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The Role of STXBP5, VAMP8, and SNAP23 in Endothelial Exocytosis
by
Qiuyu Zhu
Submitted in Partial Fulfillment of the
Requirements for the Degree
Doctor of Philosophy
Supervised by Professor Charles J. Lowenstein
Department of Pharmacology and Physiology
School of Medicine and Dentistry
University of Rochester
Rochester, New York
2015
ii
Biographical Sketch
The author was born in Tengzhou, Shandong Province, People’s Republic of China. He
attended Xi’an Jiaotong University in 2003. He completed the Seven-Year Program of
Clinical Medicine (equivalent to MD) with his internship and residency in Internal
Medicine, and he graduated with a BS/MS dual degree in Clinical Medicine in 2010. He
began his doctoral studies in Physiology at the University of Rochester in 2010. He
earned the Master of Science in Physiology in 2012. He was awarded the Howard
Hughes Medical Institute "Med-into-Grad" Fellowship in Cardiovascular Science from
2010 to 2012 and received cardiovascular-oriented clinical and translational research
training. He was awarded an American Heart Association (AHA) Predoctoral Fellowship
from 2013 to 2015. He received the 2014 Travel Award for Young Investigators from the
AHA’s Council on Arteriosclerosis, Thrombosis, and Vascular Biology (ATVB), as well
as the Travel Funds for PhD Students awarded by the University Dean of Graduate
Studies in 2014, and he presented part of this thesis work at the Sol Sherry Distinguished
Lecture in Thrombosis at AHA’s Scientific Sessions 2014. He pursued his research in
endothelial exocytosis and thrombosis under the direction of Charles J Lowenstein, MD.
The following publications are a result of work conducted during this doctoral study:
1. Nadtochiy SM, Zhu Q, Urciuoli W, Rafikov R, Black SM, and Brookes PS.
Nitroalkenes confer acute cardioprotection via adenine nucleotide translocase 1. The
Journal of Biological Chemistry. 2012;287(5):3573-80.
iii
2. Zhu Q, Yamakuchi M, Ture S, de la Luz Garcia-Hernandez M, Ko KA, Modjeski
KL, LoMonaco MB, Johnson AD, O'Donnell CJ, Takai Y, Morrell CN, and
Lowenstein CJ. Syntaxin-binding protein STXBP5 inhibits endothelial exocytosis and
promotes platelet secretion. The Journal of Clinical Investigation.
2014;124(10):4503-16.
3. Zhu Q, Bao C, Ture S, Ferlito M, Morrell CN, Yamakuchi M, Lowenstein CJ.
VAMP8 Mediates Endothelial Granule Exocytosis. Blood. (in revision for
publication)
4. Zhu Q, Yamakuchi M, Lowenstein CJ. SNAP23 Regulates Endothelial Exocytosis.
PLOS One (in review for publication)
Published Abstracts:
1. Zhu Q, Yamakuchi M, Ture S, de la Luz Garcia-Hernandez M, Ko KA, Modjeski
KL, LoMonaco MB, Johnson AD, O'Donnell CJ, Takai Y, Morrell CN, and
Lowenstein CJ. STXBP5 Regulates Endothelial Exocytosis, Platelet Secretion and
Thrombosis. American Heart Association 2014 Scientific Sessions
2. Zhu Q, Bao C, Ture S, Ferlito M, Morrell CN, Yamakuchi M, and Lowenstein CJ.
VAMP8 Mediates Endothelial Granule Exocytosis. Arteriosclerosis, Thrombosis, and
Vascular Biology 2014 Scientific Sessions
iv
Acknowledgments
I would like to thank my mentor Dr. Charles J Lowenstein. During the past five years, he
has not only been an inspiring role model, but also more importantly, as an original
thinker who imparted wisdom and unlimited intellectual stimulation to my doctoral study.
His mentoring has prepared me well to move forward in my career, which I appreciate
more than I can express. His word and deed will continue to illuminate and encourage my
future pursuit towards the great scientific unknown.
I have benefited tremendously from the invaluable guidance and support from my thesis
committee members Drs. Robert Dirksen, Ingrid Sarelius, and Craig Morrell, as well as
our collaborators Drs. Christopher O’Donnell (NIH, NHLBI, and Harvard Medical
School), Andrew Johnson (NIH and NHLBI), and Sidney Whiteheart (University of
Kentucky). I want to thank Professor James Palis for chairing my thesis defense
committee.
I want to thank all the Lowenstein lab members, previous and current, for making my
personal ecosystem equally productive and fun. This includes an enormous debt of
gratitude to Dr. Munekazu Yamakuchi, who systematically taught me the methodology in
vascular biology, and imbued me with an exceptional standard of scientific rigor and
work ethic. Dr. Maria de la Luz Garcia-Hernandez offered substantial help to my project,
as well as constant support as a bench-side mentor. Michael LoMonaco gave me valuable
technical insights and made sure that everything “under the hood” always runs smoothly.
v
Previous lab members Drs. Kenji Matsushita, Shusuke Yagi, Takashi Ito, Amit
Dhamoon, and Vijay Vanchinathan, showed me by example how scientists would study,
think, and work.
I have been blessed to work with the most extraordinary and dynamic colleagues in the
Morrell lab, who enriched my life in science and contributed to the body of this work. I
want to extend my thanks to perceptive consultant Dr. Craig N. Morrell, who guided me
through many critical steps of my study. Sara Ture assisted in technically challenging in
vivo experiments, and her exemplary work taught me many lessons in physiology. I want
to thank Dr. Scott Cameron, Dr. Angela Aggrey, David Field, and Lesley Chapman for
offering valuable suggestions and support, and for sharing frustrations and successes in
research. Thank you, Kristina Modjeski, for your advice and friendship.
I am indebted to the Department of Pharmacology and Physiology, and to the Aab CVRI
for the environments I have been privileged to work in, and for offering me many great
exemplars and teachers. This includes Dr. Joseph Miano, for his advice, guidance, and
stimulating discussions. This also includes Dr. Burns Blaxall and Dr. Paul Brookes with
whom I have apprenticed, and who encouraged my passions in science.
Thanks also to the CVRI microsurgical core and Dr. Kyung Ae Ko, whose
meticulousness and exceptional surgical skills helped address many challenging
questions.
vi
I want to thanks Linda Lipani and Sandra Morgan for their administrative effort and
dedication.
Last but not least, I want to thank all my friends and my family for their unending support
to my scientific aspirations. Most specially, I want to thank my wife, soul-mate and
intellectual partner, Cindy Yingya Zhou, for her devotion, inspiration, and selfless love.
vii
Abstract
Thromboembolic diseases are major causes of morbidity and mortality. Endothelial
exocytosis plays a critical role in thrombosis. The exocytic machinery that regulates
granule secretion from endothelial cells (EC) is not completely defined. We hypothesized
that specific members of the soluble N-ethylmaleimide sensitive factor attachment
protein receptor (SNARE) superfamily and their regulatory partners mediate endothelial
granule exocytosis and thrombosis.
EC exocytosis releases von Willebrand factor (VWF), a major factor in thrombogenesis.
Recent genome-wide association studies (GWAS) identified syntaxin-binding protein 5
(STXBP5) as a candidate gene linked to changes in plasma VWF levels. We found
STXBP5 is expressed in human ECs and interacts with SNARE proteins. STXBP5
inhibits endothelial exocytosis in vitro. Stxbp5 KO mice had higher plasma VWF levels
and more P-selectin translocation than wild-type mice, suggesting STXBP5 inhibits
endothelial exocytosis in vivo. However, Stxbp5 KO mice also displayed impaired
hemostasis and thrombosis. Platelets from Stxbp5 KO mice had defects in secretion and
activation. Therefore, STXBP5 is a novel SNARE regulatory partner that inhibits
endothelial exocytosis, but promotes platelet secretion and thrombosis.
In secretory cells, a ternary complex that mediates exocytosis is comprised of three
SNARE molecules, including isoforms of the syntaxin (STX), vesicle-associated
membrane protein (VAMP), and synaptosomal-associated protein (SNAP) families.
viii
Human ECs express proteins of the SNARE superfamily, but the exact identity of EC
SNAREs that regulate endothelial exocytosis has been unclear. We explored the SNARE
molecules required for endothelial exocytosis. We found VAMP8 co-localized with
Weibel-Palade bodies, the major endothelial granules. VAMP8 interacted with
components of the exocytic machinery. Knock down of VAMP8 expression inhibited
endothelial exocytosis. Vamp8 KO mice had decreased endothelial exocytosis. These data
suggest that VAMP8 plays a critical role in endothelial exocytosis.
In addition, we identified SNAP23 as the predominant endothelial SNAP isoform that
mediates endothelial exocytosis. SNAP23 was localized to the plasma membrane.
Knockdown of SNAP23 decreased endothelial exocytosis. SNAP23 also interacted with
the endothelial exocytic machinery. These data suggest that SNAP23 is another key
component of the endothelial SNARE machinery.
Taken together, the present work identified the role of STXBP5, VAMP8, and SNAP23
in EC exocytosis. Given the importance of EC exocytosis to vascular thrombosis and
inflammation, the insights gained from these studies may lead to novel targets for the
controlling of thromboembolic diseases.
ix
Contributors and Funding Sources
This work was supported by a dissertation committee consisting of Professor Charles J.
Lowenstein (advisor) of the Department of Medicine, Professors Robert T. Dirksen and
Ingrid H. Sarelius of the Department of Pharmacology and Physiology, and Professor
Craig N. Morrell of the Department of Medicine, Aab Cardiovascular Research Institute.
The defense committee was chaired by Professor James Palis of the Department of
Pediatrics, Hematology and Oncology.
All work for the dissertation was completed by the author except for:
Chapter 2: GWAS data was analyzed in collaboration with Drs. Andrew D. Johnson and
Christopher J. O’Donnell (NHLBI and NHLBI’s Framingham Heart Study, Framingham,
MA). The initial expression and RNAi studies in HUVEC was performed by Dr.
Munekazu Yamakuchi (Figure 2-1A and Figure 2-2A-C). Mammalian expression
plasmid pMEX neo-3×FLAG-tomosyn (8)-ATG was provided by Dr. David James
(Garvan Institute of Medical Research, Darlinghurst, NSW 2010, Australia), and FLAG-
STXBP5-N436S was designed and generated by Dr. Munekazu Yamakuchi. XS-VWF
protein was a generous gift of Dr. Sriram Neelamegham (Department of Chemical and
Biological Engineering, State University of New York at Buffalo, Buffalo, NY). Drs.
Yoshimi Takai (Department of Biochemistry and Molecular Biology, Kobe University
Graduate School of Medicine, Kobe, Japan) and Jun Miyoshi (Osaka Medical Center for
Cancer and Cardiovascular Disease, Osaka University, Osaka, Japan) generated and
x
provided the Stxbp5 KO mice. Bone-marrow transplantation was performed under the
direction of Dr. Maria de la Luz Garcia-Hernandez. Dr. Kyung Ae Ko performed FeCl3-
induced carotid thrombosis. Kristina L. Modjeski conducted analysis of ex vivo platelet
studies.
Chapter 3: Clare Bao performed in vitro studies depicted in Figure 3-2 A-C, Figure 3-3
B, Figure 3-4 B, D-E, and Figure 3-5 right panel.
In addition, the in vivo mesenteric thrombosis model, platelet rolling, leukocyte rolling,
and microsphere rolling experiments were all performed by Sara Ture. Michael B.
LoMonaco provided substantial technical assistance and conducted some of the VWF
exocytosis assays.
This work was supported by NIH/NHLBI R21 HL108372, R01 HL074061, R01
HL78635 (to Charles J. Lowenstein), by the HHMI "Med-into-Grad" Fellowship in
Cardiovascular Sciences at the Aab Cardiovascular Research Institute, University of
Rochester (to Charles J. Lowenstein and Qiuyu Zhu), and by American Heart Association
grants 0835446N (to Munekazu Yamakuchi) and 13PRE17050105 (to Qiuyu Zhu).
xi
Table of Contents
Biographical Sketch ............................................................................................................ ii
Acknowledgments.............................................................................................................. iv
Abstract ............................................................................................................................. vii
Contributors and Funding Sources..................................................................................... ix
Table of Contents ............................................................................................................... xi
List of Tables .................................................................................................................... xv
List of Figures .................................................................................................................. xvi
List of Schemes ................................................................................................................ xix
List of Abbreviations ........................................................................................................ xx
Introduction ....................................................................................................... 1
1.1 Venous thromboembolism: morbidity and mortality ............................................... 2
1.2 VTE pathophysiology: the good, the bad, and the ugly ........................................... 6
1.2.1 Endothelial cells: the good ............................................................................ 6
1.2.2 Platelets: the bad ........................................................................................... 8
1.2.3 Hypercoagulation, stasis, and beyond: the ugly.......................................... 11
1.3 Exocytosis and the SNARE hypothesis ................................................................. 15
1.3.2 Components of SNAREs ............................................................................ 17
xii
1.3.3 SNARE regulators ...................................................................................... 23
1.4 Endothelial exocytosis ........................................................................................... 28
1.4.1 Endothelial granules.................................................................................... 28
1.4.2 Major compounds released by endothelial exocytosis................................ 31
1.4.3 Types of VWF release ................................................................................ 39
1.4.4 Regulators of endothelial exocytosis .......................................................... 42
1.5 Thesis overview ..................................................................................................... 44
Dual Role of STXBP5 in Endothelial Exocytosis, Thrombosis, and Platelet
Secretion ........................................................................................................................... 46
2.1 Introduction ............................................................................................................ 47
2.1.1 Novel genetic variants are associated with human VWF levels ................. 47
2.1.2 STXBP5: A novel regulator of exocytosis.................................................. 47
2.1.3 STXBP5 SNP and plasma VWF .................................................................. 49
2.2 Results .................................................................................................................... 54
2.2.1 STXBP5 is expressed in human ECs and murine tissues ........................... 54
2.2.2 STXBP5 inhibits endothelial exocytosis in vitro ........................................ 55
2.2.3 Genetic variation in STXBP5 affects VWF exocytosis in vitro ................. 56
2.2.4 STXBP5 does not co-localize with endothelial WPBs ............................... 57
2.2.5 STXBP5 interacts with endothelial exocytic machinery ............................ 57
xiii
2.2.6 STXBP5 inhibits endothelial exocytosis in vivo ........................................ 60
2.2.7 STXBP5 affects hemostasis and thrombosis .............................................. 62
2.2.8 STXBP5 promotes platelet secretion and activation .................................. 63
2.3 Discussion .............................................................................................................. 96
VAMP8 Mediates Endothelial Exocytosis .................................................... 105
3.1 Introduction .......................................................................................................... 106
3.2 Results .................................................................................................................. 108
3.2.1 Endothelial cells express a distinct set of SNAREs that mediate granule
exocytosis ................................................................................................................ 108
3.2.2 VAMP8 and VAMP3 co-localize with VWF ........................................... 108
3.2.3 VAMP8 and VAMP3 interact with the SNAREs STX4 and SNAP23 .... 109
3.2.4 VAMP8 and VAMP3 mediates endothelial exocytosis in vitro ............... 110
3.2.5 VAMP8 mediates endothelial exocytosis in vivo ..................................... 111
3.3 Discussion ............................................................................................................ 126
SNAP23 Regulates Endothelial Exocytosis .................................................. 131
4.1 Introduction .......................................................................................................... 132
4.2 Results .................................................................................................................. 133
4.2.1 SNAP23 is expressed in human ECs and murine tissues ......................... 133
4.2.2 SNAP23 regulates endothelial exocytosis ................................................ 133
xiv
4.2.3 SNAP23 is primarily localized on cell membrane in EC ......................... 134
4.2.4 SNAP23 interacts with endothelial exocytic machinery .......................... 135
4.3 Discussion ............................................................................................................ 145
Conclusions and Perspectives ....................................................................... 149
5.1 Summary .............................................................................................................. 150
5.2 Working model for endothelial exocytosis .......................................................... 152
5.3 Future directions .................................................................................................. 156
5.3.1 STXBP5 SNP and VWF ............................................................................ 156
5.3.2 STXBP5 in ECs and in platelets ............................................................... 158
5.4 Significance and perspectives .............................................................................. 161
Materials and Methods .................................................................................. 163
References ....................................................................................................................... 181
xv
List of Tables
Table 1 Description of STXBP5 SNPs associated with plasma VWF levels. ................... 51
Table 2 List of antibodies ............................................................................................... 177
Table 3 Primers used for SYBR Green RT-qPCR .......................................................... 180
xvi
List of Figures
Figure 1-1 Model of VWF-platelet interaction in vascular injury. ................................... 38
Figure 2-1 STXBP5 expression and transcript variants. ................................................... 65
Figure 2-2 STXBP5 inhibits endothelial exocytosis in vitro. ........................................... 67
Figure 2-3 STXBP5 and constitutive release of VWF...................................................... 69
Figure 2-4 STXBP5 does not affect HUVEC content of VWF. ....................................... 70
Figure 2-5 Knockdown of STXBP5 does not affect the number of WPB granules per cell.
........................................................................................................................................... 71
Figure 2-6 Knockdown of STXBP5 does not affect the size of WPB granules. .............. 72
Figure 2-7 Knockdown of STXBP5 does not affect the shape of WPB granules. ........... 73
Figure 2-8 STXBP5 inhibits HUVEC release of P-selectin. ............................................ 75
Figure 2-9 STXBP5 SNP and release of VWF. ................................................................ 76
Figure 2-10 Particles containing STXBP5 are not co-localized with endothelial granules
containing VWF. ............................................................................................................... 77
xvii
Figure 2-11 STXBP5 co-sediments, co-localizes, and co-precipitates with STX4 and
SYT1. ................................................................................................................................ 78
Figure 2-12 STXBP5 does not co-localize with markers for ER, Golgi, lysosomes, or
endosomes. ........................................................................................................................ 82
Figure 2-13 STXBP5 does not interact with NSF and Munc family members. ............... 83
Figure 2-14 STXBP5 does not co-localize with SNAP23, VAMP8, or VAMP3. ........... 84
Figure 2-15 STXBP5 does not co-localize with Caveolin-1, Munc18-3, Munc18-2, or
NSF. .................................................................................................................................. 86
Figure 2-16 STXBP5 does not co-localize with Rab27, Myosin5a or MyRIP. ................ 87
Figure 2-17 Stxbp5 affects plasma VWF in mice. ............................................................ 88
Figure 2-18 Stxbp5 inhibits endothelial exocytosis in mice. ............................................ 89
Figure 2-19 Stxbp5 increases thrombosis in mice. ........................................................... 92
Figure 2-20 Stxbp5 regulates platelet secretion and activation. ....................................... 93
Figure 2-21 Stxbp5 in platelets increases thrombosis in mice. ......................................... 95
Figure 3-1 Expression of SNAREs in endothelial cells. ................................................. 113
xviii
Figure 3-2 Subcellular localization of VWF and SNAREs in endothelial cells. ............ 115
Figure 3-3 Interaction of SNAREs in endothelial cells. ................................................. 116
Figure 3-4 VAMP8 and VAMP3 mediate endothelial exocytosis.................................. 118
Figure 3-5 Knockdown of VAMP3 or VAMP8 does not affect endothelial granule
number or VWF content. ................................................................................................ 120
Figure 3-6 Knockdown of VAMP3 or VAMP8 does not affect WPB morphology. ...... 122
Figure 3-7 Vamp8 mediates endothelial exocytosis and leukocyte rolling in vivo. ....... 124
Figure 4-1 SNAP23 is expressed in human endothelial cells and murine tissues. ......... 138
Figure 4-2 SNAP23 is important for endothelial exocytosis. ......................................... 140
Figure 4-3 Subcellular localization of SNAP23 in endothelial cells. ............................. 142
Figure 4-4 SNAP23 interacts with endothelial exocytic machinery. .............................. 144
xix
List of Schemes
Scheme 1-1 A common pathophysiology of VTE and atherothrombosis. ......................... 5
Scheme 1-2 Virchow's triad. ............................................................................................. 14
Scheme 1-3 Topologic model of neuronal SNARE complex. .......................................... 20
Scheme 1-4 Transition states of SNARE complex and membrane fusion. ...................... 21
Scheme 2-1 Schemes of STXBP5 and Lgl family member structures. ............................ 52
Scheme 2-2 STXBP5 differentially influences exocytosis in platelets and endothelial
cells. ................................................................................................................................ 104
Scheme 5-1 Working model of endothelial exocytosis regulation. ................................ 155
xx
List of Abbreviations
ADAMTS13 ADAM metallopeptidase with thrombospondin type 1 motif, 13
ADP adenosine diphosphate
ANOVA analysis of variance
APC activated protein C
AT antithrombin
ATP adenosine triphosphate
ATPase triphosphatase
cAMP 3'-5'-cyclic adenosine monophosphate
Cas9 CRISPR-associated 9
CHARGE Cohorts for Heart and Aging Research in Genomic Epidemiology
CRISPR clustered regularly interspaced short palindromic repeats
DVT deep vein thrombosis
EC endothelial cell
EGF epidermal growth factor
ENCODE the Encyclopedia of DNA Elements
ER endoplasmic reticulum
ET-1 endothelin-1
GABA gamma-aminobutyric acid
GMP-140 granule membrane protein 140
GP glycoprotein
GPCR G-protein coupled receptors
xxi
GST glutathione S-transferase
GWAS genome-wide association studies
HAEC human aortic endothelial cell
HBMEC human brain microvascular endothelial cell
HCT hematocrit
HDMVEC human dermal microvascular endothelial cell
HUS hemolytic uremic syndrome
HUVEC human umbilical vein endothelial cell
ICAM1 intercellular adhesion molecule 1
IL-8 interleukin-8
LD linkage disequilibrium
LFA-1 lymphocyte function-associated antigen-1
Lgl lethal giant larvae
Mac-1 macrophage-1 antigen
MAF minor allele frequency
MFI median fluorescence intensity
MP micro particle
NEM N-ethylmaleimide
NETs neutrophil extracellular traps
NO nitric oxide
NSF N-ethylmaleimide-sensitive factor
PADGEM platelet activation-dependent granule to external membrane protein
PAF platelet activating factor
xxii
PAI plasminogen activator inhibitor
PDGF platelet-derived growth factor
PE pulmonary embolism
PF4 platelet factor 4
PGI2 prostacyclin
PKA protein kinase A
PKC protein kinase C
proVWF VWF pro-polypeptide
PSGL-1 P-selectin glycoprotein ligand-1
PTS post-thrombotic syndrome
ROS reactive oxygen species
SDS sodium dodecyl sulfate
SM protein Sec1/Munc18 protein
SNAP (1) soluble NSF attachment protein
(2) synaptosomal-associated protein
SNAP23 synaptosomal-associated protein, 23-KD
SNAP25 synaptosomal-associated protein, 25-KD
SNARE soluble N-ethylmaleimide sensitive factor attachment protein receptor
SNP single nucleotide polymorphism
SNV single nucleotide variant
STX syntaxin
STXBP5 syntaxin-binding protein 5
SYT synaptotagmin
xxiii
TALEN transcription activator-like effector nuclease
TF tissue factor (thromboplastin)
TFPI tissue factor pathway inhibitor
TGF-β1 transforming growth factor β1
t-PA tissue plasminogen activator
t-SNARE target membrane-SNARE
TTP thrombotic thrombocytopenic purpura
TXA2 thromboxane A2
VAMP vesicle-associated membrane protein
VCAM1 vascular cell adhesion molecule 1
VEGF vascular endothelial growth factor
VLD VAMP-like domain
VSMC vascular smooth muscle cell
v-SNARE vesicle-SNARE
VTE venous thromboembolism
VWD von Willebrand disease
VWF von Willebrand factor
WPB Weibel-Palade body
ZFN zinc finger nuclease
1
Introduction
2
1.1 Venous thromboembolism: morbidity and mortality
Venous thromboembolism (VTE) is a disease caused by the formation of thombi in deep
veins, including deep vein thrombosis (DVT) and pulmonary embolism (PE).
VTE is a major cause of morbidity (1, 2). The annual incidence of VTE is 0.1% - 0.27%,
affecting 300,000-600,000 individuals in the United States each year; 5% of the general
population will have at least one occurrence of VTE in their lifetime (3-5). Typical
symptoms of DVT include pain, swelling, and ulcer of extremities, which severely limit
patients’ physical activity. Severe forms of DVT, such as phlegmasia cerulea dolens,
result in massive thrombotic occlusion of deep veins, ischemia and gangrene, which can
be limb-threatening. PE is caused by the dislodgement of a venous clot that travels to
and clogs vasculature in the lung. It is estimated that 20% of patients with PE die on the
first day of occurrence (6). Debilitating long-term complications of VTE which harm
patients' quality of life include post-thrombotic syndrome (PTS) in 23 - 60% of patients
following DVT (7), and pulmonary hypertension in 1% - 4% of PE patients (8, 9).
The mortality from VTE is high. Before the implementation of anticoagulant therapy,
VTE was often fatal (up to 30% mortality in the first month) (10). VTE is the second
leading cause of death in cancer patients (11). VTE is the third leading cardiovascular
disease after acute coronary syndrome and stroke (12). Autopsy data shows PE is the
third most common cause of death in hospitalized patients in the United States (13), with
nearly one quarter of cases presenting as sudden death (5). However, because of the
3
difficulty in prediction and detection, VTE is under diagnosed, and the true prevalence is
probably even higher.
Anticoagulant therapy has enormously reduced the morbidity and mortality from VTE.
However, current treatment for VTE remains inadequate. Anticoagulant therapy increases
the risk of bleeding which has now emerged as a cause of death in about 25% of VTE
patients (14, 15). Anticoagulant therapy is also associated with other side effects
including thrombocytopenia, osteoporosis (heparin), and skin necrosis (coumarin) (16).
Moreover, the long-term prognosis of VTE remains dismal despite continued
anticoagulant therapy, primarily due to the high recurrence rate both during and after
anticoagulation therapy (approximately 7% at 6 months, and up to 40% in the first 10
years) (17, 18), which often necessitates indefinite anticoagulant treatment.
Risk factors for VTE are divided into acquired and hereditary causes. Well-established
risk factors include aging, immobilization, pregnancy, cancer, surgery or trauma, obesity,
diabetes mellitus, hypertension, hyperlipidemia, cigarette smoking, oral contraceptives,
and previous VTE (5, 19). In addition, genetic predisposition has been linked to VTE
(20). VTE appears more frequently in black and white populations than in East Asians or
Hispanics. Inherited rick factors include factor V Leiden, prothrombin G20210A
mutation, protein C or S deficiency, activated protein C (APC) resistance, antithrombin
(AT) deficiency, plasma von Willebrand factor (VWF), and elevated levels coagulation
factors VIII, IX, or XI (5, 19, 21).
4
Recently, a new paradigm has been proposed to consider VTE as part of a “pan-
cardiovascular syndrome” that comprised of coronary artery diseases, cerebral vascular
diseases, and peripheral vascular diseases (22). Studies show patients with VTE have
higher risk of arterial thrombosis than those without VTE (23-25). VTE shares a common
pathophysiology with atherothrombosis: both are predisposed by common risk factors,
and share mechanistic pathways in common such as endothelial injury, inflammation, and
hypercoagulability (Scheme 1-1).
Appreciation of VTE pathophysiology may eventually furnish new therapeutic targets.
Identifying the risk factors for VTE and unravelling the details of its molecular pathways
not only provides a mechanistic framework for the prevention and treatment VTE, but
also helps our understanding of the scourge of arterial thrombotic diseases, such as
coronary artery diseases and stroke, which are of growing worldwide importance.
5
Scheme 1-1 A common pathophysiology of VTE and atherothrombosis.
Adapted from Piazza et al. Circulation. 2010;121(19):2146-50 (22).
6
1.2 VTE pathophysiology: the good, the bad, and the ugly
The pathophysiology of VTE is a multifactorial process that includes interactions
between endothelial dysfunction and platelet activation and aggregation, which together
set up a cascade of pro-thrombotic events (1).
1.2.1 Endothelial cells: the good
Endothelial cells (ECs) line the inner surface of vessel wall, maintaining the integrity of
the vasculature. An adult contains about 1×1013 ECs forming a nearly 1-kilogram “organ”
(26). Under normal conditions, EC preserves vascular metabolic homeostasis and patency
through multiple mechanisms:
1) Barrier function. EC acts as an antithombotic blood-tissue barrier separating the
endothelial matrix from blood and prevents the activation of coagulation (1, 27).
Potent pro-thrombotic proteins such as tissue factor (TF; thromboplastin) and
collagen in the subendothelial matrix initiate physiologic hemostasis when the
endothelial barrier is breached (28, 29).
2) Synthesis and release of anti-thrombotic factors. EC inhibit thrombosis by
producing nitric oxide (NO) (30, 31), prostacyclin (PGI2) (32), CD39 (33), and other
anti-thrombotic factors (34-36). EC inhibits the generation and activation of thrombin
by expressing tissue factor pathway inhibitor (TFPI) (37, 38). When pathologic
insults injure the vessel wall, EC decreases production of anti-thrombotic factors and
7
increases production and release of pro-thrombotic factors such as VWF,
thromboxane A2 (TXA2), and expands binding sites for coagulation factors which
facilitate thrombus formation (34). The loss of balance between anti-thrombotic
activity and pro-thrombotic activity characterizes EC dysfunction (36, 39), portending
an elevated risk of cardiovascular events such as acute coronary syndrome and
sudden death (40-42).
3) Regulation of hemodynamics. EC regulates vascular tone by balancing the
production of vasoconstrictors and vasodilators. EC produces vasoconstrictors such as
endothelin-1 (ET-1), the most potent known vasoconstrictor, and platelet activating
factor (PAF). EC also produces labile vasodilators such as NO (30, 43-46) and PGI2
(34, 47).
4) Fibrinolysis. EC promotes fibrinolysis and maintains blood fluidity by a surface-
connected fibrinolytic system. EC produces and secretes tissue plasminogen activator
(t-PA) (48) and its receptors (49), and plasminogen activator inhibitors (PAIs) (50).
5) Regulation of vascular inflammation. EC produce adhesion molecules and
cytokines that participate and regulate the inflammatory process (51, 52).
Cardiovascular risk factors and clinical syndromes including aging, obesity, smoking,
hypertension, and hypercholesterolemia, are partly related to endothelial dysfunction
(53). Conversely, interventions that reduce cardiovascular risk factors improve
endothelial function and restore vascular homeostasis. Therefore, endothelium has been
proposed as a “barometer” for cardiovascular risk (54). Pathological insults to the
8
vasculature disrupt the anti-thrombotic function of EC and set up the stage for a sequence
of pro-thrombotic events discussed below.
1.2.2 Platelets: the bad
Platelets are the second most abundant cells in human blood (150 - 400 × 109/L) after red
blood cells. They are formed by an elaborate membranous system in their precursor
megakaryocytes (the demarcation membrane system), and shed as cytoplasmic fragments
from megakaryocytes. Therefore, platelets are anucleate and have long been considered
as fragments of cells. Advances in electron microscopy technology have revealed the
elaborate internal structure of platelets. Platelets are essential for physiological
hemostasis and thrombosis. In addition, they also mediate angiogenesis, wound repair,
cancer, and infection (55-58). Roles for platelets as important immune and inflammatory
cells have recently been recognized (59-64).
Circulating, quiescent platelets normally do not adhere to the vessel wall because of the
antithrombotic blood-tissue interface maintained by ECs. However, circulating platelets
rapidly respond to vascular damage (e.g. hemorrhage or ruptured arterial plaque) by
forming platelet plugs (thrombus). Defects of platelet number (thrombocytopenia) or
function can lead to a severe bleeding diathesis. The function of platelets in mediating
hemostasis and thrombosis depends on multiple properties of platelets:
9
1) Platelet surface receptors. Platelet glycoprotein Ib (GPIb) constitutes the GPIb-V-
IX membrane receptor complex that binds to VWF, which mediates the first step in
platelet adhesion to injured vasculature. Platelet GPIb and GPVI binds to collagen in
the sub-endothelial matrix. Collagen binding to GPIb or GPVI activates a cascade of
signals inside a platelet that include elevations in intracellular Ca2+ levels, arachidonic
acid metabolism by cyclooxygenase isoforms, and release of TXA2 (65-67). Platelet
GPIIb/IIIa (also known as integrin αIIbβ3) is a platelet receptor for VWF, fibrinogen
and fibronectin (68-70). GPIIb/IIIa on the surface of a resting platelet does not
interact with ligands, but activation of the platelet leads to a conformational change in
GPIIb/IIIa enabling it to bind to VWF and fibronectin.
2) Platelet metabolites. Platelets contain enzymes that metabolize arachidonic acid into
vasoactive substances including TXA2 that promotes thrombosis.
3) Platelet granules. Platelets have three major types of granules containing over 300
vasoactive substances that modulate hemostasis, thrombosis, and vascular
inflammation (71, 72):
a. α granules, the most abundant granules, contain peptides and proteins,
including pro-coagulation proteins such as factor V, VWF, fibronectin and
fibrinogen, growth factors such as platelet-derived growth factor (PDGF),
epidermal growth factor (EGF), chemokines such as platelet factor 4 (PF4)
and transforming growth factor β1 (TGF-β1), and adhesion molecules such as
P-selectin. These proteins constitute the bulk of the platelet secretome.
10
b. δ granules or dense granules contain small molecules such as polyphosphates,
serotonin, and calcium, which are important signaling molecule and potent
platelet-activators.
c. Lysosomes contain proteolytic enzymes such as β-hexosaminidase, cathepsin
D and E, and other enzymes. There are only a few lysosomes per platelet.
Lysosomes are considered important for clot- and wound-remodeling (73, 74).
How do platelets mediate thrombus formation? This function is carried out by a complex
process that transforms quiescent platelets into a platelet plug. This process is artificially
separated into three stages: adhesion, activation, and aggregation. First, injury to the
vessel wall exposes the sub-endothelial matrix. Injured endothelium release VWF;
platelet surface receptors recognize VWF and sub-endothelial collagen: the GPIb-V-IX
complex binds to VWF; and the GPVI receptor binds to collagen. Receptor binding not
only anchors platelets to the injury site, but also triggers a cascade of intraplatelet signals
that are essential to activation and aggregation (65-67). Second, platelet activation occurs
right after platelet receptor binding to VWF, collagen, or TF. Ligand interaction with
these platelet G-protein coupled receptors (GPCR) activates a cascade of intracellular
signals including an increase in intracellular Ca2+ which in turn leads to an array of
downstream effects: GPIIb/IIIa activation, granule secretion, and platelet spreading.
GPIIb/IIIa activation initiates outside-in and inside-out signaling and increases
production of pro-thrombotic factors such as TXA2; secretion of granule contents
enhances local platelet aggregation and attracts more circulating platelets to the injury
site; and morphological changes generate forces to plug and contract the wound. Finally,
11
platelets aggregate with other platelets as a result of GPIIb/IIIa activation. Activated
platelets also interact with coagulation factors to propagate the coagulation cascade,
which generates thrombin and fibrin. These three stages facilitate and overlap with each
other. The result is hemostasis and thrombosis.
1.2.3 Hypercoagulation, stasis, and beyond: the ugly
Virchow identified a triad of factors that contribute to VTE: abnormalities in blood flow,
injury of the vessel wall, and blood hypercoagulability (Scheme 1-2) (75, 76). Many
thromboembolic diseases involve one or more factors of this triad. Regardless of the
dispute about its origin (77), Virchow's triad integrates the basic roles of three
coordinated components in thrombosis:
1) Endothelial injury lays the foundation for platelet adhesion and coagulation.
Endothelial cells can be activated by numerous stimuli into a pro-thrombotic and pro-
inflammatory phenotype. EC release a plethora of pro-thrombotic substances by
exocytosis to trigger platelet adhesion and activation. The exposed sub-endothelial
matrix anchors VWF and platelets, while exposed TF activates the coagulation
cascade. Activated ECs synthesize and release PAF in large quantities, which triggers
inflammation and thrombosis. P-selectin recruits leukocytes that participate in
inflammation by producing reactive oxygen species (ROS) and proteases, further
boosting thrombosis (78).
12
2) Static flow induced by immobility, compression or heart failure increases the
thrombotic risk. Normal laminar flow maintains EC morphology and function and
diminish local VWF, thrombin and fibrin to prevent platelet adhesion and thrombosis.
Stasis increases the risk of local pro-thrombotic factor accumulation and fibrin
formation and polymerization (79).
3) Blood hypercoagulability can result from the antiphospholipid syndrome, cancer and
oral contraceptives. High levels of circulating pro-thrombotic factors, such as
circulating ECs, VWF, thrombomodulin, and D-dimers promote blood cell aggregates
(78). Increase of blood coaguability is also seen when there is a higher than normal
hematocrit (HCT) or erythrocytes (polycythemia vera) (80, 81), higher than normal
number of leukocytes (81), higher than normal number of platelets (thrombocytosis),
and their inappropriate activation (80, 82, 83), which all increase the risk of clotting.
Recently studies have shown thrombus formation can be triggered by discoid platelets
on undisrupted ECs indirectly through local production of ROS, adding yet another
mechanism by which elevated blood hypercoagulability (thrombocytosis) can
increase thrombosis risk (84).
However, Virchow’s triad is not the complete picture of blood clot formation. The
mechanisms responsible for VTE are manifold. In reality, in a great proportion of
patients, factors from Virchow’s concept are not detectable. Additional elements that are
important to thrombosis have been identified, including ABO blood groups, fibrinolysis,
neutrophil extracellular traps (NETs) (85), and infection (86) to name just a few. For
instance, procoagulant circulating micro particles (MPs), membrane fragments shed by
13
blood cells and the vasculature, are newly regarded as a risk factor for thrombosis (87,
88). Activated platelets and ECs are regarded as the main sources of MPs. TF-positive
MPs may explain the increased rates of VTE in cancer patients (87).
14
Scheme 1-2 Virchow's triad.
15
1.3 Exocytosis and the SNARE hypothesis
Exocytosis is the first response shared by many cells in the thrombogenic pathway. EC
responds to injury by exocytosis. Platelet activation depends on granule secretion (74).
Activated leukocytes release pro-inflammatory enzymes by exocytosis (89). Enhanced
understanding of the mechanisms that regulate exocytosis is critical for improving the
prevention and treatment of thromboembolic diseases.
Exocytosis is the regulated release of granule contents by intracellular vesicle fusion with
the plasma membrane. Exocytosis is an intracellular protein transport mechanism that
exists in all eukaryotic cells, and is conserved from yeast to human (90, 91). Protein
trafficking maintains the internal structure of a cell, and allows it to communicate with
adjacent cells and its environment. Endothelial cells, leukocytes, and platelets all
participate in the pathology of VTE by exocytosis.
By 1970, the seminal work by George Palade (1974 Nobel laureate) and others had
already made it evident that secretory proteins are transported by a series of vesicles from
the endoplasmic reticulum (ER) to the Golgi apparatus to the cell surface where they are
released to the extracellular space (92). The fusion of vesicles with a target membrane
was obviously done with extremely high specificity, to ensure that the correct cargos is
always delivered to its correct destination. However, many questions remain. One such
question is: How does each type of vesicle fuse with the right target membrane at the
right time?
16
Satisfactory answers to this question were not available until the last two decades. In the
1970s, the prevailing belief held that the anatomical arrangement of the endomembrane
systems dictated the trafficking specificity of cargos. One example is the close proximity
between the ER and the Golgi, where vesicles budded from ER appear to be force-fed
into the Golgi, but not into other intracellular organelles. Thus, it was the anatomical
proximity of membrane compartments that appeared vital to vesicle targeting specificity.
However, Rothman et al. proposed a simple hypothesis that specific membrane fusion
was governed by the intrinsic chemical specificity of membranes, not by the intricate
patterns of anatomy. Rothman and colleagues proved that membrane transport can be
reconstituted in a cell-free membrane-fusion assay (93); targeted, but not non-specific
membrane fusion, was faithfully replicated using a minimum set of proteins (94-96).
Using this assay, it was discovered that the membrane fusion machinery consists of three
components later to be called SNAREs (soluble N-ethylmaleimide-sensitive factor (NSF)
attachment protein receptors) proteins (97).
Based on this evidence, Rothman and others proposed the SNARE hypothesis to explain
vesicle fusion at the right place and the right time: SNARE proteins are vesicle and target
membrane markers; specific membrane fusion is mediated by SNARE proteins on vesicle
and target membranes; and distinct SNAREs on vesicles and distinct SNAREs on target
membranes determined the membrane fusion specificity (97). The SNARE hypothesis
was supported by many studies. One strong piece of evidence is that the neurotoxins
tetanus toxin or botulinus toxin block exocytosis by specifically degrading SNARE
proteins (98-100).
17
The work by James E. Rothman, Randy W. Schekman and Thomas C. Südhof jointly
won them the 2013 Nobel Prize in Physiology or Medicine “for their discoveries of
machinery regulating vesicle traffic, a major transport system in our cells” (101). Their
work provided a unifying conceptual framework for vesicle transport and membrane
fusion that integrated many proteins into an evolutionarily-conserved molecular machine.
SNAREs has been confirmed as a universal machinery that mediates membrane fusion in
a variety of cell types (102-113).
1.3.2 Components of SNAREs
SNAREs are membrane proteins. The SNARE hypothesis proposes that SNARE
machinery consists of three components: a SNARE on the vesicle (v-SNARE) binds
specifically to two cognate SNAREs on the target membrane (t-SNAREs), forming a
SNARE complex. The specificity of a transport vesicle for its target membrane is
mediated by the specific interaction between one unique v-SNARE and two unique t-
SNAREs. Much of our understanding of the SNARE machinery comes from presynaptic
neurons (114, 115), the v-SNARE on presynaptic vesicles is vesicle-associated
membrane protein 2 (VAMP2), whereas the t-SNAREs on presynaptic membrane are
synaptosomal-associated protein, 25-KD (SNAP25), and syntaxin 1 (STX1).
The crystal structure of a SNARE complex reveals that each SNARE protein possess at
least one α-helix. The SNARE complex is formed by the parallel arrangement of these α-
helices into a highly twisted four-helix bundle (116) (Scheme 1-3). The center of this
18
four-helix coiled-coil contains conserved leucine-zipper-like layers, with hydrophobic
residues buried inside. These hydrophobic residues consist of one arginine (R) and three
glutamine (Q) residues, contributed from each of the four helices. Therefore, SNARE
motifs used to be classified as R-SNAREs (from v-SNAREs) and Qa-, Qb-, Qc-SNAREs
(form t-SNAREs). These residues are highly conserved and sensitive to mutations (117-
120).
Vesicle fusion is mediated by the combined action of many four-helix SNARE
complexes (Scheme 1-4). Unfolded SNAREs can spontaneously assemble into four-helix
trans-SNARE complex (SNAREpin), a process called “priming”, which brings vesicles
into a “docked” position (121). The assembly is thermodynamically favorable, resistant
to sodium dodecyl sulfate (SDS) (122) and heat (123) treatment. Multiple SNAREpins
aggregate at the vesicle-target membrane interface, exerting an inward force that pulls
membrane into close proximity; yet spontaneous fusion is prevented by “clamping”
proteins such as complexin. Upon Ca2+ stimulation, the “clamps” are removed by specific
Ca2+ sensors. The energy from multiple SNAREpins eventually overwhelms the
resistance from the membranes and drives the membrane to fuse (124, 125). SNAREpins
drive fusion cooperatively; upon membrane fusion they transforms into a low-energy,
fully zippered cis-SNARE (126).
Disassembly of the SNARE complex is important for the vesicle transport cycle. Initial
experiments with cell-free membrane assays by Glick et al found that membrane
transport was blocked by N-ethylmaleimide (NEM), a sulfhydryl alkylating reagent
19
(127). NEM treatment resulted in accumulation of fused vesicles, suggesting some
regulator sensitive to NEM is required to recycle the membrane fusion machinery (128).
This soluble, N-ethylmaleimide-sensitive protein was soon purified based on its ability to
restore membrane transport following NEM treatment and was named “N-
ethylmaleimide-sensitive factor” (NSF) (128). NSF is a well conserved cytosolic
triphosphatase (ATPase). NSF hydrolysis of adenosine triphosphate (ATP) is required for
SNARE disassembly and recycling. NSF binds to soluble NSF attachment protein
(SNAP, different from synapsomal-associated protein) which in turn interacts with the
SNARE complex (129). SNARE proteins were named because they interact with SNAP
(thus SNARE stands for soluble NSF attachment protein receptors). SNARE, SNAP, and
NSF form a complex on the cell membrane which sediments at 20S (97). Unfolding of
substrate SNARE proteins is coupled to NSF hydrolysis of ATP via a “spring-loaded”
mechanism (130-132).
20
Scheme 1-3 Topologic model of neuronal SNARE complex.
Shown is the four-helix coiled-coil bundle contributed by v-SNARE VAMP2 (blue), and
t-SNAREs syntaxin 1 (red) and SNAP25 (green). Transmembrane domains of syntaxin 1
and SNAP25 are shown in yellow. SNAP25 has no transmembrane domain; instead,
SNAP25 is anchored to the target membrane by a loose unstructured loop connecting its
two helices.
Adapted from Sutton et al. Nature. 1998;395(6700):347-53(116).
21
Scheme 1-4 Transition states of SNARE complex and membrane fusion.
(A) Tans-SNARE complex. One helix on the vesicle membrane bundles with three
cognate helices on the target membrane to form a fourth-helix trans-SNARE complex.
This assembled fourth-helix bundle “zippers” towards its transmembrane anchors, but is
not completely zippered, generating an inward force that brings the vesicle membrane
into close proximity to the target membrane.
(B) Cis-SNARE complex. Eventually the inward force pulled membranes to fuse, and the
SNAREs are in parallel alignment, a low-energy state called a cis-SNARE complex.
22
(C) Membrane transition in SNARE-mediated fusion, including (a) separated
membranes, (b) fusion stalk formation, (c) membrane bilayer contact, and (d) fusion pore.
Adapted from Sudhof et al. Science. 2009;323(5913):474-7 (133) and Jahn et al. Annu
Rev Biochem. 1999;68(863-911) (134).
23
1.3.3 SNARE regulators
The SNARE machinery ensures specificity of membrane transport: the correct cargo is
delivered to the correct location. Membrane fusion specificity is encoded in SNARE
proteins per se as an intrinsic physico-chemical property (135), which can be faithfully
reconstituted using isolated SNARE proteins and liposomes (136). However, it has been
well-documented that vesicles inside cells traffic and fuse in a highly efficient and
choreographed manner, with both temporal and spatial precision. How was it achieved?
Cell-free liposome fusion assay demonstrates SNARE proteins alone cannot achieve the
same degree of speed and temporal precision in membrane fusion (137, 138). Despite its
simplicity, SNARE machinery requires additional regulators, and its controlling
mechanism proved to be a rather complicated system and only began to be appreciated.
In neurons and other cell types, several SNARE regulatory proteins have been identified.
1.3.3.2 Synaptotagmin and complexin
The SNARE machinery relies on two families of proteins, synaptotagmin (SYT) and its
cofactor complexin, as “triggers” to achieve calcium-dependent time precision (139).
Synaptotagmins are a membrane protein family characterized by an N-terminal
transmembrane region, a variable central linker, and two C-terminal C2 domains (C2A
and C2B) which are autonomous Ca2+-binding modules (140). The best characterized
member, SYT1, is a synaptic vesicle Ca2+-binding protein. Ca2+ binding to C2 domains of
24
SYT1 enhances its affinity to membrane phospholipids and SNAREs and triggers fast
vesicle release. SYT1 is required and only required for Ca2+-triggered fast
neurotransmitter release. Knock-out of SYT1 in mice impairs Ca2+-triggered neurovesicle
release. SYTs are believed to be universal Ca2+-sensors for triggered-exocytosis in many
different forms, and are evolutionarily conserved. Mammals express 16 synaptotagmins.
Different SYT homologs are expressed in different cell types, with different affinities for
Ca2+, which may explain the varied Ca2+-sensing kinetics in different vesicle transport
pathways (141-146).
However, SYTs do not act alone. Complexin is an essential co-factor of SYT1.
Complexin is a soluble protein originally discovered as a partner for trans-SNARE
complex and was subsequently named “complexin” (147). Similar to SNARE proteins,
complexin contains an α-helix that fits into the groove formed by the four-helix bundle in
the SNARE complex (148). Complexin is also evolutionarily conserved, but its role is
paradoxical. On the one hand, complexin inhibits SNARE-mediated fusion, as evidenced
in vitro and ex vivo (149-152). On the other hand, complexin knockout neurons also
exhibit similar phenotype as SYT1-null cells: loss of rapid and synchronous exocytosis,
suggesting it is required for Ca2+-triggered release (153). In addition, complexin null
mice exhibit motor and cognitive function deficits similar to those seen in Huntington's
disease, suggesting it is required for normal neurological function (154).
SYT and complexin act in concert to trigger Ca2+-induced synaptic exocytosis (139) and
both are essential for this process. But the detailed mechanism has not been completely
25
elucidated. The current hypothesis is built on the crystal structure of the complexin-
SNARE complex (148). The major part of complexin is an α-helix, which competes with
synaptotagmin for binding to the same site at assembled SNARE complexes: a unique
groove in the four-helix bundle formed by syntaxin and VAMP. Complexin “clamping”
of the SNARE complex reinforced vesicle docking and poised them for simultaneous,
Ca2+-triggered synchronous release. Ca2+ binding to synaptotagmin displaces the
clamping of complexin and frees SNARE complex, allowing assembled SNARE
complex to zipper freely. Thus complexin clamps the SNARE complex before
membrane fusion occurs, and synaptotagmin releases the complexin clamp and permits
membrane fusion to occur. This model, however, does not explain why complexin is
counter-intuitively required for Ca2+-induced exocytosis and how knockout of complexin
abolishes Ca2+-triggered synchronous release (139). Novel mechanisms have been
proposed to complement this evolving model (155).
1.3.3.3 Munc18 and Munc13
High efficiency of SNARE-mediated fusion in cells is likely conferred by Munc18 and
Munc13 proteins that prepare the core SNARE engine for assembly.
Munc18 is a member of the Sec1/Munc18 (SM) proteins that are conserved from yeast to
mammals. Unc18 (C. elegans) and Sec1 (yeast) are absolutely essential for C. elegans
movements and yeast secretion (156). Similarly, Munc18 is essential for synaptic vesicle
release in mammals. The first-reported and best characterized mammalian Munc18
26
member, Munc18-1 (also called STXBP1, Munc18a or neuronal Sec1) is a neural-
specific protein. Munc18-1 deficiency in cells abolishes synaptic membrane fusion (157).
Knockout of Munc18-1 alone in mice results in complete loss of neurotransmitter
secretion, rendering the brain “synaptically silent” (158), a phenotype that even exceeds
the knockout of a SNARE protein (159); in humans, Munc18-1 mutations cause early
infantile epileptic encephalopathy (160) called “STXBP1 encephalopathy” (161-164).
These results suggest Munc18-1 is not a by-stander but an essential component and
central actor in the membrane fusion machinery. Expression of Munc18 homologs has
been detected in various tissues, supporting it as a common component of vesicle fusion
machinary in many cell types (165, 166).
Exactly how Munc18-1 participates in membrane fusion is still unclear (133). Munc18-1
may promotes vesicle fusion by cooperating with SNAREs. In a reconstituted lipid
bilayer system, Munc18-1 strongly accelerated the fusion reaction through direct contact
with both v- and t-SNAREs (167). Crystal structure reveals Munc18-1 forms a complex
with STX1A in neurons (168). Munc18-1 guides syntaxin as a cheparone towards
productive SNARE complex formation, thus facilitates vesicle docking (157, 169).
Munc18-1 may also facilitate the topological arrangements of SNAREpins, clasp
SNAREpins circumferentially at the fusion interface, prevent their diffusion. These
functions collectively enhance the efficiency of SNARE-mediated fusion (133, 170).
Munc13 is a presynaptic member of the CATCHR protein family (members involved in
vesicle tethering) and is homologous to Unc13 of C. elegans. Munc13 is required for
27
certain types of synaptic transmission (171, 172). Glutamatergic neurons from Munc13-1
null mice show arrest of synaptic-vesicle cycles and loss of transmitter release in
response to stimulation (172). Munc13 function together with Munc18-1 to cooperate the
assembly of SNARE complexes (173, 174). Munc13 may release syntaxin from the clasp
of Munc18-1, accelerating its transition from the closed syntaxin-Munc18 complex to the
SNARE complex (173). Munc13 lowers the energy barrier for synaptic vesicle fusion
(175). Moreover, NSF disassembly of the SNARE complex requires Munc18-1 and
Munc13 (174). Interestingly, unlike its absolutely-required participation in glutamatergic
synaptic secretion (171), Munc13 is not essential for dense-core vesicle release, but only
controls dense-core vesicle localization and release efficiency (176). Similarly, synaptic
release from some GABA (gamma-aminobutyric acid)-releasing neurons are not affected
by depletion of Munc13-1 (172). Therefore, Munc13 orchestrates membrane fusion with
the SNAREs complex in a transmitter- or cell-specific manner.
28
1.4 Endothelial exocytosis
EC maintain the integrity of the vasculature. Due to its privileged location at the interface
between blood and organs, EC constantly senses changes in the blood and in the
subendothelial matrix, and actively modulates its interaction with other cells (including
blood cells, vascular smooth muscle cells and pericytes) or the blood flow. In response to
injury, EC undergo exocytosis, releasing numerous hemostatic and inflammatory
mediators into the blood steam that mediate interaction with platelets and leukocytes
(177-179).
1.4.1 Endothelial granules
In 1964, Ewald Weibel and George Palade discovered a remarkable cytoplasmic
component in rat and human arterial endothelia by electron microscopy (180). They
described the prominent morphologic features of this new organelle:
“A hitherto unknown rod-shaped cytoplasmic component which consists of a bundle of
fine tubules, enveloped by a tightly fitted membrane, was regularly found in endothelial
cells of small arteries in various organs in rat and man. It is about 0. 1 μ thick, measures
up to 3 μ in length, and contains several small tubules, ~ 150 A thick, embedded in a
dense matrix, and disposed parallel to the long axis of the rod.” (Weibel ER, and Palade
GE. J Cell Biol. 1964;23(101-112))
29
Despite the discrepancy in their distribution, these organelles were ubiquitously found in
the endothelia:
“… cytoplasmic components were found with great regularity in endothelial cells of
small branches of the pulmonary artery in rats. Much less frequently, rod-shaped bodies
were also found in the endothelium of alveolar capillaries.” (Weibel ER, and Palade GE.
J Cell Biol. 1964;23(101-112))
As Weibel and Palade concluded in their report in 1964, “the nature and significance of
these cytoplasmic components are yet unknown” (180). But they did notice that “the rod-
shaped body described in this article has been observed with regularity in numerous
vascular endothelia of the rat, man, and Amblystoma. This indicates that it must be a
structure of some functional significance which for the moment remains obscure” (180).
Thanks to the EC isolation and culture technique developed in the 1970s (181, 182), these
rod-shaped structure with numerous internal bundles were further studied in cultured ECs
(183). These granules, now termed Weibel-Palade bodies (WPBs), turned out to be a
specific marker of EC. In 1982, Wagner et al. found WPBs are storage organelles for
VWF (184). In 1989, P-selectin that mediates leukocytes rolling was found on the
membrane of WPBs (185, 186). With these and other components in WPB discovered,
one major structural basis for endothelium interactions with blood cells was gradually
delineated. WPBs essentially equip EC with a rapid means of response to stimulation and
vascular injury. A variety of stimuli, including hypoxia, physical trauma, or inflammatory
mediators, trigger EC to release the contents of WPBs (178).
30
WPBs are not the only type of endothelial granules that undergo regulated release. A few
other compounds have been reported to form specialized granules distinct from WPBs in
ECs, and undergo regulated release in response to stimuli. These compounds include
protein S, multimerin, t-PA (outside of WPBs), TFPI, and certain types of cytokines,
including CCL2 and CXCL1 (187-190). Some of them are important mediators of
thrombosis and inflammation, but the processing and release of these granules are poorly
understood. Perhaps these distinct populations of granules allow the same cell to respond
to diverse stimuli in a more controlled manner (190).
31
1.4.2 Major compounds released by endothelial exocytosis
Today, a growing list of components have been found to be present in WPBs,
participating in a broad range of biological processes including hemostasis, thrombosis,
inflammation, vasoconstriction and angiogenesis (191).
Perhaps the potent vasoactive properties of WPB contents determines that they must be
contained in such granules, and only released in a highly-regulated manner: exocytosis.
The compounds stored in WPBs make the granule “the perfect first aid kit after an insult
to the vasculature” (192).
1.4.2.1 Von Willebrand factor
VWF is synthesized exclusively by EC and by megakaryocytes (193, 194). Plasma VWF
is predominantly contributed by ECs (195).
VWF is the major and defining constituent of WPBs (184, 196). Expression of VWF
propolypeptide in a variety of nonendothelial cells results in rod-shaped granules that
closely resemble WPBs, suggesting VWF itself drives the formation of WPB structure
(197). The internal bundles in WPBs described by Weibel and Palade in 1964 were in
fact densely-packed VWF multimers (184). Once released into the blood stream by
exocytosis, these densely-packed VWF multimers are unfurled by local shear forces,
exposing its A1 domain which binds to platelet GPIb receptor, A3 domain which anchors
to collagen in the subendothelial matrix, and C1 domain which binds to activated platelet
32
GPIIbIIa. The unfurled VWF multimer can be cleaved by the protease called
ADAMTS13 (ADAM metallopeptidase with thrombospondin type 1 motif, 13) on its A2
domain (198), rendering smaller, non-thrombogenic VWF fragments that were
commonly found in the plasma (199, 200). The biosynthesis, secretion, and clearance of
VWF have been review recently in elegant detail (201).
The functional importance of VWF is demonstrated by the bleeding diathesis caused by
the quantitative or qualitative deficiency of VWF: von Willebrand disease (VWD). VWD
was first reported in 1926 by an internist from Finland, Erik von Willebrand (202) in
several members of a family. The proband, a 13-year-old girl, bled to death at her fourth
menstruation. Erik von Willebrand traveled to the girl’s hometown to study the disease in
depth. He mapped the family pedigree and found that 23 of the 66 family members had
bleeding problems. The pedigree and symptoms were both distinct from typical
hemophilia. Erik von Willebrand called the disease “hereditary pseudo hemophilia”. The
molecular basis of VWD was discovered by Southern blot in 1987 (203). This plasma
glycoprotein involved in the disease was then named for Dr. von Willebrand, together
with the bleeding disorder itself. VWD was found to be the most common inherited
bleeding disorder, with a prevalence of up to 1% (204).
VWF has two vital functions: it mediates platelet adhesion to the vessel wall during
primary hemostasis, and is the chaperone of factor VIII (205). Therefore, VWF is
essential for hemostasis and thrombosis (Figure 1-1). Recently, VWF was found to
participate in the formation of NETs (85). VWF binds to histone in vitro (206), and VWF
33
is associated with NETs in vivo (207, 208). These studies suggest VWF may have yet
undiscovered roles in immune response and thrombogenesis (86).
Plasma VWF levels are affected by both non-genetic and genetic factors. Non-genetic
determinants of plasma VWF include well-established risk factors for endothelial
dysfunction such as smoking and diabetes, as well as other less studied factors such as
aging (209, 210), exercise (211) and alcohol (212). Plasma VWF level is mainly
genetically determined. VWF levels vary with race and sex (213, 214), and its heritability
estimates from 0.31 to 0.75 (215, 216). Genetic defects are the major cause of
quantitative or qualitative VWF abnormalities in VWD individuals. While in severe
VWF patients, VWF gene mutations are common (217, 218), in milder cases the genetic
determinants are more variable and complex. Moreover, plasma VWF levels are highly
variable even in VWD individuals with identical VWF gene mutations (217), and their
clinical manifestation can be quite heterogeneous (217-219). Studies have shown that
plasma VWF level is subject to incomplete penetrance and variations in other genes. For
instance, plasma VWF level is influenced by the blood group gene ABO, with 25% lower
VWF level in O group than in non-O groups (220). The protein carring A, B and H blood
group antigens is also found on VWF A2 domain, close to the ADAMTS13 cleavage site.
The modification by ABO blood group antigen on VWF molecule directly influences its
cleavage by ADAMTS13, resulting in varied plasma VWF levels in different blood
groups (221, 222). Polymorphisms in CLEC4M gene affect the clearance of VWF, and
contribute to the variation in plasma VWF levels (223).
34
Plasma VWF levels are strongly correlated with thrombotic risk, especially VTE. For
patients with unusually large VWF multimers, as seen in hemolytic uremic syndrome
(HUS) and thrombotic thrombocytopenic purpura (TTP) resulted from ADAMTS13
mutation, there is an increased risk of platelet thrombi formation in small vessels where
shear rates is high (224-227). Increased VWF levels are strongly associated with risk for
coronary heart disease especially acute coronary syndrome (228-230). Clinical studies
show plasma level of VWF is significantly higher in VTE patients (231), and higher
VWF level independently increases VTE risk by up to 27% (232). Plasma VWF is also
frequently used as an indicator of endothelial dysfunction (233-235). Although VWF is
regarded as a novel therapeutic target for thromboembolic diseases (236, 237), regulation
of its release is still not well understood.
1.4.2.2 P-selectin
In 1824, Dutrochet first reported the adherence of white blood cells to the vessel wall and
their emigration to tissues (238). Classic works by Wagner, Cohnheim, Metchnikoff,
Addison, and the Clarks (75, 239-241) described the process in more details. In the early
20th century, Fahraeus (242) and Mogilnicki (243) provided quantitative account of
leukocyte rolling on the vessels. However, the mechanism and its significance were
largely unknown.
In 1972, Atherton and Born developed the mouse mesentery model (recapitulated and
refined in this thesis) to quantitatively investigate the adhesion of circulating leucocytes
35
to blood vessel walls in response to a variety of stimuli, and noticed that rolling
leucocytes were increased after the application of E. coli culture filtrate, suggesting the
phenomenon may implicate a defensive mechanism against infection (240). Atherton and
Born hypothesized that some “adhesive substance” existed on the interface between
leukocytes and the vessel wall.
It was not until 25 years later that Bonfanti and McEver found that P-selectin is a
component of endothelial WPBs, and Bevilacqua et al. found that P-selectin released
from WPBs was the “adhesive substance” that recruited leukocytes (244). In mice that
have defective WPB formation (VWF knockout), the recruitment of leukocytes onto the
blood vessel is essentially abolished, resulting in inflammatory response defects similar
to P-selectin knockout mice (245). P-selectin-mediated leukocyte rolling on EC is now
regarded as a hallmark of inflammation, the self-defense mechanism of the vessels
against harmful stimuli such as infection.
P-selectin (also called CD62P, platelet activation-dependent granule to external
membrane protein (PADGEM), or granule membrane protein 140 (GMP-140)) is a
selectin family membrane protein expressed in ECs (exclusively in WPBs) and in platelet
α granules. P-selectin rapidly externalizes onto the EC surface upon stimulation.
Leukocyte rolling along the vessel lumen is mediated by the interaction between P-
selectin and its ligand PSGL-1 (P-selectin glycoprotein ligand-1) expressed on most
leukocytes. P-selectin display on the EC surface is often transient (< 2min) and is readily
recycled into the cytoplasm and back to the WPBs (246). Therefore, the initial “rolling”
36
of leukocytes on P-selectin is labile and often transient. Firm adhesion and transmigration
of leukocytes involve additional adhesion molecules on ECs (e.g., intercellular adhesion
molecule 1(ICAM1) and vascular cell adhesion molecule 1 (VCAM1)), and β2 integrins
on leukocytes such as CD11a/CD18 (lymphocyte function-associated antigen-1, LFA-1)
and CD11b/CD18 (macrophage-1 antigen, Mac-1) (75).
1.4.2.3 Interleukin-8 (IL-8)
Activated ECs release IL-8, a chemotactic cytokine that stimulates neutrophil
phagocytosis and migration towards inflammatory sites (247). Expression and release of
IL-8 is contingent upon stimulation. Unstimulated EC does not store IL-8 in WPB,
whereas stimulation with IL-1β turns on its production and release from WPBs (247).
Intriguingly, IL-8 is present in the WPBs even after stimulation is removed, suggesting
storage of IL-8 in WPB may serve as the endothelial “memory” of inflammation,
enabling EC to rapidly respond to the next insult without de novo protein synthesis (248).
1.4.2.4 Other constituents in WPBs
Other important constituents in WPBs include tPA, a major initiator of fibrinolysis (249);
ET-1, a vasoconstrictor enabling EC to regulate hemodynamics (250-252); CD63 which
mediates cell-cell interaction (253); and angiopoietin-2, which sensitize ECs to
inflammation and induces angiogenesis (254, 255). Endothelial granules also contain
other pro-inflammatory and pro-thrombotic mediators that activate inflammation and
thrombosis in response to vascular injury (178, 191, 256). Together, release of these
37
compounds trigger a cascade of events that modulates hemostasis, thrombosis,
inflammation, hemodynamics, and angiogenesis (178, 257, 258) . Endothelial exocytosis
is thus a novel therapeutic target for thrombotic and inflammatory diseases (229, 230,
259).
38
Figure 1-1 Model of VWF-platelet interaction in vascular injury.
Adapted from Mannucci PM. New Engl J Med. 2004;351(7):683-94.(260)
39
1.4.3 Types of VWF release
Current studies suggest the major components of WPBs, VWF, is released from the ECs
via one of the following three pathways: unstimulated, WPB-dependent “basal” release;
unstimulated, WPB-independent “constitutive” release; or stimulated WPB exocytosis.
Co-existence of multiply secretion pathways in the same cell for the same cargo is not
uncommon. For instance, in neurons, in addition to action potential-evoked exocytosis,
two additional forms of neurotransmitter release exist: asynchronous release and
spontaneous “mini” release (261). Diversified vesicle release pathways enable cells to
more flexibly adjust to the changing environment with precise and optimized response.
In the 1990s, only two types of endothelial VWF secretion was proposed: constitutive
and stimulated secretion (262, 263). It was initially estimated that over 90% of VWF in
cultured EC would be secreted constitutively, with less than 10% secreted by the
regulated exocytosis (264). However, later studies revealed that high molecular weight
VWF (the majority of endothelial VWF content) is released almost exclusively by the
regulated pathway, and constitutive secretion of VWF (mostly immature VWF
fragments) was negligible (265). More recent studies suggest that in the absence of
stimuli, the majority of resting VWF release (approximate 80%) is from WPBs (termed
“basal secretion” to distinguish from “constitutive section”) (266); WPB-independent
“constitutive” release is only significant when the biogenesis machinery of WPB is
compromised, for example, by NH4Cl treatment or AP-1/clathrin coat depletion (266,
267). G protein-dependent signaling may differentially regulate basal and evoked VWF
40
secretion (268). Other mechanisms that regulate these different secretion pathways
remains to be discovered.
1.4.3.2 Basal release
The majority (80%) of unstimulated VWF release is though this route (266).
Study by Giblin et al. demonstrates in cultured ECs, most VWF is not sorted into the
constitutive secretory pathway, but into a post-Golgi compartment; part of it can be
subsequently secreted without stimulation. Since the only known post-Golgi secretory
organelle that contains VWF is WPBs, it was postulated that the majority of sorted VWF
ends in WPBs; WPBs then either spontaneously release their cargo, or undergo regulated
exocytosis in the presence of secretagogues (266). The mechanism that determines the
latter two different fates of WPBs is unknown. However, quantitative analysis suggests
although basal release accounts for 80% of unstimulated VWF release, the majority of
intracellular VWF content (high molecular multimers) is released by stimulated
exocytosis that far exceeds the amount by unstimulated VWF release (265).
WPBs that undergo basal secretion might differ from those secreted by exocytosis. These
WPBs are released shortly after they are formed, which are immature in their membrane
and VWF content (269, 270), whereas WPBs in exocytosis release almost all mature,
large multimers. It is postulated that the post-Golgi secretory organelles in basal secretion
are an immature form of WPBs (266).
41
1.4.3.3 Constitutive release
The hallmark of constitutive secretion is the lack of post-Golgi materials (263).
Consistent with the prediction, VWF released from this pathway is VWF pro-polypeptide
(proVWF), an unprocessed precursor of mature VWF. This pathway accounts for only an
insignificant amount of unstimulated VWF release (265), but becomes significant when
the machinery of basal release pathway or WPB biogenesis is impaired (266, 267).
1.4.3.4 Regulated exocytosis
Endothelial exocytosis can be triggered by a variety of stimulations, including physical
damage (hypoxia, radiation, stretch, temperature, and trauma), chemical stimulation
(ATP, adenosine diphosphate (ADP), epinephrine, histamine, serotonin, and
leukotrienes), peptides (thrombin, vascular endothelial growth factor (VEGF),
complements), and lipids (oxidized low-density lipoprotein, sphingosine-1 phosphate,
and ceramide) (178, 271, 272).
These agonists induce WPB exocytosis by increasing the intracellular concentration of
two different second messengers: Ca2+ or 3'-5'-cyclic adenosine monophosphate (cAMP)
(271). Most stimuli induce exocytosis through raising intracellular Ca2+ (including
thrombin and histamine). Calmodulin mediates this signaling pathway (273, 274). A few
secretagogues, such as serotonin, epinephrine and vasopressin, trigger endothelial
exocytosis by increasing cAMP, with protein kinase A (PKA) as the effector (275, 276).
Purine nucleotides stimulation of EC involves both Ca2+ and cAMP (277). It is
42
noteworthy that agonists effecting through these two different signaling pathway also
induce distinct changes on cytoskeleton remodeling and EC barrier function (278). Ca2+-
raising agonists diminish endothelial cell barrier function (279, 280), whereas cAMP-
raising agonists promote barrier function of EC (281-283).
1.4.4 Regulators of endothelial exocytosis
Much understanding of the SNARE mechanism in controlling exocytosis comes from
studies of yeast and neurons (131, 284-288). The exocytic machinery that drives vesicle
trafficking and membrane fusion in EC is similar to that found in neurons and yeast (90,
114, 134, 289, 290). V-SNARE molecules on endothelial granule surface interact with
specific t-SNAREs on the plasma membrane surface, forming a SNARE complex that
bridges the two membranes and directs vesicle fusion with the plasma membrane (291,
292).
Recent studies have defined some of the endothelial proteins that control the vesicle
secretory pathway in EC. Members of the SNARE family were found in human ECs,
including STX, VAMP, and SNAP homologs (102, 108, 109), although the exact identity
and function of endothelial SNAREs remains unclear. One exception is perhaps STX4,
which is required for endothelial exocytosis (108, 109).Various small GTPases and their
effectors such as Rab 3A, Rab3B, Rab3D, Rab11, Rab15, Rab27a, Rab27b, Rab 33a,
Rab37, RalA, and Rap1 play an important role in WPB maturation and exocytosis (102,
108, 178, 257, 293-305). MyRIP (Slac2c) serves as a bridge between Rab27a and myosin
43
Va/MyosinVIIa to control the location of endothelial granules (269, 296, 298, 305, 306).
Slp4a (granuphilin) binds to Rab8, Rab27a, Rab3A-D, and syntaxins, and promotes VWF
release (177, 296, 307). Munc13-4 was found in human EC and co-localizes with WPBs,
presumably acting as an effector of Rab27A (305). NO and thioredoxin regulate
exocytosis by chemically modifying NSF (108, 308). An assortment of secretagogues or
pathologic stimuli can stimulate granule secretion from human ECs through discrete
signaling pathways (190, 309-315). Members of the synaptotagmin family function as a
conserved sensor for Ca2+ and triggers regulated exocytosis in most cell types (139).
Unidentified factors relating to autophagy also affect exocytosis (316).
However, the detailed mechanisms controlling the endothelial SNARE machinery remain
partially unknown. Specifically, critical SNARE regulators in endothelial cell remains to
be identified, and the exact mechanism of how these regulators are coupled to the
SNARE machinery is to be characterized.
44
1.5 Thesis overview
Understanding regulation of exocytosis in ECs will address critical questions in the
pathogenesis of VTE. To elucidate the molecular mechanism of endothelial exocytosis,
the primary question is to identify and characterize the endothelial SNARE protein
members and their regulators. The thesis will approach this important question from the
following perspectives:
First, high-throughput platforms such as GWAS have discovered novel potential genetic
“hot spots” for VTE. These potential genetic determinants remain to be validated and
characterized, among which important regulatory molecules of endothelial exocytosis,
such as SNARE regulatory partners, may be identified and characterized.
Second, few endothelial SNARE candidates have been investigated in detail to date. The
identities of key endothelial SNAREs, especially VAMP homologs and SNAP homologs,
are unclear.
Therefore, we hypothesized that specific members of the SNARE superfamily and
their regulatory partners mediate endothelial exocytosis and thrombosis.
Chapter 2 harvests GWAS data to identify a novel protein STXBP5 that is associated
with human plasma VWF levels, and explores its role as a regulator of endothelial and
platelet exocytosis, as well as hemostasis and thrombosis.
45
Chapter 3 identifies the v-SNARE that mediates WPB release, and finds that VAMP8 is
an important endothelial v-SNARE.
Chapter 4 identifies the endothelial SNAP isoform that mediates WPB release: SNAP23.
The work of this thesis not only defined the role of the SNARE components and one
important SNARE regulator in endothelial exocytosis, but also expanded the potential
pathways to regulate endothelial function, hemostasis and thrombosis that will provide
novel perspectives for the prevention and treatment of VTE.
46
Dual Role of STXBP5 in Endothelial
Exocytosis, Thrombosis, and Platelet Secretion
47
2.1 Introduction
2.1.1 Novel genetic variants are associated with human VWF levels
The pathophysiology of VTE is complex, and much remains to be discovered about
cellular pathways that contribute to thrombosis. In Chapter 2, we used genetic
approaches to discover novel pathways that regulate thrombosis.
Recently, GWAS by the Cohorts for Heart and Aging Research in Genomic
Epidemiology (CHARGE) Consortium identified novel genetic variants that are
associated with altered plasma VWF levels in humans (317-319). These genetic loci
might indicate novel genes whose products regulate endothelial exocytosis. One genetic
locus with the highest significance is located within the gene that encodes syntaxin-
binding protein 5 (STXBP5). This association was strikingly reaffirmed by studies on
venous thrombosis in humans (320). The prominent relationship between human plasma
VWF alterations and genetic variation of STXBP5 prompted us to investigate the
potential role of STXBP5 in endothelial exocytosis.
2.1.2 STXBP5: A novel regulator of exocytosis
STXBP5 was first discovered by Fujita et al. as a protein interacting with syntaxin1A in
neurons and was named tomosyn (“tomo” means friend in Japanese; tomosyn means
“friend” of “syn”taxin) (321). It is enriched in rat brain and is also found in the heart,
48
spleen, lung, liver, skeletal muscle, kidney, and testis (321, 322). The gene encoding
STXBP5 (originally called tomosyn and later called tomosyn-1) is located on mouse
chromosome 10 and human chromosome 6. Murine Stxbp5 gene has at least 3 transcript
variants: Stxbp5-b, Stxbp5-m, and Stxbp5-s. Human STXBP5 gene has at least 5 predicted
transcript variants: STXBP5-1, STXBP5-2, STXBP5-X1, STXBP5-X2, and STXBP5–X3.
The gene Stxbp5L (originally called tomosyn-2) is located on mouse chromosome 16 and
human chromosome 3. Murine Stxbp5L has at least 4 transcript variants: Stxbp5L-xb,
Stxbp5L-b, Stxbp5L-m and Stxbp5L-s – all with distinct distribution patterns (322, 323).
STXBP5 belongs to the Lgl (lethal giant larvae) family which is conserved from yeast to
human. Lgl family members include Lgl, Sro7, Sro77, and STXBP5. Their common
feature are N-terminal WD40 repeats composed of β-propeller structures, which often
serve as scaffolds for protein-complex assembly (324)(Scheme 2-1). Lgl family members
have been involved in vesicle trafficking. Lgl, Sro7, and STXBP5 directly interact with
SNARE protein (321, 325-327).
All STXBP5 isoforms possess an N-terminal domain that contains WD40 repeats, a
variable tail domain, and a C-terminal domain that includes an R-SNARE-like motif
homologous to VAMP (VAMP-like domain, VLD) (322, 323, 328) (Scheme 2-1). In
neurons, VLD mediates the interaction of STXBP5 with syntaxin1A as a competitor to
VAMP2, and blocks formation of the heterotrimeric SNARE complex composed of
syntaxin1A, VAMP2, and SNAP25 (329, 330). STXBP5 inhibits neuron release of
neurotransmitters and endocrine cell secretion of insulin or other vesicles (331-337).
49
However, the role of STXBP5 in the vasculature has never been studied, and the relation
of STXBP5 with thrombosis has not been explored.
2.1.3 STXBP5 SNP and plasma VWF
The CHARGE Consortium identified genetic variations within the STXBP5 locus that
associate with plasma VWF, factor VII, and factor VIII levels (Table 1). The single
nucleotide polymorphism (SNP) with the highest genome-wide significance level (P =
6.9 × 10−22) for VWF plasma levels in the meta-analysis is rs9390459 (hg19
chr6:g.147680359G>A), a synonymous variation in the coding region for gene STXBP5
(317). Therefore, we attempted to prioritize nonsynonymous STXBP5 variants in high
linkage disequilibrium (LD) with this candidate SNP. The top non-synonymous variation,
rs1039084 (hg19 chr6:g.147635413A>G), encodes STXBP5 asparagine (N) to serine (S)
substitution at the 436 residue (hereinafter STXBP5-N436S), and is in high LD with
rs9390459 (R2 = 0.87, D’ = 0.97 from International HapMap Project; R2 = 0.93, D’ =
0.97 from 1000 Genomes Project, data from: http://www.broadinstitute.org/mpg/snap/)
(338). This N436S substitution encoded by rs1039084 is in one of the WD40 repeat
domains of STXBP5. The minor allele of rs1039084 is associated with decreased VWF
levels, higher bleeding score and decreased venous thrombosis in humans (319, 339,
340).
The CHARGE Consortium data yields two important pieces of information. First, a
cluster of SNPs within a single genetic loci identifies a candidate gene. For example,
50
many SNPs associated with altered VWF levels cluster within the STXBP5 gene.
Second, an individual SNP within a gene identifies a critical functional element within a
gene product. For example, rs1039084 alters STXBP5 residue 436 from Asn to Ser,
which suggests that this N436 is an important functional amino acid of STXBP5. Given
the association between STXBP5 genetic variations and plasma VWF levels, its structural
similarity with SNAREs, and its role in regulating neurotransmitter release, we
hypothesized that STXBP5 regulates endothelial cell exocytosis.
51
Table 1 Description of STXBP5 SNPs associated with plasma VWF levels.
refSNP Location
(hg19) Variant MAF1
Protein2
Position Residue change
rs9390459
chr6:147680359 G>A 0.4958 815 Leu > Leu
rs1039084
chr6:147635413 A>G 0.4571 436 Asp > Ser
1. MAF, minor allele frequency. Source: 1000 Genomes
2. RefSeq: NP_001121187.1
52
Scheme 2-1 Schemes of STXBP5 and Lgl family member structures.
(A) STXBP5 is composed of an N-terminal WD40 repeat domain, a C-terminal VAMP-
like domain, and a tail domain in between. The VAMP-like domain of STXBP5 enable it
to bind to the coiled-coil SNARE-domains such as one on syntaxin1.
(B) Structural resemblance of Lgl family proteins. The Lgl family proteins share
structural characteristics in N-terminal WD40 repeats composed of β-propellers. The tail
53
domain and the C-terminal VAMP-like domain are absent in Lgl. P indicates
phosphorylation sites.
Adapted from Yamamoto et al. Presynaptic Terminals. Springer Japan; 2015:129-40
(341).
54
2.2 Results
2.2.1 STXBP5 is expressed in human ECs and murine tissues
We first defined the expression of STXBP5 in human cells and in murine tissue. We
performed immunoblotting for STXBP5 on cultured human aortic endothelial cells
(HAEC), human umbilical vein endothelial cells (HUVEC), and human dermal
microvascular endothelial cells (HDMVEC). STXBP5 is expressed as a 130 kDa protein
in all three human endothelial cell types (Figure 2-1A) (321). Stxbp5 mRNA is expressed
in murine tissues, including lung, spleen, and aorta, as measured by qPCR (Figure 2-1B).
Stxbp5 mRNA is also found in murine brain, supporting studies identifying a role for
Stxbp5 in neurovesicle release (321, 330).
We next characterized the STXBP5 isoforms expressed by human tissues and cells. We
performed RT-PCR on RNA from HUVEC, human brain, human platelets, and HEK293
cells using primers flanking the splice regions. We identified 5 transcript variants of
STXBP5, all of which have been predicted but not previously characterized: STXBP5-1
(NM_139244.4), STXBP5-2 (NM_001127715.2), STXBP5-X1 (XR_245502.1), STXBP5-
X2 (XR_245503.1), and STXBP5-X3 (XR_245504.1) (Figure 2-1C). To quantitate the
relative abundance of each splice variant, we also performed qPCR. The STXBP5
transcript variant expressed at highest levels by endothelial cells is STXBP5-1, followed
by STXBP5-2, and other transcript variants are expressed at lower levels (Figure 2-1D).
The STXBP5 splice variant profile in endothelial cells resembles the profile in platelets
55
(Figure 2-1D). However, human brain expresses one predominant isoform, STXBP5-1,
and very little of the other variants were detected in brain (Figure 2-1D). Other studies
confirm that the same isoform STXBP5-1 is expressed in mammalian brain (321, 322).
2.2.2 STXBP5 inhibits endothelial exocytosis in vitro
We next explored the role of STXBP5 in endothelial exocytosis. We knocked down
expression of endogenous STXBP5 by RNA interference in HAEC, HUVEC, and
HDMVEC, stimulated the cells with the physiological agonist histamine, and then we
measured the amount of VWF released into the media using an ELISA. siRNA directed
against STXBP5 decreased expression of STXBP5 in HAEC, HUVEC, and HDMVEC
(Figure 2-2A-C, upper panel). Knockdown of STXBP5 expression does not affect the
constitutive secretion of VWF under resting conditions (Figure 2-2A-C, white bars).
However, knockdown of STXBP5 expression significantly increases histamine-induced
VWF release into media in all three different EC types (Figure 2-2A-C, black bars).
Similarly, knockdown of STXBP5 in HUVEC increases VWF release induced by ATP
(Figure 2-2D), and by calcium ionophore A23187 (Figure 2-2E), but did not change
VWF release at resting condition.
To further characterize the effect of STXBP5 on constitutive release of VWF, we
measured VWF levels in the media of HUVEC transfected with siControl of siSTXBP5
over an extended period of 12 hours, and found no change in constitutive release (Figure
2-3). STXBP5 does not affect total cellular VWF content (Figure 2-4).
56
We also tested the idea that STXBP5 affects VWF release by regulating granule number
and morphology. We counted the number of WPB granules in HUVEC transfected with
siControl or with siSTXBP5, and found that STXBP5 does not affect the number of
WPBs per cell (Figure 2-5). Analysis of the size of individual WPB revealed that
STXBP5 does not affect the size of these granules (Figure 2-6). Finally, we measured the
morphology of WPB by quantifying the aspect ratio of individual WPB granules:
STXBP5 does not affect the aspect ratio distribution of granules (Figure 2-7).
We also explored the effect of STXBP5 upon externalization of P-selectin, another
component of WPB translocated by endothelial exocytosis (186). Knockdown of
STXBP5 increases the endothelial externalization of P-selectin upon histamine
stimulation, as measured by HL-60 cell adherence (Figure 2-8).
Taken together, these results suggest that STXBP5 inhibits endothelial exocytosis.
2.2.3 Genetic variation in STXBP5 affects VWF exocytosis in vitro
We also tested the effect of genetic variation upon STXBP5 function. We knocked down
STXBP5 in endothelial cells and then rescued STXBP5 expression with either STXBP5
(WT) or STXBP5 (N436S), a variant of STXBP5 associated with altered VWF levels in
humans (319). STXBP5 (N436S) inhibits VWF secretion more effectively than wild-type
STXBP5 (Figure 2-9).
57
2.2.4 STXBP5 does not co-localize with endothelial WPBs
Since genetic variants in STXBP5 are associated with alterations in VWF levels, we next
defined the subcellular location of STXBP5 in relation to endothelial granules which
contain VWF as well as other pro-inflammatory and pro-thrombotic mediators. We found
that STXBP5 is located in the cytoplasm, primarily in a punctate pattern (Figure 2-10).
Immunostaining for VWF revealed the typical rod-shaped morphology of WPB (Figure
2-7 and Figure 2-10). Using confocal microscopy, we found that STXBP5 is minimally
co-localized with VWF (Figure 2-10). The morphology of the particles containing
STXBP5 is different from the WPB granules containing VWF, and the overlap is minor
(Figure 2-10). We searched for subcellular location of STXBP5: confocal microscopy did
not reveal a definitive co-location with markers for the ER, Golgi, lysosome, or
endosome compartments (Figure 2-12).
2.2.5 STXBP5 interacts with endothelial exocytic machinery
We next searched for links between STXBP5 and the exocytic machinery in endothelial
cells. Prior studies showed that STXBP5 can inhibit neurotransmitter release from
neurons and insulin release from pancreatic beta-cells by forming complexes with the
SNARE proteins, including syntaxin isoforms and SNAP isoforms (321, 342-344). We
and others have previously shown that endothelial SNARE molecules are likely STX4,
VAMP3, VAMP8, and SNAP23; some of them play an important role in regulating VWF
exocytosis (102, 108, 109, 345). Accordingly, we searched for an interaction between
58
STXBP5 and endothelial SNARE molecules. We first used sucrose density gradient
ultracentrifugation to specify the cellular components that co-sediment with STXBP5.
We loaded HUVEC lysates on top of a 5%-40% discontinuous sucrose density gradient,
performed ultracentrifugation, and analyzed fractions by immunoblotting. We found that
STXBP5 and STX4 partially co-sediment, suggesting these two proteins might be able to
form a complex in HUVEC, and might interact directly or indirectly with each other
(Figure 2-11A). However, STXBP5 does not co-sediment with others SNAREs that are
involved in endothelial exocytosis, such as SNAP23 (Figure 2-11A).
To further characterize the interaction between STXBP5 and STX4, we
immunoprecipitated HUVEC lysates with antibody to STXBP5 and immunoblotted the
precipitants with antibody to STX4. STXBP5 and STX4 co-precipitate (Figure 2-11B).
However, STXBP5 does not co-precipitate with STX11 or SNAP23 (Figure 2-11B).
We then searched for co-localization of STXBP5 and STX4 by immunofluorescence
staining of HUVEC. Confocal microscopy showed STXBP5 and STX4 are both
distributed in a punctate pattern, and a subset of STXBP5 co-localizes with STX4
(Pearson’s coefficient = 0.80 ± 0.03, Figure 2-11C). Co-localization of STXBP5 and
STX4 was mainly found in the cytoplasm (Figure 2-11C, upper panel). In a subset of
cells, a fraction of STX4 also co-localizes with a relatively sparser fluorescent punctation
of STXBP5 on the cell membrane (Figure 2-11C, lower panel). Taken together, these
data show that STXBP5 interacts with a complex containing STX4 in endothelial cells.
59
Given the pivotal role that STX4 plays in endothelial vesicle trafficking (108, 109), their
interaction may underline the inhibitory function of STXBP5 in endothelial exocytosis.
We also searched for an association between STXBP5 and SYT1. STXBP5 and SYT1
partially co-sediment (Figure 2-11A). STXBP5 and SYT1 co-precipitate, especially in
cells that have been stimulated with histamine or A23187 (Figure 2-11B). We also
searched for co-localization of STXBP5 and SYT1 by immunofluorescence staining of
HUVEC. In resting HUVEC, STXBP5 and SYT1 are both distributed in a punctate
pattern, and a subset of STXBP5 co-localizes with SYT1 (Pearson’s coefficient = 0.70 ±
0.08, Figure 2-11D). Furthermore, both STXBP5 and SYT1 co-localize to plasma
membranes of HUVEC after stimulation (Pearson’s coefficient = 0.69 ± 0.08, Figure
2-11D, lower row). These data reinforce observations that STXBP5 interacts with SYT1
in neurons (321, 346).
We also searched for interactions of STXBP5 and other molecules that regulate vesicle
trafficking in endothelial cells. STXBP5 does not co-precipitate with STX11 or SNAP23
(Figure 2-11B), or with NSF, Munc18-2, or Munc18-3 (Figure 2-13). Furthermore,
STXBP5 fails to co-localize significantly with other SNARE molecules, such as
SNAP23, VAMP8, or VAMP3 (Figure 2-14). STXBP5 also does not co-localize with
caveolin 1, Munc18-2, Munc18-3, and NSF (Figure 2-15). Finally, STXBP5 does not co-
localize with Rab27a, Myosin 5a, or MyRIP (Figure 2-16).
60
Taken together, our data suggest that STXBP5 interacts with a complex containing STX4
and SYT1 in endothelial cells.
2.2.6 STXBP5 inhibits endothelial exocytosis in vivo
In order to test whether or not STXBP5 regulates endothelial exocytosis in vivo, we
compared the plasma level of VWF in mice that were Stxbp5 deficient (Stxbp5 KO) and
in wild-type (Stxbp5 WT) mice. VWF levels are higher in Stxbp5 KO mice than in Stxbp5
WT mice (Figure 2-17). These data match our in vitro data showing that VWF release is
greater from endothelial cells with knockdown of STXBP5 compared to control cells
(Figure 2-2). In addition, this difference in VWF levels is not due to the ADAMTS13
enzyme that cleaves VWF in the blood, since ADAMTS13 activity is unchanged in
Stxbp5 WT mice and Stxbp5 KO mice (Figure 2-17B).
To explore the physiological relevance of STXBP5, we measured the effect of STXBP5
upon platelet interactions with endothelial cells in vivo. Platelet rolling or adherence to
the vessel walls is mediated by VWF released by exocytosis (347). We hypothesized that
deficiency of STXBP5 would permit an increase in WPB exocytosis, resulting in an
increase in platelet adherence to venule walls. Anesthetized Stxbp5 KO mice or Stxbp5
WT mice were transfused with fluorescent antibodies to label platelets. The mesentery
was externalized, superfused with 10 µM Ca2+ ionophore A23187 to stimulate WPB
exocytosis, and intravital microscopy was used to record interactions of fluorescently
labeled platelets with mesenteric venules. Platelets that remained static over multiple
61
frames or were captured by the endothelium in consecutive frames and then released in a
stop-and-go fashion were classified as adherent platelets.
In Stxbp5 WT mice, A23187 rapidly induces platelet adhesion to the venule wall (Figure
2-18A). In Stxbp5 KO mice, A23187-induced platelet interaction with the venule wall
was significantly increased (Figure 2-18A). Quantification of adherent platelets showed
that STXBP5 deficiency significantly increased A23187-induced platelet interactions
with the venule walls, but not the interaction in the resting condition (Figure 2-18B). This
increase in platelet adherence to the venule would be predicted if STXBP5 deficiency led
to increased exocytosis of WPBs and greater release of VWF.
Finally we used an assay that directly measures endothelial exocytosis, so that we would
eliminate the possible effects of STXBP5 upon platelets. Mice were perfused with
fluorescently labeled microspheres conjugated with antibody to P-selectin, and adhesion
to resting and A23187-stimulated mesenteric venules was imaged by intravital
microscopy. In Stxbp5 WT mice, A23187 treatment increases microsphere adherence to
venules (Figure 2-18 C-D). However, in Stxbp5 KO mice, microsphere adherence to
venules increases even more after A23187 stimulation (Figure 2-18 C-D). This suggests
that STXBP5 limits endothelial display of P-selectin in vivo. Taken together, these data
show that STXBP5 inhibits endothelial exocytosis in vivo.
62
2.2.7 STXBP5 affects hemostasis and thrombosis
We proceeded to test the effect of STXBP5 upon thrombosis. We expected that mice
deficient in Stxbp5 would have increased thrombosis, since they have increased plasma
VWF levels and platelet-endothelium interaction upon stimulation. We first measured the
time for hemostasis in a murine tail bleeding model. Contrary to our expectations, mice
lacking Stxbp5 displayed prolonged bleeding compared to wild-type mice (Figure
2-19A). We next measured the time for thrombosis in a murine mesenteric thrombosis
model. Stxbp5 KO mice had severely delayed time to formation of thrombus and time to
vessel occlusion, compared to Stxbp5 WT mice (Figure 2-19B). We then measured the
time for flow cessation in a murine carotid artery thrombosis model. Although carotid
artery flow ceases approximately 8 min after arterial injury in the wild-type mice, carotid
arteries remain patent for more than 30 min after arterial injury in the Stxbp5 KO mice
(Figure 2-19C). Taken together, these data suggest that mice lacking Stxbp5 have a
defect in thrombosis.
These results were surprising: mice lacking Stxbp5 have elevated plasma levels of VWF,
increased endothelial exocytosis, and increased platelet rolling along the vessel wall – but
also have decreased thrombosis. One explanation for this apparent paradox is that
STXBP5 also regulates platelet secretion.
63
2.2.8 STXBP5 promotes platelet secretion and activation
To test the hypothesis that Stxbp5 promotes platelet secretion, we first discovered that
murine platelets express Stxbp5 protein (Figure 2-20A). These data match our data
showing that human platelets express mRNA for STXBP5 isoforms (Figure 2-1 C-D).
We then explored the role of Stxbp5 in platelet activation and secretion, using platelets
from wild-type and Stxbp5 KO mice. To study platelet exocytosis of alpha-granules, we
measured externalization of P-selectin by flow cytometery. To study platelet secretion of
dense granules, we measured release of ATP by a luciferase based assay. To study
platelet activation, we measured activation of the integrin GPIIbIIIa by flow cytometery.
Platelets from Stxbp5 KO mice have defective P-selectin externalization, ATP release,
and GPIIbIIIa activation (Figure 2-20B). These data suggest that Stxbp5 facilitates
platelet exocytosis and activation.
To confirm these data, we analyzed wild-type or Stxbp5 KO mice that had received bone
marrow transplantation of either genotypes. We measured plasma VWF levels in Stxbp5
WT recipient mice receiving bone marrow from Stxbp5 WT donors or Stxbp5 KO donors,
and in Stxbp5 KO recipient mice receiving bone marrow from either donors. Plasma
levels of VWF were not changed by bone marrow transplantation (Figure 2-21A).
However, bleeding times increased in mice receiving Stxbp5 KO bone marrow, and
decreased in mice receiving Stxbp5 WT bone marrow (Figure 2-21B). Furthermore,
Stxbp5 WT mice that received Stxbp5 KO bone marrow failed to form an occlusive
thrombus in a carotid artery thrombosis model, but the non-occlusive phenotype of
64
Stxbp5 KO recipients were completely abolished by transplantation of Stxbp5 WT bone
marrow (Figure 2-21C). Taken together, these data show that Stxbp5 in endothelial cells
determines the plasma VWF level, but the thrombosis phenotype is determined by Stxbp5
in bone marrow cells (such as platelets), not by the host cells (such as endothelial cells).
65
Figure 2-1 STXBP5 expression and transcript variants.
(A) STXBP5 expression in human endothelial cells. Lysates of cultured human aortic
endothelial cells (HAEC), human umbilical vein endothelial cells (HUVEC), and human
dermal microvascular endothelial cells (HDMVEC) were probed by immunoblotting with
66
antibody to STXBP5 (top). Blotting to β-actin was used as a loading control (bottom).
Representative of 3 separate experiments.
(B) Stxbp5 expression in murine tissue. RNA was isolated from Stxbp5 WT mouse
tissues, and Stxbp5 mRNA expression was measured by qPCR and normalized to mRNA
level in brain (n=3 mice ± S.D).
(C) Human STXBP5 transcript variants. STXBP5 transcript variants were detected by RT-
PCR using primers flanking splice region in human brain, HUVEC, human platelets, and
HEK293 cells. The products were separated by agarose gel, sequenced, and compared
with NCBI Reference Sequences. The length of each PCR product were: 370 bp
(STXBP5-X1), 322 bp (STXBP5-X2), 307 bp (STXBP5-2), 262 bp (STXBP5-X3), and 199
bp (STXBP5-1). The similarities between PCR products and NCBI entries were 99%
(STXBP5-X1), 95% (STXBP5-X2), 100% (STXBP5-2), 100% (STXBP5-X3), and 100%
(STXBP5-1).
(D) Relative abundance of STXBP5 transcript variants in human brain, HUVEC, and
human platelets were measured by qPCR using variant-specific Taqman probes, with
brain STXBP5-1 set as 100% (n = 3 ± S.D).
67
Figure 2-2 STXBP5 inhibits endothelial exocytosis in vitro.
HUVEC was transfected with siRNA against STXBP5 (siSTXBP5) or control siRNA
(siControl), stimulated with an agonist, and the amount of VWF released into the media
at resting condition and after 30-min stimulation was measured by an ELISA (n=3 S.D.
68
* P < 0.05 for agonist treatment vs. control). Immunoblots at the top show siRNA
knockdown of STXBP5 expression in EC, with endogenous actin as loading control.
Shown are noncontiguous parts from the same gel.
(A-C) Knockdown of STXBP5 increases histamine-induced release of VWF, but has no
effect upon constitutive VWF release in (A) HAEC, (B) HUVEC, and (C) HMVDEC.
(D-E) Knockdown of STXBP5 in HUVEC increases VWF release induced by (D) ATP,
and by (E) calcium ionophore A23187.
69
Figure 2-3 STXBP5 and constitutive release of VWF.
Constitutive release of VWF was measured in 3-h intervals over 12 h in resting HUVEC
transfected with control siRNA or STXBP5 siRNA for 72 hours (n = 3 ± S.D. No
significant difference between siControl and siSTXBP5 was detected over the time
course).
C u ltu re t im e
VW
F r
ele
as
e(m
U/m
l)
3 h 6 h 9 h 1 2 h
0
1
2
3
s iC o n tro l
s iS T X B P 5
70
Figure 2-4 STXBP5 does not affect HUVEC content of VWF.
Total protein content in lysates of HUVEC transfected with siControl or siSTXBP5 was
measured and normalized by bicinchoninic acid assay, and lysates contain 1 μg total
protein were measured VWF content by ELISA (n = 4 ± S.D. No significant difference
was detected).
71
Figure 2-5 Knockdown of STXBP5 does not affect the number of WPB granules per
cell.
HUVEC were transfected with siSTXBP5 or siControl, or sham transfected, and the
number of WPB was counted by ImagePro Plus software (n = 3 replicate plates ± S.D.;
granules were counted in over 240 cells per plate. No significant difference among
groups were detected by one-way analysis of variance (ANOVA). Scale bar = 50 μm).
72
Figure 2-6 Knockdown of STXBP5 does not affect the size of WPB granules.
HUVEC were transfected with siSTXBP5 or siControl, and the size of individual WPB
was measured by ImagePro Plus (n = 3 replicate plates ± S.D.; granules were counted in
over 240 cells per plate. No significant difference between siControl and siSTXBP5 in all
groups).
P>0.05 for all groups
In d iv id u a l W P B S iz e (p ix e l2)
% T
ota
l W
PB
Po
pu
lati
on
3 - 8 8 - 1 3 1 3 - 1 8 1 8 - 2 3 2 3 - 2 8 2 8 - 3 3 3 3 - 3 8 3 8 - 4 3 4 3 - 4 8 > = 4 8
0
5
1 0
1 5
2 0
6 5
7 0
7 5S iC o n tro l
S iSTXBP5
73
Figure 2-7 Knockdown of STXBP5 does not affect the shape of WPB granules.
HUVEC was transfected with siSTXBP5 or siControl. The aspect ratio of individual
WPB was measured by ImagePro Plus, and its distribution was plotted as histogram (n =
3 replicate plates ± S.D.; granules were counted in over 240 cells per plate. No significant
difference between siControl and siSTXBP5). Shown at the lower panel is one
representative image of WPB and one example of measurement.
74
75
Figure 2-8 STXBP5 inhibits HUVEC release of P-selectin.
(A) Knockdown of STXBP5 in HUVEC increases P-selectin externalization as measured
by HL-60 cell adherence after 10 μM histamine stimulation. HL-60 cells were labeled
with calcein AM to allow visualization by a fluorescent microscope. HUVEC treated
with an antibody to P-selectin (anti-CD62) serve as a negative control. Shown are
representative bright-field and corresponding fluorescent images from multiple fields
from 3 wells per transfection per treatment (10 × water immersion lens; scale bar = 150
μm).
(B) Knockdown of STXBP5 in HUVEC increases P-selectin externalization as measured
by HL-60 cell adherence. Cells treated with antibody to P-selectin (Anti-CD62) serve as
negative control (n = 3 ± S.D. * P < 0.05 vs siControl).
76
Figure 2-9 STXBP5 SNP and release of VWF.
HUVEC were transfected with siSTXBP5 to knock down endogenous STXBP5
expression, and some co-transfected with an expression vector for STXBP5(WT) or
STXBP5(N436S). Release of VWF was measured after treatment with media or
histamine (n = 4 ± S.D. N.S. Not significant. * multiplicity adjusted P <0.05 by two-way
ANOVA followed by the Tukey post test).
77
Figure 2-10 Particles containing STXBP5 are not co-localized with endothelial
granules containing VWF.
(A) HUVECs were imaged by confocal microscopy after staining with antibodies to
STXBP5 (green), VWF (red), and DNA (blue) (Scale bar = 20 μm; Objective: oil 40 ×;
confocal z-resolution: 0.40 µm).
(B) Enlargement of insets in (A). STXBP5 is minimally co-localized with WPBs (Scale
bar = 5 μm; Objective: oil 40 ×; confocal z-resolution: 0.40 µm). Representative of more
than 3 separate experiments.
78
Figure 2-11 STXBP5 co-sediments, co-localizes, and co-precipitates with STX4 and
SYT1.
79
(A) STXBP5 co-sediments with STX4 and SYT1 by sucrose density gradient
fractionation. HUVEC lysate was ultracentrifuged through discontinuous 5%-40%
sucrose gradient, and the fractions were analyzed by SDS-PAGE (T, total proteins of the
lysate before fractionation; P, pellet after fractionation). Representative of 3 separate
experiments.
(B) STXBP5 co-precipitates with STX4 and SYT1. HUVEC lysate treated with media or
histamine or A23187 were immunoprecipitated with antibody to STXBP5 or mouse IgG1;
precipitants were probed with antibody to SYT1 or STX4 or STX11 or SNAP23.
Representative of 3 similar experiments.
(C) STXBP5 co-localizes with STX4 by confocal microscopy. HUVEC were stained for
STXBP5 (green), STX4 (red), and DNA (blue). STXBP5 partially co-localizes with
STX4. Limited cell membrane localization of STX4 and STXBP5 is detectible in a subset
of cells shown in the lower panel (Upper panel: scale bar = 20 μm; Objective: oil 40 ×;
confocal z-resolution: 0.40 µm; lower panel: scale bar = 30 μm; Objective: oil 60 ×;
confocal z-resolution: 0.32 µm). Pearson’s coefficient (green vs red) = 0.80 ± 0.03.
(D) STXBP5 co-localizes with SYT1. Resting and stimulated HUVEC were stained for
STXBP5 (green), SYT1 (red), and DNA (blue). STXBP5 partially co-localizes with
SYT1, and both STXBP5 and SYT1 partially translocate to the membrane after
stimulation. (Scale bar = 20 μm; Objective: oil 60 ×; confocal z-resolution: 0.32 µm).
80
Pearson’s coefficient (green vs red) =0.70 ± 0.08 for resting cells; 0.69 ± 0.08 for
histamine-stimulated cells.
81
82
Figure 2-12 STXBP5 does not co-localize with markers for ER, Golgi, lysosomes, or
endosomes.
Confocal microscopy was used to localize STXBP5, markers for endoplasmic reticulum,
Golgi apparatus, lysosome, or early endosome. STXBP5 does not co-localize with these
organelle markers. Scale bar = 20 μm.
83
Figure 2-13 STXBP5 does not interact with NSF and Munc family members.
Lysates of resting or stimulated HUVEC were precipitated with antibody to STXBP5 or
IgG, and precipitants were immunoblotted with antibody to NSF or Munc18-2 or
Munc18-3.
84
Figure 2-14 STXBP5 does not co-localize with SNAP23, VAMP8, or VAMP3.
Confocal microscopy was used to localize STXBP5 (green), SNAP23, VAMP8, VAMP3
(red), and DNA (blue) in HUVEC. STXBP5 does not co-localize with these SNAREs.
Scale bar = 20 μm.
85
86
Figure 2-15 STXBP5 does not co-localize with Caveolin-1, Munc18-3, Munc18-2, or
NSF.
Confocal microscopy was used to localize STXBP5 (green), Caveolin-1, Munc18-3,
Munc18-2, NSF (red), and DNA (blue) in HUVEC. STXBP5 does not co-localize with
these proteins in HUVEC. Scale bar = 20 μm.
87
Figure 2-16 STXBP5 does not co-localize with Rab27, Myosin5a or MyRIP.
Confocal microscopy was used to localize STXBP5 (green), Rab27, Myosin5a, MyRIP
(red), and DNA (blue) in HUVEC. STXBP5 does not co-localize with these proteins in
HUVEC. Scale bar = 20 μm.
88
Figure 2-17 Stxbp5 affects plasma VWF in mice.
(A) Plasma levels of VWF are higher in Stxbp5 KO mice than in WT mice. Plasma VWF
levels were measured by an ELISA from retro-orbital bleeding (n = 8~12 S.D.; * P <
0.05 vs. WT). (B) Plasma ADAMTS13 activity is similar in Stxbp5 KO and Stxbp5 WT
mice. ADAMTS13 activity in mouse plasma was measured by a fluorescent assay and
expressed as percentage of pooled WT plasma.
89
Figure 2-18 Stxbp5 inhibits endothelial exocytosis in mice.
(A) Intravital microscopy of platelet-EC interactions in vivo. Platelets were fluorescently
labeled in Stxbp5 WT and Stxbp5 KO mice. 10 µM ionophore A23187 was superfused
onto the mesenteric venule after 120s baseline recording. Platelet adhesion to the
90
mesenteric venule was continuously recorded by intravital microscopy. Shown are
representative images at baseline (90s) and after A23187 stimulation (345s and 600s).
Magnification 20×.
(B) Stxbp5 deficiency increases platelet-EC interactions in vivo. Platelet adhesion in (C)
was calculated by averaging the number of platelets that remained static over at least two
consecutive frames (frame rate: approximately 31 frames/second). Quantification was
expressed as adherent platelets per minute in an area of 51200 pixel2 (n = 6 mice per
genotype S.D. * P < 0.05 vs. Stxbp5 WT).
(C) Stxbp5 deficiency increases microsphere-EC interactions in vivo. Mice were perfused
with fluorescent microspheres conjugated with antibody to P-selectin, and adhesion of
microspheres was imaged 2 min before and 8 min after A23187 superfusion on
mesenteric venule. Green lines mark borders of vessel within which adherent
microspheres were counted. Magnification 20×.
(D) Quantification of adherent microspheres in (E) (n = 3-4 mice per genotype S.D. * P
< 0.05 vs. Stxbp5 WT).
91
92
Figure 2-19 Stxbp5 increases thrombosis in mice.
(A) Stxbp5 regulates bleeding time in mice. Tail bleeding times in WT and Stxbp5 KO
mice were measured after distal 5 mm tail amputation (n = 10-12 ± S.D. *P < 0.01 by
Kolmogorov-Smirnov test).
(B) Stxbp5 regulates mesenteric thrombosis in mice. Intravital microscopy was used to
measure thrombosis of mesenteric arterioles in WT and Stxbp5 KO mice, including the
time to form (left) a 50-pixel diameter thrombus, and (right) full occlusion (n = 3 - 6 ±
S.D. *P < 0.05 vs Stxbp5 WT by Kolmogorov-Smirnov test).
(C) Stxbp5 regulates carotid thrombosis in mice. Carotid arterial flow was measured by a
Doppler flow probe after FeCl3 injury, and flow after FeCl3 wash-off was plotted as
percentage of baseline flow before injury. Stxbp5 KO mice failed to form vessel
occlusion (n = 7 ± S.D. *P<0.05 by repeated measures two-way ANOVA).
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Figure 2-20 Stxbp5 regulates platelet secretion and activation.
(A) Stxbp5 expression in platelets from Stxbp5 WT and Stxbp5 KO mice as measured by
immunoblotting. Gapdh was used as loading control.
(B) Stxbp5 promotes platelet secretion. Platelets were isolated from Stxbp5 WT and
Stxbp5 KO mice, and stimulated with PBS (resting) or thrombin at indicated
concentrations. Median fluorescence intensity (MFI) changes as measured by flow
cytometry following P-selectin externalization (left), integrin GPIIbIIIa activation
(middle), and bioluminescence change following ATP release (right) were measured (n =
3 mice per treatment ± S.D. *P < 0.05 vs. Stxbp5 WT).
94
95
Figure 2-21 Stxbp5 in platelets increases thrombosis in mice.
(A) Transplantation of bone marrow does not affect plasma VWF levels. Bone marrow
from Stxbp5 WT or Stxbp5 KO mice were transplanted into lethally-irradiated Stxbp5 WT
or Stxbp5 KO recipients, respectively. Plasma VWF levels were measured 6 weeks after
transplantation (n = 5 – 9 ± S.D. NS, not significant).
(B) Transplantation of Stxbp5 KO bone marrow increases Stxbp5 WT tail bleeding time,
whereas transplantation of Stxbp5 WT bone marrow decreases Stxbp5 KO tail bleeding
time. Bone marrow from Stxbp5 WT or Stxbp5 KO mice were transplanted into lethally-
irradiated Stxbp5 WT or Stxbp5 KO recipients. Tail bleeding time was measured 6 weeks
after transplantation (n = 6 – 9 ± S.D. *P < 0.05 by Kolmogorov-Smirnov test).
(C) Transplantation of Stxbp5 KO bone marrow decreases carotid arterial thrombosis in
Stxbp5 WT recipients, whereas transplantation of Stxbp5 WT bone marrow increases
thrombosis in Stxbp5 KO recipients. Bone marrow from Stxbp5 WT or Stxbp5 KO mice
were transplanted into lethally irradiated Stxbp5 WT or Stxbp5 KO recipients. Carotid
arterial flow was measured 6 weeks after transplantation with FeCl3 injury (n = 4 – 7 ±
S.D. * multiplicity adjusted P < 0.05 by repeated measures two-way ANOVA with Tukey
multiple comparisons).
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2.3 Discussion
The major finding of Chapter 2 is that STXBP5 regulates endothelial and platelet
exocytosis. STXBP5 interacts with components of the exocytic machinery and inhibits
release of VWF in endothelial cells and in mice. STXBP5 also promotes platelet
secretion and thrombosis in mice. Thus, our current study provides strong functional
evidence for the regulatory role on circulating VWF and thrombosis by a candidate gene
identified by GWAS (317-320, 339).
We find that STXBP5 inhibits endothelial exocytosis. Others have found that STXBP5
inhibits secretion from other cell types. STXBP5 was originally identified as a protein
(called tomosyn) that inhibits neurotransmitter release from neurons (321). Subsequent
studies showed that STXBP5 decreased exocytosis in yeast, C. elegans synapses,
pancreatic β-cells and neurosecretory cells (329, 331-337). We found that STXBP5
inhibits endothelial exocytosis of granules. Cells with decreased STXBP5 have increased
VWF release into the media and increased cell membrane display of P-selectin; and mice
lacking Stxbp5 have elevated plasma VWF levels and greater display of P-selectin
(Figure 2-2, Figure 2-8, and Figure 2-18).
STXBP5 might regulate endothelial exocytosis through several potential mechanisms.
One proposed model is that STXBP5 blocks formation of a ternary SNARE complex by
competing with VAMP isoforms to interact with SNAP and syntaxin isoforms. For
example, in neurons the VAMP-like domain in the carboxy-terminus of STXBP5 forms a
97
SNARE complex-like structure with SNAP25 and syntaxin 1A. This STXBP5-SNARE
complex is stable: VAMP2 cannot displace STXBP5, resulting in inhibition of SNARE
complex assembly (328-330, 348). Alternatively, another proposal is that STXBP5
enhances oligomerization of the ternary SNARE complexes, locking the complexes into a
structure that cannot proceed with membrane fusion. For example, in neurons the N-
terminal WD40 repeat domain of STXBP5 functions as a scaffold that enhances
oligomerization of pre-assembled SNARE complexes and reduces vesicle priming and
turnover (332). In addition to these STXBP5-SNARE interaction models, Sro7, the yeast
STXBP5 homolog, acts as an allosteric regulator of exocytosis. The tail domain of Sro7
interacts with the Rab GTPase Sec4, raising the possibility that STXBP5 might be able to
interact with some other factors that control exocytosis (324, 349). Our observation that
STXBP5 interacts with a complex containing STX4 (Figure 2-11) favors the STXBP5-
SNARE interaction model. STX4 is a t-SNARE that binds to cognate v-SNAREs. The
inhibition of VWF exocytosis is probably a consequence of displacement of the
endothelial v-SNARE by STXBP5, forming a “dead-end” complex and sequestering the
formation of productive SNARE complexes.
STXBP5 can interact with either syntaxin or SNAP isoforms, as reported previously
(321, 325, 329, 350). However, our data seem to favor the STXBP5-syntaxin binary
interaction over the STXBP5-syntaxin-SNAP ternary interaction or STXBP5-SNAP
interaction in ECs, since co-localization and co-IP studies failed to detect an interaction
between STXBP5 and SNAP23. Indeed, syntaxin has been shown to be responsible for
STXBP’s inhibition of exocytosis and subcellular localization (350). The VLD of
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STXBP5 interacts with syntaxins in various cell types (321, 328-332, 335, 337, 350-353).
In PC12 cells, overexpressed STXBP5 showed a primarily cytosolic localization similar
to endothelial cells, but overexpression of syntaxin directs approximately 55% of
cytosolic STXBP5 to the plasma membrane (350). In contrast, whether a binary
interaction between STXBP5 and an SNAP isoform directs exocytosis has been
controversial. For instance, glutathione S-transferase (GST)-syntaxin-1a could bind to
STXBP5 overexpressed in COS7 cells, but GST-SNAP25 could not bind to STXBP5
under the same condition (321). Other studies, however, demonstrate STXBP5 interacts
with SNAP homologs in yeas (325), adipocytes (354), and PC12 cells (331), but whether
this interaction is formed indirectly via a ternary complex containing syntaxin is
unknown. Although we cannot exclude the existence of such a ternary complex in EC,
our data agrees with a recent study showing STXBP5 interacts mainly with syntaxin, and
STXBP5-SNAP interaction is not abundant and relies on the presence of syntaxin (350).
Another possible mechanism for STXBP5 inhibition of exocytosis is through its
interaction with a calcium sensor, SYT1. SYT1 regulates SNARE-mediated vesicle
fusion by bridging membrane fusion through a calcium-dependent mechanism (355-358).
SYT1 has been shown to be recruited to pseudogranules containing heterologously
expressed VWF (359). Others have shown that STXBP5 interacts with SYT1 through
WD40 repeat domain in the amino-terminus of STXBP5, blocking the ability of SYT1 to
mediate calcium-dependent neurotransmitter release (346). We also found that STXBP5
interacts with SYT1 with high affinity after histamine stimulation (Figure 2-11). This
raises the possibility that calcium triggers exocytosis in part by relieving the inhibitory
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effects of STXBP5, either by displacing STXBP5 from t-SNAREs or by promoting an
interaction between STXBP5 and SYT1.
Our data raise some interesting issues. First, our data suggest STXBP5 inhibits secretory
release of VWF but not constitutive release of VWF (Figure 2-2). Recent studies suggest
that endothelial cells release VWF through three distinct pathways: stimulated secretion,
unstimulated basal release, and unstimulated constitutive release (266). However, the
proteins that mediate unstimulated release have not been defined, and it is unknown
which proteins regulate both the unstimulated release pathway and the stimulated
pathway. Our data suggest that STXBP5 acts upon one or more proteins in the stimulated
secretory pathway. For example, our data show that STXBP5 interacts with STX4.
Furthermore, others have shown that in neurons, STXBP5 interacts with SYT1 whose
regulatory role in exocytosis depends on calcium (346).
One striking finding of our study is that STXBP5 has opposing effects on exocytosis
from endothelial cells and platelets: STXBP5 inhibits endothelial exocytosis but activates
platelet exocytosis. Platelets from Stxbp5 KO mice have impaired secretion of alpha-
granules as measured by P-selectin externalization, defective exocytosis of dense
granules as measured by ATP release, and blunted activation as measured by GPIIbIIIA
conformational changes (Figure 2-20B). Furthermore, mice lacking Stxbp5 display severe
defects in bleeding time, mesenteric arteriole thrombosis, and carotid artery thrombosis.
Bone marrow transplant experiments suggest that this hemostasis defect is due to bone
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marrow-derived cells, presumably platelets, lacking STXBP5, not endothelial cells
lacking STXBP5 (Figure 2-21).
Why does STXBP5 promote platelet exocytosis but suppress endothelial exocytosis? We
considered the idea that platelets and endothelial cells express different isoforms of
STXBP5, but our PCR studies show that both cell types express predicted STXBP5
transcript variants in similar relative abundance (Figure 2-1D). Another possibility is that
STXBP5 interacts with different SNARE members in different cell types. Although
STXBP5 interacts with STX4 and SYT1 in endothelial cells (Figure 2-11), STXBP5 does
not significantly interact with STX4 or SYT1 in human platelets (data not shown), or
with STX4 in rat brain (321, 322). Others have reported divergent effects of STXBP5 in
differing cell systems. For example, STXBP5 inhibits neurotransmitter release from
neurons (331-337). But STXBP5 actually increases the readily releasable pool of vesicles
in PC12 cells and in rat β-cell line INS-1E; this stimulation of exocytosis depends in part
on the phosphorylation of STXBP5 (336, 353). Detailed studies of molecular
mechanisms are needed to reconcile the dual stimulatory and inhibitory role of STXBP5
in different cell types.
Mice lacking Stxbp5 have increased plasma levels of VWF but decreased thrombosis.
Why? One possible explanation is that STXBP5 has opposing effects upon endothelial
and platelet exocytosis. In Stxbp5 null mice, the increased plasma VWF is caused by the
absence of endothelial STXBP5 that normally limits endothelial release of VWF. Since
the major source of VWF in mouse plasma is from endothelial cells, not from platelets
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(195), mice lacking Stxbp5 in both endothelial cells and platelets demonstrate increased
plasma VWF. Our data with bone marrow transplanted mice supports this conclusion,
since plasma VWF levels are unchanged by bone marrow from Stxbp5 KO donors
(Figure 2-21). In Stxbp5 null mice, the decreased thrombosis is caused by the absence of
platelet STXBP5 that normally boosts platelet activation. The enhanced plasma VWF is
not sufficient to overcome the diminished platelet activation, and the net result of
STXBP5 absence from both endothelial cells and platelets is defective thrombosis.
Human GWAS show that multiple genetic variants associated with altered VWF levels
lie within the STXBP5 allele (317-319, 340). One SNP highly correlated with altered
VWF levels in human, rs1039084, is located in exon 23 of STXBP5 (319). This non-
synonymous SNP rs1039084 changes 1307 A to G in STXBP5, changing codon 436 from
Asn to Ser (N436S). Human studies have found that this mutation STXBP5 (N436S) is
associated with lower levels of VWF in the plasma (319).
We found that this STXBP5 (N436S) mutant decreases endothelial exocytosis more than
STXBP5-WT (Figure 2-9). Residue N436 lies within the second N-terminal WD40 repeat
domain of STXBP5. WD40 repeats can form scaffolds which assemble protein
complexes, and it has been previously shown that the STXBP5 WD40 repeat domain
interacts with SNAREs STX1 and SNAP25 in neurons (332). How might the STXBP5
(N436S) mutation enhance inhibition of exocytosis? Prior studies show that the integrity
of the WD40 domain is critical for STXBP5’s inhibition of exocytosis (331). PC12 cells
expressing mutant rat m-STXBP5 lacking two fragments (aa 537–578 and aa 897–917) in
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the WD40 domain lost the inhibition of exocytosis compared with cells expressing the
wild type m-STXBP5 (335). These mutants also showed altered binding and diffusion
kinetics of STXBP5 on the PC12 cell membrane compared to wild-type m-STXBP5,
possibly via altered interactions with syntaxin or with other unknown membrane proteins
(350). In addition, using a liposome co-flotation assay, both the N-terminal domain and
C-terminal domain (VLD) of STXBP5 are required for the binding to the syntaxin
monomer, suggesting features unique to the full-length STXBP5. It is possible that the
mutation STXBP5 (N436S) alters the function of its WD40 domain and enhances the
affinity of STXBP5 for STX4 or SYT1 in endothelial cells, blocking the formation of a
ternary SNARE complex or blunting the Ca2+ triggering of membrane fusion. Another
possibility is that the mutation decreases the conformational changes that might permit
STXBP5 to release SNAREs to form the ternary SNARE complex. Or perhaps the
STXBP5 (N436S) mutation serves as a gain-of-function mutation that mediates the
interaction of STXBP5 with additional partners.
Genetic variation within STXBP5 is linked to the risk of venous thromboembolic events
in human subjects (320). The effect of the mutation STXBP5 (N436S) upon platelet
activation is unknown. We found that the mutation STXBP5 (N436S) enhances the
function of STXBP5 in endothelial cells, with enhanced suppression of exocytosis
(Figure 2-9). We would expect that the mutation would also enhance the function of
STXBP5 in platelets, with further promotion of platelet activation. However, in human
studies, the SNP rs1039084 for STXBP5 (N436S) is associated with a decreased
incidence of venous thromboembolic disease (320). It is possible that different partners
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interacting with STXBP5 in platelets and endothelial cells determines the effect of the
N436S mutation. Further studies are needed to determine the effect of mutations upon
STXBP5 function in platelets.
In conclusion, our data identify a key role for STXBP5 as a regulator in endothelial
exocytosis and thrombosis. STXBP5 inhibits stimulated endothelial release of VWF and
translocation of P-selectin in vitro. STXBP5 co-localizes with endothelial vesicles and
forms complexes with endothelial SNARE STX4. Moreover, Stxbp5 deficiency in mice
significantly increases plasma VWF levels and P-selectin translocation. However,
STXBP5 also promotes platelet secretion, and mice lacking Stxbp5 have increased
bleeding (Scheme 2-2). Our studies are relevant to humans, since human GWAS data
associates STXBP5 genetic variants with plasma VWF levels and thombosis. Further
characterization of the regulatory functions of STXBP5 in exocytosis may lead to novel
insights into vascular diseases such as VTE and VWD.
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Scheme 2-2 STXBP5 differentially influences exocytosis in platelets and endothelial
cells.
Adapted from Lillicrap D. The Journal of Clinical Investigation. 2014;124(10):4231-3.
(360).
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VAMP8 Mediates Endothelial Exocytosis
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3.1 Introduction
The ternary SNARE complex is typically comprised of one member from each of the
following families: syntaxin (t-SNARE), SNAP (t-SNARE) and VAMP (v-SNARE).
Unique combinations of three SNARE molecules play a role in directing granules to
specific target membranes.
The molecular components of the ternary SNARE machinery has been intensively
studied in neurons and many neuroendocrine and immune cells. Neurotransmission is
regulated in part by VAMP2 on synaptic vesicles interacting with STX1 and SNAP25 on
the pre-synaptic plasma membrane (361). Compound exocytosis in mast cells is mediated
in part by VAMP8 on secretory granules in conjunction with SNAP23 and STX4(362-
366). The three SNAREs controlling alpha-granule exocytosis from platelets include
STX11(367), VAMP8 on alpha-granule membranes (368-372), and SNAP23 that
distributes predominantly on platelet membranes as well as on membranes of granules
and the platelet open canalicular system (367-369, 373-378).
The exact identities of endothelial v-SNAREs are ambiguous and poorly understood. A
prior study found VAMP3 and VAMP8 associated with endothelial WPBs (102).
However, the function of these VAMPs was not fully characterized. More importantly,
the importance of endothelial v-SNARE in regulating exocytosis and thrombosis in vivo
is unknown.
107
We hypothesized that endothelial exocytosis is regulated by unique v-SNARE molecules.
Chapter 3 shows that VAMP8, in addition to VAMP3, is a key SNARE in endothelial
cells that drives granule exocytosis and affects plasma VWF and thrombosis.
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3.2 Results
3.2.1 Endothelial cells express a distinct set of SNAREs that mediate
granule exocytosis
We hypothesized that a distinct subset of SNAREs would mediate endothelial exocytosis.
We searched for endothelial expression of SNAREs by immunoblotting endothelial cell
lysates, and compared it to the known expression of SNAREs in brain, platelets, PC12 or
HeLa cells (as positive controls) or absence of SNAREs in human vascular smooth
muscle cells (VSMC, as a negative control). We found that HUVEC express certain
SNAREs such as STX4, SNAP23, VAMP3 and VAMP8 (Figure 3-1A-B), but have
undetectable level of other SNAREs such as STX1, SNAP25, VAMP1, and VAMP2.
Intriguingly, different endothelial cell types (such as HUVEC, HAEC, and human brain
microvascular endothelial cell (HBMEC)) express different levels of VAMP3 and
VAMP8 (Figure 3-1C). We focused our attention on four of these SNAREs, VAMP3,
VAMP8, STX4 and SNAP23, since they can interact with each other in other biological
systems (365, 379, 380).
3.2.2 VAMP8 and VAMP3 co-localize with VWF
We next performed confocal microscopy to see which SNARE is associated with
endothelial granules containing VWF, the WPBs (180). We found that VWF is expressed
in a granular pattern throughout endothelial cells (Figure 3-2A-D, green). Neither STX4
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nor SNAP23 co-localizes with VWF (Figure 3-2A-B, red). However, VAMP8 and
VAMP3 are both expressed in a granular pattern that partially overlaps with VWF
(Figure 3-2C-D, red and merge). To quantify the extent of SNARE co-localization with
VWF, we analyzed the confocal images using FV10-ASW 4.0 software (Olympus). The
Pearson’s correlation coefficient for SNAP23 and VWF is only 0.21 ± 0.04, and for
STX4 and VWF is only 0.15 ± 0.02. However, the Pearson’s correlation for VAMP8 and
VWF is 0.49 ± 0.05, for VAMP3 and VWF is 0.40 ± 0.03. Thus the confocal microscopy
data suggests that VAMP3 and VAMP8 co-localize with VWF in endothelial cells.
Although VAMP3 and VAMP8 co-localize with WPBs respectively, these two SNAREs
do not seem to co-localize with each other, as the granular structures that stain positive
for each molecule are distributed in distinct cellular compartments and are minimally
overlapped with a Pearson’s correlation coefficient of 0.27 ± 0.02 (Figure 3-2E).
3.2.3 VAMP8 and VAMP3 interact with the SNAREs STX4 and
SNAP23
SNAREs drive membrane fusion by forming a complex of three interacting SNAREs.
We next sought to determine whether or not VAMP8 or VAMP3 can interact with STX4
or SNAP23, which are target membrane SNAREs expressed in endothelial cells.
Immunoprecipitation of VAMP8 followed by immunoblotting of precipitates reveals that
VAMP8 interacts with STX4 and SNAP23 in endothelial cells (Figure 3-3A). Similar
interaction were detected between VAMP3 and STX4, or VAMP3 and SNAP23 (Figure
3-3B). Conversely, immunoprecipitation of STX4 revealed comparable interactions
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among STX4 with SNAP23, VAMP3 or VAMP8 (Figure 3-3C). These results suggest
that VAMP8 and VAMP3 interact with SNAREs STX4 and SNAP23 in endothelial cells.
3.2.4 VAMP8 and VAMP3 mediates endothelial exocytosis in vitro
To examine the role of VAMP8 and VAMP3 in exocytosis, we knocked down VAMP8
or VAMP3 expression by transfecting HUVEC with control siRNA or siRNA directed
against VAMP8 or VAMP3 (Figure 3-4A). Knockdown of VAMP8 does not affect
expression of other SNARE components or the thrombin receptor (Figure 3-4B). We then
stimulated these transfected HUVEC with histamine or thrombin and measured VWF
release into culture media. Histamine or thrombin increase VWF release in control
siRNA-treated cells, as expected (Figure 3-4 C, E, siControl). However, knockdown of
VAMP3 or VAMP8 decreases VWF exocytosis, and knockdown of both VAMP3 and
VAMP8 further decreases exocytosis (Figure 3-4 C, E). Importantly, knockdown of
VAMP3 or VAMP8 does not affect WPB granule number per cell (Figure 3-5), VWF
protein expression (Figure 3-5), and individual WPB size or shape (Figure 3-6).
Since siRNA directed against VAMP8 might have off-target effects, silencing genes
other than VAMP8, we next performed a rescue experiment. We used siRNA to
knockdown VAMP8 in endothelial cells, and then over-expressed VAMP8 in these cells
(The vector encoding VAMP8 lacks the siRNA target sequence, so the siRNA for
VAMP8 targets endogenous VAMP8 but not exogenous VAMP8). Endothelial cells
normally express VAMP8, siRNA knocks down VAMP8, and transfection of a VAMP8
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vector restores VAMP8 expression (Figure 3-4 D). We then treated these cells with
medium or thrombin and measured exocytosis. Thrombin activates exocytosis in control
transfected cells (Figure 3-4 E, left). Silencing VAMP8 blocks exocytosis (Figure 3-4 E,
middle). Over-expression of VAMP8 restores exocytosis (Figure 3-4 E, right). These
experiments support the hypothesis that VAMP8 mediates endothelial exocytosis.
3.2.5 VAMP8 mediates endothelial exocytosis in vivo
To examine the role of VAMP8 in exocytosis in mice, we examined wild-type mice
(Vamp8 WT) and in Vamp8 null mice (Vamp8 KO). Vamp8 is expressed at highest levels
in lung, pancreas, and spleen; and Vamp8 is expressed at lower levels in the liver and
kidney (Figure 3-7A). Plasma levels of VWF are higher in Vamp8 WT mice than in
Vamp8 KO mice (Figure 3-7B). We then measured exocytosis in vivo using intravital
microscopy. We injected Rhodamine 6G into mice to label their leukocytes, and then
performed intravital microscopy of mesenteric venules to measure leukocyte-endothelial
interactions. At baseline, there are fewer rolling leukocytes in Vamp8 KO mice than in
Vamp8 WT mice (Figure 3-7C-D). After stimulation of exocytosis by superfusing
A23187, an ionophore that induces endothelial exocytosis by directly causing
intracellular calcium surge, the number of rolling leukocyte increases, but the rolling cell
increase is much greater in the wild-type mice than in the knockout mice (Figure 3-7C-
D). To further investigate whether Vamp8 KO mice have defects in VWF exocytosis, we
gave mice subcutaneous epinephrine injection to induce VWF secretion, and collected
blood before and 30 min after epinephrine challenge. In Vamp8 WT mice, VWF levels
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increase significantly in response to epinephrine injection (Figure 3-7E; fold increase of
1.50 ± 0.42, mean ± S.D.) However, Vamp8 KO mice fail to respond significantly to
epinephrine stimulation (Figure 3-7E; fold increase of 1.24 ± 0.36, mean ± S.D.) This
lack of VWF release after epinephrine stimulation is consistent with our in vitro
observation (Figure 3-4 C). Finally, bleeding times are prolonged in Vamp8 KO mice
compared to Vamp8 WT mice (Figure 3-7F).
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Figure 3-1 Expression of SNAREs in endothelial cells.
(A) Differential expression of VAMP8 and VAMP3 in different human endothelial cells
assessed by immunoblotting.
(B) SNAREs expression in mouse brain and HUVEC measured by immunoblotting.
(C) VAMP8 expression in lysates of HUVEC, human platelets, HeLa cells, PC12 cells,
or human VSMC measured by immunoblotting.
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115
Figure 3-2 Subcellular localization of VWF and SNAREs in endothelial cells.
HUVEC was stained with antibodies to VWF and various SNAREs and imaged with
confocal microscopy. Shown are representative images of green (Alexa Fluor 488, left
column), red and blue (Alexa Fluor 594 or Cy3, DAPI, middle column), and merged
(right column) stains highlighting co-localization of the antibody-binding sites (yellow
signals). Images were taken on an Olympus IX81 confocal microscope at room
temperature. Original magnification ×60. The confocal images were analyzed by FV10-
ASW 4.0 software (Olympus). The stained intracellular antigens are:
(A) VWF and Syntaxin 4;
(B) VWF and SNAP23;
(C) VWF and VAMP8;
(D) VWF and VAMP3;
(E) VAMP8 and VAMP3.
VWF co-localizes with VAMP8 with calculated Pearson's coefficient of 0.49 ± 0.05 (C).
VWF co-localizes with VAMP3 with calculated Pearson's coefficient of 0.40 ± 0.03 (D).
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Figure 3-3 Interaction of SNAREs in endothelial cells.
HUVEC lysates were immunoprecipitated (IP) with antibodies to VAMP8, VAMP3,
STX1, or STX4, and immunoblotted (IB) with antibodies to SNARE proteins. Input
represents 2% of total cell lysate; equal amount of lysate were used for all IP lanes. Co-
immunoprecipitation was detected for
(A) VAMP8 with STX4 and SNAP23;
(B) VAMP3 with STX4 and SNAP23; and
(C) STX4 with SNAP23, VAMP3, and VAMP8.
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118
Figure 3-4 VAMP8 and VAMP3 mediate endothelial exocytosis.
HUVEC were transfected with control siRNA (siControl) or siRNA for VAMP3
(siVAMP3) or VAMP8 (siVAMP8), and treated with 10 uM histamine or 1 U/mL
thrombin. The release of VWF into media was measured by ELISA.
(A) Immunoblots show siRNA knockdown of VAMP3 and VAMP8 expression.
(B) Knockdown of VAMP8 does not affect expression of a subset of endothelial SNARE
molecules or the thrombin receptor PAR1, as shown by immunoblotting.
(C) Measurements of VWF release in siRNA-treated cells (n = 4-6 ± S.D. *P < .05 vs.
siControl with histamine). Knockdown of VAMP3, VAMP8, or both, decreases
exocytosis.
(D) HUVEC was transfected with siRNA for endogenous VAMP8 or control siRNA, and
then co-transfected with a vector expressing c-myc tagged VAMP8 (resistant to siRNA).
Immunoblots show siRNA knockdown of endogenous VAMP8 expression (middle lane)
and rescue of VAMP8 expression after knockdown (right lane).
(E) The transfected cells in (C) were treated with thrombin, and the release of VWF was
measured. Knockdown of VAMP8 decreases exocytosis and rescue of VAMP8 restores
endothelial exocytosis (n = 6 ± S.D. *P = 0.004 vs. siControl with thrombin; P = 0.14
between siControl with thrombin vs siVAMP8 + VAMP8-vector with thrombin,
suggesting no significant difference between control cells and VAMP8-rescued cells).
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Figure 3-5 Knockdown of VAMP3 or VAMP8 does not affect endothelial granule
number or VWF content.
(A) Endothelial granules with VWF in HUVEC transfected with siControl, siVAMP3, or
siVAMP8. Shown are stacked confocal images over the entire HUVEC monolayer,
representative of > 5 dishes, with VWF stained with an anti-VWF antibody (green) and
DNA stained with DAPI (blue). Scale bar = 50um.
(B) Knockdown of VAMP3 or VAMP8 does not affect WPB number per cell. HUVEC
was transfected with siControl, siVAMP3, or siVAMP8, cultured for 72 hs, followed by
immunofluorescence staining of WPBs using an anti-VWF antibody. The number of
WPBs were counted over stacks of confocal images that cover the HUVEC monolayer in
multiple culture dishes (siControl, n = 6 dishes ± S.D., 345 cells; siVAMP3, 5 dishes ±
S.D., 347 cells; siVAMP8, 6 dishes ± S.D., 390 cells. NS, not significant.).
(C) HUVEC was transfected with siControl, siVAMP3, siVAMP8, or siVAMP3 and
siVAMP8, cultured for 72 hs, followed by cell lysis. After protein normalization with
BCA assay for each sample, 7.6 ug total cell lysate in 100 ul volume was measured for
VWF content by an ELISA. (n = 4-6 ± S.D. NS, not significant vs. No Treatment).
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Figure 3-6 Knockdown of VAMP3 or VAMP8 does not affect WPB morphology.
(A) The size of WPB granules is not changed by VAMP3 or VAMP8 knockdown.
HUVEC were transfected with siControl, siVAMP3, or siVAMP8, imaged using a
confocal microscope as in Figure S3, and the size of individual WPB was measured over
z-stacks of optical sections by ImagePro Plus (n = 5-6 replicate plates ± S.D.; granules
were counted in over 300 cells per plate. No significant difference among siControl,
siVAMP3, and siVAMP8 by one-way ANOVA).
(B) Knockdown of VAMP3 or VAMP8 does not affect the shape of WPB granules.
HUVEC were transfected with with siControl, siVAMP3, or siVAMP8, imaged using a
confocal microscope as in Figure S3. Optical sections were captured to generate z-stacks
over the entire HUVEC monolayer. The aspect ratio of individual WPB was measured by
ImagePro Plus, and its distribution was plotted as histogram (n = 5-6 replicate plates ±
S.D.; granules were counted in over 300 cells per plate. No significant difference among
siControl, siVAMP3, and siVAMP8 by one-way ANOVA).
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Figure 3-7 Vamp8 mediates endothelial exocytosis and leukocyte rolling in vivo.
(A) Expression of Vamp8 in mice. Immunoblot of Vamp8 in wild-type mice (Vamp8 WT,
top) and Vamp8 knockout mice (Vamp8 KO, bottom) shows Vamp8 expression in lung,
pancreas, and spleen, and lower levels of Vamp8 expression in liver and kidney.
(B) Vamp8 regulates plasma VWF levels in mice. VWF was measured in the plasma of
wild-type mice and Vamp8 null mice. VWF levels are higher in wild-type mice than in
Vamp8 KO mice (n = 11 - 21 ± S.D. *P < .05).
(C) VAMP8 mediates endothelial interactions with leukocytes. Intravital microscopy
was used to measure leukocyte rolling along mesenteric venules in non-treated mice for 2
min and for 8 min after stimulation with the calcium ionophore A23187. Video were
captured with an electron-multiplying CCD video camera (QUANTEM, 512SC) and
were analyzed using Image-Pro Analyzer 6.2 software (Media Cybernetics). Original
magnification ×20. Shown are epresentative videomicrographs of wild-type and Vamp8
KO mice before and after treatment with A23187.
(D) Quantitation of leukocyte rolling from data above (n = 6 ± S.D. *P < .05 for WT vs.
KO). The average number of rolling leukocytes per unit area (512 pixel × 100 pixel)
were counted with Image-Pro Analyzer 6.2 software (Media Cybernetics) for
quantification. Vamp8 mediates endothelial interactions with leukocytes, and absence of
Vamp8 decreases endothelial-leukocyte interactions.
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(E) Plasma VWF levels in wild-type (n = 13) and Vamp8 KO mice (n=15) before and
after epinephrine injection. Values are expressed relative to the level of VWF in the
pooled plasma of 10 wild-type mice and 10 Vamp8 KO mice, respectively. *P < .05 for
WT epinephrine vs. baseline. NS, not significant.
(F) Vamp8 regulates bleeding time. The tail bleeding time in wild-type and Vamp8 KO
mice was measured. Vamp8 KO mice have a prolonged bleeding time compared to wild-
type mice (n = 9 - 14 ± S.D. *P < .05).
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3.3 Discussion
Chapter 3 identifies VAMP8 as a critical component of the machinery that controls
exocytosis of endothelial granules. Endothelial cells express VAMP8, VAMP8 co-
localizes with endothelial granules, and VAMP8 interacts with additional SNAREs in
endothelial cells. Finally, knockdown of VAMP8 inhibits exocytosis of endothelial cells
in vitro; and mice lacking Vamp8 have impaired secretion of VWF in vivo.
Although endothelial cells express a variety of SNAREs that mediate vesicle trafficking,
the set of three SNAREs that regulate endothelial granule fusion with the plasma
membrane is not full characterized, except for that STX4 has been shown to play a role in
endothelial exocytosis (108, 109). Endothelial cells express the SNAP isoform SNAP23
which is likely one of the three components of the SNARE complex (110-113). Others
have shown that endothelial cells express VAMP family members (102-107). But the
precise identity of the v-SNARE that is associated with endothelial granules and mediates
endothelial exocytosis is unclear.
VAMP8 regulates endothelial exocytosis in vitro and in vivo
Our data show that VAMP8 mediates endothelial exocytosis in vitro. VAMP8 is
associated with endothelial granules, and VAMP8 interacts with other endothelial
SNAREs. Knockdown of VAMP8 decreases endothelial release of VWF, and rescue of
VAMP8 expression increases VWF secretion. Furthermore, our in vivo data support a
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role for VAMP8 in mediating exocytosis. Mice lacking Vamp8 have lower levels of
plasma VWF and epinephrine-induced VWF release, and since most plasma VWF and
epinephrine-induced VWF release is from endothelial cells rather than platelets (195),
these data support the idea that Vamp8 mediates endothelial exocytosis in vivo.
Furthermore, mice lacking Vamp8 have a hemostasis defect, supporting the work of
others who show that endothelial derived VWF is a major contributor to hemostasis
(195). Finally, mice lacking Vamp8 have decreased leukocyte trafficking, further
supporting the central role of VAMP8 in regulating endothelial exocytosis.
Although our study supports a major role for VAMP8 regulation of endothelial
exocytosis in vitro and in vivo, one prior study suggests that VAMP8 does not regulate
endothelial exocytosis (102). But several aspects of this prior study are not as robust as
our current data. First, the prior study does not trigger exocytosis with a physiological
agonist applied to intact cells; instead the prior study adds exogenous calcium to
permeabilized cells. Second, the prior study does not demonstrate that VAMP8
expression or function is impaired in experimental assays before measuring exocytosis.
Finally, the prior study does not employ in vivo studies of VAMP8. In contrast, our data
use intact endothelial cells, genetic silencing techniques, and knockout mice in vivo to
show that VAMP8 mediates exocytosis and controls VWF levels.
VAMP8 in Platelet Exocytosis
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Our studies also support the work of others which show that VAMP8 plays a prominent
role in regulating secretion of alpha-granules, platelet granules that resemble endothelial
WPB. Platelets express VAMP8 (370-372, 381, 382), VAMP8 is associated with alpha-
granules (382), platelets from mice lacking VAMP8 secrete less VWF and externalize
less P-selectin than platelets from wild-type mice (371), and mice lacking VAMP8 have
delayed thrombus formation(372). In addition, humans with genetic variation in the gene
encoding VAMP8 can have increased platelet reactivity and an increased rate of arterial
thrombosis (381, 383). However, it is unclear if thrombosis in this subset of patient is
due to abnormalities in endothelial or platelet secretion (384) .
VAMP3 and Endothelial Granule Trafficking
Our data show that VAMP3 regulates endothelial exocytosis in vitro. VAMP3 co-
localizes with granules, interacts with SNAREs, and is required for secretion of VWF.
Our data support the findings of one prior study showing VAMP3 regulates endothelial
exocytosis in vitro (102). However, the function of VAMP3 in vesicle trafficking remains
controversial, since some studies show VAMP3 regulates exocytosis and others show
VAMP3 regulates endocytosis and vesicle recycling. VAMP3 may play one or several
roles in endothelial granule trafficking. VAMP3 might play a role in granule biogenesis
by transporting VWF and other cargo between ER and Golgi or between Golgi and
granule; but this possibility is not supported by our data or data of others (since we find
that endothelial cells contain normal numbers of granules and normal levels of VWF
even after VAMP3 knockdown). VAMP3 might mediate granule exocytosis by
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functioning as a vesicle SNARE on WPBs and interacting with target membranes; data
from our studies and from other studies support this idea in vitro (for example, some in
vitro studies show that VAMP3 controls exocytosis of vesicles from epithelial cells,
macrophages, beta-cells, platelets, oligodendrocytes, and endothelial cells) (102, 370,
385-392). But studies of VAMP3 knockout mice contradict this idea (for example,
platelets from VAMP3 knockout mice have normal alpha-granule release and beta-cells
from VAMP3 knockout mice have normal insulin secretion) (387, 393, 394). VAMP3
might control endocytosis of granule components from the plasma membrane to early
endosomes, retrieving SNAREs and other proteins for further cycles of exocytosis; some
data support this pathway (for example, VAMP3 mediates recycling of integrin-
containing vesicles from plasma membrane to the trans-Golgi network in human cancer
cells)(395, 396).VAMP3 might also modulate vesicle recycling from an early endosome
compartment to the trans-Golgi network; several studies support this suggestion (for
example, VAMP3 mediates transport of the mannose 6-phosphate receptor from
endosome to Golgi)(397). Finally, VAMP3 might regulate VWF trafficking by
controlling endothelial autophagy (316, 398, 399).
Model of Endothelial Granule Release
Our data and the findings of others support a model of exocytosis specific for endothelial
cells. The G-protein Rab27a along with MyRIP and myosin Va targets granules to the
plasma membrane (269, 296, 298, 305, 306, 400).VAMP8 on the granule membrane
interacts with two t-SNAREs on the inner surface of the plasma membrane, potentially
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STX4 and SNAP23. This interaction of the three SNAREs brings the granule and plasma
membrane into apposition. However, complexins may bind to the three SNAREs,
blocking membrane fusion.(356, 401) When an agonist such as thrombin activates the
endothelial cell, intracellular calcium currents rise, synaptotagmin releases the complexin
clamp, the SNARE complex drives fusion of the granule and plasma membrane, and the
contents of the granule are released (356, 400-402) NSF then disassembles the ternary
SNARE complex, and the individual SNARE components are recycled (108, 257, 300-
304, 403).
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SNAP23 Regulates Endothelial Exocytosis
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4.1 Introduction
Among the three types of SNARE molecules that regulate endothelial exocytosis, little is
known about the precise identity of the SNAP homolog. SNAP25 is present almost
exclusively in the brain. In endothelial cells, SNAP25 is not detectable, suggesting that a
homolog of SNAP25 mediates the endothelial SNARE complex (108, 178, 295).
SNAP23, a ubiquitously-expressed homolog of SNAP25, shares 59% identity to
SNAP25. SNAP23 can regulate exocytosis in several distinct cell types. SNAP23 is
localized to the plasma membrane in adipocytes and interacts with multiple syntaxin
isoforms (syntaxin 2, 3, 4, and 5) (404). SNAP23 regulates GLUT4 translocation,
neuroendocrine cell exocytosis, and mast cell degranulation (404-408), suggesting
SNAP23 appears to fulfill the function of SNAP25 in non-neuronal tissues in forming
SNARE complex. SNAP23 has been found in human endothelial cells (102, 108).
SNAP23 interacts with Cav-1 and plays an important role in endothelial caveolae
transcytosis (110). However, studies of the role of SNAP23 in endothelial exocytosis are
limited: partial knockdown of SNAP23 led to a non-significant decrease in exocytosis
(102). Therefore, in an effort to resolve the ambiguities surrounding the function of
SNAP23, we explore the role of SNAP23 in endothelial exocytosis.
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4.2 Results
4.2.1 SNAP23 is expressed in human ECs and murine tissues
We first searched for endothelial expression of SNAP isoforms. Using microarray
hybridization techniques, we found HUVEC express RNA for SNAP23, SNAP25,
SNAP29, SNAP47, and SNAP91 (Figure 4-1A). RNA expression of these homologs
were confirmed by RT-qPCR (Figure 4-1B). Transcription of these SNAP homologs in
HUVEC was further confirmed using ENCODE RNA-Seq data (Figure 4-1C). The
SNAP homolog expressed at highest levels in endothelial cells is SNAP23 (Figure 4-1 A-
B).
We next characterized the expression of SNAP23 in human endothelial cells and murine
tissues by Western blot. We found SNAP23 is expressed in different human endothelial
cell types, including HBMEC, HAEC, and HUVEC (Figure 4-1D). SNAP23 protein is
expressed in murine lung, brain, aorta, pancreas and liver (Figure 4-1E). Our results
extend previous studies showing SNAP23 mRNA is ubiquitously expressed, although its
tissue abundance appears different in mouse and human (409, 410).
4.2.2 SNAP23 regulates endothelial exocytosis
We next explored the role of SNAP23 in endothelial exocytosis. VWF is the major
component of endothelial granules that is released by endothelial exocytosis (184). We
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knocked down the expression of endogenous SNAP23 in HDMVEC and HUVEC,
stimulated the cells to trigger exocytosis, and then measured the amount of VWF released
into the media by an ELISA. Expression of SNAP23 was significantly reduced by siRNA
(Figure 4-2A). The expression of other SNARE proteins in endothelial cells were not
affected by siRNA against SNAP23, including STX4, VAMP3, and VAMP8 (Figure
4-2A). The total intracellular VWF content was also unaffected by SNAP23 knockdown
(Figure 4-2B). We found that knockdown of SNAP23 significantly reduced VWF
exocytosis induced by physiological agonists histamine and thrombin, as well as by the
Ca2+ ionophore A23187 (Figure 4-2C). Knockdown of SNAP23 decreases exocytosis
between by approximately 29% in HDMVEC to 58% in HUVEC (Figure 4-2C). Taken
together, these results suggest SNAP23 is important for Ca2+-dependent endothelial
exocytosis.
4.2.3 SNAP23 is primarily localized on cell membrane in EC
We then studied the subcellular localization of SNAP23 in endothelial cells by confocal
microscopy. We focused on the location of SNAP23 relative to the endothelial granules
called WPBs that contain VWF as well as other pro-thrombotic and pro-inflammatory
compounds. SNAP23 is primarily localized to the plasma membrane (Figure 4-3A).
VWF is localized to the typical cigar-shaped WPB granules (Figure 4-3A). There is no
significant overlap between SNAP23 and VWF staining (Figure 4-3A).
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Most SNAP23 is localized to the plasma membrane, and a minor portion of SNAP23 is
also found in the cytoplasm, consistent with previous studies (411-413).
Immunofluorescent staining on cells cultured at sub-confluent and confluent conditions
showed the subcellular distribution of SNAP23 seems to be dependent on cell
confluence. Fully confluent cells have prominent cell membrane staining of SNAP23 and
less cytoplasmic SNAP23, whereas cells at sub-confluent condition showed more
SNAP23 in the cortical region and the cytoplasm (Figure 4-3A). To confirm this
observation, we cultured HUVEC at sub-confluent and confluent conditions and isolated
cytosol and membrane fractions, and immunoblotted fractions for SNAP23. SNAP23 is
mostly found on the membrane fraction of subconfluent cells and confluent cells (Figure
4-3B). A significant amount of SNAP23 was also detected in the cytosol at sub-
confluent conditions but less SNAP23 in the cytosol of confluent cells (Figure 4-3B).
Taken together, these data suggest SNAP23 is primarily localized on the plasma
membrane in endothelial cells, and its cytosolic distribution depends on cell confluence.
4.2.4 SNAP23 interacts with endothelial exocytic machinery
Since SNAP23 is important for endothelial exocytosis, we next searched for a link
between SNAP23 and the endothelial exocytic machinery. In order to search for the
interaction partners of SNAP23, we first performed sucrose density gradient
fractionation. We separated HUVEC lysates through a 5% - 30% - 40% discontinuous
sucrose gradient, and probed 17 fractions for SNARE proteins involved in endothelial
exocytosis. SNAP23 co-sediments with STX4 in fractions 3 to 7 and 15 to P (Figure
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4-4A). SNAP23 partially co-sediments with VAMP3 and VAMP8 (Figure 4-4A). The
sucrose density gradient fractionation provided indirect evidence that SNAP23 may
interact with endothelial SNARE molecules. To confirm their interaction in a complex,
we immunoprecipitated HUVEC lysates with antibody to SNAP23 or isotype IgG, and
then probed precipitants for SNARE proteins. STX4, VAMP3, and VAMP8 were all
detectible in the precipitant, in resting and stimulated condition (Figure 4-4B). Taken
together, these data suggest SNAP23 interacts in a complex with components of the
endothelial exocytic machinery containing STX4, VAMP3, and VAMP8, both at resting
and stimulated conditions.
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Figure 4-1 SNAP23 is expressed in human endothelial cells and murine tissues.
(A) Heat map of the gene expression values for SNAP homologs in 6 samples of HUVEC
by microarray. Relative expression values were normalized to that of GAPDH.
(B) Relative expression of SNAP homologs in 3 individual donors of HUVEC as assayed
by RT-qPCR. Expression was normalized as percentage of SNAP23 expression.
(C) ENCODE data on the UCSC genome browser depicting the expression of HUVEC
SNAP homologs as assayed by RNA-seq.
(D) SNAP23 is expressed in HBMEC, HAEC, and HUVEC as measured by Western
blot.
(E) SNAP23 is expressed in multiple murine tissues as measured by Western blot. Data
are represented as mean ± SD. See also Table 1.
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Figure 4-2 SNAP23 is important for endothelial exocytosis.
(A) siRNA directed against SNAP23 (siSNAP23) knocks down SNAP23 protein levels
as measured by Western blot. The siRNA against SNAP23 has no effect on the
expression of other SNARE proteins including STX4, VAMP3, and VAMP8. GAPDH
was used as loading control.
(B) SNAP23 knockdown does not affect VWF expression in HUVEC. Total VWF
content was measured by an ELISA in control siRNA (siControl) and siSNAP23 treated
cells (n = 6; NS, non-significant).
(C) SNAP23 knockdown decreases endothelial exocytosis. HDMVEC and HUVEC were
treated with siControl or siSNAP23, stimulated with serum-free medium only (resting),
or 10 μM histamine, or 1 U/ml thrombin, or 10 μM Ca2+ ionophore A23187 for 30 min;
and then VWF released into the media was measured by an ELISA (n = 4 – 7; * P < 0.05
vs. siControl; NS, non-significant vs. siControl). Data are represented as mean ± SD.
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Figure 4-3 Subcellular localization of SNAP23 in endothelial cells.
(A) Subcellular localization of SNAP23 in sub-confluent (upper panel) and confluent
(lower panel) HUVEC. Immunofluorescent staining was performed on HUVEC with
antibodies against SNAP23 (red) and VWF (green), DNA was stained with DAPI (blue),
and the cells were imaged by confocal microscopy (objective 60× oil, scale bar = 50 μm,
confocal z resolution = 0.32 μm).
(B) Western blot analysis of cell fractions from sub-confluent and confluent HUVEC
using markers for membrane (Caveolin-1) and cytosol (GAPDH). These fractions were
also probed for SNAP23. Whole cell lysates were used for total protein. SNAP23 is
depleted from cytosol fraction when reaching confluence.
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Figure 4-4 SNAP23 interacts with endothelial exocytic machinery.
(A) SNAP23 co-sediments with STX4, VAMP3 and VAMP8 by sucrose density gradient
fractionation. HUVEC lysate was ultracentrifuged through a 5% - 40% discontinuous
sucrose gradient, and then the gradient was aliquoted into 17 fractions and analyzed by
SDS-PAGE (T, total proteins in the lysate; P, pellet after fractionation). β-actin was used
as control for fraction separation. Representative of 3 separate experiments.
(B) SNAP23 co-precipitates with STX4, VAMP3 and VAMP8. HUVEC stimulated with
serum-free medium only (Rest), or 10 μM histamine, or 10 μM Ca2+ ionophore A23187
for 30 min were immunoprecipitated with antibody to SNAP23 or isotype IgG. The
precipitants were probed with antibody to STX4, VAMP3 and VAMP8. Input represents
5% total protein. Representative of 3 similar experiments.
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4.3 Discussion
The identity of the SNAP homolog which functions as an endothelial t-SNARE is
unclear. In this study, we found that SNAP23 is the most highly expressed SNAP isoform
in different types of human endothelial cells; SNAP23 is localized to the endothelial cell
membrane; SNAP23 forms complexes with other endothelial SNARE molecules; and
most importantly, SNAP23 deficiency impairs endothelial exocytosis. These results
collectively suggest that SNAP23 plays a critical role in regulating endothelial cell
exocytosis.
Our work and the studies of others show that human endothelial cells express a
distinctive subset of SNARE molecules (102, 108, 178) Endothelial cells express
VAMP3 and VAMP8 of the v-SNARE family, STX4 of the syntaxin family, and several
SNAP isoforms including SNAP23. These results demonstrate that endothelial cells
contain family members of exocytic machinery also found in neurons and yeast. We also
found that a subset of endothelial SNAREs interact with each other: endothelial cells
contain SNARE complexes consisting of SNAP23, STX4, and VAMP3 or VAMP8
(Figure 4-4). This SNARE complex corresponds to SNARE complexes found in
neurons, composed of SNAP25, STX1a, and VAMP2 (133, 414). We also found that
SNAP23 co-sediments with STX4 and VAMP3 and VAMP8 (Figure 4-4A). Our work
supports the studies of others that show SNAP23 and STX4 form clusters in endothelial
cells, and the results of others showing SNAP25 and STX4 form clusters in
neuroendocrine cells (415, 416).
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In resting cells, SNAP23 interacts with STX4, VAMP3, and VAMP8 (Figure 4-4).
Stimulation of endothelial cells with histamine or calcium ionophore increases the
interaction of SNAP23 with STX4, VAMP3, and VAMP8 (Figure 4-4B). These data
support the idea that SNAP23 functions as one component of the ternary SNARE
complex in endothelial cells. Our work partially contrasts with the work of others (102),
who show that SNAP23 is localized to plasma membrane but has little effect on
endothelial exocytosis. One possible explanation for this discrepancy is that knockdown
of SNAP23 expression was incomplete in other studies (102).
Among the members of the SNAP family, only two isoforms have been regarded as
critical components in exocytosis, SNAP25 and SNAP23. SNAP25 is expressed in
neuronal and neuroendocrine tissues, whereas SNAP23 is ubiquitously expressed in non-
neuronal cells (409). Endothelial cells express lower levels of other SNAP isoforms
(Figure 4-1A-C).
We show that SNAP23 is localized to endothelial plasma membranes (Figure 4-3). Our
data suggest that SNAP23 plays a role similar to SNAP25 in neurons, serving as a t-
SNARE. However, a minor fraction of SNAP23 was also found in the cytosol (Figure
4-3), similar to previous studies (110). It has been previously confirmed that the plasma
membrane localization of SNAP family proteins depends on the palmitoylation of a
cysteine-rich domain (417). It is plausible that a sub-fraction of SNAP23 proteins is
detectable in the cytosol before palmitoylation occurs. Cytosolic levels of SNAP23 are
decreased in confluent cells (Figure 4-3B), suggesting that palmitoylation of SNAP23
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may be dynamically regulated and is not constitutive. In addition, SNAP homologs were
found in the cytoplasm as intracellular t-SNARE mediating intracellular membrane
fusion, such a possibility for SNAP23 cannot be ruled out (418).
Our data show SNAP23 is expressed in multiple murine tissues, confirming previous
observations (409, 410, 419). We and others found SNAP23 is most highly expressed in
the murine lung (Fig. 1E) (419). Why is SNAP23 highly expressed in the lung? One
explanation is SNAP23 is important for epithelial cell or pneumocyte exocytosis. Prior
studies have shown that regulated mucin secretion from airway epithelial cells requires
SNAP23 (420). Syntaxin 2 and SNAP23 are important for regulated surfactant secretion
from alveolar type II cells (421). SNAP23 is enriched in lipid rafts in type II alveolar
cells which serves as a functional platform for surfactant secretion (422). SNAP23 also
interacts with annexin A7 in a protein kinase C (PKC)-dependent manner which
facilitates membrane fusion during surfactant secretion (423). Another possibility is
SNAP23 is involved in the endothelial exocytosis of the lung vasculature. Still another
possibility is SNAP23 is important for the response of immune cells in the lung. SNAP23
is important for phagosome formation and maturation in macrophages (424), mast cell
degranulation (363, 405), neutrophil exocytosis (425), secretion of antibodies by human
plasma cells (426), and is found on the plasma membrane of several different human
inflammatory cells including eosinophils, basophils, neutrophils, and peripheral blood
mononuclear cells (427, 428). Interestingly, SNAP23 can be induced by various cytokines
as an immediate-early cytokine responsive gene in a murine myelomonocytic cell line.
Thus SNAP23 might be expressed by one or more cell types in the lung.
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In summary, in Chapter 4, we have shown that human endothelial cells express SNAP23,
SNAP23 interacts with other endothelial SNARE, and SNAP23 plays a crucial role in
endothelial exocytosis of VWF. This matches a model in which a three membered
SNARE complex forms before endothelial exocytosis, consisting of a VAMP on the
membrane of endothelial granules, along with STX4 and SNAP23 on the plasma
membrane (133, 361, 414).
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Conclusions and Perspectives
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5.1 Summary
EC mediates vascular injury response by exocytosis. WPBs are EC-specific granules for
exocytosis. EC exocytosis release VWF, a major initiator in thrombogenesis, and P-
selectin, which mediates vascular inflammation.
STXBP5 is a novel candidate gene linked to changes in plasma vWF levels by GWAS. In
Chapter 2, we found STXBP5 is expressed in ECs and interacts with SNARE proteins.
STXBP5 inhibits endothelial exocytosis in vitro and in vivo. However, despite higher
plasma VWF levels, Stxbp5 KO mice displayed impaired hemostasis and thrombosis, and
Stxbp5 KO platelets had severe defects in secretion and activation. These results suggest
STXBP5 inhibits endothelial exocytosis, but promotes platelet secretion and thrombosis.
STXBP5 is a novel SNARE regulatory partner with dual regulatory functions.
Human ECs express SNARE proteins, including STX, VAMP and SNAP homologs. But
the identities of endothelial SNAREs have been unclear. In Chapter 3, we found VAMP8,
in addition to VAMP3, plays a critical role in endothelial exocytosis. VAMP8 co-
localized with WPBs. VAMP8 interacted with other EC SNAREs. Knock down of
VAMP8 or VAMP3 expression inhibited endothelial exocytosis. More importantly,
Vamp8 KO mice had decreased endothelial exocytosis and hemostasis.
In Chapter 4, we found SNAP23 is another key component of the endothelial SNARE
machinery. We identified SNAP23 as the most abundant endothelial SNAP homolog that
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is important for endothelial exocytosis. SNAP23 was found on endothelial cell
membrane. Knockdown of SNAP23 decreased endothelial exocytosis. SNAP23 also
interacted with other important endothelial SNARE proteins.
In summary, the present work investigated the role of STXBP5, VAMP8, and SNAP23 in
endothelial exocytosis, provided functional relevance for a novel genetic risk factor
identified by GWAS, and improved our understanding on the regulatory mechanism of
endothelial exocytosis.
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5.2 Working model for endothelial exocytosis
Our current work identified a novel regulator of endothelial exocytosis, STXBP5, and
explored the functional implication of its genetic variants on plasma VWF levels and
thrombosis. Our work also identified important endothelial SNARE members and
characterized their role in endothelial exocytosis and thrombosis. These findings
provided a novel mechanistic view of the endothelial cell exocytic machinery.
We propose a working model that integrates the major elements examined by this project
(Scheme 5-1)
In this simplified model, the trafficking and fusion of WPBs is mediated by at least four
components. Chapter 3 and 4 have suggested that the components of the exocytic
machinery include VAMP8 or VAMP3 as the v-SNARE, and STX4 and SNAP23 as the
t-SNAREs. Cognate SNAREs interact to form a trans-SNARE complex. Formation of
this trans-SNARE complex tethers WPB to the endothelial cell membrane. Presumably,
intracellular Ca2+ elevation activates the Ca2+ sensor SYT1, which triggers the trans-
SNARE complex to change its conformation into a cis-SNARE complex, resulting in
membrane fusion and VWF release (Scheme 5-1A).
Chapter 2 identified a novel regulator of exocytosis, STXBP5. We propose that STXBP5
acts as a brake on the endothelial exocytic machinery through two potential mechanisms
(Scheme 5-1B-C):
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1) STXBP5 may interact with STX4, sequestering STX4 from the other SNAREs,
blocking formation of the trans-SNARE complex (Scheme 5-1B, marked as “1”).
2) STXBP5 may interact with the Ca2+ sensor SYT1, blunt Ca2+ signaling, resulting in
less Ca2+ sensitive vesicles and less VWF release (Scheme 5-1C, marked as “2”).
Either or both of these mechanisms are possible. Each of these steps will limit vesicle
fusion with the cell membrane and reduce VWF exocytosis.
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Scheme 5-1 Working model of endothelial exocytosis regulation.
(A) The endothelial exocytic machinery is presumably composed of t-SNAREs and v-
SNARE. In this case, VAMP8 represents the v-SNARE whereas STX4 and SNAP23
represents the t-SNAREs. WPB docks to the cell membrane as a result of trans-SNARE
complexes formation. SYT1 triggers trans-SNARE “zippering” in response to
intracellular Ca2+, causing vesicle fusion and VWF exocytosis.
(B) STXBP5 inhibits endothelial exocytosis by interacting with STX4. STXBP5 binds to
STX4 and sequesters it, so that less t-SNARE is available for productive trans-SNARE
complex formation.
(C) STXBP5 inhibits endothelial exocytosis by interacting with SYT1. Occupancy of
SYT1 by STXBP5 blocks its Ca2+ sensing. The effect of Ca2+ to bind to SYT1 and trigger
exocytosis is therefore dampened.
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5.3 Future directions
Although our data demonstrated the importance of several novel regulators of endothelial
exocytosis, we have only begun to understand the intricate regulatory mechanisms in
endothelial cells. Many questions about the regulation of endothelial exocytosis remain to
be answered.
5.3.1 STXBP5 SNP and VWF
The most fascinating question is how genetic variation in STXBP5 affects STXBP5
function and plasma VWF levels.
The SNP with the highest genome-wide significance level for plasma VWF in the meta-
analysis of the CHARGE Consortium data is rs9390459, which is a synonymous
variation (317). The top non-synonymous variation, rs1039084 (encoding STXBP5-
N436S), is in high pairwise LD with rs9390459 (338). Is the top candidate rs9390459 a
proxy variation for rs1039084? Our data seem to support this prediction, because
although rs9390459 is synonymous, the STXBP5-N436S variant does seem to inhibit
VWF exocytosis more potently than wild-type STXBP5 (in this case, both the major and
minor allele of rs9390459 encode the same amino acid). Although the technical difficulty
in platelet transgenic studies prevented us from observing a phenotype in platelets using
the STXBP5-N436S mutant, our data (Figure 2-9) is consistent with the clinical data
showing the minor allele G of rs1039084 (corresponding to 436 serine) is associated with
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lower plasma VWF, decreased DVT risk, and higher bleeding score (319, 339). This
consistency strongly suggests genetic variation in STXBP5 is a novel risk factor for
plasma VWF level and thrombotic diseases, and warrants further mechanistic
investigation.
The asparagine to serine substitution at STXBP5 436 position may change its interaction
partners or subject it to additional post-translational modification. It has been shown that
STXBP5 is phosphorylated by PKA at serine-724 (located in the hyper-variable linker
region between the N-terminal WD40 repeats and the C-terminal VAMP-like domain),
which reduces its interaction with STX1 and increases SNARE complex formation (336).
The N436S substitution is located in one of the WD40 repeat domains. It is unknown
whether this position is involved in STXBP5 interaction with other proteins or is a
potential target for post-translational modification such as phosphorylation.
One approach to answer this question is to use genome editing techniques in cells and in
mice. The rapid developing field of genome-editing technology has provided
investigators precise and high-efficient tools, such as zinc finger nucleases (ZFNs),
transcription activator-like effector nucleases (TALENs), and clustered regularly
interspaced short palindromic repeats (CRISPR)/CRISPR-associated 9 (Cas9) (429, 430).
We have started to use the CRISPR/Cas9 system to introduce a single nucleotide variant
(SNV) into the genomic DNA encoding STXBP5 in human EC to see if changing one
nucleotide, such as rs1039084 (A>G), is able to produce a phenotypic change.
Considering human and mouse STXBP5 share 98% identity in protein sequence, we have
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also successfully generated mice containing the human SNP rs1039084 (Stxbp5-N437S
mice) using the CRISPR/Cas9 system (431, 432). We propose to characterize the
phenotype of endothelial exocytosis, platelet function, hemostasis and thrombosis in
these cell and mouse models. Moreover, the novel gene editing technology has also
enabled us to elucidate the potential effects of other genetic variants in STXBP5 as well
as in many other genetic loci.
5.3.2 STXBP5 in ECs and in platelets
Another intriguing question is how STXBP5 can display a dual positive and negative role
in exocytosis.
We initially hypothesized STXBP5 might produce different splice variants in ECs and
platelets, but qPCR shows the splice variant profiles were similar in ECs and platelets
(Figure 2-1D).
Another possible explanation is STXBP5 may have different interaction partners in ECs
and platelets. Ye et al. shows that in platelets STXBP5 interacts with STX11 and
SNAP23, and to a lesser extent with STX2 and STX4; STXBP5 also interacts with the
platelet cytoskeleton (433). Our preliminary co-immunoprecipitation experiment shows,
however, in HUVEC the interaction between STXBP5 and STX11 or SNAP23 were not
detectible (data not shown). It has been shown that STX11 is required for platelet
secretion (367), so one would expect that binding of STXBP5 may presumably block
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productive SNARE complex formation. However, it is conceivable that platelet syntaxin
homologs may display different binding affinities or kinetics for STXBP5, so that their
interaction does not lead to “dead-end” SNARE complex formation, but instead
facilitates the assembly of SNARE complexes. For example, STXBP5 perhaps binds to t-
SNAREs, ensure their proper localization, or orchestrates the arrangements of SNARE-
complex (with or without participation of platelet cytoskeleton). Since STXBP5 interacts
with both syntaxins and SNAP23 in platelets, it is also possible that STXBP5 may
enhance the efficiency of syntaxin-SNAP23 or SNARE-Munc18 complex clustering
(415, 416), which in neurons facilitate SNARE-mediated vesicle fusion.
An interesting observation is that the paradoxical regulation by STXBP5 in exocytosis
closely resembles that of complexin. In neurons, complexin facilitates Ca2+-triggered fast,
synchronized synaptic release, but inhibits spontaneous fusion. Unfortunately, although
several models for the role of complexin have been proposed, none has been
experimentally tested (434). One would speculate that STXBP5 and complexin to share
some common properties. In fact, these two protein do have quite a number of common
features. They are both conserved from yeast to human. They both carry a coiled α-helix
that binds to syntaxin (complexin helix inserts into the groove between VAMP and
syntaxin helices on trans-SNARE complex; STXBP5 binds to syntaxin and displaces
VAMP with its VAMP-like domain). They both interact with synaptotagmin 1 and
regulate Ca2+-triggered exocytosis.
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Unlike STXBP5, the crystal structure of complexin has been well documented (148). The
crystal structure shows complexin stabilizes the fully assembled SNARE complex, which
is essential for Ca2+-triggered fast, synchronized exocytosis. Complexin regulation of
fusion depends on its concentration (435). But how its concentration is controlled
remains unknown. Could STXBP5 behave just like complexin, where its concentration
dictates its inhibition or promotion of exocytosis? Or does it compete with complexin in
binding SNAREs and/or synaptotagmin? Given the significant similarities between
complexin and STXBP5, the above two possibilities are not mutually exclusive.
Note complexin only binds to assembled SNARE complex which is on the cell
membrane; in contrast STXBP5 is primarily located in the cytoplasm (Figure 2-10),
although a subset of STXBP5 is also found on the cell membrane via interaction with
syntaxin (Figure 2-11C) (350). Like other nonintegral membrane proteins, it is postulated
that STXBP5 is in equilibrium between cytosolic and membrane pools in certain cell
types. The synaptic protein unbinding rate of STXBP5 (0.139/s) (350) is slower than the
rate of complexin's off rate from the SNARE complex (0.3/s) (436, 437), suggesting a
higher affinity of STXBP5 with SNARE proteins. Could it be that STXBP5 acts as a
cytoplasmic (and to a lesser extent membranous) “buffer” for un-docked or recycled
SNARE protein or synaptotagmin, whereas complexin binds SNAREs more weakly and
only on the cell membrane, and it is the stochastic ratio of their relative abundance that
determines the vesicle fusion probability? Perhaps further studies on the STXBP5 domain
structure and functions, including its crystal structure, would eventually help answer
these questions.
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5.4 Significance and perspectives
Nearly a century has passed since Finnish physician Erik von Willebrand first described
the VWD case. Today our knowledge on the pathogenesis of hemorrhagic and thrombotic
disorders has greatly improved, but our diagnosis and therapy is still far from adequate
(238, 438).
VTE is a multifactorial disorder. However, a substantial proportion of VTE patients
present with no detectable acquired risk factors of thrombosis, suggesting that genetic
factors might be responsible for certain group of individuals. GWAS have revolutionized
the discovery of genomic factors associated with complex disease traits. Our study on the
role of STXBP5 served as a promising case which a new potentially targetable genetic
risk factor for VTE is identified by GWAS and is further validated by cell and animal
models. Our study on the role of VAMP8 and SNAP23, in addition, further
complemented the framework of the endothelial exocytosis machinery and how it can
affect hemostasis and thrombosis.
GWAS-based findings will continue to accelerate the discovery of translatable clinical
predictive or prognostic targets. Novel genome-editing techniques, such as
CRISPR/Cas9, offered remarkable technical possibilities to mine the mass of GWAS data
for additional trait associations, and to model and characterize genetic risk factors for
thromboembolic diseases. While the road to apply these techniques to treating diseases is
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long, our current work opened the door to further important research that will usher in
safer and more effective therapies for both bleeding and thrombotic diseases.
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Materials and Methods
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Antibodies and reagents. Primary and secondary antibodies used for this study are listed
in Table 2. Lipofectin reagent and Opti-MEM I media were purchased from Invitrogen.
Protease Inhibitor Cocktail (Complete Mini EDTA-free) was purchased from Roche.
Calcium ionophore A23187, FeCl3, histamine, thrombin, epinephrine and ATP were
purchased from Sigma-Aldrich. Thrombin was purchased from Enzyme Research
Laboratories. The cDNA for VAMP8 was cloned into a pCMV-Myc vector (Clontech).
Individual siRNA was from Ambion (Life Technologies). The sequence for siSTXBP5 is
5’-AGTGGGAACTCAGACTGGTGCTTTA-3’. The sequence for siVAMP3 is: 5’-
GCCACTGGCAGTAATCGAAGA-3’. The sequence for siVAMP8 is: 5'-
GAGGAAAUGAUCGUGUGCGGAACCU-3'. The siRNA sequence for siSNAP23 is
5’-GACACCAACAGAGAUCGUAUUGAUA-3’.
Cell culture and transfection. HUVEC, HAEC, HDMVEC, and endothelial cell culture
medium containing endothelial cell growth supplement (VascuLife EnGS) were obtained
from Lifeline Cell Technology. Endothelial cells were maintained collagen I coated
plates as described (439), and passages 3-6 were used for all in vitro studies. HL-60
promyelocytic cell line was purchased from American Type Culture Collection. Human
platelets were isolated from whole blood of healthy donors free of non-steroidal anti-
inflammatory drugs for at least 10 days as previously described (301). Transfection was
performed at 80-90% confluence using Lipofectin reagent following the manufacturer’s
protocol, with either 0.5 μg/ml plasmid DNA or 20 nM siRNA oligonucleotides for 5
hours. Cells were stimulated or imaged at least 72 h after transfection. Cell confluence
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was visually determined when cells were in contact and the entire culture surface had no
visible space among individual cells for at least 48 h.
VWF release assay. Confluent HUVEC, HAEC, or HDMVEC were stimulated with 10
μM histamine, or 10 μM A23187, or 10 μM ATP, or sham treatment (PBS) for 30 min at
37°C to stimulate VWF release. After stimulation of the cells, VWF concentration in the
cell media was quantified by a VWF ELISA kit (American Diagnostica Inc. and Sekisui
Diagnostics, LLC). In separated experiments, HUVEC transfected with control- or
STXBP5-siRNA were lysed without stimulation and intracellular VWF contents were
measured in equal amount of protein lysate. We found the antibody to VWF in this
ELISA cross-reacts with mouse VWF antigen. Separate measurements were performed in
diluted mouse plasma collected by retro-orbital bleeding from Stxbp5 WT mice and
Stxbp5 KO littermates.
Endothelial cell-leukocyte interaction. Adhesion of leukocytes to HUVEC was performed
as described (308, 314, 315). HUVEC were transfected with control siRNA or siRNA
against STXBP5 in a 12-well plate and were then cultured until 100% confluent. Prior to
the adhesion assay, HL-60 cells were labeled with 2 μM Calcein AM (Molecular Probes)
for 30 min at 37°C, washed, and suspended in serum-free media. Then 1×105 HL-60 in
500 μl serum-free media was added to each well in the presence or absence of 10 μM
histamine. The plate was kept at 4°C for 15 min, washed twice with HBSS, refilled with
media, and immediately imaged with an Olympus BX51 upright microscope with a 10×
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water immersion objective. Cells pre-incubated with an azide-free antibody raised against
full-length P-selectin (AK4) was used as negative control.
ADAMTS13 assay. We used an assay to measure ADAMTS13 in murine plasma (440).
Dr. Neelamegham (Department of Chemical and Biological Engineering, State
University of New York at Buffalo, Buffalo, NY 14260, USA) generously provided a
recombinant polypeptide, XS-VWF, which contains a truncated fragment of the VWF-A2
domain cleavable by ADAMTS13. This polypeptide is flanked by a CFP variant
Cerulean and an YFP variant Venus that exhibit fluorescence/Förster resonance energy
transfer (FRET) properties. ADAMTS13 cleavage of XS-VWF keeps the fluophores
apart, resulting in an increase of Cerulean emission and a decrease of FRET efficiency.
ADAMTS13 activity is quantified by the FRET ratio, defined as the ratio of Cerulean
emission versus Venus emission at 420 nm. We found mouse plasma efficiently cleaves
XS-VWF. In this assay, 15 μl WT or Stxbp5 KO mouse plasma were incubated with 10 μl
XS-VWF substrate in 100 μl cleavage buffer (50 mM Tris, pH 8.0 and 12.5 mM CaCl2)
in a 96-well plate for 1 h at 37°C. The FRET ratio was measured on a plate reader and
ADAMTS13 activity was calculated as percentage of pooled WT mouse plasma.
Western blotting. Western blotting was performed as described previously (108). In brief,
endothelial cells or platelets were lysed with Laemmli sample buffer (Bio-Rad), boiled
for 5 min at 95°C, resolved on 4-20% Mini-PROTEAN TGX Precast gels (Bio-Rad), and
transferred using a Trans-Blot SD semi-dry electrophoretic transfer unit (Bio-Rad) onto
nitrocellulose membranes. After 1-hour blocking with 5% non-fat milk in PBS containing
167
0.05% Tween 20 at room temperature, the membranes were hybridized with primary
antibodies followed by HRP-conjugated secondary antibodies and enhanced
chemiluminescence detection using X-ray films.
Quantitative real-time PCR. Murine tissue was harvested immediately after transcardial
perfusion with diethylpyrocarbonate-treated PBS. Total RNA was isolated with Trizol
following the manufacture’s protocol (Invitrogen) and purified with lithium chloride
(Sigma-Aldrich). The A260/A280 ratio of all samples were between1.9-2.1 as measured
by spectrophotometry (NanoDrop; Thermo Scientific). cDNA was synthesized using an
iScript™ cDNA Synthesis kit (Bio-Rad). Quantitative real-time PCR was performed by
Taqman gene expression assay (Applied Biosystems) for 40 cycles on an iCycler thermal
cycler equipped with MyiQ PCR detection system (Bio-Rad). Three independent
experiments were performed for each organ, and in each experiment TaqMan
quantification was repeated in triplicate for each sample. Taqman probes were purchased
from Applied Biosystems. For each probe set, calibration curves were generated by 10-
fold serial dilutions of cDNA to ensure comparable PCR efficiency. Expression results
were calculated by ΔΔCT method and were normalized to the reference genes GAPDH or
Gapdh. SNAP homolog expression was quantified by ΔΔCt method using four reference
genes: B2M, GAPDH, HRPT1, and YWHAZ, and expressed as percentage relative to the
amount of SNAP23.
Identification of STXBP5 mRNA splice variants. Primers flanking the splice region of
human STXBP5 mRNAs were designed (5’-TCGCTGCAAATCTCCAACCT-3’ and 5’-
168
GGACGAGTCCGTCTTTCGAG-3’). HUVEC, human platelets, and HEK293 cDNA
were synthesized as described above. Human adult brain cDNA was purchased from
Biochain. PCR was performed using 0.25 μg cDNA per reaction with the above primers.
The products were separated by agarose gel, sequenced, and compared with validated and
predicted NCBI Reference Sequences. Five NCBI sequences were found matching the
sequencing results: NM_001127715.2, NM_139244.4, XR_245502.1, XR_245503.1, and
XR_245504.1. The quantification of each splice variant were performed using custom
Taqman probes, each spans exon boundaries specific for one splice variant
(NM_001127715.2, exons 21-23; NM_139244.4, exons 19-23; XR_245502.1, exons 21-
22; XR_245503.1, exons 20-22; XR_245504.1, exons 19-22).
Site-directed mutagenesis. Mouse pMEX neo-3×FLAG-tomosyn (8)-ATG expression
plasmid was a kind gift from Dr. David James (Garvan Institute of Medical Research,
Darlinghurst, NSW 2010, Australia) (354). A point mutation corresponding to human
SNP rs1039084 (A→G) was introduced by site-directed mutagenesis with a QuikChange
Mutagenesis Kit (Agilent Technologies). Using the plasmid as template, PCR was
performed with the following primers for mutagenesis: 5’-
AAAGGAATGGCCCATCAGCGGAGGTAATTGGGGCTT-3’ and 5’-
AAGCCCCAATTACCTCCGCTGATGGGCCATTCCTTT-3’. PCR products were
treated by DpnI to digest template plasmid and then transformed into competent cells.
The mutated plasmid was extracted using EndoFree Plasmid Maxi Kit (Qiagen) and
mutation confirmed by DNA sequencing.
169
Co-immunoprecipitation. HUVEC was lysed with coimmunoprecipitation buffer
containing 10 mM HEPES/NaOH, pH 7.4, 140 mM NaCl, 5 mM NaHCO3, 1.2 mM
NaH2PO4, 1 mM MgCl2, 10 mM glucose, 5 mM KCl, 1 mM EGTA, 1.3 mM CaCl2, and
1% (w/v) Triton X-100 supplemented with protease inhibitors cocktails on ice and
incubated for 60 min followed by centrifugation at 161,000 × g at 4 °C for 20 min. 25 μl
Protein A/G PLUS-Agarose was incubated with 2 μg antibodies or mouse IgG1 at 4 °C
overnight, washed, and then mixed with the pre-cleared cell lysate at 4 °C overnight. The
precipitants were washed with cold co-immunoprecipitation buffer 6 times and the bound
proteins were then solubilized in reducing Laemmli sample buffer at 95 °C. The elute
was resolved by SDS-PAGE followed by immunoblotting.
Confocal microscopy. HUVEC was cultured on collagen-coated 35-mm glass dishes
(MatTek). After removal of media, cells were rinsed twice with PBS and fixed
immediately with 4% PFA for 20 min. After permeabilization with 0.15% Triton X-100
for 10 min and blocking with 5% donkey serum for 1 hour at room temperature, the cells
were incubated with primary antibodies (1:100-1:330 dilution) overnight at 4 °C followed
by 1-hour incubation at room temperature with 1:2000 fluorescent secondary antibodies
and DAPI. Fluorescent microscopy was performed using an IX81 inverted confocal
microscope equipped with a high sensitivity digital camera (FV1000, Olympus). All
fluorescent images were generated using sequential line-scanning with identical settings
in each experiment at maximum z-resolution (0.40 μm for 40× objective, 0.32 μm for 60×
objective). Image analysis was carried out using FV10-ASW 3.0 (Olympus) and Image-
Pro Plus (Media Cybernetics).
170
Sucrose density gradient ultracentrifugation. HUVEC were lysed on ice with 2 ml co-
immunoprecipitation buffer supplemented with protease inhibitors cocktails and
incubated for 1 hour at 4 °C followed by centrifugation at 161,000 × g for 20 min. The
supernatant was carefully loaded on top of a discontinuous sucrose density gradient of 1
ml 5% sucrose, 6 ml 30% sucrose, and 3 ml 40% sucrose in coimmunoprecipitation
buffer (from top to bottom) in a 14 ml PET thin walled tube (Thermo Scientific). The
sucrose density gradient was then ultracentrifuged at 166,880 × g for 20 hours at 4°C on a
Discovery 100s ultracentrifugation equipped with a SureSpin 630/17 swinging-bucket
rotor (Sorvall). After ultracentrifugation, 18 equal-volume aliquots were carefully
aspirated from top to bottom, boiled in equal-volume 2× sample loading buffer, and
analyzed by 7.5% SDS-PAGE.
Microarray. Total RNA was extracted from HUVEC from six different donors by
RNeasy mini kit (Qiagen) according to the manufacturer’s protocol. The gene expression
profiling was performed using Affymetrix GeneChip at the Genomics Research Center at
University of Rochester.
Transcriptional profile by ENCODE. The Feb 2009 GRCh37/hg19 Assembly was
searched for transcription levels of SNAP homologs. Transcriptional profiles were
visualized in UCSC Genome Browser with a customized ENCODE track for HUVEC.
Stxbp5 mice management. Stxbp5 KO mice on a background of 129Sv: C57BL6: DBA/2
= 2: 1: 1 were generated as previously described (332). PCR was used for genotyping
171
with primers specific for a wild-type allele (5’-TTCTGCTCCCCGCTGCTCCTT-3’ and
5’-TCCCCGCTCCCTTCACCTTGC-3’) and a mutant allele (5’-
GGGCGCCCGGTTCTTTTTGTC-3’ and 5’-GCCATGATGGATACTTTCTCG-3’). The
PCR products for wild-type allele and mutant allele were 300 bp and 224 bp,
respectively. We used 4- to 8-week-old male Stxbp5 KO mice and Stxbp5 WT littermates
for all in vivo experiments except otherwise specified.
Platelet Isolation and activation. Murine blood was obtained by retro-orbital bleeding of
anesthetized animal into heparinized murine Tyrode's buffer (134 mM NaCl, 2.9 mM
KCl, 12 mM NaHCO3, 0.34 mM Na2HPO4, 20 mM HEPES, pH 7.0, 5 mM glucose,
0.35% bovine serum albumin) in Eppendorf tubes. The blood was then centrifuged to
yield platelet rich plasma which was then washed in new tubes containing Tyrode's buffer
with 1% PGE2 to prevent platelet activation. Platelets were then pelleted by 5 min 600 ×
g centrifugation at room temperature and the supernatant was discarded. The pelleted
platelets were gently resuspended in Tyrode's buffer and kept at room temperature for
further experiments within 2 hours. For platelet activation, diluted platelet suspension
was divided into 100 μl aliquots (3 per agonist per mouse), stimulated with 10 μl PBS or
thrombin for 10 min followed by staining with 2 μl CD62P-FITC and 4 μl JON/A-PE
antibody for 15 min, and immediately fixed with 100 ul 2% formalin. The fluorescence
intensity was measured on an Accuri C6 Flow Cytometer (BD Biosciences). The data
were analyzed with FlowJo software (Tree Star Inc.), using single-stained non-activated
WT platelets for fluorescence compensation. ATP release was measured in post-
stimulation supernatant using an ATP Bioluminescent Assay Kit (Sigma-Aldrich).
172
Platelet adherence assay. Mice were anesthetized with a cocktail of ketamine/xylazine
(80/12 mg/kg) and saline (10 ml/kg) via an i.m. injection. Anesthetized mice were
injected with a DyLight488-antibody to GPIb beta subunit to label platelets without
apparent interference to platelet-EC binding. For platelet adherence assay, the mesentery
was exteriorized on a petri dish and then placed on an inverted fluorescent intravital
microscope (Nikon, ECLIPSE Ti) with a 37°C stage warmer. An area containing target
venule (120-150 µm in diameter) was selected by measuring the vessel diameter using
the measuring tool equipped on the microscope software. After 120s baseline recording,
the mesenteric venules were superfused with 10 µM Ca2+ ionophore A23187 to activate
endothelial cells and to induce platelet rolling. Images of platelets rolling in the
mesenteric venules were continuously recorded for a total of 10 min starting at the
baseline with an electron-multiplying CCD video camera (QUANTEM, 512SC) at the
highest frame rate (approximately 31 frames per second). Using Image-Pro Plus software,
platelet adherence was determined by counting the number of platelets that remained
static for at least two consecutive images, including captured-and-go platelets. For each
mouse, we quantified platelet rolling in five equally-distributed video segments (one
during baseline and four during stimulation), each 500 frames in length, and used the
average number of rolling platelets per unit time per unit area (512 pixel × 100 pixel) for
quantification.
In vivo P-selectin exocytosis assay. Yellow-green fluorescent 1 μm microspheres
(Invitrogen) were coupled to anti-mouse CD62P antibody. Anesthetized mice were
infused with 108 microspheres intravenously, and the mesentery externalized as
173
described. Intravital microscopy was used to visualize fluorescent microspheres binding
to minimally stimulated (by surgical preparation per se) venules for 2 min followed by 8
min of stimulation by Ca2+ ionophore A23187. For each mouse, P-selectin exocytosis
was determined by counting the number of adherent microspheres on each frame of a
100-frame cropped video and averaged over resting and stimulation condition,
respectively. The adherent number of microspheres per unit area (512 pixel × 100 pixel)
was used for quantification.
Mouse tail bleeding assay. Mouse tail bleeding time was measured as described (441).
After i.p. anesthetization with ketamine and xylazine (80/12 mg/kg), the distal 5mm of
the tails of the mice were amputated and immersed immediately in 37°C saline, and the
time to visual cessation of bleeding for 30 s or continuous bleeding to 20 mins maximal
duration, whichever occurs first, was recorded.
Mouse mesenteric and carotid thrombosis model. These models were performed as
described (442, 443). For the mesenteric thrombosis model, mice were anesthetized and
platelets labeled with DyLight488-antibody to GPIb beta. A target area containing
mesenteric arterioles (120-150 µm in diameter) was externalized for imaging. The
arteriole flow was recorded for 3 min at resting condition. Then 1 mm2 of Whatman
paper saturated with 7.5% FeCl3 solution was applied to the arteriole for 3 min and the
arteriole flow was continuously recorded for a total of 30 min. The time to form a small
thrombus (50-pixel diameter) and to full vessel occlusion were recorded. Recording was
terminated at the end of 30 min if no occlusion were observed. For the carotid thrombosis
174
model, mice were sedated with 2.5% isoflurane and maintained anesthetized with 2%
isoflurane. The common carotid arteries were exposed for a baseline flow recording using
an MA1PRB Perivascular Flowprobe and a TS420 Flowmeter (Transonic Systems). Then
1×2 mm Whatmann paper soaked with 1.5 µl 7.5% FeCl3 solution was applied to the
ventral surface of the carotid upstream of the flowprobe for 3 min. Flow measurement
was resumed for a total of 30 min after FeCl3 wash-off. We define occlusion as the
absence of blood flow (0 ml/min) for 3 min.
Bone marrow transplantation. WT and Stxbp5 KO marrow donor mice were euthanized,
and femurs were isolated under sterile conditions. Bone marrow was harvested and then
aspirated repeatedly to prevent cell aggregates. Cell counts were manually performed and
107 cells were injected into each recipient mouse intravenously via the retroorbital plexus
on the same day as lethal irradiation of recipients. Stxbp5 WT 8 week old mice were used
as recipients and were lethally irradiated with an X-ray RS 2000 (Rad-sources) irradiator
delivering a single dose of 1100 rad. After marrow transplantation, mice were provided
with water supplemented with sulfatrim for 2 weeks, and allowed to reconstitute for 6
weeks prior to tail bleeding assay, blood collection, and carotid injury.
Vamp8 murine studies. Wild-type and Vamp8 KO mice were purchased from The
Jackson Laboratories (Bar Harbor, Maine). Plasma VWF was measured in platelet-poor
plasma collected into EDTA-tubes by an ELISA on 4-8 wk old mice. Bleeding time was
measured as described previously.(303) In brief, we measured the time to achieve
hemostasis after the distal 0.5 cm tip of a mouse tail (4-8 wk old) was amputated.
175
Leukocyte rolling was measured as described previously.(301) In brief, 4-5 wk old mice
were anesthetized with a cocktail of ketamine/xylazine (80/12 mg/kg) and saline (10
mL/kg) via an intramuscular injection. Anesthetized mice were injected with Rhodamine
6G (Sigma-Aldrich) in order to label leukocytes. The mesentery was exteriorized, an area
containing target venules (120-150 µm in diameter) was selectedd the venules imaged by
a Nikon ECLIPSE Ti inverted fluorescent intravital microscope (Nikon Americas.
Melville, NY). After 120 s baseline recording, the mesenteric venules were superfused
with 10 µM Ca2+ ionophore A23187 (Sigma-Aldrich) in normal saline to activate
endothelial cells which induces leukocyte rolling. Images of leukocyte rolling in the
mesenteric venules were continuously recorded for a total of 10 min starting at the
baseline with a QUANTEM 512SC electron-multiplying CCD video camera
(Photometrics, Tucson, AZ). Using Image-Pro Analyzer 6.2 software, rolling leukocytes
were determined by counting the number of leukocytes that move asynchronously with
the blood flow or remained static over at least two consecutive frames. For each mouse,
we quantified leukocyte rolling in equally-divided video segments (5 during baseline and
20 during stimulation), and used the average number of rolling leukocytes per unit area
(512 pixel × 100 pixel) for quantification. The epinephrine challenge experiment was
performed by stimulating 8-12 wk old mice with subcutaneous injection of epinephrine
(0.5 mg/kg body weight). Blood was collected before and 30 min after challenge under
2.5% isoflurane by retro-orbital punch from left and right eye, respectively.
Statistical analyses. Results were expressed as mean ± SD except for the flow cytometry
fluorescence intensity represented as median ± SD. Significance between mean values
176
was determined by the two-tailed student t test for comparison between two groups with
Gaussian distribution, and one-way ANOVA with Tukey multiple comparisons test for
comparison among three or more groups. For non-Gaussian distributed data including tail
bleeding time and mesenteric thrombosis measurement where a definite cut-off value was
assigned to some of the subjects, Kolmogorov-Smirnov test was used to compare two
groups. Responses affected by two factors were compared by two-way Tukey-corrected
ANOVA. For comparing carotid flow measurement among STXBP5 murine genotypes,
repeated measures two-way ANOVA with Tukey post test was used. Independent
variables with a value of P < .05 were considered as significant. For ANOVA post tests,
multiplicity adjusted P < .05 were considered as significant. The extent of protein co-
localization on confocal imaging was calculated using Pearson’s correlation coefficient
(444).
Study approval. All in vivo procedures and usage of mice were approved by the Division
of Laboratory Animal Medicine at the University of Rochester Medical Center (Protocol
numbers: UCAR 2010-005 and UCAR 2014-009).
177
Table 2 List of antibodies
Antibody Name Species
Immunized Cat # Source
Anti-MYRIP antibody Goat ab10149 Abcam
Anti-Syntaxin 4 antibody Rabbit ab96382 Abcam
Anti-SNAP23 antibody Rabbit ab3340 Abcam
Anti-Von Willebrand Factor
antibody Rabbit ab6994 Abcam
Anti-VAMP1 antibody Rabbit ab3346 Abcam
SELP polyclonal antibody (B01P) Mouse H00006403-
B01P Abnova
FITC Anti-Mouse CD62P Rat 553744 BD Pharmingen
Purified Mouse Anti-GM130 Mouse 610822 BD Transduction
Laboratories
Purified Mouse Anti-Syntaxin 4 Mouse 610439 BD Transduction
Laboratories
Polyclonal Rabbit Anti-Caveolin Rabbit 610059 BD Transduction
Laboratories
Calnexin (C5C9) Rabbit 2679P Cell Signaling
EEA1 (C45B10) Rabbit 3288P Cell Signaling
LAMP1 (D2D11) XP® Rabbit 9091P Cell Signaling
Integrin alphaIIbbeta3 (GPIIb/IIIa,
CD41/CD61), clone JON/A Rat M023-2 Emfret
Anti - GPIbbeta derivative Rat X488 Emfret
Cy3-AffiniPure Bovine Anti-Goat
IgG (H+L) Bovine 805-165-180
Jackson
ImmunoResearch
Laboratories
178
Alexa Fluor® 488 Goat Anti-
Rabbit IgG (H+L) Antibody,
highly cross-adsorbed
Goat A-11034 Molecular Probes
Alexa Fluor® 488 Goat Anti-
Mouse IgG (H+L) Antibody,
highly cross-adsorbed
Goat A-11029 Molecular Probes
Alexa Fluor® 594 Goat Anti-
Mouse IgG (H+L) Antibody,
highly cross-adsorbed
Goat A-11032 Molecular Probes
Alexa Fluor® 594 Goat Anti-
Rabbit IgG (H+L) Antibody,
highly cross-adsorbed
Goat A-11037 Molecular Probes
Alexa Fluor® 680 Donkey Anti-
Sheep IgG (H+L) Donkey A-21102 Molecular Probes
Rab27a Affinity Purified
Polyclonal Ab Sheep AF7245 R&D Systems
Human VAMP-8 Antibody Goat AF5354 R&D Systems
Goat IgG Horseradish Peroxidase-
conjugated Antibody Donkey HAF109 R&D Systems
SNAP 23 Antibody (H-50) Rabbit sc-50371 Santa Cruz
VAMP-1/2/3 Antibody (FL-118) Rabbit sc-13992 Santa Cruz
Tomosyn Antibody (15) Mouse sc-136105 Santa Cruz
Tomosyn Antibody (H-55) Rabbit sc-98350 Santa Cruz
SNAP25 Antibody (C-18) Goat sc-7538 Santa Cruz
VAMP2 Antibody (3E5) Mouse sc-69706 Santa Cruz
NSF Antibody (H-300) Rabbit sc-15339 Santa Cruz
Anti-Myosin Va (LF-18) antibody Rabbit M4812-.2ML SIGMA
Syntaxin 11 Rabbit 110 113 Synaptic Systems
GmbH
179
SNAP23 Rabbit 111 203 Synaptic Systems
GmbH
VAMP8 Rabbit 104 303 Synaptic Systems
GmbH
Anti -Factor VIII Related Antigen
(VWF) Rabbit F0016-13 US Biologicals
180
Table 3 Primers used for SYBR Green RT-qPCR
Gene RefSeq Forward (5'>3') Reverse (5'>3') Product
(bp)
SNAP23 NM_003825.3 ATGAGTCTCTGGAAAGTACG
AGG
CCACAGCATTTGTTGAGTTC
TG 190
SNAP25 NM_003081.3 TGTTGGATGAACAAGGAGA
ACAA CCGTCCTGATTATTGCCCCA 187
SNAP29 NM_004782.3 CCTGAACAGAATGGCACCCT TGGGGACAGGGTCTGTATCA 139
SNAP47 NM_053052.3 TGGAGGTGGCGGACAGATT AGGGTTCACAACTGGTCATG
G 129
SNAP91 NM_00124279
2.1 AGCCGGTCATGTTTGCACA
AGATCCGCTAATGGGTCCTT
T 139
B2M NM_004048.2 CCCAAGATAGTTAAGTGGG
ATCG
AGCAAGCAAGCAGAATTTG
GA 100
HPRT1 NM_000194.2 CCTGGCGTCGTGATTAGTGA
T
AGACGTTCAGTCCTGTCCAT
AA 131
YWHAZ NM_00113569
9.1
CCTGCATGAAGTCTGTAACT
GAG
GACCTACGGGCTCCTACAAC
A 100
GAPDH NM_002046.5 GGAGCGAGATCCCTCCAAA
AT
GGCTGTTGTCATACTTCTCA
TGG 197
181
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