncated complexin binds to a grove between syntaxin 1A and VAMP2 by its central α-helix and its accessory α-helix parallels the C-terminal membrane-proximal region of the fully zippered SNARE complex but not interacts with it. However, the accessory α-helix is supposed to perform the inhibitory function, perhaps creates the clamped, prefusion state 17; 18; 19 . And a crystal structure revealing complexin binding to a mimetic of prefusion SNAREpin has also been exposed recently 16 . In this model, they observed that the central α-helix of complexin was anchored to one SNARE complex, while its accessory α-helix bound to a second complex at the vacant v-SNARE binding site; therefore forms a zigzag array which is non-active for membrane fusion.

However, in both of these studies, a fragment consisting of only complexin central and accessory α-helices has been used. There are NMR studies showing that the N-terminus of complexin binds to the C-terminus of the SNARE complex and helps to stabilize the SNARE complex C-terminus and/or helps release the inhibitory function of complexin 20 .

But up to now, no clear structure information about complexin C-terminus is available. And we also suggested a binding mode between complexin and t-SNARE only previously. So crystallization study is the most straightforward and powerful way to shed light on those elusive areas about complexin N- and C-termus positions, t-SNARE binding mode and how could the balancing between different binding patterns of complexin be modulated to perform appropriate function under the corresponding conditions.

5.3 Reference

1. Chen, Y. A. & Scheller, R. H. (2001). SNARE-mediated membrane fusion. Nat Rev Mol Cell Biol 2, 98-106.

2. Chen, Y. A., Scales, S. J., Patel, S. M., Doung, Y. C. & Scheller, R. H. (1999). SNARE complex formation is triggered by Ca2+ and drives membrane fusion. Cell 97 , 165-74.

3. Sollner, T., Bennett, M. K., Whiteheart, S. W., Scheller, R. H. & Rothman, J. E. (1993). A protein assembly-disassembly pathway in vitro that may correspond to sequential steps of synaptic vesicle docking, activation, and fusion. Cell 75 , 409- 18.

4. Rizo, J. & Rosenmund, C. (2008). Synaptic vesicle fusion. Nat Struct Mol Biol 15 , 665-74.

5. Reim, K., Mansour, M., Varoqueaux, F., McMahon, H. T., Sudhof, T. C., Brose, N. & Rosenmund, C. (2001). Complexins regulate a late step in Ca2+-dependent neurotransmitter release. Cell 104 , 71-81.

6. Yoon, T. Y., Lu, X., Diao, J., Lee, S. M., Ha, T. & Shin, Y. K. (2008). Complexin and Ca2+ stimulate SNARE-mediated mem brane fusion. Nat Struct Mol Biol 15 , 707-13.

7. Tang, J., Maximov, A., Shin, O. H., Dai, H., Rizo, J. & Sudhof, T. C. (2006). A complexin/synaptotagmin 1 switch controls fast synaptic vesicle exocytosis. Cell 126 , 1175-87.

8. Huntwork, S. & Littleton, J. T. (2007). A complexin fusion clamp regulates spontaneous neurotransmitter release and synaptic growth. Nat Neurosci 10 , 1235-7.

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9. Xue, M., Reim, K., Chen, X., Chao, H. T., Deng, H., Rizo, J., Brose, N. & Rosenmund, C. (2007). Distinct domains of complexin I differentially regulate neurotransmitter release. Nat Struct Mol Biol 14 , 949-58.

10. Maximov, A., Tang, J., Yang, X., Pang, Z. P. & Sudhof, T. C. (2009). Complexin controls the force transfer from SNARE complexes to membranes in fusion. Science 323 , 516-21.

11. Giraudo, C. G., Garcia-Diaz, A., Eng, W. S., Chen, Y., Hendrickson, W. A., Melia, T. J. & Rothman, J. E. (2009). Alternative zippering as an on-off switch for SNARE-mediated fusion. Science 323 , 512-6.

12. Giraudo, C. G., Eng, W. S., Melia, T. J. & Rothman, J. E. (2006). A clamping mechanism involved in SNARE-dependent exocytosis. Science 313 , 676-80.

13. Stein, A., Weber, G., Wahl, M. C. & Jahn, R. (2009). Helical extension of the neuronal SNARE complex into the membrane. Nature 460 , 525-8.

14. Hua, S. Y. & Charlton, M. P. (1999). Activity-dependent changes in partial VAMP complexes during neurotransmitter release. Nat Neurosci 2, 1078-83.

15. Chen, X., Tomchick, D. R., Kovrigin, E., Arac, D., Machius, M., Sudhof, T. C. & Rizo, J. (2002). Three-dimensional structure of the complexin/SNARE complex. Neuron 33 , 397- 409.

16. Kummel, D., Krishnakumar, S. S., Radoff, D. T., Li, F., Giraudo, C. G., Pincet, F., Rothman, J. E. & Reinisch, K. M. Complexin cross-links prefusion SNAREs into a zigzag array. Nat Struct Mol Biol 18 , 927-33.

17. Xue, M., Lin, Y. Q., Pan, H., Reim, K., Deng, H., Bellen, H. J. & Rosenmund, C. (2009). Tilting the balance between facilitatory and inhibitory functions of mammalian and Drosophila Complexins orchestrates synaptic vesicle exocytosis. Neuron 64 , 367-80.

18. Martin, J. A., Hu, Z., Fenz, K. M., Fernandez, J. & Dittman, J. S. Complexin has opposite effects on two modes of synaptic vesicle fusion. Curr Biol 21 , 97-105.

19. Lu, B., Song, S. & Shin, Y. K. Accessory alpha-helix of complexin I can displace VAMP2 locally in the complexin-SNARE quaternary complex. J Mol Biol 396 , 602-9.

20. Xue, M., Craig, T. K., Xu, J., Chao, H. T., Rizo, J. & Rosenmund, C. Binding of the complexin N terminus to the SNARE complex potentiates synaptic-vesicle fusogenicity. Nat Struct Mol Biol 17 , 568-75.

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ACKNOWLEDGEMENTS

Here I am taking this great opportunity to express my sincere gratitude to all those people who have given me great help during my Ph.D. study.

First and foremost, I would like to thank to my major professor Dr. Yeon-Kyun Shin for his valuable guidance and advice. His inspiration was important for me to be brave enough to think, to fail and to conquer. He has always been patient and willing to help when there are difficulties on the way. His experience and insight in science helped me through all the obstructions during these years.

Also I should thank my committee members Dr. Linda, Ambriosio, Dr. Edward, Yu, Dr.Alan, DiSpirito and Dr. Diane Bassham. This dissertation would not have been possible without their support and suggestions.

I would also like to thank all of my labmates, those have been graduated or will be graduate soon. I will give my best wishes to them and hope them to have a great future and live happily ever after.

I need to thank my parents for giving me the strongest support and backing me up for all of these years studying abroad. They have sacrificed a lot for assisting my career.

Last but not the least, I would like to thank a very special person in my life. I would not be able to accomplish all I have accomplished without him. Thanks to him for making me not the person who I am, but the one who I want to be.