Identification of endothelial
Weibel-Palade bodies by
single organelle flow cytometry
Mrinal Joel
Masters thesis in cell biology
Institute for pathogy, Rikshospitalet,
University of Oslo
Department of Molecular Biosciences,
University of Oslo
Contents
Acknowledgement……………………………………………………………………..... 4
Abbreviations ………………………………………………………………………….... 5
1. Introduction…………………………………………………………………………... 6
1.1 Structural features………………………………………………………………....... 8
2. Expression and storage of von Willebrand factor………………………………….. 9
2.1 The structure of VWF……………………………………………………………… 9
2.2 VWF biosynthesis…………………………………………………………………. 11
2.3 VWF interactions and role in hemostasis………………………………………….. 13
2.4 VWF/ Factor VIII complex………………………………………………………... 14
3. Biogenesis of the WPB and storage of VWF………………………………………... 15
4. The mechanisms of WPB exocytosis………………………………………………… 17
4.1 Additional factors regulating exocytosis of the WPBs …………………………….. 19
4.1.1 Nitric oxide (NO)………………………………………………………………... 19
4.1.2 Vascular endothelial growth factor (VEGF)………………………………………. 20
4.1.3 Ceramide……………………………………………………………………….. 20
4.1.4 Sphingosine 1-phosphate………………………………………………………... 20
4.1.5 Annexin A2……………………………………………………………………... 20
4.1.6 Rab3D………………………………………………………………………….. 20
4.1.7 Syntaxin 4 and Munc 18c ……………………………………………………….. 20
5. Proteins associated with the WPB …………………………………………………...22
5.1 P-selectin……………………………………………………………………………. 22
5.2 CD63…………………………………………………………………………………24
5.3 Rab-27………………………………………………………………………………. 25
5.4 Chemokines ………………………………………………………………………… 26
5.5 Endothelin (ET)-1 and endothelin converting enzyme (ECE)……………………… 26
5.6 Angiopoietin-2 ……………………………………………………………………… 27
2
5.7 α1, 3-fucosyltransferase VI………………………………………………………… 27
5.8 Tissue plasminogen activator (t-PA)…………………………………………………27
5.9 Osteoprotegerin (OPG)……………………………………………………………… 27
6. Methodological considerations ……………………………………………………... 28
6.1 Subcellular fractionation…………………………………………………………… 28
6.1.1 Fractionation media………………………………………………………………28
6.2 ELISA………………………………………………………………………………... 29
6.3Immunocytochemistry…………………………………………………………. …….. 30
6.4Transfection…………………………………………………………………………... 30
6.5 Retroviral transduction………………………………………………………………. 31
6.6 FACS………………………………………………………………………............... 32
6.7 SOFA………………………………………………………………………………... 32
6.8 FAOS………………………………………………………………………………… 32
7. Aim of the study……………………………………………………………………….33
8. Refrences……………………………………………………………………………….34
Manuscript
3
Acknowledgement
The work presented in this thesis was carried out at the Laboratory for Immunohistochemistry
and Immunopathology (LIIPAT), Institute of Pathology, University of Oslo, Rikshospitalet
University Hospital from January 2004 to June 2005.
I would like to acknowledge Professor Per Brandtzæg for providing excellent laboratory
facilities. Furthermore, I would like to thank Dr Guttorm Haraldsen and Professor Oddmund
Bakke for giving me the opportunity to increase my knowledge in the field of cell biology.
I am extremely greatful to Dr Espen. S. Bækkevold for his incredible supervision, outstanding
guidance and enthusiasm. Thank you so much for always being so helpful and understanding
to me.
I would further like to acknowledge Inger Øynebråten and Dr. Marjan. J. T Veuger for their
remarkable support during the lab work as well as during the writing of this thesis. Thanks a
lot for always being so willing to help.
Furthermore, I am greatful to the staff at the Dept. of Gynecology and Obstetrics at
Rikshospitalet for their assistance in obtaining the umbilical cords.
Thanks to Gøril Olsen for her guidance and assistance in the FACS analysis.
I would also like to thank Kathrine Hagelsteen, Vigdis Wendel and Aaste Aursjø for their
assistance in the lab work.
Thanks to all the members of the LIIPAT for all those social events, which creates an
excellent atmosphere to work and study.
Furthermore, I would like to thank all my friends and fellow students in the room A3.M022D
for creating a friendly and cheerful environment.
Thanks to all my friends for always being there for me.
Last but not the least, I would like to thank my family, my grandpa Mr. B. A. Joel, grandma
Mrs. M. P. Joel and my mom Miss Nilima Joel for their love, support and faith in me.
4
Abbreviations
Ang-2
Angiopoeitin-2
CaM
Calmodulin
DDAVP
1-deamino-8-D-arginine vasopressin
ECs
Endothelial cells
ECE
Endothelin converting enzyme
FAOS
Flourescence activated organelle sorting
GFP
Green fluorescent protein
HA
Hemophilia
HUVECs
Human umbilical vein endothelial cells
IP3
Inositol 1, 4,5-triphosphate
MVBs
Multivesicular bodies
NSF
N- ethylmaleimide sensitive factor
NO
Nitric oxide
OPG
Osteoprotegerin
PAR
Protease activated receptor
PDI
Protein disulphide isomerase
PIP2
Phosphatidyl inositol 4,5-bisphosphate
PKA
Protein kinase A
PKC
Protein kinase C
PLC
Phospholipase C
PMA
Phorbol 12-myristate 13-acetate
PNS
Post nuclear supernatant
SNARE
Soluble NSF attachment protein receptor
SOFA
Single organelle flow analysis
S1P
Sphingosine 1-phosphate
TGN
Trans –Golgi network
TNF
Tumour necrosis factor
t-PA
tissue type plasminogen activator
VEGF
Vascular endothelial growth factor
VWD
Von willebrand disease
VWF
Von willebrand factor
WPBs
Weibel-Palade bodies
5
1. Introduction
An endothelium lines the entire vascular system and is composed of a monolayer of
endothelial cells (ECs). In adults, the endothelium consists of approximately (1-6) x 1013 ECs
which cover a surface area of about 1000m2 [2, 3].
ECs play an important role in regulating metabolite exchange by acting as a selective
permeability barrier between the bloodstream and the surrounding tissue [3]. They effectively
regulate coagulation and thrombosis by normally displaying an anticoagulant surface, while
once needed, will change properties to induce blood clotting and thrombosis. ECs are also
crucially involved in the regulation of vascular tone and blood pressure through synthesis of
nitric oxide (NO). They are also involved in the regulation of immune and inflammatory
responses by controlling extravasation of leukocytes, monocytes and lymphocytes[4] through
expression of membrane-bound adhesion molecules and soluble chemoattractants including
chemokines. Several of the properties described to this point are crucially dependant on the
ability of ECs to synthesize and store proteins in secretory granules. Such granules may then
fuse to the cell surface in response to secondry signals and empty their cargo (or expose
functional domains of transmembrane molecules) in a process called regulated secretion. The
best characterized compartment for such secretion in ECs was first described by Ewald R.
Weibel and George E. Palade in 1964, and thus named Weibel- Palade bodies (WPBs)[5].
WPBs are EC restricted rod-shaped organelles and are present in ECs of practically all blood
vessels as well as endocardium. WPBs are specialized storage and secretory vesicles for a
number of bioactive substances like hormones, enzymes, receptors and adhesive molecules
[6]. Due to selective presence of this organelle in ECs, it was early postulated that WPBs
would play some role in vascular physiology [5, 7]. However it took nearly two decades and
the development of techniques enabling the maintenance of human ECs in culture [8] before
discovering that WPBs contain von Willebrand factor (VWF)[9], a plasma protein known to
be involved in hemostasis [7, 10, 11]. In fact, VWF appears to be the predominant constituent
of WPBs, and has been found to govern many of the unusual morphological features of
WPBs[12]. Indeed, it has been shown that the expression of mRNA encoding the full length
VWF in non-endothelial cells, is sufficient to induce the formation of rod-shaped organelles
with a morphology indistinguishable from the bonafide endothelial cell WPBs [13]. However,
6
WPBs also contains numerous other proteins (reviewed later), but a systematic mapping of its
molecular components has not been performed. The purpose of this study was to establish a
method for isolation of highly purified WPBs for analytical and preparative purposes – and
thus provide a new tool to dissect the biology of WPBs.
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1.1 Structural features
Weibel-Palade bodies are elongated, membrane enclosed organelles, which are ≈0.1µm wide
and up to 4µm long [5]. These bodies are long, cylindrical rods with sharply cut-off, rather
than tapered ends [5]. The rods are surrounded by a tightly fitting unit membrane which is
about 60-80Å thick [5]. Furthermore, WPBs contain 8-12 small tubules; each of them
approximately 150Å thick embedded in a dense matrix and running parallel to the long axis of
the rod [14]. They are regularly spaced with a distance of about 250 -300Å, separated from
each other with a matrix of approximately 100Å thickness [5].WPBs usually occurs in groups
and are found to be concentrated in the perinuclear region of the cytoplasm [5]. As many as ≈
200 WPBs are detected per cell in HUVECs after 48 hours in culture [15].
a
b
Figure1: (a) Cultured HUVECs with WPBs visualized by VWF-immunostaining (b) Section of
human umbilical vein endothelial cell containing a WPB with tubules arranged parallel to its
long axis (arrow). The transverse section of WPB shows a juxtaposition of several tubules
embedded in a dense matrix[13].
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2. Expression and storage of von Willebrand factor
VWF is named after Dr. Erich von Willebrand [16], who in 1926 described a bleeding
disorder distinct from hemophilia that was later recognized to be caused by decreased
synthesis or mutations of VWF [17]. VWF promotes hemostasis by means of two separate
mechanisms. It acts as an adhesive extracellular matrix protein in the primary hemostasis,
promoting the formation and stabilization of the platelet plug [7]. It also functions as a
soluble, circulating plasma protein found in association with factor VIII, for which it appears
to behave like a chaperone [18, 19]. VWF protects factor VIII from proteolytic degradation
and directs it to sites of clot formation [7, 20, 21]. von Willebrand disease(VWD) is
considered to be the most common hereditary bleeding disorder in humans (0,8%
frequency)[22].
Synthesis of VWF occurs in two sites: in platelets and their precursor cells (megakaryocytes)
where it is stored in α-granules, and in ECs where it is either secreted constitutively or stored
in the WPB for secretion upon stimulation [23, 24].The majority of the VWF synthesized by
the ECs is secreted constitutively (i.e. directly after synthesis without any storage) and only 510% of VWF synthesized in vitro by cultured primary ECs is stored in WPBs [7, 25].
The VWF synthesized by ECs gives rise to three pools of functionally important VWF within
the body. This includes the VWF stored within the WPBs and released upon stimulation,
VWF bound to factor VIII in the blood circulation, and VWF bound to the subendothelial
basal lamina [7, 18]. Although VWF stored in the WPB is rich in ultra-large forms that are
hyper-reactive in their capacity to bind to platelets [26, 27], all the three pools appear to
contribute to the adhesion of platelets as well as formation of platelet plug during blood vessel
injury [28-31].
2.1 The structure of VWF
The primary structure of the VWF has been determined both by direct amino acid sequence
analysis and from corresponding cDNA [32]. The open reading frame in VWF cDNA predicts
a 2813 amino acid polypeptide as the primary translation product mapped 245 nucleotides
upstream of the initiator methionine [33]. Moreover, molecular cloning of the full length
9
human VWF cDNA has revealed that the mRNA is translated as a pre-pro-polypeptide
composed of a signal peptide of 22 amino acid, an unusually large propeptide of 741 amino
acids and a mature VWF molecule of 2050 amino acids[32, 34]. As the carboxyl-terminal
sequence of the precursor is identical to that of the plasma VWF, the proteolytic processing
occurs at the amino-terminal end of the precursor[32].
VWF consists of a repetitive domain structure [34] (Figure2), and four types of repeated
domains (A-D) are found, exhibiting internal homologies in 2-5 copies. To some extent, these
domains exhibit autonomous functions, which is revealed by comparative studies of their
protein sequences. Furthermore, they share similarity with a number of different, apparently
unrelated structural motifs of many proteins[34] .
Multimerisation
Dimerisation
Figure 2: Schematic view of Pre-pro-von Willebrand factor. The signal peptide (SP) is 22 aa long.
The numbers in parentheses indicate the length of the propeptide (red) and mature VWF (green).
Domains necessary for multimerization, dimerization and storage are indicated [1]. Furin (an
endopeptidase) cleavage site is indicated by the curved arrow.
10
The Propeptide of VWF is identical to a Mr 100,000 plasma glycoprotein, known as the VWF
Antigen II [18, 34-36] and is intimately involved in the maturation and trafficking of VWF to
WPBs[7]. VWF Antigen II is required for the proper post-translational multimerization and
subsequent storage of VWF [34]. Over the past few years, increasing evidence has been
obtained that VWF Antigen II might also be involved in a variety of biological processes after
it is secreted, by modulating platelet function and acting as an inflammatory mediator [34].
Another characteristic feature of the pro-VWF structure is the occurrence of the Arg- GlyAsp (RGD) sequences [34]. One of the RGD sequences is located near the C terminal region
of the propeptide while the another one is present near the C terminal end of the C1 domain
[34].The latter is found to be essential for the binding of VWF to the platelets while the
function of the former one is still unknown [34, 37].
The electron microscopic studies of VWF multimers shows that they appear as unbranched,
loosely coiled, extended thin strands [38, 39]. They are organized in a head to head and tail to
tail fashion, and the molecules are composed of globular regions connected by thin flexible
rods[18].
2.2 VWF biosynthesis
VWF is a classical example of a secretory protein , as the molecular events associated with its
biosynthesis and secretion to a very large extent are similar to the processing steps and
routing of many other proteins destined to be externalized [34]. The early work on VWF
synthesis was based on the study of the VWF endogenously expressed in cultured primary
endothelial cells [40, 41]. However, the construction and successful heterologous expression
of eukaryotic vectors encoding full length human pre-proVWF[42], made it possible to
address more precisely at the molecular level the determinants necessary for the sorting of
VWF to WPBs[7].
The molecular events associated with the biosynthesis of VWF occur in a highly ordered
manner in terms of time and space [6]. Moreover, VWF undergoes a number of posttranslational modifications including sulphation, disulphide-linked oligomerisation (involving
the cysteine residues in the N-terminus of the protein) and proteolytic cleavage of proVWF to
produce VWAgII and mature VWF (220 kDa) [7, 40]. Both the sulphation and proteolytic
11
cleavage are modifications that play an important role in many secretory proteins[7, 43].
However, such formation (or rearrangement) of disulphide bonds in the TGN is rare[7]
(Figure3).
Figure 3: Trafficking of von Willebrand factor. The left column indicates the location and
nature of the main VWF modifications (in black), as well as the stage at which P-selectin and
Rab27a are recruited (blue and green, respectively). LMW: low molecular weight: HMW:
high molecular weight[1].
After the cleavage of the signal peptide in the ER by a signal peptidase, pro-VWF undergoes
N-linked glycosylation [7, 40, 41] by the addition of asparagines(N)- linked sugars which is
thereafter followed by dimerization [18, 34] through the formation of disulphide bonds at the
C-terminal region [34, 40, 44, 45].
12
The secondary structure of VWF reveals that these intersubunit disulphide bonds are located
beyond the domain C2, in the N- terminal region (figure 2) (residues 1046-1458) and the Cterminal region (residues 2671-2813) [34, 44]. Dimerization is followed by the transport of
pro-VWF dimers to the Golgi apparatus and post-Golgi compartments, where these high
mannose oligosaccharides are processed to their complex forms, which is then followed by
sulphation and O- linked glycosylation [18, 34, 46]. During its passage through the Golgi
compartments, the processed dimers starts polymerizing, and are subsequently transported to
the trans-Golgi network where further multimerization takes place [13, 34, 47].
The VWF propeptide (VWF Antigen II) acts as an intramolecular chaperone protein by its
ability to promote interchain disulphide bonding [7, 34, 48]. The primary structure of those
peptide regions within the D domains of the propeptide are homologuos to the active site of
the proteins harbouring the protein disulphide isomerase (PDI) activity [34]. PDI is an
enzyme that catalyses the thiol-disulphide interchange reactions in the protein substrates,
leading to disulphide formation and folding of the protein [49]. Any changes in the propeptide
leads to the inhibition of PDI activity , thus inhibiting the polymerization [34, 48].
The proteolytic cleavage of VWF multimers also takes place in the TGN, leading to the
formation of mature VWF multimers and propeptide dimers [34]. A likely candidate for proVWF cleavage is Furin which is also present in TGN. After cleavage, the mature VWF and
the pro peptide of VWF remain non-covalently associated, at least within the cell [34, 50].
2.3 VWF interactions and role in hemostasis
The role of VWF in platelet adhesion was recognized already before the protein was fully
characterized [18], and ultrastructural examinations of incision wounds demonstrated defects
in platelet plug formation in patients with von Willebrand disease [51, 52].
VWF released by both the platelet α granules and WPBs enables the platelets to adhere to
exposed subendothelium as well to respond to the changes in the shear stress of the blood or
the lesions in the vascular endothelium [53]. The interaction of VWF with the platelet surface
is essential physiologically in hemostasis and pathologically in thrombosis: VWF contains
binding sites for platelet membrane glycoproteins GP Ib–IX–V and GP IIb–IIIa. The A1
domain of the mature VWF subunit contains binding site for the platelet glycoprotein GP Ib–
13
IX–V complex [10, 54] while the C2 domain close to the carboxyl terminal region of VWF
interacts with the activated GP IIb–IIIa complex.
VWF plays an important role in mediating the initial steps of platelet adhesion and
aggregation at sites of vascular injury by acting as a bridging element to structures in the
subendothelial matrix [53]. Thus, the interaction of VWF with the platelets is very important
for the initiation of any hemostatic or thrombotic process.
2.4 VWF/ Factor VIII complex
Factor VIII (FVIII) and von Willebrand factor (VWF) are distinct but related glycoproteins, in
that they circulate in plasma as a tightly bound complex (FVIII/VWF)[55]. Their deficiencies
or structural defects are responsible for the most common inherited bleeding disorders,
namely hemophilia A (HA) and von Willebrand’s disease (VWD)[55]. VWF is not only
essential for the production of FVIII but also for its stability in the plasma. Binding of VWF
to FVIII maintains the high degree of reactivity of FVIII by increasing its susceptibility to
thrombin cleavage. Once cleaved by thrombin, FVIII acts as a cofactor and helps to accelerate
the activation of factor X by activated factor IX in the coagulation cascade [55]. Binding of
FVIII to VWF is also essential for the survival of the FVIII in vivo [56, 57] as VWF protects
FVIII from useless degradation .
Since FVIII and VWF form a tightly bound, noncovalent complex in plasma, both
glycoproteins are copurified from plasma [57, 58]. FVIII is mostly produced by non-ECs but
its presence is reported in the liver sinusoidal ECs [59]. The localization of this FVIII is still
unclear in the WPBs but Rosenberg et al. [60] has shown that the transfected FVIII is sorted
to the WPBs and its expression in the liver sinusoidal ECs raises the possibility of coexpression and thus their sorting to the WPBs.
14
3. Biogenesis of the WPB and storage of VWF
One of the most remarkable features of ECs is their ability to store de novo synthesised VWF.
After endoproteolytic cleavage of pro-VWF in the TGN[50], mature VWF and its propeptide
partition between two different secretion pathways: the constitutive and the regulated
pathway. In cultured ECs, a part of the VWF and propeptide generated upon proteolytic
maturation is secreted in a constitutive manner, together with some unprocessed pro-VWF
[25]. However, a portion of mature VWF, together with its propeptide, is stored in the WPBs
where they are present in equimolar amounts [12, 61-63]. The WPB pool primarily consist of
the high molecular weight multimers which are only secreted upon proper stimulation of the
cell [6].
Insights into the mechanisms and signals involved in the biogenesis of WPBs and in directing
VWF into these storage granules are expanding. Accumulating evidence suggests that VWF
by itself is the driving force behind the formation of WPBs [6]. Apparently, mature WPBs
with densely packed VWF, surrounded by a limited membrane, are occasionally found in
close proximity of the TGN suggesting that WPBs emerge through budding from the golgi
complex [47].
Moreover, as VWF stored within the WPBs is of higher molecular weight than that secreted
constitutively, it has been postulated that WPBs selectively accumulated the higher order
oligomers, while less highly oligomerised VWF is not sorted but rather secreted constitutively
[7]. Such a mechanism would partly be in line with the “sorting by retention” model [64],
suggesting that intermolecular associations and selective coaggregation cause participating
regulated secretory proteins to be efficiently retained within maturing granules [65]. However
additional data obtained from the heterologous expression of VWF mutants have proven that
the covalent multimerization of VWF and its storage in WPBs are two completely
independent processes [13, 66].
Further studies have shown that the VWF propeptide is required for the formation of these
granules, but it cannot independently induce the formation [67]. Thus, a second hypothesis
has been proposed suggesting that the propeptide moiety of VWF contains a sorting signal
that interacts with a carrier protein or receptor in the Golgi membrane to mediate targeting of
VWF (and the propeptide itself) to storage [6, 66]. Two amino acids, R416 in the propeptide
15
and T869 in mature VWF mediate non covalent interaction between the two subunits and is
important to induce storage of mature VWF[68].
Although a number of issues are still controversial, the formation of WPBs has been partly
elucidated. Initial formation is suggested to occur at the TGN. Furthermore, it is clear that
VWF expression is necessary to trigger the formation of the WPBs, and the propeptide play
important roles by interacting non-covalently with other VWF proteins like the mature VWF.
By contrast, multimerization is neither necessary nor sufficient to induce storage and does not
seem to play any role in the biogenesis of WPBs [1]. Finally, non-covalent interactions
between VWF proteins are important for elongation and tubule formation of the WPBs [69].
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4. The mechanisms of WPB exocytosis
Regulated secretion of VWF and other constituents of the WPBs, involves the translocation
of the WPBs from the cytoplasm followed by their fusion with the plasma membrane and
release of the content [6] (Figure 4). Such release might occur within minutes in response to
agonists such as thrombin or histamine as exemplified by VWF-dependent platelet adhesion
and rolling on the EC surface: i.e. a process in which adhesion and rolling of platelets is
peaking 30-60 s after stimulation [70]. Induction of secretion from WPBs does not affect the
distribution of VWF among the constitutive and regulatory pathways, nor does it result in an
increase in biosynthesis of the protein [71].
Fig4: Machinery of stimulus induced exocytosis of WPBs by the ECs [6]. Upon the
stimulation of ECs by thrombin, IP3 pathway is activated causing the rise in the cytoplasmic
calcium ions. The stimulation of ECs agonists by vasopressin and epinephrine activates the
cAMP dependent signalling pathway. However the molecular mechanisms of the cAMP
mediated VWF secretion are still to be identified.
17
The exocytosis of WPBs can be induced by a large number of different agonists that may be
separated into two groups: those that act by elevating intracellular Ca+2 levels, and those that
act by raising cAMP levels in the cell[63, 72-81]. Secretagogues like thrombin and histamine
belong to the former type, while substances like epinephrine, adenosine or DDAVP(1deamino-8-D-arginine vasopressin) belong to the latter[82].
Exocytosis mediated by thrombin and histamine leads to the activation of the phosphatidyl
inositol (IP3) pathway, resulting in a rise of the cytoplasmic Ca2+ level, which is observed
during exocytosis of secretory organelles. As shown in figure3, thrombin signalling is
mediated through a G protein-coupled protease-activated receptor (PAR), leading to the
activation of an enzyme phospholipase C-β(PLC-β) which acts on an inositol phospholipids
phosphatidyl inositol 4,5-bisphosphate (PIP2) by hydrolyzing it to produce Inositol 1,4,5triphosphate (IP3) and diacylglycerol. IP3 leaves the plasma membrane, diffuses into the
cytosol and reaches the ER where it binds to and opens IP3 - gated Ca2+ release channels in the
ER membrane. The cellular responses to the increased [Ca+2]level is mediated through
calmodulin and small GTP binding proteins[83, 84]. Calcium in complex with calmodulin
(CaM) binds the small GTP-binding protein Ral in its GDP form, and the binding of Ral leads
to its recycling from the inactive GDP bound state to the active GTP bound form by Ral
guanine nucleotide exchange factor (Ral GEF), and is followed by the secretion of VWF[85].
It seems clear, therefore, that this small GTPase plays a key role in controlling regulated
exocytosis of VWF by ECs. Ral-GTP interacts downstream with effector molecules such as
RLIP76, a GTPase-activating protein for Cdc42 and Rac. The hydrolysis of GTP on Cdc42 or
Rac may result in cytoskeleton rearrangement, which may precede exocytosis of WPBs [6].
In addition, thrombin stimulation of the PAR includes calcium-dependent activation of
protein kinase Cα (PKC α) whose downstream compounds mediating WPB exocytosis
includes syntaxin 4 and Munc18c, two members of the SNARE (soluble NSF attachment
protein receptor) core complex critical for intracellular fusion events[86].
The molecular mechanisms associated with cAMP dependent exocytosis of WPBs involves
substances like epinephrine and vasopressin which acts on G protein-coupled receptor and the
vasopressin V2 receptor respectively [63, 78]. Stimulation of these G protein-coupled
receptors results in activation of adenylate cyclase which catalyzes the formation of the
second messenger cAMP [78, 80]. Rondaij et.al have shown in their studies that the small
18
GTP binding protein Ral which is activated during the thrombin induced VWF secretion[85]
is also activated during the epinephrine induction [82]. However, the epinephrine-induced Ral
activation was more gradual than that of thrombin [82]. The epinephrine-induced Ral
activation depends on protein kinase A(PKA) and proceeds independently of the cAMP
regulated exchange protein Epac [82]. The cAMP-induced activation of PKA which
subsequently results in the phosphorylation of the target proteins [78, 80]. Rondaij et.al thus
suggested that PKA activity is crucial for the release of VWF in epinephrine stimulation, but
not in thrombin stimulated endothelial cells [82].
Further studies, however, are required to elucidate the mechanisms behind the current finding
that epinephrine-induced VWF secretion is mediated by cAMP/PKA dependent activation of
small GTPase Ral [82]. As yet, it is not clear that how PKA contributes to the activation of
Ral [82].
One important difference between cAMP-mediated responses and those involving regulated
secretion caused by the increased Ca2+ levels, is that secretion induced by the latter involves
the release of both peripheral and central granules, whereas cAMP mediated secretion
involves only vesicles located in the periphery of the cell [87]. It has been proposed that this
latter cellular pool of VWF is of importance in maintaining plasma VWF levels [6].
4.1 Additional factors regulating exocytosis of the WPBs:
In addition to the classical compounds acting through G protein coupled receptors, several
factors modifying exocytosis of the WPBs have been identified the last couple of years. Some
of them are described below:
4.1.1 Nitric oxide (NO) is a messenger molecule involved in the regulation of vascular
inflammation in part by inhibiting exocytosis of WPBs [88]. NO modifies the activity of Nethylmaleimide –sensitive factor (NSF) [89]. NSF is a cytosolic protein which forms
homohexamers, hydrolyzes ATP, and alters the conformation of the stable SNARE complex.
The SNARE complex proteins regulate exocytosis from different cell types including
exocytosis of the WPBs [89-91]. NO inhibits NSF disassembly of SNARE complexes by
nitrosylating critical cysteine residues of the NSF [89].
19
4.1.2 Vascular endothelial growth factor (VEGF) is known to regulate processes such as
angiogenesis, vascular permeability, vasodilation and promote vascular inflammation. Recent
studies by Matsushita et al.[92] shows that VEGF regulates the WPB exocytosis by exerting
two opposing effects. VEGF triggers exocytosis through calcium and PLC-γ mediated
pathways, while inhibits exocytosis by activating the PI-3 kinase/Akt pathway which
increases NO synthesis thereby inhibits exocytosis [92].
4.1.3 Ceramide is a sphingolipid that is produced de novo from serine in ER, or from
hydrolysis of sphingomyelin by sphingomyelinase in the plasma membrane, cytosol,
lysosomes and ER. Endogenous ceramide triggers exocytosis of WPBs [93], but ceramide
also has an opposing effect as it activates endothelial NOS thereby increasing the level of NO
and inhibiting exocytosis of the WPBs [88].
4.1.4 Sphingosine 1-phosphate (S1P) is a metabolite of ceramide secreted by platelets,
monocytes, and mast cells acting as an extracellular messenger molecule. Similar to ceramide,
S1P trigger pathways that both promote and inhibit endothelial exocytosis [94]. S1P induces
WPBs exocytosis by triggering PLC-γ pathways, while inhibit it by activating the
phosphatidylinositol 3-kinase pathway, thus increasing NO synthesis [94].
4.1.5 Annexin A2 is a member of the annexin family of Ca 2+dependent lipid binding proteins
and believed to be engaged in membrane transport processes in a number of cell types.
Depletion of annexinA2 and its ligand S100A10 reduces the agonist (histamine) induced and
Ca2+-dependent secretion of VWF [95].
4.1.6 Rab3D is an isoform of Rab3 and plays an important role in the regulated secretion of
VWF [95]. Knop et al shows that Rab3D is involved in the WPB exocytosis through the Ca2+
dependant mechanism [95]. Mutations in Rab3D causes reduction in the number of WPBs.
4.1.7 Synatxin 4 and Munc 18c
Syntaxin 4 is a SNARE protein found in the association with regulatory protein
Munc18c[96]. Fu et al. showed that syntaxin 4 is involved in the exocytosis of WPBs by
facilitating SNARE complex formation, which mediates the fusion of WPBs with the
endothelial membrane [86]. Thrombin ligation of PAR-1 leads to the activation of PKC which
20
in turn phosphorylates syntaxin 4 causing its dissociation from Munc18c and leading to
SNARE complex formation [97].
21
5. Proteins associated with the WPBs
The WPBs contains a number of other proteins in addition to VWF and its propeptide [6].
These proteins are both structurally and functionally clearly distinct, and like VWF and its
propeptide, they are translocated to the cell surface upon stimulation, and may control local or
systemic biological effects, including inflammatory, hemostatic and vasoactive responses [6].
The mechanism responsible for sorting of these proteins into WPBs is poorly understood,
however accumulating evidence suggest that VWF not only plays a role in directing the
formation of WPBs, but might also play a very important role in the sorting of other WPBproteins[6].
5.1 P-selectin
The best characterized membrane protein of WPBs is P-selectin (also called GMP-140,
PADGEM, CD62P), a 140 kDa type I glycosylated transmembrane protein, which belongs to
the selectin family of c-type lectins, and is expressed in platelets and ECs [7, 98-101]. The
selectins share a modular structure with a characteristic c-type lectin domain which is largely
responsible for ligand recognition at the N-terminus [7]. It is followed by an EGF-like
domain, a series of consensus repeats and a single transmembrane spanning region with a
short cytoplasmic tail [7].
P-selectin translocated from WPBs to the cell surface mediates the initial adhesion in
inflammatory responses of leukocytes.
P-selectin shows a complex internalization in ECs. The surface exposure of P-selectin is short
lived, and disappears within 20 min after stimulation [72, 99]. These events have been tracked
in HUVECs using uptake of anti-P-selectin antibodies [102, 103], and showing that P-selectin
is rapidly re-internalized and appears in the early endosome where it colocalizes with the
transferrin receptor and transferrin [7]. Subsequently, 50% of the internalized protein is
transferred to lysosomes, where (in the presence of protease inhibitors) it can be seen to colocalize with the lysosomal marker LAMP1[7, 103]. The rest of the protein returns to the
TGN [7, 103].
22
The heterologuous expression studies of P-selectin and its chimeric proteins (which consist of
regions of P-selectin fused to various reporters) in cells containing regulated secretory
organelles shows that these chimeric proteins are sorted to these organelles[7]. In cells lacking
the secretory organelles, P-selectin is directed to the plasma membrane and ultimately to
lysosomes[7, 104-107].
The information required for the intracellular trafficking of P-selectin is located in its 35amino acid cytoplasmic tail, which is composed of a ´stop transfer` region followed by the C1
domain and the C2 domain (Figure 5).
Figure5: (A) Domain structure of P-selectin. Lectin—c-type lectin domain; EGF—epidermal
growth factor like domain; CR—consensus repeats; TM—transmembrane spanning region;
ST—‘stop transfer’ region (residues 755–761); C1 domain (residues 762–772); C2 domain
(residues 773–789)[7]. (B) WPB targeting sequences in the tail of P-selectin.
In fact, the presence of granule targeting signal within the cytoplasmic domain around
777
Y( in the YXXØ motif) in the C2 domain has been suggested[108]. However, there is also
evidence that the transmembrane region may play a secondary role in targeting[109]. Pselectin is stored in WPBs together with VWF and is degraded in its absence [103].
23
5.2 CD63
CD63 is a member of the family of lysosomal membrane proteins which include Lamp1,
Lamp2 and Lamp3 [110, 111]. These proteins share extensive glycosylation with multiple
sialylated polylactosaminoglycan residues and have significant structural similarities in their
cytoplasmic tails [110, 111]. However, unlike the Lamps, CD63 has the structure of a type III
membrane protein, with four transmembrane domains [111, 112]. CD63 is found in the
internal vesicles of late endosomes and in the lysosomes [7] and is localized to the WPBs
[110, 113].
CD63 is found in the limiting membrane of WPBs in the ECs [111], but only 5–15% of the
cellular pool of CD63 is localized to WPBs [111, 114]. However, in cells with very high
numbers of WPBs, a greater fraction of the CD63 co-localizes with VWF[7]. CD63 becomes
surface exposed upon WPB translocation and like P-selectin is subjected to rapid
internalization [111].
Little is known about the trafficking of CD63 to WPBs and late endosomes/lysosomes of
ECs[7]. However, the endocytic trafficking of CD63 has been studied in HUVECs by
determining the fate of antibodies internalized at the cell surface in resting and activated
cells[103, 114]. From these studies, it is apparent that the CD63, which appears on the plasma
membrane, first accumulates within endosomal structures and their internal vesicles.
Subsequently, CD63 is delivered to both WPBs and late endosomes/lysosomes 2–8 h after
internalization[7]. The trafficking of CD63 to granules and endocytic organelles is slower
than that of P-selectin to the same destinations [7], and it is suggested that this delay in
delivery to WPBs is due to its selective retention within the internal vesicles of multivesicular bodies (MVBs)/late endosomes, from which it then passes to the WPBs [114]. This
is different from P-selectin, which passes from endosomal structures to WPBs via an AP-1
positive compartment—presumably the TGN—as an intermediate [7, 103].
While most lysosomal proteins locate on the limiting membrane of MVBs, CD63 selectively
accumulates within their internal vesicles possibly contributing to their differential sorting to
WPBs [7]. Such targeting has been proposed to occur in platelet α granules and dense
granules formation, where CD63 accumulates within the vesicles of MVBs while P-selectin is
24
found in the limiting membrane [115, 116]. A similar segregation possibly occurs in the ECs
as well [103].
The tyrosine based motif GYXXØ found in CD63 is responsible for its delivery to the
lysosomes, and interacts with the cytoplasmic complex adaptor protein AP-3[117]. However,
it is unclear if the same targeting signal is used in the targeting of CD63 to the WPBs [7]. In
fact, it has been proposed that CD63 might use an alternative signal for its targeting to WPBs,
as GYXXØ is found in other lysosomal glycoproteins like LAMP1 and LAMP2 which are not
colocalized in WPBs [7, 114].
5.3 Rab-27
The Rab family is part of the Ras superfamily of small GTPases. Different Rab GTPases are
localized to the cytosolic face of specific intracellular membranes, where they function as
regulators of distinct steps in membrane traffic [118-120]. In the GTP-bound form, the Rab
GTPases recruit specific sets of effector proteins onto the membranes. Through their
effectors, Rab GTPases regulate vesicle formation, actin- and tubulin-dependent vesicle
movement, and membrane fusion.
It has been reported that WPBs efficiently recruit Rab 27a in a time dependant manner.
Hannah et al.[119] showed that newly formed WPBs emerging from the TGN lack Rab27a,
but is acquired 4 to 5 hours after initial appearance of the cigar-shaped organelle. Thus, WPBs
are first formed in an immature, Rab27a-negative form, with the subsequent time-dependent
Rab27a recruitment reflecting maturation of the organelle. This is in line with earlier
evidence, showing that WPBs require a considerable amount of time to get fully matured, as
several hours are required for metabolically labelled VWF to reach a compartment with the
buoyant density characteristic of mature WPBs [50, 121]. Moreover, most of the mature
WPBs appear to be Rab27a positive at steady state, thus it has been suggested that Rab27a
remains associated with the organelle until exocytosis occurs [119]. Hannah et al. also
showed that recruitment of Rab27 is defined by the organelle and not by the cell type: in
VWF-transfected HEK-293 cells significant enrichment of endogenous Rab27
immunoreactivity can be found on the WPB-like organelles that have been formed, whereas in
mock-transfected cells Rab27 appeared localized to a compact pericentriolar region [119].
Thus it appears VWF can organize its surrounding membrane, but the nature of this
mechanism has not been addressed.
25
5.4 Chemokines
Chemokines are small secreted cytokines that may activate or chemoattract leukocytes. The
neutrophil attracting CXC chemokine interleukin (IL)-8/CXCL8 and the eosinophil-attracting
chemokine eotaxin-3 are the two chemokines described so far in the WPBs[122-124].
Such prestorage of chemokines might provide a rapid pathway for specific activation of
leukocyte adhesion at sites of acute inflammation in a compartment that simultaneously
presents P-selectin to active leukocyte rolling, a prerequisite for subsequent activation of firm
adhesion[123]. IL-8 is predominantly stored in the WPBs of skin and intestinal
microvesicles[123]. VWF may play a major role in the sorting of IL-8 to the WPBs [122, 124,
125]. de Wit et al. has suggested this coexpression using cell lines with reduced VWF
synthesis which leads to the loss of expression of IL-8 [125]. Furthermore, prolonged
stimulation of primary human ECs results in the accumulation of IL-8 in the WPBs, where it
is retained for several days after the removal of the stimulus, and may be released by
secretagogues [124, 126]. Thus, such storage of IL-8 in the WPBs may serve as an EC
“memory” of a preceeding inflammatory insult, which then enables the cells to secrete IL-8
immediately without de novo protein synthesis [124].
5.5 Endothelin (ET)-1 and endothelin converting enzyme (ECE)
Endothelin-1(ET)-1 is the predominant form of endothelin and is a vasoconstrictor peptide
found in the cytoplasmic matrix as well as in WPBs [127] ET-1 is released by regulated
secretion of the WPBs towards the basolateral membrane of the ECs [128]. ET-1 is
synthesized as big endothelin-1 and processed by endothelin converting enzyme (ECE) which
convert it to the most potent form. There are two types of ECEs discovered so far, including
ECE-1 and ECE-2. Two isoforms of ECE-1, ECE-1α and ECE-1β exists that differ only in
their N-terminal sequences. Furthermore, as ECE is also localized in the WPBs this
processing might occur in the WPBs [129]. Russell et. al has also speculated that the big ET-1
is translocated from the TGN into the WPBs and is converted to ET-1 by both ECE-1α and
ECE-1β[129].
26
5.6 Angiopoietin-2
The Angiopoietins (Ang-1 and Ang-2) are soluble proteins and acts as the ligands for the
receptor tyrosine kinase Tie-2 [130-132]. Recently, Ang-2 was also discovered in the WPBs
[133], displaying release kinetics similar to VWF. The Angiopoeitins were originally found to
critically regulate vascular maturation during angiogenesis.
The identification of Ang-2 as a stored, rapidly available molecule in ECs suggested functions
of the angiopoietin/Tie-2 system beyond the established roles during angiogenesis likely to be
involved in rapid vascular homeostatic reactions such as inflammation and coagulation [133].
5.7 α1, 3-fucosyltransferase VI
α1,3-fucosyltransferases are glycosyl transferases usually found in the Golgi, but α1, 3fucosyltransferase VI is also located together with VWF in the WPBs of HUVECs [134].
Fucosyltransferase VI does not exhibit any enzymatic activity in WPBs, thus the functional
significance of this enzyme remains to be defined. However, fucosyltransferase VI may have
lectin-like functions in WPBs, in which sugar-binding properties are conserved while catalytic
functions are removed, and thus participate in the sorting of the fucosylated secretory proteins
to the WPBs [1, 134].
5.8 Tissue plasminogen activator (t-PA)
t-PA is a key enzyme in fibrinolysis found to be mainly localized in the vascular ECs, from
where it is secreted both in vivo [135] and in vitro [136]. Some investigators have found that
t-PA colocalizes with VWF in WPBs [137, 138] and released by the same agonists as VWF
[139], while another storage organelle for t-PA distinct from WPBs has been proposed [140].
5.9 Osteoprotegerin (OPG)
OPG is a member of TNF receptor family [141] and plays important roles in both skeletal and
extra-skeletal tissues. Zannettino et al. has recently discovered that OPG colocalizes with
VWF and P-selectin in the WPBs [142]. It is rapidly released in response to the proinflammatory cytokines like TNF-α and IL-1β, and thereafter facilitates the emigration of
leukocytes and platelets at the sites of inflammation and thrombous formation [142].
27
6. Methodological considerations
6.1 Subcellular fractionation is a commonly used method for organelle separation for all
types of cells and tissues. It allows the separation of the organelles based on their physical
properties, and was initially used to separate organelles derived from rat liver [143-145].
Subcellular fractionation consists of two main steps: homogenization/disruption of the cellular
material, and fractionation of the homogenate to separate the different populations of
organelles. Homogenization involves cell disruption techniques to expose the intracellular
organelles by disrupting the cell membranes. The quality of homogenization can be observed
by a phase contrast microscope, which helps in the assessment of the extent of cell disruption
[146]. The homogenized extract is then subjected to low- speed centrifugation step, which
facilitates the removal of nuclei, unbroken cells, and cell debris. The post nuclear supernatant
(PNS) thus consists of the cytosol and the other organelles in free suspension [147]. The
organelles are separated mainly by the ratio of their lipid to protein content of their
membranes at different densities[148, 149], although the difference in the composition of the
subcellular components can also affect relative densities of the fractions. Different gradient
media used for this purpose are described below.
6.1.1 Fractionation media
It was early established in the development of centrifugation -as a tool for purification of
cellular organelles, that nonelectrolyte-based media are preferable to electrolyte-based media
for reducing organelle aggregation [48, 150]. Moreover, media are generally either isoosmotic
or hyperosmotic, as compared to 0.15 M NaCl. Organelles may be regarded as “osmometers”,
allowing water moving across their limiting membranes to maintain osmotic equilibrium, thus
lysis can occur in hypoosmotic media [151].
Sucrose is the most commonly used gradient medium in subcellular fractionation, because it
is inexpensive, very soluble, and may be applied to prepare solutions that span the range of
densities of most biological organelles. Isoosmotic sucrose (0.25 M; ~9% w/v) has a density
of only 1.03 g/ml, which is at the lower-density end for biological organelles. Thus, all
isopycnic density gradient centrifugation performed in sucrose involves hyperosmotic
28
conditions, with intraorganellar water redistributing to the surrounding medium, and the
organelles shrink and increase in density [151].
Iodinated compounds increase the density of fractionation media through most of the range of
biological organelles, while maintaining reasonably low viscosity and osmolarity compared to
sucrose. Three popular and closely related compounds, metrizamide, Nycodenz, and
iodixanol, have high densities (>1.2 g/ml) exceeding that of most of the organelles and an
osmolarity of ≤400 mOsm, and reasonably low viscosity [152].
Nycodenz – the media used in this study – may be successfully used in the isolation of most
types of subcellular organelles under either isotonic or mildly hypertonic conditions[140,
153].
Ficoll 400, a chemical polymer of sucrose with epichlorohydrin, is also a commonly used
medium. Ficoll has been used to separate many different cell populations, and is frequently
applied as an additive to media containing other density-modifying agents [152].
Percoll is high-molecular weight medium based on silica particles coated with
polyvinylpyrrolidone (PVP) to reduce its adsorption to biological organelles [154]. The
gradient medium neither penetrates the organelle membranes nor influences the density of the
organelles [154]. Percoll solutions are self generating gradients, thus the gradients are
generated during the centrifugal run.
6.2 ELISA (Enzyme linked immunosorbent assay) is a highly sensitive immunoassay that
detects antigens using antibodies labelled with enzymes. Sandwich ELISAs are the most
useful of the immunosorbent assays, because they are 2 - 5 times more sensitive than those in
which antigen is directly bound to the solid phase. To detect antigen, the wells of microtiter
plates are coated with specific (capture) antibodies, blocked with a proteinaceous solution to
prevent unspecific binding, followed by incubation with test solutions containing antigen.
Unbound antigen is washed off and a different antigen-specific antibody (detecting)
conjugated to an enzyme for a developing reagent is added, followed by another incubation.
Unbound conjugate is washed off and substrate is added. The degree of substrate hydrolysis is
measured, which is proportional to the amount of antigen in the test solution [151].
29
6.3 Immunocytochemistry
This technique allows the detection and visualization of specific proteins in cells and tissues
by using specific antibodies. The method relies on proper fixation of cells to retain cellular
distribution of antigen, and to preserve cellular morphology. After fixation, the cells are
exposed to the primary antibody directed against the protein of interest. Permeabilizing
reagents may be added to ensure antibody access to intracellular epitopes. After incubation
with the primary antibody, the unbound antibody is washed off, and the bound primary
antibody may then be detected by incubation with a fluorescently tagged secondary antibody
directed against the primary antibody. After removal of excess secondary antibody, the
specimen is ready for analysis in a fluorescence microscope. Once the conditions for
observing specific immunolocalization have been identified for a given antibody and cell
type, multiple staining with antibodies conjugated to different fluorochromes can also be
employed to compare localizations [151].
6.4 Tranfection
Transfection refers to gene transfer – usually foreign DNA – by processes that make a cell
transiently permeable to DNA. A large number of methods are being used to introduce
eukaryotic DNA into cultured mammalian cells, including calcium phosphate transfection,
DEAE-dextran-mediated transfection, liposome-mediated transfection, and electroporation.
The first three procedures produce a chemical environment that results in DNA attaching to
the cell surface, where it is subsequently endocytosed by poorly characterized pathways. The
parameters for transfecting cells by these techniques vary for each different cell type, and
therefore need to be carefully optimized.
Calcium phosphate transfection method was used in this study. This relies on the uptake of
DNA as calcium phosphate-DNA complexes, and thus facilitates the binding of these
complexes to the outer membrane of a cell where after it is taken up by the cell via
endocytosis. Reporter genes are commonly used to optimize the parameters in the DNA
transfection[155]. These may not only be used to measure the activity of a gene, but also to
visualize the location of the protein that is expressed. Green fluorescent protein (GFP) is used
to directly visualize proteins in living cells, and provides a simple method for evaluating the
30
percentage of cells that have been transfected, and serve as a marker for cells within a
population that have received the transfected DNA [156].
6.5 Retroviral transduction
Retroviral transduction refers to the ability of a retroviral vector to insert itself into the host
DNA as a provirus (viral DNA integrated into the host genome), and is thus used to introduce
a nonviral gene into the mitotic cells in vivo or in vitro. Once integrated into the host cell the
vector provirus makes mRNA(s) of the gene(s) of interest which are subsequently translated
into protein with the host transcription/translation machinery.
Retroviral transduction is commonly used for the stable and efficient expression of one or
more genes into cells that cannot be easily transfected. Retroviral transduction is regarded as a
very efficient method to deliver single DNA expression constructs to a wide range of
mammalian cells. In this study, an amphotropic virus was used, which can infect more than
one species.
Packaging cell lines are used to produce virus, since the vectors are replication incompetent,
and thus do not encode the structural genes for the viral envelope or for the RTase. The
packaging cells are, however, stably transfected with these genes, making it possible to
produce replication incompetent viruses. Since the envelope and the RTase genes are not
present in the viral RNA, the virus is not able to multiply upon infection of cells which is not
a packaging cell line. Most cells may serve as packaging cell lines, but usually those that are
easily transfected are used. There are a number of packaging cell lines available, including
primate human 293 cells, COS cells, and mouse NIH 3T3 fibroblasts. In this study, Phoenix
293T cells derived from human kidney epithelial cells were applied. These are transformed
with T4 bacteriophage to make them immortal. The supernatant produced by these
retrovirally transfected cell lines comprises a virus stock, which may then used to infect the
target cells in vivo or in vitro [156].
31
6.6 FACS (fluorescence activated cell sorting) is a method in which a heterogeneous
population of suspended cells are characterized and separated based upon the intensity of
fluorescence they emit while passing single file through an illuminated volume. In the most
common scenario, one or more lasers interrogate each particle, and at a minimum, the system
measures the degree and direction of scattered light indicating the size, shape and structure of
the particle. These particles may further be stained with dyes or antibodies conjugated to
fluorochromes to obtain additional biological information. Precise optical and electronic
sensors collect the fluorescent pulses and scattered light, convert them into digital values and
relay them to a computer for analysis. Cells conjugated with fluorescence-based protein
reporters such as the GFP can also be effectively analysed and sorted to monitor both their
transfection efficiency and protein expression levels.
6.7 SOFA ( Single organelle flow analysis) is used to detect fluorescently labelled organelles
[157] and allows for rapid analysis of organelles by flow cytometry [158]. Using this
technique, vesicles can be analyzed for more than one parameter at a time, and co-localization
studies can be executed on a single organelle. SOFA also provides quantitative information,
and the possibility for statistical analysis of the data. Endosomes were amongst the first
organelles to be analyzed by SOFA [159].
6.8 FAOS (flourescence activated organelle sorting) can be used to sort labelled intracellular
organelles from a cell homogenate, or following a subcellular fractionation procedure. Flow
cytometry has been used earlier in the analysis as well as sorting of the single organelles
[149]. FAOS thus allows the isolation of highly purified organelle populations, and has been
previously been used to separate and sort apical and basolateral endocytic vesicles from
MDCK cells [160].
32
7. Aim of the study
The purpose of this work was to establish an enrichment protocol for WPBs. Although a
number of protocols are available for the generation VWF-containing fractions from cell
homogenates, isolation of highly purified WPBs for analytical and preparative purposes has
not been described. In this study, we have tried to identify populations of fluorescently
labelled WPBs by retrovirally transducing VWF-GFP fusion protein into HUVECs. These
cells were then homogenized, separated by density gradient centrifugation and the fractions
analysed by SOFA. This method may serve as a basis for fluorescence-activated organelle
sorting.
33
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Manuscript
Identification of endothelial Weibel-Palade bodies by
single organelle flow cytometry
Mrinal Joel1,2, Inger Øynebråten1, Marjan J. Veuger1, Gøril Olsen1, Oddmund
Bakke2, Guttorm Haraldsen1, and Espen S. Bækkevold1.
1
Laboratory for Immunohistochemistry and Immunopathology (LIIPAT), Institute of
Pathology, University of Oslo, Rikshospitalet University Hospital, Norway.
2
Department of Molecular Biosciences, University of Oslo, Norway
Abstract
Vascular endothelial cells contain typical, elongated vesicles called Weibel-Palade bodies
(WPBs), which serve as a storage compartment for biologically active factors involved in
hemostasis and inflammation. While the content of WPBs is dominated by the protein
von Willebrand factor (VWF), it also contains subsets of other proteins including Pselectin, CD63, interleukin-8 (IL-8) and endothelin. However, many additional proteins
are contained in WPBs, but our knowledge about the WPB proteome is scarce.
Although several protocols are available for generation of WPB-containing
fractions from cell homogenates, isolation of highly purified WPBs for analytical or
preparative purposes has not been described. By density gradient centrifugation of human
umbilical vein endothelial cell (HUVEC) homogenates, a particle was identified that
contained high concentrations of VWF-antigen. These particles resembled WPBs by
being quite dense (peaking at 1.11-1.12 g/ml in Nycodenz gradients), and VWF was
released by Triton treatment, suggesting that the isolated particle consisted of membrane
enclosed VWF. Furthermore, these particles were of shape similar to WPB and are VWFpositive. Moreover, HUVECs were retrovirally transduced with a vector encoding a
VWF-green fluorescent protein chimera (VWF-GFP) to fluorescently label WPBs, and by
single organelle flow analysis (SOFA), we found a population of GFP+ particles that
exhibited a similar density distribution.
This method may serve as a basis for fluorescence-activated organelle sorting
(FAOS), and could provide a new tool for understanding the biology of WPBs.
2
Introduction
The endothelial-specific regulated secretory organelles known as Weibel-Palade bodies
(WBPs), were first described more than 40 years ago [1], but they remain largely
uncharacterised despite their key role in hemostasis and inflammation[1-3]. In particular,
knowledge about their biosynthesis, regulation and proteome is scarce.
The content of WPBs is dominated by the hemostatic factor von Willebrand factor
(VWF)[4], which forms highly condensed multimers that appear to be visible as parallel
proteinaceous tubules in the lumen of WPBs, giving rise to the characteristic striated,
elongated, “cigar-shape” of WPBs [3]. In fact, VWF might be crucial in WPB formation,
as heterologous expression of VWF in non-endothelial cells induces WPB-like
organelles[5-7].
In addition, WPBs have been shown to contain a number of other proteins, differing both
in structure and function. The best characterized membrane molecule of WPB is the
leukocyte adhesion molecule P-selectin [8], which is translocated to the luminal surface
of endothelial cells in response to vascular injury. Furthermore, the leukocyte
chemoattractant interleukin-8 (IL-8) [9] and eotaxin-3 [10] are stored in WPBs, and
released during inflammatory reactions[9]. WPBs also harbour the adhesion molecule
CD63/lamp3 [11], and the vasoconstrictor Endothelin-1[12] as well as its processing
enzyme endothelin-converting enzyme [13]. There are also studies showing the tissuetype plasminogen activator (tPA) localized to WPBs [14], whereas other reports
concludes that tPA is released from a distinct storage granule[15] - thus this issue
remains controversial. Furthermore, an enzyme involved in endothelial selectin-ligand
synthesis, α1,3-fucosyltransferase VI, is accumulated in WPBs [16], but the functional
significance of the post-Golgi localization of this enzyme remains to be defined.
Several membrane-associated proteins involved in organelle trafficking and regulated
exocytosis have also been localized to WPB-membranes. This includes the GTPases
Rab3D and Rab27A suggested to be involved in WPB/granule formation[17] and granule
movement to docking sites, respectively[18, 19]. Furthermore, inhibition of the vesicle-
3
associated membrane protein (VAMP)-3[20] and Munc 18c[21] - two proteins critical for
intracellular fusion events - interfered with the exocytosis of the WPB, suggesting that
both factors are associated with WPBs.
However, it is reasonable to assume that many additional proteins are contained in and
associated with WPBs, and a more systematic identification is needed to explore the
biology of the WPBs and their function in endothelial cells. The recent refinement of
techniques enabling separation of thousands of proteins by 2-D gel electrophoresis, and
analysis from minute amounts of material by mass spectrometry, holds the possibility of
identifying most, if not all, proteins associated with WPBs. However, to reduce the
proteome complexity, organelle enrichment is needed, and although several protocols are
available for generation of WPB-containing fractions from cell homogenates, isolation of
highly purified WPBs for analytical or preparative purposes has not been described.
Fluorescence-activated organelle sorting (FAOS) has been shown to yield highly purified
preparations of different endosome subtypes. In fact, FAOS has been used to separate
apical and basolateral endosomes [22], as well as highly enriched fractions of mast cell
secretory lysosomes[23] .
Given the large, rod-shaped (0.2 x 2-3 µm) structure of WPBs, we considered this
organelle an attractive candidate for flow-cytometric detection and subsequent sorting.
We approached this by fluorescently labelling WPBs with a VWF-green fluorescent
protein chimera (VWF-GFP) retrovirally transduced into human umbilical vein
endothelial cells (HUVECs). These cells were then homogenized, subjected to density
gradient centrifugation, and the fractions analysed by SOFA. We found a population of
GFP+ particles in the fractions containing WPBs, constituting more than 8% of the total
number of particles in these preparations. This protocol may serve as a basis for FAOS,
and could provide a new tool to elucidate the biology of the WPBs.
4
Materials and methods
Chemicals, growth factors, and other reagents
MCDB 131, hydrocortisone, collagenase A, EDTA, Tween20, Triton X-100, sucrose,
imidazole, porcine dermal gelatine, puromycin, penicillin-streptomycin, bovine serum
albumin (BSA) and fetal calf serum (FCS) were purchased from Sigma. Fungizone was
purchased from Invitrogen, and gentamicin, trypsin and IMDM medium were from
Biowhittaker. Epidermal growth factor (EGF) and fibroblast growth factor (FGF) was
obtained from R&D systems and PeproTech, respectively. Nycodenz was obtained from
Axis-Shield. Dimethyl sulfoxide(DMSO) was purchased from Merck, and HEPES buffer
was from GIBCO.
Primary antibodies and secondary reagents
The primary antibodies are listed in Table 1. Aminomethyl coumarin acetic acid
(AMCA)-conjugated goat anti-rabbit IgG was purchased from Vector Laboratories, and
indocarbocyanine (CY3)-conjugated donkey anti-mouse IgG was from Jackson
Immunoresearch. Polyclonal rabbit anti-VWF conjugated with HRP was used as
detecting antibody used in ELISA (DAKO, Denmark).
Cell culture
Umbilical cords were obtained from the Dept. of Gynaecology and Obstetrics at
Rikshospitalet, and HUVECs were isolated by collagenase digestion as described by Jaffe
et al. [24], and cultured in MCDB 131 containing 7,5 % FCS, 10 ng/ml EGF, 1 ng/ml
FGF, 1 µg/ml hydrocortisone, 50 µg/ml fungizone, and 0.25 µg/ml gentamicin. The cells
were maintained at 37°C in a humid 95% air, 5% CO2 atmosphere, and passaged by
trypsinization.
Cell homogenization
HUVECs were trypsinized and suspended in 2.5 ml of homogenization buffer (3mM
imidazole, 0.25mM sucrose, HEPES buffer, pH 7.4). The cells were homogenized by
5
shearing through a 25G syringe 5-6 times. The homogenate was then centrifuged at 800g,
7 min to generate the post-nuclear fraction (PNS).
Nycodenz density gradient centrifugation
Density centrifugation was performed by modifying a procedure described by Emeis et
al. [15]. A Nycodenz stock solution of 30% (wt/vol) was prepared in 5 mM Tris-HCl (pH
7.4). From this stock solution, dilutions of Nycodenz of 27.5% to 5% (in increments of
2.5%) were prepared in the same buffer. A density gradient was made in 14 ml
ultracentrifuge tubes, starting with one ml of 30% Nycodenz, and then in ten steps of one
ml from 27.5% to 5%, and left to equilibrate overnight at 4°C. PNS supernatant or cell
homogenate was layered on top of the gradient, and the gradient was centrifuged for
150 min, 41,000 rpm (202,000 gav) at 4°C in a Beckman OptimaTM LE-80K
ultracentrifuge equipped with a SW-41 rotor. Subsequently, 1 ml fractions were collected,
starting at the top, and stored frozen. The density of the fractions was determined in a
refractometer (Carl Zeiss), and ranged from 1.03-1.17 g/ml.
VWF ELISA
To determine the concentration of VWF in gradient fractions, we established a solidphase sandwich ELISA. Polystyrene 96-well plates were coated overnight with 75 µl/well
of polyclonal rabbit anti-VWF diluted in PBS (9.7 µg/ml) at room temperature (RT),
blocked by incubation with 100 µl/well of 1% BSA in PBS 2 h at RT, and washed with
0,05% Tween-20 in PBS. Samples (75µl/well, undiluted and 1:10 in ELISA buffer (0.5%
BSA, 0.05% Tween-20, 1 %Triton in PBS) were loaded, and the plates were incubated
overnight at RT. The plates were then washed with washing buffer, and incubated with
polyclonal rabbit anti-VWF conjugated with HRP (75 µl/well, 1,3x103µg/ml) diluted in
ELISA buffer for 2 h at RT. After washing, HRP activity was measured using equal
quantities of TMB peroxidase substrate and peroxidase substrate solution B (KPL). The
plates were analysed with a Sunrise microtitreplate reader (TECAN).
6
Human pro-cathepsin B immunoassay
Subcellular fractions were analysed for lysosomal contamination by using a human procathepsin B ELISA kit (R&D systems) according to the manufacturer. Briefly, fractions
or standard was diluted in assay diluent, added to the plates (50µl/well), and incubated for
2 h at 4°C. The plate was then washed with the assay washing buffer, and conjugate procathepsin B conjugated to horseradish peroxidase (HRP) (200µl/well) was added, after
which the plates were incubated for 2 h at 4°C. Following washing, substrate solution
(200 µl/well) was added and incubated for 30 min at RT. Stop solution (50µl/well) was
added to each well and the plates were analysed as described above.
Immunocytochemistry
HUVECs cultured on gelatin-coated chamber slides were fixed in acetone (10 min, RT)
and air-dried (20 min, RT). The slides were then immunostained with primary antibody
(60 min, RT) washed and then incubated with the fluorochrome-conjugated secondary
antibody (AMCA-conjugated Goat anti-rabbit Ig (30 min, RT), and mounted in polyvinyl
alcohol. Microscopy specimens of density-gradient fractions were prepared by
centrifugation of 600µl of each fraction at 13000 rpm for 10 min, resuspending the pellet
in PBS, and loading 50 µl on poly-lysine-coated slides. The slides were air-dried
overnight, and prepared for immunostaining as described above using secondry antibody
(CY3)-conjugated donkey anti-mouse IgG. Negative controls were performed by
replacing the primary antibody with isotype and concentration matched irrelevant control
antibodies. Microscopy was performed with a Nikon E-800 fluorescence microscope
equipped with F-view camera (Soft-Imaging System GmbH).
Generation of retrovirus
A moloney murine leukemia based retroviral system was used generating amphitrophic
replication incompetent viruses. Phoenix-A cells (generously provided by M. H. M.
Heemskerk, Leiden University Medical Center, The Netherlands) were grown in IMDM
medium supplemented with 10% FCS and penicillin-streptomycin. Helper-free
recombinant retrovirus was produced after transfection of the retroviral vector LZRSVWF-GFP DNA (generous gift from Dr. Jan A.van Mourik, University of Amsterdam,
7
The Netherlands) into Phoenix-A cells by calcium-phosphate transfection according to
the manufacturer (Invitrogen)[25].Two days after transfection, cells were cultured in
medium with 2µg/ml puromycin. Between 10-14 days after transfection, puromycin
resistant (6x10 6) cells were plated per 10 cm petri dish in 10ml IMDM without
puromycin. After 24 hours, medium was refreshed and the next day cell supernatants
containing viruses were harvested and clarified of cell debris by centrifugation (2500rpm,
5 min) followed by 0.45µm filtration. The viral supernatants were frozen at -70°c.
Retroviral transduction
Exponentially growing HUVECs were transduced using retroviral supernatants based on
the method described by Hanenberg et al with minor modifications [26].
To facilitate virus uptake, DEAE-dextran (10µg/ml) was added to 3 ml of the thawed
viral supernatants and incubated 10 min on ice. Virus-DEAE complexes were added to
sub-confluent HUVECs grown in 6 well plates (3 ml/well), and incubated for 4-5 h at
37°c. The viral supernatants were then removed, and the cells were cultured with MCDB
131 as described above. Three to five days after transduction, GFP positive cells were
purified by FACS sorting using a FACS Diva cell sorter equipped with an argon ion
laser, TurboSort Plus option, and Diva software (Becton Dickinson). In order to ensure
that the virus was completely removed, these cells were washed 3-4 times in 5ml of PBS,
and collected after centrifugation.
SOFA
The subcellular fractions of the VWF-GFP transduced cells were analyzed using the
FACS Diva cell sorter. The 488 nm laser was tuned to 215 mW output, and a 70 µm
nozzle tip was used with a sheath pressure of 12 psi. The threshold was set on forward
light scatter, and a 10% ND filter was used. The sort gate was set using FL1 (530/30BP)
parameter. Subcellular fractions of untransduced HUVECs served as the negative
controls.
8
Results
Density gradient centrifugation of HUVECs.
To enrich WPBs for flow cytometric analysis, we performed density gradient
centrifugation of cell homogenates. In initial attempts, homogenates of fourth passage
HUVECs were separated on Percoll gradients as described by others [27] , however
subsequent analysis of VWF revealed a quite diffuse distribution throughout the gradient
(data not shown). We thus adapted a protocol using Nycodenz gradients [15], in a density
range of d = 1.03 - 1.14 g/ml. After ultracentrifugation, the fractions were diluted 1:10 in
buffer containing Triton X-100 and analysed with a VWF-ELISA. In this gradient, VWF
predominantly peaked at d = 1.11 - 1.12 g/ml (Figure 1a). In addition, VWF revealed a
second, smaller peak around d = 1.06 g/ml. To investigate whether both peaks contained
membrane enclosed VWF, ELISA was also performed on non detergent-treated fractions,
revealing no accessible VWF antigen in the denser (d = 1.11 - 1.12 g/ml) fractions
without membrane permeabilization. By contrast, the more buoyant fraction allowed
detection of VWF without permeabilization, indicating that the majority of the membrane
enclosed VWF is present in the denser fraction.
To assess whether the detected VWF could be ascribed to pathways involved in
degradation, an ELISA to the lysosomal marker pro-cathepsin B was applied. However,
this marker peaked at d = 1,03-1,04 g/ml (Figure 1b), thus distributing differently from
that of VWF.
9
Immunocytochemical analysis of density gradient fractions
To better characterize the fractions, HUVECs were fractioned as described above, before
immunocytochemical analysis using antibodies towards marker proteins of different
subcellular organelles. Immunofluorescence staining with anti-VWF on the dense (d =
1.11 - 1.12 g/ml) fraction (Figure 2c) revealed the presence of a quite homogenous
population of particles resembling the WPBs found in identically stained, intact HUVECs
(Figure 2a). At this level of resolution, the particles were indistinguishable from WPBs,
both in terms of size and VWF content. Interestingly, such particles were not present in
the buoyant (d = 1.06 g/ml) fraction (Figure 2b), further indicating that the VWF detected
with ELISA in these fractions is not associated with WPB-like structures. Moreover,
immunostainings with an irrelevant primary antibody gave no signal in the dense fraction
(shown in Figure 2d).
To assess the degree of contamination by other subcellular organelles in the WPB-rich
fractions, we stained for EEA-1 – a marker of early-endosomes[28, 29]. Although
readily detectable in the intact HUVECs (Figure 3a), no EEA-1-reactivity was observed
in the fractions containing VWF (Figure 3b,c). However, given the small size of EEA-1+
particles, we experienced difficulties identifying them even in crude homogenates, thus
these results should be interpreted with caution. Furthermore, an antibody to a Golgi
marker, Golgi 58K, which is a microtubule-binding protein also named
formiminotransferase cyclodeaminase (FTCD) [30, 31], clearly decorated perinuclear
structures in HUVECs (Figure 3d), but produced only a scattered, low signal in the WPBrich fractions (Figure 3e,f). Application of antibodies to the lysosomal-associated
membrane protein (LAMP)-2 [32, 33] showed a distinct, granular staining of HUVECs
(Figure 3g), but only a few lysosome-like particles were detected in the dense fractions
(Figure 3i). We also analysed the late-endosomal marker LBPA (lysobisphosphatidic
acid)[34]. Although late-endosomal structures were heavily stained in intact HUVECs
(Figure 3j), no reactivity was seen in either the buoyant or dense fractions (Figure k,l).
10
Retroviral transduction of HUVECs with VWF-GFP
To enable further purification of WPBs, these were fluorescently labelled by retrovirally
transduced VWF-GFP chimeric protein into HUVECs. Subsequent fluorescence
microscopy revealed a green fluorescence signal in rod-shaped organelles throughout the
cytoplasm (Figure 4a). Co-staining with anti-VWF followed by an AMCA-conjugated
secondary antibody revealed rod-shaped organelles with nearly complete colocalization
with the GFP-signal, consistent with targeting of VWF-GFP to WPBs [25].
To obtain pure populations of VWF-GFP-transduced cells, GFP-expressing cells were
isolated by FACS sorting. Our transfection method yielded 25-30% GFP+ HUVECs
(Figure 5b) compared to untransduced cells (Figure 5a). After sorting of the GFP+ cells,
populations with purity of 95-98% were obtained (Figure 5c).
Analysis of WPBs by SOFA
We next sought to determine if flow cytometry could be used to detect (and potentially
purify) fluorescently labelled WPBs. Sorted GFP+ HUVECs were fractionated on
Nycodenz gradients and analysed by SOFA, detecting GFP at 530 nm together with
forward light scatter (FSC). Identical fractions of untransduced HUVECs served as
negative controls, and were used to define the detection gate. The detection gate also
included a pulse-width analysis to exclude aggregated particles (not shown). When
applying this gate to the fractions of GFP+ HUVECs (Figure 6, right column), we found
the number of GFP+ particles in the most buoyant gradient layer (fraction, d=1.03g/ml) to
be similar to that of control homogenates. However, when analysing the increasingly
more dense fractions, we found elevated GFP-signal in fraction with densities of 1.04,
1.08, 1.09 and 1.10g/ml – and peaking in the dense fractions with a density of 1.11 and
1.12g/ml, where GFP+ particles represent >8% of the total number of events detected.
11
The percentage of GFP+ particles in each fraction was plotted (Figure 7) showing a
pattern of distribution closely resembling that of VWF (Figure 1), enriched in the dense d
= 1.11 - 1.12 g/ml fractions.
Unfractionated PNS from VWF-GFP-transduced HUVECs was also analysed by SOFA,
but the low percentage of GFP+ particles in these preparations did not allow confident
discrimination from autofluorescent particles in the PNS from untransduced cells.
12
Discussion
This study describes a novel method for identification of fluorescently tagged WPBs,
which can be applied for flow cytometric organelle sorting. This protocol requires
homogenisation of HUVECs expressing VWF-GFP, fractionation of the PNS by density
gradient centrifugation, and finally SOFA analysis. The proteins contained in WPBs play
essential roles in regulation of thrombosis and thrombolysis, platelet adherence,
modulation of vascular tone and blood flow, as well as immune and inflammatory
responses by controlling leukocyte recruitment. Our strategy thus offers a new tool to
study the protein content in this functionally important organelle.
By modifying a published protocol for ultracentrifugation of cell homogenates in a
Nycodenz gradient[15], we obtained fractions enriched for VWF, as measured by a
specific sandwich ELISA. We found that VWF was enriched in two density areas – one
dense and one buoyant. Our data indicates that only the dense fractions (d = 1.11 - 1.12
g/ml) contain VWF in sedimentable, membrane bound particles, from which the protein
could be released by Triton treatment[5]. Immunocytochemical analysis further revealed
that these particles closely resembled WPBs found in intact cells, thus strongly indicating
that these fractions indeed contain enriched WPBs. We also found VWF in buoyant (d=
1.06 g/ml) fractions, but the detection was independent of Triton treatment, and thus the
origin of the VWF in these fractions is currently unknown. Furthermore, we performed
immunostainings of the dense fractions with a panel of antibodies to different
intracellular membrane markers, including Golgi, early- and late endosomes, as well as
lysosomes, but we were unable to detect any major populations of contaminating
organelles.
However, to obtain highly purified preparations of WPBs, a specific and targeted
isolation approach is needed. Initially, we considered application of immunomagnetic
beads armed with antibodies to epitopes on the cytoplasmic face of WPBs to extract the
organelles from the enriched fractions. Several suitable reagents are available, including
an antibody to the C-terminal, cytoplasmic tail of P-selectin. However, HUVECs require
activation by IL-4 to express P-selectin in vitro[35] (and our unpublished data), and such
13
stimulation could potentially induce additional changes to the WPB protein content.
Furthermore, Rab27 is reported to coat a subset of WPBs[36], and thus be available for
antibody binding. However, our analysis of Rab27 in HUVECs revealed quite variable
staining on WPBs (data not shown). Thus, antibody-mediated isolation was not pursued.
Fluorescent protein labelling of specific organelles is a commonly used approach
for detection of intracellular structures, and VWF fused to GFP has been shown to
localize to WPBs[25]. Thus, by retroviral transduction we introduced a vector encoding
VWF-GFP into HUVECs, and confirmed the reported colocalization with endogenous
VWF in WPBs. By subcellular fractionation of the transduced cells, followed by SOFA
analysis of GFP, we observed a population of fluorescent particles that was not present in
untransduced HUVECs. Further, when the separate Nycodenz density fractions were
analysed and the percentage of fluorescent particles was estimated, we found that the
distribution of these particles closely resembled that of WPBs. In fact, in the dense
fractions, particles emitting green fluorescence constitute >8% of the recorded events.
Although only indirect data, it is tempting to speculate that the GFP+ particles could
correspond to WPBs. However, future efforts will be directed towards sorting of the
particles, followed by direct analysis of their ultrastructure and content of WPB proteins.
In the field of proteomics, there has been a long-standing interest in the analysis of subcellular fractions and organelles. More importantly, the establishment of subcellular
proteomes provides reference maps that are useful tools for researchers working on a
given organelle or cellular compartment. Fractionation allows the reduction of proteome
complexity of the sample, compared to a single step analysis. Our results indicate that
FAOS could be applied to obtain pure samples of WPBs. One technical limitation of
FAOS, however, is that depending on the abundance of the organelle, it may require large
quantities of unsorted material for preparative purposes. Estimates in HUVECs indicate
that they may contain < 200 WPBs per cell [37]. However, preliminary analysis of our
SOFA data indicates that the abundancy of GFP+ WPBs in the pre-sorted material is far
lower, with only about one event per 100 cells homogenized (data not shown). This may
reflect a tremendous loss of material in the procedures preceding SOFA. However,
analysis of the retrovirally transduced cells also revealed a quite dramatic reduction of
14
GFP in WPBs, both in terms of intensity and number of green organelles, during
passaging and expansion. In particular, cells frozen in liquid N2 after transduction,
showed very low levels of VWF-GFP after thawing and continued culturing. Thus, future
efforts will have to be made to increase the stability of transduction. However, recent
advances in proteomic technologies allow for a direct analysis of proteins or peptides
without the requirement for gel separation and extraction [38]. These highly sensitive
techniques require much less starting material, and may offer a good complement to
VWF FAOS.
In conclusion, our study provides the basis for specific isolation of WPBs from ECs by
means of flow cytometry. Purified preparations of WPBs may provide a tool for
understanding their biology, including resolving their proteome.
15
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20
Table I. Primary antibodies used for immunostaining:
Specificity
Designation
Specification
Source
VWF
A.0082
Rabbit IgG
DAKO, Denmark
VWF
F8/86
Mouse IgG1
DAKO, Denmark
Golgi
G-2404
Mouse IgG
Sigma chemicals
LAMP-2
H4B4
Mouse IgG1
Developmental studies
Hybridoma Bank, Iowa
LBPA
6C4
Mouse IgG
Dr. J. Gruenberg,
University of Geneva,
Switzerland
EEA1
Irrelevant
14
MOPC-21
Mouse IgG1
BD Biosciences, USA
Mouse IgG1
Sigma chemicals, USA
control
21
Figure legends
Figure1. Gradient centrifugation of HUVEC homogenates on a Nycodenz density
gradient. (a) The amount per fraction of VWF as measured by ELISA and optical
density (O.D.) reading. Samples diluted 1/10 with Triton is shown with triangles,
undiluted samples are shown with squares. The values are shown as mean ± SD (n=3)
from an individual HUVEC culture, and is representative of 3 independent experiments.
(b) The amount per fraction of pro-cathepsin B as measured by ELISA and optical
density (O.D) reading. The values are shown as mean ± SD (n=3)
Figure 2. Immunostaining for VWF in intact HUVECs and VWF-containing
Nycodenz fractions.
Immunostaining with VWF antibody of (a) trypsinized HUVECs, (b) buoyant (d = 1.06
g/ml) Nycodenz fraction, (c) dense (d= 1.11 g/ml) Nycodenz fraction, and (d) negative
control of the latter incubated with an irrelevant primary antibody. The
immunofluorescent pictures were acquired with a 600X original magnification.
Figure 3. Immunodetection of non-WPB organelles in intact HUVECs and VWFcontaining Nycodenz fractions.
Immunostainings of trysinized HUVECs (a, d, g, j), buoyant (d = 1.06 g/ml) Nycodenz
fractions (b,e,h, k), and dense (d= 1.11 g/ml) Nycodenz fractions (c ,f, i,l) with antibodies
specific for early endosomes (EEA1), Golgi 58K, lysosomes (LAMP-2), and late
endosomes (LBPA) as indicated. The immunofluorescent pictures were acquired with a
600X original magnification.
Figure 4. Expression of VWF-GFP in transduced HUVECs.
Immunocytochemical staining for VWF (red) in VWF-GFP-transduced HUVECs. The
picture was acquired with a 400X original magnification. Magnified selection is shown as
single-exposures are shown in the right column.
22
Figure 5. FACS sorting of VWF-GFP transduced HUVECs.
HUVECs analysed by FACS before (a) and after (b) VWF-GFP transduction, with the
gate applied for FACS sorting indicated. Secondary analysis of the sorted GFP+ fraction
is shown in (c). The percentage of GFP+ cells is indicated, and is representative of 3
independent experiments.
Figure 6. SOFA analysis of Nycodenz fractions from VWF-GFP transduced
HUVECs.
The Nycodenz fractions (density in the upper right corner) of untransduced (left column)
and VWF-GFP transduced HUVECs (right column) were analyzed by flow cytometry.
The green fluorescence was detected in FL1 (530 nm) and compared to FSC. The
percentage of events within the detection gate is given in the lower right corner.
Figure7. Distribution of GFP+ particles in Nycodenz fractions as determined by
SOFA. The percentage refers to the number of green particles within the GFP-gate
(defined in figure 6) compared to the total number of particles.
23
Figure 1.
a.)
4
3.5
undiluted
3
1/10 dilution
O.D units
2.5
2
1.5
1
0.5
0
1.01
1.03
1.05
1.07
1.09
1.11
1.13
1.15
Density (g/ml)
b.)
O.D units
1.6
1.4
1.2
1
0.8
0.6
0.4
0.2
0
1.01
1.03
1.05
1.07
1.09
Density (g/ml)
1.11
1.13
1.15
Figure 2.
a
c
b
d
Trypsinized
HUVECs
Buoyant fraction
(d=1.06g/ ml)
Dense fraction
(d= 1.1 g/ ml)
b
c
d
e
f
g
h
i
j
k
a
EEA-1
Golgi
LAMP-2
LBPA
l
Figure 3.
Figure 4.
a
VWF
GFP
VWF
VWF
GFP
GFP
Figure 5.
a)
b)
0%
GFP
GFP
c)
98%
GFP
30%
Untransduced
VWF-GFP
Transduced
Figure 6.
d=1.03g/ml
d=1.03g/ml
0.1%
1.2%
d=1.04g/ml
d=1.04g/ml
0.1%
4.9%
d=1.05g/ml
d=1.05g/ml
0.2%
4.7%
d=1.06g/ml
d=1.06g/ml
0.2%
3.6%
d=1.07g/ml
d=1.07g/ml
0.3%
d=1.08g/ml
0.3%
4.6%
d=1.08g/ml
5.2%
Figure 6.
VWF-GFP
Transduced Continued
Untransduced
d=1.09g/ml
0.6%
d=1.10g/ml
0.8%
d=1.11g/ml
0.5%
d=1.12g/ml
0.3%
d=1.13g/ml
0.1%
d=1.14g/ml
0.1%
d=1.09g/ml
5.7%
d=1.10g/ml
5.6%
d=1.11g/ml
7.5%
d=1.12g/ml
8.7%
d=1.13g/ml
5.8%
d=1.14g/ml
2.6%
O.D units
Figure 7.
10
9
8
7
6
5
4
3
2
1
0
1.01
Density (g/ml)
1.03
1.05
1.07
1.09
1.11
Density (g/ml)
1.13
1.15