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Endothelial cell-derived microparticles

Abid Hussein, M.N.

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Citation for published version (APA):Abid Hussein, M. N. (2008). Endothelial cell-derived microparticles.

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Download date: 06 Dec 2020

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M.N. Abid Hussein


M.N. Abid Hussein

PhD Thesis, University of Amsterdam –with – references – with summary in Dutch

ISBN: 978-90-9022483-1

Cover: Electronmicrograph of adherent endothelial cells (Anita N. Böing)

Printed by: PrintPartners Ipskamp

© M.N. Abid Hussein, Amsterdam, 2008


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 vrijdag 1 februari 2008, te 10.00 uur

door

Mohammed Noori Abid Hussein

geboren te Baqube, Irak

Promotiecommissie

Promotor: Prof.dr. A. Sturk

Co-promotor: Dr. R. Nieuwland

Overige leden: Prof.dr. L. Eijsman

Prof.dr. C.E. Hack

Prof.dr. M.M. Levi

Dr. J.C.M. Meijers

Prof.dr. P.H. Reitsma

Prof.dr. C.J.F. van Noorden

Faculteit der Geneeskunde, Universiteit van Amsterdam

To Rasha, Haneen and Basil

Contents

7

Contents

Chapter 1 Introduction 13

Chapter 2 Antigenic characterization of endothelial cell-derived

microparticles and their detection ex vivo.

J. Thromb. Haemost. 2003;1:2434-2443

27

Chapter 3 Phospholipid compostion of in vitro endothelial microparticles

and their in vivo thrombogenic properties.

Thromb. Res. 2007; In press

53

Chapter 4 Cell-derived microparticles contain caspase 3 in vitro and in

vivo.

J. Thromb. Haemost. 2005;3:888-896

71

Chapter 5 Simvastatin-induced endothelial cell detachment and

microparticle release are prenylation dependent.

Submitted

97

Chapter 6 Inhibition of microparticle release triggers endothelial cell

apoptosis and detachment.

Thromb. Haemost. 2007;98:1096-1107

117

Chapter 7 General discussion and summary 143

Chapter 8 Algemene discussie en samenvatting 153

Coauthors 165

Curriculum Vitae 167

Acknowledgements 169

Abbreviations

9

List of abbreviations

ACD acid citrate dextrose

ANOVA analysis of variance

APC allophycocyanine

bFGF basic fibroblast growth factor

CD cluster of differentiation

CEC circulating endothelial cells

CSF colony stimulating factor

CVD cardiovascular disease

DMSO dimethyl sulfoxide

EC endothelial cells

ECL enhanced chemiluminescence

EDTA ethylenediamine tetraacetic acid

EGF epidermal growth factor

ELISA enzyme-linked immunosorbent assay

EMP endothelial cell-derived microparticles

EPCR endothelial protein C receptor

EtOH ethanol

fc final concentration

FCS fetal calf serum

FCSi fetal calf serum, heat inactivated

FITC fluorescein isothiocyanate

FL fluorescence

FSC forward scatter

GGPP geranylgeranylpyrophosphate

GP glycoprotein

HEPES 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid

HMG-CoA 3-hydroxy-methylglutaryl coenzyme A

hpTLC high performance thin layer chromatography

HRP horseradish peroxidase

Abbreviations

10

HSA human serum albumin

HuSi human serum, heat inactivated

HUVEC human umbilical vein endothelial cells

ICAM-1 intercellular adhesion molecule-1

Ig immunoglobulin

IL interleukin

kDa kilodalton

LDL low density lipoprotein

L-PC L- -lysophosphatidylcholine

L-PE L- -lysophosphatidylethanolamine

LPS lipopolysaccharide

L-PS L- -lysophosphatidylserine

MCP-1 monocyte chemoattractant protein-1

MeOH methanol

MMP monocyte-derived microparticles

MoAb monoclonal antibody

MP microparticles

n number of experiments or subjects

NO nitric oxide

NOS nitric oxide synthase

N.S. not significant

PBS phosphate-buffered saline

PC phosphatidylcholine

PE phycoerythrin; phosphatidylethanolamine

PECAM-1 platelet-endothelial cell adhesion molecule-1

PI propidium iodide

PIn L- -phosphatidylinositol

PIP2 phosphatidyl inositol 4,5-bisphosphate

PMP platelet-derived microparticles

PS phosphatidylserine

PVDF polyvinylidene difluoride

Abbreviations

11

ROCK Rho-associated coiled kinase

s soluble

SD standard deviation

SDS sodium dodecyl sulphate

SDS-PAGE SDS-polyacrylamide gel electrophoresis

SEM scanning electron microscopy

SLE systemic lupus erythematosus

SM sphingomyelin

SSC side scatter

TBST tris-buffered saline tween

TF tissue factor

TGT thrombin generation test

TNF tumor necrosis factor

TTP thrombotic thrombocytopenic purpura

VCAM-1 vascular cell adhesion molecule-1

vWF von Willebrand factor

13

Chapter 1

Introduction

Introduction

14

1.1 The endothelium

The endothelium is a monolayer of endothelial cells lining the vascular system. In

adult humans, this layer contains about 1013 endothelial cells that cover an area between

4000-7000 m2 and weigh nearly 1 kg [1].

Endothelial cells regulate and/or affect several processes, including haemostasis,

angiogenesis, inflammation, growth and differentiation of cells, degradation of the

extracellular matrix, vascular tone and permeability [2-11]. One of the major functions of

the endothelium, however, is its role in haemostasis. Under physiological conditions,

endothelial cells inhibit coagulation by producing heparin-like glycosaminoglycans,

thrombomodulin and the endothelial protein C receptor (EPCR), and prevent platelet

activation by producing prostacyclin. Heparin-like glycosaminoglycans bind

antithrombin, thereby promoting inactivation of thrombin. By binding thrombin,

thrombomodulin facilitates activation of protein C into activated protein C, and this

activation is further amplified by EPCR [12-15]. Under pathological conditions, e.g.

during inflammation, endothelial cells can express tissue factor (TF), the initiator of

coagulation in vivo, and release von Willebrand Factor (vWF), which is essential for

platelet adhesion to the damaged vessel wall and thrombus formation [16,17].

The endothelium occupies a unique interface between circulating blood and

extravascular tissues. (Patho)physiological alterations in the blood are continuously and

directly sensed by the endothelium. For instance, during bacterial infection and/or local

inflammation, endothelial cells respond to elevated circulating levels of bacterial products

such as the lipopolysaccharides (LPS, also called endotoxin) of Gram-negative bacteria or

inflammatory cytokines such as tumour necrosis factor (TNF- ) by upregulating the

expression and production of leukocyte adhesion molecules, cytokines, growth factors and

tissue factor (TF) [18,19]. These (reversible) responses are favourable to the host and will

diminish when the attack or insult is eliminated. Alternatively, an uncontrolled

(irreversible) response of the endothelium to insults such as viruses or antigen/antibody

complexes may result in loss of physiological functions (‘dysfunction’). The underlying

intracellular mechanisms leading to such ‘dysfunction’ include (chronic) activation and/or

programmed cell death (apoptosis). Endothelial ‘dysfunction’ is manifest in (the

Chapter 1

15

development of) many clinical settings, e.g. cardiovascular diseases [20,21], systemic

lupus erythematosus (SLE) [22], diabetes [23], hypertension [24] and renal failure [25].

1.2 Assessing endothelial function and status in vitro and in vivo

In vitro, endothelial function is assessed by measuring the vasodilatory response to

physical or biochemical stimuli such as acetylcholine in isolated arteries or aorta rings

[26]. A common technique to assess endothelial function in vivo is the measurement of

the coronary artery diameter by quantitative angiography before and after infusion of

increasing doses of acetylcholine [27]. Because this method is invasive, more recently

various non-invasive methods have been developed. For example, by high resolution

ultrasound the flow-mediated vasodilatation (FMD) of the femoral or brachial artery can

be evaluated [28-30].

During the last decade, a wide array of biochemical molecules has been exploited to

assess the status of the endothelium. In many in vitro studies, the levels of soluble

endothelial markers have been measured in peripheral blood [31,32]. Well known

examples are soluble (s) P-selectin (platelet-selectin), sICAM-1 (intercellular adhesion

molecule-1), sVCAM-1 (vascular cell adhesion molecule-1), sE-selectin (endothelial-

selectin), sThrombomodulin and sPECAM-1 (platelet-endothelial cell adhesion molecule-

1; summarized by Horstman et al. [33]). Except sE-selectin, the other markers may also

originate from other cell types. Therefore, using these markers to assess the status of the

endothelium requires careful interpretation. At present, both sE-selectin and vWF are

considered to be specific markers reflecting the activation status of the endothelium in

vivo. With regard to assessing the apoptotic status of the endothelium, however, no

markers are available.

1.3 Endothelial cell-derived microparticles (EMP)

Microparticles (MP) are vesicles released from the plasma membrane of many cell

types during activation and/or apoptosis. Endothelial cells release EMP, e.g. in response to

either TNF- or LPS [34-37]. In vivo, EMP have been reported in peripheral blood from

patients suffering from various diseases, such as acute coronary syndrome [38], acute

Introduction

16

ischemic stroke [39], atrial fibrillation [40], metabolic syndrome [41], multiple sclerosis

[42], paroxysmal nocturnal haemoglobinuria [43], severe malaria complicated with coma

[44], systemic inflammatory response syndrome [45], systemic lupus erythematosus [46],

type 2 diabetes [47], vasculitis [48], and venous thromboembolism [49]. To which extent

circulating EMP reflect the status of the endothelium in vivo is unknown, and this may be

further complicated by their cumbersome identification.

Despite the many reports on EMP in the various clinical conditions, consensus on

their straightforward identification is lacking because they (i) constitute only a minor

fraction of the total population of MP in vivo, (ii) share many antigens with the more

abundant platelet-derived MP (PMP), and (iii) may originate from any of the many

endothelial cell subpopulations lining the various types of vessels. Table 1 (page 18)

shows that a wide array of CD-markers has been used in order to identify EMP in human

plasma samples.

1.4 Putative source of EMP

The endothelium consists of a confluent (adherent) monolayer of endothelial cells

that separates the blood from the extravascular tissues. Already in 1978, Hladovec

reported the presence of small numbers of (detached) circulating endothelial cells in

patients with acute myocardial infarction and angina pectoris [50]. Since then, circulating

endothelial cells (CEC) have been reported in at least 20 different diseases or disease

states, including sickle cell anaemia, acute coronary syndromes, systemic lupus

erythematosus and hypertension (summarized by Blann et al. [51]). In vitro, endothelial

cell cultures invariably also contain small numbers of detached cells. These cells display

apoptotic features, such as exposure of negatively charged phospholipids, caspase 3

activity and DNA fragmentation [52-54]. Whether or not EMP (at least partially) originate

from detached cells, however, is unknown.

1.5 Putative functions of EMP

Most studies suggest that cell-derived MP play a role in coagulation. By exposing

negatively charged phospholipids, MP facilitate binding of coagulation factors and

Chapter 1

17

promote the formation of coagulation factor complexes. MP may also expose TF, the

initiator of coagulation in vivo. For example, MP from LPS-treated monocytes expose TF

and initiate coagulation in vitro [55]. Similarly, EMP from LPS- or TNF- -treated

endothelial cells expose TF. Although such TF-exposing EMP are also coagulant in vitro,

it is equally unknown whether such EMP are indeed coagulant in vivo.

Regardless of the many functions of MP, it is unknown why all types of eukaryotic

cells release vesicles, including MP, into their environment. It is tempting to speculate that

release of vesicles may have beneficial effects for the parent cells. In an interesting study

published in 1990, Hamilton et al. showed that complement C5b-9-treated endothelial

cells were protected from lysis by shedding C5b-9-enriched EMP [56]. These findings

indicated that the release of vesicles may contribute to cellular survival by eliminating

external stress.

1.6 Structure of the thesis

The initial aim of this thesis was to establish one or more reliable markers to identify

EMP, which preferentially may reflect the status of the (parent) endothelial cells. In

Chapter 2, the antigenic phenotype of EMP from activated endothelial cells was

characterized and in Chapter 3 we assessed the phospholipid composition of such EMP,

and investigated their ability to trigger coagulation in vivo. In Chapter 4, the question

was addressed as to which extent EMP originate from adherent or detached endothelial

cells, and in Chapter 5 we studied the effects of the cholesterol-lowering drug simvastatin

on the relationship between EMP, adherent and detached endothelial cells. Finally, in

Chapter 6 we further explored the putative role of EMP formation in endothelial cell

survival.

Introduction

18

Table 1. Markers used to identify EMP ex vivo.

Condition Markers Reference

Acute coronary syndromes CD31+ CD146+

[38]

CD31+/CD42- CD51+

[57]

CD31+/CD42- [58] Acute myocardial infarction CD105+/MCP-1+

CD105+/CD62E+ [59]

Allogenic hematopoitic stem cell recipients Annexin V+/CD62E+ [60] Dengue virus infection CD62E+ [46] Diabetes mellitus type 1 CD51+ [61]Diabetes mellitus type 2 Annexin V+/CD144+ [47]

CD31+ CD144+

[62] [63]

Healthy individuals CD105+ [64] Hemato-oncological patients CD31+/CD51+ [65] Hypercholesterolemia CD31+/CD42- [66] Hypertension (gestational) CD31+/CD42-

CD62E+ [67]

Impaired systemic artery elasticity CD31+/CD42+ [68] Lupus anticoagulant CD31+/CD51+ [34] Malaria (falciparum) CD51+ [44] Meningococcal sepsis Annexin V+/CD62E+ [69] Multiple organ dysfunction Annexin V+/CD144+ [70] Multiple sclerosis CD54+

CD62E+ [71]

CD31+/CD42- CD51+

[42]

CD31+ [72] Paroxysmal nocturnal haemoglobinuria CD105+/CD45-/CD41- [43]

CD105+/CD45-/CD54+ Postprandial hypertriglyceridemia CD31+/CD42-

CD51+ [73]

Pregnancy (preeclampsia) Annexin V+/CD62E+/ CD144+

[74]

CD31+/CD42- CD62E+

[75]

Severe systemic inflammatory response syndrome

CD31+/CD54+ [45]

Sickle cell disease Annexin V+/CD144+ [76]

Chapter 1

19

Condition Markers Reference

Stable coronary disease CD105+/MCP-1+ CD105+/CD62E+

[59]

ST-segment elevation myocardial infarction

CD31+ [77]

Systemic lupus erythematosus CD62E+ [46] Thrombotic thrombocytopenia purpura CD31+/CD42-

CD51+ [35]

CD31+/CD42- CD62E+

[78]

Vasculitis CD62E+ CD105+

[48]

Venous thromboembolism CD31+/CD42- CD62E+

[49]

Numerous antibodies (single or combination thereof) have been used by different

laboratories to identify EMP in human plasma samples. EMP were identified either by

positive staining (e.g. CD62E+ or CD144+) or by exclusion (e.g. CD31+/CD42-).

Antibodies used were: CD31: platelet endothelial cell adhesion molecule-1 (PECAM-1);

CD34: glycoprotein (GP) 105-120; CD41: GPIIb ( IIb); CD42b: GPIb; CD45: leukocyte

common antigen (LCA); CD51: ; CD54: intercellular adhesion molecule-1 (ICAM-1);

CD62E: E-selectin; CD105: endoglin; CD144: vascular endothelial cadherin; CD146: S-

endo-1; MCP-1: monocyte chemoattractant protein-1.

Introduction

20

References

1. Galley HF, Webster NR. Physiology of the endothelium. Br. J. Anaesth. 2004;93:105-13.

2. Palmer RM, Ashton DS, Moncada S. Vascular endothelial cells synthesize nitric oxide from L-arginine. Nature 1988;333:664-6.

3. Furchgott RF, Zawadzki JV. The obligatory role of endothelial cells in the relaxation of arterial smooth muscle by acetylcholine. Nature 1980;288:373-6.

4. Levin ER. Endothelins. N. Engl. J. Med. 1995;333:356-63.

5. Mantovani A, Bussolino F, Introna M. Cytokine regulation of endothelial cell function: from molecular level to the bedside. Immunol. Today 1997;18:231-40.

6. Mantovani A, Sozzani S, Vecchi A, Introna M, Allavena P. Cytokine activation of endothelial cells: new molecules for an old paradigm. Thromb. Haemost. 1997;78:406-14.

7. Mantovani A, Bussolino F, Dejana E. Cytokine regulation of endothelial cell function. FASEB J. 1992;6:2591-9.

8. Moriwaki H, Kume N, Sawamura T, Aoyama T, Hoshikawa H, Ochi H, Nishi E, Masaki T, Kita T. Ligand specificity of LOX-1, a novel endothelial receptor for oxidized low density lipoprotein. Arterioscler. Thromb. Vasc. Biol. 1998;18:1541-7.

9. Bussolino F, Bocchietto E, Silvagno F, Soldi R, Arese M, Mantovani A. Actions of molecules which regulate hemopoiesis on endothelial cells: memoirs of common ancestors? Pathol. Res. Pract. 1994;190:834-9.

10. Wight TN. Cell biology of arterial proteoglycans. Arteriosclerosis 1989;9:1-20.

11. Wight TN. The extracellular matrix and atherosclerosis. Curr. Opin. Lipidol. 1995;6:326-34.

12. Weksler BB, Marcus AJ, Jaffe EA. Synthesis of prostaglandin I2 (prostacyclin) by cultured human and bovine endothelial cells. Proc. Natl. Acad. Sci. U. S. A 1977;74:3922-6.

13. Van de WM, Collen D, Conway EM. Thrombomodulin-protein C-EPCR system: integrated to regulate coagulation and inflammation. Arterioscler. Thromb. Vasc. Biol. 2004;24:1374-83.

14. Michiels C. Endothelial cell functions. J. Cell Physiol 2003;196:430-43.

Chapter 1

21

15. Esmon CT. Molecular events that control the protein C anticoagulant pathway. Thromb. Haemost. 1993;70:29-35.

16. Colucci M, Balconi G, Lorenzet R, Pietra A, Locati D, Donati MB, Semeraro N. Cultured human endothelial cells generate tissue factor in response to endotoxin. J. Clin. Invest. 1983;71:1893-6.

17. Ruggeri ZM. Glycoprotein Ib and von Willebrand factor in the process of thrombus formation. Ann. N. Y. Acad. Sci. 1994;714:200-10.

18. Adams DH, Shaw S. Leucocyte-endothelial interactions and regulation of leucocyte migration. Lancet 1994;343:831-6.

19. Mantovani A, Dejana E. Cytokines as communication signals between leukocytes and endothelial cells. Immunol. Today 1989;10:370-5.

20. Cai H, Harrison DG. Endothelial dysfunction in cardiovascular diseases: the role of oxidant stress. Circ. Res. 2000;87:840-4.

21. Suwaidi JA, Hamasaki S, Higano ST, Nishimura RA, Holmes DR, Jr., Lerman A. Long-term follow-up of patients with mild coronary artery disease and endothelial dysfunction. Circulation 2000;101:948-54.

22. Ferro D, Pittoni V, Quintarelli C, Basili S, Saliola M, Caroselli C, Valesini G, Violi F. Coexistence of anti-phospholipid antibodies and endothelial perturbation in systemic lupus erythematosus patients with ongoing prothrombotic state. Circulation 1997;95:1425-32.

23. Jensen T, Bjerre-Knudsen J, Feldt-Rasmussen B, Deckert T. Features of endothelial dysfunction in early diabetic nephropathy. Lancet 1989;1:461-3.

24. Clozel M, Kuhn H, Hefti F, Baumgartner HR. Endothelial dysfunction and subendothelial monocyte macrophages in hypertension. Effect of angiotensin converting enzyme inhibition. Hypertension 1991;18:132-41.

25. Brodsky SV, Yamamoto T, Tada T, Kim B, Chen J, Kajiya F, Goligorsky MS. Endothelial dysfunction in ischemic acute renal failure: rescue by transplanted endothelial cells. Am. J. Physiol Renal Physiol. 2002;282:F1140-F1149.

26. Holzmann S. Endothelium-induced relaxation by acetylcholine associated with larger rises in cyclic GMP in coronary arterial strips. J. Cyclic. Nucleotide. Res. 1982;8:409-19.

27. Ludmer PL, Selwyn AP, Shook TL, Wayne RR, Mudge GH, Alexander RW, Ganz P. Paradoxical vasoconstriction induced by acetylcholine in atherosclerotic coronary arteries. N. Engl. J. Med. 1986;315:1046-51.

Introduction

22

28. Celermajer DS, Sorensen KE, Gooch VM, Spiegelhalter DJ, Miller OI, Sullivan ID, Lloyd JK, Deanfield JE. Non-invasive detection of endothelial dysfunction in children and adults at risk of atherosclerosis. Lancet 1992;340:1111-5.

29. Aengevaeren WR. Beyond lipids - the role of the endothelium in coronary artery disease. Atherosclerosis 1999;147 Suppl 1:S11-S16.

30. Corretti MC, Anderson TJ, Benjamin EJ, Celermajer D, Charbonneau F, Creager MA, Deanfield J, Drexler H, Gerhard-Herman M, Herrington D, Vallance P, Vita J, Vogel R. Guidelines for the ultrasound assessment of endothelial-dependent flow-mediated vasodilation of the brachial artery: a report of the International Brachial Artery Reactivity Task Force. J. Am. Coll. Cardiol. 2002;39:257-65.

31. De MG, Bravi MC, Costanzo A, Laurenti O, Cassone FM, Armiento A, De LO, Ferri C. Effects of insulin on in vitro vascular cell adhesion molecule-1 expression and in vivo soluble VCAM-1 release. Diabetologia 1999;42:1235-9.

32. Leeuwenberg JF, Smeets EF, Neefjes JJ, Shaffer MA, Cinek T, Jeunhomme TM, Ahern TJ, Buurman WA. E-selectin and intercellular adhesion molecule-1 are released by activated human endothelial cells in vitro. Immunology 1992;77:543-9.

33. Horstman LL, Jy W, Jimenez JJ, Ahn YS. Endothelial microparticles as markers of endothelial dysfunction. Front. Biosci. 2004;9:1118-35.

34. Combes V, Simon AC, Grau GE, Arnoux D, Camoin L, Sabatier F, Mutin M, Sanmarco M, Sampol J, Dignat-George F. In vitro generation of endothelial microparticles and possible prothrombotic activity in patients with lupus anticoagulant. J. Clin. Invest. 1999;104:93-102.

35. Jimenez JJ, Jy W, Mauro LM, Horstman LL, Ahn YS. Elevated endothelial microparticles in thrombotic thrombocytopenic purpura: findings from brain and renal microvascular cell culture and patients with active disease. Br. J. Haematol. 2001;112:81-90.

36. Kagawa H, Komiyama Y, Nakamura S, Miyaka T, Miyazaki Y. Expression of functional tissue factor on small vesicles of lipopolysaccharide-stimulated human vascular endothelial cells. Thromb. Res. 1998;91:297-304.

37. Simak J, Holada K, Vostal JG. Release of annexin V-binding membrane microparticles from cultured human umbilical vein endothelial cells after treatment with camptothecin. BMC. Cell Biol. 2002;3:11.

38. Mallat Z, Benamer H, Hugel B, Benessiano J, Steg PG, Freyssinet JM, Tedgui A. Elevated levels of shed membrane microparticles with procoagulant potential in

Chapter 1

23

the peripheral circulating blood of patients with acute coronary syndromes. Circulation 2000;101:841-3.

39. Simak J, Gelderman MP, Yu H, Wright V, Baird AE. Circulating endothelial microparticles in acute ischemic stroke: a link to severity, lesion volume and outcome. J. Thromb. Haemost. 2006;4:1296-302.

40. Ederhy S, Di AE, Mallat Z, Hugel B, Janower S, Meuleman C, Boccara F, Freyssinet JM, Tedgui A, Cohen A. Levels of circulating procoagulant microparticles in nonvalvular atrial fibrillation. Am. J. Cardiol. 2007;100:989-94.

41. Arteaga RB, Chirinos JA, Soriano AO, Jy W, Horstman L, Jimenez JJ, Mendez A, Ferreira A, de ME, Ahn YS. Endothelial microparticles and platelet and leukocyte activation in patients with the metabolic syndrome. Am. J. Cardiol. 2006;98:70-4.

42. Minagar A, Jy W, Jimenez JJ, Sheremata WA, Mauro LM, Mao WW, Horstman LL, Ahn YS. Elevated plasma endothelial microparticles in multiple sclerosis. Neurology 2001;56:1319-24.

43. Simak J, Holada K, Risitano AM, Zivny JH, Young NS, Vostal JG. Elevated circulating endothelial membrane microparticles in paroxysmal nocturnal haemoglobinuria. Br. J. Haematol. 2004;125:804-13.

44. Combes V, Taylor TE, Juhan-Vague I, Mege JL, Mwenechanya J, Tembo M, Grau GE, Molyneux ME. Circulating endothelial microparticles in malawian children with severe falciparum malaria complicated with coma. JAMA 2004;291:2542-4.

45. Ogura H, Tanaka H, Koh T, Fujita K, Fujimi S, Nakamori Y, Hosotsubo H, Kuwagata Y, Shimazu T, Sugimoto H. Enhanced production of endothelial microparticles with increased binding to leukocytes in patients with severe systemic inflammatory response syndrome. J. Trauma 2004;56:823-31.

46. Abid Hussein MN, Meesters EW, Osmanovic N, Romijn FP, Nieuwland R, Sturk A. Antigenic characterization of endothelial cell-derived microparticles and their detection ex vivo. J. Thromb. Haemost. 2003;1:2434-43.

47. Tushuizen ME, Nieuwland R, Rustemeijer C, Hensgens BE, Sturk A, Heine RJ, Diamant M. Elevated endothelial microparticles following consecutive meals are associated with vascular endothelial dysfunction in type 2 diabetes. Diabetes Care 2007;30:728-30.

48. Brogan PA, Shah V, Brachet C, Harnden A, Mant D, Klein N, Dillon MJ. Endothelial and platelet microparticles in vasculitis of the young. Arthritis Rheum. 2004;50:927-36.

Introduction

24

49. Chirinos JA, Heresi GA, Velasquez H, Jy W, Jimenez JJ, Ahn E, Horstman LL, Soriano AO, Zambrano JP, Ahn YS. Elevation of endothelial microparticles, platelets, and leukocyte activation in patients with venous thromboembolism. J. Am. Coll. Cardiol. 2005;45:1467-71.

50. Hladovec J. Circulating endothelial cells as a sign of vessel wall lesions. Physiol. Bohemoslov. 1978;27:140-4.

51. Blann AD, Woywodt A, Bertolini F, Bull TM, Buyon JP, Clancy RM, Haubitz M, Hebbel RP, Lip GY, Mancuso P, Sampol J, Solovey A, Dignat-George F. Circulating endothelial cells. Biomarker of vascular disease. Thromb. Haemost. 2005;93:228-35.

52. Frisch SM, Ruoslahti E. Integrins and anoikis. Curr. Opin. Cell Biol. 1997;9:701-6.

53. Levkau B, Herren B, Koyama H, Ross R, Raines EW. Caspase-mediated cleavage of focal adhesion kinase pp125FAK and disassembly of focal adhesions in human endothelial cell apoptosis. J. Exp. Med. 1998;187:579-86.

54. Zoellner H, Hofler M, Beckmann R, Hufnagl P, Vanyek E, Bielek E, Wojta J, Fabry A, Lockie S, Binder BR. Serum albumin is a specific inhibitor of apoptosis in human endothelial cells. J. Cell Sci. 1996;109:2571-80.

55. Satta N, Toti F, Feugeas O, Bohbot A, Dachary-Prigent J, Eschwege V, Hedman H, Freyssinet JM. Monocyte vesiculation is a possible mechanism for dissemination of membrane-associated procoagulant activities and adhesion molecules after stimulation by lipopolysaccharide. J. Immunol. 1994;153:3245-55.

56. Hamilton KK, Hattori R, Esmon CT, Sims PJ. Complement proteins C5b-9 induce vesiculation of the endothelial plasma membrane and expose catalytic surface for assembly of the prothrombinase enzyme complex. J. Biol. Chem. 1990;265:3809-14.

57. Bernal-Mizrachi L, Jy W, Jimenez JJ, Pastor J, Mauro LM, Horstman LL, De Marchena E, Ahn YS. High levels of circulating endothelial microparticles in patients with acute coronary syndromes. Am. Heart J. 2003;145:962-70.

58. Bernal-Mizrachi L, Jy W, Fierro C, Macdonough R, Velazques HA, Purow J, Jimenez JJ, Horstman LL, Ferreira A, De Marchena E, Ahn YS. Endothelial microparticles correlate with high-risk angiographic lesions in acute coronary syndromes. Int. J. Cardiol. 2004;97:439-46.

Chapter 1

25

59. Heloire F, Weill B, Weber S, Batteux F. Aggregates of endothelial microparticles and platelets circulate in peripheral blood. Variations during stable coronary disease and acute myocardial infarction. Thromb. Res. 2003;110:173-80.

60. Pihusch V, Rank A, Steber R, Pihusch M, Pihusch R, Toth B, Hiller E, Kolb HJ. Endothelial cell-derived microparticles in allogeneic hematopoietic stem cell recipients. Transplantation 2006;81:1405-9.

61. Sabatier, F., Darmon, P., Hugel, B., Combes, V., Sanmarco, M., Velut, J. G., Arnoux, D., Charpiot, P., Freyssinet, J. M., Oliver, C., Sampol, J., and Dignat-George, F. Type 1 and type 2 diabetic patients display different patterns of cellular microparticles. Diabetes 2002;51:2840-45.

62. Morel O, Jesel L, Hugel B, Douchet MP, Zupan M, Chauvin M, Freyssinet JM, Toti F. Protective effects of vitamin C on endothelium damage and platelet activation during myocardial infarction in patients with sustained generation of circulating microparticles. J. Thromb. Haemost. 2003;1:171-7.

63. Koga H, Sugiyama S, Kugiyama K, Watanabe K, Fukushima H, Tanaka T, Sakamoto T, Yoshimura M, Jinnouchi H, Ogawa H. Elevated levels of VE-cadherin-positive endothelial microparticles in patients with type 2 diabetes mellitus and coronary artery disease. J. Am. Coll. Cardiol. 2005;45:1622-30.

64. Simak J, Holada K, D'Agnillo F, Janota J, Vostal JG. Cellular prion protein is expressed on endothelial cells and is released during apoptosis on membrane microparticles found in human plasma. Transfusion 2002;42:334-42.

65. Inbal A, Lubetsky A, Shimoni A, Dardik R, Sela BA, Eskaraev R, Levi I, Tov NS, Nagler A. Assessment of the coagulation profile in hemato-oncological patients receiving ATG-based conditioning treatment for allogeneic stem cell transplantation. Bone Marrow Transplant. 2004;34:459-63.

66. Pirro M, Schillaci G, Paltriccia R, Bagaglia F, Menecali C, Mannarino MR, Capanni M, Velardi A, Mannarino E. Increased Ratio of CD31+/CD42- microparticles to endothelial progenitors as a novel marker of atherosclerosis in hypercholesterolemia. Arterioscler. Thromb. Vasc. Biol. 2006;26:2530-5.

67. Gonzalez-Quintero VH, Smarkusky LP, Jimenez JJ, Mauro LM, Jy W, Hortsman LL, O'Sullivan MJ, Ahn YS. Elevated plasma endothelial microparticles: Preeclampsia versus gestational hypertension. Am. J. Obstet. Gynecol. 2004;191:1418-24.

68. Wang JM, Huang YJ, Wang Y, Xu MG, Wang LC, Wang SM, Tao J. Increased circulating CD31+/CD42- microparticles are associated with impaired systemic artery elasticity in healthy subjects. Am. J. Hypertens. 2007;20:957-64.

Introduction

26

69. Nieuwland R, Berckmans RJ, McGregor S, Böing AN, Romijn FPHTM, Westendorp RGJ, Hack CE, Sturk A. Cellular origin and procoagulant properties of microparticles in meningococcal sepsis. Blood 2000;95:930-5.

70. Joop K, Berckmans RJ, Nieuwland R, Berkhout J, Romijn FPHTM, Hack CE, Sturk A. Microparticles from patients with multiple organ dysfunction syndrom and sepsis support coagulation through multiple mechanisms. Thromb. Haemost. 2001;85:810-20.

71. Jy W, Minagar A, Jimenez JJ, Sheremata WA, Mauro LM, Horstman LL, Bidot C, Ahn YS. Endothelial microparticles (EMP) bind and activate monocytes: elevated EMP-monocyte conjugates in multiple sclerosis. Front. Biosci. 2004;9:3137-44.

72. Sheremata WA, Jy W, Delgado S, Minagar A, McLarty J, Ahn Y. Interferon-beta-1a reduces plasma CD31+ endothelial microparticles (CD31+EMP) in multiple sclerosis. J. Neuroinflammation. 2006;3:23-7.

73. Ferreira AC, Peter AA, Mendez AJ, Jimenez JJ, Mauro LM, Chirinos JA, Ghany R, Virani S, Garcia S, Horstman LL, Purow J, Jy W, Ahn YS, de ME. Postprandial hypertriglyceridemia increases circulating levels of endothelial cell microparticles. Circulation 2004;110:3599-603.

74. VanWijk MJ, Nieuwland R, Boer K, van der Post JA, VanBavel E, Sturk A. Microparticle subpopulations are increased in preeclampsia: possible involvement in vascular dysfunction? Am. J. Obstet. Gynecol. 2002;187:450-6.

75. Gonzalez-Quintero VH, Jimenez JJ, Jy W, Mauro LM, Hortman L, O'Sullivan MJ, Ahn Y. Elevated plasma endothelial microparticles in preeclampsia. Am. J. Obstet. Gynecol. 2003;189:589-93.

76. Shet AS, Aras O, Gupta K, Hass MJ, Rausch DJ, Saba N, Koopmeiners L, Key NS, Hebbel RP. Sickle blood contains tissue factor-positive microparticles derived from endothelial cells and monocytes. Blood 2003;102:2678-83.

77. Morel O, Hugel B, Jesel L, Mallat Z, Lanza F, Douchet MP, Zupan M, Chauvin M, Cazenave JP, Tedgui A, Freyssinet JM, Toti F. Circulating procoagulant microparticles and soluble GPV in myocardial infarction treated by primary percutaneous transluminal coronary angioplasty. A possible role for GPIIb-IIIa antagonists. J. Thromb. Haemost. 2004;2:1118-26.

78. Jimenez JJ, Jy W, Mauro LM, Horstman LL, Soderland C, Ahn YS. Endothelial microparticles released in thrombotic thrombocytopenic purpura express von Willebrand factor and markers of endothelial activation. Br. J. Haematol. 2003;123:896-902.

27

Chapter 2

Antigenic characterization of endothelial cell-

derived microparticles and their detection ex

vivo

Mohammed N. Abid Hussein, Eelco W. Meesters, Nada Osmanovic, Fred P.H.Th.M.

Romijn, Rienk Nieuwland and Augueste Sturk

J. Thromb. Haemost. 2003;1:2434-2443

Antigenic phenotype of endothelial microparticles

28

ABSTRACT

Background: Endothelial activation and dysfunction are associated with several diseases.

However, hardly any specific markers are available. Microparticles (MP) from endothelial

cells (EC; EMP) were reported in patient groups and healthy individuals. The antibodies

used to detect EMP, however, were mainly directed against antigens without EC

specificity.

Objectives: We evaluated the antigens on EC and EMP to establish proper markers for

EMP detection.

Methods: EMP were isolated from supernatants of resting and interleukin (IL)-1

activated human umbilical vein EC (HUVEC; n=3; 0-72 hour), stained with annexin V

and monoclonal antibodies, and analyzed by flow cytometry. Human platelet-MP (PMP),

the main MP population in plasma, were prepared in vitro. EMP and PMP were studied in

plasma from systemic lupus erythematosus (SLE) patients (n=11) and healthy individuals

(n=10).

Results: Platelet-endothelial cell adhesion molecule-1 (PECAM-1), and 3 were

constitutively exposed on HUVEC, but (almost) absent on EMP ( 15% positive for

and 3), or only exposed on a subpopulation (PECAM-1; 30-60%). Activated HUVEC

( 80%) and (subpopulations of) EMP exposed E-selectin and tissue factor. PMP strongly

exposed PECAM-1, 3 and glycoprotein (GP)Ib (CD42b), but not or E-selectin. GPIb

and P-selectin (CD62P) were absent on EMP. Plasma samples contained 0.5% MP

staining for E-selectin and/or . Plasma from one SLE patient contained E-selectin

exposing MP (21%), but little -positive MP.

Conclusions: EC release EMP in vitro. The antigenic phenotype of EMP released from

resting and IL-1 -stimulated EC differs among each other as well as from resting and

stimulated EC, respectively. E-selectin exposed on IL-1 -stimulated EC is a valid marker

for EMP detection ex vivo to establish endothelial cell activation.

Chapter 2

29

INTRODUCTION

In physiological conditions, endothelial cells (EC) play an important role in

homeostasis of the blood. This homeostasis is lost during pathological conditions, at least

in part by increased exposure of procoagulant and proadhesive antigens on their surface.

For example, only activated EC expose tissue factor (TF), the initiator of coagulation in

vivo, and E-selectin, which facilitates reversible adhesion of white blood cells as part of

the inflammatory response [1-4]. Endothelial dysfunction is associated with several

disease states such as preeclampsia, thrombotic thrombocytopenic purpura (TTP),

diabetes, systemic lupus erythematosus (SLE), lupus anticoagulant, atherosclerosis,

inflammation, hypertension, and coronary artery disease [5-10]. At present, there are only

a few markers for the detection of endothelial activation and/or dysfunction ex vivo such

as von Willebrand Factor (vWF) and soluble (s) E-selectin [11-13].

In vitro, activated EC show surface blebbing and the subsequent shedding of small

vesicles (microparticles; MP) [14-18]. Recent studies report the presence of endothelial

cell-derived microparticles (EMP) in peripheral blood from patients with lupus

anticoagulant, TTP, acute coronary syndromes, and even in blood of healthy individuals

[14,15,19]. However, in most studies the identification of EMP was based on the presence

of surface antigens that are not exclusively exposed on EC, such as the platelet-endothelial

cell adhesion molecule-1 (PECAM-1; CD31) or the vitronectin receptor ( 3; : CD51,

3: CD61). In addition, a recent study showed that MP from human erythrocytes

significantly differed in their antigenic composition from their corresponding parent cells

[20]. Thus, a comprehensive characterization of the antigenic composition of EMP is

required to accurately identify such vesicles in mixed populations of MP of various

cellular origins as present in, for example, the venous blood of healthy individuals and

patients. The aim of the present study was to compare the antigenic phenotype of EC and

EMP under resting and activation conditions to establish reliable markers to quantify

EMP.


30

MATERIALS AND METHODS

Reagents and assays

Medium M199, penicillin, streptomycin, and L-glutamine were obtained from

GibcoBRL, Life Technologies (Paisley, UK). Immunoglobulin (Ig)G1-fluorescein

isothiocyanate (FITC) and IgG1-phycoerythrin (PE) (clone X40), CD31-PE (clone WM-

59, IgG1), CD34-PE (clone My10, IgG1), and CD61-PE (clone VI-PL2, IgG1) were

obtained from Becton Dickinson ((BD) San Jose, CA, USA). CD42b-PE (clone CLB-

MB45, IgG1), CD42b-FITC (clone CLB-MB45, IgG1), fetal calf serum (heat inactivated

during 30 minutes at 56 ºC; FCSi), normal mouse serum, and human serum albumin

(HSA) were obtained from the Central Laboratory of the Netherlands Red Cross

Bloodtransfusion Service (CLB; Amsterdam, The Netherlands). CD61-FITC (clone

Y2/51, IgG1) was from Dako A/S (Glostrup, Denmark). CD51-FITC (clone AMF7, IgG1),

CD62P-FITC (clone CLB-Thromb/6, IgG1), CD54-PE (clone 84H10, IgG1) and CD62P-

PE (clone CLB-Thromb/6, IgG1) were from Immunotech (Marseille, France). CD62E-

FITC (clone 1.2B6, IgG1) was obtained from Serotec Ltd. (Kidlington, UK), CD62E-PE

(clone HAE-1f, IgG1) from Ancell (Lausen, Switzerland), CD106-FITC (clone 1.G11B1,

IgG1) from Calbiochem (La Jolla, CA, USA), CD141-PE (clone B-A35, IgG1) from

Diaclone (Besançon, France), anti-tissue factor-FITC (4508CJ, IgG1) from American

Diagnostics Inc. (Greenwich, CT, USA), and CD144-FITC (clone BMS158F1, IgG1) from

MedSystems Diagnostics GmbH (Vienna, Austria). Recombinant human interleukin (IL)-

1, human recombinant basic fibroblast growth factor (bFGF), and epidermal growth factor

(EGF) were from GibcoBRL (Gaithersburg, MD, USA). Annexin V-(allophycocyanin;

APC) was from Caltag Laboratories (Burlingame, CA, USA), collagenase (type 1A) from

Sigma (St. Louis, MO, USA), ethylenediamine tetra-acetic acid (EDTA) from Merck

(Darmstadt, Germany), heparin (400 U/mL) from Bufa BV (Uitgeest, The Netherlands),

calcium ionophore A23187 from Calbiochem (Darmstadt, Germany), and trypsin from

Difco Laboratories (Detroit, MI, USA). Human serum was provided by the Blood Bank

Center (Leiden University Medical Center) and was heat inactivated during 30 minutes at

56 ºC (HuSi). Tissue culture flasks were from Greiner Labortechnik (Frickenhausen,

Germany) and gelatin from Difco Laboratories (Sparks, MD, USA).

Chapter 2

31

Isolation and culture of human umbilical vein endothelial cells

Human umbilical vein endothelial cells (HUVEC) were collected from human

umbilical cord veins by minor modifications of previously described protocols [21,22].

Briefly, umbilical cords were filled with 1 mg/mL collagenase in M199 and subsequently

incubated in phosphate-buffered saline (PBS) (154 mmol/L NaCl, 1.4 mmol/L phosphate;

pH 7.5) for 20 minutes at 37 °C. Detached cells were collected by perfusion with medium

M199 supplemented with 10% HuSi. The cell suspension was centrifuged for 10 minutes

at 180 g and 20 °C. Cells were resuspended in M199 (37 °C) supplemented with 10%

HuSi, 2 mmol/L L-glutamine, 1 mg/mL penicillin, 0.1 mg/mL streptomycin, 0.5 μg/mL

fungizone, 10 ng/mL EGF, 20 ng/mL bFGF, and 5 U/mL heparin. HUVEC were cultured

in 25-cm2 tissue culture flasks coated with 0.75% gelatin (passage 0). Upon confluency,

the HUVEC were transferred to 75-cm2 tissue culture flasks coated with 0.75% gelatin

(passage 1). Cells were detached by trypsinisation (0.05% w/v trypsin and 2.7 mmol/L

EDTA in PBS, pH 7.4), and transferred twice more to 75-cm2 tissue culture flasks coated

with 0.75% gelatin (passage 3).

HUVEC stimulation and flow cytometric analysis

Upon confluency at passage 3, HUVEC were kept for 3-4 days in a resting state

before stimulation with IL-1 (5 ng/mL). IL-1 was prepared as stock solution (10

g/mL) in M199 and added (5 L) to 10 mL culture medium. After incubation for various

time intervals, culture supernatants were collected for MP analysis and the cells were

harvested by trypsinisation. After 4 minutes, trypsin was neutralized by 1% FCSi in PBS

(pH 7.4). The obtained HUVEC suspension was washed twice by centrifugation for 10

minutes at 180 g and 4 °C, and resuspended in PBS/FCSi. HUVEC were then kept on

melting ice for 15 minutes, centrifuged for 10 minutes at 180 g and 4 °C and resuspended

in PBS/FCSi. Monoclonal antibodies (MoAbs) (5 L) were added to 45 L cell

suspension. For HUVEC staining, the final MoAb concentrations used were 0.5 g/mL

for IgG1-FITC and IgG1-PE, 0.06 g/mL for CD31-PE, 0.5 g/mL for CD34-PE, 20

g/mL for CD51-FITC, 0.03 g/mL for CD54-PE, 2 g/mL for CD61-FITC, 1 g/mL for

CD62E-FITC and anti-TF-FITC, and 10 g/mL for CD144-FITC. Dilutions of CD106-

FITC and CD141-PE were 1:50 (v/v) and 1:10 (v/v), respectively. The cell suspension


32

was incubated with MoAbs in the dark for 30 minutes and 4 °C. After incubation, the

HUVEC were washed by addition of 1 mL PBS/FCSi, centrifuged for 10 minutes at 180 g

and 4 °C, and resuspended in 300 L PBS/FCSi (on melting ice). In each sample, 5,000

cells were analyzed in a FACScan flow cytometer with CellQuest software (BD; San Jose,

CA, USA) [22].

Isolation of MP

At various time intervals, culture supernatants were harvested and centrifuged for 10

minutes at 180 g and 20 °C to remove whole cells. Subsequently, aliquots of the cell-free

culture supernatant (250 L each) were snap-frozen in liquid nitrogen and stored at – 80

°C. Alternatively, plasma samples from citrate-anticoagulated venous blood of SLE

patients and healthy controls (with their informed consent) were collected and handled as

described previously [23]. Aliquots (250 L each) were snapfrozen in liquid nitrogen and

stored at – 80 °C. Before use, all samples were kept on melting ice to allow thawing for 1

hour. After thawing, samples were centrifuged for 30 minutes at 17,570 g and 20 °C.

Then, 225 L of (MP-free) supernatant were removed. The remaining 25 L (MP-

enriched) suspension was diluted with 225 L PBS containing 10.9 mmol/L trisodium

citrate. MP were resuspended and again centrifuged for 30 minutes at 17,570 g and 20 °C.

Again, 225 L of supernatant was removed and MP were resuspended in the remaining 25

L. For flow cytometry detection of platelet-MP (PMP) and EMP from SLE patients and

controls, this MP suspension (25 L) was diluted 4-fold with PBS/citrate (75 L; pH 7.4).

Preparation of PMP in vitro

Venous blood (8.4 mL) from three healthy controls was collected (with their informed

consent) into 1.6 mL 3.2% acid citrate dextrose solution (ACD; 85 mmol/L trisodium

citrate, 11 mmol/L glucose, 7 mmol/L citric acid; pH 4.4). Blood was centrifuged for 15

minutes at 180 g and room temperature, and platelet-rich plasma was collected. Platelet-

rich plasma was centrifuged for 20 minutes at 1,000 g and room temperature. The

supernatant was removed and the platelet pellet was gently resuspended in 10 mL Tyrode

buffer (136.9 mmol/L NaCl, 11.9 mmol/L NaHCO3, 5.6 mmol/L glucose, 1.0 mmol/L

MgCl2, 2.7 mmol/L KCl and 0.36 mmol/L NaH2PO4; pH 6.5) containing HSA (0.25%

Chapter 2

33

w/v) and EDTA (2.0 mmol/L). This platelet suspension was again centrifuged for 20

minutes at 1,000 g and room temperature, and the supernatant was removed. The platelet

pellet was resuspended in 0.5 mL Tyrode buffer (pH 7.4) containing 2 mmol/L CaCl2

instead of EDTA. After further diluting the platelet suspension with 3.5 mL of Tyrode

buffer (pH 7.4), platelets were removed, counted, and adjusted to approximately 2.0 x

105/ L. Subsequently, platelets were activated by addition of calcium ionophore A23187

(2.5 mol/L final concentration) at 37 °C (non-stirring conditions). After 20 minutes,

EDTA (5 mmol/L) was added, platelets were removed by centrifugation for 20 minutes at

1,000 g and room temperature. The supernatant, containing PMP, was used for flow

cytometric analysis.

Flow cytometric analysis

MP samples were analyzed in a FACSCalibur flow cytometer (BD; San Jose, CA).

Forward scatter (FSC) and side scatter (SSC) were set at logarithmic gain and MP were

identified as described previously [24,25]. MP were identified on FSC, SSC, and binding

of a MoAb. More than 80% of the events identified using these criteria also stained for

annexin V (data not shown). Fluorescence thresholds for MoAb were set in terms of

binding of isotype-matched control antibodies (all IgG1). Fluorescence was measured in

the FL-1 channel (FITC), FL-2 channel (PE) and FL-4 channel (APC). MP (5 L) were

diluted with 35 L PBS containing 2.5 mmol/L CaCl2 (pH 7.4) and 5 L of (1:500 diluted

in PBS) normal mouse serum. After incubation for 15 minutes at room temperature, 5 L

annexin V-APC (0.66 g/mL final concentration) and 5 L MoAb or isotype-matched

control antibody (IgG1) were added. For MP analysis, antibody concentrations used were

0.5 g/mL for IgG1-FITC and IgG1-PE, 0.06 g/mL for CD31-PE 0.5 g/mL for CD34-

PE, 1 g/mL for CD42b-FITC and CD42b-PE, 10 g/mL for CD51-FITC, 0.03 g/mL for

CD54-PE, 1 g/mL for CD61-FITC, 1.6 g/mL for CD62E-PE, 2.5 g/mL for CD62P-

FITC, 0.0625 g/mL for CD62P-PE, 0.5 g/mL for anti-TF-FITC and CD144-FITC.

Dilution of CD61-PE was 1:100 (v/v CD106-FITC) and CD141-PE were both diluted

1:50 (v/v). The mixture of MP, normal mouse serum and MoAbs was then incubated for

15 minutes in the dark at room temperature. To remove the excess of free MoAb, 200 L

PBS/calcium buffer was added and the suspension was centrifuged for 30 minutes at


34

17,570 g at 20 °C. Finally, 200 L of supernatant were removed, and MP were

resuspended with 300 L PBS/calcium. All samples were analyzed for 2 minutes.

Patients and healthy controls

In the present study, 11 SLE patients (all women) were included, all of whom fulfilled

the revised criteria of the American College of Rheumatology for the diagnosis of SLE

[26]. Their age was 42 years (median; range 23-64). The SLE Disease Activity Index [27]

was 9 (median; range 0-22). As controls, 10 age-matched women were included. The

study fulfilled the guidelines of the Medical Ethical Committee of the Slotervaart

Hospital.

Statistical analysis

Data were analyzed with Prism (3.02) for Windows. For direct comparison of the

binding of MoAbs to HUVEC and EMP, paired t-tests were used. To compare the

differences in MoAb binding to plasma samples from SLE patients and healthy

volunteers, the Mann-Whitney U test was used. Two-tailed significance levels (P) are

presented. Differences were considered statistically significant at P<0.05.

RESULTS

Antigenic exposure of resting and IL-1 -activated HUVEC

HUVEC were incubated for various time intervals up to 72 hours with and without

IL-1 (5 ng/mL). Figure 1 shows representative dot plots of the surface antigen exposure

of PECAM-1 and E-selectin. Both resting (Figure 1A) and activated (Figure 1B) HUVEC

exposed PECAM-1. In contrast, E-selectin was exposed only on the activated HUVEC

(Figure 1D versus 1C). The overall data are summarized in Figure 2 and in Table 1. As is

evident from Figure 2, PECAM-1 (CD31), (CD51), and 3 (CD61) are exposed on all

HUVEC independent of their activation status (Figure 2B, D, F). About 20% of the

resting HUVEC exposed TF (Figure 2J), whereas E-selectin was not exposed (Figure 2H).

Three hours after addition of IL-1 (the first measuring period), HUVEC exposed TF and

E-selectin. The exposure of both antigens was transient and gradually diminished after 12

Chapter 2

35

hours. Because the maximal antigen exposure of inducible antigens on the HUVEC

occurred 12-24 hours after addition of IL-1 , we summarized the overall data for all

studied antigens in Table 1 at that activation period. From Table 1, it is apparent that the

antigens PECAM-1, , 3, GP105-120 (CD34), vascular endothelial cadherin (CD144),

and to a lesser extent intercellular adhesion molecule-1 (ICAM-1; CD54), vascular cell

adhesion molecule-1 (VCAM-1; CD106) and thrombomodulin (CD141), were all exposed

on most of the HUVEC regardless of their activation status, although the surface exposure

of ICAM-1 and VCAM-1 increased upon cell activation. For other antigens studied, i.e.

E-selectin and TF, the surface exposure was inducible upon activation of the HUVEC.


100 102 103 1041010

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- IL-1 + IL-1

Figure 1. Surface exposure of PECAM-1 and E-selectin on resting and IL-1 -activated

HUVEC. HUVEC were incubated with (B, D) or without (A, C) IL-1 (5 ng/ml) for 12

hours. PECAM-1 (CD31) and E-selectin (CD62E) are shown as examples of a

constitutive and inducible cell surface antigen, respectively. All data were obtained within

one typical experiment. The line indicates the fluorescence threshold, which was set using

an isotype-matched control antibody.

36

Chapter 2

Figure 2. Surface exposure of antigens on resting and IL-1 -activated HUVEC. HUVEC were activated at t=0 by addition of IL-1(5 ng/mL). At various time intervals (0-72 hours), the exposure of the indicated antigens was determined by flow cytometry. In the left panel, representative histograms for PECAM-1 (CD31; A), (CD51; C),

3 (CD61; E), E-selectin (CD62E; G) and TF (CD142; I), at the 12 hours time interval, are shown. The thin dotted lines indicate the binding of isotype-matched control antibody to resting- and activated cells, the filled curves the exposure of the antigens on resting HUVEC, and the thick black (open) curves the antigen exposure on the activated cells. In the right panels (B, D, F, H and J), the percentages of HUVEC that stained positive for the indicated antigens for resting- and activated cells at various activation periods are depicted as ( ) and ( ), respectively. The data in the panels on the right are presented as mean ± SD (n=3).

Tissue Factor

E-selectin

3

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ells

J

37

38

Tabl

e 1.

Ant

igen

exp

osur

e on

HU

VEC

and

EM

P in

the

abse

nce

or p

rese

nce

of IL

-1.

HU

VE

C

EM

P

Ant

igen

- I

L-1

+ IL

-1P

P- I

L-1

+ IL

-1

PEC

AM

-1 (C

D31

) 98

, 99,

99

95, 9

9, 9

9 0.

422

19, 2

4, 2

7†58

, 58,

37

0.09

0 (C

D51

) 99

, 99,

99

95, 9

9, 9

9 0.

422

7, 2

2, 8

†24

, 13,

14

0.59

8 3 (

CD

61)

99, 9

9, 9

9 95

, 99,

99

0.57

9 4,

8, 8

†1,

8, 3

0.20

7 E-

sele

ctin

(CD

62E)

6,

2, 6

85

, 97,

94

0.00

2 6,

5, 1

†58

, 52,

34

0.01

6 TF

(CD

142)

17

, 11,

23

76, 9

6, 7

4 0.

024

3, 2

, 3†

17, 7

, 5

0.19

1 IC

AM

-1 (C

D54

)‡67

, 83,

32

96, 9

8, 8

5 0.

100

5, 6

, 2

26, 4

3, 3

5 0.

024

GP1

05-1

20 (C

D34

) 90

, 92,

64

80, 8

3, 6

5 0.

229

10, 8

, 12†

40, 2

0, 2

5†0.

088

VC

AM

-1 (C

D10

6)

56, 6

2, 6

9 91

, 96,

93

0.01

2 10

, 10,

5†

7, 5

, 3†

0.06

3 Th

rom

bom

odul

in (C

D14

1)

77, 5

1, 3

1 53

, 29,

33

0.22

1 1,

5, 0

.3†

0.2,

0.2

, 0.3

†0.

335

VE-

cadh

erin

(CD

144)

84

, 78,

83

75, 5

7, 5

3 0.

081

3, 1

, 2†

2, 1

, 2†

0.42

2

Pair

ed t-

test

. The

dat

a of

the

thre

e in

divi

dual

exp

erim

ents

are

pre

sent

ed a

s pe

rcen

tage

s of

HU

VEC

and

EM

P po

sitiv

e fo

r th

e in

dica

ted

antig

ens

at 1

2 ho

urs

time

inte

rval

or

24 h

ours

† .‡ ICAM

-1 (

CD

54)

was

cha

ract

eriz

ed o

n H

UVE

C a

nd E

MP

from

thre

e ot

her

umbi

lical

cord

s.

Chapter 2

39

Antigen exposure on EMP

Next, the antigen exposure on EMP, obtained from resting and activated HUVEC,

was analyzed 12 hours after addition of IL-1 . Figure 3 shows that a subpopulation of

EMP from resting and stimulated HUVEC exposed PECAM-1 (Figure 3A, B).

Approximately 20%-30% of the EMP, released from resting HUVEC, exposed PECAM-

1. Upon activation, the percentage of PECAM-1-exposing EMP increased to almost 60%

(Figure 3B). Figure 3C, D show the exposure of E-selectin on EMP from resting (Figure

3C) and activated (Figure 3D) HUVEC. Whereas EMP from resting HUVEC hardly

stained for E-selectin (Figure 3C), EMP from HUVEC strongly stained for this antigen

upon cell activation. Figure 4 summarizes the exposure of PECAM-1, , 3, E-selectin

and TF on EMP from resting and IL-1 -activated HUVEC. The antigens that were

constitutively exposed on the HUVEC (PECAM-1, and 3), were exposed only on a

subpopulation of the EMP (PECAM-1, ) or even absent ( 3). The antigens that were

inducible on the HUVEC, i.e. E-selectin and TF, were virtually absent on EMP derived

from resting HUVEC, but as also shown in Figure 3, the EMP from activated HUVEC

strongly exposed E-selectin up to 72 hours. On these EMP, TF was present but barely

detectable by flow cytometry. Table 1 presents the percentages of EMP that exposed the

indicated antigens. The antigenic phenotype of the EMP differed remarkably from the

HUVEC and depended on the activation status of the parent cells. Not only and 3, but

also GP105-120, VCAM-1, CD141 and CD144 were not or hardly detectable on the EMP,

regardless of the activation status of the HUVEC. Interestingly, not only E-selectin but

also ICAM was exposed on the EMP, albeit to a lesser extent. As with E-selectin, this

marker only occurred on the EMP from activated HUVEC.


- IL-1 + IL-1

100 102 103 104101

A

PECAM-1

100

102

103

104

101

Side

scat

ter

100 102 103 104101

B

PECAM-1

100

102

103

104

101

Side

scat

ter

100 102 103 104101

C

E-selectin

100

102

103

104

101

Side

scat

ter

100 102 103 104101

D

E-selectin

100

102

103

104

101

Side

scat

ter

- IL-1 + IL-1

100 102 103 104101100 102 103 104101

A

PECAM-1

100

102

103

104

101

Side

scat

ter

100 102 103 104101100 102 103 104101

B

PECAM-1

100

102

103

104

101

Side

scat

ter

100 102 103 104101100 102 103 104101

C

E-selectin

100

102

103

104

101

Side

scat

ter

100 102 103 104101100 102 103 104101

D

E-selectin

100

102

103

104

101

Side

scat

ter

Figure 3. Antigen exposure on EMP. Microparticles were isolated from culture

supernatants of resting and IL-1 activated HUVEC, 12 hours after addition of IL-1 .

PECAM-1 was exposed on a subpopulation of EMP during resting (panel A) and

activated (B) conditions, whereas E-selectin was detectable only upon activation with IL-

1 (D versus C). The fluorescence thresholds were set independently for EMP from

resting and activated HUVEC with an isotype-matched control antibody for the indicated

antigen. Data from a representative experiment are shown.

40

Chapter 2

Figure 4. Antigens on EMP from resting and IL-1 activated HUVEC. EMP were isolated from culture supernatants of resting and IL-1 activated HUVEC. Representative examples of the exposure of PECAM-1 (A), (C), 3 (E), E-selectin (G) and TF (I), at 12 hours time interval, are shown as histograms (left panels). The thin dotted lines show the binding of isotype-matched control antibody to EMP from resting and activated HUVEC. The filled curves and the thick (open) curves show the antigen exposure on EMP from resting and activated HUVEC, respectively. At various time intervals (0-72 hours), the exposure of the indicated antigens was determined by flow cytometry (right panels). The exposure of the antigens on EMP from resting- or IL-1 -activated HUVEC are depicted as ( ) and ( ),respectively. The data are presented as mean ± SD (n=3).

104100 102 1031010

20

100A

Cou

nts

104100 102 1031010

20

100C

Cou

nts

104100 102 1031010

20

100E

Cou

nts

104100 102 1031010

20

100G

Cou

nts

104100 102 1031010

20

40

60

80

40

60

80

40

60

80

40

60

80

40

60

80

100I

Cou

nts

Time (hours)

F

0 12 24 36 48 60 720

20

40

60

80

100

% P

ositi

ve M

P

J

0 12 24 36 48 60 72Time (hours)

0

20

40

60

80

100

% P

ositi

ve M

P

H

0 12 24 36 48 60 72Time (hours)

0

20

40

60

80

100

% P

ositi

ve M

P

B

0 12 24 36 48 60 72Time (hours)

0

20

40

60

80

100

% P

ositi

ve M

PD

0 12 24 36 48 60 72Time (hours)

0

20

40

60

80

100%

Pos

itive

MP

Tissue Factor

E-selectin

3

PECAM-1104100 102 1031010

20

100A

Cou

nts

104100 102 1031010

20

100C

Cou

nts

104100 102 1031010

20

100E

Cou

nts

104100 102 1031010

20

100G

Cou

nts

104100 102 1031010

20

40

60

80

40

60

80

40

60

80

40

60

80

40

60

80

40

60

80

40

60

80

40

60

80

40

60

80

40

60

80

40

60

80

40

60

80

40

60

80

40

60

80

100I

Cou

nts

Time (hours)

F

0 12 24 36 48 60 720

20

40

60

80

100

0

20

40

60

80

100

% P

ositi

ve M

P

J

0 12 24 36 48 60 72Time (hours)

0

20

40

60

80

100

% P

ositi

ve M

P

J

0 12 24 36 48 60 72Time (hours)

0

20

40

60

80

100

0

20

40

60

80

100

% P

ositi

ve M

P

H

0 12 24 36 48 60 72Time (hours)

0

20

40

60

80

100

% P

ositi

ve M

P

H

0 12 24 36 48 60 72Time (hours)

0

20

40

60

80

100

0

20

40

60

80

100

% P

ositi

ve M

P

B

0 12 24 36 48 60 72Time (hours)

0

20

40

60

80

100

0

20

40

60

80

100

% P

ositi

ve M

PD

0 12 24 36 48 60 72Time (hours)

0

20

40

60

80

100

0

20

40

60

80

100%

Pos

itive

MP

Tissue Factor

E-selectin

3

PECAM-1

41


42

Comparison of the antigenic profile of PMP prepared in vitro with EMP

Subsequently, we compared the antigenic profile of EMP and PMP prepared in vitro

(Figure 5). Both EMP and PMP exposed PECAM-1 (Figure 5A and B, respectively). In

contrast to the EMP, PMP strongly stained for GPIb (CD42b), GPIIIa (CD61) and P-

selectin (CD62P) (Figure 5C, G, K versus D, H, L, respectively). (CD51) was nearly

absent on EMP (Figure 5E) and PMP (Figure 5F). The only marker that positively and

selectively identified EMP was E-selectin (Figure 5I), which was absent on PMP (Figure

5J).

Detection of EMP and PMP in plasma samples of SLE patients and healthy

individuals

Based on our current observations, we reinvestigated the presence of EMP and PMP

in plasma samples from patients with SLE and healthy individuals. In order to detect EMP

and PMP in these samples, MP were isolated and stained with combinations of MoAbs

directed against and E-selectin, and GPIb and GPIIIa, respectively. had previously

been used to quantify MP [14]. As shown in Table 2, most by far of the cell-derived MP

in plasma samples studied from SLE patients and controls (median 66%) strongly stained

for GPIb (CD42b), GPIIIa (CD61), or a combination of these two MoAbs. In contrast,

hardly any MP stained for either , E-selectin, or a combination of these two MoAbs

(Figure 6B, D, F). Also no ICAM-1 positive events were detected in these samples (data

not shown). Interestingly, an E-selectin-positive subpopulation of MP occurred in plasma

from the SLE-patient who had the highest SLE Disease Activity Index (22). Of the E-

selectin-positive EMP, only 12% double-labeled for (Figure 6A, C, E). These findings

indicate that MP of endothelial origin indeed occur in vivo.

Chapter 2

Figure 5. Comparison

of the antigenic profile

of PMP and EMP.

EMP were isolated

from culture

supernatant of

activated HUVEC, 12

hours after addition of

IL-1 . PMP were

prepared by stimulation

of isolated platelets

with calcium ionophore

A23187. Both EMP

(left panels) and PMP

(right panels) were

stained with MoAbs

directed against

PECAM-1 (CD31; A,

B), GPIb (CD42b; C,

D), (CD51; E, F), 3

(CD61; G, H), E-

selectin (CD62E; I, J)

and P-selectin

(CD62P; K, L). The

filled curves depict the

binding of IgG control

antibody, and the thick

(open) curves show the

binding of the indicated

MoAbs.

PMP

104100 102 1031010

20

40

60

80

100B

Cou

nts

EMP

104100 102 1031010

20

40

60

80

100A

Cou

nts

PECAM-1

104100 102 103101

DC

ount

s

0

20

40

60

80

100

104100 102 103101

C

Cou

nts

GPIb0

20

40

60

80

100

104100 102 103101

F

Cou

nts

0

20

40

60

80

100

104100 102 103101

E

Cou

nts

0

20

40

60

80

100

104100 102 103101

H

Cou

nts

0

20

40

60

80

100

104100 102 103101

G

Cou

nts

3

0

20

40

60

80

100

104100 102 103101

J

Cou

nts

0

20

40

60

80

100

104100 102 103101

I

Cou

nts

E-selectin0

20

40

60

80

100

104100 102 103101

L

Cou

nts

0

20

40

60

80

100

104100 102 103101

K

Cou

nts

P-selectin0

20

40

60

80

100

PMP

104100 102 1031010

20

40

60

80

100B

Cou

nts

EMP

104100 102 1031010

20

40

60

80

100A

Cou

nts

PECAM-1

104100 102 103101

DC

ount

s

0

20

40

60

80

100

104100 102 103101

C

Cou

nts

GPIb0

20

40

60

80

100

104100 102 103101

F

Cou

nts

0

20

40

60

80

100

104100 102 103101

E

Cou

nts

0

20

40

60

80

100

104100 102 103101

H

Cou

nts

0

20

40

60

80

100

104100 102 103101

G

Cou

nts

3

0

20

40

60

80

100

104100 102 103101

J

Cou

nts

0

20

40

60

80

100

104100 102 103101

I

Cou

nts

E-selectin0

20

40

60

80

100

104100 102 103101

L

Cou

nts

0

20

40

60

80

100

104100 102 103101

K

Cou

nts

P-selectin0

20

40

60

80

100

43


44

Table 2. Percentages of EMP and PMP in plasma samples from SLE patients (n=11) and

healthy controls (n=10).

MP MoAb SLE patients

(n=10)

SLE patient

(n=1)

Controls

(n=10) P†

EMP (CD51)†† 1 (0.3-2.0) 4 1 (0.4-3.0) 0.666

E-selectin (CD62E)†† 1 (0.2-8.0) 21 1 (0.2-2.0) 0.803

CD51 + CD62E§ 0.2 (0.01-0.6) 2.6 0.3 (0.04-2.0) 0.393

PMP GPIb (CD42b)†† 67.5 (54-83) 11 67 (40-83) 0.341

3 (CD61)†† 77 (64-94) 39 81 (55-95) 0.621

CD42b + CD61§ 66 (52-80) 7 66 (38-84) 1.000

Data from an SLE patient that showed E-selectin-positive MP ex vivo. †Mann-Whitney U

test on SLE patients (n=10) versus healthy controls (n=10). ††Data presented are the

percentages events positive for the antigen versus all events in the flow cytometric

analysis. All values are expressed as median (range). §Plotting the surface antigen

exposure of the two antigens against each other.

Chapter 2

SLE patient (E-selectin positive)

SLE patient(E-selectin negative)

100 102 103 104101

A

E-selectin

100

102

103

104

101

Side

scat

ter

100 102 103 104101

B

E-selectin

100

102

103

104

101

Side

scat

ter

100 102 103 104101

C

100

102

103

104

101

Side

scat

ter

100 102 103 104101

D

100

102

103

104

101

Side

scat

ter

100 102 103 104101

E

100

102

103

104

101

E-se

lect

in

100 102 103 104101

F

100

102

103

104

101

E-se

lect

in

SLE patient (E-selectin positive)

SLE patient(E-selectin negative)

100 102 103 104101100 102 103 104101

A

E-selectin

100

102

103

104

101

Side

scat

ter

100 102 103 104101100 102 103 104101

B

E-selectin

100

102

103

104

101

Side

scat

ter

100 102 103 104101100 102 103 104101

C

100

102

103

104

101

Side

scat

ter

100 102 103 104101100 102 103 104101

D

100

102

103

104

101

Side

scat

ter

100 102 103 104101

E

100

102

103

104

101

E-se

lect

in

100 102 103 104101100 102 103 104101

F

100

102

103

104

101

E-se

lect

in

Figure 6. Exposure of E-selectin on MP from an SLE-patient. EMP were isolated from a

plasma sample of an SLE patient (left panels), who showed E-selectin-positive EMP (A),

hardly -exposing MP (C) and insignificant double label for both E-selectin and (E).

For comparison, the right panels show representative dot plots from one out of ten SLE

patients who hardly stained for E-selectin (B), (D) or the combination of E-selectin and

.

45


46

DISCUSSION

The present finding that the antigenic phenotype of EMP differs considerably from

the HUVEC suggests that a sorting of membrane proteins occurs during membrane

vesiculation. Interestingly, a recent study showed that calcium ionophore-activated

erythrocytes release microparticles that antigenically differ from their parent cells [20].

The selective sorting of membrane proteins into MP is likely to be a general phenomenon,

which is not cell type specific.

In vitro cultured endothelial cells not only release MP upon activation with IL-1 , but

also upon activation with tumour necrosis factor (TNF)- [14,15,17], TNF- with

cycloheximide or camptothecin [18], or lipopolysaccharide [16]. The phenotype of the

released EMP may be dependent on the agonist used to activate the parent cells. For

instance, PMP expose the fibrinogen receptor GPIIb-IIIa ( IIb 3) when released in vitro

upon activation of platelets. This receptor is in its fibrinogen-binding conformation upon

platelet activation by thrombin plus collagen, but not upon activation by the complement

C5b-9 complex [28]. Whether the antigenic composition of EMP is agonist-dependent,

remains to be investigated. Because the stimuli involved in EMP release in vivo are

unknown, the antigenic phenotype of EMP in vitro versus in vivo may then differ as well.

In accordance with Combes et al [14], we also found that resting HUVEC

constitutively exposed antigens such as PECAM-1, and 3, and that EMP derived there

from exposed PECAM-1. Thus far, EMP have been identified ex vivo, i.e. in plasma

samples, using combinations of MoAbs directed against PECAM-1 plus , or PECAM-1

plus 3 [14,15,19]. In our present study, however, we found and 3 only exposed on

a minor subpopulation ( ) or absent ( 3) on EMP prepared in vitro. Our present findings

confirm earlier reports on the occurrence of EMP in vivo. However, earlier studies in

which PECAM-1 plus , or PECAM-1 plus 3 were used to detect EMP ex vivo, may

have overestimated their presence, since most of the MP ex vivo are of platelet origin

which expose high levels of PECAM-1 and 3.

In agreement with Combes et al [14], we also found no evidence for E-selectin-

exposing EMP in plasma samples of SLE patients and healthy individuals. There was one

exception, however. One patient, actually the one with the highest SLE disease activity

Chapter 2

47

index in our study, had a subpopulation of E-selectin-exposing MP, suggesting that in this

particular patient the endothelium may have been more activated than in the other

patients. However, this could not be confirmed because we measured the plasma

concentration of vWF and found it in this patient to be not significantly higher than in

other SLE patients (data not shown). We have no explanation yet for the low percentage

PMP in the plasma of this SLE patient when compared to other SLE patients. Our present

findings suggest that part of the soluble E-selectin, which is known to be elevated in

plasma of patients with SLE, is MP-associated. Whether this E-selectin originates from

the parent cell during MP formation or resembles originally soluble E-selectin

subsequently bound to the MP from other cells, is also open for discussion. Only some

12% of the E-selectin-positive MP exposed , which supports our in vitro data that this is

not a proper marker for EMP detection ex vivo. Since we found E-selectin exposing MP

in plasma from only one out of 11 SLE patients, we also analysed plasma samples from

severely Dengue virus infected patients. These patients are know to suffer from increased

vascular permeability [29] and sera from such patients contain antibodies that directly

trigger endothelial damage [30]. We found that plasma from two of these three patients

contained a subpopulation of 8% and 17% of E-selectin exposing MP. About 90% of this

subpopulation did not double stain for (data not shown). These data support our finding

that may not be a proper marker to detect EMP.

Like E-selectin, TF was strongly inducible on the HUVEC. The exposure of TF

peaked at 12 hours and diminished afterwards. Despite the fact that only a small

subpopulation of EMP exposing TF was observed, reconstitution of these EMP strongly

generated TF- and factor VII(a)-mediated thrombin generation in plasma (data not

shown). This indicates that functional TF must be exposed on these vesicles. Possibly, the

antigenic density, i.e. the number of exposed TF molecules, is too low to be detected by

flow cytometry. Of course, this may also hold true for all other surface markers in the

present study.

We also studied the exposure of GPIb (CD42b) on HUVEC (data not shown). Two

different MoAbs failed to detect exposure of this antigen. However, a third MoAb gave

conflicting results. Therefore, we are uncertain about the exposure of GPIb on HUVEC.

The present study shows that (i) EMP are released from HUVEC, (ii) the antigenic surface


48

composition of EMP differs from HUVEC, and (iii) the surface composition of EMP is

highly dependent on the activation status of the parent cells. EMP and PMP share various

important surface antigens, which implies that the measurement of EMP by flow

cytometry should be performed carefully, since most by far of the MP found in plasma

samples of patients and controls are of platelet origin. E-selectin can be used as a specific

marker to detect EMP ex vivo but may underestimate their presence because only a

subpopulation of the EMP stained positively for E-selectin and only upon activation of the

parent cell. Whether ICAM-1 can also be used as a specific marker for EMP detection ex

vivo remains to be established, since this adhesion receptor also occurs on lymphocytes

and monocytes.

Chapter 2

49

References

1. Bevilacqua MP, Pober JS, Mendrick DL, Cotran RS, Gimbrone MA, Jr. Identification of an inducible endothelial-leukocyte adhesion molecule. Proc. Natl. Acad. Sci. U. S. A 1987;84:9238-42.

2. Maynard JR, Dreyer BE, Stemerman MB, Pitlick FA. Tissue-factor coagulant activity of cultured human endothelial and smooth muscle cells and fibroblasts. Blood 1977;50:387-96.


4. Bevilacqua MP, Stengelin S, Gimbrone MA, Seed B. Endothelial leukocyte adhesion molecule 1: an inducible receptor for neutrophils related to complement regulatory proteins and lectins. Science 1989;243:1160-5.




8. Moake JL, Rudy CK, Troll JH, Weinstein MJ, Colannino NM, Azocar J, Seder RH, Hong SL, Deykin D. Unusually large plasma factor VIII:von Willebrand factor multimers in chronic relapsing thrombotic thrombocytopenic purpura. N. Engl. J. Med. 1982;307:1432-5.

9. Werns SW, Walton JA, Hsia HH, Nabel EG, Sanz ML, Pitt B. Evidence of endothelial dysfunction in angiographically normal coronary arteries of patients with coronary artery disease. Circulation 1989;79:287-91.

10. Zeiher AM, Drexler H, Wollschlager H, Just H. Endothelial dysfunction of the coronary microvasculature is associated with coronary blood flow regulation in patients with early atherosclerosis. Circulation 1991;84:1984-92.

11. Albornoz L, Alvarez D, Otaso JC, Gadano A, Salviu J, Gerona S, Sorroche P, Villamil A, Mastai R. Von Willebrand factor could be an index of endothelial


50

dysfunction in patients with cirrhosis: relationship to degree of liver failure and nitric oxide levels. J. Hepatol. 1999;30:451-5.

12. Belch JJ, Shaw JW, Kirk G, McLaren M, Robb R, Maple C, Morse P. The white blood cell adhesion molecule E-selectin predicts restenosis in patients with intermittent claudication undergoing percutaneous transluminal angioplasty. Circulation 1997;95:2027-31.

13. Borawski J, Naumnik B, Pawlak K, Mysliwiec M. Endothelial dysfunction marker von Willebrand factor antigen in haemodialysis patients: associations with pre-dialysis blood pressure and the acute phase response. Nephrol. Dial. Transplant. 2001;16:1442-7.




17. Sabatier F, Roux V, Anfosso F, Camoin L, Sampol J, Dignat-George F. Interaction of endothelial microparticles with monocytic cells in vitro induces tissue factor-dependent procoagulant activity. Blood 2002;99:3962-70.

18. Simak J, Holada K, Vostal JG. Release of annexin V-binding membrane microparticles from cultured human umbilical vein endothelial cells after treatment with camptothecin. BMC. Cell Biol. 2002;3:11-21.

19. Mallat Z, Benamer H, Hugel B, Benessiano J, Steg PG, Freyssinet JM, Tedgui A. Elevated levels of shed membrane microparticles with procoagulant potential in the peripheral circulating blood of patients with acute coronary syndromes. Circulation 2000;101:841-3.

20. Salzer U, Hinterdorfer P, Hunger U, Borken C, Prohaska R. Ca(++)-dependent vesicle release from erythrocytes involves stomatin- specific lipid rafts, synexin (annexin VII), and sorcin. Blood 2002;99:2569-77.

Chapter 2

51

21. Beekhuizen H, van Furth R. Growth characteristics of cultured human macrovascular venous and arterial and microvascular endothelial cells. J. Vasc. Res. 1994;31:230-9.

22. Beekhuizen H, van de Gevel JS, Olsson B, van Benten IJ, van Furth R. Infection of human vascular endothelial cells with Staphylococcus aureus induces hyperadhesiveness for human monocytes and granulocytes. J. Immunol. 1997;158:774-82.

23. Joop K, Berckmans RJ, Nieuwland R, Berkhout J, Romijn FPHTM, Hack CE, Sturk A. Microparticles from patients with multiple organ dysfunction syndrom and sepsis support coagulation through multiple mechanisms. Thromb. Haemost. 2001;85:810-20.

24. Berckmans RJ, Nieuwland R, Böing AN, Romijn FP, Hack CE, Sturk A. Cell-derived microparticles circulate in healthy humans and support low grade thrombin generation. Thromb. Haemost. 2001;85:639-46.

25. Nieuwland R, Berckmans RJ, Rotteveel-Eijkman RC, Maquelin KN, Roozendaal KJ, Jansen PGM, ten Have K, Eijsman L, Hack CE, Sturk A. Cell-derived microparticles generated in patients during cardiopulmonary bypass are highly procoagulant. Circulation 1997;96:3534-41.

26. Hochberg MC. Updating the American College of Rheumatology revised criteria for the classification of systemic lupus erythematosus. Arthritis Rheum. 1997;40:1725.

27. Bombardier C, Gladman DD, Urowitz MB, Caron D, Chang CH. Derivation of the SLEDAI. A disease activity index for lupus patients. The Committee on Prognosis Studies in SLE. Arthritis Rheum. 1992;35:630-40.

28. Sims PJ, Wiedmer T, Esmon CT, Weiss HJ, Shattil SJ. Assembly of the platelet prothrombinase complex is linked to vesiculation of the platelet plasma membrane. Studies in Scott syndrome: an isolated defect in platelet procoagulant activity. J. Biol. Chem. 1989;264:17049-57.

29. Halstead SB. Pathogenesis of dengue: challenges to molecular biology. Science 1988;239:476-81.

30. Lin CF, Lei HY, Shiau AL, Liu CC, Liu HS, Yeh TM, Chen SH, Lin YS. Antibodies from dengue patient sera cross-react with endothelial cells and induce damage. J. Med. Virol. 2003;69:82-90.

53

Chapter 3

Phospholipid composition of in vitro

endothelial microparticles and their in vivo

thrombogenic properties

Mohammed N. Abid Hussein, Anita N. Böing, Éva Biró, Frans J. Hoek, Gerard M.T.

Vogel, Dirk G. Meuleman, Augueste Sturk and Rienk Nieuwland

Thromb. Res. 2007; In press

Phospholipid composition and thrombogenic properties

54

ABSTRACT

Background: Microparticles from activated endothelial cells (EMP) are well known to

expose tissue factor (TF) and initiate coagulation in vitro. TF coagulant activity is

critically dependent on the presence of aminophospholipids, such as phosphatidylserine

(PS) and phosphatidylethanolamine (PE), but it is unknown whether or not TF-exposing

EMP are enriched in such aminophospholipids. Furthermore, despite the fact that EMP

have been reported in several pathological conditions, direct evidence for their (putative)

coagulant properties in vivo is still lacking.

Objectives: We investigated the phospholipid composition of endothelial MP (EMP) and

their thrombogenic properties in vivo.

Methods: Human umbilical vein endothelial cells (HUVEC; n=3) were incubated with or

without interleukin (IL)-1 (5 ng/mL; 0-72 hours). Phospholipid composition of EMP

was determined by high-performance thin layer chromatography. The association between

EMP, TF antigen and activity was confirmed in vitro (ELISA, Western blot and thrombin

generation). Thrombogenic activity of EMP in vivo was determined in a rat venous stasis

model.

Results: Levels of TF antigen increased 3-fold in culture medium of IL-1 -treated cells

(P<0.0001). This TF antigen was associated with EMP and appeared as a 45-47 kDa

protein on Western blot. In addition, EMP from activated cells were enriched in both PS

(P<0.0001) and PE (P<0.0001), and triggered TF-dependent thrombin formation in vitro

and thrombus formation in vivo. In contrast, EMP from control cells neither initiated

coagulation in vitro nor thrombus formation in vivo.

Conclusions: EMP from activated endothelial cells expose coagulant tissue factor and are

enriched in its cofactors PS and PE.

Chapter 3

55

INTRODUCTION

Tissue factor (TF), a 45-47 kDa transmembrane receptor, initiates coagulation [1],

triggers cell migration [2] and trafficking of mononuclear phagocytes across the

endothelium [3], regulates angiogenic properties of tumor cells [4], acts as a chemotactic

factor for vascular smooth muscle cells [5], and protects endothelial cells from apoptosis

[6,7]. TF is widely distributed within the body. Extravascular cell types constitutively

express TF [8,9], and cells at the blood interface (endothelial cells) or circulating within

the blood (monocytes) inducibly express TF [10-13].

TF can also be present on cell-derived microparticles (MP) in vivo. MP isolated from

pericardial wound blood [14], synovial fluid [15] or venous blood from a patient with

meningococcal septic shock complicated by fulminant disseminated intravascular

coagulation [16] initiate TF-dependent thrombin generation in vitro. In addition, we

demonstrated that MP from (pericardial) wound blood trigger TF-mediated thrombus

formation in vivo [17]. As yet, other MP have not been demonstrated to have such activity

in vivo.

Endothelial cell-derived MP (EMP) from TNF - or LPS-activated endothelial cells

expose procoagulant TF in vitro [18,19], but whether such EMP have any biological

activity in vivo is unknown. This question is becoming increasingly relevant since

elevated numbers of EMP are now known to occur in various pathological conditions,

including systemic lupus erythematosus [20], thrombotic thrombocytopenia purpura [21],

vasculitis of the young [22], paroxysmal nocturnal haemoglobinuria [23] and multiple

sclerosis [24]. EMP in healthy subjects were reported to correlate with the serum

triglyceride concentration, suggesting that EMP may reflect endothelial dysfunction or

injury [25].

Aminophospholipids like phosphatidylserine (PS) and phosphatidylethanolamine (PE)

are well established cofactors for the procoagulant activity of membrane-exposed TF [26-

28].

Recently, we showed that the phospholipid composition of platelet-derived MP

changes upon activation [29]. Whether or not the phospholipid composition of EMP

changes during activation of endothelial cells, however, is unknown.


56

The aims of the present study were to study the presumed procoagulant properties of

EMP in vivo and to determine whether phospholipid composition changes during

endothelial cell activation may support this TF activity.


Reagents and assays

Medium M199, penicillin, streptomycin, amphotericin B and L-glutamine were

obtained from GibcoBRL, Life Technologies (Paisley, Scotland). IgG1-FITC and IgG1-PE

(clone X40) were obtained from Becton Dickinson ((BD) San Jose, CA). Annexin V-

(allophycocyanin; APC) was from Caltag Laboratories (Burlingame, CA). Human serum

albumin (HSA) and monoclonal antibodies (MoAbs), directed against factor VIIa (VII-1

[1.46 mg/mL], VII-15 [0.53 mg/mL]) and anti-factor XII (OT-2 [0.71 mg/mL), were from

Sanquin (Amsterdam, The Netherlands). Anti-TF for western blotting (4503, clone TF9-

10H10, IgG1) and anti-TF for in vivo studies (4502, polyclonal IgG) were from American

Diagnostica Inc. (Greenwich, CT). Anti-mouse IgG-horseradish peroxidase (HRP)

conjugate was from Bio-Rad (Hercules, CA). Recombinant human interleukin-1 (IL-

1 ), human recombinant basic fibroblast growth factor and epidermal growth factor were

from GibcoBRL (Gaithersburg, MD). Collagenase (type 1A) was from Sigma (St. Louis,

MO), EDTA from Merck (Darmstadt, Germany), heparin (400 U/mL) from Bufa BV

(Uitgeest, The Netherlands), and trypsin from Difco Laboratories (Detroit, MI). Human

serum was provided by the Blood Bank Center of the Leiden University Medical Center

(Leiden, The Netherlands) and was heat inactivated during 30 minutes at 56 ºC (HuSi).

Tissue culture flasks were from Greiner Labortechnik (Frickenhausen, Germany) and

gelatin from Difco Laboratories (Sparks, MD). Reptilase was from Roche (Mannheim,

Germany) and the chromogenic substrate Pefachrome TH-5114 from Pentapharm Ltd.

(Basel, Switzerland). Heparinase (Hepzyme) was from Dade Behring GmbH (Marburg,

Germany). Human brain thromboplastin was a gift from Prof. Dr. R. Bertina (Department

of Haematology, Leiden University Medical Center, Leiden, The Netherlands).

Pentobarbital sodium (Nembutal) was obtained from Sanofi (Toulouse, France). L- -

lysophosphatidylcholine (L-PC; 38-0104), sphingomyelin (SM; 56-1080), L- -

Chapter 3

57

phosphatidylcholine (PC; 37-0106), L- -PS (37-0160), L- -phosphatidylinositol (PIn; 37-

0134) and L- -PE (37-0126) were from Larodan (Malmö, Sweden), L- -

lysophosphatidylethanolamine (L-PE; L4754) and cholesterol (C8667) from Sigma (St.

Louis, MO), and L- -lysophosphatidylserine (L-PS; 850092P) from Avanti Polar Lipids

Inc. (Alabaster, AL). Chloroform, ethylacetate, acetone, methanol, ethanol,

dichloromethane, isopropanol and acetic acid (all HPLC grade) were from Merck

(Darmstadt, Germany). All other chemicals were of analytical quality.

Isolation, culture and treatment of human umbilical vein endothelial cells (HUVEC)

HUVEC were collected from human umbilical cord veins and cultured as described

previously [20].

Isolation of EMP

At the indicated activation time intervals, culture supernatants were collected and

centrifuged (10 minutes at 180 g and 20 °C) to remove detached cells. Aliquots (250 L

each) of supernatants were frozen in liquid nitrogen and stored at – 80 °C. Samples were

thawed on melting ice for 1 hour and centrifuged for 30 minutes (17,570 g and 20 °C) to

pellet EMP. Then, 225 L supernatant was removed and the EMP-enriched pellet was

washed once with 225 L PBS/10.9 mmol/L trisodium citrate (pH 7.4). Finally, EMP

were resuspended in the remaining 25 L.

Flow cytometric analysis

EMP were analyzed in a FACSCalibur flow cytometer (BD). Forward scatter (FSC)

and side scatter (SSC) were set at logarithmic gain and EMP were identified and

quantified by their FSC and SSC characteristics and binding of annexin V as described

previously [20].

Western blotting

Culture supernatants (5 mL) were collected after 24 hours of incubation without or

with IL-1 . Detached cells were removed by centrifugation (10 minutes at 180 g and 20

°C). EMP were pelleted (1 hour at 17,570 g and 20 °C) and washed once in PBS/citrate.


58

The final pellet was resuspended in 24 L PBS, to which 6 L (5-fold concentrated)

sample buffer was added ( -mercaptoethanol (12.5% v/v), bromophenol blue (0.025%

v/v), glycerol (25% v/v), SDS (10% w/v) and Tris base (312.5 mM; pH 6.8)). Samples

were heated before electrophoresis (5 minutes, 100 °C). Proteins were separated on 10%

polyacrylamide gel and transferred to a nitrocellulose membrane (Schleicher & Schuell;

Dassel, Germany). Subsequently, blots were incubated (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); 60 minutes), (mouse) anti-human-TF (1 g/mL; 60 minutes)

and (goat) anti-mouse IgG-HRP conjugate (1:3000; 45 minutes). Between the incubation

steps, blots were washed three times with TBST for 5-10 minutes. All antibodies were

diluted with blocking buffer. The bands were detected using an enhanced

chemiluminescence kit (ECL; Amersham Biosciences; Buckinghamshire, UK) and

exposed to Fuji Medical X-ray film.

TF ELISA

TF in conditioned media was determined by ELISA (American Diagnostica Inc.;

Greenwich, CT).

Thrombin generation assay

The procoagulant properties of EMP in vitro were studied in a thrombin generation

test (TGT) as described previously [30]. In a control experiment, we found no effect of

freeze-thawing on the ability of microparticles to initiate thrombin generation (data not

shown). The ability to inhibit TF-initiated coagulation of the anti-human factor VII used

in this study is comparable to that of anti-human TF previously used in our thrombin

generation assay [16,30].

Rat venous stasis model

Rats were anesthetized and subsequently the abdomen was opened and the vena cava

inferior was isolated. All side branches distal to the left renal vein were obliterated.

Afterwards, rabbit thromboplastin suspension or saline (as positive and negative control,

Chapter 3

59

respectively) or EMP were injected into the dorsal penile vein. Thromboplastin

suspension was prepared by diluting Simplastin 50-fold (v/v) with saline. After injection,

blood was allowed to circulate freely for 10 seconds. Then the vena cava was ligated

beneath the left renal vein. After maintaining stasis for 10 minutes, the vena cava was

ligated near the fusion of the iliac veins, and then opened longitudinally. The formed

thrombus was removed and weighed [17,31]. Briefly, aliquots (250 L each) of (cell-free)

conditioned medium from both activated (IL-1 , 5 ng/mL, 48 hours) or resting HUVEC

were thawed on melting ice and incubated with heparinase to degrade heparin, an essential

cofactor of Fibroblast Growth Factor for endothelial cell culture. EMP were isolated and

washed in PBS/citrate by centrifugation (30 minutes at 17,570 g and 20 °C). Before

injection, EMP were resuspended in 75 L PBS/citrate buffer (pH 7.4) or 37.5 L

antibody plus 37.5 L PBS/citrate buffer. Antibodies used were polyclonal rabbit anti-

human TF and anti-human factor XII. Male Wistar Hsd/Cpb; WU rats (n=32, body weight

300-350 g) were obtained from Harlan (Horst, The Netherlands). All procedures were

approved by the Ethics Committee of Animal Welfare of Organon in accordance with

Dutch guidelines.

Phospholipid extraction and high-performance thin layer chromatography (hpTLC)

EMP were isolated from aliquots of cell-free culture supernatants (1 mL; n=3) as

reported earlier [32]. Lipids were extracted and phospholipids were separated and

quantified as described previously [29,33,34].


To determine whether activation of endothelial cells significantly affected the overall

numbers of EMP and TF antigen levels in conditioned medium in time, area under curves

were calculated and differences were post-analyzed using (two-tailed) paired t-test

(GraphPad Prism for Windows, release 3.02 (San Diego, CA)). In case of a significant

difference, data per time interval were further analyzed by two-tailed paired t-test. For

individual phospholipids, the overall differences in time (3-72 hours) between EMP from

unstimulated versus stimulated endothelial cells were determined by calculating the "area

under curve" represented by the data shown in Table 1, followed by Mann-Whitney test


60

(two-tailed; MedCalc). When a significant difference of the “area under curve” was found

to be present, also paired t-tests were performed to determine at which time points the

differences were significant. Data from in vitro thrombin generation were analyzed by

two-tailed paired t-test. Data on thrombus formation were analyzed using the Kruskal-

Wallis test followed by Dunn’s post test to correct for multiple comparisons. All

differences were considered statistically significant at P<0.05. Values are expressed as

mean ± SD.

RESULTS

EMP from IL-1 -treated endothelial cells expose TF and trigger cogulation in vitro

After 72 hours the numbers of EMP in conditioned medium from control

(unstimulated) cultures had increased gradually about 6-fold compared to (conditioned

medium from) 3 hour control cultures (Figure 1A). In contrast, upon activation with IL-

1 , the numbers of EMP increased already about 13-fold after 12 hours of culture

compared to the 3 hours time interval, and these numbers remained virtually constant up

to 72 hours of culturing. In IL-1 -treated cultures, the overall increase of EMP numbers in

time differed significantly compared to control (P=0.016). For individual time intervals, a

significant difference was observed at 24 hours (P=0.04), but not at 3 hours (P=0.503) or

72 hours (P=0.07). Also, EMP numbers at the 48 hours activation time interval were

comparable to 24 or 72 hours control conditions (P=0.174 and P=0.324; data not shown).

Concurrently, the overall concentrations of the TF antigen in conditioned media of IL-1 -

activated cells increased in time (Figure 1B; P=0.037). This increase was significant at 24

hours (P=0.0024), but not at 3 hours (P=0.931) or 72 hours (P=0.241). After removal of

EMP by high-speed centrifugation, the concentrations of TF (antigen) in the supernatants

were below the detection limit (12.5 ng/L), indicating that all non-endothelial cell-bound

TF is associated with EMP (data not shown). The EMP-associated TF appeared as a single

45-47 kDa protein band on Western blot (insert Figure 1B). Only EMP from activated

endothelial cells initiated thrombin formation in vitro (Figure 1C), and this was inhibited

by anti-human factor VII(a) (P=0.014) but not by anti-human factor XII(a) (Figure 1D;

P=0.896). Both the total capacity of the EMP to generate thrombin and the factor VII-TF

dependency remained unchanged between 12 and 72 hours of culture. Thus, as shown

Chapter 3

previously for endotoxin or TNF-treated endothelial cells, also IL-1 -treated endothelial

cells release TF-exposing EMP, which triggers (TF-dependent) thrombin generation in

vitro.

A

Culture time (hours)

Num

bero

f EM

P 10

3

3 12 24 48 720

4

8

12


TF a

ntig

en le

vel (

ng/L

)

B

3 12 24 48 720

100

200

300

47 kDa

- + TF

C

Thro

mbi

nco

ncen

tratio

n(n

mol

/L)

Time (minutes)0 5 10 15

0

50

100

150

200

250

300 D

0 12 24 36 48 60 72


0

1

2

3

4

5

Are

aun

derc

urve

A


Num

bero

f EM

P 10

3

3 12 24 48 720

4

8

12


TF a

ntig

en le

vel (

ng/L

)

B

3 12 24 48 720

100

200

300

47 kDa

- + TF

C

Thro

mbi

nco

ncen

tratio

n(n

mol

/L)

Time (minutes)0 5 10 15

0

50

100

150

200

250

300 D

0 12 24 36 48 60 72


0

1

2

3

4

5

Are

aun

derc

urve

Figure 1. EMP from IL-1 -activated endothelial cell expose TF and are coagulant in vitro. HUVEC were incubated with or without IL-1 (5 ng/mL; control samples were collected at 3 hours, 24 hours and 72 hours; n=3). At the indicated time intervals conditioned media from control ( ) and IL-1 -activated endothelial cells ( ) were collected and analyzed. A. Numbers of EMP identified by FSC, SSC and binding of annexin V. B. TF antigen in conditioned medium containing the EMP (upon removal of the EMP, the conditioned medium did not contain detectable quantities of TF, indicating all TF to be EMP-associated); the insert shows a representative Western blot of EMP lysates from unstimulated (-) and activated (+) endothelial cells; human brain thromboplastin (TF) was used as a positive control. C. EMP from unstimulated ( ) and IL-1 -activated endothelial cells ( ) were reconstituted in defibrinated, MP-free normal plasma to assess their thrombin generating capacity. Data from a representative thrombin generation experiment. D. Thrombin generation without ( ) or with anti-human factor VII ( ) or anti-human factor XII ( ). The ability of EMP to generate thrombin was expressed as the area under the curve during 15 minutes of thrombin generation (n=3). P 0.05 (EMP without antibody versus EMP incubated with anti-human factor VII).

61


Thrombus formation by EMP in vivo

In vivo, injection of Simplastin (positive control; a commercially available mixture of

TF and lipids from rabbit brain tissues (Organon Teknika Corp.; Durham, NC)) gave

thrombi of 61.8 mg 12.6 (n=4). Injection of saline (negative control) gave thrombi of

1.5 mg 1.9 (n=4; data not shown). Upon injection of EMP from activated endothelial

cells, thrombi were formed (Figure 2; 35.1 mg 12.9, P 0.01). Preincubation with anti-

human TF significantly blocked thrombus formation (5.6 mg 9.3, P 0.05). In contrast,

preincubation with anti-factor XII had less effect (26.9 mg 9.2, P>0.05) and was not

statistically significant. In line with our in vitro observation, no thrombus formation was

observed in rats that received EMP from unstimulated endothelial cells (0.5 mg 0.7).

These data show that only EMP from activated human endothelial cells are strongly

thrombogenic in vivo in a TF-dependent manner.

EMP

EMP+anti-T

F

EMP+anti-F

XII

EMP0

20

40

60

80

100 **

*

N.S.

EMP + IL-1 EMP - IL-1

Thro

mbu

s wei

ght (

mg)

EMP

EMP+anti-T

F

EMP+anti-F

XII

EMP0

20

40

60

80

100 **

*

N.S.

EMP + IL-1 EMP - IL-1

Thro

mbu

s wei

ght (

mg)

Figure 2. Thrombus formation by EMP in vivo. EMP from unstimulated and IL-1 -activated endothelial cells (48 hours) were injected into rats to assess their ability to trigger thrombus formation in vivo. EMP fractions from three different endothelial cell cultures were used. From each individual culture, EMP from unstimulated endothelial cells were injected into 2 rats (EMP - IL-1 ). From the corresponding EMP of activated cells, EMP not preincubated with antibodies (EMP + IL-1 ) were injected into 2 rats, EMP preincubated with anti-human TF (EMP+anti-TF) in 2 rats, and EMP preincubated with anti-human factor XII (EMP+anti-FXII) also in 2 rats. Thrombus weights for individual rats are indicated. N.S. (not significant; P 0.05); P 0.05; P 0.01.

62

Chapter 3

63

Phospholipid composition of EMP

The most prominent phospholipids in EMP from both unstimulated and stimulated

endothelial cells were PC and SM (Table 1). EMP from activated endothelial cells

contained significantly increased amounts of both PS (P<0.0001) and PE (both P<0.0001)

as compared to EMP from control cells. Upon activation of endothelial cells, the total

amount of phospholipids in isolated EMP fractions tended to increase, although this

increase did not reach significance (P=0.2). Similarly, the cholesterol:phospholipid ratio

of EMP was unchanged upon activation (P=0.4).

Table 1. Phospholipid composition of EMP from unstimulated (-) or activated (+)

endothelial cells.

Phospholipid IL-1 Culture time (hours)

3 12 24 48 72

- 8 1 8 1 11 4 10 13 2 L-PC

+ 8 1 6 2 9 3 7 9 3

- 17 1 16 2 23 2 19 18 1 SM

+ 17 1 14 1 21 3 17 17 1

- 57 2 54 6 45 3 57 51 4 PC

+ 56 1 51 1 40 2 45 42 2

- 4 3 7 1 4 1 2 6 1 PS

+ 3 1 10 3 8 3 6 11 1

- 5 2 4 1 8 1 3 4 1PIn

+ 7 0 4 0 9 1 3 3 0

- 9 2 11 6 10 3 7 7 1 PE

+ 11 0 14 0 14 3 16 16 1

Data are expressed as % of total phospholipid (mol/mol). *P<0.0001 (area under curve), #P<0.05 (individual time points). Activation with IL-1 did not affect the relative amounts of L-PC (P=0.400), SM (P=0.100), PC (P=0.100) or PIn (P=0.700). Data are shown as mean SD (n=3-5) except for the 48 hours time interval, since EMP from this collection point had been arbitrarily chosen to be used in the rat venous stasis model and therefore insufficient material was available for further analysis.


64

DISCUSSION

Previous studies demonstrated that EMP from TNF- or LPS-treated endothelial cells

expose TF and trigger thrombin generation in vitro [18,19]. Similarly, our present data

show that also EMP from IL-1 -activated endothelial cells expose TF and trigger

thrombin generation in vitro. More interestingly, however, is that such EMP become

enriched in both PS and PE, and trigger thrombus formation in vivo by a TF-initiated

pathway.

In this study we present data that TF exposed by EMP from activated endothelial cells

is responsible for the coagulant activity in vitro and in vivo. This is based upon control

studies with EMP from non-activated HUVEC and from inhibitory studies with antibodies

against the extrinsic pathway. It could be argued that the absence of a coagulant effect of

EMP from the control situation is due to the fact that 2 to 3-fold lower numbers of EMP

are present in the culture medium, i.e. a lower availability of procoagulant phospholipids.

In our experiments we did not correct for that difference by taking larger volumes of

medium, because the EMP numbers vary somewhat between experiments. However, with

the EMP from the activated HUVEC, i.e. EMP exposing TF, inhibition of the extrinsic

coagulation pathway completely abolished their ability to initiate coagulation at the same

EMP concentration. Evidently, the exposure of procoagulant phospholipids is insufficient

to trigger coagulation, although it may promote the TF-associated coagulant activity and

facilitate the binding of coagulation factors.

Jimenez et al. studied the numbers and antigenic phenotype of EMP from

microvascular- and macrovascular endothelial cells after activation (TNF- ) or induction

of apoptosis (serum deprivation) [35]. They showed that EMP from microvascular- and

macrovascular endothelial cells differed in antigenic composition. Moreover, they showed

that the antigenic composition of EMP from both microvascular as well as macrovascular

EMP was differentially affected upon activation or induction of apoptosis. For instance,

whereas the numbers of annexin V-binding EMP, i.e. EMP exposing PS on their surface,

from microvascular endothelial cells was increased during growth factor deprivation

compared to EMP from activated (microvascular) endothelial cells, culture supernatants

from macrovascular endothelial cells hardly contained any annexin V-binding EMP, i.e.

not even when these cells had been subjected to growth factor deprivation resulting in

Chapter 3

65

apoptosis. In the present study, we used a different kind of endothelial cell (HUVEC), we

used a different inducer to activate (IL-1 ), and determined the antigenic composition of

EMP after freeze-thawing. Therefore, the antigenic composition of EMP in these two

studies, including the binding of annexin V, can not be directly compared. As for the PS

exposure, we used EMP after snap freezing in liquid nitrogen, storage at – 20 °C and

subsequent thawing, which increases exposure of PS on the EMP. Thus, the EMP used in

our present study can promote the coagulation process by enabling the formation of

prothrombinase- and tenase complexes on their surface, but the presence of TF is

necessary to initiate the coagulation cascade.

Recently, del Conde et al. showed that monocyte-derived MP may fuse with activated

platelets, thereby transferring their TF [36]. It was suggested that MP predestined for

fusion are likely to be enriched in fusion-promoting phospholipids like PS. Our present

data confirm their hypothesis for EMP. Thus, differences in phospholipid composition (of

MP) may not only affect their procoagulant properties but also their ability to deliver TF

to target cells. The changes in phospholipid composition are likely to be cell-type and/or

agonist dependent. Previously, we showed that upon platelet activation, the PS content of

platelet-derived MP (PMP) was unaffected, whereas their cholesterol and sphingomyelin

content increased [29].

Disseminated intravascular coagulation is a frequent complication of endotoxic shock.

Drake et al. demonstrated systemic fibrin deposition in a lethal Escherichia coli sepsis

baboon model. They failed, however, to demonstrate a significant occurrence of TF on

endothelial cells [37]. They concluded, that "compared with endothelial cells in culture,

there is in vivo significantly greater control of TF expression than expected". Our present

data suggest that the absence of TF on endothelial cells can be explained by the release of

TF-exposing EMP from these cells into the circulation. This explains on the one hand the

systemic fibrin deposition and on the other hand the unexpected absence of TF on the

endothelium in vivo.

Taken together, the present findings demonstrate that TF-exposing EMP are enriched

in aminophospholipids, and that such EMP are highly thrombogenic in vivo.


66

References

1. Nemerson Y. Tissue factor and hemostasis. Blood 1988;71:1-8.

2. Ruf W, Mueller BM. Tissue factor signaling. Thromb. Haemost. 1999;82:175-82.

3. Randolph GJ, Luther T, Albrecht S, Magdolen V, Muller WA. Role of tissue factor in adhesion of mononuclear phagocytes to and trafficking through endothelium in vitro. Blood 1998;92:4167-77.

4. Zhang Y, Deng Y, Luther T, Muller M, Ziegler R, Waldherr R, Stern DM, Nawroth PP. Tissue factor controls the balance of angiogenic and antiangiogenic properties of tumor cells in mice. J. Clin. Invest. 1994;94:1320-7.

5. Sato Y, Asada Y, Marutsuka K, Hatakeyama K, Sumiyoshi A. Tissue factor induces migration of cultured aortic smooth muscle cells. Thromb. Haemost. 1996;75:389-92.

6. Sorensen BB, Rao LV, Tornehave D, Gammeltoft S, Petersen LC. Antiapoptotic effect of coagulation factor VIIa. Blood 2003;102:1708-15.

7. Versteeg HH, Spek CA, Richel DJ, Peppelenbosch MP. Coagulation factors VIIa and Xa inhibit apoptosis and anoikis. Oncogene 2004;23:410-7.

8. Tedgui A, Mallat Z. Smooth muscle cells: another source of tissue factor-containing microparticles in atherothrombosis? Circ. Res. 2000;87:81-2.

9. Schecter AD, Giesen PL, Taby O, Rosenfield CL, Rossikhina M, Fyfe BS, Kohtz DS, Fallon JT, Nemerson Y, Taubman MB. Tissue factor expression in human arterial smooth muscle cells. TF is present in three cellular pools after growth factor stimulation. J. Clin. Invest. 1997;100:2276-85.


11. Drake TA, Ruf W, Morrissey JH, Edgington TS. Functional tissue factor is entirely cell surface expressed on lipopolysaccharide-stimulated human blood monocytes and a constitutively tissue factor-producing neoplastic cell line. J. Cell Biol. 1989;109:389-95.

12. Siddiqui FA, Desai H, Amirkhosravi A, Amaya M, Francis JL. The presence and release of tissue factor from human platelets. Platelets 2002;13:247-53.

Chapter 3

67

13. Maynard JR, Dreyer BE, Stemerman MB, Pitlick FA. Tissue-factor coagulant activity of cultured human endothelial and smooth muscle cells and fibroblasts. Blood 1977;50:387-96.

14. Nieuwland R, Berckmans RJ, Rotteveel-Eijkman RC, Maquelin KN, Roozendaal KJ, Jansen PGM, ten Have K, Eijsman L, Hack CE, Sturk A. Cell-derived microparticles generated in patients during cardiopulmonary bypass are highly procoagulant. Circulation 1997;96:3534-41.

15. Berckmans RJ, Nieuwland R, Tak PP, Böing AN, Romijn FP, Kraan MC, Breedveld FC, Hack CE, Sturk A. Cell-derived microparticles in synovial fluid from inflamed arthritic joints support coagulation exclusively via a factor VII-dependent mechanism. Arthritis Rheum. 2002;46:2857-66.


17. Biró É, Sturk-Maquelin KN, Vogel GM, Meuleman DG, Smit MJ, Hack CE, Sturk A, Nieuwland R. Human cell-derived microparticles promote thrombus formation in vivo in a tissue factor-dependent manner. J. Thromb. Haemost. 2003;1:2561-8.







68



25. Ferreira AC, Peter AA, Mendez AJ, Jimenez JJ, Mauro LM, Chirinos JA, Ghany R, Virani S, Garcia S, Horstman LL, Purow J, Jy W, Ahn YS, de ME. Postprandial hypertriglyceridemia increases circulating levels of endothelial cell microparticles. Circulation 2004;110:3599-603.

26. Smeets EF, Comfurius P, Bevers EM, Zwaal RF. Contribution of different phospholipid classes to the prothrombin converting capacity of sonicated lipid vesicles. Thromb. Res. 1996;81:419-26.

27. Neuenschwander PF, Bianco-Fisher E, Rezaie AR, Morrissey JH. Phosphatidylethanolamine augments factor VIIa-tissue factor activity: enhancement of sensitivity to phosphatidylserine. Biochemistry 1995;34:13988-93.

28. Smith SA, Morrissey JH. Properties of recombinant human thromboplastin that determine the International Sensitivity Index (ISI). J. Thromb. Haemost. 2004;2:1610-6.

29. Biró É, Akkerman JW, Hoek FJ, Gorter G, Pronk LM, Sturk A, Nieuwland R. The phospholipid composition and cholesterol content of platelet-derived microparticles: a comparison with platelet membrane fractions. J. Thromb. Haemost. 2005;3:2754-63.

30. Berckmans RJ, Nieuwland R, Böing AN, Romijn FP, Hack CE, Sturk A. Cell-derived microparticles circulate in healthy humans and support low grade thrombin generation. Thromb. Haemost. 2001;85:639-46.

31. Vogel GM, Meuleman DG, Bourgondien FG, Hobbelen PM. Comparison of two experimental thrombosis models in rats effects of four glycosaminoglycans. Thromb. Res. 1989;54:399-410.

32. Abid Hussein MN, Nieuwland R, Hau CM, Evers LM, Meesters EW, Sturk A. Cell-derived microparticles contain caspase 3 in vitro and in vivo. J. Thromb. Haemost. 2005;3:888-96.

33. Bligh EG, Dyer WJ. A rapid method of total lipid extraction and purification. Can. J. Biochem. Physiol 1959;37:911-7.

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69

34. Weerheim AM, Kolb AM, Sturk A, Nieuwland R. Phospholipid composition of cell-derived microparticles determined by one-dimensional high-performance thin-layer chromatography. Anal. Biochem. 2002;302:191-8.

35. Jimenez JJ, Jy W, Mauro LM, Soderland C, Horstman LL, Ahn YS. Endothelial cells release phenotypically and quantitatively distinct microparticles in activation and apoptosis. Thromb. Res. 2003;109:175-80.

36. 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;106:1604-11.

37. Drake TA, Cheng J, Chang A, Taylor FB, Jr. Expression of tissue factor, thrombomodulin, and E-selectin in baboons with lethal Escherichia coli sepsis. Am. J. Pathol. 1993;142:1458-70.

71

Chapter 4

Cell-derived microparticles contain caspase 3

in vitro and in vivo

Mohammed N. Abid Hussein, Rienk Nieuwland, Chi M. Hau, Ludo M. Evers, Eelco W.

Meesters and Augueste Sturk

J. Thromb. Haemost. 2004;3:888-896


72

ABSTRACT

Background: Microparticles (MP) from endothelial cells (endothelial microparticles;

EMP) circulate in disease states, but the processes such as apoptosis or cell activation

underlying their release are unclear.

Objective: We investigated whether adherent (viable) or detached (apoptotic) endothelial

cells are the possible source of EMP in vitro, i.e. under control and interleukin (IL)-1

activation conditions, and in vivo.

Methods: Adherent and detached endothelial cells, and EMP, were isolated from human

umbilical vein endothelial cell cultures (n=6), treated without or with IL-1 (5 ng/mL; 24

hour). Cell fractions were analyzed by flow cytometry for annexin V binding, propidium

iodide and caspase 3 staining (n=3). Caspase 3 in EMP was studied using Western blot

(n=6) and flow cytometry (n=6). Plasma from healthy subjects and SLE patients (both

n=3) were analyzed for caspase 3-containing (E)MP.

Results: Detached but not adherent cells double-stained for annexin V and propidium

iodide, confirming the apoptotic conditions of the detached cells and the viable nature of

the adherent cells. Caspase 3 was solely present in the detached cells and procaspase 3 in

the adherent cells. Caspase 3 was present in EMP from both control and IL-1 -treated

cultures. Counts of EMP and detached cells, but not adherent cells, highly correlated

(r=0.959, P<0.0001). In vivo circulating MP from nucleated (endothelial cells,

monocytes) and anucleated cells (platelets, erythrocytes) contained caspase 3.

Conclusions: EMP contain caspase 3 and may be mainly derived from detached

(apoptotic) endothelial cells in vitro. The presence of caspase 3 in MP from anucleated

cell types, however, suggests that its presence may not necessarily be related to apoptosis

in vivo but may be associated with caspase 3 activation unrelated to apotosis.

Chapter 4

73

INTRODUCTION

The physiological status of the endothelium is one of the important factors in

maintaining a proper hemostasis. Dysfunction of endothelial cells may result from

excessive activation or programmed cell death (apoptosis) induced by a host of factors. In

various pathologies, dysfunction of the endothelial cells may play a role, such as

hypertension [1], cardiovascular diseases [2], diabetes [3], systemic lupus erythematosus

(SLE) [4], renal failure [5] and vasculitis [6], and it is thought to play a role in the

development of coronary atherosclerosis [7]. Thus, monitoring of the endothelial cells

status is clinically pivotal, but so far only soluble E-selectin and von Willebrand factor are

thought to be specific biochemical markers that reflect the activation/dysfunction status of

the endothelial cells [8-10].

Recently, the presence of microparticles (MP) from endothelial cells (EMP) has been

reported in plasma from patients suffering from, for example, lupus anticoagulant [11],

multiple sclerosis [12], preeclampsia [13], acute coronary syndromes [14], thrombotic

thrombocytopenic purpura [15], paroxysmal nocturnal hemoglobinuria [16], severe

systemic inflammatory response syndrome [17] and severe malaria complicated with

coma [18]. Recently, we showed that E-selectin identifies approximately 50% of EMP

from activated endothelial cells in vitro and that such vesicles do also occur in vivo [19].

Others also demonstrated the occurrence of such E-selectin-positive EMP in plasma from

patients suffering from active systemic vasculitis [20]. The presence of EMP in the

circulation as a marker of endothelial dysfunction has very recently been reviewed by

Horstman et al [21].

In vitro, small fractions of endothelial cells appear as ‘floaters’ that display typical

biochemical and morphological features of apoptosis [22-25]. This is a well-known

phenomenon, called anoikis (Greek for ‘homelessness’), i.e. the induction of apoptosis

when a cell loses contact with the underlying matrix [26]. As a matter of fact, resistence to

apoptosis in spite of the loss of this matrix contact may be involved in successful

metastasis of cancer cells [27]. Circulating, i.e. detached endothelial cells have also been

reported in various disease states [28-31]. Whether or not there is indeed a relationship

between the detached, apoptotic endothelial cells and EMP, however, is unknown.

Therefore, in the present study we collected and analyzed adherent and detached


74

endothelial cells separately in order to investigate their possible relationship to EMP

formation.


Reagents and assays

Medium M199, penicillin, streptomycin, and L-glutamine were obtained from

GibcoBRL, Life Technologies (Paisley, Scotland). Immunoglobulin (Ig)G1-fluroscein

isothiocyanate (FITC), IgG1-phycoerythrin (PE) (clone X40), CD14-PE (clone M P98,9,

IgG2b), CD61-PE (clone VI-PL2, IgG1) and CD31-PE (clone WM-59, IgG1) were

obtained from Becton Dickinson ((BD) San Jose, CA, USA). In some experiments, CD61-

PE (clone Y2/51, IgG1) from Miltenyi Biotec (Bergisch Gladbach, Germany) was used.

No differences were observed between these two monoclonal antibodies (MoAbs). Anti-

Glycophorin A-PE (clone JC159, IgG1) was from Dako A/S (Glostrup, Denmark). Human

serum and fetal calf serum (both heat inactivated during 30 minutes at 56 ºC; HuSi and

FCSi, respectively) were from BioWhittaker (Walkersville, MD, USA). Human serum

albumin (HSA) was obtained from the Central Laboratory of the Netherlands Red Cross

Bloodtransfusion Service (CLB; Amsterdam, The Netherlands), CD54-PE (clone 84H10,

IgG1) from Immunotech (Marseille, France), and CD62E-PE (clone HAE-1f, IgG1) from

Ancell (Lausen, Switzerland). Recombinant human interleukin-1 (IL-1 ) was either

from GibcoBRL (Gaithersburg, MD, USA) or Sigma (St. Louis, MO, USA). Human

recombinant basic fibroblast growth factor (bFGF), and epidermal growth factor (EGF)

were derived from Invitrogen life technologies (Carlsbad, CA, USA). Collagenase (type

1A), and propidium iodide were from Sigma (St. Louis, MO, USA). Annexin V-

allophycocyanin (APC) was from Caltag Laboratories (Burlingame, CA), heparin (400

U/mL) from Leo Pharma BV (Breda, The Netherlands) and trypsin from Difco

Laboratories (Detroit, MI, USA). The following antibodies were used for Western blot

analysis: anti-human procaspase 3 MoAb from Transduction Laboratories (Lexington,

KY, USA), anti-human caspase 3 polyclonal antibody from Cell Signaling Technology

(Beverly, MA, USA), anti-human -tubulin MoAb and anti-mouse HRP conjugate from

Bio-Rad (Hercules, CA, USA), and anti-rabbit IgG HRP conjugate from Promega

Chapter 4

75

(Madison, WI, USA). Tissue culture flasks were from Greiner Labortechnik

(Frickenhausen, Germany) and gelatin from Difco Laboratories (Sparks, MD, USA).

Isolation, culture and treatment of human umbilical vein endothelial cells

Human umbilical vein endothelial cells (HUVEC) were collected from human

umbilical cord veins as described previously [19]. Briefly, umbilical cords were digested

with collagenase for 20 minutes at 37 °C. Detached cells were perfused with medium

M199 supplemented with 10% HuSi. The cell suspension was centrifuged for 10 minutes

at 180 g and 20 °C, and cells were resuspended in culture medium. HUVEC were cultured

in tissue culture flasks coated with 0.75% gelatin (passage 0). Upon confluency at passage

3, HUVEC were kept for 3-4 days in a resting state. Afterward, the culture supernatant

was refreshed and the cells were either untreated (control) or incubated with IL-1 (5

ng/mL) for 24 hours. Conditioned media (10 mL) were harvested and centrifuged for 10

minutes at 180 g and 20 °C to isolate detached cells. The pellets containing the detached

cells were carefully resuspended in 1% FCSi in PBS (pH 7.4). In parallel, the adherent

endothelial cells were harvested by trypsinization. After 4 minutes, trypsin was

neutralized by PBS/FCSi. Both cell suspensions were separately centrifuged for 10

minutes at 180 g and 4 °C, resuspended in PBS/FCSi, kept on melting ice for 15 minutes,

and then again centrifuged for 10 minutes at 180 g and 4 °C. Because adherent cells

outnumbered detached cells in all experiments, detached cells were resuspended in 0.5 mL

PBS/FCSi and the adherent cells in 1 mL PBS/FCSi. The absolute number of cells (N)

was estimated using the following formula: N = (events counted by flow cytometry /

aspiration volume) x 60 (dilution factor) x cell suspension volume.

Labeling of endothelial cells with annexin V and propidium iodide (PI)

Upon incubation without or with IL-1 , detached and adherent endothelial cells were

collected separately in PBS/FCSi as described in the previous section. The cell

suspensions were centrifuged for 10 minutes at 180 g and 20 °C and pellets were

resuspended in ice-cold binding buffer (10 mmol/L HEPES, 150 mmol/L NaCl, 8 mmol/L

KCl, 1.4 mmol/L CaCl2, 1.0 mmol/L MgCl2; pH 7.4). The cells were washed twice with

the ice-cold buffer. The pellets were then carefully resuspended in the buffer (100 L).


76

Annexin V-FITC (diluted 1:200 v/v) was added and the mixture was incubated in the dark

for at least 10 minutes at 4 °C. The excess of unbound annexin V was removed by

addition of 1 mL binding buffer and centrifugation of the mixture for 10 minutes at 180 g

and 20 °C. The pellets were resuspended in 300 L binding buffer and 5 L PI (5 g/mL

final concentration) was added to each sample immediately prior to flow cytometric

measurement. The fluorescence thresholds were set in terms of binding of annexin V and

PI to adherent cells harvested from untreated (control) HUVEC. In one preliminary

experiment, 5,000 events were analyzed by flow cytometry for the adherent cells, and

detached cells were analyzed for 1 minute. In two other experiments, 1,500 events were

analyzed for both detached and adherent cells.

Flow cytometric analysis of caspase 3 in endothelial cells

Both detached and adherent endothelial cells were analyzed by flow cytometry for the

presence of caspase 3 using the apoptosis kit I (BD). Detached and adherent cells were

collected, washed twice with cold PBS, resuspended in fixation and permeabilization

solution and subsequently incubated for 20 minutes on ice in this solution. The cells were

then pelleted (10 minutes at 180 g and 20 °C) and washed twice with detergent buffer

[perm/wash; 1:10 (v/v)]. The pellet was resuspended with the detergent buffer (100 L)

and incubated with either control antibody or rabbit anti-caspase 3 MoAb (5 L) for 30

minutes at room temperature. Afterwards, the cells were washed with detergent buffer (1

mL) and finally resuspended in 500 L detergent buffer before analysis. Detached cells

were analyzed for 1 minute and for adherent cells 1,500 events were analyzed.

Isolation of (E)MP

Aliquots (1 mL) of the cell-free culture supernatant were snap-frozen in liquid

nitrogen and stored at – 80 °C. Before use, samples were thawed on melting ice for 1

hour, then centrifuged for 1 hour at 17,570 g and 20 °C. Then, 900 L of (MP-free)

supernatant was removed. The remaining 100 L (MP-enriched) suspension was diluted

with 900 L PBS (154 mmol/L NaCl, 1.4 mmol/L phosphate) containing 10.9 mmol/L

trisodium citrate. MP were resuspended and again centrifuged for 1 hour at 17,570 g and

20 °C. Again, 900 L of supernatant was removed and MP were resuspended in the

Chapter 4

77

remaining 100 L. Plasma samples from citrate-anticoagulated venous blood of SLE

patients and healthy controls (with their informed consent) were collected and handled as

described previously [19]. For flow cytometry detection of (E)MP from SLE patients and

healthy individuals, the MP suspension was diluted 4-fold with PBS/citrate (pH 7.4).

Flow cytometric analysis of (E)MP

MP samples were analyzed in a FACSCalibur flow cytometer (BD; San Jose, CA,

USA). Forward scatter (FSC) and side scatter (SSC) were set at logarithmic gain and MP

were identified as described previously by their FSC and SSC characteristics and binding

of annexin V [19]. MP (5 L) were diluted with 35 L PBS containing 2.5 mmol/L CaCl2

(pH 7.4). Then, 5 L annexin V-APC was added (0.66 or 0.5 g/mL final concentration;

two different batches of APC-labeled annexin V were used, and both batches were titrated

for optimal staining). In the control samples of the MP, annexin V-positive events were

identified by placing a threshold in a MP sample (5 L) diluted with PBS containing 10.9

mmol/L trisodium citrate (40 L; pH 7.4) and 5 L of annexin V, i.e. without Ca2+. The

mixture of MP and annexin V was then incubated for 15 minutes in the dark at room

temperature. To remove the excess of free annexin V, 200 L PBS/calcium buffer was

added and the suspension was centrifuged for 30 minutes at 17,570 g and 20 °C. Finally,

200 L of supernatant was removed, and MP were resuspended with 300 L

PBS/calcium. All samples were analyzed for 1 minute in the flow cytometer.

Western blotting

Cell-free culture supernatants (5 mL) were collected after 24 hours. After removal of

detached cells, EMP were isolated by centrifugation for 1 hour at 17,570 g and 20 °C and

resuspended in PBS/citrate. Subsequently, EMP were pelleted (1 hour at 17,570 g and 20

°C) and resuspended in 24 L PBS. To this EMP suspension, 6 L of 5-fold concentrated

sample buffer containing -mercaptoethanol (12.5% v/v), bromophenol blue (0.025%

v/v), glycerol (25% v/v), SDS (10% w/v) and Tris base (312.5 mM; pH 6.8), was added.

Detached and adherent endothelial cells were separately isolated, washed and collected in

PBS/FCSi (0.5 and 1.0 mL, respectively). In two experiments, 330 L of detached cells

suspension and 830 L of adherent cells suspension were used to pellet the cells. In the


78

other four experiments the amount of detached and adherent cells suspensions used to

pellet the cells were 290 L and 790 L, respectively. Subsequently, sample buffer was

used to dissolve the pellets of the detached cells (final volume 30 L) and adherent cells

(final volume 60 L). Fixed volumes (30 L) of cell lysates in sample buffer were applied

for electrophoresis. Before electrophoresis, all samples were heated for 5 minutes at 100

°C. Electrophoresis was carried out in 15% polyacrylamide gel. The proteins were

transferred to nitrocellulose membrane (Schleicher & Schuell; Dassel, Germany). Blots

were incubated for 60 minutes 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 subsequently incubated with (rabbit) anti-human caspase 3 polyclonal antibody

(1:1,000) for 24 hours at 4 °C, followed by anti-rabbit IgG-HRP conjugate (1:7,500) for at

least 45 minutes at room temperature. In addition, the same blots were incubated with

(mouse) anti-human procaspase 3 MoAb (1:1,000) for 2 hours at room temperature and

(goat) anti-mouse IgG-HRP conjugate (1:3,000) for 60 minutes at room temperature.

Finally, the blots were also incubated with anti- -tubulin MoAb for 60 minutes at room

temperature and (goat) anti-mouse IgG-HRP conjugate (1:3,000) for 60 minutes at room

temperature. After each incubation step, the blots were washed three times with TBST for

5-10 minutes. All antibodies were diluted with blocking buffer. The bands were detected

using an enhanced chemiluminescence kit (ECL; Amersham Biosciences;

Buckinghamshire, UK) and exposed to Fuji Medical X-ray film.

Flow cytometric analysis of caspase 3 in (E)MP

The presence of caspase 3 in subpopulations of MP was studied by flow cytometry

using apoptosis kit I (BD) with slight modification of the manufacturer’s protocol. Briefly,

EMP were isolated as described in the previous section. The EMP suspension (25 L) was

diluted with 500 L detergent (diluted 1:10 in distilled water prior to use) and centrifuged

for 30 minutes at 17,570 g and 20 °C. Then, 500 L of (MP-free) supernatant was

removed and EMP were resuspended with 75 L PBS/citrate (pH 7.4). EMP (5 L) were

diluted with 35 L detergent. Rabbit anti-caspase 3 antibody plus CD31-PE (PECAM-1),

CD54-PE (ICAM-1) or CD62E-PE (E-selectin) (5 L each) were added. Isotype-matched

Chapter 4

79

control antibodies (IgG1) were also used to set fluorescence thresholds. The mixture was

then incubated for 30 minutes in the dark at room temperature. To remove the excess of

unbound MoAb, 500 L detergent (1:1,000 diluted in distilled water) was added and the

suspension was centrifuged for 30 minutes at 17,570 g and 20 °C. Finally, 500 L of

supernatant was removed, and MP were resuspended with additional 300 L detergent

(1:1,000). For flow cytometric analysis of MP from SLE patients and healthy controls, the

MoAb concentrations used were 0.16 g/mL for Glycophorin A-PE and 0.25 g/mL for

CD14-PE (LPS-receptor). Dilution of CD61-PE ( 3) was 1:100 (v/v).

Patients and controls

SLE patients (n=3; women) were included in this study who fulfilled the revised

criteria of the American College of Rheumatology for the diagnosis of SLE [32]. Their

age was 45, 51 and 55 years. The SLE Disease Activity Index [33] was 22, 20 and 4,

respectively. As controls, three age-matched women were included. The study fulfilled the

guidelines of the Medical Ethical Committee of the Slotervaart Hospital (Amsterdam, The

Netherlands).


Data were analyzed with Prism (3.02) for Windows. All data were analyzed with

paired t-test (two-tailed analysis). Data were considered statistically significant at P 0.05.

Correlations were determined using Pearson’s correlation test (two-tailed analysis).


80

RESULTS

Numbers of EMP and endothelial cells

EMP were isolated from control or IL-1 -treated endothelial cells (n=6) and

identified by their characteristics FSC and SSC, and their binding of annexin V. Figure 1A

shows that treatment with IL-1 resulted in a double but statistically insignificant

increase in EMP numbers (P=0.118). It should be mentioned that in five out of the six

experiments the numbers of EMP were elevated upon IL-1 -treatment compared with

control.

In control cultures, 6% 3.5 (mean SD) of the total endothelial cell number

occurred as detached cells (Figure 1B). These percentages were calculated by setting the

total number of adherent plus detached cells per experiment at 100%. Upon treatment with

IL-1 , this fraction increased to 23.1% 23.7 (P=0.162 compared to control). Again,

similar to EMP, in five out of the six experiments the percentages of detached cells were

increased upon IL-1 treatment. In one of the experiments, however, the number of

detached cells was exceptionally high (65.2%) compared to control (3.2%), and this

number deviated more than 3-fold from the average. When this number was omitted from

statistical analysis, there was still no significant difference (P=0.282). The overall

correlations for control (n=6) plus IL-1 -treatment (n=6) between the numbers of EMP

and detached as well as adherent cells are provided in Figure 1C and D, respectively. The

numbers of EMP correlated highly with the numbers of detached cells (Figure 1C;

r=0.959, P<0.0001), but not with adherent cells (Figure 1D; r=0.087, P=0.787). Again, in

one sample an exceptionally high number of EMP was observed. When this sample was

omitted, correlations were r=0.825 (P=0.001) and r=0.104 (P=0.760), respectively. This

sample was different from the one in which we observed an exceptionally elevated

number of detached cells.

Chapter 4 N

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Figure 1. Analysis of EMP and endothelial cell numbers. Endothelial cells were

incubated without (control) or with IL-1 (5 ng/mL) for 24 hours. Adherent and detached

cells as well as EMP were isolated as described (n=6). (A) Numbers of EMP, identified

on their characteristic FSC and SSC, and binding of annexin V (P 0.05). (B) Numbers of

adherent cells (gray bars; P 0.05) or detached cells (black bars; P 0.05). The

correlation between the numbers of EMP and the numbers of detached cells is shown in C

(r=0.959, P 0.0001). The correlation between the numbers of EMP and the numbers of

adherent cells is shown in D (r=0.087, P=0.787). For panels C and D, data from control

and IL-1 -treated cultures are included.

Annexin V and PI staining of endothelial cells

Adherent and detached endothelial cells were isolated from control (Figure 2A, B)

and from IL-1 (Figure 2C, D) treated cells (5 ng/mL; 24 hours), and analyzed for

binding of annexin V in combination with PI staining. Cells that stain for both annexin V

and PI (i.e. events in the upper right quadrant of Figure 2A-D) are generally considered to

be in the late stage of apoptosis [34]. Representative dot plots are shown. Most of the

81


detached endothelial cells (Figure 2B, D) double-stained for annexin V and PI (control

versus IL-1 : 68.6% 19.1 and 72.8% 8.9, respectively; P=0.621). In contrast, only a

minor fraction of the adherent cells (Figure 2A and C; upper right quadrant) were positive

for annexin V and PI (control versus IL-1 : 1.0% 0.0 and 1.6% 1.2, respectively;

P=0.422). Thus, adherent cells remained viable after treatment with IL-1 . Therefore,

under both control and IL-1 conditions, viable and apoptotic cells coexist in endothelial

cell cultures. IL-1 , however, did not affect either cell detachment or the apoptosis status

of adherent and detached endothelial cell fractions.

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Figure 2. Annexin V and PI staining of endothelial cells. Endothelial cells were incubated

without (control; A, B) or with IL-1 (5 ng/mL; C, D) for 24 hours. Detached (B, D) and

adherent endothelial cells (A, C) were isolated, labeled with annexin V and PI, and

analyzed by flow cytometry (n=3). The dot plots shown were obtained within one

representative experiment. Whereas most of the detached endothelial cells stained for

both annexin V and propidium iodide, only a minor fraction of the adherent cells was

positive for both markers.

82

Chapter 4

Presence of caspase 3 in endothelial cells

To confirm the presence of apoptotic cells even under control conditions, both

detached and adherent endothelial cells were analyzed flow cytometrically for the

presence of caspase 3. The majority of the detached cells, both under control condition

[Figure 3B; 60.0% 23.3 (n=3)] and after treatment with IL-1 for 24 hours [Figure 3D;

55.4% 17.1 (n=3)] stained for caspase 3. Treatment with IL-1 did not affect the

fraction of detached cells that stained for caspase 3 (P=0.728). In contrast, only minor

fractions of adherent cells stained for caspase 3 under control condition (Figure 3A; 0.3%

0.3) or after treatment with IL-1 (Figure 3C; 2.7% ± 2.9, P=0.314 compared with

control). Thus, cellular fractions of adherent or detached endothelial cells that stain for

caspase 3 were not affected by treatment with IL-1 .

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Figure 3. Intracellular detection of caspase 3 by flow cytometry. Endothelial cells were incubated without (control; A, B) or with IL-1 (5 ng/mL; C, D) for 24 hours. The detached (B, D) and adherent cells (A, C) were collected separately, labeled with anti-caspase 3 MoAb, and analyzed by flow cytometry (n=3). The dot plots shown were obtained within one representative experiment. Both in the absence and presence of IL-1 , the majority of the detached cells stained for caspase 3 whereas only minor fractions of the adherent cells stained positive.

83


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Detection of caspase 3 in EMP by Western blot

Because the numbers of EMP highly correlated with the numbers of detached

endothelial cells, which contained caspase 3, we hypothesized that EMP may also contain

caspase 3. Therefore, the presence of caspase 3 and its precursor, procaspase 3, were

studied by Western blot (Figure 4). EMP from control as well as IL-1 -treated endothelial

cells contained substantial amounts of the 17 kDa (and to a lesser extent the 19 kDa) form

of caspase 3. In contrast, procaspase 3 (32 kDa) was not detectable. EMP were isolated

from fixed volumes of conditioned medium (5 mL) and the absolute numbers of EMP

were determined for the conditions studied to be approximately 2-fold different (Figure

1A), which was reflected in the intensity of staining of -tubulin as a marker to indicate

the amounts of EMP loaded per lane. In four out of the six experiments, the 17 kDa

caspase 3 band was clearly more pronounced in EMP lysates from IL-1 -treated cells

compared to control. In the other two experiments, these bands showed similar intensities.

These data are roughly in line with the observed increase in EMP numbers upon IL-1

treatment (Figure 1A). Whether or not this treatment with IL-1 affects the quantity of

caspase 3 in EMP, however, remains to be established.

To confirm the presence of caspase 3 in detached endothelial cells and absence in

adherent cells, cell lysates were also subjected to Western blotting. Detached endothelial

cells from control as well as IL-1 -treated cell cultures contained the 17 kDa form of

caspase 3 but not its 32 kDa proform. In contrast, adherent cells contained only

procaspase 3. Because only a minor fraction of the endothelial cells was detached (Figure

1B), the higher number of cells in the lysates from adherent cells compared with the

detached cells explains why the tubulin band of the adherent cells is much more

pronounced than that of the detached cells on the Western blots shown in Figure 4.

Treatment with IL-1 did not affect the caspase 3 positivity of the EMP or the detached

cells, nor the procaspase positivity and absence of caspase 3 in the adherent cells.

Chapter 4

EMP Detached Adherent

Caspase 3

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Figure 4. Western blot of (pro)caspase 3 in EMP. EMP were isolated from culture

supernatants of endothelial cells incubated in the absence (control) or presence of IL-1

(5 ng/mL) for 24 hours. EMP were isolated from fixed quantities of volume and tubulin

was used as an additional marker to estimate the quantities of protein loaded per lane.

EMP contained the 17 and 19 kDa forms of caspase 3 but not the 32 kDa proform. In

parallel, adherent and detached endothelial cell lysates were also analyzed for the

presence of (pro)caspase 3. The blots shown represents one out of six separate

experiments and were all obtained within one representative experiment.

Detection of caspase 3 in subpopulations of EMP

To investigate whether the presence of caspase 3 is restricted to particular

subpopulations of EMP, we labeled EMP with anti-caspase 3 MoAb in the absence or

presence of saponin (Figure 5A and B, respectively). This figure confirms that EMP

contain caspase 3 and illustrates that permeabilization is essential for the detection of this

intravesicular protein. Because permeabilization impaired the binding of annexin V, we

omitted annexin V from the experiments described below. From the areas under the curve

it is apparent that saponin affected the numbers of events analyzed by the flow cytometer,

particularly in the (IgG1) control condition, but not the caspase 3 positivity itself.

Previously, we showed that ICAM-1 (CD54) and E-selectin (CD62E) are exposed

only on EMP from IL-1 -treated but not control cells, whereas PECAM-1(CD31)-

exposing EMP occur in both conditions [19]. EMP from IL-1 -treated cells (n=6) were

incubated with anti-caspase 3 MoAb without (Figure 5C) or in combination with MoAbs

85


86

directed against either PECAM-1 (Figure 5D), ICAM-1 (Figure 5E) or E-selectin (Figure

5F). The (representative) dot plots show that virtually all PECAM-1-positive EMP contain

caspase 3 (87.2% 5.0; n=6). ICAM-1- and E-selectin-exposing EMP also contained

caspase 3 (89.2% 2.4 and 88.4% 3.9, respectively).

Detection of microparticle-associated caspase 3 in human plasma

Recently, we and others demonstrated that E-selectin specifically detects EMP in

human plasma samples [19,20]. To study the possible occurrence of caspase 3 in such

vesicles in vivo, total MP fractions were isolated from plasma samples of three SLE

patients and three healthy individuals, and stained with anti-caspase 3 plus MoAbs

directed against either E-selectin (Figure 6, top row), platelet glycoprotein IIIa ( 3; CD61;

second row), erythrocyte glycophorin A (third row) or monocyte LPS-receptor (CD14;

bottom row). MP from all plasma samples stained for caspase 3. Plasma samples from two

SLE patients also contained a subpopulation of E-selectin-exposing EMP, which strongly

double stained for caspase 3. Most by far of the caspase 3-containing MP originated from

platelets and to a lesser extent from erythrocytes. Plasma samples of the two ‘E-selectin-

positive’ SLE patients also contained monocyte-derived MP that double stained for

caspase 3.

Chapter 4

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Figure 5. Flow cytometric detection of caspase 3 in EMP. EMP were isolated and labeled

with either IgG1-control antibody (gray) or anti-caspase 3 MoAb (dark line) in the

absence (A) or presence (B) of saponin. (C-F) Dot plots, obtained within one experiment

representative of five similar experiments, from EMP obtained after treatment of

endothelial cells with IL-1 , that were incubated with anti-caspase 3 MoAb alone (C) or

in combination with PECAM-1 (CD31, D), ICAM-1 (CD54, E) or E-selectin (CD62E, F).

87


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Figure 6. The presence of caspase 3 in MP in vivo. MP were isolated from plasma

samples of three SLE patients (shown in columns 4, 5 and 6) and three healthy individuals

(shown in columns 1, 2 and 3), labeled with MoAbs and analyzed by flow cytometry. Each

column represents dot plots from a single person, labeled with anti-caspase 3 in

combination with either (i) anti-E-selectin (top row), (ii) anti- 3 (CD61) (second row),

(iii) anti-glycophorin A (third row) or (iv) anti-CD14 (bottom row). All analyzed plasma

samples contained MP that stained for caspase 3. Most of these vesicles were derived

from platelets ( 3) and to a lesser extent from erythrocytes (glycophorin A). Plasma from

two SLE patients contained a population of E-selectin-exposing (E)MP that strongly

double stained for caspase 3. Plasma from these two patients also contained MP from

monocytes that also double stained for caspase 3 (CD14).

88

Chapter 4

89

DISCUSSION

In the present study we investigated the possible relationship between adherent and

detached endothelial cells and the formation of EMP. In line with previous studies, we

found that detached endothelial cells are apoptotic, similar to other cells losing their

association with the matrix [22-26], i.e. most of the detached cells stained for annexin V

and PI, and contained caspase 3. Also most of the EMP contained caspase 3, in vitro and

in particular in vivo. In addition, EMP numbers correlated highly with the numbers of

detached cells, suggesting that the majority of EMP originate from these cells. In vivo, the

presence of caspase 3 was not only restricted to MP originating from nucleated cells, since

significant fractions of MP from platelets and erythrocytes also contained detectable

amounts of caspase 3. Evidently, MP formation is not linked to full-blown apoptosis, i.e.

nuclear fragmentation, cell death and disintegration. Nevertheless, the present findings are

only based on a limited number of in vitro and ex vivo experiments. Also, compared to

age- and sex-matched healthy controls, only plasma from two of the three SLE patients

studied contained significantly different (sub)populations of MP. Therefore, additional

studies will be required to substantiate our present findings.

Platelets are known to contain procaspase 3 and other proteins involved in the

apoptosis process [35]. The occurrence of caspase 3 in MP from platelets may suggest that

this enzyme is somehow involved in the process of membrane vesiculation. Caspase 3

elicits other cellular functions than solely apoptosis, such as maintaining the cellular

morphology [36]. Thus, the presence of caspase 3 in these MP may not be necessarily

linked to apoptosis at all, but simply coincide with caspase 3 activation without ongoing

apoptosis.

Hamilton et al. demonstrated that endothelial cells are protected from complement-

induced lysis by shedding EMP containing the C5b-9 complex [37]. In other words, the

release of EMP protected the cells against (extracellular) stress. It has been reported, that

(cultured) cells also undergo constitutive apoptosis [38,39]. If so, one may hypothesize

that vesiculation protects cells not only against extracellular stress but also against

intracellular stress, e.g. by releasing caspase 3-containing EMP. Thus, at present we

cannot exclude that a fraction of the caspase 3-containing EMP originates from adherent

endothelial cells. Alternatively, a careful examination of the dot plots presented in Figure


90

5 (upper left in panels Figure 5D-F) suggests that a small fraction of EMP is also present

that did not contain caspase 3. Such EMP may have originated from adherent cells. Such a

small fraction, however, was absent in E-selectin-exposing MP in vivo.

Clancy et al. demonstrated elevated levels of activated circulating endothelial cells in

SLE patients [40]. They suggested that these levels may represent a marker of endothelial

injury. In the light of their findings, our in vivo observation of caspase 3 in E-selectin-

exposing MP suggests that most of such vesicles may have originated from circulating,

apoptotic endothelial cells which had also become activated as evidenced by the exposure

of E-selectin on the EMP.

The present findings show that MP may contain caspase 3. Whether or not these cell-

derived MP subsequently transfer caspase 3 to other cells and thus contribute to the

induction of endothelial cell dysfunction (as recently shown for MP from cultured

endothelial cells [41], women with preeclampsia [42], patients with myocardial infarction

[43], or T lymphocytes [44]), remains to be demonstrated.

Chapter 4

91

References


2. Cai H, Harrison DG. Endothelial dysfunction in cardiovascular diseases: the role of oxidant stress. Circ. Res. 2000;87:840-4.



5. Brodsky SV, Yamamoto T, Tada T, Kim B, Chen J, Kajiya F, Goligorsky MS. Endothelial dysfunction in ischemic acute renal failure: rescue by transplanted endothelial cells. Am. J. Physiol Renal Physiol. 2002;282:F1140-F1149.

6. Filer AD, Gardner-Medwin JM, Thambyrajah J, Raza K, Carruthers DM, Stevens RJ, Liu L, Lowe SE, Townend JN, Bacon PA. Diffuse endothelial dysfunction is common to ANCA associated systemic vasculitis and polyarteritis nodosa. Ann. Rheum. Dis. 2003;62:162-7.

7. Suwaidi JA, Hamasaki S, Higano ST, Nishimura RA, Holmes DR, Jr., Lerman A. Long-term follow-up of patients with mild coronary artery disease and endothelial dysfunction. Circulation 2000;101:948-54.

8. Albornoz L, Alvarez D, Otaso JC, Gadano A, Salviu J, Gerona S, Sorroche P, Villamil A, Mastai R. Von Willebrand factor could be an index of endothelial dysfunction in patients with cirrhosis: relationship to degree of liver failure and nitric oxide levels. J. Hepatol. 1999;30:451-5.

9. Belch JJ, Shaw JW, Kirk G, McLaren M, Robb R, Maple C, Morse P. The white blood cell adhesion molecule E-selectin predicts restenosis in patients with intermittent claudication undergoing percutaneous transluminal angioplasty. Circulation 1997;95:2027-31.

10. Borawski J, Naumnik B, Pawlak K, Mysliwiec M. Endothelial dysfunction marker von Willebrand factor antigen in haemodialysis patients: associations with pre-dialysis blood pressure and the acute phase response. Nephrol. Dial. Transplant. 2001;16:1442-7.


92




14. Mallat Z, Benamer H, Hugel B, Benessiano J, Steg PG, Freyssinet JM, Tedgui A. Elevated levels of shed membrane microparticles with procoagulant potential in the peripheral circulating blood of patients with acute coronary syndromes. Circulation 2000;101:841-3.



17. Ogura H, Tanaka H, Koh T, Fujita K, Fujimi S, Nakamori Y, Hosotsubo H, Kuwagata Y, Shimazu T, Sugimoto H. Enhanced production of endothelial microparticles with increased binding to leukocytes in patients with severe systemic inflammatory response syndrome. J. Trauma 2004;56:823-31.

18. Combes V, Taylor TE, Juhan-Vague I, Mege JL, Mwenechanya J, Tembo M, Grau GE, Molyneux ME. Circulating endothelial microparticles in malawian children with severe falciparum malaria complicated with coma. JAMA 2004;291:2542-4.



Chapter 4

93

21. Horstman LL, Jy W, Jimenez JJ, Ahn YS. Endothelial microparticles as markers of endothelial dysfunction. Front. Biosci. 2004;9:1118-35.

22. Araki S, Shimada Y, Kaji K, Hayashi H. Apoptosis of vascular endothelial cells by fibroblast growth factor deprivation. Biochem. Biophys. Res. Commun. 1990;168:1194-200.

23. Hase M, Araki S, Kaji K, Hayashi H. Classification of signals for blocking apoptosis in vascular endothelial cells. J. Biochem. 1994;116:905-9.

24. Levkau B, Herren B, Koyama H, Ross R, Raines EW. Caspase-mediated cleavage of focal adhesion kinase pp125FAK and disassembly of focal adhesions in human endothelial cell apoptosis. J. Exp. Med. 1998;187:579-86.


26. Frisch SM, Ruoslahti E. Integrins and anoikis. Curr. Opin. Cell Biol. 1997;9:701-6.

27. Douma S, Van Laar T, Zevenhoven J, Meuwissen R, Van Garderen E, Peeper DS. Suppression of anoikis and induction of metastasis by the neurotrophic receptor TrkB. Nature 2004;430:1034-9.

28. Mutunga M, Fulton B, Bullock R, Batchelor A, Gascoigne A, Gillespie JI, Baudouin SV. Circulating endothelial cells in patients with septic shock. Am. J. Respir. Crit Care Med. 2001;163:195-200.

29. Bull TM, Golpon H, Hebbel RP, Solovey A, Cool CD, Tuder RM, Geraci MW, Voelkel NF. Circulating endothelial cells in pulmonary hypertension. Thromb. Haemost. 2003;90:698-703.

30. Lefevre P, George F, Durand JM, Sampol J. Detection of circulating endothelial cells in thrombotic thrombocytopenic purpura. Thromb. Haemost. 1993;69:522.

31. Solovey A, Lin Y, Browne P, Choong S, Wayner E, Hebbel RP. Circulating activated endothelial cells in sickle cell anemia. N. Engl. J. Med. 1997;337:1584-90.

32. Hochberg MC. Updating the American College of Rheumatology revised criteria for the classification of systemic lupus erythematosus. Arthritis Rheum. 1997;40:1725.


94

33. Bombardier C, Gladman DD, Urowitz MB, Caron D, Chang CH. Derivation of the SLEDAI. A disease activity index for lupus patients. The Committee on Prognosis Studies in SLE. Arthritis Rheum. 1992;35:630-40.

34. Pitti RM, Marsters SA, Ruppert S, Donahue CJ, Moore A, Ashkenazi A. Induction of apoptosis by Apo-2 ligand, a new member of the tumor necrosis factor cytokine family. J. Biol. Chem. 1996;271:12687-90.

35. Wolf BB, Goldstein JC, Stennicke HR, Beere H, Amarante-Mendes GP, Salvesen GS, Green DR. Calpain functions in a caspase-independent manner to promote apoptosis-like events during platelet activation. Blood 1999;94:1683-92.

36. Rohn TT, Cusack SM, Kessinger SR, Oxford JT. Caspase activation independent of cell death is required for proper cell dispersal and correct morphology in PC12 cells. Exp. Cell Res. 2004;295:215-25.

37. Hamilton KK, Hattori R, Esmon CT, Sims PJ. Complement proteins C5b-9 induce vesiculation of the endothelial plasma membrane and expose catalytic surface for assembly of the prothrombinase enzyme complex. J. Biol. Chem. 1990;265:3809-14.

38. Knepper-Nicolai B, Savill J, Brown SB. Constitutive apoptosis in human neutrophils requires synergy between calpains and the proteasome downstream of caspases. J. Biol. Chem. 1998;273:30530-6.

39. Simak J, Holada K, D'Agnillo F, Janota J, Vostal JG. Cellular prion protein is expressed on endothelial cells and is released during apoptosis on membrane microparticles found in human plasma. Transfusion 2002;42:334-42.

40. Clancy RM. Circulating endothelial cells and vascular injury in systemic lupus erythematosus. Curr. Rheumatol. Rep. 2000;2:39-43.

41. Brodsky SV, Zhang F, Nasjletti A, Goligorsky MS. Endothelium-derived microparticles impair endothelial function in vitro. Am. J. Physiol Heart Circ. Physiol. 2004;286:H1910-H1915.

42. VanWijk MJ, Svedas E, Boer K, Nieuwland R, VanBavel E, Kublickiene KR. Isolated microparticles, but not whole plasma, from women with preeclampsia impair endothelium-dependent relaxation in isolated myometrial arteries from healthy pregnant women. Am. J. Obstet. Gynecol. 2002;187:1686-93.

43. Boulanger CM, Scoazec A, Ebrahimian T, Henry P, Mathieu E, Tedgui A, Mallat Z. Circulating microparticles from patients with myocardial infarction cause endothelial dysfunction. Circulation 2001;104:2649-52.

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44. Martin S, Tesse A, Hugel B, Martinez MC, Morel O, Freyssinet JM, Andriantsitohaina R. Shed membrane particles from T lymphocytes impair endothelial function and regulate endothelial protein expression. Circulation 2004;109:1653-9.

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Chapter 5

Simvastatin-induced endothelial cell

detachment and microparticle release are

prenylation dependent

Michaela Diamant, Maarten E. Tushuizen, Mohammed N. Abid Hussein, Chi M. Hau,

Anita N. Böing, Augueste Sturk and Rienk Nieuwland

Submitted

Simvastatin-induced endothelial detachment

98

ABSTRACT

Background: Statins reduce cardiovascular disease risk and affect endothelial function by

cholesterol-dependent and independent mechanisms. Recently, circulating (detached)

endothelial cells and endothelial microparticles (EMP) have been associated with

endothelial functions in vitro and in vivo.

Objective: We investigated whether simvastatin affects endothelial detachment and

release of EMP.

Methods: Human umbilical vein endothelial cells (HUVEC) were incubated with

clinically relevant concentrations of simvastatin (1.0 and 5.0 μmol/L), with or without

mevalonic acid (100 μmol/L) or geranylgeranylpyrophosphate (GGPP; 20 μmol/L) for 24

hours, and analyzed by flowcytometry and Western blot.

Results: Simvastatin increased detachment from 12.5% ± 4.1 to 26.0% ± 7.6 (1.0 μmol/L;

P=0.013) and 28.9% ± 2.2 (5.0 μmol/L; P=0.002). Concurrently, EMP release increased

2.5-fold (P=0.098 and P=0.041, respectively). Adherent cells showed no signs of

simvastatin-induced apoptosis (caspase 3, annexin V, propidium iodide), suggesting that

cell detachment and EMP release are not necessarily due to apoptosis. In contrast, the

majority of detached cells was apoptotic, although the fraction of detached cells that

showed signs of apoptosis ( 70%) was unaffected by simvastatin. Similar to these

detached cells, EMP contained caspase 3. Furthermore, detached cells and EMP contained

caspase 8 but not caspase 9. By restoring either cholesterol biosynthesis and prenylation

(mevalonate) or prenylation alone (GGPP), all simvastatin-induced effects on detachment

and EMP release could be reversed.

Conclusions: Simvastatin promotes detachment and EMP release by inhibiting

prenylation, presumably via a caspase 8-dependent mechanism. We hypothesize that by

facilitating detachment and EMP release, statins may improve the overall condition of the

vascular endothelium.

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99

INTRODUCTION

Statins are widely prescribed lipid-lowering drugs that significantly reduce

cardiovascular morbidity and mortality in many different patient populations, as

demonstrated in multiple large primary and secondary prevention trials [1-9]. By

inhibiting 3-hydroxy-methylglutaryl coenzyme A (HMG-CoA) reductase, the rate-limiting

enzyme of cholesterol biosynthesis, statins reduce both total and low-density lipoprotein-

associated cholesterol (LDL-associated cholesterol). Their beneficial effects in

cardiovascular disease (CVD) patients have been largely attributed to their efficacy to

lower LDL-associated cholesterol [4]. Statins, also have additional pleiotropic

(cholesterol-independent) effects, many of which are mediated by the vascular

endothelium [10-17]. However, data from several, mainly in vitro studies, may be difficult

to interpret since statins were used at pharmacological and possibly cytotoxic

concentrations or in combination with a variety of agonists like TNF- , endotoxin or

thrombin [13,16]. The existence of pleiotropic effects of statins in vivo, separate from

their cholesterol-lowering potential, was recently substantiated by Landmesser et al, who

showed improvement of endothelial dysfunction in patients with chronic heart failure after

simvastatin therapy but not after treatment with the cholesterol absorption inhibitor

ezetimibe, given at a dose that lowered LDL-associated cholesterol to a similar extent

[18].

Pleiotropic effects of statins seem to be mainly caused by inhibition of protein

prenylation. Prenylation is a post-translational mechanism of protein modification, in

which intermediates of the mevalonate pathway, like geranylgeranylpyrophosphate

(GGPP), are attached to proteins. Geranylgeranylated proteins, including the small G

proteins Rho, Rac and Rab, bind to cell membranes and are required for transmembrane

signaling [19]. As a consequence, G proteins are involved in the regulation of cell growth,

differentiation, gene expression, cytoskeletal assembly and cell motility, formation of

microparticles (MP) or “apoptotic bodies”, protein and lipid trafficking, nuclear transport

and host defense [19]. Thus, by preventing formation of mevalonate, statins block

cholesterol biosynthesis and transmembrane signaling.

Previously, we showed that cultures of viable and unstimulated human umbilical vein

endothelial cells (HUVEC) contain small numbers of detached cells ('floaters') undergoing


100

apoptosis as well as endothelial cell-derived MP (EMP) [20]. Since in the afore-

mentioned in vitro studies no or hardly any attention was paid to detachment and/or

release of EMP, we hypothesized that to gain insight into the full response of endothelial

cells to statins, not only the adherent cell fraction but also the corresponding fraction of

detached cells and EMP have to be analyzed. Therefore, in order to study the true impact

of statins on the endothelium, we determined the effect of simvastatin on human

endothelial cells by concurrently analyzing adherent cells, detached cells and EMP.


Reagents and assays

Medium M199, penicillin, streptomycin, Isocove’s modified dulbecco’s medium and

L-glutamine were obtained from GibcoBRL, Life Technologies (Paisley, Scotland).

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). Human

recombinant basic fibroblast growth factor (bFGF), and epidermal growth factor (EGF)

were obtained from Invitrogen life technologies (Carlsbad, CA). Collagenase (type 1A),

geranylgeranylpyrophosphate (GGPP), mevalonolactone (mevalonate) and propidium

iodide (PI) were from Sigma (St. Louis, MO). APC-labeled annexin V was from Caltag

Laboratories (Burlingame, CA). Heparin (400 U/mL) was from Leo Pharma BV (Breda,

The Netherlands), trypsin from Difco Laboratories (Detroit, MI), and simvastatin from

Calbiochem (Darmstadt, Germany). Tissue culture flasks were from Greiner Labortechnik

(Frickenhausen, Germany) and gelatin from Difco Laboratories (Sparks, MD). Stock

solutions of simvastatin, mevalonate and GGPP were prepared in ethanol, ethanol and

methanol, respectively. Antibodies against (pro-)caspase 9, (pro-)caspase 8 and caspase 3

for Western blotting were obtained from Cell Signaling (Beverly, MA). Anti-procaspase 3

was from Transduction Laboratories (San Diego, CA). Secondary antibodies used for

Western blot, i.e. goat-anti-mouse HRP conjugate and anti-rabbit HRP conjugate, were

from Biorad (Hercules, CA) and Promega (Madison, USA), respectively. FITC-labeled

annexin V was from Immuno Quality Products (Groningen, The Netherlands).

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101

Isolation, culture and treatment of HUVEC

HUVEC were collected from human umbilical cord veins as described previously

[20]. Briefly, umbilical cords were digested with collagenase for 20 minutes at 37 °C.

Detached cells were obtained by perfusion of the umbilical cord with medium M199

supplemented with 10% HuSi. The cell suspension was centrifuged for 10 minutes at 180

g and 20 °C, and cells were resuspended in culture medium. HUVEC were cultured in

tissue culture flasks coated with 0.75% gelatin (passage 0). Upon confluency at passage 3,

HUVEC were kept for 3-4 days in a resting state. Then, the culture supernatant was

refreshed and (where indicated) cultures were treated for 24 hours without any addition

(control), ethanol (0.2% v/v), methanol (0.2% v/v), ethanol and methanol (both 0.2% v/v),

simvastatin (1.0 μM and 5.0 μM final concentration (fc)), mevalonate (100 μM fc), GGPP

(20 μM fc), and combinations of simvastatin plus mevalonate or GGPP. Administration of

10-40 mg simvastatin results in (peak) plasma concentrations of 1-6 ng/mL, which is in

line with the 0.4-2.1 ng/mL used in our present study [21,22].

Flow cytometric analysis of endothelial cells

Conditioned media (10 mL per 75 cm2 flasks) were harvested after 24 hours. First,

media were centrifuged for 10 minutes at 180 g and 20 °C to isolate detached endothelial

cells and to obtain the cell-free conditioned medium for EMP isolation. The detached cell

pellets were carefully resuspended in 1% FCSi in PBS (pH 7.4). In parallel, the adherent

endothelial cells were harvested by trypsinization. After 4 minutes, trypsin was

neutralized by PBS/FCSi. Both cell suspensions were separately centrifuged for 10

minutes at 180 g and 4 °C, resuspended in PBS/FCSi, kept on melting ice for 15 minutes,

and then again centrifuged for 10 minutes at 180 g and 4 °C. The detached cells were

resuspended in 0.5 mL PBS/FCSi and the adherent cells in 1 mL PBS/FCSi. For

intracellular staining for caspase 3, the active caspase 3-FITC MoAb apoptosis kit I was

used (BD Pharmingen; San Diego, CA). From the before mentioned suspension of

detached and adherent cells, 100 L were diluted with 1 mL of ice-cold PBS (pH 7.4).

This suspension was centrifuged for 10 minutes at 180 g. After removal of the

supernatant, the cells were again diluted with 1 mL of ice-cold PBS and pelleted (10

minutes at 180 g). After removal of the supernatant, cells were resuspended in 500 L


102

cytofix/cytoperm and incubated for 20 minutes on melting ice. To remove the

cytofix/cytoperm, cells were pelleted (10 minutes at 180 g) and supernatant was removed.

Then the cells were washed twice with 10-fold diluted perm/wash, and finally

resuspended in 100 L 10-fold diluted perm/wash. From this suspension, two aliquots of

50 L each were incubated for 30 minutes at room temperature with either anti-caspase 3-

FITC (5 L) or Ig-FITC (5 L). After incubation, 1 mL of 10-fold diluted perm/wash was

added to each aliquot, and the suspension was centrifuged for 10 minutes at 180 g. The

supernatant was removed and the pellets were resuspended in 300 L 10-fold diluted

perm/wash. All samples were analyzed in a FACSCalibur flow cytometer (BD; San Jose,

CA). Percentages of adherent and detached cells were compared to the total cell count

(i.e. adherent plus detached cells)/culture flask (100%). Labeling with annexin V and PI to

determine the apoptosis status of the endothelial cells was performed as described

previously [20].

Isolation of EMP

Aliquots (1 mL) of the cell-free conditioned media were frozen in liquid nitrogen and

then stored at – 80 °C. Before use, samples were thawed on melting ice for 1 hour, and

then centrifuged for 1 hour at 17,570 g and 20 °C. Then, 950 μL of (MP-free) supernatant

was removed. The remaining 50 L of EMP suspension was divided into two aliquots of

25 L each, of which one aliquot was used for regular flow cytometry and the other

aliquot for intravesicular caspase 3 staining.

For regular flow cytometry, 25 L EMP suspension was diluted with 225 μL PBS

(154 mmol/L NaCl, 1.4 mmol/L phosphate) containing 10.9 mmol/L trisodium citrate.

EMP were resuspended and again centrifuged for 30 minutes at 17,570 g and 20 °C.

Again, 225 μL of supernatant was removed and EMP (25 L) were finally diluted with 25

L PBS/citrate buffer. For intravesicular staining of caspase 3, the 25 L EMP suspension

was diluted with 225 μL 100-fold diluted perm/wash. EMP were resuspended and

centrifuged for 30 minutes at 17,570 g and 20 °C. Again, 225 μL of supernatant was

removed and EMP (25 L) were diluted with 25 L 100-fold diluted perm/wash.

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103

Flow cytometric analysis of EMP

EMP samples were analyzed in a FACSCalibur flow cytometer (BD; San Jose, CA).

Forward scatter (FSC) and side scatter (SSC) were set at logarithmic gain and EMP were

characterized as described previously by binding of annexin V. EMP (5 μL aliquots) were

diluted with 45 μL PBS containing 2.5 mmol/L CaCl2 (pH 7.4). Annexin V-APC (5 μL of

20-fold diluted) was added. In the control samples of the MP, annexin V-positive events

were identified by placing a threshold in a MP sample (5 μL) diluted with PBS containing

10.9 mmol/L trisodium citrate (45 μL; pH 7.4) and 5 μL of annexin V, i.e. without Ca2+.

The mixture of MP and annexin V was then incubated for 15 minutes in the dark at room

temperature, and finally diluted with 900 L PBS containing either calcium or citrate. For

intravesicular staining of caspase 3, EMP were incubated for 30 minutes with the

indicated antibodies and APC-labeled annexin V in the dark at room temperature. The

labeling was stopped by addition of 900 μL of 100-fold diluted perm/wash before flow

cytometric analysis.

Western blotting

For Western blotting experiments, 400 μL of the detached cell suspensions and 450

μL of the adherent cell suspension were diluted with 5-fold concentrated sample buffer

containing -mercaptoethanol. EMP were harvested by centrifugation from 5 mL of cell-

free conditioned medium, and finally resuspended in a mixture of 24 μL PBS and 6 μL 5-

fold concentrated sample buffer. Before electrophoresis, all samples were heated for 5

minutes at 100 °C. Electrophoresis was carried out on 8-16% gradient polyacrylamide gel

(Biorad; Hercules, CA). The 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

subsequently incubated with anti-caspase 3 (1:1,000 v/v), anti-(pro)-caspase 8 (1:1,000

v/v) or anti-(pro)caspase 9 (1:1,000 v/v) for 24 hours at 4 °C, followed by incubation with

either anti-rabbit IgG-HRP conjugate (1:7,500 v/v; used in combination with the anti-

caspase 3 antibody) or goat-anti-mouse HRP conjugate (1:3,000 v/v; used in combination

with the other mentioned antibodies) for 1 hour at room temperature. After each


104

incubation step, PVDF membranes were washed three times with TBST for 5-10 minutes.

All antibodies were diluted with blocking buffer. The bands were detected using Lum-

Light Plus Western Blotting Substrate (Roche; Mannheim, Germany) and exposed to Fuji

Medical X-ray film.


Data were analyzed with GraphPad Prism for Windows (release 3.02; San Diego,

CA). Differences were analyzed by t-test for independent samples and were considered to

be significant at P<0.05. All data are presented as mean ± SD. Data were obtained from at

least 3 independent experiments, i.e. using endothelial cell cultures from 3 or more

different umbilical veins. Data were compared to ethanol- (EtOH) and methanol (MeOH)-

treated endothelial cell cultures.

RESULTS

Simvastatin induces endothelial cell detachment

Upon incubation with simvastatin, the detached cell fraction (Figure 1A) increased

from 12.5% ± 4.1 (ethanol plus methanol control) to 26.0% ± 7.6 (1.0 μmol/L simvastatin;

P=0.013) and 28.9% ± 2.2 (5.0 μmol/L simvastatin; P=0.002). Cell detachment was not

affected by incubation with mevalonate (100 μmol/L; P=0.207) or GGPP (20 μmol/L;

P=0.205) alone, but both compounds completely prevented simvastatin-induced

detachment. Thus, simvastatin-induced endothelial cell detachment can be reversed not

only by restoring cholesterol biosynthesis (mevalonate) but also by restoring prenylation

(GGPP).

Caspase 3 in adherent and detached endothelial cell fractions

Figure 1B shows flow cytometry dot plots of caspase 3 in adherent (Figure 1B; A-D)

and detached (Figure 1B; E-H) endothelial cells. Compared to staining with control

antibody (IgG-FITC), a negligible number of adherent cells stained for caspase 3 (Figure

1B; B versus A), and this was similar in the presence of 1.0 or 5.0 μmol/L simvastatin

(Figure 1B; C and D, respectively). In contrast, a substantial number of detached cells

Chapter 5

stained for caspase 3 (Figure 1B; F versus E), and these numbers increased 2.5 to 3-fold in

the presence of either 1.0 or 5.0 μmol/L (Figure 1B; G and H, respectively) simvastatin.

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Figure 1. A. Percentage of detached cell after HUVEC were incubated (24 hours) without any additions (control), ethanol (0.2% v/v) plus methanol (0.2% v/v), simvastatin (1.0 μmol/L and 5.0 μmol/L), mevalonate (100 μmol/L) without or with simvastatin, and GGPP (20 μmol/L) without or with simvastatin. Experiments were performed with at least three different HUVEC cultures and all data were compared to control, i.e. ethanol plus methanol. B. Shown are representative dot plots from a typical experiment. HUVEC incubated with vehicle (ethanol, 0.2% v/v; A, B, E, F), simvastatin (1.0 μmol/L; C, G) or simvastatin (5.0 μmol/L; D, H). Adherent (shown in panels A-D) and detached (panels E-H) cells were separately isolated and stained with FITC-labeled IgG control antibody (panels A and E) or anti-caspase 3 antibody (panels B-D and F-H).

105


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Figure 2A shows the increase in absolute numbers of detached cells containing

caspase 3 in the presence of 1.0 or 5.0 μmol/L simvastatin (P=0.039 and P=0.041,

respectively). The simvastatin-induced increases were completely blocked by co-

incubation with mevalonate or GGPP. Since the fractions of both detached and adherent

cells staining for caspase 3 were not affected by simvastatin (Figure 2B), these data may

implicate that simvastatin may facilitate detachment rather than induce apoptosis. Figure

2C confirms the presence of caspase 3 in detached cell lysates from simvastatin-treated

cultures (upper right). Since detached cells were isolated from a fixed volume of

conditioned medium, the observed increase in caspase 3 is due to the increased number of

detached cells. In contrast, in adherent cell lysates no caspase 3 could be detected (upper

left). Co-incubation with either mevalonate or GGPP completely prevented the

simvastatin-induced increase in caspase 3 formation (data not shown). Procaspase 3 was

detectable only in adherent (lower left) but not in detached cells (lower right).

Since the presence of caspase 3 is no absolute evidence that cells are undergoing

apoptosis, adherent and detached cells were stained for annexin V (early apoptosis) and PI

(late apoptosis) in a control experiment from both control and 1.0 μmol/L simvastatin-

treated cultures to determine whether or not simvastatin affects their apoptotic status

(Figure 3). On average, only 3.8% 1.6 of adherent cells stained for annexin V and PI in

this experiment, irrespective of the presence (Figure 3B and 3D) or absence (Figure 3A

and 3C) of simvastatin. The total number of detached cells staining for annexin V or PI

increased in the presence of simvastatin (Figure 3F and 3H versus 3E and 3G,

respectively). The relative proportion of these (detached) cells staining for annexin V and

PI, however, was unaffected by simvastatin (74% 10). Thus, simvastatin may not have a

direct effect on the apoptotic status of adherent endothelial cells, and the occurrence of

caspase 3 in this experiment closely parallels staining for annexin V and PI as markers of

apoptosis.

Chapter 5

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Figure 2. A. Numbers of caspase 3-containing detached cells. B. Fractions of adherent

(open bars) and detached (shaded bars) cells containing caspase 3. HUVEC were

incubated (24 hours) without any additions (control), ethanol (0.2% v/v) plus methanol

(0.2% v/v), simvastatin (1.0 μmol/L and 5.0 μmol/L), mevalonate (100 μmol/L) without or

with simvastatin, and GGPP (20 μmol/L) without or with simvastatin. Data are mean

percentages of adherent cells and detached cells containing caspase 3. C. Western blot of

caspase 3 and procaspase 3 in adherent and detached endothelial cell lysates from a

single, representative experiment.

107


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Figure 3. Effect of simvastatin on the apoptotic status of detached and adherent cells.

HUVEC were incubated without (A, C, E, G) or with (B, D, F, H) simvastatin (1.0

μmol/L) for 24 hours. Adherent (A-D) and detached cells (E-H) were stained with annexin

V (A, B, E, F) or PI (C, D, G, H).

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Effect of simvastatin on EMP formation

Upon incubation with either 1.0 or 5.0 μmol/L simvastatin, the numbers of caspase 3-

containing EMP increased (Figure 4A; C and D, respectively) when compared to control

(Figure 4A; A) or vehicle (Figure 4A; B). The numbers of annexin V-binding and caspase

3-containing EMP increased 2.5-fold in the presence of simvastatin (Figure 4B). This

increase tended to be statistically significant at 1.0 μmol/L simvastatin and reached

statistical significance at 5.0 μmol/L simvastatin (P=0.098 and P=0.041, respectively).

Co-incubation with either mevalonate or GGPP completely abolished the statin-induced

EMP release. The insert of Figure 4B confirms the presence of caspase 3 in EMP lysates.

Both mevalonate and GGPP completely prevented the observed (simvastatin-induced)

increase in caspase 3 formation (data not shown). Thus, prenylation counteracts the

simvastatin-induced release of (caspase 3-containing) EMP.

Role of caspases in statin-induced cell detachment

Active caspase 3 (17 kDa) is a cleavage product of the inactive 32 kDa precursor

(procaspase 3). Induction of programmed cell-death, either via death receptors

(‘extrinsic’) or via leakage of mitochondrial cytochrome C (‘intrinsic’), ultimately leads to

cleavage of procaspase 3 by either caspase 8 (‘extrinsic’) or caspase 9 (‘intrinsic’). Figure

5 shows that only procaspases 8 (57 kDa) and 9 (47 kDa) were detectable in adherent cell

lysates, and their relative quantities seemed unaffected by simvastatin. In contrast,

detached cells and to a lesser extent EMP contained detectable quantities of caspase 8 (43

kDa) after incubation with simvastatin, but not caspase 9 (35-37 kDa). Caspase 8 was not

detectable when cells were co-incubated with mevalonate or GGPP (data not shown).

Since caspase 8 but not caspase 9 is detectable in detached cell lysates from simvastatin-

treated cultures, we hypothesize that cleavage of procaspase 8 may explain the observed

increase in caspase 3 under these conditions.


Figure 4. A.Flowcytometric histograms of caspase 3-containing EMP from a single, representative experiment. The light grey and black curves show IgG (control antibody) and (anti-) caspase 3, respectively. EMP were isolated from HUVEC conditioned media after 24 hours of incubation without addition (control; A), vehicle (0.2% v/v; B) or simvastatin (1.0 and 5.0 μmol/L; C and D, respectively). B. Absolute numbers of caspase 3-containing EMP. EMP were isolated from conditioned media as outlined in Methods. The insert shows the effect of simvastatin on the amounts of 17 kDa caspase 3 in EMP lysates.

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Chapter 5

Adherent cells Detached cells EMP

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Figure 5. (Pro-) Caspase 8 and 9 in adherent cells, detached cells and EMP of HUVEC

incubated (24 hours) without any additions (control), ethanol (0.2% v/v) plus methanol

(0.2% v/v), simvastatin (1.0 μmol/L) or simvastatin (5.0 μmol/L). Shown are

representative Western blots from a typical experiment.

DISCUSSION

Incubation of human endothelial cells with simvastatin at clinically relevant doses

triggered endothelial cell detachment as well as EMP release. Lysates from both detached

cells and EMP contained substantial quantities of caspases 3 and 8, whereas caspase 9

remained below the detection limit, suggesting that caspase 8 formation underlies the

formation of caspase 3 under these conditions. Previously, statins were shown to induce

apoptosis of keratinocytes via ligand-independent activation of caspase 8 via a death

receptor [23]. Whether or not formation of caspase 8 precedes detachment or is a

consequence of detachment (‘anoikis’), however, remains to be determined.

Caspase 3 cleaves focal-adhesion kinase, thus eliminating essential cellular survival

signals and thereby facilitating detachment. In addition, caspase 3 cleaves kinases like

Rho-associated coiled kinase (ROCK)-I and p21-kinase, resulting in the formation of

constitutively active kinases which directly contribute to formation of “apoptotic bodies”

[24,25]. Therefore, we hypothesize that -similar to keratinocytes- simvastatin may trigger

caspase 8-mediated endothelial detachment and EMP release.

111


112

Our present data indicate that simvastatin-induced detachment and EMP release can

be circumvented by restoring prenylation. Also, other investigators showed that statins

induce apoptosis of endothelial cells in vitro, and in most studies these effects were

counteracted by restoring prenylation. The extent by which statins induce apoptosis,

however, seems to be dependent on the type of endothelial cell studied. With particular

regard to HUVEC, however, statins induce a wide variety of effects, including enhanced

expression of tissue factor and adhesion receptors, an increased release of EMP and

augmented production and bioavailability of endothelium-derived NO [10,12,13,16,26-

30]. In some of these studies, data are difficult to interpret since pharmacological and

potentially cytotoxic concentrations of statins were used or statins were used only in

combination with other endothelial agonists like TNF- or endotoxin. In most of these

studies, however, solely adherent endothelial cells were studied. Our present data suggest

that to appreciate the full scope of the statin effects on endothelial cells, all individual

components of the incubation well, i.e. adherent cells, detached cells and MP, should be

taken into account. In this regard, it should be mentioned that in some studies the

apoptotic effects of statins on (adherent) endothelial cells could only be observed in the

presence of additional inducers like TNF- , whereas in the present study the pro-apoptotic

effect of statin-treatment alone became apparent when not only adherent cells but also

detached cells and EMP were analyzed.

Whereas some potentially harmful adverse affects of statins have been reported on

endothelial cells in vitro, these drugs have many beneficial effects on the endothelium in

vivo. Statins increase the number and survival of circulating endothelial progenitor cells,

facilitate re-endothelialization, inhibit endothelial senescence and increase cell

proliferation by affecting cell cycle genes [31]. Thus, in spite of these seemingly

contradictory reports, the existence of a discrepancy between in vitro versus in vivo

effects of statins may be questioned. In our present study, we showed that adherent

endothelial cells seemed unaffected by treatment with simvastatin despite increased

numbers of both 'floaters' and EMP. Possibly, by facilitating detachment and EMP release,

statins may improve the overall condition of the vascular endothelium.

Previously, we reported a strong correlation between the numbers of detached

endothelial cells and EMP [20]. Also, our present data indicate a close association

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113

between cell detachment and EMP release. From such data, however, we may not

conclude that EMP are released from detached cells since other investigators showed that

EMP are released from still adherent cells during the process of detachment [24,32].

In summary, based on the present data we hypothesize that statins may facilitate cell

detachment and EMP release in order to preserve the overall condition and anti-

atherogenic properties of the vascular endothelium. We suggest that, in order to gain full

insight into the effects of compounds on endothelial cell biology, evaluation of adherent

cells, detached cells as well as EMP should be adopted as a general methodological

principle.


114

References

1. Collins R, Armitage J, Parish S, Sleigh P, Peto R. MRC/BHF heart protection study of cholesterol-lowering with simvastatin in 5963 people with diabetes: a randomised placebo-controlled trial. Lancet 2003;361:2005-16.

2. Heart Protection Study Collaborative Group. MRC/BHF heart protection study of cholesterol lowering with simvastatin in 20,536 high-risk individuals: a randomised placebo-controlled trial. Lancet 2002;360:7-22.

3. Combination Pharmacotherapy and Public Health Research Working Group. Combination pharmacotherapy for cardiovascular disease. Ann.Intern.Med. 2005;143:593-9.

4. LaRosa JC, Grundy SM, Waters DD, Shear C, Barter P, Fruchart JC, Gotto AM, Greten H, Kastelein JJ, Shepherd J, Wenger NK. Intensive lipid lowering with atorvastatin in patients with stable coronary disease. N. Engl. J. Med. 2005;352:1425-35.

5. Scandinavian Simvastatin Survival Study Group. Randomised trial of cholesterol lowering in 4444 patients with coronary heart disease: the Scandinavian Simvastatin Survival Study. Lancet 1994;344:1383-9.

6. Downs JR, Clearfield M, Weis S, Whitney E, Shapiro DR, Beere PA, Langendorfer A, Stein EA, Kruyer W, Gotto AM, Jr. Primary prevention of acute coronary events with lovastatin in men and women with average cholesterol levels: results of AFCAPS/TexCAPS. Air Force/Texas Coronary Atherosclerosis Prevention Study. JAMA 1998;279:1615-22.

7. Baigent C, Keech A, Kearney PM, Blackwell L, Buck G, Pollicino C, Kirby A, Sourjina T, Peto R, Collins R, Simes R. Efficacy and safety of cholesterol-lowering treatment: prospective meta-analysis of data from 90,056 participants in 14 randomised trials of statins. Lancet 2005;366:1267-78.

8. Colhoun HM, Betteridge DJ, Durrington PN, Hitman GA, Neil HA, Livingstone SJ, Thomason MJ, Mackness MI, Charlton-Menys V, Fuller JH. Primary prevention of cardiovascular disease with atorvastatin in type 2 diabetes in the Collaborative Atorvastatin Diabetes Study (CARDS): multicentre randomised placebo-controlled trial. Lancet 2004;364:685-96.

9. Vijan S, Hayward RA. Pharmacologic lipid-lowering therapy in type 2 diabetes mellitus: background paper for the American College of Physicians. Ann. Intern. Med. 2004;140:650-8.

10. Ferrara DE, Pierangeli SS. Diverse effects of statins on endothelial cells? Thromb. Haemost. 2005;93:186-8.

Chapter 5

115

11. Feng C, Ye C, Liu X, Ma H, Li M. Beta4 integrin is involved in statin-induced endothelial cell death. Biochem. Biophys. Res. Commun. 2004;323:858-64.

12. Tramontano AF, O'Leary J, Black AD, Muniyappa R, Cutaia MV, El Sherif N. Statin decreases endothelial microparticle release from human coronary artery endothelial cells: implication for the Rho-kinase pathway. Biochem. Biophys. Res. Commun. 2004;320:34-8.

13. Dimitrova Y, Dunoyer-Geindre S, Reber G, Mach F, Kruithof EK, de MP. Effects of statins on adhesion molecule expression in endothelial cells. J. Thromb. Haemost. 2003;1:2290-9.

14. Kaneta S, Satoh K, Kano S, Kanda M, Ichihara K. All hydrophobic HMG-CoA reductase inhibitors induce apoptotic death in rat pulmonary vein endothelial cells. Atherosclerosis 2003;170:237-43.

15. Kozai T, Eto M, Yang Z, Shimokawa H, Luscher TF. Statins prevent pulsatile stretch-induced proliferation of human saphenous vein smooth muscle cells via inhibition of Rho/Rho-kinase pathway. Cardiovasc. Res. 2005;68:475-82.

16. Eto M, Kozai T, Cosentino F, Joch H, Luscher TF. Statin prevents tissue factor expression in human endothelial cells: role of Rho/Rho-kinase and Akt pathways. Circulation 2002;105:1756-9.

17. Weis M, Heeschen C, Glassford AJ, Cooke JP. Statins have biphasic effects on angiogenesis. Circulation 2002;105:739-45.

18. Landmesser U, Bahlmann F, Mueller M, Spiekermann S, Kirchhoff N, Schulz S, Manes C, Fischer D, de GK, Fliser D, Fauler G, Marz W, Drexler H. Simvastatin versus ezetimibe: pleiotropic and lipid-lowering effects on endothelial function in humans. Circulation 2005;111:2356-63.

19. Takai Y, Sasaki T, Matozaki T. Small GTP-binding proteins. Physiol Rev. 2001;81:153-208.


21. Backman JT, Kyrklund C, Kivisto KT, Wang JS, Neuvonen PJ. Plasma concentrations of active simvastatin acid are increased by gemfibrozil. Clin. Pharmacol. Ther. 2000;68:122-9.

22. Sugimoto K, Ohmori M, Tsuruoka S, Nishiki K, Kawaguchi A, Harada K, Arakawa M, Sakamoto K, Masada M, Miyamori I, Fujimura A. Different effects


116

of St John's wort on the pharmacokinetics of simvastatin and pravastatin. Clin. Pharmacol. Ther. 2001;70:518-24.

23. Gniadecki R. Depletion of membrane cholesterol causes ligand-independent activation of Fas and apoptosis. Biochem. Biophys. Res. Commun. 2004;320:165-9.

24. Sebbagh M, Renvoize C, Hamelin J, Riche N, Bertoglio J, Breard J. Caspase-3-mediated cleavage of ROCK I induces MLC phosphorylation and apoptotic membrane blebbing. Nat. Cell Biol. 2001;3:346-52.

25. Rudel T, Bokoch GM. Membrane and morphological changes in apoptotic cells regulated by caspase-mediated activation of PAK2. Science 1997;276:1571-4.

26. Li X, Liu L, Tupper JC, Bannerman DD, Winn RK, Sebti SM, Hamilton AD, Harlan JM. Inhibition of protein geranylgeranylation and RhoA/RhoA kinase pathway induces apoptosis in human endothelial cells. J. Biol. Chem. 2002;277:15309-16.

27. De Staercke C, Phillips DJ, Hooper WC. Differential responses of human umbilical and coronary artery endothelial cells to apoptosis. Endothelium 2003;10:71-8.

28. Bombeli T, Karsan A, Tait JF, Harlan JM. Apoptotic vascular endothelial cells become procoagulant. Blood 1997;89:2429-42.

29. Hebert MJ, Gullans SR, Mackenzie HS, Brady HR. Apoptosis of endothelial cells is associated with paracrine induction of adhesion molecules. Am. J. Pathol. 1998;152:523-32.

30. Rikitake Y, Liao JK. Rho GTPases, statins, and nitric oxide. Circ. Res. 2005;97:1232-5.

31. Ray KK, Cannon CP. The potential relevance of the multiple lipid-independent (pleiotropic) effects of statins in the management of acute coronary syndromes. J. Am. Coll. Cardiol. 2005;46:1425-33.


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Chapter 6

Inhibition of microparticle release triggers

endothelial cell apoptosis and detachment

Mohammed N. Abid Hussein, Anita N. Böing, Augueste Sturk,

Chi M. Hau and Rienk Nieuwland

Thromb. Haemost. 2007;98:1096-1107

Inhibition of microparticle release

118

ABSTRACT

Background: Endothelial cell cultures contain caspase 3-containing microparticles

(EMP), which are reported to form during or after cell detachment.

Hypthesis: 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)-1 (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-1 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-1 -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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INTRODUCTION

Like other eukaryotic cells, endothelial cells release microparticles (MP; EMP:

endothelial microparticles) in vitro [1-3] 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

vitro [8]. 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 EMP [9,10]. 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

EMP [11], 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 3 [8]. 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-1 (IL-1 ), without or with widely used inhibitors of

microparticle release, i.e. Y-27632 and calpeptin [9,12,13].


Reagents and assays

Medium M199, penicillin, streptomycin and L-glutamine 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, USA). Human serum albumin (HSA) was obtained from Sanquin

(Amsterdam, The Netherlands). Recombinant human interleukin (IL)-1 was from Sigma

(St. Louis, MO, USA). Human recombinant basic fibroblast growth factor and epidermal


120

growth factor were from Invitrogen life technologies (Carlsbad, CA, USA). Collagenase

(type 1A) and staurosporin were from Sigma (St. Louis, MO, USA). Heparin (400 U/mL)

was obtained from Leo Pharma BV (Breda, The Netherlands), trypsin from Difco

Laboratories (Detroit, MI, USA), calpeptin from Calbiochem (La Jolla, CA, USA), and Y-

27632 from Tocris (Ellisville, MO, USA). Y-27632 is a specific inhibitor of Rho-

associated serine/threonine kinases I and II (i.e. ROCK I (p160ROCK, ROK ) and ROCK

II (Rho-kinase, ROK )), enzymes which are directly involved in the release of apoptotic

blebs [9,12]. Calpeptin inhibits calpain, a Ca2+-dependent protease, that plays a role in

(E)MP formation [13]. For Western blot analysis, anti-human caspase 3 monoclonal

antibody from Alexis Biochemicals (San Diego, CA, USA) 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 (Sparks, MD, USA).

Isolation, culture and treatment of human umbilical vein endothelial cells (HUVEC)

HUVEC were collected as described previously [3]. 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-1 (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 M; two hours

preincubation) and/or calpeptin (200 M; 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-1 , 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-1 (24 hours). Detached cells were resuspended in 10 mL fresh

culture medium (without staurosporin or IL-1 ), and numbers of detached endothelial

cells and EMP were determined at fixed time intervals (3-48 hours) by flow cytometry.

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Flow cytometric analysis of HUVEC

Conditioned media were collected and centrifuged (10 minutes, 180 g 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 PBS/FCSi (10% v/v). Both cell suspensions were centrifuged (10 minutes,

180 g and 4 °C) and pellets were resuspended in PBS/FCSi (1% v/v), and then again

centrifuged (10 minutes, 180 g 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 previously [8]. Intracellular caspase 3 was detected using the active caspase-3

MoAb apoptosis kit I from BD Pharmingen (San Diego, CA, USA). Samples were

analyzed in a FACSCalibur flowcytometer (Becton Dickinson; San Jose, CA, USA). 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, 17,570 g and 20 °C). Then, 975 L of supernatant was

removed and the pellet was resuspended in 225 L 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, 17,570 g and 20 °C), 225 L supernatant was removed and MP were diluted and

resuspended by adding 75 L PBS/citrate or perm/wash (0.1% v/v).

Flow cytometric analysis of EMP

EMP were analyzed in a FACSCalibur flow cytometer as described previously [3]. To

detect intravesicular caspase 3, MP (5 L) were diluted with 35 L 0.1% perm/wash

solution containing 2.5 mmol/L CaCl2 plus either anti-caspase 3-FITC (BD) or control

antibody, Ig-FITC (IQP; Groningen, The Netherlands). For annexin V staining, MP (5


122

L) were diluted with 35 L PBS containing 2.5 mmol/L CaCl2 (pH 7.4). Annexin V-

APC (Caltag Laboratories; Carlsbad, CA, USA; 5 L 20-fold prediluted) was added. To

remove the excess of unbound annexin V, 200 L PBS/calcium buffer (or 200 L 0.1%

perm/wash containing CaCl2 for intravesicular staining of caspase 3) was added and the

suspension was centrifuged for 30 minutes at 17,570 g and 20 °C. Finally, 200 L of

supernatant was removed, and EMP were resuspended with 300 L PBS/calcium or 300

L 0.1% perm/wash containing CaCl2. Previously, we demonstrated that numbers of EMP

(N) and detached cells highly correlate [8]. 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-1 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 L

(detached cells) and 800 L (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 L) and adherent cells (final volume 40 L). From the detached

cell lysate, 10 L 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, 17,570 g and 20

°C) and resuspended in 10 L PBS plus 10 L 2-fold concentrated reducing sample

buffer. Per EMP sample, 10 L 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, USA). 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

Chapter 6

123

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).

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

cultures contain 17 kDa caspase 3 [8]. 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-

1 (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.


All data were analyzed with GraphPad Prism for Windows, release 3.02 (San Diego,

CA, USA). Data from preliminary experiments regarding differences in numbers of

adherent cells, detached cells or EMP between control, staurosporin and IL-1 conditions

were analyzed by Wilcoxon matched pairs test (one-tailed analysis). Values are expressed

as median (range).


124

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-1 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-1 ) were calculated in the absence or presence

of 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-1 , numbers of detached cells (Figure 1C) and annexin V-binding

EMP (Figure 1E) increased (n=6). Neither staurosporin nor IL-1 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.

Chapter 6

020406080

100

%A

popt

osis

StaurosporinControl IL-1

B

D


20406080

100

% A

popt

osis

AP=0.015

Num

ber o

f adh

eren

t cel

ls

106

0123456

Control Staurosporin IL-1

E

Num

bero

f EM

P 10

6

P=0.031

P=0.031

02468

1012


C

P=0.031

P=0.015

Num

ber o

f det

ache

dce

lls

106


020406080

100

%A

popt

osis


B

D


20406080

100

% A

popt

osis

AP=0.015P=0.015

Num

ber o

f adh

eren

t cel

ls

106

0123456


E

Num

bero

f EM

P 10

6

P=0.031

P=0.031

02468

1012


C

P=0.031

P=0.015

P=0.031

P=0.015

Num

ber o

f det

ache

dce

lls

106


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-1 (5 ng/mL) for 24 hours. Adherent cells (A, B), detached

cells (C, D) and EMP (E) were isolated and analyzed as described in 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

(open bars; ‘early apoptosis’) or by staining for both annexin V and PI (dashed bars;

‘late apoptosis’). Bars indicate mean SD (n=3). In E, both annexin V-positive (dotted

bars) and annexin V-negative EMP (lower bars) are shown (n=6).

125


126

Effects of inhibitors of microparticle release on endothelial 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-1 ) 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-1 -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-1 . 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-

1 -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-1 , Figures 3 and 4, respectively), as well as

the presence of caspase 3.

Tabl

e 1.

Effe

cts o

f inh

ibito

rs o

f mic

ropa

rtic

le re

leas

e on

det

achm

ent a

nd E

MP

rele

ase

at 2

4 ho

urs.

Con

trol

P

Stau

rosp

orin

P

IL-1

P

Num

ber

of E

MP

x 10

6

No

inhi

bito

rs

1.1

0.2

-

4.5

2.7

-

3.1

1.7

-

Y-27

632

1.4

0.4

P

0.05

2.

8 0

.7

P0.

05

2.5

1.2

P

0.05

Cal

pept

in

1.4

0.6

P

0.05

3.

6 0

.4

P0.

05

2.4

1.8

P

0.05

Y-27

632

+ C

alpe

ptin

1.

4 0

.5

P0.

05

5.1

0.8

P

0.05

2.

6 1

.5

P0.

05

Det

ache

d ce

lls (%

from

tota

l)

No

inhi

bito

rs

5.7

2.2

15

.4

10.

1 19

.5

13.

6

Y-27

632

15.7

1

.3

P0.

05

29.1

4

.6

P0.

05

37.7

8

.2

P0.

05

Cal

pept

in

17.7

8

.2

P0.

05

71.2

8

.0

P0.

001

43.6

1

6.6

P0.

05

Y-27

632

+ C

alpe

ptin

26

.9

2.1

P

0.00

1 81

.4

7.2

P

0.00

1 56

.8

10.

3 P

0.05

Endo

thel

ial c

ells

wer

e in

cuba

ted

in th

e ab

senc

e (c

ontr

ol)

or p

rese

nce

of s

taur

ospo

rin

(200

nM

) or

IL-

1 (

5 ng

/mL)

, with

or

with

out Y

-

2763

2, c

alpe

ptin

or b

oth

(n=

3). A

fter 2

4 ho

urs,

deta

ched

cel

ls an

d EM

P w

ere

isol

ated

as d

escr

ibed

in M

ater

ials

and

Met

hods

. Det

achm

ent

is e

xpre

ssed

as %

of t

he to

tal n

umbe

r of c

ells

, i.e

. the

num

ber

of a

dher

ent a

nd d

etac

hed

cells

pre

sent

. Diff

eren

ces

betw

een

cond

ition

s with

and

with

out i

nhib

itor(

s) w

ere

anal

yzed

by

pair

ed t-

test

as

desc

ribe

d in

the

Stat

istic

al a

naly

sis

sect

ion

of M

ater

ials

and

Met

hods

. Dat

a ar

e

pres

ente

d as

mea

n ±

SD.

127


128

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.

Chapter 6

3 6 12 24 480

20

40

60

80

100 A

3 6 12 24 480

20

40

60

80

100B

3 6 12 24 480

20

40

60

80

100E

Time (hours)

0

2

4

6

8

3 6 12 24 48

H

% D

etac

hed

cells

% A

nnex

inV

-pos

itive

cel

ls%

Ann

exin

V-p

ositi

ve c

ells

Cas

pase

3-p

ositi

ve E

MP

x 10

6

0

10

20

30

40

50C

3 6 12 24 48

3 6 12 24 480

20

40

60

80

100F

Time (hours)3 6 12 24 48

0

20

40

60

80

100I

% C

aspa

se 3

-pos

itive

cel

ls%

Cas

pase

3-p

ositi

ve c

ells

% C

aspa

se 3

-pos

itive

EM

P

3 6 12 24 48 3 6 12 24 48

Caspase 3

+-

D

+-

Caspase 3

G

+-

Caspase 3

Time (hours)

J

3 6 12 24 48 3 6 12 24 48

3 6 12 24 48 3 6 12 24 48

3 6 12 24 480

20

40

60

80

100 A

3 6 12 24 480

20

40

60

80

100B

3 6 12 24 480

20

40

60

80

100E

Time (hours)

0

2

4

6

8

3 6 12 24 48

H

% D

etac

hed

cells

% A

nnex

inV

-pos

itive

cel

ls%

Ann

exin

V-p

ositi

ve c

ells

Cas

pase

3-p

ositi

ve E

MP

x 10

6

3 6 12 24 480

20

40

60

80

100 A

3 6 12 24 483 6 12 24 480

20

40

60

80

100

0

20

40

60

80

100 A

3 6 12 24 480

20

40

60

80

100B

3 6 12 24 483 6 12 24 480

20

40

60

80

100

0

20

40

60

80

100B

3 6 12 24 480

20

40

60

80

100E

3 6 12 24 483 6 12 24 480

20

40

60

80

100

0

20

40

60

80

100E

Time (hours)

0

2

4

6

8

3 6 12 24 48

H

0

2

4

6

8

0

2

4

6

8

3 6 12 24 483 6 12 24 48

H

% D

etac

hed

cells

% A

nnex

inV

-pos

itive

cel

ls%

Ann

exin

V-p

ositi

ve c

ells

Cas

pase

3-p

ositi

ve E

MP

x 10

6

0

10

20

30

40

50C

3 6 12 24 48

3 6 12 24 480

20

40

60

80

100F

Time (hours)3 6 12 24 48

0

20

40

60

80

100I

% C

aspa

se 3

-pos

itive

cel

ls%

Cas

pase

3-p

ositi

ve c

ells

% C

aspa

se 3

-pos

itive

EM

P

0

10

20

30

40

50C

3 6 12 24 480

10

20

30

40

50

0

10

20

30

40

50C

3 6 12 24 483 6 12 24 48

3 6 12 24 480

20

40

60

80

100F

3 6 12 24 483 6 12 24 480

20

40

60

80

100

0

20

40

60

80

100F

Time (hours)3 6 12 24 48

0

20

40

60

80

100I

3 6 12 24 483 6 12 24 480

20

40

60

80

100

0

20

40

60

80

100I

% C

aspa

se 3

-pos

itive

cel

ls%

Cas

pase

3-p

ositi

ve c

ells

% C

aspa

se 3

-pos

itive

EM

P%

Cas

pase

3-p

ositi

ve c

ells

% C

aspa

se 3

-pos

itive

cel

ls%

Cas

pase

3-p

ositi

ve E

MP

3 6 12 24 48 3 6 12 24 483 6 12 24 483 6 12 24 48 3 6 12 24 483 6 12 24 48

Caspase 3

+-

D

+-

Caspase 3

G

+-

Caspase 3

Time (hours)

J

3 6 12 24 48 3 6 12 24 483 6 12 24 483 6 12 24 48 3 6 12 24 483 6 12 24 48

3 6 12 24 48 3 6 12 24 483 6 12 24 483 6 12 24 48 3 6 12 24 483 6 12 24 48

Figure 2. Effects of inhibitors of microparticle release in endothelial cell cultures without external stress. 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

M) plus calpeptin (200 M). A. Fractions of detached cells. B and E show adherent (B) and detached (E) endothelial cell fractions binding annexin V as an indicator of apoptosis. C and F show adherent (C) and detached (F) endothelial cell fractions staining for intracellular caspase 3, whereas 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. H-J show the absolute numbers of caspase 3-containing EMP (H), the fractions of caspase 3-containing EMP (I) and western blots of EMP lysates of 17 kDa caspase 3 (J).

129


130

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 (3J, 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

Chapter 6

release. Also, the inhibitors of EMP release caused caspase 3 accumulation in detached

cells.

+-

Caspase 3

+-

Caspase 3

+-

Caspase 3

Time (hours)

D

G

J

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 24 480

20

40

60

80

100 A

% D

etac

hed

cells

% A

nnex

inV

-pos

itive

cel

ls%

Ann

exin

V-p

ositi

ve c

ells

Cas

pase

3-p

ositi

ve E

MP

x 10

6

3 6 12 24 480

20

40

60

80

100B

3 6 12 24 480

20

40

60

80

100E

Time (hours)

0

2

4

6

8

3 6 12 24 48

H

Time (hours)

% C

aspa

se 3

-pos

itive

cel

ls%

Cas

pase

3-p

ositi

ve c

ells

% C

aspa

se 3

-pos

itive

EM

P

0

10

20

30

40

50C

3 6 12 24 48

3 6 12 24 480

20

40

60

80

100F

3 6 12 24 480

20

40

60

80

100I

+-

Caspase 3

+-

Caspase 3

+-

Caspase 3

Time (hours)

D

G

J

3 6 12 24 48 3 6 12 24 483 6 12 24 483 6 12 24 48 3 6 12 24 483 6 12 24 48

3 6 12 24 48 3 6 12 24 483 6 12 24 483 6 12 24 48 3 6 12 24 483 6 12 24 48

3 6 12 24 48 3 6 12 24 483 6 12 24 483 6 12 24 48 3 6 12 24 483 6 12 24 48

3 6 12 24 480

20

40

60

80

100 A

3 6 12 24 483 6 12 24 480

20

40

60

80

100

0

20

40

60

80

100 A

% D

etac

hed

cells

% A

nnex

inV

-pos

itive

cel

ls%

Ann

exin

V-p

ositi

ve c

ells

Cas

pase

3-p

ositi

ve E

MP

x 10

6

3 6 12 24 480

20

40

60

80

100B

3 6 12 24 483 6 12 24 480

20

40

60

80

100

0

20

40

60

80

100B

3 6 12 24 480

20

40

60

80

100E

3 6 12 24 483 6 12 24 480

20

40

60

80

100

0

20

40

60

80

100E

Time (hours)

0

2

4

6

8

3 6 12 24 48

H

0

2

4

6

8

0

2

4

6

8

3 6 12 24 483 6 12 24 48

H

Time (hours)

% C

aspa

se 3

-pos

itive

cel

ls%

Cas

pase

3-p

ositi

ve c

ells

% C

aspa

se 3

-pos

itive

EM

P%

Cas

pase

3-p

ositi

ve c

ells

% C

aspa

se 3

-pos

itive

cel

ls%

Cas

pase

3-p

ositi

ve E

MP

0

10

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30

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50C

3 6 12 24 480

10

20

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50

0

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3 6 12 24 483 6 12 24 48

3 6 12 24 480

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20

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3 6 12 24 483 6 12 24 480

20

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0

20

40

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80

100I

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 M) plus calpeptin (200 M). A shows the fractions of detached cells. B and E show adherent (B) and detached (E) endothelial cell fractions binding annexin V. C and F show adherent (C) and detached (F) endothelial cell fractions staining for intracellular caspase 3, whereas 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. H-J show the absolute numbers of caspase 3-containing EMP (H), the fractions of caspase 3-containing EMP (I) and Western blots of EMP lysates of 17 kDa caspase 3 (J).

131


132

Effects of inhibitors of microparticle release in endothelial cell cultures in the

presence of additional external stress: IL-1

The IL-1 -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-1 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-1 , 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.

Chapter 6

+-

Caspase 3

+-

Caspase 3

Caspase 3

+-

Time (hours)

D

J

G

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

Time (hours)

% D

etac

hed

cells

% A

nnex

inV

-pos

itive

cel

ls%

Ann

exin

V-p

ositi

ve c

ells

Cas

pase

3-p

ositi

ve E

MP

x 10

6

3 6 12 24 480

20

40

60

80

100 A

3 6 12 24 480

20

40

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100B

3 6 12 24 480

20

40

60

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100E

0

2

4

6

8

3 6 12 24 48

H

Time (hours)

% C

aspa

se 3

-pos

itive

cel

ls%

Cas

pase

3-p

ositi

ve c

ells

% C

aspa

se 3

-pos

itive

EM

P

0

10

20

30

40

50C

3 6 12 24 48

3 6 12 24 480

20

40

60

80

100F

3 6 12 24 480

20

40

60

80

100I

+-

Caspase 3

+-

Caspase 3

Caspase 3

+-

Time (hours)

D

J

G

3 6 12 24 48 3 6 12 24 483 6 12 24 483 6 12 24 48 3 6 12 24 483 6 12 24 48

3 6 12 24 48 3 6 12 24 483 6 12 24 483 6 12 24 48 3 6 12 24 483 6 12 24 48

3 6 12 24 48 3 6 12 24 483 6 12 24 483 6 12 24 48 3 6 12 24 483 6 12 24 48

Time (hours)

% D

etac

hed

cells

% A

nnex

inV

-pos

itive

cel

ls%

Ann

exin

V-p

ositi

ve c

ells

Cas

pase

3-p

ositi

ve E

MP

x 10

6

3 6 12 24 480

20

40

60

80

100 A

3 6 12 24 480

20

40

60

80

100B

3 6 12 24 480

20

40

60

80

100E

0

2

4

6

8

3 6 12 24 48

H

Time (hours)

% D

etac

hed

cells

% A

nnex

inV

-pos

itive

cel

ls%

Ann

exin

V-p

ositi

ve c

ells

Cas

pase

3-p

ositi

ve E

MP

x 10

6

3 6 12 24 480

20

40

60

80

100 A

3 6 12 24 483 6 12 24 480

20

40

60

80

100

0

20

40

60

80

100 A

3 6 12 24 480

20

40

60

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100B

3 6 12 24 483 6 12 24 480

20

40

60

80

100

0

20

40

60

80

100B

3 6 12 24 480

20

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100E

3 6 12 24 483 6 12 24 480

20

40

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100

0

20

40

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80

100E

0

2

4

6

8

3 6 12 24 48

H

0

2

4

6

8

0

2

4

6

8

3 6 12 24 483 6 12 24 48

H

Time (hours)

% C

aspa

se 3

-pos

itive

cel

ls%

Cas

pase

3-p

ositi

ve c

ells

% C

aspa

se 3

-pos

itive

EM

P

0

10

20

30

40

50C

3 6 12 24 48

3 6 12 24 480

20

40

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100F

3 6 12 24 480

20

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100I

Time (hours)

% C

aspa

se 3

-pos

itive

cel

ls%

Cas

pase

3-p

ositi

ve c

ells

% C

aspa

se 3

-pos

itive

EM

P%

Cas

pase

3-p

ositi

ve c

ells

% C

aspa

se 3

-pos

itive

cel

ls%

Cas

pase

3-p

ositi

ve E

MP

0

10

20

30

40

50C

3 6 12 24 480

10

20

30

40

50

0

10

20

30

40

50C

3 6 12 24 483 6 12 24 48

3 6 12 24 480

20

40

60

80

100F

3 6 12 24 483 6 12 24 480

20

40

60

80

100

0

20

40

60

80

100F

3 6 12 24 480

20

40

60

80

100I

3 6 12 24 483 6 12 24 480

20

40

60

80

100

0

20

40

60

80

100I

Figure 4. Effects of inhibitors of microparticle release in endothelial cell cultures in the presence of external stress: IL-1 . Endothelial cell cultures (n=3) were incubated with additional external stress (IL-1 ) up to 48 hours, in the absence (open symbols) or presence (closed symbols) of Y-27632 (30 M) plus calpeptin (200 M). A shows the fractions of detached cells. B and E show adherent (B) and detached (E) endothelial cell fractions binding annexin V. C and F show adherent (C) and detached (F) endothelial cell fractions staining for intracellular caspase 3, whereas 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. H-J show the absolute numbers of caspase 3-containing EMP (H), the fractions of caspase 3-containing EMP (I) and Western blots of EMP lysates of 17 kDa caspase 3 (J).

133


134

The origin of EMP, attached or detached cells?

To further study the complex relationship between adherent cells, cell detachment and

release of EMP, we performed SEM on adherent and detached endothelial cells in the

three conditions studied (Figure 5). Adherent cells showed confluent, apparently healthy

monolayers in the three conditions studied (not shown). At higher magnifications,

however, some differences became apparent. Although adherent cells from control

cultures showed an intact monolayer (Figure 5A), some (adherent) cells showed extensive

blebbing (Figure 5E). In IL-1 -treated cultures, a few adherent cells showed signs of

retraction and showed extensive blebbing (Figure 5B). Futhermore, some adherent cells

were undergoing detachment (Figure 5F). In staurosporin-treated cultures, several

adherent cells showed a very different, ‘spongy’, morphology and again rather ‘local’

blebbing (Figure 5C). Detached cells (Figures 5D and 5H) showed the ‘spongy’

appearance. No apparent differences were present between detached cells in the three

conditions studied (data not shown). Some but not all detached cells showed extensive

blebbing (Figure 5H).

The SEM findings indicate EMP formation to occur on the surface of adherent as well

as detached endothelial cells in the three conditions studied.

135

Fig

ure

5. S

EM o

f ad

here

nt a

nd d

etac

hed

endo

thel

ial

cells

. SE

M w

as p

erfo

rmed

as

desc

ribed

in

Mat

eria

ls a

nd M

etho

ds o

f ad

here

nt

endo

thel

ial c

ells

from

con

trol

cul

ture

s (A

, E),

and

IL-1

-trea

ted

(B, F

) or

stau

rosp

orin

-trea

ted

(C, G

) cul

ture

s. D

etac

hed

cells

are

sho

wn

from

con

trol

cul

ture

s (D

, H).

AB

D

EF

GH

C

10 μ

M10

μM

10 μ

M10

μM

10 μ

M10

μM

10 μ

M10

μM


When the data on the number of EMP and the extent of cell detachment were

combined (Figure 6A), a very strong correlation was observed (r=0.825, P<0.0001),

suggesting that the majority of EMP originates from detaching and/or detached cells. To

further investigate this issue, we cultured isolated detached cells from control, IL-1 - or

staurosporin-treated cultures, and determined EMP formation in time (Figure 6B). In all

three conditions, 50% of the EMP had already been formed within 3 hours, and after 24-

48 hours EMP formation reached a plateau.

136

Control IL-1 Staurosporin0

100

200

300

400

500

Ann

exin

V-b

indi

ng E

MP

(%)

3 hours

12 hours24 hours48 hours

B

0 1 2 3 4 5Number of EMP x 106

60

20

40

60

80

100

Det

ache

d ce

lls (%

from

tota

l)

r = 0.825

P < 0.0001

A


100

200

300

400

500

Ann

exin

V-b

indi

ng E

MP

(%)

3 hours


B


100

200

300

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500

0

100

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300

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Ann

exin

V-b

indi

ng E

MP

(%)

3 hours


B


60

20

40

60

80

100

Det

ache

d ce

lls (%

from

tota

l)

r = 0.825

P < 0.0001

A


60

20

40

60

80

100

Det

ache

d ce

lls (%

from

tota

l)

r = 0.825

P < 0.0001

A

Figure 6. The origin of EMP, attached or detached cells? A. The numbers of EMP and % of detached cells, as presented in Table 1, show the strong correlation between EMP release and detachment. B shows EMP release from detached cells isolated from control cultures and from cultures treated with either IL-1 or staurosporin (n=3 for each condition). EMP data are expressed as % of the EMP count determined at 3 hours (for each condition).

Chapter 6

137

DISCUSSION

Our present study shows that there is a highly complex relationship between adherent

endothelial cells, detached endothelial cells and EMP formation. In both control cultures

and those treated with staurosporin or IL-1 , the SEM studies indicated low numbers of

adherent as well as detached cells to be invariably present with extensive signs of

blebbing. The correlation we observed between the numbers of EMP and detached cells

indicates the close relation between these two processes. The culture of detached cells

indicates that most EMP are formed from detached cells immediately upon or within a

few hours after detachment, with more EMP formation after prolonged detachment. These

data fit with the observation that a substantial fraction of the detached cells only becomes

apoptotic (annexin V-positive) after 6-12 hours (Figure 2E, 3E and 4E).

In the present study, we hypothesized that if indeed some of the EMP are released

directly from adherent cells to dispose of caspase 3 and thus prevent cell detachment, then

inhibition of this release should lead to intracellular accumulation of caspase 3 in adherent

endothelial cells. As a consequence, such an accumulation would cause an increased

tendency of adherent cells to detach. All experimental data in the present study can be

incorporated in the model presented in Figure 7. In the control condition (top), adherent

cells (green; center) show a basal formation of some active caspase 3 (C3, red) from its

inactive precursor procaspase 3 (PC3, black). Once sufficient caspase 3 is formed, caspase

3 is released into EMP, resulting in a cell solely containing procaspase 3 (adherent cell,

left). Other adherent cells in which caspase 3 is formed (arrow 1), release no or

insufficient EMP (adherent cell, right), resulting in accumulation of caspase 3, cell

detachment and release of microparticles or apoptotic bodies either (arrow 2) during or

after detachment. In the presence of Y-27632 and calpeptin (second panel), the release of

EMP from adherent cells becomes disturbed, and caspase 3 accumulates in adherent cells.

As a consequence the equilibrium (arrow 3) shifts towards accumulation of caspase 3,

detachment and release of microparticles of apoptotic bodies during or after detachment.

In cultures treated with staurosporin (third panel), the conversion of procaspase 3 into

caspase 3 is promoted. Increased formation of caspase 3 in adherent cells is expected to

trigger an increased release of (caspase 3-containing) EMP, but also (arrow 4) an

increased detachment and formation of microparticles of apoptotic bodies during or after


138

detachment is to be expected. In cultures treated with staurosporin, Y-27632 and

calpeptin, the adherent cells become unable to release microparticles, expected to result in

massive accumulation of caspase 3 in adherent cells and a very strong shift to the right,

i.e. massive cell detachment and release of microparticles of apoptotic bodies during or

after detachment. Confirmation of this model is provided by the finding that the inhibitors

of EMP formation, e.g. from 8.3 to 2.9 EMP/detached cell in control cultures, cause

accumulation of caspase 3 even in the detached cells (Figures 2F, 3F and 4F).

Our hypothesis of cell survival by EMP formation is substantiated by two other

findings. First, cells deficient in functional caspase 3 activity, including hepatocytes,

thymocytes or MCF-7 cells, do not or hardly release any MP, suggesting that caspase 3

contributes to or facilitates its own removal via formation of MP [15,16]. Second, when

we incubated endothelial cells with z-VAD, a general inhibitor of caspases, a 20 kDa

inactive form of caspase 3 accumulated in adherent endothelial cells. Concurrently, EMP

release became less, and 17 kDa caspase 3 could no longer be detected in EMP fractions

and detachment decreased (data not shown).

Interestingly, not only MP but also exosomes from non-apoptotic cells contain a

caspase 3-like activity [17]. These authors postulated that packaging active caspase in

exosomes may be a mechanism to ensure cell survival. Additional supportive evidence

that EMP formation/shedding contributes to cellular survival comes from an earlier study

showing that various cell types, including endothelial cells, escape from complement-

induced lysis by releasing complement C5b-9-enriched EMP [11]. Thus, two different

types of cell-derived vesicles, microparticles and exosomes, both facilitate removal of

potentially dangerous bio-molecules, and thus act as ‘garbage sacs’.

It should be noted that the inhibitors used in the present study, i.e. the ROCK

inhibitor Y-27632 and the calpain inhibitor calpeptin, exert various effects on

(endothelial) cells. For instance, O’Connell et al. showed that PIP2 incubated in platelet

membranes inhibited activation-induced microparticle formation >90%, presumably by

interaction of PIP2 with various membrane proteins. Incubation of permeabilized platelets

with purified calpain reduced PIP2 levels, whereas in the presence of calpeptin the PIP2

levels increased [18]. This indicates that calpeptin may influence PIP2 levels by calpain

independent from its calcium-dependent protease activity.

Chapter 6

139

Endothelial cells can also detach by caspase-independent mechanisms, and cell death

of detached cells is then a consequence of detachment (anoikis) of originally viable

endothelial cells. For example, Hasmim and coworkers showed that expression of

integrin cytoplasmic domains in endothelial cells induced caspase-independent

detachment that was followed by anoikis [19]. Our present data also indicate that detached

cells only become apoptotic some time after detachment.

Taken together, we postulate that the release of EMP is a general mechanism to

enable cells to dispose potentially harmful and redundant compounds, thereby supporting

cellular survival. Unfortunately, this hypothesis can not presently be tested directly,

because we have no specific markers available to distinguish EMP originating from

adherent cells and detaching/detached cells.


140

Figure 7. Schematic model of the complex relationship between adherent endothelial cells, detachment and release of EMP. This model shows the hypothesis that attached cells escape from accumulation of caspase 3, cell detachment and apoptosis by EMP formation. The model incorporates the data presented in Figures 1-4 and Table 1. Shown are four conditions, i.e. control and staurosporin-treated cultures in the absence or presence of Y-27632 plus calpeptin. Adherent cells are green, detached cells are orange, EMP from adherent cells are green and from detached cells orange. In the presence of Y-27632 plus calpeptin, release of caspase 3-containing EMP becomes inhibited per cell, resulting in accumulation of caspase 3 in adherent cells, cell detachment and accumulation of caspase 3 in detached cells. The number of arrows (1) indicate the extent of shift towards accumulation of caspase 3 and detachment, (2) shows the release of EMP during detachment and/or from detached cells, (3), (4) and (5) represent increased detachment (depicted by the thickness of the arrows) as well as release of EMP during detachment or from detached cells.

C3

PC3 C3PC3 C3

(C3)

PC3

C3C3

Control

(1)

(2)

(2)

?

PC3 C3PC3 C3

C3

PC3

C3C3

X

Control + Y27632 / Calpeptin

X(3)

X

X?

C3

C3

(1)

C3

PC3 C3PC3 C3

(C3)

PC3

C3C3

C3C3 C3

Staurosporin

(4)(1)

?

PC3 C3 PC3 C3

C3

PC3

C3

X

C3 C3

C3Staurosporin + Y27632 / Calpeptin

X(5)

C3X

?

C3

(1)

X

C3

PC3 C3PC3 C3

(C3)

PC3

C3C3

Control

(1)

(2)

(2)

?

PC3 C3PC3 C3

C3

PC3

C3C3

(3)

X

X?

X

Control + Y27632 / Calpeptin

X

C3

C3

(1)

C3

PC3 C3PC3 C3

(C3)

PC3

C3C3

C3C3 C3

Staurosporin

(4)(1)

?

PC3 C3 PC3 C3

C3

PC3

C3

X

C3 C3

C3Staurosporin + Y27632 / Calpeptin

X(5)

C3X

?

C3

(1)

X

Chapter 6

141

References







7. Mallat Z, Hugel B, Ohan J, Lesèche G, Freyssinet JM, Tedgui A. Shed membrane microparticles with procoagulant potential in human atherosclerotic plaques. Circulation 1999;99:348-53.


9. Sebbagh M, Renvoize C, Hamelin J, Riche N, Bertoglio J, Breard J. Caspase-3-mediated cleavage of ROCK I induces MLC phosphorylation and apoptotic membrane blebbing. Nat. Cell Biol. 2001;3:346-52.


11. Hamilton KK, Hattori R, Esmon CT, Sims PJ. Complement proteins C5b-9 induce vesiculation of the endothelial plasma membrane and expose catalytic


142

surface for assembly of the prothrombinase enzyme complex. J. Biol. Chem. 1990;265:3809-14.

12. Coleman ML, Sahai EA, Yeo M, Bosch M, Dewar A, Olson MF. Membrane blebbing during apoptosis results from caspase-mediated activation of ROCK I. Nat. Cell Biol. 2001;3:339-45.

13. Miyoshi H, Umeshita K, Sakon M, Imajoh-Ohmi S, Fujitani K, Gotoh M, Oiki E, Kambayashi J, Monden M. Calpain activation in plasma membrane bleb formation during tert-butyl hydroperoxide-induced rat hepatocyte injury. Gastroenterology 1996;110:1897-904.

14. van Berkel AM, van MJ, Groen AK, Bruno MJ. Mechanisms of biliary stent clogging: confocal laser scanning and scanning electron microscopy. Endoscopy 2005;37:729-34.

15. Janicke RU, Sprengart ML, Wati MR, Porter AG. Caspase-3 is required for DNA fragmentation and morphological changes associated with apoptosis. J. Biol. Chem. 1998;273:9357-60.

16. Zheng TS, Schlosser SF, Dao T, Hingorani R, Crispe IN, Boyer JL, Flavell RA. Caspase-3 controls both cytoplasmic and nuclear events associated with Fas-mediated apoptosis in vivo. Proc. Natl. Acad. Sci. U. S. A 1998;95:13618-23.

17. de Gassart A, Geminard C, Fevrier B, Raposo G, Vidal M. Lipid raft-associated protein sorting in exosomes. Blood 2003;102:4336-44.

18. O'Connell DJ, Rozenvayn N, Flaumenhaft R. Phosphatidylinositol 4,5-bisphosphate regulates activation-induced platelet microparticle formation. Biochemistry 2005;44:6361-70.

19. Hasmim M, Vassalli G, Alghisi GC, Bamat J, Ponsonnet L, Bieler G, Bonnard C, Paroz C, Oguey D, Ruegg C. Expressed isolated integrin beta1 subunit cytodomain induces endothelial cell death secondary to detachment. Thromb. Haemost. 2005;94:1060-70.

143

Chapter 7

General discussion and summary


144

Aim of the thesis In this thesis, we characterized endothelial cell-derived microparticles (antigenic and

phospholipid composition; Chapters 2 and 3, respectively), investigated their relationship

with endothelial detachment (Chapters 4 and 5), and studied their putative functions

(thrombus formation and cellular survival; Chapters 3 and 6, respectively).

Identification of endothelial cell-derived microparticles (EMP) The cellular origin of microparticles (MP) is usually established by measuring the

exposure of cell-specific antigens. Accordingly, in most studies monoclonal antibodies are

used that either recognize such (cell-specific) antigens (positive identification), or are

directed against antigens exposed on MP of non-endothelial origin (exclusion). Both

approaches are based on the assumption that similar (cell-specific) antigens are exposed

on the parent cells and their MP. An example of positive identification is the exposure of

the LPS-receptor CD14 on both monocytes and their MP, hence both can be identified

using a CD14 antibody. As CD14 is a specific antigen solely present on monocytes, MP

from monocytes can be specifically detected by e.g. flow cytometry in a MP population

from various cellular sources. An example of exclusion is the use of anti-CD42

(glycoprotein Ib) in order to distinguish between EMP (CD42-) and PMP (CD42+) in

plasma samples. The lack of consensus between different investigators to detect EMP ex

vivo is clearly illustrated in Table 1 (Introduction; page 18), which shows more than 20

different antibodies or combinations thereof that have been used to identify EMP, either

by positive identification (e.g. CD146, CD62E and CD144) or exclusion (e.g.

CD31+/CD42-).

The detection of EMP is complicated due to various reasons. In Chapter 2 we

showed that the integrin 3 (vitronectin receptor; CD51, CD61) is abundantly exposed

on (human umbilical vein) endothelial cells. In contrast, 3 is only detectable on minor

subpopulations of EMP under these conditions. In addition, we demonstrated that platelet-

endothelial cell adhesion molecule-1 (PECAM-1; CD31), which is often used to identify

EMP ex vivo, is also exposed on the much more abundant platelet-derived MP (PMP).

Furthermore, EMP in our model system, i.e. human umbilical vein endothelial cells

(HUVEC) treated with a single agonist (interleukin-1 ; IL-1 ), display a substantial

Chapter 7

145

antigenic variation. Thus, in vivo, where various types of endothelial cells coexist in a

continuously changing and dynamic environment, a multitude of different populations of

EMP are expected to exist, which may show substantial variation in their antigenic

profile. Therefore, the quest for a single and universal marker or even a combination of

markers that identify all EMP ex vivo may not prove successful.

E-selectin reflects the activation status of endothelial cells E-selectin is a cell adhesion receptor that is only produced and expressed by activated

endothelial cells. Chapter 2 shows that a subpopulation (<60%) of EMP from IL-1 -

activated human umbilical vein endothelial cells exposes E-selectin. This exposure not

only confirms the endothelial origin, but also reflects the activation status of the parent

cells during release. Since E-selectin is not exposed on PMP, this marker can be used to

identify EMP originating from activated endothelial cells in biological samples. It should

be mentioned that in the experiments described in Chapter 2 approximately 50% of the

E-selectin antigen was associated with EMP (data not shown). Thus, soluble (s) E-selectin

may consist of coexisting forms of both EMP-associated and non-EMP-associated (‘truly

soluble’) E-selectin. Previously we studied the occurrence of MP-associated E-selectin in

plasma samples of patients suffering from sepsis and multiple-organ dysfunction. In these

samples substantially elevated levels of the sE-selectin antigen were observed, but the

fraction of sE-selectin associated with (E)MP was negligible [1]. At present it is unknown

to which extent selectins or other soluble adhesion receptors are associated with (E)MP in

vivo.

Phospholipid composition of EMPIn Chapter 3 it is shown that EMP from IL-1 -activated endothelial cells are

enriched in the aminophospholipids phosphatidylserine (PS) and phosphatidy-

lethanolamine (PE). This implicates that sorting of phospholipids into EMP is dependent

on the activation status of their parent cells. Since we determined the phospholipid

composition of total EMP fractions, we do not know whether all EMP have a similar

phospholipid composition. This may be relevant, since in Chapter 2 we showed that upon

activation with IL-1 , less than 20% of the EMP stained for the TF antigen. Whether or


146

not the phospholipid composition of TF-exposing EMP differs from that of non-TF-

exposing EMP, remains to be investigated.

To which extent the observed changes in the phospholipid composition of EMP in

Chapter 3 are paralleled by changes in surface expression of these phospholipids is

equally unclear.

Relationship between EMP, detached and adherent endothelial cells As outlined in the Introduction (page 16), detached endothelial cells, adherent

endothelial cells and EMP coexist in vitro and in vivo. An unanswered question, however,

is to which extent EMP originate from detached and/or adherent endothelial cells. In

Chapter 4 we studied the relation between EMP, detached and adherent endothelial cells

in vitro. We found a strong positive correlation between the numbers of detached

endothelial cells and EMP, and both contained caspase 3 which was not detectable in

adherent cells. This led us to postulate that most EMP may have originated from detached

endothelial cells under these conditions. However, we did not exclude the possibility that

a fraction of EMP, either containing or devoid of caspase 3, may also have originated

from adherent endothelial cells.

To further elucidate the complex relation between EMP, detached endothelial cells

and adherent endothelial cells, we studied the effects of the cholesterol-lowering drug

simvastatin. In vivo, simvastatin is thought to improve the “overall condition” of the

endothelium, but in vitro it produces conflicting results. Chapter 5 shows that whereas

clinical doses of simvastatin did not affect adherent endothelial cells, marked elevations of

both detached endothelial cells and EMP were observed. Both detachment and EMP

release were reversed by restoring cholesterol biosynthesis or prenylation, again

indicating that detachment and EMP release are closely related processes. From these data

it can be concluded that a comprehensive insight into the true status of the endothelial

cells -and possibly of the endothelium- can only be obtained when also detached

(circulating) endothelial cells as well as EMP are taken into account.

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Possible functions of EMP Thrombus formation

Many studies focussed on the occurrence of (E)MP in several diseases, and because

of the increased presence of especially PMP in such conditions and their procoagulant ex

vivo, a thrombogenic role was proposed. Few studies investigated the possible functions

of EMP ex vivo. In Chapter 3 we investigated the ability of EMP to trigger thrombus

formation in a rat venous stasis model. We demonstrated that EMP from activated

endothelial cells trigger thrombus formation in a TF-dependent manner. Since such EMP

were also enriched in both PS and PE, both cofactors of TF-initiated coagulation, they

may be especially prone to initiate coagulation. In line with these data, we previously

demonstrated that TF-exposing MP from human pericardial wound blood, mainly

consisting of PMP and erythrocyte-derived MP, also trigger TF-dependent thrombus

formation in the same animal model [2].

Cell survival

Chapter 4 shows that EMP from viable cultures of human endothelial cells

invariably contain caspase 3. This finding suggests that continuous shedding of caspase 3-

containing vesicles may be an intrinsic component of an ongoing physiological process.

Similarly, in a recent study conditioned media of viable cells were also shown to contain

exosomes, which contained a caspase 3-like activity [3]. These authors postulated that

packaging active caspase in exosomes may be a mechanism to ensure cell survival. In

Chapter 6 it is hypothesized that at least part of caspase 3-containing EMP may directly

originate from adherent endothelial cells. By disposing the potentially dangerous caspase

3 via EMP release, endothelial cells may ensure their survival. The data presented in this

chapter demonstrate that part of the EMP are indeed likely to originate from adherent

cells. Furthermore, inhibition of EMP release results in accumulation of caspase 3,

detachment and apoptosis, thereby confirming our hypothesis.


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Other putative functions

Recently, Del Conde et al. showed that TF-exposing MP from monocytes may fuse

with activated platelets, thereby transferring coagulation-active TF [4]. Because

membrane fusion is promoted by PS, they postulated that TF-exposing MP are enriched in

PS. Our data in Chapter 3 support their assumption. If true, then transferring TF may also

pass on the various functions of TF other than coagulation, including angiogenesis, signal

transduction and protection against apoptosis.

Isolated fractions of MP from plasma samples of preeclamptic patients, acute

myocardial infarction or end-stage renal failure impair endothelium-dependent

vasodilatation [5-7]. These isolated fractions contain MP originating from various cell

types, including endothelial cells. Recent evidence suggests that especially EMP may play

a role in this impairment. First, isolated EMP (but not PMP) from patients suffering from

end-stage renal failure decreased endothelial NO release [7]. Second, in vitro generated

EMP impair endothelium-dependent vasodilatation [8-10]. Third, circulating levels of

EMP correlate with the loss of flow-mediated dilation in both patients with end-stage

renal failure and in patients with diabetes type 2 [7,11].

The mechanism(s) by which EMP or other types of MP exert their effect on the

endothelium is unknown. One possible mechanism is that membrane fusion between

(E)MP and endothelial cells may result in delivering intravesicular components, such as

caspase 3, into the cytosol. Previously, we showed that isolated fractions of MP from

preeclamptic patients did not affect gene expression of endothelial cells [12]. This result

was not anticipated, given the rapidly accumulating evidence that EMP exert various

effects on endothelial cells and the endothelium. This may be explained by our use of

venous endothelial cells (HUVEC) versus arterial endothelial cells that are used in

contractility studies. Another possibly explanation may be the absence (our experiments)

or presence (contractility studies) of shear, which may be a prerequisite for the interaction

between EMP and endothelial cells. Recently, an intriguing study showed the transfer of

mRNA from embryonic stem cell-derived MP to hematopoietic progenitor cells, thereby

promoting their cell growth and differentiation [13].

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Clinical relevance and future directions Throughout the literature, the occurrence of EMP in disease states has been associated

with endothelial impairment (dysfunction). In this thesis we show that the release of EMP

may also be beneficial to the endothelial cells (Chapter 6), and we demonstrate that

increases of EMP release do not necessarily indicate endothelial impairment (Chapter 5).

Evidence is accumulating that not only endothelial cells but also other cell types use

vesicles as transportation vehicles. Such vehicles can be used to remove waste or other

(potentially) harmful compounds, and contribute to intercellular communication.

Gram-negative bacteria release vesicles containing communication signals and toxins

directed against other bacteria and host cells, implicating that the release of vesicles may

be a highly conserved phenomenon. Experiments with the gram-negative bacterium

Pseudomanas aeruginosa showed that a signal molecule that was packaged into vesicles

facilitated its own packaging [14,15]. Our present findings regarding the packaging of

caspase 3 in (E)MP is in line with this concept. From the literature it is known that a

human breast cancer cell line, MCF-7, is deficient of caspase 3. Since these cells do not

release MP, the presence of caspase 3 seems to be a prerequisite for MP release. When

MCF-7 cells are transfected with a caspase 3 construct, MP release was clearly

demonstrable [16]. Similar studies performed recently in our laboratory showed that these

MCF-7-derived MP also contain caspase 3. Altogether, similar to the prokaryotic signal

molecule, also caspase 3 seems to be involved in its own packaging into MP. These

findings support the concept that different cell types may use MP as vehicles in order to

remove caspase 3. To which extent also other types of vesicles, such as exosomes,

contribute to this process remains to be investigated.

Throughout the time course of the thesis our focus shifted from merely identification

and functional properties of EMP to the more central question why (endothelial) cells

release MP or other types of vesicles into their environment. The data presented in this

thesis support the notion that the (clinical) relevance of circulating EMP as biomarkers,

i.e. markers ‘simply’ reflecting the status of the endothelium, may be insufficient. For

instance, although simvastatin-treated endothelial cell cultures contained increased

numbers of EMP, the adherent endothelial cells remained viable throughout the

experiments (Chapter 5). Moreover, evidence is provided that at least in vitro, EMP are


150

closely associated with detachment, thus further complicating the presumed ‘direct’

relationship between (adherent and/or detached) endothelial cells and EMP.

During the last decade, studies on various types of cell-derived vesicles, including

MP and exosomes, have regained scientific and clinical attention. The current view,

especially within the medical field, has changed from regarding cell-derived vesicles as a

mere artefact to a genuine acceptance of their existence in vivo. Future developments are

to be expected with regard to standardization of isolation and detection protocols of

(E)MP. The relevance of measuring (E)MP in clinical samples as markers reflecting e.g.

cellular dysfunction, disease state, coagulation or inflammation, needs further

confirmation. Based on recent literature and data presented in this thesis, it is proposed

that the release of cell-derived vesicles may represent a conserved and fundamental cell

biological phenomenon essential for survival and communication.

Taken together, it is not only important to isolate vesicles and to study their myriad of

functions in vitro and in vivo, but equally important to tackle the more fundamental

question why cells release such ‘multi-purpose vesicles’ into their environment.

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References

1. Osmanovitch N, Romijn FPHTM, Joop K, Sturk A, Nieuwland R. Soluble selectins in sepsis: microparticle-associated, but only to a minor degree. Thromb. Haemost. 2000;84:731-2.






7. Amabile N, Guerin AP, Leroyer A, Mallat Z, Nguyen C, Boddaert J, London GM, Tedgui A, Boulanger CM. Circulating endothelial microparticles are associated with vascular dysfunction in patients with end-stage renal failure. J. Am. Soc. Nephrol. 2005;16:3381-8.

8. Densmore JC, Signorino PR, Ou J, Hatoum OA, Rowe JJ, Shi Y, Kaul S, Jones DW, Sabina RE, Pritchard KA, Jr., Guice KS, Oldham KT. Endothelium-derived microparticles induce endothelial dysfunction and acute lung injury. Shock 2006;26:464-71.

9. Esposito K, Ciotola M, Schisano B, Gualdiero R, Sardelli L, Misso L, Giannetti G, Giugliano D. Endothelial microparticles correlate with endothelial dysfunction in obese women. J. Clin. Endocrinol. Metab. 2006;91:3676-9.



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12. Lok CA, Böing AN, Reitsma PH, van der Post JA, van BE, Boer K, Sturk A, Nieuwland R. Expression of inflammation-related genes in endothelial cells is not directly affected by microparticles from preeclamptic patients. J. Lab Clin. Med. 2006;147:310-20.

13. Ratajczak J, Miekus K, Kucia M, Zhang J, Reca R, Dvorak P, Ratajczak MZ. Embryonic stem cell-derived microvesicles reprogram hematopoietic progenitors: evidence for horizontal transfer of mRNA and protein delivery. Leukemia 2006;20:847-56.

14. Winans SC. Microbiology: bacterial speech bubbles. Nature 2005;437:330.

15. Mashburn LM, Whiteley M. Membrane vesicles traffic signals and facilitate group activities in a prokaryote. Nature 2005;437:422-5.


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Chapter 8

Algemene discussie en samenvatting


154

Doel van dit proefschrift In dit proefschrift zijn de antigene “opmaak” (hoofdstuk 2) en fosfolipiden-

samenstelling (hoofdstuk 3) van micropartikels (MP) van endotheelcellen (EMP)

onderzocht. Daarnaast is de relatie bestudeerd tussen de vorming van micropartikels door

endotheelcellen en hun loslaten van de kweekbodem (hoofdstukken 4 en 5). Ook is het

vermogen van EMP onderzocht bij het initiëren van een trombus in vivo (hoofdstuk 3) en

is hun bijdrage onderzocht aan het mogelijk beschermen van de endotheelcel tegen

apoptose (hoofdstuk 6).

Identificeren van EMP De cellulaire herkomst van MP wordt meestal vastgesteld door de aanwezigheid van

unieke en veelal celspecifieke oppervlakte merkers te meten. In de meeste

wetenschappelijke studies worden dan ook monoklonale antistoffen gebruikt die zijn

gericht tegen dergelijke specifieke antigenen (positieve identificatie). Ook kunnen

antistoffen worden gebruikt die juist gericht zijn tegen antigenen die niet op EMP worden

geëxposeerd (exclusie). Zowel positieve identificatie als exclusie experimenten gaan uit

van de veronderstelling dat dezelfde antigenen aanwezig zijn op de cellen en de van deze

cellen afkomstige MP.

Een voorbeeld van een merker die wordt gebruikt voor positieve identificatie is de

endotoxine receptor (CD14), die alleen aanwezig is op monocyten en op MP van deze

cellen (monocyten-MP; MMP). Zowel monocyten als MMP kunnen dus worden

geïdentificeerd door gebruik te maken van een antistof tegen CD14. Omdat CD14

exclusief wordt geëxposeerd op MMP, kunnen deze positief worden herkend in een

mengsel van MP afkomstig van verschillende soorten cellen. De meestal hiervoor

gebruikte techniek is flowcytometrie. Een voorbeeld van exclusie is het gebruik van een

antistof gericht tegen glycoproteïne Ib (CD42). Gebruikmakend van een anti-CD42

antistof kunnen, bijvoorbeeld in een plasmamonster, CD42-negatieve (EMP) en CD42-

positieve MP (MP die van bloedplaatjes afkomstig zijn; PMP) van elkaar worden

onderscheiden. Het gebrek aan consensus tussen onderzoekers over de wijze waarop EMP

in menselijke lichaamsvloeistoffen het best kunnen worden geïdentificeerd, blijkt

duidelijk uit Tabel 1 (pagina 18). Deze tabel laat zien dat meer dan 20 verschillende

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antistoffen of combinaties van antistoffen zijn gebruikt om EMP te identificeren, hetzij

door positive identificatie (bijvoorbeeld CD146, CD62E en CD144) of door exclusie

(bijvoorbeeld CD31+/CD42-).

Om verschillende redenen zijn EMP moeilijk te identificeren. In hoofdstuk 2 wordt

aangetoond dat de vitronectine receptor, ook wel integrine v 3 (CD51, CD61) genoemd,

is geëxposeerd op menselijke navelstrengendotheelcellen, terwijl deze receptor niet of

nauwelijks op EMP aanwezig is. Om EMP te identificeren worden vaak antistoffen

gebruikt die gericht zijn tegen de adhesie receptor PECAM-1 (CD31; platelet-endothelial

cell adhesion molecule-1). In hoofdstuk 2 wordt aangetoond dat deze receptor niet alleen

op EMP maar ook op PMP aanwezig is. Dit maakt identificatie van EMP in humane

plasmamonsters lastig, omdat PMP veel algemener zijn dan EMP. Ook laat dit hoofdstuk

zien dat EMP in het door ons gebruikte model systeem, dat willen zeggen gekweekte

humane navelstrengendotheelcellen die worden gestimuleerd met de ontststekin-

gsmediator interleukine-1 (IL-1 ), een grote variatie laten zien in hun antigene opmaak.

Omdat in vivo verschillende soorten endotheelcellen voorkomen en deze zich in een

steeds veranderende omgeving bevinden, mag worden verondersteld dat ook ex vivo tal

van populaties EMP voorkomen die onderling een aanzienlijke variatie kunnen vertonen

in hun antigene opmaak. Samengevat, de zoektocht naar een universele merker of

combinaties van merkers die ex vivo alle EMP zou kunnen identificeren, is dan ook

uitzonderlijk complex.

E-selectine weerspiegelt endotheelcel activatie E-selectine (CD62E) is een cel-specifieke adhesie receptor die alleen op

gestimuleerde endotheelcellen tot expressie wordt gebracht. Hoofdstuk 2 laat zien dat een

subpopulatie van EMP, afkomstig van IL-1 -gestimuleerde endotheelcellen, E-selectine

exposeert. Deze expositie bevestigt niet alleen hun cellulaire herkomst, maar geeft tevens

aan dat de endotheelcellen geactiveerd waren tijdens het afsnoeren van EMP. E-selectine

wordt niet op PMP gevonden. Deze merker kan dus worden gebruikt om EMP, afkomstig

van geactiveerde endotheelcellen, positief te identificeren. Uit de experimenten,

beschreven in hoofdstuk 2, bleek dat ongeveer 50% van het E-selectine antigeen in het

kweeksupernatant geassocieerd was met EMP. Met andere woorden, het niet-


156

celgebonden, zogenaamd “soluble” E-selectine omvat ten minste twee vormen die

tegelijkertijd aanwezig kunnen zijn: het EMP-geassocieerde en het niet-EMP-

geassocieerde (het echt “soluble”) E-selectine. In een eerdere studie is door ons de

aanwezigheid onderzocht van MP-geassocieerd E-selectine in plasmamonsters van

patiënten met sepsis en meervoudig orgaanfalen. Deze plasmamonsters bevatten

verhoogde concentraties van het E-selectine antigeen. De hoeveelheid E-selectine die

geassocieerd was met EMP in deze patiënten bleek echter klein te zijn [1]. In welke mate

selectines of andere niet cel-gebonden receptoren geassocieerd zijn met (E)MP in vivo is

onbekend.

Fosfolipidensamenstelling van EMP In hoofdstuk 3 wordt aangetoond dat EMP, afkomstig van IL-1 -gestimuleerde

endotheelcellen, verrijkt zijn in fosfatidylserine (PS) en fosfatidylethanolamine (PE). De

fosfolipidensamenstelling van EMP is dus mede afhankelijk is van activatie van

endotheelcellen. Omdat de fosfolipidensamenstelling van alle EMP is gemeten, kan niet

worden uitgesloten dat er verschillen bestaan in fosfolipidensamenstelling tussen EMP.

Dit kan relevant zijn, omdat bijvoorbeeld in hoofdstuk 2 is aangetoond dat minder dan

20% van de EMP detecteerbare hoeveelheden van het weefselfactor antigeen exposeert na

stimulering van de endotheelcellen met IL-1 . In hoeverre de fosfolipidensamenstelling

van weefselfactor-exposerende EMP verschilt van EMP die geen weefselfactor exposeren,

moet nader worden onderzocht. Ook blijft de vraag onbeantwoord in hoeverre de

waargenomen veranderingen in de totale fosfolipidensamenstelling ook het lipide

oppervlak van de MP betreffen (hoofdstuk 3).

Relatie tussen EMP, niet-adherente en adherente endotheelcellen Zoals in de inleiding van dit proefschrift al is aangegeven (pagina 16), komen zowel

in vitro als in vivo niet-adherente (“losse”) endotheelcellen, adherente (“vaste”)

endotheelcellen en EMP naast elkaar voor. In hoeverre EMP afkomstig zijn van niet-

adherente endotheelcellen en/of van adherente endotheelcellen, is onbekend. In hoofdstuk

4 hebben wij in vitro de relatie onderzocht tussen EMP, niet-adherente en adherente

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endotheelcellen. Er bleek een sterke positieve correlatie te zijn tussen de aantallen niet-

adherente endotheelcellen en EMP. Ook bleken, in tegenstelling tot adherente

endotheelcellen, zowel niet-adherente endotheelcellen als EMP caspase 3 te bevatten.

Naar aanleiding van deze resultaten hebben wij gepostuleerd dat de meeste EMP

afkomstig zijn van niet-adherente endotheelcellen. Toch sluiten deze resultaten niet uit dat

een deel van de EMP, wel of niet caspase 3 bevattend, afkomstig is van adherente

endotheelcellen.

Om meer inzicht te krijgen in de ingewikkelde relatie tussen EMP, niet-adherente en

adherente endotheelcellen, zijn de effecten onderzocht van het cholesterolverlagende

medicijn simvastatine. Simvastatine wordt verondersteld in vivo de conditie van het

endotheel te verbeteren, maar in vitro zijn de resultaten tegenstrijdig. In hoofdstuk 5

wordt aangetoond dat simvastatine, in concentraties die ook in de kliniek worden

toegepast, adherente endotheelcellen niet lijkt te beïnvloeden. Wel werd een opvallende

toename waargenomen van de aantallen niet-adherente endotheelcellen en EMP. Beide

effecten konden weer ongedaan worden gemaakt door de biosynthese van cholesterol te

herstellen of de prenylering van eiwitten weer mogelijk te maken. Deze resultaten

bevestigen dat het loslaten van endotheelcellen en het afsnoeren van EMP nauw-

geassocieerde processen zijn. Deze resultaten laten ook zien dat om een volledig inzicht te

krijgen in de conditie van de adherente endotheelcellen, en dus mogelijk ook van het

endotheel in vivo, niet alleen naar adherente endotheelcellen moet worden gekeken, maar

ook naar de niet-adherente, circulerende endotheelcellen en EMP.

Mogelijke functies van EMPTrombus vorming

Tot nu toe hebben veel studies zich gericht op de aanwezigheid van (E)MP in diverse

ziekten. Omdat in veel studies een toename van PMP is gevonden en deze PMP

stollingsbevorderend zijn ex vivo, wordt algemeen verondersteld dat (E)MP ook een rol

spelen bij de stolling in vivo. In hoofdstuk 3 is onderzocht of EMP het ontstaan van een

trombus kunnen initiëren in een proefdiermodel (veneus stase model; rat). Hierbij is

aangetoond dat EMP van gestimuleerde endotheelcellen in staat zijn weefselfactor-

afhankelijke trombusvorming te initiëren. Omdat deze EMP ook verrijkt zijn in PS en PE,


158

beide cofactoren van weefselfactor-gemedieerde stolling, mag worden aangenomen dat

dergelijke EMP speciaal zijn toegerust om in vivo de stolling te initiëren. Deze bevinding

komt overeen met eerdere onderzoeksresultaten van onze groep, waarin werd aangetoond

dat ook weefselfactor-exposerende MP, afkomstig uit humaan wondbloed, weefselfactor-

afhankelijke stolling initiëren in hetzelfde proefdiermodel [2].

Cellulaire overleving

Hoofdstuk 4 laat zien dat in het kweekmedium van navelstrengendotheelcellen altijd

caspase 3-bevattende EMP aanwezig zijn. Dit zou geïnterpreteerd kunnen worden als het

resultaat van een continu fysiologisch proces. Onlangs is in een soortgelijke studie

aangetoond dat kweekmedia van diverse soorten gezonde cellen ook een ander type

“vesicles” kunnen bevatten. Deze “vesicles”, exosomen, blijken ook een caspase 3-achtige

activiteit te bevatten [3]. De auteurs van dit artikel hebben verondersteld dat het

verpakken van een actief caspase in exosomen een mechanisme kan zijn dat bijdraagt aan

cellulaire overleving. In hoofdstuk 6 wordt verondersteld dat een deel van de caspase 3-

bevattende EMP afkomstig kan zijn van adherente endotheelcellen. Door het voor de cel

potentieel gevaarlijke caspase 3 te verwijderen middels het afsnoeren van caspase 3-

bevattende EMP, zouden endotheelcellen mogelijk beter kunnen overleven. De resultaten

in dit hoofdstuk tonen aan dat het aannemelijk is dat een aantal EMP inderdaad

rechtstreeks afkomstig is van adherente endotheelcellen. Tevens wordt aangetoond dat het

remmen van het afsnoeren van EMP leidt tot het accumuleren van caspase 3, het loslaten

van endotheelcellen en het ondergaan van geprogrammeerde celdood (apoptose).

Andere mogelijk functies

Onlangs hebben del Conde en medewerkers aangetoond dat de membranen van

weefselfactor-exposerende MMP mogelijk versmelten met geactiveerde bloedplaatjes,

waardoor het stollingsbevorderend weefselfactor wordt overgedragen [4]. Omdat de fusie

van membranen wordt bevorderd door PS, werd door hem verondersteld dat de

weefselfactor-exposerende MP verrijkt zouden kunnen zijn in PS. Onze resultaten, zoals

beschreven in hoofdstuk 3, bevestigen deze veronderstelling. Als dit model inderdaad

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valide is, dan mag worden aangenomen dat de overdracht van weefselfactor ook andere

functies van dit eiwit, zoals bevordering van angiogenese, signaal overdracht en

bescherming tegen apoptose, kan overdragen.

MP fracties, geïsoleerd uit plasmamonsters van patiënten met preeclampsie, acuut

myocard infarct of nierfalen, verstoren de endotheel-afhankelijke vaatverwijding [5-7]. Er

zijn meerdere aanwijzingen dat EMP een mogelijk rol spelen in dit proces. Ten eerste

wordt de NO-gemedieerde vaatverwijding wel geremd door geïsoleerde EMP uit

plasmamonsters van patiënten met nierfalen, maar niet door PMP afkomstig uit dezelfde

monsters [7]. Ten tweede remmen in vitro gemaakte EMP de endotheel-gemedieerde

vaatverwijding [8-10]. Ten derde is er een verband tussen de aantallen circulerende EMP

en de afname van de flow-gemedieerde vaatverwijding in patiënten met nierfalen en in

patiënten met diabetes type 2 [7,11].

Via welke mechanismen EMP en andere typen MP het endotheel beïnvloeden, is

onbekend. Een mogelijk mechanisme zou kunnen zijn dat membranen van (E)MP en

endotheelcellen versmelten, waardoor “intravesiculaire” moleculen, zoals bijvoorbeeld

caspase 3, kunnen worden overgebracht naar het cytosol. In een eerdere studie hebben wij

aangetoond dat geïsoleerde fracties van MP uit plasmamonsters van preeclamptische

patiënten in vitro de genexpressie van endotheelcellen niet beïnvloeden [12]. Mogelijke

verklaringen voor het ontbreken van een effect in dit systeem zouden kunnen zijn dat in

onze experimenten veneuze endotheelcellen zijn gebruikt, terwijl arteriële endotheelcellen

zijn gebruikt in studies naar endotheel-afhankelijke vaatverwijding. Onze eerdere

experimenten zijn bovendien uitgevoerd onder statische omstandigheden, dus zonder

“shear” (schuifkracht). Mogelijk is deze kracht nodig voor een interactie tussen EMP en

endotheelcellen. Een onlangs beschreven mechanisme is overdracht van mRNA door MP

afkomstig van embryonale stamcellen aan hematologische stamcellen, waardoor celdeling

en differentiatie van deze cellen worden bevorderd [13].

Klinische relevantie en toekomstige ontwikkelingenIn de literatuur wordt de aanwezigheid van EMP bij diverse ziekten in verband

gebracht met een verstoorde functie van het endotheel. In dit proefschrift wordt

aangetoond dat het afsnoeren van EMP ook een voordeel kan opleveren voor


160

endotheelcellen (Hoofdstuk 6) en dat een toename van EMP (dus) niet noodzakelijker-

wijs een merker is voor endotheel activatie of dysfunctie (hoofdstuk 5).

Er is steeds meer bewijs dat niet alleen endotheelcellen maar ook andere soorten

cellen hun “vesicles” gebruiken als transportmiddel. Deze “vesicles” vervoeren afval of

andere –voor de cel- schadelijke stoffen, en dragen bij aan de communicatie tussen cellen

onderling.

Gram-negatieve bacteriën snoeren MP af die zowel communicatie signalen als

toxinen bevatten. Deze toxinen zijn gericht tegen andere soorten bacteriën en

gastheercellen. Vermoedelijk is de afgifte van “vesicles” aan de omgeving een sterk

geconserveerd proces. Experimenten met een gram-negatieve bacterie, Pseudomanas

aeruginosa, hebben laten zien dat een intravesiculair signaal molecuul zelf het verpakken

in de “vesicles” van deze bacterie bevordert [14,15]. Onze resultaten met betrekking tot

het aanwezig zijn van caspase 3 in EMP passen mogelijk ook in dit model. Uit de

literatuur is bekend dat de borsttumor cellijn MCF-7 caspase 3-deficiënt is. Omdat MCF-7

cellen geen MP afsnoeren, zou de aanwezigheid van caspase 3 dus een voorwaarde

kunnen zijn voor het afsnoeren van MP. Na transfectie met een caspase 3-construct

snoeren MCF-7 cellen ook MP af [16]. Soortgelijke studies, uitgevoerd in ons

laboratorium, hebben aangetoond dat MP van MCF-7 cellen, na transfectie met caspase 3

construct, ook zelf caspase 3 bevatten. Mogelijk is dus ook caspase 3 betrokken bij het

verpakken in MP. Dergelijke resultaten bevestigen het model dat diverse soorten cellen

hun MP gebruiken als “afvalbak” om caspase 3 te verwijderen. In hoeverre niet alleen MP

maar ook andere soorten “vesicles”, zoals exosomen, bijdragen aan het verwijderen van

caspase 3 moet nader worden onderzocht.

Tijdens het onderzoek dat heeft geleid tot dit proefschrift is de interesse geleidelijk

verschoven van het identificeren en de functies van EMP naar een onderliggende en

wellicht veel fundamentelere vraag, namelijk waarom (endotheel) cellen “vesicles”

afgeven aan hun omgeving. Dit proefschrift toont aan dat het concept van circulerende

EMP als merkers die de conditie van het endotheel weerspiegelen, mogelijk een

onderschatting is van hun biologische en klinische relevantie. Bijvoorbeeld de

experimenten met simvastatine hebben aangetoond dat adherente endotheelcellen vitaal

lijken te blijven ondanks een toename van EMP en niet-adherente endotheelcellen

Chapter 8

161

(hoofdstuk 5). Bovendien hebben onze resultaten laten zien dat er een verband bestaat

tussen aantal EMP en het loslaten van de endotheelcellen in vitro. Het zal duidelijk zijn

dat deze resultaten de veronderstelde “directe” relatie die zou bestaan tussen adherente en

niet-adherente endotheelcellen en EMP complexer maakt.

De laatste tien jaar is de wetenschappelijke en klinische interesse voor de diverse

soorten blaasjes, MP en exosomen, sterk toegenomen. In het medisch onderzoek wordt de

aanwezigheid van blaasjes in vivo niet meer beschouwd als een in vitro artefact, maar als

een vaststaand feit. Er zijn verschillende toekomstige ontwikkelingen te verwachten met

betrekking tot het standaardiseren van de isolatie en detectieprotocollen van (E)MP. In

hoeverre het meten van aantallen (E)MP in klinische monsters daadwerkelijk als maat kan

dienen voor het niet goed functioneren van cellen, de ernst van de ziekte, stolling of

ontsteking, moet nader worden onderzocht. Op basis van de beschikbare weten-

schappelijke literatuur en onze resultaten zoals beschreven in dit proefschrift, wordt

verondersteld dat het afgeven van “vesicles” aan de omgeving een in de evolutie

geconserveerd proces is, dat kan bijdragen aan de overleving van cellen.

Samengevat, het lijkt niet alleen van belang te zijn om “vesicles” te isoleren en hun

talrijke functies te bestuderen in vitro en in vivo, maar het is wellicht even belangrijk om

de meer fundamentele vraag te beantwoorden waarom cellen dergelijke “multi-purpose

vesicles” afgeven aan hun omgeving.


162

References

1. Osmanovitch N, Romijn FPHTM, Joop K, Sturk A, Nieuwland R. Soluble selectins in sepsis: microparticle-associated, but only to a minor degree. Thromb. Haemost. 2000;84:731-2.






7. Amabile N, Guerin AP, Leroyer A, Mallat Z, Nguyen C, Boddaert J, London GM, Tedgui A, Boulanger CM. Circulating endothelial microparticles are associated with vascular dysfunction in patients with end-stage renal failure. J. Am. Soc. Nephrol. 2005;16:3381-8.

8. Densmore JC, Signorino PR, Ou J, Hatoum OA, Rowe JJ, Shi Y, Kaul S, Jones DW, Sabina RE, Pritchard KA, Jr., Guice KS, Oldham KT. Endothelium-derived microparticles induce endothelial dysfunction and acute lung injury. Shock 2006;26:464-71.

9. Esposito K, Ciotola M, Schisano B, Gualdiero R, Sardelli L, Misso L, Giannetti G, Giugliano D. Endothelial microparticles correlate with endothelial dysfunction in obese women. J. Clin. Endocrinol. Metab. 2006;91:3676-9.


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163


12. Lok CA, Böing AN, Reitsma PH, van der Post JA, van BE, Boer K, Sturk A, Nieuwland R. Expression of inflammation-related genes in endothelial cells is not directly affected by microparticles from preeclamptic patients. J. Lab Clin. Med. 2006;147:310-20.

13. Ratajczak J, Miekus K, Kucia M, Zhang J, Reca R, Dvorak P, Ratajczak MZ. Embryonic stem cell-derived microvesicles reprogram hematopoietic progenitors: evidence for horizontal transfer of mRNA and protein delivery. Leukemia 2006;20:847-56.

14. Winans SC. Microbiology: bacterial speech bubbles. Nature 2005;437:330.

15. Mashburn LM, Whiteley M. Membrane vesicles traffic signals and facilitate group activities in a prokaryote. Nature 2005;437:422-5.


Coauthors

165

Coauthors

Academic Medical Center, Amsterdam

Laboratory of Experimental Clinical Chemistry

Éva Biró

Anita N. Böing

Chi M. Hau

Frans J. Hoek

Rienk Nieuwland

Nada Osmanovic

Augueste Sturk

Experimental Vascular Medicine

Ludo M. Evers

Leiden University Medical Center, Leiden

Central Laboratory for Clinical Chemistry

Fred P.H.Th.M. Romijn

Slotervaart Hospital, Amsterdam

Department of Internal Medicine

Eelco W. Meesters

University Hospital Vrije Universiteit, Amsterdam

Department of Endocrinology

Michaela Diamant

Maarten E. Tushuizen

Coauthors

166

Organon BV, Oss

Pharmacology, Section General Pharmacology

Dirk G. Meuleman

Gerard M.T. Vogel

Curriculum Vitae

167

Curriculum Vitae

Mohammed N. Abid Hussein werd geboren op 19 augustus 1962 in Baqube, Irak. Hij

volgde zijn opleiding “Chemical Engineering” aan de Universiteit van Bagdad, waar hij

zijn diploma behaalde in 1984. Na een aantal jaren te hebben gewerkt in het bedrijfsleven,

kwam hij naar Nederland in 1992. Van 1993 tot 1995 werkte hij als onderzoeksanalist bij

de “Critical Care Division” van het “Royal Victoria Hospital” te Montreal, Canada. In

2000 behaalde hij zijn doctoraalexamen Farmacochemie aan de Vrije Universiteit te

Amsterdam. Aansluitend starte hij met zijn promotieonderzoek bij de afdeling Klinische

Chemie van het Leids Universitair Medisch Centrum te Leiden. Vanaf 2001 is hij

werkzaam bij het Laboratorium Experimentele Klinische Chemie van het Academisch

Medisch Centrum te Amsterdam.

Acknowledgments

169

Acknowledgments

This section is dedicated to thank several people whose help contributed

tremendously to realizing this thesis before taking its final shape.

To start with, is my promoter Guus Sturk. Dear Guus, your guidance and ideas

throughout the research period were indispensable for this thesis. You were always there

in difficult moments such as interpreting intricate data, without which major questions

would have not been tackled and problems would have not been resolved. Encouraging

and having faith in me to finalize this thesis, you were always. Ultimately Guus,

describing your assistance is hard to put in words.

The second person I would emphatically like to thank is my co-promoter Rienk

Nieuwland. Dear Rienk, you contributed to this thesis through your relentless and

insuperable effort in assisting and supervising my work from day one. You never run out

of ideas on how to put things into logical perspective. Undoubtedly, this thesis might not

have existed without your contribution. Thank you so much Rienk.

A cordial thank to my colleagues at LEKC Dennis Snoek, Éva Biró, Frans Hoek,

Loes Pronk, Maarten Tushuizen, Marc van de Zee, Marianne Schaap, René Berckmans

and Yung Yung Ko. They were extremely helpful in providing assistance, be it technical

or else. A special thank to Anita Böing and Chi Hau for the diligent and pleasurable joint

work.

Many thanks to Fred Romijn (Leiden), Ludo Evers (AMC), Eelco Meesters

(Slotervaart Hospital), Gerard Vogel and Dirk Meuleman (Organon) for their various

contribution to my research.

My thanks to any one who may have assisted in my research whose name I may have

forgotten to mention.

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