Copyright
by
Kerry L. Fuson
2009
The Dissertation Committee for Kerry L. Fuson Certifies that this is the approved
version of the following dissertation:
Structural Characterization of Synaptotagmin I
Committee:
R. Bryan Sutton, Supervisor
Andres Oberhauser
Darren Boehning
Mark Hellmich
Xiaodong Cheng
__________________
Dean, Graduate School
Structural Characterization of Synaptotagmin I
by
Kerry L. Fuson, B.S., M.S.
Dissertation
Presented to the Faculty of the Graduate School of
The University of Texas Medical Branch
in Partial Fulfillment
of the Requirements
for the Degree of
Doctor of Philosophy
The University of Texas Medical Branch
February, 2009
Dedication
This dissertation is dedicated to my wonderful wife, DeeOna, who has stood
beside me and helped me get to where I am today. You have been with me every step of
the way, through good times and bad. Thank you for the unconditional love, guidance,
and support that you have always given me, helping me to succeed and instilling in me
the confidence that I am capable of doing anything I put my mind to. Thank you for
everything. I love you!
Acknowledgements
First and foremost I offer my sincerest and heartfelt gratitude to my supervisor,
Dr. Bryan Sutton, who has supported me throughout my time at UTMB with his patience,
understanding and knowledge. It is difficult to overemphasize my gratitude to him. I
attribute the level of my Ph.D. degree to his encouragement, support and effort. Without
him this work would not have been completed or written. I simply could not wish for a
better supervisor.
I am indebted to my many student colleagues for providing a
stimulating and fun environment in which to learn and grow.
The Department of
Biochemistry and Molecular Biology has provided me with the support and equipment I
have needed to do my research and complete my studies. The gulf coast consortium
provided me with a fellowship (HAMBP) that funded my studies. I especially thank
Debora Botting for her help and support. She never lacked in her willingness to help and
I honestly don’t think the department would function without her.
Dr. Andres
Oberhauser and his lab provided me with the use of the AFM and lent his expertise in the
synaptotagmin mechanical force study. His willingness and ability to teach has also
provided me with an understanding of the principals behind the technique. Miguel
Montes provided me with a never ending supply of pure protein. His ability to purify
protein is second to none. Miguel also provided me with stimulating conversations that
helped me grow and move forward as a student. A special thanks to my committee
members for giving me solid advice and constructive criticism. Especially, I would like
to give my special thanks to my wife DeeOna whose patients and love enabled me to
complete this work.
v
Structural Characteristics of Synaptotagmin I
Publication No._____________
Kerry L. Fuson, Ph.D.
The University of Texas Medical Branch, 2009
Supervisor: R. Bryan Sutton
Synaptotagmin I is the most abundant Ca+2 binding protein present on synaptic
vesicles accounting for 7% of total vesicle protein and is widely accepted as the Ca2+
sensor in fast synchronous neurotransmitter release. The protein is composed of one transmembrane domain, an unstructured linker followed by two C2A domains identified as
C2A and C2B. Each C2 domain is composed of an 8 stranded β-sandwich joined by a 9
amino acid linker. The Ca+2 binding pocket is composed of three loops located at the apex
of the protein. In the Syt I C2AB structure, we see evidence of a domain structural change
in the absence of Ca2+. Analysis of interacting residues between C2A and C2B show a
network of highly conserved residues within the C2 domain that regulates
Ca+2/phohspholipid binding in C2A. Analysis of the Syt I C2A structure, as well as,
previous C2A structures shows a strong H-bond between Tyr 180 and His 237 in C2A. By
removing this H-bond, disorder of Loop 3 is increased and the thermodynamic stability of
the C2 domain decreases. Our hypothesis is that the absolute position of the Ca2+ binding
loops of C2 domains affects Ca+2 affinity and, and ultimately domain stability. We used
several different biochemical approaches to test the hypothesis.
vi
We assessed the
importance of Loop 3 mutations using X-ray crystallography methods, bulk
thermodynamic measurements using lifetime fluorescence, and analyzed the mechanical
properties of the C2-domains using single molecule force spectroscopy.
We studied the mechanical stability of the C2A and C2B domains of human Syt1
using single-molecule atomic force microscopy. We found that stretching the C2AB
domains of Syt1 resulted in two distinct unfolding force peaks. The larger force peak of
~100pN was identified as C2B and the second peak of ~50pN as C2A. Further, a
significant fraction of C2A domains unfolded through a low force intermediate that was
not observed in C2B.
We conclude that these domains have different mechanical
properties. We hypothesize that a relatively small stretching force may be sufficient to
deform the effector-binding regions of C2A domain and modulate the affinity for
SNAREs, phospholipids and Ca+2.
vii
TABLE OF CONTENTS
Acknowledgments....................................................................................................v
Abstract .................................................................................................................. vi
Contents ............................................................................................................... viii
List of Tables ........................................................................................................ xii
List of Figures ...................................................................................................... xiii
List of Abbreviations .............................................................................................xv
CHAPTER 1 INTRODUCTION
Synaptic Vesicle Fusion ..................................................................................1
Proteins involved in neurotransmitter release ................................................2
C2 domains .....................................................................................................4
Synaptotagmin family of proteins ..................................................................6
Synaptotagmin functions as Ca2+ sensor .........................................................7
Phospholipid binding ......................................................................................9
Past structural studies ....................................................................................10
HYPOTHESIS AND GOALS: ...................................................................................11
CHAPTER 2 PURIFICATION, CRYSTALLIZATION AND X-RAY DIFFRACTION ANALYSIS
OF HUMAN SYNAPTOTAGMIN 1 C2A-C2B.
Introduction ...................................................................................................12
Cloning, Expression and Purification ...........................................................14
Crystallization and Data Collection ..............................................................16
X-ray Diffraction ..........................................................................................16
Results and Discussion .................................................................................17
CHAPTER 3 X-RAY CRYSTAL STRUCTURE OF SYNAPTOTAGMIN C2AB
Introduction ...................................................................................................21
Experimental Procedures ..............................................................................24
Protein Expression and Purification.....................................................24
Diffraction Data Collection..................................................................24
Structure Determination and Refinement ............................................26
viii
Results and Discussion .................................................................................26
Overall Structure of Syt1 C2A-C2B ....................................................27
Variable Loop Conformations of Syt1 C2A-C2B ...............................27
Modulation of Loop 3 in C2A .............................................................29
Interdomain Interactions ......................................................................30
Consequences of the Tandem Arrangement ........................................31
Implications for Other Synaptotagmin Orthologues ............................32
Conclusion ....................................................................................................32
CHAPTER 4 THE C2
DOMAINS OF HUMAN
MECHANICAL PROPERTIES.
SYNAPTOTAGMIN 1
HAVE DISTINCT
Introduction ...................................................................................................38
Experimental Procedures ..............................................................................40
Cloning and expression of C2A and C2A-C2B ...................................40
Cloning and expression of C2AB and I27 ...........................................41
Atomic Force Microscopy ............................................................................42
Equilibrium Denaturation of I27 and C2AB domains .........................42
Single Protein Mechanics ....................................................................42
Steered Molecular Dynamics (SMD) Simulations .......................................43
Results ...........................................................................................................44
Chemical denaturation of C2AB and Titin I27 ....................................44
Mechanical stability of domain C2AB ................................................44
Mechanical stability of C2A domains..................................................45
Speed dependence of C2A vs. C2AB ..................................................46
Molecular basis for differences between C2A and C2B ......................47
Discussion .....................................................................................................49
The stability of human Syt1 C2 domains .............................................49
C2B has a higher mechanical stability than C2A ................................50
Mechanical unfolding intermediate in C2A .........................................50
Potential effects of mechanical forces on Syt1 C2 domain Ca+2 binding
loops .....................................................................................................51
Figures...........................................................................................................53
CHAPTER 5 STRUCTURAL EFFECTS OF MUTATIONS ON SYT I C2A
Introduction ...................................................................................................58
ix
Experimental Procedures ..............................................................................63
Cloning and Mutagenesis of Syt I C2A ...............................................63
Expression and Purification .................................................................64
Crystallization ...............................................................................................69
WT, D232N and D238N ......................................................................69
Y180F ..................................................................................................71
P21 space group ...........................................................................71
I4 space group .............................................................................71
Diffraction Data Collection..................................................................71
D232N and D238N .....................................................................72
Structure Determination and Refinement ............................................73
Results and Discussion .................................................................................74
The three dimensional structure of synaptotagmin C2A WT ..............78
The three dimensional structure of synaptotagmin C2A Y180F. ........79
P21 crystallographic setting for rat syt1 C2A .............................79
I4 crystallographic setting for syt1 C2A .....................................80
Comparison of synaptotagmin I C2A WT and Y180F crystal structures. ....83
The three dimensional structure of synaptotagmin I C2A D232N and D238N.
..............................................................................................................84
D232N ..................................................................................................84
D238N ..................................................................................................88
Lifetime Fluorescence Denaturation of Syt I C2A Mutants .........................89
Inter-molecular crystal contacts ....................................................................92
CHAPTER 6 CONCLUSIONS AND FUTURE DIRECTIONS
96
Appendix Ramachandran plots of C2A mutants. ................................................100
References ............................................................................................................105
Vita .....................................................................................................................117
x
List of Tables
Table 2.1; Crystal parameters, data-collection and processing statistic
18
Table 3.1: Crystallographic data and refinement statistics
23
Table 5.1; Summary of crystallographic statistics for each protein.
69
xi
List of Figures
Figure 1.1: Cartoon representation of a presynaptic nerve terminal. ....................2
Figure 1.2: The SNARE complex. ........................................................................3
Figure 1.3: Topologies of C2 domains type I and II. ............................................4
Figure 1.4: Model of the crystal structure of Synaptotagmin I C2A .....................6
Figure 1.5: Synaptotagmin I C2A inserted into a lipid bilayer. ............................7
Figure 2.1: 20% PAGE gel summarizing human synaptotagmin C2A-C2B
purification. .......................................................................................17
Figure 2.2: Crystals of human synaptotagmin 1 C2A-C2B. ...............................17
Figure 2.3; Self-rotation function calculated using the human synaptotagmin 1
crystals. .............................................................................................18
Figure 3.1: Open and closed conformations of synaptotagmin. ..........................21
Figure 3.2: Superposition of the isolated Syt1 C2A domain ...............................31
Figure 3.3: Stereoview of the C2A-C2B interdomain interactions. ....................32
Figure 3.4: ClustalW alignment of the reported human synaptotagmin paralogues.
...........................................................................................................33
Figure 3.5: The rotameric position of His-237 depends on whether C2A is coupled
to C2B. ..............................................................................................34
Figure 4.1: Equilibrium denaturation for Syt1 C2AB and titin I27 domains. .....36
Figure 4.2: Mechanical properties of C2AB. ......................................................43
Figure 4.3: Mechanical properties of C2A. .........................................................45
Figure 4.4: Mechanical properties of C2A vs. C2B domains..............................47
Figure 4.5: Steered molecular dynamic simulation (SMD) of C2A and C2B
domains. ............................................................................................49
Figure 5.1: Ion exchange chromatograph of Syt I C2A WT. ..............................62
Figure 5.2: 20% SDS PAGE gel of human synaptotagmin I C2A WT. ..............63
xii
Figure 5.3: 20% SDS PAGE gel of human synaptotagmin C2A Y180F. .............64
Figure 5.4: MALDI TOF mass spectrometry graph of purified syt I C2A Y180F.64
Figure: 5.5: Crystals of Syt I p202 C2A Y180F ...................................................67
Figure: 5.6: Syt I C2A D232N..............................................................................67
Figure: 5.7: Syt I C2A WT ...................................................................................67
Figure 5.8: 0.97Å electron density map of synaptotagmin I WT C2A domain. .69
Figure; 5.8: Structural alignment of Syt I C2A WT, Y180F without Ca2+ and Y180F
with Ca+2. ..........................................................................................69
Figure 5.9A: Loop three of Syt I C2A Y180F without bound cation ....................72
Figure 5.9B: Loop three of Syt I C2A Y180F with bound cation..........................72
Figure 5.9C: Loop three of Syt I C2A in the P21 crystal form...............................72
Figure 5.9D: Alignment of loop 3 of all three Y180F structures ...........................72
Figure 5.10: Model of Syt I C2A WT at 0.89Å resolution....................................73
Figure 5.11: Model of Syt I C2A Y180F crystallized in the P21 space group. .....74
Figure 5.12: Model of Syt I C2A Y180F in the I4 space group. ...........................75
Figure 5.13: Comparison of average B-factors for the two, C2A Y180F (I4 crystal
form), molecules in the asymmetric unit. .........................................76
Figure 5.14: Loop3 of Syt I C2A Y180F from the I4 space group. ......................77
Figure 5.15: Structural alignment of C2A WT and Y180F. ..................................78
Figure 5.16: Model of Syt I C2A D232N at a resolution of 1.2Å. ........................80
Figure 5.17: Loop 3 of Syt I C2A D232N. ............................................................81
Figure 5.18: Model of Syt I C2A D238N at a resolution of 0.95Å. ......................83
Figure 5.19: Lifetime thermal denaturation plot of Syt I C2A ..............................86
Figure 5.20A:Crystal
contacts
of
Syt
I
C2A
Y180F
I4
crystal
form.
...........................................................................................................88
Figure 5.20B: Crystal contacts of Syt I C2A Y180F P2 crystal form.
88
xiii
List of Abbreviations
AFM, atomic force microscopy
ASU, asymmetric unit
EPR, electron paramagnetic resonance
GST, glutathione S-transferase
H-bond, hydrogen bond
NMR, nuclear magnetic resonance
PCR, polymerase chain reaction
SNARE, soluble NSF (N-ethylmaleimide-sensitive factor)
attachment protein receptors
Syt, synaptotagmin.
PKC, Protein Kinase C
PS, phosphatidylserine
C2 domain, domain named after the second regulatory domain
PKC.
HEPES, (4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid)
WT, wild type
MBP, maltose binding protein
TEV, Tobacco etch virus
RT, room temperature
AD1, early stop codon that deletes the C2B domain only
AD4, null allele caused by an early stop codon that deletes the
transmembrane and cytoplasmic domains of Syt I
AD3, Y364N mutation in Syt I C2B
SNAP 25, (synaptosome-associated protein of 25,000 daltons)
POPC, 1-Palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine
xiv
POPS, 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphoserine
PIP2, phosphatidylinositiol-4,5-bisphosphate
CNS, central nervous system
FRET, fluorescence resonance energy transfer
HSQC,
Heteronuclear
xv
Single
Quantum
Coherence
CHAPTER 1: INTRODUCTION
This dissertation describes the structural characterization of synaptotagmin I C2A,
C2B, and C2AB. In the first study (Chapter 2), the expression and purification of
synaptotagmin I C2AB and C2A is described. In chapter 3, the X-ray crystal structure of
synaptotagmin I C2AB was determined. The second study (Chapter 4) is the single
molecule analysis of the stability of Syt I C2AB. In the third study, (Chapter 5), the
structural effects of mutations on Syt I C2A are studied.
SYNAPTIC VESICLE FUSION
Membrane fusion is the process whereby two separate lipid bilayers merge to
become one. The best studied biological fusion system is exocytosis. Exocytosis in
neuronal synapses is triggered by Ca2+ and mediates inter-neuronal-signaling.
Neurotransmitter is encapsulated into synaptic vesicles, which are then clustered at the
lipid membrane of the synaptic terminal. Once clustered, the vesicles are in a metastable
state and are subsequently primed for fast Ca2+-triggered fusion with the plasma
membrane (Fig. 1.1). When an action potential arrives at the pre-synaptic terminal,
exocytosis is triggered by an influx of Ca2+ ions through voltage-activated Ca2+ channels
(1). This influx of Ca2+ ions triggers a series of events that ultimately results in the
release of the neurotransmitter from a vesicle into the synaptic cleft. This is among the
fastest known fusion events.
The time between increases in pre-synaptic Ca2+
concentration and a post-synaptic response can be as short as 60 ± 200 µs (2-4). To
accomplish this biological feat, a highly specialized membrane fusion pathway has
evolved at presynaptic nerve terminals. Decades of intense research has led to the
discovery of the many proteins involved in neurotransmitter release.
1
Figure 1.1: Cartoon representation of a presynaptic nerve terminal. Vesicles filled with
neurotransmitter are clustered at the lipid membrane of the synaptic terminal. SNARE
proteins (synaptobrevin, syntaxin, and SNAP-25) form a parallel four helix bundle at the
docking interface. Syt I is a vesicle associated protein that interacts with the SNARE
complex and is the Ca2+ sensor in the neurotransmitter release process. When an action
potential reaches the presynaptic terminus, an influx of Ca2+ occurs. The Ca2+ influx
causes a series of events that ultimately result in the release of neurotransmitter.
PROTEINS INVOLVED IN NEUROTRANSMITTER RELEASE
The membrane fusion machinery is driven by a set of vesicle and target membrane
proteins called SNAREs (soluble NSF (N-ethylmaleimide-sensitive factor) attachment
protein receptors.
The SNARE superfamily of proteins are essential for many
intracellular membrane-fusion events (5). During membrane fusion, the SNARE proteins
combine to form a four helix bundle called a SNARE complex (Fig. 1.2). (6-8). SNARE
proteins can be classified into two categories:
Vesicle or (v) SNAREs and target
membrane or (t) SNAREs. v-SNAREs are incorporated into the membranes of transport
vesicles, and t-SNAREs are located in the membranes of target cell membranes. The
proteins synaptobrevin/VAMP are v SNAREs and syntaxin and SNAP-25 are t SNAREs.
The proteins interact to form a four-helix bundle with SNAP-25 contributing two helices,
2
and synaptobrevin and syntaxin, with their membrane anchors embedded in the vesicle
and plasma membrane, respectively, contribute one helix each (9, 10). The complex
promotes close phospholipid bilayer / vesicle proximity but are not sufficient for
transmitter release alone (11). Synaptotagmin I is a multi-domain vesicle associated
protein that mediates docking, recycling and fusion of vesicles with target membranes.
Figure 1.2: Figure taken from (10). The SNARE complex. The SNARE complex is
composed of synaptobrevin, shown in blue, syntaxin, shown in red and SNAP-25,
shown in green. Synaptobrevin is associated with the vesicle while syntaxin is
associated with the plasma membrane. Cleavage sites for botulism toxin are shown
for each protein. The SNARE complex brings the vesicle containing neurotransmitter
into proximity of the plasma membrane creating a docking interface.
Synaptotagmin family of proteins
Syt I is part of a family of proteins made up of 17 identifiable members.
Presumably, these proteins mediates docking, recycling and fusion of vesicles with target
membranes in many different cell types (12). The expression of synaptotagmin 1 (Syt1)
3
is highest in neuronal tissues, although some synaptotagmins show a widespread tissue
distribution (13, 14). Syt1 plays an essential role in fast synaptic vesicle exocytosis (15),
and is widely considered to be the Ca2+ sensor in calcium dependent neurotransmitter
release (16, 17). It is the most abundant Ca2+ binding protein present on synaptic vesicles
accounting for 7% of total vesicle protein (18, 19). Syt I contains two tandem C2
domains that bind Ca2+, interact with phospholipid and interact with the SNARE
complex, all of which are important components of the neurotransmitter release
machinery (20). The full-length human protein contains approximately 280 amino acids
and is roughly 65 kDa.
Figure 1.3: Model of the crystal structure of Synaptotagmin I C2A at a resolution of
0.89Å without Ca2+ bound. The green represents the β-sandwich structural motif, the
yellow loops at the apex of the molecule labeled loops 1, 2 and 3 are the Ca 2+ binding
loops. Synaptotagmin I C2A binds 3 calcium ions coordinated by five acidic residues,
four aspartic acids and one serine shown. The lipid interacting residues are shown at the
4
top of loop 1 and loop 3. Histidine 237 and Tyrosine 180 are also shown; there is a
hydrogen bond between the residues. The distance between these atoms is 2.6 Å. The
interaction between Tyr 180 and His 237 acts as a support for loop 3 and is critical for the
function of C2A.
C2 domains of synaptotagmin
Synaptotagmin is a multi-domain protein consisting of two tandem C2 domains.
C2 domains in general are named after the second conserved regulatory domain in PKC,
are ubiquitous structural motifs. A large number of proteins containing C2 domains have
been identified, and many are involved in signal transduction or membrane trafficking.
The C2 domain represents the second most abundant lipid binding domain (21). Most,
but not all, C2 domains bind Ca2+ and this interaction is often required for the protein to
interface with lipid membrane (21, 22). The structures for many C2 domains have been
determined as both single domains and as intact proteins (23-28).
Synaptotagmin I is a type I membrane protein (fig 1.3) composed of one transmembrane domain inserted into the synaptic vesicle, a presumably unstructured linker
followed by two tandem C2 domains identified as C2A and C2B (29-31). Each C2
domain is folded in a greek key motif composed of an 8 stranded β-sandwich joined by a
short 9 amino acid linker. Both share considerable structural similarity with the C2
domain of protein kinase C (32), and convey Ca+2-dependent phospholipid-binding
activity to the protein (33, 34).
Synaptotagmin I interacts with Ca+2 ions via four
aspartates and one serine in loops 1 and 3 of each C2 domain. C2A has the ability to bind
3 Ca+2 ions while C2B can bind 2 (23, 35). Synaptotagmins affinity for calcium ion is
fairly low (36), but is markedly enhanced by the interaction of the C2 domains with lipid
bilayers containing acidic phospholipids such as phosphatidylserine (PS) (37).
5
Figure 1.4: Topologies of C2 domains type I and II. Figure taken from (38). C2 domains
occur in two folding variants, Type 1 and Type 2. These two topologies are circularly
permutated relative to each other, and differ only in the connectivity of β-strands 1 and 8.
Despite the differences in β -strand connectivity, most C2 domains share conserved
primary sequences in essential regions of the domain. Synaptotagmin I follows a type I
domain folding pattern.
SYNAPTOTAGMIN FUNCTIONS AS CA2+ SENSOR
Syt I is localized to neuronal secretory vesicles and controls Ca2+-dependent
fusion in the central nervous system (CNS). (19, 39). Syt1 clearly plays a crucial role in
neurological function. Drosophila and mice lacking Syt1 or that contain a point mutation
in the Ca2+-binding C2 domains display a deficiency in fast synchronous neurotransmitter
release (15, 36, 40, 41). The C2 domains of Syt1 cooperate and penetrate membranes in
response to Ca2+ binding and are believed to lower the energy barrier for membrane
fusion (42, 43). C2B binds phosphatidylinositiol-4,5-bisphosphate (PIP2) strongly in a
Ca2+ dependent manner (44). PIP2 is enriched in plasma membrane microdomains at
sites of exocytosis (45). This interaction could steer the membrane penetration activity of
Syt1 towards the target membrane. Recombinant Syt1 exhibits Ca2+-dependent binding
to syntaxin 1, a component of the synaptic vesicle fusion complex (SNARE complex),
6
with half-maximal binding occurring at ~200 μM Ca2+ in agreement with the high, local
Ca2+ concentrations estimated to be required for transmitter release at many synapses
(13).
PHOSPHOLIPID BINDING
Figure1.5: Synaptotagmin I C2A inserted into a lipid bilayer. Loops 1 and 3 undergo
Ca2+ dependent insertion into the lipid membrane. The Ca2+ binding loops are shown in
yellow and the two lipid interacting residues (Phe 234 and Met 173) are shown in red.
Syt I inserts into the membrane approximately 5 Å. While C2A is shown, C2B also binds
membrane in a similar fashion. Both domains preferentially bind to phosphatidylcholine
(PC) and phosphatidylserine (PS) lipds with micromolar efficiency (43). Also shown are
residues Tyr 180 and His 237. The interaction between residues could act as a support
mechanism for loop 3.
Ca2+-triggered synaptic vesicle exocytosis is a critical event in fast synchronous
neurotransmitter release.
Syt I contains two C2 domains (C2A and C2B) that are
independently folded Ca2+-dependent membrane binding domains (38, 46). The C2A
domain is able to bind three calcium ions whereas the C2B domain can bind two. To
bind phospholipid, multiple calcium ions must bind to the top loops in the C2 domain
fold. Binding Ca2+ has little effect on the overall structure of the individual C2 domains;
however, binding stabilizes the Ca2+ coordinating loops, as well as creates a favorable
environment for phospholipid interaction (23, 47). In the absence of Ca2+, there is
predominately a negative electrostatic potential in the Ca2+ binding pocket that prevents
the protein from efficiently associating with phospholipid. When calcium ion binds, the
7
negative potential is quenched resulting in a more positive potential that allows
phospholipid membrane association. The process is known as the electrostatic switch
(47). Loops 1 and 3 are key in interacting with Ca2+ and phospholipids, since the residues
for coordinating Ca2+ and insertion into the membrane are located on these loops (48).
The C2 domains of Syt1 preferentially bind membrane surfaces composed of acidic
phospholipids phosphatidylcholine (PC) and phosphatidylserine (PS) in a Ca2+-dependent
manner with a micromolar lipid dissociation constant (34, 43, 46).
PAST STRUCTURAL STUDIES
A considerable amount of structural analysis, both X-ray and NMR has been done
on synaptotagmin (23, Cheng, 2004 #1204, 47, 49-51). The initial X-ray crystal structure
of the C2A domain of Syt1 was solved to a resolution of 1.9 Å (23). They reported that
the C2 domain forms a greek key motif that they dubbed the C2 key. The key consists of
an eight-stranded ß-sandwich constructed around a conserved four-stranded motif. In
addition, they reported that the Ca2+ binding pocket consists of a cup-shaped depression
between two polypeptide loops located at the N- and C-termini of the C2-key. Later, the
crystal structure of synaptotagmin III C2AB was published (51). The 3.2 Å structure
consisted of both cytosolic C2 domains. In this structure, the two C2 domains do not
interact but appear to be two separate proteins tethered together. It was widely believed
that the domains acted separately and did not interact with one another. The Rizo group
(47) solved the NMR structure of synaptotagmin I C2A in the Ca2+ free and the Ca2+
bound form.
The study attempted to elucidate the structural consequences of Ca2+
binding to the C2A domain. They concluded that three Ca2+ molecules are bound to C2A
and that these ions are coordinated by five aspartate side chains and one serine side chain.
The also concluded that Ca2+ binding does not result in a significant conformational
change in the domain. They concluded that the main effect of Ca2+ binding to the C2A
domain is a change in its electrostatic potential rather than the structure (47). The Cheng
8
lab group (50) reported on the crystal structure of Syt1 C2B domain. They crystallized
the domain in the ion bound form as well as the unbound form at resolution of 1.5 Å and
1.04 Å respectively. They reported that the C2B domain binds two calcium ions that are
coordinated by four highly conserved aspartic acid residues (D303, D309, D363, and
D365) and two main chain oxygens (M302 and Y364).
They did not observe a
significant structural change between the Ca2+ bound and Ca2+ unbound forms of the
protein. Similar to the C2A case, no global structural change was seen in C2B; however,
small local changes were observed at the Ca2+ binding pocket. They noted that Ca2+
binding had a stabilizing effect on loop 1 of the binding pocket. In the unbound form, no
electron density could be observed for the loop where as the loop was clearly visible in
the Ca2+ bound form. Their result is consistent with previous NMR studies. (49) reported
that cation-binding does not impact on the global structure of the C2B domain, but rather
stabilizes the conformation of loop 1. These studies were consistent in that none saw a
significant structural change either in the individual C2 domains or between the tandem
C2 domains upon Ca2+ binding. The Arac lab group (52) used NMR to determine if the
domains were interacting in the presence as well as the absence of Ca2+. They used 1H15
N HSQC spectra of the C2A domain alone, the C2B domain alone and both domains
linked together. They saw no difference in the spectra between the individual domains
and the linked domain whether Ca2+ was present or absent. They concluded that there is
no interaction between the domains. The majority of these studies focused specifically
on the individual domains themselves and not the tandem C2 domains. There are a few
studies, however that used the connected C2 domains and indicate an interaction between
the two domains (53-55).
Fluorescence resonance energy transfer was used on synaptotagmin I C2AB to
probe Ca2+ induced structural changes. (54)
The study and found that at Ca2+
concentrations required for fusion, a conformational change occurs that brings the two C2
domains in Syt1 closer together. This study, however, has been questioned (52) who
9
found no interaction using NMR techniques. They contend that the results were likely
from misfolded protein caused by the attachment of the coumarin label to a partially
buried cysteine or due to the hydrophobicity of the probe itself. Garcia and colleagues
(53) used partial proteolysis to show that Syt1 undergoes a conformational change as a
function of Ca2+ binding.
They report that the tandem C2 domains do undergo a
structural change when bound to Ca2+. We recently published the crystal structure of
Syt1 C2AB in the absence of Ca2+ (55). The structure shows an interaction between C2A
and C2B in the absence of Ca2+. The structure is further discussed in chapter 2.
While a considerable amount of structural information has been gathered on the
individual domains of Syt I, the work has largely been done from the assumption that
synaptotagmin is a very static molecule.
The majority of studies report no
conformational change in the C2 domains and no interaction between the domains. A
highly conserved motif in the C2 family denoted as the "...SDPYVKV..." motif has been
known, but it’s relevance has yet to be reported on. This study presents evidence that the
C2 domains of synaptotagmin are not static domains, but dynamic proteins that interact
with one another and have important regulatory motifs. We show that residues in the
"...SDPYVKV..." motif in Syt I are involved in regulating the position of loop 3 of the
calcium ion binding pocket as well as how the protein interacts with calcium ion.
Hypothesis and Goals:
First, crucial residues in the Ca+2 binding pocket of C2 domains can regulate the
position of loop 3 to alter how the domain interacts with Ca+2 and phospholipid. Second,
the domains of Synaptotagmin I C2A and C2B are distinct domains and have distinct
mechanical properties. This property is likely an essential requirement for Ca+2 dependent
exocytosis.
Aim 1: Determine crystal structure of Syt1 C2A-C2B.
10
Aim 2: Investigate the Ca+2 binding pocket residues (D232, D238, Y180). Do
these residues aid in shaping Loop 3?
Aim 3: Investigate the mechanical properties of Syt1 C2A and Syt1 C2AB.
Chapter 2: Purification, crystallization and X-ray
diffraction analysis of human synaptotagmin 1 C2AC2B.
(Studies in this chapter are based on studies published in the paper: Montes, M., Fuson,
K.L., Sutton, R.B., Robert, J.J., (2006) Purification, crystallization and X-ray diffraction
analysis of human synaptotagmin 1 C2A-C2B. Acta Crystallogr Sect F Struct Biol Cryst
Commun. 62(Pt 9):926-9, and with permission.)
INTRODUCTION
Ca2+-dependent release of neurotransmitters into the synaptic space is one of the
most fundamental concepts in modern neuroscience. While this phenomenon has been
thoroughly described at the cellular level, identification of the molecular agents
responsible for this activity has been challenging. Over the years, several candidate
proteins have been posited as the Ca2+ receptor; these include an annexin (56), protein
kinase C (57) and calmodulin (57, 58). However, the vesicle-localized protein
synaptotagmin has recently gained acceptance as the Ca+2 receptor in neurons (20, 59).
In general, the synaptotagmin family mediates docking, recycling and fusion of
vesicles with target membranes (60). While all isoforms possess a single transmembrane
span, the main function of this protein is mediated through its peripheral association with
acidic components of target phospholipid membranes. Some isoforms of synaptotagmin
have evolved a sensitivity to Ca2+ gradients (61), while others have dispensed with this
activity (62). To date, there have been 17 paralogs of synaptotagmin identified within the
human genome (63). The cellular distribution of these isoforms ranges from ubiquitous
expression (in most cell types) to exclusive localization in the brain, as is the case for
synaptotagmin 1.
11
The molecular weight of the complete human synaptotagmin 1 protein is
approximately 65 kDa. The short amino-terminal extracellular portion of the molecule is
glycosylated and serves as the receptor for Botulinum neurotoxins (subtypes B and G;
(29-31). The single transmembrane helix of synaptotagmin 1 is followed by a long
tethering linker which is rich in arginine and lysine residues. This linker is followed by
the tandem C2 domains that are localized to the carboxy-terminal end of the molecule.
These are labeled C2A and C2B. Both share considerable structural similarity with the
C2 domain of protein kinase C (32) and impart Ca2+-dependent phospholipid-binding
activity to the protein (33, 34).
Human synaptotagmin 1 C2A-C2B possesses ~280 amino acids. Each C2 domain
is composed of an eight-stranded β-sandwich (23, 49, 64) and the two domains are joined
by a short approximately five-amino-acid linker. To bind phospholipids, multiple calcium
ions must bind in a cleft at one end of the fold. The divalent-cation-binding loops colocalize with residues implicated in phospholipid association. While there are published
reports of Ca2+-dependent structural changes in synaptotagmin 1 (53), there is little
structural evidence to support large-scale reorganizations of the C2 domains.
A considerable amount of structural analysis, both X-ray and NMR (23, 47, 49,
50), has been devoted to the isolated C2 domains of synaptotagmin, yet little has been
accomplished on the structure of the functional C2A-C2B pair. However, some evidence
from NMR studies and X-ray structural studies suggests that the two domains do not
interact (51, 52). Here, we report the crystallization and preliminary X-ray diffraction
results of this domain in order to gain insight into the possible intercommunication
between C2 domains and to address the possible role of oligomerization in synaptotagmin
biology
12
Cloning, expression and purification
Human synaptotagmin 1 (Syt1) C2A-C2B was obtained via PCR from a human
hippocampal Quick-Clone cDNA library (Clontech). An 858-nucleotide fragment
comprising Syt1 C2A-C2B residues 140–422 was directly cloned into PCR2.1
(Invitrogen), excised using BamHI and XhoI and subsequently subcloned into pGEX4T-1
(GE-Healthcare). DNA-sequence analysis confirmed insertion in the correct reading
frame with respect to the GST fusion partner. BL21 cells containing the Rosetta plasmid
were transformed with the pGEX4T-1-SytI plasmid and the resultant colonies were
grown in 10 l ECPM1 medium (65) in a BioFlo 3000 fermentor at 310 K with 100 µg
ml−1 ampicillin as the selection agent. Heterologous expression of the synaptotagmin
fusion protein was initiated with 400 µM IPTG and expression took place for 4 h at 298
K. The culture was immediately centrifuged at 277 K and the pellets were frozen in liquid
nitrogen and stored at 193 K.
Frozen cells were thawed in cold lysis buffer (50 mM sodium phosphate buffer,
50 mM Tris, 300 mM NaCl, 5 mM DTT, 10 mM EDTA, 0.5 mM PMSF, 1% Triton X100 pH 7.4). A hand blender was used to homogenize the cells. They were disrupted
using a Microfluidizer M110-EH (Microfluidics). The lysate was then centrifuged at 40
000 rev min−1 for 60 min at 277 K. All subsequent chromatography columns and
chromatography media were obtained from GE-Healthcare. Buffer exchanges were
performed using G-25 Sephadex medium packed in an XK 26/20 column. The lysate
supernatant was filtered using 0.2 µm syringe filters and loaded at a flow rate of 300 cm
h−1 onto Glutathione Sepharose 4 Fast Flow packed in an XK 26/20 column equilibrated
with lysis buffer. The column was then washed with the following buffers at a flow rate
of 600 cm h−1 until ΔA 280 < 5 mAU: lysis buffer, wash buffer (50 mM sodium phosphate
buffer, 50 mM Tris, 300 mM NaCl, 5 mM DTT pH 7.4), nuclease buffer (50 mM sodium
phosphate buffer, 200 µg ml−1 DNase, 20 µg ml−1 RNase, 2 mM MgCl2 pH 7.4), wash
buffer, high-salt buffer (wash buffer + 1 M NaCl pH 7.4) and finally wash buffer. The
13
fusion protein was eluted with elution buffer (wash buffer + 15 mM reduced glutathione
pH 8.0). The eluant was buffer-exchanged into 50 mM sodium phosphate, 150 mM NaCl
pH 7.4. Human α-thrombin (Haematologic Technologies Inc) was added to the eluant at
50 NIH units per milligram of fusion protein. The protease cleavage reaction was allowed
to proceed for 36 h at 297 K with occasional gentle mixing. The cleavage product was
spun, filtered using a 0.2 µm syringe filter, buffer-exchanged into wash buffer and loaded
onto the Glutathione Sepharose 4 Fast Flow column at 300 cm h−1. The flowthrough was
collected and buffer-exchanged into IEXA buffer (50 mM HEPES, 150 mM NaCl pH 7.2)
and loaded onto an equilibrated SP Sepharose HP XK16/20 column at 480 cm h−1,
washed until the A
280
was stable and developed using a gradient from IEXA buffer to
IEXB buffer (IEXA + 1 M NaCl pH 7.2) over 20 column volumes. The desired fractions
were combined and concentrated using 10 kDa molecular-weight cutoff Ultra Centrifugal
Filter Devices (Millipore). Gel filtration was performed using Superdex 75 Prep Grade in
an XK 16/60 column equilibrated with 25 mM sodium phosphate, 50 mM Tris, 100 mM
NaCl pH 7.2. The pure protein fractions were mixed and concentrated to 17 mg ml−1
using YM-10 centrifugal filters (Millipore) for immediate crystallization setup.
The protein quantitation and overall purity at each step was assessed using the
Bradford method and SDS–PAGE (Fig. 1). MALDI–TOF was performed on purified
samples in the University of Texas Medical Branch Mass Spectrometry Laboratory. The
average calculated molecular weight for the fragment 140–422 of human synaptotagmin
C2A-C2B after thrombin cleavage is 32 487.47 Da; MALDI analysis yielded a weight of
32 635.1 Da. The identity of the resultant protein as synaptotagmin 1 C2A-C2B was also
confirmed by its mostly β-sheet circular-dichroism signature (data not shown) and
Western blot analysis (data not shown) using commercially available polyclonal Ab
produced against the C2A specific peptide from synaptotagmin 1 (Synaptic Systems,
GmbH).
14
Crystallization and data collection
Initial crystallization conditions were obtained by screening with crystallization
kits from Hampton Research (Laguna Niguel, CA, USA) and Molecular Dimensions Inc.
(Apopka, FL, USA). All experiments were performed using the hanging-drop and sittingdrop vapor-diffusion method at 293 K. 2 µl of 17 mg ml−1 protein solution was mixed with
2 µl reservoir solution over wells containing 500 µl precipitant solution. After one week,
small crystals appeared in Hampton SaltRx Screen formulation 12 (3.2 M sodium chloride,
100 mM sodium acetate pH 4.6). Crystallization conditions were optimized to 3 M sodium
chloride, 75 mM sodium acetate pH 4.5. Microseeding techniques yielded crystals of
dimensions 0.1 × 0.1 × 0.05 mm that were suitable for diffraction studies (Fig. 2 ). While
the precipitant alone was rather acidic, the resultant 2 µl protein solution + 2 µl precipitant
had a pH of ~5.8.
X-ray diffraction
The crystals for data collection were harvested from sitting drops when 0.1 mm in
size. These crystals were picked up in 0.1–0.2 mm nylon loops, soaked in 5 M sodium
chloride, 75 mM sodium acetate pH 4.5 for 10 min and then flash-frozen in liquid
nitrogen. Initial X-ray diffraction experiments were performed at 100 K with an oscillation
range of 1° per frame over 30 min using a MacScience MO6XHF X-ray generator and a
MacScience DIP2030H-VLM dual 30 cm diameter imaging-plate detector at the
University of Texas Medical Branch X-ray Crystallography Core facility. Final X-ray data
sets were collected at beamline 11-1 at the Stanford Synchrotron Radiation Laboratory
(SSRL; Stanford, CA, USA) at 100 K. All data were processed with MOSFLM (66) and
15
CCP4 (67). The optimal data-collection strategy was computed using the ‘Strategy’ option
in MOSFLM. Structure factors were calculated with TRUNCATE (68).
RESULTS AND DISCUSSION
Crystals grown in sodium chloride diffracted to 2.7 Å at SSRL. The refined
mosaicity was low (0.6°). We tried to soak these crystals in solutions containing divalent
cations, such as ~100 µM Ca2+ or Pb2+; however, the diffraction limit decreased and the
mosaicity increased markedly. Attempts to co-crystallize this protein with calcium ion
yielded crystals that diffracted to ~25 Å resolution (personal communication from Dr
Byron DeLaBarre, Stanford University). These results could be indicative of a substantial
conformational change within these crystals upon divalent cation binding.
The crystal belong to space group P212121, with unit-cell parameters a = 82.37, b
= 86.31, c = 147.2 Å (Table 2.1). In this case, the Matthews coefficient is ambiguous. For
two molecules in the asymmetric unit, the Matthews coefficient is calculated as 4.09 Å3
Da−1. This implies ~70% solvent content in the crystal. However, three molecules per
asymmetric unit yields a Matthews coefficient of 2.73 Å3 Da−1 and 55% solvent content.
Two molecules per asymmetric unit is consistent with the rather high solvent content
found in the related synaptotagmin III structure (51); however, the value corresponding to
three molecules per asymmetric unit is more characteristic of well diffracting protein
crystals. To resolve this ambiguity, a self-rotation function was calculated using the
available X-ray data (Fig. 2.3). At κ = 180°, there were two major peaks that could not be
attributed to crystallographic symmetry. Both these peaks were found ~45° about the
crystallographic c axis. There were no major peaks observed at κ = 120°. Therefore, we
conclude that this crystal form possesses a non-crystallographic dimer of synaptotagmin
molecules. The self-rotation function was calculated using GLRF (69). The Patterson
integration radius was 40 Å and data were used in the resolution range 15.0–4.0 Å.
16
While crystallographic and NMR structures of C2A (23, 70) and C2B (49, 50)
exist, the functional synaptotagmin protein comprises the two domains in tandem.
Therefore, the structural information obtained from this crystal structure at atomic
resolution will certainly augment our understanding of Ca2+-dependent exocytosis and
synaptotagmin biology in general. Also, since this crystal represents the first instance of
synaptotagmin as a homodimer, it is hoped that we will gain insight into the structural
mechanism of synaptotagmin oligomerization and perhaps the interdomain interactions
between C2A and C2B.
Figure 2.1: 20% PAGE gel summarizing human synaptotagmin C2A-C2B purification.
Lane 1, molecular-weight markers (kDa); lane 2, pre-induction cell lysate; lane 3, postIPTG induction cell lysate; lane 4, glutathione-Sepharose purified fusion protein; lane 5,
GST-Syt1-C2A-C2B cut with thrombin; lane 6, thrombin cut protein re-purified over
glutathione-Sepharose; lane 7, overloaded sample of Mono S/gel-filtered final protein.
17
Figure 2.2: Crystals of human synaptotagmin 1 C2A-C2B. The largest crystals are
approximately 0.1 mm in size.
Values in parentheses are for the highest resolution shell (2.85–2.70 Å).
Synchrotron radiation
Beamline 11-1, SSRL, λ = 0.979 Å
Detector
ADSC Quantum 4 CCD, Dfx = 400 mm
Space group
P212121
Unit-cell parameters (Å) a = 82.37, b = 86.31, c = 147.2
Resolution range (Å)
50.0–2.70
Total reflections
116148
Unique reflections
29346
Completeness (%)
99.5 (99.9)
Rmerge (%)
8.6 (44)
I/σ(I)
13 (2.8)
3
Unit-cell volume (Å )
†
1046000
Solvent content (%)
70
Multiplicity
4.0 (4.0)
Table 2.1; Crystal parameters, data-collection and processing statistics
†Assuming
two molecules in the asymmetric unit.
18
Figure 2.3; Self-rotation function calculated using the human synaptotagmin 1 crystals.
A self-rotation search with κ = 180° was used to identify the non-crystallographic
twofold axis of symmetry. Contours start at 2 standard deviations, with intervals of 0.5.
Calculations and plots were performed and produced using GLRF (69).
19
CHAPTER 3: X-RAY STRUCTURE OF SYT I C2AB
(Studies in this chapter have been published in the paper: Fuson, K.L., Montes, M.,
Robert, J.J., Sutton, R.B. (2007) Structure of human synaptotagmin 1 C2AB in the
absence of Ca2+ reveals a novel domain association. Biochemistry 46: 13041-13048, and
with permission.)
INTRODUCTION
Ca2+-dependent release of neurotransmitter into the synaptic space is one of the
fundamental tenets of modern neurobiology. This aspect of neuronal function has been
studied for many years, yet the molecular details are only now being elucidated. The
process begins with a neurotransmitter-filled synaptic vesicle progressing from a docked
position to a primed, fusion-competent position next to the target membrane. Once the
vesicle membrane and the target membrane fuse, neurotransmitter is released into the
synaptic space. The merger of these two distinct bilayers, and the concomitant formation
of a lipidic fusion pore, is mediated by Ca2+ and a number of proteins specific for the
task. These include the protein complex formed between vesicle-localized R-SNAREs1
such as synaptobrevin and the Q-SNAREs such as SNAP-25 and syntaxin. This protein
complex either acts as a staging platform for other constituents required for vesicle fusion
(71) or may directly participate in the actual fusion event itself (11). Regardless of the
specific mechanism, the fusion of vesicle membranes with presynaptic membranes
requires a calcium ion sensor or trigger. The C2A-C2B domains of synaptotagmin 1
(Syt1) possess the Ca2+-dependent phospholipid-binding activity (20) and the SNAREbinding activity predicted for such a trigger (72).
Syt1 is a vesicle-localized transmembrane protein with two tandem C2 domains
(C2A and C2B) at the C-terminus of the protein. There are two possible explanations for
the tandem C2 domain architecture of Syt1. Either the two domains are joined by a
flexible linker and are completely independent or there is some degree of cooperativity
20
between the domains that is essential to the overall function of the protein. These two
scenarios are not mutually exclusive, as evidence exists for both. NMR analysis of Syt1
C2A-C2B concluded that there was no detectable interaction between the domains (52).
Also, the crystal structure of Syt3, a homologue of Syt1, displays no evidence of
interdomain interaction (51) (Figure 3.1B). On the other hand, the C2 domains of Syt1
are known to function synergistically when compared to isolated C2 domains (73).
Subsequent experiments concluded that a close physical coupling of the tandem domains
is critical for function (74).
Each C2 domain of synaptotagmin 1 is formed from eight -strands arranged
around a Greek key topology, known as the C2 key (23). The curvature of the domains is
governed by a series of tandem -bulges, some of which are unique to the C2 domain.
Both the C2A and C2B domains of Syt1 have extended loops at one end of the domain
(loops 1-3) that have been specialized for Ca2+/phospholipid binding (75). Three ions can
bind in a cuplike depression formed from these loops, but only loops 1 and 3 contribute
calcium ion-binding residues. Loop 2 contributes to the overall shape of the Ca2+-binding
pocket, but does not directly participate in the coordination of cations. Hydrophobic
residues at the apexes of loop 1 and loop 3 embed the domain in the target membrane (76,
77). In the isoforms of Syt that bind Ca2+, five conserved acidic residues on loops 1 and 3
coordinate the binding of Ca2+. Both X-ray studies and NMR analysis of the cationbinding properties of the Syt1 C2A domain confirm that the C2A domain can
accommodate three calcium ions (26, 75), while the C2B domain binds two (50, 78) (49,
50). However, the isolated C2 domains show no gross structural change when the Ca2+free and Ca2+-bound states are compared (79); therefore, any structural change associated
with full-length synaptotagmin function must rely on relative domain motions and not
intradomain changes.
21
Figure 3.1: Open and closed conformations of synaptotagmin. (A) Stereoview of Syt1
C2A-C2B in the closed configuration. The C2A domain is colored as green ribbons. The
C2B domain is colored as blue ribbons. The six known Ca2+-binding residues are shown
as sticks. (B) Stereodiagram of synaptotagmin 3 C2A-C2B in the open configuration
(PDB code 1DQV). The C2A domain in Syt3 is rendered as green ribbons. The C2B
domain in Syt3 is rendered as blue ribbons. Yellow spheres represent magnesium ions.
EXPERIMENTAL PROCEDURES
Protein expression and purification
Human synaptotagmin 1 C2A-C2B was obtained via PCR from a human
hippocampal Quick-Clone cDNA library (Clontech). An 858-nucleotide fragment
comprising Syt1 C2A-C2B residues 141-422 was directly cloned into PCR2.1
(Invitrogen), excised using BamH1 and Xho1, and subsequently subcloned into pGEX4T1 (GE-Healthcare). DNA sequence analysis confirmed insertion in the correct reading
frame with respect to the GST fusion partner. The cytosolic fragment of human
22
synaptotagmin 1, containing only the cytosolic C2A and C2B, was overexpressed in
Escherichia coli as a GST fusion protein (80) and purified.
Diffraction data collection
Data were collected at the Stanford Synchrotron Radiation Laboratory (SSRL) on
beamline 11-1 at a wavelength of 0.97 Å (Table 3.1). The data sets were collected at 100
K using an ADSC image plate detector. Data were integrated, reduced, and scaled using
HKL2000 (81). The crystals were indexed in the orthorhombic space group P212121; data
statistics are summarized in Table 3.1.
Structure Determination and Refinement
Crystals were grown in the absence of Ca2+ as described elsewhere (80). The
structure of synaptotagmin 1 C2A-C2B was solved via molecular replacement using
Phaser (82) using diffraction data collected from one crystal to 2.7 Å (Table 3.1). Initial
model coordinates were obtained using the C2A domain (PDB code 1BYN) and the C2B
domain (PDB code 1TJX). Model building was done with Coot (83). Refinement of the
model was carried out with CNS with a random subset of all data set aside for calculation
of Rfree (10%). Manual adjustments to the models were carried out with Coot. After
refinement of the protein was complete, solvent molecules were assessed followed by
manual adjustments. Chloride ions were assigned on the basis of the chemical
environment, the refined B factor vs that of the other water molecules, and the available
coordination potential. The structure of C2A-C2B was verified by examining a simulated
annealing omit map generated with CNS v1.2 (Figure S1 in the Supporting Information).
There were no residues in disallowed regions of the Ramachandran map (Table 3.1). All
figures were rendered with Pymol (84).
23
Table 3.1: Crystallographic data and refinement statistics
Crystallographic Data
Beamline 11-1, SSRL, λ = 0.979 Å
Detector
ADSC Quantum 4 CCD, D =400 mm
Space Group
P2 2 2
Unit-Cell Parameters
a= 82.4 Å b= 86.3 Å c= 147.2 Å
Resolution Range (Å)
50 - 2.7
Total Reflections
116148
Unique Reflections
29346
Completeness (%)*
99.5 (99.9)
R merge (%)*
8.6 (44)
fx
1 1 1
I/σ (I)*
13 (2.8)
3
Unit cell Volume (Å )
Solvent content *
Multiplicity*
1,046,000
70 %
4.0 (4.0)
Refinement Statistics
Residue range for Refinement (Å)
50 – 2.7
Total reflections used
29304
r.m.s.d bond lengths (Å)
0.007
r.m.s.d. angles (°)
1.3
Crystallographic R-factor (%)
23
R-free ¶ (%)
25.3
*Numbers in parenthesis correspond to the highest resolution shell of data, which was
2.85-2.7 Å
¶Ten percent of the truncated data set was excluded from refinement to calculate R-free
RESULTS AND DISCUSSION
To examine whether the C2 domains of Syt1 interact with each other prior to a
Ca2+-dependent switch, we solved the 2.7 Å crystal structure (Table 3.1) of the complete
cytosolic portion of Syt1 C2AB (Figure 3.1A). The protein was purified in the presence
of EDTA, and no additional sources of divalent cations were added at the crystallization
stage. We show that the two C2 domains of Syt1 are capable of forming specific
interdomain interactions between amino acids on the H-A helix of C2B and residues on
loops 1-3 of C2A. These interactions distort loop 3 of C2A relative to the conformation
of the same loop in the isolated C2A domain. The two Syt1 C2AB molecules in the
24
asymmetric unit show the same interactions; further, there are no crystal contacts that
would artificially distort loop 3 of C2A. Therefore, this orientation likely represents the
interdomain interactions that are present in Syt1 prior to the Ca2+ flux.
Overall Structure of Syt1 C2A-C2B
As this is the first atomic structure of Syt1 C2A-C2B, we have defined the extent
of the C2 domains, as well as the linker, on the basis of their structural contributions to
the overall fold of the protein. The residue numbering used throughout will correspond to
the well-described rat Syt1 protein sequence, unless otherwise noted. The human Syt1
primary sequence is essentially identical to the rat/mouse sequence with the exception of
one additional amino acid at the extreme N-terminus. We define a residue as part of the
C2 domain "motif" if it interacts with other amino acids in the fold either by contributing
a hydrogen bond to peptide backbone residues or by packing next to other secondary
structural elements within the C2 domain. By this definition, the C2A domain extends
from residue Leu-142 to Gln-263. The nine-residue linker extends from Ser-264 to Lys272. The C2B domain of Syt1 extends from Leu-273 to Lys-423. The two C2 domains of
human Syt1 share ~33% primary sequence identity; however, C2B possesses additional
residues that correspond to two -helices near its C-terminus (H-A and H-B) (49, 50).
Primary sequence alignment of all the known human isoforms of synaptotagmin show
that while the H-A helix is a common feature of most C2B domains, the H-B helix is
present in only a subset of the paralogues (Figure 3.4).
Variable Loop Conformations of Syt1 C2A-C2B
Syt1 crystallizes as a dimer in the asymmetric unit. While we do not ascribe
physiological meaning to the dimer itself, it does allow us to compare the loop structures
between the two molecules of the asymmetric unit and the isolated C2 domains, within
the limits of our resolution range and refinement protocol. In this Syt1 C2A-C2B crystal
25
structure, the tertiary folds of both C2B domains are similar. Further, the fold of the C2B
domains in our crystal is similar to that of the isolated C2B domain (49, 50). The rmsd
between the isolated C2B structure and the C2B domain in the context of the tandem C2
domains is 0.58 Å over all C residues. We conclude that no part of C2B undergoes
significant structural change in the context of C2A-C2B.
When the two C2A domains present in the crystallographic asymmetric unit are
compared to the isolated Syt1 C2A domain, there are significant differences in loops 1
and 3 (Figure 3.2B). These differences are partly due to the hingelike behavior of the
loops in the unliganded state of the C2A domains in this crystal. Because of this hinge
motion, Asp-172, one of the Ca2+-binding residues on loop 1, was traced near the surface
of the domain in one molecule, while the other conformation was directed in toward the
Ca2+-binding pocket (Figure 3.2A). The flexible nature of this loop was not observed for
C2B, as both loops 1 and 3 appear to be well ordered in this crystal form.
The structural variations observed in loop 3 of C2A are evident when C2A-C2B is
compared to either the isolated Syt1 C2A crystal structure (with no Ca2+) (23) or Syt1
C2A with saturating Ca2+ (47) (Figure 3.2A). Regardless of the Ca2+ occupancy of the
isolated C2A domain, the absolute position of loop 3 resembles that of the C2B domains;
consequently, the Ca2+-associating residues are oriented to bind calcium ion. However, in
the C2A-C2B domain structure, this loop is restrained in an alternate conformation that is
not consistent with the typical Ca2+ coordination geometry previously observed in the
C2A domain. Typical calcium ion coordination by proteins utilizes a pentagonal
bipyramidal array with an average 2.4 Å separation to oxygen atoms (85). This type of
coordination is not possible while loop 3 is restrained by C2B. Therefore, given this
geometry, one would not expect C2A to bind Ca2+ in the presence of C2B while in the
closed conformation.
26
Modulation of Loop 3 in C2A
The stabilization of loop 3 in this alternate position involves key residues in the
"...SDPYVKV..." sequence in C2A. This motif is highly conserved across many C2
sequences, so the potential to modulate loop 3 may be a common feature of the C2
domain (Figure 3.4). The Ca2+-binding conformation of loop 3 in the isolated C2A
domain is stabilized by a hydrogen bond between His-237 (H-bond acceptor) and the
central Tyr in the sequence motif listed above (Tyr-180, the H-bond donor) (Figure
2.5B). However, in the human Syt1 C2A-C2B structure, an alternate interaction of His237 with Thr-406 in the C2B domain allows loop 3 in C2A to collapse (Figure 3.5A).
The importance of this Tyr on -sheet 3 at this special position is supported by genetic
evidence.
In the Drosophila Syt1 C2B domain, the mutation of Tyr-364 to Asn (known as
the AD3 mutation) abolishes Ca2+-evoked neurotransmitter release (86, 87). By raising
extracellular Ca2+ from 400 M to 6 mM, exocytosis in these mutants can be partially
rescued (88). This implies that the divalent cation-binding machinery is still present in
C2B, but the affinity for Ca2+ has been drastically altered. The marked phenotype of this
mutation clearly illustrates that there is a direct link between Tyr-364 and Ca2+ affinity.
To understand the relationship between our structure of human Syt1 C2AB and this
mutation in Drosophila, we computed a homology model of Drosophila Syt1 C2B based
on the primary structure alignment of all available Syt1 orthologues and our X-ray
structure of human Syt1 C2B (data not shown). In this model, Tyr-364 in Drosophila
Syt1 C2B is homologous to the Tyr-180 position in the human Syt1 C2A domain. The
model also predicts that Ser-423 (on loop 3) serves as the H-bond acceptor of Tyr-364 ( sheet 3). We hypothesize that the phenotype of the AD3 mutation results from the
removal of the hydrogen bond between Tyr-364 on -sheet 3 and Ser-423 on loop 3 of
Drosophila C2B, thus collapsing loop 3 of C2B. The collapse of loop 3 repositions the
Ca2+-binding residues, thereby increasing the amount of Ca2+ needed to rescue function.
27
A homologous process may be occurring in human Syt1 C2A by the close proximity of
C2B. Since C2B reorients His-237, the H-bonding acceptor to Tyr-180, the same loss of
Ca2+ affinity could be occurring in C2A (Figure 3.5B). Therefore, the AD3 mutation may
mimic the behavior of the wild-type C2A domain under the influence of C2B.
Interdomain Interactions
The conformation of loop 3 in C2A is restrained exclusively by the interdomain
interactions described below. As there are no significant contacts made by
crystallographically related molecules, we conclude that the interactions we observe are
possible in the native protein. Almost all of the amino acid interactions that link C2A
with C2B are located on the H-A -helix of C2B and loops 1-3 of the divalent cationbinding pocket of C2A (Figure 3.3). Arg-388 on the H-A helix of C2B forms a saltbridge interaction with Asp-178 located on loop 1 of C2A. In previous structural analysis,
Asp-178 contributes a single carbonyl oxygen to the highest affinity Ca2+-binding site
(23). When Asp-178 is mutated to Asn in the isolated C2A domain, Ca2+ binding is
disrupted (35, 40). Since the observed interaction between Asp-178 and Arg-388 directly
links C2B to calcium ion-binding residues in C2A, it is tempting to conclude that this
interaction may represent a switch or trigger to link Ca2+ occupancy to Syt1 function.
However, when this mutation was introduced into Drosophila, no significant phenotype
was observed (89). While Asp-178 is certainly important to the overall function of Syt1,
it is clear that this residue is not solely responsible for the interactions with C2B. A
second interdomain interaction observed between C2A and C2B is contributed by Asp392 of C2B. This residue forms a bifurcated salt bridge between Asp-392 and Arg-199 on
loop 2 in C2A and between Asp-392 and Arg-233 on loop 3. Arg-199 is also a notable
residue as it participates in several activities of C2A (interactions with phospholipids (77)
and SNARE components) and is a contributor to the electrostatic switch of C2A (90).
28
Consequences of the Tandem Arrangement
Most C2 domains share two common functions: The first is as an adapter module
that links a client protein or a neighboring motif to a phospholipid bilayer through the
hydrophobic residues at the tips of the C2 domain (91), and the second function is
mediated through the effector-binding polybasic regions (92, 93). In the closed
conformation, the association with C2B affects residues that mediate both of these
activities. The polybasic region of C2A is composed of four contiguous Lys residues at
the amino terminus of the -strand and three basic amino acids at the carboxy terminus of
-strand 4. While the biological function of the polybasic domain of C2A in Syt1 function
is not as clear as that of C2B, this region of C2A is almost completely occluded by its
interaction with C2B. Upon forming the closed conformation, ~890 Å2 of solventaccessible surface area is buried per C2 domain. This degree of domain contact is
comparable to that of other observed protein dimer interfaces and is indicative of
associations that require weaker interactions (94). Thus, the closed conformation of Syt1
may obstruct premature interactions with effectors such as phospholipids, syntaxin, or the
voltage-gated Ca2+ channel (90, 95, 96). Another notable feature of C2B is the highly
conserved "WHxL" sequence in C2B. This motif has been implicated in docking events
and neurexin binding of Syt1 in the synapse (97). In the closed conformation of Syt1
C2A-C2B, ~40% of the solvent-exposed surface area of the "...WHTL..." motif is
occluded by the C2A-C2B interface. Most of this surface area is lost by burying Trp-405
within the interface. If this sequence does mediate the docking of synaptotagmin to the
plasma membrane through a specific receptor, then it would be unable to do so while in
this closed conformation.
Implications for Other Synaptotagmin Orthologues
The human genome possesses genes for at least 17 distinct synaptotagmin
proteins (Figure 3.4). The excessive redundancy of the Syt orthologues is either due to
the critical nature of Syt in regulated exocytosis (98) or as part of an intricate addressing
29
system that allows for the temporal and spatial fine-tuning of fusion between specific
vesicles and specific membrane targets in various cell types (7). The residues that we
observe at the interface between C2A and C2B are present in many of the other isoforms,
but not all. Therefore, it is conceivable that this C2A-C2B domain orientation is unique to
Syt1. The position of C2A relative to C2B may vary by homologue depending on the
particular biological requirements of the cell.
CONCLUSION
The C2A and C2B domains of Syt1 are related by a ~108
rotation about the
linker axis. This is in contrast to the more "open" conformation of Mg2+-bound Syt3
C2A-C2B where there are no interdomain interactions between domains and the
phospholipid-binding loops directly face one another (Figure 3.1B) (51). NMR analysis
of Syt1 C2A-C2B concluded that, like Syt3, the domains of Syt1 can be separate and
noninteracting (52). However, we clearly observe extensive interactions between
domains. To rationalize these seemingly disparate results, we hypothesize that Syt1
possesses an open conformation and a closed conformation that are critical to its function.
The closed form may be required for regulating the activity of Syt1 prior to fulfilling its
role in exocytosis, while the open form may be required for effector interaction during
the final stages of exocytosis.
We propose that, while still in this closed conformation, C2B·Ca2+ initially
localizes to the membrane surface and may begin to associate with phospholipid
membranes and other factors required for exocytosis (99). As the Ca2+/phospholipidbinding loops of C2B are already in a conformation amenable to phospholipid
interaction, it is reasonable to assume that C2B will behave as a phospholipid-binding
module to initially localize the Ca2+ sensor to the target membrane. This may explain
why C2B appears more essential to the Syt1 protein, as it could be the first domain
capable of responding to cytosolic Ca2+ gradients (100). At some point, the C2A domain
30
must detach from C2B under the influence of a trigger. We predict that the trigger will be
a threshold Ca2+ concentration, but phospholipid interaction, effector protein binding, or
some combination of the three may initiate the transformation of the Syt1 molecule to an
open conformation. Next, the divalent cation-binding loops 1 and 3 of C2A will attain
their final conformation as calcium ions occupy binding sites (99). The final phase of
synaptotagmin action is presently unclear, as it may involve a direct mechanism of
phospholipid demixing that leads to fusion, or the protein will set the stage for another
component to facilitate the final step. This model of Syt1 C2AB also suggests a
mechanism whereby the two C2 domains of synaptotagmin may act on the target
membrane in series rather than in parallel, thereby providing a true structural trigger.
Consequently, C2A may only attain its final Ca2+-occupied, membrane-binding
competent structure during the final few microseconds prior to exocytosis. Defining the
structural basis for the interactions of synaptotagmin 1, as well as other isoforms, with
SNARE components and phospholipid bilayers will be the focus of future studies.
Figure 3.2: (A) Superposition of the isolated Syt1 C2A domain (PDB code 1RSY,
yellow), the Ca2+-saturated Syt1 C2A domain (PDB code 1BYN, cyan), and C2A (in
this crystal structure, green). The C2B domain is shown in gray. The Ca2+-binding
31
residues for the C2A domain are shown as sticks. The more distal position of Asp-172 in
the C2AB structure is highlighted. The N- and C-termini of the synaptotagmin C2A-C2B
protein are labeled as N and C. (B) Rmsd plot over all CR atoms between each of the
C2A domains in the C2A-C2B crystal structure and the isolated C2A domain (1RSY).
The red curve is the C2A domain of molecule A vs 1RSY, and the blue curve represents
molecule B vs 1RSY. The residue ranges corresponding to loops 1 and 3 are shown by
the bars above the major peaks. NCS restraints were used throughout the refinement with
the exception of residues of loops 1 and 3 in C2A, where considerable differences in the
path of the loops were observed.
Figure 3.3: Stereoview of the C2A-C2B interdomain interactions. C2A is rendered in
green ribbons, and C2B is rendered in gray on the right. Interacting residues are labeled
according to the rat Syt1 numbering scheme. Also shown is the intradomain interaction
between Asp-232 in C2A and the backbone amide of Phe-234.
32
Figure 3.4: ClustalW alignment of the reported human synaptotagmin paralogues. The
green arrows correspond to ß-sheet structures in the C2 domains. The green tubes
represent R-helical regions in the C2B domain. The top half of the alignment shows
residues Glu-169 to Gly-242 in human Syt1 C2A; the bottom half only shows residues
Gly-385 to Lys-422 in C2B. Residues highlighted in red correspond to acidic residues in
Syt1 C2A known to coordinate Ca2+. Red residues in Syt2-17 share homologous
positions and, in most cases, contribute to Ca2+ binding in those isoforms. The residues
highlighted in the yellow box are those in ß-strand 3 implicated in the modulated loop 3
structure of Syt1. The other residues homologous to the “SDPYVKV” motif within the
yellow box may play similar roles in other paralogues. Residues in blue were observed
forming salt bridges between C2A and C2B in Syt1. Residues boxed in red comprise the
conserved “WHxL” motif.
33
Figure 3.5: The rotameric position of His-237 depends on whether C2A is coupled to
C2B. (A) The His-237 residue can pair with Thr-406 in C2B. C2B is rendered as gray
helices and ribbons. (B) Hydrogen-bonding partner of His-237 in the isolated C2A
domain of rat synaptotagmin 1 (1RSY).
34
CHAPTER 4: SINGLE MOLECULE ANALYSIS
C2AB
OF THE
STABILITY
OF
SYT I
(Studies in this chapter have been published in the paper: Fuson, K.L., Ma, L., Sutton,
R.B., Oberhauser A.F., The C2 Domains of Synaptotagmin 1 Have Distinct Mechanical
Properties. Biophys J. 2009 Feb;96(3):1083-90. and with permission.)
INTRODUCTION
Biological systems have evolved a relatively small set of proteins known as
SNAREs to catalyze the fusion of cargo containing phospholipid vesicles with a target
membrane. The assembly of the these proteins (synaptobrevin, syntaxin, and SNAP-25)
into a parallel four-helix bundle at the vesicle docking interface provides the free energy
required in its role as the fusion engine for exocytosis (5, 101, 102). Synaptotagmin 1
(Syt1) is a vesicle-associated protein that interacts with the SNARE complex (51, 103),
and is thought to fine-tune the probability of calcium-ion dependence of release(104).
The very nature of exocytosis, that is, the fusion of juxtaposed phospholipid
membranes, implies a great deal of mechanical force. For example, the interaction forces
between the three protein components of the SNARE complex have been measured by
single-molecule atomic force microscopy (AFM) to be in excess of 285 pN (105, 106).
Since the Syt1 protein directly interacts with the SNARE complex, it must be able to
adjust its structure to this highly mechanical, force-driven framework. Here, we used
single-molecule AFM to study the mechanical properties of the tandem C2 (C2A and
C2B) domains of human Syt1, as these domains are the Ca+2/phospholipid and SNARE
interacting portion of the Syt1 protein (Figure 4.1A).
The SNARE binding region of Syt1 localizes to the polybasic β4 strand of C2A
and the Ca+2-binding loop 1 of C2B (103), while the calcium ions localize to a cup-like
depression formed from three loops (loops 1, 2, and 3) at the apex of both C2A and C2B
(55). Only loops 1 and 3 in the C2 domain contribute the conserved acidic residues that
actually coordinate the cation.
35
Figure 4.1. Equilibrium denaturation for Syt1 C2AB and titin I27 domains. A) X-ray
structure of human Synaptotagmin 1 C2AB showing the position solvent accessible (in
yellow) and buried tryptophan residues (in orange). Left domain (green) is C2A.
Ca+2/phospholipid binding loops are labeled in C2A as Loop 1, 2, 3. The blue domain is
C2B. B) NMR structure of I27; the buried Trp is shown in orange. C) Chemical
denaturation curves for human Syt1 C2AB domains (filled squares) and titin I27 domain
(gray circles). The data were fit by a simple sigmoid. The black horizontal line
demarcates the point where 50% of the normalized fluorescent signal, [D]50%. The
estimated [D]50% are ~1.6 M for C2AB and 2.7 M for I27.
The apex of the loops 1 and 3 of the C2 domains of Syt1 possess hydrophobic
residues that insert into the phospholipid membrane(77), and may directly contribute to
36
the fusion process(107). At the core of C2 domain is a Greek-key folding motif that is
conserved among all C2 domains(23).
While the strand connectivity is markedly
different, the secondary structure and overall fold of the C2 domain is similar to other βsandwich proteins with structural and mechanical roles. These include, for example, titin
(108-110), fibronectin (111) and NCAM(112).
The primary sequences of the C2 domains of human Syt1 are 31% identical (56%
similar). The X-ray crystal structures of both the isolated C2A (23) and C2B (50)
domains superimpose with an RMSD of 2.0Å.
Despite having similar structural
characteristics, the tandem C2 domains of Syt1 differ in their biochemical and biological
functions(113). C2A and C2B domains have been shown to participate at different stages
in exocytosis(100), have different affinities for phospholipids(52), and differ in their
selectivity for highly charged inositol compounds(114).
In addition, our recent high-
resolution crystal structure of human Syt1 C2AB shows that the C2B domain of Syt1 can affect
the shape of the Ca+2 binding pocket of C2A, thus potentially modifying its Ca+2/phospholipid
binding potential(55). It is presently unclear how these disparate biological observations for
binding and function can result from the tandem domain organization of Syt1.
Since the 3D structure and the biochemical characterization of these two domains
do not provide a clear explanation for this disparity, one hypothesis to explain the
differences in biological behavior between C2A and C2B is that the biophysical
properties of each domain are fundamentally different.
Here, we compared the
mechanical properties of Syt1 C2A and C2B domains. Stretching a construct containing
a C2AB fragment resulted in two distinct unfolding force peaks. The larger force peak of
~100pN was identified as C2B. The second peak, which unfolded at ~50% lower forces
we identified as C2A (~50pN); in addition about 40% of C2A domains unfold through a
mechanical intermediate.
Hence, our data shows that C2A and C2B are have
significantly different mechanical properties.
37
This feature of the molecule may be
important for the C2 domains of Syt1 to respond asymmetrically to effectors such as
SNAREs, phospholipids and Ca+2.
EXPERIMENTAL PROCEDURES
Cloning and expression of a C2A and C2AB domain-titin I27 protein chimeras for
AFM experiments
Human synaptotagmin I C2A (residues 140 - 265) and C2AB (residues 140 - 414)
were amplified from our GST-Syt1 expression vector by PCR, using the following
primers:
5’-GCGCGCGAGAAACTGGGAAAACTTCAG-3’ (forward primer for C2A
and C2AB)
3’-GATCACTAGTACTTTGCAGGTCACGCCATTC-5’ (reverse primer for
C2A)
3’-GATCACTAGTTTACTTCTTGACGGCCAGCA-5’
(reverse
primer
for
C2AB)
The PCR products were gel purified and cloned into the pDrive direct cloning
vector (Qiagen).
Each clone was excised from pDrive with BssHII and SpeI and
subcloned into a modified version of pRSETA vector, which includes a His-tag at the Nterminus (115). Both constructs (I272-C2A-I272 and I272-C2AB-I272) were confirmed
by DNA sequencing. The expression vector was transformed into E. coli BL21(DE3)
cells (Novagen). Transformed colonies were grown overnight at 37 ºC in 5 ml LB
containing 100 µg/ml ampicillin. The overnight culture was re-inoculated into 1 L fresh
TB and the cells were grown at 37 ºC and induced with 400 μM IPTG when the OD600
reached 0.7. The cells were collected by centrifugation, quick frozen in liquid nitrogen
and stored at -80 ºC. 5 g of cells were sonicated in 50 ml lysis buffer (1X PBS pH 7.4, 1
mg/ml lysozyme, 0.1 mg/ml DNAse I), centrifuged at 15,000 rpm for 30 min at 4 ºC.
38
The proteins were purified by Ni+2 affinity chromatography, eluted with elution buffer
(1X PBS, 250 mM imidazole, pH 7.4), concentrated to 6 mg/ml and stored at 4 oC.
Cloning and expression of C2AB and I27 for chemical denaturation experiments
Human synaptotagmin I C2AB (residues 140 to 418) was amplified from a human
hippocampus QUICK-Clone cDNA library (Clontech) by PCR using the following
primers:
5’GGATCCGAGAAACTGGGAAAACTTCAGTATTCACTGGATTATG 3’
3’ TCACTATTACTTCTTGACGGCCAGCATGGC5’
The PCR reactions were gel purified and cloned into the pCR2.1 TA-cloning
vector (Invitrogen). The gene was excised from pCR2.1 with BamHI and EcoRI and
subcloned into pGEX-4T (GE Healthcare Life Sciences). The recombinant vector was
transformed into E. coli Rosetta cells (Novagen). Heterologous gene expression was
induced by adding 400 μM IPTG to a culture in 1L Terrific Broth for 4 hours at 37 °C.
C2AB was initially purified using GST affinity resin and cation exchange
chromatography. The GST tag was removed using human α-thrombin (50 units per ml
total protein) and final purification was done using a Superdex 75 gel filtration column.
The purified C2AB was concentrated to 24 mg/ml, divided into aliquots and quick frozen
in a liquid nitrogen bath. Samples were stored at -80 ºC. The titin I27 was cloned and
expressed as previously described (116). The purity of the proteins was confirmed by
SDS-PAGE.
SINGLE-MOLECULE ATOMIC FORCE MICROSCOPY
The mechanical properties of single proteins were studied using a home-built
single molecule AFM as previously described (111, 116-119). The spring constant of
39
each individual cantilever (MLCT-AUHW: silicon nitride gold-coated cantilevers; Veeco
Metrology Group, Santa Barbara, CA) was calculated using the equipartition theorem
(120). The cantilever spring constant varied between 10-50 pN/nm and rms force noise
(1-kHz bandwidth) was ~15 pN. Unless noted, the pulling speed of the different force–
extension curves was in the range of 0.4–0.6 nm/ms.
EQUILIBRIUM DENATURATION OF I27 AND C2AB DOMAINS
The experiments were carried out at 28°C in PBS buffer. Protein concentration
was 1–2 M. The stability of I27 and C2AB domains was determined by using
equilibrium guanidinium chloride (GdmCl) denaturation. The C2AB protein has an
emission maximum ~ 345 nm (data not shown), suggesting that the tryptophan residues
are relatively exposed to the solvent.
Unfolding was monitored by change in
fluorescence at 345 nm for C2AB and at 320nm for I27 (excitation 290 nm) using an LS50B spectrofluorimeter.
SINGLE PROTEIN MECHANICS
In a typical experiment, a small aliquot of the purified proteins (~1-50 µl, 10-100
µg/ml) was allowed to adsorb to a clean glass coverslip (for ~10 min) and then rinsed
with PBS pH 7.4. Proteins were picked up randomly by adsorption to the cantilever tip,
which was pressed down onto the sample for 1-2 seconds at forces of several nanonewtons and then stretched for several hundred nm.
STEERED MOLECULAR DYNAMICS (SMD) SIMULATIONS
We simulated the unfolding of C2AB using steered molecular dynamics (SMD)
techniques as implemented in NAMD (121, 122). Coulombic forces were restricted
using the switching function from 10 Å to a cutoff at 12 Å. The CHARM22 force field
was employed throughout. C2AB (pdbcode:2R83) was solvated in water sphere with a
40
boundary of 15 Å. The system was charge neutralized by adding Na+ and Cl- ions. The
total ionic strength of the system corresponded to a final concentration of 0.1 M. This
simulation contains a total of 13762 atoms. The system was then minimized with 1000
steps of conjugate gradient minimization from an initial temperature of 310K. This step
was followed by a 400ps MD simulation to equilibrate the entire system (protein, water,
and ions). For the SMD portion of the simulation, a spring constant (κ) of 10 kBT Å-2 was
used. Simulated force was applied by fixing one termini of the protein and moving the
SMD atom with constant velocity along a predetermined vector. The trajectories were
recorded every 2 fs and analyzed with VMD. The C2AB fragment of Syt1 was stretched
at a constant velocity of 0.001 Å ps-1 and was followed for ~260 Å. We ran three
simulations of the extension of C2AB in both pulling directions (N->C and C->N) with
similar results.
41
Results
C2AB HAS A LOWER THERMODYNAMIC STABILITY THAN TITIN I27
As a first step to analyze the stability of the C2AB domains, we used chemical
denaturation with guanidinium chloride (GdmCl) and steady-state fluorescence
techniques to determine the thermodynamic stability of the domains. As a reference, we
used the titin I27 domain, which has been extensively studied using both chemical and
mechanical denaturation techniques (116, 123-125). Further, both the I27 domain and
the C2 domain are similarly sized β-sheet domains that are constructed around a central
Greek key motif (23, 126). In the crystal structure of C2AB (Figure 4.1A), two of its
three tryptophan residues are exposed to the solvent (in yellow), so the main contribution
to the fluorescence intensity and emission arises from the most buried tryptophan in C2B
(in orange). The I27 domain of titin has a single, buried Trp residue (Figure 4.1B). As
shown by the denaturation curve in Figure 4.1C, the C2AB protein readily denatures
when exposed to GdmCl (black squares).
The fluorescent signal rapidly changes
between ~1 and 2 M GdmCl, with a [D]50% ~1.6 M. In contrast, the denaturation curve
for I27 shows [D]50% ~2.7 M GdmCl (Figure 4.1C; (127)). The simplest explanation for
this ~2-fold difference in [D]50% is that the C2 domains are thermodynamically less stable
than titin I27 domain.
MECHANICAL STABILITY OF DOMAIN C2AB
To measure the single-molecule mechanical properties of C2AB domains of Syt1,
we constructed a protein chimera consisting of a C2AB module flanked on the N- and Ctermini by two I27 domains (I27)2-C2AB-(I27)2. Figure 4.2A shows that stretching of
the (I27)2-C2AB-(I27)2 polyprotein results in a force-extension curve with a
characteristic sawtooth pattern with several force peaks.
42
Figure 4.2. Mechanical properties of C2AB. A) Typical force-extension curve
obtained after stretching a I272-C2AB-I272 protein. The I27 domains unfold at forces of
~190pN and give an increase in contour length, ΔLc, of ~28nm. These were identified by
measuring the spacing between force peaks using the worm-like chain (WLC) equation
(thin black lines); in this recording we see four I27 domains unfolded and the first force
peaks must correspond to the unfolding of C2 domains. B) For C2 domains we measured
a ΔLc of ~43 + 8nm (n=98). C) An unfolding force histogram shows that C2 domains
have unfold at forces of 62 + 30pN (n=279; shown in red) a value that is 3-fold lower
than that for I27 (188 + 29pN, n=264; shown in black). D) A plot of unfolding force vs.
the pulling speed shows that the 3-fold difference between C2 and I27 domains is
maintained over a wide range of pulling speeds.
We found that most recordings showed two levels of unfolding forces (Figure
4.2A); low force peaks (~10 - 150pN) and high force peaks (150 - 250pN). To establish
a molecular fingerprint for each domain in the protein chimera, we analyzed the spacing
between peaks in the sawtooth patterns. We used the worm-like chain (WLC) model for
polymer elasticity, which predicts the entropic restoring force (F) generated upon the
43
extension (x) of a polymer (128, 129). The thin red and black lines in Fig. 2A correspond
to fits of the WLC equation to the curve that precedes each force peak. The I27 domains
are known to unfold at forces of ~200pN and give an increase in contour length, Lc, of
~28nm upon unfolding (108, 116). Hence, in this recording the last four force peaks
must correspond to I27 and the first two force peaks to C2 domains. As shown in Figure
2B, we observed a wide range of Lc values (from ~25-60nm) for C2 domains, with a
mean Lc of 43 ± 8nm (n=98).
The data shown in Figure 2C corresponds to the
combined unfolding forces for C2A and C2B domains. The C2A domain has 124 amino
acids, while the C2B domain has 150 amino acids. This yields a predicted value of the
fully extended polypeptide chains of ~40nm and ~50nm, respectively. So, the measured
Lc is close to the expected combined average value, ~45nm.
An unfolding force
histogram (Figure 2C) revealed that C2 domains unfold at forces of 62 ± 30pN (n=279;
shown in red) a value that is 3-fold lower than that for I27 (188 ± 29pN, n=264; shown in
black). Further, a speed-dependence plot comparing I27 and C2 domains confirms that
this 3-fold lower unfolding force is maintained over a wide range of pulling speeds
(Figure 4.2D).
These data demonstrate that C2 domains have a lower mechanical
stability than titin I27 domains.
MECHANICAL STABILITY OF C2A DOMAINS
While we can confidently discriminate between C2 domains and I27 with this
data, the identity of the individual C2 domains in these recordings cannot be established
at this point. To unambiguously identify the force peaks from each C2 domain, we
constructed a polyprotein chimera that contains one repeat of C2A and two flanking titin
I27 domains.
44
Figure 4.3. Mechanical properties of C2A. A) Two examples of force-extension
curves obtained after stretching I272-C2A-I272 proteins. The thin lines correspond to fits
to the WLC equation. In the example shown on the right, the C2A force peak precedes a
low-force peak. B) Histogram of increases in contour lengths measured for the C2A
domain. Most domains unfold in an all-or-none fashion were the Lc is ~40nm (39.6 +
4.5nm, n=51). However, we also observed about 38% (31 out of 82 recordings) of
domains unfolded through an intermediate which contributes to an increase in contour
length of ~7nm (7.4 + 3.5nm, n=31; blue bars). C) An unfolding force histogram shows
that C2A domains have unfolding forces of 51 + 14pN (n=78).
Figure 4.3A shows two examples of force-extension curves obtained after
stretching I272-C2A-I272 proteins. We found that most C2A domains unfolded in a twostate manner with a Lc of ~40nm (39.6 ± 4.5nm, n=51). This corresponds well to the
predicted Lc (124aa x 0.35nm/aa = 43nm) and those measured for C2A polyproteins
(130). The recording shown on the left panel is a representative example. In these
recordings the C2A unfolded at forces of ~50pN (51 ± 14pN; n=78; Figure 4.3C).
45
However, we also observed that about 40% of C2A domains unfolded in more complex
pattern, as shown in Figure 4.3A, right panel. In these cases, we find that the domain
unfolds in two steps; the first elongates the protein by ~7nm (7.4 + 3.5nm, n=31; blue
bars) and the second by ~36nm. We interpret this pattern as the C2A domains unfolding
through a mechanical unfolding intermediate: native-to-intermediate state (N-> I)
followed by an intermediate-to-unfolded state (I-> U). The origin of this unfolding
intermediate remains unclear, but it may correspond to the first two beta strands of C2A
unfolding prior to the core (see below).
SPEED DEPENDENCE OF C2A VS. C2B
Now that we have established the mechanical fingerprint for C2A, we can reanalyze the C2AB data shown in Figure 4.2. The C2A domain unfolds at forces of
~50pN and sometimes through a mechanical unfolding intermediate.
This is the
mechanical fingerprint of the C2A domain. For example, stretching the (I27)2-C2AB(I27)2 protein results in a sawtooth pattern with 6 force peaks (Figures 4.2A and 4.4A).
The first force peak we identify as C2A, the second as C2B and the last four as I27
domains. Using this mechanical ‘fingerprinting’ approach, we find that the C2B unfolds
at higher forces than C2A. Figure 4.4B shows a histogram for unfolding forces for the
peaks identified as C2A (grey bars) and C2B (red bars); the average unfolding forces are
~50pN (49+18pN, n=91) and ~100pN (106+23pN, n=110), respectively.
We now can study the mechanical properties of each domain separately under
different conditions. One important variable is how the unfolding forces vary with the
pulling speed. Figure 4A shows an experiment done at 5nm/ms which 10x faster than the
typical pulling speed. The sawtooth pattern is similar to that obtained at 0.5 nm/ms
except that the unfolding forces are about ~1.5x higher. A plot of the pulling speed
dependence (Figure 4.4C) shows that a 100-fold increase in pulling speed increases the
46
unfolding forces by 60pN for C2A and 96pN for C2B. This result further demonstrates
that Syt1 C2A and C2B domains have distinct mechanical properties.
Figure 4.4. Mechanical properties of C2A vs. C2B domains. A) Force-extension curve
obtained after stretching a I272-C2AB-I272 protein at 5nm/ms. We identify the first peak
as the mechanical unfolding of C2A and the second as the unfolding of C2B domain. B)
Unfolding force histogram for C2A (in black) and C2B domains (in red). The average
unfolding forces are ~50pN (49+18pN, n=91) and ~100pN (106+23pN, n=110),
respectively. C) Plot of unfolding forces vs. pulling speed for C2A (black squares), C2B
(red triangles) and I27 (open circles). A 100-fold increase in pulling speed increases the
unfolding forces by 60pN for C2A, 96pN for C2B and 93pN for I27.
47
MOLECULAR BASIS FOR DIFFERENCES BETWEEN C2A AND C2B
To formulate a domain-level explanation for the differences between C2A and
C2B domains, we simulated the extension of C2AB using SMD (steered molecular
dynamics) (122, 131). For this, we fixed the Cα position of residue 140 in C2A and
extended the chain by applying an external pulling force on residue 414 in C2B. These
forces were applied by harmonically restraining the C-terminal Cα atom of C2B and
moving that point at a constant velocity along a defined vector. In Figure 5, we pulled
the human Syt1 C2AB structure at a constant velocity of 0.001 Å ps-1 for 200 Å. The
initial events noted in this simulation are β1 and β2 decoupling from the one side of the
β-sheet in C2A. As β2 is linked to Loop 1, the shape of the Ca+2 binding pocket was
severely distorted. Loop 1 is one of the three Ca+2/phospholipid binding loops in Syt, as
it has two of the five Ca+2 binding residues of C2A (Asp 172 and Asp 178) and one of the
hydrophobic residues known to interact with phospholipids (Met 174). The C2B domain
was not distorted to the same degree over the same time period. We obtained the similar
results regardless of pulling direction and speed. In this series of simulations we find
that, C2A denatures first, followed by C2B, which is consistent with our forcespectroscopy results.
48
Figure 4.5. Steered molecular dynamic simulation (SMD) of C2A and C2B domains.
Two snapshots of the extension C2AB (residues 140-414) pulled at a constant velocity of
0.001Å ps-1 over 20nm. The fixed atom was in residue 414 in C2B. The top structure
represents the initial conformation of C2AB and the bottom structure represents the
conformation after 200 Å in the simulation. The red sphere corresponds to the fixed atom
and the green sphere and arrow corresponds to the moving atom. The Ca+2 binding
residues of C2A are highlighted as sticks. Figure was rendered using VMD and Tachyon.
49
DISCUSSION
The C2A and C2B domains of Syt1 are similar with respect to their overall
sequence similarity and their 3D structures. In addition, they bind Ca+2 and phospholipid
with similar affinities, yet they play different roles in exocytosis (113). This observation
implies that there are marked differences between the two domains that are crucial to
function, yet are not obvious with respect to their primary or tertiary structure. To probe
these differences, we analyzed the C2 domains of Syt1 using single molecule force
microscopy. Our analysis shows that 1) the C2 domains are thermodynamically less
stable compared to other β-sheet domains, and 2) the C2A and C2B domains have
different mechanical stabilities. The C2B domain is relatively strong and unfolds at
~100pN in an all-or-none manner. The C2A domain, on the other hand, is significantly
weaker and tends to unfolds in two steps through a mechanical unfolding intermediate.
THE STABILITY OF HUMAN SYT1 C2 DOMAINS
To examine the stability of the C2AB portion of Syt1 relative to control domains,
we analyzed the C2AB domains of Syt1 and the I27 domain of titin using steady-state
chemical denaturation. Our analysis of the C2AB domains of Syt1 shows that the C2
domains of Syt1 are thermodynamically less stable than the I27 domain of titin. Since
the C2A domain lacks an appropriate environmentally sensitive fluorophore, we relied on
the partially buried Trp in C2B. This experiment showed that the C2B domain, as a part
of C2AB, is significantly less stable than titin I27 domains. This is an interesting result
as the I27 domain and both C2 domains are of similar size and overall fold.
Our force spectroscopy results show that C2 domains are also less stable than I27
titin domains. The relatively low resistance to mechanical forces of C2 domains lies in
their topologies. The C2 domain topology lacks the force-bearing terminal β-strand
architecture of the I27 domain (131). The hydrogen bonds holding C2 domains together
50
are parallel to the axis of extension and thus, are in a “zipper-like” configuration (131).
This implies that the bonds must break sequentially causing the strands to separate at
lower force.
C2B HAS A HIGHER MECHANICAL STABILITY THAN C2A
Since the two C2 domains of Syt1 possess similar topology, one would expect
both to possess similar mechanical properties; however, they do not. We found that C2A
unfolds at ~50pN whereas C2B at ~100pN.
One possible explanation for these
differences is the presence of the H-A helix within the C2B fold (Figure 4.5). Based on
the SMD simulations, we find that the H-A helix stabilizes β8 by contributing additional
H-bonds and hydrophobic interactions with core packing residues. β8 makes backbone
H-bonds with β1 to form one edge of one β-sheet of the C2B domain, so stabilizing this
interaction would add strength to the entire domain.
Sixteen different isoforms of
synaptotagmin have been identified within the human genome and all have residues that
correspond to the helix-A (H-A) in their C2B domains (63). So, it is likely that the
difference in mechanical responsiveness between C2A and C2B domains is a common
property of all synaptotagmin isoforms.
MECHANICAL UNFOLDING INTERMEDIATE IN C2A
We observed that ~40% of C2A domains unfolded in more complex pattern, as
shown in Figure 3A, right panel. In these cases, we find that the domain unfolds in two
steps, the first elongates the protein by ~7nm (7.4 + 3.5nm, n=31; blue bars) and the
second by ~36nm. We interpret this pattern as the C2A domains unfolding through a
mechanical unfolding intermediate: native-to-intermediate state (N-> I) followed by an
intermediate-to-unfolded state (I-> U).
Mechanical unfolding intermediates have been observed in other -sandwich
folds such as FNIII (132) and filamin (133). By analyzing both the wild-type and mutant
forms of
10
FNIII, it was found that the unfolding intermediate observed in
51
10
FNIII was
due to the A and G β-strands detaching from the domain followed by the unfolding of the
remainder of the protein. The authors concluded that the intermediate in FNIII could
protect the domain from complete unfolding in response to an applied force (132). In
contrast, our data suggests that the unfolding intermediate is not very stable, since the
initial peak has a low force value (<30pN). C2A may readily respond to stretching forces
by partially unfolding the first ~20aa, which may correspond to 1 and 2 in Figure 5.
Interestingly, since 2 is linked to the Ca+2-binding loop 1, this unfolding event might be
important to modulate Ca+2/phospholipid binding in C2A, but not C2B.
POTENTIAL
EFFECTS OF MECHANICAL FORCES ON
SYT1 C2
DOMAIN
CA+2
BINDING
LOOPS
How can the different stabilities between C2A and C2B fit into the exocytotic
pathway? While synaptotagmin has been identified as a Ca+2 trigger for exocytosis (20,
134, 135), there is also evidence that it participates in a more direct way by mediating the
final step in fusion pore formation (42, 99, 136). A force applied to C2A could easily
perturb the structure of loop 1 in C2A after it binds to the target membrane, thus
delocalizing any calcium ions within the Ca+2 binding pocket.
simulations of
10
For example, SMD
FNIII suggested that a force applied to one terminus of the domain
results in a deformation of the integrin binding loop. They speculated that a shortening of
this loop could potentially reduce the accessibility to its binding partners, thereby
modulating its affinity (137). In the case of Syt1, the divalent cation-binding pocket of
C2A provides the negatively charged quenching field to cancel the charge from the Ca+2.
This quenching field would be dependent on the tertiary structure of the domain (90).
Modification of this quenching field would spontaneously cause a buildup of a strong
localized electric field within the membrane at the site of exocytosis (138, 139). Based
on our SMD results shown in Figure 4.5, we hypothesize that a relatively small stretching
force may be sufficient to deform the Ca+2-binding loop and SNARE bindings regions of
52
C2A. These built-in mechanical sensitive switches may be important in modulating the
affinity for Syt1 binding partners.
53
Chapter 5: Structural Effects of Mutations on Syt I
C2A
INTRODUCTION
In 1967 Katz and Miledi (140) demonstrated that Ca2+ influx into the presynaptic
terminus triggered neurotransmitter release; however, the molecular mechanisms
underlying the event remained elusive. Synaptotagmin I is now widely accepted as the
Ca+2 sensor in fast synchronous neurotransmitter release.
The synaptotagmin family is made up of 17 proteins that mediate docking,
recycling and fusion of vesicles with target membranes throughout the human body (12).
The expression of most synaptotagmins is highest in neuronal tissues, although some
synaptotagmins show a widespread tissue distribution (13, 14).
Synaptotagmin I is the most abundant Ca2+ binding protein present on synaptic
vesicles accounting for 7% of total vesicle protein (18). Synaptotagmin I contains two
C2 domains that bind Ca+2, interact with phospholipid and interact with the SNARE
complex, which is an important component of the neurotransmitter release machinery
(18, 20). Previous studies have reported on synaptotagmin’s ability to oligomerize with
itself in a Ca+2 dependent fashion (88, 93, 141-143) and concluded that oligomerization is
required for synaptotagmin function. However, to date, the relevance of either homooligomerization (144, 145) or hetero-oligomerization (93, Osborne, 1999 #625, Fukuda,
2000 #500, Fukuda, 2001 #446, 146) to synaptotagmin function has not been fully
realized.
Three synaptotagmin mutants created in Drosophila (AD1, AD3, and AD4) have
been useful in understanding synaptotagmin function and have been characterized at the
genetic and biochemical level. Each has defects in specific molecular interactions (88,
97, 147). Syt (AD4) is a null allele caused by an early stop codon that deletes the
transmembrane and a large fraction of the first cytoplasmic domains of the protein. As
54
only a few residues of the extracellular domain remain intact, this mutation disrupts all
synaptotagmin interactions (147). Syt (AD1) inserts a stop codon that deletes the C2B
domain only while leaving the C2A domain intact. The result of this mutation is a
decrease in the levels of wild-type synaptotagmin at synapses and a reduction in Ca2+dependent binding of synaptotagmin to SNAREs, as well as a decrease in selfoligomerization (88, 145). AD1 mutants have severe defects in synaptic transmission
that cannot be rescued by higher levels of extracellular Ca2+ (88, 148).
The syt (AD3) is the best studied allele of the three mutants and can be partially
rescued with increased internal Ca+2 concentrations (149).
(149) studied both the AD1 and AD3 mutations. The AD3 mutation
encodes a Y364N change in the C2B domains of Drosophilia Syt1. This mutation does
not abolish SNARE or phospholipid binding, but instead disrupts Ca2+-dependent
neurotransmitter release. This tyrosine residue is highly conserved among other Syt
isoforms, and between the C2A and C2B domains. The sequences surrounding the
tyrosine (SDPYVK) are also highly conserved throughout the family, as well as between
the tandem C2 domains (13, 97). Deletion of the PYVK sequence in the C2A domain of
Syt II (a very similar protein to Syt1) results in the abolishment of Ca2+-dependent
phospholipid binding. (150). The PYVK sequence of the C2B domain is expected to be
similarly involved in Ca2+-dependent phospholipids binding.
In (97), synaptotagmin AD3 mutants were tested for their ability to self
oligomerize in a Ca+2 dependent manner.
Fukuda generated AD3 mutants of
synaptotagmin I and II C2AB. They found that the mutation results in a loss of ability to
oligomerize in a Ca+2 dependent manner. Their conclusion is that lack of oligomerization
is the reason for loss of Ca+2 dependent neurotransmitter release. (151) further tested the
oligomerization of synaptotagmin using a knock-in approach.
They infected
synaptotagmin deficient hippocampal neurons with virus engineered to express the SytI
mutants.
They used three mutants that have previously been shown to inhibit
55
oligomerization at different levels. Y311N, K326/327A and K327A. Y311N, which
corresponds to the AD3 mutation, has been shown to significantly decrease
oligomerization (88, 145).
polylysine region of C2B.
The K326A and K327A mutations are located on the
The K326A mutation has been shown to reduce
oligomerization by 50% (93, 152) and the double mutant (K326/327A) shows no
detectable oligomerization (93, 144, 152). Their hypothesis was that if oligomerization is
required for neurotransmitter release, there should be a correlation between the amount of
oligomerization and neurotransmitter release.
Since the K326A mutant inhibits
oligomerization by 50% and the double mutant completely inhibits it, this should also
hold true for transmitter release. They found that oligomerization levels do not correlate
with release rates. All three mutants rescued synaptotagmin deficient neurons to similar
levels even the double mutation which has shown no propensity to oligomerize. They
also tested the ability of each mutant to bind Ca+2. All three mutants showed decreased
ability to bind Ca+2 with the AD3 and double mutation being most pronounced. They
show a roughly two fold reduction in Ca+2 affinity. They conclude that a decrease in Ca+2
binding and not the inhibition of oligomerization is responsible for the defect in the
mutants ability to release neurotransmitter.
(153) tested synaptotagmins ability to bind SNARE, specifically SNAP 25. The
used sequence alignment, mutagenesis, computerized docking and binding assays
(immunoprecipitation) to create a structurally based model for syt / SNARE binding.
They focused on evolutionary conserved residues in SNAP 25 and syt I, They concluded
that syt I C2B binds SNAP 25 of the SNARE complex. Mutation of several basic
residues prevented syt binding to SNARE and disrupted Ca+2 triggered neurotransmitter
release. The mutations had no effect on the stability of the SNARE complex or it’s
ability to form. They also tested the AD3 mutation in C2B. They found that the single
point mutation was sufficient for disrupt interaction between SNAP 25 and Syt. They
also made mutations Y311F and Y311N. The Y311F mutation had little effect on
56
SNARE binding of Syt, but the Y311N mutation strongly decreased the ability of syt to
bind SNARE. They conclude that the mutation causes a conformational change in C2B
preventing SNARE association and disrupting neurotransmitter release.
We recently published the X-ray crystal structure of synaptotagmin I C2AB in the
absence of Ca+2 (55). In the structure we see, for the first time, an interaction between
C2A and C2B (Fig. 5.1). Interestingly, one interaction occurs between Tyrosine 180 in
C2A and Threonine 307 in C2B. The Y180 residue in C2A is the comparable residue in
the AD3 mutation of C2B (Y311). The interaction of Tyr180 and Thr307 causes a
pronounced collapse of loop 3 in C2A. The physiology consequences of this collapse has
not yet been investigated; however, in silico prediction of the available Ca+2 binding sites
in C2A and C2B yield interesting results. Using a computational method based on graph
theory and geometry (154), we were able to predict the potential Ca+2 binding sites in
both the isolated C2A domain and the isolated C2B domain. The isolated C2A domain
(1rsy or 1byn) was predicted to bind 3 Ca+2; and both were coincident with the 3 known
Ca+2 binding sites of C2A. The C2B domain was predicted to bind 2 calcium ions; and
both predicted ions were coincident with the known C2B Ca+2 loci. However, when we
used the same criteria on the C2A domain of Syt 1 C2A, within the context of C2AB, no
ions were predicted for C2A, and 2 Ca+2 were predicted for C2B. It appears that the
collapse of loop 3 of C2A domain, as a consequence of its interactions with C2B,
eliminated the calcium ion binding sites by changing the geometry of the oxygen atoms
needed for efficient ion coordination.
Therefore, there is a difference in the shape of Loop 3 in the presence of Ca+2 and
in the absence of Ca+2. Since Loop 3 is instrumental in binding Ca+2 and interacting with
phospholipids (76), it is possible that this loop3 is part of a regulatory mechanism to
“switch off” Ca+2 binding in one domain, but not the other. To examine this hypothesis
closer, we can examine the status of these residues within the known structures of C2A.
In the isolated C2A X-ray structure (23), we find that tyrosine 180 is hydrogen bonded to
57
histidine 237 (figure 12), which makes up part of loop 3. This hydrogen bond could
stabilize the structure of loop 3 by providing the steric bulk of a large Tyr residue.
Therefore, this H-bonding could be thought of as a switch to position loop 3, and its
associated acidic residues, in a favorable conformation to bind Ca+2 and interact with
phospholipids. To test the hypothesis, we mutated the tyrosine 180 to a phenylalanine
(Y180F) to remove the hydrogen bond potential, but to maintain the steric qualities of the
original Tyr residue. We hypothesize that the interaction between Y180 and H237 is
essential for synaptotagmin function and explains why a loss of function is seen in the
AD3 mutation.
Two other mutations in C2A have also been extensively examined in the
literature: D232N and D238N (Figure 5.16 and 5.18). Both are thought to be simple
defects in Ca+2 binding; however, the phenotypes of these residues are more complicated.
Asp 232 and Asp 238 contribute to the primary Ca+2 binding site of the C2A domain of
Syt1. Mice with knock-in mutations of aspartate-to-asparagine at two different locations
in the Ca2+-binding site of Syt1 (D232N or D238N) were analyzed.
Although the
mutations are close in proximity to each other and effect residues responsible for Ca2+
coordination, they show different phenotypes.
The D232N mutation considerably
increases Ca2+-dependent SNARE complex binding, but leaves phospholipid binding
unaffected. The D238N mutation does not appreciably affect SNARE complex binding,
and causes a decrease in phospholipid binding.
In contrast, the D232N mutation
increased Ca2+-triggered release, whereas the D238N mutation decreased release. (98).
An explanation for the observed phenotypes of these two mutants has thus far eluted
researchers. We examined both these mutants using X-ray crystallography and can now
relate the phenotypes to changes in the structure of Loop 3, as we have measured in the
Y180F mutation.
58
EXPERIMENTAL PROCEDURES
Cloning and Mutagenesis
Human Syt1 C2A was obtained via PCR from a previous construct of
synaptotagmin I C2AB (80) that was obtained from a human hippocampal Quick-Clone
cDNA library (Clontech). A 125-nucleotide fragment comprising Syt1 C2A residues
141-266 was directly cloned into PCR2.1 (Invitrogen), excised using Nde1 and Xho1, and
subsequently subcloned into p202. It was cloned into pGEX KG (155) using Nco1 and
Xho1. DNA sequence analysis confirmed insertion in the correct reading frame with
respect to the GST fusion partner. The PCR primers used for cloning are:
p202
5’-CATATGGAGAAACTGGGAAAACTTCAG-3’ (forward primer)
3’-GACCTGCAAAGTGCTTAATAGTGACTCGAG-5’ (reverse primer)
pGEX-KG
5’-CCATGGGAGAAACTGGGAAAACTTCAG-3’ (forward primer)
3’-GACCTGCAAAGTGCTTAATAGTGACTCGAG-5’ (reverse primer)
Mutants were created using the QuickChange Site-Directed Mutagenesis kit
(Stratagene) protocol.
Primers were generated using the online mutagenesis primer
design program, PrimerX (http://www.bioinformatics.org/primerx/). The mutagenesis
primers are as follows:
Y180N
5’-GGGGGCACATCTGATCCTAATGTGAAAGTGTTTCTGCTAC-3’
5’-GTAGCAGAAACACTTTCACATTAGGATCAGATGTGCCCCC-3’
Y180F
5’-GGGCACATCTGATCCTTTTGTGAAAGTGTTTCTGC-3’
5’-GCAGAAACACTTTCACAAAAGGATCAGATGTGCCC-3’
59
Protein expression and purification
The cytosolic fragment of human synaptotagmin 1, containing only the C2A
domain, was cloned into the pGEX-KG bacterial expression plasmid and transformed
into chemically competent Escherichia coli BL21-DE3 cells. A 20 ml culture was grown
overnight in the presence of 100 mg/ml ampicillin in Terrific Broth (TB). Four liters of
TB was inoculated using 5 ml per liter of the overnight culture with 100 mg/ml
ampicillin. The culture was grown at 37°C until an O.D. 600 of 1.0 was reached. Upon
reaching the O.D., the temperature was reduced to 32°C. Protein expression was induced
at an O.D. 600 of 1.0 with the addition of 400 µM IPTG. This culture was grown for 4 hr
at 32°C while shaking in a 2 liter Fernbach Flask. After 4 hr, cells were collected by
centrifugation at 5000 rpm in a Sorvall GSA centrifugation rotor for 30 min. Typically, 5
g (wet weight) of bacteria could be collected per liter of culture. The cells were quick
frozen in liquid nitrogen and stored at -80°C until ready for further processing. The cells
were thawed on ice and resuspended in 10 ml of lysis buffer (50 mM Tris [pH 7.7], 200
mM NaCI, 5% glycerol, 1 mM DTT) per 5g cells. PMSF was added to a concentration of
0.5 mM and cells were immediately lysed using a Microfluidics M-110EH-30
Microfluidizer® Processor. After lysis, 5 mg Dnase I along with 1 mM MgCl2 and 1 mM
MnCl2 were added and the mixture was incubated on ice for 30 min. The total cell lysate
was centrifuged in a SS-34 rotor at 4°C for 30 min at 10,000 rpm. The supernatant was
then filtered through a 0.45 µm syringe filter. The filtered supernatant fraction was
applied to a 10 ml GST-Sepharose column that was prepared according to the instructions
provided by GE Healthcare. The GST column was washed extensively with PBS until
the O.D. 280 of the flow through buffer was 0.02 or lower. The fusion protein was eluted
with 15 mM reduced glutathione in 50 mM Tris (pH 8.0). All chromatography to this
point was performed at room temperature. Approximately 75 mg of fusion protein per
liter would elute from the glutathione column. The protein was extensively dialyzed
against thrombin cleavage buffer (50 mM Tris [pH 8.0], 200 mM NaCl, 2.5 mM CaCl2)
60
at 4°C. The fusion protein was cleaved with thrombin for 1 hr at 25°C. Approximately
300 µg of thrombin was used to cleave 75 mg of fusion protein. The cleaved proteins
were concentrated using Amicon Ultra-15 Centrifugal Filter Unit with Ultracel-10
membrane. The Syt I C2A domain was separated from the GST and the remaining
uncleaved fusion protein with a mono S HR 5/5 ion exchange column in (figure 5.2 and
5.3, right) and final purification was done using a GE Healthcare Superdex-200 gel
filtration column in 100 mM BES (pH 7.4), 200 mM NaCl, 1 mM EGTA, and 1 mM
EDTA. Purity was assayed on a GE Healthcare 20% PhastGel as one band at the
appropriate molecular weight (Figure 5.3). Protein concentration was measured with the
Pierce BCA Protein Assay kit (Bradford assay), as well as O.D. 280. Figures 5.2 thru 5.3
shows the syt I C2A protein in various stages of purification. While the figures show
either WT or the Y180F mutant, the purification for the WT, as well as the mutant was
done the same and there were no noticeable differences between the samples with the
exception of the Y180N mutant, which would not concentrate.
Once purified and
concentrated, the protein samples were quick frozen in liquid nitrogen and stored at 80°C until use. For use, samples were removed from the -80°C and placed on ice. The
sample was then spun at 4°C, 15,000 rpm for 20 min and a protein concentration
determination was done prior to use.
.
61
Figure 5.1; Ion exchange chromatograph of Syt I C2A WT. The purification was done
using a Mono S 5/5 column (GE Healthcare) on a ÄKTA™ purifier FPLC (Fast protein
liquid chromatography) unit (GE Healthcare). The GST purified protein was filtered
through a 0.2 µm syringe filter and bound to the column in 100 mM BES (pH 7.4), 10
mM NaCl. The bound protein was eluted using a gradient in the same buffer with 1M
NaCl. A 20% SDS PAGE gel of the fractions is shown in figure 5.2 right.
Figure 5.2; 20% SDS PAGE gel of human synaptotagmin I C2A WT. The gel on the left
shows the expression, initial purification and enzymatic removal of the selection tag.
Lane one shows the molecular weight markers (GE Healthcare low molecular weight
marker). Lane two and three shows a whole cell lysate before induction and post
induction respectively. 1 ml of cell culture was diluted to 0.7 O.D. 600 and spun at 5,000
kg for 5 min. The supernatant was poured off and the pellet was resuspended in 100 µl
2M urea and 100 µl 2X Phast sauce (20 mM Tris [pH 8.0], 2 mM EDTA, 5% SDS, 10 %
v/v ß-ME). Lane four is the cell lysis pellet. The lysis pellet was prepared the same as
the pre induction/post induction samples except that cell pellet size was estimated, lane
62
five is the uncut elution from GST resin and lane six is the fusion protein cut with
thrombin protease. Samples from lane five and six were spun at 4°C, 15,000 rpm for 20
min prior to gel loading to ensure the protein is soluble. The gel on the right shows
protein samples from the ion exchange purification step (figure 5.1). The samples were
taken from different peaks from the purification step. Lane one is the low molecular
weight markers (GE Healthcare), lane two is fraction number two, lane three is fraction
number three, lane four is fraction number four, lane five is fraction number nine and
lane six is the thrombin digested protein shown in lane six of the left gel used as a
control. The ion exchange chromatogram is shown in figure 5.1 and the fractions are
labeled. After ion exchange, the protein was further purified using gel filtration (gel not
shown).
Figure 5.3; 20% SDS PAGE gel of human synaptotagmin C2A Y180F. The gel on the
left shows the purification steps of the Y180F mutant. Lane one is the molecular weight
markers, lane two is the uncut fusion protein after elution from GST resin and lane three
shows the cut protein with the selection tag (GST) and Y180F mutant. The gel on the
right shows a dilution of the protein from 26 mg/ml to 1.5 mg/ml. The gel on the right is
after ion exchange, gel filtration and protein concentration.
63
Figure 5.4; MALDI TOF mass spectrometry graph of purified syt I C2A Y180F. After
gel filtration and concentration, the sample was diluted to 1 mg/ml. 10 µl of the 1 mg/ml
sample was prepared using a ZipTipC18 (Milipore) to remove salt. The protein was
applied to and eluted from the ZipTip using standard protocols provided by the company.
The calculated mass of syt I C2A Y180F is 16,190 kDa and the experimental mass is
16,193 kDa (above). The calculated and experimental masses agree to a high degree.
The data were collected on a Ciphergen proteinchip® system. While this graph shows
the mass spec of the Y180F mutant, the graphs of the other purified samples show similar
results.
Crystallization
All protein samples were concentrated to 25 mg/ml and spun 15,000 rpm for 20
min at 4°C in a microfuge immediately prior to crystal screen set up. Protein samples
were kept in ice throughout the crystallization setup. Hanging drop (2+2) crystal trials
were conducted using a sparse matrix crystallization strategy.
Two microliters of
purified protein was mixed with two microliters of the crystallization mother liquor
containing the precipitant on a siliconized glass cover slip (Hampton Research). Each
glass cover slip was placed on a well of a pre-greased Limbro 24 well plate. Each well
contained 500 µl of crystallization mother liquor. Initial crystal conditions were obtained
using commercially available screens purchased from Molecular Dimensions and
Hampton Research.
Once the initial crystallization condition was determined, the
condition was further refined using the online program Make Tray (Hampton Research,
http://hamptonresearch.com/make_tray.aspx). The program allows a user to enter initial
conditions and then creates a recipe for a screen that explores the chemical space around
64
that condition. All crystals were grown at 17°C in a environmentally stable vibration
resistant incubator. A typical protein crystal took approximately 1 week to grow to
sufficient size for data collection. For X-ray data collection, the crystals were quick
frozen in their own mother liquor using liquid nitrogen and stored at 100 Kelvin until
transferred to the goniometer.
WT, D232N and D238N
Synaptotagmin I C2A WT, D232N and D238N were crystallized using previously
published techniques (23). Diffraction quality crystals of the syt I C2A WT protein were
grown in 100 mM HEPES (pH 7.1), 2.3 M LiSO4. Crystals were grown at 17°C and took
about one week to grow to sufficient size. Crystals of each synaptotagmin I C2A protein
are shown in Figures 5.5-5.7.
Y180F
Synaptotagmin I C2A Y180F was crystallized in two different space groups, P21
and I4.
P21 space group
The P21 crystals were crystallized from protein purified using the pGEX-KG
vector (155), while the I4 crystal form was from protein expressed using the p202 vector
(kindly provided by Dr. Jim Sacchetini, TAMU). The difference in the two vectors is the
N-terminal tail. Whereas the monoclinic crystal form possesses an additional 14 amino
acids at the N-terminus due to the expression vector, the p202 vector is a derivative of
pET28b that contains an MBP (maltose binding protein) selection tag in addition to the
His 6 tag and a TEV cleavage sequence instead of thrombin. The p202 vector results in
an extra five residues on the C-teminus (…GDITH…).
The pGEX-KG vector has been modified from the original pGEX vector (GE
Healthcare) to contain a glycine-rich linker (PGISGGGGGILDSM) immediately
following the thrombin cleavage site. The linker increases thrombin cleavage efficiency,
but also adds extra residues to the C-terminus of the target protein. Crystals in the I4
65
space group were grown in 1.2 M sodium malonate pH 6.75, figure 5.6; while the P21
crystal were grown in lithium sulfate.
I4 space group
The I4 tetragonal crystals were obtained using the p202. The C2A Y180F I4
crystals were slightly more difficult to crystallize than those in the P21 space group.
Preliminary crystallization trials resulted in crystals too small to collect diffraction data;
however, using a seeding protocol, diffraction quality crystals were obtained. One round
of crystal seeding was required. Initial crystals were obtained in 2.3 M malonate, 100
mM HEPES pH 7.1. One crystal of the first round of crystallization was added to 150 µl
the same mother liquor as above, physically crushed, vortexed at RT for 5 min and spun
at 16,000 X g for 10 min at RT. The second round of crystallization included adding 5 µl
of the crushed crystal solution to the 500 µl crystallization reservoir. 2 + 2 hanging drops
(2 µl protein + 2µl mother liquor mixed together) were used for the crystallization setup.
The crushed crystal solution was used for the protein. The procedure resulted in large
diffraction quality crystals. Figure 5.5 are crystals of C2A Y180F grown in 2.3 M
malonate pH 7.1 in 100 mM HEPES.
Fig 5.5; Crystals of Syt I p202 C2A Y180F
grown in 1.2 M Na Malonate pH 6.7.
Fig 5.6 Syt I C2A D232N grown in 2.3
M LiSO4 and 100 mM HEPES
pH 7.0
66
Fig. 5.7 Syt I C2A WT grown in 2.3 M LiSO4 and
100 mM HEPES pH 7.0
Diffraction data collection
Data were collected at the Stanford Synchrotron Radiation Laboratory (SSRL) on
beamline 11-1 at a wavelength of 0.97 Å (Table 5.1). The data sets were collected at 100
K using an ADSC image plate detector. The data was integrated, reduced, and scaled
using HKL2000 (81). The crystals were indexed in the monoclinic space group P21 for
the wild type, D232N, D238N and Y180F and tetragonal space group I4 for the Y180F
with Ca+2; data statistics are summarized in Table 5.1.
D232N and D238N
Syt I D232N data were collected using the automated data collection system at
SLAC, Web-Ice (156). Web-Ice is a web browser interfaced application designed to
optimize automated crystallographic data collection at synchrotrons.
The program
screens the crystal and suggests optimal experimental parameters to optimize the
collection of the data set. The crystals were indexed in the orthorhombic space group
P21, data statistics are summarized in Table 5.1.
Structure Determination and Refinement
The structure of all synaptotagmin 1 C2A mutants was solved via molecular
replacement with Phaser (82) using diffraction data collected from one crystal (Table
5.1). Initial model coordinates were obtained using the C2A domain (PDB code 1RSY).
67
Model building was done with Coot (83). Initial refinement of the model was carried out
with ccp4i with a random subset of all data set aside for calculation of Rfree (10%).
Refinement was completed using phenix.refine (157). Manual adjustments to the models
were carried out with Coot. After refinement of the protein was complete, solvent
molecules were assessed followed by manual adjustments. Ions were assigned on the
basis of the chemical environment, the refined B factor vs that of the other water
molecules, and the available coordination potential. The structure of C2A was verified
by examining a simulated annealing omit map generated with CNS v1.2. For simulated
annealing, 5% of the data was removed. There were no residues in disallowed regions of
the Ramachandran map (Table 5.11). All figures were rendered with Pymol (84).
68
Syt I C2A WT
Crystallographic
Data
Detector
Beamline 9-1,
SSRL, λ = 0.817
Å
ADSC Quantum
315R CCD,
D =400 mm
fx
Space Group
Syt I C2A
Y180F I4
Beamline 9-1,
SSRL, λ =
0.818Å
ADSC Quantum
315R CCD,
D =400 mm
fx
P2 2 2
Syt I C2A
Y180F P21
Beamline 9-1,
SSRL, λ = 0.818
Å
ADSC Quantum
315R CCD,
D =400 mm
fx
Syt I C2A
D232N
Beamline 9-1,
SSRL, λ = 0.818
Å
ADSC Quantum
315R CCD,
D =400 mm
fx
Syt I C2A
D238N
Beamline 9-1,
SSRL, λ = 0.818
Å
ADSC Quantum
315R CCD,
D =400 mm
fx
I4
P21
P21
P21
a= 41.73 Å b=
38.54 Å c=
44.29 Å
50 – 0.89
a=99.6 Å b=99.6
Å c=77.2 Å
50-1.7
a=41.8 Å
b=38.4 Å
c=44.1 Å
50-1.29
a=41.8 Å
b=38.4 Å
c=44.1 Å
50-1.2
a=41.8 Å
b=38.4 Å
c=44.1 Å
50-0.95
678,475
248,712
206,622
399,537
394,188
Unique Reflections
29,346
41,452
34,437
43,103
65,452
Completeness (%)*
99.5 (99.9)
99.1 (98.0)
99.9 (99.6)
99.2
98.0
R merge (%)*
3.3 (60)
5.1 (57)
6.0 (15.5)
4.0 (31)
8.2 (45)
I/σ (I)*
51 (2.0)
82 (4.2)
34 (9.3)
39.5 (4.6)
70.4 (2.5)
Unit cell Volume
70,582
766,219
70,167
69,947
70,513
(Å )
Solvent content *
43.5 %
56.8%
50.6%
50.6%
50.6%
Multiplicity*
7.0 (4.6)
14.9 (12)
6.9 (5.2)
6.2 (4.1)
7.8 (3.7)
1 1 1
Unit-Cell
Parameters
Resolution Range
(Å)
Total Reflections
3
Refinement Statistics
Residue range for
Refinement (Å)
Total reflections
used
r.m.s.d bond
lengths (Å)
r.m.s.d. angles (°)
50 – 0.89
50-1.7
50-1.3
50-1.2
50-0.95
97,100
41,107
22,378
43,103
64,381
0.020
0.004
0.005
0.009
0.009
1.75
0.83
1.30
1.37
1.30
Crystallographic Rfactor (%)
R-free ¶ (%)
14.0
18.0
22.4
14.4
18.1
15.6
20.3
23.5
17.0
19.2
Table 5.1; Summary of crystallographic statistics for each protein.
69
Figure 5.8; 0.97Å electron density map of synaptotagmin I WT C2A domain.
RESULTS AND DISCUSSION
The “…SDPYVKV…” motif is highly conserved in the synaptotagmin family
(Figure 3.4). A single point mutation (AD3) at the tyrosine residue in C2B results in a
protein that has a reduced capacity to bind Ca+2. The mutation affects Ca+2 binding;
however, the mutation is distant from the Ca+2 binding pocket. This is a paradox, since
structural changes are not thought to occur in C2 domains of synaptotagmin (47).
Previous experiments were unable to discern why a mutation distant from the Ca+2
binding pocket affects Ca+2 binding. To examine the structural effect of point mutations
in the motif in the C2A domain, we created two point mutants, Y180N to mimic the AD3
mutation in the C2A domain, and Y180F to keep the steric bulk of the residue, but
remove the hydrogen bonding potential. We successfully crystallized the Y180F mutant;
however, the Y180N protein proved difficult to manipulate and is very unstable in
solution. This observation could reflect the aberrant phenotype observed with the AD3
(Y311N) mutation in Drosophila. Concentrations of Y180N reached a maximum of 5
mg/ml, which proved too low for crystallization. However, the Y180F mutation was well
behaved in solution and was amenable to crystallographic analysis.
We solved the
structure of the wild type and Y180F mutant to resolutions of 0.89Å and 1.3Å
respectively (Table 5.1).
70
Figure; 5.8: Structural alignment of Syt I C2A WT, Y180F without Ca and Y180F with
Ca+2. The green spheres represent Ca+2. The WT molecule is shown in blue. The
Y180F structure with Ca+2 is shown in green with yellow Ca+2 binding loops and Y180F
without Ca+2 is shown in peach.
In Figure 5.8, the Ca+2 coordinating residues for each protein are shown along
with His 237 and residue 180 which is a Tyr in the wt but has been mutated to Phe in the
Y180F mutant. The His residue is in a different rotomer in the Y180F mutant compared
to the WT. Both Y180F structures came from the same crystal, the I4 space group (figure
5.6). In this structure, there are two molecules in the asymmetric unit, one bound to ion
and one unbound. The figure shows C2A in both the Ca+2 bound state as well as the
unbound state. The majority of the structure is identical between the different structures
with an average RMSD of 0.99Å over the entire molecule; however, the Ca+2 binding
loops differ considerably. The largest difference between the loops is seen in loop 1;
however, this portion of the Ca+2 binding pocket is highly disordered in all of the C2A
structures except the Y180F mutant with cation bound (figure 5.12). Loop 3 is stable in
each of the structures but differs in its absolute position between the different mutants.
The WT structure, shown in green/yellow, and the Y180F mutant with bound ion, shown
in blue, have very similar loop 3 positions. The Y180F mutant without bound ion, shown
71
in peach, however has a collapsed loop 3 similar to what is seen in the synaptotagmin I
C2AB structure in chapter 3. The addition of ion could give stability to the loops and
affect lipid interaction.
The three dimensional structure of synaptotagmin C2A WT
During this project, we took advantage of the synchrotron source and cryofreezing conditions to collection ultra-high resolution of the C2A domain of rat Syt1.
The rat synaptotagmin I C2A WT domain was solved to 0.89Å resolution and agrees with
previously published structures. We solved the structure with Se-met phasing techniques
and refined the structure with REFMAC and phenix.refine. A portion of the electron
density is shown in figure 5.8. The structure is presently the highest resolution crystal
structure of the synaptotagmin I C2A domain known. The best previous resolution is 1.9
Å (23).
Figure 5.10 shows a model of the Syt C2A WT molecule at a resolution of 0.89Å.
The protein was crystallized in the P21 space group. The model is colored by b-factor.
The highest b-factors, the most dynamic portions of the protein, are yellow and red. Red
indicates the most dynamic regions. The lowest b-factors, the least dynamic and most
stable portions of the protein, are shown in green and blue. Blue indicates the most stable
regions of the molecule. The most dynamic portions of the molecule include the Ca+2
binding loops, loop1, as well as the polybasic region. The calcium ion coordinating
residues as well as tyrosine 180 and histidine 237 are highlighted. The calcium ion
coordinating residues are all pointing toward the center of the binding pocket in a
position ready to accept divalent cation. A hydrogen bond exists between tyrosine 180
and histidine 237.
Our hypothesis is that this interaction is acting as a regulatory
mechanism for loop3. The hydrogen bond stabilizes the loop in a position where it can
readily interact with calcium ion and phospholipid.
72
Figure 5.9A: Loop three of Syt I C2A
Y180F w/o Ca2+
Figure 5.9B: Loop three of Syt I C2A
Y180F with Ca2+
Figure 5.9C; Loop three of Syt I C2A
Y180F in the P21 crystal form.
Figure 5.9D; Alignment of all three loops
Yellow is the I4 with ion, peach is the I4
without ion and blue is the P21 without
ion.
Figure 5.9 A thru D shows loop three of the Y180F mutant in the I4 and P21
crystal form. A shows C2A Y180F (I4 space group) without Ca+2 bound, B is with Ca+2
bound (I4 space group), C is loop 3 of Y180F crystallized in the P21 space group. Good
electron density is seen for loop 3 in all of the structures.
73
Figure 5.10: Model of Syt I C2A WT at 0.89Å resolution. The model is colored by bfactor. Blue and green colors represent low b-factors and orange are red represent higher
b-factors with red being the highest.
The three dimensional structure of synaptotagmin C2A Y180F.
P21 crystallographic setting for rat syt1 C2A
To investigate whether the human synaptotagmin I C2A Y180F mutant has an
effect on the absolute position of loop 3, we used X-ray crystallography to examine this
mutation of C2A at atomic detail. The Y180F mutant was crystallized in two different
space groups, I4 and P21. The tetragonal (I4) crystal form was solved using molecular
replacement techniques using Phaser (158) and is shown in figure 5.12; the molecule
crystallized in the P21 space group is shown in figure 5.11. The Y180F structure in the
P21 space group was solved to a resolution of 1.3 Å, and the I4 was solved at a resolution
of 1.7 Å and the structural statistics are summarized in Table 5.1. Due to the molar
74
concentration of sulfate the crystallization media, no Ca2+ is present within the divalent
cation binding pocket of the C2A that crystallized in P21. However, we observed that
loop3 of the Ca+2 binding pocket is in a different position compared to the WT molecule
that was crystallized in the same space group.
Figure 5.11; Model of Syt I C2A Y180F crystallized in the P21 space group. The
structure was solved to 1.3Å. The model is colored by b-factor. Blue and green colors
represent low b-factors and orange are red represent higher b-factors with red being the
highest. Loop1 is the most dynamic portion of the molecule similar to the non cation
bound molecule in the Y180F I4 crystal form. Also shown is phenylalanine 180 and
histidine 237. In the Y180F mutant, there is no interaction between the residues.
I4 crystallographic setting for syt1 C2A
The I4 crystal structure of Y180F C2A was refined using data to 1.7Å resolution.
In the I4 space group, there are two molecules in the asymmetric unit that rotated with
respect to each other by approximately 180 degrees. One molecule in the asymmetric unit
(chain B) is partially occupied with cation (probably Mg2+ contamination from the
75
crystallization media), whereas the other chain (chain A) is empty. Figure 5.12 shows the
X-ray structure of both Y180F molecules in the asymmetric unit.
This is a rather
confusing result, since one would expect that both molecules will be exposed to the same
divalent cation concentration, and therefore, they should be in similar conformations. One
explanation for this unusual observation are the differences in the hydrogen bonding
partners within the Ca2+ binding loops between the two molecules in the asymmetric unit.
If we plot the C-alpha B-factors of chain B (chain with divalent cation) as a function of
residue number (Figure 5.13), we see that loop 3 has the highest b-factor and loop1 has a
relatively low b-factor. In chain-A, the opposite is measured. The Ca2+ binding loops for
chain-A are located at the bottom of the figure. Loop3 has a low overall b-factor, while
loop1 has a high b-factor.
Figure 5.12: Model of Syt I C2A Y180F in the I4 space group. There are two molecules
in the asymmetric unit. The molecules are rotated approximately 180 degrees with
respect to each other. The molecule labeled chain-B contains cation while chain-A does
not. The models are colored by b-factor. Blue and green colors represent low b-factors
and orange are red represent higher b-factors with red being the highest. The Ca2+
binding loops of chain-B are shown at the top of the figure. In this chain, loop3 has the
76
highest b-factor and loop1 has a relatively low b-factor. In chain-A, the opposite is seen.
The Ca2+ binding loops for chain-A are located at the bottom of the figure. Loop3 has a
low b-factor and loop1 has a high b-factor.
Figure 5.13: Comparison of average B-factors for the two, C2A Y180F (I4 crystal form),
molecules in the asymmetric unit. The blue line represents the A chain and the magenta
line represents the B chain. The A molecule has a flexible loop 1 while the other
molecule has a flexible loop 3.
A reasonable explanation is that the binding of Asp 172 (one of the Ca2+ binding
residue on loop1, Figure 5.14) to divalent cation lowers the overall temperature factor of
the loop 1 due to the linkage of Asp 172 on Loop 1. Loop3, on the other hand, is
disordered when divalent cation is present because Asp 232 is binding to cation, and is no
longer available to stabilize Loop 3 (Figure 5.14). Following the same rationale, when
ion is not present, loop 1 is disordered as usual. Asp232 is now free to interact with other
neighboring residues (Ser 235 backbone carbonyl). This interaction reduces disorder in
the loop which lowers the temperature-factors. When ion is bound, the coordinating
residues are interacting with the divalent cation and cannot form a hydrogen bond with
neighboring residues. The effect is to increase disorder which increases b-factors.
From our results, it is clear that the rotameric position of Asp232 is dictated by the
presence or absence of cation. In addition, the rotameric position of this residue can
77
result is molecules with different overall shapes leading to our I4 crystal form where two
molecules in the asymmetric unit have different cation binding modes. Given the genetic
evidence that D232N presents a different phenotype relative to the other Ca+2 binding
residues, it is possible that Asp232 is the molecular basis of Ca+2 sensitivity and lipid
binding cooperativity in the C2 domains of Syt1.
Figure 5.14: Loop3 of Syt I C2A Y180F from the I4 space group. Only loop3 is shown.
The molecule on the left is not occupied with cation and the figure on the right is
occupied with cation. The figure on the left shows D232 interacting with the backbone of
other residues in loop3 which decreases the disorder of the loop. The figure on the right
shows D232 interacting with cation and not the backbone. The interaction with cation
and not backbone cause an increase in disorder of loop3.
Comparison of synaptotagmin I C2A WT and Y180F crystal structures.
Like the C2A WT structure, the structure of the Y180F mutant contains the
classic greek key motif. The overall RMSD between the WT structure and the Y180F
structure is 0.4 Å, with the highest deviation at loop 3 of the Ca+2 binding pocket. Solid
electron density was observed for loop 3 in both the WT and Y180F structures indicating
a stable conformation for the loop. The ribbon diagrams of both the WT structure and the
78
Y180F structure are overlaid in figure 5.15. The diagram shows the change in the
absolute position of loop 3. In the Y180F mutant, Histidine 237 is no longer hydrogen
bonded to Tyrosine 180. This lack of interaction leads to a change in the absolute
position of loop 3. The change in position causes the Ca+2 binding pocket to open to a
position where the Ca+2 coordinating residues are not in a position to bind Ca+2
efficiently.
Figure 5.15; Structural alignment of C2A WT and Y180F. The wild type molecule is
shown in light blue and the Y180F mutant is shown in magenta. Residue 180 and 237 are
highlighted. In the wild type, there is a hydrogen bond between tyrosine 180 and
histidine 237. In the mutant, the bond is disrupted and the rotomer of the histidine is
different. Disruption of the hydrogen bond causes a collapse so loop3 of the calcium ion
binding pocket.
79
The three dimensional structure of synaptotagmin I C2A D232N and D238N.
D232N
The Syt I C2A D232N mutant has previously been associated with an apparent
gain of function in that it initiates neurotransmitter release at lower Ca+2 concentrations;
however, it’s affinity for calcium ion is decreased. The three Ca2+-binding sites of the
synaptotagmin 1 C2A domain are formed by six amino acid side chains, five aspartate
residues and one serine residue located on the Ca+2 binding loops (35). Aspartate 232 is
one of the residues responsible for the coordination of Ca+2. Previous studies have shown
that mutation of this residue to an asparagine severely altered the Ca2+-binding properties
of the domain (36). The same study generated knock-in mice carrying the D232N
mutation. The mice showed no outward abnormalities; however, electrophysiological
recordings from cultured hippocampal neurons from newborn knock-in mice showed that
the D232N mutant was activated at lower Ca+2 concentrations than the WT protein (98).
The mechanism for the apparent gain of function, but decreased Ca+2 ion affinity of the
D232N mutant has previously evaded researchers. Our X-ray crystal structure of the
D232N mutant gives some insight into the molecular reason behind the observed gain of
function. In our structure, the mutant residue asparagine 232, is hydrogen bonded to the
backbone of loop3 whereas in the WT structure, residue 232 is directed toward the Ca+2
binding pocket (Figure 5.16). The D232N mutation has the effect of adding stability to
loop 3 which can allow it to bind Ca+2 at a lower affinity, but still act as an effective Ca+2
sensor. Our data demonstrate that a single amino acid substitution in synaptotagmin-1
C2A D232N can have an effect on how the molecule interacts with Ca+2 and gives insight
into inter-molecular mechanisms of Ca+2 binding and neurotransmitter release.
80
Figure 5.16: Model of Syt I C2A D232N at a resolution of 1.2Å. The model is colored
by b-factor. Blue and green colors represent low b-factors and orange are red represent
higher b-factors with red being the highest. Loop1 is the most dynamic portion of the
molecule similar to the non cation bound molecule in the Y180F I4 crystal form. Also
shown is phenylalanine 180 and histidine 237. There is a hydrogen bond between these
two residues (distance is 2.6Å).
81
Figure 5.17: Loop 3 of Syt I C2A D232N. The figure shows loop 3 of the C2A D232N
mutant. In this structure, the Asn is interacting with the neighboring carbonyl of residue
236 (Lys).
Figure 5.17 shows loop3 of the calcium binding pocket of Syt1 C2A. The residue
N232 is interacting with the carbonyl of K236. This interaction effectively decreases
disorder in the loop. Previous studies have shown that this mutant’s ability to bind
calcium is diminished; however, it induces neurotransmitter release at lower Ca2+
concentrations.
This is counter intuitive in that one would expect neurotransmitter
release to also be adversely effected due to a decrease in affinity for calcium ion. One
possible explanation for this is the way asparagine 232 hydrogen bonds to lysine 236.
Similar to what is seen in the Y180F mutant, where the b-factors differ between the
cation bound state and the unbound state (figure 5.13), the interaction between residue
232 and the neighboring residues has an effect on the stability of loop3. It is possible that
N232 is able to interact with calcium ion and the neighboring loop3 residues
simultaneously.
If this is occurring, the stability of loop3 would increase and the
increased stability could enhance its ability to interact with phospholipid. In the wild
82
type C2A domain, a similar hydrogen bonding is seen; however, because the residue
cannot simultaneously interact with calcium and the neighboring residues, it requires a
higher concentration of calcium ion to induce neurotransmitter release. Using this model,
the stability of loop3 affects the activity of the molecule. By binding Ca2+, loop3
becomes more stable allowing the protein to efficiently interact with lipid membrane. By
having a more stable loop3 a lower concentration of calcium ion can achieve the same
result, in the D232N mutant, as higher concentrations in the wt protein.
D238N
D238 is one of the six residues that coordinate Ca+2 ion in synaptotagmin I C2A.
Previous work has demonstrated that the single amino acid substitution selectively
decreases the apparent Ca2+ affinity of synaptotagmin-1 during phospholipid binding, and
decreases the amount of neurotransmitter release triggered by Ca2+ (98). We used X-ray
crystallography to examine this mutation of C2A at atomic detail. The D238N mutant
was crystallized in the P21 space group. The structure was solved to a resolution of
0.95Å using molecular replacement techniques using Phaser (158) and is shown in figure
5.18. The structural statistics are summarized in Table 5.1. Very little structural change
is seen in the D238N mutation when compared to the wild type protein. From our
structural analysis, we can conclude that D238N is a simple removal of a oxygen used for
Ca+2 coordination, whereas, the D232N mutation is more more complicated. Not only
does it participate in Ca+2 binding, it also acts to stabilize the shape of Loop3. Therefore,
this residue may act as a switch in that it links shape of Loop3 (which possesses residues
known to interact with phospholipids) to Ca+2 occupancy.
83
Figure 5.18: Model of Syt I C2A D238N at a resolution of 0.95Å. The model is colored
by b-factor. Blue and green colors represent low b-factors and orange are red represent
higher b-factors with red being the highest. Loop1 is the most dynamic portion of the
molecule similar to the non cation bound molecule in the Y180F I4 crystal form. Also
shown is phenylalanine 180 and histidine 237. There is a hydrogen bond between these
two residues (distance is 2.6Å).
LIFETIME FLUORESCENCE DENATURATION OF SYT I C2A
Steady state tryptophan fluorescence is one of the most common techniques that is
used to accurately quantitate the thermodynamic affects of ligand binding on a
macromolecule. This technique relies on monitoring the change in fluorescence of a Trp
residue from a primarily hydrophobic environment to a more aqueous environment as a
function of denaturant. This technique is often used to measure protein stability, and its
contribution to ligand binding.
The C2A domain of synaptotagmin has one Trp residue. Further, this lone Trp is
always solvent exposed, so this technique is not suited to measure changes in ΔG in
either mutations, Ca+2 binding, or phospholipid binding in this particular domain. The
84
only successful determination of thermodynamic parameters for C2 domains was made
by Shao (90), where they measured the heat denaturation of C2A in the presence and
absence of Ca+2 by circular dichroism. This technique generally works well; however, it
is unable to measure contribution that phospholipid binding has on stabilizing C2
domains. This is primarily because the light scattering caused by the large phospholipid
vesicles would impair obtaining an accurate CD signal from the protein. Lifetime
fluorescence techniques do not suffer from these deficiencies. This technique relies on
changes in fluorescence caused increasing collisions between the Trp reporter group with
backbone atoms. Further, this technique is insensitive to vesicle scatter.
The thermodynamic stability of C2A, C2A Y180F and C2A Y180N (in the presence of
Ca2+ and phospholipids) was analyzed using lifetime fluorescence. Figure 5.19 shows
plots of the lifetime thermal denaturation of the WT domain along with each of the
mutants (Y180N and Y180F). The Tm (melting temperature) increases with the addition
of calcium ion similar to the observation of Shao, et al (90). We have observed a small
increase in Tm with the addition of phospholipid (PS/PC), but there is a substantial
increase in stability with both Ca2+ and phospholipid. This result highlights that the C2
domain is a Ca2+ dependent phospholipid binding motif.
As we expected, the overall stability of the Y180F domain is very similar to the
WT domain (Fig 5.19) with Ca2+ and with phospholipid. This is expected since we are not
directly impacting either the Ca2+ binding or phospholipid binding components of the
domain to a large degree. The major difference we measure was in the shape of the
denaturation curve in the presence of Ca2+ and lipid. This indicates that the cooperativity
between Ca2+ and phospholipid has changed as a function of the Y180F mutation.
We also looked at the Y180N mutation in C2A (Figure 5.19). This mutation is
homologous to the original AD3 Drosophila mutation that was described in 1993 (87).
Unexpectedly, this mutation resulted in complete loss of cooperativity to Ca+2 or
phospholipids, and the stability of the entire domain was compromised. It is likely that a
85
similar effect occurred in Drosophila to impart a complete loss of Ca2+ dependent
exocytosis.
What is meant by Ca+2-dependent phospholipid binding? After all, there are
plenty of Ca+2 binding proteins and plenty of phospholipid biding proteins, but how does
a protein link the binding of one ligand to the binding of another completely different
ligand? It is possible that our thermodynamics measurements of these mutations of C2A,
as well as our crystallographic snapshots answers this question. In this scenario, the
rotameric position of Asp232 is the master logic switch in C2A linking Ca+2 occupancy
with the stability of Loop 3 and its phospholipid binding residues (Phe 234). When the
divalent cation binding pocket is empty, Asp232 forms H-bonds with the backbone amide
of Ser235. This strong interaction stabilizes Loop 3, but the domain is unable to bind
Ca+2 at full occupancy. In addition, His 237-Tyr180 also adds to the ΔG of stabilization
of Loop 3 in the absence of cation. However, when Ca+2 is present, the Asp232 rotamer
changes to bind Ca+2, thereby ignoring its previous bond to Ser235. Without further
stabilization, this loop would no doubt collapse (Figure 5.8 ‘I4+Ca+2). His235+Y180
provide a degree of stabilization to allow the domain to keep Phe234 in a position to bind
phospholipid, but only when Ca+2 is present.
86
Figure 5.19: Lifetime thermal denaturation plot of Syt I C2A. The protein was denatured
in the presence of EGTA (black sqares), Ca2+ (red circles), Lipid (blue triangles) and Ca2+
/ lipid (purple triangles). The C2A WT domain gains stability with the addition of
calcium ion and gains more stability with the addition of Ca2+ and lipid. The reason for
the increase in stability is most likely due to support added to the calcium ioin binding
loops with the addition of Ca2+ and lipid. The Syt I C2A Y180F mutant shows similar
stability to the WT; however, the Y180F has less cooperativity for calcium ion. The Syt I
C2A Y180N is structurally unstable and shows no difference in stability in the presence
of calcium ion or lipid.
INTER-MOLECULAR CRYSTAL CONTACTS
There is a possibility that inter-molecular crystal contacts are causing the
movement seen in the Ca+2 binding loops; however, we do not believe this is the case.
The crystal environment does have an effect on the absolute position of the Ca+2 binding
loops between the two crystal forms (monoclinic and tetragonal), but this is a matter of
extent of change, not whether changes occurs.
87
We analyzed the crystal contacts between adjacent molecules in the crystal to
determine the extent of interactions between neighboring molecules (figure 5.20 A and
B). The Syt I C2A Y180F molecule was crystallized in two different space groups,
monoclinic P21 and tetragonal I4. The P21 crystal form shown in Figure 5.20B has one
molecule in the asymmetric unit whereas the I4 form has two molecules per asymmetric
unit (Figure 5.20A). Interestingly, one of the molecules in the asymmetric unit has
divalent cation within the calcium ion-binding pocket, while the other does not. From Bfactor analysis of this crystallographic data, we have identified the ions as most likely
Mg+2. We did not purposefully add Mg+2 to the crystallization media, so it must be a
contaminant of the sodium malonate that we used for the crystallization media. Crystal
contacts between neighboring molecules in the crystal were analyzed to determine the
effect the contacts have on the position of the Ca+2 binding loops. The different crystal
forms were also compared two previously published X-ray crystallography and NMR
structures (PDB codes 1RSY and 1BYN respectively). Very few contacts are made and
those that are effect the position of the side chain rotomer and not the absolute position of
the loop. This can best be seen using the Y180F structure which was crystallized in two
different space groups. Figure 5.20 shows contacts between Syt I C2A Y180F loop 3 and
the neighboring molecule in the crystal lattice. The absolute position of loop 3 remains
constant between the space groups in the absence of Ca+2 ion; however, the position of
the loop does change when Ca+2 is present.
88
Figure 5.20A Crystal contacts of Syt I C2A Y180F I4 crystal form.
Figure 5.20B Crystal contacts of Syt I C2A Y180F P2 crystal
form.
Figure 5.20; Crystal contacts between neighboring molecules in the crystal. Figure A
and B shows the crystal contacts between loop 3 and the neighboring molecule in the
Y180F mutant in the I4 space group (A) and the P21 space group (B). There were no
crystal contacts between the other Ca+2 binding loops, loop 1 and 2, and adjacent
molecules. The inter-molecular contacts between the molecules is limited and is largely
between highly mobile rotomers (guanidino- groups of Arg) or Lys of adjacent
molecules.
89
Figure; 5.21; Alignment of the I4-molecule of Syt I C2A Y180F with ion with the solution
structure of C2A (PDB code 1byn). 1byn is the NMR solution structure of Syt I C2A wt
with Ca+2 ion bound. 1byn is in magenta and Y180F is in green/yellow. The Ca+2
binding loops are shown in yellow. Ca+2 ions are represented with black plus signs. The
figure shows that there is very little difference between the mutant structure with ion
bound and the solution structure of the WT molecule with bound ion.
As seen in Figure 5.21, very little structural difference is measured between the
solution structure and the crystal structure indicating a nominal effect of crystal packing
on the position of the Ca+2 binding loops. If crystal packing was an influence on the
position of the Ca+2 binding loops, one would expect to see a marked difference between
the crystal structure and the NMR solution structure.
90
Chapter 6 Discussion and Future Directions
Synaptotagmins make up a family of membrane-trafficking proteins that are
characterized by an N-terminal transmembrane domain, a linker of variable size, and two
C-terminal C2 domains denoted as C2A and C2B.
There are currently 17 known
synaptotagmin isoforms expressed in different tissues throughout the body.
Synaptotagmin I is localized to neuronal tissues and is the calcium ion sensor in fast
synchronous neurotransmitter release.
Analysis of the structure of synaptotagmin 1
C2AB reveals a conserved Tyr residue in the “SDPYVK” sequence. The Tyr residue
appears to modulate the absolute position of Loop 3 in both C2A and C2B. Mutation of
this residue in Syt1 C2B dramatically alters Ca2+ affinity (88), even though it is distant
from the Ca2+ binding site. The majority of studies assume that the two domains act like
balls on a string and that the two domains act in a very similar way. We show that this
assumption is not true. The two C2 domains of synaptotagmin I are distinct molecules
and have different properties.
In chapter 3, we present the crystal structure of synaptotagmin I C2AB. The 2.7
Å crystal structure of the cytosolic domains of human synaptotagmin 1 in the absence of
Ca2+ reveals a novel closed conformation of the protein. The shared interface between
C2A and C2B is stabilized by a network of interactions between residues on the Cterminal alpha-helix of the C2B domain and residues on loops 1-3 of the Ca2+-binding
region of C2A. These interactions alter the overall shape of the Ca2+-binding pocket of
C2A, but not that of C2B. Further analysis of the structure led us to the hypothesis that
Tyr 180 could regulate the position of Loop 3 in synaptotagmin by modulating the
position of Loop 3 (55). To test this hypothesis, we created two mutants, Y180N and
Y180F.
In chapter 4, the mechanical properties of Syt I C2A and C2B were analyzed
using single-molecule atomic force microscopy. We found that stretching the C2AB
91
domains of Syt1 resulted in two distinct unfolding force peaks. One larger force peak of
approximately 100, identified as C2B and a second peak of approximately 50 pN,
identified as C2A. Interestingly, a significant fraction of C2A domains unfolded through
a low force intermediate that was not observed in C2B. We conclude that these domains
have different mechanical properties. We hypothesize that a relatively small stretching
force may be sufficient to deform the effector-binding regions of the C2A domain and
modulate the affinity for soluble N-ethylmaleimide-sensitive factor (NSF) attachment
protein receptors (SNAREs), phospholipids, and Ca+2.
In chapter 5, we detail the structural effects of mutations on Syt I C2A. Two
mutants were created to better understand the role tyrosine 180 plays in the regulation of
the position of loop 3 and hence, gave us a more structural view of the cooperative nature
between Ca+2 and phospholipid..
Y180N was created in order to mimic the AD3
mutation. In Drosophila, this mutation, in C2B, causes the loss of Ca2+ dependent
neurotransmitter release (86, 87). Since C2A and C2B have the same overall tertiary
structure, we concluded that the same effect seen in C2B would also occur in C2A. The
Y180N mutant was difficult to work with however. We were able to purify it, however;
concentrating it to proved impossible.
We were able to achieve concentration of
approximately 5 mg/ml, but no more. This concentration is much too low to grow
crystals. Although the crystallographic studies proved impossible on this mutant, we
were able to obtain lifetime denaturation data. The data shown in figure 5.19 shows that
the mutant domain is highly unstable and shows no cooperativity for Ca2+ or phospholipid
interaction.
The Y180F mutant was created to keep the steric bulk of the residue but remove
the hydrogen bonding potential between residue 180 and neighboring residues,
specifically histidine 237. We were successful in purifying and concentrating this mutant
to purity and concentrations suitable for crystallographic studies. Previous structures of
Syt I C2A domains have crystallized in one crystal form, P21. The Y180F mutant
92
crystallized in two different crystal packing forms, P21 and I4. In both the P21 and I4
crystal forms, the position of loop 3 is in a different position than what is seen in the WT
protein. The position of loop 3 in Y180F is very close to what is seen in the crystal
structure of C2AB we reported in chapter 3.
The P21 arrangement in the Y180F mutant is the same space group seen in the
WT structure. Both the wt and the Y180F (P21 crystal form) structures are unbound to
Ca2+; however, the position of loop 3 is in a different position. The Y180F I4 space
group however is interesting in that of the two molecules in the asymmetric unit, one is
bound to ion while the other is unbound. The position of loop 3 is different in each
molecule of the asymmetric unit. The position of loop 3 in the unbound molecule is very
similar to the loop 3 position in the Y180F P21 space group whereas the position of loop
3 in the ion bound molecule is virtually identical the position of loop 3 in the WT
molecule.
Future studies on synaptotagmin could expand on the studies we have done. We
have learned much about how the tandem domains of Syt I interact with one another as
well as structural characteristics of the individual domains that effect calcium ion binding
and phospholipid interaction.
Using what we have learned in these studies, future
experiments could be designed to answer important questions that have been brought up
as a result of what we have learned.
We have presented the first structural evidence that the tandem C2 domains of Syt
I interact (chapter 3).
What we don’t know is how this interaction fits into how
synaptotagmin works and also how the interaction fits into neurotransmitter release as a
whole. Also, we have shown an interaction between the domains for Syt I only. There
are currently 17 known synaptotagmins.
Is the interaction uniform across all
synaptotagmins or is it specific for Syt I? Is the interaction a regulation mechanism? It’s
possible that the interaction deactivates C2A until calcium ion influx occurs, or some
93
other event still undetermined, at which time the domain becomes active and performs its
function. These are questions that still need to be answered.
We revealed important regulatory mechanisms in C2A that effect calcium ion and
phospholipid interaction. To design our experiments, we used the knowledge that there is
a highly conserved motif across all C2 domains and that previous studies have shown that
mutations in this motif effects synaptotagmins function as a calcium receptor. We have
shown that this regulatory mechanism is important for Syt I C2A however, we don’t
know if this is mechanism is common throughout the C2 domain family.
Similar
experiments on other C2 domains still need to be done. One example is dysferlin. This
protein is involved in limb-girdle muscular dystrophy and contains 6 tandem C2 domains.
Very little structural knowledge is currently known about the protein.
The same
experiments done in this dissertation could be applied to dysferlin which could help
better understand the disease and potentially help with a treatment.
We also discovered that the two C2 domains of Syt I have distinct mechanical
properties. This is an indication that each domain could play a specific role during
neurotransmitter release. What individual roles do they have? It could also be that the
different mechanical properties play a part in the regulation mechanism that was hinted at
in the interaction of C2A and C2B.
What we have learned about synaptotagmin in these studies can be used to design
many new studies. Outlined here are only a few of the possibilities. One of the most
important things we leaned is that synaptotagmin, and probably C2 domains in general,
are much more complicated than ‘simple balls’ on a string that bind cation and interact
with phospholipid membrane. Using what we now know, new avenues of research can
be explored.
94
Appendix: Ramachandran plots of C2A mutants.
Ramachandran plot of synaptotagmin I C2A WT. All residues are either in the favored or
allowed region.
95
Ramachandran plot of syt I C2A Y180F (I4 crystal form). All residues are either in the
favored region or allowed region.
96
Ramachandran plot of syt I C2A D232N. All residues are in the favored region.
97
Ramachandran plot of syt I C2A D238N. All residues are in the favored region.
98
Ramachandran plot of syt I C2A Y180F (P21 crystal form). All residues are either in the
favored region or allowed region.
99
REFERENCES
1.
Katz, R. L., and K. E. Eakins. 1969. The actions of neuromuscular blocking
agents on extraocular muscle and intraocular pressure. Proc R Soc Med 62:12171220.
2.
Llinas, R., I. Z. Steinberg, and K. Walton. 1981. Presynaptic calcium currents in
squid giant synapse. Biophys J 33:289-321.
3.
Sabatini, B. L., and W. G. Regehr. 1996. Timing of neurotransmission at fast
synapses in the mammalian brain. Nature 384:170-172.
4.
Almers, W. 1994. Synapses. How fast can you get? Nature 367:682-683.
5.
Jahn, R., and R. H. Scheller. 2006. SNAREs--engines for membrane fusion. Nat
Rev Mol Cell Biol 7:631-643.
6.
Ferro-Novick, S., and R. Jahn. 1994. Vesicle fusion from yeast to man. Nature
370:191-193.
7.
McNew, J. A., F. Parlati, R. Fukuda, R. J. Johnston, K. Paz, F. Paumet, T. H.
Sollner, and J. E. Rothman. 2000. Compartmental specificity of cellular
membrane fusion encoded in SNARE proteins. Nature 407:153-159.
8.
Schekman, R. 2004. Merging cultures in the study of membrane traffic. Nat Cell
Biol 6:483-486.
9.
Hanson, P. I., J. E. Heuser, and R. Jahn. 1997. Neurotransmitter release - four
years of SNARE complexes. Curr Opin Neurobiol 7:310-315.
10.
Sutton, R. B., D. Fasshauer, R. Jahn, and A. T. Brunger. 1998. Crystal structure of
a SNARE complex involved in synaptic exocytosis at 2.4 A resolution. Nature
395:347-353.
11.
Weber, T., B. V. Zemelman, J. A. McNew, B. Westermann, M. Gmachl, F.
Parlati, T. H. Sollner, and J. E. Rothman. 1998. SNAREpins: minimal machinery
for membrane fusion. Cell 92:759-772.
12.
Craxton, M. 2007. Evolutionary genomics of plant genes encoding N-terminalTM-C2 domain proteins and the similar FAM62 genes and synaptotagmin genes
of metazoans. BMC Genomics 8:259.
13.
Li, C., B. Ullrich, J. Z. Zhang, R. G. Anderson, N. Brose, and T. C. Sudhof. 1995.
Ca(2+)-dependent and -independent activities of neural and non-neural
synaptotagmins. Nature 375:594-599.
14.
Sudhof, T. C. 2002. Synaptotagmins: why so many? J Biol Chem 277:7629-7632.
100
15.
Geppert, M., Y. Goda, R. E. Hammer, C. Li, T. W. Rosahl, C. F. Stevens, and T.
C. Sudhof. 1994. Synaptotagmin I: a major Ca2+ sensor for transmitter release at
a central synapse. Cell 79:717-727.
16.
Sudhof, T. C. 1995. The synaptic vesicle cycle: a cascade of protein-protein
interactions. Nature 375:645-653.
17.
Sudhof, T. C., and J. Rizo. 1996. Synaptotagmins: C2-domain proteins that
regulate membrane traffic. Neuron 17:379-388.
18.
Perin, M. S., V. A. Fried, G. A. Mignery, R. Jahn, and T. C. Sudhof. 1990.
Phospholipid binding by a synaptic vesicle protein homologous to the regulatory
region of protein kinase C. Nature 345:260-263.
19.
Takamori, S., M. Holt, K. Stenius, E. A. Lemke, M. Gronborg, D. Riedel, H.
Urlaub, S. Schenck, B. Brugger, P. Ringler, S. A. Muller, B. Rammner, F. Grater,
J. S. Hub, B. L. De Groot, G. Mieskes, Y. Moriyama, J. Klingauf, H. Grubmuller,
J. Heuser, F. Wieland, and R. Jahn. 2006. Molecular anatomy of a trafficking
organelle. Cell 127:831-846.
20.
Chapman, E. R. 2002. Synaptotagmin: a Ca(2+) sensor that triggers exocytosis?
Nat Rev Mol Cell Biol 3:498-508.
21.
Cho, W., and R. V. Stahelin. 2006. Membrane binding and subcellular targeting
of C2 domains. Biochim Biophys Acta 1761:838-849.
22.
Nishizuka, Y. 1988. The molecular heterogeneity of protein kinase C and its
implications for cellular regulation. Nature 334:661-665.
23.
Sutton, R. B., B. A. Davletov, A. M. Berghuis, T. C. Sudhof, and S. R. Sprang.
1995. Structure of the first C2 domain of synaptotagmin I: a novel
Ca2+/phospholipid-binding fold. Cell 80:929-938.
24.
Essen, L. O., O. Perisic, R. Cheung, M. Katan, and R. L. Williams. 1996. Crystal
structure of a mammalian phosphoinositide-specific phospholipase C delta.
Nature 380:595-602.
25.
Perisic, O., S. Fong, D. E. Lynch, M. Bycroft, and R. L. Williams. 1998. Crystal
structure of a calcium-phospholipid binding domain from cytosolic phospholipase
A2. J Biol Chem 273:1596-1604.
26.
Sutton, R. B., and S. R. Sprang. 1998. Structure of the protein kinase Cbeta
phospholipid-binding C2 domain complexed with Ca2+. Structure 6:1395-1405.
27.
Pappa, H., J. Murray-Rust, L. V. Dekker, P. J. Parker, and N. Q. McDonald. 1998.
Crystal structure of the C2 domain from protein kinase C-delta. Structure 6:885894.
101
28.
Dai, H., D. R. Tomchick, J. Garcia, T. C. Sudhof, M. Machius, and J. Rizo. 2005.
Crystal structure of the RIM2 C2A-domain at 1.4 A resolution. Biochemistry
44:13533-13542.
29.
Nishiki, T., Y. Kamata, Y. Nemoto, A. Omori, T. Ito, M. Takahashi, and S.
Kozaki. 1994. Identification of protein receptor for Clostridium botulinum type B
neurotoxin in rat brain synaptosomes. J Biol Chem 269:10498-10503.
30.
Rummel, A., T. Karnath, T. Henke, H. Bigalke, and T. Binz. 2004.
Synaptotagmins I and II act as nerve cell receptors for botulinum neurotoxin G. J
Biol Chem 279:30865-30870.
31.
Dong, M., D. A. Richards, M. C. Goodnough, W. H. Tepp, E. A. Johnson, and E.
R. Chapman. 2003. Synaptotagmins I and II mediate entry of botulinum
neurotoxin B into cells. J Cell Biol 162:1293-1303.
32.
Perin, M. S., P. A. Johnston, T. Ozcelik, R. Jahn, U. Francke, and T. C. Sudhof.
1991. Structural and functional conservation of synaptotagmin (p65) in
Drosophila and humans. J Biol Chem 266:615-622.
33.
Chapman, E. R., and R. Jahn. 1994. Calcium-dependent interaction of the
cytoplasmic region of synaptotagmin with membranes. Autonomous function of a
single C2-homologous domain. J Biol Chem 269:5735-5741.
34.
Davletov, B. A., and T. C. Sudhof. 1993. A single C2 domain from
synaptotagmin I is sufficient for high affinity Ca2+/phospholipid binding. J Biol
Chem 268:26386-26390.
35.
Ubach, J., X. Zhang, X. Shao, T. C. Sudhof, and J. Rizo. 1998. Ca2+ binding to
synaptotagmin: how many Ca2+ ions bind to the tip of a C2-domain? Embo J
17:3921-3930.
36.
Fernandez-Chacon, R., O. H. Shin, A. Konigstorfer, M. F. Matos, A. C. Meyer, J.
Garcia, S. H. Gerber, J. Rizo, T. C. Sudhof, and C. Rosenmund. 2002.
Structure/function analysis of Ca2+ binding to the C2A domain of synaptotagmin
1. J Neurosci 22:8438-8446.
37.
Zhang, X., J. Rizo, and T. C. Sudhof. 1998. Mechanism of phospholipid binding
by the C2A-domain of synaptotagmin I. Biochemistry 37:12395-12403.
38.
Nalefski, E. A., and J. J. Falke. 1996. The C2 domain calcium-binding motif:
structural and functional diversity. Protein Sci 5:2375-2390.
39.
Matthew, W. D., L. Tsavaler, and L. F. Reichardt. 1981. Identification of a
synaptic vesicle-specific membrane protein with a wide distribution in neuronal
and neurosecretory tissue. J Cell Biol 91:257-269.
102
40.
Fernandez-Chacon, R., A. Konigstorfer, S. H. Gerber, J. Garcia, M. F. Matos, C.
F. Stevens, N. Brose, J. Rizo, C. Rosenmund, and T. C. Sudhof. 2001.
Synaptotagmin I functions as a calcium regulator of release probability. Nature
410:41-49.
41.
Mackler, J. M., J. A. Drummond, C. A. Loewen, I. M. Robinson, and N. E. Reist.
2002. The C(2)B Ca(2+)-binding motif of synaptotagmin is required for synaptic
transmission in vivo. Nature 418:340-344.
42.
Martens, S., M. M. Kozlov, and H. T. McMahon. 2007. How Synaptotagmin
Promotes Membrane Fusion. Science.
43.
Rufener, E., A. A. Frazier, C. M. Wieser, A. Hinderliter, and D. S. Cafiso. 2005.
Membrane-bound orientation and position of the synaptotagmin C2B domain
determined by site-directed spin labeling. Biochemistry 44:18-28.
44.
Schiavo, G., Q. M. Gu, G. D. Prestwich, T. H. Sollner, and J. E. Rothman. 1996.
Calcium-dependent switching of the specificity of phosphoinositide binding to
synaptotagmin. Proc Natl Acad Sci U S A 93:13327-13332.
45.
Aoyagi, K., T. Sugaya, M. Umeda, S. Yamamoto, S. Terakawa, and M.
Takahashi. 2005. The activation of exocytotic sites by the formation of
phosphatidylinositol 4,5-bisphosphate microdomains at syntaxin clusters. J Biol
Chem 280:17346-17352.
46.
Nalefski, E. A., M. A. Wisner, J. Z. Chen, S. R. Sprang, M. Fukuda, K.
Mikoshiba, and J. J. Falke. 2001. C2 domains from different Ca2+ signaling
pathways display functional and mechanistic diversity. Biochemistry 40:30893100.
47.
Shao, X., I. Fernandez, T. C. Sudhof, and J. Rizo. 1998. Solution structures of the
Ca2+-free and Ca2+-bound C2A domain of synaptotagmin I: does Ca2+ induce a
conformational change? Biochemistry 37:16106-16115.
48.
Sutton, R. B. 1997. PhD Dissertation. UTSMC, Dallas, TX, Dallas.
49.
Fernandez, I., D. Arac, J. Ubach, S. H. Gerber, O. Shin, Y. Gao, R. G. Anderson,
T. C. Sudhof, and J. Rizo. 2001. Three-dimensional structure of the
synaptotagmin 1 C2B-domain: synaptotagmin 1 as a phospholipid binding
machine. Neuron 32:1057-1069.
50.
Cheng, Y., S. M. Sequeira, L. Malinina, V. Tereshko, T. H. Sollner, and D. J.
Patel. 2004. Crystallographic identification of Ca2+ and Sr2+ coordination sites
in synaptotagmin I C2B domain. Protein Sci 13:2665-2672.
51.
Sutton, R. B., J. A. Ernst, and A. T. Brunger. 1999. Crystal structure of the
cytosolic C2A-C2B domains of synaptotagmin III. Implications for Ca(+2)independent snare complex interaction. J Cell Biol 147:589-598.
103
52.
Arac, D., X. Chen, H. A. Khant, J. Ubach, S. J. Ludtke, M. Kikkawa, A. E.
Johnson, W. Chiu, T. C. Sudhof, and J. Rizo. 2006. Close membrane-membrane
proximity induced by Ca(2+)-dependent multivalent binding of synaptotagmin-1
to phospholipids. Nat Struct Mol Biol 13:209-217.
53.
Davletov, B. A., and T. C. Sudhof. 1994. Ca(2+)-dependent conformational
change in synaptotagmin I. J Biol Chem 269:28547-28550.
54.
Garcia, R. A., C. E. Forde, and H. A. Godwin. 2000. Calcium triggers an
intramolecular association of the C2 domains in synaptotagmin. Proc Natl Acad
Sci U S A 97:5883-5888.
55.
Fuson, K. L., M. Montes, J. J. Robert, and R. B. Sutton. 2007. Structure of human
synaptotagmin 1 C2AB in the absence of Ca2+ reveals a novel domain
association. Biochemistry 46:13041-13048.
56.
Pollard, H. B., C. J. Pazoles, C. E. Creutz, and O. Zinder. 1980. Role of
intracellular proteins in the regulation of calcium action and transmitter release
during exocytosis. Monogr Neural Sci 7:106-116.
57.
Hutton, J. C. 1986. Calcium-binding proteins and secretion. Cell Calcium 7:339352.
58.
DeLorenzo, R. J. 1982. Calmodulin in neurotransmitter release and synaptic
function. Fed Proc 41:2265-2272.
59.
Sudhof, T. C. 2004. The synaptic vesicle cycle. Annu Rev Neurosci 27:509-547.
60.
Yoshihara, M., and E. S. Montana. 2004. The synaptotagmins: calcium sensors
for vesicular trafficking. Neuroscientist 10:566-574.
61.
Bhalla, A., W. C. Tucker, and E. R. Chapman. 2005. Synaptotagmin isoforms
couple distinct ranges of Ca2+, Ba2+, and Sr2+ concentration to SNAREmediated membrane fusion. Mol Biol Cell 16:4755-4764.
62.
von Poser, C., K. Ichtchenko, X. Shao, J. Rizo, and T. C. Sudhof. 1997. The
evolutionary pressure to inactivate. A subclass of synaptotagmins with an amino
acid substitution that abolishes Ca2+ binding. J Biol Chem 272:14314-14319.
63.
Craxton, M. 2004. Synaptotagmin gene content of the sequenced genomes. BMC
Genomics 5:43.
64.
Ubach, J., Y. Lao, I. Fernandez, D. Arac, T. C. Sudhof, and J. Rizo. 2001. The
C2B domain of synaptotagmin I is a Ca2+-binding module. Biochemistry
40:5854-5860.
65.
Bernard, A., and M. Payton. 1995. Fermentation and Growth of Eschericia coli
for optimal protein production. Current Protocols in Protein Science 5.3:1-18.
104
66.
Leslie, A. 1992. Recent changes to the MOSFLM package for processing film and
image plate data. Joint CCP4 + ESF-EAMCB Newsletter on Protein
Crystallography.
67.
Collaborative Computational Project, N. 1994. The CCP4 Suite: Programs for
Protein Crystallography. Acta Cryst. D 50:760-763.
68.
French, S., and K. Wilson. 1978. On the treatment of negative intensity
observations. Acta Cryst. A34:517-525.
69.
Tong, L., and M. G. Rossmann. 1997. Rotation function calculations with GLRF
program. Methods Enzymol 276:594-611.
70.
Shao, X., T. C. Sudhof, and J. Rizo. 1997. Assignment of the 1H, 15N and 13C
resonances of the calcium-free and calcium-bound forms of the first C2-domain
of synaptotagmin I. J Biomol NMR 10:307-308.
71.
Dennison, S. M., M. E. Bowen, A. T. Brunger, and B. R. Lentz. 2006. Neuronal
SNAREs Do Not Trigger Fusion between Synthetic Membranes but Do Promote
PEG-Mediated Membrane Fusion. Biophys J 90:1661-1675.
72.
Rickman, C., D. A. Archer, F. A. Meunier, M. Craxton, M. Fukuda, R. D.
Burgoyne, and B. Davletov. 2004. Synaptotagmin interaction with the
syntaxin/SNAP-25 dimer is mediated by an evolutionarily conserved motif and is
sensitive to inositol hexakisphosphate. J Biol Chem 279:12574-12579.
73.
Damer, C. K., and C. E. Creutz. 1994. Synergistic membrane interactions of the
two C2 domains of synaptotagmin. J Biol Chem 269:31115-31123.
74.
Hui, E., J. Bai, and E. R. Chapman. 2006. Ca2+-triggered simultaneous
membrane penetration of the tandem C2-domains of synaptotagmin I. Biophys J
91:1767-1777.
75.
Shao, X., B. A. Davletov, R. B. Sutton, T. C. Sudhof, and J. Rizo. 1996. Bipartite
Ca2+-binding motif in C2 domains of synaptotagmin and protein kinase C.
Science 273:248-251.
76.
Chae, Y. K., F. Abildgaard, E. R. Chapman, and J. L. Markley. 1998. Lipid
binding ridge on loops 2 and 3 of the C2A domain of synaptotagmin I as revealed
by NMR spectroscopy. J Biol Chem 273:25659-25663.
77.
Chapman, E. R., and A. F. Davis. 1998. Direct interaction of a Ca2+-binding loop
of synaptotagmin with lipid bilayers. J Biol Chem 273:13995-14001.
78.
Fernandez, J. M., and H. Li. 2004. Force-clamp spectroscopy monitors the folding
trajectory of a single protein. Science 303:1674-1678.
105
79.
Rizo, J., and T. C. Sudhof. 1998. C2-domains, structure and function of a
universal Ca2+-binding domain. J Biol Chem 273:15879-15882.
80.
Montes, M., K. L. Fuson, R. B. Sutton, and J. J. Robert. 2006. Purification,
crystallization and X-ray diffraction analysis of human synaptotagmin 1 C2AC2B. Acta Cryst. F.
81.
Otwinowski, Z., and W. Minor. 1997. Processing of X-ray Diffraction Data
Collected in Oscillation Mode. In Methods in Enzymology, Macromolecular
Crystallography, part A. C. Carter, and R. Sweet, editors. Academic Press, New
York. 307-326.
82.
McCoy, A. J., R. W. Grosse-Kunstleve, L. C. Storoni, and R. J. Read. 2005.
Likelihood-enhanced fast translation functions. . Acta Cryst. D61:458-464.
83.
Emsley, P., and K. Cowtan. 2004. Coot: model-building tools for molecular
graphics. Acta Crystallogr. D60:2126-2132.
84.
DeLano, W. L. 2002. The PyMOL Molecular Graphics System. DeLano
Scientific, Palo Alto, CA, USA.
85.
Strynadka, N. C., and M. N. James. 1989. Crystal structures of the helix-loophelix calcium-binding proteins. Annu Rev Biochem 58:951-998.
86.
Okamoto, T., T. Tamura, K. Suzuki, and Y. Kidokoro. 2005. External Ca2+
dependency of synaptic transmission in drosophila synaptotagmin I mutants. J
Neurophysiol 94:1574-1586.
87.
DiAntonio, A., K. D. Parfitt, and T. L. Schwarz. 1993. Synaptic transmission
persists in synaptotagmin mutants of Drosophila. Cell 73:1281-1290.
88.
Littleton, J. T., J. Bai, B. Vyas, R. Desai, A. E. Baltus, M. B. Garment, S. D.
Carlson, B. Ganetzky, and E. R. Chapman. 2001. Synaptotagmin mutants reveal
essential functions for the C2B domain in Ca2+-triggered fusion and recycling of
synaptic vesicles in vivo. J Neurosci 21:1421-1433.
89.
Robinson, I. M., R. Ranjan, and T. L. Schwarz. 2002. Synaptotagmins I and IV
promote transmitter release independently of Ca(2+) binding in the C(2)A
domain. Nature 418:336-340.
90.
Shao, X., C. Li, I. Fernandez, X. Zhang, T. C. Sudhof, and J. Rizo. 1997.
Synaptotagmin-syntaxin interaction: the C2 domain as a Ca2+-dependent
electrostatic switch. Neuron 18:133-142.
91.
Edwards, A. S., and A. C. Newton. 1997. Regulation of protein kinase C betaII by
its C2 domain. Biochemistry 36:15615-15623.
106
92.
Grass, I., S. Thiel, S. Honing, and V. Haucke. 2004. Recognition of a basic AP-2
binding motif within the C2B domain of synaptotagmin is dependent on
multimerization. J Biol Chem 279:54872-54880.
93.
Chapman, E. R., R. C. Desai, A. F. Davis, and C. K. Tornehl. 1998. Delineation
of the oligomerization, AP-2 binding, and synprint binding region of the C2B
domain of synaptotagmin. J Biol Chem 273:32966-32972.
94.
Jones, S., and J. M. Thornton. 1996. Principles of protein-protein interactions.
Proc Natl Acad Sci U S A 93:13-20.
95.
Charvin, N., C. L'Eveque, D. Walker, F. Berton, C. Raymond, M. Kataoka, Y.
Shoji-Kasai, M. Takahashi, M. De Waard, and M. J. Seagar. 1997. Direct
interaction of the calcium sensor protein synaptotagmin I with a cytoplasmic
domain of the alpha1A subunit of the P/Q-type calcium channel. Embo J 16:45914596.
96.
Cohen, R., B. M. Schmitt, and D. Atlas. 2005. Molecular identification and
reconstitution of depolarization-induced exocytosis monitored by membrane
capacitance. Biophys J 89:4364-4373.
97.
Fukuda, M., J. E. Moreira, V. Liu, M. Sugimori, K. Mikoshiba, and R. R. Llinas.
2000. Role of the conserved WHXL motif in the C terminus of synaptotagmin in
synaptic vesicle docking. Proc Natl Acad Sci U S A 97:14715-14719.
98.
Pang, Z. P., O. H. Shin, A. C. Meyer, C. Rosenmund, and T. C. Sudhof. 2006. A
gain-of-function mutation in synaptotagmin-1 reveals a critical role of Ca2+dependent soluble N-ethylmaleimide-sensitive factor attachment protein receptor
complex binding in synaptic exocytosis. J Neurosci 26:12556-12565.
99.
Kertz, J. A., P. F. Almeida, A. A. Frazier, A. K. Berg, and A. Hinderliter. 2007.
The Cooperative Response of Synaptotagmin I C2A. A Hypothesis for a Ca2+Driven Molecular Hammer. Biophys J 92:1409-1418.
100.
Nishiki, T., and G. J. Augustine. 2004. Dual roles of the C2B domain of
synaptotagmin I in synchronizing Ca2+-dependent neurotransmitter release. J
Neurosci 24:8542-8550.
101.
Rothman, J. E., and T. H. Sollner. 1997. Throttles and dampers: controlling the
engine of membrane fusion. Science 276:1212-1213.
102.
Liu, W., V. Montana, J. Bai, E. R. Chapman, U. Mohideen, and V. Parpura. 2006.
Single molecule mechanical probing of the SNARE protein interactions. Biophys
J 91:744-758.
103.
Lynch, K. L., R. R. Gerona, E. C. Larsen, R. F. Marcia, J. C. Mitchell, and T. F.
Martin. 2007. Synaptotagmin C2A Loop 2 Mediates Ca2+-dependent SNARE
107
Interactions Essential for Ca2+-triggered Vesicle Exocytosis. Mol Biol Cell
18:4957-4968.
104.
Sugita, S., O. H. Shin, W. Han, Y. Lao, and T. C. Sudhof. 2002. Synaptotagmins
form a hierarchy of exocytotic Ca(2+) sensors with distinct Ca(2+) affinities.
Embo J 21:270-280.
105.
Yersin, A., H. Hirling, P. Steiner, S. Magnin, R. Regazzi, B. Huni, P. Huguenot,
P. De los Rios, G. Dietler, S. Catsicas, and S. Kasas. 2003. Interactions between
synaptic vesicle fusion proteins explored by atomic force microscopy. Proc Natl
Acad Sci U S A 100:8736-8741.
106.
Li, F., F. Pincet, E. Perez, W. S. Eng, T. J. Melia, J. E. Rothman, and D. Tareste.
2007. Energetics and dynamics of SNAREpin folding across lipid bilayers. Nat
Struct Mol Biol 14:890-896.
107.
Frazier, A. A., C. R. Roller, J. J. Havelka, A. Hinderliter, and D. S. Cafiso. 2003.
Membrane-bound orientation and position of the synaptotagmin I C2A domain by
site-directed spin labeling. Biochemistry 42:96-105.
108.
Rief, M., M. Gautel, F. Oesterhelt, J. M. Fernandez, and H. E. Gaub. 1997.
Reversible unfolding of individual titin immunoglobulin domains by AFM.
Science 276:1109-1112.
109.
Li, H., W. A. Linke, A. F. Oberhauser, M. Carrion-Vazquez, J. G. Kerkvliet, H.
Lu, P. E. Marszalek, and J. M. Fernandez. 2002. Reverse engineering of the giant
muscle protein titin. Nature 418:998-1002.
110.
Fowler, S. B., R. B. Best, J. L. Toca Herrera, T. J. Rutherford, A. Steward, E.
Paci, M. Karplus, and J. Clarke. 2002. Mechanical unfolding of a titin Ig domain:
structure of unfolding intermediate revealed by combining AFM, molecular
dynamics simulations, NMR and protein engineering. J Mol Biol 322:841-849.
111.
Oberhauser, A. F., C. Badilla-Fernandez, M. Carrion-Vazquez, and J. M.
Fernandez. 2002. The mechanical hierarchies of fibronectin observed with singlemolecule AFM. J Mol Biol 319:433-447.
112.
Carl, P., C. H. Kwok, G. Manderson, D. W. Speicher, and D. E. Discher. 2001.
Forced unfolding modulated by disulfide bonds in the Ig domains of a cell
adhesion molecule. Proc Natl Acad Sci U S A 98:1565-1570.
113.
Bai, J., and E. R. Chapman. 2004. The C2 domains of synaptotagmin--partners in
exocytosis. Trends Biochem Sci 29:143-151.
114.
Mikoshiba, K., M. Fukuda, K. Ibata, H. Kabayama, and A. Mizutani. 1999. Role
of synaptotagmin, a Ca2+ and inositol polyphosphate binding protein, in
neurotransmitter release and neurite outgrowth. Chem Phys Lipids 98:59-67.
108
115.
Steward, A., J. L. Toca-Herrera, and J. Clarke. 2002. Versatile cloning system for
construction of multimeric proteins for use in atomic force microscopy. Protein
Sci 11:2179-2183.
116.
Carrion-Vazquez, M., A. F. Oberhauser, S. B. Fowler, P. E. Marszalek, S. E.
Broedel, J. Clarke, and J. M. Fernandez. 1999. Mechanical and chemical
unfolding of a single protein: a comparison. Proc Natl Acad Sci U S A 96:36943699.
117.
Bullard, B., W. A. Linke, and K. Leonard. 2002. Varieties of elastic protein in
invertebrate muscles. J Muscle Res Cell Motil 23:435-447.
118.
Miller, E., T. Garcia, S. Hultgren, and A. F. Oberhauser. 2006. The mechanical
properties of E. coli type 1 pili measured by atomic force microscopy techniques.
Biophys J 91:3848-3856.
119.
Oberhauser, A. F., P. E. Marszalek, H. P. Erickson, and J. M. Fernandez. 1998.
The molecular elasticity of the extracellular matrix protein tenascin. Nature
393:181-185.
120.
Florin, E. L., M. Rief, T. Lehmann, M. Ludvig, C. Dornmair, V. T. Moy, and H.
E. Gaub. 1995. Sensing specific molecular interactions with the atomic force
microscope. Biosens. Bioelectron 10:895-901.
121.
Lu, H., B. Isralewitz, A. Krammer, V. Vogel, and K. Schulten. 1998. Unfolding
of titin immunoglobulin domains by steered molecular dynamics simulation.
Biophys J 75:662-671.
122.
Phillips, J. C., R. Braun, W. Wang, J. Gumbart, E. Tajkhorshid, E. Villa, C.
Chipot, R. D. Skeel, L. Kale, and K. Schulten. 2005. Scalable molecular dynamics
with NAMD. J Comput Chem 26:1781-1802.
123.
Wright, C. F., A. Steward, and J. Clarke. 2004. Thermodynamic characterisation
of two transition states along parallel protein folding pathways. J Mol Biol
338:445-451.
124.
Wright, C. F., K. Lindorff-Larsen, L. G. Randles, and J. Clarke. 2003. Parallel
protein-unfolding pathways revealed and mapped. Nat Struct Biol 10:658-662.
125.
Li, H., A. F. Oberhauser, S. B. Fowler, J. Clarke, and J. M. Fernandez. 2000.
Atomic force microscopy reveals the mechanical design of a modular protein.
Proc Natl Acad Sci U S A 97:6527-6531.
126.
Hamill, S. J., A. Steward, and J. Clarke. 2000. The folding of an immunoglobulinlike Greek key protein is defined by a common-core nucleus and regions
constrained by topology. J Mol Biol 297:165-178.
109
127.
Fowler, S. B., and J. Clarke. 2001. Mapping the folding pathway of an
immunoglobulin domain: structural detail from Phi value analysis and movement
of the transition state. Structure 9:355-366.
128.
Bustamante, C., J. F. Marko, E. D. Siggia, and S. Smith. 1994. Entropic elasticity
of lambda-phage DNA. Science 265:1599-1600.
129.
Marko, J. F., and E. D. Siggia. 1995. Stretching DNA. Macromolecules 28:87598770.
130.
Carrion-Vazquez, M., A. F. Oberhauser, T. E. Fisher, P. E. Marszalek, H. Li, and
J. M. Fernandez. 2000. Mechanical design of proteins studied by single-molecule
force spectroscopy and protein engineering. Prog Biophys Mol Biol 74:63-91.
131.
Lu, H., and K. Schulten. 1999. Steered molecular dynamics simulations of forceinduced protein domain unfolding. Proteins 35:453-463.
132.
Li, L., H. H. Huang, C. L. Badilla, and J. M. Fernandez. 2005. Mechanical
unfolding intermediates observed by single-molecule force spectroscopy in a
fibronectin type III module. J Mol Biol 345:817-826.
133.
Schwaiger, I., A. Kardinal, M. Schleicher, A. A. Noegel, and M. Rief. 2004. A
mechanical unfolding intermediate in an actin-crosslinking protein. Nat Struct
Mol Biol 11:81-85.
134.
Koh, T. W., and H. J. Bellen. 2003. Synaptotagmin I, a Ca2+ sensor for
neurotransmitter release. Trends Neurosci 26:413-422.
135.
Yoshihara, M., and J. T. Littleton. 2002. Synaptotagmin I functions as a calcium
sensor to synchronize neurotransmitter release. Neuron 36:897-908.
136.
Wang, C. T., J. Bai, P. Y. Chang, E. R. Chapman, and M. B. Jackson. 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.
137.
Krammer, A., H. Lu, B. Isralewitz, K. Schulten, and V. Vogel. 1999. Forced
unfolding of the fibronectin type III module reveals a tensile molecular
recognition switch. Proc Natl Acad Sci U S A 96:1351-1356.
138.
Rosenheck, K. 1998. Evaluation of the electrostatic field strength at the site of
exocytosis in adrenal chromaffin cells. Biophys J 75:1237-1243.
139.
Luitel, P., D. F. Schroeter, and J. W. Powell. 2007. Self-electroporation as a
model for fusion pore formation. J Biomol Struct Dyn 24:495-503.
140.
Katz, B., and R. Miledi. 1967. The release of acetylcholine from nerve endings by
graded electrical pulses. Proceedings of the Royal Society of London. Series B,
Biological Sciences 167:23-38.
110
141.
Sugita, S., Y. Hata, and T. C. Sudhof. 1996. Distinct Ca(2+)-dependent properties
of the first and second C2-domains of synaptotagmin I. J Biol Chem 271:12621265.
142.
Chapman, E. R., S. An, J. M. Edwardson, and R. Jahn. 1996. A novel function for
the second C2 domain of synaptotagmin. Ca2+-triggered dimerization. J Biol
Chem 271:5844-5849.
143.
Damer, C. K., and C. E. Creutz. 1996. Calcium-dependent self-association of
synaptotagmin I. J Neurochem 67:1661-1668.
144.
Desai, R. C., B. Vyas, C. A. Earles, J. T. Littleton, J. A. Kowalchyck, T. F.
Martin, and E. R. Chapman. 2000. The C2B domain of synaptotagmin is a
Ca(2+)-sensing module essential for exocytosis. J Cell Biol 150:1125-1136.
145.
Fukuda, M., H. Kabayama, and K. Mikoshiba. 2000. Drosophila AD3 mutation of
synaptotagmin impairs calcium-dependent self-oligomerization activity. FEBS
Lett 482:269-272.
146.
Osborne, S. L., J. Herreros, P. I. Bastiaens, and G. Schiavo. 1999. Calciumdependent oligomerization of synaptotagmins I and II. Synaptotagmins I and II
are localized on the same synaptic vesicle and heterodimerize in the presence of
calcium. J Biol Chem 274:59-66.
147.
DiAntonio, A., and T. L. Schwarz. 1994. The effect on synaptic physiology of
synaptotagmin mutations in Drosophila. Neuron 12:909-920.
148.
Davis, A. F., J. Bai, D. Fasshauer, M. J. Wolowick, J. L. Lewis, and E. R.
Chapman. 1999. Kinetics of synaptotagmin responses to Ca2+ and assembly with
the core SNARE complex onto membranes. Neuron 24:363-376.
149.
Littleton, J. T., M. Stern, M. Perin, and H. J. Bellen. 1994. Calcium dependence
of neurotransmitter release and rate of spontaneous vesicle fusions are altered in
Drosophila synaptotagmin mutants. Proc Natl Acad Sci U S A 91:10888-10892.
150.
Fukuda, M., T. Kojima, and K. Mikoshiba. 1996. Phospholipid composition
dependence of Ca2+-dependent phospholipid binding to the C2A domain of
synaptotagmin IV. J Biol Chem 271:8430-8434.
151.
Borden, C. R., C. F. Stevens, J. M. Sullivan, and Y. Zhu. 2005. Synaptotagmin
mutants Y311N and K326/327A alter the calcium dependence of
neurotransmission. Mol Cell Neurosci 29:462-470.
152.
Fukuda, M., T. Kojima, J. Aruga, M. Niinobe, and K. Mikoshiba. 1995.
Functional diversity of C2 domains of synaptotagmin family. Mutational analysis
of inositol high polyphosphate binding domain. J Biol Chem 270:26523-26527.
111
153.
Rickman, C., J. L. Jimenez, M. E. Graham, D. A. Archer, M. Soloviev, R. D.
Burgoyne, and B. Davletov. 2006. Conserved prefusion protein assembly in
regulated exocytosis. Mol Biol Cell 17:283-294.
154.
Deng, H., G. Chen, W. Yang, and J. J. Yang. 2006. Predicting calcium-binding
sites in proteins - a graph theory and geometry approach. Proteins 64:34-42.
155.
Guan, K. L., and J. E. Dixon. 1991. Eukaryotic proteins expressed in Escherichia
coli: an improved thrombin cleavage and purification procedure of fusion proteins
with glutathione S-transferase. Anal Biochem 192:262-267.
156.
Gonzalez, A., P. Moorhead, S. E. McPhillips, J. Song, K. Sharp, J. R. Taylor, P.
D. Adams, N. K. Sauter, and S. M. Soltis. 2008. Web-Ice: integrated data
collection and analysis for macromolecular crystallography. Journal of Applied
Crystallography 41:176-184.
157.
Adams, P. D., R. W. Grosse-Kunstleve, L. W. Hung, T. R. Ioerger, A. J. McCoy,
N. W. Moriarty, R. J. Read, J. C. Sacchettini, N. K. Sauter, and T. C. Terwilliger.
2002. PHENIX: building new software for automated crystallographic structure
determination. Acta Crystallogr D Biol Crystallogr 58:1948-1954.
158.
McCoy, A. J. 2007. Solving structures of protein complexes by molecular
replacement with Phaser. Acta Crystallogr D Biol Crystallogr 63:32-41.
112
Fuson, Kerry L
Principal Investigator/Program Director (Last, First, Middle):
BIOGRAPHICAL SKETCH
NAME
POSITION TITLE
Kerry L. Fuson
Graduate Assistant
EDUCATION/TRAINING (Begin with baccalaureate or other initial professional education, such as nursing, and include postdoctoral training.)
DEGREE
(if applicable)
YEAR(s)
Univ. of Texas of the Permian Basin, Odessa, TX
Texas State Univ. at San Marcos, San Marcos, TX
B.Sc.
M.S.
1995-1997
2000-2002
Biology
Microbiology
Univ. of Texas Medical Branch at Galveston,
Galveston, TX
Ph.D
2003-2009
Biochemistry
INSTITUTION AND LOCATION
FIELD OF STUDY
A. Positions.
1990 – 1994 United States Army. Cavalry Scout, Attained rank of Specialist
Promotable (E-4P).
1997 – 1998 7th Grade Science Teacher. Presidio Independent School District,
Presidio,
TX.
1998 – 2000 Graduate Student, Sul Ross State University and Texas A&M University,
Corpus Christi, TX. (Transferred to Texas State University).
2000 – 2002 Graduate Teaching Assistant, Texas State University
2002 – 2003 Research Associate I, Department of Physiology and Biophysics,
University of Texas Medical Branch, Galveston, TX.
2002 – 2003 Adjunct Instructor of Microbiology and Pathology. San Jacinto
Community College, Houston, TX.
2004 – 2009 Graduate Student. Department of Human Biological Chemistry &
Genetics (HBC&G), University of Texas Medical Branch, Galveston, TX.
2009 – Present Postdoctoral Researcher, Texas Tech University, Lubbock, TX.
B.
Honors and Professional Membership.
The Texas Brach American Society for Microbiology Orville Wyss Award for
Outstanding Scientific Achievement (March 2001).
George H. Myer Award in Microbiology for Excellence in Microbiology (April 2001).
Pre-doctoral fellowship from the Keck Houston Area Molecular Biophysics Program
(HAMBP), (2006-2009).
Member of the Biophysical Society
1
2007 - Poster - The mechanical properties of human synaptotagmin 1 C2 domains
(Won team poster competition)
Invited Speaker – Feb. 15 2008 – Keck Seminar: Keck Annual Research Conference
Poster Winners, Part 2. Title: The mechanical properties of human synaptotagmin 1 C2
domains.
Invited Speaker – 13th Annual Structural Biology Symposium May 16-17, 2008.
Invited Speaker – Center for Membrane Protein Research, Texas Tech Health Science
Center. January 23, 2009
C. Courses
1997 Summer Research Internship in Molecular Biology, University of Texas, Austin,
TX.
2001 Molecular Biology and Protein Expression, University of Texas Medical Branch,
Galveston, TX.
April 23-27, 2007 - RapiData 2007 Tutorial Course in X-ray crystallography:
Brookhaven National Laboratory, Long Island, NY
D. Selected peer-reviewed publications (in chronological order).
Fuson, K.L., Ma, L., Sutton, R.B., Oberhauser A.F., The C2 Domains of Synaptotagmin
1 Have Distinct Mechanical Properties. Biophys J. 2009 Feb;96(3):1083-90.
Fuson, K.L., Montes, M., Robert, J. J., Sutton R. B., Structure of Human Synaptotagmin
1 C2A-C2B in the Absence of Ca2+ Reveals a Novel Domain Association, Biochemistry.
2007 Nov 13;46(45);13041-8.
Montes M, Fuson KL, Sutton RB, Robert JJ. Purification, crystallization and X-ray
diffraction analysis of human synaptotagmin 1 C2A-C2B. Acta Cryst. F 62, Page 926-9,
Aug 2006
Chada S, Sutton RB, Ekmekcioglu S, Ellerhorst J, Mumm JB, Leitner WW, Yang HY,
Sahin AA, Hunt KK, Fuson KL, Poìndexter N, Roth JA, Ramesh R, Grimm EA,
Mhashilkar AM. MDA-7/IL-24 is a unique cytokine--tumor suppressor in the IL-10
family. Int Immunopharmacol. 2004 May;4(5):649-67. Review.
2