Microparticles: mediators of cellular and environmental homeostasis

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UvA-DARE (Digital Academic Repository)

Böing, A.N.

Publication date

2011

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Final published version

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Böing, A. N. (2011). Microparticles: mediators of cellular and environmental homeostasis.

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icroparticles

ediators of cellular and

environmental homeostasis

Anita N. Böing

A.N

. Böing 2011

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Microparticles

mediators of cellular and environmental

homeostasis

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Microparticles, mediators of cellular and environmental homeostasis Anita N. Böing

PhD Thesis, University of Amsterdam – with summary in Dutch ISBN: 978-94-91211-12-6

Cover: Microparticles released from 29EGFP-expressing MCF-7 cells (transmission electron microscopy; Edwin van der Pol)

Printed by: Ipskamp Drukkers

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Microparticles

mediators of cellular and environmental

homeostasis

academisch proefschrift

ter verkrijging van de graad van doctor aan de Universiteit van Amsterdam op gezag van de Rector Magnificus

prof. dr. D.C. van den Boom

ten overstaan van een door het college voor promoties ingestelde commissie,

in het openbaar te verdedigen in de Agnietenkapel op woensdag 27 april 2011, te 14:00 uur

door

Anita Nathalie Böing

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Promotiecommissie

Promotor: Prof. dr. A. Sturk

Co-promotor: Dr. R. Nieuwland

Overige Leden: Prof. dr. J.W.N. Akkerman Prof. dr. M. Diamant

Prof. dr. A.G.J.M. van Leeuwen Prof. dr. J.C.M. Meijers Prof. dr. C.J.F. van Noorden Prof. dr. H. Pannekoek

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Contents

List of abbreviations

ACD acid citrate dextrose

ANOVA one-way analysis of variance APC allophycocyanine asTF alternatively spliced form of TF

AUC area under curve

B2M β-2-microglobulin

BD Becton Dickinson

bFGF basic fibroblast growth factor

BMI B lymphoma Mo-MLV insertion region

BMI body mass index

BSA bovine serum albumin

CD cluster of differentiation

CDKN cyclin dependent kinase inhibitor β-COP coatomer protein complex subunit-β

Cy3 cyanine dye 3

DAPI 4',6-diamidino-2-phenylindole

ECL enhanced chemiluminescence kit

EGFP enhanced green fluorescence protein 29EGFP 29 kDa caspase-3 with EGFP

EMP endothelial cell-derived microparticles

ESCRT endosomal sorting complex required for transport F factor

FCSi fetal calf serum heat inactivated

FITC fluorescein isothiocyanate

Flt-1 fms-like tyrosine kinase-1

FSC forward scatter

GAM goat-anti-mouse GAR goat-anti-rabbit

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Abbreviations

HEK human embryonic kidney

HELLP hemolysis, elevated liver enzymes, low platelets hpTLC high-performance thin layer chromatography

HRP horseradish peroxidise

HSA human serum albumin

HUVEC human umbilical vein endothelial cells IL interleukin

ISSHP international society for the studies of hypertension in pregnancy

KDR kinase-insert domain region

LMP leukocyte-derived microparticles

L-PC l-α-lysophosphatidylcholine L-PE l-α-lysophosphatidylethanolamine L-PS l-α-lysophosphatidylserine MCP monocyte chemoattractant protein MIF (macrophage) migration inhibitory factor MLPA multiplex ligation-dependent probe amplification

MMP monocyte-derived microparticles

MoAbs monoclonal antibodies

MP microparticles

MYC early-response (proto-oncogene) gene myc NCBI national center for biotechnology information

NFκB nuclear factor of kappa light chain enhancer in B-cells

PAK p-21 activated kinase

PARN poly-A specific ribonuclease

PBS phosphate-buffered saline

PC l-α-phosphatidylcholine PC3 procaspase-3

PCR polymerase chain reaction

PDE phosphodiesterase

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Abbreviations PE phosphatidylethanolamine

PE phycoerythrin PGDF platelet-derived growth factor PI l-α-phosphatidylinositol

PI propidium iodide

PIP2 phosphatidyl inositol 4,5-biphosphate

PM plasma membrane

PMP platelet-derived microparticles

PS phosphatidylserine PSGL-1 p-selectin glycoprotein ligand-1

PTP protein-tyrosine phosphatase

PVDF polyvinylidene difluoride

RA receptor antagonist

ROCK1 rho-associated coiled coil kinase 1

RT reverse transcriptase

s soluble

SDS sodium dodecyl sulphate

SDS-PAGE SDS-polyacrylamide gel electrophoresis

SEM scanning electron microscopy

SERP serine proteinase inhibitor

SM sphingomyelin SN supernatant

SPSS statistical package of the social science software for Windows

SSC sideward scatter

TBST tris-buffered saline-tween

TF tissue factor

TFc coagulant form of TF

TFnc non-coagulant form of TF

TGF-β transforming growth factor beta

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Abbreviations

TNF tumor-necrosis factor

TNF-R1 tumor-necrosis factor receptor 1 VEGF vascular endothelial growth factor WU rats male wistar Hsd/Cpb rats

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Chapter 1

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Chapter 1

Background

Microparticles are small cell-derived vesicles, presumed to range in size between 100 nm and 1000 nm, and released from the cell (plasma) membrane. The presence of cell-derived microparticles in blood was first observed by Chargaff et al. in 19461_{. He showed that}

‘plasma, freed from intact platelets, generates thrombin on recalcification and (that) the rate of this thrombin generation can be reduced by prior high-speed centrifugation of the plasma’. In 1967, Wolf et al. showed that high-speed centrifugation of platelet-free plasma resulted in a pellet, which triggered thrombin generation after recalcification of the plasma2_.

Originally, Wolf called this coagulant material “platelet dust”, which name was changed into microparticles by Crawford et al. in 19713_.

Especially since the 1990’s, the research on microparticles has increased tremendously. Nowadays, we know that probably all eukaryotic cell types, including blood cells such as platelets, monocytes, granulocytes, erythrocytes and endothelial cells, release microparticles. Furthermore, the occurrence of microparticles and other types of cell-derived vesicles is not limited to blood, but they are also present in other human body fluids, such as cerebrospinal fluid4_{, synovial fluid}5_{, urine}6;7 _{and mother milk}8_{. In these body}

fluids, microparticles coexist with cells in physiological and pathological conditions, but the numbers, cellular origin, composition and functions of the vesicles is different in health and disease. In this thesis the main focus is on the various functions of microparticles. Figure 1 gives an overview of the functions of microparticles.

Figure 1. The various functions of microparticles.

microparticles

angiogenesis (Chapter 5) coagulation (Chapters 7-9)

communication inflammation (Chapter 6) waste management (Chapters 2-4)

cell cell microparticles angiogenesis (Chapter 5) coagulation (Chapters 7-9) communication inflammation (Chapter 6) waste management (Chapters 2-4) cell

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Introduction and aim of the thesis

1 Why do cells release microparticles?

To answer the question why cells release microparticles, we may learn from bacteria. Gram-negative bacteria release the so-called outer membrane vesicles, containing signalling molecules, into the environment to facilitate the communication between bacteria9_{and between bacteria and eukaryotic cells}10-12_{. Furthermore, these outer membrane}

vesicles can contain bacterial virulence factors, e.g. cytolethal distending toxin, which can be delivered to (eukaryotic) host cells to kill these cells, thereby promoting bacterial survival10-12_{. Similarly, DNA-encoding virulence genes can be transferred to other}

bacteria9_{. Thus, outer membrane vesicles facilitate communication and promote bacterial}

survival.

Waste management

Cell-derived microparticles from eukaryotic cells, similar to the bacterial outer membrane vesicles, play a role in the protection against “stress” induced by either the environment (extracellular stress) or e.g. by accumulation of dangerous or redundant compounds within the cell (intracellular stress). For instance, platelets release C5b-9 complex-enriched microparticles upon incubation with the complement complex C5b-9, which can be considered as a form of extracellular stress, presumably to protect platelets from complement-induced lysis13_{. In addition, during incubation with cytostatic drugs cancer}

cells release microparticles containing elevated concentrations of these drugs compared to the cancer cells themselves14;15_{. This release of microparticles can be considered as a form}

of protection against intracellular stress. Furthermore, microparticles from healthy and viable endothelial cells contain caspase-3, which is one of the main executioner enzymes of programmed cell death (apoptosis)16_{. Since endothelial cells contain no detectable amounts}

of caspase-3 but only the inactive proenzym, procaspase-3, we hypothesized that the potentially dangerous caspase-3 is continuously removed from the endothelial cells via the release of caspase-3 containing microparticles16_{. If so, then microparticles may indeed act}

like “garbage cans”. Whether the release of caspase-3 containing microparticles contributes to cellular survival was investigated in Chapter 2 of this thesis.

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Chapter 1

the formation of a constitutively active ROCK1, which in turn facilitates myosin light chain phosphorylation and thus membrane blebbing17_{. Whether caspase-3 contributes not only to}

membrane blebbing but also to the subsequent release of microparticles was investigated in Chapter 3 of this thesis. In Chapter 4 we investigated whether microparticles from platelets, i.e. a-nucleated cells that per definition can not undergo full apoptosis including nuclear DNA fragmentation, contain caspase-3.

Exchange of genetic information

Recently, microvesicles (a term used to describe microparticles plus exosomes, particles in the presumed size range of 10 – 100 nm and released from the cell when their multivesicular bodies fuse with the outer cell membrane) were shown to contain mRNA and microRNA’s. Valadi et al. showed that mRNA in microvesicles from murine mast cells could be transferred to, and expressed by, human mast cells18_{. Also, murine embryonic}

stem cells support self-renewal and expansion of adult stem cells by vesicle-mediated transfer of RNA19_{, and microvesicles from endothelial progenitor cells activate an}

angiogenic program in endothelial cells by the transfer of mRNA20_{. Thus, similar to}

vesicles from prokaryotes, the vesicles from eukaryotic cells are able to support the exchange of genetic information between cells. In the various studies, however, a clear distinction between the role of microparticles versus exosomes was not made, so it remains to be investigated which type of microvesicle actually performs those functions.

Microparticles and angiogenesis

Several studies have shown that microparticles promote or modulate angiogenesis. For instance, platelet microparticles containing important angiogenesis-promoting growth factors, including vascular endothelial growth factor (VEGF), basic fibroblast growth factor (bFGF) and platelet-derived growth factor (PGDF), induce angiogenesis in vitro and in vivo21;22_{. In contrast, endothelial microparticles impair angiogenesis at high}

concentrations23_{, although the precise underlying mechanisms were not investigated in that}

study.

As mentioned, one of the important growth factor in angiogenesis is VEGF. VEGF is produced by several organs and cells, and during pregnancies VEGF is also produced by

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the placenta. VEGF present in plasma can bind to VEGFR-1 (fms-like tyrosine kinase

(Flt-1)) and VEGFR-2 on endothelial cells, thereby triggering angiogenesis. The VEGF receptor Flt-1 can be spliced alternatively, leading to the secretion of soluble (s) Flt-1. When sFlt-1 is present in plasma it will bind to VEGF, thereby lowering the concentration of plasma VEGF24_{. As a consequence, less VEGF will be available for the binding to VEGF receptors}

on endothelial cells, thus resulting in suppression of angiogenesis. In preeclamptic patients, the plasma levels of sFlt-1 are known to be elevated compared to women with normal pregnancies25_{. In Chapter 5, we investigated whether the elevated concentrations of sFlt-1}

are present as a truly “soluble” protein or whether sFlt-1 is also associated with microparticles in preeclamptic patients.

Microparticles and inflammation

Microparticles, generated in vitro and originating from various cell types, can stimulate the inflammatory process in several ways. They trigger the cellular production of a plethora of inflammatory mediators, such as interleukins, monocyt chemoattractant proteins (MCP) 1 and 2, tumor-necrosis factor (TNF), and matrix degrading enzymes, by transferring water-soluble second messengers26-29_{, and binding to specific adhesion receptors on cells}30-33_{. The}

ability of microparticles to affect inflammation was supported by a study of Berckmans et al. He showed that microparticles from synovial fluid of patients with rheumatic arthritis induce the release of inflammatory mediators from autologous synovial fibroblasts in vitro34_.

Both inflammation and endothelial dysfunction play a role in development of preeclampsia, but the exact underlying mechanism behind the development of preeclampsia is unknown. Since microparticles of leukocytes are elevated in preeclamptic patients35_{, and}

leukocyte-derived microparticles induce inflammation in vitro and ex vivo, we investigated whether microparticles of preeclamptic patients affect the mRNA expression of a set of inflammation genes in human umbilical vein endothelial cells in Chapter 6.

Microparticles and coagulation

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Chapter 1

thereby promoting the formation of the intrinsic Factor X (FX) converting tenase complex (IXa and VIIIa)- and the FII (prothrombin) converting prothrombinase complex (Xa and Va). Platelet-derived microparticles are enriched in binding sites for activated factor V (FVa), FVIIIa and FIXa, and provide a suitable membrane surface to promote thrombin generation13;38_.

Apart from negatively charged phospholipids and thus binding sites for (activated) coagulation factors, microparticles can also expose tissue factor (TF). TF is a 45 kDa transmembrane protein, the main initiator of coagulation in vivo and is produced by various types of extravascular cells, including smooth muscle cells. Upon vascular damage, blood will contact the extravascular TF, resulting in (extrinsic) coagulation activation via activation of FX by the extrinsic tenase complex (TF, FVIIa). In plasma of healthy subjects, only very low amounts of TF (antigen) are detectable, and this TF can be present as a truly soluble (non-membrane bound) protein or associated with microparticles. Under pathological conditions, both endothelial cells, monocytes and possibly other leukocytes produce TF in response to endotoxin and other pro-inflammatory mediators, and this TF can be released from the cell surface on microparticles in vitro39-41_.

Microparticles and in vivo coagulation

Previously, we demonstrated that pericardial (wound) blood from patients undergoing open heart surgery contains high numbers of microparticles exposing TF when compared to systemic blood samples from the same patients42_{. These pericardial microparticles triggered}

TF-dependent thrombin generation in vitro42_{, and were prothrombogenic in a rat venous}

stasis model in vivo43_{. In addition, we showed that high numbers of coagulant TF-exposing}

microparticles were present in blood from a patient suffering from meningococcal septic shock and disseminated intravascular coagulation, whereas a patient with meningococcal septic shock without disseminated intravascular coagulation lacked such microparticles44_,

suggesting that the presence of these TF-exposing microparticles is associated with (development of) disseminated intravascular coagulation in some patients. Furthermore, there is growing evidence that extrinsic coagulation activation and the development of venous thromboembolism in cancer patients is associated with the presence of TF-exposing microparticles that at least in part originate from the tumor45;46_{. Thus, TF-exposing}

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1

microparticles can be present in human blood and these microparticles are capable of

triggering TF-dependent coagulation in vitro and in vivo43;46;47_.

The coagulant activity of TF is enhanced when negatively charged phospholipids such as phosphatidylserine (PS) or phosphatidylethanolamine (PE) are present48-52. Since nothing is known about the phospholipid composition of TF-exposing microparticles, and whether their phospholipid composition differs from non-TF-exposing microparticles, we investigated the phospholipid composition of TF-exposing microparticles from resting and activated endothelial cells, and their ability to trigger thrombus formation in vivo in Chapter 7.

Indirect mechanisms of microparticle-induced coagulation

As mentioned above, microparticles can directly initiate and facilitate the coagulation process by exposure of PS and/or TF (Figure 2A). In addition, microparticles may also affect coagulation more indirectly. Microparticles from activated platelets expose P-selectin which can bind to P-selectin glycoprotein ligand-1 (PSGL-1) on monocytes. This interaction triggers de novo synthesis of TF by monocytes (Figure 2B), and this TF is released from the cell on TF-exposing microparticles (Figure 2B)53;54_{. Furie and coworkers}

showed in a laser injury model that platelets adhere to the damaged vessel wall and then become activated. These activated platelets expose P-selectin, which in turn can capture circulating leukocyte-derived microparticles exposing PSGL-1 (from monocytes, but possibly also granulocytes) as well as TF. In this way, the circulating TF is thought to be delivered at the damaged vessel wall, where in turn this TF becomes coagulant and coagulation can be initiated (Figure 2C)55_.

Microparticles, TF and rafts

Previously, del Conde et al. showed that TF-exposing microparticles released from monocytic cells fuse with platelets, thereby directly delivering TF to the platelet surface. In this manner, TF as the initiator of coagulation and the coagulation factors to be activated are brought together (Figure 2D)56_{. He hypothesized that TF-exposing microparticles}

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raft-Chapter 1

however, is still debated by several investigators57;58_{. Therefore, in Chapter 8, we}

investigated whether TF is a raft associated protein in purified plasma membranes and whether TF-exposing microparticles contain rafts.

Figure 2 gives an overview of the currently known effects of microparticles on the activation of the coagulation cascade. Arbitrarily, we distinguish direct (A) and indirect (B-D) mechanisms. First, figure A shows that microparticles from platelets (PMP) directly stimulate coagulation by exposing phosphatidylserine (PS; ), which binds (activated) coagulation factors. In addition, microparticles from especially leukocytes and endothelial cells (LMP and EMP, respectively) can expose tissue factor (TF; ), the initiator of the coagulation cascade in vivo. Second, figure B shows that coagulation can be initiated indirectly by PMP from activated platelets by binding to monocytes. These microparticles expose not only PS, but also P-selectin ( ) and are thus capable of binding to monocytes via P-selectin glycoprotein ligand-1 (PSGL-1; ). This interaction upregulates the expression of TF in the monocyte (B). Subsequently, the TF-exposing monocytes release highly procoagulant TF-exposing microparticles (MMP; B). Third, another indirect mechanism of microparticle-induced coagulation has been proposed for thrombus formation in an endothelial injury model. Figure C shows that, upon endothelial injury, platelets adhere to the damaged vessel wall and become activated. Activated platelets expose PS and P-selectin. The latter can bind PSGL-1, which is exposed on circulating microparticles from monocytes. Since these microparticles also expose TF, TF becomes localized at the site of the thrombus to be formed, and coagulation can be initiated and propagated. Fourth, figure D shows that TF-exposing microparticles may fuse with the outer membranes from activated platelets, thereby delivering coagulant TF on the surface of the activated platelet. This delivery results in the ultimate co-localization of TF and coagulation factor complexes on the activated platelet surface, making platelets the mediator in the activation and propagation of the coagulation cascade.

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1

Figure 2. Mechanisms of microparticle-induced coagulation.

The coagulation cascade Microparticles and coagulation

LMP/EMP

A

B

C

D

monocyte platelet endothelial cell act. platelet act. platelet PMP MMP PMP X Xa Va prothrombin thrombin

TF

_VIIa PS PS XIIa XIa IXa VIIIa

PS

MMP

The coagulation cascade Microparticles and coagulation

LMP/EMP

A

B

C

D

monocyte platelet endothelial cell act. platelet act. platelet PMP MMP PMP X Xa Va prothrombin thrombin

TF

_VIIa PS PS XIIa XIa IXa VIIIa

PS

MMP

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Chapter 1 TF and PDI

TF can be present in two different conformational forms, a coagulant active form and a cell signalling active form, the latter lacking any coagulant activity. Protein disulfide isomerase (PDI) was shown to oxidize and reduce the disulfide bond between Cys186 and Cys209 in the extracellular domain of TF, thereby switching the function of TF from cell signalling to coagulation and vice versa59_{. Since then, the role of PDI in the regulation of the TF}

coagulant activity has been strongly debated, since it is still unclear whether PDI can reach the cysteins deep inside the TF-VIIa complex60_.

PDI is also present in microparticles. Microparticles from platelets contain PDI61_.

Whether or not PDI in microparticles can influence the coagulant activity of TF present on microparticles or even cells, however, is unknown. We investigated in Chapter 8 to which extent PDI and TF are present in rafts from microparticles.

Alternatively spliced TF

In 2003, an alternatively spliced (as) form of TF was described, which lacks the transmembrane domain due to a deletion which results in a frameshift and thus a peptide sequence different from TF. This asTF is produced by several cell types and organs, and is present in blood of healthy human individuals62_{. Initially, asTF was claimed to possess}

coagulant activity, which could not be confirmed in a more recent studie63_{. In Chapter 9,}

we investigated whether asTF is associated with microparticles and whether such asTF has any coagulant activity.

Structure of the thesis

The main question in this thesis is to establish whether and how microparticles balance cellular and environmental homeostasis. In Chapter 2, the question whether the release of microparticles from human umbilical endothelial cells contributes to cellular survival is investigated. In Chapter 3, caspase-3 deficient cells were transfected with caspase-3 constructs to study the effects of these constructs on the release of microparticles and on the sorting of caspase-3 into these microparticles. To further investigate the presence of caspase-3 in microparticles, in Chapter 4 microparticles present in aging platelet concentrates were studied.

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In Chapter 5, 6 and 7, various functions of microparticles were investigated. In

Chapter 5 the extent of “soluble” VEGFR-1 (sFlt-1) association with microparticles in plasma from pregnant women suffering from preeclampsia was studied. In Chapter 6, the expression levels of inflammation-related genes were investigated in human endothelial cells in response to microparticles from preeclamptic patients, and in Chapter 7 the phospholipid composition and in vivo coagulant properties of TF-exposing microparticles from endothelial cells were studied.

In Chapter 8, the presence of the coagulant and non-coagulant forms of TF in purified plasma membranes and microparticles was studied in detail, and their association with rafts and PDI. Finally, in Chapter 9 the question was addressed whether the alternatively spliced form of TF is associated with microparticles and whether this alternatively spliced form has any coagulant activity.

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39. Abid Hussein MN, Meesters EW, Osmanovic N et al. Antigenic characterization of endothelial cell-derived microparticles and their detection ex vivo. J.Thromb.Haemost. 2003;1:2434-2443. 40. Kagawa H, Komiyama Y, Nakamura S et al. Expression of functional tissue factor on small

vesicles of lipopolysaccharide-stimulated human vascular endothelial cells. Thromb.Res. 1998;91:297-304.

41. Satta N, Toti F, Feugeas O et al. Monocyte vesiculation is a possible mechanism for dissemination of membrane-associated procoagulant activities and adhesion molecules after stimulation by lipopolysaccharide. J.Immunol. 1994;153:3245-3255.

42. Sturk-Maquelin KN, Nieuwland R, Romijn FP et al. Pro- and non-coagulant forms of non-cell-bound tissue factor in vivo. J.Thromb.Haemost. 2003;1:1920-1926.

43. Biro E, Sturk-Maquelin KN, Vogel GM et al. Human cell-derived microparticles promote thrombus formation in vivo in a tissue factor-dependent manner. J.Thromb.Haemost. 2003;1:2561-2568.

44. Nieuwland R, Berckmans RJ, McGregor S et al. Cellular origin and procoagulant properties of microparticles in meningococcal sepsis. Blood 2000;95:930-935.

45. Tesselaar ME, Romijn FP, van dL, I et al. Microparticle-associated tissue factor activity: a link between cancer and thrombosis? J.Thromb.Haemost. 2007;5:520-527.

46. Zwicker JI, Liebman HA, Neuberg D et al. Tumor-derived tissue factor-bearing microparticles are associated with venous thromboembolic events in malignancy. Clin.Cancer Res. 2009;15:6830-6840.

47. Dvorak HF, Van DL, Bitzer AM et al. Procoagulant activity associated with plasma membrane vesicles shed by cultured tumor cells. Cancer Res. 1983;43:4434-4442.

48. Bach R, Rifkin DB. Expression of tissue factor procoagulant activity: regulation by cytosolic calcium. Proc.Natl.Acad.Sci.U.S.A 1990;87:6995-6999.

49. Greeno EW, Bach RR, Moldow CF. Apoptosis is associated with increased cell surface tissue factor procoagulant activity. Lab Invest 1996;75:281-289.

50. Le DT, Rapaport SI, Rao LV. Relations between factor VIIa binding and expression of factor VIIa/tissue factor catalytic activity on cell surfaces. J.Biol.Chem. 1992;267:15447-15454. 51. Le DT, Rapaport SI, Rao LV. Studies of the mechanism for enhanced cell surface factor

VIIa/tissue factor activation of factor X on fibroblast monolayers after their exposure to N-ethylmaleimide. Thromb.Haemost. 1994;72:848-855.

52. Wolberg AS, Monroe DM, Roberts HR, Hoffman MR. Tissue factor de-encryption: ionophore treatment induces changes in tissue factor activity by phosphatidylserinedependent and -independent mechanisms. Blood Coagul.Fibrinolysis 1999;10:201-210.

53. Celi A, Pellegrini G, Lorenzet R et al. P-selectin induces the expression of tissue factor on monocytes. Proc.Natl.Acad.Sci.U.S.A 1994;91:8767-8771.

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54. Furie B, Furie BC. P-selectin induction of tissue factor biosynthesis and expression. Haemostasis 1996;26 Suppl 1:60-65.

55. Falati S, Liu Q, Gross P et al. Accumulation of tissue factor into developing thrombi in vivo is dependent upon microparticle P-selectin glycoprotein ligand 1 and platelet P-selectin. J.Exp.Med. 2003;197:1585-1598.

56. Del Conde I, Shrimpton CN, Thiagarajan P, Lopez JA. Tissue-factor-bearing microvesicles arise from lipid rafts and fuse with activated platelets to initiate coagulation. Blood 2005;106:1604-1611.

57. Dietzen DJ, Page KL, Tetzloff TA. Lipid rafts are necessary for tonic inhibition of cellular tissue factor procoagulant activity. Blood 2004;103:3038-3044.

58. Mandal SK, Pendurthi UR, Rao LV. Cellular localization and trafficking of tissue factor. Blood 2006;107:4746-4753.

59. Ahamed J, Versteeg HH, Kerver M et al. Disulfide isomerization switches tissue factor from coagulation to cell signaling. Proc.Natl.Acad.Sci.U.S.A 2006;103:13932-13937.

60. Bach RR, Monroe D. What is wrong with the allosteric disulfide bond hypothesis? Arterioscler.Thromb.Vasc.Biol. 2009;29:1997-1998.

61. Raturi A, Miersch S, Hudson JW, Mutus B. Platelet microparticle-associated protein disulfide isomerase promotes platelet aggregation and inactivates insulin. Biochim.Biophys.Acta 2008;1778:2790-2796.

62. Bogdanov VY, Balasubramanian V, Hathcock J et al. Alternatively spliced human tissue factor: a circulating, soluble, thrombogenic protein. Nat.Med. 2003;9:458-462.

63. Censarek P, Bobbe A, Grandoch M, Schror K, Weber AA. Alternatively spliced human tissue factor (asHTF) is not pro-coagulant. Thromb.Haemost. 2007;97:11-14.

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Chapter 2 Inhibition of microparticle release triggers

endothelial apoptosis and detachment

Anita N. Böing, Mohammed N. Abid Hussein, Auguste Sturk, Chi M. Hau and Rienk Nieuwland

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Abstract

Introduction. Endothelial cell cultures contain caspase-3-containing microparticles (EMP), which are reported to form during or after cell detachment. We hypothesize that also adherent endothelial cells release EMP, thus protecting these cells from caspase-3 accumulation, detachment and apoptosis.

Methods. Human umbilical vein endothelial cells (HUVEC) were incubated with and without inhibitors of microparticle release (Y-27632, calpeptin), both in the absence or presence of additional “external stress”, i.e. the apoptotic agent staurosporin (200 nmol/L) or the activating cytokine interleukin (IL)-1α (5 ng/mL).

Results. Control cultures contained mainly viable adherent cells and minor fractions of apoptotic detached cells and microparticles in the absence of inhibitors. In the presence of inhibitors, caspase-3 accumulated in adherent cells and detachment tended to increase. During incubation with either staurosporin or IL-1α in the absence of inhibitors of microparticle release, adherent cells remained viable, and detachment and EMP release increased slightly. In the presence of inhibitors, dramatic changes occurred in staurosporin-treated cultures. Caspase-3 accumulated in adherent cells and >90% of the cells detached within 48 hours. In IL-1α-treated cultures no accumulation of caspase-3 was observed in adherent cells, although detachment increased. Scanning EM studies confirmed the presence of EMP on both adherent and detached cells. Prolonged culture of detached cells indicated a rapid EMP formation as well as some EMP formation at longer culture periods. Conclusions. Inhibition of EMP release causes accumulation of caspase-3 and promotes cell detachment, although the extent depends on the kind of “external stress”. Thus, the release of caspase-3-containing microparticles may contribute to endothelial cell survival.

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Inhibition of microparticle release

2 Introduction

Like other eukaryotic cells, endothelial cells release microparticles (MP; EMP: endothelial microparticles) in vitro1-3 _{and in vivo}4-7_{. To which extent EMP originate from adherent or}

from detached endothelial cells, however, is a still unanswered question. Previously, we reported a correlation between the numbers of detached cells and EMP in vitro8_{. Other}

investigators provided indications that EMP are released from adherent endothelial cells during detachment, and that endothelial cells "rapidly lost adhesion" immediately after release of EMP9;10_{. Thus, EMP are presumed to originate from detaching and detached}

endothelial cells. However, Hamilton et al. showed that endothelial cells escape from complement-induced lysis by releasing C5b-9-enriched EMP11_{, suggesting that EMP}

release may contribute to survival by eliminating externally imposed stress.

Recently, we demonstrated that EMP from endothelial cell cultures contain substantial quantities of active (17 kDa) caspase-38_{. These data prompted us to hypothesize}

that adherent endothelial cells may also release caspase-3-containing EMP, and thus escape from internally imposed stress, detachment and apoptosis. If true, then inhibition of EMP release is expected to result in intracellular accumulation of caspase-3 in adherent cells, with increased cell detachment and apoptosis. To test this hypothesis, we treated endothelial cells with a sub-lethal concentration of the apoptotic agent staurosporin or the activator interleukin-1α (IL-1α), without or with widely used inhibitors of microparticle release, i.e. Y-27632 and calpeptin9;12;13_.

Materials and Methods

Reagents and assays

Medium M199, penicillin, streptomycin and L-glutamin were from GibcoBRL (Life Technologies; Paisley, UK). Human serum and fetal calf serum (both heat inactivated during 30 minutes at 56 ºC; HuSi and FCSi, respectively) were from BioWhittaker (Walkersville, MD). Human serum albumin (HSA) was obtained from Sanquin (Amsterdam, The Netherlands). Recombinant human IL-1α was from Sigma (St. Louis, MO). Human recombinant basic fibroblast growth factor and epidermal growth factor were from Invitrogen life technologies (Carlsbad, CA). Collagenase (type 1A) and staurosporin

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Netherlands), trypsin from Difco Laboratories (Detroit, MI), calpeptin from Calbiochem (La Jolla, CA), and Y-27632 from Tocris (Ellisville, MO). Y-27632 is a specific inhibitor of Rho-associated serine/threonine kinases 1 and 2 (i.e. ROCK1 (p160ROCK, ROKβ) and ROCK2 (Rho-kinase, ROKα)), enzymes which are directly involved in the release of apoptotic blebs9;12_{. Calpeptin inhibits calpain, a Ca}2+_{-dependent protease, that plays a role}

in (E)MP formation13_{. For Western blot analysis, anti-human caspase-3 monoclonal}

antibody from Alexis Biochemicals (San Diego, CA) and polyclonal goat-anti-mouse HRP conjugate (DAKO; Glostrup, Denmark) were used. Tissue culture flasks were from Greiner Labortechnik (Frickenhausen, Germany) and gelatin was from Difco Laboratories.

Isolation, culture and treatment of human umbilical vein endothelial cells (HUVEC) HUVEC were collected as described previously3. Upon confluency at passage 3 in 25 cm2 culture flasks, HUVEC were kept for 2 days in a resting state. Culture supernatant was refreshed and cultures were treated without or with staurosporin (200 nM, a sub-lethal concentration in the culture conditions used), or IL-1α (5 ng/mL, a concentration providing extensive endothelial cell activation such as cell surface exposure of E-selectin). Where indicated, cultures were co-incubated with Y-27632 (30 µM; two hours preincubation) and/or calpeptin (200 µM; one hour preincubation, or, when used in combination with Y-27632, added after one hour of incubation with Y-27632). Stock solutions of staurosporin, IL-1α, Y-27632 and calpeptin were prepared in ethanol, medium M199, PBS and DMSO, respectively. Control cultures were incubated with DMSO and ethanol.

In three experiments, we studied whether detached cells release EMP. Detached cells were harvested from 10 mL culture medium from HUVEC cultures treated without or with staurosporin or IL-1α (24 hours). Detached cells were resuspended in 10 mL fresh culture medium (without staurosporin or IL-1α), and numbers of detached endothelial cells and EMP were determined at fixed time intervals (3-48 hours) by flow cytometry.

Flow cytometric analysis of HUVEC

Conditioned media were collected and centrifuged (10 minutes, 180g and 20 °C) to isolate detached cells. Pellets were resuspended in PBS containing 1% (v/v) FCSi (pH 7.4). Adherent cells were detached by trypsinization. After 4 minutes, trypsin was neutralized by

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2

PBS/FCSi (10% v/v). Both cell suspensions were centrifuged (10 minutes, 180g and 4 °C)

and pellets were resuspended in PBS/FCSi (1% v/v), and then again centrifuged (10 minutes, 180g and 4 °C). Detached cells were resuspended in 0.5 mL PBS/FCSi (1% v/v) and adherent cells in 1.0 mL PBS/FCSi (1% v/v). Cells were labeled with annexin V-FITC (IQP; Groningen, The Netherlands) and propidium iodide (PI; a gift from Dr. E. Reits, Department of Cell Biology and Histology, AMC, The Netherlands) as described previously8_{. Intracellular caspase-3 was detected using the Active Caspase-3 mAb}

Apoptosis Kit I from BD Pharmingen (San Diego, CA). Samples were analyzed in a FACSCalibur flowcytometer (Becton Dickinson; San Jose, CA). The cell number was estimated per culture flask using flow cytometry.

Isolation of EMP

Aliquots (1 mL) of the cell-free culture supernatants were snapfrozen in liquid nitrogen and stored at – 80 °C. Before use, samples were thawed on melting ice for 1.5 hour, and then centrifuged (1 hour, 18,890g and 20 °C). Then, 975 µL of supernatant was removed and the pellet was resuspended in 225 µL PBS (154 mmol/L NaCl, 1.4 mmol/L phosphate) containing 10.9 mmol/L trisodium citrate, or in perm/wash (0.1% v/v) for intravesicular caspase-3 staining. MP were resuspended and again centrifuged (30 minutes, 18,890g and 20 °C), 225 µL supernatant was removed and MP were diluted and resuspended by adding 75 µL PBS/citrate or perm/wash (0.1% v/v).

Flow cytometric analysis of EMP

EMP were analyzed in a FACSCalibur flow cytometer as described previously3_{. To detect}

intravesicular caspase-3, MP (5 µL) were diluted with 35 µL 0.1% perm/wash solution containing 2.5 mmol/L CaCl2 plus either anti-caspase-3-FITC (BD) or control antibody,

Ig-FITC (IQP). For annexin V staining, MP (5 µL) were diluted with 35 µL PBS containing 2.5 mmol/L CaCl2 (pH 7.4) and annexin V-APC (Caltag Laboratories; Carlsbad, CA; 5 µL

20-fold prediluted) was added. To remove the excess of unbound annexin V, 200 µL PBS/calcium buffer (or 200 µL 0.1% perm/wash containing CaCl2 for intravesicular

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resuspended with 300 µL PBS/calcium or 300 µL 0.1% perm/wash containing CaCl2.

Previously, we demonstrated that numbers of EMP (N) and detached cells highly correlate8_.

Therefore, the efficacy of inhibitors to inhibit EMP release was expressed either as ratio per detached cell, i.e. NEMP/Ndetached cell, or, where indicated, as percentage from the control of that particular condition, i.e. untreated, or staurosporin or IL-1α without inhibitors: ([(NEMP, control/Ndetached cells, control) - (NEMP, inhibitor/Ndetached cells, inhibitor)] / (NEMP, control/Ndetached cells, control)) x 100%.

Western blotting

Detached and adherent endothelial cells were separately isolated, washed and collected in PBS/FCSi (0.5 and 1.0 mL, respectively). From these suspensions, 300 µL (detached cells) and 800 µL (adherent cells) were used to isolate cells. Subsequently, 2-fold concentrated reducing sample buffer was used to dissolve the pellets of the detached cells (final volume 20 µL) and adherent cells (final volume 40 µL). From the detached cell lysate, 10 µL was applied to SDS-PAGE, and from the adherent cell lysates, volumes were adjusted to 5 x 104

cells per lane. After removal of detached cells, EMP were isolated from the cell-free culture supernatants by centrifugation (1 hour, 18,890g and 20 °C) and resuspended in 10 µL PBS plus 10 µL 2-fold concentrated reducing sample buffer. Per EMP sample, 10 µL was applied to SDS-PAGE. Prior to electrophoresis, all samples were preheated (5 minutes at 100 °C). Electrophoresis was carried out in 8-16% gradient SDS-PAGE gels (BioRad; Hercules, CA). Proteins were transferred to PVDF membrane (BioRad). Blots were incubated for 1 hour at room temperature with blocking buffer (Tris-buffered saline-Tween (TBST); 10 mmol/L Tris-HCl, 150 mmol/L NaCl, 0.05% (v/v) Tween-20; pH 7.4), containing 5% (w/v) dry milk powder (Protifar; Nutricia, Vienna, Austria)). The blots were incubated with monoclonal anti-human caspase-3 (1:1,000; v/v) overnight at 4 °C, followed by incubation with polyclonal goat-anti-mouse HRP conjugate (1:30,000; v/v) for 1 hour at room temperature. Between incubation steps, blots were washed three times with TBST for 5-10 minutes. All antibodies were diluted with 2.5% (w/v) blocking buffer. The bands were visualized on Fuji Medical X-ray film by using Lumi-Light Plus Western Blotting Substrate (Roche; Mannheim, Germany).

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2

Previously we showed that detached cell lysates from control and IL-1α-treated

cultures contain 17 kDa caspase-38_{. The absence of detectable amounts of caspase-3 in}

detached cell lysates by Western blot in our present experiments should be interpreted as "below detection level" rather than being completely absent, since lesser numbers of detached cells were available due to the necessity of downscaling of the culture conditions compared to our previous studies as a consequence of the number of experimental conditions to be tested simultaneously.

Scanning Electron Microscopy (SEM)

HUVEC (third passage) were cultured on gelatin-coated coverslips. At 90% confluence, cells were incubated overnight without or with staurosporin (200 nM) or IL-1α (5 ng/mL). Specimens were prepared essentially as described by van Berkel et al.14. Briefly, cells were fixed in McDowell’s fixative for 45 minutes, washed (phosphate buffer), dehydrated and dried with hexamethyldisilazane. Detached cells were captured on poly-L-lysin-coated coverslips (30 minutes) and then treated as described above. Dried coverslips were mounted on stubs and coated with 10 nm gold, and imaged with a Philips SEM 525.

Statistical analysis

All data were analyzed with GraphPad Prism for Windows, release 3.02 (San Diego, CA). Data from preliminary experiments regarding differences in numbers of adherent cells, detached cells or EMP between control, staurosporin and IL-1α conditions were analyzed by Wilcoxon matched pairs test (one tailed analysis). Values are expressed as median (range). Data regarding annexin V/PI labeling, i.e. the extent of apoptosis, of adherent cells and detached cells were analyzed by paired t-test (one tailed analysis). Differences in the percentages of adherent cells or detached cells upon incubation with inhibitors in the absence or presence of staurosporin or IL-1α were analyzed with one-way analysis of variance (ANOVA). The method of Dunnett or Bonferroni was used to correct for multiple comparisons. Correlations were determined using Pearson’s correlation test (two tailed analysis). For the time dependent experiments (0-48 hours), the areas under curve per condition (control, staurosporin or IL-1α) were calculated in the absence or presence of

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Chapter 2

inhibitors of EMP release, and these data were compared using paired t-test (one-tailed). Differences were considered statistically significant at P<0.05.

Results

Basal conditions: endothelial cell cultures in the absence of inhibitors of microparticle release

Figure 1 shows that in response to external stress, i.e. incubation with either staurosporin or IL-1α, numbers of detached cells (Figure 1C) and annexin V-binding EMP (Figure 1E) increased (n=6). Neither staurosporin nor IL-1α affected the exposure of aminophospholipids (binding of annexin V: early apoptosis, open bars) or nuclear fragmentation (PI staining and annexin V binding: late apoptosis, dashed bars) of adherent (Figure 1B) or detached (Figure 1D) cells. Whereas minor fractions of adherent cells stained for annexin V or PI (Figure 1B), approximately 90% of detached cells were undergoing early or late apoptosis at this 24 hour culture period (Figure 1D). In sum, in the three conditions studied, endothelial cell cultures contain mainly viable adherent cells, low numbers of apoptotic detached cells and some EMP. Detachment and EMP release increased in response to external stress, but the apoptotic status of the adherent and detached cells was unaffected.

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2

Figure 1. Basal conditions: endothelial cell cultures in the absence of inhibitors of

microparticle release.

Endothelial cells were incubated without (control) or with staurosporin (200 nM) or IL-1α (5 ng/mL) for 24 hours. Adherent cells (A, B), detached cells (C, D) and EMP (E) were isolated and analyzed as described in Materials and Methods, and their numbers were estimated by flow cytometry. Bars indicate median (range). Adherent cells (B) and detached cells (D) were analyzed for their apoptotic status by annexin V binding (white bars; ‘early apoptosis’) or by staining for both annexin V and PI (grey bars; ‘late apoptosis’). Bars indicate mean and SD (n=3). In E, both annexin V-positive (grey

P=0.031 0 20 40 60 80 100 % Apoptosi s Staurosporin Control IL-1 B D N u m ber of adherent cel ls  10 6 A P=0.015 0 1 2 3 4 5 6

Control Staurosporin IL-1

E N u m ber of E M P  10 6 P=0.031 P=0.031 0 2 4 6 8 10 12

Control Staurosporin IL-1 C N u m b er of detached c e ll s  10 6 P=0.015

Control Staurosporin IL-1 0 1 2 3 4 5 6 Staurosporin Control IL-1 0 20 40 60 80 100 % A poptosi s P=0.031 0 20 40 60 80 100 % Apoptosi s Staurosporin Control Staurosporin IL-1 Control IL-1 B D N u m ber of adherent cel ls  10 6 A P=0.015 0 1 2 3 4 5 6

Control Staurosporin IL-1 Control Staurosporin IL-1

E N u m ber of E M P  10 6 P=0.031 P=0.031 0 2 4 6 8 10 12

Control Staurosporin IL-1 C N u m b er of detached c e ll s  10 6 P=0.015

Control Staurosporin IL-1 Control Staurosporin IL-1 0 1 2 3 4 5 6 Staurosporin Control Staurosporin IL-1 Control IL-1 0 20 40 60 80 100 % A poptosi s

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Chapter 2

Effects of inhibitors of microparticle release on endothelial cell detachment and EMP release

The effects of two widely used inhibitors of microparticle release (Y-27632, calpeptin) on cell detachment and EMP release were tested in endothelial cell cultures in the absence (control, untreated) or presence of external stress (staurosporin, IL-1α) for 24 hours (Table 1). In control cultures, the numbers of EMP were unaffected by either Y-27632, calpeptin or their combination. In contrast, detached endothelial cell fractions increased from 5.7% ± 2.2 to 26.9% ± 2.1. Also in staurosporin- or IL-1α-treated cultures, only minor effects of Y-27632 and/or calpeptin were observed on EMP release, but detachment increased dramatically from 15.1% ± 5.1 to 81.4% ± 7.2 with staurosporin and 56.8% ± 10.3 with IL-1α. When we assume that most EMP originate from detaching or detached endothelial cells (Figure 3A), then it can be calculated from the data shown in Table 1 that in the presence of the combination of inhibitors, the ratio of EMP/detached cell decreases from 8.3 ± 0.7 to 2.9 ± 1.1 in control cultures (P<0.05), from 15.1 ± 5.1 to 1.9 ± 1.0 in staurosporin-treated cultures (P<0.001), and from 7.3 ± 0.5 to 2.0 ± 1.0 in IL-1α -treated cultures (P<0.001). Evidently, the inhibitors do inhibit EMP formation but not below a certain basal level.

To gain a more detailed insight into the complex relationship between EMP release, induction of apoptosis and cell detachment, we determined the time dependence of the effects of inhibitors of microparticle release both in the absence (Figure 2) and presence of additional external stress (staurosporin or IL-1α, Figures 3 and 4, respectively), as well as the presence of caspase-3.

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Table 1. Effects of inhibitors of microparticle release on detachment and EMP release at 24 hours.

Control P Staurosporin P IL-1α P Number of EMP x 106

No inhibitors 1.1 ± 0.2 - 4.5 ± 2.7 - 3.1 ± 1.7 -

Y-27632 1.4 ± 0.4 P0.05 2.8 ± 0.7 P0.05 2.5 ± 1.2 P0.05

Calpeptin 1.4 ± 0.6 P0.05 3.6 ± 0.4 P0.05 2.4 ± 1.8 P0.05

Y-27632 + Calpeptin 1.4 ± 0.5 P0.05 5.1 ± 0.8 P0.05 2.6 ± 1.5 P0.05

Detached cells (% from total)

No inhibitors 5.7 ± 2.2 15.4 ± 10.1 19.5 ± 13.6

Y-27632 15.7 ± 1.3 P<0.05 29.1 ± 4.6 P0.05 37.7 ± 8.2 P0.05

Calpeptin 17.7 ± 8.2 P<0.05 71.2 ± 8.0 P<0.001 43.6 ± 16.6 P0.05

Y-27632 + Calpeptin 26.9 ± 2.1 P<0.001 81.4 ± 7.2 P<0.001 56.8 ± 10.3 P<0.05

Endothelial cells were incubated in the absence (control) or presence of staurosporin (200 nM) or IL-1α (5 ng/mL), with or without Y-27632, calpeptin or both (n=3). After 24 hours, detached cells and EMP were isolated as described in Materials and Methods. Detachment is expressed as % of the total number of cells, i.e. the number of adherent and detached cells present. Differences between conditions with and without inhibitor(s) were analyzed by paired t test as described in the Statistical analysis section of Materials and Methods. Data are presented as mean ± SD.

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Chapter 2

Effects of inhibitors of microparticle release in endothelial cell cultures without additional external stress

In control cultures in the absence of inhibitors of EMP release (open symbols throughout Figures 2-4), adherent cell fractions binding annexin V (2B) or containing caspase-3 (2C) remained constant in time, and caspase-3 was not detectable on Western blot (2D, left). Detachment increased slightly (2A) and from 12 hours onwards >80% of detached cells bound annexin V (2E), but due to the low numbers of detached cells at 3 and 6 hours these fractions varied considerably. Detached cell fractions staining for caspase-3 ranged between 10-30% (2F), and caspase-3 was not detectable on blot (2G, left). Numbers of caspase-3-containing EMP increased in time (2H) and virtually all EMP contained caspase-3 (2I). The occurrence of caspase-3 in EMP was confirmed at 24 and 48 hours by Western blot (2J, left). The annexin V findings indicate that some 50% of detached cells are not yet apoptotic in the first few hours after detachment.

In the presence of inhibitors (closed symbols throughout Figures 2-4), adherent cell fractions binding annexin V (2B) tended to increase (P=0.186). Fractions staining for caspase-3, however, increased slightly (2C; P=0.03) and a faint (17 kDa) caspase-3 band became visible at 48 hours (2D, right panel). Detachment tended to increase (2A; P=0.07). Similar to adherent cells, detached cell fractions binding annexin V were unaffected (2E; P=0.377), but those containing caspase-3 increased (2F; P=0.003). The latter could not be confirmed on blot (2G, right), which probably is due to the low numbers of detached cells. The total EMP numbers increased similar to the cultures without inhibitor treatment (2H; P=0.497), and fractions of caspase-3-containing EMP (2I; P=0.096) were unaffected by the inhibition treatment. Caspase-3 in EMP was visible at both 24 and 48 hours (2J, right).

In sum, adherent endothelial cells in control cultures, i.e. without external stress, showed a modest accumulation of caspase-3 in the presence of inhibitors of microparticle release. Cell detachment tended to increase, evidence was obtained for the presence of caspase-3 in detached cells, and detached cells were not immediately apoptotic upon their detachment. Finally, the numbers of EMP were comparable in the absence and presence of these inhibitors.

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Figure 2. Effects of inhibitors of microparticle release in endothelial cell cultures.

Endothelial cell cultures (n=3) were incubated without external stress up to 48 hours, in the absence (open symbols) or presence (closed symbols) of Y-27632 (30 µM) plus calpeptin (200 µM). Figure A shows the fractions of detached cells. Figures B and E show adherent (B) and detached (E) endothelial cell fractions binding annexin V as an indicator of apoptosis. Figures C and F show adherent (C) and detached (F) endothelial cell fractions staining for intracellular caspase-3, whereas figures D and G show the ‘total amounts’ of 17 kDa caspase-3 detectable in lysates of adherent- (D) and detached (G) endothelial cells in the absence (-) or presence (+) of Y-27632 plus calpeptin. Figures H-J show the absolute numbers of containing EMP (H), the fractions of

caspase-3-Caspase-3

Caspase-3

Time (hours) Time (hours)

Time (hours) ( 10 6) 6 12 24 48 3 6 12 24 48 3 6 12 24 48 3 6 12 24 48 3 6 12 24 48 3 6 12 24 48 3 A B C E F H I J D - + - + - + G Caspase-3 Caspase-3 Caspase-3

Time (hours) Time (hours)

Time (hours) ( 10 6) 6 12 24 48 3 6 12 24 48 36 12 24 48 3 6 12 24 4836 12 24 48 3 6 12 24 48 3 6 12 24 48 3 6 12 24 48 3 6 12 24 48 3 6 12 24 48 3 6 12 24 48 3 6 12 24 48 3 6 12 24 48 3 6 12 24 48 3 A B C E F H I J D - + - + - + - + - + - + G

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Effects of inhibitors of microparticle release in endothelial cell cultures in the presence of additional external stress: staurosporin

In the absence of inhibitors of microparticle release, detached cell fractions increased compared to the cultures without external stress (3A versus 2A; P=0.041; open symbols). Compared to cultures without external stress (Figures 2B and 2C), adherent cell fractions in staurosporin-treated cultures staining for annexin V or caspase-3 (Figures 3B and 3C, respectively) were not increased (P=0.390 and P=0.199, respectively). Also on blot, caspase-3 was not detectable (3D, left). At prolonged culture periods, detached cell fractions binding annexin V (3E) increased (P=0.016) compared to control cultures (2E), but caspase-3-containing fractions were unaffected (3F versus 2F; P=0.461). Depending on the experiment, in some lysates a weak caspase-3 band was visible at 12 hours (3G, left). The numbers of caspase-3-containing EMP increased compared to untreated cultures (3H versus 2H; P=0.017), and virtually all EMP contained caspase-3 (3I). On blot, already from 12 hours onwards, caspase-3 was clearly visible in EMP lysates (3J, left), which evidently is earlier than in control cultures (2J, left).

In the presence of inhibitors, more than 90% of endothelial cells detached within 48 hours (3A; P=0.01, compared to staurosporin alone; closed symbols). After 48 hours, >80% of the few remaining adherent cells stained for annexin V (3B; P=0.02 versus staurosporin alone), whereas 20% contained caspase-3 (3C; P=0.04). Accumulation of caspase-3 in adherent cell fractions was confirmed on blot (3D, right). The absence of caspase-3 on Western blot at 48 hours is most likely explained by the insufficient numbers of adherent cells due to the extensive cell detachment. Detached cell fractions staining for annexin V were unaffected (3E; P=0.241), but the fractions of caspase-3-containing detached cells strongly increased (3F; P<0.001). The latter was confirmed by Western blot (3G, right). The absolute numbers of caspase-3-containing EMP increased slightly (3H; P=0.02), and again virtually all EMP contained caspase-3 (3I). Again, the presence of caspase-3 in EMP could be confirmed on Western blot (3J, right). Thus, exposure of endothelial cell cultures to mild external stress, i.e. a low concentration of the apoptosis-inducer staurosporin, triggered accumulation of caspase-3 in adherent cells and massive detachment in the presence of inhibitors of microparticle release. Also, the inhibitors of EMP release caused caspase-3 accumulation in detached cells.

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2

Figure 3. Effects of inhibitors of microparticle release in endothelial cell cultures in the presence of external stress: staurosporin.

Endothelial cell cultures (n=3) were incubated with additional external stress (staurosporin) up to 48 hours, in the absence (open symbols) or presence (closed symbols) of Y-27632 (30 µM) plus calpeptin (200 µM). Figure A shows the fractions of detached cells. Figures B and E show adherent (B) and detached (E) endothelial cell fractions binding annexin V. Figures C and F show adherent (C) and detached (F) endothelial cell fractions staining for intracellular caspase-3, whereas figures D and G show the ‘total amounts’ of 17 kDa caspase-3 detectable in lysates of adherent- (D) and detached (G) endothelial cells in the absence (-) or presence (+) of Y-27632 plus calpeptin. Figures H-J show the

A B C D E F G 3 - + - + Caspase- 3 Caspase- 3 H I J ( 10 6)

Time (hours ) Time (hours)

Time (hours) + -Caspase- 3 3 6 12 24 48 3 6 12 24 48 3 6 12 24 48 3 6 12 24 48 3 6 12 24 48 3 6 12 24 48 A B C D E F G 3 - + - + - + - + Caspase- 3 Caspase- 3 H I J ( 10 6)

Time (hours ) Time (hours)

Time (hours) + -Caspase- 3 3 6 12 24 48 3 6 12 24 48 3 6 12 24 48 3 6 12 24 48 3 6 12 24 48 3 6 12 24 48

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Chapter 2

Effects of inhibitors of microparticle release in endothelial cell cultures in the presence of additional external stress: IL-1α

The IL-1α-induced increase in detachment was comparable to control cultures in the absence of inhibitors of microparticle release (Figures 4A versus 2A; P=0.146; open symbols). Compared to the control cultures, i.e. the endothelial cell cultures without external stress, fractions of annexin V-binding adherent cells were lower (4B versus 2B; P=0.03) and those of caspase-3-containing adherent cells were comparable (4C versus 2C; P=0.145). On blot no caspase-3 was detectable (4D, left). Detached cell fractions staining for annexin V or caspase-3 were both comparable to untreated cultures (4E versus 2E and 4F versus 2F, respectively; P=0.359 and P=0.448). No caspase-3 was detectable in detached cell lysates (4G, left). EMP release was comparable to untreated cultures (4H versus 2H; P=0.407), and again most if not all EMP contained caspase-3 (4I). Faint caspase-3 bands were visible after 24 and 48 hours on Western blot (Figure 4J, left).

In the presence of inhibitors of microparticle release, detachment increased (4A versus 2A; P=0.02). Fractions of adherent cells staining for annexin V increased (4B; P=0.04), but those staining for caspase-3 remained low and were unchanged compared to IL-1α alone (4C; P=0.115). On Western blot, a faint caspase-3 band became visible in some lysates (4D, right panel). Detached cell fractions staining for annexin V were unaffected (4E; P=0.157). Although caspase-3-containing detached cell fractions increased (4F; P=0.02), no or hardly any caspase-3 was detectable on blot (4G, right). The numbers of caspase-3-containing EMP increased insignificantly (4H; P=0.139). Again, most EMP contained caspase-3 (4I). The presence of caspase-3 was confirmed by blot at 24 and 48 hours (4J, right).

Taken together, the overall responses induced by IL-1α, both in the presence and absence of inhibitors, closely paralleled the changes occurring in control cultures in time (Figure 2), with the exception of some increased cell detachment.