Fingerprinting Apoptotic Cell Surfaces

Fingerprinting Apoptotic Cell Surfaces:

Alterations of Glycocalyx and Membrane Composition

Den Naturwissenschaftlichen Fakultäten

der Friedrich-Alexander-Universität Erlangen-Nürnberg

zur

Erlangung des Doktorgrades

vorgelegt von

Sandra Franz

aus Chemnitz

Als Dissertation genehmigt von den Naturwissenschaftlichen Fakultäten der Universität Erlangen-Nürnberg

Tag der mündlichen Prüfung: 13.04.2007

Vorsitzender der Prüfungskommision: Prof. Dr. E. Bänsch

Erstberichterstatter: Prof. Dr. L. Nitschke

Zweitberichterstatter: Prof. Dr. Dr. h.c. J. R. Kalden

Drittberichterstatter: Prof. Dr. W. Jahnen-Dechent

Danksagung

Herrn Prof. Dr. Dr. h.c. J. R. Kalden möchte ich herzlich für die Betreuung der Arbeit und die Bereitstellung eines Arbeitsplatzes am Institut für klinische Immunologie und Rheumatologie danken. Ich erhielt vielseitige Einblicke in diese spannende Wissen- schaftsgebiet.

Ebenso möchte ich mich bei Herrn Prof. Dr. G. Schett für die Möglichkeit bedanken, meine Dissertation am Institut für Klinische Immunologie und Rheumatologie fortzuführen und erfolgreich zu beenden.

Herzlichen Dank auch an Herrn Prof. Dr. L. Nitschke für die Bereitschaft meine Arbeit von Seiten der Naturwissenschaftlichen Fakultät II zu vertreten.

Besonders bedanken möchte ich mich bei Herrn Prof. Dr. Martin Herrmann für die Bereitstellung des sehr interessanten Themas, seine intensive Unterstützung, die vielen Anregungen und zahlreichen sehr ergiebigen Diskussionen. Ich durfte mich stets in allen Themenbereichen seiner Arbeitsgruppe einbringen und erhielt die Möglichkeit auf zahlreichen Kongressen meine Ergebnisse zu präsentieren.

Ebenfalls herzlich danken möchte ich Herrn Prof. Dr. W. Jahnen-Dechent für die Übernahme des Drittgutachtens.

Mein Dank gilt auch Herrn Dr. R. E. Voll, Herrn Prof. Dr. H.-M. Jäck und Herrn Prof. Dr. L. Nitschke für die guten Kooperationen, die zahlreichen wissenschaftlichen Diskussionen und für das stete Interesse am Verlauf meiner Arbeit.

Der DFG möchte ich für die Förderung durch ein Promotionsstipendium im GRK 592 "Lymphozyten: Differenzierung, Aktivierung und Deviation" danken. In diesem Zusammenhang möchte ich mich ganz besonders bei Herrn Prof. Dr. H.-M. Jäck für die ausgezeichnete wissenschaftliche Ausbildung durch das GRK 592 bedanken. Ebenso möchte ich Herrn Dr. M. Lutz und Herrn Dr. A. Pahl für die Übernahme meiner Betreuungskommission und die vielen hilfreichen Disskussionen danken.

Ein ganz herzliches Dankeschön an die ehemaligen, alten und neuen Mitarbeiter der AG Herrmann: Pitti, Karina, Uwe, Tom, Udo, Benni, Luis, Ahmed, Silke, Andi, Alex, Gerhard, Birgit, Rüdiger, Kerstin, Connie, Christina für die tolle Zusammenarbeit (bis in die Nacht vor dem Facs vereint!), den guten Austausch, die stete Hilfsbereitschaft und auch die schöne Stunden außerhalb des Labors.

Ebenso herzlichen Dank an die Mitarbeiter und Freunde an der Medizinischen Klinik III für die schöne Zeit, die gute Zusammenarbeit und Hilfe; insbesondere an Marlies, Marvin, Dirk, Beate, Sven sowie an die „Vollis“ Babsi, Damian, Silke, Vilma, Kirsten, Sabine, Eva.

Bedanken möchte ich mich auch bei den Kollegiaten des GRK 592, besonders für die vielen außerordentlichen Stammtische mit dem „harten Kern“.

Mein besonders herzlicher Dank gilt meinem Freund Matthias und meinen Eltern für ihre unermüdliche Unterstützung und dass sie mir immer den Rücken frei gehalten haben.

Contents

1 Zusammenfassung 3

2 Introduction 11 2.1 Cell death by necrosis 12 2.2 Cell death by apoptosis 13 2.3 Clearance of apoptotic cells 16 2.4 Cell death and autoimmunity 20 2.5 Scope of the thesis 22

3 Results 23 3.1 During execution of apoptosis various cell populations with distinct morphological features are consecutively generated 23 3.2 Staining with certain lectins change morphology and membrane integrity of dying cells 25 3.3 The non lytic lectins GSL II, NPn, and UEA I bind differentially to viable and dying cells 27 3.4 Cell shrinkage precedes exposure of additional lectin binding sites during apoptosis 29 3.5 Loss of membrane integrity further increases the lectin binding sites 32 3.6 PS exposure precedes lectin binding in the time course of apoptosis 35 3.7 Lectins recognising epitopes from immature glycoproteins show increased binding to shrunken, late apoptotic cells 35 3.8 Before shrinkage apoptotic cells lose sialic acid epitopes of mature glycoproteins as detected by siglec binding 37 3.9 Loss of sialic acid during apoptosis is not blocked in the presence of sialidase inhibitor 38 3.10 Inhibition of glycoprotein processing in viable cells results in surface glycosylation similar to that of shrunken apoptotic cells 40 3.11 During apoptosis a relocation of ER and Golgi membranes toward cell surfaces can be observed 42 3.12 After shrinkage apoptotic cells expose the ER-resident chaperon calnexin 42 3.13 PS exposure precedes lectin binding and calnexin exposure in the time course of apoptosis 47 3.14 In apoptosing Ag8.H transfectants, dysfunctional immunoglobulin µ chains get access to the cell surface 47 3.15 CT-B binds to freshly isolated neutrophils but not to eosinophils 51 3.16 During the ageing process of PMN, GM1 disappears from the surfaces of full size apoptotic cells and resurfaces on shrunken apoptotic cells 51 3.17 During apoptosis intracellular ganglioside GM1 is relocated to the plasma membrane of neutrophil granulocytes 53 3.18 Exposure of NPn binding sites, calnexin, and GM1 on apoptosing PMN is abrogated when apoptotic membrane blebbing is blocked 55

3.19 In late stages of apoptosis KDEL receptor transgenic HeLa cells expose ER membrane on their surfaces 56 3.20 Exposure of immature glycoproteins is not sufficient to promote clearance of viable cells 59

4 Discussion 61 4.1 NPn, GSL II, and UEA I are suitable lectins for analysing the glycosylation status of dying cells 61 4.2 After shrinkage apoptotic cells expose immature glycoproteins 63 4.3 Before shrinkage apoptotic cells lose mature glycoproteins 67 4.4 During apoptosis internal membranes are translocated to the cell surface to substitute plasma membrane that are shed during the blebbing process 70 4.5 Carbohydrates of immature glycoproteins as “back-up-eat-me” signal of apoptotic cells that have escape early PS dependent clearance? 73 4.6 Hypothesis how apoptotic cells keep their silence and avoid immunogenicity 74

5 Materials 77 5.1 Cells and culture conditions 82 5.2 Animals 83 5.3 FITC labelling 84 5.4 Induction of apoptosis and necrosis 84 5.5 Detection of Apoptosis and Necrosis 84 5.6 Flow cytometry and microscopy analysis 86 5.7 Analysis of cell surface glycosylation 87 5.8 Analysis of surface sialic acid levels in the presence of sialidase inhibitor 88 5.9 Inhibition of glycoprotein processing 88 5.10 Cell labelling with ER-Tracker and Golgi-Stain 88 5.11 Detection of calnexin exposure 89 5.12 Detection of exposure of immunoglobulin µ chains on Ag8.H transfectants 89 5.13 Detection of GM1 exposure on neutrophil granulocytes 90 5.14 Inhibition of apoptotic membrane blebbing 90 5.15 Live-cell imaging of KDEL receptor-GFP transgenic HeLa cells undergoing apoptosis 91 5.16 Phagocytosis assays 91 5.17 Statistical analysis 92

6 References 93

7 Figure Index 105

8 Abbreviation Index 107

1 Zusammenfassung

Die Clearance apoptotischer Zellen ist ein fein abgestimmter Prozess, der auf komplexen Interaktionen zwischen Phagozyten und sterbenden Zellen basiert. Im Idealfall detektieren die Phagozyten apoptotische Zellen in einer frühen Phase des Zelltods und nehmen sie sofort auf, so dass ein Übergang der apoptotischen Zellen in die sekundäre Nekrose vermieden wird. Andernfalls würde aus den sekundär nekrotischen Zellen verändertes intrazelluläres Material freigesetzt, welches dann zu Entzündungen und Autoimmunerkrankungen führen kann. Die wichtigsten Aufgaben einer apoptotischer Zelle während des Clearance-Prozesses sind: (I) der Umgebung ihr Sterben zu signalisieren und ihre Oberfläche für die stille Erkennung und Beseitigung durch Phagozyten zu markieren, und (II) die Ionenselektivität ihrer Plasmamembran aufrecht zu erhalten bis sie phagozytiert wurde. Für ihre Beseitigung sezernieren apoptotische Zellen so genannte „Find-me“ Signale, welche die Phagozyten zu ihrem Wirkungsort dirigieren und exponieren „Eat- me“ Signale, welche den Phagozytoseprozess initiieren. Das am besten bekannte „Eat-me“ Signal ist Phosphatidylserin (PS), das in der frühen Phase der Apoptose vom inneren zum äußeren Blatt der Zytoplasmamembran verlagert wird. Verschiedene Phagozyten-Rezeptoren (der Vitronektin Rezeptor

[αvβ3 Integrin], der β2-Glycoprotein-I [β2-GPI] Rezeptor sowie die Rezeptor- Tyrosin Kinase Mer) binden an PS über das jeweils dazugehörige Brückenmolekül (milk-fat-globue-EGF-factor 8 [MFG-E8], β2-GPI, growth-arrest specific gene 6 [gas6]). Die apoptotischen Zellen werden dann unmittelbar von den Phagozyten aufgenommen. Es entkommen jedoch auch einige apoptotische Zellen dieser frühen PS abhängigen Clearance. In diesem Fall treten sie in späte Phasen der Apoptose ein, die durch das Schrumpfen der Zellen und den Verlust von Plasmamembran aufgrund des umfangreichen Abschnürens von Blebs (von Plamamembran umschlossene Vesikel)

1 Zusammenfassung

charakterisiert sind. Es ist bekannt, dass apoptierende Zellen die Ionenselektivität ihrer Plasmamembran für eine lange Zeit aufrechterhalten. Demzufolge müssen sie diesen massiven Verlust an Plasmamembran ausgleichen. Die hierfür verantwortlichen Mechanismen sind allerdings unbekannt. Um sekundäre Nekrose und Autoimmunität zu vermeiden muss zudem eine zweite Reihe an Signalen an der Oberfläche der sterbenden Zellen erscheinen, welche die Clearance der spät apoptotischen Zellen unterstützen. Verschiedene nicht PS spezifische Brückenmoleküle, wie zum Beispiel C reaktives Protein (CRP), Komplement Protein C1q, Surfactant Protein A und D (SP-A und SP-D), Mannose bindendes Lektin (MBL) und andere Lektine sind bekannt an spät apoptotische Zellen zu binden. Ihre spezifischen Epitope auf der apoptotische Oberfläche sind allerdings nicht bekannt.

Ziel dieser Arbeit war es Veränderungen in der Glykokalyx und in der Zusammensetzung der Plasmamembran von apoptotischen Zellen zu analysieren, um potentielle „Eat-me“ Signale besonders von spät apoptotischen Zellen zu identifizieren. Da verschiedene Lektine besonders spät apoptotische Zellen opsonisieren und deshalb massive Änderungen des Glykosilierungsmusters auf den Zelloberflächen voraussagen, wurde die Oberflächenglykosilierung von apoptotischen Zellen in verschiedenen Phasen des Zelltods untersucht. Die Bindung von 23 Fluoreszenz markierten Lektinen an die Oberflächen apoptotischer Zellen wurde analysiert. Lektine, die bekannt sind an Mannose-, N-Acetylglucosamin- und Fucose-Reste zu binden, zeigten die höchste Bindungsaktivität. In vitalen Zellen umfasst dieses Glykosilierungsmuster gewöhnlich die terminalen Zuckerreste von nicht vollständig prozessierten (unreifen) Glykoproteinen. In apoptotischen Zellen war die Zugänglichkeit dieser Zuckerstrukturen auf der Zelloberfläche auf späte Phasen des Zelltods beschränkt. Die Zellen waren bereits geschrumpft, besaßen aber immer noch eine ionenselektive Plasmamembran (Ausschluss von Propidium Iodid). Im Gegensatz dazu gingen Sialinsäuren, die terminalen Zuckerreste von vollständig prozessierten (reifen) Glykoproteinen, auf den Oberflächen der apoptotischen Zellen verloren. Dieser Verlust wurde bereits vor

1 Zusammenfassung

dem Schrumpfen der sterbenden Zelle beobachtet und wird höchst wahrscheinlich durch das Entlassen von Blebs von der Zelloberfläche hervorgerufen. Aufgrund dieser Beobachtungen wurde die Hypothese aufgestellt, dass während der Apoptose intrazelluläre Membranen und die darin enthaltenen unreifen Glykoproteine aus dem Endoplasmatischen Reticulum (ER) oder dem Golgi Komplex an die Zelloberfläche verlagert werden, um verloren gegangene Plasmamembran zu ersetzen.

Um diese Hypothese zu prüfen, wurden die Plasmamembranen apoptotischer Zellen nach Proteinen untersucht, die in vitalen Zellen im ER zurückgehalten werden. Das Chaperon Calnexin, eine dysfunktionale schwere Immunglobulin- Kette (µHC) und der KDEL Rezeptor sind Proteine, die in der ER-Membran verankert sind und nicht auf vitalen Zellen exponiert werden, aber auf den Oberflächen von geschrumpften, spät apoptotischen Zellen nachgewiesen werden konnten. Besonders wichtig anzumerken ist, dass die spät apoptotischen Zellen auch hier noch eine ionenselektive Plasmamembran besaßen. Calnexin und die unreifen Zuckerstrukturen erschienen im Verlauf der Apoptose mit einer ähnlichen Kinetik an der Zelloberfläche. Zudem kolokalisierten sie innerhalb vitaler Zellen und an der Oberfläche apoptotischer Zellen. Neben der Proteinzusammensetzung wurde auch die Lipidstruktur der Plasmamembranen von apoptotischen Zellen untersucht. GM1 ist ein Ganglioside, dass an der Zelloberfläche in Lipid Rafts und intrazellulär im ER angereichert vorkommt. Wie bereits bei Sialinsäure enthaltenden Glykostrukturen beobachtet, wurde die Oberflächenexpression von GM1 auf früh apoptotischen Zellen merklich weniger. In späten Phasen der Apoptose erschien GM1 auf den Oberflächen geschrumpfter, von einer ionenselektiven Plasmamembran umgebenen Zellen wieder. Es konnte gezeigt werden, dass auf den spät apoptotischen Zellen intrazelluläres GM1 aus dem ER auf der Zelloberfläche zugänglich wird. Wenn das apoptotische Abschnüren von Blebs von der Zelloberfläche (auch als Blebbing bezeichnet) mit Hilfe des ROCK- (Rho assozierte Proteinkinase) Inhibitors Y-27632 unterbunden wurde, waren auf der Oberfläche der spät apoptotischen Zellen interessanterweise weder

1 Zusammenfassung

unreife Glykostrukturen, noch aus dem ER stammende Proteine und Lipide zu detektieren.

All diese Ergebnisse untermauern die Hypothese, dass der infolge des Blebbing-Prozesses hervorgerufene Verlust an Plasmamembran durch die Verlagerung von aus dem ER stammenden Membranen an die Zelloberflächen ausgeglichen wird. Um den Apoptose spezifischen Membranfluß vom ER zur Plasmamembran zu verfolgen, wurden apoptierende KDEL Rezeptor-GFP transgene HeLa-Zellen an einem Migrationsmikroskop verfolgt. Es wurde beobachtet, dass sich im Verlauf der Apoptose Plasmamembran von der Zelloberfläche abschnürt und direkt durch Einlagerung von ER-Membran in die Oberfläche ergänzt wird. In der späten Apoptose war die ursprüngliche Plasmamembran fast vollständig auf der Zelloberfläche verschwunden und ER- Membran formte jetzt die äußere Hülle der Zelle. Schließlich wurde auch die ER-Membran in Form von Blebs von der Zelloberfläche abgeschnürt.

In vivo werden apoptotische Zellen normalerweise erkannt und phagozytiert noch bevor sie späte Phasen der Apoptose erreichen. Es wird vorgeschlagen, dass die Verlagerung von ER-Membranen an die Zelloberfläche eine Strategie apoptotischer Zellen ist, die Ionenselektivität ihrer Plasmamembran aufrecht zu erhalten und so lang wie möglich ein Auslaufen zu verhindern, wenn die frühe Clearance über Erkennung von PS fehlgeschlagen ist. Zusätzlich könnte die Bewegung von internen Membranen an die Zelloberfläche einen energiesparenden Weg apoptotischer Zellen darstellen, um gleichzeitig eine Vielzahl vorgeformter, später „Eat-me“ Signale an der Oberfläche zu exponieren, die im vitalen Zustand im Inneren der Zellen verborgen sind.

1 Summary

The clearance of apoptotic cells is a fine-tuned process based on a complex interaction between apoptotic cells and phagocytes. At best the phagocytic machinery detects and swallows all apoptotic cells in a way that progression to secondary necrosis is avoided. Otherwise, inflammation and autoimmune diseases may occur. The major tasks of apoptotic cells during the clearance process are: (I) to signal their demise to the environment and to flag themselves for their silent uptake, and (II) to maintain their membrane integrity until they are phagocytosed. For their removal apoptotic cells secrete “find-me” signals directing phagocytes to their site of action and expose “eat-me” signals initiating the process of phagocytosis. The best known “eat-me” signal is phosphatidylserine (PS), which is translocated from the inner to the outer leaflets of the cytoplasmic membrane early in the apoptotic process. Various phagocyte receptors (e.g. the vitronectin receptor [αvβ3 integrin], the β2- glycoprotein-I [β2-GPI] receptor, and the receptor-tyrosin kinase Mer) bind to PS via their corresponding bridging molecules (milk-fat-globue-EGF-factor 8 [MFG-

E8], β2-GPI, and growth-arrest specific gene 6 [gas6]). Most apoptotic cells are phagocytosed instantaneously in a silent fashion. However, some dying cells escape early PS dependent clearance and enter late stages of apoptosis characterised by cell shrinkage and plasma membrane loss due to extensive blebbing. Apoptosing cells are known to remain their membrane integrity for a long time. Hence, it is unclear how apoptotic cells compensate for this massive loss of plasma membranes. To avoid progression to secondary necrosis and autoimmunity a second line of signals which support clearance of late apoptotic cells has to emerge on the cells surfaces. Several non-PS specific bridging molecules including C reactive protein (CRP), complement C1q, surfactant proteine A and D (SP-A/-D), mannose binding lectin (MBL), and other lectins

1 Summary

are known to bind late apoptotic cells. However, their targets on the apoptotic surface are unknown.

The aim of this work was to analyse alterations in the glycocalyx and the composition of plasma membranes of cells undergoing apoptosis to identify potential “eat me” signals especially of late apoptotic cells. Changes in the surface glycosylation of cells undergoing apoptosis were monitored in various phases of cell death, since several lectin ligands specifically opsonising late apoptotic cells predict massive altered surface glycosylation. When we analysed the binding to apoptosing cells of 23 fluorescence labelled lectins, those known to bind mannose, N-acetylglucosamine, and fucose displayed the highest activity. This carbohydrate pattern usually comprises terminal sugar moieties of incompletely processed (immature) glycoproteins. The accessibility of these sugar structures was restricted to late apoptotic cells after shrinkage but with still an ion selective cytoplasmic membrane (propidium iodide [PI] exclusion). By contrast, sialic acids, terminal sugar residues of completely processed (mature) glycoproteins, got lost on the membranes of apoptosing cells. This loss was already observed before cell shrinkage and is, in all probability, caused by the release of plasma membrane surrounded blebs from the cell’s surface. Therefore, we hypothesized that during apoptosis intracellular membranes derived from the endoplasmic reticulum (ER) or the Golgi complex containing immature glycoproteins are translocated to the cells’ surfaces to substitute loss of plasma membrane.

We screened the plasma membranes of apoptotic cells for proteins that in viable cells are sequestered in the ER to prove this hypothesis. The chaperone calnexin, a dysfunctional immunoglobulin µ heavy chain (µHC), and the KDEL receptor, all ER resident proteins that are not exposed on viable cells, were detected on the surfaces of shrunken late apoptotic cells. Importantly, the latter still had an ion selective plasma membrane. Calnexin and the immature glycostructures occurred on the cell surface with a similar kinetic during the time course of apoptosis. Furthermore, they co-localised inside viable cells and on the surfaces of apoptotic ones. Additionally the plasma membranes’ lipid

1 Summary

composition of cells undergoing apoptosis was analysed. GM1 is a ganglioside that is enriched in lipid rafts on the cell surface and intracellularly in the ER, respectively. As sialic acid containing glycostructures, surface exposure of GM1 was observed to substantially fade early in apoptosis. GM1 reappeared on the surfaces of shrunken cells in late apoptosis. Again, the latter still equipped with an ion selective plasma membrane. We conclude that GM1 derived from the intracellular ER stores gets access to the cell surface. Interestingly, when the apoptotic blebbing process was abrogated using the ROCK- (Rho-associated protein kinase) inhibitor Y-27632 neither augmentation of immature glycostructures, nor exposure of ER derived proteins or lipids on the surfaces of apoptotic cells was to be detected.

Together all findings substantiated the hypothesis that the loss in consequence of the blebbing process of plasma membrane gets substituted by surface exposure of ER-derived membranes. Finally, the apoptosis related ER to plasma membrane trafficking was monitored performing live-cell imaging with KDEL receptor-GFP transgenic HeLa cells. It was recorded that during apoptosis plasma membrane blebbed from the cell surface and were directly replenished by surface incorporation of ER membranes. In late apoptosis, the genuine plasma membrane was almost completely lost from the cell surface and ER membranes now formed the outer envelope of the cells. At last, ER membranes were also released via blebs from the cell surfaces.

In vivo clearance of apoptotic cells is normally accomplished before the cells enter late stages of apoptosis. We suggest that the translocation of ER membranes to the cell surface is a strategy of apoptotic cells to maintain the ion selectivity of the plasma membrane and to prevent leakage as long as possible if the early PS dependent clearance has failed. Additionally, the movement of internal membranes to the cell surface may represent an energy-saving way of apoptotic cells for the concomitant exposure of a multitude of preformed late “eat-me” signals normally sequestered inside the cell.

2 Introduction

The subject of cell death has been fascinating scientist of different disciplines for a long time. Although most of them assume the birth of the subject in the 1970s, in the mid of the last century physiological cell deaths were already in the focus of histologists and developmental biologists (Clarke and Clarke, 1996). Virchow described cell death first in 1858 (Virchow, 1858) and Flemming observed 1879 during his study of mammalian ovarian follicles morphological changes in cells that we would today refer to as typical apoptotic (Flemming, 1885). The history of the clearance of dying cells has its beginning in 1882 as Mechnikov observed the absorption of the tadpole’s tail by neighbouring cells. For his discovery of phagocytosis Mechnikov was awarded 1908 the Novel prize (Metschnikoff, 1883). Short 100 years later, Kerr and colleagues framed some observations of the scientist of the nineteenth and early twentieth century into the concept of apoptosis (Kerr et al., 1972). Afterwards, genetic studies in the nematode C. elegans revealed apoptosis as a genetically controlled process that is conserved from worms to mammalians. Brenner, Sulston, and Horvitz were awarded in 2002 the Novel prize for their contributions in analysing the genetics of cell death.

However, the ongoing boom in cell death research of the last two decades emphasises the key role of cell death in any multicellular organisms. Apoptosis is not an incidental part of life but a highly regulated and controlled process. Pathophysiological conditions such as autoimmune diseases, cancer, or neurodegenerative disorder are often associated with disequilibrium in cell death control mechanisms.

2.1 Cell death by necrosis

The term necrosis (in Greek Νεκρός = death) is used to describe accidental death of cells and living tissues. Some authors name necrosis also as >accidental cell death< or >non programmed cell death<. However, necrosis follows acute (sudden anoxia or lack of nutrients) or extreme (heat, irradiation, toxins, mechanical or oxidative stress) injuries and can therefore be viewed as a sort of violent cell death. Mechanistically, necrosis appears disordered and messy and its morphological features are quite distinct from those of apoptosis (Walker et al., 1988). Rapid metabolic collapse leads to swelling of the entire cytoplasm (oncosis), early loss of plasma membrane integrity, and ultimate cell rupture (Lieberthal and Levine, 1996). Furthermore, the nucleus in necrotic cells is pycnotic and the chromatin does not get fragmented. The main difference compared to apoptosis is the release of intracellular components into the environment as a consequence of the cell damage. Hence, inflammation and immune response are characteristic by-products of necrotic cell death (Savill, 1998; Wyllie et al., 1980). The fundamental differences between apoptosis and necrosis are illustrated in Figure 1.

Necrosis is often viewed as antonym of apoptosis, but recent findings shed doubt on this view. Various toxins can induce both apoptosis and necrosis depending on the intensity of the stimulus (Bonfoco et al., 1995; Kroemer, 1995). In some models anti-apoptotic mechanisms were also effective against necrosis (Kane et al., 1993). Apoptotic signalling pathways were revealed to trigger necrosis (Holler et al., 2000; Lemasters et al., 1999; Vercammen et al., 1998). By manipulating these pathways a switch between both forms of cell death could be induced. Furthermore, several serum proteins mediating apoptotic cell clearance were identified as opsonins of necrotic cells (Bickerstaff et al., 1999; Gaipl et al., 2001; Nauta et al., 2003). Hence, necrosis, along with apoptosis, was considered as an active and controlled form of programmed cell death (Proskuryakov et al., 2002). In contrast, Manjo et al proposed necrosis not being a form of dying but the final end of any cell death process (Majno and Joris, 1995). Albeit the classification of necrosis and apoptosis is controversially

2.2 Cell death by apoptosis

discussed, it is generally assumed that the physiological consequences of both are quite different for the whole organism (Proskuryakov et al., 2002). In the case of necrosis, inflammation is provoked by intracellular constituents spilling out of the leaky plasma membrane. During apoptosis these components are safely enclosed by membranes and stay unnoticed by the body and the immune system.

2.2 Cell death by apoptosis

Apoptosis (in Greek απόπτωσις = falling of the leaves) is an intrinsic cellular suicide mechanism that follows characteristic features like cell shrinkage, margination of chromatin, membrane budding and fragmentation of the cell in apoptotic bodies (Kerr et al., 1972). According to this features Kerr, Wyllie and Currie coined the term apoptosis in 1972, which is today often used as synonym for programmed cell death. Furthermore, they pointed out that apoptosis is an important aspect of life. Apoptotic cell death occurs during fundamental biological processes such as embryogenesis, development, and normal tissue turnover. For example, maintenance of homeostasis in a human body includes the removal of about 10 billion cells a day to make room for new cells that arise by mitosis. Such massacre must not induce an immune response emphasising the importance of a rapid and efficient phagocytosis of the cells dying by apoptosis. Furthermore the process has to be tightly regulated as too little or too much apoptosis are linked to pathology like cancer, neurodegenerative or autoimmune diseases.

Apoptosis can be induced by the action of various extrinsic factors (death signals as Fas ligand, tumor necrosis factor α [TNFα]; or cessation of survival signals) on the cell and also by intrinsic signals (drugs, toxins, heat, radiation). At present, three pathways are known transforming apoptotic stimuli into the execution of the death program. All three mechanisms lead to the activation of caspases that are responsible for the morphological changes characterisic for apoptotic cell death. In the death receptor pathway Fas and tumor necrosis

2.2 Cell death by apotosis

SECONDARY APOPTOSIS NECROSIS early late

viable

NECROSIS

Figure 1: Schedule of apoptotic and necrotic cell death. Swelling of cytoplasm and organelles and subsequent cell rupture are characteristic features of necrotic cell death. In contrast, apoptotic cells shrink due to extensive generation of blebs. Chromatin condensation, DNA fragmentation, and formation of apoptotic bodies are further morphological hallmarks of apoptosis. When apoptotic cells are not cleared in time they can no longer maintain their membrane selectivity and become secondary necrotic.

receptor 1 (TNFR1) modulate via their associated death domains activation of caspase-8 (Hsu et al., 1995; Scaffidi et al., 1998). The mitochondrial pathway is often triggered in response to DNA damage and results in release of cytochrome c from mitochondria. The latter forms with the apoptosis protease activating factor1 (Apaf-1) and procaspase-9 the so-called apoptosome further modulating apoptosis (Li et al., 1997). Both pathways converge at the level of caspase-3 activation. However, normal apoptosis development in mice lacking

2.2 Cell death by apotosis

of caspase-8 and caspase-9 indicated the existence of an additional death pathway (Hengartner, 2000). Indeed, the endoplasmic reticulum (ER) mediated mechanisms seems to be independent of the two former pathways. ER stress including disruption of ER calcium homeostasis or accumulation of excess proteins in ER results in activation of caspase-12 (Nakagawa et al., 2000).

However, in the propagation of the apoptotic process mitochondria take a central part. They are strongly involved in determination the cell’s “point of no return”. Hence, pro- and anti-apoptotic members of the bcl-2 family fight for the cell’s fate at the surfaces of the mitochondria and compete for the release of cytochrome c (Hengartner, 2000). When the pro-apoptotic fraction has won the sequential activation of caspases follows.

In cells undergoing apoptosis, caspases target three groups of proteins: 1) regulatory proteins including several kinases or anti-apoptotic proteins (Widmann et al., 1998), 2) nuclear proteins like transcription factors, DNA repair proteins or lamin (Lazebnik et al., 1994; Rao et al., 1996), and 3) structural proteins such as actin, fodrin, gelsolin (Kothakota et al., 1997; Stegh et al., 2000; Villa et al., 1998). In the latter case, the cytoskeleton is destroyed step by step leaving the plasma membrane without the underlying net of actin, focal adhesion proteins, ERM (ezrin/radixin/moesin) molecules, plectin, intermediate filaments, and microtubules. The plasma membrane loses its rigidity without the support of the cytoskeleton. Two effects are indicative of this early apoptotic process where the cytoskeletal anchors are degraded. The plasma membrane gets acid labile (Heyder et al., 2003) and its fluidity increases significantly as can be concluded by increases in the mobility of phosphatidylserine (PS) (Appelt et al., 2004). Furthermore, early during apoptosis the transient surface exposure by viable cells of PS becomes persistent. The aminophospholipid translocase that flips back errant PS via an ATP-dependent mechanism loses activity, whereas the phospholipid flip-flop increases (Bratton et al., 1997). Both phenomena have been shown to depend on caspase activation or production of oxidants (Zwaal et al., 2005). Aside PS exposure, other morphological changes

2.3 Clearance of apoptotic cells

occur such as chromatin condensation and cytoplasmic contraction. Biochemically, DNA gets cleaved into oligonucleosomal fragments by various endonucleases. The characteristic DNA laddering is considered as a late event of apoptosis (Cohen et al., 1992). Extreme cell shrinkage due to extensive cytoplasm “boiling” and plasma membrane “blebbing” is also assigned to late apoptosis. Finally, a cell undergoing apoptosis is disintegrated into many apoptotic bodies and a lot of smaller blebs partially containing organelles or nuclear fragments (Coleman et al., 2001). Most importantly, these vesicles are still surrounded by intact membranes and consequently protect surrounding tissues from their potential cytotoxic contents. The final common event of the apoptotic program is to eliminate the apoptotic corpses from the tissue in a silent and efficient manner, to avoid inflammation and immune response.

2.3 Clearance of apoptotic cells

The clearance of apoptotic cells is a fine-tuned process based on a complex interaction between apoptotic cells and phagocytes. The importance of this process is highlighted by the fact that 8 out of 14 genes affecting apoptosis in C. elegans regulate the phagocytosis of the dying cells (Ellis et al., 1991).

The clearance of apoptotic cells is mediated by professional phagocytes (macrophages, immature dendritic cells [DC]) or non-professional phagocytes (neighbouring cells). Professional phagocytes have been shown to be highly motile and phagocytic and to immediately ingest apoptotic cells. By contrast, amateur phagocytes are rather sessile and reluctantly engulf apoptotic cells (Parnaik et al., 2000). However, for recognition by phagocytes apoptotic cells have to signal the readiness for their demise to the environment. It is assumed that apoptotic cells can direct phagocytes to their site of action by secreting so- called “find-me” signals. The phospholipid lysophosphatidylcholine (LPC) has recently been identified as such a potential attraction signal (Lauber et al., 2003). Furthermore, apoptotic cells express surface markers, so-called “eat-me” signals that are interpreted by phagocytes as positive signal for uptake. These

2.3 Clearance of apoptotic cells

“eat-me” signals encompass lipids, carbohydrates, and proteins (Liu et al., 2006). They either newly appear on the cell surface or arise from existing molecules modified during apoptosis. The most renowned “eat-me” signal is PS that is increasingly exposed early in the apoptotic process (Fadok et al., 1992). LPC (Kim et al., 2002) and oxidised proteins as well as lipids like the oxidised low-density lipoprotein (OxLDL) (Bird et al., 1999; Chang et al., 1999) at the apoptotic cell surface are also important signals for phagocyte recognition. Furthermore, it has been shown that efficient uptake of apoptotic cells requires exposition of both PS and its oxidized counterpart (Ox-PS) (Kagan et al., 2002). Intracellular adhesion molecule-3 (ICAM-3) was shown to function as a phagocytic marker of apoptotic leukocytes (Moffatt et al., 1999). From studies demonstrating apoptotic cell clearance via a carbohydrate-depended mechanism it is assumed that alterations in the glycosylation pattern of apoptotic cells serve as additional “eat-me” signals (Dini et al., 1992; Duvall et al., 1985).

Specific receptors on phagocytes recognise these signals either via direct binding or, more generally, via bridging molecules. The soluble adaptor molecules milk-fat-globule-EGF-factor 8 (MFG-E8) (Hanayama et al., 2002), and growth-arrest specific gene 6 (gas6) (Nakano et al., 1997) bind PS on apoptotic cells and bridge the latter to phagocytes via their corresponding receptors, the vitronectin receptor (αvβ3 integrin) (Rubartelli et al., 1997), and the receptor-tyrosin kinase Mer (Scott et al., 2001). Annexin I (Ax I) (Arur et al.,

2003), β2-glycoprotein-I (β2-GPI) (Balasubramanian et al., 1997) and serum- derived protein S (Anderson et al., 2003) were also identified as PS specific bridging molecules. By contrast, the phagocyte receptors class B scavenger receptor type I (SR-BI) and CD36 can tightly bind PS and might, therefore, directly interact with the apoptotic cell surface (Rigotti et al., 1995). The lectin- like OxLDL receptor-1 (LOX-1), the OxLDL receptor in endothelial cells, was also suggested to mediate phagocytosis of apoptotic cells recognising PS (Oka et al., 1998). Additionally, non PS specific phagocyte receptors and bridging molecules are also implicated in the recognition of apoptotic cells. Very recently,

2.3 Clearance of apoptotic cells

nucleolin, a multifunctional shuttling protein found in nucleus, cytoplasm and on the surface of some cell types, was identified as an early apoptotic cell-binding protein with a specificity for polylactosaminoglycans (Hirano et al., 2005). The vitronectin receptor αvβ3 and the thrombospondin (TSP) receptor (CD36) cooperate in binding TSP-1 that is known to interact with apoptotic cells (Savill et al., 1992). CD14, known as a receptor for bacterial lipopolysaccharide (LPS), mediates recognition and phagocytosis of apoptotic cells (Devitt et al., 1998; Schlegel et al., 1999). Various plasma proteins opsonise apoptotic cells, such as IgM, the pentraxins C-reactive protein (CRP) and the serum amyloid P component (SAP) (Ciurana and Hack, 2006). CRP and SAP promote uptake of apoptotic cells through the Fc receptors FcγRI and/or FcγRIII (Mold et al., 2002). Natural IgM antibodies that recognise LPC on apoptotic cells were proposed to be in part responsible for complement deposition on dying cells leading to their safe clearance (Kim et al., 2002). The collectins surfactant protein A (SP-A) and D (SP-D) (Schagat et al., 2001), the mannose binding lectin (MBL) (Nauta et al., 2003) as well as the complement protein C1q (Gaipl et al., 2001) bind to apoptotic surfaces and mediate cell clearance via a mechanism involving calreticulin and CD91 on phagocytes (Ogden et al., 2001; Vandivier et al., 2002). Noteworthy, apoptotic cell recognition and uptake via PS is an early clearance event, whereas binding of most non PS specific bridging molecules (C1q, SP-A, SP-D, MBL, SAP, CRP) occur in late apoptotic stages. Figure 2 illustrates the state-of-the-art of apoptotic cell clearance.

The multitude of interactions between phagocytes and apoptotic cells is often summarised in the so-called “tethering and tickling” model. Certain “eat me” signals first ensure the tethering of apoptotic cell and phagocyte, then the PS signal triggers the intracellular cascade for engulfment of the cell (tickling) (Somersan and Bhardwaj, 2001). However, according to a new concept tickling is not induced by receptor-signal interaction but by depolarisation of the macrophages (Simon Brown, personal communication). Very recently, alterations in the charge of the inner surface of the plasma membrane of

2.3 Clearance of apoptotic cells

β2GP1 β2GP1 AxI Receptor ?

MER PS or oxPS gas6 ανβ3 integrin MFG-E8 CD36 LOX-1

SR-BI IgM LPC ?

ICAM-3 CD14 ? lectin oxLDL ? ? altered nucleolin carbohydrates MBL CD91+ C1q calreticulin ? ? SP-A/D ανβ3 integrin TSP ? CD36 ? SAP Fcγ ? Receptor CRP APOPTOTIC CELL PHAGOCYTE

Figure 2: Phagocyte receptors, adaptor molecules and apoptotic “eat-me” signals that are involved in the apoptotic clearance process. Early clearance of apoptotic cells is mainly mediated via recognition of phosphatidylserine (PS). Apoptotic cells that escaped PS dependent uptake are concomitantly opsonised by several non PS specific adaptor molecules (C1q, SP-A, SP-D, MBL, CRP, lectins) which mediate late apoptotic cell clearance. This indicates that a second line of, yet unidentified, “eat-me” signals emerges on the surfaces of late apoptotic cells. Interactions of phagocyte receptors, adaptor molecules and apoptotic “eat-me” signals are discussed in the text.

2.4 Cell death and autoimmunity

macrophages during the course of phagocytosis have been shown. Furthermore, signalling molecules such as K-Ras, Rac1, and c-Src were rapidly released from membrane subdomains where the surface charge was altered (Yeung et al., 2006).

Unpublished data demonstrate changes of the membranes electrical polarisation of macrophages getting in contact with apoptotic cells. The membranes of the latter had also altered physicochemical features compared to viable cells. The depolarisation effects have been shown to benefit binding and engulfment of the cells (Simon Brown, personal communication). Moreover, it should be noted that Fc receptor-ligand binding is associated with depolarisation of macrophage membranes (Young et al., 1983). Fc receptor mediated phagocytosis is also implicated in the clearance process facilitating this new concept.

Mechanisms leading to recognition and uptake of apoptotic cells are very complex and to date only partially enlightened and understood. Nevertheless, it is generally accepted that clearance of apoptotic cells elicits neither inflammation nor immune response (Savill, 1997).

2.4 Cell death and autoimmunity

Apoptotic cells have been demonstrated to be removed in a silent fashion due to the release of anti-inflammatory cytokines (Fadok et al., 1998; Voll et al., 1997). Nevertheless, apoptotic cells are potential reservoirs of autoantigens that once released drive an immune response (Casciola-Rosen et al., 1994). This is true for apoptotic cells which have not been cleared in time and therefore turned to secondary necrosis. The cells’ membranes are then disintegrated and potential cytotoxic components can contact immune competent cells and trigger an immune response (Rovere et al., 2000). Dendritic cells may acquire the modified apoptotic self-materials and present them to autoreactive T cells, which, consecutively, get activated. There is growing evidence that defects in

2.4 Cell death and autoimmunity

the apoptotic cell clearance cells accompanied by secondary necrosis may contribute to the pathogenesis of systemic lupus erythematosus (SLE), an autoimmune disease mainly targeting wide spread nuclear autoantigens. DNA- protein complexes that escape from secondary necrotic cells have been considered to initiate and propagate the multisystem autoimmunity of SLE (Bickerstaff et al., 1999; Napirei et al., 2000). Several murine models demonstrated autoimmune disorders in consequence of defective apoptotic cell removal. For example, lupus like syndromes have been described for mice lacking functional C1q (Botto et al., 1998) or the secreted form of IgM (Boes et al., 2000; Ehrenstein et al., 2000), both bridging molecules in the clearance process. Mice defective in the phagocyte receptor MER also display lupus-like autoimmunity (Scott et al., 2001). In all mice a huge amount of uncleared apoptotic cells were found. In humans, a deficiency in the phagocytic activity of macrophages and PMN in a subgroup of SLE patients was shown (Herrmann et al., 1998; Landry, 1977). A study analysing lymph node biopsies revealed a reduced number of tingible body macrophages usually containing engulfed apoptotic nuclei in the germinal centres (GC) of some SLE patients. Apoptotic cells were accumulated in these lymph nodes. Even more, they were found associated with follicular dendritic cells (FDC) (Baumann et al., 2002). Notably, in the GC it is of major importance to remove apoptotic corpses very fast and efficiently. Otherwise, apoptotic-cell-derived autoantigens would serve as antigen for affinity maturation of B cells (Wellmann et al., 2005).

The following hypothesis for the induction of autoantibodies in SLE is currently discussed (Gaipl et al., 2003): In the GC apoptotic centrocytes escape their engulfment by tingible body macrophages and become secondary necrotic. Intracellular autoantigens of the leaky cells get immobilized by binding to FDC and, thereby, provide a survival signal for autoreactive B cells. Histone-specific T cells in the mantle zone of the lymph nodes may provide the signal for the persistence of these autoreactive B cells. Consecutively the latter proliferate, and differentiate into memory and anti-dsDNA antibody producing plasma cells.

2.5 Scope of the thesis

Taken together, these findings support the concept of apoptotic cells being potential reservoirs of autoantigens that might initiate and drive the humoral autoimmune response in SLE.

2.5 Scope of the thesis

In healthy conditions, apoptotic corpses are rapidly ingested and degraded by phagocytes. For their removal, apoptotic cells secret “find-me” signals and expose “eat me” signals as described in paragraph “clearance of apoptotic cells”. Several non-PS specific bridging molecules are known to bind apoptotic cells. However, their targets on the apoptotic surface are mostly unknown. Interestingly, the majority of these molecules including C1q, SP-A, SP-D, MBL, CRP, and several lectins opsonise only late apoptotic cells indicating a second line of, yet unidentified, “eat-me” signals, which emerge on the surfaces of apoptotic cells that have escaped earlier clearance mechanisms.

The aim of this work was to analyse alterations in the glycocalyx and the composition of plasma membranes of cells undergoing apoptosis to identify potential “eat me” signals especially of late apoptotic cells. The surface glycosylation status of dying cells was characterised using 23 fluorescence labelled lectins. Viable, early and late apoptotic and necrotic cells were compared for their lectin binding. Analysis of the glycocalyx revealed that immature glycoproteins get access to apoptotic cell surfaces. The hypothesis was proven that intracellular membranes derived from ER or Golgi containing these immature glycoproteins are translocated to the cell surfaces during apoptosis. The plasma membranes of apoptotic cells were screened for proteins that in viable cells are restricted to the ER. Furthermore, alterations in the lipid composition of apoptotic cells’ plasma membranes were analysed. Finally, it was proven that sugar structures identified as typical for apoptotic cells play a role in the clearance process.

3 Results

First, the glycosylation status of dying cells in various stages of apoptosis and in necrosis was analysed. Apoptosis in the human cell lines Jurkat, Raji, and U937 and in human peripheral blood lymphocytes (PBL) was induced by either irradiation with UV-B or treatment with staurosporine. Polymorphonuclear cells (PMN) underwent spontaneously apoptosis during in vitro culture. Primary necrosis in the cells was induced by heat or the membranes of the cells were permeabilised using detergent.

3.1 During execution of apoptosis various cell populations with distinct morphological features are consecutively generated

Cell death is characterised by morphological changes. The flow cytometer routinely measures two basic morphologic attributes of cells. The cell “size” and “granularity”, which are reflected by the forward (FSc) and the side scatter (SSc) values, respectively. Changes of the dying cells’ morphology were determined according to their FSc / SSc properties (Figure 3 a). Additionally, the viable, apoptotic or necrotic status of the cells was detected by staining with annexin V (AxV) in the presence of propidium iodide (PI) (Figure 3 b).

Untreated Jurkat cells and freshly isolated PBL and PMN did show no binding of AxV and PI (Figure 3 b). The morphology of these viable cells is displayed in population 1 (Figure 3 a, p.1). Apoptosis-related morphological changes can be seen after irradiation of Jurkat cells and PBL, or in vitro culture of ageing PMN. Irrespective of the apoptosis stimulus and the cell type, at a certain stage of apoptosis the cells lose volume caused by the release of membranous surface vesicles (blebs). The dying cells constantly lose FSc due to this shrinkage process. Therefore, two populations of apoptosing cells were

3.1 During execution of apoptosis various cell populations with distinct morphological features are consecutively generated

a) b) Jurkat wo UV-B, 3h UV-B, 6h 30`56°C wo UV-B 30`56° 100 80 60 4 40

1 2 3 2 cells [%] 20 0 p.1 p.2 p.2 p.3 p.4 PBL 3h 6h wo UV-B, 6h UV-B, 12h 30`56°C wo UV-B 30`56° 100 80 60 40 4

1 2 3 2 cells [%] 20 0 p.1 p.2 p.2 p.3 p.4 6h 12h PMN culture, culture, 0h 12h 26h 30`56°C 0h culture 30`56° 100 80 60 40 4 20 1 2 3 2 cells [%] 0

log SSc p.1 p.2 p.2 p.3 p.4 FSc p.1: viable 12h 26h p.2: full size, early apoptotic AxV - / PI - AxV + / PI - p.3: shrunken, late apoptotic p.4: primary necrotic AxV + / PI +

Figure 3: Detection of morphological changes of dying Jurkat, PBL, and PMN by their FSc / SSc properties by flow cytometry analysis. a) Time course of morphological changes in UV-B stressed Jurkat and PBL and ageing PMN as well as morphological attributes of viable and heat necrotised Jurkat, PBL, and PMN are displayed in FSc / SSc dot plots. During apoptotic schedule two cell populations, marked with 2 and 3, were defined. The viable populations of untreated or freshly isolated cells are marked with 1; the populations of necrotic cells are indicated as p.4. b) Viable, apoptotic, or necrotic status of populations 1, 2, 3, and 4 of Jurkat, PBL, and PMN, respectively, were detected by AxV / PI staining. Note: Although population 2 barely showed an altered cellular morphology some cells of population 2 already bound AxV, but still excluded PI. Therefore cells of population 2 are referred to as full size, early apoptotic cells, whereas cells of population 3 are referred to as shrunken, late apoptotic cells.

3.2 Staining with certain lectins change morphology and membrane integrity of dying cells

distinguished in Figure 3 a: a cell population which maintained their size (p.2), and a population of shrunken cells (p.3). The latter comprise apoptotic as well as secondary necrotic cells, reflecting a late apoptosis stage. Binding of AxV and exclusion or penetration of PI confirmed the apoptotic or secondary necrotic status of the cells in population 3, respectively (Figure 3 b, p.3). The apoptosing population of not shrunken cells (p.2) showed no noticeable alterations of the cellular morphology when compared with the viable population (p.1). However, some AxV binding cells were also detected in the full size population, reflecting early apoptosis (Figure 3 b, p.2). Hence, cells of population 2 were referred to as full size, early apoptotic cells, and cells of population 3 as shrunken, late apoptotic cells.

The typical morphology of necrotic cells can be observed in population 4 (Figure 3 a, p.4) when cells were treated with 56°C for 30 min. Necrotic PMN showed reduced FSc and decreased SSc properties, whereas the granularity of necrotic PBL and Jurkat cells is markedly increased. The necrotic status of the cells was confirmed by their binding of AxV and penetration of PI (Figure 3 a, p.4).

3.2 Staining with certain lectins change morphology and membrane integrity of dying cells

For the characterization of the glycosylation status of cells undergoing apoptosis the cell surface binding of 23 lectins was analysed. Three cell lines (Raji, Jurkat, U937), PBL, and PMN were subjected to the analysis of the surface carbohydrate content.

Several plant lectins are known to exert toxic effects on mammalian cells. This is reflected in altered morphology and loss of membrane integrity of the lectin treated cell. Therefore, changes of the cells FSc / SSc properties upon contact with a lectin were interpreted as toxic effects. Figure 4 shows FSc / Fluorescence (FL) dot plots of apoptotic Jurkat, Raji, and U937 cells after

3.2 Staining with certain lectins change morphology and membrane integrity of dying cells staining with the lectins derived from Narcissus pseudonarcissus (NPn), Griffonia simplicifolia (GSL II), Ulex europaeus (UEA I), Datura stramonium (DSL), and Ricinus communis I (RCA I). In comparison to AxV, staining with DSL and RCA I did change the cellular morphology. The cell populations clearly show a shift toward lower FSc (marked with an arrow). In contrast, NPn, GSL II, and UEA I exerted no remarkable effects on the cells morphology.

AxV GSL II NPn UEA I DSL RCA I 2 2 2 2 2 2 Jurkat 1 1 1 1 1 1 2 2 2 2 2 1 2 1 Raji

1 1 1 1 2 2 2 2 2 2 1 1 U937 1 1 1 1

log FL FSc p.1: full size, early apoptotic p. 2: shrunken, late apoptotic

Figure 4: Analysis of the cellular morphology of human cell lines (Raji, Jurkat, U937) after staining with certain lectins by flow cytometry. Binding of AxV and of the lectins GSL II, NPn, UEA I, DSL, and RCA I to apoptosing Raji, Jurkat, and U937 were detected by FSc / FL dot blots. The myeloid cell lines were induced to undergo apoptosis by UV-B irradiation and then cultured for 24 hours. Note: Morphology of cells after GSL II, NPn, and UEA I staining were close to those of AxV staining. In contrast, in the DSL and RCA I columns the cellular morphology is changed. The shift of the cell population toward lower FSc (marked with an arrow) was interpreted as toxic effect. Therefore, DSL and RCA I (marked with a star) were not used for further analysis. Note: Binding of GSL II, NPn, and UEA I to the shrunken, late apoptotic populations (p.2) of the myeloid cell lines were increased, when compared to the appropriate population of full size, early apoptotic cells (p.1). Staining with AxV served as control.

3.3 The non lytic lectins GSL II, NPn, and UEA I bind differentially to viable and dying cells

Table 1 gives an overview of the toxicity of all lectins that were tested. Most of the lectins exhibited membrane destructive properties and consecutively changed the morphology of at least one of the cell types. All cytotoxic lectins were excluded from further analysis. The lectins derived from Narcissus pseudonarcissus (NPn), Griffonia simplicifolia (GSL II), Ulex europaeus (UEA I) (all indicated in bold letters) neither exert toxic effects on cell lines (Jurkat, Raji, and U937) nor on primary cells (PBL and PMN) and can therefore be referred to as non lytic lectins. This makes NPn, GSL II, and UEA I suitable for the analysis of apoptosis related changes of the plasma membranes glycosylation status.

3.3 The non lytic lectins GSL II, NPn, and UEA I bind differentially to viable and dying cells

As an example for the binding of non lytic lectins to viable and dying cells Jurkat, PBL and PMN were stained with NPn and GSL II, respectively (Figure 5). Both lectins detected ligands on viable (p.1), apoptotic (p.2 and p.3) and necrotic cells (p.4). However, in all samples necrotic cells displayed the most lectin binding sites followed by late apoptotic cells. The same results were observed for UEA I (not shown). This suggests that the amount of NPn, GSL II, and UEA I binding epitopes increases on apoptotic and necrotic cells. Table 2 indicates the mean fluorescence intensity (MFI) for the binding of NPn, GSL II, and UEA I to viable and dying Jurkat, PBL, and PMN.

No binding on the surfaces of viable and apoptotic cells, or intracellular in necrotic cells were observed when cells were stained with fluorescein isothiocyanate (FITC) labelled goat IgG antibodies for control purposes (not shown).

3.3 The non lytic lectins GSL II, NPn, and UEA I bind differentially to viable and dying cells

Table 1: List of all lectins used in this work and their toxicity to Raji, U937, Jurkat, PBL, and PMN.

Lectins Toxicity

Raji U937 Jurkat PBL PMN

Concanavalin A (Con A) y y y y y Datura stramonium lectin (DSL) y y y y y Dolichos biflorus agglutinin (DBA) n y y n n Erythrina cristagalli lectin (ECL) y n y y y Griffonia simplicifolia lectin I (GSL I) n n y n n Griffonia simplicifolia lectin II (GSL II) n n n n n Jacalin (J) y y y y y Lens culinaris agglutinin (LCA) y y y y y Lycopersicon esculentum lectin (LEL) y y y y y Maackia amurensis lectin (MAL) y y y y y Narcissus pseudonarcissus lectin (NPn) n n n n n Peanut agglutinin (PNA) y n y n n Phaseolus vulgaris erythroagglutinin (PHE) y y y y y Phaseolus vulgaris leucoagglutinin (PHL) y y y y y Pisum sativum agglutinin (PSA) y y y y y Ricinus commuins agglutinin I (RCA I) y y y y y Solanum tuberosum lectin (STL) y y y y y Sophora japonica agglutinin (SJA) n n y y n Soybean agglutinin (SA) y n y n n Ulex europaeus agglutinin I (UEA I) n n n n n Vicia villosa lectin (VVL) n n y n n Wheat germ agglutinin (WGA) y y y y y Wheat germ agglutinin (WGA), succinylated y y y y y

Toxicity of the lectins was analysed by the FSc / SSc properties of the cells after lectin staining in comparison to AxV/PI staining. Lectins that obviously changed the cellular morphology of Raji, U937, Jurkat, PBL, or PMN were interpreted as toxic. y = toxic; n = not toxic

3.4 Cell shrinkage precedes exposure of additional lectin binding sites during apoptosis

Table 2: Binding of non lytic lectins shown in Figure 2 to viable and dying cells.

Jurkat PBL PMN Lectin vit apo nek vit apo nek vit apo nek NPn 3,7 18,8 186,8 1,5 5,7 174,3 2,9 23,6 61,0 GSL II 15,5 30,8 132,2 1,3 7,5 101,0 4,5 18,9 94,5 UEA I 21,5 34,5 137,4 2,7 14,7 133,8 7,3 19,7 134,5

The MFI of the specific binding to viable (vit), apoptotic (apo), and necrotic (nec) Jurkat, PBL, and PMN of the lectins NPn, GSL II, and UEA I is displayed.

3.4 Cell shrinkage precedes exposure of additional lectin binding sites during apoptosis

Figure 5 and 6 demonstrate that the loss of cellular volume, reflected by a decrease in the forward scatter, preceded the increased binding to apoptotic cells of NPn, GSL II, and UEA I. Ageing PMN after 21 h or 30 h in vitro culture as well as lymphocytes (Jurkat, PBL) treated with staurosporine or UV-B and additional culture for up to 12 h were prone to cell death. The populations of full size cells and shrunken cells (p.2 and p.3 in Figure 5, respectively) can, therefore, considered being apoptotic. However, the binding of the lectins to early apoptotic cells, which had preserved their size (Figure 6 and p.2 in Figure 5) was similar or weaker than the binding to viable cells (p.1 in Figure 5). Importantly, the binding by shrunken, late apoptotic Jurkat, PBL, and PMN (Figure 6 and p.3 in Figure 5) was significantly (see legend Figure 6) increased for all three lectins, when compared to the binding of viable cells. Taken together, cell shrinkage preceded the exposure of additional lectin epitopes on the surfaces of apoptosing cells indicating a late apoptotic event. Secondary necrotic cells as detected by their permeability for PI were excluded from analyses.

3.4 Cell shrinkage precedes exposure of additional lectin binding sites during apoptosis

Jurkat PBL NPn GSL II NPn GSL II 1 wo 1 3 2 UV-B, 12h 3 2 3 2 STS, 12h 3 2 4 30`56°C 4 log SSc log SSc count FSc log FL FSc count log FL PMN NPn GSL II p.1: viable 0h 1 p. 2: full size, early apoptotic

culture, 21h p. 3: shrunken, late apoptotic 3 2 p. 4: primary necrotic

30`56°C 4 log SSc FSc count log FL

Figure 5: Analysis of NPn and GSL II binding to viable and dying Jurkat, PBL, and PMN by flow cytometry. The population of viable (p.1), apoptotic (p.2 and p.3), and necrotic (p.4) Jurkat, PBL, and PMN, respectively, are displayed in FSc / SSc dot blots. Binding to the populations 1, 2, 3, and 4 of NPn and GSL II is presented in histograms. Note: Viable cells of Jurkat, PBL, and PMN show a substantial binding of NPn and GSL II. On dead cells the lectin binding is considerably increased. Necrotic cells possess the most binding capabilities for NPn and GSL II followed by shrunken, late apoptotic cells. In case of apoptosing cells, only cells which exclude PI were analysed.

3.4 Cell shrinkage precedes exposure of additional lectin binding sites during apoptosis

Jurkat PBL PMN

UV-B *** UV-B * 30h * GSL II STS *** STS * 0 1 2 3 4 5 6 012 024 6 8 10

UV-B ** UV-B ** 30h

NPn * STS *** STS ** 0 1 2 3 4 5 6 012 024 6 8 10

UV-B *** UV-B *** 30h * UEA I STS *** STS *** 0 1 2 3 4 5 6 012 024 6 8 10

relative MFI

full size, early apoptotic shrunken, late apoptotic

Figure 6: Comparison of GSL II, NPn, and UEA I binding to viable and full size, early and shrunken late apoptotic Jurkat, PBL, and PMN, respectively. Apoptosis in Jurkat and PBL was induced by various stimuli. Cells irradiated with UV-B (UV-B) or treated with staurosporine (STS) were cultured for 12 hours. PMN underwent spontaneous apoptosis during in vitro culture for 30 hours. Note: Irrespective of apoptosis stimulus and cell type, only the population of shrunken, late apoptotic cells showed a significantly increased binding of GSL II, NPn, and UEA I in comparison to the viable population (relative MFI of 1, indicated as dashed line). The relative MFI is calculated from the ratio of the MFI of the appropriate apoptotic populations and the MFI of the viable population. Only cells which excluded PI were analysed. *** p<0.001; ** p<0.01; * p<0.05

3.5 Loss of membrane integrity further increases the lectin binding sites

Figures 5 and 7 show a significantly higher binding to primary necrotic cells (p.4 in Figure 5) of GSL II, NPn, and UEA I than to shrunken, apoptotic cells (p.3 in Figure 5). This effect was more prominent in PBL than in the Jurkat cells or PMN.

Jurkat PBL PMN

100 100 100 *** *** *** * *** ** *** *** *** *** *** *** * ** *** 10 10 10 relative MFI

1 1 1 GSL II NPn UEA I GSL II NPn UEA I NPn GSL II UEA I shrunken, late apoptotic primary necrotic UV-B STS culture, 30h 30` 56°C

Figure 7: Comparison of GSL II, NPn, and UEA I binding to shrunken, late apoptotic and primary necrotic Jurkat, PBL, and PMN, respectively. Jurkat and PBL irradiated with UV-B (UV-B) or treated with staurosporine (STS) were cultured for 12 hours. PMN did age during 30 hours in vitro culture. Primary necrosis in all cells was induced by heat shock (30´56°C). Note: Binding to primary necrotic cells of GSL II, NPn, and UEA I is significantly higher than to shrunken, apoptotic ones. The relative MFI is calculated from the ratio of the MFI of the appropriate populations of dead cells and the MFI of the viable cells. In case of apoptosing cells, only cells which exclude PI were analysed. *** p<0.001; ** p<0.01; * p<0.05

In Figure 9 the time course of GSL II and NPn binding to Jurkat cells after apoptosis induction by UV-B is shown. Viable Jurkat cells (PI-, p.1) became

3.5 Loss of membrane integrity further increases the lectin binding sites

apoptotic (PI-, p.2 and p.3) followed by secondary necrosis (PI+, p.4). The lectin binding to the distinct cell population is displayed in FSc / FL dot blots. About 5h after UV-B irradiation increased binding to shrunken, apoptotic cells (p.3) of NPn and GSL II was observed. This is more prominent after 24 h in vitro culture of the irradiated cells. At this time point also a population of secondary necrotic cells (p.4) has emerged. The lectin binding to the secondary necrotic cells was increased when compared to shrunken, apoptotic cells (p.3) but similar to that of primary necrotic cells (p.5).

The binding of GSL II, NPn, and UEA I to necrotic and apoptotic Jurkat cells was also analysed by confocal microscopy (Figure 8). In necrotic cells the lectins showed a uniform intracellular staining, whereas the lectin staining appeared in spots at the surfaces of apoptotic cells. The membranes of the apoptotic cells were still intact as proven by their impermeability for PI (not shown).

Jurkat GSL II NPn UEA I A B C apoptotic

D E F necrotic

Figure 8: Analysis of GSL II, NPn, and UEA I binding to apoptotic and primary necrotic Jurkat by confocal microscopy. GSL II, NPn, and UEA I bind in a patchy pattern to the surfaces of apoptotic cells whereas in necrotic cells a uniform intracellular staining of the lectins was observed.

3.5 Loss of membrane integrity further increases the lectin binding sites

GSL II

PI - PI +

wo 1 3 UV-B, 2 5h 4 PI + 3 UV-B, 4 30`56°C 24h 5 2 log FL FSc p.1: viable NPn p.2: full size, early apoptotic PI - PI + p.3: shrunken, late apoptotic 1 p.4: secondary necrotic wo p.5: primary necrotic

3 UV-B, 2 5h 4 PI + 3 UV-B, 4 30`56°C 24h 5 2 log FL FSc

Figure 9: Time course of NPn and GSL II binding in the presence of PI to apoptosing Jurkat analysed by flow cytometry.

The time course of lectin binding is presented in FSc / log FL dot blots. Jurkat cells that became secondary necrotic during apoptosis were permeable for PI (PI+). This way, they were distinguished from apoptotic cells (PI-). During apoptosis three populations were defined according to their cellular morphology and permeability for PI: a full size, early apoptotic population (p.2, high FSc / PI-), a shrunken, late apoptotic population (p.3, low FSc / PI-), and a secondary necrotic population (p.4, low FSc / PI+). Note: Binding of GSL II and NPn to full size, early apoptotic cells is similar to viable cells (p.1), but markedly increased on shrunken, late apoptotic cells. Secondary necrotic cells show the same binding capacity for GSL II and NPn as primary necrotic (p.5) cells which is observable higher than for shrunken, apoptotic cells.

3.6 PS exposure precedes lectin binding in the time course of apoptosis

Figure 10 shows a typical apoptotic time course. The loss of the mitochondrial membrane potential (∆Ψm) (staining with the fluorescent dye 3.3'- dihexyloxacarbocyanine iodide [DiO6C(3)]), the exposure of phosphatidylserine (PS) (staining with AxV), and DNA degradation (staining with PI in the presence of detergent) were used as markers for early and late apoptosis. Early during the progress of apoptosis, the mitochondrial membrane potential (∆Ψm) decreases and the dying cells expose PS at the outer surfaces of their plasma membranes. The degradation of DNA reflected by the appearance of nuclei with sub-G1 DNA content is a rather late event of apoptosis.

The three lectins GSL II, NPn, and UEA I displayed a continuous increase of staining during the time course of apoptosis (Figure 10). Comparing the lectin staining with the other markers of apoptotic cell death (AxV, ∆Ψm, subG1), it was observed that the decrease of the mitochondrial membrane potential (∆Ψm) and the AxV signal preceded the increase of binding of GSL II, NPn, and UEA I (Figure 10 a). The kinetic of the binding to apoptotic cells of GSL II, NPn, and UEA I, and the appearance of cells with subG1-DNA content was remarkably similar (Figure 10 b). This indicates once more that the exposure of new lectin binding sites is a late event of apoptotic cell death.

3.7 Lectins recognising epitopes from immature glycoproteins show increased binding to shrunken, late apoptotic cells

Figures 3 and 4 clearly show the increased binding to shrunken, late apoptotic cells of NPn, GSL II, and UEA I. These lectins recognise mannose- (NPn), N- acetylglucosamine- (GSL II), and fucose-containing epitopes (UEA I), respectively. Interestingly, mannose, N-acetylglucosamine, and fucose residues can be found as terminal carbohydrates of immature glycoproteins during their

3.7 Lectins recognising epitopes from immature glycoproteins show increased binding to shrunken, late apoptotic cells processing. This finding indicates that epitopes of incompletely processed glycoproteins get access to the surfaces of shrunken, late apoptotic cells.

a) UV-B STS 100 100

10 10 relative MFI

1 1 0 4 8 12 16 04 81216 time (hours) b) STS 100 100

80 80

]

% 60 60 [

40 40 cells

20 20

0 0 0 4 8 12 16 0 4 8 12 16 time (hours) GSL II AxV NPn Δψm UEA I subG1

Figure 10: Time course of apoptosis in Jurkat. Changes in GSL II, NPn, and UEA I binding, changes in AxV binding, the mitochondrial membrane potential (ΔΨm), and the lowered DNA content (subG1) of Jurkat undergoing apoptosis were analysed at various time points. Note: a) Increased binding of AxV and loss of mitochondrial membrane potential preceded increased lectin binding. b) Time dependent increase of GSL II, NPn, and UEA I positive cells, as well as of cells with lower DNA content display similar kinetics but are delayed in comparison to AxV binding. Only cells which excluded PI were analysed.

3.8 Before shrinkage eapoptotic cells lose sialic acid epitopes of mature glycoproteins as detected by siglec binding

3.8 Before shrinkage apoptotic cells lose sialic acid epitopes of mature glycoproteins as detected by siglec binding

In completely processed proteins mannose, N-acetylglucosamine, and fucose residues are covered by sialic acid. Therefore, the amount of sialic acid should not be increased or even decreased on the surfaces of shrunken apoptotic cells when compared with viable cells. To prove this hypothesis PMN, PBL and Jurkat cells undergoing apoptosis were stained with sialic acid specific lectins, which are also referred to as siglecs.

The experiments revealed that the amount of accessible sialic acid epitopes on the plasma membranes decreased from viable to apoptotic cells. Interestingly, the decrease of siglec binding sites was observed in early apoptotic stages. Figure 11 shows the binding to ageing PMN of the siglecs derived from Maackia amurensis (MAL I) and Sambucus nigra (SNA). The populations of early apoptotic PMN (p.2) which had preserved their size bound markedly less MAL I and SNA than freshly isolated PMN (p.1). Later in apoptosis, the population of shrunken PMN (p.3) showed also a reduced binding of MAL I and SNA. In UV-B irradiated Jurkat cells and PBL a significant decrease of siglec binding during apoptosis could also be observed too (Figure 12). Most important, in all cells the loss of sialic acid epitopes preceded cell shrinkage. This indicates that early in the apoptotic process mature glycoproteins already get lost from the dying cells surfaces.

3.9 Loss of sialic acid during apoptosis is not blocked in the presence is sialidase inhibitor

PMN MAL I SNA 1 1 0h 1

2 2 8h 2 2 2 23h 3 3 3 2 log count SSc FSc log FL

p.1: viable p.2: full size, early apoptotic p.3: shrunken, late apoptotic

Figure 11: Analysis of MAL I and SNA binding to ageing PMN by flow cytometry. The population of viable (p.1) and apoptotic (p.2 and p.3) PMN are displayed in FSc / SSc dot blots. Binding to the populations 1, 2, and 3 of MAL I and SNA is presented in histograms. Note: Viable PMN possess many binding capabilities for MAL I and SNA, but on both full size as well as shrunken apoptotic cells MAL I and SNA binding is highly reduced. Only cells, which excluded PI are shown.

3.9 Loss of sialic acid during apoptosis is not blocked in the presence of sialidase inhibitor

Two mechanisms could lead to the surface loss of sialic acid: (I) a defective maturation of glycoproteins or (II) a sialidase activity to mature glycoproteins. To determine whether sialidases contribute to the loss of sialic acids during apoptosis a sialidase inhibitor was added to Jurkat cells before induction of apoptosis. The binding of MAL I and SNA to viable Jurkat and apoptotic Jurkat in the presence and absence of sialidase inhibitor was analysed. As shown in Figure 13 the decrease in MAL I as well as SNA binding was not inhibited by

3.9 Loss of sialic acid during apoptosis is not blocked in the presence is sialidase inhibitor the sialidase inhibitor. In a control experiment the sialidase inhibitor did block the activity of sialidase added to untreated Jurkat cells (not shown).

PMN 25 MAL I 25 SNA 20 20 * 15 15 ***

MFI 10 *** 10 *** 5 5 0 0 0h 23h 0h 23h

PBL 7 MAL I SNA 6 80 5 60 * 4 * MFI 3 40 * 2 20 1 0 0 wo UV-B wo UV-B

Jurkat 50 MAL I 30 SNA 40 25 * 20 30 * 15 MFI 20 10 * * 10 5 0 0 wo UV-B wo UV-B

viable full size, early apoptotic shrunken, late apoptotic

Figure 12: Comparison of MAL I and SNA binding to viable, full size apoptotic and shrunken apoptotic Jurkat, PBL, and PMN, respectively. Jurkat and PBL irradiated with UV-B were cultured for 12 hours. PMN did age during 23 hours in vitro culture. Note: Loss of MAL I and SNA binding sites preceded shrinkage. Except for MAL I in PBL, binding to full size but also shrunken apoptotic cells of MAL I and SNA is significantly reduced when compared to viable cells (wo). Only cell which excluded PI are shown. *** p<0.001; * p<0.05

3.10 Inhibition of glycoprotein processing in viable cells results in surface glycosylation similar to that of shrunken apoptotic cells

Jurkat

80 MAL I 14 SNA 70 12 60 10 50 8 40 MFI MFI 6 30 20 4 10 2 0 0 wo UV-B, 4h UV-B, 8h wo UV-B, 4h UV-B, 8h PBS sialidase inhibitor

Figure 13: Analysis of MAL I and SNA binding to apoptosing Jurkat in the presence of sialidase inhibitor. Viable (wo) and UV-B irradiated Jurkat cells were stained with the lectins in the absence and presence of sialidase inhibitor Note: Loss of MAL I and SNA binding sites was not blocked in the presence of sialidase inhibitor.

3.10 Inhibition of glycoprotein processing in viable cells results in surface glycosylation similar to that of shrunken apoptotic cells

A cocktail of the glycoprotein-processing inhibitors castanospermine, 1- deoxymannojirimycin, and swainsonine were used to block protein glycosylation in the different processing pathways. The processing inhibitors did not exert toxic effects on the cells as exemplarily shown for Jurkat cells in Figure 14 a. Jurkat cells that were cultured in vitro in the presence of the processing inhibitors (p.1, processing inhibitors) showed no morphological changes in comparison to untreated cells (p.1, wo). Furthermore, these cells did not bind AxV and were negative for PI confirming their viable status. The analysis of the glycosylation status of Jurkat cells treated with the processing inhibitors revealed increased binding of NPn and GSL II and reduced binding of MAL I and SNA when compared with untreated cells (Figure 14 b, wo and processing inhibitors). Interestingly, shrunken apoptotic yet PI impermeable cells expose

3.10 Inhibition of glycoprotein processing in viable cells results in surface glycosylation similar to that of shrunken apoptotic cells

a) Jurkat 100

wo ] 80 % 1 [ 60 40 cells 20 UV-B, 8h 0 3 2 p.1 p.2 p.3 p.1 wo UVB inhibitors processing inhibitors AxV - / PI - AxV + / PI - 1 AxV + / PI +

log SSc p.1: viable p.2: full size, early apoptotic FSc p.3: shrunken, late apoptotic

b) ** Jurkat 30 25 20 * 15 * MFI * ** 10 * 5 ** ** 0 NPn GSL II MAL I SNA wo, UV-B, processing inhibitors, viable shrunken apoptotic viable

Figure 14: Analysis of lectin binding to Jurkat cells treated with glycoprotein processing inhibitors by flow cytometry. a) The cell populations after 24 hours in vitro culture without treatment (wo, p.1), after UV-B irradiation (UV-B, p.2 and p.3), and after 24 hours in vitro culture in the presence of a processing inhibitor cocktail (processing inhibitor, p.1) are shown as FSc / SSc dot blots. The viability of the populations 1, 2, and 3 were analysed by staining with AxV/PI. Note: Treatment with the processing inhibitors exerted no toxic effects on the cells. b) Lectin binding to untreated, irradiated, and processing inhibitor treated Jurkat are compared. Note: NPn and GSL II show significantly increased binding to inhibitor treated cells and to shrunken apoptotic ones. Furthermore, MAL I and SNA binding is significantly decreased in both cases. * p<0.05

3.11 During apoptosis a relocation of ER and Golgi membranes toward cell surfaces can be observed additional NPn, GSL II, but less MAL I and SNA binding sites on their surfaces as well (Figure 14 b, UV-B). These data substantiate the hypothesis that in early apoptotic stages completely processed glycoproteins get lost from the cells surfaces and later in apoptosis, after cell shrinkage incompletely processed glycoproteins get access to the plasma membrane.

3.11 During apoptosis a relocation of ER and Golgi membranes toward cell surfaces can be observed

The fluorescent dyes ER-Tracker™ Blue-White DPX and Golgi-Stain BODIPY TR C5-ceramide specifically stain the ER and Golgi, respectively. Both, ER- Tracker and Golgi-Stain penetrate intact cells and locate to the appropriate cell compartment without exerting toxic effects to the cells. Therefore, the dyes were used to determine the distribution of ER and Golgi during apoptosis by fluorescence microscopy. Ageing PMN were analysed that had been stained as freshly isolated cells. Viable PMN showed a characteristic uniform staining pattern of the ER and Golgi, respectively (Figure 15 A, C, and E, G). After 20h in vitro culture the staining of ER and Golgi had drastically changed in apoptotic cells. Both the ER-Tracker and the Golgi stain were now observed at the plasma membrane indicating a redistribution of ER and Golgi toward the cells surface (Figure 15 B, D, and F, H).

3.12 After shrinkage apoptotic cells expose the ER-resident chaperon calnexin

The chaperone calnexin is an integral membrane protein located primarily in the ER. The C-terminus of the ER resident protein contains the characteristic ER retention motif. Viable cells contain calnexin sequestered in their ER membranes. Therefore, no signal could be detected by flow cytometry, when the surfaces of freshly isolated PMN were stained with an antibody against a

3.12 After shrinkage apoptotic cells expose the ER resident chaperone calnexin

PMN ER-Tracker Golgi-Tracker 0h 20h 0h 20h viable apoptotic viable apoptotic A B E F

C D G H

Figure 15: Examination of ageing PMN stained with ER-Tracker and Golgi Stain by fluorescence microscopy. Freshly isolated PMN (0h) and those after an ageing process of 20 hours in vitro culture (20h) were analysed. Viable and apoptotic PMN stained with either the ER- Tracker (A - D) or the Golgi Stain (E - H) are displayed. Note: In viable PMN a uniform staining of ER (C) and Golgi (G) can be observed. In apoptotic PMN, the dyes have a different appearance – they are translocated to the cell’s surfaces (B + D).

luminal calnexin domain (α calnexin) (Figure 16, p.1). In contrast, calnexin could be detected in necrotic cells, as α calnexin got access to the intracellular compartment (Figure 16, p.4). Interestingly, during apoptosis the ER resident protein emerge a t the plasma membrane of the dying cells. More importantly, only shrunken, late apoptotic cells did expose calnexin on their surfaces (Figure 16, p.3). Early apoptotic PMN which had preserved their size were negative for α calnexin (Figure 16, p.2) as viable cells (p.1). The amount of intracellular calnexin is markedly higher than that on late apoptotic cells indicating that not the total ER pool of calnexin is reaching the cell surface. Secondary necrotic cells as detected by their permeability for PI were excluded from analyses.

3.12 After shrinkage apoptotic cells expose the ER resident chaperone calnexin

The relocation of ER resident calnexin to the surfaces of aging PMN was confirmed by fluorescence microscopy (Figure 18 a). Comparing the distribution of calnexin and the ER-Tracker, co-localisation in viable as well as in apoptotic cells were found. In viable PMN, both molecules were detected intracellular in the ER (Figure 18 a, A - C). In apoptotic ones, ER-Tracker and calnexin partially located at the plasma membrane (Figure 18 a, D - F).

PMN

1 control 0h 1 α calnexin

2 3 culture, 20h 3 2

4 intracellular 4 log SSc count FSc log FL

p.1 viable p.3 shrunken, late apoptotic p.2 full size, early apoptotic p.4 necrotic

Figure 16: Analysis of calnexin exposure on ageing PMN by flow cytometry. The population of freshly isolated PMN (p.1, 0h), of apoptotic PMN (p.2 and p.3) after 20 hours in vitro culture (20h) and of necrotic PMN (p.4) that were obtained by permeabilisation of the plasma membranes (intracellular) are displayed in FSc / SSc dot blots. Calnexin exposure on the populations 1, 2, 3, and 4 were analysed in histograms. Note: Anti-calnexin does not bind to the surface of viable (p.1) and early apoptotic cells (p.2) but to the plasma membrane of shrunken apoptotic cells (p.3). Necrotic cells (p.4) bound m arkedly more anti-calnexin, when compared with shrunken apoptotic ones. In populations 1, 2, and 3 only cells which excluded PI are shown.

Lymphocytes undergoing apoptosis were also observed to expose ER resident calnexin on their surfaces after shrinkage. As exemplarily shown for Jurkat cells

3.12 After shrinkage apoptotic cells expose the ER resident chaperone calnexin

in Figure 17 calnexin could not be found on viable cells and on full size apoptotic cells. In contrast, on shrunken late apoptotic Jurkat cells calnexin was detectable on the surfaces of still PI impermeable cells. The results were confirmed using confocal microscopy (Figure 18 b). Late apoptotic cells with an intact plasma membrane, as shown by their impermeability for PI, exclusively displayed binding of α calnexin on their surfaces (Figure 18 b, A - C). In secondary necrotic cells an intracellular staining of the cells was additionally to be observed (Figure 18 b, D - F). Viable cells did not show a surface signal (not shown).

Jurkat

100 ***

* ** MFI 1

0 wo UV-B STS intracellular

viable shrunken, late apoptotic full size, early apoptotic necrotic

Figure 17: Analysis of calnexin exposure on viable, apoptotic, and necrotic Jurkat. Apoptosis was induced by irradiation (UV-B) or staurosporine treatment (STS) followed by 8 or 16 hours in vitro culture, respectively. Necrotic Jurkat were obtained by permeabilisation of their plasma membranes (intracellular). Note: Calnexin can not be detected on viable and full size apoptotic cells, but on shrunken apoptotic ones and, to the strongest extent, in necrotic cells. In case of apoptosing Jurkat PI positive cells were excluded from analysis. *** p<0.001; ** p<0.01; * p<0.05

3.12 After shrinkage apoptotic cells expose the ER resident chaperone calnexin

a) PMN ER- α calnexin Tracker merge A B C

intracellular

D E F surface, apoptotic

b) Jurkat c) Jurkat secondary surface, apoptotic necrotic apoptotic intracellular A D A D

α calnexin α calnexin

B E B E

PI UEA I

C F C F

merge merge

Figure 18: Analysis of calnexin exposure on PMN and Jurkat by microscopy. a) PMN double stained with anti-calnexin (green) and the ER-Tracker (red) were analysed by fluorescence microscopy. Note: Calnexin and ER-Tracker co-localise intracellular in permeabilised cells (A - C) as well as on the surfaces of late apoptotic PMN (D - F). Both dyes were not detected on the surfaces of viable PMN (not shown). b) Binding of anti-calnexin (green) in the presence of PI (red) to apoptosing Jurkat cells was analysed by confocal microscopy. Note: Anti-calnexin binds in a patchy pattern to the surfaces of apoptotic, PI impermeable cells. In secondary necrotic, PI positive cells calnexin was also found intracellular. c) Apoptosing Jurkat double stained with anti- calnexin (green) and UEA I (red) were analysed by confocal microscopy. Note: Calnexin epitopes and UEA I binding sites co-localise on the surfaces of apoptotic cells and intracellular.

3.13 PS exposure precedes lectin binding and calnexin exposure in the time course of apoptosis

Furthermore, the distribution of calnexin and UEA I binding sites of Jurkat cells were analysed by confocal microscopy (Figure 18 c). Interestingly, calnexin and UEA I epitopes co-localised intracellular of permeabilised cells (Figure 18 c, A - C) as well as at the plasma membrane of apoptotic ones (Figure 18 c, D - F).

3.13 PS exposure precedes lectin binding and calnexin exposure in the time course of apoptosis

Figure 19 shows a typical apoptotic time course of ageing PMN. Early during the progress of apoptosis, the cells exposed PS at the outer surfaces of their plasma membranes as detected by AxV-staining with a continuous increase up to 85%. In the same way, the binding to apoptosing PMN of the lectins GSL II and NPn, as well as of α calnexin increased continuously. The kinetics of the surface exposure of the lectin binding sites and of calnexin were basically the same. Importantly, in the time course of apoptosis the exposure of PS clearly preceded both, the lectin binding and the calnexin exposure. These data indicate that in late stages of apoptosis not only ER resident calnexin is translocated to the cell surfaces but, concomitantly also immature intracellular glycoproteins emerge at the plasma membrane.

3.14 In apoptosing Ag8.H transfectants, dysfunctional immunoglobulin µ chains get access to the cell surface

Approximately half of all productively rearranged µ heavy chains (µHC) are unable to pair with surrogate light chain and conventional immunoglobulin light chain (ten Boekel et al., 1997). These dysfunctional µHCs are retained by the ER-resident heavy chain binding protein (BiP) and can, therefore, not reach the cell surface (Haas and Wabl, 1983; Kline et al., 1998).

3.14 In apoptosing Ag8.H transfectants, dysfunctional immunoglobulin µ chains gat access to the cell surface

PMN 100

80 AxV GSL II 60 NPn α calnexin cells [%] 40

0 0 10 20 30 40 50 time (hours)

Figure 19: Time course of apoptosis for ageing PMN. Surface binding of GSL II, NPn, anti-calnexin (α calnexin), and AxV to ageing PMN was analysed at various time points. Note: Time dependent increase of GSL II, and NPn positive cells, as well as of cells binding α calnexin display similar kinetics but are delayed when compared to AxV binding. PI positive cells were excluded from analysis.

The Ag8.H cell line expressing neither Igα nor µHC was transfected with Igα and with a non-pairing dysfunctional µHC. Since the latter is retained in the ER the transfectant Ag8.Hα µdys cells were used to substantiate the observations of ER-resident components getting access to the surfaces of late apoptotic cells.

As shown in Figure 20 a by flow cytometry, the dysfunctional µHC in Ag8.Hα

µdys cells was exclusively detectable by intracellular staining but not on the cell surface. Fluorescence microscopic analysis verified that viable Ag8.Hα µdys cells did not show surface exposure of their µHC (Figure 21 a, A - C). After treatment with UV-B and in vitro culture for 12 hours, the dysfunctional µHC were detected on the surfaces of apoptotic Ag8.Hα µdys cells (Figure 21 a, D - F

3.14 In apoptosing Ag8.H transfectants, dysfunctional immunoglobulin µ chains gat access to the cell surface and G - I). As typical for cells undergoing apoptosis they showed high granularity and occasional blebs. The membranes of those cells were still intact as proven by their impermeability for PI. In contrast, the membranes of necrotic

Ag8.Hα µdys cells were permeable for PI. The antibodies against the µHC were able to enter necrotic cells and did bind the ER-resident µHC (Figure 21 a, J - L). Importantly, the exposure of the dysfunctional µHC on apoptotic Ag8.Hα

µdys cells was only observed after cell shrinkage, as detected by flow cytometry

(Figure 21 b, Ag8.Hα µdys).

The Ag8.H cell line was also transfected with Igα and with a pairing functional

µHC. The intracellular expression and surface exposure of µHC in Ag8.Hα µfct is displayed in Figure 20 b. After the induction of apoptosis significantly more of the µHC were found on the cell surfaces, but only of shrunken, late apoptotic cells (Figure 21 b, Ag8.Hα µfct).

Taken together all findings clearly show that during apoptosis epitopes that are sequestered inside viable cells (ER resident calnexin, ER resident µHC as well as intracellular, incompletely processed glycoproteins from ER and Golgi) get access to the cell surfaces. The exposure of these epitopes is only observed after cell shrinkage on late apoptotic but still PI negative cells.

a) Ag8.Hα µdys b) Ag8.Hα µfct intracellular surface intracellular surface

count log FL (µHC) control: Ag8.H

Figure 20: Analysis of expression of µHC in Ag8.Hα transfectants by flow cytometry.

Surface and intracellular binding of anti-µHC to Ag8.Hα µdys cells and Ag8.Hα µfct are displayed in histograms. Note: In Ag8.Hα µdys transfectant the dysfunctional µHC were solely detected intracellular in permabilised cells but not on the plasma membranes of viable cells. In contrast, in Ag8.Hα µfct transfectant the µHC were found intracellular and on the surfaces. Staining of Ig-negative Ag8.Hα cells with anti-µHC served as control.

3.14 In apoptosing Ag8.H transfectants, dysfunctional immunoglobulin µ chains gat access to the cell surface

a) Ag8.Hα µdys b) µHC PI A B C *** 40 wo viable 30 D E F

MFI 20 totic

p ***

o G H I UV-B

p 10 a

0 J K L Ag8.Hα µdys Ag8.Hα µfct full size, early apoptotic 30`56°C necrotic shrunken, late apoptotic

Figure 21: Analysis of µHC exposure in Ag8.Hα transfectants undergoing apoptosis.

Ag8.Hα µdys and Ag8.Hα µfct transfectant were treated as follows: untreated cells (wo), apoptosis induction by irradiation with UV-B followed by 10 hours in vitro culture (UV- B), necrosis induction by heat shock (30´56°C) or permeabilisation of the cells (intracellular). a) Binding of anti-µHC in the presence of PI to viable, apoptotic, and necrotic Ag8.Hα µdys cells was analysed by fluorescence microscopy. Note: Viable cells (A - C) show no binding of anti-µHC. After apoptosis induction, µHC were detected on the surfaces of dying Ag8.Hα µdys cells with an intact plasma membrane (D - F). In necrotic cells (G - I), anti-µHC mainly binds to intracellular targets. b) Binding of anti-

µHC to full size, early apoptotic and shrunken, late apoptotic Ag8.Hα µdys and

Ag8.Hα µfct cells is compared. Note: Significant exposure of the dysfunctional µH C and significantly increased exposure of the functional µHC were only detected on shrunken, late apoptotic Ag8.Hα transfectants. Only cells which excluded PI were analysed. *** p<0.001

3.15 CT-B binds to freshly isolated neutrophils but not to eosinophil

3.15 CT-B binds to freshly isolated neutrophils but not to eosinophils

The ganglioside GM1 is a glycolipid that associates with lipid rafts at the cell surfaces. Cholera toxin subunit B (CT-B) specifically binds to GM1 and was therefore used to analyse GM1 exposure of ageing PMN.

Eosinophils and neutrophils were differentiated by staining with anti-CD16 (neutrophils) and anti-CD49d (eosinophils) antibodies, respectively. Staining of freshly prepared PMN with FITC-labeled CT-B was analysed. As shown in Figure 22 binding of CT-B was observed only to neutrophils and not to eosinophils. Microscopic examination of freshly isolated PMN showed special binding in a patchy pattern of CT-B (Figure 23 a). The PMN were also photographed in several layers with 1 µm intervals (Figure 23 b). Fluorescence at the plasma membrane but also intracellular fluorescence was detected indicating an uptake of CT-B by the PMN.

3.16 During the ageing process of PMN, GM1 disappears from the surfaces of full size apoptotic cells and resurfaces on shrunken apoptotic cells

During in vitro culture PMN underwent spontaneous apoptosis. Morphological changes of the ageing PMN could be observed as changes of the cells FSc / SSc properties. In the sequence of apoptosis the cells lost volume caused by the release of membranous surface vesicles (blebs). The cell shrinkage is reflected by a lower FSc of the cells. Therefore, a population of full size, early apoptotic cells (Figure 24 a, p.2) and a population of shrunken, late apoptotic cells (Figure 24 a, p.3) were distinguished. The apoptotic status of the two populations was proven by staining with AxV in the presence of PI (not shown).

Before the start of apoptotic processes almost all freshly isolated PMN did bind CT-B. Only 5% of the cells in the viable population were CT-B negative (Figure 24, p.1, 0h). After 5 h in vitro culture the amount of CT-B negative cells

3.16 During the ageing process of PMN, GM1 disappears from the surfaces of full size apoptotic cells and resurfaces on shrunken apoptotic cells

PMN 91 7,5 100 7,6 92 CD16-PE CD49d-PE PE-Control

FITC-Control CT-B-FITC CT-B-FITC

Figure 22: Analysis of CT-B binding to neutrophil and eosinophil granulocytes by flow cytometry. Double staining of PMN with CT-B and either anti-CD16 or anti-CD49d are presented in FL FITC / FL PE dot blots. Note: Binding of CT-B was observed only to CD16- positive cells (neutrophils, 91% of PMN) and not to CD49d-positive cells (eosinophils, 7.5% of PMN).

Freshly isolated PMN a) b) 1 µm2 µm3 µm

4 µm5 µm6 µm

Figure 23: Examination of CT-B binding to freshly isolated PMN by fluorescence microscopy. a) Note: CT-B binds in a patchy pattern to the surfaces of viable PMN. A composite picture is shown. b) CT-B binding to PMN was analysed in several layers at 1 µm intervals. Note: Fluorescence was detectable at the plasma membrane but also to a lesser extend inside the cells.

3.17 During apoptosis intracellular ganglioside GM1 is relocated to the plasma membrane of neutrophil granulocytes had drastically increased. The population of early apoptotic cells could clearly be separated into two groups: a group of CT-B binding cells and a group of none CT-binding cells. The latter comprised 30% of the cells (Figure 24, p.1, 5h). During the ongoing ageing process the non–CT-B binding population increased continuously and became dominant after 22 h in vitro culture (Figure 24, p.1, 22h). At this time point, a significant amount of cells were found in the shrunken apoptotic cell gate. Interestingly, almost all shrunken apoptotic cells did bind CT-B (Figure 24, p.2, 22h). As time advanced, the cell population of full size apoptotic cells decreased and the population of shrunken apoptotic PMN increased. After 45 h in vitro culture, 80% of the cells were found in population 2. Over 95% of these cells were positive for CT-B (Figure 24, p.2, 45h). As shown by the MFI values in Figure 24 b shrunken apoptotic cells bound more amounts of CT-B than viable cells did. This indicates that GM1 from intracellular stores might reach the shrunken apoptotic cells surfaces.

3.17 During apoptosis intracellular ganglioside GM1 is relocated to the plasma membrane of neutrophil granulocytes

The distribution of the ganglioside GM1 in viable and apoptotic neutrophils was analysed by fluorescence microscopy (Figure 25). In viable cells GM1 was detected at the plasma membrane and in intracellular stores. GM1 showed a uniform intracellular staining in viable cells (Figure 25, A) while it appeared in a patchy pattern at the plasma membranes (Figure 25, D). In apoptotic cells the GM1 staining pattern had drastically changed. GM1 lost its uniform intracellular distribution and was found to cap in a distinct area of the plasma membrane (Figure 25, G). Comparing the location of GM1 and the ER-Tracker showed that both molecules co-localised inside viable cells (Figure 25, A - C) but not at the plasma membrane of viable cells (Figure 25, D - F) Importantly, co- localisation of GM1 and ER-Tracker were also found at the plasma membrane of apoptotic cells (Figure 25, G - I).

3.17 During apoptosis intracellular ganglioside GM1 is relocated to the plasma membrane of neutrophil granulocytes

a) PMN p. 2: p. 1: shrunken viable or apoptotic full size apoptotic

0h 1

5h 1

22h 2 1

45h 2 1 log SSc count count log FL FSc log FL b) culture 0h 5h 22h 45h

cells p.1 98,4 98,1 47,9 19,6 [%] p.2 1,6 1,9 52,1 80,4

CT-B p.1 95,7 70,3 35,0 9,3 cells p.2 91,0 95,1 MFI p.1 10,6 6,3 3,2 2,1 CT-B p.2 18,9 20,5

Figure 24: Time course of CT-B binding to ageing PMN analysed by flow cytometry. a) The population of viable PMN (p.1) and ageing PMN at various time points (p.1 and p.2) are displayed in FSc / SSc dot blots. Binding to populations 1 and 2 of CT-B was analysed and is displayed in histograms. Note: During apoptosis binding to full size apoptotic PMN of CT-B get increasingly lost. On shrunken, apoptotic PMN CT-B binding reappears. Only PMN which excluded PI were analysed. b) Table lists the amounts of cells and the amount of CT-B positive cells in populations 1 and 2, as well as the MFI of CT-B binding to populations 1 and 2.

3.18 Exposure of NPn binding sites, calnexin, and GM1 on apoptosing PMN is abrogated when apoptotic membrane blebbing is blocked

PMN CT-B ER-Tracker merge ABC intracellular, viable

DE F surface, viable

GHI surface, apoptotic

Figure 25: Examination of PMN stained with ER-Tracker and CT-B by fluorescence microscopy. Freshly isolated and aged PMN (20 hours in vitro culture) were labelled with the ER- Tracker (red), Aditionally surface and intracellular binding of CT-B (green) was analysed. Note: CT-B and ER-Tracker co-localised intracellular in permeabilised cells (A - C), and on the surfaces of apoptotic cells (G - I). On the surfaces of viable PMN only CT-B was detected in a patchy pattern (D + F). The ER-Tracker did not stain the surface of viable cells (E + F).

Taken together, ER-Tracker and GM1 (Figure 25) but also Golgi-Stain and calnexin (shown in Figure 15 and Figure 16, respectively) lost its uniform intracellular distribution and were found translocated the surfaces of apoptotic cells. This suggests that in later phases of apoptosis the membranes derived from ER and/or Golgi capture the cell surfaces.

3.18 Exposure of NPn binding sites, calnexin, and GM1 on apoptosing PMN is abrogated when apoptotic membrane blebbing is blocked

The relocation of internal membranes on the surfaces of apoptotic cells was only observed after cell shrinkage in late stages of apoptosis. This suggests that

3.18 Exposure of NPn binding sites, calnexin, and GM1 on apoptosing PMN is abrogated when apoptotic membrane blebbing is blocked apoptotic membrane blebbing and the resulting loss of plasma membrane and cell volume are important events preceding the exposure of internal membranes. To improve bleb formation and cell shrinkage as prerequisite for the access of internal membranes to the cell surfaces, apoptotic cell blebbing of PMN undergoing spontaneous apoptosis was abrogated using the Rho- associated protein kinase- (ROCK) inhibitor Y-27632. Figure 26 a shows apoptotic PMN after 24 hours in vitro culture in the presence or absence of Y- 27632. Mainly one population of not shrunken early and late apoptotic cells (p.4) were found in presence of the ROCK inhibitor. Staining with AxV and PI confirmed that the cells underwent apoptosis (Figure 26 a, p.4). In Figure 26 b the binding of NPn, anti-calnexin, and CT-B to apoptosing PMN treated with the ROCK inhibitor Y-27632 or not were analysed. Even if the cells were dead; in the presence of the bleb inhibitor no exposure of mannose containing epitopes, calnexin, and GM1 was observed. This is in striking contrast to cells not treated with Y-27632. The results suggest the exposure of ER derived membranes as consequence of membrane blebbing, to substitute plasma membrane that was lost due to the blebbing process.

3.19 In late stages of apoptosis KDEL receptor transgenic HeLa cells expose ER membrane on their surfaces

KDEL receptor-GFP transgenic HeLa cells were used to monitor the relocation of internal membranes to apoptotic cell surfaces in consequence of bleb formation and loss of plasma membrane (Figure 27). The KDEL receptor is an ER/cisGolgi compartment-located integral membrane protein (Cabrera et al., 2003). The plasma membranes of the transgenic HeLa cells were stained red (Vybrant DiI dye) whereas the ER membranes were seen as green fluorescent (KDEL receptor-GFP). In viable cells ER membranes are located inside the cells (Figure 27 a). This composition of ER membrane completely changed in apoptosing cells. KDEL receptor-GFP transgenic HeLa cells were induced to

3.19 In late stages of apoptosis KDEL receptor transgenic HeLa cells expose ER membrane on their surfaces

a) PMN Y-27632 100 0h 24h 24h 80 60 40

cells [%] 20 1 3 2 4 0

log SSc p.1 p.2 p.3 p.4 FSc p.1: viable 0h 24h 24h p.2: full size, early apoptotic Y-27632 p.3: shrunken, late apoptotic AxV - / PI - AxV + / PI - p.4: full size, early and late apoptotic AxV + / PI +

Figure 26: Analysis of NPn, anti- b) NPn calnexin, and CT-B binding to 3 * * apoptosing PMN treated with the 2 ROCK inhibitor Y-27632 by flow 1 cytometry. 0 0h 24h 24h PMN were analysed that underwent Y-27632 spontaneous apoptosis in the presence α calnexin or absence of the ROCK inhibitor Y- 3 ** ** 27632 during 24 hours in vitro culture. 2 a) Populations of freshly isolated and MFI 1 aged PMN are presented in FSc / SSc

0 dot blots. The viable, apoptotic or 0h 24h 24h Y-27632 necrotic status of the cells were CT-B analysed by staining with AxV and PI. * * 8 Note: In the absence of Y-27632 two 6 4 apoptotic populations were found: a full 2 size one (p.2), and a shrunken one 0 0h 24h 24h (p.3). In the presence of Y- 27632 bleb Y-27632 formation is abrogated and, therefore, only one population of not shrunken early and late apoptotic cells was detected (p.4). b) Binding to populations 1, 2, 3, and 4 of NPn, anti-calnexin, and CT-B is compared. Note: Apoptotic PMN that had aged in the presence of the blebbing inhibitor showed a similar binding of NPn, anti-calnexin, and CT-B as viable PMN. Only cells, which excluded PI were analysed. * p<0.05; ** p<0.01

3.19 In late stages of apoptosis KDEL receptor transgenic HeLa cells expose ER membrane on their surfaces

KDEL receptor-GFP-transgenic HeLa cells

a) KDEL R-GFP DII merge A B C D

240 min 255 min 270 min 285 min

UV-B

300 min 315 min 330 min 345 min

360 min 375 min 390 min 405 min

red: plasma membrane (DII) green: ER (KDEL R-GFP)

Figure 27: Monitoring of apoptosing KDEL receptor-GFP-transgenic HeLa cells by fluorescence microscopy. a) Location of the ER membrane (green fluorescent KDEL receptor-GFP) in viable transgenic HeLa cells is shown. The plasma membrane is labelled with DiI (red). Note: The ER was clearly found inside the cell near the nucleus. b) UV-B induced apoptosis in KDEL receptor-GFP-transgenic HeLa cells was monitored in the time laps as indicated. The distribution of the plasma membrane (red) and the ER membrane (green) were analysed. Note: After 240 minutes plasma membrane starts blebbing followed by translocation of ER membranes to the surfaces of the cells (300 min). Later on, also ER membrane derived blebs are released.

3.20 Exposure of immature glycoproteins is not sufficient to promote clearance of viable cells undergo apoptosis by irradiation with UV-B. The distribution of the plasma membrane and the ER membrane were monitored in time-lapse as shown in Figure 27 b. The plasma membrane started blebbing already 240 minutes after UV-B. ER membrane was translocated toward the blebbing plasma membrane and, ultimately, occupied the cells’ surfaces where the plasma membrane was completely lost (300 min). Interestingly, 315 minutes after irradiation the KDEL receptor positive membranes were also released via blebbing. These results clearly show that during apoptosis ER membrane is transported to the cells’ surfaces to substitute plasma membrane that got lost during the blebbing process.

3.20 Exposure of immature glycoproteins is not sufficient to promote clearance of viable cells

A contribution of lectins to the clearance process of apoptotic cells is well established. We performed in vitro phagocytosis assays to investigate whether the glycosylation pattern of shrunken late apoptotic cells serves as signal for their recognition and uptake. Untreated and UV-B irradiated PBL as well as PBL treated with a cocktail of processing inhibitors were applied as prey for murine peritoneal macrophages. After a 3 hours incubation period at 37°C of macrophages and prey, and prey alone the cell counts in the supernatants were determined by flow cytometry. Afterwards the amount of cleared cells was calculated and presented as clearance index in Figure 28 a. The clearance index of viable untreated cells (wo) which served as negative controls and that of viable PBL carrying an apoptotic glycosylation pattern (induced by glycosylation inhibitors) did not differ significantly. The positive control group of irradiated PBL was cleared significantly better than the viable cells.

Since it is known that soluble serum factors are mediators in the apoptotic cell uptake the prey was incubated with 10% autologous human serum for 1 hour at 37°C before addition to the macrophages. However, opsonisation with

3.20 Exposure of immature glycoproteins is not sufficient to promote clearance of viable cells autologous serum had no effect on the phagocytosis of the prey. The clearance indices of viable cells and viable cells with the apoptotic surface glycosylation did not significantly differ (Figure 28 b, wo and processing inhibitors) and were both significantly lower than the clearance index of apoptotic cells (UV-B).

a) PBL b) opsonised PBL

40 *** 40 *

30 30

20 20

10 10 clearance IDX clearance IDX 0

0 g g wo wo UVB UV-B inhibitors rocessin inhibitors rocessin p p

Figure 28: In vitro clearance assays with human PBL. Viable (wo), apoptotic (UV-B) PBL, and viable PBL with an apoptotic surface glycosylation pattern (processing inhibitors) were used as prey in the phagocytosis assays. a) Clearance (presented as clearance index [IDX]) of the three distinct prey by murine peritoneal macrophages is shown. Note: Viable and processing inhibitor treated PBL were cleared similar and only to a less extent. Clearance of apoptotic PBL was significantly better when compared with viable ones. b) Clearance of prey that was opsonised with active autologous serum is shown. Note: Opsonisation of the distinct prey did not enhance their clearance.

4 Discussion

The aim of this work was to analyse alterations in the glycocalyx and the composition of plasma membranes of cells undergoing apoptosis to identify potential “eat me” signals especially of late apoptotic cells. It was found that early in apoptosis mature glycostructures as well as the ganglioside GM1 got lost from the cell surface. Later in apoptosis when the dying cells were shrunken due to extensive membrane blebbing the exposure of internal structures such as immature glycosides, ER resident proteins and ER derived GM1 on the cell surface was detected. Importantly, the late apoptotic shrunken cells had still an intact membrane. It was shown that during the blebbing process ER membranes enter the cell surface and substitute lost plasma membrane. However, although the involvement of carbohydrates in apoptotic cell clearance is known it could not yet been demonstrated that immature glycostructures found on shrunken apoptotic cells play a role in the recognition process.

4.1 NPn, GSL II, and UEA I are suitable lectins for analysing the glycosylation status of dying cells

Lectins are proteins from many sources (plants, viruses, microorganisms and animals), which all share the common property of binding to defined sugar structures (Rudiger and Rudiger, 1978). Typically, lectin targets are carbohydrate moieties of complex glycoconjugates in virtually all cell walls or biological membranes. Lectin binding to these targets may change the physiology of the membrane and may cause agglutination, mitosis, or other biochemical changes in the cell. Because of their strong specificity toward a particular carbohydrate structure lectins provide researchers with a powerful tool to explore a multitude of biological structures and processes.

4.1 NPn, GSL II, and UEA I are suitable lectins for analysing the glycosylation status of dying cells

Many members of the lectin protein family are known to agglutinate but also to lyse cells (De Muelenaere, 1965; Greer and Pusztai, 1985; Stirpe et al., 1982). Most of the 23 lectins that were tested revealed such cytotoxic effects. Exemplarily shown for DSL and RCA I, many lectins (ConA, ECL, Jacalin, LCA, LEL, MAL, PHE, PHL, PSA, STL, WGA) exhibited membrane destructive properties and consecutively changed the morphology of model cell lines (Jurkat, U937, Raji) and primary cells (PBL, PMN). A few other lectins (GSL I, PNA, SJA, SBA, and VVL) differentially destroyed one or two of the cell types. It is of major importance to preserve the membrane integrity during the staining procedure. Otherwise lectins could get access to internal lectin binding epitopes falsifying the data on surface glycosylation. Therefore, all membrane disturbing lectins were not useful for reliable evaluations of apoptosis related changes of the plasma membranes glycosylation status and, hence, were excluded from the panel.

Beside the criteria not to change the morphology of the cells, a lectin has to fulfil two more criteria to be suited for analysing changes in the glycocalyx of viable and dying cells. First, the lectin should differentially bind to viable and dying cells, and second, this effect should be conserved for many cell types. The lectins derived from Narcissus pseudonarcissus (NPn), Griffonia simplicifolia (GSL I I), and Ulex europaeus (UEA I) fulfilled all these criteria when tested with the cell lines Jurkat, U937, Raji as well as primary PBL and PMN. The morphology of the cells did not change during the lectin staining procedure and showed the same features as cells stained with Annexin V (AxV). Furthermore, the three lectins produced higher binding signals on dying cells than to viable ones. Necrotic cells displayed even more lectin binding sites than apoptotic ones. Finally, PMN, PBL, and three different cell lines showed similar results. Together, the results establish GSL II, NPn, and UEA I as tools to detect apoptosis related alterations of the surface glycosylation.

4.2 After shrinkage apoptotic cells expose immature glycoproteins

Alterations in the glycosylation of plasma membrane structures during apoptosis have been suggested a long time ago (Morris et al., 1984). Many experiments were performed to analyse sugar moieties exposed on the surfaces of distinct apoptotic cell types including murine thymocytes (Beaver and Stoneman, 1999; Morris et al., 1984; Russell et al., 1998), neonatal rat liver cells and fibroblasts (Dini et al., 1992), several human colon carcinoma cell lines (Azuma et al., 2002; Rapoport and Pendu, 1999) and human PBL (Falasca et al., 1996) and neutrophils (Hart et al., 2000).

In this work alterations in surface glycosylation of the human hematopoietic cell lines Jurkat, Raji, U937 and human primary PBL and PMN undergoing apoptosis were studied via the following non lytic lectins: NPn, GSL I, and UEA I. It was observed that the lectin binding increased during apoptosis in all cell types. Furthermore, the increased binding was restricted to populations of late apoptotic cells that were already shrunken but still had ion selective membranes (PI impermeable). Latter was shown comparing the lectin binding kinetics during the time course of apoptosis with early (exposure of PS, loss of ∆Ψm) and late (DNA fragmentation) markers of apoptotic cell death. Early after apoptosis induction, the cells augmented PS exposure on their surfaces and the membrane potential of their mitochondria (∆Ψm) declined, but no loss of cell volume was to be observed. The lectin binding capabilities of those early apoptotic cells did not change. Later in apoptosis, when the cells had shrunken, PS exposure and the loss of ∆Ψm increased further. Interestingly, these shrunken cells then showed an increase in the binding of NPn, GSL II, and UEA I, although they were still impermeable for PI. The loss of ∆Ψm and the exposure of PS as typical markers for early apoptosis preceded the augmentation of lectin specific epitopes on the cell surface. Furthermore, cells with increased lectin binding and cells with fragmented DNA, which is typical for late apoptosis, occurred with a similar kinetic in the time course of apoptosis. This indicates that, first the enhanced lectin binding is not a result of e.g. a

4.2 After shrinkage apoptotic cells expose immature glycoproteins

chemical modification induced by the agent but, second is due to plasma membrane changes occurring during late phases of apoptosis.

According to the binding specificity of NPn, GSL II, and UEA I an increase in the number on late apoptotic cells of mannose- (Man), N-acetylglucosamine- (GlcNAc), and fucose- (Fuc) containing epitopes, can be concluded. These findings are in part in agreement with other studies. For example in rat hepatocytes increased exposure of mannose, N-acetylglucosamine, fucose, and also galactose (Gal) residues have been reported (Dini et al., 1992). Apoptotic PBL were shown to expose a great number of mannose, galactose, and N-acetylglucosamine moieties, whereas, in contrast to the results of this work, the number of fucose residues did not change when compared to viable PBL (Falasca et al., 1996). However, the authors of these two studies did not indicate the stage of apoptosis when they analysed the dying cells surface glycosylation. In murine thymocytes a decrease of N-acetylglucosamine containing structures in early apoptosis was observed. Later in apoptosis the amount of N-acetylglucosamine as well as of fucose residues increased (Beaver and Stoneman, 1999; Russell et al., 1998). The expression of membrane glycoproteins containing α-d-mannose and β-d-galactose was shown on late apoptotic murine leukemic cells L1210 and fibroblast L929 as well as on apoptotic human MCF-7 and Jurkat cells (Bilyy et al., 2004). Studies of the glycosylation in the late phase of apoptosis from several colon carcinoma cell lines revealed an increase of galactose containing carbohydrates but a decrease of fucosylated structures. The amount of mannose residues remained unchanged (Azuma et al., 2002; Rapoport and Pendu, 1999).

Summarising data from the present work and from former studies, carbohydrates that become increasingly accessible on surfaces of apoptotic cells mainly encompass fucose, galactose, N-Acetylgalactosamine (GalNAc), N- acetylglucosamine, and mannose (Table 3). However, there are controversial data. Few studies reported unchanged or decreased exposure of some of these sugars during apoptosis (Azuma et al., 2002; Azuma et al., 2000; Beaver and

4.2 After shrinkage apoptotic cells expose immature glycoproteins

Stoneman, 1999; Hart et al., 2000; Morris et al., 1984; Rapoport and Pendu, 1999; Russell et al., 1998). These conflicting results are often suggested to be due to the different cell types under investigation. It is also important to exactly define the stage of apoptosis when the glycosylation status is analysed. In this work, time kinetic analyses of lectin binding during apoptosis revealed increased exposure of carbohydrates exclusively in late phases of apoptotis. This is consistent with results from other reports (Beaver and Stoneman, 1999; Russell et al., 1998). It should be mentioned that often lectins with a different fine specificity were used for the detection on apoptotic cells of specific carbohydrates. Lectin binding to a certain target is not only determined by the specific sugar residue but also by the sugar’s conformation, its position in the sugar chain, and how it is linked to neighbouring carbohydrates. Additionally, the sugar’s surrounding, including neighbouring carbohydrates and amino acids, plays an essential role for the affinity of the lectin sugar interaction. Hence, the affinity between a lectin and its target varies considerably upon small changes in the target’s structure. This would explain the conflicting data from Hart et al. showing decreased expression of mannose, GLcNAc, galactose and fucose moieties on apoptotic (CD16low) neutrophils (Hart et al., 2000). For detecting these sugar epitopes they used ConA, WGA, PNA, and TGP respectively. Furthermore, it has been shown in this work that some lectins exert strong cytotoxicity on various cell types including the lectins ConA, WGA, and PNA, that had been used as carbohydrate epitope detecting agent. The authors did not test if the lectins damaged the neutrophils during the staining procedure.

Augmentation of mannose, N-acetylglucosamine, and fucose residues on the surfaces of shrunken apoptotic cells have been presented in this work for five human cell types (Jurkat, Raji, U937, PBL and PMN) after apoptosis induction via three different stimuli. This suggests the exposure of these carbohydrates to be a general event of late apoptosis in various cell types. Interestingly, these sugar structures are usually found as terminal sugar residues on immature glycoproteins, which are modified during protein transport through the ER and

4.2 After shrinkage apoptotic cells expose immature glycoproteins

# # # Dini, 1992 Dini, 1992 Dini, 1992 Dini, 1992 Hart, 2000 Hart, 2000 Hart, 2000 Hart, 2000 Hart, 2000 Bilyy, 2004 Reference Bilyy, 2004 Bilyy, 2004 Morris, 1984 Morris, 1984 Morris, 1984 Beaver, 1999 Azuma, 2002 Azuma, 2002 Azuma, 2000 Azuma, 2002 Beaver, 1999 Azuma, 2002 Azuma, 2000 Russell, 1998 Falasca, 1996 Falasca, 1996 Falasca, 1996 Falasca, 1996 Russell , 1998 Rapoport, 1999 Rapoport, 1999 Rapoport, 1999 Rapoport, 1999 Rapoport, 1999 Rapoport, 1999

)

MAL II , PNA SNA VAA , ( , LFA LPA LEA PLA AAL TGP TGP PNA PNA SNA SNA SNA PNA PHA NPn UEA UEA RCA MAL RCA RCA PSL MAA WGA WGA WGA WGA WGA UEA I GSL II Con A Con A Con A Con A SNA , RCA MAA RCA I LPA ow ow ow ow

l l l l l

y y y y y y y y y n.d. n.d. n.d. n.d. n.d. late late n.d. late late n.d. n.d. n.d. late late late n.d. n.d. n.d. late late late late earl earl earl earl earl earl earl earl earl CD16 CD16 CD16 CD16 CD16 Stage of Apoptosis Lectin SW-48 , SW-48 , HT-29

U937 U937 U937 , , , , i i i j j j HT-29 tes tes tes , y y y HT-29 tes tes tes , Ra Ra Ra , , , murine P815 y y y

, Jurkat rat 208F fibroblasts rat 208F fibroblasts , , , moc moc moc MCF-7 tes SW-48 SW-1116 , y y y SW-48 , , moc moc moc y , PBL y y y PBL PBL PBL PMN HT-29 HT-29 PMN Cells PMN PMN HT-29 HT-29 Jurkat Jurkat Jurkat Jurkat Jurkat PBL SW-1116 SW-707 , , , , , murine L929 fibroblasts murine murine L929 fibroblasts murine SW-1116 , , , moc PRO HT-29 y Jurkat rat th rat th rat th PMN PBL PBL SW-707 , PBL , , rat 208F fibroblasts , murine th murine th murine th neonatal rat liver cells MCF-7 MCF-7 LS-174T , , SW-707 , PMN PMN PMN murine th LS-174T PRO , Jurkat Jurkat neonatal rat liver cells neonatal rat liver cells PRO

sure d d d d d d d d d ed ed ed ed ed ed reported to change their surface exposure on cells undergoing apoptosis. increas increas increas increas increas increas decrease decrease decrease decrease decrease unchange unchange unchange unchange

Fucose GalNAc GlcNAc Mannose Sialic acid Galactose Carbohydrate Surface Expo Carbohydrate Surface Table 3: Carbohydrates

4.3 Before shrinkage apoptotic cells lose mature glycoproteins

Table 3: The table summarises results from this work (marked with #) and from various other studies. Carbohydrates reported to change their surface exposure during apoptosis mainly encompass fucose, galactose, N-acetylgalactosamine (GalNAc), N- acetylglucosamine (GlcNAc), mannose, and sialic acid. Controversial results may be due to the cell types that had been analysed, the stage of apoptosis, or the use of lectins with different fine specificities as carbohydrate detecting agents.

Golgi compartments of cells (Figure 29). In necrotic cells great amounts of mannose, N-acetylglucosamine, and fucose residues were found. When protein processing had been blocked using several processing inhibitors proteins that preserved their immature glycosylation form were transported to the surfaces. Immature sugar structures could be detected by increased binding of NPn and GSL II. In non apoptotic cells the majority of these sugars are located intracellulary. In late apoptotic stages they get exposed on the cell membrane suggesting that internal membranes containing incompletely processed proteins are translocated to the cells’ surfaces under these conditions. The incorporation of new, intracellular derived membranes in the plasma membrane of apoptotic cells have been proposed for a long time (Duvall et al., 1985). At the same time, a second suggestion has been made explaining the apoptotic phenomenon of alterations in the glycocalyx. According to this hypothesis the increase of mannose, N-acetylglucosamine, galactose and fucose epitopes on apoptotic cells is achieved by cleavage of terminal residues, such as sialic acids, of the glycoproteins sugar chains (Duvall et al., 1985). This will be discussed in the next chapter.

4.3 Before shrinkage apoptotic cells lose mature glycoproteins

Sialic acids on the cell surfaces typically comprise the non reducing ends of sugar chains linked to glycosphingolipids or to mature N- and O-linked glycoproteins. Most commonly, they are appended to an underlying galactose

4.3 Before shrinkage apoptotic cells lose mature glycoproteins

or GalNAc via a α-2.3- or α-2.6-glycosidic linkage. The lectins derived from Maackia amurensis (MAL I) and Sambucus nigra (SNA) specifically bind to carbohydrate chains containing sialic acid; MAL I particularly to the αNeu5Ac(β2-3)Gal(1-4)GlcNAc/Glc sequence (Imberty et al., 2000) and SNA preferred to the αNeu5Ac(α2-6)Gal/GalNAc sequence (Shibuya et al., 1987). MAL I and SNA, therefore, have a high affinity for structures of mature glycoproteins. During apoptosis binding to the cell surfaces of both lectins decreased indicating a loss of sialic acid containing structures. The desialylation process seems to be a universal phenomenon of apoptosis since it was found in various cells undergoing apoptosis, including Jurkat, PBL, and PMN (shown in this and other works (Azuma et al., 2000; Hart et al., 2000)) as well as several colon carcinomas (Azuma et al., 2002; Rapoport and Pendu, 1999) (summarised in Table 3). It is assumed that the loss of sialic acids is caused by sialidases released from the apoptosing cells. Indeed, increased sialidase activity has been reported in etoposide treated Jurkat cells undergoing apoptosis (Azuma et al., 2000). The translocation of an intracellular sialidase into the plasma membrane has been suggested to be responsible for desialisation as observed in activated neutrophils (Cross and Wright, 1991) has been suggested. The secession of sialic acids is associated with increased adhesiveness and decreased surface charge (Gallin, 1980). Therefore, desialisation of the plasma membrane of apoptotic cells may facilitate recognition and/or uptake by phagocytes of apoptotic cells. However, in this work it is shown that apoptosis related loss of sialic acids was not significantly blocked in the presence of a broad acting sialidase inhibitor. This indicates that sialidases only partially contribute to the surface desialisation occurring during execution of apoptosis.

Reduced binding of MAL I and SNA have been observed in both shrunken apoptotic cells and apoptotic cells that still had maintained their sizes. Desialylation can be considered as an early apoptotic event occurring when apoptotic cells start blebbing. We suggest that plasma membrane containing

4.3 Before shrinkage apoptotic cells lose mature glycoproteins

mature sialic acid glycosides are shedded during the blebbing process resulting in a net “desialylation” of apoptotic cell surfaces by loss of mature surface proteins. Furthermore, internal membranes containing immature glycosides replenish the lost membranes leading to increased accessibility of mannose, N- acetylglucosamine, and fucose residues on the cell surfaces. However, experiments showing apoptotic blebs with high degree of sialylation or demonstrating abrogation of desialylation on apoptotic cells in the presence of a bleb inhibitor would substantiate this hypothesis.

Immature Glycoproteins Mature Glycoprotein

MAL I SNA NPn GSL II

UEA I

N-Acetylglucosamine Mannose Fucose

Galactose Sialic Acid

Figure 29: Carbohydrate chains of immature and mature N-linked glycoproteins. During the processing of N-linked glycoproteins intermediates with terminal N- Acetylglucosamin, mannose, and fucose residues are formed. These carbohydrates can be recognised by the lectins GSL II, NPn, and UEA I, respectively. In mature glycoproteins these residues are covered by galactose and sialic acid. The latter is detectable by the lectins MAL I and SNA.

4.4 During apoptosis internal membranes are translocated to the cell surface to substitute plasma membrane that are shed during the blebbing process

Results from the first part of this work clearly indicate the exposure of ER and / or Golgi derived membranes on the surfaces of late apoptotic cells. Indeed, in fluorescence microscopic examinations a redistribution of ER and Golgi membranes toward the cells surfaces of apoptosing PMN could be observed. However, there are several reports showing fragmentation of the Golgi apparatus during apoptosis (Lane et al., 2002; Mancini et al., 2000), while the ER was observed to become distended (Sesso et al., 1999). Furthermore, a close association of ER membranes with the plasma membrane (Latz et al., 2004) and with microtubules which, naturally, protrude to the plasma membrane (Terasaki et al., 1986) has recently been described. It has also been shown that the loss of surface membrane of phagocytes occurring during particle internalisation is substituted by the ER (Gagnon et al., 2002). Therefore, the ER membrane is a prime candidate for the incorporation into the plasma membrane.

Calnexin is an ER-resident chaperone normally retained within the cell compartment. In this work, this ER derived protein was found on the surfaces of apoptotic cells. However, a small fraction of calnexin expressed on the surfaces of various cells such as mastocytoma cells, murine splenocytes, fibroblast cells, and human HeLa cells had already been reported by Okazaki and colleagues (Okazaki et al., 2000). Therefore, cell lines and primary cells were selected that did not expose calnexin if viable. In apoptotic cells calnexin was only found after cell shrinkage, and in the time course of apoptosis, exposure of PS clearly preceded that of calnexin. Experiments with the transfected Ag8.H cell lines confirmed the exposure of ER derived proteins on apoptotic surfaces. During apoptosis the dysfunctional µHC, which is normally retained in the ER (Haas and Wabl, 1983; Kline et al., 1998), translocated to the cell surface. The functional µHC expressed on the surfaces of viable cells was augmented during

4.4 During apoptosis internal membranes are translocated to the cell surface to substitute plasma membrane that are shed during the blebbing process

apoptosis. Both observations were restricted to shrunken apoptotic cells. Together these findings provide evidence that ER membrane is translocated to the cell surfaces in late apoptosis. As co-localisation of calnexin and lectin binding sites in viable cells and, more important, on apoptotic cells was observed it can be assumed that immature glycoproteins anchored in the ER membrane are transported along with the latter to the late apoptotic cell surfaces.

Redistribution to the plasma membrane of ER derived GM1 was also observed in late apoptosis. GM1 is a glycosphingolipid that associates with lipid rafts (Kenworthy et al., 2000) at the cell surfaces and was therefore found on freshly isolated neutrophils in a patchy pattern. During early neutrophil apoptosis plasma membrane GM1 was substantially depleted. This might happen by covalent modification, blebbing of GM1-containing membrane rafts, or relocation of the molecule to intracellular membranes. Apoptotic neutrophils were demonstrated to undergo specific alterations in their receptor profiles (Hart et al., 2000). It has been suggested that these alterations may contribute to functional downregulation and phagocyte recognition of apoptotic neutrophils. However, mechanisms leading to early apoptotic related molecular surface changes such as loss of GM1 are still elusive. The blebbing of membrane rafts containing GM1 is a likely mechanism that would also explain the simultaneous loss of sialic acid glycosides. Interestingly, in late apoptosis, GM1 re-appeared on the surfaces but only of shrunken neutrophils. At this apoptotic stage cells are thought to have stopped synthesis of new membrane components. Given that apoptotic cells lose plasma membrane through shedding of blebs it can be assumed that GM1 re-exposure relies on substitution of lost plasma membrane by intracellular membranes (e.g. from the ER). This is corraborated by the facts that GM1 is also found in the ER of viable neutrophlis and that both ER and GM1 co-localised on the cell surfaces in late apoptosis.

The findings that ER resident proteins and lipids translocate to the surfaces of apoptotic cells after shrinkage provide evidence that plasma membrane

4.4 During apoptosis internal membranes are translocated to the cell surface to substitute plasma membrane that are shed during the blebbing process

blebbing from the cell surface is replenished by ER membranes, which then form the outer envelope of the late apoptotic cells. Experiments with the bleb inhibitor Y-27632 (Sebbagh et al., 2001) additionally confirmed this hypothesis of membrane exchange. When the blebbing process was abrogated no augmentation of immature glycoproteins, no exposure or re-exposure of ER derived proteins and lipids, respectively, on the surfaces of apoptotic cells was observed. This suggests the surface capping of ER derived membranes as a consequence of membrane blebbing. Live-cell imaging of KDEL receptor-GFP transgenic HeLa cells allowed us to monitor the apoptosis related ER to plasma membrane trafficking. First the plasma membrane blebbed from the surface and then ER membranes incorporated in “plasma-membrane-free” areas. Due to this mechanism the dying cell was always surrounded by an intact outer membrane. After a while, ER membranes were also released via blebbing. Just recently, active remodelling of ER derived membranes into apoptotic blebs was reported (Lane et al., 2005). In the blebs ER membranes were found as a cortical layer close to the plasma membrane but not incorporated in the latter. Although the authors did present convincing data of the cortical formation of the ER they did not show such an ER layer surrounded by a plasma membrane. Maybe they did not consider the massive loss of plasma membrane during blebbing. The surface area of three dimensional bodies increases if divided into multiple smaller particles. This is also the case for apoptotic cell blebbing. As apoptotic cells do not lose their membrane integrity upon blebbing, new membranes have to be provided by the dying cells at the best from their internal stores. From the apoptotic cells’ point of view it makes sense to use membranes that already exist since biosynthesis of new membranes would consume energy, but ATP is limited during apoptosis (Chiarugi, 2005).

4.5 Carbohydrates of immature glycoproteins as “back-up-eat-me” signal of apoptotic cells that have escape early PS dependent clearance?

Phagocytes express a large repertoire of receptors to detect their prey including lipid receptors, scavenger receptors, but also several lectins. Sugar recognition would, therefore, be a reasonable mechanism to swallow apoptotic material. In deed, Kupffer cells (Falasca et al., 1996), macrophages (Duvall et al., 1985), fibroblasts (Hall et al., 1994), and endothelial cells (Dini et al., 1995) recognise apoptotic cells via carbohydrate-specific receptors. The sugar-lectin interaction has been shown classically by inhibition of phagocytosis in the presence of exogenous sugars. Saccharides blocking clearance varied according to the type of both apoptotic and phagocytic cells. For example, N-acetylglucosamine, mannose, and galactose were identified as inhibitory sugars in liver cell phagocytosis (Dini et al., 1992), whereas neutrophil clearance was inhibited by mannose, fucose, and glucose (Hall et al., 1994). However, beside carbohydrate-specific receptors soluble lectins have also been demonstrated to bind late apoptotic cells and to mediate their removal. The mannose binding lectin as well as surfactant protein A and D, all members of the collectin family of proteins, stimulate the ingestion of apoptotic cells via the calreticulin/CD91 mechanism (Ogden et al., 2001; Vandivier et al., 2002). The lectins L-ficolin and H-ficolin participate in apoptotic cell clearance through complement activation (Kuraya et al., 2005). L-ficolins was reported to recognise N-acetylglucosamine whereas N-acetyl-D-galactosamine, N-acetylglucosamine, and D-fucose are suggested as target for H-ficolin (Holmskov et al., 2003). Together all reports support the role of surface sugars found on late apoptotic cells in the clearance process.

Under normal conditions apoptotic cells are recognised and phagocytosed via PS very early in the cell death program. If this immediate clearance fails the dying cells enter late stages of apoptosis characterised by membrane blebbing and cell shrinkage. Interestingly, many adaptor molecules including soluble

4.6 Hypothesis how apoptotic cells keep their silence and avoid immunogenicity

lectins only bind to late apoptotic cells with a shrunken morphology (Hart et al., 2004). This suggests altered glycosylation structures acting as “back-up-eat- me-signal” of late apoptotic cells that had escape early PS dependent clearance. However, in this work clearance via recognition of immature sugar structures could not be demonstrated. In the phagocytosis assays viable lymphocytes mimicking apoptotic surface glycosylation achieved by treatment with processing inhibitors were used as prey. Recognition and phagocytosis of apoptotic cells are complex processes and many molecules on the part of apoptotic cells and phagocytes are involved. Recently, it has been shown that phagocytosis of Fas-triggered Jurkat cells was inhibited in the absence of oxidised PS on the apoptotic cell surface while PS was externalized(Kagan et al., 2002). This suggests not the exposure of a certain molecule but rather the interplay of several “eat me” markers as the important signal of apoptotic cells to become recognised and phagocytosed by macrophages. The lymphocytes treated with processing inhibitors did augment immature glycoproteins on their surfaces but other surface changes, which possibly make in conjunction with the sugar alterations the complex recognition signal, were missing. This explains why clearance of these cells failed in our in vitro assay.

4.6 Hypothesis how apoptotic cells keep their silence and avoid immunogenicity

In this work, apoptotic cells have been demonstrated to loss sialic acids characterising immature glycoproteins early in the cell death process. Later in apoptosis, after shrinkage the cells expose sugar structures of immature glycoproteins as well as the ER resident proteins calnexin and a dysfunctional µHC and ER derived gangliosid GM1. The surface capping of ER membranes to substitute plasma membrane lost due to the blebbing process was concluded and could be verified in live-cell imaging using KDEL receptor-GFP transgenic HeLa cells. Since the ER membranes forming the outer envelop of apoptotic cells was only observed in late stages of apoptosis we hypothesise the following

4.6 Hypothesis how apoptotic cells keep their silence and avoid immunogenicit

(Figure 30): Apoptotic cells translocate internal ER derived membranes to their surfaces to substitute lost plasma membrane and in consequence to maintain their membrane ion selective. Hence, in late apoptotic stages the dying cells are still surrounded by an intact membrane increasingly composed of ER. Thereby, apoptotic cells prevent lysis and the release of immunogenic intracellular material. Furthermore, by the exposure of ER membranes new structures reach the cell surfaces that are not accessible on viable cells. These structures act as late “eat-me” signals explaining the concomitant binding of a multitude of adaptor molecules to late apoptotic cells. Recognition sites on apoptotic cells can (I) be synthesised de novo by a newly expressed enzymatic activity (like the induction of tissue transglutaminase and generation of lysophospholipids), can (II) be generated by (accidental) chemical modification (like oxidation of phospholipids), or can (III) just be generated by the exposure of preformed epitopes which are usually sequestered inside viable cells (like phosphatidylserine). An apoptotic cell soon shuts down its power production and must, therefore, deal parsimoniously with its ATP reservoir. From this point of view, the exposure of preformed structures that are sequestered inside viable cells is an economical method for the concomitant generation of several “eat- me” signals on surfaces of dying cells.

Taken together, translocation of ER membranes to the cell surface comprises an important energy saving strategy of apoptotic cells to expose “back-up-eat- me” signals if they have escaped early clearance. More importantly, this procedure prevents cell lyses and keeps the cells calm until their proper removal by phagocytes.

4.6 Hypothesis how apoptotic cells keep their silence and avoid immunogenicity

et et val. nals lectin SP-A /D g ? ? late “eat me“ si late “eat me“ ? costructure y l g late clearance: ? C1q immature ? ? APOPTOSIS dding and blebbing resulting in a massive loss of plasma in a massive loss dding and blebbing resulting ectivity of their plasma membrane and initiate their PTX-3 CRP mbrane. Thereby, structures sequestered inside of viable cells g inside of viable sequestered mbrane. Thereby, structures s. late apoptotic cells and to be involved in late clearance. in late clearance. be involved cells and to late apoptotic ß2gp1 Gas6 E8 early late rface membranes and preventing cell lysis the late apoptotic cells translocate ER lysis the cell preventing rface membranes and PS which is bound by several adaptor molecules initiating apoptotic cell remo apoptotic cell molecules initiating several adaptor PS which is bound by -me” signal lost plasma me s su MFG- own to opsonise ed late “eat to substitute heir surface AxI ER derived structures (including immature glycostructures) may serve as late “eat me” signal ER derived structures (including immature glycostructures)

early clearance: ER viable rly clearan ion selectivity of their of their ion selectivity daptor molecules kn Phosphatidylseri ne Early in apoptosis cells expose on t Early in apoptosis cells Cells that escape this eamembrane. Maintaining membrane (green) to their surfaces These cell surface. access to the ce undergo extensive membrane bu by several a recognised Figure 30: Hypothesis how lateFigure apoptotic cells maintain ion sel clearance by exposure of ER deriv

5 Materials

Reagents, auxiliary materials, and instruments used in this work are summarized in the Tables 4, 5 and 6, respectively. Not listed are chemicals, materials and instruments which are considered to be part of the general laboratory equipment.

Table 4: Reagents

Reagents Source

1-Deoxymannojirimycin Calbiochem, Merck, Darmstadt, Germany 2.3-Dehydro-2-deoxy-N-Acetyl- Calbiochem, Merck, Darmstadt, neuraminic Acid Germany Ampuva Fresenius, Bad Homburg, Germany Amylum Sigma, Munich, Germany Anti-µHC FITC Southern Biotech, Birmingham, AL, USA Anti-C1q FITC DAKO, Glostrup, Denmark Anti-calnexin FITC BD Pharmingen, Heidelberg, Germany Anti-CD14 linked magnetic Miltenyi, Bergisch Gladbach Germany microbeads Anti-CD16 PE BD Pharmingen, Heidelberg, Germany Anti-CD49d PE BD Pharmingen, Heidelberg, Germany AxV Responsif, Erlangen, Germany

BODIPY TR C5 ceramide Molecular Probes Invitrogen, Karlsruhe, Germany Castanospermine, Calbiochem, Merck, Darmstadt, Germany Concanavalin A Rhodamine Vector Laboratories, Linaris, Wertheim, Germany

5 Materials

Reagents Source

CT-B FITC Sigma, Munich, Germany Cytofix/CytoPerm PlusTM Kit BD Pharmingen, Heidelberg, Germany Datura stramonium lectin FITC Vector Laboratories, Linaris, Wertheim, Germany

DiO6C(3) Sigma, Munich, Germany DMEM:F12 Gibco Invitrogen, Karlsruhe, Germany DMSO Roth, Karlsruhe, Germany Dolichos biflorus agglutinin Vector Laboratories, Linaris, Rhodamine Wertheim, Germany EDTA Sigma, Munich, Germany ERTracker™ BlueWhite DPX Molecular Probes Invitrogen, Karlsruhe, Germany Erythrina cristagalli lectin FITC Vector Laboratories, Linaris, Wertheim, Germany Ethanol Sigma, Munich, Germany FCS Gibco Invitrogen, Karlsruhe, Germany FluoroTag fluorescein isothiocyanate Sigma, Munich, Germany (FITC) conjugation kit G418 Roth, Karlsruhe, Germany Glutamine Gibco Invitrogen, Karlsruhe, Germany Griffonia simplificolia lectin I Vector Laboratories, Linaris, Rhodamine Wertheim, Germany Griffonia simplificolia lectin II FITC Vector Laboratories, Linaris, Wertheim, Germany Hank`s buffered Salt Solution (HBSS) Pharmacy, Clinic University Erlangen, Erlangen, Germany HEPES Merck, Darmstadt, Germany Human C1q Calbiochem, Merck, Darmstadt, Germany Human PTX3, biotinylated Prof. A. Manfredi, Clinical Immunology, San Raffaele Scientific Institute, Milano, Italy Hypoxanthine Sigma, Munich, Germany IgG FITC BD Pharmingen, Heidelberg, Germany

5 Materials

Reagents Source

Jacalin FITC Vector Laboratories, Linaris, Wertheim, Germany Lens culinaris agglutinin Rhodamine Vector Laboratories, Linaris, Wertheim, Germany Lycopersicon esculentum lectin FITC Vector Laboratories, Linaris, Wertheim, Germany Lymphoflot Biotest, Dreieich, Germany Maackia amurensis lectin I FITC Vector Laboratories, Linaris, Wertheim, Germany Mycophenolic acid Sigma, Munich, Germany

NaN3 Merck, Darmstadt, Germany Narcissus pseudonarcissus lectin Vector Laboratories, Linaris, Wertheim, Germany PBS Gibco Invitrogen, Karlsruhe, Germany Peanut agglutinin Rhodamine Vector Laboratories, Linaris, Wertheim, Germany Penicillin Gibco Invitrogen, Karlsruhe, Germany PFA Sigma, Munich, Germany Phaseolus vulgaris erythroagglutinin Vector Laboratories, Linaris, Rhodamine Wertheim, Germany Phaseolus vulgaris leucoagglutinin Vector Laboratories, Linaris, Rhodamine Wertheim, Germany PI Sigma, Munich, Germany Pisum sativum agglutinin Rhodamine Vector Laboratories, Linaris, Wertheim, Germany Ricinus commuins agglutinin I Vector Laboratories, Linaris, Rhodamine Wertheim, Germany Ringer`s solution Delta Select, Pfullingen, Germany RPMI 1640 Gibco Invitrogen, Karlsruhe, Germany Sambucus nigra lectin FITC Vector Laboratories, Linaris, Wertheim, Germany

Sialidase from Vibrio cholerae Calbiochem, Merck, Darmstadt, Germany Sodium citrate Merck, Darmstadt, Germany

5 Materials

Reagents Source

Sodium pyruvate Gibco Invitrogen, Karlsruhe, Germany Solanum tuberosum lectin FITC Vector Laboratories, Linaris, Wertheim, Germany Sophora japonica agglutinin Vector Laboratories, Linaris, Rhodamine Wertheim, Germany Soybean agglutinin Rhodamine Vector Laboratories, Linaris, Wertheim, Germany Staurosporine Alexis, Lausen, Switzerland Streptavidin FITC Southern Biotech, Birmingham, AL, USA Streptomycin Gibco Invitrogen, Karlsruhe, Germany Swainsonine Calbiochem, Merck, Darmstadt, Germany Thymidine Sigma, Munich, Germany Triton-X100 Sigma, Munich, Germany Trypan blue Sigma, Munich, Germany Ulex europaeus agglutinin I FITC Vector Laboratories, Linaris, Wertheim, Germany Ulex europaeus agglutinin I Vector Laboratories, Linaris, Rhodamine Wertheim, Germany Vicia villosa lectin FITC Vector Laboratories, Linaris, Wertheim, Germany Vybrant™ DiI cell labelling solution Molecular Probes Invitrogen, Karlsruhe, Germany Wheat germ agglutinin Rhodamine Vector Laboratories, Linaris, Wertheim, Germany Wheat germ agglutinin Rhodamine, Vector Laboratories, Linaris, succinylated Wertheim, Germany Xanthine Sigma, Munich, Germany Y27632 Calbiochem, Merck, Darmstadt, Germany

5 Materials

Table 5: Auxiliary Materials

Material Source

Cell culture flasks (50 ml, 250 ml, Greiner bio-one, Frickenhausen, 600 ml) Germany Cell culture plates (6-wells, 12-wells, Greiner bio-one, Frickenhausen, 24-wells, 48-wells) Germany Cellstar PP-tubes (15 ml, 50 ml) Greiner bio-one, Frickenhausen, Germany Combitips plus (0.5 ml; 2.5 ml; 5 ml; Eppendorf, Hamburg, Germany 12.5 ml) EpT.I.P.S. standard (100 µl white, Eppendorf, Hamburg, Germany 200 μl yellow, 1000 μl blue) Falcon 5ml polystyrene round-bottom Becton Dickinson, Bedford, MA, USA tube Gilson micro volume pipettes (P 10, Gilson, Wisconsis, USA P 20, P 100, P 200, P 1000) Laboratory film Parafilm “M” American National Can, USA Lab-Tek chambered coverglass, Nalge Nunc International, Roskilde, german borosilicate, 8 chambers Denmark Micro test tubes Safe-Lock (0.5 ml; Eppendorf, Hamburg, Germany 1.5 ml) Nunc cryo tube vials Nalge Nunc International, Roskilde, Denmark Pipetboy comfort INTEGRA Biosciences, Chur, Switzerland Pipette tips (200 μl yellow, 1000 μl Greiner bio-one, Frickenhausen, blue) Germany Serological pipettes (2 ml, 5 ml, 10 ml, Greiner bio-one, Frickenhausen, 20 ml) Germany Tissue culture dish, PS (94 x 16 mm) Greiner bio-one, Frickenhausen, Germany Vaccum driven disposable filtration Millipore Co., Bedford, USA system

5.1 Cells and culture conditions

Table 6: Instruments

Instruments Source

Auto MACS Milteny, Bergisch-Gladbach, Germany Axiovert 200M inverted microscope, Zeiss, Oberkochen, Germany equipped with a ApoTome and incubator XL-3 for life microscopy Cell culture incubator HERA cell Heraeus, Hanau, Germany Centrifuge Rotina 46 RS Hettich, Tuttlingen, Germany Clean work bench HERA safe Heraeus, Hanau, Germany Coulter XLTM software, version 3 Coulter Electronics Inc., Miami, USA Flow cytometer EPICS XL-MCL / Coulter Electronics Inc., Miami, USA SYSTEM II Inverted System Microscope Olympus Olympus Optical Co., Hamburg, IX70 Germany Leica DMR confocal microscope and Leica, Wetzlar, Germany software version 2.00 Microcentrifuge MiniSpin Eppendorf, Hamburg, Germany Thermomixer comfort Eppendorf, Hamburg, Germany TILLvisION v3.3 software T.I.L.L. Martiensried, Germany Photonics GmbH UV table TFP – M / WL Vilber Lourmat, Marne-La-Vallee Cedex 1, France

5.1 Cells and culture conditions

Primary cells were isolated from venous blood that was drawn from normal healthy volunteers according to institutional guidelines. Blood was anticoagulated by addition 20 U/ml heparin. Peripheral blood mononuclear cells (PBMC) were isolated using Ficoll density-gradient centrifugation. Isolated PBMC were washed twice with phosphate buffered saline (PBS) and remaining platelets were removed by centrifugation through a cushion of fetal calf serum. Peripheral blood lymphocytes (PBL) were obtained by removing monocytes from PBMC by immunomagnetic separation of CD14+ monocytes using

5.1 Cells and culture conditions

anti CD14Ab linked microbeads according to the manufacturer’s instruction. Polymorphonuclear cells (PMN) were isolated by Ficoll density-gradient centrifugation. After centrifugation PMN were collected from the thin cell layer immediately above the erythrocytes. Residual erythrocytes were removed by hypotonic lysis that was performed twice. Viability of PBL and PMN was tested by trypan blue exclusion. It was, without exception, greater than 90%. The primary cells were cultured in R10 (RPMI 1640 supplemented with 10% heat inactivated FCS, 1% glutamine, 1% HEPES, and 1% penicillin-streptomycin) in a humidified atmosphere at 37°C containing 5.5% CO2.

The nonadherent hematopoetic cell lines Jurkat, Raji, and U937 were obtained from American Type Culture Collection (ATCC). Stable transfectants

Ag8.Hα µdys and Ag8.Hα µfct were kindly provided by Prof. H.M Jäck (Division of Molecular Immunology, Department of Internal Medicine III, University of Erlangen-Nürnberg). KDEL receptor-GFP transgenic HeLa cells were kindly provided by Dr. R. Voll (IZKF Research Group 2, Nikolaus-Fiebiger-Center of Molecular Medicine, University of Erlangen-Nürnberg). All cells were cultured in a humidified atmosphere at 37°C containing 5.5% CO2. Jurkat, Raji, U937, and the stable HeLa transfectant were maintained in R10; the medium of Hela cells was additionally supplemented with 1 mg/ml G418 for clonal selection. Stable Ag8.H transfectants were maintained in RPMI 1640 supplemented with 50 U/ml penicillin, 50 µg/ml streptomycin, 5% FCS, 1 mM sodium pyruvate, 2 mM L- glutamine, 1 mg/ml G418, 1.25 µg/ml mycophenolic acid, 250 µg/ml xanthine, 100 µM hypoxanthine and 16 µM thymidine.

5.2 Animals

For the phagocytosis experiments with viable, apoptotic, and processing inhibitor-treated PBL, 6-week-old female Balb/c mice were used. Those mice were obtained from Charles River Wiga (Sulzfeld, Germany).

5.4 Induction of apoptosis and necrosis

5.3 FITC labelling

The lectin derived from Narcissus pseudonarcissus (NPn) as well as annexin V (AxV) were labelled using the FluoroTag fluorescein isothiocyanate (FITC) conjugation kit according to the manufacturer’s instruction. After passage through a gel filtration column to remove unbound FITC, AxV was diluted to 1 mg/ml with Ringer’s solution; NPn was diluted in PBS to a final concentration of 2 mg/ml.

5.4 Induction of apoptosis and necrosis

PMN underwent spontaneous progressive apoptosis during in vitro culture. Apoptosis in PBL, Raji, Jurkat, U937, and the three transgenic cell lines were triggered by either UV-B irradiation or treatment with staurosporine. Raji and U937 were stressed with UV-B at a dose of 270 mJ/cm². PBL, Jurkat cells and all transfecants were irradiated at a dose of 180 mJ/cm2. For apoptosis induction by staurosporine the cells were cultured in R10 containing 2 µM staurosporine or 0.5 µM staurosporine in the case of Jurkat cells. After apoptosis stimulation all cells were cultured under standard conditions.

Primary necrosis in all cells was induced by incubation of the cells at 56°C for 30 minutes.

5.5 Detection of Apoptosis and Necrosis

Monitoring of apoptosis and necrosis was performed by microscopic examination and by flow cytometry using several methods: (I) annexin V (AxV) staining in the presence of propidium iodide (PI) (Koopman et al., 1994; Vermes et al., 1995); (II) detection of cells with sub-G1 DNA content (Nicoletti et al., 1991); (III) monitoring of the mitochondrial membrane potential (Petit et al., 1995); and (IV) morphological changes of the cells [forward scatter (FSc) versus side scatter (SSc)] (Berndt et al., 1998; Hagenhofer et al., 1998).

5.5 Detection of apoptosis and necrosis

The anionic phospholipid phosphatidylserine (PS) is exposed by dying cells early in the apoptotic program. The Ca2+ dependent protein AxV binds PS with a high affinity and is established for detecting apoptotic and necrotic cells. FITC- labelled human AxV was used to flag PS on the surfaces of dying cells. Approximately 1x105 cells were incubated with 500 µl Ringer's solution containing 0.25 µg AxV-FITC and 0.5 µg PI. After 30 minutes at 4°C the samples were immediately analysed by flow cytometry. Dead cells (apoptotic and necrotic cells) exposing PS did bind AxV reflected by increased FL 1 values. PI was added to distinguish between apoptotic and necrotic cells. The plasma membranes of the latter are holey and therefore permeable for PI. Hence, AxV binding PI negative and positive cells are referred to as apoptotic and necrotic, respectively.

Cells dying by apoptosis degrade their DNA. Therefore the nuclear DNA content was measured as the cells plasma membranes were permeabilised with detergent in a hypotonic buffer containing PI. 1x105 cells were treated with 500 µl PI/Triton staining solution (0.1% sodium citrate, 0.1% Triton-X100, and 1 µg/ml PI) and incubated at 4°C in the dark for at least 24 hours before analysis by flow cytometry. During the prolonged incubation the intracellular nucleases completely degraded the cellular RNA as detected by the addition of 1 mg/ml RNase A in a pilot study. Apotosis was detected by nuclei with sub-G1 DNA content reflected by decreased FL-4 values.

The loss of the mitochondrial membrane potential (ΔΨm) is a characteristic hallmark of apoptotic cells. To measure ΔΨm 500 µl freshly prepared (40 mM stock in DMSO diluted 1:106 with Ringer´s solution) fluorescent cationic lipophilic dye 3.3'-dihexyloxacarbocyanine iodide [DiO6C(3)] was added to 1x105 cells. After 30 min incubation at room temperature, the cells were chilled on ice to stop the reaction. Afterwards, the samples were measured by flow cytometry. The loss of ΔΨm was reflected by reduced FL1 values. Heat necrotized cells served as negative controls.

5.7 Flow cytometry and microscopy analysis

During apoptosis cells undergo characteristic morphological changes. As morphological changes typical for apoptosis an increased side scatter (SSc; granularity) and decreased forward scatter (FSc; cell volume) was quantified by flow cytometry. According to the FSc / SSc properties of the cells two populations of apoptotic cells can be gated: a population of full size cells and a population of shrunken cells. The reliability of two apoptotic cell gates was confirmed by staining with FITC labelled AxV in the presence of PI.

Necrosis is characterised by a dye permeable cytoplasmic membrane. It was detected by staining with PI of cells in isotonic tissue R10 or Ringer's solution and calculated by flow cytofluorometry detecting the FL4 fluorescence. Heat necrotised cells served as positive controls.

5.6 Flow cytometry and microscopy analysis

The analyses by flow cytometry were performed with an EPICS™ cytofluorometer (Coulter Hialeah, USA). Excitation was at 488 nm, the FITC fluorescence was recorded on FL1 sensor (515 - 545 nm BP), the PE (R- Phycoerythrin) or Rhodamine fluorescence on FL2 sensor (575 nm BP), and the PI fluorescence on FL4 sensor (600 nm LP). Electronic compensation was used to eliminate bleed through of fluorescence. A minimum of 10.000 cells were measured. Data analysis was performed with Coulter XL™ software, version 3.

Fluorescence microscopy was performed with an IX70 inverted fluorescence microscope (Olympus, Hamburg, Germany). Blue fluorescence of the ER- Tracker™ Blue-White DPX was visualised using excitation at 374 nm and a band-elimination filter D 460/50. FITC fluorescence was acquired with excitation at 460 nm and a band-elimination filter D 520/20. Excitation at 520 nm and a 575-640 filter were used to record red fluorescence of PI or Rhodamine dyes.

Confocal microscopy was performed with a Leica DMR confocal microscope (Leica, Wetzlar, Germany), software version 2.00.

5.7 Analysis of cell surface glycosylation

The surface glycosylation of cells was detected using 23 fluorescence labelled lectins indicated in Table 1. 100 µl cell suspension (1x105 cells) was incubated with 2 µl of the lectins (2 mg/ml) for 30 minutes at 4°C. After adjustment to 500 µl with PBS containing 0.1% PI, the fluorescence was measured by flow cytometry. PI was added to exclude necrotic cells. The staining with 4 µg FITC labelled goat IgG antibody served as control.

Since most lectins exerted toxic effects on the cells (Table 1) the following lectins were used for further analysis of the cells glycosylation status:

1. GSL II-FITC (α- or β-linked N-acetylglucosamine at the non-reducing terminal of oligosaccharides (Hennigar et al., 1986))

2. NPn-FITC (terminal and internal 1.6-linked α-mannosyl units or galactomannans (Kaku et al., 1990))

3. UEA I-FITC or -Rhodamine (α-linked L-fucose residues (Allen et al., 1977))

4. MAL I-FITC (N-acetylsialic acid linked α 2.3 to galactose (Sata et al., 1989))

5. SNA-FITC (N-acetylsialic acid linked α 2.6 to galactose (Shibuya et al., 1987))

MAL I and SNA had to be used in a 25 and 50 times lower concentration, respectively, than above-mentioned. The undiluted lectins were severely affecting shape and viability of the cells. However, the lectin-dilutions used in the experiments were not toxic during the 30 minutes incubation period.

5.9 Inhibition of glycoprotein processing

5.8 Analysis of surface sialic acid levels in the presence of sialidase inhibitor

To estimate the sialidase inhibitor activity Jurkat cells were treated with 1 U/ml sialidase in the presence of various concentrations (0 µM, 5 µM, 10 µM, 20 µM) of sialidase inhibitor (2.3-Dehydro-2-deoxy-N-Acetylneuraminic acid). To determine if sialidases contribute to surface desialisation of apoptosing cells Jurkat in R10 containing 10 µM sialidase inhibitor were induced to undergo apoptosis. For detecting surface sialisation cells were washed and then stained with MAL I or SNA.

5.9 Inhibition of glycoprotein processing

Castanospermine, 1-deoxymannojirimycin, and swainsonine are inhibitors of glycoprotein processing (Elbein, 1984; Fuhrmann et al., 1984; Saul et al., 1983). Jurkat cells were cultured in R10 in the presence of either 50 µg/ml castanospermine or 1 µg/ml swainsonine or an inhibitor cocktail (50 µg/ml castanospremine, 100 µg/ml 1-Deoxymannojirimycin, and 1 µg/ml swainsonine) for 24 hours. PBL were cultured in vitro for 24 hour in R10 containing the inhibitor cocktail. The glycosylation status of the cells was proven by staining with the lectins NPn, GSL II, UEA I, MAL I, and SNA as described above.

5.10 Cell labelling with ER-Tracker and Golgi-Stain

The endoplasmic reticulum (ER) in live cells was labelled using ER-Tracker™ Blue-White DPX. The cell-permeant, photostable Dapoxyl dye is highly selective for ER. Cells were pelletised and resuspend in prewarmed staining solution (R10 containing 1 µM ER-Tracker). After incubation for 60 minutes at 37°C cells were washed twice with R10. Finally, the cells were adjusted to a working concentration of 1 Mio/ml in R10.

5.11 Detection of calnexin exposure

The B ODIPY TR C5 ceramide were used for fluorescent marking of the Golgi complex. The fluorescent sphingolipid penetrates the cell and produces selective staining of the Golgi complex and is, therefore, referred to as Golgi- Stain in this work. Pelletised cells were incubated for 30 minutes at 4°C with 20 µM Golgi-Stain in Hank`s buffered Salt Solution (HBSS) containing 10 mM HEPES. Afterwards the cells were washed twice with R10 and incubated for further 30 minutes at 37°C in fresh R10. Finally, the cells were adjusted to a working concentration of 1 Mio/ml.

5.11 Detection of calnexin exposure

The surface expression of calnexin was detected using an FITC-labelled antibody against an ER luminal domain of calnexin (α-calnexin). The surface staining of the cells was performed as follows: 1x105 cells were incubated at 4°C with 2 µg α-calnexin. After 30 min the cells were washed with PBS and then fixed with 500 µl PBS containing 1% (w/v) paraformaldehyde (PFA). To detect necrotic cells 0.5 µg PI was added to all samples. For intracellular staining the cells were permeabilised using the Cytofix/CytoPerm Plus™ Kit according to the manufacturers’ instruction and then stained as mentioned. To verify calnexin to be resident in the ER cells were first labelled with the ER-Tracker and then stained with α-calnexin. To analyse co-localisation of lectin binding sites and calnexin cells were simultaneously stained with the appropriate lectin and α- calnexin. The staining with 2 µg FITC labelled goat IgG antibody served as control.

5.12 Detection of exposure of immunoglobulin µ chains on Ag8.H transfectants

For surface staining of the Ag8.H transfectants 1x105 cells were incubated for

30 min at 4°C in PBS supplemented with 2% FCS, 0.1% NaN3 and 0.5 µg of an antibody directed against the heavy chain of IgM (α-µHC). The cells were then

5.13 Detection of GM1 exposure on neutrophil granulocytes

washed with PBS and fixed with 500 µl PBS containing 1% (w/v) PFA and 0.1% PI. The latter was added to exclude necrotic cells. For cytoplasmic staining, 1x105 cells were first fixed in 4% PFA dissolved in PBS at RT for 10 min, permeabilised with 0.1% Tween 20 at 37°C for 15 min and then stained at RT for 15 min with α-µHC.

5.13 Detection of GM1 exposure on neutrophil granulocytes

The subunit B of cholera toxin (CT-B) specifically binds the ganglioside GM1 (Van Heyningen et al., 1976). To analyse GM1 exposure on neutrophil and eosinophil granulocytes 1x105 PMN were incubated for 30 min at room temperature (RT) with 0.2 µg FITC-labelled CT-B and 2 µl PE-conjugated antibodies against either CD16 or CD49d. After staining, cells were fixed with PBS/PFA (1%). Since CT-B was observed to slightly penterate the cell during incubation at RT, CT-B staining in the following experiments was performed at 4°C. To follow GM1 exposure on neutrophil granulocytes undergoing apoptosis the ageing PMN were incubated for 30 min at 4°C with 0.5 µg CT-B and then fixed with 500 µl PBS containing 1% (w/v) PFA and 0.1% PI. The latter was added to exclude necrotic cells. To analyse co-localisation of ER and GM1 PMN were first labelled with the ER-Tracker followed by CT-B staining. Intracellular distribution of GM1 was detected using the Cytofix/CytoPerm Plus™ Kit to permeabilise the cells.

5.14 Inhibition of apoptotic membrane blebbing

The pyridine derivative, Y-27632, was identified to efficiently inhibit the catalytic activity of ROCK I (Uehata et al., 1997) The latter was described to play a role in the bleb formation of apoptotic cells (Coleman et al., 2001; Sebbagh et al., 2001). Therefore, the ROCK inhibitor Y-27632 was used to abrogate apoptotic cell blebbing of PMN undergoing spontaneous apoptosis. PMN were in vitro cultured in R10 containing 50 µM of the ROCK inhibitor for up to 24 hours.

5.15 Live-cell imaging of KDEL receptor GFP transgenic HeLa cells undergoing apoptosis

5.15 Live-cell imaging of KDEL receptor-GFP transgenic HeLa cells undergoing apoptosis

Plasma membranes of KDEL receptor transgenic HeLa cells were labelled using the Vybrant™ DiI cell labelling solution. Adherent transgenic HeLa cells in a Lab-Tek 8 chambered coverglass were incubated with serum free medium (RPMI 1640 supplemented with 1% glutamine, 1% HEPES, and 1% penicillin- streptomycin) containing 5 µM DiI. After 45 min at 37°C the cells were repeated washed and resuspend in a special culture medium (D-MEM:F-12 supplemented with 10% heat inactivated FCS and 1% penicillin-streptomycin) that is recommended for life microscopy.

Image aquisition of apoptosing KDEL receptor-GFP-transgenic HeLa cells was done with a Zeiss AxioCam Mrm digital camera attached to a Zeiss Axiovert 200M inverted microscope. The microscope was equipped with a Zeiss ApoTome and an incubator Zeiss XL-3 for life microscopy. Image acquisition was done under 5% CO2 at 37°C in a humidified atmosphere for 16 hours. Images of full sensor resolution (1.3 MP) were captured every 5 minutes until the end of experiment using a 40x objective (Zeiss Plan-Neofluar 40x/0,75) and the following filter sets for fluorescence (all from Zeiss): red: Filter set 20 (exc.: BP 546/12; emm. 575-640); green: Filter set 38 (exc.: BP 470/40; emm. 525/50).

5.16 Phagocytosis assays

Peritoneal macrophages of mice were obtained 4 days after i.p. injection of 1.0 ml amylum (2% in PBS) by peritoneal lavage. In brief, after carefully removing the skin ice cold culture medium (R10) was injected into the peritoneal cavity (~5 - 7 ml per mouse). The abdomen was gently pressed to bring the

5.16 Phagocytosis assays

cells into suspension and then the culture medium was aspirate with a small needle. The cells were washed and erythrocytes were removed by incubating the cells in erylysis buffer (0.15 M NH4Cl, 20mM HEPES in aqua) for 10 min at RT. The macrophages (2.5 x 105 cells in 500 µl R10) then were seeded into 5 ml falcon tubes and in vitro cultured. After 24 hours nonadherent cells were removed by washing two times with R10.

In the phagocytosis experiments viable PBL (negative control), apoptotic PBL (positive control), and viable PBL displaying an apoptotic surface glycosylation served as prey, respectively. 100 µl of the prey (adjusted to a concentration of 1 Mio/ml in R10) were added to 5 ml falcon tubes containing either peritoneal macrophages in 500 µl R10 or 500 µl R10 alone. Prey in the absence or presence of macrophages was incubated for 3 h at 37°C and then analysed by flow cytometry. The clearance index was calculated as follows: (cell count of prey in the tubes containing no macrophages – cell count of non phagocytosed prey in the tubes containing macrophages) / cell count of prey in the tubes containing no macrophages x 100.

5.17 Statistical analysis

All results presented in this work are representative of at least two independent experiments and all samples were analysed at least in triplicates. Statistical analysis was performed using the two-tailed Student’s t-test for unpaired samples. P values less than 0.05 were considered statistically significant.

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7 Figure Index

Figure 1: Schedule of apoptotic and necrotic cell death. 14 Figure 2: Phagocyte receptors, adaptor molecules and apoptotic “eat-me” signals that are involved in the apoptotic clearance process. 19 Figure 3: Detection of morphological changes of dying Jurkat, PBL, and PMN by their FSc / SSc properties during flow cytometry analysis. 24 Figure 4: Analysis of the cellular morphology of human cell lines (Raji, Jurkat, U937) after staining with certain lectins by flow cytometry. 26 Figure 5: Analysis of NPn and GSL II binding to viable and dying Jurkat, PBL, and PMN by flow cytometry. 30 Figure 6: Comparison of GSL II, NPn, and UEA I binding to viable and full size, early and shrunken, late apoptotic Jurkat, PBL, and PMN, respectively. 31 Figure 7: Comparison of GSL II, NPn, and UEA I binding to shrunken, late apoptotic and primary necrotic Jurkat, PBL, and PMN, respectively. 32 Figure 8: Analysis of GSL II, NPn, and UEA I binding to apoptotic and primary necrotic Jurkat by confocal microscopy. 33 Figure 9: Time course of NPn and GSL II binding in the presence of PI to apoptosing Jurkat by flow cytometry. 34 Figure 10: Time course of apoptosis in Jurkat. 36 Figure 11: Analysis of MAL I and SNA binding to ageing PMN by flow cytometry. 38 Figure 12: Comparison of MAL I and SNA binding to viable, full size apoptotic and shrunken apoptotic Jurkat, PBL, and PMN, respectively. 39 Figure 13: Analysis of MAL I and SNA binding to apoptosing Jurkat in the presence of sialidase inhibitor. 40 Figure 14: Analysis of lectin binding to Jurkat cells treated with glycoprotein processing inhibitors by flow cytometry. 41 Figure 15: Fluorescence microscopic examination of ageing PMN stained with ER-Tracker and Golgi Stain. 43 Figure 16: Analysis of calnexin exposure on ageing PMN by flow cytometry. 44 Figure 17: Analysis of calnexin exposure on viable, apoptotic, and necrotic Jurkat. 45 Figure 18: Microscopical analysis of calnexin exposure on PMN and Jurkat. 46 Figure 19: Time course of apoptosis for ageing PMN. 48 Figure 20: Analysis of expression of µHC in Ag8.Hα transfectants by flow cytometry. 49

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7 Figure Index

Figure 21: Analysis of µHC exposure in Ag8.Hα transfectants undergoing apoptosis. 50 Figure 22: Analysis of CT-B binding to neutrophil and eosinophil granulocytes by flow cytometry. 52 Figure 23: Fluorescence microscopic examination of CT-B binding to freshly isolated PMN. 52 Figure 24: Time course of CT-B binding to ageing PMN ana ly se d by f low cytometry. 54 Figure 25: Fluorescence microscopic examination of PMN stained w it h E R -T r ac ker and CT-B. 55 Figure 26: Analysis of NPn, anti-calnexin, and CT-B binding to apoptosing PMN treated with the ROCK inhibitor Y-27632 by flow cytometry. 57 Figure 27: Monitoring of apoptosing KDEL receptor-GFP-transgenic HeLa cells by fluorescence microscopy. 58 Figure 28: In vitro clearance assays with human PBL. 60 Figure 29: Carbohydrate chains of immature and mature N-linked glycoproteins. 69 Figure 30: Hypothesis how late apoptotic cells maintain ion selectivity of their plasma membrane and initiate their clearance by exposure of ER derived late “eat-me” signals. 76

Table 1: List of all lectins used in this work and their toxicity to Raji, U937, Jurkat, PBL, and PMN 28 Table 2: Binding of non lytic lectins shown in Figu re 2 to v ia bl e an d dy in g c e lls . 29 Table 3: Expression of carbohydrates on the surfaces of cells undergoing apoptosis. 66 Table 4: Reagents 77 Table 5: Auxiliary Materials 81 Table 6: Instruments 82

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8 Abbreviation Index

αvβ3 integrin the vitronectin receptor

β2-GPI β2-glycoprotein-I ∆Ψm mitochondrial membrane potential µHC µ heavy chains AAL Aleuria aurantia lectin Apaf-1 apoptosis protease activating factor 1 ATP adenosine 5’-tripohosphate Ax I annexin I AxV annexin V Con A Concanavalin A CRP C-reactive protein CT-B cholera toxin subunit B DBA Dolichos biflorus agglutinin DC dendritic cell DiO6C(3) 3.3'- dihexyloxacarbocyanine iodide DSL Datura stramonium lectin ECL Erythrina cristagalli lectin ER endoplasmic reticulum ERM ezrin/radixin/moesin Fc R Fc receptor FDC follicular dendritic cell FITC fluorescein isothiocyanate FL fluorescence FSc forward scatter Fuc fucose Gal galactose

107

8 Abbreviation Index

GalNAc N-Acetylgalactosamine gas6 growth-arrest specific gene 6 GC germinal centre GlcNAc N-acetylglucosamine GSL I Griffonia simplicifolia lectin I GSL II Griffonia simplicifolia lectin II HBSS Hank`s buffered Salt Solution ICAM-3 intracellular adhesion molecule 3 IDX clearance index J Jacalin LCA Lens culinaris agglutinin LEA, LEL Lycopersicon esculentum lectin LFA Limax flavus agglutinin LOX-1 lectin-like OxLDL receptor 1 LPA Limulus polyphemus agglutinin LPC lysophosphatidylcholine LPS lipopolysaccharide MAA, MAL Maackia amurensis lectin MAL I Maackia amurensis lectin I Man mannose MBL mannose binding lectin MFG-E8 milk-fat-globue-EGF-factor 8 MFI mean fluorescence intensity NPn Narcissus pseudonarcissus lectin OxLDL oxidised low-density lipoprotein PBL peripheral blood lymphocytes PBMC peripheral blood mononuclear cells PFA paraformaldehyde PHE Phaseolus vulgaris erythroagglutinin PHL Phaseolus vulgaris leucoagglutinin PI propidium iodide

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8 Abbreviation Index

PLA Phaseolus limensis agglutinin PMN polymorphonuclear cells PNA Peanut agglutinin PS phosphatidylserine PSA Pisum sativum agglutinin RCA I Ricinus communis agglutinin I ROCK Rho-associated protein kinase SA, SBA Soybean agglutinin SAP serum amyloid P SJA Sophora japonica agglutinin SLE systemic lupus erythematosus SNA Sambucus nigra agglutinin SP-A/-D surfactant proteine A and D SR-BI class B scavenger receptor type I SSc side scatter STL Solanum tuberosum lectin STS staurosporine TGP Lotus Tetragonolobus lectin TNF tumor necrosis factor TSP thrombospondin UEA I Ulex europaeus lectin I UV-B Ultra violet light -B VAA Viscum album agglutinin VVL Vicia villosa lectin WGA Wheat germ agglutinin

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Publications and Presentations

Publications

1 Sarter K., Mierke C., Beer A., Frey B., Führnrohr B., Schulze C., Franz S. (2007) Sweet Clearance: Involvement of Cell Surface Glycans in the Recognition of Apoptotic Cells. Autoimmunity in press (Review)

2 Schiller M., Blank N., Franz S., Gaipl U.S., Ho A.D., Lorenz H.M. (2007) Apoptotic bodies derived from apoptotic lymphoblasts contain a distinct pattern of antigens. Autoimmunity in press

3 Munoz, L.E.*, Franz, S.*, Pausch, F., Furnrohr, B., Sheriff, A., Vogt, B., Kern, P.M., Baum, W., Stach, C., von Laer, D., Brachvogel, B., Poschl, E., Herrmann, M., Gaipl, U.S. (2007) The influence on the immunomodulatory effects of dying and dead cells of Annexin V. J Leukoc Biol 81, 6-14.

4 Franz, S., Herrmann, K., Fuhrnrohr, B., Sheriff, A., Frey, B., Gaipl, U.S., Voll, R.E., Kalden, J.R., Jack, H.M., Herrmann, M. (2007) After shrinkage apoptotic cells expose internal membrane-derived epitopes on their plasma membranes. Cell Death Differ 14, 733-742

5 Franz, S., Gaipl, U.S., Munoz, L.E., Sheriff, A., Beer, A., Kalden, J.R., Herrmann, M. (2006) Apoptosis and autoimmunity: when apoptotic cells break their silence. Curr Rheumatol Rep 8, 245-7. (Review)

6 Gaipl, U.S., Kuhn, A., Sheriff, A., Munoz, L.E., Franz, S., Voll, R.E., Kalden, J.R., Herrmann, M. (2006) Clearance of apoptotic cells in human SLE. Curr Dir Autoimmun 9, 173-87. (Review)

7 Franz, S., Frey, B., Sheriff, A., Gaipl, U.S., Beer, A., Voll, R.E., Kalden, J.R., Herrmann, M. (2006) Lectins detect changes of the glycosylation status of plasma membrane constituents during late apoptosis. Cytometry A 69, 230-9.

8 Rodel, F.*, Franz, S.*, Sheriff, A., Gaipl, U., Heyder, P., Hildebrandt, G., Schultze-Mosgau, S., Voll, R.E., Herrmann, M. (2005) The CFSE distribution assay is a powerful technique for the analysis of radiation- induced cell death and survival on a single-cell level. Strahlenther Onkol 181, 456-62.

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9 Gaipl, U.S., Voll, R.E., Sheriff, A., Franz, S., Kalden, J.R., Herrmann, M. (2005) Impaired clearance of dying cells in systemic lupus erythematosus. Autoimmun Rev 4, 189-94. (Review)

10 Grossmayer, G.E., Munoz, L.E., Gaipl, U.S., Franz, S., Sheriff, A., Voll, R.E., Kalden, J.R., Herrmann, M. (2005) Removal of dying cells and systemic lupus erythematosus. Mod Rheumatol 15, 383-90.

11 Munoz, L.E., Gaipl, U.S., Franz, S., Sheriff, A., Voll, R.E., Kalden, J.R., Herrmann, M. (2005) SLE--a disease of clearance deficiency? Rheumatology (Oxford) 44, 1101-7. (Review)

12 Gaipl, U.S., Franz, S., Voll, R.E., Sheriff, A., Kalden, J.R., Herrmann, M. (2004) Defects in the disposal of dying cells lead to autoimmunity. Curr Rheumatol Rep 6, 401-7. (Review)

13 Korn, A., Frey, B., Sheriff, A., Gaipl, U.S., Franz, S., Meyer-Pittroff, R., Bluemelhuberh, G., Herrmann, M. (2004) High hydrostatic pressure inactivated human tumour cells preserve their immunogenicity. Cell Mol Biol (Noisy-le-grand) 50, 469-77.

14 Frey, B., Franz, S., Sheriff, A., Korn, A., Bluemelhuber, G., Gaipl, U.S., Voll, R.E., Meyer-Pittroff, R., Herrmann, M. (2004) Hydrostatic pressure induced death of mammalian cells engages pathways related to apoptosis or necrosis. Cell Mol Biol (Noisy-le-grand) 50, 459-67.

15 Sheriff, A., Gaipl, U.S., Franz, S., Heyder, P., Voll, R.E., Kalden, J.R., Herrmann, M. (2004) Loss of GM1 surface expression precedes annexin V-phycoerythrin binding of neutrophils undergoing spontaneous apoptosis during in vitro aging. Cytometry A 62, 75-80. * equally contributed

Short Publications

1 Franz, S., Frey, B., Herrmann, K., Beer, A., Gaipl, U.S., Sheriff, A., Voll, R.E., Kalden, J.R., Herrmann, M. (2005) Late apoptotic cells expose incompletely processed proteins on their surfaces. Immunobiology 210(6 – 8), 558.

2 Frey, B., Franz, S., Korn, A., Meyer-Pittroff, R., Gaipl, U.S., Meister, S., Voll, R.E., Bluemelhuber, G., Herrmann, M. (2005) High hydrostatic pressure treated tumour cells. Cell death pathways and immunogenicity of treated cells. Immunobiology 210(6 – 8), 478.

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3 Gaipl, U.S., Franz, S., Appelt, U., Voll, R.E., Sheriff, A., Kalden, J.R., Herrmann, M. (2005) Intrinsic and heterogeneous clearance defects in some patients with autoimmune disease systemic lupus erythematosus (SLE). Immunobiology 209(S1), 8.

4 Franz, S., Gaipl, U.S., Sheriff, A., Voll, R.E., Kalden, J.R., Herrmann, M. (2004) The role of carbohydrates in the clearance process of apoptotic cells. Immunobiology 209(4 - 6), 449.

5 Franz, S., Appelt, U., Sheriff, A., Gaipl, U.S., Voll, R.E., Kalden, J.R., Herrmann, M. (2004) Annexin V binding to dying cells is highly cooperative. Immunobiology 209(4 - 6), 448.

6 Franz, S., Gaipl, U.S., Appelt, U., Heyder, P., Voll, R.E., Kalden, J.R., Herrmann, M. (2004) The role of a defective clearance in the phatogenesis of systemic lupus erythematosus. Arthritis Research & Therapy 6(S1), 34

Book Chapters

1 Gaipl, U.S., Sheriff, A., Franz, S., Munoz, L.E., Voll, R.E., Kalden, J.R., Herrmann, M. (2006) Inefficient clearance of dying cells an autoreactivity. In Current conncepts in autoimmunity and chronic inflammation, Volume 305 (A. Radbruch and P. E. Lipsky, eds), Springer, Berlin Heidelberg New York 161 - 176.

2 Dransfield, I., Franz, S., Wilkinson, K., McColl, A., Herrmann, M., Hart, SP. (2006) Cell surface molecular changes associated with apoptosis In Progress in inflammation research, (M.J. Parnham, ed.), Volume Resolution of inflammation (A.G. Rossi and D.A. Sawatsky, eds), Birkhäuser-Verlag AG, Basel Boston Berlin

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Talks

1 Franz, S. Fingerprinting Apoptotic Cell Surfaces: Alterations of Glycocalyx and Membrane Composition. Montagsseminar der Medizinischen Klinik 3, 12-2006, Erlangen

2 Franz, S. Wunderwerk Immunsystem – Wie sich der Körper gegen Krankheitserreger schützt. Naturwissenschaftliche Tage am Dietrich-Bonhoeffer-Gymnasium, 02- 2006, Oberasbach

3 Franz, S. ”Eat me" signals of late apoptotic cells are derived from internal membranes. 6. Berichtssymposium des Graduiertenkollegs 592 "Lymphozyten: Differenzierung, Aktivierung und Deviation", 01-2006, Heiligenstadt

4 Franz, S., Frey, B., Herrmann, K., Gaipl, U.S., Sheriff, A., Herrmann, M. Late apoptotic lymphocytes expose incompletely processed proteins on their surfaces. 36th Annual Meeting of the DGFI German Society of Immunology,09- 2005, Kiel

5 Franz, S., Herrmann, M. Late apoptotic cells expose incompletely processed proteins on their surfaces. Rheumatology Day, 07-2005, Erlangen

6 Franz, S. Late apoptotic lymphocytes expose incompletely processed proteins on their surfaces. 5th Retreat of the GK520 (Würzburg) & GK592 (Erlangen), 07-2005, Markt Taschendorf

7 Franz, S. Do apoptotic cells expose premature proteins on their surfaces during the late phase of apoptosis? Freitagsseminar der Medizinischen Klinik 3, 01-2005, Erlangen

8 Franz, S. Do apoptotic lymphocytes expose premature proteins on their surfaces during the late phase of apoptosis? 5. Berichtssymposium des Graduiertenkollegs 592 "Lymphozyten: Differenzierung, Aktivierung und Deviation", 11-2004, Heiligenstadt

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9 Franz, S. Surface changes of dying lymphocytes 4th Retreat of the GK520 (Würzburg) & GK592 (Erlangen), 07-2004, Markt Taschendorf

10 Franz, S., Gaipl, U. S., Appelt, U., Heyder, P., Voll, R.E., Herrmann, M. Heterogeneous defects of phagocytosis and clearance in certain SLE patients. European Workshop for Rheumatology Research, 02-2004, Berlin

11 Franz, S., Gaipl, U. S., Voll, R.E., Herrmann, M. Surface changes during apoptotic and necrotic cell death of lymphocytes. Freitagsseminar der Medizinischen Klinik 3, 12-2003, Erlangen

12 Franz, S. Oberflächenveränderungen auf Lymphozyten im Zuge des apoptotischen und nekrotischen Zelltods. 4. Berichtssymposium des Graduiertenkollegs 592 "Lymphozyten: Differenzierung, Aktivierung und Deviation", 11-2003, Heiligenstadt

13 Franz, S., Gaipl, U. S., Heyder, P., Herrmann, M. Membrane changes of dying cells; e.g. alterations in the binding of lectins. Freitagsseminar der Medizinischen Klinik 3, 07-2003, Erlangen

Post ers

1 Franz, S., Frey, B., Herrmann, K., Beer, A., Gaipl, U.S., Sheriff, A., Voll, R.E., Kalden, J.R., Herrmann, M. After shrinkage apoptotic cells expose internal membrane-derived epitopes on their plasma membranes. 1st International GK-Symposium: Regulators of Adaptor Immunity, 09- 2006, Erlangen

2 Franz, S., Frey, B., Herrmann, K., Beer, A., Gaipl, U.S., Sheriff, A., Voll, R.E., Kalden, J.R., Herrmann, M. Late apoptotic cells expose incompletely processed proteins on their surfaces. 36th Annual Meeting of the DGFI German Society of Immunology,09- 2005, Kiel

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3 Franz, S., Appelt, U., Sheriff, A., Gaipl, U.S., Voll, R.E., Kalden, J.R., Herrmann, M. Annexin V binding to dying cells is highly cooperative. 2nd Tutzing Antiphospholipid Conference, 04-2005, Tutzingen

4 Franz, S., Gaipl, U.S., Sheriff, A., Voll, R.E., Kalden, J.R., Herrmann, M. The role of carbohydrates in the clearance process of apoptotic cells. Joint Annual Meeting of the German and Dutch Societies for Immunology, 10-2004, Maastricht

5 Franz, S., Gaipl, U.S., Appelt, U., Heyder, P., Voll, R.E., Kalden, J.R., Herrmann, M. The role of a defective clearance in the phatogenesis of systemic lupus erythematosus. European Workshop for Rheumatology Research, 02-2004, Berlin

6 Franz, S., Gaipl U.S., Heyder P, Beyer T.D., Voll R.E., Kalden J.R., Herrmann M. Membrane changes of dying cells, e.g. alterations in the binding of lectins 3.Erlanger IZKF Workshop, 07-2003, Erlangen

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Lebenslauf Sandra Franz

Persönliche Daten

Anschrift Werner-von-Siemens-Straße 7 91052 Erlangen Telefon 09131-205972 E-Mail [email protected] Geburtstag 20.04.1976 Geburtsort Chemnitz Staatsangehörigkeit deutsch Familienstand ledig

Wissenschaftliche Ausbildung

05/2003 – 04/2007 Doktorarbeit am Institut für Klinische Immunologie und Rheumatologie, Friedrich-Alexander-Universität Erlangen-Nürnberg (FAU) „Fingerprinting Apoptotic Cell Surfaces: Alterations of Glycocalyx and Membrane Composition“ 08 – 09/2005 Trainee Aufenthalt MRC Centre for Inflammation Research, University of Edinburgh, Prof. I. Dransfield

Ausbildung

11/1995 – 12/2002 Studium des Lehramts am Gymnasium für Biologie und Chemie, FAU Studienabschluss: erste Staatsprüfung (Note: 1,48) 06/1992 – 06/1995 Gymnasium in Nürnberg Abschluss: Allgemeine Hochschulreife (Note: 2,5) 09/1990 – 05/1992 Gymnasium in Chemnitz 09/1982 – 08/1992 Allgemeine polytechnische Schule in Chemnitz

Förderungen und Preise

07/2006 Posterpreis, Interdisziplinäres Zentrums für Klinische Forschung, Erlangen 05/2003 – 04/2006 Promotionsstipendium, Graduiertenkolleg 592 „Lymphozyten“ der DFG

Erlangen, 05.02.2007

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