complement factor h dysfunction in atypical hemolytic - helda

Academic Dissertation

Complement factor H dysfunction in atypical hemolytic uremic syndrome

Markus Lehtinen

Department of Bacteriology and Immunology

Haartman Institute

University of Helsinki

Finland

To be publicly discussed with permission of the Medical Faculty of the University of Helsinki, in the Small Auditorium of the Haartman Institute on March 18th, 2011, at 12 o’clock noon.

Supervisor:

T. Sakari Jokiranta MD, PhD, Adjunct Professor Department of Bacteriology and Immunology Haartman Institute University of Helsinki

Reviewers:

Olli Lassila MD, PhD, Professor Department of Medical Microbiology and Immunology University of Turku

Sarah Butcher PhD, Professor Institute of Biotechnology University of Helsinki

Opponent:

Anna Blom PhD, Professor Department of Laboratory Medicine University of Lund

©2011 by Markus Lehtinen

Printed at Unigrafia OY

ISBN 978-952-92-8586-0 (paperback)

ISBN 978-952-10-6817-1 (pdf)

To my family Minna, Iina, and Enna

Contents

Original publications ............................................................................................................................... 7

Abbreviations .......................................................................................................................................... 8

Abstract ................................................................................................................................................. 10

1 Review of the literature ................................................................................................................ 12

1.1 Main functions of the complement system .......................................................................... 12

1.2 Activation of the complement system .................................................................................. 12

1.3 Amplification and terminal pathway of complement ........................................................... 15

1.4 Regulation of the complement system ................................................................................. 16

1.5 Complement in innate immune response ............................................................................ 22

1.6 Complement in adaptive immunity ...................................................................................... 24

1.7 Complement in cell death ..................................................................................................... 25

1.8 Crosstalk of complement with coagulation system .............................................................. 27

1.9 Atypical hemolytic uremic syndrome ................................................................................... 29

2 Aims of the study .......................................................................................................................... 32

3 Materials and Methods ................................................................................................................. 33

3.1 Materials ............................................................................................................................... 33

3.2 Methods ................................................................................................................................ 33

4 Results ........................................................................................................................................... 39

4.1 Rational mutagenesis of FH19-20 ......................................................................................... 39

4.2 Self - non-self discrimination by FH19-20 ............................................................................. 48

4.3 aHUS mutations disturb target recognition function of FH19-20 by various mechanisms .. 52

5 Discussion...................................................................................................................................... 56

5.1 Sequence and structure analysis in mutagenesis of FH19-20 .............................................. 56

5.2 Self – non-self discrimination by FH19-20 ............................................................................ 56

5.3 Molecular pathogenesis of aHUS .......................................................................................... 58

5.4 Dysfunction of FH19-20 in aHUS ........................................................................................... 58

6 Acknowledgments ......................................................................................................................... 61

7 References .................................................................................................................................... 64

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Original publications

I Lehtinen MJ, Meri S, and Jokiranta TS. Interdomain contact regions and angles between adjacent short consensus repeat domains. Journal of Molecular Biology. 344(5):1385-96, 2004.

II Jokiranta TS, Jaakola VP, Lehtinen MJ, Pärepalo M, Meri S, and Goldman A. Structure of complement factor H carboxyl-terminus reveals molecular basis of atypical haemolytic uremic syndrome. EMBO Journal. 25(8):1784-94, 2006.

III Lehtinen MJ, Rops AL, Isenman DE, van der Vlag J, and Jokiranta TS. Mutations of factor H impair regulation of surface-bound C3b by three mechanisms in atypical hemolytic uremic syndrome. Journal of Biological Chemistry. 284(23):15650-8, 2009.

IV Kajander T, Lehtinen MJ, Hyvärinen S, Bhattacharjee A, Leung E, Isenman DE, Meri S, Goldman A, and JokirantaTS. Dual interaction of factor H with C3d and glycosaminoglycans in host-nonhost discrimination by complement. Proceedings of the National Academy of Sciences USA. Early Edition Feb 1, 2011.

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Abbreviations

Ab Antibody aHUS Atypical hemolytic uremic syndrome AP Alternative pathway BCR B cell receptor C Complement C1-Inh C1 inhibitor C1qR C1q receptor C3aR C3a receptor C4bp C4 binding protein C5aR C5a receptor CA Cofactor activity CD Cluster of differentiation CP Classical pathway CR Complement receptor CRP C-reactive protein CS-A Chondroitin sulfate A DAA Decay accelerating activity DAF Decay accelerating factor (CD55) DC Dendritic cell DDD Dense deposit disease EC Endothelial cell Esh Sheep erythrocyte F Factor FB Factor B FcγR Fc gamma receptor FD Factor D FH Factor H FHL-1 Factor H like protein 1 FHR Factor H related protein FI Factor I GAG Glycosaminoglycan GBM Glomerular basement membrane GEnC Glomerular endothelial cell GlcNAc N-acetyl-glucosamine HS Heparan sulfate IC Immune complex iC3b Inactive C3b iC4b Inactive C4b ICR Interdomain contact region Ig Immunoglobulin IL Interleukin LP Lectin pathway LPS Lipopolysaccharide MAC Membrane attack complex MASP MBL-associated serine protease MBL Mannan binding lectin MCP Membrane cofactor protein (CD46) mGEnC mouse glomerular endothelial cell

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NF-kappa B Nuclear factor kappa B NMR Nuclear magnetic resonance P Properdin PBS Phosphate buffered saline PS Phosphatidylserine PTX3 Long pentraxin 3 RBC Red blood cell RCA Regulators of complement activation RLA Radioligand assay SA Sialic acid SAP Serum amyloid P component SCR Short consensus repeat domain SDS-PAGE Sodium dodecyl sulfate – polyacrylamide gel electrophoresis SPR Surface Plasmon resonance TD T-cell dependent TGF-β Transforming growth factor β TF Tissue factor THBD Thrombomodulin TI T-cell independent TLR Toll-like receptor TMB Tetramethylbenzidine TNF-α Tumor necrosis factor α vWF von Willebrand factor Å Ångström (0.1 nm)

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Abstract

Complement is a group of plasma proteins that recognizes and attacks invading microbes leading to opsonization, lysis, and induction of inflammation. It also facilitates clearance of apoptotic and necrotic cells by opsonization and enhances B-cell responses to antigens. Complement is initiated via different activation pathways that converge at the C3 step, leading to C3b deposition on the target. C3b deposition initiates the amplification of the complement that leads to release of anaphylatoxins and to the formation of lytic membrane attack complexes (MAC). One of the complement activation pathways - the alternative pathway (AP) - can in principle activate on any surface, self or non-self, and is therefore tightly regulated by the endogenous regulatory proteins. In atypical hemolytic uremic syndrome (aHUS) the AP regulation on self surfaces is insufficient and leads to complement attack against erythrocytes, platelets, and endothelial cells resulting usually in end-stage renal disease. In this disease, impaired regulation of the AP activation and complement amplification result from either point mutations in AP proteins or autoantibodies that interfere with their function.

The AP protein that is most often dysfunctional in aHUS is factor H (FH). FH is the main regulator of AP activation in plasma, and on the self surfaces it is one of the key regulators inhibiting AP activation and complement amplification. FH is composed of 20 homologous short consensus repeat domains (SCRs) that are also found in many other complement regulators. The N-terminal domains 1-4 (FH1-4) mediate together with factor I the inactivation of C3b on the self surfaces. The C-terminal domains 19 and 20 (FH19-20) are critical for the ability of FH to discriminate between C3b opsonized self and non-self surfaces. Self surfaces are rich in anionic sialic acids (SA) and glycosaminoglycans (GAGs) such as heparan sulfate that are not usually present in non-pathogenic microbes. FH19-20 contains binding sites for both the C3d part of C3b and self surface polyanions that enhances avidity of FH to C3b on self surfaces and thus C3b inactivation. Factor H mutations that have been described from aHUS patients segregate to FH19-20 and impair self surface recognition by FH.

The aims were to study rational mutagenesis of SCR domains, molecular mechanism of FH19-20 mediated self – non-self discrimination and to investigate why point mutations in FH19-20 lead to aHUS. To address these questions crystallographic structures of FH19-20 and FH19-20 in complex with C3d (FH19-20:C3d) were solved and rational mutagenesis to FH19-20 was planned based on sequence and structure analysis of SCRs and FH19-20. Point mutations that had aHUS association or were located in the FH19-20-C3d or heparin binding interfaces were introduced into recombinant FH19-20. Functional defects caused by these mutations were studied by analyzing the binding of the FH19-20 mutant proteins to C3d, C3b, heparin, and mouse glomerular endothelial cells (mGEnCs) using radioligand assays, affinity chromatography, enzyme immunoassays, and surface plasmon resonance.

The results revealed two independent binding interfaces between FH19-20 and C3d - the FH19 site and the FH20 site. Superimposition of the FH19-20:C3d complex on the previously published C3b and FH1-4:C3b structures showed that the FH20 site on C3d was partially occluded, but the FH19 site was fully available. Furthermore, the binding of FH19-20 via the FH19 site to C3b did not block the binding of the functionally important FH1-4 domains and kept the FH20 site free. The results of

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structural analysis also showed that the simultaneous binding of the FH19-20 via the FH19 site to C3b and via the FH20 site to C3d was possible.

It was shown by binding assays that when FH19-20 was bound to C3b via the FH19 site, FH20 domain could bind to heparin. In addition the heparin binding and mGEnC binding sites on FH20 were found to be nearly identical indicating that the FH20 mediates binding to cells and heparin similarly. It was further shown that FH19 site and FH20 were both necessary to recognize non-activator cell model of self-surfaces. These findings indicate that simultaneous binding of FH19 site to C3b and FH20 to anionic self structures are the key interactions in self surface recognition by FH and explains how FH discriminates between self and non-self.

The aHUS associated mutations on FH19-20 were found to disrupt binding of the FH19 or FH20 site to C3d/C3b, or to disrupt binding of FH20 to heparin or mGEnC. Any of these dysfunctions leads to loss of FH avidity to C3b bearing self surfaces explaining the molecular pathogenesis of aHUS cases where mutations are found within FH19-20.

In conclusion, the results have revealed the molecular mechanism of FH19-20 mediated self – non-self discrimination by FH. It is based on simultaneous binding of FH19 site to C3b and binding of FH20 to anionic self structures that increases FH avidity to C3b and enhances downregulation of complement activation. The aHUS-associated mutations in FH19-20 disrupt the self – non-self discrimination mechanism of FH that can lead to thrombocytopenia, microangiopathic hemolytic anemia, and acute renal failure - the triad of typical characteristics in aHUS. Understanding of the molecular pathogenesis of aHUS will help in designing correct treatment for the patients and in developing new therapeutic approaches for aHUS.

Review of the literature

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1 Review of the literature

1.1 Main functions of the complement system

The complement system is composed of about 35 plasma or membrane proteins that form a cascade-like activation, amplification, effector, and control system (Müller-Eberhard 1988). During an infection, microbes trigger complement system via lectin pathway (LP), classical pathway (CP), and alternative pathway (AP) that all converge to the amplification of the cascade and opsonization of the microbe with C3b. Opsonization facilitates phagocytosis and stimulates B-cell responses, and the amplification leads to release of anaphylatoxins that attract and activate the cells of the innate immune system (Hugli et al. 1978; Fearon et al. 2000; Köhl 2006). The end point of the complement cascade is the formation of membrane attack complex (MAC) that can insert into target membranes and lead to lysis of the microbe (Cole et al. 2003).

Another major function of complement is to facilitate clearance of apoptotic and necrotic cells (Poon et al. 2010). Complement activation leads to opsonization of the non-viable cells, but the activation is restricted and does not lead to lysis of self cells and generation of heavy inflammatory response. In addition complement facilitates clearance of immune complexes (IC) from circulation by solubilizing and transporting them to the reticuloendothelial system (Schifferli et al. 1986).

1.2 Activation of the complement system

Complement is continuously monitoring blood and lymph for the presence of its targets e.g. microbes and necrotic cells. The targets are recognized mainly by three activation pathways CP, LP, and AP. Recent results have suggested that properdin (P) may also act as a target-recognition molecule leading to complement activation (Hourcade 2006). These pathways recognize different structures on the targets but all lead to C3b deposition on the surface (Figure 1). Complement can also be activated extrinsically by coagulation system proteases such as thrombin, which has been shown to cleave C3 (Spath et al. 1976; Huber-Lang et al. 2006).

Figure 1. Schematic presentation of the activation of complement cascade. After recognizing the target all the three pathways converge to covalent attachment of C3b on the surface.


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1.2.1 Alternative pathway AP activation requires only the plasma proteins factor B (FB), factor D (FD), and C3 to initiate the attack on any surface (Figure 2) (Schreiber et al. 1978). The activation is not really triggered but is rather continuous by spontaneous conversion of C3 to metastable C3* that reacts with water and forms C3b like C3(H2O)(Lachmann et al. 1975; Pangburn et al. 1983). C3(H2O) binds FB and forms a C3(H2O)B complex that is a target for protease FD (Pangburn et al. 1981). FD cleaves FB in a complex that leads to release of Ba fragment and to formation of the enzyme C3(H2O)Bb. The Bb fragment is attached transiently in C3(H2O)Bb that has a half-life of 90 seconds restricting the operation time of the enzyme and complement activation (Medicus et al. 1976; Pangburn et al. 1986). Properdin (P) can act as positive regulator of complement by binding to C3(H2O)Bb that stabilizes the enzyme (Fearon et al. 1975). C3H2OBb is a C3 convertase enzyme that cleaves C3 to C3a and to metastable C3b* that has a short-lived and highly reactive thioester moiety (Law et al. 1977; Sim et al. 1981). Importantly, C3b* can react with –OH or –NH groups that are universally present in almost every biological macromolecule, giving explanation to the versatile recognition capability of the AP (Pangburn et al. 1980a; Pangburn et al. 1980b). A covalent bond is formed between C3b and the target that directs the downstream complement cascade events to that site.

Figure 2. Alternative pathway activation. Spontaneous hydrolysis of C3 to C3* (box 1) initiates the formation of C3(H2O)Bb that converts C3 to metastable C3b* (box 2). C3b* can attach to surface or react with water and form fluid phase C3b(H2O)Bb that can cleave more C3 (box 3). Properdin has been omitted from the figure for clarity. The dotted arrows indicate enzymatic activity.

1.2.2 Classical pathway The action of the AP is complemented by CP and LP that utilize pattern recognition molecules to identify their targets. The CP is initiated by the collectin C1q that recognizes multiple ligands like lipid A that is part of lipopolysaccharide (LPS) on microbial surfaces (Loos et al. 1974). CP is activated indirectly by binding of C1q to clusters of IgG1 or IgG3, or IgM on microbial surfaces or in immune complexes. CP is also activated on dying cells that are opsonized with C-reactive protein (CRP) or


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display phosphatidylserine, chromatin, and/or DNA on their surface that are all C1q ligands (Agnello et al. 1970; Claus et al. 1977; Robey et al. 1985; Paidassi et al. 2008).

C1q monomer is composed of collagen-like tail and fibrinogen like head domain that form a hexameric C1q that resembles a bunch of tulips (will be referred as C1q) (Strang et al. 1982). C1q is bound to two C1r and to two C1s proteases that together form C1 complex (C1qr2s2) in plasma or on the target (Gigli et al. 1976). Binding of C1q to the target auto-activates C1r that in turn activates C1s (Sim et al. 1977). Active C1s enzyme cleaves plasma C4 to C4a and C4b (Sim et al. 1977). C4a that is released to plasma in proteolysis is not anaphylatoxic and is not known to have any other function either (Gorski et al. 1979; Meuer et al. 1981). C4b on the other hand has highly reactive and short-lived thioester (like C3b) that mediates covalent binding of C4b to the target (Dodds et al. 1996). C4b acts as an acceptor for C2 that is cleaved by C1r to fragments C2a and C2b (Sim et al. 1977). The complex C4b2b (C4b2a in old nomenclature), i.e. the classical pathway C3 convertase, cleaves plasma C3 to C3b and C3a (Cooper 1975).

1.2.3 Lectin pathway The LP is initiated by mannan binding lectin (MBL), or one of the L-, H-, or M-ficolins that are homologous to C1q (Thiel 2007). LP recognizes microbial sugar structures, like lipoteichoic acid (LTA) and peptidoglycan of gram positive bacteria, GlcNAc and GalNAc of gram negative bacteria, and mannose/mannan in yeast cell walls, leading to complement attack (Ikeda et al. 1987; Lynch et al. 2004; Matsushita 2009; Zhang et al. 2009). It also recognizes DNA and C-reactive protein (CRP) on the surfaces of dying cells and facilitates their clearance (Claus et al. 1977; Ng et al. 2007). The functions of the CP and LP are thus similar, but the ligands on the targets are different, and importantly only the CP recognizes antibodies (Wallis et al. 2010).

MBL or one of the ficolins forms a complex with one of the MBL-associated serine proteases 1, 2, or 3 (MASPs) of which MASP-2 has been shown to cleave C4 and C2 and to deposit C3b on the target (Takayama et al. 1994; Thiel et al. 1997; Matsushita et al. 2000a). MASP-1 can apparently cleave C2 and C3 but not C4 and also to activate of MASP-2 (Chen et al. 2004; Takahashi et al. 2008). This indicates a distinct function for MASP-1 from MASP-2, but its role is still elusive. The function of MASP-3 is unknown, but it does not cleave C2 or C4 (Dahl et al. 2001; Zundel et al. 2004). MASPs are homologous to C1r and C1s but do not bind C1q. Also C1r and C1s do not bind MBL or ficolins that makes the activation of the pathways clearly distinct but functionally complementary (Harmat et al. 2004; Wallis et al. 2010).

1.2.4 Properdin pathway The recently rediscovered properdin pathway utilizes properdin as a pattern recognition molecule (Pillemer et al. 1954; Hourcade 2006). Properdin has been shown to bind to dying cells, certain microbes and malignant cells where it acts as a receptor for C3b, C3bB and C3bBb, initiating the amplification of the complement system (Kemper et al. 2008). The significance of the properdin pathway is still elusive, but it has raised a lot of attention as a novel activation pathway.


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1.3 Amplification and terminal pathway of complement

Activation via AP, CP, and LP leads to the main amplification step of complement that is necessary for proceeding to the terminal pathway and in the generation of many antimicrobial effector functions (Suankratay et al. 1998; Brouwer et al. 2006; Harboe et al. 2008).

1.3.1 Amplification C3b deposition on the surface starts the main amplification step of the complement that uses the components of the alternative pathway C3, FB and FD (Figure 3). Target bound C3b acquires FB from plasma that is cleaved by FD and the complex C3bBb is formed (Lesavre et al. 1979). C3bBb, i.e. the alternative pathway C3 convertase, is an enzyme that cleaves C3 to C3a and C3b and thus amplifies C3b deposition on the surface. C3bBb, like fluid phase C3(H2O)Bb, is transient with half life of 90 s (Medicus et al. 1976; Pangburn et al. 1986). C3a is released to the environment where it informs inflammatory cells (e.g. macrophages and neutrophils) that complement activation is taking place (Cochrane et al. 1968). The high density of C3b molecules and C3bBb complexes enhances binding of the multimeric properdin to C3b and C3bBb, thus increasing the half-life of the C3bBbP approximately 10-fold and further enhancing complement activation (Fearon et al. 1975). Although the classical pathway C3-convertase C4b2b can also bind properdin, the role of CP is considered to be less significant in amplification (Harboe et al. 2004).

Figure 3. The main amplification step of the complement system and the terminal pathway. Complement activation leads to deposition of C3b on the surfaces. The generation of C3bBb(P) amplifies C3 conversion to C3b (box 1) and leads to propagation of the cascade by binding back to C3bBb(P) on the surface (box 2). C3b2Bb(P) acts as C5 convertase that initiates the terminal pathway (box 3). The dotted arrows indicate enzymatic activity.

1.3.2 Terminal pathway The terminal pathway of complement is initiated when ample amounts of C3b is created and some of it binds back to C3bBb(P) or C4b2b(P) complexes that change their substrate specificity to C5 and cleave it to C5a and C5b (Takata et al. 1987; Kinoshita et al. 1988; Rawal et al. 2001). Importantly, C5a that is released to plasma is a powerful anaphylatoxin and chemotaxin, and more potent than C3a released earlier in the cascade (Meuer et al. 1981). Unlike C3b or C4b, C5b is not capable of covalent binding to the surface but acts as a platform for sequential C6 and C7 binding (Gewurz et al. 1968; Podack et al. 1978a; Law et al. 1980). C5b-7 inserts into the membrane of the target cell and acquires C8 from plasma (Hammer et al. 1975; Podack et al. 1980). C8 can protrude into the lipid bilayer and act as an initiator of attachment of 16 to 18 C9 molecules that create a cylindrical pore in


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the membrane (Podack et al. 1982; Tschopp et al. 1982). This C5b-9 or membrane attack complex (MAC) allows diffusion of small molecules through the membrane that leads to rapid lysis of the target cell (Podack et al. 1984b). Formation of MAC together with opsonization of the target with C3b and the release of C3a and C5a are the main consequences of the amplification and terminal pathway.

1.4 Regulation of the complement system

The complement system is capable of initiating and augmenting the inflammatory response and lysing cells as described above. In order to prevent unwanted damage to self cells the complement system needs to be tightly regulated. Regulatory proteins may act during activation, amplification, and effector steps of the cascade that enables fine tuning of the activation in different physiological circumstances.

1.4.1 Regulator proteins Many complement regulatory proteins are encoded by the regulators of complement activation (RCA) gene cluster in chromosome 1 (Carroll et al. 1988; Rodríguez de Córdoba et al. 1999). All the RCA proteins are composed of 4 to 30 homologous short consensus repeat (SCR) domains that have probably arisen by gene duplication events (Pardo-Manuel et al. 1990; Krushkal et al. 2000). SCR domains are small, approximately 60 amino acid protein domains that typically have short beta-strands forming a bead-like structure stabilized by two characteristic disulphide bridges (Figure 4) (Norman et al. 1991). The SCR domains are usually arranged in proteins in a head-to-tail fashion making RCA proteins resemble beads-in-string (Perkins et al. 1991). The binding sites in SCR- bearing proteins may span over several domains and therefore the orientation between the SCRs is critical for the function of the protein.

The RCA proteins include fluid phase proteins C4b binding protein (C4bp), factor H (FH), factor H-like protein 1 (FHL-1), and six factor H related (FHR) proteins 1, 2, 3, 4A, 4B, and 5, and membrane bound regulators complement receptor 1 (CR1, CD35), membrane cofactor protein (MCP, CD46), and decay accelerating factor (DAF, CD55). Complement receptor 2 (CR2, CD21) is also encoded by the same locus, but has not been shown to regulate complement activation. FHL-1 is an alternatively spliced and truncated gene product of FH that has the first seven SCRs and four extra amino acids in the C-terminus (Scwaeble et al. 1987). Genes for other complement regulators are scattered in the genome and are composed mainly of other types of domains. These include membrane proteins CD59 (protectin) and thrombomodulin (THBD), and plasma proteins carboxypeptidase B, sMAP, MAP-1, vitronectin, clusterin, and C1 inhibitor (C1-Inh) (Reid et al. 1989).

All the membrane bound regulators (RCA and non-RCA) are quite ubiquitously expressed on self cells to protect them, but for example MCP is not expressed on erythrocytes (Cole et al. 1985). Fluid phase regulators protect also self-cells, but importantly also act on non-cellular self-structures. Furthermore, fluid phase regulators restrict spreading of complement activation from the site of attack and prevent the activation of complement in plasma.


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Figure 4. Schematic presentation of the exon structure of DAF gene and the crystal structure of four SCRs in DAF. Typically one exon codes a single SCR domain but some domains are coded by two exons. In proteins, SCRs are aligned like beads on a string. For example, FH is composed of 20 SCRs, CR1 of 30 SCRs, and MCP and DAF of 4 SCRs. DAF PDB ID: 1OJV (Lukacik et al. 2004).

1.4.2 Means of regulation The regulator proteins function by competing with the complement components for the binding sites. In addition, they may have cofactor activity (CA) or decay accelerating activity (DAA) (Figure 5). The decay of the transient C3 and C5 convertases can be accelerated by for example FH and DAF (DAA). Factor I (FI) is a soluble protease that cleaves C3b and C4b to their inactive forms iC3b and iC4b by releasing C3f or C4f fragments to the fluid phase (Harrison et al. 1980a; Harrison et al. 1980b). FI needs C4bp, CR1, MCP, FH, or FHL-1 as a cofactor to perform the proteolysis. These proteins are said to have cofactor activity (CA). CR1 is not particularly efficient in generating iC3b, but is the only regulator that acts as a cofactor for the third cleavage that generates C3dg and C3c, which is released to fluid phase (Ross et al. 1982; Seya et al. 1985). C3dg can be further cleaved by plasma proteases to C3d that is thought to have similar biological activity than C3dg.

Figure 5. Principles of decay accelerating and cofactor activities. A. Proteins that have decay accelerating activity can accelerate the decay of C3 and C5 convertases and thus inhibit complement activation. An example of DAA is illustrated with DAF and C3bBb. B. Proteins that have cofactor activity act as cofactors for protease FI that inactivates C3b. An example of CA is illustrated with MCP. The DAA and CA functions are similar in fluid phase and on surfaces.

1.4.3 Factor H and self – non-self discrimination The properties of surfaces that are different in humans and microbes determine whether complement activation is terminated or not. It has been shown that C3b prefers to attach to certain


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carbohydrates and amino acid hydroxyl groups, but this is not sufficient for target discrimination (Tack et al. 1980; Meri et al. 1990b; Sahu et al. 1994; Sahu et al. 1995). Rather, the surface properties determine the rate of C3b deposition. Thus target discrimination is mediated by FH that forms a ternary complex with C3b and the surface (Carreno et al. 1989; Meri et al. 1990b). Accordingly, it has been shown that the presence of negatively charged heparin or sialic acids on the self cell surfaces leads to approximately 10-fold increase in the avidity of FH to C3b-surfaces and to higher cofactor activity of FH (Weiler et al. 1976; Fearon 1978; Pangburn et al. 1978; Kazatchkine et al. 1979b; Meri et al. 1990b).

1.4.4 Anionic self surface structures Heparin, a highly sulfated form of heparan sulfate (HS), has been used widely in complement research as a model of the cell surface glycosaminoglycans (GAGs). HS is a major contribution to the negative charge of the endothelial cell surface glycocalyx and extracellular matrices, like glomerular basement membrane (GBM) (Rops et al. 2004b; Weinbaum et al. 2007). Soluble heparin is released and synthesized only in mast cells, but HS in the glycocalyx of the cells has heparin-like domains where FH is thought to bind.

Chondroitin sulfate A (CS-A) is the main GAG on platelets (Okayama et al. 1986). Upon platelet activation, the fully sulfated form of CS-A is exposed on the surface and is also released from α-granules (MacPherson 1972; Ward et al. 1979; Okayama et al. 1986). Factor H also protects platelets from complement attack (Devine et al. 1987).

Sialic acids or neuraminic acids also contribute to the biology and the high negative charge of the self surfaces in addition to GAGs. Sialic acids are a diverse group of nine-carbon sugars with a negative charge that are present as terminal sugars in glycans of lipids and proteins (Angata et al. 2002). Sialic acid biology and specificity of the interactions with proteins are still very much elusive (Varki 2008). However, target surfaces bearing sialic acids, especially erythrocytes, are recognized by FH, which subsequently restricts complement activation on these surfaces (Fearon 1978; Pangburn et al. 1978).

1.4.5 C3b, GAG and sialic acid binding sites on FH The major functions of FH have been located on certain domains by several structural and functional methods (Figure 6). The domains 19 and 20 of factor H (FH19-20) have been shown to be crucial for self – non-self discrimination (Pangburn 2002; Ferreira et al. 2006), to bind C3d, iC3b and C3b (Sharma et al. 1996; Jokiranta et al. 2000; Pangburn et al. 2004), and to bind heparin (Gordon et al. 1995; Blackmore et al. 1998; Herbert et al. 2006). The sialic acid and CS-A binding sites that are important in self surface recognition have been localized to FH16-20 (Ram et al. 1998) and FH15-20, respectively (Jokiranta et al. 2005). These binding sites are probably located in FH19-20, since these domains are critical for protecting sialic acid-rich sheep erythrocytes (Pangburn 2002) and CS-A rich platelets (Jokiranta et al. 2005; Vaziri-Sani et al. 2005). FH19-20 is also a hot-spot for aHUS associated mutations in FH (Pérez-Caballero et al. 2001).

FH1-4, or minimally FH1-3, binds to C3b, possesses CA and DAA functions, and also competes with FB on binding to C3b (Alsenz et al. 1984; Gordon et al. 1995; Kühn et al. 1995a; Kühn et al. 1996a;


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Wu et al. 2009). Third binding site for C3b on FH mediates binding to the C3c part of C3b, but the results on the location have been controversial. FH8-20 has been shown to bind C3c, whereas in the same assay, FH1-5 and FH19-20 did not bind C3c, and FH2-6, FH8-11, and FH15-18 did not bind C3b (Jokiranta 2000). Others have shown that FH6-8, FH6-10, FH7-8 and FH8-15 bind C3b, whereas FH15-18, FH10-15, FH10-12, FH11-14, FH12-13, and FH13-15 do not interact with C3b ((Sharma et al. 1996; Schmidt et al. 2008)). Thus it seems that the binding site would be located at FH8 and might expand to FH7 and FH9 as well, but the role of other domains or existence of even a fourth binding site cannot be ruled out.

FH has a well defined heparin binding site on FH7 in addition to FH19-20 (Gordon et al. 1995; Blackmore et al. 1998; Herbert et al. 2006; Prosser et al. 2007). A third heparin binding site has been mapped to domain 9 and also suggested to be in domain 13 (Pangburn et al. 1991; Ormsby et al. 2006), but a recent report has contradicted the existence of these sites (Schmidt et al. 2008). The mid-part of the FH has been shown to be compact and partly folded back and thus functional sites may be hidden in the full protein (Aslam et al. 2001; Oppermann et al. 2006).

Figure 6. Schematic presentation of binding sites in FH. The SCR domains of FH are presented as circles.The solid line presents the binding site and dashed line presents possible role of additional domains in binding.

1.4.6 Regulation of complement activation Factor H is the main regulator of AP activation that is initiated in the fluid phase. Deficiency of FH leads to consumption of C3 from plasma and to Dense Deposit Disease (DDD) (Smith et al. 2007). DDD and aHUS both affect kidney function and are caused by dysfunctional factor H. However, in DDD complement is activated in plasma and glomerular basement membrane, while in aHUS complement is activated mainly on cell surfaces. Factor H functions in the fluid phase by competing with factor B for C3b binding, by acting as a cofactor in cleavage of C3b to iC3b and accelerates spontaneous decay of the fluid phase C3 convertases, C3b(H2O)Bb and C3(H2O)Bb (Weiler et al. 1976; Whaley et al. 1976; Pangburn et al. 1977; Kazatchkine et al. 1979a; Farries et al. 1990). FHL-1 performs CA and DAA like FH but the physiological significance of FHL-1 has not been shown - it could be modulation of activation, since plasma concentration of FHL-1 is about 10 % of FH concentration (Misasi et al. 1989; Kühn et al. 1995b; Kühn et al. 1996b). C4bp has also been shown to cleave fluid phase C3b and thus may fine tune AP activation (Nagasawa et al. 1977; Seya et al. 1985; Blom et al. 2001).


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The main regulator of the CP and LP is C1-Inh, which binds to C1r, C1s (Ratnoff et al. 1969), and MASPs (Matsushita et al. 2000b). C1-Inh is a suicide protease inhibitor that irreversibly binds to its targets (Laurell et al. 1978) and disassembles the C1 and probably also MBL/ficolin/MASP complexes (Ziccardi et al. 1979; Presanis et al. 2004). The activation of the LP is also regulated by MAP-1 and sMAP proteins that are alternatively spliced isoforms of the MASP-1 and MASP-2 genes, respectively (Matsushita et al. 2000a; Skjøedt et al. 2010). MAP-1 and sMAP lack the serine protease domains of the MASPs and act as competitive inhibitors for MASPs on MBL and ficolin binding. On surfaces C1-Inh, sMAP and MAP-1 function less efficiently than in the fluid phase, allowing complement activation (Wallis et al. 2010).

CP and LP activation lead to deposition of C4b and C4b2b on the cell surfaces. C4b is cleaved to inactive C4b (iC4b) by C4bp, MCP, or CR1 that act as cofactors for FI (Scharfstein et al. 1978; Iida et al. 1983; Seya et al. 1986), and the decay of the C4b2b complexes is accelerated by C4bp, DAF, and CR1 (Gigli et al. 1979; Nicholson-Weller et al. 1982; Iida et al. 1983). C4bp also binds to C4b and inhibits convertase formation (Blom et al. 1999).

1.4.7 Regulation of amplification and inhibition of MAC formation on surfaces The amplification step of the complement is redundantly regulated by many proteins (Figure 7). The decay of C3bBb is accelerated by FH, FHL-1, CR1, and DAF (Weiler et al. 1976; Whaley et al. 1976; Fearon 1979; Nicholson-Weller et al. 1982; Kühn et al. 1995a), and the deposited C3b is inactivated by FH, FHL-1, CR1 and MCP that act as cofactors for factor I mediated cleavage (Pangburn et al. 1977; Medicus et al. 1983; Seya et al. 1986; Kühn et al. 1995c). Factor H also competes with factor B on binding to C3b (Kazatchkine et al. 1979a; Farries et al. 1990).

Binding of properdin to C3bBb or C4b2b does not probably block the DAA or CA activity of the regulatory proteins, but rather inhibits their actions by stabilizing the convertase. Properdin has been shown to inhibit FH and FI mediated cleavage of C3b in C3bBbP (Medicus et al. 1976) and FH and DAF have been shown to accelerate decay of C3bBbP (Weiler et al. 1976; Whaley et al. 1976; Nicholson-Weller et al. 1982; Hourcade 2006).

The decay of C5-convertases (C3b2Bb(P) and C3bC4b2b(P)) is accelerated by FH (Kuhn 1996), CR1 (Krych-Goldberg 1999), C4bp (Rawal 2007) and DAF (Kuttner-Kondo 2001). MCP and CR1, but not FH and C4bp, can act as cofactors for FI in cleaving C3b dimers on C5 convertases (Seya 1991) and C4bp, but not FH, has CA for C3bC4b2b (Meri et al. 1990c; Rawal et al. 2007).

Recently, the action of the C5 convertase was suggested to be inhibited also by FHR-1 (Heinen et al. 2009). The functions of the other five FHR proteins are not well established but they may also act on C5 convertases (Zipfel et al. 2009). The actions of C5 convertases that lead MAC formation are inhibited on the self cell membranes by CD59 and vitronectin (S-protein), which interfere with C9 polymerization (Bhakdi et al. 1988; Meri et al. 1990a).


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Figure 7. Regulation of the main amplification loop. FH, DAF, and CR1 accelerate the decay of C3 and C5 convertases. FH, MCP, and CR1 act as cofactors for factor I in cleaving C3b to iC3b. The amplification loop and the terminal pathway are shown in grey.

1.4.8 Restriction of complement activation in the fluid phase The function and lifetime of the inflammatory mediators C3a and C5a is regulated by the soluble protease carboxypeptidase-N. Carboxypeptidase-N cleaves C3a and C5a to the C3adesArg and C5adesArg forms, which lack terminal arginines (Erdos et al. 1962; Bokisch et al. 1969). The desArg forms exhibit reduced inflammatory and chemotactic functions. Consequently, the action of carboxypeptidase-N is to create a chemical gradient that guides the inflammatory cells to the site of complement activation, and to limit the inflammatory response at more remote sites where the peptides have been taken (for example, by diffusion).

Some C3b and C4b that are not surface-bound but remain in the fluid phase are cleared mainly by the fluid phase regulators FH and C4bp (Seya et al. 1995). In addition, the CR1 that is expressed on red blood cells (RBCs) captures C4b, C3b, iC3b, C3b2-IgG, and C3b-ICs (Fearon 1979; Kinoshita et al. 1986; Schifferli et al. 1986). CR1 can also process iC3b to C3dg, an ability unique to this regulator (Ross et al. 1982).

C5b, C5b6, and C5b-7 generated by amplification may drift from the original site of the activation and insert into membranes of bystander cells or form MAC in the fluid phase. These actions are regulated by vitronectin and clusterin, which bind to soluble C5b-7 (sC5b-7), sC5b-8, and sC5b-9 leading to inhibition of MAC formation (Podack et al. 1978b; Podack et al. 1979; Podack et al. 1984a; Jenne et al. 1989; Tschopp et al. 1993).

Taken together, the main functions of complement regulators are (1) to limit the activation in the fluid phase and on viable self-cells and noncellular self-surfaces, (2) to regulate the extent of activation by controlling the amplification, (3) to limit complement attack to the target or its immediate vicinity by inactivating the components released into fluid phase, and (4) to limit the effects of the anaphylatoxins, and inflammation in general, to the region where activation occurs.


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1.5 Complement in innate immune response

In an infection, the microbe is opsonized with C1q, MBL, ficolins, C3b, iC3b, and C3dg, and anaphylatoxins C3a and C5a are generated in response to activation (Figure 8, box 1). Neutrophils, macrophages, and dendritic cells that are important in the innate immune response bear C3a and C5a receptors (C3aR and C5aR) and receptors for complement components (Gasque 2004; Haas et al. 2007; Klos et al. 2009). It is noteworthy that the complement activation products C3a and C5a cannot inform the cellular immunity about the type of pathogen because all the recognition pathways lead to these same products - independently of the properties of the target surface. It seems that C5a non-specifically enhances the proinflammatory response, while toll-like receptor (TLR) and cytokine receptor activation on immune system cells determines the type of the response. The interplay between the signaling pathways of C5a receptor (C5aR), TLRs, and cytokine receptors has been recognized recently (Hajishengallis et al. 2010).

Figure 8. Complement in the initiation of innate immune response. Box 1. Activation of complement on microbes opsonizes it and creates anaphylatoxin C5a. Box 2. C5a binds to C5aR on the macrophages, monocytes, neutrophils, and DCs. C5aR activation attracts cells to the site of infection and augments TLR signaling in macrophages and DCs. Box 3. TNF-α and IL-1 that are synthesized mainly by macrophages activate endothelial cells. Box 4. C5a has synergistic effects on TNF-α and IL-1 induced activation of endothelium and expression of selectins. Box 5. CR1, CR2, C1qR, and FcγR bind to opsonins on the microbe that facilitates phagocytosis. Receptors and cells are indicated with an oval and bolded circles, respectively. Soluble proteins are not circled. Arrows schematically indicate diffusion of C5a or cytokines from the source of their production.


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1.5.1 Actions of anaphylatoxins Complement activation leads to formation of C3adesArg and C5adesArg which induce leukocytosis in 20 minutes, some 30 minutes before cytokines take action, thus providing the first signal of an infection (Menkin 1949; Rother 1972; Kajita et al. 1990; Jagels et al. 1994). At the infection focus, C5a and C5adesArg act as powerful chemotactic agents for neutrophils, monocytes, macrophages, and dendritic cells (DCs) to indicate the presence of an infection and to recruit them to the site of activation, i.e. the site of invader (Figure 8, box 2) (Shin et al. 1968; Snyderman et al. 1970; Snyderman et al. 1975; Fernandez et al. 1978). The importance of C3a or C3adesArg in inducing chemotaxis has been controversial and poorly studied, but in the light of current information they don’t induce neutrophil chemotaxis, but contribute to eosinophil chemotaxis (Daffern et al. 1995). The physiological reason for the lack of this activity on neutrophils may be that C3a is also generated during removal of late apoptotic/necrotic cells, where excess inflammation and recruitment of neutrophils should be avoided.

Tissue-resident macrophages are important in phagocytosis and in sensing infection. The important early response to infection by macrophages is to generate pro-inflammatory TNF-α and IL-1 via activation of the NF-kappa B pathway that leads to activation of the cellular innate immune responses. Complement activation and generation of C5a is triggered by e.g. yeast cell wall zymosan and bacterial LPS that also bind to, respectively, TLR2/6 and TLR4 on the macrophage surface. Triggering of C5aR and TLRs 2/6 or TLR4 in mice have been shown synergistically to induce the NF-kappa B pathway and the production of pro-inflammatory cytokines TNF-α, IL-1, IL-6 and IL-12 (Zhang et al. 2007). C5a has also been shown to induce production and release of TNF-α and IL-1 from LPS-stimulated and non-stimulated human macrophages (Goodman et al. 1982; Cavaillon et al. 1990; Pushparaj et al. 2009) and to activate macrophages dependent on NF-kappa B (Kastl et al. 2006) (Figure 8, box 3). It seems that C5a contributes very early in infection to enhance cellular immunity together with TLRs (Hajishengallis et al. 2010).

The effects of TNF-α and IL-1 on the endothelium is to activate it via the NF-kappa B pathway (Schleef et al. 1988) (Figure 8, box 4). C5a acts synergistically with cytokines to activate endothelium to express neutrophil ligand P-selectin on its surface (Geng et al. 1990; Foreman et al. 1994; Mulligan et al. 1997), and later in infection leukocyte ligand E-selectin (Springer 1994). Neutrophils that are attached to the endothelium may be activated to release hydrolytic enzymes that can cause damage to the endothelium (Varani et al. 1989; Hardy et al. 1994).

1.5.2 Opsonization of microbes While complement has attracted inflammatory cells to the site of infection, it has also opsonized the microbe to facilitate its phagocytosis. Professional phagocytes like macrophages, dendritic cells, and neutrophils use an array of receptors to attach to the pathogen (Figure 8, box 5) (Stuart et al. 2005). These include C1q receptors (C1qR), Fc gamma receptors (FcγR), integrins CR3 (CD11b/CD18) and CR4 (CD11c/CD18), and complement receptor 1 (CD35) (Underhill et al. 2002). The most important phagocytic receptors of the complement system are CR1 and CR3. CR1 binds C3b, iC3b, C3dg, C4b, and iC4b, and also C1q and MBL on the target surface (Fearon 1979; Kinoshita et al. 1986; Klickstein et al. 1997; Ghiran et al. 2000; Krych-Goldberg et al. 2001). CR3 recognizes iC3b and also microbial components such as beta-glucan and LPS (Ross et al. 1985; Wright et al. 1989; Thornton et al. 1996).


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Complement receptors 1 and 3 potentiate FcγR mediated phagocytosis by clustering the receptors that trigger common signaling pathways (Ehlenberger et al. 1977; Bobak et al. 1987; Fries et al. 1987; Krauss et al. 1994; Zhou et al. 1995). The functions of CR4 and C1qRs are not well established, but they seem to participate in recognizing the target and mediating signals to phagocytes (Myones et al. 1988; Gasque 2004; Ghebrehiwet et al. 2004).

Phagocytes, especially DCs, sense the infection and type of pathogen by several TLRs. The deposition of the complement components alone on the cells is not sufficient for phagocytosis (Wright et al. 1983), but inflammatory mediators like TNF-α and LPS can provide additional stimuli to induce CR- dependent phagocytosis by enhancing expression of the receptors (Berger et al. 1984; Wright et al. 1985). At least CR3 seems to act in concert with TLR4 and CD14 in regulating the NF-kappa B pathway (Perera et al. 2001). Taken together, the activation of complement, opsonization of the microbe, and production of C5a leads to phagocytosis and triggering of anti-microbial effector mechanisms of the cells.

1.6 Complement in adaptive immunity

1.6.1 C3dg as an adjuvant in B-cell response Generation of the antibody response against microbes is augmented by complement C3 (Pepys 1974). On complement activation, C3b is bound to microbes and is processed by the actions of CR1 and factor I to the C3dg fragment, which remains covalently bound to the antigen. B-lymphocytes express complement receptor 2 (CR2, CD21) and the B-cell receptor (BCR) that recognize the C3dg and the antigen, respectively. CR2 is part of the CD21/CD19/CD81 BCR co-receptor complex that is able to augment BCR signaling (Carter et al. 1988; Carter et al. 1992). The B-cell response is thus greatly enhanced when C3dg is bound to the co-receptor complex and antigen to the BCR (Fearon et al. 2000).

T-cell dependent (TD) antigens (typically non-repetitive proteins) with one epitope for BCR, and one C3dg attached to it, do not efficiently cluster receptors and trigger the B-cell response. The presence of two or more C3dg molecules on the antigen clusters the BCR and its co-receptor, increasing the B-cell response 10 000 fold (Dempsey et al. 1996). T-cell independent (TI) antigens such as polysaccharides bear identical epitopes and are large enough to cluster receptors and activating B-cells without T-cell help, but the binding of multiple C3dgs to TI antigens can still further increase the B-cell activation.

TD (and TI) antigen presentation takes place in lymph nodes and in spleen where T-cell help and antigen-presenting cells are available to provide second signals to B-cells. In tissues, DCs expressing CRs capture antigens that may bear C3b, iC3b or C3dg and migrate into lymph nodes to present antigen to T- and B-cells (Banchereau et al. 1998). Antigen does not need to be transported by DCs to lymph nodes where resident macrophages, DCs, and follicular DCs can capture C3b/iC3b/C3dg-antigen complexes from lymph and present the antigen (Batista et al. 2009). The complement receptors CR1, CR2, CR3 as well as FcγRs play critical roles in antigen capturing, trafficking and presentation to B-cells in lymph nodes and spleen (Pozdnyakova et al. 2003; Ferguson et al. 2004; Gonzalez et al. 2010).


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B-cells endocytose the antigen, cleave it to peptide fragments, and display some of these fragments on MHC-II molecules and present the complex to helper T-cells that have been activated by DCs to display CD40L. CD40-CD40L and TCR-MHCII interactions activate the B-cells that start the formation of germinal centers. Follicular DCs in germinal centers bear CR1, CR2, and CR3 that retain the C3dg-antigen complex in the follicles and positively affect B-cell affinity maturation and memory B-cell generation (Reynes et al. 1985).

1.6.2 Complement in T-cell responses An emerging field in complement research is the role of complement components in T-cell activation, regulation, and direction of the T-cell responses (Kemper et al. 2007). For example, there is evidence that synthesis of C3 by DCs is required for full T-cell activation and TH1 polarization (Peng et al. 2006). In addition, a critical role for C3aR in DC antigen uptake and T-cell stimulation has been suggested (Li et al. 2008; Peng et al. 2008) and also C3a and C5a that were produced by DCs were found to provide costimulatory and survival signals to naïve CD4 cells (Strainic et al. 2008). All these studies indicate an important role for C3 components, and C3aR and C5aR in the interplay between DCs and T-cells. Taken together, the complement system is now becoming linked to T-cell biology as well highlighting the importance of complement as an initiator and regulator of both the innate and adaptive immune responses.

1.7 Complement in cell death

Uncontrolled complement activation on self cells produces sublytic and lytic depositions of MAC that can induce apoptosis or cause necrosis (Mevorach et al. 1998; Cole et al. 2003). If MAC inserts into the membrane it increases the influx of calcium to the cell, speeds up cell death and leads to swelling of the cell and rupture of the membrane (Kim et al. 1987). These events lead to cell death that has apoptotic and necrotic features (Shimizu et al. 2000). Importantly, for renal diseases and aHUS, even sublytic MAC deposition has been shown to induce apoptosis in renal and endothelial cells (Figure 9, box 1) (Sato et al. 1999; Hughes et al. 2000; Nauta et al. 2002).

1.7.1 Activation of complement Dying cells present marker-molecules on the surface, like phospholipids and nucleic acids that are recognized by soluble opsonins and on phagocytes by various receptors (Poon et al. 2010). The soluble opsonins include MBL, C1q and ficolins that can bind to DNA and phospholipids on the membrane. Also, IgM and the pentraxins serum amyloid protein (SAP) (Hicks et al. 1992), CRP (Mold et al. 1999; Gershov et al. 2000), and long pentraxin 3 (PTX3) (Bottazzi et al. 1997) opsonize dying cells and are recognized by C1q (Korb et al. 1997; Paidassi et al. 2008). C1q and CP activation are key players in the clearance of dying cells (Gullstrand et al. 2009), and their dysfunction is associated with autoimmune diseases such as systemic lupus erythematosus and rheumatoid arthritis (Botto et al. 1998; Okroj et al. 2007).

CP is activated on late apoptotic and necrotic cells but not much on early apoptotic cells (Figure 9, box 2) (Nauta et al. 2003). Complement activation on cells in late apoptosis is enhanced by the


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down-regulation of membrane-bound complement regulators DAF and CD59 or by shedding of MCP and CD59 (Jones et al. 1995; Elward et al. 2005; Cole et al. 2006). If complement activation proceeds to the terminal pathway, the activation of complement would lead to lysis of the cell and induction of inflammation (Figure 9, box 3). However, the complement activation is restricted by simultaneous acquisition of FH and C4bp that bind directly to the apoptotic or necrotic cells via e.g. DNA or by utilizing opsonins such as CRP (Mold et al. 1984; Sjöberg et al. 2006; Trouw et al. 2007) or PTX3 (Deban et al. 2008). This restricts the launching of the terminal pathway and generation of highly pro-inflammatory C5a, but not C3a (Figure 9, box 4). C3a has traditionally been considered to be pro-inflammatory, but recent evidence suggests that C3a may in fact dampen the inflammatory response that would be beneficial in clearing apoptotic or necrotic cells (Hawlisch et al. 2006). For example, it has been shown that neutrophils possess theC3a receptor but C3a does not attract them (Martin et al. 1997; Kemper et al. 2008). Also, C3a inhibits the production of proinflammatory IL-1β and TNF-α production in non-adherent peripheral blood monocytes (Takabayashi et al. 1996), and IL-6 and TNF-α production in B-cells (Fischer et al. 1997).

Figure 9. Uncontrolled complement activation on endothelial cells can induce cell death. Box 1. Complement activation on endothelial cells may induce apoptosis. Box 2. As apoptosis proceeds to late phase complement is activated via classical pathway and cell surface regulation is down-modulated. Box 3. If the complement activation is not held in check the cell may be lysed or become necrotic. Box 4. Complement regulators like FH restrict the activation to C3-stage that prevents the generation of pro-inflammatory C5a, but allows opsonization of the cells with C3b and iC3b.

1.7.2 Phagocytosis The activation of complement on dying cells leads to opsonization of the cell with C1q, MBL, ficolins, and iC3b, C3b, iC4b, and C4b that all act as ligands for phagocytes. For instance, iC3b on apoptotic cells has been shown to enhance phagocytosis by macrophages (Takizawa et al. 1996). Phagocytes mediate uptake of apoptotic cells by a variety of receptors. Particularly important are CR1 and CR3, which recognize C3 and C4 fragments, and CD91/calreticulin receptors which recognize C1q, MBL, and ficolins (Ogden et al. 2001; Honoré et al. 2007; Jensen et al. 2007). Uptake of apoptotic cells induces anti-inflammatory cytokines such as TGF-β production in macrophages (Fadok et al. 1998). It has been shown that CR3 and CR4 dependent uptake of apoptotic or necrotic cells bearing iC3b dampens the inflammatory response by decreasing IL-12 production by macrophages (Mevorach et al. 1998; Kim et al. 2004), and that the expression of costimulatory molecules in DCs is decreased upon CR3 stimulation (Verbovetski et al. 2002; Morelli et al. 2003). Taken together, the activation of


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complement and other mechanisms in clearing late apoptotic or necrotic cells induces the type of inflammation that is directed to tissue regeneration, while inhibiting the adaptive immune responses to prevent generation of autoimmune diseases. The important difference in the clearance of late apoptotic or necrotic cells vs microbes is that high amounts of C5a and MAC are generated only in response against the microbes.

1.8 Crosstalk of complement with coagulation system

There are three major protease cascades in plasma - coagulation, fibrinolysis, and complement - that are homologous in terms of protein structure and function (Krem et al. 2002). The complement and coagulation systems work in an analogous fashion – both are quickly propagating enzyme cascades that need to be activated only locally and controlled efficiently. These systems are interconnected and triggered together in physiological and pathological conditions such as blood clotting and aHUS that is characterized by microvascular thrombi and dysfunctional complement system (Markiewski et al. 2007).

1.8.1 Initiation of coagulation by C5a and MAC Coagulation can be initiated via an extrinsic pathway by damaged endothelium, leading to exposure of subendothelial matrix bearing collagen, tissue factor (TF), von willebrand factor (vWf), and fibrinogen, or by increased expression of TF and vWf by endothelium (Savage et al. 1996; Karpman et al. 2006; Adams et al. 2009). C5a and MAC deposition have been shown to induce endothelial cell damage, vWf release, and TF upregulation (Figure 10, box 1) (Ikeda et al. 1997; Tedesco et al. 1997). Platelets bind to vWf on endothelium and provide negatively charged phophatidylserine surface and receptors for binding of plasma coagulation factors (F) (Figure 10, box 2) (Hoffman et al. 2001). The key protein in initiating the coagulation cascade is the endothelium-bound TF that binds factor VII (FVII) and localizes the coagulation at the site (Figure 10, box 3). The TF:FVIIa complex (“a” indicating activated protein) propagates the cascade to formation of prothrombinase FXa:FVa complex. Prothrombinase cleaves prothrombin to thrombin that binds to negatively charged surfaces on platelets and endothelium, and activates the platelets on the site of injury (Monroe et al. 1996; Diaz-Ricart et al. 2000).


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Figure 10 Simplified presentation of induction of coagulation system by complement and its feed-back mechanism. Box 1. Complement activation products C3a, C5a, and MAC can induce pro-coagulant phenotype to endothelium that leads to up-regulation of TF and vWf. Box 2. Platelets adhere to vWf and provide negatively charged surface for coagulation. Box 3. TF binds to FVIIa initiating coagulation that generates thrombin. Thrombin binds to platelets and cleaves several coagulation cascade enzymes and initiates clot formation. Box 4. Thrombin cleaves C3 and C5 that initiates complement activation. Box 5. C3a, C5a, and MAC can further activate endothelium and platelets thus creating a feed-back mechanism.

1.8.2 Activation of complement by coagulation The amplification and propagation of blood clotting on platelet surfaces is initiated by the actions of thrombin that leads to generation of the enzyme complexes FIXa:FVIIIa and FXa:FVa on the platelet surface (Hoffman et al. 2001; Adams et al. 2009). These complexes generate massive amounts of thrombin feeding back to the coagulation system. The platelets have high affinity binding sites for thrombin, FIXa, FXa, and FXI that increases their local concentration (Greengard et al. 1986; Rawala-Sheikh et al. 1990; Cirino et al. 1997). At high concentrations thrombin, FIXa, FXa, and FXIa can cleave C3 and C5 and thus initiate complement activation at the site of coagulation (Figure 10, box 4) (Spath et al. 1976; Huber-Lang et al. 2006; Amara et al. 2010). C3a, C5a and sublytic amounts of MAC can further activate platelets and endothelium creating a feed-back loop (Figure 10, box 5) (Grossklaus et al. 1976; Polley et al. 1983; Damerau et al. 1986; Wiedmer et al. 1986).

The endpoint of the coagulation cascade is the formation of a blood clot. Thrombin cleaves fibrinogen to fibrin that forms a soft clot that is further covalently linked by FXIIIa to form hard clots. In aHUS complement activation takes place ubiquitously on cell surfaces, but the renal vessels in the kidney glomeruli are primarily occluded. Microvasculature in glomeruli forms a capillary bed where the diameter of a capillary is only 3-10 µm whereas the average diameter of an erythrocyte is 7-8 µm and that of a platelet is about 2 µm (Weinbaum et al. 2007) and thus the formation of thrombi easily occludes these narrow capillaries.


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1.8.3 Interplay of complement and coagulation regulators The spreading of the local blood clotting is controlled and down regulated by the plasma protease inhibitors (e.g. anti-thrombin III) and the intact endothelium surrounding the clot. Thrombomodulin (THBD) is expressed especially abundantly on endothelium in microvasculature (Cadroy et al. 1997) and captures active thrombin that drifts from the site of coagulation. Upon binding to THBD, thrombin changes its specificity from fibrinogen to protein C and activates it (Ye et al. 1991). Activated protein C makes a complex with protein S that inactivates FVa and FVIIIa deposited on bystander endothelial surfaces. Thus the action of thrombin on intact endothelium is anti-thrombotic. THBD also enhances thrombin-mediated activation of pro-carboxypeptidase B, an inhibitor of fibrinolysis that is also able to inactivate C3a and C5a to their desArg forms (Bajzar et al. 1995; Campbell et al. 2002). Recently, THBD was connected to aHUS and found to regulate complement by binding FH and C3b, leading to enhanced C3b inactivation (Delvaeye et al. 2009). Coagulation system protein S has also been shown to participate in complement regulation by binding C4bp. In complex with C4bp, protein S cannot inactivate FVa and FVIIIa but can still mediate binding of C4bp to negatively charged glycolipids (Rezende et al. 2004). This interaction thus promotes coagulation while controlling complement CP and LP activation.

Blood clots are removed by the fibrinolysis system. The main enzyme of this process is plasmin that cleaves fibrin clots. Plasmin is also able to cleave C3 and C5, suggesting that complement activation is also kept on at the resolution stage of the clotting (Goldberger et al. 1981; Amara et al. 2010). The clotting and the complement systems are interconnected and complement activation can induce a pro-coagulant state (Shebuski et al. 2002; Esmon 2004). This is well exemplified in atypical hemolytic uremic syndrome where complement and coagulation systems are both triggered.

1.9 Atypical hemolytic uremic syndrome

Hemolytic uremic syndrome is a disease that belongs to a class of diseases called thrombotic microangiopathies and is characterized by hemolytic anemia, thrombocytopenia, and impairment of renal function (Ruggenenti et al. 2001). The typical form of HUS usually presents with diarrhea and is secondary to infections by shiga-toxin associated bacteria such as Escherichia coli serotype O157:H7 (Noris et al. 2005). The atypical form is not associated with diarrhea caused by shiga-toxin producing bacteria, but has usually a genetic background. Both forms of HUS have similar histologic lesions and are characterized by thrombotic and inflammatory state of small arteries and capillaries (Ruggenenti et al. 2001).

1.9.1 Clinical findings Typical clinical findings in aHUS patients are hemolysis and hemolytic anemia, thrombi in kidney microcirculation, endothelial swelling and detachment, subendothelial accumulation of protein and cell debris, and thickening of the arterioles and capillaries (Noris et al. 2009). The glomeruli are heavily inflamed and occluded leading to impairment of renal function that is the most common cause of death in aHUS. Renal function is quite well restored in patients with typical HUS, but atypical form develops to end stage renal disease in about 50 % of the patients (Caprioli et al. 2006). In 20 % of aHUS patients extrarenal complications, like neurological or cardiovascular, have been described (Jalanko et al. 2008; Loirat et al. 2008; Sallee et al. 2010).


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1.9.2 Incidence and genetic background The incidence of HUS is about 6 / 100 000 children under 5 years of which 10 % are classified as atypical (Noris et al. 2009). Overall, 67 % of aHUS cases with known genetic background are presented during childhood, but the disease may be triggered also at later age. Thus the disease has incomplete penetrance and the onset may need mutations in more than one gene or an environmental trigger like pregnancy or infection (Kavanagh et al. 2008).

The genetic background for all the aHUS patients has not been described, but in 50-60% of aHUS patients one or more point mutations are found in genes encoding complement proteins FB (1-4 % of patients), C3 (2-10 %), FH (20-30 %), MCP (5-15 %), and FI (4-10 %) (Noris et al. 2009; Maga et al. 2010). Mutations have also been described in complement and coagulation regulator THBD in 3-5 % of the patients (Delvaeye et al. 2009; Maga et al. 2010). These patients are often heterozygous for mutation, but still present the disease, i.e. the mutations have a dominant negative effect.

In 3-5 % of the aHUS patients a recombination event between FH and FHR-1 has occurred that leads to an abnormal FH gene (Noris et al. 2009; Maga et al. 2010). In addition, the lack of FHR-1 and/or FHR-3 predisposes to the development of auto-antibodies against FH that have been described in 6-10 % of patients (Józsi et al. 2008; Noris et al. 2009).

In Finland three patients have been described having aHUS mutation (R1215Q) in factor H gene ((Jalanko et al. 2008); Jokiranta, personal communication). Two of the patients were from a family that also had asymptomatic carriers indicating incomplete penetrance. One of the aHUS patients had also factor V Leiden mutation that slightly increases the thrombogenicity of blood.

1.9.3 Molecular pathogenesis of aHUS Pathogenesis of aHUS has been associated with defective regulation of AP activation and amplification. The C3bBb complex that initiates the amplification of complement activation is transient, but some aHUS associated mutations in the interface between the Bb and C3b increase the half life of C3bBb, leading to uncontrolled deposition of C3b (Goicoechea de Jorge et al. 2007; Roumenina et al. 2009). It would be expected that aHUS mutations could also disturb DAA and in this way increase C3b deposition, but apparently this is not the case since DAF dysfunction has not been associated with aHUS (Kavanagh et al. 2007).

Atypical HUS can also be caused by impaired inactivation of C3b that is normally inactivated on self cells by FH or MCP that act as cofactors for factor I in cleaving C3b to iC3b. Upon binding of FH or MCP to C3b, a cleavage site for factor I is formed on C3b (Wu et al. 2009). Factor I then cleaves two bonds in the C3b releasing C3f and leaving iC3b on the surface in a conformation that is no longer able to bind FB. Mutations in MCP, FH1-4, or FI can impair the inactivation of C3b (Noris et al. 2003; Richards et al. 2003; Kavanagh et al. 2005).

In FH most aHUS mutations are not segregated to FH1-4 but to FH19-20. FH19-20 is crucial for self-non-self discrimination function of FH and it has been shown that patient-derived full-length FH proteins bearing mutations W1183L, V1197A, and R1210C are unable to protect non-activator sheep erythrocytes from lysis (Sánchez-Corral et al. 2004). Domain 19 mutations D1119G, V1134G, E1135D, Y1142C/D, W1157R, C1163W, and V1168E, and domain 20 mutations R1182S, W1183L/R, T1184R,


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L1189F/P/R, S1191L/W, G1194D, V1197A, E1198A/K, F1199S, R1203W, R1206C, R1210C, S1211P, R1215G/Q, and P1226S have been described in aHUS patients (see study IV for detailed references or www.fh-hus.org) (Warwicker et al. 1998; Ying et al. 1999; Pérez-Caballero et al. 2001; Richards et al. 2001; Sánchez-Corral et al. 2002; Neumann et al. 2003; Ståhl et al. 2008; Maga et al. 2010; Sullivan et al. 2010; Westra et al. 2010).

1.9.4 Treatment Atypical HUS has usually been treated with plasmapheresis, transplantation, or recently with monoclonal antibody against C5 (Kavanagh et al. 2010; Köse et al. 2010; Loirat et al. 2010). The severity of aHUS is dependent on the affected protein that has also an effect on the choice of treatment. Plasmapheresis is used to treat acute episodes of aHUS, but in some patients this treatment is only transiently beneficial. Patients carrying MCP mutations have a good prognosis after kidney transplantation, since the defective protein in the kidney is replaced with functional molecules in the transplant. For the soluble proteins factor H, factor B, factor I, and C3 kidney transplantation is not a good option since these proteins are mainly produced in the liver. An option for these patients is combined liver-kidney transplantation after extensive plasma exchange. This procedure has been successfully done in Finland to three patients, of which all are still alive, and abroad to a few patients also with good results (Jalanko et al. 2008; Kavanagh et al. 2010). An alternative to transplantation has recently been shown to be the anti-C5 monoclonal antibody Eculizumab, which prevents generation of C5a and propagation of the complement cascade to the terminal pathway (Gruppo et al. 2009; Nürnberger et al. 2009; Zimmerhackl et al. 2010). This antibody seems to be a very promising option in the treatment of aHUS, but knowledge of the molecular pathogenesis of aHUS is critical in developing further treatment options for aHUS patients.

Aims of the study

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2 Aims of the study

I. To study the basis of rational mutagenesis to SCR domains and FH19-20.

II. To analyze the molecular mechanism of the FH19-20 mediated self non-self discrimination by FH.

III. To elucidate the pathogenic mechanism of aHUS that is caused by mutations in FH19-20.

Materials and methods

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3 Materials and Methods

3.1 Materials

3.1.1 Proteins and carbohydrates In study II the wild-type FH19-20 without expression tags was cloned by MSc Maria Pärepalo (Pärepalo 2002). FH19Del-20 (Q1137A/ Q1139A/ Y1142A) and FH19-20Del (T1184G/ K1202A/ R1203A/ Y1205A) used in study IV were cloned, expressed and purified by MSc Satu Hyvärinen (Study IV).

C3d that was used in the crystallization in study IV and in functional assays had surface exposed cysteine (C17, part of the reactive thioester) mutated to alanine. C3dg wild-type, and the C3dg mutant proteins D36A, E37A, E117A, D122A, E160A, D163A, and K291A were used in binding assays of study IV. C3d and C3dg proteins were expressed in E. coli without tags and were received from Professor David Isenman and Elisa Leung (Nagar et al. 1998; Isenman et al. 2010).

Rabbit polyclonal antibodies against FH19-20 and factor H (K2), and monoclonal anti FH19-20 antibody VIG8 (Prodinger et al. 1998) were received from Dr. Jens Hellwage. Commercial polyclonal anti-FH antibody was from Calbiochem (Merck).

Heparin from porcine intestinal mucosa and dextran were from Sigma-Aldrich and heparin decamer from Neoparin Ltd.

3.1.2 Microbial strains and cells Cloning of the FH19-20 mutant constructs was done in E. coli XL10-Gold® cells (Stratagene) and the expression of FH19-20 proteins in Pichia pastoris X-33 strain (Invitrogen). Conditionally immortalized mouse glomerular endothelial cells (mGEnC-1) that were used in binding assays have been described by Dr. Angelique Rops (Rops et al. 2004a) and were used in the Nijmegen Centre for Molecular Life Sciences, The Netherlands, under supervision of adjunct professor Johan van der Vlag.

3.1.3 Bioinformatical databases The following web resources were used to obtain sequence and structure information: European Bioinformatics Institute (www.ebi.ac.uk) (McWilliam et al. 2009), National Center for Biotechnology Information (www.ncbi.nlm.nih.gov) (Benson et al. 1997), Protein Data Bank (www.pdb.org) (Berman et al. 2000).

3.2 Methods

3.2.1 General laboratory methods for proteins The proteins were resolved based on their size using standard SDS-PAGE techniques and visualized using Coomassie Brilliant blue, silver staining, or by performing Western blotting where detection was done with primary antibodies and horseradish peroxidase-conjugated secondary antibodies


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(Jackson Immunoresearch). Concentrations of the proteins were measured using absorbance at 280 nm wavelength, BCA assay (Thermo Scientific), and comparing intensity of the protein bands with a standard in the gel using ImageJ program (http://rsb.info.nih.gov/ij/). Buffer exchanges were performed using dialysis membranes or PD-10 buffer exchange columns (GE Healthcare).

3.2.2 Protein iodination Proteins were iodinated using the Iodogen method in a hood of a class B radioactive work area (Fraker et al. 1978). Iodogen tubes were prepared by mixing Iodogen to chloroform and dried. Iodination was performed by incubating 20-100 µg of protein with 0.2-1.0 mCi of 125-I for 10-20 minutes in PBS. After the reaction the mixture was transferred to a glass tube for unreacted iodine ions to form molecular iodine I2. The free iodine was separated from proteins using PD-10 buffer exchange columns. The specific activity of the proteins was typically 1.0 - 4.0 x 106 cpm/μg (16.7 – 66.7 kBq/μg).

3.2.3 Mutagenesis, cloning, and screening of FH19-20 mutant proteins Mutagenesis was performed using QuikChange Multi site-directed mutagenesis kit (Stratagene). The mutagenesis primers were designed based on the structure of the FH19-20 protein and criteria presented by the manufacturer of the mutagenesis kit. The mutagenesis primers were obtained from Sigma-Aldrich and are presented in studies II and III. These were used to induce mutations D1119G, D1119G/Q1139A, Q1139A, W1157L, R1182A, W1183L, T1184R, K1186A, K1188A, E1198A, R1203A, R1206A, R1210A, and R1215Q to FH19-20. The pPICZαB vector (Invitrogen) with the inserted FH19-20 sequence was used as a template in mutagenesis PCR reactions. E. coli cells were transformed with a PCR product using a heat shock transformation as instructed by the manufacturer. The clones were selected using Zeocin® (InvivoGen) antibiotic selection on Luria-Bertani medium and then sequencing from 3’ and 5’ ends over the FH19-20 region in the Haartman Institute sequencing core facility. The plasmids harboring the mutation(s) were transferred to Pichia pastoris for expression using electroporation according to manufacturer’s instructions (BioRad). The transformed cells were selected using Zeocin® and small scale culture expression in 10 ml. The media was screened for the presence of FH19-20 proteins using Western blotting with polyclonal anti-FH19-20 antibody. The clones that were recognized by the antibody and had the highest expression level were selected for large scale expression.

3.2.4 Expression and purification of FH19-20 mutant proteins The FH19-20 proteins were expressed using the Pichia pastoris expression system as described by the manufacturer (Stratagene). All the incubation steps were performed at 28 °C under continuous shaking (225 rpm). Briefly, 5 ml of BMGY (Buffered Methanol, Glycerol, Yeast extract) medium with 100 µg/ml of Zeocin® was inoculated and incubated for approximately 17 hours. The culture was used to inoculate 50 ml of pre-tempered BMGY that was incubated for 16 hours. The cells were harvested by 5 min centrifugation at 3000 g at 22 °C. The cells were suspended to 1 liter of BMM (Buffered Methanol Medium) medium for expression. 10 ml of methanol was added to the culture after 24 hrs and 48 hrs of expression. The expression was stopped after 72 hrs and the subsequent


35

steps were performed at 4 °C. The supernatant was collected by centrifuging the cells twice at 5000 g for 5-10 min. The supernatant was filtered using 0.8 µm/0.2 µm filters (Pall). The pH of the filtrate (pH < 5) was adjusted to 6.0-6.5 with Na2HPO4 due to stability of the Hi-Trap heparin column (pH should be >5) and diluted 1+1 with H2O. To prevent microbial growth Na-azide (0.02% final concentration) was added. The proteins were purified from the filtrate using 1 ml Hi-Trap heparin columns (GE Healthcare) and eluted with 1 M NaCl in phosphate buffered saline (PBS: 137 mM NaCl, 2.8 mM KCl, 9.7 mM phosphate, pH 7.4). The protein preparations were further purified with Sephadex 75 (10/300 GL) gel filtration column using the ÄKTA® purifier system (GE Healthcare).

3.2.5 Generation of C3b The C3b protein was prepared from plasma purified C3 (kindly provided by MSc Karita Haapasalo)

using trypsin cleavage. 1 mg /ml of C3 in PBS was warmed in a water bath to 37 °C. Trypsin (Sigma-Aldrich) was added to the solution to make it 1 % (w/v), mixed, and incubated for 3 min. Soy bean trypsin inhibitor (SBTI)(Sigma-Aldrich) was added to the solution to make it 2 % (w/v) and mixed to stop the reaction. The result was monitored with SDS-PAGE gels, where a shift in the alpha chain of C3 indicated cleavage of C3a from C3 and generation of alpha’-chain. Trypsin, SBTI, C3a, and C3b were separated with HiLoad 16/60 Sephadex 200 gel filtration column attached to an ÄKTA® purifier system (GE Healthcare).

3.2.6 Radioligand assays The proteins were attached to Nunc Polysorp Break Apart plates at 5-20 μg/ml in ½ PBS (68.5 mM NaCl, 1.4 mM KCl, 4.85 mM phosphate, pH 7.4) by incubating for 17 hours at 4 °C. The wells were blocked with 0.5 - 2.0 % bovine serum albumin (BSA, Sigma-Aldrich) in ½ or full PBS for 2 hours at 22 °C and washed 1-2 times with ½ or full PBS. In inhibition experiments the radioligand and inhibitor were usually premixed. In direct binding experiments the radioligand was pipetted onto the wells bearing the ligand. The plates were incubated for 2 - 3 hours at 22 °C and washed with ice cold ½ or full PBS once or twice. The radioactivity of the separated wells was measured with a Wizard gamma counter (Perkin-Elmer).

3.2.7 Surface plasmon resonance Surface plasmon resonance assays were performed on a Biacore 2000 equipment using carboxymethylated dextran chips CM4 and CM5 (GE Healthcare). For the amine coupling procedure the proteins were dialyzed against 20 mM sodiumacetate buffer (pH 4.5). The optimal amount of protein coupled to the chip in kinetic assays was decided based on the surface density calculator (www.biacore.com). The reference flow cell was activated and deactivated using the amine coupling procedure. The assays were performed at 22° C in ½ or full PBS using a flow rate of 30 µl /min in kinetic experiments.


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3.2.8 Heparin affinity chromatography Heparin affinity chromatography was performed with the ÄKTA® purifier system (GE Healthcare) and Hitachi LaChrom HPLC (Merck). The proteins were injected into a 1 ml Hi-Trap Heparin column (GE Healthcare) in ½ PBS and eluted using a linear salt gradient up to 0.5 - 1.0 M NaCl in PBS. The elution was monitored using absorbance at 280 nm and by Western blotting the elution fractions with anti-FH19-20 antibodies. The ionic strength of the elution buffer was monitored in real time with a conductivity meter in ÄKTA® or measured from the elution fractions using a conductivity meter CDM210 (Radiometer Analytical SAS).

3.2.9 Enzyme immunoassay on mGEnC-1 The mGEnC-1 were grown to confluence at 37° C under 5.0 % CO2 atmosphere. The cells were detached, washed with PBS and activated with 10 ng/ml of TNF-α (Peprotech, Rocky Hill, NJ, USA) on 96 well plates for 18 hours. The cells were washed with PBS and the FH19-20 proteins were added in 2 % BSA/PBS onto the cells and incubated for 2 hrs at 37° C under 5.0 % CO2 atmosphere. The proteins were detected with polyclonal rabbit anti-FH19-20 antibody and horse radish peroxidase (HRP) conjugated donkey anti-rabbit IgG F(ab’)2 (Jackson Immunoresearch). Antibody incubations were performed in 2 % BSA/PBS and the cells were washed with 0.05 % Tween 20 in PBS between the incubations. The HRP-conjugated antibodies were detected using 100 µl of tetramethylbenzidine (TMB) substrate solution (SFRI Laboratories) for 15 min at 22° C after which the reaction was stopped with 100 µl of 2 M H2SO4. The absorbance (at 450 nm) of the wells was measured immediately after stopping the reaction.

3.2.10 General bioinformatical tools Molecular modeling was performed at Finnish IT Center for Science (www.csc.fi) using Insight II (Accelrys Inc.). Sequences were analyzed using tools of Expert Protein Analysis System (http://au.expasy.org/) (Gasteiger et al. 2003), Baylor College of Medicine Search Launcher (http://searchlauncher.bcm.tmc.edu/) (Smith et al. 1996), and PFAM (Finn et al. 2009). The sequences were aligned mainly with BLAST at NCBI or EBI (Altschul et al. 1990), and ClustalW at EBI (Thompson et al. 1994) and resulting multiple sequence alignments visualized and analyzed using GeneDoc program (Nicholas et al. 1997). The molecular images were created with PDBviewer (Guex et al. 1997).

3.2.11 Determination of SCR consensus sequences All the SCR containing proteins in the SwissProt-database release 40 (24.10.2001) (www.ebi.ac.uk) were obtained using SRS Sequence Retrieval System. The proteins and the corresponding SCRs were divided into three datasets RCA1: (CFAH (P08603), FHR1 (Q03591), FHR2 (P36980), FHR3 (Q02985), FHR4 (Q92496), and FHR5 (Q9BXR6)), RCA2: (C4BPA (P04003), C4BPB (P20851), CR1 (P17927), CR2 (P20023), DAF (P08174), and MCP (P15529)), and non-RCA: (APOH (P02749), C1S (P09871), C1R (P00736), CFAB (P00751), CO2 (P06681), CO6 (P13671), CO7 (P10643), MASP1 (P48740), F13B (P05160), IL2RA (P01589), LYAM1 (P14151) , LYA2 (P16581), LYAM3 (P16109), MASP2 (O00187), PAPP1 (Q13219), PGCA (P16112), CSPG2 (P13611), SRPX (P78539), and SE6L1 (Q9BYH1)). The codes


37

present the UniProt (previously SwissProt) entry names in the form ABBREVIATION_HUMAN (accession code). Representative experimental structures of the SCR domains (CR1 domains 15, 16 (PDB code 1GKG); CR1 17 (1GKN); CR2 1, 2 (1GHQ); DAF 2 (1NWV); DAF 3, 4 (1H03); FH 15 (1HFI); FH 16 (1HCC); MCP 1, 2 (1CKL)) coded by the RCA cluster were obtained from Protein Data Bank (Barlow et al. 1993; Casasnovas et al. 1999; Smith et al. 2002; Uhrinova et al. 2003; Williams et al. 2003). Multiple sequence alignment was created using three-dimensional structures of the SCRs that were aligned using Combinatorial Extension (CE) method (Shindyalov et al. 1998). For a pair of structures CE determines aligned fragment pairs that are used heuristically to create final optimal alignment between the structures. Pairwise alignments from CE were used to generate multiple sequence alignment of experimentally solved SCR domains with emphasis on the conservation of the extended beta-strand regions denoted in PDB. Based on the degree of amino acid conservation, the columns in the alignment were scored from 1 to 9 (12/12 hits yielded score of 9 ((X/12)*9 = score)). This sequence profile was loaded to ClustalX-program (Thompson et al. 1997), where SCR domain sequences without known structure were heuristically added one by one to the profile to create a full alignment. Multiple sequence alignment was manually refined with emphasis on the conserved cysteines and the secondary structure elements. The consensus sequences were produced from the multiple sequence alignment using a GeneDoc program (Nicholas et al. 1997).

3.2.12 Analysis of interdomain interactions The interdomain contact regions were calculated from experimentally-solved SCR domain pairs deposited in RCSB Protein Data Bank (FH 15-16 (1HFH); MCP 1-2 (1CKL); VCP 2-3 and 3-4 (1VVC); APOH 1-2, 2-3, and 3-4 (1C1Z); CR1 15-16 and 16-17 (1GKN, and 1GKG)) (Barlow et al. 1993; Casasnovas et al. 1999; Schwarzenbacher et al. 1999; Szakonyi et al. 2001; Smith et al. 2002). Sting Millenium suite (Neshich et al. 2003) was used to calculate the contacts between the SCR domain pairs. The interdomain angles were measured using coordinates imported from modeling program package Insight II (Accelrys Inc) to Microsoft Excel, where data were processed. The conserved cysteines were used as reference points to calculate the tilt, twist, and skew angles that define the interdomain orientation. Tilt defined the depth of the angle between the consecutive domains. Skew defined the angle in which the C-terminal domain was tilted relative to the N-terminal domain. Twist defined how much the C-terminal domain was rotated relative to the N-terminal domain. The interdomain contact regions were extracted from SCR domain sequences in SwissProt-database release 40 (24.10.2001) (ebi.ac.uk/swissprot) and their properties calculated using self made Perl scripts (See study I for further details).

3.2.13 X-ray crystallography The FH19-20 and FH19-20D1119G/Q1139A, proteins were purified to 99 % purity and used in crystallization experiments. The crystallographic experiments and structure determination were performed in Professor Adrian Goldman’s laboratory by MSc Arnab Bhattacharjee, PhD Veli-Pekka Jaakola, and PhD Tommi Kajander.


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3.2.14 Curve fitting and statistical analyses Curve fitting was done using non-linear regression models implemented in GraphPad Prism®. For HPLC data Unicorn® program of ÄKTA® system was used to analyze the elution curves (GE Healthcare).

The statistical analyzes were performed using Microsoft Excel® and GraphPad Prism®. The comparison of mean values was done using combination of F-tests and t-tests, and ANOVA. The results were presented with mean using standard deviation to indicate the error.

Results

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4 Results

4.1 Rational mutagenesis of FH19-20

4.1.1 Identification of conserved amino acids in SCRs The SCR domains have a similar fold with each other but they mediate a variety of functions in different biological systems. This indicates that the conserved amino acids in the SCR domain family are important for the fold of the domain but not necessarily for their function per se. By avoiding mutagenesis of the conserved structural amino acids it is more likely that the fold of the SCR is conserved upon mutant protein expression and that the mutation has only local effect in the binding interface. To identify the structurally important amino acids, consensus sequence of SCR domains and SCR-SCR interdomain contact regions (ICRs) were identified and analyzed.

Sequence data of 177 SCR domains and 13 experimentally solved SCR structures were obtained from publicly available databases to elucidate the structurally important amino acids in the SCR domain family (Lehtinen et al. 2003; Lehtinen et al. 2004). Based on the location of the genes coding SCR domains, the SCR-containing proteins were further divided to regulators of complement activation group 1 (RCA1) at locus 1q31.3 (48 SCRs of FH, FHL-1 and FHRs 1 to 5), RCA2 group at 1q32.2 (64 SCRs of CR1, CR2, C4bp, DAF, and MCP), and to non-RCA group of domains whose genes are scattered in the genome (total of 65 SCR domains). Multiple sequence alignment of the SCRs based on the experimental SCR structures was created and analyzed. The consensus sequences of the SCR domains in RCA1, RCA2, and non-RCA groups had some clearly identifiable features (Figure 11).

The four characteristic cysteines that make the disulphide bridges in the SCRs were the only fully conserved amino acids in this analysis (sites 1, 42, 76, 102 in Figure 11). Some of the SCR domains in the non-RCA group had two additional cysteines (sites 16 and 38) that were located in the beginning and at the end of the functionally important hypervariable loop. Most of the semi-conserved sites correlated with the location of the beta-strands in the SCR structures except the proline rich areas at the beginning (sites 2-5) and at the end (sites 97-100) of the consensus sequences. The differences between the RCA1 and RCA2 group consensus sequences were mainly due to insertions after the Cys at site 42 and between Gly at site 80 and Trp at site 86. Factor H domains 19 and 20 that were subjected to mutagenesis in the further studies had typical sequences for SCR domains, except the N-terminus of FH20 that was found to have some unusual features. It lacked the typical proline rich region after the first cysteine (sites 2-5) and the NG motif at sites 11-12 in the hypervariable loop (Figure 11).

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Figure 11. Structure based consensus sequences of SCR domains. Consensus sequences of all SCR domains (first line) and the subgroups of RCA1, RCA2, and non-RCA are shown with FH19 and FH20 sequences. Fully conserved cysteines are marked with white font and black background. Amino acids marked with white and grey background are conserved in 25-50 % and in 50-99 % of SCR domains within the group, respectively. Dash indicates a gap and a dot indicates occurrence of at least one amino acid on that position in the multiple sequence alignment. Numbers above the alignment indicate the site of the amino acid in the multiple sequence alignment.

4.1.2 Identification of interdomain contact regions between SCRs One goal of study I was to identify ICRs, characterize them and to examine their impact on orientation between the neighbouring SCR domains. SCR domains in proteins are arranged like beads-on-a-string connected by short linker peptides. This conformation allows adjacent domains to interact with each other and to form contiguous binding sites. The amino acids that mediate the interdomain interactions should not be subjected to mutagenesis in binding site mapping studies because disruption of the interface may lead to loss-of-function due to change in the interdomain orientation instead of disruption of the interface. Most of the experimental SCR structures have been solved for pairs of SCR domains that made it possible to study the interactions of neighbouring domains in silico. Previously published experimental structures of the SCR domain pairs were analyzed using Sting Millenium suite (Neshich et al. 2003) in order to detect the amino acid residues that made contacts to adjacent domains and formed the linker area between the SCRs. Four sequence stretches N#1, C#1, C#2 and the interdomain linker were identified in conserved locations that formed the ICRs. These locations can be used to predict ICRs in SCR domain sequences, as was done for FH19-20 (Figure 12). Sequence analyses of the ICRs revealed that they were more hydrophilic than human protein sequence on average and that the increasing interdomain linker length correlated with increasing hydrophilicity. It was concluded in study I that in general the ICRs are probably flexible and that the short linkers form more compact and rigid interface between the adjacent domains, as would be expected.

To analyze the impact of ICRs to orientations of the neighboring SCR domains, tilt, twist, and skew angles were calculated from experimental structures that describe the orientation between the domains (See methods for explanation of the angles). The twist and skew angles were found to

Results

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depend on each other due to steric restriction by the linker and the N#1 region. The interdomain angles were also compared to amino acid sequences of the ICRs but no correlation was found.

Figure 12. Predicted and experimental interdomain contact regions in FH19-20. The predicted ICRS are shown with curved arrows above the FH19 and below the FH20 sequences and are mapped to FH19-20 structure on the left (PDB ID: 2G7I). The experimental ICRs are shown with grey background of the amino acids in the alignment and mapped to FH19-20 structure on the right. N#1-regions are colored yellow in the structures, linkers green, C#1-regions red, and C#2-regions blue.

4.1.3 Crystal structure of FH19-20 Computer based predictions are a valuable aid in designing point mutations to functionally important amino acids, however, the understanding of the impact of the mutations to the structure of the protein requires a detailed atomic structure. In study II the crystal structure of FH19-20 (FH amino acids G1109 - K1230) was solved at 1.8 Å resolution (Figure 13). The structure in the crystal of FH19-20 was similar to previously solved SCR domain pair structures with two bead-like domains arranged head-to-tail.The interdomain orientation was also typical. However, FH20 had two exceptional features, a short alpha helix (R1171 - Y1177) in the hypervariable loop and a buried water molecule that interacted with C1167, K1188 and S1191. Interestingly, the alpha helix site was located in the non-typical hypervariable loop of FH20 identified in the SCR consensus sequence analysis (Figure 11). In the crystal, the FH19-20 monomers formed a tight D2 tetramer structure (A, B, C, and D monomers) with two dimerization interfaces (AxB and AxC) in one monomer (Figure

Results

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4.1.3) (Table 1). The heparin binding sites of FH19-20 monomers A and D, and monomers B and C point to same direction, but these pairs do not have dimerization interfaces in the crystal. Dimerization (and oligomerization) of FH19-20 has been shown in vitro and suggested to be enhanced by polyanions, but the physiological meaning of this has remained unclear (Perkins et al. 1991; Pangburn et al. 2009). Based on the structure, polyanion binding should not enhance dimerization but could be involved in stabilizing tetramers.

Figure 13. The crystal structure of FH19-20. A. Monomer of FH19-20 with otherwise typical appearance of SCR domain pair except the alpha helix on SCR20. B. The tetramer of FH19-20 as it was crystallized. One monomer (e.g. A) contacts two monomers (B and C) that are upside down compared to A. This means that heparin binding to FH19-20 cannot stabilize dimers but may stabilize tetramers.

Table 1. Dimerization interfaces in the crystal structure of FH19-20. AxB interface (880 Å2) is composed of interfacing residues: K1108, D1119-A1129, Q1137-Y1142, A1185-Y1190, R1192, and E1195. AxC interface (982Å2) is composed of interfacing residues: K1108-I1120, F1123-V1127, Q1139-Y1142, G1155, Q1156, H1165-I1169, R1171, E1172, and Q1187-Y1190. H = hydrogen bond, SB = salt bridge. Interface AxB Interface AxC Residue in A Bond Residue in B Residue in A Bond Residue in C R1192 H T1121 N1117 H N1117 S1126 2H A1185 N1117 H D1116 V1127 H A1185 Q1139 H Q1139 Q1156 H E1172 H1165 H, SB D1116 Y1190 H P1112

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4.1.4 Crystal structure of FH19-20 and C3d complex In study IV, the complex structure of FH19-20 with C3d was solved at 2.3 Å resolution (Figure 14). The wild-type FH19-20 did not form crystals with C3d but crystallized easily on its own as a tetramer. The mutant FH19-20D1119G/Q1139A protein was used in crystallization procedure because it was found to have decreased potential to form the tetramer. Overall, the structures of FH19-20 and C3d were almost identical with the previously solved structures of FH19-20 or C3d alone, only FH19-20 had some local changes upon binding to C3d. Also, there were no significant changes in the interdomain orientation of FH19-20 D1119G/Q1139A in the FH19-20:C3d complex compared to that found in the FH19-20 tetramer. The D1119G and Q1139A mutations were modeled back to original residues on the complex structure and found to fit well into the interface, and thus FH19-20 D1119G/Q1139A will be later, and was in the study IV, referred to as FH19-20. The existence of the two binding sites was confirmed by analyzing binding of the C3dg point mutations in FH19 and FH20 interfaces in study IV.

In the FH19-20:C3d complex, importantly, two different interfaces were found between FH19-20 and C3d in contrast to previous reports that assumed 1:1 binding (Figure 14). The sites were called the FH19 site and the FH20 site. The FH19 site on FH19-20 (FH19-20FH19) stretched from domain 19 to the linker region and to the hypervariable loop of domain 20, and covered a buried surface area of 620 Å2. The FH19-20FH19 consisted of four peptides on FH19-20: the hypervariable loop of FH19 (P1112-P1124), the stretch around the second cysteine and the loop after it (Q1137-Y1142), the linker region (H1165-V1168) and the C-terminal part of the hypervariable loop of FH20 (K1188-Y1190). On C3d, the FH19 site (C3dFH19) is located at the side of the α/α-barrel. The interface mainly consisted of four peptides: the N-terminus of helix 4 (K112-K118), the loop around P121 (Q119-Q126), the loop between helix 6 and 7 (E167-S171) and a stretch in the middle of helix 7 (K178-F182) (C3d helices and numbering as in Nagar et al. 1998).

The FH20 site on FH19-20 (FH19-20FH20) was located exclusively on domain 20 and covered a buried surface area of 490 Å2. The binding site consisted of three peptides: the N-terminus of hypervariable loop of FH20 (R1171-T1184), the stretch around the second cysteine and the loop after it (V1200-Y1205), and the C-terminus of domain 20 (K1230-R1231). On C3d, the FH20 site (C3dFH20) was formed around a negatively-charged pocket on the concave top of the α/α-barrel. The binding site consisted of four peptides: seven amino acids in helix 1 - turn - helix 2 region (H33-L53), helix 3 and the loop after it (S94-I102), the turn between helices 5 and 6 (E160-D163), and the C-terminus of helix 12 (Q284-D292). The FH20 interface had many charged residues and was observed to be highly electrostatic. The crystal structures of the FH19-20 homotetramer and the FH19-20:C3d complex were the basis for designing mutagenesis on FH19-20 and allowed the structural interpretation of the functional effects of the mutations and intriguingly revealed the molecular basis of the target discrimination by factor H.

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Figure 14. The crystal structure of FH19-20:C3d complex revealed two binding sites. A. FH19 binding site. B. FH20 binding site. In A and B FH19-20 is shown with cartoon presentation. The residues that make bonds to C3d are shown with sticks. Green indicates SCR19, gold indicates SCR20, and red indicates linker residues between SCR19 and SCR20. C3d is shown with surface presentation. Blue color indicates the FH19 site, green color the FH20 site, and yellow color the location of thioester that mediates covalent binding to surfaces. C. FH19 and FH20 binding sites on C3d structure. C3d fold forms inner and outer α-barrel (α/α-barrel) that is clearly visible in the lower panel. FH19 binding site is located on the side of the barrel (blue coloring) and the FH20 site on the concave top (green coloring). C3d is shown with ribbon presentation and the residues in the FH19 site and in the FH20 site with sticks.

4.1.5 Mutagenesis of FH19-20 For analyzing the physiological function of FH19-20 and functional consequences of aHUS mutations, a total of 17 mutant FH19-20 proteins were created using PCR and Pichia pastoris expression system in studies II, III, and IV (Figure 15 and Table 2). The selection of the point mutations was based mostly on crystallographic structures and the association of the mutation with aHUS, but also on the previously published biochemical data on FH function, like the known involvement of arginines and

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lysines in heparin binding (Table 2). The mutations were also compared to interdomain contact regions and conserved amino acids of SCRs to avoid global changes in the structure. In studies II and III mutations D1119G, D1119G/Q1139A, Q1139A and W1157L were induced to domain 19 and R1182A, W1183L, K1186A, K1188A, E1198A, R1203A, R1206A, R1210A, and R1215Q to domain 20 to study their impact on C3d, C3b, heparin, and cell binding. In study IV a Q1137A/ Q1139A/ Y1142A triple mutation protein was designed to knock-out the FH19 site (FH19Del-20) and a T1184G/ K1202A/ R1203A/ Y1205A quadruple mutation was designed to knock-out the FH20 site (FH19-20Del). All the FH19-20 mutant proteins bound polyclonal anti-FH and anti-FH19-20, heparin (except FH19-20Del), C3d, and C3b (Studies II, III, and IV) suggesting that the antigenicity of the FH19 or FH20 had remained intact.

Figure 15. Mutations induced to FH19-20. The residues that were mutated are shown with sticks and CPK coloring.

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Table 2. Mutations in the generated mutant proteins used in studies II, III, IV and their role in structure of FH19-20. The mutations were selected based on their association with aHUS (aHUS), their positive charge that indicates a heparin binding site (+ charge), the location in the dimerization interface (dimerization), or in the C3d binding site (FH19Del-20 or FH19-20Del). None of the mutations were located in the interdomain contact regions except Y1142A in FH19Del-20. Mutation aHUS

association Conserved Secondary

structure Side chain accessible

Selection criteria

D1119G D1119G No Beta Yes aHUS, dimerization Q1137A No Yes FH19Del-20 Q1139A No Yes FH19Del-20, dimerization W1157L W1157R Yes Beta No aHUS Y1142A Yes Only tip FH19Del-20 R1182A R1182S No Yes aHUS, + charge W1183L W1183L/R No Yes aHUS T1184R T1184R No Turn Yes aHUS T1184G No Turn Yes FH19-20Del K1186A No Yes + charge K1188A No Yes + charge L1189R L1189F/R/P No Yes aHUS E1198A E1198A/K Yes Beta Only tip aHUS K1202A No Yes FH19-20Del R1203A R1203W No Turn Yes aHUS, + charge, FH19-20Del Y1205A No Yes FH19-20Del R1206A R1206C No Beta Yes aHUS, + charge R1210A R1210C No Turn Yes aHUS, + charge R1215Q R1215G/Q No Beta Yes aHUS, + charge

4.1.6 Structure function analysis of FH19-20 mutations on C3d binding In studies II and III binding of the FH19-20 mutant proteins to C3d and C3b was tested using radioligand assays (RLA) and surface plasmon resonance (SPR) technique to map the binding site for C3d/C3b without knowledge about the actual binding site (Table 3). In study IV, the FH19-20:C3d structure was solved allowing interpretation of the results of the functional analyses in studies II and III on the basis of the real interaction in FH19-20:C3d structure (Figure 16). The mutations D1119G, Q1139A, T1184R, K1188A, and R1203A lead to loss of bonds between C3d and FH19-20 indicating that these mutations should lower the affinity in the experiments as well. However, the mutant proteins FH19-20D1119G and FH19-20T1184R did not have statistically-decreased binding in all assays (Table 3). The FH19-20 binding to C3d is mediated by two low affinity binding sites that possibly mask the effect of a single point mutation in one site.

Six mutations W1157L, R1182A, W1183L, E1198A, R1206A, and R1210A were not directly in the FH19-20:C3d interfaces but caused decreased C3d or C3b binding by FH19-20. The analysis of the structure showed that W1157L and W1183L probably destabilize the fold of SCR19 or SCR20, respectively. R1182A, E1198A, R1206A, and R1210A lead to change of charge on SCR20 and probably disrupt the electrostatics-based orientational steering that can significantly effect the affinity of protein-protein interactions (Janin 1997; Sinha et al. 2002; Persson et al. 2009). The mutant proteins

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FH19-20R1215Q, FH19-20L1189R, and FH19-20K1186A had wild-type C3d/C3b binding, consistent with the location of the mutations far away from the C3d binding sites in the complex structure.

Figure 16. Effects of FH19-20 mutations on binding to C3d/C3b in functional assays and their location on FH19-20:C3d structure. FH19-20 shown in ribbon and two C3d’s in surface presentation. Color scheme is as in Figure 14. The mutations that were analyzed in functional assays are shown in sticks. The interpreted mechanical effects of the mutations on loss of FH19-20-C3d interaction: blue destabilization of SCR, red loss of a bond in the interface, purple loss of electrostatic complementarity in FH19-20-C3d interface. Residues marked with grey had no effect on binding to C3d in functional assays.

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Table 3. Functional consequences of FH19-20 mutations on binding to C3d/C3b and interpretation of the mechanism on the basis of the FH19-20:C3d structure. All the results published in studies II, III, and IV, except *(Lehtinen et al. 2008), # (Bhattacharjee et al. 2010), and () unpublished. ↓ decreased binding, ↑increased binding, and ↔ wild-type like binding. Mutation(s)

aHUS association

Bond in binding

site

C3d binding

RLA

C3d binding

SPR

C3b binding

RLA

Mechanism

D1119G D1119G FH19 ↔ ↓* ↔ Loss of H-bond D1119G/Q1139A

FH19 ↓ ↓* ↓ Loss of 2 H-bonds

Q1139A FH19 ↓ ↓* ↓ Loss of H-bond W1157L W1157R ↓ ↓* ↓ Destabilization R1182A R1182S (↓) ↓ ↓ Possibly loss of

electrostatic complementarity

W1183L W1183L/R (↓) ↓ ↓ Possibly disruption of loop V1168-S1191

T1184R T1184R FH20 ↔ ↔* ↓ Loss of 2 H-bonds K1186A (↔) n.d. ↔ None K1188A FH19 (↓) ↓ ↓ Loss of salt bridge L1189R L1189F/R/P ↔ ↔* ↔ None E1198A E1198A/K (↓) ↔ ↓ None or possibly loss of


R1203A R1203W FH20 ↓# ↓* ↓# Loss of 2 H-bonds and 3 salt bridges

R1206A R1206C ↓ ↓* ↓ Possibly loss of electrostatic

complementarity R1210A R1210C ↓ ↓* ↓ Possibly loss of


R1215Q R1215G/Q ↔ ↔* ↔ None

4.2 Self - non-self discrimination by FH19-20

4.2.1 FH19 site is exposed and FH20 site is partially occluded on C3b In study IV, the FH19-20:C3d structure revealed that there were two binding interfaces between FH19-20 and C3d. The affinities of FH19Del-20 and FH19-20Del to C3d and C3b were measured using SPR. It was found that the FH19 site had similar affinities to both (Table 4), but the FH20 binding site had roughly four times higher affinity to C3d than to C3b. This discrepancy was structurally explained by analyzing the availability of FH19 and FH20 sites on the C3b and C3b:FH1-4 structures (PDB IDs: 2I07 and 2WII). The FH20 site in the C3d part of C3b (D1007-Q1021) was occluded by the MG1 domain belonging to the C3c part of C3b (residues F40-K43; R80-V98; Q631-A636). Although the FH20 site is partially buried in the C3b crystal it is available on soluble C3b, since FH19Del-20 clearly bound to it (Table 4). Thus it seems that C3c might act as a competitive inhibitor for FH19-20 on FH20 site binding.

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Wu and co-workers compared the contacts between C3d and C3c in C3b and FH1-4:C3b structures and found that the FH1-4 binding tightens up the C3d-C3c interface by introducing hydrogen bonds to the interface that also leads to change in orientation of C3d (12 degree rotation) (Wu et al. 2009). In addition, FH4 was bound partially onto the FH20 site on C3d in the FH1-4:C3b complex that could further block the FH19-20 binding. These findings indicated that FH1-4 binding to C3b favors binding of FH19-20 to the FH19 site since the C3c binding to FH20 is enhanced.

Table 4. Relative affinities of FH19 and FH20 sites on C3d and C3b binding. C3d and C3b columns: The numbers present relative decrease in affinity compared to FH19-20 that is 1.0. Fold change column: For each protein the number presents decrease of affinity on C3b binding compared to C3d binding. Protein C3d Fold change C3b FH19-20 1.0 3.0 3.0 FH19Del-20 2.3 4.4 10.2 FH19-20Del 6.3 1.3 8.2

4.2.2 FH20 mediates binding to heparin and mGEnCs The heparin binding site of FH19-20 was mapped to FH20 in studies II and III using heparin affinity chromatography and the mutant proteins (Table 5). Heparin binding was found to be mediated by residues R1182, K1186, K1188, R1203, R1206, R1210, and R1215. These results were corroborated by the results of Herbert and co-workers, who analyzed the perturbations in the NMR-spectra upon addition of heparin tetrasaccharide to FH19-20 solution (Herbert et al. 2006). In that study, the heparin binding site was also mapped to lysine and arginine residues on FH20 (Table 5), but in addition, the residues E1145, L1164, H1165 and T1193 in the hinge between FH19 and FH20 were affected by heparin binding. The authors suggested that these changes were most probably structural and might have an effect on the interdomain orientation. The heparin binding site was also confirmed by Ferreira and co-workers who used FH19-20 mutant proteins in heparin affinity chromatography and gel mobility shift assays (Table 5) (Ferreira et al. 2009). These data were collectively used in study IV to map the heparin binding site on FH19-20 of the FH19-20:C3d complex structure (Figure 17). The heparin binding site was found to extensively overlap with the FH20 binding site (Table 5), but not with the FH19 site that shared only the K1188A residue with the heparin site. These findings were analyzed experimentally using the FH19Del-20 and FH19-20Del mutant proteins in a radioligand assay where binding of only FH19Del-20 to C3d was inhibited by heparin. Importantly, this result showed that the FH19 site is not affected by heparin binding and can bind simultaneously to C3d or C3b while heparin is bound to FH20.

Heparin binding has been used to model FH binding to HS glycosaminoglycans that are present also on glomerular endothelial cells and glomerular basement membrane. Binding of the FH19-20 mutant proteins to mouse glomerular endothelial cells (mGEnCs) was analyzed in study III, and it was found that the binding site of FH19-20 to mGEnCs and was almost identical with the heparin binding site on FH20 (Figure 17 and Table 5). The only differences were that the mutation K1186A increased binding to mGEnC-1 but decreased binding to heparin and that the mutation R1215Q had no effect on mGEnC binding but decreased binding to heparin.

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Figure 17. Heparin and mGEnC binding sites on FH20. FH19-20 shown in ribbons bind to surface contoured C3d via FH19 site (Coloring as in Figure 14). The view is from the surface that is indicated by the thioester shown in yellow in the left lower corner. Red residues shown in sticks indicate heparin binding sites and blue residues indicate mutations that increase affinity to heparin. The bold labels indicate residues that similarly affected binding to mGEnC and heparin.

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Table 5. Comparison of functional consequences of FH19-20 binding to heparin and cells. ↓ decreased binding, ↑increased binding, ↔ wild-type like binding, + perturbation in NMR spectra, and # (Bhattacharjee et al. 2010). Residue or Mutation

Heparin binding

(Study III)

Heparin binding

(Ferreira et al. 2009)

Heparin binding by NMR

(Herbert et al. 2006)

mGEnC-1 binding

(Study III)

Overlap with C3d binding

sites (Study IV)

E1145 + L1164 + H1165 + E1172R ↑ R1182A ↓ + ↓ R1182S ↓ + W1183R ↑ FH20 T1184R ↑ ↑ ↑ FH20 K1186A ↓ + ↑ K1188A ↓ + ↓ FH19 K1188Q ↓ + FH19 L1189F ↑ L1189R ↑ ↑ ↑ T1193 + E1198A ↑ ↑ E1198K K1202 + FH20 R1203A ↓# + FH20 R1203S ↓ + FH20 R1206A ↓ ↓ R1210A ↓ ↓ R1210C ↓ R1210S ↓ R1215G ↓ + R1215Q ↓ + ↔ K1230A ↓ + FH20 R1231A + FH20

4.2.3 Simultaneous binding of FH19 site to C3b and FH20 to anionic self cell structures explains target recognition by FH

Based on FH19-20:C3d complex, its superimpositions onto C3b and FH1-4:C3b structures, and biochemical data, it was deduced that FH19-20 binding via FH19 site to C3d/C3b left the FH20 free to bind heparin or mGEnC (Figure 18). This indicated that FH19 site and FH20 are needed for self surface recognition. This was confirmed experimentally by analyzing the binding of FH19-20 to C3b-coated sheep erythrocytes (Esh-C3b) that have been widely used to model self surfaces. FH19Del-20 and FH19-20Del proteins had decreased binding to Esh-C3b, as expected. In addition, the binding of FH19-20 to the FH19 site on FH1-4:C3b complex did not interfere with FH1-4 binding that mediates the CA and DAA functions of FH. This was also experimentally confirmed by performing CA and DAA assays with FH19Del-20 and FH19-20Del mutant proteins in study IV.

It was shown in study IV by structural analyses that C3b (via FH19 site) and C3d (via FH20 site) could bind simultaneously to a single FH19-20 molecule (Figure 18) and in studies III and IV that C3d and

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heparin binding sites on FH20 were overlapping. The FH20 site on C3d is anionic like heparin suggesting that FH19-20 could bind to self surface bound C3d similarly to polyanions.

The dual interaction of FH19 site to C3b and FH20 to heparin or cells is likely to present the physiological complex that recognizes self surfaces. These results are in very good agreement with all the previously published biochemical data about target recognition that had indicated simultaneous binding of FH to C3b and to polyanions on self surfaces (Pangburn et al. 2008).

Figure 18. Molecular basis of self – non-self discrimination by FH. A. Binding of FH19-20 to C3b:FH1-4 via FH19 site does not interfere with FH1-4 binding and allows simultaneous binding to heparin or cells. B. C3d binds to the same region site on FH20 that mediates binding to heparin and cells. The SCR domains are shown in ribbon and are colored blue in FH1-4, green in FH19, and gold in FH20. The residues that mediate binding to heparin are presented with red space fill on FH20. C3b is presented in surface contour with dark grey indicating C3d and light grey C3c parts of C3b. In B the C3d binding to heparin or cell binding site is shown with ribbons and in light blue color.

4.3 aHUS mutations disturb target recognition function of FH19-20 by various mechanisms

4.3.1 Disruption of the FH19 site Mutations D1119G, Y1142C/D, and V1168E cause loss of at least one bond in the interface and clearly must disrupt or weaken the binding to C3d (Figure 19). In vitro FH19-20D1119G bound weakly to C3d/C3b in some of the experiments (Table 3)(Ferreira et al. 2009). The mutations V1134G, E1135D, W1157R, and C1163W in domain 19 probably destabilize the fold leading to impaired binding of C3d (Figure 19), but this has not been proven experimentally.

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4.3.2 Disruption of the FH20 site Mutations T1184R and R1203W lead to loss of bonds in the FH20 site interface and thus should decrease the affinity of FH19-20 to C3d (Figure 20). Experimental results on the R1203A mutation clearly support this, but the experimental result on T1184R was controversial (Table 3). T1184R had no effect on binding of FH19-20 to C3d but decreased binding to C3b.

Mutations R1182S, E1198A/K, R1206C and R1210C or alanine mutations in these residues have been shown to decrease affinity to C3d/C3b (Table 3) (Sánchez-Corral et al. 2002; Ferreira et al. 2009). However, these mutations are not located in the FH20 binding site, but close to it. These mutations lead to loss of positive charge in FH19-20 that may disturb long-range electrostatic interactions between positively charged FH19-20FH20 and negatively charged C3dFH20, possibly explaining the loss of C3d/C3b binding in vitro.

4.3.3 Destabilization of FH20 may impair binding to C3dFH19, C3d FH20, or polyanions The mutations L1189F/R/P, W1183L/R, S1191L/W, G1194D, V1197A, F1199S, S1211P, and P1226S in FH20 are more or less buried and probably disturb the fold of the SCR20 and thus may disturb heparin or cell binding, or the FH19 or FH20 site. Some aHUS mutations in SCR20 are close to the FH19 site since it also expands to SCR20. For example, the residues K1188 and Y1190 that make contacts to C3dFH19 are located next to L1189F/R/P and S1191L/W and could disturb the FH19 site, but this has not been experimentally confirmed.

W1183L was shown to disturb only C3d/C3b binding but not heparin or mGEnC binding (Table 3 and 5). Patient-derived full length FH with W1183L and V1197A have been shown by others to decrease affinity to C3b (Sánchez-Corral et al. 2002).

4.3.4 Disruption of heparin or cell binding The mutations R1182S, R1203W, R1206C, R1210C, and R1215G/Q probably disturb binding to heparin and cells. FH19-20 mutant proteins R1182A/S, R1203A/S, R1206A, R1210A/S/C, and R1215G/Q have been shown to impair mGEnC and heparin binding by us (Table 5) and others (Manuelian et al. 2003; Sánchez-Corral et al. 2004; Ferreira et al. 2009). Exceptionally, R1215Q had only impaired heparin binding, but not mGEnC binding (Table 5).

4.3.5 Function of FH19 site and FH20 site is disturbed by enhanced heparin binding In studies II and III, the aHUS associated mutations T1184R, L1189R and E1198A were found to increase the affinity of FH19-20 to heparin and mGEnC (Table 5). Mutation W1183R has also been shown to increase affinity to heparin by others (Ferreira et al. 2009). In study III these mutations were observed to extend the positively charged heparin binding site on FH19-20 by either adding positive charge or by taking away negative charge (Figure 21). Furthermore, it was shown in study IV that enhanced heparin binding by the mutant T1184R lead to impaired binding of FH19-20 to C3d. Mutations W1183R, L1189R, and E1198A/K may have similar effects on the FH19-20 function. Taken together, the location of the mutations on the structure explains the increased affinity to heparin and decreased affinity to C3d that were observed experimentally.

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4.3.6 All aHUS mutations disturb FH19 site binding to C3b or FH20 binding to polyanions or cells

In conclusion, aHUS mutations may cause (i) loss of bond(s) in FH19-20 - C3d and/or heparin interfaces, (ii) defective long-range electrostatic interactions between C3d and FH19-20, (iii) enhanced binding to heparin that can inhibit binding of FH19-20 to C3d, or (iv) destabilization of the structure of the domains 19 or 20 that leads to disturbed binding to C3d and/or heparin. In addition, mutations may disturb binding of FH19-20 to the FH20 site on C3d that was suggested to participate in down-regulation of complement activation on self surfaces in study IV. It is clear that the aHUS mutations have various combinations of effects on C3d and heparin or cell binding, but the common denominator for the aHUS mutations seems to be impaired binding of the FH19 site to C3b and FH20 to heparin or cells that are crucial interactions for self surface recognition by factor H.

Figure 19. Atypical HUS mutations on FH19 site. FH19-20 is shown with ribbons and C3d with surface contour. FH19 is indicated with green and FH20 with golden color. The aHUS mutations are shown with sticks. Mutations to residues marked with red cause loss of bond(s) in the FH19 site and those marked with blue may cause destabilization of the SCR fold.

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Figure 20. Atypical HUS mutations on FH20 site. FH19-20 is shown with ribbons and C3d with surface contour. FH19 is indicated with green and FH20 with golden color. The aHUS mutations are shown with sticks. Mutations in residues marked with red cause loss of bond(s) in the FH20 site and those marked with blue may cause destabilization of the SCR fold. Mutations in residues marked with green probably disturb the electrostatic complementarity of the FH20 site.

Figure 21. Atypical HUS mutations on heparin or cell binding. FH19-20 shown with ribbons is binding to the FH19 site on C3d that is shown with surface contour. FH19 is indicated with green and FH20 with golden color. The aHUS mutations are shown with sticks. Mutations in residues marked with red decrease affinity to heparin and those marked with blue increase affinity to heparin that interferes with FH19 site and FH20 site binding to C3d.

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5 Discussion

5.1 Sequence and structure analysis in mutagenesis of FH19-20

In this thesis, structure-based multiple sequence alignment of SCRs was employed to identify conserved and structurally important amino acids of SCR family. Mutations in these residues would likely lead to destabilization of the SCR fold and thus not be informative in binding site mapping or functional studies. For example, W1157 was located in the core of SCR19 and in the SCR consensus sequence it was highly conserved (Figure 11 site 86). The mutation W1157L was shown to decrease binding to C3b (Table 3), but it was not located in the FH19 binding site (Figure 16). W1157L probably destabilizes the fold of the SCR and thus the experimental result on W1157L binding was misleading for mapping the C3d binding site on FH19-20 in study III.

Mutations to surface-exposed residues are more informative in terms of binding site mapping, since they are more likely to change structure only locally. This has been shown crystallographically for Q1139A and R1203A mutations on FH19-20 (Bhattacharjee et al. 2010) and with NMR assays for many others mutations in FH19-20 (Herbert et al. 2006; Ferreira et al. 2009). Q1139A and R1203A mutations also reduced binding to C3d in functional assays and were located in the FH19 and FH20 sites, respectively. On the other hand, some mutations outside the binding interface, like R1206 and R1210, also decreased affinity of FH19-20 to C3d in functional assays, indicating that there is always uncertainty in interpreting the results of these assays.

In study I the interdomain contact regions between SCR domain pairs were characterized based on the experimental SCR structures and found to be located in conserved sites. These sites were used in study I to design a model that could be used to predict ICRs. In this thesis, the ICRs were predicted for FH19-20 and found to be nearly the same compared to the actual ICRs in the crystal structure of FH19-20 (Figure 12) indicating that the model has true predictive value. Mutations to amino acids located in the ICRs were avoided in designing mutagenesis to FH19-20 in studies II, III, and IV to prevent possible changes in orientations between the two SCR domains. For example, the FH19 binding site was found to extend over the ICR between SCR19 and SCR20 in study IV and thus a change in the orientation between the SCRs would also lead to disruption of the FH19 site.

Taken together, bioinformatical analyses have clearly predictive value in designing mutagenesis to SCR domains that are not experimentally solved. This was shown by comparing the predicted ICRs and conserved amino acids to experimental structures and the results of functional assays.

5.2 Self – non-self discrimination by FH19-20

The molecular basis of the FH-mediated target discrimination has been a puzzling question in the complement field for decades. It has been shown already in the late 1970’s that negatively-charged surfaces bearing sialic acids and heparin were critical for protection of erythrocytes from complement attack by FH (Fearon 1978; Pangburn et al. 1978; Kazatchkine et al. 1979b). In 1990 the self - non-self discrimination of C3b deposited surfaces was found to be mediated by the polyanion binding site on FH (Meri et al. 1990b). Later, C3d/C3b and heparin binding sites were mapped on the FH19-20 region (Gordon et al. 1995; Sharma et al. 1996; Blackmore et al. 1998; Jokiranta et al. 2000)

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and FH19-20 was found to be critical for factor H activity on self surfaces (Pangburn 2002). However, at the same time, the C3d binding and heparin sites on FH19-20 were shown to be overlapping (Hellwage et al. 2002). This observation was enigmatic since previous results have suggested simultaneous binding of FH to C3b and heparin, which would strengthen the avidity of FH to C3b on self surfaces.

5.2.1 Molecular basis of self – non-self discrimination In study IV, it was found by crystallography that FH19-20 had two distinct binding sites for C3d, the FH19 site and the FH20 site. The validity of the crystal structure was confirmed by analyzing the binding of the FH19-20 wild-type and point mutant proteins to C3d and C3b, and binding of C3dg wild-type and point mutant proteins to FH19-20. The results of the functional assays were consistent with the binding interfaces in the crystal structure indicating that the interactions were not crystallographic artifacts. In addition, aHUS mutations have been described in the FH19 and FH20 site in both FH19-20 and C3d (Frémeaux-Bacchi et al. 2008; Maga et al. 2010).

When the FH19-20:C3d complex was superimposed on experimentally solved C3b and C3b:FH1-4 structures, it was found that only the FH19 site was fully available and that this conformation left FH20 free to bind heparin, mGEnC or C3d. Importantly, heparin did not inhibit binding of the FH19 site to C3d, and the FH19 and FH20 site were both found to be necessary in recognizing Esh-C3b. These results indicated that the FH19 site can bind to C3b and FH20 on self surfaces simultaneously. FH19-20 binding via FH19 site did not overlap with the FH1-4 binding on FH1-4:C3b structure and did not interfere with CA and DAA functions of FH1-4 in vitro. Thus it can be concluded that this complex is likely to present the physiological target recognition complex between FH and C3b (Figure 18) that would explain the increased affinity of FH to C3b on self surfaces (Carreno et al. 1989; Meri et al. 1990b). Affinity of FH19-20 to C3b via the FH19 site probably remains the same in self and non-self surfaces if hypothetical conformational changes in C3b on different surfaces are excluded. Thus, the determining factor in self- non- self discrimination by FH is the affinity of FH20 to anionic structures on self surfaces.

5.2.2 The role of C3dg as marker of self In study IV, structural analyses showed that FH19-20 could bind to C3b via the FH19 site and to an additional C3d via the FH20 site (Figure 18) that has interesting implications for complement regulation on surfaces by FH. C3b is converted to C3dg on self surfaces that have membrane bound CR1 catalyzing the cleavage from iC3b to C3dg, and on those surfaces which are in close contact with erythrocytes that have abundantly CR1 on the surface. The surface bound C3dg could compensate for the loss or unavailability of polyanion binding sites or could even create additional binding sites for FH. Thus C3d on the self surface may enhance binding of FH to self surfaces and lead to complement inactivation through amplified down-regulation of AP attack. Interestingly, DNA has also been shown to bind FH19-20 (Leffler et al. 2010) that indicates that GAGs, sialic acids, C3d, and other anionic cell surface molecules could utilize the same binding site on FH20 in downregulation of complement activation.

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5.3 Molecular pathogenesis of aHUS

The aHUS mutations on FH19-20 caused various combinations of dysfunction. Mutations were shown to cause loss of bond(s) in the FH19 site and FH20 site, to disrupt heparin or cell binding sites, or to inhibit the FH19 site and FH20 site binding to C3d by enhancing binding to heparin. The mutations that were not located in the binding interfaces probably lead to destabilization of the fold and thereby cause disruption of FH19-20 function. Despite all the different effects of the aHUS mutations on FH19-20 structure they disrupt the same function of FH - self surface recognition - by interfering with simultaneous binding of the FH19 site to C3b and the FH20 site to polyanions or cells.

The decreased binding of the FH20 site to C3d may also play a role in the pathogenesis of aHUS if the binding of FH to the FH20 site on C3d will be shown to amplify downregulation of complement on self cells as was suggested in study IV. This model is supported by two aHUS patient mutations, R49L and A101V in the FH20 site on C3d (Frémeaux-Bacchi et al. 2008; Maga et al. 2010).

Ferreira and co-workers found that aHUS associated mutations W1183R, T1184R, L1189F/R, S1191L, and S1191L/V1197A enhanced binding of FH19-20 to C3b (Ferreira et al. 2009). These results are contradictory to the results presented in this thesis. They studied the interaction by using only SPR binding assays that may be sensitive to charge changes on proteins due to the high negative charge of carboxymethylated dextran matrix. FH19-20 may have some affinity on the matrix since it recognizes highly negative dextran sulfate.

Taken together, the aHUS-associated mutations disrupt FH19 binding to C3b and FH20 binding to anionic self surface structures by different mechanisms. The consequence of the aHUS mutations on FH19-20 function is however the same - decreased avidity of FH-C3b interaction on the self surfaces that is the basic pathogenic mechanism of aHUS.

5.4 Dysfunction of FH19-20 in aHUS

C3b that is deposited on self or non-self surfaces does not seem to discriminate between them (Meri et al. 1990b) and thus it is probable that FH19-20 binding affinity to the exposed FH19 site on C3b is the same on all surfaces. Thus, the binding affinity of FH20 to self surfaces is crucial for inhibiting complement activation. This means that FH20 needs to recognize all the different types of cell surfaces in contact with plasma like erythrocytes, platelets and glomerular endothelial cells that are damaged in aHUS.

5.4.1 Complement activation on CS-A rich platelets causes thrombocytopenia It has been noted that thrombi in glomerular capillaries cannot alone explain thrombocytopenia in aHUS, indicating that platelets are also activated also in extra-renal circulation (Walters et al. 1988). Chondroitin sulfate A is the main GAG on platelets that seem to be devoid of HS (Ward et al. 1979; Okayama et al. 1986). It has been shown that FH binds to washed human platelets with FH15-20 (Vaziri-Sani et al. 2005) and that FH with misfolded FH20 has impaired binding to platelets (Ståhl et al. 2008). In addition, FH15-20 binding to platelets can be partially blocked with heparin (Vaziri-Sani

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et al. 2005) and FH15-20 binding to heparin can be inhibited with CS-A (Jokiranta et al. 2005). Since FH15-20 has only one heparin binding site it is likely that CS-A binding is mediated by the heparin site on FH20 that would then also be important for recognizing platelets. Ståhl et al. studied platelets from aHUS patients with the V1168E, E1198K, and E1198Stop mutations on FH20 and found that C3 and C9 were deposited on their surface (Ståhl et al. 2008). Platelets from aHUS patients form aggregates and agglutinate, indicating their activation in the extra-renal circulation (Ståhl et al. 2008; Licht et al. 2009). It can be concluded that dysfunctional FH19-20 cannot control complement deposition on the platelets, inducing adherence and aggregation, thus explaining the resultant thrombocytopenia.

5.4.2 Glomerular endothelial cell damage and thrombus formation In studies III-IV it was shown that the mGEnC and heparin binding sites were nearly identical on FH20. Although the mouse GEnC binding results cannot be directly applied to humans, it has been shown that (i) mouse GEnCs have features of human GEnC like fenestrae, also in vitro (Rops et al. 2004a), (ii) the HS moieties are well conserved in mammalian cells (Esko et al. 2002), and (iii) the C-terminal domains 18-20 of human and mouse FH bind similarily to heparin and HUVECs (Cheng et al. 2006). In GBM, HS accounts for up to 80 % of total GAGs (Kanwar et al. 1981; van den Heuvel et al. 1988) and is probably the main ligand for FH20. Glomerular endothelial cell glycocalyx has about 50 - 90 % HS and 10-20 % CS side chains on proteoglycans (Oohira et al. 1983) that are probably important for FH20 binding.

Defective binding of FH to glomerular endothelium leads to generation of C3a, C5a, and MAC that activate the endothelium leading to a pro-coagulant phenotype of the GEnCs (Ikeda et al. 1997; Tedesco et al. 1997). Activated endothelium expresses TF that initiates coagulation and release of vWf that can act as a receptor for platelets (Hattori et al. 1989; Saadi et al. 1995; Savage et al. 1996). Platelets are activated by released C3a and C5a and thereafter attach more easily to activated glomerular endothelium (Polley et al. 1983). Platelet adherence to the endothelial matrix leads to propagation of the coagulation cascade and to thrombus formation in the glomeruli. Thrombin and several other coagulation cascade enzymes cleave C3 and C5 that further activates complement on the platelets and on endothelium creating a vicious cycle (Figure 10) (Huber-Lang et al. 2006). The actions of complement and coagulation systems can therefore lead to the formation of thrombi and endothelial swelling in the glomeruli - the typical histological findings in aHUS patients.

5.4.3 Complement activation on sialic acid rich erythrocytes causes hemolysis Hemolysis in aHUS probably occurs mechanically in occluded capillaries as well as non-mechanically in the circulation (Söderholm et al. 2009). Erythrocytes surfaces are rich in sialic acids and FH16-20 has been shown to bind to sialic acids (Ram et al. 1998). Although direct interactions of the FH20 domain with sialic acids have not been proven, results of functional assays suggest that heparin and sialic acid sites overlap. Ferreira et al. studied the effect of the R1215G and K1230A mutations on FH19-20 binding to Esh-C3b (Ferreira et al. 2009). They found that these mutations do not impair C3b binding and decrease the affinity of FH19-20 to Esh-C3b. Based on the FH19-20:C3d structure, these surface exposed mutations are not in the C3d binding site, indicating that these mutations probably interact with sialic acids on erythrocytes. Although the presence of other receptors on

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erythrocytes cannot be excluded, it is known that FH binding is dependent on sialic acids on erythrocytes and it can be postulated that the FH20 site probably mediates this interaction. It has been shown that FH19-20 is needed to protect erythrocytes from complement mediated lysis (Ferreira et al. 2006) and that the aHUS mutations in FH19 or FH20 cause hemolysis of human erythrocytes (Ferreira et al. 2009). Also, FH from aHUS patients with theE1172Stop, W1183L, V1197A, E1198K, and R1210C mutations in FH20 areunable to protect non-activator sheep erythrocytes from lysis (Sánchez-Corral et al. 2004; Vaziri-Sani et al. 2006; Heinen et al. 2007). Taken together, it is likely that FH20 also recognizes sialic acids and that dysfunctional FH in aHUS cannot protect erythrocytes from complement attack.

5.4.4 Pathogenesis model of aHUS In conclusion, the aHUS-associated mutations in FH19-20 decrease the avidity of FH to C3b deposited on self surfaces, leading to decreased efficiency in inactivation of C3b. C3b deposition on GBM and GEnCs leads to complement amplification and generation of C5a and MAC. These synergistically damage and activate the endothelium known to lead to pro-coagulant and pro-inflammatory phenotypes. Platelets are activated in this manner and adhere to the glomerular vessel walls and form thrombi. Thrombi and swelling of the endothelium lead to occluded capillaries also contributing to hemolysis. In the systemic circulation complement attack against RBCs and platelets leads to hemolysis and platelet activation and aggregation. Complement activation induces inflammation, coagulation and cell damage that each feed-back to activate more complement and coagulation, explaining the acute attacks often seen in aHUS patients.

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6 Acknowledgments

This work was done almost completely in Haartman Institute, University of Helsinki, between 2001 and 2011. I want to thank the administration of the Medical Faculty and the Haartman Institute for providing me the facilities to do the research, and the Academy of Finland, Biomedicum Helsinki Foundation, Emil Aaltonen Foundation, Finnish Cultural Foundation, Maud Kuistila Memorial Foundation, and Sigrid Juselius Foundation for financial support.

I am thankful and honored that Anna Blom accepted the invitation to be my opponent. I want to express my gratitude to the reviewers of my thesis, Sarah Butcher and Olli Lassila, for their expert comments.

I am most indebted to my supervisor Sakari Jokiranta who has believed in me as a scientist from the beginning. His endless optimism, encouraging attitude, support and vast expertise have been the driving forces in the making of this thesis. Sakari has taught me how to do science – what more could a PhD student want?

I am very thankful for Seppo Meri who gave me the opportunity to enter the world of science. He has shared his deep understanding in immunology with me and helped on many matters along the way.

True friends are a rarity – I want to very warmly thank Taru Meri for her friendship and support in matters of science and life. She has been invaluable.

I am grateful for all the past and present members of the Jokiranta-group for keeping up the spirit and giving support in the daily work. I am very thankful for Hanne Amdahl, Arnab Bhattacharjee, Zhu-Zhu Cheng, Karita Haapasalo-Tuomainen, Satu Hyvärinen, Harriet Hägglund, Aino Koskinen, Tanja Pasanen, Elina Pusa, Hannah Söderholm, and Maria Pärepalo for their contributions and nice moments in the lab.

This work was in great part based on high quality structural biology work done in the Institute of Biotechnology. I want to thank Adrian Goldman, Tommi Kajander, and Veli-Pekka Jaakola for very fruitful collaboration and sharing their expertise.

I am thankful for Johan van der Vlag for the opportunity to work in his group at Nijmegen, the Netherlands, and sharing his deep understanding on glycobiology and endothelial cells. I want to thank Angelique Rops for collaboration and her support during my three-months stay in the lab. I and my family were very warmly welcome to the Netherlands. I am (and we are) indebted to Johan and his family, Angelique, Marinka Bakker and her family, Casandra van Bavel, Jürgen Dieker, Martijn Kramer, Tom Nijenhuis, and Wim Tamboer.

I am honored and grateful that I had a chance to collaborate with David Isenman from the University of Toronto who gave excellent advice on studies III and IV and provided together with Elisa Leung the C3dg mutant proteins.

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Mathematician Simopekka Vänskä is acknowledged for his expert advice on interdomain angle calculations and Derek Ho for checking my thesis for the correctness of English language.

I want to sincerely thank the “more experienced” generation of the Meri-group Hanna Jarva, Sami Junnikkala, Antti Lavikainen, Antti Sommarhem, and Jorma Tissari for all the help, collaborations, scientific discussions and nice moments. I also wish to thank Antti Lavikainensis for our productive collaboration that has certainly helped me to fund this thesis project. I have shared many memorable scientific and non-scientific moments with, and I am thankful for the help I got from many members of the Meri-group: Antti Alitalo, Tobias Freitag, Nathalie Friberg, Juha Hakulinen, Mikko Holmberg, Ayman Khattab, Vesa Kirjavainen, Vesa Koistinen, Nina Lindeman, Sami Nikoskelainen, Jaana Panelius, Riina Richardson, Rauna Riva, Jari Suvilehto, Jasse Tiainen, Antti Väkevä, and Jinghui Yang.

The early “koppi” years were hilarious and most informative, and I want to thank Laura Kerosuo-Palmberg for her friendship and sharing the booth with me and Taru. I wish to thank Jussi Hepojoki and Tomas Strandin for helpful advice and great moments in the depths of the peptide lab.

I want to thank Livija Deban for our collaboration and nice discussions. I am honored that I have had a chance to know many foreign colleagues like Kyra Gelderman, David Gordon, Florian Eberle, Jens Hellwage, Mihaly Jozsi, Rebecca Ormsby, Tanya Sadlon, Leendert Trouw, and Valentina la Verde with whom I have shared good moments and scientific discussions.

I want to very warmly thank Tiina Kasanen for her support, friendship, and all the cheerful coffee breaks with Laura Kalin, Laura Mattinen, and Jukka-Pekka Palomäki. In addition I want to acknowledge Marta Biedzka-Sarek and Mikael Skurnik for their positive spirit.

I have shared an office with many members of the Arstila-group and I wish to thank Eliisa Kekäläinen, Tuisku-Tuulia Laurinolli, Anni Lehtoviita, Pirkka Pekkarinen, Laura Rossi, and Heli Tuovinen for their positive spirit and T-cell help.

I want to warmly thank Kirsi Udueze for her support, cheerful discussions, and all the help with so many practical matters. I would like to thank Marjatta Ahonen, Marko Hietavuo, Pirkko Kokkonen, Ilkka Vanhatalo and Kirsti Widing for good moments, practical and technical support, and keeping the lab, computers, and equipment running.

Very nice and professional people have worked with me in the department of bacteriology and immunology. I want to thank Petteri Arstila, Mia Eholuoto, Anu Kantele, Hilkka Lankinen, Sari Pakkanen, Nora Pöntynen, Heikki Repo, Heidi Sillanpää, Eine Virolainen, and Anni Virolainen-Julkunen for all the discussions and good moments. I wish to acknowledge Christophe Roos who gave me the inspiration to learn bioinformatics and helped me in the beginning of my career. My fellow engineers at Haartman institute Samu Myllymaa and Tuomas Raitila are thanked for their friendship.

Many wonderful persons and personalities have enriched my life in science and I have shared good moments and scientific discussions with them – I am grateful for also those that were not mentioned here.

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I want to acknowledge all my friends and relatives for their support, good moments, and bringing balance to my life.

I am deeply indebted and thankful beyond words to my wife Minna for all her unselfish support during these years. It has been quite an effort to stand by me, but you always did – I love you.

Helsinki, 2011

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