Function and Regulation of Podosomes, Important Adhesion Structures in Dendritic Cells

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Function and regulation of podosomes, important adhesion structures in dendritic cells

Van Helden, Suzanne Thesis Radboud University Nijmegen Medical Centre, The Netherlands

Cover illustration: immature DCs stained for actin and vinculin Cover design: Suzanne van Helden Layout: Stan van de Graaf

Printed by: F&N bookservice Function and regulation of podosomes, important adhesion structures in dendritic cells

Een wetenschappelijke proeve op het gebied van de Medische Wetenschappen

Proefschrift

ter verkrijging van de graad van doctor aan de Radboud Universiteit Nijmegen op gezag van de rector magnificus, prof. mr. S.C.J.J. Kortmann, volgens besluit van het College van Decanen in het openbaar te verdedigen op dinsdag 25 november 2008 om 15.30 uur precies

door

Suzanne Francina Gertruda van Helden

geboren op 6 juli 1979 te Horst Promotor: prof. dr. C.G. Figdor

Copromotor: dr. F.N. van Leeuwen

Manuscriptcommissie: prof. dr. B. Wieringa prof. dr. R. Brock dr. P.L. Hordijk, Sanquin/AMC Amsterdam

The research was performed at the department of Tumor Immonology of the Nijmegen Centre for Molecular Life Sciences, Radboud University Nijmegen Medical Centre, Nijmegen, The Netherlands. Contents

Chapter 1 General introduction 7

Chapter 2 Human and murine model cell lines for dendritic 25 cell biology evaluated

Chapter 3 A critical role for prostaglandin E2 in podosome 41 dissolution and induction of high-speed migration during dendritic cell maturation

Chapter 4 TLR4-mediated discrimination between Gram- 63 negative and Gram-positive bacteria directs migration and maturation of dendritic cells

Chapter 5 PGE2-mediated podosome loss in dendritic cells 81 is dependent on actomyosin contraction downstream of the Rho-Rho kinase axis

Chapter 6 Adhesion of dendritic cells to micro-patterned 105 substrates reveals physical aspects of podosome regulation

Chapter 7 General discussion and summary 121

Nederlandse samenvatting 135

List of abbreviations 141

Dankwoord 143

Curriculum Vitae 147

List of publications 149

Chapter 1 General introduction Chapter 1

The immune system Our body is constantly challenged by pathogens, such as bacteria, viruses and fungi. Therefore, it has developed several defense systems, such as the skin and mucosal surfaces that form a difficult to cross barrier for pathogens. If pathogens succeed in entering the body they are detected and attacked by the immune system. In addition to protecting against infections, the immune system is also important for proper clearance of dead cells. Moreover, the immune system can detect and eradicate cells with aberrant properties to prevent cancer. Consequently, a disregulated immune system can lead to life threatening infections or the formation of cancer, but also autoimmunity and allergy. Under these pathological conditions, modulation of the immune response can be beneficial. For instance, immune responses can be dampened in allergy, autoimmunity or transplantation settings, while immune responses can be enhanced to combat infections or cancer.

Figure 1. Cells of the innate and adaptive immune system The innate immune system consists of mast cells, NK cells, granulocytes and antigen presenting cells. Antigen presenting cells form the link between innate and adaptive immunity. Cells of the adaptive immune system are T cells and B cells. NK cell; natural killer cell, neutr; neutrophil, eosin; eosinophil, baso; basophil, Mφ ; macrophage, mono; monocyte, myDC; myeloid dendritic cell, pDC; plasmacytoid dendritic cell, act T cell; activated T cell and *; phagocytes.

8 General introduction

In vertebrates, the immune system consists of two branches with different characteristics, known as innate and adaptive immunity (Figure 1). Innate immunity is the first line of defense and regulated by natural killer cells, mast cells, granulocytes and phagocytes [1-5]. If this first line of defense is not sufficient, the adaptive immune system comes into action. Antigen presenting cells (APCs), which include monocytes, macrophages and dendritic cells (DCs), form the link between the innate and adaptive immune system. APCs express specific cell surface receptors that recognize molecules common to pathogens. These so-called pathogen recognition receptors (PRRs) include Toll-like receptors (TLRs), lectins and nucleotide-binding oligomerization domain containing (NOD)-like receptors [6-9]. Activation of PRRs leads to internalization of pathogens and signaling events resulting in activation of nuclear factor κB (NF-κB), which controls the expression of an array of inflammatory cytokines. Upon internalization, pathogens are degraded which leads to the generation of peptides that are presented on the surface of APCs in the context of major histocompatibility complex (MHC) molecules [10]. Peptides generated from proteins derived from internalized material are presented in the context of MHC class II molecules, which are predominantly expressed on APCs [11]. Peptides presented in MHC class II molecules are recognized by T cells expressing CD4, called helper T cells. These helper T cells activate cells of the innate immune system and stimulate antibody production by B cells [12]. However, in APCs some exogenous antigens derived via endocytic uptake are presented on MHC class I molecules, instead of MHC class II molecules. This process is known as cross-presentation [13]. Peptides derived from cytoplasmic proteins, either endogenous proteins or intracellular pathogens, are presented in the context of MHC class I molecules by all cells [14]. Peptides presented in MHC class I molecules are recognized by T cells expressing CD8, cytotoxic T cells [15]. APCs as well as their target (B and T) cells express a set of auxiliary receptor-ligand pairs, known as costimulatory molecules which are needed for proper activation of the adaptive immune response. Only simultaneous activation of costimulatory molecules and specific receptors for the peptide-MHC complexes on T cells and B cells leads to sufficient activation of these lymphocytes and consequently the induction of adaptive immune responses. Dendritic cells As mentioned earlier, all antigen presenting cells use PRRs to recognize pathogens and respond by the production of cytokines. However, of all APCs, DCs are most potent in modulating T cell

9 Chapter 1

responses and thus play a central role in linking innate and adaptive immunity [16-19]. As the most effective antigen presenting cells of the body, DCs are being explored in the clinic for boosting immunological responses against various cancers including melanoma, myeloma and prostate cancer [20-23]. In these studies, monocytes or DC-precursors are isolated from cancer patients and differentiated to DCs in vitro. Upon loading with tumor-specific antigens or a tumor lysate the DCs are given back to the patient to trigger specific T cell responses against the tumor cells [24, 25]. Although DCs are used in clinical trials world-wide, fundamental knowledge about the complex

Figure 2. Different stages in the life of DCs

DC precursors (preDC) move from the blood into the tissues. In the tissues, immature DCs (iDC) scan for antigens. Upon encountering pathogens or other antigens under inflammatory situations the iDCs undergo maturation and migrate through the lymph vessels to the lymph nodes. In the lymph nodes, mature DCs (mDC) stimulate B cells and T cells, leading to the initiation of an adaptive immune response. After fulfilling their

function the DCs undergo programmed cell death (apoptosis) (DCapop).

10 General introduction biology of these cells is lagging behind. DCs can originate from lymphoid or myeloid precursors giving rise to plasmacytoid DCs (pDCs) and myeloid DCs, respectively. The more recently identified pDCs are interferon (IFN)-producing cells found in blood and they are important mediators in immune responses against viruses [26, 27]. Myeloid DCs are found in the blood and in peripheral tissues and these cells can initiate immune responses against diverse antigens in various situations. To study myeloid DCs, monocyte-derived DCs are often used as a model. For this purpose, monocytes are cultured in the presence of IL-4 and GM-CSF for 6-7 days to obtain DCs [28, 29]. Alternatively DCs can be differentiated from CD34-positive cells (hematopoietic progenitors). Currently, no good DC-model cell lines are available, which hampers cell biological studies [30]. PRRs During their development (myeloid) DC-precursors move from the blood into the peripheral tissues, where these cells reside in an immature state (Figure 2). These immature DCs (iDCs), are equipped with a plethora of PRRs [31, 32], such as TLRs, C-type lectins and NOD-like receptors, that allow them to efficiently recognize pathogen-associated molecules on pathogens (Table 1). TLRs are the vertebrate counterparts of the Drosophila melanogaster receptor Toll [32]. To date, 13 members of this family of transmembrane receptors have been identified and they can operate both as homo- and heterodimers that can recognize a large variety of pathogen components. For instance, TLR4 recognizes lipopolysaccharide (LPS) from Gram-negative bacteria and TLR5 recognizes flagellin [33, 34]. TLR2 can form heterodimers with TLR1 and TLR6 which recognize peptidoglycans, lipoteichoic acid (LTA) and lipoproteins, components of Gram-positive bacteria [35]. TLR2 and TLR4 are also involved in recognizing fungi [36]. On the other hand, TLR3, 7 and 8 can recognize viral components, such as double stranded and single stranded RNA [37, 38]. TLRs are signaling receptors and activation most often leads to NF-κB mediated gene expression. Most TLRs make use of the adaptor protein myeloid differentiation primary response gene 88 (MyD88) [39]. In contrast, TLR3 uses the TIR domain-containing adaptor-inducing IFNβ (TRIF) adaptor, which signals to increase expression of type I IFNs [40]. TLR4 is unique since until now it is the only TLR that can activate the MyD88 as well as the TRIF dependent signaling pathway. In addition to transmembrane PRRs, DCs also express recently identified intracellular receptors for microbes, like NOD-like receptors and RIG-like helicases

11 Chapter 1

PRRs on myeloid and Ligand binding References monocyte-derived DCs

TLRs TLR1 Combines with TLR2 [112] TLR2 Peptidoglycan, lipoproteins, lipoarabino- [36, 113-116] mannan, zymosan TLR3 polyI:C, viral double stranded RNA [37] TLR4 LPS, heat shock protein 60 [33, 117, 118] TLR5 Flagellin [34] TLR6 Combines with TLR2 [6] TLR7 and 8 Imidazoquinolines, viral single [38, 119, 120] stranded RNA TLR10 ?

C-type lectins Mannose receptor Mannose, fucose, sLewisX [121] DEC-205 HIV1 [122, 123] DC-SIGN Complex mannose residues, LewisX, [124-130] viruses, fungi, ICAM-2, -3 DCIR ? [131] Dectin-1 β-glucan [132, 133] Dectin-2 High mannose, fungi [134, 135] CLEC-1 ? [42, 136] CLEC-2 ? [136] Endo-180 Mannose, fucose, N-acetylglycosamine, [137, 138] collagen MGL Terminal N-acetyl galactosamines [139, 140] DLEC ? [141] DCAL-1 CD4 T cells [142] DCAR ? [143] MICL ? [144]

Intracellular PRRs NOD1 and 2 Muropeptides derived from peptidogly- [9, 145-148] cans RIG-I and MDA-5 Viral double stranded RNA [41, 149-151]

Table 1. Pathogen recognition receptors (PRRs) in human DCs.

12 General introduction

[41]. In addition, DCs express several C-type lectins including DC- SIGN, dectin-1 and the mannose receptor [31]. C-type lectins are calcium dependent transmembrane receptors that bind carbohydrate structures on pathogens as well as on endogenous ligands. Binding of pathogens to C-type lectins leads to internalization of the receptor/ligand complex, however recent findings indicate that C-type lectins are also capable of activating intracellular signaling pathways [42, 43]. Pathogens often activate multiple PRRs and these PRRs can affect each others function. Therefore, the resulting response is determined by the specific combination of receptors present on the APC. DC maturation When iDCs are exposed to pathogens, antigens and/or pro-inflammatory signals, they undergo a dramatic phenotypic conversion known as DC maturation (Figure 2). This process transforms iDCs, specialized in antigen recognition and internalization, into antigen presenting mDCs, which are very efficient in stimulating T cells. During maturation, molecules needed for antigen recognition and uptake are downregulated, while molecules required for antigen processing and presentation, such as MHC and costimulatory molecules, are upregulated [44-46]. Moreover, in response to maturating signals, DCs start to secrete cytokines that further induce and activate T cells. In addition to upregulating MHC molecules, DCs are able to regulate the fate of MHC molecules. In iDCs the majority of the MHC molecules is maintained in an intracellular lysosomal compartment known as the MHC class II compartment. Upon maturation, MHC molecules, loaded with peptides derived from internalized antigens, are transported from the MHC class II compartment to the cell membrane [18]. DCs are more potent than other APC in stimulating T cells, in part because they are able to efficiently cross-present antigens [13]. During maturation, tissue resident iDCs have to acquire the ability to travel out of the tissue and into the lymph nodes [47] (Figure 2). Therefore, strongly adherent iDCs transform into highly migratory mDCs. Migration to the lymph nodes is mediated by the chemokines CCL19 and CCL21. In order to respond to these chemokines mDCs have to express the active chemokine receptor CCR7 [47-49]. PGE2, a prostaglandin produced from arachidonic acid by the cyclooxygenase (COX) enzymes [50-52], is important for the induction of migration during maturation [53-57], in part because it induces expression and activation of CCR7 [58, 59]. The development of migration also involves extensive changes in the cytoskeletal organization of the

13 Chapter 1

DCs [60, 61] (this thesis). During maturation DCs lose adhesion structures and develop the characteristic dendritic extensions, after which they are named. These dendritic extensions enhance contact with T cells and B cells once the DCs reach the lymph nodes. Adhesive interactions in the immune system Regulation of adhesive and migratory properties plays a prominent role in immune regulation. The ability of immune cells to enter lymph nodes or tissues relies on sets of adhesion molecules, such as integrins. The latter are composed of an α chain and a β chain. Distinct conformations determine the ability to bind ligand, where a closed, bended conformation is unable to bind ligand and the open, extend- ed conformation can bind ligand [62-65]. In addition, there are intermediate activation states resulting in intermediate affinities [66]. The activity of integrins can be further regulated by receptor clustering.[67-70]. In addition to β2- and β7-integrins, which are selectively expressed in the immune system, β1- and β3-integrins are important for proper functioning of the immune system. Integrins play important roles during various phases of the immune response and defective integrin function lead to severe diseases, such as blistering disorder, leukocyte adhesion deficiencies and bleeding disorders [71]. The αMβ2- and αXβ2-integrins are complement receptors and αMβ2-, αLβ2- and α5β1-integrins are receptors for several pathogens [72- 75]. In addition to cell-pathogen interactions, integrins are important

Figure 3. Adhesion structures, FAs and podosomes, in DCs In the upper panel a DC displaying FAs is depicted. In the lower panel a DC displaying podosomes can be seen. Staining for the cytoskeletal plaque protein vinculin (green) reveals FAs and the rings of the podosomes. Phalloidin-TexasRed staining of F-actin (red) reveals actin stress fibers connected to the FAs as well as the cores of the podosomes.

14 General introduction

Cell type Putative function References Endothelial cells matrix degradation, arterial remodeling [152, 153] Macrophage/monocyte/DC adhesion, migration, diapedesis [95, 101, 154] Osteoclast sealing zone formation, bone resorption [96, 155] Smooth muscle cell matrix degradation, migration [97, 156] Transformed fibroblast adhesion, migration [103]

Table 2. Putative podosome functions in different cells. for cell-cell interactions, such as the rolling of leukocytes on the endothelium and formation of the immunological synapse between DCs and T cells [76, 77]. Furthermore, integrins, especially the β1- integrins, mediate cell-extracellular matrix (ECM) interactions [78- 80]. The interactions with other cells and with the ECM and the regulation thereof are essential for adhesion and migration of leukocytes. Constitutive active integrins result in defective cell migration, showing that the activity of integrins has to be tightly regulated and is important for the initiation of immune responses [81]. In addition to sensing and binding to their surroundings, cells have to control their shape and move to coordinate adhesion and migration. This is controlled by the cytoskeleton, a framework of filaments that coordinates cell division and intracellular traffic, supports the plasma membrane, provides stability and controls cell shape. The cytoskeleton consists of three types of filaments, actin filaments, microtubules and intermediate filaments. For cells to move forward they have to undergo cycles of protrusion, attachment to the substrate, forward translocation of the cell body, release of adhesions and retraction at the cell rear [82]. The forces needed for these processes are generated either by polymerization of actin leading to protrusion of the plasma membrane [83], or by motor proteins, which use the energy derived from repeated cycles of ATP hydrolysis to move along filaments [84]. Nonmuscle myosin II is the actin-binding motor protein that is essential to generate contractility. Light-chain phosphorylation allows the myosin molecules to assemble into short, bipolar, thick filaments, which leads to contraction [85]. The ECM is linked to the actomyosin cytoskeleton through integrins, which can organize into specific "higher order" adhesion sites. Dependent on organization and composition, these adhesion sites are known as focal adhesions (FAs), focal complexes, fibrillar adhesions, invadopodia and podosomes [82, 86, 87]. FAs, focal complexes and fibrillar adhesions are related (integrin containing) adhesion plaques found in many cell types which connect the cell to the surrounding ECM. Focal complexes are smaller nascent adhesion struc-

15 Chapter 1

tures found at the periphery of spreading or migrating cells, which can further mature into FAs [88]. The latter are relatively stable structures connected to stress fibers, bundles of actin filaments crosslinked by α-actinin (Figure 3). FAs can further mature into fibrillar adhesions, which are mostly located to the center of the cell [82, 88, 89]. Podosomes In contrast to these adhesion plaques, podosomes and invadopodia have a very different morphology. Podosomes have a very characteristic organization consisting of a core of actin surrounded by a ring containing integrins and structural proteins, such as vinculin [90, 91] (Figure 3). Podosomes are very dynamic structures that can merge, split and display high actin turnover in the core [92]. The appearance of podosomes and of the related invadopodia is restricted to particular cell types (Table 2). Invadopodia are a feature of invasive (tumor) cells [90] whereas podosomes can be found in endothelial cells, Src- transformed fibroblasts and cells of the myeloid lineage, such as monocytes, macrophages, osteoclasts and DCs [93]. In contrast to the adhesion plaques, podosomes are active sites of matrix degradation which allow the cells to invade into tissues or cross tissue boundaries [94, 95]. In addition, podosomes play an important role during osteoclast-mediated bone resorption [96, 97]. Podosomes are regulated by the small Rho GTPases, Rho, Rac and Cdc42, and regulators thereof [94, 98-100]. Actin regulatory pathways affect the formation and turnover of podosomes, which is not surprising since the actin core is a prominent part of podosomes. The absence of proteins regulating actin turnover, such as Wiskott-Aldrich Syndrome protein (WASP) or the actin nucleating Arp2/3 complex, results in podosome disruption [99, 101]. In addition, podosomes are regulated by tyrosine phosphorylation, which is underscored by the finding that cells transformed with the tyrosine kinase Src can form podosomes [102, 103]. In addition to the actin cytoskeleton, podosomes are regulated by microtubules [104, 105] and their interactions with the cell substrate [106]. However, the precise mechanisms behind this regulation remain largely unknown. Adhesion and migration in DCs Although iDCs also display FAs, podosomes are the most prominent adhesions found in these cells [99] (this thesis). In contrast, migratory mDCs completely lack podosomes and FAs but instead display numerous dendritic extensions, which serve to enhance the cells surface and hence facilitate the interactions with T cells [107]. These

16 General introduction findings indicate that both the DC cytoskeleton and the organization of DC adhesion change dramatically during the process of DC maturation. Aim of this thesis DCs link innate and adaptive immunity and are key players in the induction of efficient immune responses. Functional maturation of DCs is critical for the induction of efficient immune responses [108]. For instance, the induction of migration during DC maturation was shown to be essential for the induction of T cell responses both in mice and in human cancer patients receiving DC-vaccination [109- 111]. However, the cytoskeletal changes that occur during DC maturation and its consequences for cell adhesion or the induction of migration remain poorly understood. More insight into these processes will help improve our understanding of the function of DCs in inflammation, infection and autoimmunity and may improve DC-vaccination strategies in cancer patients. Therefore, the aim of this thesis is to investigate the cytoskeletal changes that occur in human monocyte-derived DCs in response to maturation signals, and study its consequences for adhesive and migratory behavior. In chapter 2 we compare different APC cell lines as models to study DC-biology. DCs are post-mitotic cells that have to be isolated and differentiated from blood, spleen or buffy-coat material. The production of DCs involves the use of expensive cytokines and is rather time consuming. Moreover, donor-to-donor variations often can give rise to variable results. Hence, a good model cell line would circumvent some of these problems. Although no cell line can fulfill all require- ments for a DC-model, we describe a number of cell lines that can be used to investigate different aspects of DC-biology. In chapter 3 the loss of cell adhesion structures and the acquisition of migration during DC-maturation are investigated in detail. Podosome dissolution in response to maturation stimuli is mediated by prostaglandins produced by the DCs, and the prostaglandin PGE2 induces very rapid podosome loss. Podosome loss as well as inactivation of α5β1-integrin are needed to acquire migration. In chapter 4 the effects of different classes of bacteria on adhesive and migratory properties and other aspects of DC-maturation are investigated. Gram-negative bacteria are superior to Gram-positive bacteria in inducing podosome loss, migration and inducing expression of costimulatory molecules, CCR7 and cytokines. This is mediated by activation of TLR4 and predominantly TRIF-dependent. Moreover, these effects involve the production of prostaglandins

(possibly PGE2) by DCs.

17 Chapter 1

In chapter 5, we investigate the signal transduction pathways

involved in PGE2 mediated podosome dissolution. PGE2 activates the EP2 and EP4 receptors on DCs, which raises cAMP levels. This results in activation of the small GTPase RhoA and inactivation of Rac1 and Cdc42. Active RhoA activates its downstream effector Rho kinase. Activation of the RhoA-Rho kinase axis leads to actomyosin contraction, which is necessary for podosome loss and leads to the (transient) formation of FAs. A model linking actomyosin contractility to podosome loss is put forward. In chapter 6, we explore the regulation of podosome formation on different kinds and shapes of substrates. DCs can adhere to and form adhesion structures on a broad range of substrates. FA-formation, but not DC-adhesion or podosome-formation, is affected by the hydrophobicity of the substrate. Interestingly, when DCs are seeded on substrates with varying topology, podosomes specifically form in those areas where height differences occur. These changes in substrate topology most probably induce a curvature of the cell membrane triggering local podosome formation. We postulate that podosomes are mechanosensitive structures regulated by mechanical stress on the outside (chapter 6) and cytoskeletal tension on the inside of the cell (chapter 5). References 1. Erb, K.J., J.W. Holloway, and G. Le Gros, Mammalian NLR proteins; discriminating foe Mast cells in the front line. Innate immunity. from friend. Immunol Cell Biol, 2007. 85(6): p. Curr Biol, 1996. 6(8): p. 941-2. 495-502. 2. Weissmann, G., J.E. Smolen, and H.M. 10. Neefjes, J.J., T.N. Schumacher, and H.L. Korchak, Release of inflammatory mediators Ploegh, Assembly and intracellular transport from stimulated neutrophils. N Engl J Med, of major histocompatibility complex mole- 1980. 303(1): p. 27-34. cules. Curr Opin Cell Biol, 1991. 3(4): p. 601- 3. Trinchieri, G., Natural killer cells wear different 9. hats: effector cells of innate resistance and 11. Cresswell, P., Assembly, transport, and func- regulatory cells of adaptive immunity and of tion of MHC class II molecules. Annu Rev hematopoiesis. Semin Immunol, 1995. 7(2): Immunol, 1994. 12: p. 259-93. p. 83-8. 12. Kaye, J., et al., Selective development of 4. Hirai, K., et al., Regulation of the function of CD4+ T cells in transgenic mice expressing a eosinophils and basophils. Crit Rev Immunol, class II MHC-restricted antigen receptor. 1997. 17(3-4): p. 325-52. Nature, 1989. 341(6244): p. 746-9. 5. Greenberg, S. and S. Grinstein, Phagocytosis 13. Belz, G.T., F.R. Carbone, and W.R. Heath, and innate immunity. Curr Opin Immunol, Cross-presentation of antigens by dendritic 2002. 14(1): p. 136-45. cells. Crit Rev Immunol, 2002. 22(5-6): p. 6. Kaisho, T. and S. Akira, Critical roles of Toll- 439-48. like receptors in host defense. Crit Rev 14. Monaco, J.J., A molecular model of MHC Immunol, 2000. 20(5): p. 393-405. class-I-restricted antigen processing. 7. Wright, J.R., Immunoregulatory functions of Immunol Today, 1992. 13(5): p. 173-9. surfactant proteins. Nat Rev Immunol, 2005. 15. Emmrich, F., U. Strittmatter, and K. 5(1): p. 58-68. Eichmann, Synergism in the activation of 8. McGreal, E.P., J.L. Miller, and S. Gordon, human CD8 T cells by cross-linking the T-cell Ligand recognition by antigen-presenting cell receptor complex with the CD8 differentiation C-type lectin receptors. Curr Opin Immunol, antigen. Proc Natl Acad Sci U S A, 1986. 2005. 17(1): p. 18-24. 83(21): p. 8298-302. 9. Kaparakis, M., D.J. Philpott, and R.L. Ferrero, 16. Banchereau, J. and R.M. Steinman, Dendritic

18 General introduction

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Chapter 2 Human and murine model cell lines for dendritic cell biology evaluated

Suzanne F.G. van Helden, Frank N. van Leeuwen and Carl G. Figdor

Immunology Letters, 2008, 117: 191-197 Chapter 2

Abstract Dendritic cells (DCs) are specialized antigen presenting cells that link innate and adaptive immune responses. As key mediators of T cell dependent immunity, DCs are considered primary targets for initiat- ing immune responses in infectious diseases and cancer. Conversely, DCs can also play an important role in the induction of tolerance in organ transplantation, autoimmune disorders and allergy. While DCs have been used in clinical trials worldwide during the past decade, many of the highly specialized cell biological characteristics of DCs remain poorly understood. Small numbers of DCs can be isolated as terminally differentiated, post-mitotic cells from either blood or spleen. Alternatively, DC-precursors, such as monocytes or bone marrow-derived stem cells, can be isolated and differentiated into DCs in vitro. The relative low numbers of cells that can thus be obtained, combined with difficulties manipulating these terminally differentiated primary cells in vitro and in vivo, have seriously hampered studies aimed at exploring the cell biology of DCs. Good model cell lines therefore provide invaluable tools to study DC biology. So far most DC models are myeloid leukemia-derived cell lines that can be differentiated in vitro towards a DC phenotype. Here, we compared the phenotypical and functional characteristics of frequently used mouse and human DC-model cell lines. We conclude that, although none of these cell lines fully recapitulates all cell biological or immunological features of primary DCs, some of these cell lines provide valuable tools to study specific aspects of DC biology. Introduction Antigen presenting cells, such as B cells, macrophages and dendritic cells (DCs), are responsible for the induction of cellular immune responses, by internalizing and processing antigens for presentation to T cells. Of all antigen presenting cells, DCs are by far the most effective inducers of T cell-based immunity [1]. As DCs are also rapidly activated in response to bacterial or viral antigens, these cells play an essential role in linking innate and adaptive immune responses. DCs originate from myeloid precursors that during their development move from the blood into the (peripheral) tissues. These tissue-resident DCs, generally known as immature DCs (iDCs), are equipped with specialized receptors, such as Toll-like receptors that allow them to efficiently recognize and respond to pathogens through pathogen-associated molecules. In addition, C- type lectins and Fc receptors present on these cells facilitate internalization and processing of pathogens or other foreign antigens. When iDCs are exposed to pathogens, antigens and/or pro-inflam-

26 Model cell lines for DC biology evaluated matory signals, they undergo a dramatic conversion known as maturation. During maturation DCs upregulate major histocompatibility complex (MHC) and costimulatory molecules, activate their antigen processing machinery and become highly migratory, which enables these cells to travel to the lymph nodes were they interact with B and T cells to initiate an adaptive immune response. Since DCs are so well equipped to initiate adaptive immune responses, they are considered prime targets for modulating immune responses against cancer, but also vehicles that can be used to dampen responses in case of autoimmunity, allergy or transplantation. Although laboratories worldwide are exploring therapeutic applications of DCs, many aspects of DC-biology are still poorly understood. This is to a large part due to the fact that studying DC-biology in vitro is difficult for a number of reasons. First of all, the most commonly used DCs are isolated from either spleen [2] or blood [3, 4] and post-mitotic, limiting the amount of cells that can be obtained. For instance, myeloid DCs, which can be isolated directly from peripheral blood [5], make up less than 1 % of all mononuclear cells. Consequently, isolation of these myeloid DCs by flow cytometry or magnetic bead sorting is rather inefficient and expensive, while relatively small numbers of cells can be obtained. Moreover, the life span of such cultures is limited since DCs are terminally differentiated and cannot divide. Larger numbers of DCs can be derived by in vitro differentiation of bone marrow stem cells [6], peripheral blood mononuclear cells [7] or monocytes [8]. Isolation of these DC-precursors from the bone marrow, whole blood or buffy-coat material is laborious while their subsequent differentiation towards DCs is time consuming, requiring six to nine days. Moreover, transfection or electroporation of recombinant DNA or short hairpin RNAs into DCs is generally very ineffective and associated with significant cell death. Finally, experiments using these primary cells are often complicated by donor to donor variations in cytokine expression levels, cell surface molecule expression levels and the ability to stimulate T cells. Hence, there is a significant need for model cell lines that accurately replicate cell biological aspects of DC biology such as antigen uptake, processing and presentation and that can be induced to undergo maturation in response to pathogens or cytokine stimulation. Here we compare a number of cell lines of either mouse or human origin, for their use as DC models. As mouse models, we evaluated Raw264.7, J774 and D1 cells, while the cell lines THP-1, HL60 and MUTZ-3 cells were compared for their ability to serve as human DC equivalents. We examined the presence of DC-specific molecules and describe the ability of each of these cell lines to respond to mat-

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uration stimuli. Moreover, we have compared the ability of these cells to take-up and present antigens as well as examined adhesive and migratory properties either in unstimulated cells or after receiving a differentiation signal. Finally, we evaluated ease of culture and transfectability of these different DC models. Expression of MHC, costimulatory and DC-specific molecules DC maturation, induced by exposure to pathogens or inflammatory stimuli, is characterized by upregulation of MHC and costimulatory molecules and accompanied by extensive changes in the cytoskeletal organization [9-13]. Raw264.7 and J774 cells are widely used murine cell lines with macrophage properties derived from tumors in Balb/c mice [14, 15]. Although not as efficient in antigen presentation as DCs, significant similarities exist between DC and macrophages regarding their response after exposure to pathogens or foreign antigens. When compared to DCs, these macrophage cell lines express costimulatory molecules and MHC class I and II ([16, 17] and our own observations). Moreover, Raw264.7 cells efficiently upregulate surface expression of CD80, CD86, CD40 and MHC class II molecules in response to lipopolysaccharide (LPS) [16, 17]. In addition, we observed that Raw264.7, differentiated by exposure to IL-4 and GM-CSF for 7 days upregulated surface expression of CD11c, a DC- specific marker in mice (Table 1). When these IL-4 and GM-CSF treated cells are further stimulated with LPS for 24 h, upregulation of the costimulatory molecule CD40 is observed. J774 cells also express MHC class II and costimulatory molecules [18] and respond to LPS by upregulating CD86 expression [19]. We similarly stimulated J774 cells with IL-4 and GM-CSF with or without LPS but observed no change in the expression pattern of MHC and costimulatory molecules and no expression of CD11c (Table 1). The third mouse cell line that we evaluated is D1, a growth factor dependent cell line derived from spleen cells of C57BL/6 mice [20]. Even without cytokine stimulation, these cells closely resemble mouse iDCs, based on expression of DC-specific markers as well as MHC- and costimulatory molecules. Moreover, D1 cells acquire a mature phenotype upon exposure to inflammatory stimuli [20]. D1 cells express MHC class I and II molecules, the costimulatory molecules CD80, CD86 and CD40, as well as CD11c [20]. After maturation MHC class II, CD86 and CD40 are further upregulated [20] (Table 1). In addition to MHC, costimulatory and DC-specific molecules, Raw264.7, J774 and D1 cells express receptors for recognition and up-take of pathogens [19-25]. Of these three murine cell lines examined, the

28 Model cell lines for DC biology evaluated

iD1 mD1 Raw Raw Raw J774 J774 J774 IL-4 IL-4 IL-4 IL-4 GM-CSF GM-CSF GM-CSF GM-CSF LPS LPS MHC class II + m + i + + + + + + MHC class I + + + + + + + + CD80 + + low low low - - - CD86 low + low low low low low low CD40 + + low low + low low low CD11c + + low + + - - - Table 1. Expression of MHC and costimulatory molecules and CD11c in D1, Raw264.7 and J774 cells. Immature (iD1) and mature D1 (mD1) cells, unstimulated Raw264.7 (Raw) cells or J774 cells were compared. In addition, Raw264.7 cells and J774 cells were cultured with IL-4 and GM-CSF (both 20 ng/ml) for 7 days (Raw+IL-4/GM-CSF and J774+IL-4/GM-CSF). These stimulated cells were further stimulated with LPS for 24 h (Raw+IL-4/GM-CSF+LPS and J774+IL-4/GM-CSF+LPS). +; expressed, low; low expression and -; not expressed, m; membrane, i; intracellular receptor repertoire in D1 cells most closely resembles that found in DCs. Taken together, these data suggest that although all three cell lines express, or can be stimulated to express, MHC molecules and costimulatory molecules, only Raw264.7 and D1 cells can be stimulated to express DC-specific molecules. We conclude that of these mouse models, D1 cells most closely resemble primary DCs. It should be noted however that D1 cells are more difficult to grow when compared to the other cell lines. THP-1, HL-60 and MUTZ-3 cells are all human leukemia cell lines with monocyte-like properties, which makes them potentially good surrogates for human DCs. THP-1 is a monoblastic leukemia cell line established from a 1-year-old male [26]. While lacking expression of costimulatory and MHC class II molecules ([27] and our own observations), THP-1 cells express MHC class I molecules [28] and can be induced to express the costimulatory molecule CD86 in response to LPS [29]. Moreover, combined stimulation of THP-1 cells with IL-4 and protein kinase C activators such as bryostatin or phorbol 12- myristate 13-acetate (PMA) leads to expression of CD80, CD86, CD40 and the DC-specific C-type lectin Dendritic Cell-Specific ICAM3 Grabbing Non-integrin (DC-SIGN) ([30] and our own observations) (Table 2). A second myeloid leukemia cell line, HL-60, established from a 35-year-old female is arrested at the myeloblast differentiation stage and can be differentiated to monocytes or granulocytes [31-34]. HL-60 cells, express MHC class I molecules [35], but lack expression of MHC class II and costimulatory molecules [27]. Stimulation with the calcium ionophore ionomycin was shown to

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induce expression of CD80, CD86 and CD83 [36], which indicates that this stimulation results in a more mature phenotype. No (enhanced) expression of costimulatory molecules or MHC molecules was induced by treatment with the phorbol-ester PMA (Table 2).

HL-60 HL-60 MUTZ-3 iMUTZ-DC mMUTZ-DC THP-1 THP-1 PMA PMA IL-4 MHC class I + + + + + + + MHC class II - - + + + - - CD80 - - low + + - + CD86 - - + + + - + CD40 nd nd + + + + + CD14 - - + - - low - CD83 - - - - + - - DC-SIGN - - - + + - +

Table 2. Expression of MHC and costimulatory molecules, monocyte marker CD14, DC- maturation marker CD83 and DC-specific molecule DC-SIGN in HL-60, MUTZ-3 and THP-1 cells. Unstimulated HL-60, MUTZ-3 or THP-1 cells were compared. In addition, HL-60 cells were stimulated with 0.1 µM PMA for 24h and THP-1 cells were stimulated with 0.1µM PMA and 500 U/ml IL-4 for 24h. MUTZ-3 cells were differentiated with GM- CSF (100 ng/ml) and IL-4 (1000 U/ml) for 7 days to obtain iMUTZ-DCs and further α matured with MCM, TNF and PGE2 for 2 days to obtain mMUTZ-DCs. +; expressed, low; low expression, -; not expressed and nd; not determined.

MUTZ-3 is a human myeloid leukemia cell line established from a 29- year-old male [37]. Similar to primary monocytes, MUTZ-3 cells can be differentiated in six or seven days using GM-CSF and IL-4 to acquire an iDC-phenotype (iMUTZ-DCs) [38]. Moreover, maturation can be induced by treatment with LPS or cytokines to acquire a phenotype (mMUTZ-DCs) closely resembling that of mature primary DCs [38]. Undifferentiated MUTZ-3 cells express CD34, CD14, CD86, CD40 and MHC class I an II molecules ([38] and our own observations). Upon differentiation with IL-4 and GM-CSF, CD14 and CD34 are downregulated, while the cells start to express CD80 and DC-SIGN, and upregulate CD86, CD40 and MHC class II molecules ([38] and our own observations) (Table 2). Upon further stimulation with LPS or cytokine combinations CD80, CD86, CD40 and MHC class II molecules are further upregulated and the cells start to express the maturation marker CD83 and the chemokine receptor CCR7 [38, 39]. All three human cell lines express receptors for recognition and up-take of pathogens [29, 30, 38, 40-42]. The

30 Model cell lines for DC biology evaluated expression pattern of these receptors in (stimulated) THP-1 cells more closely resembles that of DCs when compared to HL-60 cells, however the receptor profile of MUTZ-3 cells most closely approaches that of primary DCs. Taken together we conclude that HL-60 cells express MHC class I but lack expression of MHC class II, costimulatory and DC-specific molecules, and do not significantly respond to signals reported to enhance differentiation/maturation. THP-1 cells express costimulatory, MHC class I and II molecules and can be induced to acquire some DC like features, such as expression of costimulatory molecules and DC-SIGN. However, in comparison to these two cell lines, differentiation and maturation of MUTZ-3 cells most accurately mirrors normal DC development. On the negative side, MUTZ-3 cells are heterogeneous and more difficult to propa- gate when compared to HL-60 and THP-1. Antigen uptake and presentation iDCs are strongly endocytotic cells that use phagocytosis and pinocytosis to take up antigens and sample their environment. In response to maturation, internalized antigens are processed and presented to T cell as peptides in the context of MHC class I/II molecules. Both Raw264.7 and J774 cells efficiently take up antigen by phagocytosis [23, 43] and pinocytosis [44, 45]. Consequently, these cells are widely used model cell lines to study phagocytosis. In addition, release of cytokines such as IL-6, TNFα, IL-1β [46, 47] and IL- 12 [48, 49] , in response to LPS, occurs efficiently in both cell lines. However, while LPS and/or phosphatidic acid-activated Raw264.7 cells can stimulate alloreactive T cells [17], J774 cells cannot. Similar to these two macrophage cell lines, (immature) D1 cells efficiently take up antigens by fluid phase-, as well as receptor-mediated endocytosis, while after maturation the endocytotic capacity decreases [20, 50]. Furthermore, maturation of D1 cells leads to the redistribution of MHC class II molecules from intracellular vesicles to the plasma membrane [20]. Similar to primary mature DCs, mature D1 cells produce IL-12p75 in response to TNF [20] and efficiently stimulate T cells [20]. In addition, D1 cells can present peptides derived from exogenous antigens in the context of MHC class I molecules [51-53], a mechanism known as crosspresentation, which is considered to be essential for the induction of cytotoxic T cell (CTL) responses. To summarize, Raw264.7, J774 as well as D1 cells can take up antigen, however only the Raw264.7 and D1 cells have the capacity to process and present antigen to T cells. While Raw264.7 provide and excellent model for studying phagocytosis, D1 cells are superior to the Raw264.7 cells in their ability to stimulate T cells and

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therefore more closely resemble primary mouse DCs. Hence, D1 cells are the model of choice for studying antigen presentation. The human cell line THP-1 [54] as well as differentiated MUTZ-3 [40] or HL-60 cells [55] are capable of taking up antigen by endocytosis. THP-1 cells secrete T cell recruiting and differentiating cytokines, such as IL-6 and IL-8 [56], leading to moderate stimulation of T cells [57, 58]. HL-60 cells produce IL-6 [59] and IL-8 [60] and can also stimulate T cells upon activation, albeit moderately [61, 62]. The relative low T cell stimulating capacity of THP1 and HL 60 cells is reflected by the absence of MHC class II molecules. Unstimulated MUTZ-3 cells are also weak inducers of T cell stimulation, while iMUTZ-DCs, and even more so mMUTZ-DCs, produce IL-12 and are efficient T cell stimulators [38, 63]. Upon external loading with MHC class I-specific peptides MUTZ-DCs can induce potent CTL responses [63]. THP-1 cell are able to crosspresent antigens in MHC class I molecules and this is enhanced upon differentiation [64]. MUTZ-3 cells are likely to be suitable to study crosspresentation, since these cells are much more efficient in stimulating T cells than THP-1 cells. In conclusion, all three human cell lines compared here are able to take up antigens, however the MUTZ-DCs are by far the most efficient in stimulating T cells [40, 65].

Fig. 1. Morphology of J774 and Raw264.7 cells upon stimulation with cytokines and LPS. J774 or Raw264.7 cells adhere on fibronectin-coated coverslips and upon stimulation with mouse GM-CSF and IL-4 (both 20 ng/ml) for 7 days these cells spread and become more adhesive as visualized by an actin staining. Additional stimulation with LPS for 24 h results in even further spreading. Note, how particularly the Raw264.7 cells develop elaborate filopodium-like protrusions after stimulation.

Adhesion and migration In addition to endocytotic capacity, cytokine production and antigen presenting capabilities, regulation of adhesive and migratory proper-

32 Model cell lines for DC biology evaluated ties is an important aspect of DC-behavior. Immature DCs are strongly adherent cells that spread and form podosomes [13, 66]. These specialized adhesion structures are found in Src transformed cells, as well as myeloid cells, such as osteoclasts, monocytes, DCs and macrophages. Podosomes consist of an actin-dense core surrounded by a ring containing structural proteins, such as vinculin, paxillin and myosin II [67]. Podosomes are thought to play a role in crossing tissue boundaries. During maturation DCs loose podosomes and transform from slowly migrating cells into cells capable of high-speed migration [13]. Raw264.7 and J774 cells are strongly adherent cells and after exposure to IL-4/GM-CSF and LPS these cells undergo extensive spreading and become even more adherent (Figure 1), making these cells extremely suitable for microscopy studies. This response is most pronounced in the Raw264.7 cells. Under normal culture conditions, neither Raw264.7 nor J774 cells form podosomes, and neither can such structures be induced in response to IL-4/GM-CSF and LPS. However, at least in Raw264.7 cells, podosome formation can be induced by the addition of RANK ligand, an osteogenic factor that promotes differentiation of these cells towards osteoclasts [68]. In spite of their phenotypic resemblance to mouse DCs, iD1 cells are less adherent than primary mouse DCs and podosomes cannot be found ([20] and our own observations) suggesting that iD1 are not suitable to study this aspect of DC-biology. However, by similarity to mouse-derived DCs, iD1 cells are slowly migrating cells that switch to high-speed migration in response to maturation [20]. Hence, to study podosomes turnover in a mouse model, Raw264.7 cells differentiated towards osteoclasts can be used. However, D1 cells more strongly resemble primary mouse DCs, as these cells can be induced to acquire a highly migratory phenotype in response to maturation stimuli. The human cell lines, THP-1, HL-60 and MUTZ-3 are suspension cells and thus unable to form podosomes. Upon differentiation into iMUTZ-DCs podosomes can be observed in a very small portion of these cells (our own observations) (Figure 2). However, when THP- 1 cells are stimulated with a combination of IL-4 and PMA or when HL-60 cells are stimulated with PMA alone, most of these cells adhere and form podosomes ([69] and our own observations) (Fig 2). Similar to podosomes observed in primary human DCs, PMA- induced podosomes in THP1 and HL-60 cells dissolve within minutes after the addition of the inflammatory mediator prostaglandin E2

(PGE2), a potent migratory stimulus for these cells [13]. Moreover, THP-1 show directed migration towards several chemokines important for DC migration, while HL-60 cells can only do so when differ-

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Fig. 2. Morphology of different human cell lines compared to monocyte-derived DCs. A. Comparison of adhesive behavior and podosome formation between human monocyte-derived DCs and various DC model cell lines, seeded on fibronectin. Monocytes were isolated from buffy-coat material and plastic adherent monocytes were cultures with GM-CSF (800 U/ml) and IL-4 (500 U/ml) for 6 days to obtain iDCs. To induce maturation iDCs were stimulated with monocyte-conditioned medium (MCM, α µ 50% v/v), TNF (10 ng/ml) and PGE2 (10 g/ml) for 2 days. MUTZ-3 cells were differentiated with GM-CSF (100 ng/ml) and IL-4 (1000 U/ml) for 7 days to obtain iMUTZ-DCs α and further matured with MCM, TNF and PGE2 for 2 days to obtain mMUTZ-DCs. HL- 60 cells were left untreated or stimulated with 0.1 µM PMA for 24 h and THP-1 cells were left untreated or stimulated with 0.1 µM PMA and 500 U/ml IL-4 for 24 h. Cells were seeded on fibronectin-coated coverslips and stained for vinculin (green) and actin (red). HL-60, THP-1, MUTZ-3 and iMUTZ-DCs are poorly adherent cells that do not develop specific adhesion structures. Monocytes, iDCs, but also stimulated HL-60 and THP-1 cells are strongly adherent and develop podosomes that can be seen as vinculin rings (green) surrounding actin dots (red). mDCs and mMUTZ-DCs show reduced adherence accompanied by a loss of podosomes.

B. Podosome dissolution induced by PGE2. iDCs, HL-60 stimulated with PMA for 24 h and THP-1 cells stimulated with PMA/IL-4 for 24 h were seeded on fibronectin and stim- µ ulated with 10 g/ml PGE2 for 5 min and stained for vinculin (green) and actin (red). Podosomes that were present in these cells (panel A) rapidly dissolve in response to stimulation, while the cells remain attached. This response is somewhat less pronounced in the THP-1 cells.

entiated towards monocytes [70]. In contrast to primary DCs, however, neither of these two cell types switch to a high-speed migration mode in response to maturation [71]. Relative to primary DCs,

34 Model cell lines for DC biology evaluated

MUTZ-DCs are poorly adhesive, even in the immature state, but similar to DCs, MUTZ-DCs acquire a more migratory phenotype in response to maturation (our own observations). Taken together these results demonstrate, that THP-1 and HL-60 cells are useful models to study podosome biogenesis and turnover, while MUTZ-3 cells more accurately reflect the induction of high-speed migration that occurs during DC maturation. Culture and manipulation In addition to closely resembling DCs, a good model cell line should be easy to culture and readily transfectable. J774 and Raw264.7 cells are easy to culture cell lines, which can either be subjected to retroviral transduction [72] or transfected with calcium phosphate [73, 74], DEAE dextran [24] or other transfection reagents such as lipo- fectamine [75, 76]. D1 cells are dependent on conditioned medium [20] and need to be maintained under carefully controlled conditions in order to prevent loss of their specific DC-like features. Although retroviral transduction of D1 cells has been reported [77], to date, successful transfection of these cells has not been established. As for human DC models, MUTZ-3 cells are dependent on the addition of conditioned medium and these cells need to be cultured and differentiated towards MUTZ-DCs under carefully defined culture conditions. The differentiation of MUTZ-3 cells is as time consuming and expensive as differentiation of DCs from peripheral blood mononuclear cells or monocytes. By contrast, THP-1 and HL-60 cells are easy to culture suspension cells. Furthermore, induction of a DC- morphology in THP-1 or HL-60 cells is less time consuming and does not require the use of expensive cytokines. All three human cell lines can be efficiently tranduced with retroviral vectors [38, 78]. There are some reports were THP-1 or HL-60 cells are transfected by electroporation [79], however the transfection efficiency is low. MUTZ-3 cells can be transfected with low efficiency using Amaxa nucleofection but this is associated with significant cell death (our own observations). Conclusions and perspectives It can be concluded that we have yet to find the perfect DC-model, as none of the cell lines described here completely mirrors all cell biological/immunological aspects of DCs. Strengths and weaknesses of each of the particular model cell lines are summarized in Table 3. In the mouse setting, D1 cells most closely resemble primary DCs, however culture conditions and poor transfectability of these cells stand in the way of large-scale biochemical experiments. For those

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Mouse Human

D1 J774 Raw264.7 HL-60 MUTZ-3 THP-1

adherent - ++ ++ + 1 - + 2 endocytosis + ++ ++ + 3 + 5 + cytokines ++ + + + ++ + chemotaxis + + + + 1 + +

migration ++ + (slow) + (slow) + 1 ++ 5 + antigen presentation/T cell stimu- ++ - + 6 + ++ + lation DC-phenotype ++ - + 5 - ++ 5 + 2 podosomes - - + 4 + 1 - + 2 transducable + + + + + + transfectable - + + - - -

easy to culture -(sup R1) + + + - (sup 5637) +

Table 3. Overview of DC-functions displayed by D1, J774, Raw264.7, HL-60, MUTZ-3 and THP-1 cells. 1; upon PMA stimulation, 2; upon PMA/IL-4 stimulation, 3; upon

Me2SO stimulation, 4; upon RANKL stimulation, 5; upon IL4/GM-CSF stimulation (and maturation) and 6; upon LPS or phosphatidic acid stimulation. ++; very good, +; good and -; poor. Sup R1: supernatant of R1 cells needed, sup 5637: supernatant of 5637 cells needed

types of experiments, Raw 264.7 cells may be a more suitable model. In the human setting THP-1 and HL-60 cells are robust cell lines that can be used when large numbers of cells are needed, or when the focus is on studying podosome formation or turnover. However, while both THP-1 and HL-60 cells mimic some aspects of iDC-biology, MUTZ-3 cells are the only model cell line suitable for investigating mDC-function, although the need to carefully control culture conditions is a complicating factor. With the exception of strong adhesion and podosome formation in the immature state, MUTZ-3 derived DCs most closely resemble primary DCs. Particularly when it comes to immunological experiments involving antigen presentation and T cell stimulation, these cells are the system of choice. In spite of their limitations, a number of DC model cell lines can be successfully used to study important aspects of DC biology in vitro. We believe that such in vitro approaches will continue to be important, as poor transfectability and donor to donor variations preclude the use of primary cells in some experiments. We expect that the proper use of DC model cell lines may help reveal important new

36 Model cell lines for DC biology evaluated insights into DC biology that can subsequently be applied to improve application of DCs in the clinic.

Acknowledgements We want to thank dr. R. Scheper for providing the MUTZ-3 cells and dr. L Machesky for providing the HL-60 cells and M. Oud for experimental support. SvH is supported by research funding from the Foundation for Fundamental Research on Matter (FOM 01FB06) and the Dutch Cancer Society (KWF KUN 2006-3699). FvL is supported by research funding from the Dutch Cancer Society (KWF KUN 2006-3485). References 1. Banchereau, J. and R.M. Steinman, Dendritic receptor expression. Immunol Rev, 2000. cells and the control of immunity. Nature, 177: p. 134-40. 1998. 392(6673): p. 245-52. 11. Gunzer, M., et al., Antigen presentation in 2. Steinman, R.M. and Z.A. Cohn, Identification extracellular matrix: interactions of T cells of a novel cell type in peripheral lymphoid with dendritic cells are dynamic, short lived, organs of mice. I. Morphology, quantitation, and sequential. Immunity, 2000. 13(3): p. tissue distribution. J Exp Med, 1973. 137(5): 323-32. p. 1142-62. 12. West, M.A., et al., Enhanced dendritic cell 3. van Voorhis, W.C., et al., Human dendritic antigen capture via toll-like receptor-induced cells. Enrichment and characterization from actin remodeling. Science, 2004. 305(5687): peripheral blood. J Exp Med, 1982. 155(4): p. p. 1153-7. 1172-87. 13. van Helden, S.F., et al., A critical role for 4. Crow, M.K. and H.G. Kunkel, Human dendrit- prostaglandin E2 in podosome dissolution ic cells: major stimulators of the autologous and induction of high-speed migration during and allogeneic mixed leucocyte reactions. dendritic cell maturation. J Immunol, 2006. Clin Exp Immunol, 1982. 49(2): p. 338-46. 177(3): p. 1567-74. 5. Reid, S.D., G. Penna, and L. Adorini, The con- 14. Raschke, W.C., et al., Functional trol of T cell responses by dendritic cell sub- macrophage cell lines transformed by sets. Curr Opin Immunol, 2000. 12(1): p. 114- Abelson leukemia virus. Cell, 1978. 15(1): p. 21. 261-7. 6. Inaba, K., et al., Generation of large numbers 15. Ralph, P. and I. Nakoinz, Phagocytosis and of dendritic cells from mouse bone marrow cytolysis by a macrophage tumour and its cultures supplemented with cloned cell line. Nature, 1975. 257(5525): p. granulocyte/macrophage colony-stimulating 393-4. factor. J Exp Med, 1992. 176(6): p. 1693-702. 16. Saxena, R.K., V. Vallyathan, and D.M. Lewis, 7. Romani, N., et al., Proliferating dendritic cell Evidence for lipopolysaccharide-induced dif- progenitors in human blood. J Exp Med, ferentiation of RAW264.7 murine 1994. 180(1): p. 83-93. macrophage cell line into dendritic like cells. J 8. Kiertscher, S.M. and M.D. Roth, Human Biosci, 2003. 28(1): p. 129-34. CD14+ leukocytes acquire the phenotype and 17. Lee, Y.N., et al., Phosphatidic acid positively function of antigen-presenting dendritic cells regulates LPS-induced differentiation of when cultured in GM-CSF and IL-4. J Leukoc RAW264.7 murine macrophage cell line into Biol, 1996. 59(2): p. 208-18. dendritic-like cells. Biochem Biophys Res 9. De Vries, I.J., et al., Effective migration of Commun, 2004. 318(4): p. 839-45. antigen-pulsed dendritic cells to lymph nodes 18. Zur Lage, S., et al., Activation of in melanoma patients is determined by their macrophages and interference with CD4+ T- maturation state. Cancer Res, 2003. 63(1): p. cell stimulation by Mycobacterium avium sub- 12-7. species paratuberculosis and Mycobacterium 10. Sallusto, F. and A. Lanzavecchia, avium subspecies avium. Immunology, 2003. Understanding dendritic cell and T-lympho- 108(1): p. 62-9. cyte traffic through the analysis of chemokine 19. O'Farrell, A.M., et al., IL-10 inhibits

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Hu, Z.B., et al., Establishment and character- member contribute to the myeloid-specific ization of two novel cytokine-responsive transcription of the Fc gamma RIIIA promoter. acute myeloid and monocytic leukemia cell EMBO J, 1994. 13(16): p. 3852-60. lines, MUTZ-2 and MUTZ-3. Leukemia, 1996. 25. Bakker, A.B., et al., Myeloid DAP12-associat- 10(6): p. 1025-40. ing lectin (MDL)-1 is a cell surface receptor 38. Masterson, A.J., et al., MUTZ-3, a human cell involved in the activation of myeloid cells. line model for the cytokine-induced differenti- Proc Natl Acad Sci U S A, 1999. 96(17): p. ation of dendritic cells from CD34+ precur- 9792-6. sors. Blood, 2002. 100(2): p. 701-3. 26. Tsuchiya, S., et al., Induction of maturation in 39. Kim, K.D., et al., Impaired responses of cultured human monocytic leukemia cells by leukemic dendritic cells derived from a human a phorbol diester. Cancer Res, 1982. 42(4): p. myeloid cell line to LPS stimulation. Exp Mol 1530-6. 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Carpentier, V., et al., Characterization and sone-induced suppression of IFN-gamma. J cellular localization by monoclonal antibodies Interferon Cytokine Res, 2000. 20(11): p. of the 60 kDa mannose specific lectin of 927-34. human promyelocytic cells, HL60. Glycoconj 30. Puig-Kroger, A., et al., Regulated expression J, 1994. 11(4): p. 333-8. of the pathogen receptor dendritic cell-specif- 43. Suram, S., et al., Regulation of cytosolic ic intercellular adhesion molecule 3 (ICAM-3)- phospholipase A2 activation and cyclooxyge- grabbing nonintegrin in THP-1 human nase 2 expression in macrophages by the leukemic cells, monocytes, and beta-glucan receptor. J Biol Chem, 2006. macrophages. J Biol Chem, 2004. 279(24): p. 281(9): p. 5506-14. 25680-8. 44. Han, C.Y., S.Y. Park, and Y.K. Pak, Role of 31. Collins, S.J., R.C. Gallo, and R.E. 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38 Model cell lines for DC biology evaluated

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39 Chapter 2

beta-chemokines. Cytokine, 1997. 9(7): p. 521-8. 72. Ganz, T., et al., Posttranslational processing and targeting of transgenic human defensin in murine granulocyte, macrophage, fibroblast, and pituitary adenoma cell lines. Blood, 1993. 82(2): p. 641-50. 73. Buras, J.A., B.G. Monks, and M.J. Fenton, The NF-beta A-binding element, not an over- lapping NF-IL-6-binding element, is required for maximal IL-1 beta gene expression. J Immunol, 1994. 152(9): p. 4444-54. 74. Mazzone, T., L. Pustelnikas, and C.A. Reardon, Post-translational regulation of macrophage apoprotein E production. J Biol Chem, 1992. 267(2): p. 1081-7. 75. Liu, Q.P., et al., Negative regulation of macrophage activation in response to IFN- gamma and lipopolysaccharide by the STK/RON receptor tyrosine kinase. J Immunol, 1999. 163(12): p. 6606-13. 76. Pierce, R.A., et al., Monocytic cell type-specific transcriptional induction of collagenase. J Clin Invest, 1996. 97(8): p. 1890-9. 77. Gasperi, C., et al., Retroviral gene transfer, rapid selection, and maintenance of the immature phenotype in mouse dendritic cells. J Leukoc Biol, 1999. 66(2): p. 263-7. 78. Unger, R.E., et al., The nef gene of simian immunodeficiency virus SIVmac1A11. J Virol, 1992. 66(9): p. 5432-42. 79. Hickstein, D.D., et al., Identification of the promoter of the myelomonocytic leukocyte integrin CD11b. Proc Natl Acad Sci U S A, 1992. 89(6): p. 2105-9.

40 Chapter 3 A critical role for PGE2 in podosome dissolution and induction of high-speed migration during dendritic cell maturation

1Suzanne F.G. van Helden, 3Daniëlle J.E.B. Krooshoop, 1Karin C.M. Broers, 2Reinier A.P. Raymakers, 1Carl G. Figdor and 1Frank N. van Leeuwen

1Department of Tumor Immunology, NCMLS, Radboud University Nijmegen Medical Centre, Nijmegen, The Netherlands 2Department of Hematology, NCMLS, Radboud University Nijmegen Medical Centre, Nijmegen, The Netherlands 3Department of Infectious Diseases and Immunology, Faculty Veterinary Medicine, University of Utrecht, Utrecht, The Netherlands

The Journal of Immunology, 2006, 177: 1567-1574 Chapter 3

Abstract Dendritic cells (DCs) are professional antigen presenting cells of the immune system that play a key role in regulating T cell-based immunity. The capacity of DCs to activate T cells depends on their maturation state as well as their ability to migrate to the T cell areas of draining lymph nodes. Here we investigated the effects of DC maturation stimuli on the actin cytoskeleton and β1-integrin-dependent adhesion and migration. Podosomes, specialized adhesion structures found in immature DCs, rapidly dissolve in response to matu- α ration stimuli such as TNF and PGE2, whereas the TLR agonist LPS induce podosome dissolution only after a long lag time. We demonstrate that LPS-mediated podosome disassembly as well as the onset of high-speed DC migration is dependent on the production of prostaglandins by the DCs. Moreover, both of these processes are inhibited by Ab-induced activation of β1-integrins. Together, these results show that maturation-induced podosome dissolution and loss of α5β1-integrin activity allow human DCs to undergo the transition from an adhesive to a highly migratory phenotype. Introduction Dendritic cells (DCs) are the most potent antigen presenting cells of the immune system [1]. Upon antigen uptake and exposure to inflammatory stimuli these cells undergo a remarkable transition from a tissue resident, endocytic cell type to a highly migratory antigen presenting cell type, a process known as DC maturation [1, 2]. This phenotypical conversion not only involves up-regulation of MHC and costimulatory molecules, but is also accompanied by extensive changes in cytoskeletal organization and cell adhesion [3-6]. The ability of DCs to migrate is essential for the induction of immune responses. Upon antigen recognition and exposure to inflammatory signals, DCs migrate out of the tissues, via the blood or lymph vessels into the lymph nodes, were they activate T cells. We recently demonstrated both in preclinical mouse-models [7] and in human cancer patients [3], that DC-vaccine efficacy directly correlates with DC maturation and the capacity of these cells to migrate into the lymph nodes. The migration of mature DCs (mDCs) can be defined as high-speed migration [8], comparable to the migration of T cells [9]. This type of migration is characterized by short lived and low affinity interactions with the substrate. Immature DCs (iDCs), on the other hand, show an adhesive and low-speed migratory behavior which is dependent on strong interactions with the extracellular matrix (ECM), similar to fibroblasts [8]. β1-integrins are the main adhesion molecules responsible for these DC-ECM interactions [10,

42 PGE2 in DC maturation

11]. Although the fibronectin binding integrins α4β1 and α5β1 are both expressed by DCs, we and others have shown that adhesion of DCs to fibronectin is predominantly α5β1 mediated [3, 12, 13]. In their high affinity state, integrins form multimeric complexes in the plasma membrane that enhance binding to the ECM. The integrin cytoplasmic domains in turn recruit a large number of structural and signaling proteins into higher order complexes that connect integrins to the actomyosin cytoskeleton. In most cells, such adhesive contacts are known as focal adhesions [14]. However, cells of myeloid origin, such as macrophages, osteoclasts and DCs, form adhesion complexes distinct from focal adhesions, known as podosomes [15]. Unlike focal adhesions, podosomes are not linked to contractile actin stress fibers but rather consist of an actin-dense core surrounded by a ring of cytoskeletal proteins [16, 17]. Although a large variety of stimuli, including cytokines, CD40 ligation or TLR ligands induce DC maturation [18-20], these stimuli not always bring out all the characteristics mDCs need to fulfill their function [21]. Particularly migration appears to be insufficiently developed in response to some stimuli. In order to effectively manipulate DC migration in clinical applications, a more detailed knowledge of how maturation factors mobilize DCs is needed. Here we investigated the effects of different maturation stimuli on cell adhesion and migration in DCs. We find that high-speed DC migration, induced by maturation signals, requires dissolution of podosomes as well as loss of β1-integrin activity. In the presence of a β1-activating Ab, both podosome dissolution and cell migration are effectively inhibited. Moreover, we demonstrate that prostaglandins such as PGE2, produced by DCs during maturation, mediate podosome disassembly as well as the onset of high-speed migration.

Results Induction of high-speed migration in response to DC maturation stimuli The ability of DCs to migrate is essential for the efficient induction of T cell responses. In earlier work, we have shown that the capacity of DCs to migrate on fibronectin in vitro, correlates well with their migratory behavior in vivo [3]. Consistent with these previous findings, the majority of the iDCs were slowly migrating with a mean speed of 0.5 ± 0.1 µm/min (Figure 1A). In contrast, DCs matured with a maturation cocktail α (MCM, PGE2, TNF ), routinely used in clinical applications [3, 22], were highly migratory, showing random migration at a mean speed of

43 Chapter 3

Figure 1: mDCs migrate on fibronectin and migration is initiated after 16 to 24 h. (A/B) Immature and mature DCs were added to fibronectin-coated plates and monitored for 60 min. Plots of 50 individual tracks, aligned at their starting positions, are shown. The mean speed ±SEM (µm/min) is depicted in the upper right corner as well as the percentage of migrating cells ±SD. (C) DCs were incubated for different time periods (0, 5, α 16, 24, 41, and 48 h) with a combination of MCM, PGE2 and TNF , added to fibronectin- coated plates and migration was monitored for 60 min. The percentage of migrating cells ±SD is depicted. Significant differences (p<0.05) compared to the percentage of migrating DCs after 48 h of maturation are indicated by an asterisk. 5 ± 0.5 µm/min (Figure 1B). The effects of maturation on cell migration were noticeable after 16 h of stimulation whereas a full migratory capacity was obtained after 24 h of stimulation (Figure 1C).

The activity of α5β1-integrin is down-regulated during DC maturation DC maturation is associated with a significant loss in adhesion to fibronectin [3]. The expression of active β1, as determined by an Ab (12G10) which exclusively recognizes the active conformation of this integrin [23], was almost completely absent on mDCs. At the same time, expression levels of β1-integrins were unaffected during maturation (Fig 2A). These data suggest that the regulation of β1-integrin activity, rather than expression, controls DC adhesion. More importantly, adhesion of mDCs to fibronectin could be completely restored using a β1-activating Ab (TS2/16), while the adhesion of iDCs could

44 PGE2 in DC maturation

Figure 2: Active β1-integrin is down-regulated upon maturation. (A) Active β1-integrin expression on DCs decreases upon maturation. Immature DCs and mDCs were stained using monoclonal antibodies recognizing β1-integrin (AIIB2) or a specific activation epitope of the β1-integrin (12G10). Expression was determined by flow cytometry. Open graphs represent isotype controls. (B) Adhesion of mDCs to fibronectin can be enhanced by Ab-induced activation of β1-integrins. Calcein-AM- labeled iDCs and mDCs were added with or without preincubation with a blocking Ab against α5-integrin (SAM-1) to fibronectin- coated plates, and incubated in the presence or absence of an anti-β1-integrin activating Ab (TS2/16). Cells were lysed and the fluorescence was quantified using a cytofluometer. Results are depicted as the mean percentage of adhesion of triplicate wells SD. Adhesion of iDCs and mDCs incubated with SAM-1 before and after activation with TS2/16 is significantly decreased in comparison to non-treated control cells. Adhesion of mDCs using TS2/16 is significantly increased in comparison to adhesion of mDCs without a β1-integrin activating Ab (p<0.05). not be further enhanced (Fig 2B). Ab-induced adhesion was still α5β1 mediated, since an α5-blocking Ab (SAM-1) effectively interfered with this interaction (Fig 2B). These data demonstrate that, while the α5β1-integrin expressed on mDCs is still capable of mediating adhesion to fibronectin, its activity is specifically down-regulated during DC maturation.

Immature DCs display podosomes enriched in active β1-integrin The dramatic changes in cell adhesion and migration, as they occur during DC maturation, are also reflected in the morphology and cytoskeletal organization of these cells. When plated on fibronectin, iDCs exhibit extensive cell spreading accompanied by the formation of numerous podosomes (Fig 3A, upper panel). In sharp contrast, mDCs were much less spread and no longer carry podosomes (Fig 3A, lower panel). Importantly, both the β1- and α5-chain of the fibronectin-binding α5β1-integrin were present in podosomes and the active form of the β1-integrin, as detected with the 12G10 Ab, was

45 Chapter 3

Figure 3: Immature DCs form podosomes enriched in active α5β1-integrin. (A) Immature DCs form podosomes on fibronectin. Immature and mature DCs were plated on fibronectin-coated coverslips and stained with an anti-vinculin Ab (green) and phalloidin-Texas Red (red) to detect F-actin. In the lower right corner of the overlay of iDCs a higher magnification of part of the image is shown. (B) Active β1-integrin is enriched in podosomes in iDCs. Immature DCs seeded on fibronectin-coated coverslips were stained with monoclonal antibodies against α5- (SAM-1), β1- (AIIB2), and active β 1-integrin (12G10) (green). F-actin was detected using phalloidin-Texas Red (Red). highly enriched in podosomes (Figure 3B). These results indicate that adhesion of iDCs to fibronectin is mediated by active α5β1-integrin molecules present in these podosomes.

DC maturation signals induce rapid disassembly of podosomes Since DC maturation leads to dramatic changes in adhesive behavior and migration, we explored the effect of maturation-inducing factors on the actin cytoskeleton. In the immature state most DCs dis-

46 PGE2 in DC maturation

Figure 4: Rapid dissolution of podosomes in DCs in response to maturation stimuli. Immature DCs were seeded on fibronectin-coated coverslips and stimulated for different periods with maturation stimuli. Subsequently, the cells were stained with an anti- vinculin Ab (green) and phalloidin-Texas Red to stain F-actin (red). (A) Immature DCs were left untreated or stimulated for 5 min or 16 h with a combination of MCM, PGE2 and TNFα. Representative images are depicted. (B) Immature DCs were either left α untreated or stimulated for 5 min or 16 h with a combination of MCM, PGE2 and TNF α , MCM alone, TNF alone, PGE2 alone or LPS. The number of cells containing podosomes was counted in 7 images per condition per experiment. The percentage of cells expressing podosomes ±SEM is depicted. An average of 3 experiments is shown. Conditions that are significantly different from the control situation (p<0.05) are indicated with an asterisk. (C) Immature DCs were transfected with βactinGFP, seeded on fibronectin and imaged for 60 min prior to and 20 min after the addition of TNFα with a

30 second interval or for 20 min with a 20 second interval without PGE2 and 17 min with a 10 second interval with PGE2. Fluorescent images were taken at different time points α after TNF addition (upper panel) and after PGE2 addition (lower panel)(see also Supplementary video 1 and Supplementary video 2). played podosomes. However, in most cells, podosomes were lost within 5-10 min in response to either the maturation cocktail or PGE2 alone, while stimulation with TNFα induced podosome dissolution in

47 Chapter 3

Figure 5: PGE2 is able to mediate complete podosome dissolution at a concentration of 10 µg/ml. Immature DCs were seeded on fibronectin-coated coverslips and left untreated (control) or stimulated for 5 min or 16 h with different concentrations of

PGE2. Subsequently, the cells were stained with an anti-vinculin Ab and phalloidin-Texas Red to stain F- actin. The number of cells containing podosomes was counted in 7 images per condition. The percentage of cells expressing podosomes ±SEM is depicted. One representative experiment is shown. Conditions that are different from the control situation (p< 0.05) are indicated with an asterisk.

Figure 6: The lifetime of individual podosomes is less than 10 minutes. The lifetime of individual podosomes in unstimulated cells was determined by counting in how many frames individual podosome are observed (Supplementary video 2). The lifetime of 50 podosomes was determined and the distribution of podosome lifetimes is depicted. a subset of the DCs (Figure 4A and 4B). Prolonged stimulation did not further affect the number of cells displaying podosomes (Figure 4B). A moderate effect of MCM on podosome dissolution was observed after prolonged stimulation (16 h). These results identify

PGE2 as the most active component in the maturation cocktail. A µ concentration of 10 g/ml PGE2, which is routinely used in the maturation cocktail, was the lowest concentration to induce efficient podosome dissolution (Figure 5). Surprisingly, LPS, a TLR ligand and potent DC maturation factor, failed to induce podosome dissolution after 5-10 min, while prolonged stimulation resulted in a nearly complete loss of podosomes (Figure 4B). These findings demonstrate that, while most maturation stimuli tested induce dissolution of podosomes, the extent to which this occurs as well as timing differs between agonists. To examine the kinetics of podosome dissolution in response to maturation stimuli, we expressed a β-actinGFP con- struct in iDCs and performed live cell imaging. We observed a rapid turnover of individual podosomes in iDCs expressing β-actinGFP (lifetime of individual podosomes was less than 10 min), while the total number of podosomes remained relatively constant (Figure 6). α Addition of PGE2 or TNF to iDCs caused a rapid and complete dissolution of podosomes within minutes after stimulation (Figure 4C and movie 1 and 2). Together our results show that podosomes in DCs are highly dynamic and that podosome dissolution is a first effect of DC maturation.

48 PGE2 in DC maturation

Activation of β1-integrins interferes with podosome dissolution and high-speed migration We have shown that β1-integrins are prominently present in podosomes and DC maturation is initiated by a loss of these adhesion sites, as well as a global decrease in β1-integrin activity. Therefore we examined if and how stimulation of β1-integrin activity using a β1-integrin activating Ab (TS2/16) affects cytoskeletal changes and the onset of high-speed migration in response to DC maturation. Stimulation with TS2/16 effectively interfered with PGE2- induced podosome dissolution (Fig 7A) as well as the induction of rapid migration after prolonged incubation (Fig 7B). These results demonstrate that loss of α5β1-integrin activity during DC maturation is key to the induction of high-speed migration.

Figure 7: Podosomes dissolution and migration can be inhibited by activating α5β1-integrin. α β (A/B) Podosome dissolution in response to PGE2 can be blocked by activation of 5 1- integrin. Immature DCs (control) were seeded on fibronectin-coated coverslips and left β untreated or stimulated 5 minutes or 16 h with PGE2, PGE2 with the 1-integrin activating Ab (TS2/16) or with TS2/16 alone. Subsequently, the cells were stained with phalloidin-Texas Red to stain F-actin. The anti-vinculin Ab could not be used since it has the same isotype as the TS2/16 Ab used to activate β1-integrins. (A) Representative µ images of unstimulated DCs or cells stimulated for 16 h with 10 g/ml PGE2 in the presence or absence of TS2/16 Ab are shown. (B) The number of cells containing podosomes was counted in 7 images per condition. The percentage of cells expressing podosomes ±SEM is depicted. A representative experiment is shown. Conditions that are significantly different from the control situation (p<0.05) are indicated with an aster- α β isk. (C) PGE2-induced migration is inhibited by activating 5 1-integrin. Immature DCs (control) or DCs matured with PGE2 with or without TS2/16 were added to fibronectin- coated plates and migration was monitored for 60 min. A representative experiment is shown. The percentage of migrating cells ±SD is depicted. Significant differences (p<0.05) are indicated with an asterisk.

49 Chapter 3

LPS induced podosome dissolution and high-speed migration are dependent on the production of prostaglandins The disassembly of podosomes in response to LPS occurs with a lag phase of 16 h (Figure 4B) and therefore cannot be a direct consequence of TLR activation. We investigated whether podosome dis-

solution induced by LPS involves production of PGE2 by the DCs. Therefore, iDCs were seeded on fibronectin and stimulated with LPS for 16 h, either in the presence or absence of indomethacin. This

drug inhibits the production of PGE2 and other PGs by inhibiting cyclooxygenases, enzymes involved in the production of PGs [24]. Indeed, an effective inhibition of LPS-induced podosome dissolution was observed in the presence of indomethacin (Figure 8A and 8B).

Figure 8: LPS induced podosome disassembly and migration are mediated by prostaglandins. (A/B) LPS mediated podosome dissolution is PG mediated. Immature DCs were seeded on fibronectin-coated coverslips and left untreated or stimulated for 16 h with LPS in the presence or absence of indomethacin. Subsequently, cells were stained with an anti-vinculin Ab (green) and phalloidin-Texas Red to stain F-actin (red). (A) Representative images of DCs either unstimulated (control) or stimulated for 16 h with 2 µg/ml LPS in the presence or absence of 50 µM indomethacin. (B) Immature DCs were left unstimulated (control) or stimulated with 2 µg/ml LPS with or without indomethacin (10 µM) or with only indomethacin. The number of cells containing podosomes was counted in 7 images per condition per experiment. The percentage of cells expressing podosomes ±SEM is depicted. Significant differences (p<0.05) are indicated by an asterisk. An average of 3 experiments is shown. (C) Migration of LPS- matured DCs is blocked by indomethacin. DCs were matured with 2 µg/ml LPS with or without indomethacin (50 µM). During maturation indomethacin was refreshed every 12 h. Immature (control) and mature DCs were added to fibronectin-coated plates and migration with or without indomethacin was monitored for 60 min. The percentage of migrating cells ±SD is depicted. Significant differences (p<0.05) are indicated by an asterisk.

50 PGE2 in DC maturation

In the absence of LPS, no effect of indomethacin on podosomes was seen. Similarly we observed that indomethacin affects the migratory capacity of LPS-matured DCs (Figure 8C). These results identify

PGs, such as PGE2, as important mediators of maturation-induced DC migration.

PGE2 and LPS induce podosome dissolution in myeloid DCs To examine whether our findings also apply to DCs generated ex vivo, human myeloid DCs were isolated from peripheral blood by magnetic sorting of CD1c-positive/CD19-negative cells (Figure 9A). These myeloid DCs, although much smaller than the monocyte- derived DCs, similarly formed podosomes on fibronectin, which rapidly dissolved in response to PGE2 stimulation (Figure 9B and 9C). Furthermore, PGE2-induced podosome dissolution was efficiently

Figure 9: Podosome dissolution in myeloid DCs. (A) Myeloid DCs (CD1c/BDCA positive cells) were isolated from PBMCs. The obtained myeloid DCs were analyzed by flow cytometry. These myeloid DCs were 99% pure (CD19-CD1c+), express high levels of CD11c and a subset of these myeloid DCs express CD14. (B) Myeloid DCs were seeded on fibronectin-coated coverslips and stimulated with PGE2 for 5 min. Subsequently, the cells were stained with an anti-vinculin Ab (green) and phalloidin-Texas Red to stain F-actin (red). Representative images are depicted. (C) Myeloid DCs were seeded on fibronectin-coated coverslips and stimulated with PGE2 for 5 min in the presence or absence of TS2/16 Ab. Subsequently, the cells were stained with phalloidin-Texas Red to stain F-actin (red). The number of cells containing podosomes was counted in 15 images per condition. The percentage of cells expressing podosomes ±SEM is depicted. Conditions that are significantly different (p<0.05) are indicated with an asterisk. (D) Myeloid DCs were seeded on fibronectin- coated coverslips and stimulated with LPS in the presence or absence of indomethacin. Subsequently, the cells were stained with an anti-vinculin Ab (green) and phalloidin- Texas Red to stain F-actin (red). The number of cells containing podosomes was counted in 7 images per condition. The percentage of cells expressing podosomes ±SEM is depicted. Conditions that are significantly different (p<0.05) are indicated with an asterisk. One representative experiment is shown.

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blocked by stimulation with the β1-integrin activating Ab TS2/16

(Figure 9C), while stimulation with TS2/16 without PGE2 had no effect on podosomes. We also investigated the effect of LPS on podosome dissolution in these myeloid DCs. Similar to monocyte- derived DCs, no immediate effects of LPS on podosome dissolution were observed, wereas a complete loss of podosomes was seen after 16 h stimulation with LPS (Figure 9D). Again, LPS-induced podosome dissolution was effectively inhibited in the presence of indomethacin (Fig 9D). These results indicate that both in monocyte- derived DCs as well as myeloid DCs, the effects of LPS on podosome dissolution are dependent on the production of PGs by the DCs.

Movies can be found at http://www.ncmls.eu/NCMLS/Visual /movie.asp Discussion In this paper we report how maturation-induced podosome dissolution and loss of α5β1-integrin activity allow DCs to switch from a strongly adhesive to a highly migratory phenotype. Although a sub- stantial amount of reports describe the biogenesis of podosomes in different cell types [17, 25] mechanisms responsible for podosome disassembly have remained elusive. Consistent with earlier findings in osteoclasts [26], we show that these structures are very dynamic. Although podosomes are generally considered to be features of migratory cells [25, 27], we find a strong inverse correlation between the presence of podosomes and DC migratory capacity. This apparent discrepancy can be easily explained by the fundamental differences in the way immature and mature DCs interact with the substrate to produce migration. While iDCs display very strong membrane protrusive activity, as indicated by the presence of lamellipodia and membrane ruffles, strong interactions with the substrate (i.e. podosomes) only allow low-speed migration (0.5 µm/min). Mature DCs display short-lived, weak interactions with the substrate resulting in high-speed migration, similar to the migration of T cells [9]. Consistent with the notion that podosomes restrict high-speed migration, we observed that the active form of the α5β1-integrin is enriched in these structures. Moreover, DC maturation is accompanied by a nearly complete loss of α5β1-integrin mediated adhesion as well as dissolution of podosomes. As a consequence, mDCs show migration speeds that are 10-fold higher (5 µm/min) than those observed in iDCs. We also show that loss of fibronectin binding, as it occurs during DC maturation, is due to loss of α5β1-integrin activity rather than

52 PGE2 in DC maturation expression. Furthermore, activation of the α5β1-integrin, using an activating Ab, prevents podosome disassembly and restricts high- speed migration in mDCs. Although preinbucation with this integrin- activating Ab effectively inhibits podosome disassembly, the Ab failed to induce new podosomes in fully mature DCs (data not shown). This suggests that, although down-regulation of integrin activity during DC maturation is important for the induction of high-speed migration, α5 β1-integrin activation cannot fully reverse the effects of DC maturation on the cytoskeleton. While there are many examples where integrin activation of leukocytes triggers the induction of cell migration [28-30], we show here that down-regulation of integrin activity may be equally important. Similarly, the importance of integrin de-activation in the induction of cell migration was recently demonstrated for αLβ2-integrin (LFA-1). Mice expressing a constitutively active variant of LFA-1 showed defective cell migration and compromised immune function due to impaired de-adhesion from its ligand ICAM-1 [31]. α A maturation cocktail, consisting of MCM, PGE2 and TNF , was used to induce DC maturation. This combination effectively induces all aspects of DC maturation and is used in clinical DC-vaccination studies [22, 32]. Here we show that PGE2 is an essential component in the induction of full migratory capacity by this cytokine cocktail. DCs matured in the presence of PGE2 not only migrate more efficiently than without, we also observed that PGE2 alone is sufficient to induce DC migration. The importance of PGE2-mediated signaling in DC migration and consequently the induction of a proper immune response is also supported by mouse models. Mice deficient in EP4, a PGE2 receptor, show impaired skin immune responses due to impaired migration and maturation of Langerhans cells [33].

Moreover it was shown in human DCs that PGE2 enhances both the expression and the sensitivity of CCR7 [34-36], the receptor for the chemokines CCL19 and CCL21 that controls DC migration into lym- phatic vessels and lymph nodes [37, 38]. Consistent with these findings, preliminary results in two melanoma patients participating in a DC-vaccination trial suggest that the percentage DCs that migrate to the lymph nodes is higher when DCs are matured in the presence of

PGE2 (data not shown). These findings emphasize the importance of PGE2 in promoting efficient DC maturation and migration, a prerequisite for the induction of an efficient immune response in vivo.

The effect of PGE2 on podosome dissolution is immediate, as well as the partial effect of TNFα. This partial response may be due to a lack of TNFα receptor expression in part of the DCs. In contrast, the effects of LPS are only observed after a long lag time. Importantly, LPS-induced podosome disassembly as well as the induction of

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high-speed migration can be efficiently inhibited by blocking the production of PGs by the DCs. Furthermore, in response to LPS stimu-

lation DCs start to produce PGE2 after 6 h and this production is maximal after 24 h [39, 40], which coincides with the loss of podosomes and the induction of migration. These results demonstrate that pathogen-derived molecules, such as LPS, induce podosome dissolution and high-speed migration by stimulating the production of

PGE2 by DCs during an inflammatory response. Ex vivo myeloid DCs form podosomes on fibronectin similar to monocyte-derived DCs. In addition, myeloid DCs also dissolve their

podosomes rapidly in response PGE2. Furthermore, this PGE2- induced podosome dissolution can be inhibited by activating β1-integrins, indicating that also in myeloid DCs regulation of the activity of β1-integrins is important in adhesion and migration. Moreover, also in myeloid DCs the LPS-induced podosome dissolution is depending on PG production by the DCs. These findings indicate that ex vivo DCs behave in the same way as the monocyte-derived DCs and fur-

ther emphasize the importance of PGE2 for the proper function of DCs. What could be the function of podosomes in iDCs? Podosomes are found in cells that have the capacity to cross tissue boundaries. The observation that podosomes are sites of active matrix degradation [41, 42] and that matrix metalloproteinases are needed for transendothelial migration of leukocytes [43], supports the idea that podosomes are required for crossing tissue barriers. In osteoclasts podosomes are involved in bone resorption and remodeling [44], while in macrophages podosomes are considered regulators of cell adhesion and migration [45]. In addition to a role during transmigration, podosomes in iDCs may contribute to the movement of iDCs within peripheral tissues, while patrolling for antigens. Disassembly of podosomes observed in DCs upon stimulation with maturation inducing agents is probably unique for DCs, and an inte- gral part of the DC maturation process. In mouse DCs, West et al have described increased antigen macropinocytosis leading to enhanced antigen presentation on MHC class I and II molecules and a simultaneous disappearance of podosomes in response to LPS [6]. The observed disappearance of podosomes was rapid and transient, while we find that podosome dissolution in response to LPS is only observed after prolonged stimulation. A transient disappearance of podosomes could have gone unnoticed in our system, however after 5, 10, 15 min or 1 h stimulation with LPS no effect on podosomes was observed. Therefore, it is likely that this difference is species related similar to other processes [46]. Podosome dissolution during

54 PGE2 in DC maturation

DC maturation may serve to redeploy actin and actin-binding or regulatory proteins to other processes in the cell that require high actin turnover. Podosome biogenesis has been linked to lysosome exocy- tosis and late endosomal/lysosomal proteins (p61Hck, CD63 and LYAAT-1) localize to podosomes [47-49]. Therefore it has been put forward that late endosomal vesicles fusing with the plasma membrane contribute to podosome formation [49]. During podosome dissolution the opposite process could be taking place in order to fuel increased vesicle transport as it occurs during DC maturation.

How does PGE2 induce podosome disassembly? The effects of PGE2 on podosomes occur within minutes, which implies that these are a direct consequence of PGE2 receptor activation rather than changes in gene expression. Spatial and temporal regulation of actomyosin contractility was shown to be important for the assembly/disassembly of focal adhesions and podosomes [25, 50]. Two receptors for PGE2 (EP2/4) are expressed on DCs affecting DC migration [33, α 34, 51]. Both these receptors couple to G s to increase cAMP levels, leading to activation of protein kinase A [52]. Changes in activity regulate actomyosin contractility, either by affecting the activity of the small GTPase RhoA, a key regulator of myosin II function, or by inactivation of myosin light chain kinase (reviewed by Burridge et al and references therein [53]). Future experiments will therefore be aimed at determining how these signal transduction pathways contribute to DC adhesion and migration. Materials and methods Chemicals and antibodies The following antibodies were used: anti-HLA class I (W6/32), anti- HLA-DR/DP (Q5/13), anti-CD80 (all from Becton Dickinson & Co., Franklin Lakes, NJ), anti-CD14 and anti-CD83 (both Beckman Coulter, Mijdrecht, The Netherlands), anti-CD86 (Pharmingen, San Diego, CA), anti-CD209/DC-SIGN (AZN-D1) [54], anti-β1-integrin activating epitope (TS2/16) [55], anti-β1-integrin blocking epitope (AIIB2) (Developmental Studies Hybridoma Bank, Iowa City, IA), anti- β1-integrin recognizing an active conformation (12G10) [23] (kind gift of M. Humphries, Manchester, UK), anti-α5-integrin (SAM-1), anti- vinculin (Sigma, St. Louis, MO), mIgG1 (Becton Dickinson & Co.), mIgG2a, mIgG2b, rIgG1 isotype (Pharmingen). Alexa Fluor 488- labeled secondary antibodies were from Molecular Probes (Molecular Probes, Leiden, The Netherlands) and FITC-labeled secondary antibodies from Zymed (Zymed, San Francisco, CA) and Cappel (Cappel, MP Biomedicals, Irvine, CA). Texas Red-conjugated

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phalloidin (Molecular Probes) was used to stain F-actin. Indomethacin was obtained from Sigma (Sigma).

Preparation of DCs DCs were generated from PBMCs as described previously [22, 56]. Monocytes were derived from buffy coats or from a leukapheresis product. Plastic-adherent monocytes were cultured in X-VIVO 15 medium (BioWhittaker, Walkersville, MD) supplemented with 2% pooled human serum (PAA laboratories, Linz, Austria), IL-4 (500 U/ml) and GM-CSF (800 U/ml) (both Schering-plough, International, Kenilworth, NJ) [57] or in RPMI 1640 medium (Gibco; Life Technologies, Breda, The Netherlands) supplemented with 10% (v/v) FCS (Greiner, Kremsmuenster, Austria), IL-4 (500 U/ml) and GM- CSF (800 U/ml) [18]. Immature DCs were harvested on day 7. DCs were matured with LPS (2 µg/ml) (Sigma), rTNFα (75 ng/ml) (kind gift of Dr. Adolf, Bender, Vienna, Austria) or a combination of monocyte- α conditioned medium (MCM) (50% v/v), rTNF (10 ng/ml) and PGE2 (10 µg/ml) (Pharmacia & Upjohn, Puurs, Belgium), for 2 days. Mature DCs were harvested on day 9. Expression of MHC class I/II, costimulatory molecules and DC-specific markers on DCs were measured by flow cytometry (data not shown) and the expression of MHC molecules, costimulatory molecules and DC-markers was similar to what was described before [3]. Immature DCs expressed MHC class I and II, the costimulatory molecule CD86, the DC-specific marker CD209/DC-SIGN, low levels of the costimulatory molecule CD80 and lacked expression of the maturation marker CD83. Maturation resulted in the induction of CD83, up-regulation of MHC class I and II, CD86 and CD80 and a down modulation of CD209/DC-SIGN expression.

Isolation of myeloid DCs Myeloid DCs (CD19-CD1c+) were isolated from PBMCs derived from buffy coats. PBMCs were labeled with a FITC conjugated mAb to CD3 (Dako, Glostrup, Denmark) and T cells and B cells were deplet- ed with anti-FITC and anti-CD19 beads (both Miltenyi Biotec, Bergisch Gladbach, Germany). Myeloid DCs were then isolated from the CD3-CD19- cells using a biotinylated anti-CD1c (BDCA-1) Ab and anti-biotin beads (both Miltenyi Biotec) following the manufacturer's instructions. To assess purity, isolated cells were labeled with streptavidinCy5 (Jackson Immunoresearch, Cambridgeshire, UK) and a PE-conjugated anti-CD19Ab (Miltenyi Biotec), PE-conjugated anti-CD14 (Beckman Coulter) or anti-CD11c (Becton Dickinson & Co.) Abs. Myeloid DCs were identified as cells that expressed CD1c

56 PGE2 in DC maturation and CD11c but were negative for CD19. Myeloid were up to 99% pure and 0.3% of the PBMCs were myeloid DCs.

Flow cytometry Cells (1 x 105) were incubated with 2% (v/v) human serum in PBA

(PBS containing 1% (w/v) BSA and 0.05% (w/v) NaN3) for 15 min at 4°C. After washing with cold PBA the cells were incubated with primary Ab (5 µg/ml) for 30 min at 4°C. Subsequently, the cells were incubated with FITC-labeled goat anti-mouse. Cells were washed and resuspended in 100 µl PBA. Fluorescence was measured using a FACSCalibur™ with CellQuest software (Becton Dickinson & Co.).

Fluorescence microscopy Coverslips were coated with fibronectin (20 µg/ml) (Roche, Mannheim, Germany) in PBS for 1 hour at 37°C. Cells were seeded on fibronectin-coated coverslips, left to adhere for 1 to 12 h and fixed in 3.7% (w/v) formaldehyde in PBS for 10 min. Cells were permeabilized in 0.1% (v/v) Triton X-100 in PBS for 5 min and blocked with 2% (w/v) BSA in PBS. The cells were incubated with primary Ab for 1 h. Subsequently the cells were incubated with Alexa Fluor 488-labeled secondary antibodies for 45 min. Subsequently, cells were incubated with Texas Red-conjugated Phalloidin for 30 min. Images were collected on a Leica DMRA fluorescence microscope with a 63x PL APO 1.3 NA oil immersion lens (or a 40x PL FLUOTAR 1.0 NA oil immersion lens for overview images) and a COHU high performance integrating CCD camera (COHU, San Diego, CA). Pictures were analyzed with Leica Qfluoro version V1.2.0 (Leica) and Adobe Photoshop 7.0 (Adobe Systems, Mountain View, CA) software.

Migration assay 96-wells flat bottom plates (Costar, Corning, Acton, MA) were coated with fibronectin, washed and blocked with 0.01% (w/v) gelatin (Sigma) in PBS for 30 min at 37°C. Per well, 3 x 103 cells were added and mineral oil was pipetted on top of the medium to prevent pH changes and evaporation of the medium. The cells were monitored with a previously described migration system [58]. Cells were recorded for 1 h followed by analysis of individual cells. The speed was defined as traversed path during the entire experiment divided by the imaging time.

Adhesion assay 96-wells flat bottom plates (Costar) were coated with fibronectin and blocked with gelatin. Cells (7x 106/ml) were labeled with Calcein-AM

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(25 µg/ml) in PBS (Molecular Probes) for 30 min at 37°C, either untreated or preincubated for 10 min at RT with blocking Ab (10 µg/ml), and seeded on the fibronectin-coated plates (3 x 104/well) for 45 min at 37°C in the presence or absence of activating Ab (1 µg/ml). Non-adherent cells were removed by washing with 0.5% (w/v) BSA

in TSM (150 mM NaCl, 10 mM Tris/HCl, 2mM MgCl2, 1 mM CaCl2, pH 8.0) at 37°C. Adherent cells were lysed in 100 µl lysisbuffer (50 mM Tris, 0.1% (w/v) SDS) and fluorescence was quantified using a cytofluometer (PerSeptive Biosystems, Framingham, MA). Results are expressed as the mean percentage of adhesion of triplicate wells.

Transfection and live imaging of DCs DCs (3 x 106) were electroporated with pCMV-β-actinGFP on day 5 using Amaxa nucleofection (Amaxa biosystems, Cologne, Germany). Transfected DCs were seeded on fibronectin-coated Willco glass bottom dishes (Willco Wells BV, Amsterdam, The Netherlands). Cells were imaged at 37°C in RPMI 1640 without phenol red (Gibco) supplemented with 10% (v/v) FCS using a Zeiss LSM 510-meta micro-

scope equipped with a type S heated stage CO2 controller and PlanApochromatic 63x 1.4 NA oil immersion DIC lens (Carl Zeiss GmbH, Jena, Germany). Podosomes were imaged for 20 min with

20-second intervals without PGE2 and for 17 min with 10-second µ intervals with PGE2 (10 g/ml). In addition, podosomes were imaged for 60 min without TNFα, and 20 min in the presence of TNFα (75 ng/ml) at 30-second intervals. Cells were imaged using Zeiss LSM Image Browser version 32 (Carl Zeiss) and images were processed with Image J version 1.32j software (National Institutes of Health, http://rsb.info.nih.gov/ij).

Statistical analysis An ANOVA test or a two-tailed student-t test was used for statistical analysis. Acknowledgements We thank M. Humphries for providing the 12G10 Ab. We thank F. de Lange from the Microscopic Imaging Center of the NCMLS and B. Joosten for assistance with microscopic imaging. We thank I. de Vries, N. Meeuwsen-Scharenborg, A. de Boer, M. Brouwer and M. van de Rakt for providing DCs. This study was supported by research

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62 Chapter 4 TLR4-mediated discrimination between Gram-negative and Gram-positive bacteria directs migration and maturation of DCs

1Suzanne F.G. van Helden, 1Machteld M. Oud, 2Reinier A.P. Raymakers, 3Mihai G. Netea, 4Frank N. van Leeuwen and 1Carl G. Figdor

1Department of Tumor Immunology, NCMLS, Radboud University Nijmegen Medical Centre, Nijmegen, The Netherlands 2Department of Hematology, NCMLS, Radboud University Nijmegen Medical Centre, Nijmegen, The Netherlands 3Department of Internal Medicine, Radboud University Nijmegen Medical Centre, Nijmegen, The Netherlands 4Laboratory of Pediatric Oncology, NCMLS, Radboud University Nijmegen Medical Centre, Nijmegen, The Netherlands

Submitted Chapter 4

Abstract Recognition of pathogenic bacteria, mediated by TLRs expressed on antigen presenting cells, represents a critical step in the induction of adaptive immune responses against such pathogens. Investigating the effects of different pathogens on dendritic cell (DC) maturation, we found that Gram-negative bacteria are much more potent inducers high-speed migration, cytokine secretion and expression of CCR7 and costimulatory molecules than Gram positive bacteria. In addition, only Gram-negative bacteria, are capable of inducing podosome dissolution in DCs. Similar to whole bacteria, purified ligands for TLR4, but not TLR2, induce podosome loss. The effects of Gram-negative bacteria on DC maturation and podosome dissolution are mainly dependent on the adaptor protein TRIF and involve the production of prostaglandins by the DCs. We conclude that DCs differ in their capacity to respond to these different classes of bacteria and this could account for the distinct pathology of these bacteria. Introduction Pathogenic bacteria are an important cause of morbidity and mortal- ity worldwide. The two classes, Gram-negative and Gram-positive bacteria, preferentially localize to distinct anatomical sites in the body. In the lower gastrointestinal tract mostly Gram-negative bacteria are found, while the upper gastrointestinal tract and the respiratory airways are mainly colonized with Gram-positive bacteria. The different composition and structure of the cell wall of Gram-negative and Gram-positive bacteria results in recognition by different pathogen recognition receptors (PRRs). TLRs are a family of PRRs, which at least 11 members [1], recognizing very different pathogen associated molecular patterns (PAMPs) [2], leading to activation of nuclear factor κB (NF-κB)-dependent and interferon (IFN)-regulatory factor (IRF)-dependent signaling pathways [3]. Gram-negative bacteria are mainly recognized by TLR4 through their lipopolysaccharide (LPS), whereas structural components of Gram-positive bacteria, such as peptidoglycans, lipoteichoic acid (LTA) and lipoproteins activate TLR2 [4-6], which forms heterodimers with TLR1 or TLR6. TLRs can modulate the function of each other and the cross talk between PRRs can modify the resulting immune responses [3]. Dendritic cells (DCs) are the most potent antigen presenting cells of the immune system [7] and they express different PRRs, such as C- type lectins, NOD-like receptors and Toll-like receptors (TLRs) [8-10]. Stimulation of PRRs, especially TLRs, on DCs bridges innate to adaptive immune responses by inducing signaling that will induce cytokine production and DC-maturation and thereby induce efficient

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T cell stimulation by mDCs. Most studies on the effects of PRR triggering in DCs have been performed using purified or synthetic ligands, while the effects of whole pathogens are largely unknown. As DCs come in contact with Gram-negative or Gram-positive bacteria, depending on the anatomical site, we investigated if and how Gram- negative and Gram-positive bacteria differ in their ability to stimulate DCs. By comparing Gram-positive with Gram-negative bacteria, we find that the latter are far more efficient inducers of DC-maturation. Previously we showed that podosomes, specialized adhesion structures found in DCs, dissolve in response to LPS [11]. Here we find that only Gram-negative bacteria are capable of inducing podosome dissolution and high speed migration in DCs. These effects are dependent on TLR4, TRIF, MyD88 and the production of prostaglandins by the DCs. We conclude that the preferred ability of Gram-negative bacteria to stimulate DCs is determined by TLR4 ligands expressed on the surface of these pathogens.

Results Gram-negative bacteria are superior to Gram-positive bacteria in inducing DC-maturation. We investigated the effects of the Gram-negative bacteria Neisseria meningitidis and Salmonella enteritidis, as well as the Gram-positive bacteria Staphylococcus aureus and Streptococcus pneumoniae, on the induction of cytokine secretion and expression of CD80, CD83 and CCR7. LPS or Gram-negative bacteria induced high levels of TNFα, IL-6 and IL-8 by the DCs, while exposure to Gram-positive bacteria resulted in much lower levels of these pro-inflammatory cytokines (Figure 1A). In addition, the expression of the costimulatory molecules, CD80 and CD83, and CCR7, a chemokine receptor required for migration to the lymph nodes, was investigated. Gram- negative bacteria were capable of inducing upregulation of CD80, CD83 as well as CCR7 to the same level (or even higher) as LPS (Figure 1B). However, the up-regulation of these markers in response to Gram-positive bacteria was lower (Figure 1B). The differential effects of these pathogens on CCR7 expression prompted us to also explore effects of pathogen-stimulation on migratory behavior. We observed that whereas only 20-40% of the unstimulated iDCs or DCs stimulated with Gram-positive bacteria showed cell migration in vitro, exposure to Gram-negative bacteria induced high- speed migration in about 70% of the DCs, levels comparable to those obtained in response to LPS (Figure 1C). Taken together, these data

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Figure 1. Gram-negative bacteria are superior to Gram-positive bacteria in inducing DC- maturation. iDCs were left untreated or incubated with LPS, N. meningitidis (meningo), S. enteritidis (salmon), S. aureus (staphyl) or S. pneumoniae (strept) for 42-48h. (A) Supernatants were harvested from the stimulated DCs and the levels of human TNFα, IL-6 and IL-8 were measured with an inflammatory cytokine beads array. An average of 5 experiments (±SEM) is depicted. (B) The expression of the costimulatory molecules CD80 and CD83, and the chemokine receptor CCR7 was analyzed by flow cytometry. The average mean fluorescence intensity (MFI) (±SEM) of 4 experiments is depicted. (C) Random migration on fibronectin-coated plates was monitored for 60 min. The percentage of migrating cells (±SEM) is depicted.

show that Gram-negative bacteria are superior to Gram-positive bacteria in inducing DC-maturation and migration.

Gram-negative, not Gram-positive, bacteria induce podosome loss in DCs. Previously, we showed that podosome dissolution in response to maturation stimuli is required for the induction of high-speed migration [11]. Therefore, we tested whether the differential effects of Gram-negative and Gram-positive bacteria on cytokine secretion, expression of CD80, CD83 and CCR7 and induction of high-speed migration are reflected by the ability to induce podosome dissolution in DCs. Indeed, whereas Gram-negative bacteria induced podosome dissolution, Gram-positive bacteria produced no significant effect on the number of cells displaying podosomes (Figure 2A, B). The reduced ability Gram-positive pathogens to stimulate DCs is not

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Figure 2. Gram-negative bacteria induce podosome loss, while Gram-positive bacteria do not. (A/B) iDCs were left untreated or stimulated with LPS, N. meningitidis (meningo), S. enteritidis (salmon), S. aureus (staphyl) or S. pneumoniae (strept) at moi 20 for 16h. (A) Representative images are depicted. Scale bar indicates 20 µm. (B) The average number of cells displaying podosomes of 4 experiments is shown. (C/D) iDCs were incubated for 45-60 min with FITC-labeled bacteria. (C) The amount of DCs that had bound (or internalized) bacteria was analyzed by flow cytometry. A representative experiment is shown. (D) Internalization was confirmed by making z-stacks using a Zeiss LSM 510- meta microscope. Representative images are depicted. Scale bar indicates 10 µm. due to a difference in binding or internalization. Both Gram-negative and Gram-positive bacteria showed similar binding to DCs, as determined by flow cytometry using FITC-labeled bacteria (Figure 2C). In addition, using confocal microscopy we generated Z stacks to demonstrate that Gram-negative as well as Gram-positive bacteria are efficiently internalized by the DCs (Figure 2D). We conclude therefore that the reduced ability of Gram positive bacteria to stimulate DC maturation is not caused by a lack of binding or internalization by the DCs.

Pure TLR ligands mimic the effect of whole bacteria. Gram-negative bacteria are mainly recognized by TLR4, while Gram- positive bacteria predominantly activate TLR2. Therefore, we investigated whether activating these receptors with purified ligands mimics the effect of whole bacteria. Indeed, LPS, a TLR4 ligand, was able

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Figure 3. TLR4, but not TLR2 activation, induces podosome dissolution in iDCs. (A/B) iDCs were left untreated or stimulated with LPS, Pam3cys, FSL-1 or a combination of Pam3cys and FSL-1 for 16h. (A) Representative images are depicted. Scale bar indicates 20 µm. (B) The average number of cells displaying podosomes of 4 experiments is shown. (C) Supernatants were harvested from unstimulated, LPS, Pam3cys or FSL-1 stimulated DCs and the levels of human TNF and IL-6 were measured. An average of 8 experiments (±SEM) is depicted. (D) iDCs were left untreated or stimulated with different concentrations of LPS for 16h. The average number of cells displaying podosomes of 3 experiments is shown. (E) iDCs were left untreated or stimulated with LPS or a suboptimal concentration of LPS (6 ng/ml) with or without Pam3cys or FSL-1 for 16h. The average number of cells displaying podosomes of 3 experiments is shown.

to induce podosome loss, while the TLR2/1 ligand Pam3cys and the TLR2/6 ligand FSL-1, either alone or in combination, did not affect podosomes (Figure 3A, B). DCs were not completely unresponsive to these TRL2 ligands, as stimulation with Pam3cys or FSL-1 induced the production of TNF and IL-6, albeit at lower levels in comparison to LPS (Figure 3C). A potential synergy between de TLR4 and 2 ligands was investigated by combining a suboptimal concentration of LPS (6 ng/ml), derived from LPS serial dilutions (Figure 3D), with TLR2 ligands. However, no synergy between LPS and TLR2 ligands was observed (Figure 3E). These data demonstrate that purified TLR4 and TLR2 ligands differ in their capacity to induce podosome dissolution, which mirrors the effects observed with whole pathogens that express either of these ligands.

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Podosome loss induced by Gram negative bacteria is TLR4 dependent. Intracellular Gram-negative bacteria, Bartonella quintana and Chlamydia pneumoniae, which carry a structurally different LPS that is incapable of triggering TLR4 signaling [12-14], were also tested. Stimulation of iDCs with B. quintana or C. pneumoniae had no effect on the amount of podosomes (Figure 4A), consistent with the notion that Gram-negative bacteria induce podosome loss by triggering TLR4 signaling through LPS exposed on the cell surface of these bacteria. Moreover, addition of the TLR4 antagonist Bartonella quintana LPS [14] completely blocked the LPS- or Gram-negative bacteria-induced podosome loss (Figure 4B), showing that the response is indeed TLR4 mediated. In order to unequivocally demonstrate the importance of TLR4 stimulation in podosome-dissolution, we examined (bacterially-induced) podosome dissolution in mouse bone marrow-derived DCs (BDMCs) from either wild type or TLR deficient mice. Similar to human monocyte-derived DCs (MODCs), podosome dissolution was observed in normal BDMCs in response to LPS and Gram-negative bacteria but

Figure 4. LPS- and Gram-negative bacteria-induced podosome dissolution is mediated by TLR4. (A) iDCs were left untreated or stimulated with LPS, B. quintana or C. pneumoniae at moi 20 for 16h. The average number of cells displaying podosomes of 3 experiments is shown. (B) iDCs were left untreated or stimulated with LPS with or without the TLR4 antagonist (TLR4a) B. quintana LPS or the TLR4 antagonist alone for 16h. The average number of cells displaying podosomes of 4 experiments is shown. (C/D) Immature BMDCs from wild-type (WT), TLR2-/-, TLR4-/- mice were left untreated or stimulated with LPS, Pam3cys, FSL-1, N. meningitidis (menigo), S. enteritidis (salmon), S. aureus (staphyl) or S. pneumoniae (strept) for 16h. (C) Representative images are depicted. Scale bar indicates 20 m. (D) The average number of cells displaying podosomes of 4 experiments is shown.

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not in response to Pam3cys, FSL-1 or Gram-positive bacteria (Figure 4C, D). Moreover, we observed that the LPS- and Gram-negative bacteria-induced responses were unaffected in TLR2-/- BMDCs, whereas these were completely abrogated in TLR4-/- BMDCs (Figure 4C, D). At the same time, these TLR4-/- BMDCs showed

rapid podosome dissolution in response to PGE2 stimulation, similar to MODCs [11], WT and TLR2-/- BMDCs (data not shown), showing that signaling downstream of TLR4 is unaffected in these knockout cells. Together, these data show that both LPS- and Gram-negative bacteria-induced podosome dissolution is completely dependent on TLR4 signaling.

TLR4-mediated podosome loss is mainly TRIF dependent. To date, two major signaling routes downstream of TLR4 have been described. The first depends on the adaptor protein myeloid differentiation primary response gene 88 (MyD88), whereas the second depends on the adaptor protein TIR domain-containing adaptor- inducing IFNβ (TRIF) [15]. To investigate the involvement of these signaling routes, we made use of MyD88-/- mice and Lps2 mice, which carry a natural mutation resulting in TRIF deficiency [16]. BMDCs from these mice were able to form podosomes and showed

podosome dissolution in response to PGE2, while TLR2 ligands and Gram-positive bacteria had no effect (Figure 5A, B and data not shown). However, the LPS-, S. enteritidis- as well as the N. meningitidis-induced responses were partly abrogated in Lps2 BMDCs (Figure 5A, B). In MyD88-/- BMDCs, LPS and S. enteritidis-induced podosome loss, although not significant, seemed somewhat impaired (Figure 5A, B). In addition, the N. meningitidis-induced

Figure 5. LPS- and Gram-negative bacteria-induced podosome loss in mediated by TRIF and MyD88. (A/B) Immature BMDCs from wild-type (WT), lps2 (TRIF-/-) and MyD88-/- mice were left untreated or stimulated with LPS, Pam3cys, FSL-1, N. meningitidis (menigo), S. enteritidis (salmon), S. aureus (staphyl) or S. pneumoniae (strept) for 16h. (A) Representative images are depicted. Scale bar indicates 20 µm. (B) The average number of cells displaying podosomes of 4 experiments is shown.

70 Gram- vs Gram+ bacteria in DC maturation response was partly abrogated (Figure 5A, B). These findings suggest that these responses are mainly TRIF mediated and that TRIF and MyD88 dependent signaling might synergize in these responses.

Gram-negative bacteria-induced podosome loss is mediated by production of prostaglandins.

We have shown earlier that PGE2 can induce rapid podosome loss and that the LPS response is dependent on prostaglandin production by the DCs [11]. Therefore, we investigated whether podosome loss, induced by Gram-negative bacteria, also involved the production of prostaglandins. Indomethacin, a COX inhibitor and known to interfere with prostaglandin production [17], interfered with podosome loss induced by Gram-negative bacteria (Figure 6A), suggesting that this response is, at least in part, mediated by the production of prostaglandins by the DCs. To further investigate a role for prostaglandins, we tested whether tissue culture medium conditioned by DCs that had been stimulated with different TLR ligands or bacteria, was capable of inducing (rapid) podosome loss in other DCs. Medium obtained from unstimulated, Pam3cys- or S. aureus-stimulated DCs induced no podosome dissolution (Figure 6B). However, supernatant of DCs stimulated with

Figure 6. Gram-negative bacteria-induced podosome loss is mediated by production of prostaglandins by the DCs. (A) iDCs were left untreated or stimulated with LPS, N. meningitidis (meningo) or S. enteritidis (salmon) at moi 20 for 16h with or without indomethacin (indo) or indomethacin alone. The average number of cells displaying podosomes of 3 experiments is shown. (B) iDCs were left untreated or stimulated with LPS, Pam3cys, S. enteritidis (salmon) or S. aureus (staphyl) for 16h. Supernatants of these stimulated DCs were harvested and added for 30 min to DCs from a different donor. The average number of cells displaying podosomes of 4 experiments is shown. (C) iDCs were left untreated or stimulated with LPS, Pam3cys, S. enteritidis (salmon) with or without indomethacin (indo) or indomethacin alone for 16h. Supernatants of these stimulated DCs were harvested and added for 30 min to DCs from a different donor. The average number of cells displaying podosomes of 2 experiments is shown.

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PGE2 (data not shown), LPS or S. enteritidis induced podosome dissolution in at least part of the DCs (Figure 6B). Moreover, supernatants from DCs stimulated with LPS or S. enteritidis in the presence of indomethacin were unable to induce podosome loss in other

DCs (Figure 6C). Although the levels were low, we could detect PGE2 in the supernatants of DCs stimulated with LPS, but not when DCs were stimulated with Pam3cys or FSL-1 (data not shown). Together these findings support the notion that prostaglandins produced by the

DC, possibly PGE2, are needed for podosome turnover in response to TLR4 stimulation. Discussion Bacterial infections are an important cause of morbidity and mortali- ty worldwide. The interaction between colonizing microorganisms and the host has important effects on the normal development of the immune system or the intestinal tract. This fact is also mirrored by the specific microbial flora of anatomic segments, such as the abun- dance of Gram-positive bacteria as colonizers of the mouth, nose and the respiratory tract, and the colonization of the lower intestinal tract by Gram-negative bacteria. APCs are cells of the innate host defense system that ingest and present bacterial antigens, regulating the type of interaction of a colonizing or infecting microorganism with the immune system. The prototype APC is the DC, and in the present study we have investigated the interactions of specific microorganisms with these cells. Surprisingly, our data show a significant difference between the various bacterial classes, with Gram-negative bacteria being superior to Gram-positive bacteria in inducing DC maturation. Effective maturation of DCs is essential for the induction of adaptive immune responses. During maturation adhesive, antigen recognizing iDCs are transformed into highly migratory mDCs that efficiently stimulate T cells. DC maturation is accompanied by relocalization of MHC molecules to the plasma membrane, upregulation of costimulatory molecules as well as the production of cytokines. Gram-negative bacteria are much more potent in inducing the secretion of proinflammatory cytokines such as TNFα, IL-6 and IL-8, [18, 19], and upregulation of expression of the costimulatory molecules CD80 and CD83. In addition, DC maturation involves elaborate changes in cytoskeletal organization and integrin-mediated cell adhesion [11, 20], which allows mature DCs to migrate to and into the lymph nodes and stimulate T cells. Although Gram-positive and Gram-negative bacteria do not differ in

72 Gram- vs Gram+ bacteria in DC maturation their ability to be bound and internalized by DCs, we find that the latter are much more efficient in stimulating the cytoskeletal changes required for the induction of DC migration. For instance, only Gram- negative bacteria trigger podosome dissolution in DCs, and they more strongly induce upregulation of CCR7, a chemokine receptor directing DC migration to the lymph nodes [21]. Hence, we conclude, that Gram-negative and Gram-positive bacteria differ in their ability to regulate adhesive and migratory properties of DCs. Bacteria bind and activate different PRRs on DCs, for instance Gram-negative bacteria mostly signal through TLR4, while Gram- positive bacteria mainly activate TLR2. Although TLR2 ligands were shown to affect DCs [22, 23] and induce cytokine secretion, we and others [24] found that the cytokine secretion in response to intact Gram-positive bacteria was very low or absent. This discrepancy may be due to the relatively high concentration of pure TLR ligands (10 µg/ml) used, in comparison to the amounts of TLR ligands that can be presented to the DCs when complete bacteria are used. However, even with purified ligands, cytokine responses observed with purified TLR2 ligands were significantly lower when compared to LPS stimulation. Moreover, stimulation with LPS [11], but not Pam3cys or FSL-1, resulted in podosome loss in DCs, suggesting that irrespective of concentration or context, TLR4 is a much more effective inducer of DC maturation than TLR2. Our experiments using a TLR4 antagonist and the TLR4-/- BMDCs, show that the effects of Gram-negative bacteria are completely dependent on TLR4 signaling. Previously, it was shown that a mutant strain of N. meningitidis lacking LPS is less potent in inducing proinflammatory cytokines and largely incapable of inactivating DCs [25, 26]. We show that the effects of Gram-negative bacteria can be mimicked by LPS, whereas Gram-negative bacteria possessing an LPS unable to activate TLR4 [12-14] are unable to induce podosome loss in DCs. Together these results show that the effects of Gram-negative bacteria on DC maturation are largely TLR4-mediated and suggest that TLR4 antagonist might be used in clinical applications to treat Gram-negative bacteria induced infection and sepsis. TLR4 can activate MyD88 [27, 28] as well as TRIF dependent signaling [15, 29]. LPS- or Gram-negative-induced podosome loss is partly abrogated in lps-2 BMDCs, suggesting that the TRIF-dependent signaling pathway is involved in podosome dissolution in DCs. Previously, it was suggested that the effects of live of heat-killed Escherichia coli on DCs are mediated through TLR4 and TRIF [30]. This is in line with the finding that activation of TLR2, which only uses the MyD88 dependent pathway [31, 32], fails to induce podosome

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loss. The importance of the MyD88 dependent pathway is less straightforward. Although the LPS- or S. enteritidis-induced podosome loss appeared somewhat impaired in MyD88-/- BMDCs, this effect was not significant. However, the N. meningitidis-induced podosome loss was significantly impaired in MyD88-/- BMDCs. This could either mean that MyD88 is only involved when TLR4 is triggered by certain strains of Gram-negative bacteria or that there is some involvement of MyD88 in the responses of DCs to TLR4 triggering. The latter seems more likely since we have observed that triggering of TLR3, which only uses the TRIF dependent pathway, is unable to induce podosome loss in DCs (data not shown), suggesting that both pathways are involved. This suggests that both pathways may cooperate during DC-maturation. Until now, TLR4 is the only TLR known to signal both through the MyD88- and TRIF- dependent pathways, which may account for the fact that TLR4 ligands are the most potent inducers of DC maturation. The need for activation of both pathways could be an explanation for the synergy often observed between TLR-ligands signaling via MyD88 and TLR- ligands activating TRIF-dependent signaling [33, 34]. The ability of conditioned supernatants to induce podosome loss and the effects of COX inhibition on podosome dissolution, show that Gram-negative bacteria induced podosome loss is mediated by the production of prostaglandins, similarly to the LPS induced responses [11]. This could be mediated through PKC and p38MAPK signaling downstream of TLR4, which is known to induce COX enzymes [35].

PGE2, needed for the induction of high-speed migration during DC- maturation [36], is in itself very efficient in inducing podosome dissolution in DCs [11]. In addition, LPS, but not Pam3cys or FSL-1, stim-

ulation led to PGE2 production by DCs (data not shown), suggesting that Gram-negative bacteria may also trigger PGE2 secretion. It was shown already that Gram-negative bacteria and not Gram-positive

bacteria can induce production of PGE2 in monocytes [37]. Together, these findings emphasize the importance of prostaglandin production

by DCs and suggest that PGE2 might be the prostaglandin involved. In addition to the bacteria described in this study, the Gram-negative bacterium Klebsiella pneumoniae (ATCC 10031), the Gram-positive strain Mycobacterium BCG, and additional strains of S. pneumoniae, with and without capsules, give similar results (data not shown). The LPS used here is derived from E. coli, suggesting that this strain will induce the same responses. This indicates that our findings are likely to apply to a broad range of bacteria. The finding that Gram-negative bacteria induced enhanced DC maturation suggests that these bacteria might be more effective in the induction of T cell dependent

74 Gram- vs Gram+ bacteria in DC maturation responses. Indeed, the induction of adaptive immunity appears to be important in the prevention of diseases involving Gram-negative bacteria. Immunocompromised patients, especially in combination with mucositis, are at high risk to develop infections and Gram-negative sepsis [38]. These patients routinely receive antibiotics directed against Gram-negative bacteria (selective gut decontamination) [39]. Alternatively, it could mean that although DCs are predominantly activated by Gram-negative bacteria, Gram-positive bacteria activate other APCs, monocytes and macrophages. Although there it has been suggested that macrophages produce more cytokines in response to Gram-negative bacteria [24], we find that macrophages produce cytokines in response to Gram-negative as well as Gram- positive bacteria (data not shown). In addition, in vivo the activation of other APCs could also induce activation of DCs and thus induce efficient T cell responses. In conclusion, we find that Gram-negative and Gram-positive bacteria differ in their ability to activate DCs. The differential effects of Gram-negative and Gram-positive bacteria on DC maturation could account for the different pathology of these bacteria. Materials and methods Chemicals and antibodies The following antibodies were used: anti-hCD80 and mIgG1 (Becton Dickinson & Co., Franklin Lakes, NJ), anti-hCD83 (Beckman Coulter, Mijdrecht, The Netherlands), mIgG2a and mIgG2b (Pharmingen, San Diego, CA), anti-hCCR7 (R&D systems, Minneapolis, MN) and anti- vinculin (Sigma, St. Louis, MO). Alexa Fluor 488-labeled secondary antibodies were from Molecular Probes (Molecular Probes, Leiden, The Netherlands), FITC-labeled secondary antibodies from Zymed (Zymed, San Francisco, CA), and PE-labeled secondary antibodies from Beckman Coulter and Becton Dickinson & Co. Texas Red-conjugated phalloidin (Molecular Probes) was used to stain F-actin. Indomethacin and LPS (which was purified before use) were obtained from Sigma. Pam3cys and FSL-1 were obtained from EMC microcollections (EMC microcollections GmbH). Salmonella enteritidis [40] and Streptococcus pneumoniae were a clinical isolates. Neisseria meningitidis H44/76 was isolated from a patient with invasive meningococcal disease. Staphylococcus aureus was obtained from the ATCC (ATCC 43300). Bartonella quintana Oklahoma strain [14] and Chlamydia pneumoniae strain TW-183 [41] were used. Bartonella quintana derived LPS is used as TLR4 antagonist [14]. Streptococcus pneumoniae OXC14 was kindly provided by P. Hermans (Radboud University Nijmegen Medical Centre, Nijmegen,

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The Netherlands). All bacteria were heat killed and used at a mode of infection (moi) of 20. To label bacteria with FITC, they were washed in PBS and incubated in 0.5 mg/ml FITC for 60 min. FITC- labeled bacteria were washed and stored in PBS at -20 C.

Preparation of human DCs DCs were generated from PBMCs as described previously [42, 43]. Monocytes were derived from buffy coats or from a leukapheresis product. Plastic-adherent monocytes were cultured in RPMI 1640 medium (Gibco; Life Technologies, Breda, The Netherlands) supplemented with 10% (v/v) FCS (Greiner, Kremsmuenster, Austria), IL-4 (500 U/ml) and GM-CSF (800 U/ml). Immature DCs were harvested on day 6. Expression of MHC class I/II, costimulatory molecules and DC-specific markers on DCs were measured by flow cytometry (data not shown) and the expression of MHC molecules, costimulatory molecules and DC-markers was similar to what was described before [44]. Immature DCs expressed MHC class I and II, the costimulatory molecule CD86, the DC-specific marker CD209/DC-SIGN, low levels of the costimulatory molecule CD80 and lacked expression of the maturation marker CD83.

Mice C57BL/6 mice were obtained from Charles River WIGA Gmbh. TLR2-/- [6], TLR4-/-, and MyD88-/- [45] mice were obtained from S. Akira (Osaka University, Osaka, Japan). The MyD88-/- mice were backcrossed more than 8 times on the C57BL/6 background. Lps2 mice were obtained from B. Beutler (The Scripps Research Institute, La Jolla, USA).

Preparation of mouse DCs Bone marrow was isolated from femurs of C57BL/6 WT, TLR2-/- or TLR4-/- mice. Bone marrow cells were cultured in RPMI 1640 medium supplemented with 10% (v/v) FCS, 50 µM β-mercaptoethanol (Sigma), mouse IL-4 (10 ng/ml) (DNAX, Schering-Plough, Palo Alto, CA) and GM-CSF (10 ng/ml) (PeproTech EC Ltd., London, UK). Immature BMDCs were harvested at day 7. Expression of MHC class I/II, costimulatory molecules and DC-specific markers on DCs were measured by flow cytometry. The iBMDCs expressed high levels of MHC class I/II, CD11c and CD11b, moderate levels of CD80, CD86, CD40 and GR-1 and no expression of F4/80 could be detected (data not shown).

Flow cytometry

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Cells (1 x 105) were incubated with 2% (v/v) human serum in PBA

(PBS containing 1% (w/v) BSA and 0.05% (w/v) NaN3) for 15 min at 4 C. After washing with cold PBA the cells were incubated with primary Ab (5 µg/ml) for 30 min at 4°C. Subsequently, the cells were incubated with PE-labeled goat anti-mouse. Cells were washed and resuspended in 100 µl PBA. Fluorescence was measured using a FACSCalibur™ with CellQuest software (Becton Dickinson & Co.).

Fluorescence microscopy Coverslips were coated with fibronectin (20 µg/ml) (Roche, Mannheim, Germany) in PBS for 1 hour at 37°C. Cells were seeded on fibronectin-coated coverslips, left to adhere for 1 to 6 hrs. After stimulation or being left unstimulated the cells were fixed in 3.7% (w/v) formaldehyde in PBS for 10 min. Cells were permeabilized in 0.1% (v/v) Triton X-100 in PBS for 5 min and blocked with 2% (w/v) BSA in PBS. The cells were incubated with primary Ab for 1 hour. Subsequently the cells were incubated with Alexa Fluor 488-labeled secondary antibodies for 45 min. Subsequently, cells were incubated with Texas Red-conjugated Phalloidin for 30 min. Images were collected on a Leica DMRA fluorescence microscope with a 63x PL APO 1.3 NA oil immersion lens (or a 40x PL FLUOTAR 1.0 NA oil immersion lens for overview images) and a COHU high performance integrating CCD camera (COHU, San Diego, CA). Pictures were analyzed with Leica Qfluoro version V1.2.0 (Leica) and Adobe Photoshop 7.0 (Adobe Systems, Mountain View, CA) software.

Podosome assay Cells were seeded on fibronectin-coated coverslips, stimulated as indicated and stained with an anti-vinculin mAb and phalloidin-Texas Red to detect F-actin. The number of cells displaying podosomes was counted in 7 images per condition per experiment and n indicates the number of independent experiments. An average with SEM is given.

Cytokine measurement Concentrations of human TNF, IL-6 and IL-8 were measured using a human inflammation cytokine bead array (CBA) (Becton Dickinson & Co.) according to the instructions of the manufacturer. Concentrations of human TNFα in were measured using specific RIAs as described previously [46]. Concentrations of human IL-6 were measured by a commercial ELISA kit (Sanquin, Amsterdam, The Netherlands).

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Migration assay 96-wells flat bottom plates (Costar, Corning, Acton, MA) were coated with fibronectin, washed and blocked with 0.01% (w/v) gelatin (Sigma) in PBS for 30 min at 37°C. Per well, 3 x 103 cells were added and mineral oil was pipetted on top of the medium to prevent pH changes and evaporation of the medium. The cells were monitored with a previously described migration system. Cells were recorded for 1 hour followed by analysis of individual cells. The speed was defined as traversed path during the entire experiment divided by the imaging time.

Statistical analysis An ANOVA test or a two-tailed student-t test was used for statistical analysis. P<0.05 was considered significant. Acknowledgements We thank I. Verschueren, Dr. S. Nierkens, Dr. G. Adema and Dr. L. Joosten for providing mice and helping with the BMDC-isolation. We thank C. de Jongh, dr. H. Bootsma and dr. P. Hermans for growing and providing the Streptococcus pneumoniae OXC14. We thank L. Jacobs and T. Jansen for help with the TNFα RIA and the IL-6 ELISA. We thank Dr. G. Schreibert and N. Meeuwsen-Scharenborg for help with the CBA. This study was supported by research funding from the Foundation for Fundamental Research on Matter (FOM 01FB06) and the EU (Cancerimmunotherapy LSHC-CT-2006 518234). References 1. Yarovinsky, F., et al., TLR11 activation of den- 1998. 392(6673): p. 245-52. dritic cells by a protozoan profilin-like protein. 8. Figdor, C.G., Y. van Kooyk, and G.J. Adema, Science, 2005. 308(5728): p. 1626-9. C-type lectin receptors on dendritic cells and 2. Takeda, K. and S. Akira, TLR signaling path- Langerhans cells. Nat Rev Immunol, 2002. ways. Semin Immunol, 2004. 16(1): p. 3-9. 2(2): p. 77-84. 3. Trinchieri, G. and A. Sher, Cooperation of Toll- 9. Tada, H., et al., Synergistic effect of Nod1 and like receptor signals in innate immune Nod2 agonists with toll-like receptor agonists defense. Nat Rev Immunol, 2007. 7(3): p. on human dendritic cells to generate inter- 179-90. leukin-12 and T helper type 1 cells. Infect 4. Elson, G., et al., Contribution of Toll-like Immun, 2005. 73(12): p. 7967-76. receptors to the innate immune response to 10. Kanneganti, T.D., M. Lamkanfi, and G. Nunez, Gram-negative and Gram-positive bacteria. Intracellular NOD-like receptors in host Blood, 2007. 109(4): p. 1574-83. defense and disease. Immunity, 2007. 27(4): 5. Underhill, D.M., et al., The Toll-like receptor 2 p. 549-59. is recruited to macrophage phagosomes and 11. van Helden, S.F., et al., A critical role for discriminates between pathogens. Nature, prostaglandin E2 in podosome dissolution 1999. 401(6755): p. 811-5. and induction of high-speed migration during 6. Takeuchi, O., et al., Differential roles of TLR2 dendritic cell maturation. J Immunol, 2006. and TLR4 in recognition of gram-negative 177(3): p. 1567-74. and gram-positive bacterial cell wall compo- 12. Heine, H., et al., Endotoxic activity and chem- nents. Immunity, 1999. 11(4): p. 443-51. ical structure of lipopolysaccharides from 7. Banchereau, J. and R.M. Steinman, Dendritic Chlamydia trachomatis serotypes E and L2 cells and the control of immunity. Nature, and Chlamydophila psittaci 6BC. Eur J

78 Gram- vs Gram+ bacteria in DC maturation

Biochem, 2003. 270(3): p. 440-50. cytokine production induced by group B 13. Cao, F., et al., Chlamydia pneumoniae-- Neisseria meningitidis: interleukin-12 produc- induced macrophage foam cell formation is tion depends on lipopolysaccharide expres- mediated by Toll-like receptor 2. Infect sion in intact bacteria. Infect Immun, 2001. Immun, 2007. 75(2): p. 753-9. 69(7): p. 4351-7. 14. Popa, C., et al., Bartonella quintana 27. Horng, T., et al., The adaptor molecule TIRAP lipopolysaccharide is a natural antagonist of provides signalling specificity for Toll-like Toll-like receptor 4. Infect Immun, 2007. receptors. Nature, 2002. 420(6913): p. 329- 75(10): p. 4831-7. 33. 15. Yamamoto, M., et al., Cutting edge: a novel 28. Yamamoto, M., et al., Essential role for Toll/IL-1 receptor domain-containing adapter TIRAP in activation of the signalling cascade that preferentially activates the IFN-beta pro- shared by TLR2 and TLR4. Nature, 2002. moter in the Toll-like receptor signaling. J 420(6913): p. 324-9. Immunol, 2002. 169(12): p. 6668-72. 29. O'Neill, L.A. and A.G. Bowie, The family of 16. Beutler, B., et al., Lps2 and signal transduc- five: TIR-domain-containing adaptors in Toll- tion in sepsis: at the intersection of host like receptor signalling. Nat Rev Immunol, responses to bacteria and viruses. Scand J 2007. 7(5): p. 353-64. Infect Dis, 2003. 35(9): p. 563-7. 30. De Trez, C., et al., TLR4 and Toll-IL-1 recep- 17. Seibert, K., et al., Mediation of inflammation tor domain-containing adapter-inducing IFN- by cyclooxygenase-2. Agents Actions Suppl, beta, but not MyD88, regulate Escherichia 1995. 46: p. 41-50. coli-induced dendritic cell maturation and 18. Zachariae, C.O., et al., Phenotypic determi- apoptosis in vivo. J Immunol, 2005. 175(2): p. nation of T-lymphocytes responding to 839-46. chemotactic stimulation from fMLP, IL-8, 31. Wang, Q., et al., Micrococci and peptidogly- human IL-10, and epidermal lymphocyte can activate TLR2-->MyD88-->IRAK-- chemotactic factor. Arch Dermatol Res, 1992. >TRAF-->NIK-->IKK-->NF-kappaB signal 284(6): p. 333-8. transduction pathway that induces transcrip- 19. Lotz, M., et al., B cell stimulating factor tion of interleukin-8. Infect Immun, 2001. 2/interleukin 6 is a costimulant for human thy- 69(4): p. 2270-6. mocytes and T lymphocytes. J Exp Med, 32. Oshiumi, H., et al., TICAM-1, an adaptor mol- 1988. 167(3): p. 1253-8. ecule that participates in Toll-like receptor 3- 20. Burns, S., et al., Maturation of DC is associ- mediated interferon-beta induction. Nat ated with changes in motile characteristics Immunol, 2003. 4(2): p. 161-7. and adherence. Cell Motil Cytoskeleton, 33. Napolitani, G., et al., Selected Toll-like recep- 2004. 57(2): p. 118-32. tor agonist combinations synergistically trig- 21. Dieu, M.C., et al., Selective recruitment of ger a T helper type 1-polarizing program in immature and mature dendritic cells by dis- dendritic cells. Nat Immunol, 2005. 6(8): p. tinct chemokines expressed in different 769-76. anatomic sites. J Exp Med, 1998. 188(2): p. 34. Warger, T., et al., Synergistic activation of 373-86. dendritic cells by combined Toll-like receptor 22. Michelsen, K.S., et al., The role of toll-like ligation induces superior CTL responses in receptors (TLRs) in bacteria-induced matura- vivo. Blood, 2006. 108(2): p. 544-50. tion of murine dendritic cells (DCS). 35. Deva, R., et al., Candida albicans induces Peptidoglycan and lipoteichoic acid are selectively transcriptional activation of inducers of DC maturation and require TLR2. cyclooxygenase-2 in HeLa cells: pivotal roles J Biol Chem, 2001. 276(28): p. 25680-6. of Toll-like receptors, p38 mitogen-activated 23. Hertz, C.J., et al., Microbial lipopeptides stim- protein kinase, and NF-kappa B. J Immunol, ulate dendritic cell maturation via Toll-like 2003. 171(6): p. 3047-55. receptor 2. J Immunol, 2001. 166(4): p. 2444- 36. Kabashima, K., et al., Prostaglandin E2-EP4 50. signaling initiates skin immune responses by 24. Karlsson, H., et al., Pattern of cytokine promoting migration and maturation of responses to gram-positive and gram-nega- Langerhans cells. Nat Med, 2003. 9(6): p. tive commensal bacteria is profoundly 744-9. changed when monocytes differentiate into 37. Hessle, C.C., B. Andersson, and A.E. Wold, dendritic cells. Infect Immun, 2004. 72(5): p. Gram-negative, but not Gram-positive, bacte-

2671-8. ria elicit strong PGE2 production in human 25. Pridmore, A.C., et al., A lipopolysaccharide- monocytes. Inflammation, 2003. 27(6): p. deficient mutant of Neisseria meningitidis 329-32. elicits attenuated cytokine release by human 38. Donnelly, J.P., Bacterial complications of macrophages and signals via toll-like recep- transplantation: diagnosis and treatment. J tor (TLR) 2 but not via TLR4/MD2. J Infect Antimicrob Chemother, 1995. 36 Suppl B: p. Dis, 2001. 183(1): p. 89-96. 59-72. 26. Dixon, G.L., et al., Dendritic cell activation and 39. Ramsay, G. and R.H. van Saene, Selective

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gut decontamination in intensive care and surgical practice: where are we? World J Surg, 1998. 22(2): p. 164-70. 40. Netea, M.G., et al., Salmonella septicemia in rheumatoid arthritis patients receiving antitumor necrosis factor therapy: association with decreased interferon-gamma production and Toll-like receptor 4 expression. Arthritis Rheum, 2003. 48(7): p. 1853-7. 41. Netea, M.G., et al., Non-LPS components of Chlamydia pneumoniae stimulate cytokine production through Toll-like receptor 2- dependent pathways. Eur J Immunol, 2002. 32(4): p. 1188-95. 42. Thurner, B., et al., Generation of large numbers of fully mature and stable dendritic cells from leukapheresis products for clinical application. J Immunol Methods, 1999. 223(1): p. 1-15. 43. De Vries, I.J., et al., Phenotypical and functional characterization of clinical grade dendritic cells. J Immunother, 2002. 25(5): p. 429- 38. 44. De Vries, I.J., et al., Effective migration of antigen-pulsed dendritic cells to lymph nodes in melanoma patients is determined by their maturation state. Cancer Res, 2003. 63(1): p. 12-7. 45. Adachi, O., et al., Targeted disruption of the MyD88 gene results in loss of IL-1- and IL-18- mediated function. Immunity, 1998. 9(1): p. 143-50. 46. Drenth, J.P., et al., Endurance run increases circulating IL-6 and IL-1ra but downregulates ex vivo TNF-alpha and IL-1 beta production. J Appl Physiol, 1995. 79(5): p. 1497-503.

80 Chapter 5 PGE2-mediated podosome loss in dendritic cells is dependent on actomyosin contraction downstream of the RhoA- Rho kinase axis

1Suzanne F.G. van Helden, 1Machteld M. Oud, 1Ben Joosten, 2Niels Peterse, 1Carl G. Figdor and 2Frank N. van Leeuwen

1Department of Tumor Immunology, NCMLS, Radboud University Nijmegen Medical Centre, Nijmegen, The Netherlands 4Laboratory of Pediatric Oncology, NCMLS, Radboud University Nijmegen Medical Centre, Nijmegen, The Netherlands

Journal of Cell Science, 2008,121:1096-1106 Chapter 5

Abstract Podosomes are dynamic adhesion structures found in dendritic cells (DCs) and other cells of the myeloid lineage. We previously showed

that prostaglandin E2 (PGE2), an important proinflammatory mediator produced during DC maturation, induces podosome disassembly within minutes after stimulation. Here we demonstrate that this response is mediated by cAMP elevation, occurs downstream of Rho

kinase and is dependent on myosin II. Whereas PGE2 stimulation leads to activation of the small GTPase RhoA, decreased levels of Rac1-GTP and Cdc42-GTP are observed. These results show that

PGE2 stimulation leads to activation of the RhoA-Rho kinase axis to promote actomyosin-based contraction and subsequent podosome dissolution. Because podosome disassembly is accompanied by de novo formation of focal adhesions (FAs), we propose that the disassembly/formation of these two different adhesion structures is oppositely regulated by actomyosin contractility and relative activities of RhoA, Rac1 and Cdc42. Introduction Based on stability, composition and subcellular localization, different classes of cell-matrix adhesions can be recognized. In most cell types, focal adhesions (FAs), which are present at the end of actin stress fibers, are the most prominent. However, cells of the myeloid lineage, such as macrophages, osteoclasts and dendritic cells (DCs), form podosomes, specialized punctuate adhesions clearly distinct from FAs [1]. Podosomes consist of an actin-dense core surrounded by a ring of cytoskeletal proteins also present in FAs, such as vinculin, paxillin and talin, known to connect integrins to the actin cytoskeleton [2]. Unlike FAs, podosomes are sites of active matrix degradation that promote cell migration and invasion, although, in osteoclasts they have a more specialized role in bone resorption [2]. Moreover, it has been suggested that podosomes facilitate transendothelial migration [3, 4]. DCs are highly specialized antigen presenting cells that play a central role in the induction of T cell-mediated immunity. In response to antigen uptake and exposure to inflammatory stimuli DCs undergo a dramatic phenotypic conversion from a tissue-resident, antigen-cap- turing cell to a highly migratory antigen presenting cell, a process known as DC maturation [5]. Not surprisingly, this transition is accompanied by extensive changes in cell adhesion and cytoskeletal organization [6-8]. A prominent feature of immature DCs (iDCs), which strongly interact with the extracellular matrix, is the presence of podosomes [6, 9]. Mature DCs (mDCs) on the other hand, which

82 PGE2 signals via Rho kinase in DCs are loosely adherent [6], no longer form podosomes on fibronectin (FN) and instead display characteristic actin-rich extensions (den- drites) that maximize the contact area for T cell interactions [10]. Although podosomes are generally considered a feature of migratory and invasive cells, migration speeds in (podosome bearing) iDCs are 10-fold lower than those observed in mDCs [9]. The migratory capacity of DCs on FN in vitro correlates well to the migratory capacity in vivo and the high-speed migration is essential for efficient induction of immune responses in cancer patients treated with DC- vaccination [7]. The presence of podosomes appears to be incom- patible with the high-speed migration observed in mDCs, which could explain why these structures are lost during DC maturation. Podosomes are influenced by both actin and microtubule dynamics [1, 11] and regulated by coordinated activity of the Rho GTPases Rho, Rac and Cdc42 [11]. However, the mechanisms that lead to podosome formation/dissolution are still poorly understood. Previously we found that stimulation of iDCs with maturation inducing agents, such as lipopolysaccharide, leads to a loss of podosomes and that this response is dependent on the production of prostaglandins by the DCs [9]. Moreover, addition of prostaglandin

E2 (PGE2) was sufficient to induce podosome dissolution within minutes after stimulation. These results illustrate that (early) DC maturation involves extensive changes in cytoskeletal organization and cell adhesion, which requires coordinate interactions between signaling molecules, integrins and the actomyosin cytoskeleton. Because

PGE2 plays a key role in the DC maturation process [12, 13] and little is known about the signal-transduction pathways responsible for cytoskeletal remodelling during DC maturation, we have examined in detail how PGE2 signaling affects podosome turnover in DCs. Results iDCs rapidly dissolve podosomes and form FAs in response to

PGE2 stimulation. Immature DCs, seeded on FN, form prominent podosomes. We have shown that PGE2 causes a nearly complete dissolution of podosomes (from 89 ± 1% in the control situation to 10 ± 2% of the

DCs displaying podosomes after 5 min of PGE2 stimulation) within minutes after stimulation, an effect that is maintained in the continued

(16 h) presence of PGE2 (Fig. 1A, B, C, D). In addition, in part of the DCs we observed adhesion structures mostly at the cell periphery (Fig. 1A), which either represented FAs or (more peripherally located) precursors of FAs known as focal complexes [14]. Podosome dissolution in response to PGE2 was accompanied by rapid de novo

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Figure 1. PGE2 stimulation mediated by EP2 and EP4 receptors or by cAMP elevation, leads to podosome dissolution in DCs.

(A/B/C/D) PGE2 stimulation of iDCs leads to podosome loss and the appearance of FAs. (A/B) iDCs seeded on FN-coated coverslips were left untreated or stimulated with

PGE2 for 5 minutes (min) or 16 hours (h) and stained with an anti-vinculin mAb (green) and phalloidin-Texas Red (to detect F-actin, red). (A) Representative images are depicted. (B) The number of cells displaying podosomes or FAs was counted in 7 images per condition per experiment and an average (with s.e.m.) of 6 experiments is shown. (C)

iDCs seeded on FN-coated coverslips were left untreated or stimulated with PGE2 for 5 minutes (min) and stained with an anti-zyxin antibody (green) and phalloidin-Texas Red (to detect F-actin, red). (D) iDCs seeded on FN-coated coverslips were left untreated or

stimulated with PGE2 for 1-5 min and stained with an anti-vinculin mAb and phalloidin- Texas Red (to detect F-actin). The number of cells displaying podosomes was determined in 7 images per condition per experiment and an average (with s.e.m.) of 4 experiments is shown. Asterisks indicate significant differences (P<0.05).

formation of these structures (from 56 ± 3% in the control situation to

84 ± 2% of the DCs displaying FAs after 5 min of PGE2 stimulation) (Fig. 1A, B). The cytoskeletal protein zyxin, an established marker for mature FAs [15], was prominently present both in podosome-rings as well as in the in the newly formed adhesion structures (Fig. 1C), suggesting that the latter represent FAs rather than focal complexes.

After prolonged exposure to PGE2 the amount of cells displaying FAs gradually decreased (Fig. 1A, B).

84 PGE2 signals via Rho kinase in DCs

The transition from podosomes to FAs was further confirmed using Interference Reflection Microscopy (IRM). In iDCs podosomes were seen as dark dots and in part of the cells FAs were visible as dark stripes (Fig. 2A, B and Movie 3, Movie 3 and Fig. S1A in supplementary material). Immunofluorescence detection of F-actin (staining the podosome core) and vinculin (to reveal the podosome ring as well as FAs) was used to confirm that these IRM patterns accurately detect podosomes and FAs in these cells (Fig. 2A). In some DCs we observed bundles of actin filaments, stress fibers, connected to the FAs (Fig 2A lower panel). The average size of the dark spots (0.75 µm) closely corresponds to the average size of the actin core (0.78 µm), whereas the vinculin ring was slightly larger (0.91 µm) (Fig. 2C, D, E, F and Fig. S1A in supplementary material), suggesting that the structures observed with IRM represent the actin cores of the podosomes. By live cell imaging of iDCs using IRM we determined that podosomes dissolve within minutes after stimulation with PGE2, while FAs are forming within the same time frame (Fig. 2B and 3 and

4 in supplementary material). In addition, stimulation with PGE2 appeared to induce a contractile response, which can be seen in the DIC part of the movies (Movie 3 and 4 in supplementary material). IRM imaging revealed that podosome dissolution was consistently preceded by the appearance of bright rings surrounding the podosome cores (Fig. 2B and Movie 3 and 4 in supplementary material), suggesting that changes in the podosome ring structure pre- cede dissolution of the actin-rich core. In order to look at the onset of podosome dissolution in more detail, we fixed cells during live-cell imaging at the moment that the formation of these bright rings was apparent, followed by staining with various podosome components that either localize to the podosome ring or the podosome core. Hence we found that in podosomes undergoing disassembly, as indicated by the presence of the bright rings in the IRM image, the actin core was still largely intact, whereas vinculin could no longer be detected (Fig. S1B in supplementary material). In contrast, the podosome-ring component zyxin could still be observed in these podosomes undergoing disassembly (Fig. S1B in supplementary material). We conclude therefore that the rapid loss of podosomes, induced by PGE2 stimulation, involves the sequential loss of podosome components, accompanied by the formation of FAs.

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86 PGE2 signals via Rho kinase in DCs

Figure 2. Podosome loss and FA formation in DCs upon PGE2 stimulation, visualized with IRM. Also see Movies 1, 2 and Fig. S1A,B in the supplementary material. (A) IRM images compared to fluorescent images (detected with anti-vinculin mAb, GaM- ATTO633 and phalloidin-Texas Red, to detect F-actin) identify adhesion structures as podosomes (seen as dark dots with IRM and as actin dots surrounded by vinculin rings in the fluorescent images) and FAs (seen as dark stripes with IRM and as vinculin stripes in the fluorescent images). (B) Images from an IRM movie (Movie 1 in the supplementary material) showing podosomes before stimulation and loss of podosomes in response to PGE2. Note how the dissolution of podosomes is (transiently) preceded by the formation of bright rings surrounding the podosome core. FA formation in response to stimulation can be observed in the upper-right part of the images. (C) Overlay of the fluorescence intensity of the vinculin staining (broken line) and the actin staining (solid line) from straight-line selections with ImageJ through the middle of the podosome. The actin peak flanked by two vinculin peaks reflects the actin core surrounded by the vinculin-containing ring. (D,E,F) The size of the adhesive contacts observed by live-imaging IRM, the actin core and the vinculin ring were determined in 50 podosomes from ten different cells by making straight-line selections with ImageJ through the middle of the podosome. A representative curve of one podosome is shown and the average size is depicted in the upper-right corner of the curves with errors depicted as SEM. (D) The fluorescence intensity of the actin staining from a straight-line selection through the middle of a podosome. The width of the peak at half the height was used to determine the width of the actin core (0.78 ± 0.01µm). (E) The fluorescence intensity of the vinculin staining from a straight-line selection through the middle of a podosome. The distance from one peak to the other peak was measured to obtain the size of the vinculin ring (0.91 ± 0.02 µm). (F) The light intensity in the IRM image from a straight-line selection through the middle of a podosome. The width of the peak at half the height was used to determine the width of the adhesive contact (0.75 ± 0.02 µm).

Figure 3. PGE2 induced podosome loss is mediated by the EP2 and EP4 receptors. (A) iDCs and mDCs express the EP2 and EP4 receptors. On iDCs and mDCs a RT- β PCR for the PGE2 receptors EP1-4 was performed. Controls include -actin (actin) and minus RT (-). A specific product is detected for EP2 and EP4, but not for EP1 and EP3 receptors. (B) EP2 and EP4 antagonists block PGE2-induced podosome dissolution. iDCs seeded on FN-coated coverslips were left untreated or stimulated with PGE2 in the presence or absence of AH6809 (5 µM for 30 minutes) or AH23848 (50 µM for 30 minutes), or with AH6809 or AH23848 alone and stained with anti-vinculin mAb and phalloidin-Texas Red (to detect F-actin). The number of cells displaying podosomes was counted in 7 images per condition per experiment and an average (with SEM) of 4 experiments is shown. (C) EP2 and EP4 agonists and cAMP elevation mimic PGE2 in inducing podosome dissolution. iDCs seeded on FN-coated coverslips were left untreat- µ µ ed or stimulated with PGE2, butaprost (buta, 2 M for 15 min), PGE1-OH (5 g/ml for 30 min), sulprostone (sulp, 2µ M for 15 min), IBMX (10 µM for 15 min) or IBMX with forskolin (IBMX/fors, 10 µM for 15 min) and stained for vinculin and F-actin. The number of cells displaying podosomes was counted in 7 images per condition per experiment and an average (with SEM) of 3 experiments for the EP agonists (buta, PGE1-OH and sulp) and 7 experiments for cAMP elevation (IBMX and fors) is shown. Asterisks indicate significant differences (P<0.05).

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Figure S1. Podosome dissolution imaged in IRM mode after PGE2 stimulation. (A) Podosome cores are seen in IRM mode as dark spots. IRM image and the IRM image merged with the vinculin (green) or actin (red) staining. Representative images are depicted. (B) Vinculin is already absent form the podosome-rings at the time the bright rings appear, while zyxin is still present. IRM images alone, combined with vinculin (green) or zyxin (green) and/or actin (red) and vinculin (green) or zyxin (green) combined with the actin (red) staining are shown. Representative images are depicted. (C) Myosin IIA localizes to the areas of the bright rings. IRM images alone or combined with myosin IIA (green) and/or actin (red) staining or myosin IIA (green) combined with actin (red) staining. Representative images are depicted. The white arrow indicates a region with bright rings and prominent myosin IIA staining.

PGE2-induced podosome dissolution is mediated by the EP2 and EP4 receptors and can be mimicked by elevating cAMP levels.

To date, four PGE2 receptors (EP1-EP4) have been identified [16]. To investigate which PGE2 receptors are expressed in DCs, we analyzed expression of these receptors by RT-PCR on iDCs and mDCs. Hence, we detected expression of EP2 and EP4, but not EP1 and EP3, receptors in these cells (Fig. 3A). Consistent with these obser-

88 PGE2 signals via Rho kinase in DCs vations, both AH6809 - an antagonist of EP1, EP2 and EP3 [17] - and AH23848, which selectively blocks EP4 [18], alone had no effect on the amount of cells displaying podosomes, although they effectively interfered with PGE2-induced podosome dissolution (Fig. 3B). In addition, sulprostone, a specific agonist for the EP1/EP3 [19], had no effect, whereas butaprost or PGE1-OH, selective agonists for the EP2 and EP4 respectively [19], rapidly induced podosome dissolution (Fig. 3C). Furthermore, we observed that addition of IBMX, forskolin or a combination thereof, resulting in elevated cAMP levels, could induce podosome dissolution (Fig. 3C and data not shown).

PGE2 mediated podosome loss is dependent on myosin IIA function

Because PGE2 appears to induce a contractile response in DCs (as shown by DIC images Movie 3 and 4 supplementary material), we further explored a role for myosin II in PGE2-induced podosome dissolution. We determined by Western blot analysis that myosin IIA is the predominant isoform expressed in human iDCs, whereas expression of myosin IIB was low and myosin IIC was undetectable (Fig. 4A). In most cells, we observed a fine punctuate pattern of myosin IIA distribution that was enriched in the podosome-ring structures or podosome-rich areas of the cell, whereas myosin IIB was nearly undetectable (Fig. 4B). Importantly, inhibition of myosin II function, by preincubation with 10 µ M of the myosin II specific inhibitor blebbistatin [20] blocked PGE2- induced podosome loss (from 10 ± 2% with PGE2 to 60 ± 4% with PGE2 and blebbistatin); FAs failed to develop (Fig. 4C and Movie 5 in supplementary material). Although it has been reported that myosin II function is also required for the formation (and maintenance) of podosomes, incubation with blebbistatin had no effect on the amount of podosomes in unstimulated cells nor did it significantly affect myosin IIA distribution at the indicated concentration (Fig. 4D). Although in unstimulated control cells myosin IIA was enriched in podosome rings, in cells stimulated with PGE2 a more fibrillar organization of myosin IIA was observed (Fig. 4D). This PGE2- dependent reorganization of myosin IIA into fibrils was not observed when cells were pre-treated with blebbistatin. In addition, the formation of fibrillar myosin IIA coincided with the dissolution of podosomes and fibrillar myosin IIA assemblies appeared to specifically localize to those areas of the cell in which podosomes were actively dissolving (as indicated by the IRM ring structures) (Fig. S1C in supplementary material), suggesting that an increase in myosin IIA activity con- tributes to podosome dissolution at these sites. Together these find-

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Figure 4. Myosin II is enriched in podosomes and myosin II function is needed for PGE2- induced podosome dissolution. (A) Myosin IIA is the predominant isoform in DCs. Cell lysates from iDCs and N1E-115 cells (N115, positive control expressing all isoforms) were analyzed for expression of myosin (myo) IIA, IIB and IIC. (B) Myosin IIA is enriched in podosome-rings, whereas myosin IIB is nearly undetectable. iDCs were plated on FN-coated coverslips and subsequently stained with an anti-myosin IIA or IIB antibody (green) and phalloidin-Texas Red (to detect F-actin, red). (C,D) iDCs were plated on

FN-coated coverslips and left untreated or stimulated with PGE2 for 5 min in the presence or absence of blebbistatin (bleb, 10 µM for 30 min) or treated with blebbistatin

alone. (C) Inhibition of myosin II function blocks PGE2-induced podosome dissolution. iDCs were stained with anti-vinculin antibodies and phalloidin-Texas Red. The number of cells displaying podosomes was counted in 7 images per condition per experiment and an average (with SEM) of 3 experiments is shown. Asterisks indicate significant differences (P<0.05). (D) Inhibition of myosin II function blocks PGE2-induced myosin IIA redistribution. iDCs were stained with anti-myosin IIA antibodies and phalloidin-Texas Red.

90 PGE2 signals via Rho kinase in DCs

ings indicate that PGE2-mediated podosome dissolution and the subsequent formation of FAs require myosin II-mediated contraction.

PGE2 stimulation activates RhoA and reduces Rac1 and Cdc42 activity Small GTPases of the Rho family (Rho, Rac and Cdc42) are key regulators of adhesion dynamics that control the formation and turnover of filamentous actin as well as microtubules and modulate myosin II- based contractility [21, 22]. Therefore, we examined the effects of

PGE2 stimulation on Rho GTPase activity using GTPase pull down assays. The number of cells needed for such experiments and the high protease activity observed in DC-lysates precluded the use of primary DCs for this purpose. Instead we used HL-60 cells, a monocytic leukaemia cell line that can be induced to form podosomes in response to PMA. We determined by RT-PCR that, similar to DCs, HL-60 cells predominantly express EP2 and EP4 receptors and respond to PGE2 by dissolving podosomes within the same timescale as observed in DCs. In HL-60 cells, this response

Figure 5. HL-60 cells form podosomes upon PMA stimulation and respond to PGE2. (A/B) HL-60 cells form podosomes upon stimulation with PMA, which dissolve after stimulation with PGE2 or an EP4-agonist. HL-60 cells seeded on FN-coated coverslips were left untreated or stimulated with PMA for 24h and left untreated or stimulated with

PGE2 for 5 min and stained with an anti-vinculin mAb and phalloidin-Texas Red (to detect F-actin). (A) Representative images are depicted. (B) The number of cells displaying podosomes was counted in 7 images per condition per experiment and an average (with SEM) of 3 experiments is shown. (C) HL-60 cells show expression of EP2 and EP4, while EP1 and EP3 receptors are nearly undetectable as determined by RT-PCR. Asterisks indicate significant differences (P<0.05).

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appeared to be mediated primarily by the EP4 receptor, because these cells responded well to the EP4 agonist PGE-OH (Fig. 5), but not butaprost (data not shown) When measuring RhoA activity, using a Rhotekin-GST fusion protein, we observed a 2.6 fold increase in RhoA-GTP levels in response to

Figure 6. PGE2 receptor stimulation leads to activation of RhoA and inactivation of Rac1. (A) Activation of the small GTPase RhoA in response to PGE2. GTP-RhoA levels were determined in lysates (pd: pull down, TL: total lysate) of HL-60 cells using a

Rhotekin-GST pull down assay. In cells treated with PGE2 for 5 min, Rho-GTP levels showed a 2.6 ± 0.9 (s.d.) fold increase relative to untreated cells (n=7). (B) Loss of Rac1

GTPase activity in response to PGE2. Rac1-GTP levels were determined in lysates of HL-60 cells using a biotinylated Pak1 peptide. In cells stimulated with PGE2 Rac1 activity was decreased, 0.5 ± 0.2 (s.d.)-fold relative to untreated cells (n=6). (C) Loss of

Cdc42 GTPase activity in response to PGE2. Cdc42-GTP levels were determined in lysates of HL-60 cells using a biotinylated Pak1 peptide. In cells stimulated with PGE2 Cdc42 activity was decreased, 0.7 ± 0.2 (s.d.)-fold relative to untreated cells (n=6).

(D/E) Activation of the small GTPase RhoA in response to PGE1-OH or 8-Bromo-cAMP. GTP-RhoA levels were determined in lysates (pd: pull down, TL: total lysate) of HL-60 cells using a Rhotekin-GST pull down assay. (D) In cells treated for 5 min with an EP4

agonist (PGE1-OH), Rho-GTP levels showed a 1.6 ± 0.5 (s.d.) fold increase relative to untreated cells (n=8). (E) In cells treated for 5 min with 8-Bromo-cAMP (8-Bromo, 200 µM), which elevates cAMP levels, Rho-GTP levels showed a 2.5 ± 0.9 (s.d.) fold increase relative to untreated cells (n=5). Asterisks indicate significant differences (P<0.05).

92 PGE2 signals via Rho kinase in DCs

PGE2 (Fig. 6A). We similarly determined Rac1 and Cdc42 activity in response to PGE2 stimulation using a biotinylated Pak1 peptide. Stimulation with PGE2 led to a decrease in Rac1 activity (0.5 fold of control) and a comparable decrease in Cdc42 activity (0.7 fold of control) (Fig. 6B and C), whereas the total amount of Rac1 and

Cdc42 remained constant. These results indicate that PGE2-induced podosome dissolution involves activation of RhoA accompanied by inactivation of Rac1 and Cdc42. In addition, the activity of RhoA was determined upon stimulation with butaprost or PGE1-OH, EP2 and EP4 agonists respectively. Stimulation with butaprost did not change the amount of active RhoA

(1.1 fold of control) (data not shown), whereas PGE1-OH led to an increase in RhoA activity (1.6 fold of control) (Fig. 6D). These findings are consistent with our observation that PGE2-mediated podosome dissolution in these cells appears to be predominantly mediated by EP4. Furthermore, the effect of the cAMP-analog 8- Bromo-cAMP on RhoA activity was investigated. Hence we determined that a 5 minute stimulation with 8-Bromo-cAMP led to 2.5 fold increase in RhoA activity (Fig. 6E). Together, these findings suggest that raising cAMP levels, by stimulation of EP-receptors, leads to activation of RhoA.

PGE2-induced podosomes dissolution is mediated by Rho kinase. We subsequently investigated a role for Rho kinase, a prominent downstream effector of RhoA, in PGE2-induced podosome dissolution. The Rho kinase inhibitor Y27632 [23] effectively blocked podosome dissolution induced by PGE2 in DCs (from 22 ± 3% with PGE2 to 69 ± 5% with PGE2 and Y27632) (Fig. 7A) as well as HL-60 cells (data not shown). In addition, EP2 and EP4 agonist-induced podosome loss was effectively blocked by Y27632 (Fig. 7A). Podosome dissolution induced by IBMX or IBMX/forskolin was similarly blocked by Rho kinase inhibition (from 13 ± 5% with IBMX to 80 ± 4% with IBMX and Y27632) (Fig. 7B). The distribution of podosomes in response to Y27632 treatment alone was unaffected, whereas when cells were treated with Y27632 in combination with

PGE2, podosomes were still present (Fig. 7C). We conclude that PGE2 receptors that are present on dendritic cells as well as HL-60 monocytic leukaemia cells signal to activate the RhoA-Rho kinase axis and promote actomyosin-based contractility, leading to podosome dissolution and the formation of FAs.

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Figure 7. Podosome loss induced by PGE2, EP-agonists and elevated cAMP-induced podosome loss is mediated by Rho kinase.

(A) Rho kinase inhibition blocks PGE2- and EP-agonist induced podosome dissolution. iDCs seeded on FN-coated coverslips were left untreated or stimulated with PGE2 for 5 µ µ min, butaprost (buta, 2 M for 15 min) or PGE1-OH (5 g/ml for 30 min) in the presence or absence of Y27632 (20 µM for 1 h) or treated with Y27632 alone and stained with anti-vinculin mAb and phalloidin-Texas Red to detect F-actin. The number of cells displaying podosomes was determined in 7 images per condition per experiment and an average (with SEM) of 3 experiments is shown. (B) IBMX and IBMX/forskolin-induced podosome dissolution is efficiently blocked by Y27632. iDCs seeded on FN-coated coverslips were left untreated or stimulated with IBMX (10 µM for 5 min) or IBMX with forskolin (IBMX/fors, both 10 µM for 5 min) in the presence or absence of Y27632 (20 µM for 1 h) or with Y27632 alone, and stained with anti-vinculin mAb and phalloidin- Texas Red (to detect F-actin). The number of cells displaying podosomes was counted in 7 images per condition per experiment and an average (with SEM) of 3 experiments is shown. Asterisks indicate significant differences (P<0.05). (C) Rho kinase inhibition

blocks podosome loss in response to PGE2, EP2/EP4-agonists or cAMP elevation. Representative images are depicted.

Movies can be found at http://www.ncmls.eu/NCMLS/Visual /movie.asp Discussion In response to antigen uptake and exposure to inflammatory stimuli

iDCs mature into highly migratory mDCs [5]. PGE2, a proinflammatory mediator and maturation-inducing factor is essential for the induction of full migratory capacity in DCs [9, 12, 24]. Immature DCs display podosomes while part of the cells also displays vinculin-containing adhesions at the cell periphery. The presence of zyxin in these structures identifies these as FAs [15]. Upon stimulation with

94 PGE2 signals via Rho kinase in DCs

PGE2, iDCs rapidly dissolve podosomes and start to form FAs, suggesting that signals inducing podosome dissolution favor the formation of FAs. When, after prolonged exposure to PGE2, the transition from strongly adhesive iDCs to highly migratory mDCs is made, the amount of cells displaying FAs gradually decreases. Because transfection of fluorescently-labeled proteins in primary

DCs is difficult, we applied IRM to follow the effects of PGE2 on DC adhesion structures in living cells. With this technique, which has been previously used to visualize either FAs or podosomes [25-27], structures close to the glass, such as podosomes and FAs, come out dark whereas structures further away from the glass appear lighter. By combining IRM with immunofluorescence detection after fixation, we confirmed that the dark spots observed by IRM correspond in size and localization to the actin-rich cores of podosomes. The latter may seem counter-intuitive, as the (integrin containing) podosome rings are the primary sites for adhesion; however, the fact that podosomes are highly protrusive structures driven by actin polymerization in the core may explain why the podosome core appears closer to the glass than the surrounding ring structure [3, 4, 28]. Interestingly, podosome dissolution is invariably preceded by the transient appearance of bright rings in the IRM image, surrounding the actin-rich cores. By in situ fixation of podosomes undergoing dissolution, we showed that at the time these bright rings appear, vinculin is no longer present in these podosome-remnants, whereas the actin-rich cores, as well as associated zyxin are still clearly detectable. Together these results suggest that initial changes affect the podosome ring prior to (complete) dissolution of the actin core. Mouse knockout studies have revealed an important role for EP4 in Langerhans cell migration [29], while in humans, both EP2 and EP4 have been linked to DC function [30]. In line with earlier findings [31], expression of EP2/EP4 but not EP1/EP3 was detected in DCs. The effects of specific EP4-(ant)agonists implicate this receptor in PGE2- induced podosome loss. Similarly, the effects of a specific EP2 agonist and the lack thereof in response to an EP1/EP3 agonist similarly support a role for the EP2 receptor in PGE2-mediated podosome loss. These pharmacological studies, combined with the observed expression pattern, suggest that these PGE2-induced responses in DCs are mediated by EP2/EP4 receptors both of which signal through Gs to elevate cAMP levels [16]. Indeed we observed that raising cAMP levels mimics the effects of PGE2 on podosome dissolution. Although podosome formation has been studied in some detail, much less is known about signaling mechanisms that trigger

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podosome dissolution. Small GTPases of the Rho family (RhoA, Rac1 and Cdc42) are key regulators of adhesion dynamics that control the formation and turnover of filamentous actin as well as microtubules, and modulate myosin II-based contractility [21, 22]. In endothelial cells, podosome organization is regulated by RhoA and Cdc42 [32] and in Src-transformed fibroblasts active RhoA localizes to podosomes [33]. Inhibition of RhoA maintains the podosome belt at the osteoclast periphery despite depolymerization of microtubules by nocodazole [34], suggesting that RhoA activity is needed to induce podosome loss. Consistent with these observations in osteo-

clasts, we found that PGE2-induced podosome dissolution involves activation of RhoA. Ourselves and others have shown that activation of RhoA is often accompanied by a loss of Rac1 activity, whereas active forms of these two Rho GTPases often produce opposite

effects on the cytoskeleton [35, 36]. Indeed, PGE2-mediated activation of RhoA is accompanied by a decrease in activity of both Rac1 and Cdc42. It was shown earlier that Rac1 as well as Cdc42 are needed for podosome formation in DCs [11], while the Cdc42/Rac1- regulated kinases Pak1 and Pak4 either induce podosome formation or reduce podosome turnover [37-39]. Another effector of Cdc42, the Wiskott Aldrich Syndrome protein (WASp), also plays an essential role in the formation and maintenance of podosomes in both in human macrophages [40] and endothelial cells [32], whereas WASp deficient DCs and macrophages lack podosomes [6, 40]. Together,

these data suggest that PGE2-mediated activation of RhoA combined with a simultaneous inactivation of Rac1 and Cdc42 contribute to the dissolution of podosomes in DCs and HL-60 cells. In most cell types cAMP-mediated activation of PKA antagonizes the effects of RhoA on actomyosin contractility, leading to cytoskeletal

relaxation [41-43]. In DCs however, PGE2 stimulation appears to induce a contractile response while activating RhoA. Similar to PGE2, we observed cytoskeletal contraction and increased RhoA-GTP levels in response to EP-agonists as well as to stimulation with the cAMP-analog 8-Bromo-cAMP. In addition, stimulation with a PKA activator resulted in increased levels of RhoA-GTP in HL-60 cells,

whereas PKA inhibition interfered with PGE2-induced podosome loss in DCs (results not shown). Together these results indicate that acti-

vation of RhoA as well as PGE2-induced podosome dissolution is mediated by cAMP-PKA signaling. In most adherent cells, the formation of stress fibers and stable FAs is triggered by activation of RhoA and its downstream effector Rho kinase, promoting actomyosin contractility [21]. Rho kinase acts by promoting phosphorylation of the myosin II regulatory light chain [44,

96 PGE2 signals via Rho kinase in DCs

45]. In addition, Rho kinase activity can enhance stress fiber formation by inactivation of the actin severing protein cofilin [46], whereas direct activation of mDia by RhoA stimulates actin polymerization [21, 47]. Therefore, we assessed the effects of Rho kinase inhibition on

PGE2-induced podosome dissolution. In smooth muscle cells, phorbol-dibutyrate-induced podosome formation appears to be Rho kinase independent [48, 49]. In our experiments however, inhibition of Rho kinase not only effectively interfered with podosome dissolution induced by PGE2, but also interfered with that induced by EP- agonists and by elevated cAMP levels. This effect of Rho kinase was observed in primary DCs as well as in HL-60 cells, demonstrating that PGE2-induced podosome dissolution is dependent on Rho kinase activity both in DCs and these monocytic leukaemia cells. This is the first demonstration that (cAMP elevating) EP2/EP4 receptors can signal to activate the RhoA-Rho kinase axis, and our results indicate that inhibition of RhoA signaling downstream of cAMP is not a universal principle. A number of studies have implicated myosin II in podosome-regulation. For instance, formation of podosomes was shown to involve (local) inhibition of myosin II in smooth muscle cells as well neuroblastoma cells [49, 50], whereas in macrophages [51] or fibroblasts [52] myosin II-based activity is needed for the formation and maintenance of these structures. Although these different experimental outcomes may be in part dependent on cell type or culture conditions, we propose that basal myosin IIA activity is required for the formation and maintenance of podosomes, whereas a sudden (stimulus induced) increase in myosin II activity may trigger podosome dissolution. This idea is supported by our findings that relatively low concentrations of blebbis- µ tatin (10 M) effectively block PGE2-induced podosome loss without affecting podosome turnover, whereas higher concentrations (40-100 µM) interfere with the formation and/or maintenance of these structures [51, 52]. Hence we conclude that, dependent on cell type and experimental conditions, local changes in myosin II activity can either trigger formation of podosomes or contribute to their demise. In mammalian cells three myosin II isoforms have been identified, known as myosin IIA, myosin IIB and myosin IIC [53-55]. Here we show that myosin IIA, the predominant isoform expressed in DCs, localizes to podosome rings, whereas only trace amounts of myosin IIB are present. Although we cannot exclude a role for myosin IIB in podosome dissolution in DCs, the prominent expression and localization of myosin IIA to podosome-rich areas of the cells, suggest that this myosin isoform is primarily involved in PGE2-mediated

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podosome dissolution. Pharmacological inhibition of myosin II function, using blebbistatin, demonstrated a requirement for myosin II in

PGE2-induced podosome dissolution. In addition, PGE2 stimulation in DCs leads to a noticeable change in myosin II distribution, from a punctuate cytosolic pattern to a more fibrillar distribution. These fibrillar structures are reminiscent of the myosin II ribbons described in fibroblasts [56] and most likely reflect increased association with F- actin and/or a change in solubility of myosin filaments. We have shown previously, in mouse neuroblastoma cells, that pharmacological inhibition of myosin II leads to a loss of focal adhesions accompanied by the formation of podosomes [50]. In addition, WASp-interacting-protein deficient DCs lack podosomes, but still form FAs [25]. Furthermore, in avian multinucleated giant cells disruption of the microtubules by nocodazole induced the loss of podosomes accompanied by the formation of stress fibers and FAs and activation of RhoA and inactivation of Rac1, whereas repoly- merization of microtubules induced loss of FAs and reformation of podosomes [57]. These findings suggest that, in different cell types, podosomes and FAs are oppositely affected by Rho GTPase activity and actomyosin contractility. Hence we propose that localized

Figure 8. Regulation of adhesion dynamics in dendritic cells by PGE2. PGE2 signaling, mediated by EP2 and EP4 receptors leads to elevation of the level of cAMP, activation of RhoA and inactivation of Rac1. PGE2-mediated activation of RhoA promotes Rho kinase activity and, subsequently, myosin II dependent contraction, leading to podosome dissolution and FA formation. By contrast, pathways leading to activation of Rac1 are known to promote podosome formation and induce cytoskeletal relaxation accompanied by a loss of FAs.

98 PGE2 signals via Rho kinase in DCs changes in actin dynamics in combination with the contractile state of a cell, determine whether FAs or podosomes are formed.

Besides enabling high-speed migration, PGE2-induced podosome dissolution could have additional functions. It may serve to redeploy actin to other processes in the cell, such as vesicle movement, which has been linked to podosome dynamics [58, 59]. Furthermore, microtubules were shown to influence FA and podosome dynamics, an effect that could be mediated by (microtubule dependent) vesicle transport [51, 60]. In mouse DCs a loss of podosomes was associated with a transient increase in antigen uptake [8]. Similarly, we observed a moderate increase in antigen uptake in human DCs shortly after stimulation with PGE2 (data not shown). Thus in addition to enabling high-speed migration, podosome dissolution in DCs could transiently enhance antigen uptake and thereby add to the induction of immune responses. As summarized in the model shown in Fig. 8, our data show that

PGE2 mediates podosome dissolution by myosin II-mediated contractility downstream of both RhoA and Rho kinase activation. This is the first report showing activation of the RhoA-Rho kinase axis downstream of receptors (EP2/EP4) known to couple to cAMP elevation. Moreover, our data provide novel insights in the dynamics of the DC- cytoskeleton and its regulation during the induction of DC-maturation. Materials and methods Chemicals and antibodies Antibodies used were: mIgG1 (Becton Dickinson & Co.), GaM- ATTO633, anti-vinculin and anti-myosin IIB (Sigma), anti-myosin IIA (Biomedical Technologies Inc.), anti-myosin IIC (obtained from dr. R. Adelstein (NIH), anti-RhoA and anti-zyxin (Santa Cruz biotechnology), anti-Rac1 (Upstate biotechnology, Millipore), IRDye 800 CW- labeled secondary antibodies (LI-COR Biosciences), and Alexa Fluor 488-labeled secondary antibodies and Texas Red-conjugated phalloidin (Molecular Probes). Chemicals used were: blebbistatin, Y27632 and PMA (Calbiochem), FN, butaprost, sulprostone, AH23848, AH6809, IBMX, forskolin and 8-Bromo-cAMP (Sigma), and PGE1-OH (Cayman chemical company). PGE2 was used at 10 µg/ml (Pharmacia & Upjohn).

Isolation of monocytes, preparation of DCs and cell culture DCs were generated from peripheral blood mononuclear cells as described previously [7]. Expression of major histocompatibility complex class I/II, costimulatory molecules and DC-specific markers

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were measured by flow cytometry on iDCs d6 and expression was similar to what was described before (data not shown) [7]. HL-60 cells (generous gift from Dr. L. Machesky) were cultured in IMDM medium (Invitrogen) supplemented with 10% (v/v) FCS (Greiner). N1E-115 cells were cultured in DMEM medium (Invitrogen) supplemented with 10% (v/v) FCS.

Fluorescence microscopy Fluorescence microscopy was performed as described previously [9]. A Leica DMRA fluorescence microscope with 63x PL APO 1.3 NA or 40x PL FLUOTAR 1.0 NA oil immersion lens and COHU high performance integrating CCD camera were used. Pictures were analyzed with Leica Qfluoro version V1.2.0 and Adobe Photoshop 7.0 software.

Interference Reflection Microscopy (IRM) IRM was performed using a Zeiss LSM 510-meta microscope with 488 nm laser with a band pass filter at 470-500 nm or 633 nm laser with a long pass filter at 560 nm and a Plan-Apochromatic 63x 1.4 NA oil immersion DIC lens (Carl Zeiss GmbH). Cells were seeded on FN-coated Willco glass bottom dishes (Willco Wells BV) and either fixed and stained for vinculin and actin before fluorescent and IRM images were obtained, or cells were live-imaged in IRM mode with 20-second intervals at 37°C in RPMI 1640 without phenol red

(Invitrogen) during addition of PGE2. In addition, cells were live- imaged in IRM mode and fixed while podosomes were dissolving

upon PGE2 stimulation and stained for actin and vinculin, myosin IIA or zyxin and fluorescent and IRM images were obtained. Cells were imaged using Zeiss LSM Image Browser version 32 and images were processed with Image J version 1.32j software (National Institutes of Health, http://rsb.info.nih.gov/ij).

RT-PCR RNA was isolated from iDCs, mDCs and HL-60 cells with Trizol reagent (Gibco) according to the instruction of the manufacturer. A RT-PCR was performed using a T3 Thermocycler at 20°C for 10 min, 42°C for 45 min and 95°C for 10 min. An annealing temperature of 55°C with 30 cycles was used. The following primers were used: EP1 forward, 5'-ATCATGGTGGTGTCGTGCATC-3'; EP1 reverse, 5'- GGTCCAGG ATCTGGTTCCAG-3'; EP2 forward, 5'-ATGACCAT- CACCTTCGCC-3'; EP2 reverse, 5'-CAAAGACCCAAGGGTCAATT- 3'; EP3 forward, 5'-GTATGCGAGCCACATGAAGA-3'; EP3 reverse, 5'-CAGAGGCGAAGAAAAGGTTG-3'; EP4 forward, 5'-TCCTGGCTT

100 PGE2 signals via Rho kinase in DCs

TTGAGCACTTT-3'; EP4 reverse, 5'-CCTCTGCTGTGTGCCAAATA- 3'; actin forward, 5'-GCTACGAGCTGCCTGACGG-3'; actin reverse, 5'-GAGGCCAGGATGGAGCC-3'.

RhoA-, Rac1- and Cdc42-activation assay and myosin II isoform detection RhoA, Rac1 and Cdc42 pull down experiments were performed as described previously [61]. Samples were analyzed by Western blotting using a mouse anti-RhoA or anti-Rac1 or a rabbit anti-Cdc42 antibodies. Cell lysates were analyzed for myosin II isoform expression by Western blotting using rabbit anti-myosin IIA, -myosin IIB or -myosin IIC antibodies. Samples were analyzed on an Odyssey infrared imaging system (LI-COR Biosciences).

Statistical analysis ANOVA or the two-tailed t test were used for statistical analysis. Significant differences or significant differences from control are indicated with asterisks (p<0.05). Acknowledgements We thank Dr. L. Machesky for providing the HL-60 cells and Dr. R. Adelstein for providing myosin IIC antibodies. We thank dr. I. de Vries, N. Meeuwsen-Scharenborg, A. de Boer, M. van de Rakt and M. Kerkhoff for providing DCs. This study was supported by research funding from the Foundation for Fundamental Research on Matter (FOM 01FB06) and the Dutch Cancer Society (KWF KUN 2006- 3699). References 1. Linder, S. and M. Aepfelbacher, Podosomes: 7. De Vries, I.J., et al., Effective migration of adhesion hot-spots of invasive cells. Trends antigen-pulsed dendritic cells to lymph nodes Cell Biol, 2003. 13(7): p. 376-85. in melanoma patients is determined by their 2. Buccione, R., J.D. Orth, and M.A. McNiven, maturation state. Cancer Res, 2003. 63(1): p. Foot and mouth: podosomes, invadopodia 12-7. and circular dorsal ruffles. Nat Rev Mol Cell 8. West, M.A., et al., Enhanced dendritic cell Biol, 2004. 5(8): p. 647-57. antigen capture via toll-like receptor-induced 3. Calle, Y., et al., Inhibition of calpain stabilizes actin remodeling. Science, 2004. 305(5687): podosomes and impairs dendritic cell motility. p. 1153-7. J Cell Sci, 2006. 119(Pt 11): p. 2375-85. 9. van Helden, S.F., et al., A critical role for 4. Carman, C.V., et al., Transcellular diapedesis prostaglandin E2 in podosome dissolution is initiated by invasive podosomes. Immunity, and induction of high-speed migration during 2007. 26(6): p. 784-97. dendritic cell maturation. J Immunol, 2006. 5. Banchereau, J. and R.M. Steinman, Dendritic 177(3): p. 1567-74. cells and the control of immunity. Nature, 10. Mellman, I., S.J. Turley, and R.M. Steinman, 1998. 392(6673): p. 245-52. Antigen processing for amateurs and profes- 6. Burns, S., et al., Maturation of DC is associ- sionals. Trends Cell Biol, 1998. 8(6): p. 231- ated with changes in motile characteristics 7. and adherence. Cell Motil Cytoskeleton, 11. Burns, S., et al., Configuration of human den- 2004. 57(2): p. 118-32. dritic cell cytoskeleton by Rho GTPases, the

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protein antigens: role of PGE2 in chemokine turnover in migrating cells -- over and over and cytokine expression by MoDCs. Int and over again. Nat Cell Biol, 2002. 4(4): p. Immunol, 2005. 17(12): p. 1561-72. E97-100. 25. Chou, H.C., et al., WIP regulates the stability 39. Webb, B.A., et al., PAK1 induces podosome and localization of WASP to podosomes in formation in A7r5 vascular smooth muscle

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cells in a PAK-interacting exchange factor- 54. Katsuragawa, Y., et al., Two distinct nonmus- dependent manner. Am J Physiol Cell cle myosin-heavy-chain mRNAs are differen- Physiol, 2005. 289(4): p. C898-907. tially expressed in various chicken tissues. 40. Linder, S., et al., Wiskott-Aldrich syndrome Identification of a novel gene family of verte- protein regulates podosomes in primary brate non-sarcomeric myosin heavy chains. human macrophages. Proc Natl Acad Sci U S Eur J Biochem, 1989. 184(3): p. 611-6. A, 1999. 96(17): p. 9648-53. 55. Shohet, R.V., et al., Cloning of the cDNA 41. Dong, J.M., et al., cAMP-induced morpholog- encoding the myosin heavy chain of a verte- ical changes are counteracted by the activat- brate cellular myosin. Proc Natl Acad Sci U S ed RhoA small GTPase and the Rho kinase A, 1989. 86(20): p. 7726-30. ROKalpha. J Biol Chem, 1998. 273(35): p. 56. Verkhovsky, A.B., T.M. Svitkina, and G.G. 22554-62. Borisy, Myosin II filament assemblies in the 42. Lang, P., et al., Protein kinase A phosphoryla- active lamella of fibroblasts: their morphogen- tion of RhoA mediates the morphological and esis and role in the formation of actin filament functional effects of cyclic AMP in cytotoxic bundles. J Cell Biol, 1995. 131(4): p. 989- lymphocytes. EMBO J, 1996. 15(3): p. 510-9. 1002. 43. Manganello, J.M., et al., Protein kinase A- 57. Ory, S., O. Destaing, and P. Jurdic, mediated phosphorylation of the Galpha13 Microtubule dynamics differentially regulates switch I region alters the Rho and Rac activity and triggers Rho-inde- Galphabetagamma13-G protein-coupled pendent stress fiber formation in macrophage receptor complex and inhibits Rho activation. polykaryons. Eur J Cell Biol, 2002. 81(6): p. J Biol Chem, 2003. 278(1): p. 124-30. 351-62. 44. Amano, M., et al., Phosphorylation and acti- 58. Cougoule, C., et al., Activation of the lyso- vation of myosin by Rho-associated kinase some-associated p61Hck isoform triggers the (Rho-kinase). J Biol Chem, 1996. 271(34): p. biogenesis of podosomes. Traffic, 2005. 6(8): 20246-9. p. 682-94. 45. Kimura, K., et al., Regulation of myosin phos- 59. McNiven, M.A., M. Baldassarre, and R. phatase by Rho and Rho-associated kinase Buccione, The role of dynamin in the assem- (Rho-kinase). Science, 1996. 273(5272): p. bly and function of podosomes and invadopo- 245-8. dia. Front Biosci, 2004. 9: p. 1944-53. 46. Maekawa, M., et al., Signaling from Rho to 60. Ezratty, E.J., M.A. Partridge, and G.G. the actin cytoskeleton through protein kinas- Gundersen, Microtubule-induced focal adhe- es ROCK and LIM-kinase. Science, 1999. sion disassembly is mediated by dynamin 285(5429): p. 895-8. and focal adhesion kinase. Nat Cell Biol, 47. Geneste, O., J.W. Copeland, and R. 2005. 7(6): p. 581-90. Treisman, LIM kinase and Diaphanous coop- 61. van Leeuwen, F.N., et al., Rac activation by erate to regulate serum response factor and lysophosphatidic acid LPA1 receptors actin dynamics. J Cell Biol, 2002. 157(5): p. through the guanine nucleotide exchange 831-8. factor Tiam1. J Biol Chem, 2003. 278(1): p. 48. Hai, C.M., et al., Conventional protein kinase 400-6. C mediates phorbol-dibutyrate-induced cytoskeletal remodeling in a7r5 smooth muscle cells. Exp Cell Res, 2002. 280(1): p. 64- 74. 49. Burgstaller, G. and M. Gimona, Actin cytoskeleton remodelling via local inhibition of contractility at discrete microdomains. J Cell Sci, 2004. 117(Pt 2): p. 223-31. 50. Clark, K., et al., TRPM7, a novel regulator of actomyosin contractility and cell adhesion. EMBO J, 2006. 25(2): p. 290-301. 51. Kopp, P., et al., The kinesin KIF1C and microtubule plus ends regulate podosome dynamics in macrophages. Mol Biol Cell, 2006. 17(6): p. 2811-23. 52. Collin, O., et al., Spatiotemporal dynamics of actin-rich adhesion microdomains: influence of substrate flexibility. J Cell Sci, 2006. 119(Pt 9): p. 1914-25. 53. Golomb, E., et al., Identification and characterization of nonmuscle myosin II-C, a new member of the myosin II family. J Biol Chem, 2004. 279(4): p. 2800-8.

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104 Chapter 6 Adhesion of dendritic cells to micro-patterned substrates reveals physical aspects of podosome regulation

1Suzanne F.G. van Helden, 1Joost te Riet, 1Machteld M. Oud, 2Heike Glauner, 3Ruth Diez-Ahedo, 4Frank N. van Leeuwen, 3Maria F. Garcia Parajo, 2Roland Brock and 1Carl G. Figdor.

1Department of Tumor Immunology, NCMLS, Radboud University Nijmegen Medical Centre, Nijmegen, The Netherlands 2Department of Biochemistry, NCMLS, Radboud University Nijmegen Medical Centre, Nijmegen, The Netherlands 3Department of BioNanoPhotonics, IBEC-Institut de Bioenginyeria de Catalunya and CIBER BBN, Barcelona, Spain. 4Laboratory of Pediatric Oncology, NCMLS, Radboud University Nijmegen Medical Centre, Nijmegen, The Netherlands Chapter 6

Abstract Dendritic cells (DCs) are professional antigen presenting cells and play a key role in the initiation of adaptive immune responses. During their life DCs encounter and interact with very different tissue environments. How physical as well as chemical parameters of their microenvironment affect DCs, in particular cell-substrate adhesion, is largely unknown. Here, we investigate how DCs respond to changes in substrate composition, physicochemical properties or topology. We find that DCs adhere to and form specific adhesion structures, known as podosomes, on a variety of substrates. Clearly, hydrophobicity or substrate composition do not appear to have much effect on DC adhesion or podosome formation in vitro. To investigate the effects of substrate area dimensions on cell adhesion, surfaces consisting of permissive and non-permissive substrates are needed. We show that hydrogels are non-permissive for DC-adhesion and, in combination with microcontact printing, can be used to create substrate-areas with well-defined sizes. This should allow us to determine the minimal contact area required for efficient DC adhesion. Interestingly, we observed that podosome formation is strongly triggered by changes in substrate topology. Specifically, podosomes preferentially form on the edges of height patterns, suggesting that mechanical stress induced by the slopes of these patterns triggers the local formation of podosomes. Introduction

Dendritic cells (DCs) are the most potent antigen presenting cells of the immune system [1]. Immature DCs (iDCs) specialized in antigen recognition and uptake are adhesive, while antigen presenting mature DCs (mDCs) are highly migratory [2-5]. The adhesive and migratory properties are tightly regulated during the DC life cycle and dependent on interactions with elements of the micro-environment, such as the extracellular matrix (ECM). The ECM provides stability and is able to sequester growth factors and cytokines. The main components of the ECM are glycosaminoglycans often linked to proteins to form proteoglycans and fibrous proteins, such as fibronectin, collagen and laminin. DCs adhere to collagen and laminin, although fibronectin is the preferred substrate in vitro [6, 7]. Regulation of expression and activity of adhesion receptors is important for DCs [8- 12]. For example, DCs express the fibronectin binding integrins α4β 1 and α5β1, but the adhesion to fibronectin is exclusively α5β1-integrin dependent [3, 13] while the activity of this receptor is downregulated during DC maturation [8].

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In addition to regulating the activity of adhesion receptors and the spacing of ligands [14], the organization of adhesion receptors into specialized adhesion structures is important. When seeded onto 2- dimensional surfaces, most adherent cells form (integrin containing) adhesion structures, such as focal adhesions (FAs) or podosomes. FAs are relatively stable structures connected to actin stress fibers and present in a broad range of cell types [15, 16]. Podosomes are more dynamic structures [8, 17] found in Src-transformed cells, endothelial cells and cells of the myeloid lineage, such as osteoclasts, monocytes, macrophages and DCs [18, 19]. Podosomes have a characteristic morphology consisting of an actin core surrounded by a ring containing integrins and structural proteins, such as vinculin. Although FAs can also be found in iDCs, the main adhesion structures in these cells are podosomes [5, 15, 19]. We have shown earlier [8] that DC maturation involves the rapid dissolution of podosomes, which is presumably required to accommodate the high speed migration observed in these cells. Hence, regulation of cell- substrate adhesion and adhesion structures is carefully controlled during the DC life cycle. The adhesive properties of cells, including DCs, are influenced by intrinsic factors, such as differentiation or the contractile state of the cell, while external stimuli also affect adhesive behavior. It is becom- ing increasingly clear that also the physical aspects of the cellular environment can affect adhesive behavior. For instance, the flexibility of the substrate influences the adhesive properties of cells [20, 21]. In addition, also the hydrophobicity of the substrate is known to influence cell adhesion, with hydrophobic surfaces being less permissive for adhesion than hydrophilic surfaces [22]. However, how such aspects influence the adhesive properties of DCs is largely unknown. As DCs encounter a range of different tissue environments during their life cycle, we explored how DC adhesion is affected by substrate composition, - topology and - shape.

Results DCs form podosomes on different components of the ECM. Knowing that DCs form podosomes and FAs on fibronectin, we examined whether DCs are also able to form these structures on other ECM components or substrates. Therefore, iDCs were seeded on fibronectin, laminin or gelatin. Indeed we found that DCs generated podosomes as well as FAs on these ECM components (Figure 1A). In addition to these integrin substrates, iDCs were seeded on uncoated glass and poly-L-lysine (PLL), a positively-charged

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Figure 1. DCs form podosomes on different substrates (A/B) DCs were seeded on coverslips coated with fibronectin, gelatin or laminin, uncoated or PLL coated coverslips. The cells were fixed and stained with an anti-vinculin mAb (green) and phalloidin-Texas Red (red) to detect F-actin. (A) Representative images are depicted. (B) The number of cells displaying podosomes was counted in 7 images per condition per experiment and an average (with SD) of 2 experiments is shown.

aminoacid used to promote cell adhesion to poorly adhesive surfaces. Surprisingly, DCs not only attached but also formed prominent podosomes and FAs on glass and PLL (Figure 1A). Moreover, the amount of DCs able to form podosomes or FAs was similar on all substrates (Figure 1B and data not shown). Podosome formation was not affected by the presence or absence of serum in the medium (data not shown). These findings show that DCs can form podosomes as well as focal adhesions on 2-D surfaces irrespective of matrix composition or charge.

Hydrophobicity influences FA-formation, but not podosome formation in DCs. The hydrophobicity of a substrate affects the ability of cells to adhere with hydrophobic surfaces generally being non-permissive. To test the effect of hydrophobicity on DC adhesion, we tested a range of

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Figure 2. FA formation, but not adhesion and podosome formation, are regulated by substrate hydrophobicity (A/B) DCs were seeded on materials of different hydrophobicity. From hydrophobic to hydrophilic: Teflon, PS, PEN and PMMA. The cells were fixed and stained with an anti- vinculin mAb (green) and phalloidin-Texas Red (red) to detect F-actin. (A) Representative images are depicted. (B) The number of cells displaying podosomes or FAs was counted in 7 images per condition. The average amount of cells displaying podosomes or FAs (with SD) is shown. Significant differences from Teflon are indicated with asterisks (p<0.05). substrates with different hydrophobicity. From hydrophobic to more hydrophilic we used: Teflon, polystyrene (PS), poly(ethylene naphthalate) (PEN) and poly(methyl methacrylate) (PMMA). Surprisingly, the DCs attached and formed podosomes and FAs on each of these substrates (Figure 2A). The amount of cells that formed podosomes was comparable in all conditions, however FAs were much more frequently observed when cells were seeded on hydrophobic surfaces (Fig. 2B). These results indicate that hydrophobicity does not affect DC adhesion to 2-D surfaces per se, but may affect the type of adhesion structures formed.

Hydrogels are non-permissive for DCs, but upon spotting or microcontact printing become locally permissive allowing DCadhesion and podosome formation. In addition to hydrophobicity, we wanted to assess how the relative size of the adhesive surfaces may affect cell adhesion and the formation of specific adhesion structures in DCs. Therefore, a surface composed of adhesive and non-adhesive areas is needed. For DCs this is problematic as they adhere even to hydrophobic substrates, such as Teflon, which are non-permissive for most cell types. Hydrogels consist of a type of poly(ethylene glycol) (PEG) coating

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Figure 3. DCs adhere specifically to spotted and microcontact-printed areas of hydrogels (A) Solutions of fibronectin, PLL with PMA or PLL with zymosan-FITC (green) were spotted onto hydrogels. DCs were seeded onto these hydrogels and fixed and stained with an anti-vinculin mAb (green) (except when zymosan-FITC was present) and phalloidin-Texas Red (red) to detect F-actin. The edge of the spotted area is depicted with a dotted line. Representative images are depicted. (B) A solution of fibronectin with KLH-Alexa488 (green) was spotted onto hydrogels. DCs were seeded onto these hydrogels and fixed and stained with phalloidin-Texas Red (red) to detect F-actin. Representative images are depicted. (C) The method of microcontact printing. A PMMA- stamp with an array of dots of different size and spacing (gray) is incubated with the protein mixture to be printed (green). Excess solution is removed and the stamp covered with the remaining protein solution is placed onto the hydrogel (blue), which is coated on a glass slide (darker blue). The stamp can be removed and a pattern of printed protein is left on the hydrogel. DCs (red) can be seeded on these printed hydrogels. (D) DCs were seeded on the fibronectin/rIgG1-FITC (green) printed hydrogel and fixed and stained with phalloidin-Texas Red (red) to detect F-actin. A representative image with 25 µm dots is depicted in the left panel and representative images form 10 µm dots are depicted. A higher magnification of part of the image is shown in the upper left corner of the overlay.

110 DCs on micro-patterned substrates that prevents adhesion of cells. Therefore, we tested if DCs adhesion would be impaired on uncoated hydrogel. Indeed we found that DCs were unable to adhere to the uncoated hydrogel, but efficiently adhered to parts of the hydrogel spotted with fibronectin, PLL or PLL in combination with FITC-labeled zymosan (Figure 3A). Proteins and even big particles, such as the pathogen-mimic zymosan, can be absorbed onto hydrogels (Figure 3A). In addition, hydrogels were spotted with fibronectin mixed with Alexa488-labeled keyhole limpet hemocyanin (KLH). Similarly to the other proteins, this fluorescent mixture was absorbed onto the hydrogel. The fluorescence disap- peared specifically underneath podosome clusters (Figure 3B), suggesting that these proteins are degraded underneath podosomes. In addition to spotting proteins, we wanted to test whether microcontact printing would work on hydrogels, since this would allow a precise spacing of adhesive substrates. The process of microcontact printing is depicted in Figure 3C. A poly(dimethyl siloxane) (PDMA)-stamp, in our case with an array of different sized dots with different spacing, was incubated with (inking) fibronectin mixed with rIgG1-FITC for visualization. Excess protein solution is removed and the stamp covered with the fibronectin mixture can be put onto the hydrogel leav- ing a pattern of different sizes of fibronectin/rIgG1-FITC dots on the hydrogel. Then DCs can be added and left to adhere. Indeed, this method was highly effective as the DCs only formed podosomes on the well-defined printed areas (Figure 3D). These findings indicate that hydrogels are a promising tool to study effects of the size of the permissive area and the spacing thereof on adhesion and podosome formation.

DCs can sense and respond to changes in substrate topology. To explore potential effects of substrate topology on cell adhesion, we used substrates consisting of higher and lower parts of variable width. Teflon substrates with a height of 0.5 µm and width of 2, 5, 10 and 20 µm were generated. Surface topology was evaluated by atomic force microscopy (AFM), in order to confirm that the dimensions of the substrates resemble the expected height and width (Figure 4A). We observed that the DCs adhere to these substrates and the podosomes specifically aligned on the edges of these Teflon ridges (Figure 4B). Within these podosomes-alignments substruc- tures were still recognizable as individual podosomes. On the 2 µm wide substrates, a double row of podosomes was formed while very few podosomes were found on other parts of the substrate (Figure 4B left panel). When the distance between the Teflon ridges became wider than 2 µm, podosomes and FAs were observed on the flat

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areas. (Figure 4B). In addition to Teflon, the same effect was observed on PS, PEN and PMMA 3-D substrates (Figure 4C), con- firming that the material itself had little effect on podosome alignment. Furthermore, Teflon substrates of other heights were tested to investigate how DC-podosome formation was affected by height differences. Substrates containing ridges of 1 and 0.1 µm were tested in addition to the 0.5 µm high substrates used above. Alignment of podosomes at the edges of these substrates was observed in all cases (Figure 4D), showing that DCs can sense a height difference of 0.1 µm. In addition, the same effect was observed with PMMA 3- D substrates (data not shown), again showing that the type of mate-

Figure 4. DCs align podosomes on the angles of substrates with higher and lower parts (A) The topology and shape of Teflon substrates as determined by AFM. (B) DCs were seeded on Teflon substrates of 0.5 µm high and 2, 5, 10 or 20 µm wide. Cells were fixed and stained with an anti-vinculin mAb (green) and phalloidin-Texas Red (red) to detect F-actin. Representative images are depicted. (C) DCs were seeded on PS, PEN and PMMA substrates with different topologies. The cells were fixed and stained with an anti-vinculin mAb (green) and phalloidin-Texas Red (red) to detect F-actin. Representative images of the 10 µm wide lines are depicted. (D) DCs were seeded on Teflon substrates of 1 and 0.1 µm high. The cells were fixed and stained with an anti- vinculin mAb (green) and phalloidin-Texas Red (red) to detect F-actin. Representative images of 5 µm wide substrates are depicted.

112 DCs on micro-patterned substrates rial does not affect the ability of the DCs to respond to these changes in substrate topology. In addition to the ridges used above, other shapes were also tested. Rods of 5 by 2 µm with rounded edges were used in 2 orientations, octagons with squares of 25 µm on each side each and circles of 2 µm width and a diameter of 50 µm were scanned by AFM (Figure 5A). Also on these substrates DCs aligned podosomes on the edges (Figure 5B). In addition, we observed that the DCs maximized their contact area to the edges of the octagons with squares and circles by spreading along the edge. The podosome alignment observed on these substrates suggest that topography but not the shape of the substrate determines where podosomes are formed. Taken together, we postulate that the sharp angle at these topographic boundaries induces curvature of the cell membrane resulting in mechanical stress, which triggers podosome formation at these sites.

Figure 5. DCs align podosomes on differentially shaped substrates with high and low parts (A) The shape and topology of Teflon substrates determined by AFM scanning. Rods of 5 by 2 µm in 2 orientations, octagons with on each side squares of 25 µm and circles of 2 µm width and a diameter of 50 µm. (B) DCs were seeded on Teflon shapes. The cells were fixed and stained with an anti-vinculin mAb (green) and phalloidin-Texas Red (red) to detect F-actin. Representative images are depicted.

Discussion The adhesive properties of cells are influenced by cell intrinsic factors, but also by the type of substrate they encounter [14, 15, 23-25]. Here we show that ECM components support DC-adhesion, podosome- and FA-formation. In addition, DCs adhere and form podosomes and FAs on PLL and glass, showing that DCs can adhere and form adhesion structures on 2-D surfaces irrespective of the type of substrate. Whether integrins are involved in adhesion to

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glass and PLL is unclear at this point. However, it is likely that there is some role for integrins since integrin-containing adhesion structures, such as podosomes and FAs, are formed upon ligand binding by integrins. It would be interesting to investigate the involvement of integrins in the adhesion and formation of these structures on non- ECM components by using blocking antibodies. Hydrophobicity is one of the physical aspects known to influence cell adhesion. For instance, fibroblasts were shown to adhere best to the least hydrophobic substrates on a range of substrates with similar chemical properties but different hydrophobicity, [22]. In addition, hydrophobic substrates, such as Teflon, are often used to avoid cell adhesion. Surprisingly, DC adhesion is does not appear to be much affected by the hydrophobicity of the substrate. The number of cells with podosomes, the most prominent adhesion structure in DCs, is unaffected, although the number of cells displaying FAs increases somewhat on hydrophobic substrates. In addition to hydrophobicity, the size and spacing of the adhesive area can influence cell adhesion. To be able to investigate these properties, substrates consisting of adhesive and non-adhesive areas are needed. Teflon or other hydrophobic surfaces are often used as non-permissive substrate, however for DCs this is not feasi- ble as these cells can still efficiently adhere to these surfaces. Therefore, we have sought for alternative substrates. Hydrogels are designed for use in DNA and protein microarrays and consist of poly- mer with NHS-ester-groups that can react with amines [26]. These hydrogels show a very low non-specific binding thus minimizing background. Untreated hydrogels were non-adhesive even for DCs and can be used to produce a substrate containing adhesive (fibronectin or PLL) and non-adhesive (untreated) areas. These adhesive areas allow DC-adhesion and podosome formation. In addition, the disappearance fluorescently-labeled mixtures underneath podosomes, which are known to be sites of active matrix degradation [27-30], shows local degradation on hydrogels. Together, these data show the potential of hydrogels for studying DC adhesion on partially permissive substrates. To control the size and pattern of the adhesive part, microcontact printing is often used [31-33]. We tested microcontact printing on hydrogels and found that this method works well, providing us with the opportunity to start investigating the effect of substrate area size on adhesion and podosome formation. By applying stamps with sub- micron size patterns the minimal size required for individual podosomes to be formed could be determined. Another aspect that can influence cell adhesion is the topology of the

114 DCs on micro-patterned substrates substrate [23, 34]. To investigate this we used substrates in which ridges were introduced by hot embossing. Surprisingly, we observed that DCs aligned their podosomes at the topological boundaries on these substrates. Variations in the height of these ridges had no effect on the podosome alignment and we observed that DCs could sense a height difference of 100 nm. The preferred podosome-alignment on the topological boundaries, suggests that the angle of the substrate induces a curvature of the cell membrane resulting in mechanical stress leading to local podosome formation. It will be interesting to see if this podosome-alignment is conserved on substrates with different angles. By linking the ECM to the inside of the cell (cytoskeleton), integrins and integrin-containing adhesion structures play a key role in sensing mechanical stresses [25]. The regulation of basic cellular functions, such as differentiation, apoptosis, adhesion and migration, is dependent on the balance between forces generated by the ECM on the outside of the cells and forces generated by myosin II-mediated contraction inside the cell [15, 25]. Cells could sense external forces in different ways; stretching can lead to an altered organization of an adhesion structure, to conformational changes in components of the structures, or to opening of specific (stretch-activated) ion channels [24, 35-37]. It is well known that FAs increase in size in response to force and shrink upon relaxation [38]. As the size of FAs is propor- tional to the force, FAs are considered mechanosensing structures. The force influencing FAs can be generated externally, through applied force or shear stress [39-41], or by myosin II-mediated contraction [42-44]. An increase in myosin II-mediated contraction leads to the formation of FAs, while relaxation induces FA disassembly [15, 45, 46]. The effects of contractility on podosomes are less well understood. In addition to FA loss, relaxation of the cytoskeleton was shown to induces podosome formation in smooth muscle cells and neuroblastoma cells [45, 46]. Interestingly, we previously demonstrated that an increase in myosin II-mediated contractility, through activation of the RhoA-Rho kinase axis, leads to podosome loss in DCs [47]. This indicates that podosomes and FAs may be inversely regulated by contractility. The finding that contractility influences podosomes makes it likely that also forces generated outside the cell can influence these structures. The effects of substrate topography on podosome formation show that podosomes are regulated by and can respond to mechanical stress which suggests that podosomes, similar to focal adhesions, are mechanosensing structures. The external forces applied by the substrate on a cell may lead to opening or closing of (stress-activated) ion channels and this in turn could

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induce responses that lead to the local formation of a podosome. To our knowledge, this is the first report describing effects of mechanical stress on podosome formation. The mechanical effects on formation and regulation of podosomes could have implications for DC- behavior in different tissues and be important when DC cells encounter tissue boundaries (for instance when moving from the peripheral tissues into the lymph vessels). It would be interesting to see if the effects of physical aspects of substrates on podosomes in DCs are conserved between myeloid cells and other cells that form podosomes. Information on how physical aspects of substrates, especially those leading to mechanical stress, affect podosome formation/turnover will provide much needed insight into the function and regulation of these structures in DCs or other podosome-bearing cell types. Materials and methods Chemicals and antibodies The following antibodies were used: rIgG1-FITC (BD Bioscience Pharmingen, San Diego, Ca), anti-vinculin (Sigma, St. Louis, MO), Alexa Fluor 488-labeled secondary antibody (GaM) and Texas Red- conjugated phalloidin were from Molecular Probes (Molecular Probes, Leiden, The Netherlands). The following chemicals were used: fibronectin (Roche, Mannheim, Germany), gelatin, laminin and PLL (Sigma), PMA and KLH (Calbiochem, Merck, Nottingham, UK), Teflon, PS, PEN, and impact modified PMMA (Goodfellow Gmbh, Bad Nauheim, Germany). Zymosan (Sigma) was coupled to FITC (Sigma) and KLH was labeled with Alexa Fluor 488 labeling kit (GE Healthcare, General Electric Company, London, UK). Hydrogels are p-slides form Nexterion (Schott, Mainz, Germany).

Preparation of human DCs DCs were generated from PBMCs as described previously. Monocytes were derived from buffy coats or from a leukapheresis product. Plastic-adherent monocytes were cultured in RPMI 1640 medium (Gibco; Life Technologies, Breda, The Netherlands) supplemented with 10% (v/v) FCS (Greiner, Kremsmuenster, Austria), IL-4 (500 U/ml) and GM-CSF (800 U/ml). Immature DCs were harvested on day 6. Expression of MHC class I/II, costimulatory molecules and DC-specific markers on DCs were measured by flow cytometry (data not shown) and the expression of MHC molecules, costimulatory molecules and DC-markers was similar to what was described before [3]. Immature DCs expressed MHC class I and II, the costimulatory molecule CD86, the DC-specific marker CD209/DC-SIGN, low levels

116 DCs on micro-patterned substrates of the costimulatory molecule CD80 and lacked expression of the maturation marker CD83. Substrate preparation Different types of substrates: Coverslips were coated with fibronectin (20 µg/ml) in PBS for 1 hour at 37 C, gelatin (0.01% w/v) in PBS for 30 minutes at 37 C, laminin (20 µg/ml) in PBS for 1 hour at 37 C, PLL (100 µg/ml) in PBS for 30 minutes at 37 C or left untreated. Substrates with parts with different heights: The substrates with different heights of Teflon, PS, PEN and PMMA were made with hot embossing [48]. Hydrogel spotting: Drops (0.5 µl) of PBS with fibronectin (200 µg/ml), PLL (100 µg/ml) with 0.2 µM PMA (to enhance podosome formation), zymosan-FITC (6 mg/ml)/PLL (100 µg/ml) or KLH-Alexa488 (200 µg/ml)/fibronectin (200 µg/ml) were spotted on hydrogels. The spotted hydrogels were washed with PBS and 40000 DCs in 100 µl RPMI 1640 medium with cytokines were seeded. Microcontact printing: A silicon wafer was made with photolithogra- phy. PDMS Sylgard 184 silicone elastomer was mixed with a crosslinking agent containing a Pt-catalysator (both from Dow Corning, Midland, MI) in the ratio 10:1. Gas bubbles were removed in the exsiccator for approximately 30 min. The wafer was placed with the structured surface faced up and a few ml of the PDMS was poured onto it. The mixture was degassed again. Polymerization was achieved during incubation at 60-65°C for approx. 20 h. After polymerization, the stamp was peeled off the wafer. The stamps were incubated with fibronectin (100 µg/ml)/rIgG1-FITC (5 µg/ml) for 1 hour at RT in the dark. The solution is removed and the stamp is washed with mQ and dried with oil-free N2 using waferguard pistol (Millipore, Billerica, MA). The stamp is placed on the hydrogel and removed again. The printed area of the hydrogel is washed with 200 µl PBS and 9x104 DCs in 150 µl RPMI 1640 with cytokines were seeded.

Atomic force microscopy Surface topography was quantitatively evaluated using a Dimension atomic force microscope (AFM; Dimension 3100, Veeco, Santa Barbara, CA). Tapping in ambient air was performed with two types of cantilevers; for the lines and low patterns, a 118 ?m long high aspect ratio silicon cantilever (NW-AR5T-NCHR, NanoWorld AG, Wetzlar, Germany) was used, the rest of the patterns were analyzed with 100 µm long gold-coated silicon cantilevers (NSG-10, NT-MDT, Moscow, Russia). Both types have a nominal radius of curvature of the AFM probe tip less than 10 nm. Height images of each field/sam-

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ple were captured in ambient air at 50% humidity at tapping frequen- cies of 271 kHz and 261 kHz for the NSG-10 and NW-AR5T-NHCR cantilevers respectively. The analyzed field was scanned at a scan rates between 0.2-0.8 Hz and at a 512x256 or 512x512 scanning resolution. Nanoscope imaging software (version 6.13r1, Veeco) was used to analyze the resulting images.

Fluorescent microscopy DCs were seeded on substrates and left to adhere for 16 h. The cells were fixed in 3.7% (w/v) formaldehyde in PBS for 10 min. Cells were permeabilized in 0.1% (v/v) Triton X-100 in PBS for 5 min and blocked with 2% (w/v) BSA in PBS. The cells were incubated with primary Ab for 1 hour. Subsequently the cells were incubated with Alexa Fluor 488-labeled secondary antibodies for 45 min. Subsequently, cells were incubated with Texas Red-conjugated Phalloidin for 30 min. Images were collected on a Leica DMRA fluorescence microscope with a 63x PL APO 1.3 NA oil immersion lens (or a 40x PL FLUOTAR 1.0 NA oil immersion lens for overview images) and a COHU high performance integrating CCD camera (COHU, San Diego, CA). Pictures were analyzed with Leica Qfluoro version V1.2.0 (Leica) and Adobe Photoshop 7.0 (Adobe Systems, Mountain View, CA) software.

Statistical analysis ANOVA test was used for statistical analysis. Significant differences are indicated with asterisks (p<0.05). Acknowledgements We thank Dr. C. Mills (IBEC and Nanotechnology platform, Barcelona Science Park, Spain) for help with making the 3D substrates. We thank Dr. G. Roth (University of Tübingen, Germany) for making the silicon wafer needed to prepare the PDMS stamp. References 1. Banchereau, J. and R.M. Steinman, Dendritic cyte traffic through the analysis of chemokine cells and the control of immunity. Nature, receptor expression. Immunol Rev, 2000. 1998. 392(6673): p. 245-52. 177: p. 134-40. 2. Steinman, R.M., The dendritic cell system 5. Burns, S., et al., Maturation of DC is associ- and its role in immunogenicity. Annu Rev ated with changes in motile characteristics Immunol, 1991. 9: p. 271-96. and adherence. Cell Motil Cytoskeleton, 3. De Vries, I.J., et al., Effective migration of 2004. 57(2): p. 118-32. antigen-pulsed dendritic cells to lymph nodes 6. Sadovnikova, E., et al., Adhesion capacity in melanoma patients is determined by their and integrin expression by dendritic-like cells maturation state. Cancer Res, 2003. 63(1): p. generated from acute myeloid leukemia 12-7. blasts by calcium ionophore treatment. Exp 4. Sallusto, F. and A. Lanzavecchia, Hematol, 2004. 32(6): p. 563-70. Understanding dendritic cell and T-lympho- 7. Kohl, K., et al., Subpopulations of human

118 DCs on micro-patterned substrates

dendritic cells display a distinct phenotype range of wettability. J Biomed Mater Res B and bind differentially to proteins of the extra- Appl Biomater, 2007. 81(1): p. 66-75. cellular matrix. Eur J Cell Biol, 2007. 86(11- 23. Vogel, V. and M. Sheetz, Local force and 12): p. 719-30. geometry sensing regulate cell functions. Nat 8. van Helden, S.F., et al., A critical role for Rev Mol Cell Biol, 2006. 7(4): p. 265-75. prostaglandin E2 in podosome dissolution 24. Bershadsky, A.D., N.Q. Balaban, and B. and induction of high-speed migration during Geiger, Adhesion-dependent cell dendritic cell maturation. J Immunol, 2006. mechanosensitivity. Annu Rev Cell Dev Biol, 177(3): p. 1567-74. 2003. 19: p. 677-95. 9. DeMali, K.A., K. Wennerberg, and K. 25. Ingber, D.E., Cellular mechanotransduction: Burridge, Integrin signaling to the actin putting all the pieces together again. Faseb J, cytoskeleton. Curr Opin Cell Biol, 2003. 2006. 20(7): p. 811-27. 15(5): p. 572-82. 26. Kusnezow, W. and J.D. Hoheisel, Antibody 10. Garcia, A.J., J.E. Schwarzbauer, and D. microarrays: promises and problems. Boettiger, Distinct activation states of Biotechniques, 2002. Suppl: p. 14-23. alpha5beta1 integrin show differential binding 27. Burgstaller, G. and M. Gimona, Podosome- to RGD and synergy domains of fibronectin. mediated matrix resorption and cell motility in Biochemistry, 2002. 41(29): p. 9063-9. vascular smooth muscle cells. Am J Physiol 11. Huttenlocher, A., M.H. Ginsberg, and A.F. Heart Circ Physiol, 2005. 288(6): p. H3001-5. Horwitz, Modulation of cell migration by inte- 28. Calle, Y., et al., Inhibition of calpain stabilises grin-mediated cytoskeletal linkages and lig- podosomes and impairs dendritic cell motility. and-binding affinity. J Cell Biol, 1996. 134(6): J Cell Sci, 2006. 119(Pt 11): p. 2375-85. p. 1551-62. 29. Chellaiah, M., et al., Gelsolin deficiency 12. Semmrich, M., et al., Importance of integrin blocks podosome assembly and produces LFA-1 deactivation for the generation of increased bone mass and strength. J Cell immune responses. J Exp Med, 2005. Biol, 2000. 148(4): p. 665-78. 201(12): p. 1987-98. 30. Kanehisa, J., et al., A band of F-actin contain- 13. Jancic, C., et al., Interactions of dendritic cells ing podosomes is involved in bone resorption with fibronectin and endothelial cells. by osteoclasts. Bone, 1990. 11(4): p. 287-93. Immunology, 1998. 95(2): p. 283-90. 31. Gasteier, P., et al., Surface grafting of PEO- 14. Cavalcanti-Adam, E.A., et al., Lateral spacing based star-shaped molecules for bioanalyti- of integrin ligands influences cell spreading cal and biomedical applications. Macromol and focal adhesion assembly. Eur J Cell Biol, Biosci, 2007. 7(8): p. 1010-23. 2006. 85(3-4): p. 219-24. 32. Quist, A.P., E. Pavlovic, and S. Oscarsson, 15. Geiger, B. and A. Bershadsky, Exploring the Recent advances in microcontact printing. neighborhood: adhesion-coupled cell Anal Bioanal Chem, 2005. 381(3): p. 591- mechanosensors. Cell, 2002. 110(2): p. 139- 600. 42. 33. Lele, T.P., et al., Tools to study cell mechanics 16. Petit, V. and J.P. Thiery, Focal adhesions: and mechanotransduction. Methods Cell Biol, structure and dynamics. Biol Cell, 2000. 2007. 83: p. 443-72. 92(7): p. 477-94. 34. Thery, M., et al., Cell distribution of stress 17. Destaing, O., et al., Podosomes display actin fibres in response to the geometry of the turnover and dynamic self-organization in adhesive environment. Cell Motil osteoclasts expressing actin-green fluores- Cytoskeleton, 2006. 63(6): p. 341-55. cent protein. Mol Biol Cell, 2003. 14(2): p. 35. Matthews, B.D., et al., Cellular adaptation to 407-16. mechanical stress: role of integrins, Rho, 18. Buccione, R., J.D. Orth, and M.A. McNiven, cytoskeletal tension and mechanosensitive Foot and mouth: podosomes, invadopodia ion channels. J Cell Sci, 2006. 119(Pt 3): p. and circular dorsal ruffles. Nat Rev Mol Cell 508-18. Biol, 2004. 5(8): p. 647-57. 36. Hayakawa, K., H. Tatsumi, and M. Sokabe, 19. Linder, S. and M. Aepfelbacher, Podosomes: Actin stress fibers transmit and focus force to adhesion hot-spots of invasive cells. Trends activate mechanosensitive channels. J Cell Cell Biol, 2003. 13(7): p. 376-85. Sci, 2008. 121(Pt 4): p. 496-503. 20. Collin, O., et al., Spatiotemporal dynamics of 37. Chen, B.M. and A.D. Grinnell, Integrins and actin-rich adhesion microdomains: influence modulation of transmitter release from motor of substrate flexibility. J Cell Sci, 2006. 119(Pt nerve terminals by stretch. Science, 1995. 9): p. 1914-25. 269(5230): p. 1578-80. 21. Pelham, R.J., Jr. and Y. Wang, Cell locomo- 38. Galbraith, C.G., K.M. Yamada, and M.P. tion and focal adhesions are regulated by Sheetz, The relationship between force and substrate flexibility. Proc Natl Acad Sci U S A, focal complex development. J Cell Biol, 2002. 1997. 94(25): p. 13661-5. 159(4): p. 695-705. 22. Wei, J., et al., Adhesion of mouse fibroblasts 39. Kaverina, I., et al., Tensile stress stimulates on hexamethyldisiloxane surfaces with wide microtubule outgrowth in living cells. J Cell

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Sci, 2002. 115(Pt 11): p. 2283-91. 40. Riveline, D., et al., Focal contacts as mechanosensors: externally applied local mechanical force induces growth of focal contacts by an mDia1-dependent and ROCK- independent mechanism. J Cell Biol, 2001. 153(6): p. 1175-86. 41. Helmke, B.P. and P.F. Davies, The cytoskeleton under external fluid mechanical forces: hemodynamic forces acting on the endothelium. Ann Biomed Eng, 2002. 30(3): p. 284-96. 42. Balaban, N.Q., et al., Force and focal adhesion assembly: a close relationship studied using elastic micropatterned substrates. Nat Cell Biol, 2001. 3(5): p. 466-72. 43. Chrzanowska-Wodnicka, M. and K. Burridge, Rho-stimulated contractility drives the formation of stress fibers and focal adhesions. J Cell Biol, 1996. 133(6): p. 1403-15. 44. Zamir, E. and B. Geiger, Molecular complexi- ty and dynamics of cell-matrix adhesions. J Cell Sci, 2001. 114(Pt 20): p. 3583-90. 45. Clark, K., et al., TRPM7, a novel regulator of actomyosin contractility and cell adhesion. EMBO J, 2006. 25(2): p. 290-301. 46. Burgstaller, G. and M. Gimona, Actin cytoskeleton remodelling via local inhibition of contractility at discrete microdomains. J Cell Sci, 2004. 117(Pt 2): p. 223-31.

47. van Helden, S.F., et al., PGE2-mediated podosome loss in dendritic cells is dependent on actomyosin contraction downstream of the RhoA-Rho-kinase axis. J Cell Sci, 2008. 121(Pt 7): p. 1096-106. 48. Gallant, N.D., et al., Micro- and nano-patterned substrates to manipulate cell adhesion. J Nanosci Nanotechnol, 2007. 7(3): p. 803-7.

120 Chapter 7 General discussion and summary Chapter 7

Introduction The immune system plays a key role in our body's defense against infections. In addition, it can protect against cancer by recognizing and eradicating aberrant cells such as cancer cells. However, hyper- stimulation of the immune system can lead to autoimmune diseases and allergy. The innate immune system is an evolutionary conserved system that recognizes and eliminates invading pathogens [1]. In vertebrates a second line of defense is created by the adaptive immune system. The adaptive immune system is activated by antigen presenting cells (APCs) of the innate immune system, i.e. dendritic cells (DCs), macrophages and monocytes. The DC lifecycle involves a series of developmental stages. Precursors move from the blood into the tissue, were the immature DCs (iDCs) scan for antigen. When the iDCs encounter antigen they transform into mature DCs (mDCs) and migrate to the lymph nodes where they activate T cells. DCs are considered the most potent APC, as they are the most effective inducers of T cell activation [2, 3]. To fulfill these functions, DCs have to interact with different cellular environments depending on their life-stage, and these interactions have to be tightly regulated. For instance, the ability to cross tissue boundaries and to migrate to the lymph nodes, as it occurs in response to DC maturation, is essential for the induction of efficient immune responses [4, 5]. In this thesis, we investigated the regulation of adhesive and migratory properties during DC-maturation. DC cell lines Several difficulties arise when studying the biology of DCs. DCs are primary cell that have to be isolated from blood or spleen. An alternative is the isolation of DC-precursors, such as CD34+ cells or monocytes, and subsequent differentiation in vitro. The number of cells that can be obtained is limited, as is the life span of these cultures. In addition, expensive cytokines are needed and manipulation of DCs or DC-precursors is difficult. Additional complications arise from differences between donors. Therefore, a good DC-model cell line would be very useful. In chapter 2 we have compared different mouse and human cell lines for their use as a DC-model cell line. In the mouse setting, the D1 cell line most closely resembles in vivo DCs. However, when large numbers of cells or manipulation of these cells is required, the macrophage-like Raw264.7 cell line provides a better alternative. In the human setting, the MUTZ-3 cell line is the best model available thus far, especially when studying mature DC- function. Similarly to the D1 cells however, these cells are difficult to manipulate and have to be cultured under tightly controlled situa-

122 General discussion and summary tions. MUTZ-3 cells are poorly adhesive and therefore less suited as a model for DC adhesion. For these studies, THP-1 cells or HL-60 cells provide a good alternative. Although there is not one cell line that can fulfill all aspects of DC-biology, the use of combination of cell lines, each for a certain application, will contribute to our understanding of DC-biology. DC maturation and regulation of adhesion and migration In the tissues iDCs interact with the surrounding extracellular matrix (ECM) and it turns out that iDCs are able to bind to several components of the ECM. DCs bind the ECM component fibronectin through the α5β1-integrin [5]. During maturation strongly adherent iDCs change into highly migratory mDCs (chapter 3). This is reflected both by changes in the cytoskeleton of these cells as well as the activity of α5β1-integrin. While iDCs show prominent cell-substrate adhesions, in mDCs those structures are absent. Instead mDCs display characteristic dendritic extensions that give these cells their name. Although in iDCs focal adhesions (FAs) can be found, the main adhesion structures in these cells are podosomes. In chapter 3 we describe that the active form of the α5β1-integrin is enriched in podosomes and that the activity of this integrin is downregulated during maturation. Moreover, keeping the integrin in the active form prevents the loss of podosomes and the induction of migration, showing that a tight regulation of the activity of integrins is essential for the proper DC function. We observed that upon addition of a maturation-inducing combination of monocyte-conditioned medium (MCM), TNFα and prostaglandin E2 (PGE2) DCs show a rapid loss of podosomes (chapter 3). PGE2 can substitute for this combination and is sufficient to induce podosome loss and migration. By using live-cell imaging in combination with interference refection microscopy (IRM), we have investigated the dynamics of podosome loss in DCs (chapter 5). IRM allows one to image adhesion structures without a need for fluorescently-tagged molecules [6-8]. Structures close to the glass substrate appear dark with IRM, hence podosome appear as dark spots and FAs as dark stripes. Prior to dissolution of a podosome, a bright ring surrounding the podosome-core was observed, suggesting a local change in the proximity of the cell to its substrate. In addition to a loss of podosomes, PGE2 stimulation led to contraction of the DCs accompanied by an increase in the number of FAs (chapter 5). In other cell types, the reversed process, a loss of FAs accompanied by podosome formation, was shown [9, 10], suggesting that FAs

123 Chapter 7

and podosomes are oppositely regulated. After prolonged stimulation the number of DCs with FAs decreased again, with mDCs being completely devoid of FAs. Loss of FAs is associated with increased Rac1 activity [9] and may involve the microtubule network [11]. However, much remains unknown about the function and regulation of FA loss, especially during DC maturation. The dendritic extensions displayed by mDCs can be induced by CCL19, a chemokine that regulates migration to the lymph nodes [12]. Although the regulation of this process is largely unknown, the actin-bundling protein fascin and the Rho GTPases Rac and Cdc42 appear to be involved [13, 14]. However, detailed information on the mechanisms responsible for loss of FAs and subsequent induction of dendritic extensions during DC maturation is still lacking. Reactions to bacteria and TLR4 signaling in DCs The bacterial cell wall component lipopolysaccharide (LPS), a potent DC maturation stimulus, also induced podosome dissolution in DCs (chapter 3). However, this effect occurred much more slowly than

the PGE2-induced podosome loss. LPS is specifically found on Gram-negative bacteria and binds to Toll-like receptor 4 (TLR4) [15]. Components of Gram-positive bacteria activate TLR2, which dimer- izes either with TLR1 or TLR6 [16]. In chapter 4 we have compared the effects of Gram-negative and Gram-positive bacteria on DC-maturation. We observed that expression of costimulatory molecules as well as secretion of inflammatory cytokines is higher in response to Gram-negative bacteria than to Gram-positive bacteria. Moreover, Gram-negative bacteria were found to be more effective inducers of migration and expression of CCR7, the chemokine receptor required for CCL19/21-induced migration to the lymph nodes. Furthermore, only Gram-negative bacteria are able to induce podosome dissolution. These findings show that Gram-negative bacteria are superior to Gram-positive bacteria in inducing DC-maturation. An explanation for these difference could be that evolutionarily the number of Gram- negative bacteria infections led to the development of an APC dedi- cated to respond to this class of pathogens; the DC. Other APCs, such as monocytes and macrophages, may have evolved to coordinate immune responses against Gram-positive as well as Gram-negative bacteria. Indeed, macrophages effectively secrete cytokines, such as IL-6 and TNFα, in response to both classes of bacteria. At least our findings demonstrate that DCs discriminate between Gram- positive and Gram-negative bacteria. It will be interesting to see whether DCs also differentially respond to distinct classes of other pathogens, such as viruses.

124 General discussion and summary

Stimulation with purified ligands for TLR2 did not induce podosome loss. In addition, the amount of cytokines produced in response to TLR2 ligands was significantly lower than in response to LPS. This suggests that activation of TLR4 induces better DC activation than TLR2. Responses induced by LPS or Gram-negative bacteria could be efficiently blocked by TLR4-receptor antagonists, suggesting that these responses are fully dependent on TLR4 (chapter 4). By using mouse bone marrow derived DCs (BMDCs), we showed that these responses are conserved between mouse and men. Responses induced either by LPS or Gram-negative bacteria were absent in TLR4-/- BMDCs, which further confirms the TLR4 dependency of these responses (chapter 4). TLR4 can activate signaling pathways that act either downstream of the myeloid differentiation primary response gene 88 (MyD88) or the TIR domain-containing adaptor- inducing IFNβ (TRIF) adaptor [17, 18]. We used MyD88- [19] and TRIF-deficient [20] BMDCs to investigate the role of these signaling pathways (chapter 4). The effects of LPS or Gram-negative bacteria are mainly TRIF dependent, which is in line with the inability of TLR2 activation to induce potent DC maturation. However, MyD88 appears to also be involved, suggesting that activation of both signaling pathways is needed to induce full DC maturation.

The importance of PGE2 α The combination of MCM, TNF and PGE2, to induce DC maturation, is currently used in DC vaccination trials in melanoma patients. The presence of PGE2 was shown to be essential for the induction of migration and consequently efficient anti-tumor immune responses [5, 21-24]. This effect is partly mediated by the effects on expression and activation of CCR7, the chemokine receptor needed for migration to the lymph nodes [22, 23, 25]. Interestingly, we found that

PGE2 alone is sufficient to induce rapid podosome dissolution and migration in DCs (chapter 3). Therefore, we tested whether the effects of LPS or Gram-negative bacteria on DCs were dependent on the production of prostaglandins. LPS-induced podosome dissolution and the induction of migration could be inhibited by indomethacin, a pharmacological inhibitor that blocks the production of prostaglandins by inhibition of cyclooxygenase (COX) enzymes (chapter 3). Similarly, also the Gram-negative bacteria induced effect could be inhibited by using indomethacin (chapter 4). These findings reinforce the notion that the production of prostaglandins by DCs is important for the induction of an efficient immune response. At the moment it is unclear which prostaglandins are involved in these processes. PGE2 is very potent in inducing podosome loss and

125 Chapter 7

migration, both in vitro and in vivo, and is produced by DCs in response to LPS. However, the involvement of other prostaglandins cannot be ruled out at this point.

Given the importance of prostaglandins, in particular PGE2, in the regulation of adhesion (i.e. podosome dissolution) and migration during DC-maturation, we investigated the signaling pathways involved

in PGE2-mediated podosome dissolution (chapter 5). From the four receptors known for PGE2, EP1-4 [26], we detected EP2 and EP4 on DCs. PGE2-induced responses were mediated through both of these receptors. Activation of the EP2 and EP4 receptor induces signaling

through Gs to elevated cAMP levels [26] and these PGE2-induced responses can be mimicked by elevating cAMP levels. Cell shape and the organization of the cytoskeleton are influenced by numerous processes regulated by the family of small Rho GTPases [27-29].

Therefore, the role of RhoA, Rac1 and Cdc42 in PGE2-induced podosome loss was investigated (chapter 5). The active (GTP bound) form of RhoA increases when cells were stimulated with

PGE2, EP agonists or cAMP raising drugs. At the same time, levels of active Rac1 and Cdc42 decreased. This suggests that PGE2 induces podosome loss by activation of RhoA. In most cell types cAMP-mediated activation of PKA antagonizes the effects of RhoA on actomyosin contractility, leading to cytoskeletal relaxation [30-32].

In DCs however, PGE2 stimulation appears to induce a contractile response while activating RhoA. At the moment, it is not clear how the elevation of cAMP levels in DCs leads to activation of RhoA. Probably, PKA activation is involved, however activation of the EPAC-Rap1 pathway may also play a role [33]. We showed that the function of Rho kinase, a downstream effector of RhoA, is needed for

PGE2, EP agonist or cAMP elevation-induced podosome loss. Together our findings show that PGE2 induced podosome loss occurs by activation of the RhoA-Rho kinase axis. Podosome regulation: a balance of forces Activation of the RhoA-Rho kinase axis is known to promote the formation of FAs and induce cell contraction [27, 34, 35]. In addition, Rho kinase activity enhances stress fiber formation by inactivation of the actin severing protein cofilin [35], whereas direct activation of mDia by RhoA stimulates actin polymerization [27, 36]. Contraction of the cytoskeleton is generated through the function of the motor protein myosin II [37] and Rho kinase promotes phosphorylation of the myosin II regulatory light chain [38, 39]. Of the three myosin II isoforms (A, B and C) [40-42], myosin IIA is the predominant isoform expressed in DCs. The involvement of the RhoA-Rho kinase axis and

126 General discussion and summary

the contraction observed in DCs upon PGE2 stimulation, suggest a role for myosin II-mediated contraction in these responses. Indeed, by blocking myosin II with blebbistatin, a selective inhibitor of myosin IIA ATPase activity, we were able to show the importance of myosin

II-mediated contraction in PGE2-induced podosome loss (chapter 5). In addition, we found that the localization of myosin II changes dramatically upon PGE2 stimulation from a diffusely cytosolic to a more fibrillar distribution. These fibrillar structures colocalize with the bright rings observed by IRM, most likely reflecting an increased association with F-actin and/or a change in solubility of myosin filaments. In some cells, myosin II function was needed for podosome formation and maintenance [43, 44], whereas in other cells inhibition of myosin II function was important for podosome formation [9, 10]. We propose that basal myosin IIA activity is required for the formation and maintenance of podosomes, whereas a sudden (stimulus induced) increase in myosin II activity may trigger podosome dissolution. The regulation of basic cellular functions, such as differentiation, apoptosis, adhesion and migration, is dependent on the balance between forces generated by myosin II-mediated contraction inside the cell and forces generated by the ECM outside the cells [45, 46]. Integrins organized into adhesion structures form the link between the surrounding of the cell and the cytoskeleton. Physical properties of the substrate can influence the generation of forces outside the cell. Therefore, we investigated the effects of some of these physical aspects on DC-adhesion and podosome formation (chapter 6). The ability of DCs to adhere and form podosomes and FAs appears to be independent of the type or hydrophobicity of the substrate, although DCs more frequently form FAs on hydrophobic substrates. It will be interesting to determine, by using blocking antibodies, whether integrins are important for the adhesion and formation of podosomes and FAs on substrates other than ECM components. It has been suggested that the dynamic behavior of podosomes, rather than their size is affected by the substrate [44]. Therefore it will be important to determine whether the life-time of podosomes varies on those different substrates or whether the kinetics of podosome dissolution in response to PGE2 is affected by substrate composition. To investigate effects of spacing and size of substrate on DC adhesion characteristics, we searched for a substrate that is non-adhesive for DCs but that can be treated to locally allow DC-adhesion. We found that hydrogels form such a non-permissive substrate for DCs. Moreover, we found that (ECM) proteins or even large particles can be spotted onto these hydrogels leading to local DC-substrate interactions and podosome formation. Hydrogels can be used in combination with

127 Chapter 7

microcontact printing, which allows better controlled size and spacing. We conclude therefore that hydrogels, combined with microcontact printing, provide good tools to explore the role of substrate size and spacing on DC adhesion (chapter 6). In addition, we investigated the effects of topography on DC-adhesion and podosome formation by using substrates with ridges or other forms of height structures. Surprisingly, DCs aligned podosomes on the topological boundaries of these substrates (chapter 6). This suggests that the angle of the substrate induces a curvature of the cell membrane resulting in mechanical stress, giving rise to local podosome formation. By using pharmacological inhibitors, varying the angle of the ridges or by using a stretchable substrate, regulation of podosome formation by mechanical stretching could be further investigated. The profound effects of substrate topology on podosome formation show that podosomes are regulated by and can respond to mechanical stress, which suggests that podosomes, similar to focal adhesions [47], are mechanosensing structures. In line with this, we find zyxin, which has been suggested to be a mechanosensor [48, 49], in the podosome rings. The mechanical effects on formation and regulation of podosomes could have implications for DC-behavior in different tissues and be important when DC cells encounter tissue boundaries. Together, our findings suggest that podosomes, in addition to FAs, are regulated by the contractile state of the cell as well as the forces applied to the cell by the extracellular matrix. Interestingly, podosomes appear to be oppositely regulated by force when compared to FAs. While contraction induces FA formation and relaxation leads to a loss of these adhesions [9, 10, 34], (myosin II-based) cytoskeletal contraction leads to podosome loss while cytoskeletal relaxation promotes the formation of podosomes. Hence, such a balance of forces may be key to the regulation of cell adhesion and migration in DCs and consequently for the induction of adaptive immune responses. The function of podosomes in DCs Although there is a time gap between the loss of podosomes (minutes) and the induction of migration (16 hours), we see a strong correlation between the loss of podosomes and migration (chapter 3 and 4). This suggests that podosomes prevent the induction of this high-speed, T cell-like migration needed for proper functioning of mDCs. Dissolution of podosomes is a tightly regulated process. By fixing DCs at the moment of bright ring formation and subsequent staining

128 General discussion and summary

Figure 1. Model of the morphology of podosomes In the upper left an IRM image of a DC with podosomes (dark spots) is depicted. In the upper right the same DC is stained for vinculin (green) and actin (red). An enlargement of a podosome is depicted in the lower right corner of this image. In the lower part of the figure a schematic model of a podosome is shown. A podosome consists of an actin- rich core (red) connected to a surrounding ring structure (green). In the core actin polymerization takes place. Here, the actin nucleation complex Arp2/3 and its regulator WASp, as well as the actin crosslinking protein α-actinin and the actin-bundling protein cortactin are localized. Integrins binding their extracellular ligands, such as ECM molecules, are found in the ring structure. Also the integrin binding proteins talin and paxillin, the talin- and paxillin-binding protein vinculin, zyxin and myosin II are found there. In podosomes integrins are connected to the actin cytoskeleton, mediating adhesion and migration. for various podosome components, we found that vinculin could no longer be detected at the time that the actin core was still largely intact (Chapter 5). At this time, zyxin was still present in the podosome-ring, while myosin II redistributes to fibrillar-like structures. We conclude that loss of podosomes involves the sequential loss of podosome components. Podosome formation probably

129 Chapter 7

involves a similar sequential acquisition of podosome components and detailed information on the acquisition or loss of components can shed more light on the function of podosomes. Although the role of podosomes in iDCs is still somewhat elusive, some suggestions can be made. Similar to FAs, podosomes link integrins to the actin cytoskeleton. Podosomes have a very distinct morphology with a core that contains actin and actin binding proteins surrounded by a ring that contains adaptor molecules and integrins (Figure 1). In addition, they are highly dynamic adhesion structures with a life-time of 2 to 12 minutes that often localize to the leading edge of the iDCs. Therefore, it is likely that podosomes are important for adhesion, the formation of polarity and directional crawling, resembling slow, fibroblast-like migration. This could facilitate the scavenging for antigens by iDCs in the tissues. In addition, local protein degradation takes place underneath podosomes as the result of focalized release of matrix metalloproteases [50, 51]. Although the adhesion-mediating integrins are found in the podosome ring, the dark spots observed by IRM correspond in size and localization to the actin-rich cores of podosomes, suggesting that the actin-rich core is closer to the substrate. Actin polymerization in the core could drive protrusion towards or into the substrate [52-54]. Moreover podosomes have been suggested to palpate the substrate [54] and they have been implicated in leukocyte transmigration [6]. Together, these observations suggest that podosomes allow DCs to either enter or leave the peripheral tissues and help the DCs to move from the tissue into a lymph vessel. Concluding remarks In this thesis we show the importance of adhesion regulation during DC-maturation and more specifically the need for these cells to dissolve podosomes in order to acquire migratory properties. Gram- negative bacteria are more capable of inducing DC maturation relative to Gram-positive bacteria. This difference is due to differential

activation of TLR4. Prostaglandins such as PGE2 are key mediators of DC maturation in response to either pathogen-derived molecules such as LPS or Gram-negative bacteria. A model showing TLR4 acti-

vation and subsequent PGE2 signaling is depicted in Figure 2. PGE2- induced podosome dissolution, which either involves the EP2 or the EP4 receptor, raises cAMP levels. This increase in cAMP leads to activation of the RhoA-Rho kinase pathway and consequently an increase in myosin II-mediated contraction. This contractile response leads to a rapid loss of podosomes and the (transient) formation of FA. In conclusion, our results show that regulation of the actomyosin

130 General discussion and summary

Figure 2. Regulation of podosomes loss during DC maturation Stimulation with LPS or Gram-negative bacteria activates TLR4, MyD88 and TRIF dependent signaling, resulting in COX-mediated production of prostaglandins, such as

PGE2. PGE2 activates the EP2 and EP4 receptors, resulting in cAMP elevation and PKA activation. This leads to activation of RhoA and inactivation of Rac1 and Cdc42. Active RhoA induces Rho kinase activation. Activation of the RhoA-Rho kinase axis induces myosin II-mediated contraction, which results in podosome loss and temporary FA formation. cytoskeleton, mediated by Rho GTPases in response to extracellular signals, plays an important role during the transition from a (tissue resident) iDC to a highly motile mDC. References 1. Greenberg, S. and S. Grinstein, Phagocytosis 16(23): p. 2337-44. and innate immunity. Curr Opin Immunol, 7. Kaverina, I., T.E. Stradal, and M. Gimona, 2002. 14(1): p. 136-45. Podosome formation in cultured A7r5 vascu- 2. Banchereau, J. and R.M. Steinman, Dendritic lar smooth muscle cells requires Arp2/3- cells and the control of immunity. Nature, dependent de-novo actin polymerization at 1998. 392(6673): p. 245-52. discrete microdomains. J Cell Sci, 2003. 3. Mellman, I., S.J. Turley, and R.M. Steinman, 116(Pt 24): p. 4915-24. Antigen processing for amateurs and profes- 8. Usson, Y., et al., Quantitation of cell-matrix sionals. Trends Cell Biol, 1998. 8(6): p. 231- adhesion using confocal image analysis of 7. focal contact associated proteins and interfer- 4. Eggert, A.A., et al., Analysis of dendritic cell ence reflection microscopy. Cytometry, 1997. trafficking using EGFP-transgenic mice. 28(4): p. 298-304. Immunol Lett, 2003. 89(1): p. 17-24. 9. Burgstaller, G. and M. Gimona, Actin 5. De Vries, I.J., et al., Effective migration of cytoskeleton remodeling via local inhibition of antigen-pulsed dendritic cells to lymph nodes contractility at discrete microdomains. J Cell in melanoma patients is determined by their Sci, 2004. 117(Pt 2): p. 223-31. maturation state. Cancer Res, 2003. 63(1): p. 10. Clark, K., et al., TRPM7, a novel regulator of 12-7. actomyosin contractility and cell adhesion. 6. Chou, H.C., et al., WIP regulates the stability EMBO J, 2006. 25(2): p. 290-301. and localization of WASP to podosomes in 11. Broussard, J.A., D.J. Webb, and I. Kaverina, migrating dendritic cells. Curr Biol, 2006. Asymmetric focal adhesion disassembly in

131 Chapter 7

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protein antigens: role of PGE2 in chemokine member of the myosin II family. J Biol Chem, and cytokine expression by MoDCs. Int 2004. 279(4): p. 2800-8. Immunol, 2005. 17(12): p. 1561-72. 41. Katsuragawa, Y., et al., Two distinct nonmus-

132 General discussion and summary

cle myosin-heavy-chain mRNAs are differentially expressed in various chicken tissues. Identification of a novel gene family of vertebrate non-sarcomeric myosin heavy chains. Eur J Biochem, 1989. 184(3): p. 611-6. 42. Shohet, R.V., et al., Cloning of the cDNA encoding the myosin heavy chain of a vertebrate cellular myosin. Proc Natl Acad Sci U S A, 1989. 86(20): p. 7726-30. 43. Kopp, P., et al., The kinesin KIF1C and microtubule plus ends regulate podosome dynamics in macrophages. Mol Biol Cell, 2006. 17(6): p. 2811-23. 44. Collin, O., et al., Spatiotemporal dynamics of actin-rich adhesion microdomains: influence of substrate flexibility. J Cell Sci, 2006. 119(Pt 9): p. 1914-25. 45. Geiger, B. and A. Bershadsky, Exploring the neighborhood: adhesion-coupled cell mechanosensors. Cell, 2002. 110(2): p. 139- 42. 46. Ingber, D.E., Cellular mechanotransduction: putting all the pieces together again. Faseb J, 2006. 20(7): p. 811-27. 47. Galbraith, C.G., K.M. Yamada, and M.P. Sheetz, The relationship between force and focal complex development. J Cell Biol, 2002. 159(4): p. 695-705. 48. Lele, T.P., et al., Mechanical forces alter zyxin unbinding kinetics within focal adhesions of living cells. J Cell Physiol, 2006. 207(1): p. 187-94. 49. Yoshigi, M., et al., Mechanical force mobilizes zyxin from focal adhesions to actin filaments and regulates cytoskeletal reinforcement. J Cell Biol, 2005. 171(2): p. 209-15. 50. Burgstaller, G. and M. Gimona, Podosome- mediated matrix resorption and cell motility in vascular smooth muscle cells. Am J Physiol Heart Circ Physiol, 2005. 288(6): p. H3001-5. 51. Osiak, A.E., G. Zenner, and S. Linder, Subconfluent endothelial cells form podosomes downstream of cytokine and RhoGTPase signaling. Exp Cell Res, 2005. 307(2): p. 342-53. 52. Destaing, O., et al., Podosomes display actin turnover and dynamic self-organization in osteoclasts expressing actin-green fluorescent protein. Mol Biol Cell, 2003. 14(2): p. 407-16. 53. Calle, Y., et al., Inhibition of calpain stabilises podosomes and impairs dendritic cell motility. J Cell Sci, 2006. 119(Pt 11): p. 2375-85. 54. Carman, C.V., et al., Transcellular diapedesis is initiated by invasive podosomes. Immunity, 2007. 26(6): p. 784-97.

133 Chapter 7

134 Nederlandse samenvatting Samenvatting (Summary in Dutch)

Het immuunsysteem speelt een essentiële rol in de bescherming tegen infecties. Misregulatie van het immuunsysteem kan leiden tot aanhoudende infecties, kanker, auto-immuunziekten en allergie. Het aangeboren immuunsysteem is een evolutionair geconserveerd sys- teem dat pathogenen herkent en opruimt. Een tweede verdedi- gingslijn wordt gevormd door het adaptieve immuunsysteem, dat geactiveerd wordt door antigen presenterende cellen (APCs) van het aangeboren immuunsysteem; dendritische cellen (DCs), macrofagen en monocyten. DCs zijn de sterkste APCs als het gaat om het activeren van T cellen. Tijdens hun leven doorlopen DCs meerdere fasen waarin ze verschillende functies uitoefenen. DC voorloper cellen verplaatsen zich van het bloed naar de weefsels, waar de onvolwassen DCs (iDCs) patrouilleren op zoek naar antigenen. Na interactie met zulke antigenen ondergaan de DCs een maturatiepro- ces waardoor iDCs veranderen in volwassen DCs (mDCs) die mi- greren naar de lymfeknopen waar de T cellen geactiveerd worden. Deze migratie vanuit de weefsels naar de lymfeknopen is van groot belang voor het induceren van een productieve immuun respons. In dit proefschrift is de regulatie van adhesieve en migratoire eigen- schappen tijdens DC-maturatie onderzocht. Het onderzoek naar de biologie van DCs wordt bemoeilijkt door een aantal factoren. DCs zijn primaire cellen die moeten worden ge- ïsoleerd uit het bloed of de milt. Een alternatief hiervoor is het iso- leren van voorloper cellen, zoals CD34+ cellen of monocyten, om die vervolgens te differentiëren tot DCs. Voor deze methode is het gebruik van dure cytokinen vereist. Het aantal cellen dat op deze manier verkregen kan worden is beperkt, net als de het tijdsbestek waarin deze kweken gebruikt kunnen worden. Verder is het (genetisch) manipuleren van (voorlopers van) DCs erg moeilijk. Extra complicaties ontstaan door verschillen tussen donoren. Om de com- plexe biologie van DCs beter te kunnen bestuderen zou een goede model DC-cellijn erg nuttig zijn. Daarom hebben wij verschillende muizen - en humane cellijnen vergeleken voor het gebruik als model DC-cellijn (hoofdstuk 2). Voor de muis is de D1 cellijn degene die de meeste overeenkomsten met DCs vertoont. Deze cellijn is evenwel moeilijk (genetisch) te manipuleren en groeit langzaam. De (macro- faag achtige) Raw264.7 cellijn is een goed alternatief wanneer cel- manipulatie of grote aantallen nodig zijn. Van de humane cellijnen lijkt op dit moment de MUTZ-3 cellijn het beste model om de biologie van (vooral volwassen) DCs te bestuderen. Helaas zijn ook deze cellen moeilijk genetisch te manipuleren en moeten deze onder strikt

136 Nederlandse samenvatting gecontroleerde omstandigheden gekweekt worden. Verder hechten deze cellen relatief slecht waardoor ze minder geschikt zijn voor studies naar DC-adhesie. Voor dit laatste vormen de THP-1 of HL-60 cellen een goed alternatief. Samenvattend kunnen we concluderen dat er niet één cellijn is die in alle aspecten op DCs lijkt. Desalniettemin kan het gebruikt van cellijnen, elk voor een eigen toepassing, het onderzoek naar de biologie van DCs vergemakke- lijken, zoals ook in dit proefschrift beschreven. Tijdens maturatie veranderen de sterk hechtende iDCs in snel mi- grerende mDCs (hoofdstuk 3). Dit proces gaat gepaard met veran- deringen in het cytoskelet van deze cellen. mDCs hebben dendritische uitlopers, maar geen adhesiestructuren. iDCs hebben z.g.n. 'focal adhesions' (FAs), maar de belangrijkste adhesiestructuren in deze cellen zijn podosomen. Wij hebben ontdekt dat DCs na toevoeging van een maturatie-inducerende combinatie, bestaande uit monocyt-geconditioneerd medium (MCM), TNF α en prostaglan- dine E2 (PGE2), zeer snel hun podosomen verliezen (hoofdstuk 3). Deze combinatie wordt ook gebruikt bij patiënten met huidkanker die

DC-vaccinatie ondergaan. Hierbij is gebleken dat PGE2 onmisbaar is voor het induceren van migratie en goede antitumor responsen.

Onze in vitro experimenten bevestigen dat de toevoeging van PGE2 voldoende is om podosoomdissolutie en snelle migratie te induceren in DCs. In de weefsels komen iDCs in contact met de omringende extra- cellulaire matrix (ECM) waarbij deze aan verschillende ECM componenten kunnen binden. In vitro heeft binding aan fibronectine via het α5β1-integrine de voorkeur. Wij hebben aangetoond dat actief α5β1- integrine verrijkt is in podosomen en dat de activiteit van dit integrine vermindert tijdens maturatie (hoofdstuk 3). Als het integrine in zijn actieve vorm gehouden wordt remt dit het oplossen van podosomen en het ontstaan van snelle migratie. Dit wijst erop dat een strikte regulatie van de activiteit van integrines essentieel is voor het functioneren van DCs. Hoewel het oplossen van podosomen (minuten) en het ontstaan van snelle migratie (16 uur) in de tijd van elkaar gescheiden zijn, zien we een sterke correlatie tussen het verdwijnen van podosomen en het ontstaan van snelle migratie. Waarschijnlijk zijn podosomen van belang bij de langzame, fibroblast-achtige migratie van iDCs, maar verhinderen ze de snelle, T cel-achtige migratie noodzakelijk voor het functioneren van mDCs. Ook hebben we gevonden dat lipopolysaccharide (LPS), een maturatie stimulus afkomstig van de celwand van gram negatieve bac- teriën ook podosoomdissolutie in DCs kan induceren. Dit effect

137 treedt evenwel beduidend later op dan met PGE2. LPS bindt en activeert Toll-like receptor 4 (TLR4). Componenten van Grampositieve bacteriën leiden tot activatie van TLR2, dat dimeren vormt met TLR1 of TLR6. Wij hebben de effecten van Gramnegatieve en Grampositieve bacteriën op DC-maturatie in detail onderzocht (hoofdstuk 4). Tot onze verbazing vonden we dat Gramnegatieve bacteriën of LPS, maar niet Grampositieve bacteriën of pure TLR2 liganden, podosoomdissolutie induceren. Verder blijken de Gramnegatieve bacteriën superieur in het induceren van snelle migratie en expressie van CCR7, de chemokine receptor nodig voor de migratie naar de lymfeknopen. Ook de expressie van costimulatoire moleculen en productie van inflammatoire cytokinen is beduidend hoger na stimulatie van DCs met Gramnegatieve bac- teriën dan wanneer hiervoor Grampositieve bacteriën worden gebruikt. Deze bevindingen tonen aan dat Gramnegatieve bacteriën beter zijn in het induceren van DC-maturatie dan Grampositieve bac- teriën. De effecten van LPS of Gramnegatieve bacteriën kunnen worden geblokkeerd door TLR4 te blokkeren and zijn afwezig in TLR4-/- muize beenmerg DCs (BMDCs), wat aangeeft dat dit proces volledig afhankelijk is van TLR4 en dat deze reacties geconserveerd zijn tussen muis en mens (hoofdstuk 4). TLR4 kan twee signaleringsroutes activeren, de ene verlopend via het adapter molecuul MyD88 en de andere via TRIF. Wij hebben MyD88- en TRIF-defi- ciënte BMDCs gebruikt om de rol van deze signaleringsroutes op DC maturatie verder te onderzoeken (hoofdstuk 4). Uit deze studies blijkt dat de effecten van Gramnegatieve bacteriën op maturatie en podosoom verlies, in hoofdzaak TRIF afhankelijk zijn.

Omdat PGE2 stimulatie leidt tot het snel oplossen van podosomen, is onderzocht of de effecten van LPS danwel Gramnegatieve bacteriën op DCs afhankelijk zijn van de productie van prostaglandines. Het door LPS geïnduceerde podosoomverlies en de snelle migratie blijkt te kunnen worden geremd met indomethacine, een middel dat de productie van prostaglandinen door de COX-enzymen remt (hoofdstuk 3). Op dezelfde manier kunnen de effecten geïnduceerd door Gramnegatieve bacteriën geremd worden met indomethacine (hoofdstuk 4). Deze resultaten bevestigen dat prostaglandines, geproduceerd door DCs na stimulatie met bacteriële antigenen, een belangrijke rol spelen bij de inductie van cel migratie tijdens DC maturatie.

Gezien het belang van prostaglandines zoals PGE2 voor DC migratie, zijn de signaleringsroutes betrokken bij de PGE2-geïn- duceerde podosoomdissolutie verder onderzocht (hoofdstuk 5). Van

138 Nederlandse samenvatting

de vier bekende receptoren voor PGE2, EP1-4, blijken vooral EP2 en EP4 betrokken bij PGE2-geinduceerde responsen in DCs. Activatie van EP2 en EP4 leidt tot productie van cAMP en het (pharmacolo- gisch) verhogen van cAMP concentraties in de cel kan de PGE2- geïnduceerde effecten nabootsen. Door 'Live Cell Imaging' te combineren met interferentie reflectie microscopie (IRM), zijn we in staat geweest om de dynamiek van podosoomverlies in tijd te volgen (hoofdstuk 5). IRM maakt het mogelijk om adhesiestructuren in levende cellen te volgen in de tijd zonder gebruik te maken van fluorescent-gelabelde moleculen. Gebruik makend van deze techniek worden podosomen zichtbaar als donkere stippen en FAs als donkere strepen. Behalve tot het verlies van podosomen, leidt PGE2 stimulatie ook tot een toename in het aantal DCs met FAs (hoofdstuk 5). Direct voorafgaand aan het oplossen van de podosomen ontstaat een heldere ring rondom de podosoom kern, hetgeen suggereert dat er een fysische verandering plaats vindt in deze ring structuur. FA formatie en het verlies van podosomen gaat gepaard met contractie (het samentrekken van het cytoskelet) van de DCs. Contractie van het cytoskelet wordt gegenereerd door het motoreiwit myosine II in associatie met het actine cytoskelet. Van de drie vormen van myosine II (A, B en C) komt hoofdzakelijk myosine IIA voor in DCs. Door gebruik te maken van de myosine II remmer blebbistatine hebben we het belang van myosine II(A) in PGE2-geïnduceerde podosoomdissolutie verder kunnen bevestigen (hoofdstuk 5). De vorm van de cel en de organisatie van het cytoskelet worden door verschillende processen beïnvloed, en in belangrijke mate gereguleerd door de GTPases RhoA, Rac1 en CDC42. Daarom is de betrokkenheid van deze GTPases bij de PGE2-geïnduceerde effecten op podosomen onderzocht (hoofdstuk 5). Hieruit blijkt dat activatie van RhoA end RhoA-gemedieerde contractie een essentiële rol speelt bij PGE2-geïnduceerd podosoomverlies. Uit deze resultaten blijkt dat PGE2 stimulatie , leidt tot cAMP verhoging, wat het verlies van podosomen en ontstaan van FAs induceert. Deze effecten zijn een direct gevolg van myosine II geïnduceerde contractie die ontstaat door activatie van de RhoA-Rho kinase route (hoofdstuk 5). Verder suggereren deze bevindingen dat podosomen en FAs tegengesteld gereguleerd worden door de contractiele status van de cel. De regulatie van elementaire cellulaire functies, zoals differentiatie, apoptose, adhesie en migratie, is afhankelijk van de balans tussen krachten die gegenereerd worden door de ECM aan de buitenkant van de cel en de krachten die ontstaan door myosine II geïnduceerde

139 contractie in de cel. Integrines die georganiseerd zijn in adhesie structuren vormen de verbinding tussen de omgeving van de cel en het cytoskelet. Fysische aspecten van het substraat en de cellulaire omgeving kunnen het ontstaan van krachten binnen in de cel beïn- vloeden. In hoofdstuk 6 hebben we de effecten van deze fysische aspecten op DC-adhesie en het ontstaan van podosomen onderzocht. Het vermogen van DCs om te hechten en om podosomen en FAs te vormen op tweedimensionale substraten lijkt onafhankelijk van het type substraat of het hydrofobe karakter ervan. Om de effecten van topologie en omvang van het substraat oppervlak te onderzoeken is gezocht naar een substraat waarop de DCs niet kunnen hechten, maar dat zodanig behandeld kan worden dat de DCs locaal wel kunnen hechten. Hydrogels blijken aan deze eigen- schappen te voldoen. Eiwitten en zelf grotere deeltjes geïmmo- biliseerd op deze hydrogels geven aanleiding tot DC-adhesie en podosoom vorming. Hydrogels kunnen worden gebruikt in combinatie met microcontact printing, wat een meer gecontroleerde af- meting en plaatsing van de eiwit-gecoate opervlakken toelaat. Het gebruik van hydrogels in combinatie met microcontact printing lijkt dan ook een goede methode om de rol van substraat grootte en af- stand te onderzoeken (hoofdstuk 6). Ook is het effect van reliëfverschillen in het substraat op DC-adhesie en podosoomvorming onderzocht gebruikmakend van substraten met richels en anders gevormde hoogtestructuren. DCs vormen specifiek podosomen op de randen van deze hoogtestructuren (hoofdstuk 6). Dit suggereert dat de hoek van het substraat een kromming van de celmembraan induceert wat resulteert in mecha- nische stress die leidt tot locale podosoomformatie. De effecten van reliëfverschillen op podosoomformatie suggereren dat podosomen mechanosensoren kunnen zijn, net als FAs. De invloed van mecha- nische effecten op podosoomvorming en -regulatie zou gevolgen kunnen hebben op het gedrag van DCs in de verschillende weefsels en tijdens het verlaten van de weefsels. Samenvattend tonen de bevindingen in dit proefschrift aan dat adhesiestructuren (met name podosomen) strikt gereguleerd worden tijden DC-maturatie. Deze regulatie speelt een belangrijke rol bij de inductie van snelle migratie vanuit de weefsels naar de lymfeklieren, een voorwaarde voor goede T cel activatie.

140 List of abbreviations

List of abbreviations

-/- knock out Ab antibody AFM atomic force microscopy APC antigen presenting cell BMDCs bone marrow-derived DCs BSA bovine serum albumin cAMP cyclic adenosine monophosphate CBA cytokine beads array COX cyclooxygenase CTL cytotoxic T lymphocyte DCs dendritic cells DC-SIGN DC-specific ICAM-3 grabbing nonintegrin DEAE diethylaminoethyl DNA desoxyribonucleic acid ECM extracellular matrix ELISA enzyme-linked immunosorbent assay FAs focal adhesions FCS fetal calf serum FITC fluorescein isothiocyanate FN fibronectin g gram GM-CSF Granulocyte Macrophage-Colony Stimulating Factor GTP Guanosine triphosphate h hour HLA human leukocyte antigen ICAM intercellular adhesion molecule iDCs immature DCs IFN interferon IL interleukin IRF IFN-regulatory factor IRM Interference Reflection Microscopy KLH keyhole limpet hemocyanin l liter LPS lipopolysaccharide LTA lipoteichoic acid MAL MyD88 adaptor-like MCM monocyte conditioned medium MDA-5 melanoma differentiation-associated gene 5 mDCs mature DCs MFI mean fluorescent intensity MHC major histocompatibility complex

141 List of abbreviations

min minutes MODCs monocyte-derived DCs moi mode of infection MyD88 myeloid differentiation primary response gene 88 myDCs myeloid DCs MΦ macrophage NF-κB nuclear factor κB NK cell natural killer cell NOD nucleotide-binding oligomerization domain PAMPs pathogen associated molecular patterns PBMCs peripheral blood mononuclear cells PBS phosphate buffered saline Pd pull down pDCs plasmacytoid DCs PDMA poly(dimethyl siloxane) PE phycoerythrin PEG poly(ethylene glycol) PEN poly(ethylene naphthalate)

PGE2 prostaglandin E2 PGs prostaglandins PKA protein kinase A PLL poly-L-lysine PMA phorbol 12-mystate 13-acetate PMMA poly(methyl methacrylate) polyI:C polyinosinic-polycytidylic acid PRRs pattern recognition receptors PS polystyrene RIG retinoic acid-inducible gene RNA ribonucleic acid RT-PCR reverse transcriptase polymerase chain reaction SD (s.d.) standard deviation SEM (s.e.m.) standard error of the mean Th T helper TL total lysate TLRs Toll-like receptors TNFα Tumor Necrosis Factor α TRAM TRIF-related adaptor molecule TRIF TIR domain-containing adaptor-inducing IFN WASp Wiskott Aldrich Syndrome protein WT wild-type

142 Dankwoord

Dankwoord

Nu is het dan zover, mijn proefschrift is af. Tijdens mijn verblijf op het TIL waren er veel mensen die hebben bijgedragen aan het tot stand komen van dit boekje en deze mensen wil ik hier graag bedanken.

Als eerste wil ik natuurlijk mijn promotor Carl Figdor en mijn copromotor Frank van Leeuwen bedanken voor jullie steun en begeleiding. Beste Carl, bedankt voor de kans om bij jou promotieonderzoek te mogen doen. Zelfs met je (te) drukke agenda had je vaak goede suggesties en (wilde) ideeën voor deze eigenwijze AIO. Ik heb veel van en door je geleerd en ik vind het erg leuk dat je de podosoomlijn wilt voortzetten. Beste Frank, bedankt voor al je goede adviezen en tips. Ik kon altijd even bij je binnen lopen ('Nu even niet!') en later hebben we aardig wat tijd aan de telefoon gehangen. Je enthousiasme voor celbiologie en mooie microscopieplaatjes bleek erg aanstekelijk. Veel plezier en succes met je eigen club.

Dan mijn paranimfen, Sophia en Machteld. Sophia, voor jou was het geen verrassing dat ik je als paranimf naast me wilde, want die afspraak hadden we eigenlijk al een jaar of vijf geleden gemaakt. Sinds we beiden naar Nijmegen kwamen zijn we vriendinnen en hebben inmiddels al heel wat meegemaakt samen. Tijdens onze theedrinksessies en vele etentjes kunnen we het over alles hebben (ook al gaat het wel vaak over wetenschap). Jammer dat jij net weg ging uit Amsterdam nu ik er werk. Veel plezier en succes in Londen straks, die stad moeten we samen met onze mannen maar eens goed verkennen! Machteld, je begon bij het TIL en kreeg de taak mij te ondersteunen bij het afronden van mijn promotieonderzoek (ook al hadden we elkaar niet gesproken voor je begon). Onze samenwerking was erg prettig en we kunnen lekker kletsen. Je hebt vele uren in de kweek en achter de Leica gezeten, wat met wat stevige muziek uiteindelijk best vol te houden bleek. Je hebt me enorm geholpen en ik vind het daarom erg leuk dat je mijn paranimf bent. Veel plezier met floorball en ijshockey!

Niels bedankt voor de RT-PCRs en Nannette voor je hulp in de kweek. Daniëlle en Karin B voor de hulp met het migratiesysteem. Erik B en Roger bedankt voor de hulp bij alle FACS problemen. Gosse, Liz, Stefan en Martijn bedankt voor alle hulp met (vooral het

143 Dankwoord

bureaucratische deel van) de proeven met de TLR-, MyD88- en TRIF-deficiënte muizen. Joost te R, Martijn en Arthur bedankt voor alle hulp met computerissues. Ben en Frank de L bedankt voor alle hulp (en creatieve naamgeving) met de confocaal. Joost te R bedankt voor de AFM metingen op de patterns. Louise en Jeanette voor de secretariële ondersteuning. Jolanda en iedereen van de patiëntengroep voor de vele 'left over' DCs en Gerty en Nicole voor de CBA. En iedereen van de adhesie-DLM voor input en suggesties.

Natuurlijk zijn er ook mensen van buiten het TIL betrokken bij het tot stand komen van dit proefschrift. The physics connection with Enschede and Barcelona; Maria Garcia-Parajo, Gert-Jan Bakker, Ruth Diez-Ahedo, Niek van Hulst, Marjolein Mutsaers, Vinod Subramaniam, Hans Kanger, Mathilde de Dood and Marieke Snijder- van As, thanks for al the discussions en suggestions. Maria, Ruth and Chris thanks for the patterns. The language of biology and physics is very different and this gave some misunderstandings, but I learned a lot from it. My initial project didn't work out the way we hoped, but in the end with the patterning I turned back to the bio- physics! Roland Brock, Heike Glauner en Michael Sinzinger, thank you for the pleasant collaboration concerning the hyrogels. Roland bedankt dat je in mijn manuscriptcommissie wilde zitten, ik vind het ongelooflijk hoe snel je Nederlands hebt geleerd. Mihai Netea en Reinier Raymakers, bedankt voor jullie enthousiasme en klinische input, Hoofdstuk 4 wordt vast nog een mooie pu- blicatie! Inneke Verschueren bedankt voor de hulp met de muizen, Liesbeth Jacobs en Trees Jansen bedankt voor de cytokineme- tingen. Leo Joosten bedankt voor de TRIF-deficiënte muizen.

Samen met een aantal andere TILlers heb ik een flinke tijd op de 6e verdieping doorgebracht. Alessandra, Ben, Joost, Agnieszka, Machteld, 'bezoeker' Inge en op het laatst Koen, ik vond dat we een erg gezellig groepje hadden samen. Ook wil ik alle celbiologen bedanken voor de goede sfeer en leuke borrels e.d., het was erg gezellig. Ik vond het erg leuk om mijn data ook aan een celbiologisch publiek te kunnen presenteren. Bé, bedankt dat je in mijn manuscriptcommissie wilde zitten en voor je leuke suggesties. Jack, bedankt voor je enthousiasme en hulp met de IRM experimenten, jammer dat die TIRF microscoop pas kwam toen ik op hield. Huub, bedankt voor alle hulp met de confocaal en de Leica, alle liters mowiol en de gezelligheid.

144 General introduction

Behalve het direct bijdragen aan experimenten en dit proefschrift, wil ik ook iedereen verder bedanken voor de fantastische sfeer op het lab. De mensen waar ik naast heb gezeten waren onmisbaar om goede en slechte resultaten mee te delen. Robbert, als student bij jou begon mijn verblijf op het TIL en ik heb veel van je geleerd en zelfs even kennis kunnen maken met het 'muizenwerk'. Annemiek de B, het was altijd erg gezellig om samen in de kweek te zitten. Frank de L bedankt alle gesprekken voor microscopie, fysica, klussen en hypotheken. Patricia, je bent een vrolijk persoon, met een dipje tijdens het gevecht met de erfenis van Gerard. Cândida, (met dakje want anders is het de schimmel) ik vergeet die enorme BBQ met stofzuiger-aan-blazer denk ik niet snel. Veel plezier verder down under! Maaike van H, was kort, maar wel leuk. Inge R, met jou kletsen is altijd erg gezellig en het blowkarten was gaaf! En Alessandra en Ben, bedankt voor alle gesprekken, de steun en gezelligheid, tot binnenkort in Amersfoort.

Verder wil ik alle TILlers bedanken voor de fijne tijd die ik heb gehad. Een aantal mensen en groepjes wil ik nog even apart noemen. The outdoor-borrelcommissie; Karlijn, Ben, Joost te R en nu Jori. Catanclubje; Ben, Martijn, Karlijn en Daniëlle. BOMmen met Cândida, Karlijn, Alessandra en Friederike. Ladies-4 diner; Alessandra, Aukje en Daniëlle. Anna, Agnieszka, Annemiek van S, Alessandra, Aukje, Bas, Ben, Cândida, Dagmar (binnenkort gaan we thee drinken of lekker eten), Daniel (visiting Barcelona was great! Good luck there), Daniëlle, Erik B, Frank W (ik ben benieuwd wat er met de data van Sip gebeurd), Friederike (Utrecht is nu een stuk dichterbij), Inge B, Ingrid, Jori, Joost te R, Karlijn (nu wonen we al lekker dicht bij elkaar, als jullie in je nieuwe huis gaan wordt het zelf nog beter!), Koen (leuk dat je verder gaat met de podosomen, succes!), Kris (fellow AIO of Frank, your favorite indian restaurant has also become one of our favorites. Enjoy Scotland), Liza, Maaike L, Machteld, Mandy, Martijn, Matthijs, Nicole, Robbert, Roger, Sonja, Stefan, Twan (leuk dat we coauteurs zijn), Vassillis, Veronique.

Inmiddels werk ik al een tijdje op de afdeling MCB van Sanquin. Peter, bedankt dat je in mijn manuscriptcommissie wilde zitten en voor deze leuke baan. En alle MCBers wil ik bedanken voor de leuke tijd deze eerste maanden!

145 Chapter 1

Verder wil ik de 'Brabantse groep' bedanken voor alle gezelligheid en relativering van het promoveren. Kim, we woonden op dezelfde gang en zijn daarna vriendinnen gebleven, ondanks een flinke reistijd, bedankt.

Natuurlijk wil ik op deze plek ook mijn ouders bedanken voor alle steun en vertrouwen. Ook wil ik mijn schoonfamilie bedanken voor alle interesse en steun.

Lieve Stan, jou wil ik het meest bedanken van iedereen. Bedankt voor je hulp bij het lay-outen van dit boekje en voor het lezen van al die immunologische verhalen. Maar vooral voor je vertrouwen, steun, en bovenal onze liefde. Ik vind het heerlijk om je vrouw te zijn!

146 Curriculum Vitae

Curriculum Vitae

Suzanne van Helden werd geboren op 6 juli 1979 te Horst. In 1997 behaalde zij haar VWO-diploma aan de Philips van Horne scholengemeenschap te Weert. Daarna studeerde ze Biologie aan de Radboud Universiteit Nijmegen (toen nog Katholieke Universiteit Nijmegen). Tijdens deze studie liep zij stage bij de afdeling Moleculaire Dierfysiologie (prof. dr. G.J. Martens) van de Radboud Universiteit Nijmegen en een tweede stage bij de afdeling Tumor Immunologie (prof. dr. C.G. Figdor) van het Radboud Universiteit Nijmegen Medisch Centrum. In november 2002 slaagde zij voor het doctoraal examen en begon in december 2002 als Junior Onderzoeker op de afdeling Tumor Immunologie van het Radboud Universiteit Nijmegen Medisch Centrum bij prof. dr. C.G. Figdor en dr. F.N. van Leeuwen. De resultaten van dat promotieonderzoek staan beschreven in dit proefschrift. Sinds juni 2008 is ze werkzaam als postdoc bij de afdeling Moleculaire Celbiologie van Sanquin research en Landsteiner laboratorium te Amsterdam.

147

List of publications

List of Publications

Cambi A, van Helden SFG, and Figdor CG. (2005) Roles for Integrins and Integrin-associated Proteins in the Haematopoietic System. In: Integrins in Development, ed. EHJ Danen, Landes Bioscience, Texas. van Helden SFG, Krooshoop DJEB, Broers KCM, Raymakers RAP,

Figdor CG and van Leeuwen FN. A critical role for PGE2 in podosome dissolution and induction of high-speed migration during dendritic cell maturation. J Immunol. 2006;177(3):1567-74. van Helden SFG, Oud MM, Joosten B, Peterse N, Figdor CG and van Leeuwen FN. PGE2-mediated podosome loss in dendritic cells is dependent on actomyosin contraction downstream of the RhoA-Rho kinase axis. J Cell Sci. 2008;121 (Pt7):1096-106. van Helden SFG, van Leeuwen FN and Figdor CG. Human and murine model cell lines for dendritic cell biology evaluated. Immunol Lett. 2008;117(2):191-7. van Lieshout AW, van der Voort R, Toonen LW, van Helden SFG, Figdor CG, van Riel PL, Radstake TR and Adema GJ. Regulation of CXCL16 expression and secretion by myeloid cells is not altered in rheumatoid arthritis. Ann Rheum Dis. 2008 Jul 15. van Helden SFG, Oud MM, Raymakers RAP, Netea MG, van Leeuwen FN and Figdor CG. TLR4-mediated discrimination between Gram-negative and Gram-positive bacteria directs migration and maturation of dendritic cells. Submitted.

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