Investigation of the FAT10 Conjugation Pathway

Investigation of the FAT10 conjugation pathway

Dissertation

zur Erlangung des akademischen Grades eines Doktors der Naturwissenschaften (Dr. rer. nat.) an der Universität Konstanz (Fachbereich Biologie)

vorgelegt von

Stella Ryu

Tag der mündlichen Prüfung: 19.04.12 1. Referent: Prof. Dr. Marcus Groettrup, Universität Konstanz 2. Referent: Prof. Dr. Elke Deuerling, Universität Konstanz

Table of contents

1 Danksagung 6

2 Zusammenfassung / Summary 7

2.1 Deutsch 7

2.2 English 9

3 Introduction 12

3.1 The Ubiquitin conjugation system 12 3.1.1 E1 ubiquitin activating enzymes 15 3.1.2 The ubiquitin-like modifier activating enzyme 6 (UBA6) 16 3.1.3 E2 conjugating enzymes 17 3.1.4 E3 Ligases 17 3.1.4.1 HECT E3 ligases 18 3.1.4.2 RING finger E3 ligases 18 3.1.5 The tripartite motif (TRIM) protein family 21 3.1.6 TRIM11 22

3.2 Ubiquitin-like proteins (UBLs) 24 3.2.1 Ubiquitin like modifier (ULM): an overview 24 3.2.2 The ubiquitin like modifier FAT10 26 3.2.3 FAT10 conjugation pathway 31 3.2.4 The small ubiquitin-like modifier SUMO 32

3.3 Inhibitor of apoptosis protein (IAP) family 34 3.3.1 BRUCE represents a special BIR containing protein (BIRP) 36

3.4 Autophagy 39 3.4.1 Ambra1 (activating molecule in Becn1-regulated autophagy) 44

3.5 Transcription factors 46 3.5.1 The transcription factor AP-1 47 3.5.1.1 Structural and biochemical properties of AP-1 48 3.5.1.2 Transcriptional and post-transcriptional regulation of AP-1 50 3.5.2 The transcription factor JunB 50

Table of contents

4 Aim of this study 55

5 Materials and Methods 56

5.1 Materials 56 5.1.1 Chemicals and Materials 56 5.1.2 Reagents and reaction kits 57 5.1.3 Buffers and solutions 57 5.1.3.1 Media for cell culture 57 5.1.3.2 Stock solutions 58 5.1.3.3 Antibiotics and Inductors 58 5.1.3.4 Buffers and Solutions for Agarose Gel Electrophoresis (AGE) 58 5.1.3.5 Buffers and Solutions for SDS-Polyacrylamide Gel Electrophoresis 59 5.1.3.6 IP and lysis buffer 60 5.1.3.7 Washing and elution buffer 61 5.1.3.8 Cultivation Media and Media Additives for bacteria 62 5.1.3.9 Cultivation Media and Media Additives for yeast 63 5.1.4 Cell culture 67 5.1.4.1 Eukaryotic cell culture 67 5.1.4.2 Prokaryotic cell culture 67 5.1.4.3 Yeast 67 5.1.5 Antibodies 68 5.1.6 Enzymes and other cloning components 68 5.1.7 Recombinant proteins 69 5.1.8 Vectors used for cloning 69 5.1.9 Plasmid constructs 69 5.1.9.1 Primers 70

5.2 Methods 70 5.2.1 Plasmid DNA purification 70 5.2.2 Cloning 71 5.2.2.1 PCR (Polymerase chain reaction) 71 5.2.2.2 Site-directed mutagenesis 73 5.2.2.3 Agarose gel electrophoresis 73 5.2.2.4 Restriction digest 73 5.2.2.5 Ligation 74 5.2.2.6 Preparation of chemically competent E. coli cells 74 5.2.2.7 Transformation into E. coli TOP 10 F’ 75

Table of contents

5.2.3 Expression and purification of recombinant GST-TRIM11 from E. coli 75 5.2.4 Cell culture 75 5.2.5 Transient transfection 76 5.2.6 SDS-PAGE (Sodium dodecylsulphate polyacrylamide gel electrophoresis) 77 5.2.7 Immunoblotting 77 5.2.8 Immunoprecipitation (IP) 78 5.2.9 In vitro FAT10ylation assay of JunB 79 5.2.10 Growth of yeast strains 79 5.2.11 DNA isolation from yeast with glass beads 79 5.2.12 Yeast two-hybrid assay 80 5.2.13 Luciferase reporter assays 82 5.2.14 Immunofluorescence and confocal microscopy 83

6 Results 84

6.1 Yeast two-hybrid screen with UBA6 84 6.1.1 BRUCE interacts non-covalently with UBA6 and FAT10 87 6.1.2 Endogenous FAT10 co-immunoprecipitates with BRUCE 92

6.2 Yeast two-hybrid screen with TRIM11 94 6.2.1 TRIM11 interacts specifically with JunB and Ambra1 in a yeast two-hybrid screen 94 6.2.2 Interaction of JunB with TRIM11 in human cell culture 96 6.2.3 Ubiquitin and FAT10 become isopeptide linked to to JunB 98 6.2.4 Proteasome inhibition augments conjugate formation between JunB and FAT10 102 6.2.5 JunB has no influence on the degradation rate of FAT10 105 6.2.6 Co-expression of FAT10 hardly affects the degradation of unconjugated JunB 106 6.2.7 CHX data reveal a role for proteasome dependent degradation of the conjugate between JunB and FAT10 108 6.2.8 TRIM11 becomes degraded via the proteasome 110 6.2.9 TRIM11 turnover in presence of FAT10 is slightly accelerated 112 6.2.10 Co-expression of TRIM11 does not change protein turnover rates of JunB and FAT10 113 6.2.11 Conjugate formation of endogenous JunB and FAT10 115 6.2.12 Conjugate formation of JunB and FAT10 under semi-endogenous conditions 116

Table of contents

6.2.13 In vitro auto-FAT10ylation assay 117 6.2.14 JunB is conjugated with FAT10 on lysine 237 119 6.2.15 Identification of JunB in mass spectrometry analysis 122 6.2.16 Post-translational modification: Phosphorylation of JunB on Serine 259? 124 6.2.17 FAT10 and JunB co-localize at the nuclear membrane and in the cytosol 125 6.2.18 FAT10ylation controls JunB transcriptional activities on minimal AP-1 driven reporter genes 128 6.2.19 Interaction of Ambra1 and TRIM11 in a human cell line 131 6.2.20 Ambra1 interacts non-covalently with ubiquitin and FAT10 133 6.2.21 The proteasome is involved in Ambra1 degradation 136 6.2.22 Ambra1 co-localizes with FAT10 in punctuated structures 140

7 Discussion 146

8 References 175

9 Abbreviations 202

10 Addendum 206 Danksagung

1 Danksagung

Mein Dank gilt Herrn Prof. Dr. Marcus Groettrup für die Bereitstellung des Themas und für die wissenschaftliche und menschliche Unterstützung. Seine Begeisterung für die Wissenschaft ist unendlich und ansteckend.

Danken möchte ich....

Frau Dr. Annette Aichem für die Betreuung am Biotechnologie-Institut Thurgau (BITg) und für die Unterstützung während der gesamten Doktorarbeitszeit.

der Konstanzer Research School Chemical Biology (KoRS-CB) und besonders meinem Promotionskommitee Frau Prof. Elke Deuerling und Herrn Prof. Valentin Wittman und allen Mitgliedern der Graduiertenschule.

Herrn Dr. Daniel Legler für seine freundliche Aufnahme am BITg und dafür, dass er stets ein offenes Ohr für mich hatte.

Karin Schäuble für unzählige Gespräche über Wissenschaft, nächtelangen Laborsessions und vieles mehr, Verena „die Göttin“ Wörtmann und Nicola Catone, die neben der Laborarbeit zu guten Freunden geworden sind.

natürlich der gesamten „FAT10“-Gruppe für eine tolle Zusammenarbeit.

insbesondere Valentina Spinnenhirn, Kathrin Kluge, Andrea Kniepert, Annegret Bitzer und Khalid „the champ“ Wasim für eine unvergessliche Zeit in Konstanz und wissenschaftliche Hilfe. Ihr seid mir sehr ans Herz gewachsen.

Hesso Farhan und Veronica Reiterer für die Hilfe am Mikroskop, für viele tolle Gespräche und kontinuierlichen Schokoladennachschub.

den guten Seelen des BITgs Josepha, Ilona, Conni und besonders Edith Uetz für die Unterstützung bei den Reporterassays, für die „Schweizerdeutsch“-Nachhilfe und für viel Spaß bei der Laborarbeit.

Gunter Schmidtke und Michi Basler, für viele hilfreiche Tipps.

meinen Bürokollegen Margit, Valentina, Francesco und Bruxe.

all meinen Kollegen des BITg und der Immuno-Gruppe des Groettrup-Labors, die ich leider aufgrund von Platzgründen nicht alle persönlich benennen kann.

meiner Familie und meinen Freunden, die mich während meiner gesamten Doktorandenzeit unterstützt hatten. Ganz besonders Andoni, für seine Liebe und Geduld.

Zusammenfassung / Summary

2 Zusammenfassung / Summary

2.1 Deutsch

Posttranslationale Modifikationen können die Aktivität, Funktion, Stabilität oder die intrazelluläre Lokalisation von Proteinen verändern. Posttranslationale Konjugation von einen oder mehreren Ubiquitin oder Ubiquitin-ähnlichen Proteinen zu ausgewählten Substraten, Ubiquitinierung genannt, ist eines der wichtigsten und vielfältigsten regulatorischen Mechanismen in der Biologie und erfordert das fortlaufende Zusammenspiel einer 3-Schritt Enzymkaskade. Initiierender Schritt zur Konjugation von Ubiquitin oder Ubiquitin-ähnlichen Proteinen an seine Zielproteine ist die Ausbildung eines energiereichen Thioesters an seinem C-Terminus. Diese Aktivierung erfolgt durch ein Ubiquitin-aktivierendes E1 Enzym. Das aktivierte Ubiquitin wird in einer Transesterifizierungskaskade auf eines von mehreren Ubiquitin konjugierenden E2-Enzymen übertragen. Das Ubiquitin-beladene E2 und ein spezifisches Substratprotein werden dann von einer Ubiquitin Proteinligase (E3) gebunden, welche den Transfer des aktivierten Ubiquitins zwischen seiner carboxy-terminalen Hydroxylgruppe auf die ε-Amino-Seitenkette eines internen Lysinrestes des Akzeptorproteins katalysiert. Ubiquitin-ähnliche Proteine (UBLs) wie beispielsweise SUMO, NEDD8 und ISG15 werden durch eine vergleichbare E1-E2-E3 Multienzym-Kaskade an ihre Zielproteine ligiert. Für das Ubiquitin-ähnliche Protein FAT10 ist der Konjugierungsmechanismus noch nicht vollständig erforscht.

Das IFN-γ und TNF-α induzierbare Protein FAT10 ist ein junges Mitglied der Ubiquitin- ähnlichen Proteine, welches über sein C-terminales Di-Glycin-Motiv kovalent an Zielproteine binden kann. Zudem ist es bisher das einzig identifizierte Ubiquitin-ähnliche Protein, welches Substratproteine Ubiquitin-unabhängig für den proteasomalen Abbau markieren kann. Erst vor Kurzem wurde ein FAT10 aktivierendes E1-Enzym, UBA6 und ein FAT10 konjugierendes E2 Enzym, USE1, identifiziert, welches interessanterweise zugleich das erste bekannte Substrat für FAT10 darstellt, da es in cis autoFAT10yliert wird.

Das Ziel dieser Doktorarbeit war die Charakterisierung des Konjugierungswegs von FAT10, beginnend mit der Identifizierung von UBA6 interagierenden Proteinen in einem „Yeast two- hybrid Screen“. Da zu Beginn der Doktorarbeit noch kein FAT10-spezifisches E2 Enzym bekannt war, lag der Fokus hier in der Identifikation von möglichen FAT10 E2 Enzymen.

Zusammenfassung / Summary

In einem „Yeast two Hybrid Screen“ konnte eine direkte Interaktion von UBA6 und dem C- terminalen Ende von BRUCE gezeigt werden, welches eine vollständige hochkonservierte Ubiquitin-konjugierende Domäne enthält. Diese Interaktion wurde in dieser Arbeit mittels Co- Immunopräzipitations-Experimente mit einem Expressionskonstrukt, welches für das vollständige BRUCE-Protein kodiert, besonders hinsichtlich einer möglichen FAT10 E2 Funktion weiter charakterisiert.

Ein weiteres Ziel der Doktorarbeit lag in der Identifikation von möglichen FAT10 Substraten. In einem „Yeast Two-Hybrid Screen“ mit einer möglichen FAT10 E3 Ligase TRIM11 als „bait“-Protein und einer cDNA-Bank aus humanem Thymus, konnte eine spezifische Interaktion zwischen TRIM11 sowohl mit JunB, als auch mit Ambra1 nachgewiesen werden. Die Ring-Finger E3 Ligase TRIM11 wurde zuvor in einem „Yeast Two-Hybrid Screen“ mit FAT10 und einer cDNA-Bank aus humanem Thymus identifiziert. Sowohl für JunB als auch für Ambra1 konnte eine spezifische Interaktion in HEK293 Zellen mittels Co- Immunopräzipitations-Experimenten mit TRIM11 ermittelt werden. Zudem konnte gezeigt werden, dass JunB kovalent mit Ubiquitin oder FAT10 modifiziert wird, nicht jedoch in Anwesenheit einer FAT10 Mutante, der das C-terminale Di-Glycin Motiv fehlt, welches für die Isopeptidbindung an Substratproteine verantwortlich ist. Dieses Ergebnis weist auf die Bildung eines Konjugats zwischen JunB mit beiden Ubquitin-ähnlichen Proteinen hin, was vermuten lässt, dass JunB sowohl ein Ubiquitin, als auch ein FAT10 spezifisches Substrat darstellt. Zudem konnte auch eine nicht kovalente Bindung von JunB sowohl mit Ubiquitin als auch FAT10 detektiert werden. Ferner konnte mittels „Cycloheximid-Chase“ Experimenten verdeutlicht werden, dass nicht nur die Isopeptid-gebundene Form von JunB und FAT10 für den proteasomale Abbau markiert wird, sondern auch JunB, welches nicht kovalent mit FAT10 modifiziert wurde. Proteasomale Inhibition mittels der Zugabe von MG132 führte zu einer Akkumulation des JunB-FAT10 Konjugats, welches im Falle eines FAT10 Substrats erwartet werden würde. Die Rolle von TRIM11 als FAT10 spezifische E3 Ligase mit JunB als Substrat konnte nicht eindeutig geklärt werden, da JunB in vitro in Anwesenheit von rekombinantem FAT10, UBA6 (E1), USE1 (E2) und TRIM11 (mögliche E3-Ligase) nicht FAT10yliert wurde. Überexpression von TRIM11 führte zu einer Herunterregulierung von Ubiquitin, FAT10 und JunB, als auch den JunB-Ubiquitin oder JunB-FAT10 Konjugaten auf Proteinebene. Mittels Konfokalmikroskopie konnte eine klare Co-Lokalisation von FAT10 und JunB an der nukleären Membran detektiert werden. Die Zugabe von MG132 führte zu einer Translokation von JunB in das Cytoplasma. Als vermutlich funktionale Konsequenz hatte JunB FAT10ylierung in Reporterassays eine deutlich verminderte JunB Transaktivierungs- Leistung zur Folge.

Zusammenfassung / Summary

Im Falle von Ambra1 konnte eine nicht-kovalente Bindung sowohl mit Ubiquitin, als auch mit FAT10 mittels Co-Immunopräzipitationsversuchen nachgewiesen werden. MG132 Zugabe führte zu einer Akkumulation von überexprimiertem Ambra1, welches darauf hinweist, dass Ambra1 proteasomal degradiert wird. Akkumulation nach proteasomaler Inhibition konnte allerdings nicht beobachtet werden, wenn Ambra1 und FAT10 co-exprimiert wurden, was darauf schließen lässt, dass die nicht-kovalente Interaktion mit FAT10 vermutlich zu einem anderen oder zusätzlichen Degradationsmechanismus führt. Experimente mit Konfokalmikroskopie verdeutlichen eine eindeutige Co-Lokalisation von FAT10 und Ambra1 in aggresomalen Strukturen, was vermuten lässt, dass die Interaktion von Ambra1 und FAT10 zu einer Translokalisation beider Proteine hin zu solchen Strukturen führt.

2.2 English

Posttranslational modifications are important means to alter a proteins’ activity, function, stability or its intracellular localization. The post-translational conjugation of one or more molecules of Ub and ubiquitin-like proteins (UBLs) to selected substrates, namely ubiquitination, is one of the most important and multifaceted regulatory mechanisms in biology and requires the sequential interaction of a 3-step enzyme cascade. The initiating step for ubiquitin or UBL conjugation to its target proteins is the formation of an energy-rich thioester at its C-terminus. This activation takes places through an ubiquitin activating E1 enzyme. The activated ubiquitin is then transferred in a trans-thioesterification cascade to one of multiple E2 conjugating enzymes. The ubiquitin-charged E2 enzyme and a specific substrate protein are then both bound by a ubiquitin protein ligase (E3), which catalyzes the transfer of the activated ubiquitin between its carboxy-terminal hydroxyl-group onto the ε- amino-side chain of an internal lysine residue of the acceptor protein. Canonical ubiquitin-like proteins (UBLs) such as ubiquitin, SUMO, NEDD8, and ISG15 are transferred by a similar E1-E2-E3 multi-enzyme cascade to its targets. For the ubiquitin-like modifier FAT10, the enzyme cascade has not yet been characterized completely.

The IFN-γ and TNF-α inducible modifier FAT10 is a young member of ubiquitin-like proteins, which can be conjugated to target proteins via its C-terminal diglycine motif. Moreover, it is to date the only identified ubiquitin-like protein, which can assign substrate proteins, in an ubiquitin-independent manner, for proteasomal degradation. Recently, the FAT10 activating enzyme (E1) UBA6 and a FAT10 conjugating enzyme (E2), namely USE1 was identified, which interestingly, was at the same time the first known substrate for FAT10, as it was auto-FAT10ylated in cis.

Zusammenfassung / Summary

The aim of this thesis was the characterization of the FAT10 conjugation pathway, starting with the identification of UBA6 interacting proteins in a yeast two-hybrid screen. Given that at the beginning of the doctoral thesis no FAT10 conjugating E2 enzymes were known so far, the focus here was the identification of potential FAT10 E2 enzymes. In a yeast two-hybrid approach, a direct interaction of UBA6 and the C-terminal end of BRUCE, containing the entire highly conserved ubiquitin conjugating domain, could be shown. This interaction was further characterized with a construct encoding for full length BRUCE via co- immunoprecipitation experiments, especially in terms of exhibiting a putative FAT10 E2 function.

A further aim of this doctoral thesis was the identification of putative FAT10 substrates. In a yeast two-hybrid screen with the putative FAT10 E3 ligase TRIM11 and a cDNA-library from human thymus, a specific interaction between TRIM11 and JunB as well as with Ambra1 could be observed. The RING finger containing E3 ligase TRIM11 was previously identified in a yeast two-hybrid screen with FAT10 and a cDNA-library from human thymus. For both JunB and Ambra1, a specific interaction with TRIM11 could be verified in co- immunoprecipitation assays. Moreover, it could be demonstrated that JunB becomes covalently linked to either ubiquitin or FAT10, but not in the presence of a FAT10 mutant, lacking the di-glycine motif, which is required for isopeptide linkages to substrate proteins. This result points to a conjugate formation between JunB with the ubiquitin-like modifier ubiquitin and FAT10, which indicates, that JunB is a ubiquitin as well as FAT10 specific substrate. Moreover, a non-covalent linkage of JunB with either ubiquitin or FAT10 could be detected. Furthermore, cycloheximide experiments revealed evidence that not only the isopeptide linked form of JunB and FAT10 became assigned for proteasomal degradation but also JunB, which was non-covalently modified with FAT10. Proteasome inhibition with MG132 led to an accumulation of the JunB-FAT10 conjugate, which would be expected for a FAT10 substrate. The role of TRIM11 as a FAT10 specific E3 ligase could not be solved definitely, due to the fact that JunB, in presence of recombinant FAT10, UBA6 (E1), USE1 (E2) and TRIM11 (putative FAT10 E3) did not become FAT10ylated in vitro. TRIM11 overexpression resulted in a decreased protein level of ubiquitin, FAT10, JunB and also the JunB-ubiquitin and JunB-FAT10 conjugates. A clear co-localization of FAT10 and JunB at the nuclear membrane could be detected by means of confocal microscopy. MG132 treatment caused the translocation of JunB into the cytosol. As a presumable functional consequence, JunB FAT10ylation led to a reduced JunB trans-activating capacity in reporter assays.

Zusammenfassung / Summary

In case of Ambra1, a non-covalent interaction with ubiquitin and FAT10 could be verified in co-immunoprecipitation experiments. Addition of MG132 led to an accumulation of overexpressed Ambra1, indicating that Ambra1 becomes degraded by the proteasome. However, no accumulation after proteasome inhibition was observable, when Ambra1 was ectopically co-expressed with FAT10, suggesting that the non-covalent interaction with FAT10 led to a different degradation mechanism other than the proteasome. Experiments with confocal microscopy illustrate an unambiguous co-localization of FAT10 and Ambra1 in punctuated structures, suggesting that the interaction of Ambra1 and FAT10 led to translocation of both proteins into punctuated structures.

Introduction

3 Introduction

3.1 The Ubiquitin conjugation system

Ubiquitin (Ub) is a highly conserved protein of 76 amino acids (aa), encoded on multiple genes, which was originally isolated by Goldstein and co-workers in the search for hormones derived from the thymus (Goldstein et al., 1975). It is a heat-stable protein found throughout the cells of eukaryotes that folds into a compact globular structure, a so-called ‘β-grasp fold’, wherein five β -sheets pack around a central α-helix. Although prokaryotes do not possess an Ub homolog, several prokaryotic proteins adopt a β- grasp fold, including MoaD and ThiS (Iyer et al., 2006). There are several mechanistic parallels between the ATP-dependent activation of Ub and MoaD/ThiS, although they differ broadly in function (Iyer et al., 2006; Miranda et al., 2011). Ub is either found solely distributed in the cell body or mostly covalently attached to substrate proteins. Modifications of substrate proteins with Ub alter their functions, locations or target them for destruction by the 26S proteasome (Kirkin and Dikic, 2007). The post-translational conjugation of one or more molecules of Ub and ubiquitin-like proteins (UBLs) to selected substrates, namely ubiquitination, is one of the most important and multifaceted regulatory mechanisms in biology (Hershko and Ciechanover, 1998; Jentsch, 1992; Pickart, 2004). It plays an integral role in a wide variety of functions in eukaryotic cells including signal transduction, transcription, heterochromatin formation, genome stability, protein trafficking, cell division, morphogenesis, DNA repair, endocytosis, apoptosis, autophagy and proteasome-mediated proteolysis (Hershko and Ciechanover, 1998; Hochstrasser, 2009; Jentsch, 1992; Pickart, 2004). Proteasomal degradation was not only the first consequence of ubiquitination to be identified, but is still recognized today as the most prevalent function of ubiquitin modification. The ultimate mechanisms that cells use to ensure the quality of intracellular proteins are on one hand the selective destruction of misfolded or damaged polypeptides to prevent the accumulation of non-functional, potentially toxic proteins (Goldberg, 2003). On the other hand, there is the participation in regulatory mechanisms by selectively destroying key molecules, like transcription factors (Hammond- Martel et al., 2011) or cell cycle regulators by activating cell cycle dependent kinases (Cdks), for instance (Glotzer et al., 1991; Reed, 2003). Another crucial function is the processing of antigens for later presentation on MHCI (Rock and Goldberg, 1999). Besides, several studies provided evidences that Ub conjugation is involved in the recognition and elimination of intracellular bacteria (Fujita and Yoshimori, 2011; Steele- Mortimer, 2011).

Introduction

Prior to conjugation, Ub is expressed as a pro-protein with a C-terminal extension and needs to be proteolytically processed by UBL specific proteases (ULPs) or deubiquitinating enzymes (DUBs), to expose the C-terminal di-glycine motif before yielding functional, monomeric Ub (Jentsch, 1992). Ubiquitination usually results in the formation of an isopeptide bond between the C-terminus of Ub (G76) and the ε-amino group of a substrate lysine residue. Substrate proteins can be modified by Ub in different ways. Conjugation of a single Ub to a single lysine residue (mono- ubiquitination) is a regulatory modification involved in diverse processes including endocytosis, endosomal sorting, histone regulation, transcription, virus budding and nuclear export (Haglund and Dikic, 2005; Mukhopadhyay and Riezman, 2007). Multiple mono- ubiquitination occurs, when several lysine residues of a substrate are modified by single Ub molecules. Here, a role in receptor internalization and endocytosis has been described (Mosesson et al., 2003). The ability of Ub to form isopeptide linked polymers is crucial for the versatility of the Ub system. Ub itself contains seven internal lysine residues (K6, K11, K27, K29, K33, K48, and K63) which can be potentially used as acceptors for the attachment of other Ub molecules, allowing the formation of different types of Ub chains (poly-ubiquitination). In addition, linear poly-Ub can be linked by amide bonds formed between the C-terminal glycine residue of Ub and the N-terminal methionine residue of a following Ub (Kirisako et al., 2006). Substrates with four or more lysine 48 (K48) ~ glycine 76 (G76) linked Ub moieties are usually targeted to the 26S proteasome for degradation, whereas Ub is not degraded along with the substrate but removed and subsequently recycled (Hanna and Finley, 2007; Hershko and Ciechanover, 1998). However, recent publications reported that other than K48 linked ubiquitin chains are also accepted by the proteasome (Jacobson et al., 2009; Xu et al., 2009). Chains linked through lysine 63 have a role in endocytosis and NF-kB signaling (Deng et al., 2000; Galan and Haguenauer-Tsapis, 1997; Ikeda and Dikic, 2008). In contrast, K11-linked chains are involved in endoplasmic reticulum associated degradation (ERAD) and mitosis whereas K29-linked chains may participate in Ub fusion degradation (Johnson et al., 1995). Ubiquitination can in some cases also occur on substrate serine, threonine or cysteine residues (Vosper et al., 2009; Wang et al., 2007). Moreover, beside modification of internal amino acid residues, Ub conjugation to the N- terminal residue of substrates has been described for several proteins (Breitschopf et al., 1998; Ciechanover and Ben-Saadon, 2004). The impact of Ub conjugation on its substrates is tightly controlled by a variety of specific enzymes which in turn designate the fate of ubiquitinated proteins.

Introduction

Elaborate ATP-dependent conjugation systems covalently and reversibly attach Ub and UBLs to their target proteins (overview in Figure 1). This process of conjugation and deconjugation is carried out by a stringent enzymatic cascade consisting of a three-step mechanism, whereby in general, the different UBLs have their own discrete E1–E2–E3 cascades and have distinct effects on their targets (Hershko, 1983; Schulman and Harper, 2009).

Figure 1: General conjugation pathway of ubiquitin and ubiquitin-like proteins (UBLs) Ubiquitin and some ubiquitin-like proteins (UBLs) are processed by either deubiquitinating enzymes (DUBs) or UBL-specific proteases (ULPs) to expose a C-terminal glycine. Conjugation to substrates is an ATP dependent process, facilitated with the help of an enzyme cascade composed of at least three different enzymes. The ubiquitin activating enzyme E1 binds first ATP and then the Ub/UBL which leads to the formation of an Ub- adenylate that serves as a donor of Ub to the E1 active site cysteine. A subsequent trans-thiolation reaction transfers the Ub/UBL to a conserved cysteine residue on the ubiquitin conjugating E2 enzyme. In the final step of the cascade, the Ub/UBL is transferred from the E2 to an ε-amino group of lysine residues on protein substrates. This final step is usually mediated by E3 ligases that may function in one of two distinct ways: The HECT-like E3 ligases transfer the Ub/UBL from E2 to an internal cysteine through a further trans-thiolation step before transferring it to the target, whereas the RING (U-Box) and A20 finger-type E3 ligases seem to mediate a direct transfer of the UBL to the substrate within the target lysine residues. Figure taken from (Kerscher et al., 2006).

Ub is first linked to an ubiquitin activating enzyme (E1) and becomes activated in an ATP dependent manner (see 3.1.1). Thereupon, the activated Ub is transferred to the ubiquitin conjugating enzyme (E2), where Ub become trans-thiolated to its own conserved active site cysteine (see 3.1.3).

Introduction

From an E2 enzyme Ub is transferred to the ε-amino group of a lysine residue either within the target protein or the growing poly-Ub chain, thus forming an isopeptide bond. This transfer is often assisted by a ubiquitin E3 ligase (see 3.1.4). Ubiquitin E3 ligases recognize substrates for ubiquitination, and are considered to be crucial for determining ubiquitination specificity. Ubiquitin specific proteases known as deubiquitinating enzymes (DUBs) can remove covalently attached Ub from proteins, thereby controlling substrate activity and/or abundance (Ventii and Wilkinson, 2008). In some circumstances, an E4 enzyme can act as auxiliary factor to catalyze multi-ubiquitin chain assembly in collaboration with E1, E2 and E3 (Hoppe, 2005; Koegl et al., 1999). The process of dynamically modifying proteins with Ub and other UBLs creates reversible switches between different functional states of a substrate protein, allowing fine-tuned control of numerous cellular pathways.

3.1.1 E1 ubiquitin activating enzymes

Ubiquitin-activating enzyme E1 (UBA1, UBE1) is the archetype for a family of enzymes, which catalyze the ATP-coupled activation of Ub and other UBLs required for their subsequent conjugation to cellular targets. The general physical and structural features of the E1 family are well conserved. E1s can be classified on the basis of the domain structure. The so-called canonical E1s include UBE1, NEDD8-activating enzyme (NAE), SUMO- activating enzyme (SAE), UBA6 and UBA7 owing to their related domain structures and enzymatic mechanisms, and the non-canonical E1s include Atg7, UBA4 and UBA5 (Schulman and Harper, 2009). In the first step, ATP and UBL bind together to form a UBL–acyl adenylate intermediate, releasing inorganic pyrophosphate. The C-terminus of free UBL is adenylated by an ubiquitin activating (E1) enzyme, leaving the Ub–AMP adduct bound to the enzyme. The UBL–AMP then reacts with the E1 active-site thiol to form an E1~UBL thioester. Subsequently, a second ATP and UBL bind the enzyme as in the first step to form a ternary complex that contains two UBL molecules bound to the E1 (Haas and Rose, 1982; Haas et al., 1982). Charging of an E1 with Ub or a UBL triggers conformational changes in the E1, which exposes a negatively charged groove within a Ub fold to allow the formation of a proper E1~E2 complex (Lee and Schindelin, 2008). This form of E1 is competent for transthiolation of the thioester-bound UBL to a pathway-specific E2 and is required for the downstream function of UBL conjugation (Figure 2).

Introduction

Figure 2: Enzymatic mechanism of the ubiquitin activation and conjugation cycle. Ub(A) represents ubiquitin that is associated non-covalently at the adenylation active site, and Ub(T) represents ubiquitin that is covalently linked to the catalytic Cys of an E1 enzyme through a thioester bond. Step 1 shows adenylate formation, step 2 shows thioester formation, step 3 shows double ubiquitin loading of E1 and step 4 shows ubiquitin transfer to E2. Step 2 is repeated on the E1 Ub(A)~adenylate generated in step 4 to continue the cycle. Figure taken from (Schulman and Harper, 2009).

The initial activation of Ub was for decades believed for decades to be accomplished solely by a single enzyme designated ubiquitin activating enzyme 1 (UBE1) (Ciechanover et al., 1981; Haas et al., 1982). The surprising discovery that Ub can be stimulated by two different essential E1 enzymes, namely ubiquitin-activating enzyme E1 (UBE1) and ubiquitin-like modifier activating enzyme 6 (UBA6) (Chiu et al., 2007; Pelzer et al., 2007) and that UBA6 can activate two different UBLs (Ub and FAT10) (Chiu et al., 2007) illustrates, that a unilateral assignment of an E1 enzyme to a select UBL is no longer valid, and raises the question how these modifiers compete for activation.

3.1.2 The ubiquitin-like modifier activating enzyme 6 (UBA6)

The activating enzyme UBA6 was 2007 identified as novel E1 enzyme, which can be charged by ubiquitin as well as FAT10 (Chiu et al., 2007; Pelzer et al., 2007). Human UBA6 and UBE1 have distinct preferences for E2 charging in vitro, and their specificity depends in part on their C-terminal ubiquitin-fold domains, which recruit E2s. UBE1 is phylogenetically more closely related to UBA7 (the E1 for ISG15) (Jin et al., 2007), whereas UBE1 and UBA6 show only about ~40 % sequence identity. UBA6 is uniquely responsible for transferring ubiquitin (Jin et al., 2007) as well as FAT10 to a UBA6-specific E2 enzyme USE1 (Aichem et al., 2010). UBA6 and USE1 are found from humans to zebrafish, as well as sea urchin, and are ubiquitously expressed, but they are absent from worms, flies, plants and yeast, which indicates a selective role in certain multicellular organisms.

Introduction

Deletion of the mouse uba6 gene results in embryonic lethality (Chiu et al., 2007) and can block the conjugation of FAT10 to unknown proteins (Aichem et al., 2010). However, mice that lack FAT10 are viable (Canaan et al., 2006), suggesting that the essential functions of UBA6 are not linked to FAT10 activation.

3.1.3 E2 conjugating enzymes

Ubiquitin-conjugating enzymes (E2s) are responsible for transferring UBLs to substrate proteins. Activated UBLs are subsequently transferred to an E2 enzyme, where UBLs become trans-thiolated to its own conserved active site cysteine. E2s often function with a single or limited number of E3 ligases, although in some cases no E3 is required. Thus, E2s function as key mediators of Ub chain assembly. These enzymes are able to govern the switch from Ub chain initiation to elongation, regulate the processivity of chain formation and establish the topology of assembled chains, thereby determining the consequences of ubiquitination for the modified proteins (Ye and Rape, 2009). Together, these factors determine the fate of ubiquitinated substrate proteins depending on whether they are mono or poly-ubiquitinated and on the site(s) to which Ub is conjugated. E2 family members possess a highly conserved core ubiquitin-conjugating (UBC) domain, consisting of approximately 150 amino acids (aa) containing the catalytic cysteine (Cys) residue which resides in a shallow groove (Wenzel et al., 2011). Ubiquitin E2 variant (UEV) proteins also have a UBC domain but lack an active site Cys residue (Hurley et al., 2006). After being charged with a UBL, E2s engage E3s to catalyse ubiquitin transfer to the ε-amino group of a lysine residue either within the target protein or the growing poly-Ub chain, thus forming an isopeptide bond, whereby a single E2 can interact with several different E3s.

3.1.4 E3 Ligases

E3-mediated attachment of Ub/UBL to substrates is highly regulated in response to cellular cues, and can modulate a target protein’s half-life, localization, interactions with protein or DNA partners and many other functions. Substrate selectivity of the Ub proteasome system relies primarily on the specificity of hundreds of E3 ubiquitin–protein ligases in the human genome, which mediate the transfer of activated Ub/UBL from an E2 enzyme to substrates (Varshavsky, 1997). Moreover, the E3 enzyme, in combination with the E2, is also important for determining the topology of the poly-Ub chain.

Introduction

Coupled to various cellular signaling events, ubiquitin E3 ligases ensure that the ubiquitination process is temporally controlled and tightly regulated with a high degree of substrate specificity which enables their function as key regulators in many cellular pathways (Ciechanover, 2003)

There are two major types of E3s in eukaryotes, defined by the presence of either a homologous to E6-AP C-terminus (HECT) (see 3.1.4.1) or a really interesting new gene (RING) domain (see 3.1.4.2), which are characterized by distinct Ub conjugation mechanisms (Fang and Weissman, 2004).

3.1.4.1 HECT E3 ligases HECT E3 ligases contain an approximately 350–amino acid long C-terminal region homologous to that of E6-associated protein (E6-AP), with a conserved active-site cysteine residue near the C-terminus, through which HECT domain E3 ligases form thioester intermediates with Ub before transferring it to the target protein (Huibregtse et al., 1995; Scheffner et al., 1995; Schwarz et al., 1998). N-terminal regions are highly variable and may be involved in substrate recognition (Hershko and Ciechanover, 1998). The HECT domain protein E6-AP is known for its role in binding the E6 protein of oncogenic human papilloma viruses and was first identified to be involved in the rapid degradation of p53 via the Ub dependent proteolytic pathway (Scheffner et al., 1994). In addition to ubiquitinate proteins for degradation by the 26S proteasome, HECT-E3 enzymes regulate the trafficking of many receptors, channels, transporters and viral proteins (Rotin and Kumar, 2009).

3.1.4.2 RING finger E3 ligases The really interesting new gene (RING) family, which includes the related U-box, B-box, leukocyte-association protein (LAP) domain and plant homeodomain (PHD) containing proteins, is conserved from yeast to humans. This family represents with over 600 members, the most abundant class of E3 ligases which mediates protein ubiquitination. Some of the ubiquitin ligases are single subunit ubiquitin ligases, such as Mdm2, containing the RING finger and the substrate-binding site in the same molecule (Marine and Lozano, 2010), while the majority of the RING E3s are multi-subunit complexes. Well investigated representatives are the cullin RING finger ubiquitin ligases (CRL) (Liu and Nussinov, 2009) and the high molecular mass anaphase promoting complex (APC) (Peters, 2002).

Introduction

The RING domain was originally described by Freemont and colleagues as a novel cysteine- rich sequence motif (Freemont et al., 1991). RING domain proteins compared to HECT E3s, do not have catalytic activity themselves but rather act as scaffolding proteins which facilitate the interaction between an E2 and target proteins (Joazeiro and Weissman, 2000). They bind the E2~Ub thioester together with the substrate, which brings them in close proximity to each other, often conveyed by conformational changes, and thereby permit the Ub transfer from the E2 directly to the target proteins (Deshaies and Joazeiro, 2009; Jentsch, 1992; Kerscher et al., 2006). In general, an ε-amino group of a Lys residue in the associated substrate attacks the thioester of the transiently associated charged E2, making an isopeptide bond with Ub or UBLs. The discharged E2 then dissociates from the E3, allowing a second charged E2 to interact with the E3 to facilitate a second round of Ub/UBL transfer, either by attack of a Lys residue in ubiquitin itself or by attack of a different Lys in the substrate. Multiple E2 cycles of E1-mediated Ub/UBL loading and subsequent unloading through a range of mechanisms lead to poly-ubiquitination of the substrate (Figure 3).

Figure 3: Reaction cycle of a RING E3. RING E3s are bisubstrate enzymes that catalyze the conversion of the reactants E2~Ub and substrate to the products E2 and substrate-Ub. Un-liganded E3 (a) binds substrate and E2~Ub to form the Michaelis complex (b). It is generally assumed that the two substrates do not need to bind in a predetermined order. (c) Ubiquitin is transferred from E2~Ub to substrate to yield the products, E2 and substrate-Ub. (d) For further ubiquitination to occur, E2 must dissociate to allow a fresh molecule of E2~Ub to bind (e). E2 cannot be recharged on E3 because E1 and E3 use overlapping surfaces to bind E2. The newly recruited E2~Ub transfers its cargo to yield di- ubiquitinated substrate (f). From this scheme, it is evident that the relative rates of substrate-Ub dissociation and E2~Ub recruitment/E2 dissociation can have a major impact on the number of ubiquitins that a substrate receives every time it binds to an E3. Figure taken from (Deshaies and Joazeiro, 2009).

Introduction

The ~70 aa residue RING finger domains bind two zinc ions in a unique "cross-brace" arrangement through a defined motif of eight highly conserved cysteine and histidine residues as depicted in Figure 4 (b) and (c). Unlike zinc fingers, the zinc coordination sites in a RING “finger” are interleaved and this arrangement endows the RING domain with a globular conformation, characterized by a central alpha-helix and loops of variable-length separated by several small beta-strands, suitable for protein-protein as well as protein-DNA interactions (see Figure 4 (a)) (Borden and Freemont, 1996). Structures of different RING E3s have shown, that the RING domains interact directly with E2s (Passmore and Barford, 2004).

Figure 4: The RING finger domain (a) A ribbon diagram, based on a model of the RING finger of CNOT4 bound to the E2 UBCH5B (blue; catalytic cysteine of E2 in yellow), the amino and carboxyl termini are indicated, respectively. (b) A schematic model of a RING finger domain. (c) RING-like sequence variants (coordinating residues are numbered in red, X indicates intervening amino acids followed by spacing in numbers). Figure taken from (Lipkowitz and Weissman, 2011).

Another structurally related domain which confers E3 ligase activity is the U-box domain, which does not contain any zinc coordinating residues but is still able to recruit E2 enzymes (Hatakeyama et al., 2001). In comparison, B-box domains of the TRIM subfamily of RING E3 ligases can bind one zinc atom, respectively, and adopt a similar structure reminiscent of the RING domain but are not capable to recruit E2s (Tao et al., 2008).

Introduction

3.1.5 The tripartite motif (TRIM) protein family

The largest family of higher order RING finger-containing proteins are the tripartite motif (TRIM) or RING/B-box/Coiled Coil (RBCC) proteins (Borden, 1998; Reddy et al., 1992). TRIM proteins are characterized by the presence of the tripartite motif consisting of a RING domain, one or two B-box motifs followed by a coiled-coil region, which frequently mediates hetero- or homodimerization and a variable C-terminus (Borden, 1998; Henry et al., 1998; Reddy et al., 1992; Rhodes et al., 2005). Notably, this arrangement is conserved throughout evolution, further supporting its functional relevance (Reymond et al., 2001). TRIM proteins are involved in a plethora of cellular processes such as development and cell growth, apoptosis, cell-cycle regulation and viral response. Consistently, their alteration results in many diverse pathological conditions (Meroni and Diez-Roux, 2005). To date, very little is known about the biological and molecular mechanisms mediated by the TRIM proteins. Over the past few years, several TRIM proteins have been reported to control gene expression through regulation of the transcriptional activity of numerous sequence-specific transcription factors. These proteins include the transcriptional Intermediary Factor 1 (TIF1) regulators, Trim19 or promyelocytic leukemia tumor suppressor (PML), the RET finger protein (RFP) and TRIM45, for instance (Kim et al., 1996; Le Douarin et al., 1995; Moosmann et al., 1996; Quignon et al., 1998; Zhong et al., 2000). The N-terminal region of PML harbours the typical tripartite motif which is essential for PML nuclear body formation in vivo (Jensen et al., 2001). Overexpression of TRIM45 inhibits the transcriptional activities of the transcription factors ElK-1 and AP-1, suggesting that TRIM45 acts as transcriptional repressor in MAPK signaling pathway. Among the plethora of cellular functions controlled by TRIM proteins, recent studies have demonstrated that TRIM proteins regulate signaling pathways which lead to type I interferon (IFN) induction in response to viral infection. IFNs, a group of secreted cytokines, are the main mediators of innate immunity against viral infection, by up-regulating the expression of many antiviral effectors within cells (Randall and Goodbourn, 2008). Moreover, expression of various TRIM genes is up-regulated in response to IFNs, suggesting the involvement of TRIM proteins in regulating host antiviral activities (Carthagena et al., 2009). Various TRIM proteins, for instance TRIM11, TRIM22, TRIM5α, TRIM19, TRIM25 or PML possess potential roles to trigger either autocrine or paracrine cell defense mechanisms and are possibly involved in innate immunity (Chelbi-Alix et al., 1998; Choi et al., 2006; Gack et al., 2007; Lin et al., 2004; Uchil et al., 2008). Furthermore, other TRIMs have been demonstrated to regulate signaling molecules downstream of pattern recognition receptors (PRRs) (McNab et al., 2011).

Introduction

Almost all TRIM family proteins possess a RING finger domain at their N-terminus. E3 ligase activity and function in ubiquitination processes mediated through the RING finger has been confirmed for some members of the TRIM family, including MID1/TRIM18, Efp/TRIM25, TRIM32 and TRIM22 (Duan et al., 2008; Frosk et al., 2002; Gack et al., 2007; Meroni and Diez-Roux, 2005; Trockenbacher et al., 2001). Very recently it has been shown, that some members of the TRIM superfamily possess SUMO E3 ligase activity, dependent on the TRIM motif, suggesting it to be the first widespread SUMO E3 motif (Chu and Yang, 2011). These TRIM proteins bind both the SUMO-conjugating enzyme Ubc9 and substrates and strongly enhance transfer of SUMOs from Ubc9 to these substrates like for instance tumor suppressor p53 and its principal antagonist Mdm2 or the transcription factor c-Jun. Furthermore, TRIM E3 activity may be an important contributor to SUMOylation specificity and the versatile functions of TRIM proteins (Chu and Yang, 2011). As a number of other RING finger proteins either homo- or heterodimerize, dimerization may facilitate optimal E3 ligase activity of TRIMs. There exist variants that retain all of the TRIM domains except the RING finger. In some cases these form heterodimers with RING finger- containing TRIMs via their coiled-coil domains. TRIMs without RING fingers may help to modulate substrate interactions, or serve as substrates themselves. Although the mechanisms how many TRIM proteins operate still remain to be deciphered, the highly conserved modular structure let suggest, that a common biochemical function may underlie their assorted cellular roles.

3.1.6 TRIM11

TRIM11 (52,8 kDa) is a member of the TRIM family proteins containing an N-terminal RING- finger which is a type of zinc finger motif, a B-box type 1, a B-box type 2, a coiled-coil region and a C-terminal PRY/SPRY domain, which constitute the B30.2 domain (Figure 5). Homooligomers can be formed through its coiled-coil domain (Reymond et al., 2001; Woo et al., 2006). This protein localizes to the nucleus and the cytoplasm (Ishikawa et al., 2006).

Figure 5: Schematic model of TRIM11. TRIM11 is composed of an N-terminal Ring finger domain, two B-Boxes, followed directly by a leucine coiled coil domain and a Pry and Spry domain which together constitute the B30.2 domain.

Introduction

TRIM11 is known to interact with humanin, an inhibitor of Alzheimer-like neuronal insults via its B30.2/SPRY domain and thereby to destabilize this protein. It is proposed to regulate intracellular levels of humanin by acting as an E3 ligase and thus inducing Ub mediated humanin degradation in neural tissues (Niikura et al., 2003). Moreover, humanin interacts with the apoptotic factor Bax and induces antiapoptotic effects by inhibiting the translocation of Bax to the mitochondria (Guo et al., 2003; Zapala et al., 2010). Thus, TRIM11 may upregulate apoptosis by controlling humanin stability. Another interaction partner for TRIM11 is the activator-recruited cofactor 105 kDa component (ARC105) which mediates chromatin-directed transcription activation and plays a crucial regulatory role for transforming growth factor β (TGF-β) signaling. Co-expression of TRIM11 increased ARC105 degradation which could be rescued through proteasome inhibition. Further, TRIM11 suppressed ARC105-mediated transcriptional activation induced by TGF-ß which suggests that TRIM11, together with the ubiquitin-proteasome pathway, regulates ARC105 function in TGF-ß signaling (Ishikawa et al., 2006). Besides, TRIM11 was able to interact in a yeast two hybrid screen with the homeodomain transcription factor Paired-like homeobox 2b (Phox2b), which is one of the key determinants involved in the development of noradrenergic (NA) neurons in both, the central nervous system (CNS) and the peripheral nervous system (PNS) (Hong et al., 2008). Furthermore, TRIM11 could interact with the transcription factor Paired box gene 6 (Pax6), a protein which is involved in the development and regulation of eyes and other sensory organs, brain, certain neural and epidermal tissues as well as other homologous structures. The interaction with TRIM11 mediates Pax6 degradation via the ubiquitin-proteasome system. However, abrogation of endogenous TRIM11 expression in the developing cortex increases the level of insoluble forms of Pax6 and enhances apoptosis. This indicates that an auto-regulatory feedback loop between TRIM11 and Pax6 maintains a balance between Pax6 and TRIM11 protein levels in cortical progenitors, indicating an essential role for the Pax6-dependent neurogenesis (Tuoc and Stoykova, 2008). Moreover, members of the TRIM family of E3 ligases have been shown to interfere with retroviral lifecycle and to exhibit antiviral activities (see 3.1.5). Overexpression of TRIM11 suppressed infectivity of human immunodeficiency virus 1 (HIV-1), by suppressing viral gene expression. This antiviral activity has been shown to depend on a functional TRIM11 E3 ubiquitin-protein ligase domain. Downregulation of TRIM11 enhanced virus release, suggesting that this protein contributes to the endogenous restriction of retroviruses in cells.

Introduction

Notably, while TRIM11 inhibited HIV entry, enhancing effects were observed for murine leukemia virus (MLV). Moreover, TRIM11 plays a role to regulate TRIM5 turnover, a protein which promotes innate immune signaling, via the proteasome pathway, thus counteracting the TRIM5-mediated cross-species restriction of retroviral infection at early stages of the retroviral life cycle (Uchil et al., 2008). In our laboratory a specific interaction of TRIM11 and the UBL FAT10 could be shown in a yeast two hybrid screen (A. Aichem, personal communication). Further, TRIM11 could interact in vivo with the FAT10 specific E2 enzyme USE1, and siRNA mediated TRIM11 downregulation decreased FAT10 conjugates, indicating that TRIM11 may function as a FAT10 specific RING finger E3 ligase.

3.2 Ubiquitin-like proteins (UBLs)

All eukaryotic cells contain several additional proteins related to ubiquitin, called ubiquitin-like proteins (UBL) with either sequence or structure similarities which can be further subdivided into two separate classes. The first group of UBLs are a heterogeneous group of ubiquitin domain proteins (UBDs), which bear a ubiquitin-like domain embedded in their sequences and thereby enable the binding of mono- or poly-ubiquitin chains in a non-covalent manner (Di Fiore et al., 2003; Jentsch and Pyrowolakis, 2000; Kerscher et al., 2006; Schnell and Hicke, 2003; Welchman et al., 2005). Their involvement has been described for various aspects of cellular physiology including protein degradation, receptor trafficking, DNA repair, autophagy and apoptosis (Ikeda et al., 2010). For instance, the first UBD to be published was the proteasome subunit S5A/RPN10 (Young et al., 1998). The second group presents the ubiquitin-like modifiers (ULMs). These proteins can be covalently attached to target proteins in a similar manner like Ub. To date, there are 17 known ULMs from nine phylogenetically distinct classes (NEDD8, SUMO, ISG15, FUB1, FAT10, Atg8, Atg12, Urm1, and UFM1) that have been identified to conjugate to substrates in a manner analogous to Ub.

3.2.1 Ubiquitin like modifier (ULM): an overview

Ubiquitin-like modifiers in turn function as their name already suggests in “ubiquitin-like” manner, what means, that they exert their function in being covalently attached to substrate molecules (see Table 1)

Introduction

Table 1: Ubiquitin like modifiers.

Modified and extended from (Hochstrasser, 2009). Identity with ULM Enzymes Substrates Comments and Functions ubiquitin [%]

E1: UBA3-Ula1 Cullins, p53, Positive regulator of ubiquitin E3s; heterodimer NEDD8 58 Mdm2, activation and destabilization of SCF; E2: Ubc12 synphilin-1 transcriptional regulation of p53 E3: many

E1: Aos-1-Uba1 c-Jun, IκB, p53, SUMO encoded by 3-4 genes in heterodimer Mdm2, STAT-1, vertebrates, depending on the species. SUMO-1 18 E2: Ubc9 PML, RanGAP1, Control of protein stability, function and E3: RanBP2, RanBP2, PCNA, localization, antagonist to ubiquitin, Pc2, PIAS topoisomeraseII overlap with SUMO-2/3

E1: Aos-1-Uba1 RanGAP1, Transcription regulation, cycle SUMO-2/3 16 heterodimer topoisomeraseII progression E2: Ubc9

E1: UBE1L PLCγ1, JAK1, Positive regulator of IFN-related immune ISG15 29, 27 E2:UbcH8 STAT1, ERK1/2, response, potentially involved in cell E3: Herc5, Efp serpin 2a growth and differentiation

Derived from a ribosomal protein FUB1 38 NI NI precursor

E1: UBA6 Cell cycle checkpoint for spindle p53, USE1, FAT10 29, 36 E2: USE1 assembly, ubiquitin- independent huntingtin E3: NI degradation

Three known isoforms in humans. E1: Atg7 Phosphatidyl- Contains a β-grasp fold. Atg8 10 E2: Atg3 ethanolamine Autophagy, cytoplasm-to-vacuole targeting

20 % identical to Atg8. E1: Atg7 Atg12 17 Atg5 Autophagy, cytoplasm-to-vacuole E2: Atg10 targeting

Related to the small-sulphur-carrying Ahp1 (in yeast) proteins MoaD and ThiS. Contains a β- 10 (Emboss MOCS3, grasp fold. Most ancient ULM. Role in Urm1 needle E1: UBA4 ATPBD3, CTU2, tRNA modification. Urm1 modification in alignment) CAS the response of cells to oxidative damage.

6 (Emboss E1: UBA5 Contains a β-grasp fold. UFM1 needle E2: UFC1 C20orf116 Important for the prevention of ER alignment) E3: UFL1 stress-induced apoptosis. NI, not identified.

Introduction

3.2.2 The ubiquitin like modifier FAT10

The fat10 gene was discovered in 1996 by chromosomal sequencing of the human major histocompatibility complex (MHC) class I locus (Fan et al., 1996) close to the HLA-F locus, leading to the designation of HLA-F adjacent transcript 10 (FAT10) (Liu et al., 1999). The region of human chromosome 6 encoding the MHC complex contains a diverse set of genes, including genes whose function can be directly related to immune function such as the MHC class I and II gene products, and genes encoding for members of the complement cascade, TNF-a and -ß, the transporter associated with antigen processing (TAP), and the LMP2 and 7 components of the proteasome (Pichon et al., 1996). FAT10, consisting of 165 amino acids is an ~18 kDa ULM which comprises two ubiquitin-like domains in a head to-tail formation, connected by a short linker peptide. These domains form the same three-dimensional core structure - the β-grasp fold – like Ub, revealing a common ancestry for the modification systems. Similar to Ub, it possesses a free C-terminal di-glycine motif required for covalent conjugation to USE1 (Aichem et al., 2010), p53 (Li et al., 2011) and huntingtin (Nagashima et al., 2011) and to several so far unknown target proteins (Chiu et al., 2007; Raasi et al., 2001). There exists a high degree of sequence similarity between murine and human FAT10 at mRNA and protein levels. Due to its analogy to a tandem fusion of two Ubs, it was originally called “ubiquitin D” or “diubiquitin”. The N-terminal and C-terminal ubiquitin-like domains of FAT10 are more closely related to Ub than to each other and show 29 % and 36 % sequence identity to Ub, respectively. Four of the lysine residues involved in poly-ubiquitin-chain formation – corresponding to K27, K33 and most notably K48 and K63 – are conserved in both ubiquitin- like domains of FAT10 (Figure 6). Atypical to Ub, FAT10 contains 4 cysteine residues in its sequence (Bates et al., 1997). Like Ub, FAT10 is activated by UBA6 (Chiu et al., 2007; Groettrup et al., 2008; Pelzer et al., 2007) and can be transferred to the E2 enzyme USE1 (Aichem et al., 2010). FAT10 is only expressed in vertebrates, i.e. it is evolutionary one of the youngest members of the ULM family. The importance of the regulation of FAT10 expression has been highlighted by different observations. Initially, up-regulation of FAT10 expression was shown to be restricted to mature dendritic cells (DCs) and B-cells (Bates et al. 1997). Unlike ubiquitin, which is expressed constitutively, constitutive FAT10 mRNA expression at tissue level seems to be limited to organs of the immune system. This was confirmed by Northern blot analysis, in situ hybridization as well as quantitative real-time PCR (qRT-PCR) in organs of the immune system like spleen, gut, lymphnodes and especially thymus (Liu et al., 1999; Lukasiak et al., 2008).

Introduction

Figure 6: Sequence comparison and ribbon diagram of Ubiquitin and the predicted tertiary FAT10 model structure (a) FAT10 is composed of two ubiquitin-like domains (UBLs) which are more closely related to ubiquitin than to each other. Both, the N- and C-terminal UBL show the typical β-grasp fold and displays 29 % and 36 % sequence identity to ubiquitin, respectively (Groettrup et al., 2008). (b) FAT10 encompasses two ubiquitin-like domains (UBLs), in which the C-terminal di-glycine motif is conserved in the second domain of FAT10. In addition, four of the lysines involved in poly-ubiquitin-chain formation – corresponding to K27, K33, K48 and K63 – are conserved. (c) Sequence alignment of the N- and C-terminal parts of FAT10 with ubiquitin (Ub). Conserved lysine residues are highlighted in yellow.

Moreover, an induction of FAT10 could be observed under particular conditions, such as inflammation, and has to be removed efficiently, when its inducing signals are turned off. However, FAT10 can be synergistically induced in many tissues by the proinflammatory cytokines IFN-γ and TNF-α (Liu et al., 1999; Raasi et al., 1999). Induction of the FAT10 mRNA was independent of protein neosynthesis but partially dependent on proteasome activity as treatment with proteasome inhibitors prevented induction of FAT10 with TNF-α, but not IFN-γ (Raasi et al., 1999). Although its function has not been fully elucidated, FAT10 has been implicated to play important roles in various cellular processes, for instance cancer, antigen presentation, cytokine response, apoptosis and mitosis. Studies in a murine fibroblast cell line revealed that induced expression of FAT10 resulted in massive caspase dependent cell death within 24 to 48 hours. Assumedly, the induction of apoptosis was dependent on the conjugation of FAT10 to so far unidentified target proteins (Raasi et al., 2001).

Introduction

Interestingly, fat10 is one of the most highly upregulated genes in HIV-infected renal tubular epithelial cells (RTECs). Down-regulation of FAT10 expression was shown to reduce apoptosis in RTECs infected by human immunodeficiency virus (HIV), suggesting a novel role for FAT10 in epithelial apoptosis (Ross et al., 2006). Moreover, FAT10 is a critical mediator of Viral Protein R (Vpr) induced apoptosis in human and murine RTECs, whereby the vpr gene plays an important role in FAT10 up-regulation. These proteins interact non-covalently and co-localize to mitochondria (Snyder et al., 2009). In seeming contradiction, up-regulation of FAT10 expression could be observed in several carcinomas, most notably in hepatocellular carcinoma (HCC) and in gastrointestinal and gynecological cancers (Lee et al., 2003; Lukasiak et al., 2008). In 2008, Oliva et al. identified FAT10 as a potential marker for liver preneoplasia, as it was highly overexpressed in a model of Mallory-Denk body containing chronic liver diseases, which are thought to progress to hepatocellular carcinoma (Oliva et al., 2008). Both suggested an active involvement of FAT10 in tumorigenesis based on its non covalent interaction with the spindle assembly checkpoint protein mitotic arrest deficiency 2 (MAD2), as previously shown in a yeast two hybrid assay (Liu et al., 1999). FAT10 is thought to displace MAD2 from the kinetochore during prometaphase, associated with incomplete chromosomal segregation and increased mitotic non-disjunction, resulting in a generation of cells that contain aberrant chromosome numbers, which is commonly observed in several cancers but no direct evidence was demonstrated (Ren et al., 2006). This finding strengthens the hypothesis that FAT10 plays a role in the regulation of genomic stability. FAT10 expression at transcript level undergoes cell cycle–specific changes, with the highest expression during the S phase and a low expression during G2/M phase (Lim et al., 2006). This finding further supported the hypothesis that FAT10 disturbs correct chromosomal segregation and is thus cell cycle regulated. Moreover, a presumable role for FAT10 in carcinogenesis has been proposed, as the presence of wild-type p53, which is known to play an important role in cell-cycle regulation, negatively regulates FAT10 mRNA expression and promoter activity and prevents reaching high FAT10 levels in the cell (Zhang et al., 2006), whereas mutant p53 provokes a contrary effect. Very recently Li et al. revealed evidence, that p53 becomes FAT10ylated and p53 transcriptional activity was found to be substantially enhanced in FAT10-overexpressing cells (Li et al., 2011). However, a further study identified that FAT10 possesses no transforming capability and increased FAT10 expression in several tumors is due to the up-regulation of proinflammatory cytokines (Lukasiak et al., 2008). So far, there is still a matter of debate whether it functions as a tumor suppressor or rather an oncogene.

Introduction

Hipp et al. reported in 2004, that degradation of FAT10 and its conjugates is accelerated in vitro and in vivo via its non-covalent interaction with the UBL-UBA domain protein NEDD8 ultimate buster 1-long (NUB1L), which binds to the proteasome through its UBL domain (Hipp et al., 2004; Schmidtke et al., 2006). The UBA domains of NUB1L are required for binding but not for accelerated degradation of FAT10 by the proteasome. This finding led to the assumption that NUB1L might not only act as a linker between the 26S proteasome and ULMs, but also as a facilitator of proteasomal degradation (Schmidtke et al., 2006). The degradation of FAT10 and its conjugates was initially described to be independent of Ub. Fusion of FAT10 to the N-termini of very long-lived proteins, like green fluorescent protein (GFP) for instance, enhanced their degradation rate as potently as fusion with Ub did. Therefore, it was suggested that FAT10 is the first ULM which provides a signal for proteasomal degradation of other proteins as an alternative route for Ub mediated protein degradation (Hipp et al., 2005). Further, FAT10 degradation occurred normally in E1 temperature-sensitive mutants, however it should be emphasized that Ub conjugation in this mutant is largely deficient at the restrictive temperature but a small share of poly-ubiquitin conjugate formation remained, which can lead to FAT10 ubiquitination (Hipp et al., 2005). Because no evidence for the deconjugation of FAT10 from its substrates has been obtained, it was believed that FAT10 is probably degraded, along with its substrates, in a manner similar to that seen with Ub-modified substrates when deconjugation is inhibited (Hanna and Finley, 2007). Contradictory, a very recent article assumed that FAT10 degradation by the proteasome requires its prior ubiquitination (Buchsbaum et al., 2011), based on the observation that a non-ubiquitinable lysine-less form of FAT10 is rapidly aggregated and precipitated in a insoluble fraction which is probably not sensible to the proteasome, whereas the WT protein appears to be less susceptible to aggregation. Moreover, FAT10 stabilization could be observed by using cells expressing non-polymerizable Ub and in cells harboring a thermo- labile mutation in the ubiquitin-activating enzyme, E1. The discrepancy to the previous article (Hipp et al., 2005) could origin in the different experimental setups to inactivate the E1 enzyme (Buchsbaum et al., 2011). Their own statement could be confuted by an experiment where they showed, that degradation of FAT10-GFP occurs in the presence of a nonpolymerizable mutant of ubiquitin which can be explained by the fact that the interaction of FAT10 with the proteasome is sufficient to promote, at least partially, the degradation of a downstream fused protein. This is in line with previous findings, showing that FAT10 can constitute a degradation signal without further ubiquitination (Hipp et al., 2005). .

Introduction

Along with this, some interaction partners of FAT10 have been determined to link FAT10 with aggregate formation. Kalveram et al. reported in 2008 that a cytoplasmic protein, histone deacetylase 6 (HDAC6), can interact non-covalently with FAT10 under proteasome inhibition, leading to the localization of FAT10 in aggresomes (Kalveram et al., 2008). This could provide an alternative route to ensure sequestration and subsequent removal of FAT10- conjugated proteins if FAT10 fails to subject its target proteins to proteasomal degradation. FAT10 also co-localizes with the catalytic immunoproteasome subunits LMP2 and LMP7 and its expression increases in liver cells forming Mallory-Denk bodies due to accumulation and aggregation of ubiquitinated cytokeratins (Bardag-Gorce et al., 2010). Many late-onset neurodegenerative diseases are associated with the formation of intracellular aggregates by misfolded or toxic proteins, revealing a high importance in the degradation pathways acting on such aggregate-prone cytosolic proteins including the ubiquitin-proteasome system and macroautophagy (Ross and Pickart, 2004; Williams et al., 2006). A recent report described that FAT10 molecules were covalently attached to the soluble fraction of the aggregate prone protein huntingtin and FAT10-modified huntingtin is prone to degradation by the proteasome. Moreover, completely aggregated huntingtin lacks any FAT10 and FAT10 knockdown enhanced aggregate formation. These data let suggest that FAT10 plays a role in stabilizing soluble huntingtin by facilitating the interaction with the proteasome (Nagashima et al., 2011). Several evidences for the involvement of FAT10 in the immune system are given. One hint for its immunological relevance is that the fat10 gene is encoded in the MHC locus, for instance (Fan et al., 1996). Moreover, FAT10 is constitutively expressed in immune cells and in organs of the immune system like spleen, gut, lymphnodes and especially the thymus (Lukasiak et al., 2008). Further, it is synergistically inducible in many tissues with the proinflammatory cytokines IFN-γ and TNF-α (Liu et al., 1999; Raasi et al., 1999). Interestingly, FAT10 can inhibit hepatitis B virus expression in a hepatoblastoma cell line after IFN-γ treatment (Xiong et al., 2003) and fat10 gene targeted mice demonstrated a high level of sensitivity toward lipopolysaccharide challenge and their lymphocytes are more susceptible to spontaneous apoptotic death (Canaan et al., 2006). These findings indicate that FAT10 may function as a survival factor, but the function and mechanism of its action in the immune system still remains poorly understood and still need to be investigated.

Introduction

3.2.3 FAT10 conjugation pathway

Besides Ub, several UBLs have been discovered; they all harbor structural homology to Ub, and many of them can also be conjugated to target proteins in order to modify their fate (Hochstrasser, 2009; Welchman et al., 2005). However, little is known about the regulation of UBLs, and in particular about the role of ubiquitination to control their levels. The initial activation of Ub was for decades believed to be accomplished solely by a single enzyme designated ubiquitin activating enzyme 1 (UBE1) (Ciechanover et al., 1981; Haas et al., 1982). Surprisingly, in 2007, a second E1 type enzyme, ubiquitin-like modifier activating enzyme 6 (UBA6) was identified which can activate Ub as well as the ULM FAT10 and depletion of UBA6 can block the conjugation of FAT10 to so far unknown proteins (Chiu et al., 2007; Pelzer et al., 2007). This finding raises the interesting question how this E1 enzyme can discriminate, whether an E2 enzyme needs to be charged with activated Ub or FAT10 (Groettrup et al., 2008). Activated FAT10 can be conjugated by the UBA6 specific E2 enzyme (USE1), which was recently identified in a yeast two-hybrid screen. Therefore, UBA6 as well as USE1 are examples that enzyme sharing between ULMs is much more common than initially believed. USE1 is not only the first described E2 enzyme in the FAT10 conjugation pathway but also the first physiological substrate of FAT10 conjugation, as it was efficiently auto-FAT10ylated in cis but not in trans (Aichem et al., 2010). Co-expression of NUB1L, a linker protein that facilitates the degradation of FAT10 and its conjugates, led to proteolytic down regulation of USE1 which suggest that USE1 can auto- modify itself with FAT10 and thus negatively regulates the FAT10 conjugation pathway (A. Aichem, manuscript in preparation). To date, no E3 enzyme for FAT10 has been identified. In our laboratory a specific interaction of FAT10 and the RING finger E3 ligase TRIM11 could be shown in a yeast two hybrid screen (A. Aichem, unpublished), indicating that this protein may be involved in the FAT10 conjugation pathway. This finding could further be strengthened by the fact that TRIM11 interacts in vivo with the FAT10 specific E2 enzyme USE1, and siRNA-mediated knockdown of TRIM11 reduced the amount of FAT10 conjugates in HEK293 cells, indicating that TRIM11 may function as a FAT10 specific RING finger E3 ligase (A. Aichem, unpublished).

Introduction

3.2.4 The small ubiquitin-like modifier SUMO

The small ubiquitin-like modifier protein (SUMO) was initially identified in 1996, almost two decades later than Ub, in the yeast Saccharomyces cerevisiae as a suppressor of mutations in the centromere protein MIF2 and designated suppressor of MIF2 mutations-3 (Smt3p) (Meluh and Koshland, 1995). Almost coincidentally, the mammalian homolog of the yeast protein was discovered and referred to as SUMO-1 (small ubiquitin-like modifier 1), UBL1 (ubiquitin-like 1), PIC1 (promyelocytic leukemia protein (PML)-interacting protein 1), sentrin or GMP1 (GTPase- activating protein (GAP)-modifying protein 1) (Boddy et al., 1996; Mahajan et al., 1997; Matunis et al., 1996; Okura et al., 1996). The highly conserved SUMO is ubiquitously expressed in all eukaryotic cells but is absent in bacteria and archaea. To date, four SUMO isoforms SUMO-1/Smt3H3, SUMO-2/SmtH2, SUMO-3/Smt3H1 and SUMO-4/Smt3H4 have been described in mammals. SUMO1-3 is found to be expressed in all tissues at all developmental stages, whereas expression of SUMO-4 appears to be restricted in kidney and various immune tissues, especially lymphnodes and spleen (Bohren et al., 2004; Guo et al., 2004; Seeler and Dejean, 2003). Only three of these (SUMO-1, SUMO-2 and SUMO-3) are processed in vivo to bear the C-terminal diglycine motif required for post-translational conjugation. SUMO-2 and SUMO-3 are nearly identical and are assumed to be largely redundant in their functions. SUMOs share only ~18 % sequence identity with Ub, although structure analysis by nuclear magnetic resonance (NMR) revealed that both share a common three dimensional structure, which is characterized by a tightly packed globular fold with five anti-parallel β-sheets wrapped around one α-helix (Bayer et al., 1998). Reversible attachment of SUMO, namely SUMOylation, to its targets is controlled by a strict enzymatic energy-consuming reaction cascade that is analogous to the Ub pathway. First, the C-terminal end is processed by a protease (in human these are the SENP proteases and Ulp1 in yeast) to expose a C-terminal di-glycine motif, which is required for substrate protein binding. Subsequently, SUMO becomes activated by an E1 SUMO-activating enzyme (heterodimeric SAE1 and SAE2). Then, it is transferred to the E2 enzyme ubiquitin carrier 9 (Ubc9) and finally becomes attached to an ε-amino group of a specific lysine in the target proteins mostly via one of several E3 enzymes. Further, SUMO can be deconjugated from the target protein by the action of SUMO specific proteases (Hay, 2005; Johnson and Blobel, 1997; Su and Li, 2002). During SUMOylation the target lysine residue is generally located within a recognizable consensus sequence, namely ψKxE, where ψ is an aliphatic and x an arbitrary amino acid (Sampson et al., 2001; Seeler and Dejean, 2003).

Introduction

The presence of a consensus site is not a strict requirement for SUMOylation of a target since several proteins have been found to be modified on non-consensus sites (Hoege et al., 2002; Pichler et al., 2005). Interestingly, SUMO proteins do not have the lysine residue corresponding to K48 in the Ub molecule that is required for the most common formation of poly-ubiquitin chains, suggesting that SUMO does not build the same type of multi-chains as Ub (Bayer et al., 1998). In contrast to ubiquitin where all seven lysine residues of Ub have been implicated in linkage formation (Xu et al., 2009), SUMO chains are linked mainly through a single lysine residue at position 11 at the N-terminus, which is embedded in the ψKxE consensus sequence (Bencsath et al., 2002; Bylebyl et al., 2003; Knipscheer et al., 2007; Skilton et al., 2009; Tatham et al., 2001). Protein SUMOylation is a highly regulated post-translational modification that is involved in versatile modes of regulation in widely different biological processes. Modification of target proteins with SUMO has been shown to be involved in the subcellular localization of proteins, maintenance of genome integrity, protein–protein interactions and mainly in the transactivation of transcription factors (Anckar and Sistonen, 2007; Garaude et al., 2008; Muller et al., 2004; Pichler and Melchior, 2002; Ulrich, 2008; Yang et al., 2003; Yeh, 2009; Zhao, 2007). For instance, the observation that RanGAP1, the first identified SUMO substrate and the promyelocytic leukemia protein (PML), also known as TRIM19, are targeted to distinct subcellular structures upon conjugation to SUMO-1 let suggest, that SUMOylation might play an important role in regulating the subcellular localization of proteins (Mahajan et al., 1997; Matunis et al., 1996; Seeler and Dejean, 2001). Moreover, transcription factors such as AP-1 and p53 are two of many examples where modification with SUMO regulates their transcriptional activity. For instance, SUMOylation of the heterodimeric transcription factor complex composed of c-Jun and c-Fos, which belong to the AP-1 family, decreases their transactivation potency (Bossis et al., 2005; Muller et al., 2000), whereas SUMOylation of the AP-1 family member JunB in T lymphocytes can positively regulate cytokine gene transcription and likely plays a critical role in T cell activation (Garaude et al., 2008). Furthermore, SUMOylation is influenced by other protein modifications like phosphorylation or acetylation (Urvalek et al., 2011; Yao et al., 2011). Interestingly, in addition to the structural relationship between the UBLs, Ub and SUMO, a crosstalk between SUMO and Ub-based signaling was described, sharing a multitude of functional interrelations. This include the targeting of the same attachment sites in specific substrates, such as the inhibitor of NF-kBα (IκBα), the proliferating cell nuclear antigen (PCNA) and the NF-κB essential modulator (NEMO) (Desterro et al., 1998; Hoege et al., 2002; Huang et al., 2003).

Introduction

In addition, SUMOylation can lead to ubiquitination of proteins and vice versa and thereby inhibit or support proteasome mediated protein degradation (Hay, 2005). Enzyme activities that control either SUMOylation or ubiquitination are regulated by the respective other modification (Miteva et al., 2010).

3.3 Inhibitor of apoptosis protein (IAP) family

Inhibitor of apoptosis proteins (IAPs) are an evolutionarily conserved class of multifunctional proteins (Srinivasula and Ashwell, 2008), which are defined by the presence of one to three tandem baculoviral IAP repeats (BIRs). The BIR domain is a highly conserved zinc binding fold of approximately 70 amino acids, harboring a CX2CX6WX3DX5HX6C consensus sequence (Hinds et al., 1999), which folds to a three stranded β-sheet surrounded by six short α-helices to coordinate a zinc ion chelated by three cysteine and one histidine residues (Birnbaum et al., 1994; Wu et al., 2000). The BIR domain is a prerequisite for protein–protein interaction (Hinds et al., 1999; Sun et al., 1999) and the inhibitory activity of IAPs in apoptosis (Takahashi et al., 1998). The first members of IAPs were discovered in studies with baculoviruses, where binding and inhibiting cysteine-containing aspartate-specific proteases (caspases) contributed to the efficient infection and replication cycle in the host (Birnbaum et al., 1994; Crook et al., 1993). The name IAP derived from the ability, to efficiently prevent apoptosis by binding and inhibiting active caspases and apoptotic protease activating factor 1 (APAF-1) through the BIR domains. Owing to this structural motif, IAPs are assigned to the BIR containing proteins (BIRPs) which can be classified into two groups: the IAP-like BIRPs, which are well established as apoptosis regulators and the survival-like BIRPs with important functions in cell cycle and cytokinesis (Verhagen et al., 2001). Apoptosis, or programmed cell death, is a critical cellular process in normal development and homeostasis of multicellular organisms (Thompson, 1995) and caspases are the executioners in both, intrinsic and extrinsic pathways of apoptosis by cleaving a plethora of cellular components (Riedl and Shi, 2004). Several members of the IAP family can act both, upstream and downstream of caspase activation to promote cell survival (Deveraux and Reed, 1999). The action of IAPs can be opposed by certain pro-apoptotic factors, especially second mitochondria-derived activator of caspases (Smac/DIABLO) which promotes cytochrome-c dependent caspase activation (Vucic et al., 2002) or the evolutionarily conserved trimeric serin protease HtrA2/Omi (Martins, 2002; Suzuki et al., 2004; Vande Walle et al., 2008), which unleashes caspase activity in a biphasic process.

Introduction

It frees the active forms of caspases-3, -7 and -9 by proteolytically removing their natural inhibitors (Yang et al., 2003). Smac and HtrA2 are synthesized as larger cytosolic precursors that are proteolytically cleaved in mitochondria to expose their N-terminal tetrapeptide IAP binding motifs. In response to apoptotic stimuli, the processed Smac and HtrA2 are released into the cytosol where they can bind to a hydrophobic core at the BIR domain of IAPs (Liu et al., 2000; Vaux and Silke, 2003; Wu et al., 2000). IAPs can effectively suppress apoptosis induced by a variety of stimuli, including death receptor activation, growth factor withdrawal, ionizing radiation, viral infection, and genotoxic damage. Indeed, IAPs are frequently overexpressed in many types of human cancer and are associated with chemoresistance, disease progression and poor patient survival (Hunter et al., 2007; LaCasse et al., 1998). On the contrary, loss of IAPs is in some cases associated with the development of certain types of cancer. The physical association of the two best studied IAPs, XIAP and survivin, was found to convey the ubiquitin-dependent activation of NF-κB, which than drives the expression of genes important for cell migration, cell invasion and metastasis (Mehrotra et al., 2010). Therefore, depending on the cellular context, IAPs seem to have both, pro-tumorigenic and anti-tumorigenic roles (Keats et al., 2007). Moreover, several IAPs have been shown to promote proteasomal degradation of apoptotic proteins by catalyzing their ubiquitination and thereby regulating key components in cell death signaling cascades (Bartke et al., 2004; Vucic et al., 2011). Recently, IAPs have emerged as broader regulators of cellular homeostasis, with functions extending beyond tumor development and apoptosis inhibition (Srinivasula and Ashwell, 2008), such as inflammatory signaling and immunity, mitogenic kinase signaling and mitosis (Dogan et al., 2008; Gyrd-Hansen et al., 2008; O'Riordan et al., 2008). The activity of IAP is strictly regulated by different mechanisms. For RING domain containing IAPs a decreased half-life was observed due to the fact that they catalyze self-ubiquitination and proteasome-mediated degradation in a RING-dependent manner. This provides the interesting possibility that abundance is actively self-regulated (Yang et al., 2000). In addition, some IAP antagonists, such as Reaper, Grim and Hid, also stimulate IAP auto- ubiquitination and proteasomal degradation (Yoo et al., 2002).

Introduction

3.3.1 BRUCE represents a special BIR containing protein (BIRP)

The inhibitor of apoptosis protein, BIR repeat containing ubiquitin-conjugating enzyme (BRUCE), represents a special member of the BIR containing protein (BIRP) family. It is a giant, highly conserved 528 kDa membrane associated protein which localizes mainly to the trans-golgi network (TGN) and perinuclear vesicles (Hauser et al., 1998). It bears on one hand a single N-terminal BIR domain and on the other hand a C-terminal UBC-E2-domain, respectively. In contrast, most other BIRPs contain several BIR-domains and a C-terminal RING finger domain. Sequence analysis of BRUCE’s BIR domain revealed that BRUCE belongs to the survival-like BIRPs suggesting a role in cell cycle regulation and cytokinesis. Remarkably, BRUCE is to date the only IAP described, which contains a functional C- terminal UBC-domain, characteristic for E2 enzymes of the ubiquitin proteasome system (Hauser et al., 1998). Due to the presence of this domain, BRUCE can form a thioester with Ub in vitro and mediate Ub-transfer to substrate proteins (Bartke et al., 2004; Hao et al., 2004; Hauser et al., 1998). These structural features indicate that BRUCE may combine properties of Ub-conjugating enzymes and IAP-like proteins and let suppose that the family of IAP-like proteins is functionally and structurally more diverse than previously expected. The BRUCE cDNA was originally discovered in a homology screen for Ub conjugating enzymes in mice (Hauser et al., 1998). It is expressed in almost every mouse tissues with the highest expression in brain, liver and kidney (Hauser et al., 1998). The human homologue, Apollon, is like BRUCE widely expressed in almost every adult tissue and overexpressed in some drug-resistant cancer cells, suggesting that it may have an anti- apoptotic function (Chen et al., 1999). The importance and relevance of this antiapoptotic characteristic was further investigated and ascertained in numerous studies. For instance, the expression level of BRUCE in rat brain and in cultured hippocampal neurons after treatment with the neurodegenerative component kainic acid was explored. Thereby it was shown that BRUCE could promote survival of distinct neurons, whereas kainic acid treatment leads to BRUCE downregulation, leading to increased caspase-3 activation and cell death (Sokka et al., 2005). Downregulation of BRUCE expression in brain cancer cells with antisense oligonucleotides significantly sensitized the cells to apoptosis induced by DNA damaging agents, which is consistent with the fact that cancer cells overexpressing BRUCE show resistance to apoptosis inducing agents (Chen et al., 1999). The most severe functional effect could be observed in gene-targeted BRUCE knockout mice, which led to impaired placenta development and growth retardation, resulting in embryonic lethality discernible after embryonic day 14.

Introduction

Despite the anti-apoptotic activities of BRUCE in cells, apoptosis was neither detected in mutant placenta nor in mouse embryonic fibroblasts (MEFs) (Lotz et al., 2004). Further, also deletion of the C-terminal half of BRUCE, including the UBC domain, causes activation of caspases and apoptosis in the placenta and yolk sac, leading to embryonic lethality. This apoptosis is associated with up-regulation, stabilization and nuclear localization of the tumor suppressor p53, leading to the activation of Pidd, Bax and Bak, which might be specifically accompanied by mitochondrial-triggered apoptosis (Ren et al., 2005). A unique function of BRUCE during mouse development lies in the regulation of spongiotrophoblast cell proliferation. Progressive loss of the spongiotrophoblast layer in Birc6/BRUCE mutants results in lethality at day 11.5 and 14.5 of embryonic development. This further suggests a role for BRUCE, in addition to be an apoptosis inhibitor, as cell division regulator (Hitz et al., 2005). Genetic analysis in Drosophila has demonstrated that the Drosophila homologue dBRUCE, inhibits cell death induced by the essential Drosophila cell-death activators Reaper and Grim, resulting in viable but infertile male Drosophila BRUCE mutants (Vernooy et al., 2002). This argued for a role of dBRUCE in specialized pathways leading to cell death. Moreover, the final stage of spermatid terminal differentiation in Drosophila requires the elimination of most of the cytoplasm, a process known as spermatid individualization, which is locally and temporary accompanied by an apoptosis like capase activation (Arama et al., 2003). Male sterile dBRUCE -/- flies failed to exclude the bulk cytoplasm which leads to sperm nucleus hyper-condensation and finally degeneration, indicating an uncontrolled or excessive apoptotic process. Interestingly, transient caspase activation is also involved during mammalian spermatogenesis, where spermatid cytoplasm needs to be eliminated and in consequence, there might be a need for the anti-apoptotic potential of BRUCE (Arama et al., 2003). Despite the significant role for BRUCE in blocking apoptosis, very little is known about the regulation of BRUCE expression. Jansen and colleagues could show, that the treatment of myoblasts with prostaglandin F2alpha (PGF2α) reduces cell death and promotes myotube growth during myogenesis via upregulation of BRUCE through a pathway dependent on the nuclear factor of activated T cell 2 transcription factor (NFAT) (Jansen and Pavlath, 2008). This finding describes a hitherto unrecognized relationship between NFAT signaling and regulation of IAP expression. As aforementioned BRUCE belongs, with respect to its BIR-domain, to the survival-like BIRPs suggesting a role in cell cycle regulation and cytokinesis. Pohl and Jentsch reported in 2008, that BRUCE plays a crucial role at final stages of cytokinesis and particularly controls proper midbody ring formation which is required for cell cycle continuation.

Introduction

During cell cycle, BRUCE concentrates in a pericentriolar compartment in interphase, moves partially to spindle poles in metaphase, and finally localizes to the spindle midzone and the midbody in telophase and during cytokinesis, where it serves as a platform for the membrane delivery machinery and binds mitotic regulators and components of the vesicle targeting machinery. BRUCE depletion causes cytokinesis defects and cytokinesis associated apoptosis. Notably, Ub relocalizes from midbody microtubules to the midbody ring during cytokinesis and depletion of BRUCE disrupts this process. Upon mitotic exit, BRUCE is targeted to the midbody ring via its C-terminus binding to mitotic kinesin-like protein 1 (MKLP1), which is a core component of the midbody ring. Both, BRUCE and MKLP1 are ubiquitinated and ubiquitin-specific protease Y (UBPY) was shown to serve as their deubiquitinating enzyme (Wu et al., 2004). This let suggest that BRUCE coordinates multiple steps required for abscission and ubiquitination of different protein targets may be a crucial trigger (Pohl and Jentsch, 2008). Further, Pohl and Jentsch could show that the midbody ring disposal by autophagy is a post-abscission event of cytokinesis, suggesting that autophagy is coupled to cytokinesis (Pohl and Jentsch, 2009). Moreover, a novel regulatory role for BRUCE was uncovered in the interplay with the effector death caspase1 (Dcp-1) in starvation-induced autophagy during early Drosophila melanogaster oogenesis. These findings provide new insights into the molecular mechanisms that regulate autophagic and apoptotic events in vivo (Hou et al., 2008). Interestingly, BRUCE itself can be degraded by autophagy and this controls DNA fragmentation and cell death in nurse cells during late oogenesis in Drosophila melanogaster. These results reveal autophagic degradation of an IAP as a mechanism of triggering cell death and thereby provide a mechanistic link between autophagy and cell death (Nezis et al., 2010). BRUCE is a unique and an especial member of the BIRP family, due to its size, localization and its functions. BRUCE can act as a chimeric E2-E3 Ub ligase and can mono-ubiquitinate the IAP-antagonists Smac/Diablo, HtrA2 and active caspase-9 in vitro, which promotes their degradation (Bartke et al., 2004; Hao et al., 2004; Qiu and Goldberg, 2005). Therefore, BRUCE is thought to preserve cell survival by antagonizing apoptosis induced by spontaneously released pro-apoptotic factor Smac (Hao et al., 2004). Hereby, BRUCE can associate with precursors as well as mature forms of Smac by binding to regions in addition to the IAP binding motif, which promotes degradation of Smac and inhibits the activity of caspase-9 but not the effector caspase-3 (Qiu and Goldberg, 2005). Moreover, Smac as well as HtrA2 are able to compete for BRUCE-bound caspases. In response to apoptotic stimuli, BRUCE itself can be a substrate and cleaved by caspases and HtrA2 depending on the specific stimulus and the cell type (Sekine et al., 2005).

Introduction

Besides the regulation of caspases/proapoptotic factors, BRUCE itself seems to be regulated via ubiquitin-dependent degradation. Previous studies have implicated that the TRIM protein Nrdp1/RNF41 (neuregulin degrading protein 1/ring finger protein 41), a RING finger containing ubiquitin E3 ligase initially found to be involved in the degradation of ErbB/EGFR family of receptor tyrosine kinases (Diamonti et al., 2002; Qiu and Goldberg, 2002; Qiu et al., 2004), can act as a ubiquitin E3 ligase for BRUCE and mediate its proteasomal degradation. Thereby cellular levels of BRUCE are maintained, impairing inhibition of apoptosis. Overexpression of Nrdp1 was shown to decrease cellular levels of BRUCE; therefore Nrdp1 can be important in the initiation of apoptosis by catalyzing ubiquitination and degradation of BRUCE (Qiu et al., 2004). Moreover, Nrdp1 regulates the turnover of the proteasome subunit hRpn13 and Parkin, which is a member of the E3 ubiquitin ligase family (Liu et al., 2007; Yu and Zhou, 2008). Nrdp1 itself undergoes self-ubiquitination which leads to its proteasomal degradation. Notably, Nrdp1 also specifically interacts and becomes stabilized by the deubiquitinating enzyme USP8/UBPY (ubiquitin-specific protease 8 or Y) (Wu et al., 2004), a cysteine protease implicated in cell cycle regulation, efficient downregulation of the EGF receptor and stability regulation of ESCRT-0 components Hrs and STAM2 (Cao et al., 2007; Clague and Urbe, 2006; Wright et al., 2011).

3.4 Autophagy

Autophagy, literally meaning ‘self-eating’, is an evolutionarily conserved intracellular catabolic process, in which portions of the cytoplasm are sequestered within cytosolic double- membrane vesicles called autophagosomes and subsequently delivered to the lysosome to allow degradation and recycling of the cargo (Kundu and Thompson, 2008). So far, the endoplasmic reticulum (ER), the Golgi, mitochondria, and plasma membrane have all been implicated in autophagosome formation (Hailey et al., 2010; Tooze and Yoshimori, 2010). The autophagy-lysosome system is, beside the UPS, the other major protein degradation mechanism in eukaryotes (Kirkin et al., 2009; Nedelsky et al., 2008). Autophagy embraces three major intracellular pathways in eukaryotic cells namely, macroautophagy, microautophagy, and chaperone mediated autophagy (CMA), which share a common destiny of lysosomal degradation, but are mechanistically different from one another (Cuervo, 2011; Klionsky, 2005; Li et al., 2011; Mijaljica et al., 2011) (Figure 7). The process of mammalian autophagy is divided into six principal steps: initiation, nucleation, elongation, closure, maturation and degradation or extrusion (Orsi et al., 2010).

Introduction

Figure 7: Distinct types of autophagy. Cytosolic proteins can enter the lysosome for degradation by at least three autophagic pathways. (a) Macroautophagy is usually a catabolic process in which proteins, organelles or other cytosolic components are sequestered within cytosolic double-membrane vesicles called autophagosomes and subsequently delivered to the lysosome to allow degradation and recycling of the cargo. Selective variations of this process, in which distinct substrates (aggregate proteins or organelles) are targeted for degradation, and their names, are also depicted. (b) During microautophagy the lysosomal membrane itself is envisaged as undergoing local rearrangement to directly engulf portions of cytoplasm or any constituents and are thereupon internalized after membrane scission and degraded in the lumen of the organelle. Cytosolic material can be sequestered ‘in bulk’ or selectively with the help of a cytosolic chaperone that recognizes the substrates. (c) Chaperone-mediated autophagy (CMA) is a type of autophagy distinct from the other two autophagic pathways owing to its selectivity, saturability and competitivity by which a subset of long-lived cytosolic soluble proteins are directly delivered into the lysosomal lumen via specific receptors. Soluble cytosolic proteins containing a targeting motif are recognized by the cytosolic heat shock cognate 70 (HSC70) chaperone and its co-chaperones, which deliver the substrate to the membrane of the lysosome. After docking onto the cytosolic tail of the lysosomal receptor, the substrate protein unfolds and crosses the lysosomal membrane through a multimeric complex. Substrate translocation requires a lumenal HSC70 chaperone and is followed by rapid degradation in the lysosomal lumen. CMA participates in quality control to maintain normal cell functions by clearing "old" proteins and provides energy to cells under nutritional stress. Figure taken from (Cuervo, 2011).

Introduction

During macroautophagy (autophagy), as depicted in Figure 7 (a), proteins, intact organelles (such as mitochondria) and portions of the cytosol are sequestered into a double-membrane vesicle, termed autophagosome. Subsequently, the completed autophagosome matures by fusing with an endosome and/or lysosome, thereby forming an autolysosome. This latter step exposes the cargo to lysosomal hydrolases to allow its breakdown and the resulting macromolecules are released back into the cytosol through membrane permeases for reuse (Beynon and Bond, 1986; Mortimore and Poso, 1987).

During microautophagy cytoplasmic materials are translocated into the lysosome or vacuole for degradation by direct invagination, protrusion, or septation of the lysosomal or vacuolar membrane, as depicted in Figure 7 (b) (Mijaljica et al., 2011).

By contrast, CMA translocates unfolded, soluble proteins directly across the limiting membrane of the lysosome (see Figure 7 (c)). CMA is activated as part of the cellular response to oxidative stress to target oxidized proteins to lysosomes without perturbing neighbouring unaffected proteins. Also, during prolonged starvation, the selectivity of CMA provides cells amino acids through selective degradation of expendable proteins (Kiffin et al., 2004). Through these diverging mechanisms cells ensure quality control, development and survival under nutrient deprivation and maintain cellular homeostasis by eliminating unnecessary proteins or damaged organelles (Cuervo, 2004; Klionsky, 2005; Li et al., 2011; Mizushima et al., 2008). Autophagy may be triggered under physiological conditions, such as nutrient starvation, or in response to a variety of stress stimuli, such as exposure to radiations or cytotoxic compounds. Moreover, autophagic processes have been described to play an important role in energetic balance, in cellular and tissue remodeling, aging and in the cellular defense against extracellular insults and pathogens. Basal autophagy also plays a key role in eliminating defective organelles or aggregated proteins that may be resistant to the ubiquitin-proteasome degradation pathway and thereby sustain the cellular homeostasis (Komatsu et al., 2006; Ravikumar et al., 2002). Although autophagy is basically a protective mechanism that sustains cell survival under adverse conditions, it has been demonstrated that the induction of autophagic process can contribute to a wide range of diseases, including cancer, neurodegeneration and microbial infection (Kundu and Thompson, 2008; Lee, 2009; Martinet et al., 2009). Therefore, as an intracellular self-destructive system, autophagy must be tightly regulated in order to adapt to different intracellular and extracellular stresses (see Figure 8).

Introduction

Figure 8: Macroautophagy is extensively involved in cellular homeostasis The morphological features of macroautophagy are depicted schematically. The initial sequestering compartment, the phagophore, expands into the double-membrane autophagosome. Fusion with an endosome generates the single-membrane amphisome, which subsequently fuses with a lysosome. The degraded cargo is released back into the cytosol through permeases. Some of the physiological connections between macroautophagy and human health and disease are indicated by the surrounding terms. Figure taken from (Klionsky, 2010).

Currently, there are over 34 proteins known which are required for autophagy. They were first identified in yeast and called autophagy related (Atg) proteins (Yang and Klionsky, 2010) and almost all of them are involved in the induction of autophagy, autophagosome nucleation, vesicle expansion and completion, and final retrieval of Atg proteins from mature autophagosomes. One key Atg-containing complex is the autophagy-specific class III phosphatidylinositol 3- kinase (PI3P-kinase), also known as Vps34, which produces PI3P at the site of autophagosome formation (Simonsen and Tooze, 2009). In mammals, activation of the membrane anchored kinase Vps34 is dependent on the formation of a multi-protein complex that consists of Beclin1 (Becn1), UV irradiation resistance-associated tumour suppressor gene (UVRAG), Endophilin B1 (Bif-1), activating molecule in Beclin1-regulated autophagy1 (Ambra1), and a myristylated serine kinase Vps15 (Fimia et al., 2007; Takahashi et al., 2007). The evolutionarily conserved Becn1 (the mammalian ortholog of Atg6), originally discovered as a Bcl-2-interacting protein (Liang et al., 1998) represents the first human protein shown to be indispensable for autophagy and exist in several complexes involved in autophagosome formation and maturation (Cao and Klionsky, 2007; Liang et al., 1999).

Introduction

It interacts with proteins that positively regulate autophagy, such as Atg14-like (Atg14L), UVRAG, Bif-1, run domain and cysteine-rich domain containing, Becn1-interacting protein (Rubicon), Ambra1, neuronal PDZ domain protein interacting specifically with TC10 (nPIST), (Orr, 2002), vacuole membrane protein 1 (VMP1) (Vaccaro et al., 2008), signaling lymphocyte-activation molecule (SAM) (Berger et al., 2010), inositol-1,4,5 trisphosphate receptor (IP(3)R) (Criollo et al., 2007), PTEN-induced kinase I (PINK-1) and survivin (Roca et al., 2008), to regulate the lipid kinase Vps-34 protein and promote formation of Becn1-Vps34- Vps15 (Kihara et al., 2001) core complexes. Moreover, it can interact with anti-apoptotic proteins such as Bcl-2 via its BH3-like domain to negatively regulate autophagy (He and Levine, 2010; Levine et al., 2008; Maiuri et al., 2007; Pattingre et al., 2005). The Becn1-Vps34-Vps15 complex plays a crucial role in the induction of the autophagic process by generating Phosphatidylinositol-3-phosphate (PtdIns(3)P) rich membranes, which act as platforms for Atg protein recruitment and autophagosome nucleation (Kihara et al., 2001; Simonsen and Tooze, 2009). Many of the signals modulating autophagy converge at the (PtdIns(3)P)-related kinase mammalian target of rapamycin (mTOR), which acts as a “gatekeeper” in controlling nutrient signaling and induction of autophagy (Codogno and Meijer, 2005). The inhibition of mTOR ultimately leads to the formation of autophagosomes via Atg. Two ubiquitin-like conjugation systems are involved in autophagy. The UBLs Atg8 (LC3 in mammalians), which is prior cleaved by Atg4 to expose the C-terminal glycine, and Atg12 become activated by the common E1-like enzyme Atg7. Atg8 is then conjugated by the E2 enzyme Atg3 and finally becomes lipidated with phosphatidylethanolamine (PE) and inserted in the autophagosomal membrane, where it remains after completion of autophagosome formation (Ichimura et al., 2000; Kabeya et al., 2000; Kirisako et al., 1999). Atg12 in turn becomes conjugated to Atg5 via the E2 enzyme Atg10 where it forms a protein complex with Atg16 through oligomerization of Atg16 monomers (Ichimura et al., 2000; Shintani et al., 1999). The Atg5/Atg12/Atg16 complex asymmetrically localizes to the outer side of the autophagosomal membrane throughout the elongation process and dissociates from the autophagosome upon completion (Ichimura et al., 2000).

Introduction

3.4.1 Ambra1 (activating molecule in Becn1-regulated autophagy)

Ambra1 is a ~1.300 aa highly conserved protein in vertebrates, which was identified in a large-scale mutagenesis approach based on gene trapping in mice. Interestingly, it possesses a WD40 domain at its N-terminus and WD domain containing proteins have been described to have critical roles in many essential biological functions ranging from signal transduction, transcription regulation, to apoptosis (Smith et al., 1999). Moreover, proteins containing WD40 repeats are known to serve as platforms for the assembly of protein complexes or mediators of transient interplay among other proteins (Li and Roberts, 2001). As the name implies, Ambra1 associates with the Atg protein Becn1, which was identified in a yeast two-hybrid screen and is an important mediator of autophagy. Hereby, Ambra1 promotes Becn1 interaction with its target lipid kinase VPS34, thus mediating autophagosome nucleation (Fimia et al., 2007). Ambra1 positively regulates the Becn1-dependent programme of autophagy; hence downregulation of Ambra1 expression in a human fibrosarcoma cell line results in a remarkable decrease in rapamycin and nutrient deprivation-induced autophagy and rate of cell proliferation, whereas overexpression has the opposite effect. Ambra1 plays a crucial role during embryogenesis, since null mutations in mice resulted in embryonic lethality with severe neural tube defects associated with autophagy impairment, accumulation of ubiquitinated proteins, unbalanced cell proliferation and excessive apoptotic death (Fimia et al., 2007). These results support the existence of a complex interplay between autophagy, cell growth and cell death required for neural development in mammals. Further, Ambra1 can co-immunoprecipitate with Vps34, suggesting that Becn1, Vps34 and Ambra1 are components of a multiprotein complex. This is supported by the fact that downregulation of Ambra1 also significantly reduces the ability of Becn1 to interact with its target lipid kinase Vps34 and thereby prevent autophagosome nucleation (Fimia et al., 2007). This protein complex resides at the dynein motor complex due to a direct binding between Ambra1 and the kinase dynein light chain 1 (DLC1). When autophagy is induced, Ambra1 gets phosphorylated by the unc-51-like kinase 1 (ULK1) which leads to the release of the Ambra1/Becn1 autophagy core complex from the dynein motor complex. This interaction permits translocation of the complex to the endoplasmic reticulum (ER) to initiate autophagosome nucleation (see Figure 9) (Di Bartolomeo et al., 2010). In addition, DLC1 downregulation and Ambra1 mutations in its DLC1-binding sites strongly enhance autophagosome formation, which reveal a new function for Ambra1, not only to be a co-factor of Becn1 in favouring its kinase-associated activity, but also to act as a crucial upstream regulator of autophagy initiation (Fimia et al., 2011).

Introduction

Figure 9: Dynamic interaction of Ambra1 with dynein regulates autophagy induction. Left part: In nutrient-rich conditions, mTorc1 binds to and phosphorylates Ulk1, inhibiting its activity. The Beclin1 complex is associated with dynein through a direct interaction of Ambra1 with the dynein light chain 1 or 2 (DLC1/2). This interaction keeps the Beclin1 complex apart from autophagosome formation sites, such as the endoplasmic reticulum (ER). Middle part: Autophagic stimuli, such as nutrient starvation, inhibit mTorc1 activity and induce its release from Ulk1. Active Ulk1 is then able to phosphorylate Ambra1, thus causing the dissociation of the Beclin1 complex from dynein. Right part: Once released from dynein, the Beclin1 complex translocates to the ER where it generates phosphatidylinositol-3-phosphate (PtdIns(3)P)-enriched membranes, labeled by the PtdIns(3)P binding protein DFCP1. These modified membranes allow the recruitment of several ATG proteins, which are responsible for the formation of the phagophore, the autophagosome precursor. Dynein HC: dynein heavy chain; DIC: dynein intermediate chain. Figure taken from (Fimia et al., 2011).

Moreover, the anti-apoptotic Bcl-2 family members, which were described to negatively regulate autophagy and Ambra1, interact via its BH3-like domain with Becn1 at the mitochondrial outer membrane and control autophagy (Pattingre et al., 2005). Here, Ambra1 preferentially binds the mitochondrial pool of Bcl-2 and this interaction is disrupted following autophagy induction. Further, Ambra1 can compete with both mitochondrial and ER-resident Bcl-2 (mito-Bcl-2 and ER-Bcl-2, respectively) to bind Becn1 (Strappazzon et al., 2011). This interaction is dissociated upon nutrient starvation, leading to Ambra1-Becn1 association at the ER, mitochondria or other phagophore assembly sites (PAS) to stimulate autophagy, which provides novel insight into the importance of subcellular location on the activity of autophagy proteins and the crosstalk between autophagy and apoptosis (Tooze and Codogno, 2011).

Introduction

Recently, Parkin was identified as a novel Ambra1 interaction partner using tandem affinity purification in HEK293 cells. Parkin is a member of the E3 ubiquitin ligase family that is defined by a tripartite RING1-in-between-ring (IBR)-RING2 motif which can augment binding of the E2 proteins UbcH7 and UbcH8, and the subsequent ubiquitination of the proteins synphilin-1, Sept5, and SIM2. Parkin translocates from the cytosol to depolarized mitochondria and induces their autophagic removal (mitophagy), whereby translocation is not Ambra1, but phosphatase and tensin homolog (PTEN)-induced putative kinase 1 (PINK1) dependent (Geisler et al., 2010; Matsuda et al., 2010; Vives-Bauza et al., 2010). In particular, Ambra1 is recruited to perinuclear clusters of depolarized mitochondria and activated class III PI3K in their immediate vicinity. Van Humbeeck et al could recently show, that during prolonged mitochondrial depolarization the interaction of endogenous Parkin and Ambra1 strongly increased, which is important for subsequent mitochondrial clearance of Parkin- mediated mitophagy (Van Humbeeck et al., 2011).

3.5 Transcription factors

Transcription factors regulate the specificity of transcription from DNA to mRNA alone or in a complex with other proteins, as an activator or as a repressor. Eukaryotic transcription factors can be classified into several families on the basis of conserved specific sequences among their DNA-binding domains (DBD) and the genes that they regulate (Latchman, 1997; Mitchell and Tjian, 1989). Such regions were identified on the basis that they can stimulate transcription when linked to the DNA binding domain of a completely unrelated factor and are known as activation domains (see Figure 10). Transcription of many genes, particularly in higher eukaryotes is further dependent upon multiple physiological signals (Ptashne and Gann, 1997).

Figure 10: Schematic model of gene transcription A transcription factor specifically regulates gene expression by binding to distinct consensus sequences in DNA strains (schematic model).

Introduction

3.5.1 The transcription factor AP-1

Much of our current knowledge about the characteristics of transcription factors arises from the discovery and study of activating protein 1 (AP-1). The transcription factor AP-1 was first identified as a 12-O-tetradecanoylphorbol-13-acetate (TPA)-inducible transcriptional factor that binds to an essential cis-element of the human metallothionein IIa (hMTIIa) promoter and an enhancer element of the simian virus 40 (SV40) promoter (Lee et al., 1987). This binding is required for optimal basal activity of these promoters in vivo (Haslinger and Karin, 1985) and in vitro (Lee et al., 1987). Phorbol esters such as TPA are potent tumor promoters, capable of potentiating the effect of a subcarcinogenic dose of an initiating carcinogen (Blumberg et al., 1981; Slaga, 1983; Weinstein et al., 1979). AP-1 is assembled of various dimeric basic region-leucine zipper (bZIP) proteins. It functions as homo- or heterodimer to form AP-1 transcriptional complexes composed of structurally and functionally related proteins belonging to the Jun (c-Jun, JunB and JunD), Fos (c-Fos, FosB, Fra-1 and Fra-2), Maf (c-Maf, MafB, MafA, MafG/F/K and Nrl) and ATF (ATF2, LRF1/ATF3, B-ATF, JDP1, JDP2) early multi-gene sub-families (Hess et al., 2004). The stability and activity of AP-1 directly results from the composition of its dimers (Eferl and Wagner, 2003; Shaulian and Karin, 2001). The broad combinatorial possibilities provided by the large number of AP-1 proteins determine its binding specificity and affinity and consequently, the spectrum of regulated genes (Hess et al., 2004). Each AP-1 family member is differentially expressed and regulated, which means that every cell type has an intricate mixture of AP-1 dimers, slightly diverse in their functions (Wagner, 2001). Hence, the accurate interplay between AP-1 family members plays an essential role in gene transcription because imbalances have been shown to induce malignant cell transformation (Angel and Karin, 1991). The homo-/hetero dimers mediate AP-1 regulated gene expression through binding to the palindromic TPA Responsive Element (TRE), composed of the 7 bp DNA consensus sequence 5’-TGA G/C TCA-3’ (Angel et al., 1987), which are present in promoters of many cellular and viral genes. Jun-Fos heterodimers bind preferentially to the heptameric TRE consensus sequence, whereas Jun-ATF or ATF homodimers bind with higher affinity to a related 8 bp motif 5'- TTACCTCA-3' and to the sequence 5’-TGA GC TCA-3’ known as the cyclic AMP responsive element (CRE) (Angel and Karin, 1991; Chinenov and Kerppola, 2001; Nakabeppu et al., 1988).

Introduction

AP-1 activity is induced by a plethora of physiological stimuli and environmental insults. In turn, AP-1 regulates a wide range of cellular processes, including cell migration and proliferation, differentiation and cell transformation during development and in adult tissues, autophagy and survival, but also plays a key role in regulating inflammatory processes, and apoptosis, (Ameyar et al., 2003; Eferl and Wagner, 2003; Shaulian, 2010; Shaulian and Karin, 2002; Yogev et al., 2010; Yogev and Shaulian, 2010). AP-1 is at the receiving end of complex signaling pathways in response to many pathological and physiological impulses, including cytokines, cell–matrix interactions, growth factors, stress signals, viral and bacterial infections as well as oncogenic stimuli and regulates, upon activation, numerous target genes (Colin et al., 2011; Hess et al., 2004). But even prior to stimulation a considerable basal activity of TRE-containing genes could be detected due to a certain amount of active AP-1, enabling to establish the role of AP-1 in controlling both, basal and inducible transcription of several genes (Angel et al., 1987). Moreover, the activity of AP-1 can be modulated through changes in transcription of genes encoding AP-1 subunits, control of their mRNAs stability, by interactions with other transcriptional regulators and cofactors, dimer composition, translational regulation, (Vesely et al., 2009) post-translational modifications (Garaude et al., 2008), interaction with other proteins (Herrlich and Ponta, 1994) and is further controlled by upstream kinases that link AP-1 to various signal transduction pathways (Eferl and Wagner, 2003; Radler-Pohl et al., 1993; Rincon and Flavell, 1994). The mechanism of post-translational control is most extensively documented in the case of mitogen- and cellular stress-induced hyper- phosphorylation and, in particular, activation of cJun through the Jun N-terminal kinase (JNK) cascade (Karin et al., 1997).

3.5.1.1 Structural and biochemical properties of AP-1 The ability of a single transcription factor to control a versatile collection of diverse biological processes originates primarily from its structural and regulatory complexity. A common feature of AP-1 proteins is the evolutionarily conserved bZIP domain, the collective term for a basic DNA-binding domain combined with a leucine zipper region consisting of a periodic repetition of leucine residues (Landschulz et al., 1988; O'Shea et al., 1989). This leucine-zipper provides a dimerization motif and forms an X-shaped amphipatic α-helix with the adjacent hydrophobic leucine region which requires dimerization to bind to the DNA backbone, as depicted in Figure 11 (Glover and Harrison, 1995).

Introduction

Figure 11: Crystal structure of the Jun-Fos heterodimer. The bZIP domains of Jun and Fos form an X-shaped α-helical structure, which binds to the palindromic TRE/AP-1 site (TGAGTCA) (Glover and Harrison, 1995). The bZIP domain of Jun is shown in blue and the bZIP domain of Fos in red. The DNA backbone is shown in yellow. http://www.ncbi.nlm.nih.gov/Structure/mmdb/mmdbsrv.cgi?uid=72084

Above all, the leucine zipper composition plays a crucial role for the major specificity and stability of homo- and heterodimers formed by the various Jun, Fos and ATF proteins (Eferl and Wagner, 2003). While Fos protein members (c-Fos, FosB, Fra-1, Fra-2) are not able to homo- but heterodimerize (Hai and Curran, 1991), Jun (c-Jun, JunB and JunD) homo- and heterodimers can bind to the TPA response element (TRE), thereby stimulating the transcription of target genes. These properties are significantly enhanced by the cooperative formation of transcriptionally active heterodimer complexes between Jun proteins and Fos or ATF family members with significant functional consequences (Chiu et al., 1989; Chiu et al., 1988; Cohen et al., 1989; Halazonetis et al., 1988; Nakabeppu et al., 1988; Rauscher et al.,

1988; Smeal et al., 1989; Zerial et al., 1989). Beside the well-characterized bZIP domain, Jun and Fos family members include a transactivation domain, which vary between the individual Jun and Fos proteins regarding its potential (Angel and Karin, 1991), and docking sites for several kinases (Karin et al., 1997).

Introduction

3.5.1.2 Transcriptional and post-transcriptional regulation of AP-1 Post-translational modification has been extensively studied for the Jun family members, which become phosphorylated and activated by the stress activated protein kinase (SAPK) cascade (Karin et al., 1997). SAPK, which are Jun N-terminal Kinases (JNK) and p38- kinases, belong to the Mitogen-Activated Protein Kinase (MAPK) superfamily. Activated by variant stress stimuli through a MAPK cascade, JNK can translocate to the nucleus, phosphorylate serine and threonine residues within its N-terminal transactivation domain and thereby modulate its transactivation potential (Hess et al., 2004). Further, it could be shown, that the activation of c-Jun after phosphorylation by MAPK is accompanied by a reduction in c-Jun ubiquitination and consequent stabilization of the protein, revealing an important role for regulated protein degradation in the signal-dependent control of gene expression (Musti et al., 1997). Additionally, AP-1 can be modulated by interaction with other cofactors and transcription factors like for instance the protein-protein interaction based crosstalk via Glucocorticoid Receptors (GR) (Herrlich and Ponta, 1994; Hess et al., 2004). This transactivation potential between individual Jun and Fos proteins can differ significantly. The AP-1 family members c-Jun, c-Fos and FosB are considered to be strong transactivators, whereas JunB, JunD, Fra-1 and Fra-2 possess only a weak transactivation potential. Specific circumstances can provoke, that the latter may even act as repressors of AP-1 activity by competing for binding to AP-1 sites or by forming inactive heterodimers with Jun, Fos or FosB (Angel and Karin, 1991; Hess et al., 2004).

3.5.2 The transcription factor JunB

The transcription factor JunB belongs to the AP-1 family with the predicted size of ~35.7 kDa. The expression of JunB can be triggered by numerous extracellular stimuli, such as serum, growth factors, phorbol esters (TPA) and activators of protein kinase A (PKA) due to transcriptional activation of the junB promoter. The 5´flanking region of junB contains recognition sequences for Smad and Ets, (Coffer et al., 1994), transcription factors, which can be regulated by MAP-kinases. Further, it contains an IL-6 response element (IL-6RE) (Nakajima et al., 1993) containing a STAT3 binding site and a CRE and SRE (serum response element) like site (Kitabayashi et al., 1993), a GC rich region, an inverted repeat (IR) element (de Groot et al., 1991) and a myeloid-specific IL-6RE (Sjin et al., 1999) in the proximal promoter region. Additionally, the regulation of JunB by v-src involves the CAAT and TATA box region (Apel et al., 1992).

Introduction

AP-1 transcription factors integrate both mitogenic and stress signaling pathways. Signaling pathways mediated by growth factors induce JunB through a TRE, a SRE and two Ets-linked motifs located in a region around -1000 to - 2000 in the mouse JunB promoter (Phinney et al., 1996). In contrast, Pdgfb, serum, bFGF, phorbol ester and forskolin, can induce JunB expression by a SRE and a CRE site located in the 3’ flanking region of the junB gene (Perez-Albuerne et al., 1993). Moreover, four NF-κB binding sites located downstream of the gene have been shown to mediate transcriptional induction of JunB in response to oxygen deprivation (Schmidt et al., 2007) (Figure 12).

Figure 12: Schematic model of the junB promoter.

JunB has been described to act antagonistically to c-Jun in transcriptional regulation and is proposed to be a negative regulator of cell proliferation (Chiu et al., 1989; Schutte et al., 1989). During organogenesis in mouse, c-Jun and JunB have distinct tissue-specific roles in cell proliferation and differentiation during fetal development (Wilkinson et al., 1989). In comparison to c-Jun, JunB has a ten-fold decreased activity to activate AP-1-responsive genes containing single AP-1 binding sites to induce oncogenic transformation, due to a small number of amino acid changes between its DNA-binding and dimerization motifs (Deng and Karin, 1993). But strikingly, JunB appears to be as effective as Jun in trans-activating reporter genes containing multiple AP-1 binding sites which suggest, that trans-activation by JunB may require synergistic interactions between multiple homodimers bound to adjacent sites (Angel and Karin, 1991; Chiu et al., 1989). Increased JunB expression can suppress cell proliferation by transcriptional activation of the cyclin-dependent kinase inhibitor p16INK4α and elevated JunB expression in 3T3 cells also inhibited Ras- and Src-mediated transformation and tumor growth in vivo (Passegue and Wagner, 2000). Further, the antagonism between c-Jun and JunB plays an important role in regulating cell cycle progression, through negatively regulating cyclin D1 during mitosis by increasing c-Jun activating phosphorylation and triggering the phosphorylation dependent degradation of JunB via p34cdc2-cyclin B kinase on three proline-flanked serine or threonine residues (Ser23, Thr150 and Ser186) in M and early G1 phase of the cell cycle (Bakiri et al., 2000).

Introduction

Surprisingly, high JunB expression in S phase, drop during mid- to late G2 phase due to accelerated phosphorylation-dependent degradation by the proteasome, an incident which is absolutely required for proper mitosis. Consistently, abnormal JunB expression in late G2 phase entails a variety of mitotic defects, thus JunB might contribute to tumorigenesis (Farras et al., 2008). Moreover, JunB is suggested to become phosphorylated by JNK in T cells at threonine residues 102 and 104 which is important for synergy with c-Maf transcription factor to activate IL-4 expression and T helper cell differentiation (Li et al., 1999). Beside, JNKs are responsible to regulate the E3 ubiquitin ligase Itch via phosphorylation dependent activation. Itch can bind via the WW domains to the proline-rich sequence PPxY (PY) motif of JunB which targets JunB for ubiquitination in T cells (Gao et al., 2004). Mice lacking Itch show an accumulation of JunB in helper T cells, which lead to an increased Th2 differentiation (Fang et al., 2002). One mechanism by which AKT kinase-dependent hypersensitivity to mammalian target of rapamycin (mTOR) inhibitor is controlled, is by the differential expression of cyclin D1 and c- MYC. Vartanian et al. could recently show, that AKT-kinase dependent Itch-mediated JunB degradation has a regulatory function in the transcriptional responses of cyclin D1 and c- MYC to rapamycin (Vartanian et al., 2010). Furthermore, the (HECT)-type ubiquitin ligase Smurf1 interacts with JunB through the PPxY motif and negatively regulates mesenchymal stem cell (MSC) proliferation and differentiation by controlling JunB turnover through ubiquitination and the proteasome pathway. This represents another regulatory mechanism controlling JunB function in cells (Zhao et al., 2010). Recently, Das et al. described that an increased expression of the disulfide oxidoreductase Thioredoxin (Trx), a protein involved in redox dependent cellular functions, activates NFκB and induces enhanced binding and transactivation potential of the AP-1 family members, JunB, c-Jun and Fra1, by activating JNK subgroup of MAPKs (Das, 2001; Das and Muniyappa, 2010). JunB has a unique, non-redundant function in vivo, because it is required for proper endothelial morphogenesis (Licht et al., 2006) and lack of JunB in mice causes defects in placentagenesis and vascularisation which leads to embryonic lethality between day 8.5 and 10.0 (Schorpp-Kistner et al., 1999). Moreover, it was shown, that loss of JunB expression in the epidermis of adult mice severely affects the skin and bone formation due to high systemic levels of the negatively regulated JunB-target granulocyte colony-stimulating factor (G-CSF) (Hess et al., 2003; Meixner et al., 2008).

Introduction

Interestingly, JunB loss in mice has an impact on fat metabolism, due to increased level of adipose triglyceride lipase and hormone-sensitive lipase, the key enzymes of lipolysis (Pinent et al., 2011). Further, JunB represses the IL-6 promoter, which is reflected in the fact, that epidermal loss of JunB leads to a Systemic lupus erythematosus (SLE) phenotype, a complex autoimmune disease, due to hyper IL-6 signaling (Pflegerl et al., 2009). Beside, JunB is an essential target gene of hypoxia-induced signaling via NF-κB, responsible for hypoxia-mediated vascular endothelial growth factor (VEGF) expression and tumor angiogenesis, which is hypoxia-inducible factor (HIF) independent (Schmidt et al., 2007; Textor et al., 2006; Wang et al., 2010). Thus, JunB is a critical independent regulator of the autocrine and paracrine acting VEGFα. Interestingly, a decreased JunB expression was further described for synovial fibroblasts from rheumatoid arthritis patients (Huber et al., 2003). Moreover, deletion leads to a phenotype resembling the histological and molecular hallmarks of psoriasis, including arthritic lesions (Zenz et al., 2005). JunB has revealed an important regulatory role in the hematopoietic system and tumorigenesis, since JunB inactivation in postnatal mice results in a myeloproliferative disease resembling early human chronic myelogenous leukemia (CML) (Passegue et al., 2001; Passegue et al., 2004). In agreement with this, JunB expression is diminished in some human chronic myeloid leukemia (Yang et al., 2003) and B cell leukemias (Ott et al., 2007; Szremska et al., 2003), and downregulation of JunB partially depends on junB gene methylation (Yang et al., 2003). JunB is also described as a major determinant for maintenance of dividing and adult muscle cells by inducing hypertrophy and thereby efficiently preventing atrophy (Raffaello et al., 2010). Another emergent role for JunB was characterized for the differentiation of naïve CD4+ T cells into T helper1 and 2 cells, which are classified by their specific set of cytokines. Here, JunB expression is not only essential, but has to be tightly adjusted to ensure a proper cell function, such as a Th2 immune response (Hartenstein et al., 2002). Furthermore, the post-translational modification of JunB with small ubiquitin-like modifier (SUMO) on lysine 237 plays a critical role in T cell activation. Indeed, SUMOylation of JunB regulates its ability to induce cytokine gene transcription and likely plays a critical role in T cell activation (Garaude et al., 2008). For natural killer (NK) cell activation, JunB plays a pivotal role in the negative regulation of the NKG2D-ligand, activating receptor RAE-1- epsilon expression (Nausch et al., 2006). Moreover, Textor et al. described novel functions for JunB in regulating novel target genes in mast cells, which are required for proper mast cell degranulation and mast cell mediated angiogenic processes (Textor et al., 2007). Very recently, another level of regulation is provided where JunB and c-Rel cooperatively enhance Foxp3 expression during induced regulatory T (iTreg) cell differentiation (Son et al., 2011).

Introduction

A further level of regulation implicates the AP-1 proteins with autophagy. Interestingly, unlike c-Jun, which can enhance apoptosis, the data implicating JunB in the regulation of apoptosis are less significant. In fact, it has been suggested that JunB inhibits cytokine-mediated apoptosis in an insulin-producing cell line and in purified rat primary beta cells (Gurzov et al., 2008), or in nigral neurons following axotomy (Winter et al., 2002). Conversely, Yogev et al. could show recently, that JunB and c-Jun can inhibit autophagy induced by starvation, overexpression of a short form of ARF (smARF) or after rapamycin treatment and deregulation of JunB expression, when autophagy is specifically required, tilts the fate of starved cells to apoptosis (Yogev et al., 2010; Yogev and Shaulian, 2010). Although JunB has been considered to be a tumor suppressor, indeed, it has also a cell cycle-promoting role through activating the transcription of the cell cycle regulator cyclin A (Andrecht et al., 2002). Passegue et al. demonstrated, using a knock-in strategy and a transgenic complementation approach, that JunB may substitute for c-Jun in improving survival of c-Jun-deficient embryos (Passegue et al., 2002). These data support previous studies which demonstrated that JunB is required for cell cycle re-entry after quiescence (Kovary and Bravo, 1991) and can cooperate with c-Jun in the development of mouse fibrosarcoma (Bossy-Wetzel et al., 1992), and collectively suggest that JunB is able to enhance proliferation, cooperate with c-Jun and promote neoplastic transformation, especially lymphomas. This goes along with findings revealing that in some hyperproliferative T cell lymphomas JunB was highly overexpressed (Mao et al., 2004; Rassidakis et al., 2005). Accordingly, JunB is a critical regulator of the expression of cytokines important for T lymphocyte proliferation and differentiation, namely IL-4 (Hartenstein et al., 2002) and IL-2 (Jain et al., 1992; Li et al., 1999). Beside, JunB expression and binding capacity are decreased in anergic T cells, which do not produce IL-2 (Heisel and Keown, 2001; Mondino et al., 1996; Schwartz, 1997; Sundstedt and Dohlsten, 1998). Moreover, aberrant JunB and CD30 expression is involved in the development of Hodgkin lymphoma (HL) and anaplastic large cell lymphoma (ALCL), whereby initiation and constitutive transcription of CD30 gene by aberrant JunB expression maintains hypomethylation of CD30 CpG islands, contributing to the pathogenesis of HL and ALCL (Watanabe et al., 2008). In addition, JunB, like c-Jun, binds to the promoter of the tumor promoter Dmp1 and elevates its expression. Dmp1 subsequently up-regulates the expression of p19ARF, another tumor suppressor residing in the INK4 locus, which is a major regulator of p53 activity (Sreeramaneni et al., 2005). These mechanisms may also be the basis for the enhancement of drug-induced senescence by JunB (Yogev et al., 2006).

Aim of this study

4 Aim of this study

FAT10 is beside ubiquitin the only ULM that can target proteins for proteasomal degradation (Hipp et al., 2005). Thus, FAT10 is part of one of the most important regulating systems in living cells to determine the fate of target proteins for degradation. However, the precise molecular mechanisms of the FAT10 conjugation pathway in cell biology and immunology are poorly investigated.

The objective of this study was to gain more insight into the FAT10 conjugation pathway, starting with the identification of novel UBA6 interacting proteins in a yeast two-hybrid screen. Regarding the fact that UBA6 is so far the only known FAT10 activating enzyme it could be possible that new identified partners of UBA6 can give a hint for novel E1-E2-E3 cascades for FAT10. One of the interaction partners, the inhibitor of apoptosis protein BRUCE, should be analyzed biochemically and with respect to a potential function as a FAT10 specific E2 enzyme, since BRUCE contains a highly conserved C-terminal UBC domain, characteristic for E2 conjugating enzymes. Moreover, this protein was a very interesting candidate to investigate because it has been previously shown to act as a chimeric E2/E3 ubiquitin ligase (Bartke et al., 2004).

Beyond this initial scope, a second yeast two-hybrid screen with the putative FAT10 specific E3 ligase TRIM11 was performed to identify further target proteins in the FAT10 conjugation pathway. Two of the identified interaction partners, namely the transcription factor JunB and Ambra1, should be further investigated biochemically and by microcopy techniques to delineate the underlying molecular basis in terms of being FAT10 specific substrates. Moreover, functional consequences of JunB FAT10ylation in terms of modification of transcriptional activity should be investigated.

Materials and Methods

5 Materials and Methods

5.1 Materials

5.1.1 Chemicals and Materials

3-Amino-1,2,4-triazole (3-AT) Sigma, Steinheim ß-mercaptoethanol Sigma, Steinheim Adenosine triphosphate (ATP) Sigma, Steinheim Agar Fluka, Steinheim Agarose Fluka, Steinheim Amonium persulphate (APS) Sigma, Steinheim Aprotinin Fluka, Steinheim Bromophenol blue Riedel de Haen, Steinheim Complete Mini Protease-EDTA-free Inhibitor Cocktail (PI) Roche, Mannheim Creatine Phosphate Sigma, Steinheim Cycloheximide Sigma, Steinheim Dimethyl sulfoxide (DMSO) Merck, Darmstadt Ethidium bromide Roth, Karlsruhe Fetal Calf Serum (FCS) Linaris, Wertheim-Bettingen Glycerol AppliChem, Darmstadt HA-peptide Sigma, Steinheim IFN-γ Peprotech, Hamburg Inorganic pyrophosphatase Sigma, Steinheim Leupeptin Sigma, Steinheim MG132 (Z-Leu-Leu-Leu-CHO) Sigma, Steinheim Non-essential amino acids (NEAA) Invitrogen, Carlsbad N,N,N’,N’- Tetramethylethylenediamine (TEMED) Fluka, Steinheim NP-40 Fluka, Steinheim NuPAGE SDS MES, Tris-Acetate Running Buffer Invitrogen, Carlsbad (+Antioxidans) Pepstatin Sigma, Steinheim Phenylmethylsulfonyl fluoride (PMSF) Fluka, Steinheim Polyethylene glycol 3500 (PEG 3500) Sigma, Steinheim Polysorbate (Tween 20) Sigma, Steinheim Roti-Block Roth, Karlsruhe Rotiphorese Gel 40 Roth, Karlsruhe Sodiumdesoxycholate Merck, Darmstadt Sodium dodecyl sulfate (SDS) AppliChem, Darmstadt TNF-α Peprotech, Hamburg Tris Roth, Karlsruhe Tryptone AppliChem, Darmstadt TRITON-X-100 Fluka, Steinheim Trypsin/EDTA Lonza, Verviers Yeast extract Fluka, Steinheim

Materials and Methods

Yeast nitrogenbase w/o amino acids Sigma, Steinheim

Antibiotics Ampicillin Fluka, Steinheim Kanamycin Fluka, Steinheim Penicillin/Streptomycin Sigma, Steinheim

Gels NuPAGE 4-12 % Bis/Tris Gels Invitrogen, Carlsbad NuPAGE 3-8 % Tris-Acetate Gels Invitrogen, Carlsbad

Protein and DNA markers GeneRuler 1 kb DNA Ladder Fermentas, St. Leon-Rot SeeBlue Plus2 Pre-Stained Standard Invitrogen, Carlsbad Precision Plus Protein Prestained Standard Bio-Rad, München HiMark™ Pre-stained Protein Standard Invitrogen, Carlsbad

Transfer membrane Protran nitrocellulose membrane Whatman

5.1.2 Reagents and reaction kits

NucleoSpin® Plasmid Macherey-Nagel, Düren GenElute Plasmid Miniprep Kit Sigma-Aldrich, Seelze GenElute PCR Clean-Up Kit Sigma-Aldrich, Seelze GenElute Gel Extraction Kit Sigma-Aldrich, Seelze NucleoBond PC 100 Midiprep Kit Macherey-Nagel, Düren Dual-Luciferase® Reporter (DLR™) Assay System Promega Restore Western Blot Stripping Buffer Thermo Scientific, Rockford Super Signal West Pico Chemiluminescent Thermo Scientific, Rockford Super Signal West Femto Maximum Sensitivity Substrate Thermo Scientific, Rockford

5.1.3 Buffers and solutions

5.1.3.1 Media for cell culture Dulbecco's Modified Eagle Medium (DMEM) Lonza, Verviers Earle´s Balanced Salt Solution (E7510) Sigma, Steinheim Roswell Park Memorial Institute (RPMI 1640) Lonza, Verviers

Materials and Methods

5.1.3.2 Stock solutions

Table 2: Stock solutions: Composition and Storage.

Component Composition / Storage

3-AT 1 M in H2O. Storage at -20 °C. -1 Aprotinin 10 mg ml in H2O. Storage at -20 °C. -1 Histidine 10 g l in H2O (bidest) and filtered through a 0,2 μM sterile filter. Storage at 4 °C.

IPTG 1 M IPTG in H2O (bidest) and filtered through a 0,2 μM sterile filter. Storage at -20 °C.

-1 Leucine 10 g l in H2O (bidest) and filtered through a 0,2 μM sterile filter. Storage at 4 °C.

-1 Leupeptin 1 mg ml in H2O. Storage at -20 °C.

MG132 10 mM in DMSO. Storage at -20 °C.

Pepstatin 10 mg ml-1 EtOH. Dissolved in EtOH with heat. Storage at -20 °C.

PMSF 10 μM in 10 ml Isopropanol. Storage at -20°C protected from light.

-1 Tryptophane 10 g l in H2O (bidest) and filtered through a 0,2 μM sterile filter. Storage at 4 °C.

-1 Uracil 2 g l in H2O (bidest) and filtered through a 0,2 μM sterile filter. Storage at 4 °C.

X-GAL 100 mg ml-1 in DMF. Storage at -20 °C protected from light.

5.1.3.3 Antibiotics and Inductors If not indicated divergently antibiotics and inductors are added to medium with following final concentrations.

Table 3: Antibiotics and inductors used in this work. Component Final concentration -1 Ampicillin (Ap) 100 mg l -1 Kanamycin (Km) 50 mg l -1 Chloramphenicol (Cm) 20 mg l -1 Tetracycline (Tc) 25 mg l

Stock solutions of Chloramphenicol and Tetracycline listed in Table 3 are prepared with ethanol. All other substances are dissolved in H2O and filtered through a 0,2 μM sterile filter.

5.1.3.4 Buffers and Solutions for Agarose Gel Electrophoresis (AGE) Tris Acetate EDTA (TAE) Buffer (50 x) Table 4: Composition of TAE buffer. Component Concentration Tris-Acetate 2 M Acetic acid 57,1 % EDTA 50 mM

The pH is adjusted to 8.4 with 1M NaOH.

Materials and Methods

DNA loading dye solution (6x): Table 5: Composition of DNA loading buffer (6x). Component Concentration Glycerol 60 % Tris (pH 7.6) 5 mM Bromphenolblue 0,1 % Xylene cyanol 0,1 %

Tris-EDTA (TE) buffer Table 6: Composition of TE buffer. Component Final concentration

Tris 10 mM

EDTA 1 mM pH is adjusted to 7.4 with acetic acid (100 %) 5 M NaOH and buffer is autoclaved for 25 min at 121 °C.

5.1.3.5 Buffers and Solutions for SDS-Polyacrylamide Gel Electrophoresis

SDS-PAGE Loading Dye Solution (5 x Laemmli Buffer) Table 7: Composition of 5 x Laemmli buffer. Component Concentration Tris-HCl 225 mM Glycerol (v/v) 50 % SDS (w/v) 5 % Bromophenol blue (w/v) 0,05 % 2-mercaptoethanol 4 %

Tris Glycine SDS (TGS) Running Buffer Table 8: Composition of TGS buffer. Component Concentration Glycine 192 mM Tris 25 mM -1 SDS 1 g l

Staining Solution for SDS Polyacrylamide Gels Table 9: Composition of SDS-PAGE staining solution. Component Concentration

H2O (v/v) 65 % 2-Propanol (v/v) 25 % Acetic acid (v/v) 10 % Coomassie brilliant blue G-250 2 g l-1

Materials and Methods

Destaining Solution for SDS Polyacrylamide Gels Table 10: Composition of SDS-PAGE destaining solution. Component Concentration

H2O (v/v) 65 % 2-Propanol (v/v) 25 % Acetic acid (v/v) 10 %

Towbin buffer (10x) Wet-blot Table 11: Composition of Towbin buffer (10x). Component Concentration Tris 250 mM Glycine 1,92 M pH is set to 8.3 with a 37 % HCl solution.

Towbin buffer (1x) Wet-blot Table 12: Composition of Towbin buffer (1x). Component Concentration

H2O (v/v) 70 % Towbin buffer (10x) (v/v) 10 % MeOH (v/v) 20 %

5.1.3.6 IP and lysis buffer FAT10 IP-buffer Table 13: Composition of FAT10 IP-Buffer. Component Concentration TrisHCl (pH 7.6) 20 mM NaCl 50 mM

MgCl2 10 mM ATP 4 mM DTT 0,1 mM Inorganic pyrophosphatase 5 U ml-1 Creatine-phosphate 20 mM Creatine-phosphokinase 4 μg ml-1

Materials and Methods

BRUCE-IP buffer Table 14: Composition of BRUCE-IP buffer. Component Concentration NaCl 150 mM HEPES (pH 7.2) 50 mM EDTA 2 mM NP-40 0,1 %

Lysis Buffer for bacteria

Table 15: Composition of Lysis buffer for bacteria. Component Concentration NaCl 100 mM Tris-HCl (pH 8.0) 50 mM EDTA 1 mM

5.1.3.7 Washing and elution buffer GST high-salt washing buffer Table 16: Composition of GST high-salt washing buffer. Component Concentration NaCl 300 mM EDTA 1 mM Tris-HCl (pH 8.0) 50 mM

Washing Buffer NET-TON Table 17: Composition of Washing buffer NET-TON. Component Concentration NaCl 650 mM EDTA 5 mM Tris-HCl (pH 8.0) 50 mM Triton X-100 0,5 %

Washing Buffer NET-T Table 18: Composition of Washing buffer NET-T. Component Concentration NaCl 150 mM EDTA 5 mM Tris-HCl (pH 8.0) 50 mM Triton X-100 0,5 %

Materials and Methods

GST-elution buffer Table 19: Composition of GST-elution buffer. Component Concentration Tris (pH 8.0) 50 mM Reduced Glutathione 10 mM

5.1.3.8 Cultivation Media and Media Additives for bacteria Luria Bertani (LB) Liquid Medium Table 20: Composition of LB medium. Component Concentration [g l-1] Casein peptone 10 NaCl 5 Yeast extract 5

Components listed in Table 20 are dissolved in H2O, pH is set to 7.4 with 5 M NaOH and medium is autoclaved for 25 min at 121 °C (when used in shake flask or test tube cultivations).

Luria Bertani (LB) Solid Agar Table 21: Composition of LB agar. Component Concentration [g l-1] Casein peptone 10 NaCl 5 Yeast extract 5 Agar 15

Components of Table 21 are dissolved in H2O, pH is set to 7.4 with 5 M NaOH and medium is autoclaved for 25 min at 121 °C.

SOC Medium Table 22: Composition of SOC medium. Component Concentration Casein tryptone/peptone 20 g l-1 Yeast extract 5 g l-1 NaCl 10 mM

MgSO4 10 mM KCl 2,5 mM Glucose 2 mM

Components listed in Table 22 are dissolved in H2O and medium is autoclaved for 25 min at 121 °C. After cooling down, addition of an appropriate volume of a 1M sterile-filtrated (0,2 μM) Glucose stock solution

Materials and Methods

Half-Synthetic Glycerine Medium (HSG) Table 23: Composition of HSG medium. Component Concentration Casein tryptone/peptone 20 g l-1 Yeast extract 7 g l-1 Glycerin (99.5 %) 14,9 g l-1 NaCl 2,5 g l-1 -1 K2HPO4 2,3 g l -1 KH2PO4 1,5 g l -1 MgSO4 x H20 0,14 g l

Components listed in Table 23 are dissolved in 800 ml H2O, pH is set to 7.4 with 5 M NaOH and then refilled to the final volume. Medium is autoclaved for 25 min at 121 °C.

5.1.3.9 Cultivation Media and Media Additives for yeast YPAD-Medium Table 24: Composition of YPAD-medium. Component Concentration Yeast extract 1 % Peptone 2 % Glucose 2 % Adenine sulphate 0,004 %

Components listed in Table 24 are dissolved in H2O and medium is autoclaved for 25 min at 121 °C. After cooling down, addition of an appropriate volume of a 20 % sterile-filtrated glucose stock solution. Sterile filter pore size (0,2 μM).

YPAD-Agar Table 25: Composition of YPAD-agar. Component Concentration Yeast extract 1 % Peptone 2 % Glucose 2 % Adenine sulphate 0,004 % Agar 2 %

Components listed in Table 25 are dissolved in H2O and medium is autoclaved for 25 min at 121 °C. After cooling down, addition of an appropriate volume of a 20 % sterile-filtrated glucose stock solution. Sterile filter pore size (0,2 μM).

Materials and Methods

Synthetic dropout (SD) mix (-TLH) Table 26: Composition of synthetic dropout (SD) mix (-TLH). Component Concentration Adenine, Hemisulfate 0,2 % L-Alanine 0,77 % L-Arginine 0,77 % L-Asparagine 0,77 % L-Aspartate 0,77 % L-Cysteine 0,77 % L-Glutamine 0,77 % L-Glutamate 0,77 % Glycine 0,77 % myo-Inositol 0,77 % L-Isoleucine 0,77 % L-Lysine monohydrochloride 0,77 % L-Methionine 0,77 % para-Aminobenzoic acid 0,07 % L-Phenylalanine 0,77 % L-Proline 0,77 % L-Serine 0,77 % L-Threonine 0,77 % L Tyrosine 0,77 % Uracil 0,77 % L-Valine 0,77 %

Components listed in Table 26 are mixed overnight at 4 °C.

SD/-T (-Trp) - medium Table 27: Composition of SD/-T-medium. Component Concentration Amino acid dropout mix SD (-TLH) 1 g l-1 Leucine 0,1 g l-1 Histidine 0,02 g l-1 Yeast nitrogen base w/o aa 6,7 g l-1 Glucose 10 %

Components listed in Table 27 are dissolved in H2O and autoclaved for 25 min at 121 °C. After cooling down, addition of an appropriate volume of a 20 % sterile-filtrated glucose stock solution, leucine and histidine stock solution. Sterile filter pore size (0,2 μM).

Materials and Methods

SD/-T (-Trp)-Agar Table 28: Composition of SD/-T-agar. Component Concentration Amino acid dropout mix SD (-TLH) 1 g l-1 Leucine 0,1 g l-1 Histidine 0,02 g l-1 Yeast nitrogen base w/o aa 6,7 g l-1 Glucose 10 % Agar 4 %

Components listed in Table 28 are dissolved in H2O and autoclaved for 25 min at 121 °C. After cooling down, addition of an appropriate volume of a 20 % sterile-filtrated glucose stock solution, leucine and histidine stock solution. Sterile filter pore size (0,2 μM).

SD/-L (-Leu) - medium Table 29: Composition of SD/-L-medium. Component Concentration Amino acid dropout mix SD (-TLH) 1 g l-1 Tryptophane 0,01 g l-1 Histidine 0,02 g l-1 Yeast nitrogen base w/o aa 6,7 g l-1 Glucose 10 %

Components listed in Table 29 are dissolved in H2O and autoclaved for 25 min at 121 °C. After cooling down, addition of an appropriate volume of a 20 % sterile-filtrated glucose stock solution, tryptophane and histidine stock solution. Sterile filter pore size (0,2 μM).

SD/-L (-Leu) - agar Table 30: Composition of SD/-L-agar. Component Concentration Amino acid dropout mix SD (-TLH) 1 g l-1 Tryptophane 0,01 g l-1 Histidine 0,02 g l-1 Yeast nitrogen base w/o aa 6,7 g l-1 Glucose 10 % Agar 4 %

Components listed in Table 30 are dissolved in H2O and autoclaved for 25 min at 121 °C. After cooling down, addition of an appropriate volume of a 20 % sterile-filtrated glucose stock solution, tryptophane and histidine stock solution. Sterile filter pore size (0,2 μM).

Materials and Methods

SD/-TL (-Trp/-Leu) - medium Table 31: Composition of SD/-TL – medium. Component Concentration Amino acid dropout mix SD (-TLH) 1 g l-1 Histidine 0,02 g l-1 Yeast nitrogen base w/o aa 6,7 g l-1 Glucose 10 %

Components listed in Table 31 are dissolved in H2O and autoclaved for 25 min at 121 °C. After cooling down, addition of an appropriate volume of a 20 % sterile-filtrated glucose stock solution, histidine stock solution. Sterile filter pore size (0,2 μM).

SD/-TL (-Trp/-Leu) - agar Table 32: Composition of SD/-TL-agar. Component Concentration Amino acid dropout mix SD (-TLH) 1 g l-1 Histidine 0,02 g l-1 Yeast nitrogen base w/o aa 6,7 g l-1 Glucose 10 % Agar 4 %

Components listed in Table 32 are dissolved in H2O and autoclaved for 25 min at 121 °C. After cooling down, addition of an appropriate volume of a 20 % sterile-filtrated glucose stock solution, histidine stock solution. Sterile filter pore size (0,2 μM).

Overlay mix: X-GAL - Agarose Table 33: Composition of X-GAL-agarose. Component Concentration PBS containing 100 µg ml-1 X-Gal (100 mg ml-1 in DMF) Agarose 5 g l-1

STET-Buffer for DNA isolation (yeast) Table 34: Composition of STET-buffer Component Concentration Sucrose 8 % Triton X-100 5 % EDTA 50 mM Tris (pH 8.0) 50 mM Ammonium Acetate 7,5 M

Materials and Methods

5.1.4 Cell culture

5.1.4.1 Eukaryotic cell culture

Cell culture media

HEK293 DMEM, 10% FCS 1% Penicillin/Streptomycin

HEK293T DMEM, 10% FCS 1% Penicillin/Streptomycin

HELA DMEM, 10% FCS 1% Penicillin/Streptomycin

JURKAT RPMI 1640 10% FCS 1% Penicillin/Streptomycin 0,1 mM NEAA 50 μM ß-mercaptoethanol

5.1.4.2 Prokaryotic cell culture Strain Genotype

E.coli TOP10F´ (Invitrogen) F'[lacIq Tn10(tetR)] mcrA Δ(mrr-hsdRMS- mcrBC) φ80lacZΔM15 ΔlacX74 deoR nupG recA1 araD139 Δ(ara-leu)7697 galU galK rpsL(StrR) endA1 λ-

– - - E.coli BL21 (DE3) (Invitrogen) F ompT gal dcm lon hsdSB(rB mB ) λ(DE3 [lacI lacUV5-T7 gene 1 ind1 sam7 nin5])

5.1.4.3 Yeast Strain Genotype NMY51 (Dualsystems Biotech) (MATa his3 D 200 trp1-901 leu2-3,112 LYS2 :: (lexAop)4-HIS3 ura3 :: (lexAop)8- lacZ (lexAop)8-ADE2 GAL4

Materials and Methods

5.1.5 Antibodies

Primary antibodies

Table 35: Primary antibodies

Antibody Specification Source

α-β-actin mouse, monoclonal Abcam α-Ambra1 rabbit, polyclonal Abcam α-BIRC6 mouse, monoclonal BD Transduction Laboratories α-BIRC6 rabbit, polyclonal Abcam α-FAT10 (4F1) mouse, monoclonal (Aichem et al., 2010), Enzo Lifescience α-FAT10 rabbit, polyclonal (Hipp et al., 2005), Enzo Lifescience α-FLAG (M2) mouse, monoclonal Sigma, Steinheim α-FLAG (M2)-HRP mouse, monoclonal Sigma, Steinheim α-GST mouse, monoclonal Santa Cruz α-HA-Peroxidase (Klon HA-7) mouse, monoclonal Sigma, Steinheim α-HA (Klon HA-7) rabbit, polyclonal Sigma, Steinheim α-6-HIS-Pox mouse, monoclonal Sigma, Steinheim α-JunB rabbit, polyclonal Abcam α-TRIM11 rabbit, polyclonal Sigma, Steinheim α-Alexa Fluor®488-HA Conjugate mouse, monoclonal Invitrogen α-Alexa Fluor® 546 rabbit, polyclonal Invitrogen

Secondary antibodies

Table 36: Secondary antibodies Antibody Specification Source α-mouse-HRP goat, polyclonal Jackson Immuno Research α-rabbit-HRP goat, polyclonal Jackson Immuno Research

5.1.6 Enzymes and other cloning components

Table 37: Enzymes and other cloning components. Component Source

Calf Intestine Alkaline Phosphatase (CIAP) Fermentas, St. Leon-Rot Expand High FidelityPLUS PCR System Roche, Mannheim Expand Long Template PCR System Roche, Mannheim QuikChange Site-Directed Mutagenesis Kit Stratagene/Agilent Red Taq Polymerase Sigma, Steinheim Restriction enzymes/buffers Fermentas, St. Leon-Rot T4 Ligase/buffer Fermentas, St. Leon-Rot

Materials and Methods

5.1.7 Recombinant proteins

Table 38: Recombinant proteins used in this work Component Source

FAT10 N. Catone HIS-USE1 N. Catone FLAG-UBA6 Enzo Life Sciences GST-TRIM11 this work

5.1.8 Vectors used for cloning

Table 39: Vectors used for cloning. Component Source pACT2 Clontech pLexA-N Dualsystems

5.1.9 Plasmid constructs

Table 40: Plasmid constructs used in this work.

Construct Source

pcDNA3 Invitrogen pLex-A-FAT10 Dualsystems pcDNA3-HIS-3x-FLAG-FAT10 PD Dr. G. Schmidtke pcDNA3- HIS-3x-FLAG-FAT10ΔGG PD Dr. G. Schmidtke pcDNA3-TRIM11-FLAG Dr. A. Aichem pcDNA3-HIS/-A-TRIM11 Dr. A. Aichem pcDNA3.1-HA-FAT10 PD Dr. G. Schmidtke pcDNA3.1-HA-FAT10ΔGG Dr. A. Aichem pcDNA3.1-HA-UBA6 Dr. C. Pelzer pcDNA3.1-HA-Ubiquitin PD Dr. G. Schmidtke pcDNA3.1-HA-JunB Prof. Dr. M. Piechazcyk pcDNA3.1-HA-JunB-K237R Prof. Dr. M. Piechazcyk pcDNA3.1-HA-JunB-K267R Prof. Dr. M. Piechazcyk pcDNA3.1-HA-JunB-K301R Prof. Dr. M. Piechazcyk pcDNA3.1-HA-JunB-K237/267/301R (pcDNA3.1-JunB-K3R) Prof. Dr. M. Piechazcyk pCl3-FLAG-BRUCE Prof. Dr. S. Jentsch pGEX4T3-TRIM11 Dr. A. Aichem pCMV6-JunB-MYC-FLAG OriGene pCMV6-Ambra1-MYC-FLAG OriGene

Materials and Methods

5.1.9.1 Primers To generate the bait yeast two-hybrid plasmid pLexA-UBA6-plasmid the full length UBA6 sequence from pcDNA3.1-HA-UBA6 was amplified using

AA-113-forward: 5`-cgg ggt acc gaa gga tcc gag cct gtg gcc -3` containing a KpnI- Restriction-site as forward and AA-114-reverse: 5`-cta ctt cag tca tga cac tga tta agg tac ccc cg -3`, containing a KpnI- Restriction-site as reverse primer and cloned the into plasmid pLexA-N-plasmid (Table 39) (Dualsystems Biotech, Zurich, Switzerland).

To generate the control prey yeast two-hybrid plasmid pACT2-FAT10-plasmid the full length FAT10 sequence from pcDNA3.1-HA-FAT10 was amplified using

AA-58-forward: 5`- cgg gat ccg agc tcc caa tgc ttc ctg cct c -3` containing a BamHI- Restriction-site as forward and AA-59-reverse: 5`- ggc atc tta ttg tat tgg agg gtg aga att cgg -3`, containing a EcoRI- Restriction-site as reverse primer and cloned into plasmid pACT2-plasmid (Table 39) (Clontech). The sequences of the constructs that were made in this work were verified by sequencing (GATC Biotech, Konstanz).

5.2 Methods

5.2.1 Plasmid DNA purification

For preparative purposes a single bacterial colony was picked from an LB plate and used to inoculate 6 ml of LB medium containing the appropriate antibiotic. Cultures were allowed to grow overnight at 37 °C with vigorous shaking. Bacteria were harvested by centrifugation (14000 rpm, 1 min) and plasmid DNA was isolated by alkaline lysis and binding to anion- exchange matrices using the NucleoSpin® Plasmid (Macherey-Nagel), or GenElute Plasmid Miniprep Kit (Sigma-Aldrich) (see 5.1.2) according to the manufacturer’s instructions. For preparative purposes and to yield larger amounts of plasmid DNA for transfection of mammalian cells overnight cultures of 100-400 ml were used.

Materials and Methods

Bacteria were harvested at 4 °C (4000 rpm, 30min) and plasmid DNA preparation was carried out using the NucleoBond® PC 100 Midiprep Kit (see 5.1.2). DNA concentration was determined by measuring absorption at 260 nm using the NanoDrop spectrophotometer (Witec AG). For storage, 830 μl of bacterial cultures were mixed with 130 μl Glycerol (87 %), frozen in liquid nitrogen and stored at -80 °C.

5.2.2 Cloning

5.2.2.1 PCR (Polymerase chain reaction) The polymerase chain reaction (PCR) serves as an in vitro-amplification system of DNA molecules (Mullis and Faloona, 1987). In order to generate the different constructs for protein expression in yeast as well as in mammalian cells either pcDNA3.1-HA-UBA6 or pcDNA3.1- HA-FAT10 was used as a template in a PCR reaction together with the oligonucleotides indicated above (5.1.9.1) to amplify the desired DNA fragments. PCR conditions were optimized for the different fragments to yield enough PCR products for subsequent cloning steps. The different reaction mixtures with a total volume of 50 μl and the amplification conditions that were used are listed below:

UBA6 Components Volume [ μl ] Expand long template PCR system (Roche) 10 x buffer 5 DMSO 2,5 dNTP-mix (10 mM each) 2,5 AA-113 Oligonucleotide forward (10 μM) 2 AA-114 Oligonucleotide reverse (10 μM) 2 pcDNA3.1-HA-UBA6 (350 ng µl-1) 1 DNA-Polymerase (Expand Long template) 1

H2O 34

Materials and Methods

PCR program: Touchdown PCR Process Temperature [° C] Duration [sec] Repetition Initial denaturation 94 240 1 x Denaturation 94 60 Annealing 65 60 15 x Δ 1°C Elongation 68 210 Denaturation 94 60 Annealing 50 60 20 x Elongation 68 210 Final Elongation 68 600 1

Due to the size of the uba6 gene (3159 bp), it was not possible to obtain an amplification product with the correct size using standard PCR methods. Touchdown PCR uses a cycling program with varying annealing temperatures and thereby enhances the specificity of the initial primer–template duplex formation and hence the specificity of the final PCR product. We chose an initial annealing temperature of 65 °C and in subsequent cycles (15 x), the annealing temperature is decreased in steps of 1 °C/cycle. Afterwards, a standard PCR program was applied for further 20x cycles with a final elongation step of 10 min at 68 °C.

FAT10 Components Volume [ μl ] Expand High Fidelity Plus (Roche)

5 x buffer 2 (MgCl2) 10 DMSO 2,5 dNTP-mix (10 mM each) 2,5 AA-115: Oligonucleotide forward (10 μM) 2 AA-116: Oligonucleotide reverse (10 μM) 2 pcDNA3.1-HA-FAT10 (400 ng µl-1) 1 DNA-Polymerase (Expand High Fidelity Plus ) 1

H2O 29

Materials and Methods

PCR program: Process Temperature [° C] Duration [s] Repetition Initial denaturation 94 180 1 x Denaturation 94 60 Annealing 60 30 30 Elongation 72 30 Final Elongation 72 480 1 x

5.2.2.2 Site-directed mutagenesis All plasmids containing uba6 of fat10 that had been checked through sequencing contained at least one mutation in their sequence. To remove these mutations QuikChange Site- Directed Mutagenesis Kit (Stratagene) was applied. Primers (Microsynth) were designed that flanked the site of mutation with 10-15 nt overhangs on either side and used in a PCR reaction according to the manufacturer's instructions together with the mutated constructs as templates. Amplification was carried out with 18 cycles of melting (95°C for 30 s), annealing (Tm - 5°C for 60 s) and amplification (68°C for 60 s per 1 kb of DNA). PCR products were subsequently digested with 10 U DpnI for 1-3 h, heat inactivated repaired constructs were transformed into the E. coli strain XL10 Gold and the removal of mutations was verified by sequencing.

5.2.2.3 Agarose gel electrophoresis After digestion of DNA with a restriction enzyme or purification of DNA, agarose gel electrophoresis was performed for both preparative and analytical purposes, using gels with an agarose concentration of 0,8-1,5 % and 0,5 μg ml-1 ethidium bromide in TAE buffer. Samples were mixed with 6x loading buffer (Table 5) and and the mixtures transferred into the slots of the gel. The GeneRuler 1 kb DNA Ladder (see 5.1.1) was used as a DNA marker. For electrophoretic separation 100 V were applied. DNA was visualized by UV and PCR products were extracted and purified from agarose gels using the GenElute Gel Extraction Kit (Sigma-Aldrich) (see 5.1.2).

5.2.2.4 Restriction digest The ability to cleave DNA at specific sites is one of the cornerstones of today’s methods of DNA manipulation. Restriction endonucleases are bacterial enzymes that cleave duplex DNA at specific target sequences with the production of defined fragments.

Materials and Methods

Site specific restriction endonucleases in the appropriate buffers (all Fermentas) were used to generate compatible ends that allow for ligation of DNA fragments into the respective restriction sites of the different vectors. The restriction sites had been included into the primer sequence and DNA fragments as well as the target vectors were digested with the same pairs of restriction enzymes. Digests were carried out using a standard protocol. A typical digest mixture with a final volume of 100 μl contained 5 μg of plasmid DNA or respectively at least 1 μg of PCR product, 10 μl of 10 x restriction buffer and 1-10 U of enzyme. Mixtures were incubated at 37 °C for 2-5 h, according to the enzyme. Prior to ligation, the vector DNA was dephosphorylated to prevent self- or re-ligation for 30 min at 37 °C with the Calf Intestine Alkaline Phosphatase (CIAP), see 5.1.6. Dephosphorylation increases the effective concentration of available ends and efficiency of productive ligations. The reaction was stopped by exposing the enzyme to 85 °C for 20 min. Digested DNA and dephosphorylated DNA was subjected to agarose gel electrophoresis, and bands were visualized by ethidium bromide staining, cut out and subsequently purified from the gel slice using GenElute PCR Clean-Up Kit (Sigma-Aldrich) (see 5.1.2).

5.2.2.5 Ligation Digested DNA fragments were ligated into designated vectors using 1 μl of the T4-DNA ligase and 2 μl of the matching 10x ligase buffer (Fermentas) in a total reaction volume of 20 μl. A 1:5 molar ratio of vector to insert was used whereas either 50 or 100 μg of vector were applied. Ligations were carried out in a thermocycler at 16 °C for 20 h. Afterwards, the T4 ligase was inactivated at 65 °C for 10 min.

5.2.2.6 Preparation of chemically competent E. coli cells A single colony of bacteria (E.coli TOP10 F') was picked from an LB plate and used to inoculate 5 ml LB medium. Bacteria had time to grow overnight in an incubator at 37 °C and 200 rpm. Overnight culture was diluted at least 1/100 to inoculate 100ml LB medium. The bacterial culture was allowed to grow further to an OD600 of 0.5. The culture was cooled down on ice for 30 min before it was harvested through centrifugation at 4 °C with 4000 x g in a Sorval centrifuge for 20 min. Pelleted bacteria were carefully resuspended on ice in sterile

100 mM CaCl2, pelleted again (2200 x g for 30 min, 4 °C) and resuspended in 10 ml of a chilled 10 % glycerol-solution in CaCl2. Aliquots of 100 μl were filled in pre-chilled eppendorf tubes and frozen in liquid nitrogen before use or stored at -80 °C.

Materials and Methods

5.2.2.7 Transformation into E. coli TOP 10 F’ The ligations were transformed into chemically competent E.coli strain TOP10 F'. 100 μl of competent bacteria was thawed on ice and carefully mixed with 10 μl of the ligation mixture and incubated on ice for another 30 min to allow the plasmids to attach to the bacterial surface. Cells were then heat shocked for 40 s at 42 °C in a waterbath and transferred back on ice for 2 min. 400 μl of pre-warmed SOC medium without antibiotic was added and transformed bacteria were shaken for 1 h at 37 °C to allow development of antibiotic resistance, before they were streaked out on LB plates containing the respective antibiotic to select for successfully transformed bacteria.

5.2.3 Expression and purification of recombinant GST-TRIM11 from E. coli

To purify recombinant GST-TRIM11 from E. coli BL21(DE3), 100ml HSG (Table 23) supplemented with Ampicillin (100 μg ml-1) was inoculated with a single colony of bacteria (E. coli BL21(DE3) GST-TRIM11) from LB agar plates including ampicillin and shaken at

37°C overnight. The overnight culture was diluted to a final OD600 of 0,1 in a total volume of

800 ml HSG and grown to exponential phase (OD600 2~3) followed by induction of the lac operon with 0,1 mM IPTG for 4 hours at 20 °C. Cells were harvested in a Sorvall RC6 Plus centrifuge (Thermo), at 18.000 rpm and 4 °C for 15min and pellets were resuspended in lysis buffer (Table 15) containing the protease inhibitors 0,1 mM phenylmethylsulfonyl fluoride (PMSF), 2 μg ml-1 aprotinin, 2 μg ml-1 leupeptin and 2 μg ml-1 pepstatin and disrupted 2 times in French press (2,5 kBar). Cell debris was removed by centrifugation at 18.000 rpm for 30 min at 4 °C. The supernatant containing the GST fusion protein was filtered through a 0,2 μm Filter and loaded onto a 1 ml Glutathione-SepharoseTM 4B column (GE Healthcare), with a flow rate of 1 ml min-1 using AKTAexplorer purification systems from GE Healthcare, as recommended by the supplier. 30 column volumes of PBS and GST high-salt washing buffer (Table 16) were used to wash unspecifically bound proteins off the column. GST- TRIM11 was eluted with 5 column volumes of GST-elution buffer (Table 19) containing 10 mM reduced Glutathione and subsequently dialyzed against PBS, concentrated and stored at 4 °C.

5.2.4 Cell culture

HEK293, HEK293T and HELA cells were maintained on Dulbecco´s Modified Eagle Medium (DMEM) supplemented with 10 % fetal calf serum and 100 U ml-1 penicillin, and 100 μg ml-1 streptomycin (Lonza) at 37 °C and 5 % CO2 environment.

Materials and Methods

The T-cell line Jurkat was maintained in RPMI 1640 medium supplemented with 10 % fetal calf serum, 100 U ml-1 penicillin, 100 μg ml-1 streptomycin (Lonza), 0,1mM NEAA (Invitrogen) and 50 μM ß-mercaptoethanol, at 37 °C and 5 % CO2 environment.

5.2.5 Transient transfection

HEK293, HEK293T, HELA Wildtype HEK293, HEK293T or HELA cells were cultured in DMEM supplemented with 10 % FCS and 1 % Penicillin/Streptomycin until they reached 90 % confluency. HEK293 or HELA cells were then washed once with PBS before Trypsin/EDTA was added to detach adherent cells from cell culture flasks. Trypsin was inactivated by adding cell culture medium containing 10 % FCS. The cell suspension was centrifuged for 4 min at 1250 rpm and cell pellets were resuspended in medium. The number of cells was determined using an Improved Neubauer chamber. Cells were seeded at either 1.5x106 cells/dish (10 cm2 cell culture dish) or 6x106 cells/dish (14,5 cm2 cell culture dish) and cultured for about 20h until 60-80 % confluency was reached. For transient transfection, FuGene6 (Roche) or TransIT- LT1 transfection reagent (Mirus) was used. 436,5 μl (10 cm2 cell culture dish) or 776 μl (14,5 cm2 cell culture dish) of serum and antibiotic free DMEM was mixed by vortexing with 13.5 μl Transfection Reagent (for 10 cm2 cell culture dish) or 24 μl (for 14,5 cm2 cell culture dish) and incubated at room temperature for 5 min. 4,5 μg (10 cm2 cell culture dish) or 8 μg (14,5 cm2 cell culture dish) of DNA was added, components were mixed again and after further 15 min of incubation at room temperature allocated dropwise onto the cells. 20-24 h later cells were harvested and proteins were subjected to further analysis.

JURKAT cells Jurkat (ATCC #TIB-152) cells were grown in RPMI-1640 medium containing 10 % FCS, 1 % Penicillin/Streptomycin, 0,1 mM NEAA, 50 μM ß-mercaptoethanol (referred as complete RPMI medium) and sub-cultured 24 h prior to electroporation. Cell suspension was centrifuged for 8 min at 90 x g and cell pellets were washed in pre-warmed PBS. After a second centrifugation step, cells were resuspended in 1 ml RPMI medium without complements, cells were counted and cell density was determined using an Improved Neubauer chamber. 15 million Jurkat cells (in 400 μl medium) per approach in logarithmic growth phase were transfected with the indicated amounts of plasmid by electroporation using the Gene Pulser MXcell electroporation system. Cells in each experiment were transfected with the same total amount of DNA by adding the required quantities of empty vector.

Materials and Methods

Cells were incubated for 10 min at room temperature with the DNA mix and transferred to Nucleocuvette™ Vessels (0,4 μM) and electroporated immediately at 260 mV, 960 mF in 400 μl of RPMI 1640. Electroporated cells were resuspended in 500 μl pre-warmed complete RPMI and transferred to 6-well plates, containing 1,5 ml pre-warmed complete RPMI medium per well. Cells were incubated for 48 h in a humidified 37 °C incubator with 5 % CO2 until analysis.

5.2.6 SDS-PAGE (Sodium dodecylsulphate polyacrylamide gel electrophoresis)

To separate proteins according to their size, the NuPAGE system from Invitrogen was used (see 5.1.1). Samples were prepared by mixing 4 volumes of sample with 1 volume of 5 x SDS Sample buffer either with or without 10 % ß-mercaptoethanol. Samples were boiled for 5 min at 95 °C before being loaded onto gradient NuPAGE® 4-12 % Bis-Tris gels together with the SeeBlue® Plus2 PreStained Protein Standard (Invitrogen) or Precision Plus Protein Prestained Standard (BioRad). Gels were run with NuPAGE® SDS MES running buffer at a constant current of 100 mA per gel. During electrophoresis in gradient gels, proteins migrate until the decreasing pore size impedes further progress. Once the “pore limit” is reached, the protein banding pattern does not change appreciably with time, although migration does not cease completely. To detect proteins of high molecular weights, samples were loaded on gradient NuPAGE® Novex 3-8 % Tris-Acetate Gels together with HiMark™ Pre-stained Protein Standard. Gels were run with Novex® Tris-Acetate SDS Running Buffer at a constant current of 80 mA per gel. Following electrophoresis cell lysates were subjected to Western blotting (see below).

5.2.7 Immunoblotting

Proteins resolved by SDS-PAGE were transferred onto Protran nitrocellulose membrane (Whatman) using the Criterion Blotting system from BioRad, according to the manufacturer's instruction. The transfer was conducted in Towbin buffer supplemented with 20 % methanol at 110 mA and 110 V for 50 min. Afterwards membranes were blocked using RotiBlock solution (Roth) for 1 h at room temperature or overnight at 4 °C on a rotating platform (50- 100 rpm). Subsequently, membranes were hybridized with the primary antibody diluted in

3 % BSA/PBS-T (PBS containing 0,02% Tween 20) supplemented with 0.02 % NaN3 for 1.5 h at room temperature or overnight at 4 °C with rolling, followed by extensive washing of the membrane with PBS-T (3-5 times, 10 min at RT).

Materials and Methods

Afterwards, blots were incubated for 1 h with the respective secondary HRP-coupled antibody which was being diluted in milk powder (1:5000/PBS-T). Membranes were washed 3 times again with PBS-T before the proteins recognized by the employed antibodies were detected by chemiluminescence (Super Signal Chemiluminescent, Thermo Scientific) according to the manufacturer's protocol.

5.2.8 Immunoprecipitation (IP)

HEK293, HELA HEK293 and HELA cells were transiently transfected as described above. After 24 h, cells were harvested, resuspended in 1ml (per 10 cm2 cell culture dish) of FAT10-IP buffer (Table 4) containing 1 % NP40, supplemented with Complete Mini Protease-Inhibitor Cocktail (Roche) and incubated on ice for 10 min. After sonification (2 cycles for 20 sec each) cell lysates were vortexed for 30 s and centrifuged for 30 min at 14000 rpm and 4 °C.

293T 293T cells were grown to about 80 % confluency, washed with ice-cold PBS, harvested by scraping and lysed on ice for 30 min in BRUCE-IP buffer (see Table 14) containing 1 mM PMSF and Complete Protease inhibitors (Roche). Cellular debris was removed by centrifugation at 10000 x g for 10 min and 40 μl of the supernatant was mixed with 10 μl of 5 x SDS sample buffer, boiled and used in SDS-PAGE and subsequent Western blot analysis to show protein amounts in the total lysate. To the remaining supernatant, antibody- coupled beads or specific antibodies together with protein-A-sepharose (see 5.1.5) were added and incubated at 4 °C for at least 2 h with rolling. Amounts of beads (all obtained from Sigma-Aldrich) were the following:

 Monoclonal anti-HA-Agarose (clone HA-7): 20 μl  EZview™ Red ANTI-FLAG M2 Affinity Gel: 20 μl  Protein-A-Sepharose: 40 μl+2 μl anti-FAT10 (4,4 mg ml-1) (4F1) monoclonal antibody (see Table 35)

Antibody-coupled agarose or protein-A-sepharose was pelleted and washed 3 times with NET-T before 5x SDS sample buffer was added and samples were boiled for 5 min at 95°C. Subsequent protein analysis was carried out by subjecting the samples to SDS-PAGE and Western blotting as already described above.

Materials and Methods

5.2.9 In vitro FAT10ylation assay of JunB

Cells were harvested at 24 h after transfection, washed with cold PBS, and centrifugated at 1500 rpm for 5 min at 4 °C. Cell pellets were resuspended in FAT10 IP buffer (see Table 13) including 1 % NP-40 + protease inhibitor pill (Roche) and sonified. Cell debris was removed by centrifugation at 14000 rpm for 30 min at 4 °C. Anti-FLAG-M2-conjugated agarose beads (Sigma-Aldrich) were added to the cleared lysate to immunoprecipitate JunB and the mixture was rotated overnight at 4 °C. The anti-FLAG-M2 agarose beads were collected and washed three times with FAT10 IP buffer (see Table 13) containing complete protease inhibitor mix and recombinant FAT10, FLAG-UBA6 (E1), HIS-USE1 (E2) and GST-TRIM11 (E3) was added to a final volume of 500 μl with ATP containing FAT10-IP buffer (+ protease inhibitor pill). The mixture was rotated for 2-4 h at 4 °C. Beads were washed twice with NET-TON buffer (see Table 17) and three times with NET-T buffer (Table 18) prior to boiling in SDS sample buffer with 10% β-mercaptoethanol. Samples were separated on 4-12 % BIS-Tris SDS gels analyzed by Western blot probed with anti-HA Peroxidase conjugate, anti- FLAG, anti-FAT10, anti-HIS and anti-GST antibody.

5.2.10 Growth of yeast strains

The yeast strain NMY51 was grown in either liquid medium (30 °C, with shaking for one day) or on the surface of a solid agar plate for 4 days at 30 °C. Yeast cells can grow on a minimal medium containing dextrose (glucose) as a carbon source and salts that supply nitrogen, phosphorus, and trace metals, also known as synthetic defined (SD) medium. SD dropout medium lacks a single (or several) nutrient that allows selection for maintenance of particular plasmids or selection for induction or repression of specific gene promoters (Table 26). Yeast cells grow much more rapidly in the presence of rich medium, for example yeast extract, peptone, adenine, dextrose (YPAD) is most commonly used for growing yeast under nonselective conditions (Table 24). These provide many of the metabolites that the cells would synthesize when growing under minimal growth conditions.

5.2.11 DNA isolation from yeast with glass beads

To isolate DNA from yeast, an overnight culture of 5 ml was harvested and pellets were resuspended in 100 µl STET buffer (Table 34) and vortexed briefly. 100 µl of acid washed glass beads (0,45 mm) were added and vortexed for 5 min at room temperature and another 100 µl of STET buffer was added and vortexed briefly.

Materials and Methods

Then, tubes were placed in a heat block at 100 °C for 5 min and subsequently cooled on ice for 3 min. After a short centrifugation step, supernatant was transferred to a 1,5 ml Eppendorf tube and centrifuged for 15 min with maximum speed at 4 °C. 100 µl of supernatant was transferred to a fresh Eppendorf-tube and 500 µl of 4 °C Ammonium-acetate (7,5 M) was added and mixed briefly. The tubes were stored overnight at -20 °C. 500 µl of supernatant was transferred to Eppendorf-tubes containing 1 ml ice-cold EtOH, mixed shortly and let stand for 5 min at room temperature. After centrifugation for 10 min with maximum speed at room temperature, the supernatant was discarded and the pellet was washed with 400 µl of 70 % EtOH. The reaction mix was centrifuged for 5 min with maximum speed. Supernatant was removed and pellet was dried at room temperature. The pellet was resuspended in 20 µl of TE (see Table 6) and 10 µl was used to transform competent bacteria.

5.2.12 Yeast two-hybrid assay

The yeast-2-hybrid system is a powerful in situ screening system to identify and characterize novel protein-protein interactions of cytosolic proteins (Fields and Song, 1989). The great benefit of an in situ system is that interactions are identified in intact cells, and that no further optimization or washing steps are needed. In this assay, the bait and the prey protein are fused to two parts of a transcription factor, the DNA binding domain (DBD) that mediates binding of the transcription factor to gene promoters by sequence specific DNA recognition and the activation domain (AD) that recruits the transcriptional apparatus to the gene for mRNA production (Coates and Hall, 2003). If bait and prey are co-expressed in the same yeast cell, a functional transcription factor is reconstituted upon association of the target protein with its binding partner, which can translocate to the nucleus and activate distinct downstream reporter genes. Hence, a positive interaction is often measurable by growth under selective conditions or a color signal, resulting from the enzymatic activity of β-galactosidase. By using a cDNA-library novel protein interactions can be discovered easily (Figure 13).

Materials and Methods

Figure 13: Schematic model of the yeast-two-hybrid approach. A target protein fused to the DNA binding domain of a yeast transcription factor is co-transfected with a prey consisting of the activation domain (AD) and a putative binding partner of the target protein. Upon association of the target protein with its binding partner the transcription factor is reconstituted and activates the transcription of reporter genes.

To screen for specific interacting proteins full-length uba6 (this work) and trim11 (provided by A. Aichem) cDNA were cloned as N-terminal fusion proteins to the lexA DNA-Binding Domain (DBD), into the pLexA-N plasmid (Dualsystems Biotech, Zurich, Switzerland) (see 5.1.8), which contains a TRP1 gene to enable selection in tryptophane (Trp) auxotrophic yeast strains. These baits were used to screen a human thymus cDNA library (Thymus MATCHMAKER cDNA library, Clontech), containing the GAL4 AD fused to a protein encoded by the cDNA in a fusion library. The hybrid protein is expressed in yeast host cells from the enhanced, truncated ADH1 promoter and is targeted to the yeast nucleus by the SV40 T-antigen nuclear localization sequence. Moreover, the pACT prey plasmid contains the LEU2 gene for selection in leucine (leu) auxotrophic yeast strains. The bait construct was transformed into the yeast strain NMY51 (MATa his3Δ200 trp1-901 leu2-3,112 LYS2::(lexAop)4-HIS3 ura3::(lexAop)8-lacZ (lexAop)8-ADE2 GAL4 Dualsystems Biotech, Zurich, Switzerland)) with the YEASTMAKER lithium acetate method and the absence of self-activation was tested with a X-Gal filter assay on selection plates lacking Trp. Whatman filters, which were used to make replicates of the plates, were frozen in liquid nitrogen, subsequently soaked with PBS containing 100 μg ml-1 X-Gal (Table 33) and incubated at room temperature. For the yeast two hybrid screen, the human thymus cDNA library was transformed into the yeast strain NMY51 expressing the bait, as described by the DUALhybrid kit instructions.

Materials and Methods

In addition to growth on selection plates lacking Trp, Leu and His yeast NMY51 cells have a further reporter gene named ADE2 which is not transcribed in absence of a positive protein- protein interaction, resulting in red colored yeast colonies. In case of a positive protein- protein interaction the gene is transcribed and the color of the cells turns from red to faint pink or even to white, depending on the strength of the interaction. Thus white colonies indicate a stronger protein-protein interaction than faint pink colonies. Yeast cells were grown for 4 days on selection plates lacking Trp, Leu and His supplemented with 3-amino-1,2,4- triazole (3-AT). Interaction of proteins was further tested with a X-Gal filter assay. Clones that turned blue were grown on new selection plates and exposed to a second X-Gal test. The prey plasmids of the positive interacting clones were isolated using a standard method with glass beads (see 5.2.11) and were re-transformed into NMY51 together with either the bait plasmid, or a non cognate plasmid pLexA-LaminC or pLexA-p53, respectively, to test for bait- independency. Prey plasmids from positive clones were isolated using a standard method with glass beads (see 5.2.11) and retransformed in E. coli Top10 F´-cells for amplification. A restriction digest with BglII was performed, to identify equal prey plasmids. The identity of the remaining clones was determined by sequencing.

5.2.13 Luciferase reporter assays

A mixture of an inducible transcription factor responsive firefly luciferase reporter (AP-1) and constitutively expressing Renilla (pRl-Tk-Basic) construct was used together with constructs encoding for FAT10, FAT10∆GG, JunB and JunB-K3R to transiently transfect HEK293 cells or Jurkat cells. After 24 h (HEK293 cells) or 48 h (Jurkat cells) of ectopic protein expression, cells were washed once with PBS and cell lysates were prepared with 1x passive lysis buffer, provided together with the Dual-Luciferase® Reporter (DLR™) Assay System (see 5.1.2), according to the manufacturer’s instructions. In the DLR™ Assay, the activities of firefly (Photinus pyralis) and Renilla (Renilla reniformis) luciferases are measured sequentially from a single sample. The inducible transcription factor responsive construct AP-1 (4xTRE) encodes the firefly luciferase reporter gene under the control of a minimal promoter element (TATA box) joined to tandem repeats of a specific transcriptional response element (4xTRE). This construct monitors both, increases and decreases in the activity of a key transcription factor (in this case JunB), which is a downstream target of a specific signaling pathway. The firefly luciferase reporter is measured first by adding Luciferase Assay Reagent II (LAR II) to generate a stabilized luminescent signal. After quantifying the firefly luminescence, this reaction is quenched, and the Renilla luciferase reaction is simultaneously initiated by adding Stop & Glo® Reagent to the same tube.

Materials and Methods

The Renilla luciferase reporter is constitutively expressed and acts as an internal control for normalizing transfection efficiencies and monitoring cell viability. It is also useful to confirm transfection and to verify active luciferase in the transfected cultures. The pGL3 plasmid, encoding for firefly luciferase was used to determine basal activity. All samples were measured using a Berthold Lumat LB9501.

5.2.14 Immunofluorescence and confocal microscopy

Transiently transfected HEK293 or HELA cells were plated on coverslips with a density of about 30-40 %. 20 h post-transfection, cells were washed carefully with cold PBS and fixed with 4 % paraformaldehyd. To block reactive aldehyde groups, coverslips were incubated with 50mM NH4Cl for 10 minutes at 4°C. Cells were permeabilised with 0,2 % Triton in PBS for 5 minutes at room temperature. After a further 30 minutes blocking step with 3 % BSA/PBS the microscopic preparations were incubated with the indicated antibodies over night at 4°C or 1 hour at room temperature. Following intense washing with PBS the secondary antibody was added. Afterwards coverslips were mounted in VECTASHIELD® or polyvinyl alcohol mounting Media and analyzed by confocal imaging using a 63x oil objective on a confocal microscope (Leica TCS SP5 II) at room temperature. Images were generated using pinhole size corresponding to 1 µm layers for each channel. Contrast and brightness of pictures were simultaneously adjusted with Adobe Photoshop CS2.

Results

6 Results

6.1 Yeast two-hybrid screen with UBA6

The initial activation of ubiquitin was thought for decades to be accomplished by a single enzyme designated UBE1 (Ciechanover et al., 1981; Haas et al., 1982). Surprisingly, in 2007 a second E1 type enzyme, called UBA6 has been identified, which can activate ubiquitin as well as FAT10 (Chiu et al., 2007; Jin et al., 2007; Pelzer et al., 2007). UBA6 cooperates partially with the same set of E2 enzymes and one E2 enzyme, the UBA6-specific E2 enzyme 1 (USE1) has been described which could accept ubiquitin only from UBA6 but not from UBE1 (Jin et al., 2007). One aim of this project was to identify new protein-interaction partners of UBA6 and putative ubiquitin-like modifier / E2 enzymes that rely on UBA6 for their supply of activated ubiquitin or FAT10. Given that UBE1 and UBA6 are co-expressed in many tissues (although UBE1 is up to 10 fold more abundant than UBA6), these two enzymes may act in concert or in sequence to affect various signaling pathways (Groettrup et al., 2008). One possibility is that UBE1 and UBA6 might use a different spectrum of E2-enzymes and eventually different E3- enzymes with their corresponding substrates. Regarding the fact, that UBA6 is so far the only known FAT10 activating enzyme it could be possible that new identified partners of UBA6 can give a hint for novel E1-E2-E3 cascades for this ULM. At the beginning of this doctoral thesis, no FAT10 specific conjugating E2 enzyme had been identified yet. In 2010 Aichem et al. (Aichem et al., 2010) could show, that USE1 is a bispecific E2 conjugating enzyme for ubiquitin as well as FAT10 and moreover, the first identified FAT10 specific substrate, which FAT10ylates itself in cis. To gain more insight about the FAT10 conjugation pathway and to identify further E2 enzymes for FAT10, a yeast two-hybrid screen with human UBA6 serving as bait for screening a human thymus cDNA library, was performed. The human uba6 gene was cloned into the bait vector pLexA-N as an N-terminal fusion to the lexA domain. Given that UBA6 interacts with FAT10, the full length fat10 gene was cloned into the prey vector pACT2, which served as a positive control. Absence of self-activation of the yeast two-hybrid constructs was positively tested on selection plates lacking tryptophane (SD-T) or leucine (SD-L), respectively, because expression of UBA6 or FAT10 alone did not result in ADE2 activation, resulting in redish colored colonies (Figure 14 (a), left panel), nor lacZ activation (Figure 14 (a), right panel), which was tested by a (X-Gal) filter assay.

Results

The screening stringency was optimized and determined during a pilot screen to eliminate most false positive interaction partners. To reduce the background, 1 mM 3-aminotriazole (3- AT) was included in the medium throughout the screen. A large-scale yeast two-hybrid screen uncovered a total of 114 clones, which were able to grow on selection plates lacking His, Leu and Trp, indicating already a positive protein-protein interaction. A further control for positive protein–protein interaction was the color change of yeast colonies from reddish to white, indicating that the ADE2 gene in yeast was sussessfully transcribed. Moreover, the interaction between the proteins described was positively tested in a X-Gal filter assay and judged from the color change on selective growth plates. Clones that turned blue in the X- Gal-filter assay were grown on new selection plates and X-Gal filter assay was repeated four times. The prey plasmids of 114 clones were isolated using a standard method with glass beads (see 5.2.11) and retransformed into NMY51, together with either pLexA-UBA6, to re- confirm the positive interactions of the screen or together with pLexA-LaminC or pLexA-p53, to test for bait independency. The identity of the remaining 104 clones was determined by sequencing.

Figure 14: UBA6 interacts specifically with BRUCE in a yeast two-hybrid screen. (a) Yeast NMY51 was transformed with the bait construct pLexA-UBA6 and grown on selection plates lacking tryptophane (left panel). Absence of self-activation was tested with a X-GAL filter assay (right panel). (b) Yeast NMY51 was transformed with the bait construct pLexA-FAT10 and grown on selection plates lacking leucine (left panel). Absence of self-activation was tested with an X-GAL filter assay (right panel).

Results

(c) Yeast NMY51 was co-transformed with pLexA-UBA6 and pACT2-FAT10 (positive control), or with pLexA- UBA6 and and the isolated pACT2-BRUCE (C-terminus) constructs and grown on selection plates containing 1 mM 3-AT and lacking tryptophane, leucine and histidine (left panel). An X-Gal filter assay was performed to verify the interaction of the respective proteins (right panels). A representative experiment from a total of three experiments with similar outcome is shown. (d) Yeast NMY51 was co-transformed with pLexA-FAT10 and pACT2-BRUCE-C-terminus and grown on selection plates containing 1 mM 3-AT and lacking tryptophane, leucine and histidine (left panel). An X-Gal filter assay was performed to verify the interaction of the respective proteins (right panel).

Among 104 proteins, which positively interacted with UBA6, sequencing of the plasmids revealed, that 69 plasmids encoded for full length fat10 and 3 Plasmids encoded for the C- terminal truncation of bruce, containing the last ~4000 nucleotides of the bruce gene. Interestingly, each of the three independent clones contained the entire C-terminal ubiquitin conjugating (UBC) domain, which is essential for ubiquitin conjugating E2 enzymes. Apart from being able to grow on -TLH selection plates, already indicating the interaction of the described proteins, cells turned blue in the X-Gal filter assay (Figure 14 (c), right panel) and showed the same white color as it can be seen in the positive control (UBA6 and FAT10) (Figure 14 (c), left panel). To further confirm the interaction of BRUCE and FAT10, pLexA- FAT10 was co-transformed together with the isolated prey pACT2-BRUCE plasmid containing the C-terminal half of BRUCE into the yeast strain NMY51. Growing on selective growth plates, lacking Trp, Leu and His resulted in colonies with a faint pink color (Figure 14 (d), left panel), which gives a hint, that a weak interaction indeed occurs between these proteins. The weak interaction could be also confirmed in a X-Gal-filter assay (Figure 14 (d), right panel). An overview of positively interacting proteins in the yeast two-hybrid assay with UBA6 is depicted in Table 41. .

Table 41: Classification of positively interacting proteins in yeast two-hybrid screen with UBA6

Nr. Proteins Frequency

1 FAT10 69 2 Ubiquitin C 1 3 Receptor proteins 10 4 E2 like enzymes 4 5 Proteins with zinc finger motif 1 6 Ribonucleoproteins 1 7 Protein kinases 2 8 Hypothetical proteins 16

Among the proteins which could positively interact with UBA6, the following proteins were positively tested for their interaction with FAT10 in a yeast two-hybrid assay (see Figure 15).

Results

Yeast NMY51 was co-transformed with pLexA-FAT10 together with the prey plasmids pACT2-PLZF, pACT2-Splicing factor, arg/ser-rich, pACT2-JMJD6 or pACT2-F-actin-capping protein beta SU, and grown on selection plates containing 1 mM 3-AT and lacking Trp, Leu and His (Figure 15 , left panel). An X-Gal filter assay was performed to verify the interaction of the respective proteins (Figure 15, right panels).

Figure 15: FAT10 interacts with some of the identified proteins from the UBA6 yeast two-hybrid screen. Yeast NMY51 was co-transformed with pLexA-FAT10 and pACT2-PLZF, pACT2-Splicing factor, arg/ser-rich, pACT2-JJMJD6 or pACT2- F-actin-capping protein beta SU, which where isolated from the UBA6 yeast two- hybrid screen and grown on selection plates containing 1 mM 3-AT and lacking tryptophane, leucine and histidine (left panel). An X-Gal filter assay was performed to verify the interaction of the respective proteins (right panels).

Interestingly, many proteins identified in the yeast two-hybrid screen with UBA6 as well as in the additional yeast two-hybrid interaction assay with FAT10 as a bait protein, are involved in cell cycle regulation.

6.1.1 BRUCE interacts non-covalently with UBA6 and FAT10

We decided, that the huge membrane- associated and evolutionary highly conserved protein BRUCE (528 kDa) is a very interesting protein to investigate, as it contains a catalytic ubiquitin conjugating enzyme (UBC) domain at its C-terminus, and a BIR (baculovirus inhibitor of apoptosis repeat) motif. BRUCE has been previously described to act as a chimeric ubiquitin E2/E3 ligase with Smac being a substrate (Bartke et al., 2004). Furthermore, during final stages of cytokinesis BRUCE moves from the vesicular system to the midbody ring and serves as a platform for the membrane delivery machinery and mitotic regulators, thereby ensuring controlled abscission (Pohl and Jentsch, 2008).

Results

Moreover, due to its N-terminal BIR domain, which are found within inhibitor of apoptosis proteins (IAP), it can further act as an antiapoptic IAP (Bartke et al., 2004). Therefore, BRUCE may function to mediate both, ubiquitin-dependent proteolysis and contribute to anti-apoptotic cellular pathways.

To verify the yeast-two-hybrid findings and confirm the interaction with UBA6, we examined the ability of human UBA6 to interact with the human native full-length BRUCE (kindly provided from S. Jentsch) in human cells. The putative association of UBA6 with BRUCE might argue for an UBE1- independent loading of BRUCE with either ubiquitin of FAT10 by this newly identified ortholog. Hence, we tested the possibility of interaction between BRUCE and either ubiquitin or FAT10. FLAG-tagged BRUCE together with HA-tagged FAT10, -Ubiquitin and -UBA6 were co- expressed in HEK293T cells. After 24 h ectopic protein expression, cells were washed twice in ice-cold PBS and scraped on ice from cell culture dishes. Cells were instantly lysed in ice- cold BRUCE-IP buffer (Table 14) containing complete protease inhibitor mix and lysed on ice for 30 min. Whole cell lysate were boiled without ß-mercaptoethanol (non-reducing conditions) to preserve thioester linkages or in Laemmli buffer containing 10% ß- mercaptoethanol (reducing conditions), to cleave thioester bonds. Samples were separated on 4-12 % Bis-TRIS SDS-gels and directly immunoblotted with a anti-HA-antibody to analyze expression of HA-tagged ubiquitin, FAT10 or UBA6. Following immunoprecipitation with anti- FLAG-M2-conjugated agarose against the FLAG-tag of BRUCE, samples were boiled in Laemmli buffer without or with 10 % ß-mercaptoethanol and subjected to SDS-PAGE and Western blot analysis using a anti-HA antibody.

Results

(a)

(b)

Results

Figure 16: UBA6, ubiquitin and FAT10 co-immunoprecipitate with full length BRUCE in HEK293 cells. (a) 293T cells were transiently transfected with FLAG-BRUCE, HA-UBA6, HA-FAT10 and HA-Ubiquitin as indicated. (a) After 24 h ectopic protein expression whole cell lysates were boiled in Laemmli buffer without (non- reducing conditions) or with 10 % ß-mercaptoethanol (reducing conditions) and separated on 4-12 % Bis-Tris gels and analyzed by Western blotting with a HA-reactive monoclonal antibody. (b) After immunoprecipitation against the FLAG-tag of BRUCE with anti-FLAG-M2-conjugated agarose, samples were boiled in Laemmli buffer without (non-reducing conditions) or with 10 % ß-mercaptoethanol (reducing conditions) and separated on 4-12% Bis-Tris SDS gels and subjected to Western blot analysis with a HA-reactive monoclonal antibody.

Expression of HA-tagged UBA6 (~110 kDa), FAT10 (18 kDa), ubiquitin (8 kDa) and ubiquitin conjugated proteins could be detected under non-reducing as well as under reducing conditions (Figure 16 (a)). After immunoprecipitation against the FLAG-tag of FLAG-BRUCE, and Western blot analysis with a HA-reactive antibody an upcoming ubiquitin ladder was visible under non-reducing and reducing conditions, suggesting that BRUCE interacts with ubiquitin or ubiquitin conjugates (Figure 16 (b), lane 3+4). In contrast, a non-covalent interaction of FLAG-BRUCE and HA-UBA6 as well as with HA-FAT10 could be detected by Western blot analysis under reducing conditions (Figure 16 (b) lane 6, 7 and 9), whereas under non-reducing conditions only a slight band was apparent at the height of UBA6 and almost no FAT10 was detectable, suggesting that BRUCE interacts with UBA6 and FAT10 via a thioesterbond linkage.

To detect conjugation bands at the height of 500-600 kDa 3-8 % Tris-Acetate gels for large protein separation were used.

(a)

Results

(b)

Figure 17: FAT10 becomes thioester-linked to full length BRUCE in HEK293 cells. (a) 293T cells were transiently transfected with FLAG-BRUCE, HA-UBA6, HA-FAT10 and HA-Ubiquitin as indicated. (a) After 24 h ectopic protein expression immunoprecipitation was performed with anti-FLAG-M2 affinity gel. Samples were boiled in Laemmli buffer without (non-reducing) or with 10 % ß-mercaptoethanol (reducing) and were separated on 3-8 % Tris-Acetate gels followed by Western blot analysis using a FLAG-reactive antibody to reveal FLAG-BRUCE expression. (b) After immunoprecipitation against the FLAG-tag of BRUCE with anti- FLAG-M2-conjugated agarose, samples were boiled in Laemmli buffer without (non-reducing conditions) or with 10 % ß-mercaptoethanol (reducing conditions) and separated on 4-12% Bis-Tris SDS gels and subjected to Western blot analysis with a HA-reactive monoclonal antibody. Arrow head indicates thioester-linked BRUCE conjugates.

FLAG-BRUCE expression could be detected at a molecular weight of about 530 kDa (Figure 17 (a)). Interestingly, after immunoprecipitation against the FLAG-tag of BRUCE and immunoblotting with a FLAG-reactive antibody, double bands at the height of BRUCE appear under non-reducing (Figure 17 (a), arrow head), but not under reducing conditions (Figure 17 (a), reducing) which indicates that BRUCE becomes modified via a thioester-linkage to FAT10. Moreover, at the size of BRUCE (~530 kDA) very weak conjugate bands were observable after immunoprecipitation against the FLAG-tag of BRUCE and Western blot analysis with a HA-reactive monoclonal antibody, when FLAG-BRUCE and HA-UBA6, FLAG- BRUCE and HA-FAT10 or even when all three proteins were overexpressed under non- reducing conditions (Figure 17 (b), lane 6,7 and 9). Due to the very faint band appearing, further experiments are required to ensure that BRUCE gets modified with FAT10 via a thioester-linkage.

Results

The conjugate bands almost completely disappeared under reducing (10% ß- mercaptoethanol) conditions, which additionally argue for a thioester-bond between the respective proteins. Besides, an increased amount of co-immunoprecipitated HA-UBA6 is detectable under reducing condition, which strengthens the hypothesis that BRUCE might become thioester-linked to UBA6 (Figure 17 (b), lane 6+7).

6.1.2 Endogenous FAT10 co-immunoprecipitates with BRUCE

To further investigate, if BRUCE can become thioester-linked to FAT10, when FAT10 is induced with the pro-inflammatory cytokines TNF-α and IFN-γ, a semi-endogenous co- immunoprecipitation assay with FLAG-BRUCE, HA-UBA6 and endogenous FAT10 was performed. Therefore, HEK293 cells were transfected with constructs expressing FLAG-tagged BRUCE or HA-tagged UBA6 and cells were treated with TNF-α (400 U ml-1) and IFN-γ (200 U ml-1) to induce the upregulation of endogenous FAT10. 24 h later, cells were lysed, and a part of the lysates with 10 % ß-mercaptoethanol (reducing conditions). Samples were separated on SDS-PAGEs and proteins were analyzed via Western blot with a HA-reactive antibody to reveal HA-UBA6 expression, a monoclonal FAT10 (4F1) antibody to determine FAT10 expression and a FLAG-reactive antibody, to detect FLAG-BRUCE expression (see Figure 18, input). The remaining cell lysate was used to conduct co-immunoprecipitation assays with anti-FLAG M2 affinity gel, to immunoprecipitate FLAG-tagged BRUCE. Samples were boiled in Laemmli buffer containing ß-mercaptoethanol (10 %) or without ß-mercaptoethanol followed by SDS-PAGE and Western blot analysis using anti-HA, anti-FAT10 or anti-FLAG antibodies (Figure 18 (b), IP: anti-FLAG).

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Results

(b)

Figure 18: BRUCE interacts non-covalently with endogenous FAT10

(a) 293T cells were transiently transfected with FLAG-BRUCE or HA-UBA6, as indicated. Endogenous FAT10 was upregulated through addition of the proinflammatory cytokines IFN-γ (200 U ml-1) and TNF-α (400 U ml-1) (a) After 24 h ectopic protein expression, cells were lysed in BRUCE-IP buffer containing whole protease inhibitor mix and samples were boiled in Laemmli buffer containing 10 % ß-mercaptoethanol. Samples were separated on 4- 12 % Bis- Tris or 3-8 % Tris-Acetate gels followed by Western blot analysis using anti-HA, anti-FAT10 (4F1) or anti-FLAG-reactive antibodies. (b) After immunoprecipitation against the FLAG-tag of BRUCE with anti-FLAG-M2- conjugated agarose, samples were boiled in Laemmli buffer without (non-reducing conditions) or with 10 % ß- mercaptoethanol (reducing conditions). Samples were separated on 4-12% Bis-Tris or 3-8% Tris-Acetate gels SDS gels and subjected to Western blot analysis with anti-HA, anti-FAT10 (4F1) and anti-FLAG reactive antibodies. One experiment out of two is shown.

Figure 18 (a), illustrates that all proteins were expressed. Interestingly, an increase of HA- UBA6 expression could be observed, when co-expressed together with BRUCE. Moreover, the UBA6 protein amount was even higher, when cells were treated additionally with TNF-α (400 U ml-1) and IFN-γ (200 U ml-1) to induce FAT10. Immunoprecipitation assays revealed, that HA-UBA6 becomes co-immunoprecipitated with BRUCE, as previously shown and the amount of co-immunoprecipitated HA-UBA6 decreased under non-reducing conditions (Figure 18 (b), WB: anti-HA, lane 4+5 and 8+9), which strongly suggest, that both proteins become thioester-linked to each other. Interestingly, endogenous FAT10 becomes co-immunoprecipitated with BRUCE, but no difference on protein level is visible under reducing or non-reducing conditions (Figure 18 (b), WB: anti-FAT10, lane 5+7 and 9+11). No clear conjugate bands were observable under reducing nor under non-reducing conditions, after immunoprecipitation against the FLAG-tag of BRUCE and Western blot analysis with a FLAG-reactive monoclonal antibody, when FLAG-BRUCE was co-expressed with either HA-UBA6, endogenous FAT10 or both together (Figure 18 (b), WB: anti-FLAG).

Results

Moreover, no conjugate band at the height of ~530 kDa was observable after immunoprecipitation against the FLAG-tag of BRUCE and immunoblotting with a HA- reactive, or FAT10 monoclonal antibody, neither under reducing nor under non-reducing conditions (data not shown). This let suggest, that either the conjugate is not very stable or detection limit is reached due to a lower sensitivity of the monoclonal FAT10-antibody compared to the anti-HA-antibody used in the previous experiment, because also the anti-HA antibody detected only a very faint conjugate band under non-reducing conditions (see Figure 17 (b), IP: anti-FLAG, WB: anti-HA)).

To further investigate, if activated FAT10 can be transferred from UBA6 onto BRUCE we conducted in vitro experiments with recombinant proteins. Unfortunately, in vitro experiments failed, and no transfer of FAT10 on BRUCE could be detected in presence of UBA6 (E1), FAT10 and the putative FAT10 E2 enzyme BRUCE (data not shown). Further testing will be needed to conclude, whether BRUCE is an E2 that specifically functions together with UBA6 and FAT10.

6.2 Yeast two-hybrid screen with TRIM11

In a yeast two-hybrid screen that had been performed previously to this work, TRIM11 was identified as a putative interaction partner of FAT10. Moreover, TRIM11 could interact with the FAT10 specific E2 enzyme USE1 in vivo and siRNA mediated TRIM11 downregulation led to decreased FAT10 conjugates, indicating that TRIM11 may function as a FAT10 specific RING finger E3 ligase (A. Aichem, unpublished data).

6.2.1 TRIM11 interacts specifically with JunB and Ambra1 in a yeast two- hybrid screen

To discover interacting proteins and to gain more information about the function of TRIM11, a yeast two-hybrid screen with the E3 ligase TRIM11 was performed. As TRIM11 could be a putative E3 ligase for FAT10, our focal point was the identification of FAT10 substrates. Since we measured the highest FAT10 expression in thymus, we used a human thymus cDNA library, where the prey proteins were expressed as N-terminal fusion proteins with the GAL4 activation domain and screened with TRIM11 as a bait protein, which was expressed as an N-terminal fusion with the LexA-DNA binding domain.

Results

Absence of self-activation of the bait construct pLexA-TRIM11 was positively tested on selection plates lacking tryptophane (SD-T) (Figure 19 (a) left panel). Further, expression of TRIM11 did not result in lacZ activation, which was tested by a (X-GAL) filter assay as depicted in Figure 19 (a), right panel. The screening stringency was optimized and determined during a pilot screen to eliminate most false positive interaction partners. To reduce the background, 5 mM 3-aminotriazole (3- AT) was included in the medium throughout the screen. A large-scale yeast two-hybrid screen uncovered a total of 121 clones, which were positively tested by an X-Gal test (see 5.2.12). Clones that turned blue were grown on new selection plates and the X-Gal test was repeated four times. The prey plasmids of the remaining 111 clones were isolated using a standard method with glass beads (see 5.2.11) and were retransformed into NMY51, together with either pLexA- TRIM11, to confirm the positive result of the screen or together with pLexA-LaminC or pLexA-p53, to test for bait independency. The identity of the remaining 96 clones was determined by sequencing. A large-scale yeast two-hybrid screen uncovered 3 independent cDNA clones encoding for JunB and 11 clones encoding for Ambra1. The interaction between the proteins described was apparent in an X-Gal-filter assay and judged from the color change on selective growth plates (Figure 19 (b) left panel). Apart from being able to grow on -TLH selection plates (as evidenced by white colony color on selective plates) already indicating the interaction of the proteins, they turned blue in the X-Gal test (Figure 19 (b) right panel) and they showed the same white color as it can be seen in the positive control (ISG15 and its E1 enzyme UBE1L). The interactions between TRIM11 with either JunB or Ambra1 were further characterized in this work where JunB and Ambra1 were also investigated in terms of a being a FAT10 specific substrate.

Results

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(b)

Figure 19: TRIM11 interacts specifically with JunB and Ambra1 in a yeast two-hybrid screen. (a) Yeast NMY51 was transformed with the bait construct pLexA-TRIM11 and grown on selection plates lacking tryptophane (left panel). Absence of self-activation was tested with an X-GAL filter assay (right panel). (b) Yeast NMY51 was co-transformed with pLexA-ISG15 and pACT2-UBE1L (positive control), or with pLexA- TRIM11 and pACT2-JunB or pACT2-Ambra1, respectively and grown on 5 mM 3-AT-containing selection plates lacking tryptophane, leucine and histidine (left panel). An X-Gal filter assay was performed to verify the interaction of the respective proteins (right panels). A representative experiment from a total of three experiments with similar outcome is shown.

6.2.2 Interaction of JunB with TRIM11 in human cell culture

Often, distinct circumstances have to be given to permit protein-protein interactions. Yeast cells offer a different environment than do mammalian cells; thus a positive result in a yeast two-hybrid assay does not ensure that interaction between two proteins takes place under in vivo conditions. In order to confirm the yeast two-hybrid results and to investigate, whether interactions can indeed occur in mammalian cells, human TRIM11 and the native full-length JunB were further tested for their interaction under in vivo conditions in the mammalian cell line HEK293. HEK293 cells were either transiently single-transfected with a MYC-FLAG- tagged JunB construct alone, or together with a HIS-tagged TRIM11 construct to carry out co-immunoprecipitation experiments. TRIM11, a member of the TRIM protein family of E3 ubiquitin ligases has been described to interact with several transcription factors and transcriptional co-activators and to regulate their degradation via the ubiquitin–proteasome system (Hong et al., 2008; Ishikawa et al., 2006; Tuoc and Stoykova, 2008). In case that JunB is a TRIM11 specific substrate, JunB could be, due to modifications with ubiquitin or FAT10, destabilized and rapidly degraded by the proteasome.

Results

Therefore, cells were treated in parallel with the proteasomal inhibitor MG132 for 6 h, before cells were lysed and JunB and TRIM11 expression levels in the presence or absence of the proteasome inhibitor MG132 were compared. After 24 h ectopic protein expression, JunB was immunoprecipitated from cell lysates using anti-FLAG-Agarose and expression was detected by Western blot analysis through specific antibodies (Figure 20). As a loading control for the prepared lysates, β-actin was used. All samples were analyzed under reducing conditions (10 % β-mercaptoethanol).

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(b)

Figure 20: TRIM11 interacts with JunB in vivo. HEK293 cells were transiently transfected with pCMV6-JunB-MYC-FLAG and pcDNA3-HIS/-A-TRIM11, or left untransfected, as indicated. Cells were pre-treated with/without the proteasome inhibitor MG132 for 6 hours. All samples were analyzed under reducing conditions (10% β-mercaptoethanol). After immunoprecipitation against the FLAG-Tag of JunB-MYC-FLAG samples were separated on 4-12% Bis-Tris SDS gels and subjected to Western blot analysis using a (a) FLAG-reactive antibody conjugated to horseradish peroxidase (HRP). Whole cell lysates were immunoblotted for JunB to control for differences in expression (input). (b) HIS-TRIM11 expression (input) and co-immunoprecipitated TRIM11 was detected with a peroxidase (POX) conjugated polyhistidine-reactive antibody. β-actin served as a loading control.

Results

JunB migrates as a duplet/triplet in HEK293 cells at a size of 48 kDa (highest band). JunB was detected by Western blot analysis using an anti-FLAG antibody with an expected size of 48 kDa (Figure 20 (a) lane 2-4, input). The protein level of JunB decreased when TRIM11 was co-expressed and MG132 treatment barely rescued proteasomal degradation (Figure 20 (a) lane 3 and 4). Even major apparent differences were visible regarding the protein level of immunoprecipitated JunB. The amount of immuno-precipitated JunB protein level decreased when JunB was ectopically co-expressed together with TRIM11 (Figure 20 (a) lane 6-8). This could argue for a probable role as E3 ubiquitin ligase where binding destabilizes JunB, as has been described for other TRIM11 substrates (Hong et al., 2008; Ishikawa et al., 2006; Niikura et al., 2003; Tuoc and Stoykova, 2008). Moreover, in Figure 20 (a) lane 8, a high molecular weight smear is observable, when cells were treated additionally with MG132 for 6 h. This could argue for augmented JunB conjugate formations, for instance ubiquitination when TRIM11 is overexpressed, which are rescued from proteasomal degradation. HIS-tagged TRIM11 could be detected with an anti-6-HIS-antibody at a size of 52 kDa (Figure 20 (b), lane 2-4). The expression of TRIM11 decreased when cells where double-transfected together with JunB (Figure 20 (b), lane 3 and 4) whereas the amount of total protein remained stable (loading control β-actin). The interaction between TRIM11 and JunB after immunoprecipitation of JunB and Western blot analysis against the HIS-tag of TRIM11 was hardly detectable (Figure 20 (b)). Pre-treatment with the proteasomal inhibitor MG132 for 6 h clearly augmented the amount of co-immunoprecipitated TRIM11, suggesting that TRIM11 or JunB became stabilized after proteasome inhibition.

6.2.3 Ubiquitin and FAT10 become isopeptide linked to to JunB

Several TRIM proteins are implicated in ubiquitination and act as single RING finger ubiquitin E3 ligases, which can affect the direct transfer of ubiquitin from E2~Ub/UBL to substrate proteins (Hong et al., 2008; Ishikawa et al., 2006; Tuoc and Stoykova, 2008). Preliminary work could show a specific interaction between TRIM11 and the ULM FAT10 in a yeast two hybrid screen. Moreover, TRIM11 could interact in vivo with the FAT10 specific E2 enzyme USE1 and depletion of TRIM11 via siRNA knockdown significantly reduced the amount of FAT10 conjugates, indicating that TRIM11 may function as a FAT10 specific RING finger E3 ligase (A. Aichem, unpublished). Because enzyme sharing between different ULMs is not unusual it should be tested if TRIM11 not only possesses ubiquitin E3 ligase activity but is also able to cooperate with FAT10.

Results

Hence, we were interested, if the transcription factor JunB can covalently bind both, ubiquitin, as well as FAT10 and could be, beside USE1 (Aichem et al., 2010), p53 (Li et al., 2011) and huntingtin (Nagashima et al., 2011) a further substrate in the FAT10 conjugation pathway. If TRIM11 is either a ubiquitin or FAT10 specific E3 ligase with JunB as substrate, it would be expected that co-expression of these proteins would either enhance ubiquitin or FAT10 conjugation to JunB or lead to general augmentation of ubiquitin or FAT10 conjugate formation in transfected cells. Typically, E3 ligases confer substrate specificity in the ULM conjugation pathway but that doesn’t exclude that they are able to facilitate the modification of many different substrates. We transiently transfected HEK293 cells with either a HA-tagged ubiquitin or HA-tagged FAT10 constructs alone, or together with a MYC-FLAG-tagged JunB and a HIS-tagged TRIM11 construct. After 24 h ectopic protein expression, ubiquitin and FAT10 were immunoprecipitated from cell lysates using anti-HA-agarose and expression was detected through immunoblotting with specific antibodies (Figure 21(a), (b) and (c)). β-actin served as a loading control for the prepared lysates. All samples were boiled in reducing gel sample buffer (10 % β-mercaptoethanol), to cleave non-covalent thioester-linkages, whereas isopeptide linkages remain stable.

(a)

Results

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(c)

Figure 21: Conjugate formation of JunB with either ubiquitin or FAT10 in vivo. Co-immunoprecipitation of JunB-MYC-FLAG, HA-FAT10, HA-Ubiquitin and HIS-TRIM11 as indicated. HEK293 cells were transiently transfected with pCMV6-JunB-MYC-FLAG, pcDNA3.1-HA-FAT10, pCDNA-3.1-HA-Ubiquitin and pCDNA3-HIS/-A-TRIM11, or left untransfected. All samples were analyzed under reducing conditions (10% β- mercaptoethanol). After immunoprecipitation against the HA-Tag of HA-FAT10 and HA-Ubiquitin with anti-HA- agarose samples were separated on 4-12% Bis-Tris SDS gels and subjected to Western blot analysis with a horseradish peroxidase (HRP)-conjugated HA-reactive antibody (a). The expression of HIS-TRIM11 was detected with a peroxidase (POX) conjugated polyhistidine-reactive antibody (b). The expression of JunB-MYC-FLAG was detected with a FLAG-reactive antibody conjugated to horseradish peroxidase (HRP). Conjugate formation of HA- FAT10 and JunB-MYC-FLAG could be detected after immunoprecipitation against the HA-Tag of FAT10 and Western blot analysis against the FLAG-tag of JunB (c). Arrow head indicates a putative homo/hetero-dimerized JunB-ubiquitin conjugate. β-actin served as a loading control.

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Whole cell lysates were directly immunoblotted with an anti-HA antibody to reveal the expression of HA-tagged ubiquitin (~8 kDa) and HA-tagged FAT10 (~18 kDa) as depicted in Figure 21 (a). Further, cell lysates were directly immunoblotted with an anti-6HIS antibody to illustrate expression of HIS-TRIM11 (~52 kDa) and an anti-FLAG-antibody, to detect FLAG- tagged JunB (~48 kDa) and JunB conjugates as depicted in Figure 21 (b) and (c). All proteins could be detected at their predicted size. The amount of freely distributed ubiquitin was highest, when no further protein was co-expressed. The same result was observable for monomeric FAT10 (Figure 21 (a), input). Moreover, when ubiquitin was co- expressed with JunB or TRIM11, or with both proteins together, ubiquitin conjugates shifted to higher molecular weights (apparent in ubiquitin ladder, see Figure 21 (a), input). Interestingly, immunoprecipitation against the HA-tag of ubiquitin and Western blot analysis through a HA–reactive antibody revealed, that ubiquitin-smear is almost absent in Figure 21 (a), lane 6, when ubiquitin is co-expressed together with JunB and TRIM11. Strikingly, a prominent double band appears at the height of ~110 kDa. This size would equate to homo/hetero-dimerized JunB conjugated to ubiquitin (marked by arrow head), but this hypothesis indeed needs further investigation. Interestingly, this double band is also visible in Figure 21 (c), lane 3, when ubiquitin, co-expressed with JunB, is immunoprecipitated and analyzed with an anti-FLAG reactive antibody to detect ubiquitinated JunB. Following immunoprecipitation of HA-Ubiquitin with anti-HA-agarose beads and Western blot analysis against the FLAG-tag of JunB, JunB could be on one hand detected for the most part at a size of 48 kDa, what would indicate a non-covalent interaction between JunB and ubiquitin (see Figure 21 (c) lane 3-5). The amount of co-immunoprecipitated JunB was clearly higher than the amount that unspecifically bound to the anti-HA-agarose beads apparent in Figure 21 (c) line 2, suggesting that the two proteins are mainly non-covalently linked (Figure 21 (c) lane 3). However, on the other hand a clear conjugate band at a height of ~56 kDa appears (Figure 21 (c), line 3), which size equates to mono-ubiquitinated JunB. This conjugate linkage is not reducible what strongly argues for an isopeptide linkage. Furthermore, a smear at higher molecular weights is visible, indicating that JunB becomes also poly-ubiquitinated. Conspicuous in this case is the prominent double band appearing at the height of ~110 kDa, as seen before (Figure 21 (a), lane 6, arrow head), which size equates to homo- or hetero -oligomerized JunB, conjugated to ubiquitin, but still remains to be investigated. Interestingly, co-expression of ubiquitin and JunB together with TRIM11 resulted in almost complete abrogation of Ub-JunB conjugate formation and TRIM11 overexpression led to the abolishment of ubiquitin smear (IP: Figure 21 (c) lane 4).

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Surprisingly, after co-immunoprecipitation of HA-FAT10 and immunoblotting against the FLAG-tag of JunB, a high molecular weight (~66 kDA) conjugate double band of FAT10 and JunB was visible, although a JunB (48 kDa) remained partially non-covalently linked (Figure 21 (c) lane 5). The size of the double band fits to a monoFAT10ylated form of JunB which is not reducible and gives a strong hint that modification occurs covalently. Strikingly, the conjugate double-band disappeared almost completely when TRIM11 was co-expressed (Figure 21 (c) lane 6), similar to the previous case, when ubiquitin and JunB were co- expressed together with TRIM11. Generally, overexpression of TRIM11 led to a decreased protein level of ubiquitin, FAT10 and JunB (Figure 21 (a), (b), (c)), which could argue for accelerated protein degradation when TRIM11 is present in an abundant amount. Collectively, these results show that JunB becomes ubiquitinated and FAT10ylated under the chosen conditions and provide evidence, that an interaction of JunB and ubiquitin or FAT10 occurs in vivo. So far, the putative FAT10 E3 ligase function of TRIM11 is not evidenced, and was investigated in an in vitro FAT10ylation assay at a later time point (see 6.2.13).

6.2.4 Proteasome inhibition augments conjugate formation between JunB and FAT10

To date, ubiquitin and FAT10 are the only ULMs that have been described to tag proteins for the degradation through the 26S proteasome (Hershko, 1983; Hipp et al., 2005). Like ubiquitin, FAT10 bears a free di-glycine motif at the C-terminus, which mediates the conjugation to target proteins. To elucidate, if the JunB-FAT10 conjugate is degraded via the proteasomal pathway, a MYC-FLAG- tagged JunB construct was expressed together with HA-FAT10 or together with HA-FAT10 and HIS-TRIM11 in HEK293 cells. Transfected cells were pre-treated with the proteasome inhibitor MG132 (10 µM), 6 h before cell lysis, or left untreated. Moreover, to ensure that conjugate formation of JunB and FAT10 consists of an isopeptide-linkage, we co-expressed JunB-MYC-FLAG, HIS-TRIM11 together with HA- FAT10ΔGG, which served as a negative control, since HA-tagged-FAT10ΔGG is not capable to form an isopeptide linkage to lysine residues on specific substrates.

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Figure 22: Conjugate formation of FAT10 and JunB in vivo. Co-immunoprecipitation of JunB-MYC-FLAG, HA-FAT10, HA-FAT10ΔGG and HIS-TRIM11 as indicated. HEK293 cells were transiently transfected with pCMV6-JunB-MYC-FLAG, pcDNA3.1-HA-FAT10, pCDNA-3.1-HA- FAT10ΔGG and pcDNA3-HIS/-A-TRIM11. Cells were treated either with/without the proteasome inhibitor MG132 (10 µM) for 6 hours. All samples were analyzed under reducing conditions (10% β-mercaptoethanol). After immunoprecipitation against the HA-Tag of HA-FAT10 and HA-FAT10ΔGG with anti-HA-agarose, samples were separated on 4-12% Bis-Tris SDS gels and subjected to Western blot analysis with a horseradish peroxidase (HRP)-conjugated HA-reactive antibody (a). The expression of HIS-TRIM11 was detected with a peroxidase (POX) conjugated polyhistidine-reactive antibody (b). The expression of JunB-MYC-FLAG was detected with a FLAG-reactive antibody conjugated to horseradish peroxidase (HRP). Conjugate formation of HA-FAT10 and JunB-MYC-FLAG could be detected after immunoprecipitation against the HA-Tag of FAT10 and Western blot analysis against the FLAG-tag of JunB (c). The plasmid content in all transfections was balanced with empty expression vector pcDNA3. β-actin served as a loading control.

24 h after transfection, immunoprecipitation with anti-HA agarose was performed to accumulate FAT10-conjugated proteins. Analysis by Western blotting revealed, that at the height of ~66 kDa a double band appears the size of which equates to a monoFAT10ylated form of JunB (as shown before in Figure 21), detected through a peroxidase-conjugated FLAG-reactive antibody against the FLAG-Tag of JunB (Figure 22 (c), lanes 4-7). The appearing double band led to the assumption, that different forms of JunB are modified with FAT10. Whether this is a phosphorylated or other modified form of JunB remains hitherto unclear. The amount of JunB-FAT10 conjugates could be augmented significantly, when cells were pre-treated with MG132 for 6 hours before cell lysis, suggesting an involvement of the proteasome for conjugate degradation. No conjugation formation was detectable when JunB was co-transfected together with FAT10ΔGG (Figure 22 (c) lane 8-11). This result strengthens the presumption, that FAT10 becomes isopeptide linked to an ε- lysine residue of JunB, since FAT10ΔGG, lacking the di-glycine motif is not capable to form this linkage. Co-transfection of JunB together with either FAT10 or FAT10∆GG decreased the amount of FAT10 and FAT10∆GG, while treatment with MG132 recovered the effect (Figure 22 (a) lane 3, 4 and lane 8, 9). Moreover, co-transfection of TRIM11 highly decreased the amount of JunB, FAT10 and FAT10∆GG (Figure 22 a), b) and c) lane 6-7). This effect could in turn partially be rescued with MG132 treatment (Figure 22 a-c). Further, conjugate formation of JunB and FAT10 was almost completely abolished, when TRIM11 was co-expressed, which argue in the first instance against the hypothesis, that TRIM11 is a FAT10 specific E3 ligase with JunB as substrate. It rather let suppose a role for TRIM11 in controlling JunB and JunB conjugate stability. There is a strong hint that FAT10 modification serves as a degradation signal, because the conjugate was accumulating after proteasome inhibition.

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6.2.5 JunB has no influence on the degradation rate of FAT10

Attachment of FAT10 causes the rapid degradation of long-lived proteins, which is dependent on the 26S proteasome (Hipp et al., 2005). In contrast to ubiquitin, FAT10 has a relatively short half-life because it is also subject to proteasomal degradation in its monomeric form and, additionally, it is probably not recycled but instead degraded along with its substrates (Hanna and Finley, 2007). The turnover of FAT10 was described to decrease rapidly after addition of cycloheximide (Raasi et al., 2001) showing that the half-life of FAT10 is ~1 hour. To show as a control that JunB expression does not alter FAT10 degradation rate, we next investigated in a cycloheximide chase if JunB interaction with FAT10 would have functional relevance for the degradation of FAT10 in vivo. The compound cycloheximide inhibits protein synthesis by blocking translation elongation and therefore can be used in time-course experiment followed by Western blotting of the cell lysates for the protein of interest to determine the half-life of proteins. HEK293 cells were transfected with pCMV6-JunB-MYC-FLAG and pcDNA3.1-HA-FAT10 constructs and cells were treated with cycloheximide (50 µg ml-1) for the indicated time points. Six hours before harvesting MG132 (10 μM) was added, as indicated. Whole-cell lysates were either directly immunoblotted 24 h post-transfection with an anti-HA reactive antibody or subjected to co-immunoprecipitation assays, using anti-HA agarose followed by SDS-PAGE and Western blotting with a anti-HA antibody (Figure 23).

Figure 23: Co-expression of JunB has no influence on the degradation rate of FAT10. HEK293 cells were transiently co-transfected with pCMV6-JunB-MYC-FLAG and pcDNA3.1-HA-FAT10 plasmid. Before cell lysis, cells were treated with the proteasome inhibitor MG132 (10 µM) for 6 hours and with cycloheximide (50 µg ml-1) at different time points, as indicated. After immunoprecipitation against the HA-Tag of FAT10 with anti-HA agarose, samples were separated on 4-12% Bis-Tris SDS gels and subjected to Western blot analysis using a directly peroxidase-linked anti-HA mAb to evaluate FAT10 expression. All samples were analyzed under reducing conditions (10% β-mercaptoethanol). β-actin served as a loading control. Graphs show the quantification of HA-FAT10 in the cells. HA-FAT10 ECL signals of all experiments were quantified with Quantity One Software (BioRad) and mean values ± SEM of five independent experiments are depicted as relative expression to the ECL signal of HA-FAT10 transfected cells without cycloheximide treatment, which was set to unity. 105

Results

The quantitative analysis revealed a half-life for FAT10 of approximately 1 h (Figure 23) when co-expressed together with JunB. In accordance with a previous report (Raasi et al., 2001) the half life of FAT10 is 1 h. After 2.5 h the amount of FAT10 decreased to 25 % and 5 h after CHX treatment only ~10 % FAT10 remained left. Treatment with MG132 led to an accumulation of monomeric FAT10 (~150 %) by preventing degradation via the proteasome. Taken together, these results indicate that co-expression of JunB did not change the degradation rate of FAT10.

6.2.6 Co-expression of FAT10 hardly affects the degradation of unconjugated JunB

To assure that FAT10ylation is necessary to sign JunB for proteasomal degradation we next needed to determine the protein half life of unconjugated JunB in presence and absence of FAT10. Pulse-chase experiments in Swiss 3T3 cells after serum stimulation revealed a half life of approximately three hours for JunB (Kovary and Bravo, 1991). As described in the literature, JunB levels vary at M-G1 transition, depending on the phosphorylation state of JunB. In quiescent cells JunB expression is low but reaches a peak during G0/G1 transition before returning to an intermediate level in response to mitogenic stimuli, whereas in cycling cells JunB expression strongly increases when cells progress through G1-phase and enter S-phase (Farras et al., 2008). Moreover, phosphorylation of JunB by the p34cdc2-cyclin B kinase is associated with lower JunB levels in mitotic and early G1 cells (Bakiri et al., 2000). HEK293 cells were transfected with pCMV6-JunB-MYC-FLAG and pcDNA3.1-HA-FAT10 constructs and cells were treated with cycloheximide (50 µg ml-1) for the indicated time points. Six hours before harvesting MG132 (10 μM) was added, as indicated. 24 after post- transfection cells were lysed and lysates were directly immunoblotted with a FLAG-reactive antibody coupled to horseradish peroxidase to determine JunB-MYC-FLAG expression (Figure 24).

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200 200

150 150

100 100

50 50 relative JunB expression relative JunB expression

0 when FAT10 co-expressed 0 0 2.5 5 5 + 6h MG132 0 2.5 5 5 + 6h MG132 CHX treatment [h] CHX treatment [h]

Figure 24: CHX-chase: determination of JunB turnover rate in presence or absence of FAT10 HEK293 cells were transiently transfected with pCMV6-JunB-MYC-FLAG or together with pcDNA3.1-HA-FAT10 plasmid. Before cell lysis, cells were treated with/without the proteasome inhibitor MG132 (10 µM) for 6 hours and with cycloheximide (50 µg ml-1) at different time points, as indicated. a) Samples were separated on 4-12% Bis- Tris SDS gels and subjected to Western blot analysis using a directly peroxidase-linked anti-FLAG mAb to evaluate JunB expression. All samples were analyzed under reducing conditions (10% β-mercaptoethanol). β- actin served as a loading control. Graphs show the quantification of JunB-MYC-FLAG in the cells. JunB-MYC- FLAG ECL signals of all experiments were quantified with Quantity One Software (BioRad) and mean values ± SEM of five independent experiments are depicted as relative expression to the ECL signal of JunB-MYC-FLAG- transfected cells without cycloheximide treatment, which was set to unity.

JunB could be detected at a size of ~48 kDa. As depicted in Figure 24 (input), the JunB protein level remained stable over the period of time when cells were chased. This result was not surprising because cells were not synchronized, representing an intermediate protein level in different cellular states.

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Only a very slight decrease in JunB protein level over time was visible, when cells were co- transfected together with FAT10. In this experimental setup we cannot exclude the possibility that the decrease of JunB level is owing to a non-covalent interaction with FAT10. However, pre-treating the cells with MG132 could rescue JunB turnover and led to a JunB accumulation of about 125 % in both cases. These results indicate that the turnover rate of monomeric unconjugated JunB in presence of FAT10 is hardly affected.

6.2.7 CHX data reveal a role for proteasome dependent degradation of the conjugate between JunB and FAT10

The in vivo data provided evidence that FAT10 becomes stably attached to JunB (see 6.2.3) and proteasome inhibition led to an accumulation of the conjugate (see 6.2.4). We could also show, that neither FAT10 (see 6.2.5) nor JunB turnover (6.2.6) is noticeably altered in its monomeric form, when not conjugated to each other. Since FAT10 is degraded by the 26S proteasome and targets FAT10-linked proteins for proteasomal degradation (Hipp et al., 2004; Schmidtke et al., 2009), we investigated in cycloheximide-chase experiments, if the turnover rate of the conjugate between JunB and FAT10 changes over time and is caused through proteasomal degradation. Moreover, the in vivo data already indicated that FAT10 becomes also non-covalently linked to JunB (see chapter 6.2.3 and 6.2.4.). Hence, we next aimed to determine the turnover rate of covalently and non-covalently linked JunB and FAT10. For this purpose, HEK293 cells were transiently transfected with MYC-FLAG-tagged JunB and HA-tagged FAT10 constructs and treated with cycloheximide (50 µg ml-1) for 2.5 and 5 h, or left untreated. Six hours before cells were harvested, 10 μM MG132 was added to inhibit the proteasome. 24 h after transfection, whole-cell lysates were subjected to co- immunoprecipitation assays using anti-HA agarose, followed by SDS-PAGE and Western blotting with an anti-FLAG reactive antibody.

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Figure 25: CHX-chase: Determination of JunB-FAT10 conjugate turnover rate HEK293 cells were transiently co-transfected with pCMV6-JunB-MYC-FLAG and pcDNA3.1-HA-FAT10 plasmid. Before cell lysis, cells were treated with the proteasome inhibitor MG132 (10 µM) for 6 h and with cycloheximide (50 µg ml-1) at different time points, as indicated. Whole cell lysates were subjected to Western blot analysis with a FLAG-reactive antibody coupled to HRP. After immunoprecipitation against the HA-Tag of FAT10 with anti-HA agarose, samples were subjected to Western analysis using a directly horseradish peroxidase (HRP) -linked anti- FLAG mAb to evaluate conjugate formation. All samples were analyzed under reducing conditions (10% β- mercaptoethanol). β-actin served as a loading control. ECL signals of all experiments were quantified with Quantity One Software (BioRad). A representative blot out of five independent experiments is shown. Mean values ± SEM of five independent experiments are depicted as relative expression to the ECL signal of HA-FAT10 JunB-MYC-FLAG transfected cells without cycloheximide treatment, which was set to unity.

Conjugate formation between FAT10 and JunB could be observed after immunoprecipitation against the HA-tag of FAT10 and immunoblotting against the FLAG-tag of JunB, apparent in the higher molecular weight double band at around 66 kDa (Figure 25). Regarding the conjugate stability of covalently linked FAT10 and JunB, a continuous decline during the indicated timepoints is detectable, which reveals a half-life of approximately 2.5-3 h compared to the much shorter half life of monomeric FAT10 (1 h) (Raasi et al., 2001). The FAT10-JunB complex degradation could be rescued, when cells were pre-treated with the proteasomal inhibitor MG132 which led to a considerable accumulation of the conjugate (~180 %), suggesting a continuous degradation via the proteasome. Interestingly, non- covalently bound JunB was continuously degraded and revealed a similar rate of approximately 2.5 h. Combined with previous findings, these results suggest that the role of FAT10 is the rapid destruction of its target proteins via conjugation, either covalently or surprisingly also non-covalently, and subsequent degradation.

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6.2.8 TRIM11 becomes degraded via the proteasome

The previous results did not provide clear evidence that TRIM11 acts either as a FAT10 or ubiquitin specific E3 ligase with JunB as substrate. We would expect, if TRIM11 is a special ubiquitin or FAT10 E3 ligase with JunB as substrate, conjugate formation to JunB would be enhanced, when the proteasome is inhibited at the same time. However, we could observe the opposite effect; overexpression of TRIM11 led to severe downregulation of JunB-Ub and JunB-FAT10 conjugates. There exists the possibility that TRIM11 is not an E3 ligase but rather a substrate for ubiquitin or FAT10 and attachment leads to to its proteasomal degradation. In this case a further E3 ligase could be involved to assign TRIM11 for proteasomal degradation. So far, no protein half-life could be determined for endogenous TRIM11, because specific antibodies were missing. To characterize TRIM11 we investigated the turnover rate of TRIM11 in a cycloheximide chase with a new polyclonal TRIM11 antibody (Sigma), which reliably recognized endogenous and overexpressed TRIM11. To investigate, if TRIM11 is assigned to proteasomal degradation, cells were treated with or without MG132 for 6 h. HEK293 cells were treated with cycloheximide (50 µg ml-1) for 0, 2.5, 5 and 7.5 h, or left untreated. Six hours before cells were harvested, 10 μM MG132 was added.

Figure 26: Determination of endogenous TRIM11 turnover rate HEK293 cells were treated before cell lysis with the proteasome inhibitor MG132 (10 µM) or left untreated for 6 hours and with cycloheximide (50 µg ml-1) at different time points, as indicated. Samples were subjected to Western blot analysis using a polyclonal TRIM11 antibody (Sigma) to evaluate TRIM11 expression. All samples were analyzed under reducing conditions (10% β-mercaptoethanol). β-actin served as a loading control. ECL signals of all experiments were quantified with Quantity One Software (BioRad). A representative blot out of three independent experiments is shown. Mean values ± SEM of three independent experiments are depicted as relative expression to the ECL signal of endogenous TRIM11 without cycloheximide treatment, which was set to unity.

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In HEK293 cells, a continuous decrease of endogenous TRIM11 for the indicated timepoints after cycloheximide addition could be observed and a protein half-life of approximately 7-8 hours was calculated (Figure 26). Only a slight rescue of TRIM11 could be measured when MG132 was added, which would argue against proteasomal degradation. However, cycloheximide treatment for more than 5 h led to increased cell death and additional apposition of MG132 enhanced the effect.

To compare the protein half-lifes of endogenous and overexpressed TRIM11 a second cycloheximide chase was performed. HEK293 cells were transiently transfected with a TRIM11-FLAG construct and cycloheximide (50 µg ml-1) was added for 2.5, 5, 7.5 or 10 h, or left untreated, before cell lysis. Moreover, cells were treated with or without MG132 for 6 h. After cell lysis, samples were boiled in Laemmli-Buffer containing 10% ß-mercaptoethanol followed by SDS-PAGE and Western blotting with a FLAG-reactive antibody.

Figure 27: Determination of ectopic expressed TRIM11 turnover rate HEK293 cells were transiently transfected with pcDNA3-TRIM11-FLAG plasmid. Before cell lysis, cells were treated with the proteasome inhibitor MG132 (10 µM) for 6 hours and with cycloheximide (50 µg ml-1) for different time periods, as indicated. After immunoprecipitation against the FLAG-Tag of TRIM11 with anti-FLAG agarose, samples were subjected to Western analysis using a directly coupled horseradish peroxidase (HRP) -linked anti- FLAG mAb to evaluate TRIM11 expression. All samples were analyzed under reducing conditions (10% β- mercaptoethanol). β-actin served as a loading control. ECL signals of all experiments were quantified with Quantity One Software (BioRad). A representative blot out of three independent experiments is shown. Mean values ± SEM of three independent experiments are depicted as relative expression to the ECL signal of TRIM11- FLAG transfected cells without cycloheximide treatment, which was set to unity.

FLAG-tagged TRIM11 migrates as a double band with a size of 48 and 55 kDa. A continuous decline of overexpressed TRIM11-FLAG protein level is observable for the indicated timepoints and quantitative analysis revealed a half-life of approximately 5 h (Figure 27), which is much faster than for endogenous TRIM11.

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Besides, treatment with MG132 completely blocked TRIM11-FLAG degradation and increased accumulation, which clearly substantiate an involvement of the proteasome for TRIM11-FLAG turnover. These results clearly show that ectopically expressed TRIM11 is faster degraded than endogenous TRIM11, maybe due to excess of protein. Moreover, MG132 treatment leads to accumulation of TRIM11, which demonstrate an involvement of the proteasome in TRIM11 turnover.

6.2.9 TRIM11 turnover in presence of FAT10 is slightly accelerated

In order to investigate whether TRIM11 degradation is affected in presence of FAT10, HEK293 cells were transiently transfected with a HA-tagged FAT10 and a HIS-tagged TRIM11 construct. Cycloheximide (50 µg ml-1) was added to the indicated timepoints for 2.5 and 5 h or left untreated treated. Moreover, cells were treated with or without MG132 for 6 h before cell lysis.

Figure 28: Determination of TRIM11 turnover rate in the presence of FAT10 HEK293 cells were transiently transfected with pcDNA3-HIS/-A-TRIM11and pcDNA3.1-HA-FAT10 plasmids. Before cell lysis, cells were treated with the proteasome inhibitor MG132 (10 µM) for 6 hours and with cycloheximide (50 µg ml-1) for different time periods, as indicated. Samples were subjected to Western analysis using a HIS-reactive antibody to evaluate TRIM11 expression. All samples were analyzed under reducing conditions (10% β-mercaptoethanol). β-actin served as a loading control. A representative blot out of three independent experiments is shown. Mean values ± SEM of three independent experiments are depicted as relative expression of transfected cells without cycloheximide treatment.

After cell lysis, whole-cell lysates were subjected to Western blot analysis with a anti-6HIS- reactive antibody, to illustrate TRIM11 expression. The cycloheximide data illustrate, that the HIS-TRIM11 half life in presence of FAT10 is about 4 h and therefore slightly accelerated in comparison to TRIM11-FLAG alone (see 6.2.8), where 5 h after cycloheximide treatment TRIM11 protein level decreased about 50 %. Besides, MG132 treatment enhanced accumulation of HIS-TRIM11. 112

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6.2.10 Co-expression of TRIM11 does not change protein turnover rates of JunB and FAT10

We next aimed to determine the protein turnover rate of JunB and FAT10 in presence of TRIM11. Moreover, we wanted to analyze whether TRIM11 degradation is affected in presence of JunB and FAT10. HEK293 cells were transiently transfected with a HA-tagged FAT10 and FLAG-tagged JunB construct, or together with a HIS-tagged TRIM11 construct. Cycloheximide (50 µg ml-1) was added for the indicated time periods, 2.5 and 5 h, or left untreated. Moreover, cells were treated with or without MG132 for 6 h. Aliquots of lysates from transfected cells were analyzed for expression of the respective proteins (Figure 29 (a) for FAT10, (b) for TRIM11 and (c) for JunB and JunB-FAT10 conjugates). After immunoprecipitation with anti-HA-agarose beads, we looked for immunoprecipitated FAT10 (Figure 29 (b)) and co-immunoprecipitated JunB (Figure 29 (c)).

(a)

(b)

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Figure 29: Determination of the turnover rate of the JunB-FAT10 conjugate HEK293 cells were transiently co-transfected with a pCMV6-JunB-MYC-FLAG and a pcDNA3.1-HA-FAT10 plasmid. Before cell lysis, cells were treated with/without the proteasome inhibitor MG132 (10 µM) for 6 hours and with cycloheximide (50 µg ml-1) at different time points, as indicated. After immunoprecipitation against the HA-tag of FAT10 with anti-HA agarose, samples were subjected to Western analysis using a directly horseradish peroxidase (HRP) -linked anti-FLAG mAb to evaluate JunB expression and conjugate formation. The plasmid content in all transfections was balanced with empty expression vector pcDNA3. All samples were analyzed under reducing conditions (10% β-mercaptoethanol). β-actin served as a loading control. ECL signals of all experiments were quantified with Quantity One Software (BioRad). A representative blot out of four independent experiments is shown. Mean values ± SEM of three independent experiments are depicted as relative expression to the ECL signal of transfected cells without cycloheximide treatment, which was set to unity.

Quantitave analysis resulted in a quite similar degradation rate for either FAT10 or JunB, in presence or absence of overexpressed HIS-TRIM11 (data not shown). The JunB-FAT10 conjugate turnover rate between JunB and FAT10 in presence of TRIM11 could not be calculated, because co-expression with TRIM11 almost completely abolished conjugate formation (Figure 29 (c), lane 6-9). As we could show previously, co-expression of TRIM11 resulted in severely decreased protein levels either for JunB or FAT10 and TRIM11 itself, without changing the velocity of degradation, which might suggest a role in regulating and controlling protein stability and abundance and might be involved in protein quality control as has been shown for Pax6 (Tuoc and Stoykova, 2008).

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6.2.11 Conjugate formation of endogenous JunB and FAT10

To test whether a JunB-FAT10 conjugate can also be found in absence of overexpression, we treated HEK293 cells with TNF-α and IFN-γ to induce FAT10 expression and then performed immunoprecipitation with a FAT10-specific monoclonal antibody (mAb) (designated 4F1), followed by Western blot analysis with either a FAT10-specific polyclonal antibody (pAB) or a polyclonal JunB antibody under reducing conditions (10 % β- mercaptoethanol). This cell line, despite low JunB expression, was chosen because FAT10 can easily induced with the proinflammatory cytokines TNF-α and IFN-γ.

(a)

(b)

Figure 30: Co-immunoprecipitation of endogenous FAT10 and JunB from HEK293 cells. HEK293 cells were treated with 200 U ml− 1 IFN-γ and 400 U ml− 1 TNF-α to up-regulate endogenous FAT10 expression or left untreated. After 24 h, cells were lysed and FAT10 was immunoprecipitated using a monoclonal FAT10-reactive antibody (4F1), followed by SDS-PAGE and Western blot analysis using a FAT10-reactive polyclonal Ab (a) or a JunB-reactive polyclonal Ab (ab314221) (b) under reducing conditions (10 % β- mercaptoethanol). The arrow head indicates JunB-FAT10 conjugate formation. Asterisks indicate heavy and light antibody chains of the FAT10-reactive antibody used for the immunoprecipitation.

Whole-cell lysates were also directly immunoblotted with a polyclonal FAT10 and JunB- antibody to reveal the expression of induced FAT10 and endogenous JunB. FAT10 (~18 kDa) was clearly induced, although better detectable after immunoprecipitation (Figure 30 (a)). Due to various background bands it was difficult to detect the appropriate band showing endogenous JunB (Figure 30 (b), input). However, the blot in Figure 30 (b) reveals, that endogenous JunB (~48 kDa) becomes co-immunoprecipitated with FAT10, which indicates a non-covalent interaction between the two proteins.

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Endogenous JunB-FAT10 conjugate was detectable in a very low grade, in the presence but not in the absence of FAT10 induction by proinflammatory cytokines (Figure 30 (b), lane 4, arrow head).

6.2.12 Conjugate formation of JunB and FAT10 under semi-endogenous conditions

Since conjugate formation between JunB and FAT10 was hardly detectable under completely endogenous conditions, the experimental setup was slightly modified and performed under semi-endogenous conditions. Hence we next aimed to determine whether endogenous FAT10, upregulated by TNF-α and IFN-γ, can build a stable conjugate with over-expressed JunB. HEK293 cells were transiently transfected with the expression construct pCMV6-JunB-MYC- FLAG and treated with TNF-α and IFN-γ for one day, or left untreated, to carry out co- immunoprecipitation experiments. JunB was immunoprecipitated with FLAG-M2 affinity matrix and samples immunoblotted either with a FLAG-reactive antibody to reveal JunB expression (Figure 31 (a)), or a polyclonal FAT10 antibody to detect endogenous FAT10 and FAT10 conjugates (Figure 31 (b)).

(a) (b)

Figure 31: Co-immunoprecipitation of endogenous FAT10 and transiently transfected JunB from HEK293 cells. HEK293 cells were transiently transfected with 4 µg pCMV6-JunB-MYC-FLAG plasmid and subsequently treated with 200 U ml− 1 IFN-γ and 400 U ml− 1 TNF-α to up-regulate endogenous FAT10 expression, or left untreated. After 24 h, cells were lysed and JunB was immunoprecipitated using anti-FLAG-agarose, followed by SDS-PAGE and Western blot analysis using a FLAG-reactive antibody conjugated to HRP (left panel) or a polyclonal FAT10 specific Ab (right panel) under reducing conditions (10 % β-mercaptoethanol). The arrow head indicates JunB- FAT10 conjugate formation.

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JunB expression was apparently increased in INF-γ/TNF-α treated cells (Figure 31 (a), lane 3), whereas the total protein amount remained constant (WB: anti-ß-actin). Conjugate formation between JunB and FAT10 could be detected under reducing conditions (10 % ß-mercaptoethanol) after immunoprecipitation against the FLAG-tag of JunB and Western Blot analysis against FAT10 in a high molecular weight band (~65 kDa), as indicated by the arrow head (Figure 31 (b)). Interestingly, only a single conjugate band could be detected in comparison to the previous experiments (see 6.2.3), where always a double- band appeared. However, these results indicate, that conjugate formation between JunB and FAT10 takes place under semi-endogenous condition.

6.2.13 In vitro auto-FAT10ylation assay

The experiments described above strongly indicate, that FAT10 becomes stably attached to JunB via an isopeptide linkage. The function of TRIM11 as a putative FAT10 E3 ligase remains hitherto unclear. In the performed experimental in vivo setups we could show, that co-expression of TRIM11 led to decreased protein amounts of either FAT10, ubiquitin or JunB. Moreover, overexpression of TRIM11 reduced severely the amount of JunB-FAT10 conjugate. To ensure in vitro, if TRIM11 is a FAT10 specific E3 ligase and to determine, whether TRIM11 mediates the transfer of FAT10 onto JunB, we performed in vitro activation experiments with recombinant proteins. We expected when TRIM11 acts as a FAT10 specific E3 ligase, FAT10 can be transferred on the putative substrate JunB only in presence of FAT10 specific E1, E2 and TRIM11 under conditions containing a rich source of renewable energy such as ATP, creatine phosphate, creatine phosphokinase and inorganic pyrophosphatase, which could serve as a cell free system for the FAT10ylation reaction in vitro. For this purpose immunoprecipitation was coupled with an in vitro FAT10ylation assay of FLAG-tagged JunB in the presence of the recombinant FAT10 E1 (UBA6), E2 (HIS-USE1) and the putative FAT10 specific E3 ligase GST-TRIM11. JunB-MYC-FLAG was expressed in HEK293 cells and immunoprecipitated with anti-FLAG- agarose to purify JunB for the subsequent in vitro FAT10ylation assay. JunB bound to the beads was incubated together with purified recombinant FAT10, FLAG-UBA6 and 6HIS- USE1 in the presence of ATP at 37 °C for 1 h. Finally, JunB was visualized by Western blot analysis using anti-FLAG antibodies (Figure 32 (a)), USE1 was detected with an anti-6HIS- POX antibody (Figure 32 (b)), TRIM11 was detected with a monoclonal anti-GST-antibody (Figure 32 (c)) and FAT10 was visualized with a monoclonal anti-FAT10 antibody (Figure 32 (d)).

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(a) (b)

(d) (c)

Figure 32: In vitro FAT10ylation assay of JunB HEK293 cells expressing MYC-FLAG-tagged JunB were lysed in FAT10 IP-buffer buffer supplemented with 1 % NP40 and an immunoprecipitation was performed anti-FLAG-Agarose. Beads were washed 3 times with FAT10- IP buffer and immunoprecipitated. JunB was incubated with 10 μg FAT10, 1,25 μg FLAG-UBA6, 3 μg HIS-USE1, 50 μg GST-TRIM11 and ATP. After 60 min of incubation at 30°C, the beads were washed twice with FAT10-IP- buffer and boiled with reducing Laemmli buffer containing 10 % β-mercaptoethanol, followed by Western blot analysis with (a) a FLAG-reactive antibody conjugated to horseradish peroxidase (HRP) to detect either JunB- MYC-FLAG or FLAG-UBA6. (b) HIS-USE1 was detected with a peroxidase (POX) conjugated polyhistidine- reactive antibody. (c) GST-TRIM11 was detected with an anti-GST-antibody and (d) FAT10 was detected through a monoclonal FAT10-antibody. Input contains 10 % of JunB, recombinant FAT10, HIS-USE1, FLAG-UBA6 and GST-TRIM11 of the total protein amount used for the in vitro FAT10ylation assay. Asterisks indicate light and heavy antibody chains of the FLAG-reactive antibody used for the IP of FLAG-JunB.

Although all samples were treated under reducing conditions (10 % β-mercaptoethanol), conjugate formation between UBA6 and FAT10 could be detected in the anti-FLAG-blot (Figure 32 (a)), as well as in the anti-FAT10-blot (Figure 32 (d)), indicating a covalent linkage. The in vitro conjugation assay revealed, that JunB was not apparently FAT10ylated in vitro in the presence of recombinant FAT10, UBA6, USE1 and TRIM11, because no defined conjugate band was detectable neither on the anti-FLAG nor anti-FAT10 blot (Figure 32 (a), lane 11 and (d), lane 6).

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Interestingly, a high molecular weight smear, indicating a poly-FAT10ylation of JunB was detectable when FAT10 and its E1, E2 enzyme and TRIM11 were present (Figure 32 (a) and (d)). To argue why the in vitro FAT10ylation assay with GST-TRIM11 failed seems to be very troublesome. One reason could be that the purification of recombinant GST-TRIM11 was not satisfactory and contained still impurities and degradation products (Figure 32 (c), lane 1, input). Another problem could be the poor solubility of recombinant TRIM11.

6.2.14 JunB is conjugated with FAT10 on lysine 237

FAT10ylation is a post-translational modification that conjugates FAT10 onto specific lysine residues of substrate proteins. JunB contains 19 lysine residues that could potentially be modified by FAT10. To date, no crystal structure of JunB has been solved, which makes it difficult to predict, which lysines are exposed on the protein surface. To address the question which lysine of JunB is involved in FAT10 conjugate formation, co- immunoprecipitation assays with JunB WT, JunB lysine mutants together with FAT10 in HEK293 cells where performed, where JunB lysines (K) are mutated individually (JunB- K237R, -K267R, and -K301R) or in combination (JunB-K237R/K267R/K301R, termed JunB- K3R) to arginines (R) (constructs kindly provided by M. Piechaczyk). These mutations were chosen, because aforementioned lysines are embedded in a SUMOylation consensus sequence. Garaude et al. could show 2008 that lysine 237 is the primary SUMOylation site on JunB (Garaude et al., 2008). SUMOylation generally occurs on the lysine residue within a ψKXE consensus motif, where ψ is an aliphatic amino acid and X any amino acid (see 3.2.4). Three motifs are present in JunB, surrounding lysines 237, 267, and 301 (Figure 33). To date, no FAT10 consensus sequence has been identified but the existence of SUMOylation consensus sequences and the abundance of ubiquitin binding domains (UBDs) make the existence of such domains quite likely. Hence, we were interested, if FAT10ylation occurs on the same lysine residue to which SUMO is conjugated on JunB.

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Figure 33: Diagram depicting the major structural features of JunB JunB lysines embedded in a SUMOylation consensus motif. ψ represents an aliphatic amino acid and X any amino acid. DBD: DNA-binding domain; NLS: Nuclear Localization Signal; and bZip: basic domain-leucine zipper Figure taken from (Garaude et al., 2008).

HEK293 cells were transiently transfected with a HIS-3xFLAG-tagged FAT10 construct and with HA-tagged JunB, JunB-K237R, JunB-K267R, JunB-K301 or JunB-K3R alone, or in combination with both together. Double-transfected cells were treated with MG132 (10 µM) for 6 hours before cells were lysed to prevent conjugate degradation. Overexpressed wt or mutant HA-tagged JunB (~48 kDa) in the whole cell extract was detected in Western blots using an HA-reactive antibody (Figure 34 (a)). Moreover, lysates were directly immunoblotted with an anti-FLAG antibody to reveal the expression of 3xFLAG-HIS-tagged FAT10 (~24 kDa) as depicted in Figure 34 (b), input.

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(b)

Figure 34: JunB is conjugated with FAT10 on lysine 237 HEK293 were singly transfected with pcDNA3-JunB-HA, the single mutants pcDNA3-JunB-K237R-HA, pcDNA3- JunB-K267R-HA, pcDNA3-JunB-K301R-HA and the JunB triple mutant pcDNA3-JunB-K237R/K267R/K301R, termed JunB-K3R, or double transfected together with pcDNA3-3xFLAG-HIS-FAT10. Double transfected cells were treated with MG132 (10 µM) for 6 hours, as indicated. All samples were analyzed under reducing conditions (10% β-mercaptoethanol). After immunoprecipitation against the HA-Tag of JunB, samples were separated on 4- 12% Bis-Tris SDS gels and subjected to Western blot analysis using a HA-reactive antibody conjugated to horseradish peroxidase (a) or a FLAG-reactive antibody (b). β-actin served as a loading control.

After immunoprecipitation against the HA-Tag of JunB, samples were separated on 4-12% Bis-Tris SDS gels and subjected to Western blot analysis using a HA-reactive antibody to detect JunB (Figure 34 (a)) or a FLAG-reactive antibody (Figure 34 (b)) to detect JunB- conjugates. WT JunB and all mutant forms of JunB were detectable in a band appearing at 48 kDa, indicating that JunB WT as well as mutant forms become non-covalently linked to FAT10. Co-immunoprecipitation of FAT10 together with WT-JunB, JunB-K267R and JunB- K301R resulted in a stable conjugate double band at a height of ~66 kDa. Mutation of lysine 237, but not of lysine 267 or 301, completely abolished JunB modification by over-expressed FAT10 for the upper conjugation band (Figure 34 (b), lane 4). Moreover, the higher molecular conjugate band is neither detectable with the JunB-K237R nor the triple JunBK3R mutant, giving a strong hint that lysine 237 is implicated in FAT10 conjugate formation.

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Interestingly, also the lower conjugate band is strongly diminished when FAT10 is co- expressed together with the JunB-K3R mutant (Figure 34 (b), lane 7). As a control co- expression of FAT10∆GG with either JunB or JunB-K3R completely abolished conjugate formation (data not shown). Taken together, these results provide clear evidence, that lysine 237 in JunB is probably the major acceptor site not only for the ULM SUMO, but also for FAT10.

6.2.15 Identification of JunB in mass spectrometry analysis

To further ensure that the conjugate bands consist of JunB and FAT10 and in order to identify, if FAT10ylated JunB contains post-translational modifications, mass spectrometry analysis was performed. Moreover, we were interested if the lysine involved in conjugate formation, could be confirmed. Therefore, HEK293 cells were transiently transfected either with pcDNA3.1-HA-FAT10 and pCMV6-JunB-MYC-FLAG or left untransfected. After 24 h of ectopic protein expression, JunB was immunoprecipitated from cell lysates using anti-FLAG-agarose. Samples were boiled in reducing sample buffer containing 10 % β-mercaptoethanol and proteins were separated on 4-12 % Bis-Tris-gels. The gel was stained for 10 minutes with Coomassie (see Table 9) and destained for 30 min with destaining solution (see Table 10) and the two bands appearing at the height 66 kDa which equate to JunB-FAT10 conjugates (Figure 35 (a), indicated as (2) and (3)) were cut out and analyzed by mass spectrometry. As a negative control, gel pieces were cut out from the untransfected control lane (Figure 35 (a), lane 1) at the same height where conjugate bands are appearing (Figure 35 (a) indicated as 1.1 and 1.2) and sent to mass spectrometry analysis, as well. As a further control, samples were analyzed by Western blotting and probed with a anti-HA- reactive antibody, to ensure that the bands, appearing at the expected height in the Coomassie stained gel, coincide with the JunB-FAT10 conjugate bands detected in the Western blot (Figure 35 (b)).

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a) b)

Figure 35: Mass spectrometric analysis of the JunB-FAT10 conjugate HEK293 cells were co-transfected with 6 µg pCMV6-JunB-MYC-FLAG and 6 µg pcDNA3.1-HA-FAT10 expression plasmids, respectively or left un-transfected. After immunoprecipitation against the FLAG-Tag of JunB, samples were boiled in gel sample buffer containing 10% β-mercaptoethanol (reducing conditions) and proteins were separated on 4-12% Bis-Tris SDS gels. They were either stained with Coomassie a) or subjected to Western blot analysis using a HA-reactive antibody conjugated to horseradish peroxidase to detect conjugate formation (b). The asterisks mark the signal corresponding to antibody light or heavy chains used for immunoprecipitation. Double bands appearing (2,3) in the Coomassie gel which equates to FAT10ylated JunB at the height of ~66 kDa were excised and analyzed by mass spectrometry. As negative control bands at the same height ~66 kDa of untransfected HEK293 cells were cut out (1.1; 1.2) as well and analyzed by mass spectrometry.

Mass spectrometric analysis (Thermo LTQ Orbitrap, Proteomics facility at the University of Constance, A. Marquart) revealed that the lower (Figure 35 (2)) as well as the upper conjugate band (Figure 35 (3)) contained JunB peptide fragments. However, detailed analysis of the lower band (2) revealed, that two JunB-peptide fragments could be detected, in comparison to the upper band (3), where three peptide fragments where measured, suggesting a post-translational modification for the latter case (for mass spectrometry results see addendum, chapter 10. Unfortunately no FAT10 could be identified in the analyzed samples and therefore sent to H. Urlaub group (MPI Göttingen) for further analysis by a program called Chop'N'spice, a mass spectrometric approach that has been described to allow the identification of endogenous SUMO conjugated peptides (Hsiao et al., 2009). No JunB or FAT10 was detected in the untransfected control bands, which served as negative control (1.1 and 1.2).

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6.2.16 Post-translational modification: Phosphorylation of JunB on Serine 259?

Our previous studies demonstrated that JunB becomes stably attached to FAT10 (see chapter 6.2.3, 6.2.4 and 6.2.7), and interestingly, conjugate formation resulted in a double band appearing at the height of approximately 66 kDa. To address the issue, whether a post- translationally modified form of JunB becomes conjugated to FAT10, we considered it to be likely, that phosphorylation could be the reason.

(a)

(c)

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Figure 36: Serine259 phophorylated form of JunB becomes conjugated to FAT10 HEK293 were single transfected with MYC-FLAG-tagged JunB or HA-tagged FAT10 or double-transfected. After 24 h, cells were lysed and immunoprecipitated against the HA-tag of FAT10. Samples were boiled in gel sample buffer containing 10% β-mercaptoethanol (reducing conditions) and proteins were separated on 4-12% Bis-Tris SDS gels and subjected to Western blot analysis using a (a) polyclonal JunB antibody (ab31421) or (b) serine 259 (S259) phospho-specific JunB antibody (ab30628) to evaluate JunB expression, or (c) a HA-reactive antibody to detect FAT10 expression. β-actin served as a loading control. Arrow heads indicate JunB-FAT10 conjugates. Asterisks mark unspecific bands.

HEK293 were transiently transfected with a MYC-FLAG-tagged JunB, a HA-tagged FAT10 or double-transfected. 24 h after ectopic protein expression, samples were immunoprecipitated with HA-agarose against the HA-tag of FAT10 to accumulate FAT10 and FAT10 conjugates and samples were immunoblotted with a polyclonal JunB antibody (ab31421) (Figure 36 (a)), or a serine 259 (S259) phospho-specific JunB antibody (ab30628) (Figure 36 (b)) to detect JunB, and a HA-reactive antibody (Figure 36 (c)) to visualize FAT10 expression.

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Interestingly, co-expression of JunB together with FAT10 resulted in increased JunB expression, which was detectable in the JunB-blot ((Figure 36 (a), input) as well as in the phospho-JunB(S259) blot (Figure 36 (b), input), similar to the case, when FAT10 was induced with proinflammatory cytokine INF-γ and TNF-α (see 6.2.12). In comparison, FAT10 expression decreased significantly when co-expressed together with JunB (Figure 36 (c), input). The fact, that the highest and coincidentally the most prominent JunB band is recognized by both, the polyclonal JunB and the Ser259 phospho-specific JunB antibody suggested, that this band presents a phosphorylated form of JunB. Analysis of co-immunoprecipitated JunB and immunoblotting with either the polyclonal JunB, or phospho-Ser259 specific JunB antibody illustrate, that a JunB-FAT10 conjugate is formed apparent in a high molecular weight double band at 66 kDa. Interestingly, this double band was not detectable, when samples were analyzed by immunoblotting with a Ser79 phospho- specific antibody (data not shown), suggesting a specific interaction of FAT10 with a Ser259 phosphorylated form of JunB.

6.2.17 FAT10 and JunB co-localize at the nuclear membrane and in the cytosol

Our previous experiments revealed clear evidence that FAT10 becomes stably attached to JunB and inhibition of the proteasome led to a significant accumulation of the conjugate. To further investigate the localization and fate of either JunB or FAT10 when co-expressed and to analyze the role of FAT10ylation in this process, we performed confocal laser- scanning microscopy. Therefore, we transfected HEK293 cells with either a pcDNA3.1-HA- FAT10 or pCMV6-JunB-MYC-FLAG construct alone, or co-transfected them together, treated them with or without MG132 for 6 hours and carried out a series of co-localization experiments using a directly labelled HA-coupled Alexa Fluor 488 antibody to visualize HA- FAT10 expression and a rabbit polyclonal antibody to JunB (ab31421) followed by a Alexa Fluor 546-coupled secondary goat-anti-mouse antibody, to stain for JunB.

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Figure 37: FAT10 modification leads JunB to the nuclear membrane HEK293 cells were transiently transfected with MYC-FLAG-tagged JunB and a HA-tagged FAT10 construct alone or co-transfected together and treated with or without MG132 (10 µM) for 6 h and fixed with 4 % PFA. (a) HA- FAT10 transfected cells were stained with a 6,85 µg ml-1 dilution of directly labelled HA-coupled Alexa Fluor 488 antibody (green) and (b) treated additionally with MG132. (c) JunB transfected cells were stained with a 5 µg ml-1 dilution of a rabbit polyclonal antibody to JunB (ab31421), followed by a Alexa Fluor 546-coupled secondary goat- anti-mouse antibody (red) and (d) treated additionally with MG132. (e) Double transfected cells were treated subsequently with Alexa488-HA antibody and a rabbit polyclonal Ambra1 (ab31421) antibody followed by a Alexa Fluor 546-coupled secondary goat-anti-mouse antibody and (f) treated additionally with MG132. Confocal microscopy images are shown. Scale bar: 25 µm. Images are representatives of several cells examined in three independent experiments.

In singly transfected cells, FAT10 was evenly distributed throughout the cytosol and showed varying degrees of localization to the nucleus (see Figure 37 (a)). Kalveram et al. (Kalveram et al., 2008) previously described, that FAT10 localizes in aggresomal structures under proteasomal inhibiton, which unfortunately could not be confirmed here (see Figure 37 (b)). JunB by contrast, was completely excluded from the cytosol and showed thoroughly nucleolar localization (see Figure 37 (c)). Strikingly, treatment with MG132 resulted in JunB dislocation towards the nuclear membrane and to the cytosol. Moreover, cells seem to be stressed, which is apparent in increased membrane ruffle formation (see Figure 37 (d)). Interestingly, co-expression of FAT10 together with JunB resulted in a clear co-localization of both proteins in favor to the nuclear membrane (see Figure 37 (e)). In double transfected and MG132 treated cells, cells showed again increased membrane ruffle formation. Interestingly, JunB dislocalizes from the nucleus to the cytosol (visible in modified nucleus) like previously seen in Figure 37 (d) and JunB co-localized with FAT10 near the nuclear membrane. However, for the majority of cells, JunB and FAT10 co- localization was predominantly detectable in the cytosol (see Figure 37 (f)). Unfortunately, a quantitative analysis could not be performed due to a lack of time. Our results substantiate the precedent data that JunB and FAT10 interact and we strongly suggest that modification with FAT10 regulate JunB localization and function. Moreover, we assume that interaction of JunB and FAT10 occurs preferentially at the nuclear membrane, which requires the translocation of JunB away from the nucleus towards the nucleolic membrane. Strikingly, treatment with MG132 leads to a cumulative displacement of JunB to the nuclear membrane and into the cytosol which implicate as functional consequence, that the transcription of JunB transactivating genes could be impaired. Further work will determine whether FAT10ylation is a cause or a consequence of this localization. This issue was further investigated in reporter assays to reveal the impact of FAT10ylation on JunB transcriptional activities on minimal AP-1 driven reporter genes (see chapter 6.2.18).

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6.2.18 FAT10ylation controls JunB transcriptional activities on minimal AP-1 driven reporter genes

In order to understand, if JunB FAT10ylation has an impact on biological functions of JunB we next aimed to determine, if transcription is affected through this post-translational modification. JunB belongs to the AP-1 family, which collectively describes a class of structurally and functionally related proteins, characterized by a basic DNA-binding domain and a basic leucine zipper (bZIP) dimerization motif which require dimerization before binding to a common DNA target sites, called TPA response elements (TRE) (Wagner, 2001). Previously Garaude et al. (Garaude et al., 2008) described, that post-translational modification of JunB with SUMO regulates the transcriptional activity of JunB in T lymphocytes and plays a critical role in T cell activation. In comparison to c-Jun, JunB has a ten-fold decreased activity to activate AP-1-responsive genes containing single AP-1 binding sites, due to a small number of amino acid changes between its DNA-binding and dimerization motifs (Deng and Karin, 1993). But strikingly, JunB appears to be as effective as c-Jun in trans-activating reporter genes containing multiple AP-1 binding sites which suggests, that trans-activation by JunB may require synergistic interactions between multiple homodimers bound to adjacent sites (Angel and Karin, 1991; Chiu et al., 1989). To address either potential similarities or differences in biological functions, when JunB becomes attached to FAT10, we conducted a luciferase reporter assay in Jurkat cells on reporter genes containing 4x TRE binding sites, namely AP-1 (4xTRE). Our previous results indicated that K237 was the primary site for FAT10ylation; when this site was mutated to arginine, we detected a complete abrogation of the higher conjugate band of FAT10ylated JunB. The triple mutant JunB-K3R was barely FAT10ylated, as not only the upper conjugate completely disappeared, but also the lower conjugate band was barely detectable (see chapter 6.2.14). Therefore, we decided to use JunB wt as positive, or the triple mutant JunB- K3R as negative control, to investigate the role of FAT10ylation on the transcriptional activity of JunB. Furthermore, co-transfection with FAT10∆GG served as a further negative control, where JunB FAT10ylation is prevented. Twenty million Jurkat cells were transfected via electroporation with 5 µg expression vectors encoding for HA-tagged JunB wt, HA-tagged JunB-K3R mutant, FLAG-tagged FAT10 or FLAG-tagged FAT10∆GG. In each experiment, cells were transfected with the same total amount of 12 µg DNA by adding the required quantities of pcDNA3 empty vector, along with 2 µg of a reporter gene controlled by the AP-1 (4xTRE-luc) promoter or a pGl3 vector which served as a normalization vector, as indicated. 1/600 TK-Renilla-Luc was used as a control for transfection efficiency and luciferase activity is expressed relative to Renilla luciferase.

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Two days after transfection, cell extracts were prepared in 100 μl Passive Lysis Buffer (PLB) Dual-Luciferase® Reporter (DLR™) Assay System (see 5.1.2) and luciferase activity was determined using Berthold Lumat LB9501.

Figure 38: FAT10ylation affects JunB transcriptional activity on minimal AP-1-driven reporter genes in JURKAT cells Twenty million Jurkat cells were co-transfected with 5 µg of expression vectors for HA-tagged JunB wt, JunB- K3R, FLAG-tagged FAT10 or FAT10∆GG, as indicated. In each experiment, cells were transfected with the same total amount of 12 µg DNA by adding the required quantities of pcDNA3 empty vector, together with 2 µg of a reporter gene controlled by the AP-1 (4xTRE-luc) promoter, as indicated. Transfection of 2 µg pGl3 vector together with 10 µg pcDNA3 served for normalization. 1/600 TK-Renilla-Luc was used as a control for transfection efficiency and luciferase activity is expressed relative to Renilla luciferase. Two days after transfection, luciferase activities were measured. The data are presented as the mean ± SD of five independent experiments. n.s indicates not significant. The asterisks represent the level of significance as calculated with a paired two-tail P value test.

JunB wt and JunB-K3R showed indistinguishable activity on the reference AP-1 reporter gene, driven by four canonical binding sites upstream of the TATA box, namely AP-1 (4xTRE-luc), indicating that the mutation did not affect basal functions. In comparison, neither FAT10 nor FAT10ΔGG trans-activated the AP-1 reporter gene, showing that binding of either JunB wt or JunB-K3R to the 4xTRE sites is specific. Strikingly, co-expression of JunB wt and FAT10 resulted in a significantly decreased reporter activity in comparison to JunB alone. Co-expression of either mutant JunB-K3R with FAT10 or FAT10ΔGG, or JunB wt and FAT10ΔGG (negative controls) resulted only in a slightly decreased activity in comparison JunB alone, suggesting that JunB FAT10ylation specifically provoke the reduced

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Results activity. Taken together, these results suggest that FAT10ylation repress JunB transcriptional activity on AP-1 driven promoter with minimal 4xTRE binding sites. To evaluate the effect of FAT10 conjugation for JunB transactivating capabilities in a different cell line, we performed co-transfection experiments with the AP-1 (4xTRE) reporter in HEK293 cells. 3 x 105 HEK293 cells were transiently transfected with 1,5 µg vectors expressing for HA-tagged JunB wt, HA-tagged JunB-K3R mutant, FLAG-tagged FAT10 or FLAG-tagged FAT10∆GG. In each experiment, cells were transfected with the same total amount of 4,5 µg DNA by adding the required quantities of pcDNA3 empty vector, along with 1 µg of a reporter gene controlled by the AP-1 (4xTRE-luc) promoter or a pGl3 vector which served as a normalization vector, as indicated. 1/600 TK-Renilla-Luc was used as a control for transfection efficiency and luciferase activity is presented relative to Renilla luciferase. One day after transfection, cell extracts were prepared in 500 μl Passive Lysis Buffer (PLB) Dual-Luciferase® Reporter (DLR™) Assay System (see 5.1.2) and luciferase activity was determined using Berthold Lumat LB9501.

Figure 39: FAT10ylation affects JunB transcriptional activity on minimal AP-1-driven reporter genes in HEK293 cells 3x105 HEK293 cells were co- transfected with 1,5 µg of expression vectors for HA-tagged JunB wt, JunB-K3R, FLAG-tagged FAT10 or FAT10∆GG, as indicated. In each experiment, cells were transfected with the same total amount of 4,5 µg DNA by adding the required quantities of pcDNA3 empty vector, together with 1 µg of a reporter gene controlled by the AP-1 (4xTRE-luc) promoter or a pGl3 vector which served as a normalization vector. Transfection of 1 µg pGl3 vector together with 3 µg pcDNA3 served for normalization. 1/600 TK-Renilla-Luc was used as a control for transfection efficiency and luciferase activity is expressed relative to Renilla luciferase. 24 h post-transfection, luciferase activities were measured. The data are presented as the mean ± SD of five independent experiments. n.s indicates not significant. The asterisks represent the level of significance as calculated with a paired two-tail P value test. 130

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In HEK293 cells, JunB-K3R transactivated reporter genes driven by these composite sites at the same level as JunB wt. Like previously seen in JURKAT cells, co-expression of JunB together with FAT10 resulted in significantly diminished AP-1 promoter activity and co- expression of either mutant JunB-K3R with FAT10 or FAT10ΔGG, or JunB wt and FAT10ΔGG (negative controls) resulted in a similar transactivation of AP-1 (4xTRE), as JunB wt. Taken all the experiments together it seems, that JunB-FAT10 conjugation (as seen in previous experiments) has a direct consequence on transactivating activities of JunB under these conditions.

6.2.19 Interaction of Ambra1 and TRIM11 in a human cell line

In a yeast two-hybrid screen with TRIM11 as the bait protein, the activating molecule in beclin-1-regulated autophagy (Ambra1) was among the putative interaction partners of TRIM11 (see chapter 6.2.1, Figure 19). Ambra1 was a very interesting candidate to investigate further, because sequencing revealed that eleven of 96 positive interacting clones encoded for full length Ambra1. In addition Ambra1 was identified in a Mass spectrometric analysis of FAT10 interacting proteins (A. Aichem et al., submitted). Due to the clear interaction between Ambra1 and the E3 ligase TRIM11 in the screen it was decided to further investigate the link between the two proteins, especially in terms of acting as a FAT10 specific E3 ligase, which mediates ligation of FAT10 to the putative substrate Ambra1. To investigate, if the putative interaction between TRIM11 and Ambra1 occurs under in vivo conditions, Ambra1 and TRIM11 were further tested for their interaction in the mammalian cell line HEK293. A MYC-FLAG-tagged Ambra1 and HIS-tagged TRIM11 construct were transiently transfected in HEK293 cells to carry out co-immunoprecipitation experiments. To prevent proteasomal degradation of either HIS-TRIM11 or Ambra1-MYC-FLAG, cells were treated additionally with or without MG132 for 6 h before cell lysis.

24 h after transfection, cells were lysed and whole-cell lysates were subjected to co- immunoprecipitation assays using anti-FLAG Agarose, followed by SDS-PAGE and Western blotting with anti-FLAG or anti-6HIS antibodies. Whole-cell lysates were also directly immunoblotted with anti-FLAG and anti-6HIS to reveal the expression of Ambra1-MYC-FLAG and HIS-TRIM11. As a loading control for the prepared lysates, β-actin was used. All samples were analyzed under reducing conditions (10 % β-mercaptoethanol).

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Figure 40: TRIM11 interacts with Ambra1 in HEK293 cells

HEK293 cells were transiently transfected with either pCMV6-Ambra1-MYC-FLAG or pcDNA3-HIS/-A-TRIM11, or double transfected, as indicated. Cells were pre-treated with/without the proteasome inhibitor MG132 for 6 hours. 24h after transient transfection with the respective plasmids, cells were lysed with FAT10 IP-buffer and samples were loaded on SDS gels under reducing conditions (10 % ß-mercaptoethanol) immunoblotted and designated as input. After immunoprecipitation against the FLAG-Tag of Ambra1-MYC-FLAG samples were separated on 4-12% Bis-Tris SDS gels and subjected to Western blot analysis using a (a) FLAG-reactive antibody conjugated to horseradish peroxidase (HRP) to reveal Ambra1 expression. (b) HIS-TRIM11 expression (input) and co- immunoprecipitated TRIM11 was detected with a peroxidase (POX) conjugated polyhistidine-reactive antibody. β- actin served as a loading control.

Immunoblotting with a FLAG-reactive antibody detected Ambra1 at a size of approximately 130 kDa (Figure 40 (a) lane 2-4, input). Co-expression of TRIM11 strongly decreased the Ambra1 protein level, which could be rescued with MG132 treatment (Figure 40 (a) lane 3 and 4, input). The same outcome was true for immunoprecipitated Ambra1 (Figure 40 (a) lane 6-8, IP: anti-FLAG). HIS-tagged TRIM11 could be detected with an anti-6HIS-antibody at a size of 52 kDa (Figure 40 (b), lane 2-4 (input)). The expression of TRIM11 decreased when cells were double-transfected together with Ambra1 (Figure 40 (b), lane 3). However, the total protein amount was also less in this case (Figure 40 (b), WB: anti-ß-actin, lane 3). TRIM11 expression could on the other hand be rescued again under proteasomal inhibition (Figure 40 (b), lane 4).

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A weak interaction between TRIM11 and Ambra1 could be detected after immunoprecipitation against Ambra1 and Western blot analysis against the HIS-tag of TRIM11 (Figure 40 (b), lane 7, IP: anti-FLAG), which could be slightly enhanced, when cells were pre-treated with MG132 (Figure 40 (b), lane 8, IP: anti-FLAG). These results affirm the yeast two-hybrid results, indicating that interaction between Ambra1 and TRIM11 occurs in mammalian cells.

6.2.20 Ambra1 interacts non-covalently with ubiquitin and FAT10

Apart from being an interaction partner of Ambra1, we were further interested in the role of TRIM11 as an E3 ligase, mediating the ligation of either ubiquitin or FAT10 to Ambra1. To investigate, if interaction between Ambra1 with either ubiquitin or FAT10 takes place, co- immunoprecipitation assays were performed. These experiments should address the question if Ambra1 either interacts non-covalently with ubiquitin or FAT10 or if the two proteins get linked to each other through an isopeptide bond. Therefore, Ambra1-MYC-FLAG was expressed together with HA-Ubiquitin, HA-FAT10 and HIS-TRIM11 in HEK293 cells and treated with or without MG132 for 6 hours before cell lysis. 24 h after ectopic protein expression cells were lysed and lysates were subjected to co- immunoprecipitation assays using anti-HA agarose to accumulate ubiquitin, FAT10 or conjugates accordingly, followed by SDS-PAGE and Western blotting with anti-HA or anti- FLAG antibodies. Whole-cell lysates were also directly immunoblotted with anti-HA, anti- 6HIS and anti-FLAG antibodies to reveal the expression of either HA-Ubiquitin, HA-FAT10, HIS-TRIM11 or Ambra1-MYC-FLAG.

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Figure 41: Co-immunoprecipitation of Ambra1 and Ubiquitin or FAT10 in HEK293 cells HEK293 cells were transiently transfected with pCMV6-Ambra1-MYC-FLAG, pcDNA3.1-HA-Ubiquitin, pcDNA3.1- HA-FAT10 and pcDNA3-HIS-TRIM11. After immunoprecipitation against the HA-Tag of ubiquitin and FAT10, samples were separated on 4-12% Bis-Tris SDS gels and subjected to Western blot analysis (a) with a horseradish peroxidase (HRP)-conjugated HA-reactive antibody. (b) The expression of HIS-TRIM11 was detected with a peroxidase (POX) conjugated polyhistidine-reactive antibody. (c) The expression of FLAG-Ambra1 was detected with a peroxidase-FLAG-reactive antibody. Co-immunoprecipitation of either ubiquitin or FAT10 with Ambra1 could be detected after co-immunoprecipitation against the HA-Tag of ubiquitin or FAT10 and Western blot analysis against the FLAG-tag of Ambra1. (d) Cells were treated with or without MG132 (10 µM) for 6 hours and the expression of FLAG-Ambra1 was detected with a FLAG-reactive peroxidase-coupled antibody. Co- immunoprecipitation of either ubiquitin or FAT10 with Ambra1 could be detected after co-immunoprecipitation against the HA-Tag of ubiquitin or FAT10 and Western blot analysis against the FLAG-tag of Ambra1. All samples were analyzed under reducing conditions (10 % β-mercaptoethanol). β-actin served as a loading control.

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Input controls illustrate, that all proteins were expressed (Figure 41 (a), (b) and (c)). Co- expression of ubiquitin and Ambra1 did neither change the amount of monomeric ubiquitin, nor the pattern of formed ubiquitin conjugates (Figure 41 (a), lane 2+3, input). Co-expression of ubiquitin with TRIM11, or together with Ambra1 and TRIM11, decreased the amount of monomeric ubiquitin and ubiquitin conjugates shifted to higher molecular weight bands (see Figure 41 (a), lane 4+5, input), which however could not be observed after immunoprecipitation against the HA-tag of ubiquitin (Figure 41 (a), lane 4+5, IP: anti-HA). A reduced amount of monomeric FAT10 was also detectable, when co-expressed with either TRIM11 or TRIM11 and Ambra1 (see Figure 41 (a), lane 8+9, input), which was even more obvious after immunoprecipitation against the HA-Tag of FAT10 (Figure 41 (a), lane 8+9, IP: anti-HA). Ambra1 was co-immunoprecipitated with ubiquitin and FAT10 (Figure 41 (c), lane 3-6, IP: anti-HA), but no modification with either ubiquitin or FAT10 was detectable when TRIM11 was overexpressed, suggesting, that Ambra1 becomes non-covalently linked to ubiquitin or FAT10. Interestingly, overexpression of TRIM11 decreased the protein amount of Ambra1, but did not result in covalent conjugate formation with ubiquitin or FAT10. It has been shown that the covalent modification of proteins with FAT10 serves as a signal for subsequent proteasomal degradation (Hipp et al., 2005). To further investigate if Ambra1 forms a conjugate with either ubiquitin or FAT10, the experiment was repeated under conditions of proteasome inhibition. In case that Ambra1 would be a substrate of ubiquitin or FAT10, the amount of ubiquitinated or FAT10ylated Ambra1 should increase or at least be better detectable under proteasomal inhibition. MG132 treatment could increase the amount of co-immunoprecipitated Ambra1 with ubiquitin or FAT10, respectively (Figure 41 (d), IP: anti-HA), whereas co-expression of HIS-TRIM11 decreased the amount of co- immunoprecipitated Ambra1. However, also the amount of ubiquitin, FAT10 and TRIM11 increased under conditions of proteasome inhibition (data not shown). Taken together, our data show that Ambra1 becomes co-immunoprecipitated with ubiquitin as well as FAT10, and MG132 treatment increased the amount of co-immunoprecipitated Ambra1.

6.2.21 The proteasome is involved in Ambra1 degradation

Ambra1 has been described to be an indispensible regulator of autophagy initiation and is thereby involved in the process of cellular degradation (Di Bartolomeo et al., 2010; Fimia et al., 2007; Liang et al., 1999; Sun et al., 2009), but hitherto nothing is known about how Ambra1 itself becomes degraded and no data were available about the protein half life of Ambra1.

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To determine the protein half life of Ambra1 we performed cycloheximide chase experiments. HEK293 cells were transiently transfected with a MYC-FLAG-tagged Ambra1 construct and cycloheximide (50 µg ml-1) was added for the indicated time periods for 2.5 and 5 h or left untreated. Besides, cells were treated with or without MG132 (10 µM) for 6 h.

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relative Ambra1 expression 0 0 2.5 5 5 + 6 h MG132 CHX treatment [h]

Figure 42: Determination of the turnover rate of ectopically expressed Ambra1 HEK293 cells were transiently transfected with a pCMV6-Ambra1-MYC-FLAG plasmid. Before cell lysis, cells were treated with the proteasome inhibitor MG132 (10 µM) for 6 hours and with cycloheximide (50 µg ml-1) for different time periods, as indicated. Whole cell lysates were subjected to Western blot analysis using a directly coupled horseradish peroxidase (HRP) -linked anti-FLAG mAb to evaluate Ambra1 expression. All samples were analyzed under reducing conditions (10% β-mercaptoethanol). β-actin served as a loading control. One out of two experiments with similar outcome is shown. ECL signals of all experiments were quantified with Quantity One Software (BioRad). The ECL signal of Ambra1-MYC-FLAG transfected cells without cycloheximide treatment was set to unity. Mean values ± SEM of two independent experiments are shown.

The cycloheximide data illustrate, that the protein turnover of Ambra1 only slightly decrease to ~80 % after 5 h of cycloheximide treatment. Surprisingly, MG132 treatment led to a considerable accumulation of Ambra1 to 160 %, which clearly indicate an involvement of the proteasome in Ambra1 degradation. We next aimed to determine the protein turnover rate of Ambra1 in presence of FAT10. HEK293 cells were transiently transfected with a HA-tagged FAT10 and MYC-FLAG-tagged Ambra1 construct. Cycloheximide (50 µg ml-1) was added for the indicated time periods for 2.5 and 5 h or left untreated. Moreover, cells were treated with or without MG132 (10 µM) for 6 h. Whole cell lysates from transfected cells were analyzed for expression of the respective proteins with a FLAG-reactive antibody, to detect MYC-FLAG tagged Ambra1 and an anti-HA antibody to reveal HA-FAT10 expression.

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Figure 43: Determination of Ambra1 turnover rate in the presence of FAT10 HEK293 cells were transiently transfected with (a) pCMV6-Ambra1-MYC-FLAG and pcDNA3.1-HA-FAT10 plasmids. Before cell lysis, cells were treated with the proteasome inhibitor MG132 (10 µM) for 6 hours and with cycloheximide (50 µg ml-1) for different time periods, as indicated. Whole cell lysates were subjected to Western analysis using a directly coupled horseradish peroxidase (HRP) -linked anti-FLAG mAb to evaluate Ambra1 expression and an HA-reactive antibody to evaluate FAT10 expression. All samples were analyzed under reducing conditions (10% β-mercaptoethanol). β-actin served as a loading control. One out of two experiments with similar outcome is shown. The ECL signal of Ambra1-MYC-FLAG and pcDNA3.1-HA-FAT10 transfected cells without cycloheximide treatment was set to unity. Mean values ± SEM of two independent experiments is shown.

Surprisingly, Ambra1 turnover in presence of FAT10 illustrates a faster degradation progress than in absence of FAT10 (see Figure 42). The half life of Ambra1 in presence of FAT10 is about 2.5 h and interestingly, almost no rescue with MG132 could be observed (see Figure 43). These results suggest, that co-expression of FAT10 alters the degradation rate of Ambra1 and interaction leads presumably to another degradation machinery than the proteasome. In contrast, the FAT10 half life in presence of Ambra1 was calculated to be approximately 1 h and MG132 treatment led to an increased accumulation of monomeric FAT10. These results indicate that FAT10 co-expression leads to accelerated Ambra1 protein turnover, whereby the turnover rate of monomeric FAT10 is hardly altered, when Ambra1 is co-expressed. Next, we aimed to determine the Ambra1 turnover in presence of FAT10 and TRIM11. HEK293 cells were transiently transfected with an HA-tagged FAT10, a MYC-FLAG-tagged Ambra1, together with a HIS-tagged TRIM11 construct. Cycloheximide (50 µg ml-1) was added for the indicated time periods for 2.5 and 5 h or left untreated. Further, cells were treated with or without MG132 (10 µM) for 6 h.

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Whole cell lysates from transfected cells were analyzed for expression of the respective proteins with a FLAG-reactive antibody, to detect MYC-FLAG tagged Ambra1 (Figure 44 (a)), anti-HA antibody to reveal HA-FAT10 expression (Figure 44 (b)) and an anti-6HIS-Pox antibody to visualize HIS-TRIM11 expression (Figure 44 (c)).

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Figure 44: Determination of the turnover rate of ectopically expressed Ambra1 in the presence of FAT10 and TRIM11 HEK293 cells were transiently transfected with pCMV6-Ambra1-MYC-FLAG, pcDNA3.1-HA-FAT10, together with pcDNA3-HIS/-A-TRIM11. Before cell lysis, cells were treated with the proteasome inhibitor MG132 (10 µM) for 6 hours and cycloheximide (50 µg ml-1) was added for different time periods, as indicated. Whole cell lysates were subjected to Western blot analysis using a directly coupled horseradish peroxidase (HRP) -linked anti-FLAG mAb to evaluate Ambra1 expression, an HA-reactive antibody to evaluate FAT10 expression and a 6HIS-POX antibody to determine HIS-TRIM11 expression. All samples were analyzed under reducing conditions (10% β- mercaptoethanol). β-actin served as a loading control. One out of two experiments with similar outcome is shown. The ECL signal of Ambra1-MYC-FLAG, pcDNA3.1-HA-FAT10 and pcDNA3-HIS/-A-TRIM11 transfected cells without cycloheximide treatment was set to unity. Mean values ± SEM of two independent experiments is shown.

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The cycloheximide data illustrate, that the protein turnover Ambra1 in presence of FAT10 and TRIM11 is similar compared to the degradation rate when Ambra1 alone was ectopically expressed in the cell (see Figure 42). 5 h after cycloheximide addition the protein level decreased to ~80% and strikingly, MG132 treatment led to an enhancement of Ambra1 to 140 % (see Figure 44 (a)), which again clearly indicate an involvement of the proteasome in Ambra1 degradation, in contrast to Ambra1 when co-expressed together with FAT10, where no rescue of Ambra1 after MG132 treatment could be observed (see Figure 43, lane 4). The role of TRIM11 in this process is currently under investigation and needs to be clarified. These data let strongly suggest, that Ambra1 becomes degraded through different pathways, whereby the proteasome is presumably one of the degradation machineries engaged in Ambra1 depletion.

6.2.22 Ambra1 co-localizes with FAT10 in punctuated structures

In co-immunoprecipitation assays we could clearly show, that Ambra1 co-immunoprecipitates with either ubiquitin or FAT10 in a non-covalent manner. The subcellular localization of Ambra1 has been determined to occur in cytoplasmic vesicle and autophagosomes. FAT10 has been suggested to be involved in autophagic processes given that p62/SQSTM, a autophagosomal receptor and signaling adapter was found to be covalently as well as non-covalently linked to FAT10 (A. Aichem et al., manuscript submitted). Moreover, Kalveram et al. showed a localization of FAT10 in aggresomes. FAT10 interacts and co-localizes with histone deacetylase 6 (HDAC6) in aggresomes under conditions of proteasome inhibition in a microtubule dependent manner, suggesting that HDAC6 functions as a linker between the dynein motor complex and FAT10 and FAT10ylated protein cargos (Kalveram et al., 2008). To further investigate the localization and fate of either Ambra1 or FAT10 when co- expressed together and to analyze the role of non-covalent interaction in this process, we conducted confocal laser-scanning microscopy. We transfected HEK293 cells with either a pcDNA3.1-HA-FAT10 or pCMV6-Ambra1-MYC-FLAG construct alone, or co-transfected them together. Cells were treated with or without MG132 for 6 hours to inhibit the proteasome or amino acid starved for 4 h, to induce autophagy and a series of co- localization experiments were carried out, using a directly labelled HA-coupled Alexa Fluor 488 antibody to visualize HA-FAT10 expression and a rabbit polyclonal antibody to Ambra1 (ab59141) followed by a Alexa Fluor 546-coupled secondary goat-anti-mouse antibody, to stain for Ambra1.

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Figure 45: Co-localization of FAT10 and Ambra1 293 cells were transiently transfected with pCMV6-Ambra1-MYC-FLAG or pcDNA3.1-HA-FAT10 alone or were co-transfected together and fixed with 4 % PFA. (a) FAT10 transfected cells were stained with a 6,85 µg/ml dilution of directly labelled Alexa488-HA antibody (green). (b) Ambra1 transfected cells were stained with a 5 µg/ml dilution of a rabbit polyclonal antibody to Ambra1 (ab59141) followed by an Alexa Fluor 546-coupled secondary goat-anti-mouse antibody. (c) Double transfected cells were treated subsequently with Alexa488-HA antibody and rabbit polyclonal antibody to Ambra1 (ab59141), followed by an Alexa Fluor 546-coupled secondary goat-anti-mouse antibody. Confocal microscopy images are shown. Scale bar: 25 µm. Images are representatives of several cells examined in two independent experiments.

In singly transfected cells, FAT10 was evenly distributed throughout the cytosol and nucleus (see Figure 45 (a)). Ambra1 was detectable primarily in the cytosol and showed varying degrees of localization to the nucleus. Moreover, formation of punctuated structures, presumably autophagosomes, mainly in the cytosol, was detectable in Ambra1 transfected cells (see Figure 45 (b)). Strikingly, co-expression of FAT10 and Ambra1 resulted in a clear co-localization of Ambra1 and FAT10 in punctuated structures mainly in the cytosol, suggesting that Ambra1 and FAT10 indeed interact and this leads to their trans-location into punctuated structures.

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(a) (b)

(c)

Figure 46: Co-localization of FAT10 and Ambra1 after serum starvation 293 cells were transiently transfected with pCMV6- Ambra1-MYC-FLAG or pcDNA3.1-HA-FAT10 alone or were co-transfected together, starved for 4 h in Earle´s Balanced salt solution and fixed with 4 % PFA. (a) HA-FAT10 transfected cells were stained with a 6,85 µg/ml dilution of directly labelled Alexa488-HA antibody (green). (b) Ambra1-MYC-FLAG transfected cells were stained with a 5 µg/ml dilution of a rabbit polyclonal antibody to Ambra1 (ab59141) followed by an Alexa Fluor 546-coupled secondary goat-anti-mouse antibody. (c) Doubly transfected cells were treated subsequently with Alexa488-HA antibody and rabbit polyclonal antibody to Ambra1 (ab59141), followed by an Alexa Fluor 546-coupled secondary goat-anti-mouse antibody. Confocal microscopy images are shown. Scale bar: 25 µm. Images are representatives of several cells examined in two independent experiments.

We next aimed to determine, if autophagy induction through amino acid starvation changes the localization of FAT10 or Ambra1. In singly transfected and amino acid starved cells, FAT10 was evenly distributed throughout the cytosol and nucleus (see Figure 46 (a)), showing similar distribution in comparison to non-starved cells (see Figure 45 (a)). Previous reports indicated, that autophagy induction results in an increased localization of Ambra1 in the perinuclear region and co-localization studies revealed that a high percentage of Ambra1 relocates to the endoplasmic reticulum, where it partially co-localizes with the omegasomes, the sites of autophagosome production (Di Bartolomeo et al., 2010; Fimia et al., 2011).

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Our data show, that Ambra1 was detectable primarily in the cytosol and showed increased formation of punctuated structures after amino-acid starvation (see Figure 46 (b)). Co-expression of FAT10 and Ambra1 resulted in a clear co-localization of Ambra1 and FAT10 in punctuated structures mainly in the cytosol (see Figure 46 (c)), similar to the case when FAT10 and Ambra1 were co-expressed in unstarved cells (see Figure 45 (c)).

(a) (b)

(c)

Figure 47: Co-localization of FAT10 and Ambra1 after MG132 treatment 293 cells were transiently transfected with constructs encoding for Ambra1-MYC-FLAG or HA-FAT10 or were co- transfected together and treated with MG132 (10 μM) for 6 h and subsequently fixed with 4 % PFA. (a) HA-FAT10 transfected cells were stained with a 6,85 µg/ml dilution of directly labelled Alexa488-HA antibody (green). (b) Ambra1-MYC-FLAG transfected cells were stained with a 5 µg/ml dilution of a rabbit polyclonal antibody to Ambra1 (ab59141) followed by an Alexa Fluor 546-coupled secondary goat-anti-mouse antibody. (c) Double transfected cells were treated subsequently with Alexa488-HA antibody and rabbit polyclonal antibody to Ambra1 (ab59141), followed by an Alexa Fluor 546-coupled secondary goat-anti-mouse antibody. Confocal microscopy images are shown. Scale bar: 25 µm. Images are representatives of several cells examined in two independent experiments.

Our previous data indicated that the proteasome is involved in Ambra1 as well as FAT10 degradation (see chapter 6.2.21). To further investigate, if proteasome inhibition has an impact on either FAT10 or Ambra1 localization or distribution in the cell, HEK293 transfected cells were treated with MG132 (10 μM) for 6 h.

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Results

Singly transfected cells expressing FAT10 (see Figure 47 (a)) or Ambra1 (see Figure 47 (b)) reveal an indistinguishable localization pattern compared to untreated cells (see Figure 45 (a) + (b)). Co-transfection of Ambra1 and FAT10 led to a clear co-localization in punctuated structures widely distributed in the cytosol as well as in the nucleus (see Figure 47 (c)) in contrast to untreated or starved cells, where these structures were mainly detectable in the cytosol. To investigate the subcellular distribution of either FAT10 or Ambra1 when the proteasome is inhibited and additionally autophagy is induced, we treated Ambra1, FAT10 or co-transfected cells for 6 with MG132 and subsequently starved the cells in amino acid and serum-free medium for 4 h.

(a) (b)

(c)

Figure 48: Co-localization of FAT10 and Ambra1 after amino acid starvation and MG132 treatment 293 cells were transiently transfected with pCMV6-Ambra1-MYC-FLAG or pcDNA3.1-HA-FAT10 constructs alone or were o-transfected together, treated with MG132 (10 μM) for 6 h, starved for 4 h in Earle´s Balanced salt solution and fixed with 4 % PFA. (a) HA-FAT10 transfected cells were stained with a 6,85 µg/ml dilution of directly labelled Alexa488-HA antibody (green). (b) Ambra1-MYC-FLAG transfected cells were stained with a 5 µg/ml dilution of a rabbit polyclonal antibody to Ambra1 (ab59141) followed by an Alexa Fluor 546-coupled secondary goat-anti-mouse antibody. (c) Doubly transfected cells were treated subsequently with Alexa488-HA antibody and rabbit polyclonal antibody to Ambra1 (ab59141), followed by an Alexa Fluor 546-coupled secondary goat-anti- mouse antibody. Confocal microscopy images are shown. Scale bar: 25 µm. Images are representatives of several cells examined in two independent experiments.

144

Results

In singly transfected, MG132 treated and amino acid starved cells, FAT10 was evenly distributed throughout the cytosol and nucleus (see Figure 48 (a)). Increased formation of punctuated structures was visible in Ambra1 transfected cells (see Figure 48 (b)), distributed in the cytosol. Compared to MG132 treated or serum starved cells, appearing punctae were strongly augmented in size. Co-expression of FAT10 and Ambra1 resulted in a clear co-localization of Ambra1 and FAT10 in punctuated structures mainly in the nucleus (see Figure 48 (c)), and these structures cannot consist of autophagosomes. Our results substantiate the precedent data that Ambra1 and FAT10 interact in cells and these data strongly suggest that interaction of Ambra1 and FAT10 regulates localization (co-localization of FAT10 and Ambra1 in punctuated structures) and probably function of both proteins. Unfortunately, no quantitative analysis for all microscopy experiments was carried out due to time limitations. However, all microscopy results verified the interaction of Ambra1 and FAT10, as has previously been determined in co-immunoprecipitation assays.

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Discussion

7 Discussion

FAT10, a very young member of the ubiquitin protein family, is encoded in the major histocompatibility complex class I locus and is synergistically inducible with the proinflammatory cytokines IFN-γ and TNF-α in cells of nearly every tissue origin. It consists of two ubiquitin-like domains, which are connected by a short linker, and bears a free di- glycine-motif at its C-terminus through which it can become covalently conjugated to target proteins (Chiu et al., 2007; Raasi et al., 2001). FAT10 is so interesting to investigate because it is beside ubiquitin so far the only described ubiquitin-like modifier (ULM), where conjugation to target proteins serves as a signal for proteasomal degradation (Hipp et al., 2005). Moreover, FAT10 was shown to be as potent as ubiquitin in targeting artificial fusion proteins for proteasomal degradation (Hipp et al., 2005; Hipp et al., 2004), which is independent of poly-ubiquitination (Schmidtke et al., 2009) but can be accelerated by the UBL-UBA domain protein, NEDD8 ultimate buster 1-long (NUB1L) (Hipp et al., 2005; Hipp et al., 2004; Schmidtke et al., 2009). FAT10 and FAT10 conjugates are rapidly degraded by the proteasome and most likely not recycled as is usually the case for ubiquitin (Hipp et al., 2005). In order gain a more detailed insight into the FAT10-conjugation pathway it is crucial to identify the FAT10 specific E1, E2, and E3 enzymes as well as the physiological substrates of FAT10 conjugation. The identification of enzymes involved in FAT10 conjugation and the identification of FAT10-interacting proteins or substrates is very important for the investigation of biological functions of FAT10ylation and FAT10-dependent protein degradation. With respect to the conjugation cascade, Ubiquitin-like modifier activating enzyme 6 (UBA6) has been characterized not only as a second E1 for ubiquitin but also as an E1 for FAT10 (Chiu et al., 2007; Pelzer et al., 2007). Given that UBE1 and UBA6 are co-expressed in many tissues (although UBE1 is up to 10 fold more abundant than UBA6), these two enzymes may act in concert or in sequence to affect various signaling pathways. One possibility is that UBE1 and UBA6 might use a different spectrum of E2-enzymes and eventually different E3- enzymes with their corresponding substrates. UBA6 is highly expressed in testis, therefore an organ specific function is expected. At the beginning of the doctoral thesis, no E2, E3 enzymes nor FAT10 specific substrates had been identified yet. In 2010, the UBA6 specific E2 enzyme (USE1) has been confirmed to be not only the first identified E2 enzyme for FAT10, but also the first identified substrate in the FAT10 conjugation pathway (Aichem et al., 2010). Very recently, two further substrates of FAT10, namely huntingtin (Nagashima et al., 2011) and p53 (Li et al., 2011) have been described.

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Discussion

In order to identify new interaction partners of UBA6 and putative E2 enzymes for FAT10 we performed a yeast two-hybrid screen using a human thymus cDNA library as FAT10 was found to be most highly expressed in the thymus (Lukasiak et al., 2008). Among 96 bait dependent clones, which could be confirmed by retransformation, 69 different clones encoded for full length fat10 and 3 Plasmids encoded for the C-terminal half of bruce containing the entire ubiquitin conjugating (UBC) domain, which was further investigated in terms of beeing a putative FAT10 E2 conjugating enzyme. The putative association of UBA6 with BRUCE might argue for an UBE1- independent loading of BRUCE with either ubiquitin or FAT10 by this newly identified ortholog. Hence, we tested the possibility of interaction between BRUCE and either ubiquitin or FAT10.

BRUCE interacts non-covalently with UBA6 and FAT10 To verify the yeast-two-hybrid findings and confirm the interaction with UBA6, we examined the ability of human UBA6 to interact with the human native full-length BRUCE in HEK293T cells (see 6.1.1). In co-immunoprecipitation experiments under non-reducing and reducing conditions (10 % ß-mercaptoethanol) we could detect a clear non-covalent interaction of BRUCE with UBA6, since UBA6 becomes co-immunoprecipitated to a higher amount with BRUCE under reducing conditions. UBA6 is probably a specialized E1 that can only transfer ubiquitin or FAT10 to a small number of E2s like USE1 and probably BRUCE. FAT10 could be detected after co-immunoprecipitation with BRUCE in 4-12 % Bis-Tris-gels in a higher amount under reducing conditions than under non-reducing conditions, indicating that also FAT10 becomes thioester-linked to BRUCE and presumably can be transferred from UBA6 onto BRUCE in vivo. Moreover, on 3-8 % Tris-Acetate gels, which allow the detection of high molecular weight bands, a very faint conjugate band at the height of ~530 kDa could be observed under non- reducing conditions, which strengthens the hypothesis that FAT10 becomes thioester-linked to BRUCE. Only a weak interaction between BRUCE and FAT10 could be previously detected in a yeast two-hybrid interaction assay with FAT10 and the C-terminal half of BRUCE. Further, a covalent interaction of ubiquitin and BRUCE was observed. BRUCE has been described to be a highly unusual enzyme of the ubiquitin conjugation system as it combines in a single poly-peptide ubiquitin conjugating enzyme (E2) with ubiquitin E3 ligase forming a chimeric E2/E3 ubiquitin ligase. It could be shown in vitro that BRUCE mediated the ubiquitination of the substrates Smac/Diablo, HtrA2 and active caspase-9 only in presence of functional UBE1 (Bartke et al., 2004; Hao et al., 2004).

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Discussion

To further test, if BRUCE can become thioester-linked to endogenous FAT10, a semi- endogenous co-immunoprecipitation assay with FLAG-BRUCE, HA-UBA6 and endogenous FAT10 was performed, where FAT10 was induced with the pro-inflammatory cytokines TNF- α and IFN-γ (6.1.2). Immunoprecipitation assays revealed that HA-UBA6 co- immunoprecipitates with BRUCE, as previously shown and the amount of co- immunoprecipitated HA-UBA6 decreased under non-reducing conditions, which strongly suggest, that both proteins become thioester-linked to each other. Moreover, endogenous up-regulated FAT10 co-immunoprecipitated with BRUCE, but no difference between reducing or non-reducing conditions on protein level could be observed. Further, no conjugate formation between BRUCE and FAT10 at the height of ~530 kDa neither under non-reducing nor reducing conditions could be detected, and therefore provide no evidence, that FAT10 becomes thioester-linked to the active site cysteine of the UBC domain of BRUCE. However, FAT10 linked proteins have been described to be difficult to be detect, as so far identified, but unpublished FAT10 substrates were only to 5-10 % covalently modified with FAT10. Moreover, BRUCE has been described to be one of the largest proteins and the detection limit can be reached because of limited gel resolution.

We further tested in vitro whether activated FAT10 can be transferred from UBA6 onto BRUCE. For this purpose immunoprecipitation was coupled with an in vitro FAT10ylation assay of FLAG-tagged BRUCE in the presence of the recombinant FAT10 and recombinant UBA6 (E1). Unfortunately, no transfer of FAT10 on BRUCE could be detected in the presence of UBA6 (E1), FAT10 and the putative FAT10 E2 enzyme BRUCE (data not shown). In a modified in vitro assay, HA-FAT10 was coupled on HA-agarose beads and incubated with recombinant UBA6 and FLAG-BRUCE, which was eluted from anti-FLAG M2 affinity gel with FLAG-peptide. Also in this approach, no FAT10 transfer on BRUCE could be detected. However, the amounts of thioester linked UBA6~FAT10 adducts were quite low. For further experiments it is essential to develop a reliable in vitro system to clarify the role of BRUCE as a putative FAT10 E2 conjugating enzyme. So far it can only be concluded that BRUCE is interacting with UBA6 and FAT10 in a yeast two-hybrid assay and in co-immunoprecipitation assays. To gain absolute certainty about the identity of the BRUCE-FAT10 conjugate it could also be analyzed by mass spectrometry. This is however only possible if it can be identified on a silver-stained SDS gel. To prove the possibility that BRUCE is an E2 enzyme, which mediates beside ubiquitin conjugation also charging with the ULM FAT10, a reasonable experiment would be siRNA mediated downregulation of BRUCE. Hereby it could be proven, if silencing of BRUCE leads to a reduction of either ubiquitin or FAT10 conjugates and therefore to determine, whether BRUCE has a central role in ubiquitin or FAT10 conjugate formation.

148

Discussion

Strong evidence for the function of FAT10 as a tumor suppressor comes from the finding that expression of FAT10 is able to induce caspase-dependent apoptosis in a variety of models. Studies in a murine fibroblast line (Raasi et al., 2001) and renal tubular epithelial cells (Ross et al., 2006) demonstrated that overexpression of FAT10 resulted in massive caspase dependent apoptosis dependent on the conjugation of FAT10 to so far unidentified target proteins. BRUCE contains a single BIR-domain and an ubiquitin-conjugating enzyme (UBC) E2 domain and has been described to function as an inhibitor of apoptosis protein in mammalian cells (Bartke et al., 2004; Hao et al., 2004). It can bind and inhibit activated initiator caspases-8 and -9 and executioner caspases -3, -6 and -7 and moreover, both Smac and HtrA2 are able to compete for BRUCE-bound caspases. Interestingly, BRUCE is also a substrate of caspases and the serine protease HtrA2, pointing to a role in regulating apoptosis at early stages when proteolytic activity mediated by these enzymes is still low. These interesting characteristics of both proteins leads to the hypothesis that binding of FAT10 to BRUCE might prevent caspase mediated apoptosis of FAT10 through binding to active caspases or FAT10 itself. As a next step, functional studies could be performed. An important feature of BRUCE is its association with membranes including the Golgi apparatus (Hauser et al., 1998). To test if FAT10 has an influence on the subcellular localization of BRUCE, transiently transfected cells could be analyzed with confocal immunofluorescence microscopy in the presence and absence of FAT10. BRUCE is a survival-like BIR domain containing protein (BIRP) with a role in cell cycle regulation and cytokinesis and BRUCE depletion causes cytokinesis defects and cytokinesis associated apoptosis. Especially, BRUCE has an indispensible role at final stages of cytokinesis and particularly controls proper midbody ring formation which is required for cell cycle continuation (Pohl and Jentsch, 2008). Since the expression of FAT10 as well as BRUCE is cell cycle regulated, co-localization studies should be performed with synchronized cells to exactly determine protein localization at different cellular states. The RING finger containing ubiquitin ligase Nrdp1/FLRF has been described to catalyze BRUCE ubiquitination and proteasomal degradation, which contributes to apoptosis induction. Despite the UBC E2 motif, BRUCE did not function with Nrdp1 as an E2 in its own ubiquitination (Qiu et al., 2004), but it remains possible that BRUCE may function as an E2 for ubiquitination of other proteins under other conditions. In vitro experiments with the RING finger containing ubiquitin E3 and putative FAT10 E3 ligase TRIM11 would be interesting to investigate, to test whether TRIM11 mediates either FAT10 or ubiquitin ligation to BRUCE dependent on the UBC domain of BRUCE.

149

Discussion

The ubiquitin-proteasome system (UPS) and the autophagy-lysosome pathway are the two main routes for eukaryotic intracellular protein clearance. Depending on the physiological and pathological conditions, autophagy has been shown to act as a pro-survival or pro-death mechanism in vertebrates (Levine and Klionsky, 2004). Interestingly, drosphila BRUCE (dBRUCE) was shown to regulate autophagy and cell death during early and mid-oogenesis in Drosophila (Hou et al., 2008). In this case, dBRUCE and caspase activity were shown to influence autophagy, which provided first evidence for a mechanism by which autophagy regulates dBRUCE and cell death. Moreover, BRUCE itself becomes degraded by autophagy in the degenerating nurse cells during late oogenesis whereby autophagic degradation of dBRUCE controls DNA fragmentation in nurse cells and thereby provides a mechanistic link between autophagy and cell death (Nezis et al., 2010). Recently, also a mechanistic link between FAT10, autophagy and the UPS was discovered. The autophagosomal receptor and multidomain protein p62, which is acting as a linker between ubiquitinated cargo and the autophagosome, was identified as a putative substrate of FAT10. Non-covalent and covalent interaction of FAT10 and p62 could be observed in co- immunoprecipitation assays and moreover co-localized together in p62 bodies whereby FAT10ylation of p62 led to its proteasomal degradation (A. Aichem et al. submitted). It was shown that autophagy inhibition, which was previously believed to only affect long- lived proteins, can also compromise the ubiquitin- proteasome system which was largely dependent on p62 accumulation after autophagy inhibition, whereby excess of p62 inhibited the clearance of ubiquitinated proteins destined for proteasomal degradation by delaying their delivery to the proteasome (Korolchuk et al., 2009). An interesting experiment would be to determine the interaction between FAT10 and BRUCE under conditions of either proteasome or autophagy impairment with the use of MG132 and Chloroquine. BRUCE is a peripherally membrane associated protein, localized mainly to the trans-golgi network (TGN) and perinuclear vesicles. To test if FAT10 has an influence on the subcellular localization of BRUCE when the proteasome or autophagy is either inhibited or induced, transiently transfected cells could be analyzed by confocal immunofluorescence microscopy. BRUCE is highly expressed in brain, testis, lymphatic cells and secretory organs and also found in any other tissue (Chen et al., 1999). Moreover, it is highly expressed in the mouse embryos up to E11 and then transcript levels drop (Hitz et al., 2005; Lotz et al., 2004; Ren et al., 2005). In comparison, FAT10 mRNA is highly expressed in the lymphatic organs like thymus, spleen and lymph nodes but barely expressed in the brain (Liu et al., 1999; Lukasiak et al., 2008).

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Discussion

However, expression of FAT10 is not restricted to cells of the immune system, but can be synergistically induced with the proinflammatory cytokines TNF-α and IFN-γ in cells of almost every tissue origin (Raasi et al., 2001). Interestingly, FAT10 (Ji et al., 2009; Lee et al., 2003; Lukasiak et al., 2008; Lukasiak et al., 2008) as well as BRUCE up-regulation (Chen et al., 1999) has been observed in certain types of cancers, which raises the possibility that interaction of BRUCE and FAT10 indeed occur in cells under certain physiological conditions.

Yeast two-hybrid screen with TRIM11 Before the state of this work, a yeast two-hybrid screen with FAT10 as bait used to screen a human thymus cDNA library had been performed, to identify new FAT10-interacting proteins (A. Aichem). In this assay the ring finger ubiquitin ligase TRIM11 was identified as an interaction partner of FAT10. Moreover, TRIM11 could interact in vivo with the FAT10 specific E2 enzyme USE1, and siRNA mediated TRIM11 downregulation decreased FAT10 conjugates, indicating that TRIM11 may function not only as a ubiquitin but also as a FAT10 specific ring finger containing E3 ligase (A. Aichem, unpublished). TRIM11 has already been described to act as an E3 ligase that mediates the ubiquitination of many different substrates such as humanin, a neuroprotective peptide that specifically suppresses Alzheimer's disease (AD)-related neurotoxicity (Niikura et al., 2003), the transcription factors Pax6 (Tuoc and Stoykova, 2008) and Phox2B (Hong et al., 2008) and probably ARC105 that mediates chromatin-directed transcription activation and is a key regulatory factor for transforming growth factor beta (TGF-ß) signaling (Ishikawa et al., 2006). In order to gain more information about the function of TRIM11 a yeast two-hybrid screen with human TRIM11 as bait protein was performed to screen a human thymus cDNA library because FAT10 was found to be most highly expressed in the thymus (Lukasiak et al., 2008), especially with the aim to identify new interaction partners of TRIM11 and putative substrates of FAT10. Among 121 bait dependent clones, which could be confirmed by retransformation, the large-scale yeast two-hybrid screen with TRIM11 uncovered 3 independent cDNA clones encoding for the transcription factor JunB and 9 cDNA clones encoding for Ambra1 (see 6.2.1). The interactions between TRIM11 with either JunB or Ambra1 was further characterized in this work, where JunB was also investigated in terms of a being a FAT10 specific substrate.

151

Discussion

TRIM11 and JunB interact in human cell culture The yeast two-hybrid system used in this work is only suitable for the investigation of protein- protein interactions between soluble proteins where the bait as well as the prey constructs are both translocated to the nucleus to form an active transcription factor and activate the respective reporter genes. Due to a different environment in the yeast cell, mammalian proteins are sometimes not correctly modified, for instance phosphorylated, or some proteins might specifically interact when they are co-expressed in yeast, although in reality they are never present in the same cell compartment at the same time, which can lead to false positive results. To verify the yeast two-hybrid results and to ensure, that the interaction between TRIM11 and JunB also takes place in human cells, co-immunoprecipitation assays in the human cell line HEK293 with a MYC-FLAG tagged JunB and a HIS-tagged TRIM11 construct were performed (see 6.2.2). Co-expression of JunB and TRIM11 resulted in decreased JunB protein level and proteasome inhibition rescued the effect, which could argue for a probable role as E3 ubiquitin ligase where binding destabilizes JunB, as has been described for other TRIM11 substrates (Hong et al., 2008; Ishikawa et al., 2006; Niikura et al., 2003; Tuoc and Stoykova, 2008). Interestingly, the amount of TRIM11 protein level decreased concurrently when JunB was ectopically co-expressed. A high molecular weight smear was observable, when JunB-MYC-FLAG and HIS-TRIM11 transfected cells, treated additionally with MG132 for 6 h, were immunoprecipitated against the FLAG-tag of JunB and immunoblotted against JunB. This could argue for augmented JunB conjugate formation (for instance ubiquitination when TRIM11 is over-expressed), which are rescued from proteasomal degradation. Moreover, the result of the co- immunoprecipitation assays of JunB and TRIM11 could verify the yeast-two-hybrid interaction between these two protein, while pre-treatment with the proteasome inhibitor MG132 for 6 h clearly augmented the amount of co-immunoprecipitated TRIM11, suggesting that TRIM11 or JunB become stabilized after proteasome inhibition. TRIM11 is a member of the TRIM family proteins containing an N-terminal RING- finger, a B-box type 1, a B-box type 2, a coiled-coil region and a C-terminal B30.2 domain. The B30.2 domain has been described to be essential for interaction with some of the substrate proteins, like Phox2B, humanin and Pax6 (Hong et al., 2008; Niikura et al., 2003; Tuoc and Stoykova, 2008). The amino acids lining the binding surface are highly variable among the B30.2/SPRY domains, suggesting that these domains are protein-interacting modules, which recognize a specific individual partner protein rather than a consensus sequence motif (Woo et al., 2006). Future work on the TRIM11-JunB interaction could therefore address the question if this motif is responsible for the binding of JunB or if a distinct site of TRIM11 interacts with JunB.

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Discussion

For this purpose, truncation mutants of TRIM11 (e.g. missing the B30.2/SPRY domain) should be tested in co-immunoprecipitation experiments or in vitro pull-down experiments with the use of recombinant proteins. Co-immunoprecipitation of JunB together with a TRIM11 truncation mutant, lacking the entire ring domain, revealed that these proteins still interact, indicating that the ring finger is not essential for JunB recognition (this work, data not shown). Furthermore, it should be tested, if co-expression of JunB and a mutated TRIM11 derivative in which the RING finger domain is disrupted by mutating the consensus cysteine or deleting the entire ring finger domain, plays a role in the regulation of intracellular JunB level through ubiquitin-mediated protein degradation pathways. Nevertheless it will it be essential to confirm the interaction of the endogenous proteins.

JunB becomes isopeptide-linked to ubiquitin and FAT10 Regulation of JunB activity can occur at the level of transcription, mRNA turnover, protein stability, post-translational modifications, or interactions with other transcription factors (Eferl and Wagner, 2003). JunB turnover via the ubiquitin proteasome pathway is strictly regulated and ensures controlled cell-cycle progression and regulated proliferation (Bakiri et al., 2000). Hitherto, two Ub specific E3 ligases have been described to target JunB for proteasomal degradation, namely the HECT E3 Smurf1 (Zhao et al., 2010) and the RING E3 ligase Itch (Gao et al., 2004). To investigate if JunB becomes covalently attached to either ubiquitin or FAT10 mediated by the ring E3 ligase TRIM11, co-immunoprecipitation assays with ectopically expressed proteins in the mammalian cell line HEK293 were performed (see 6.2.3). In case that TRIM11 is either a ubiquitin or FAT10 specific E3 ligase with JunB as substrate, we expected that co-expression of these proteins would augment ubiquitin or FAT10 conjugation to JunB in transfected cells. Surprisingly, a covalent as well as non-covalent interaction between either JunB and ubiquitin or JunB and FAT10 was detectable under reducing conditions, after immunoprecipitation against the HA-tag of ubiquitin and FAT10 and immunoblotting with a FLAG-reactive antibody against the FLAG-tag of JunB. Interestingly, JunB becomes mono- ubiquitinated, although a protein smear also may indicate poly-ubiquitination. In the case of FAT10, a non-covalent interaction as well as a covalent interaction with JunB was detectable. Remarkable, in this case, was that conjugate formation resulted in a high molecular double band, fitting to the size of mono-FAT10ylated JunB. The appearing double band nourished the notion that presumably different post-translationally modified forms of JunB become attached to FAT10. This possibility was further investigated in mass spectrometric analysis and with phospho-specific JunB antibodies (see 6.2.15 and 6.2.16).

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Discussion

Strikingly, the conjugate double-band disappeared almost completely when TRIM11 was co- expressed, similar to the previous case, when ubiquitin and JunB were co-expressed together with TRIM11. One possibility is that conjugate formation is mediated through TRIM11, but conjugates are rapidly degraded via the proteasome. Another hypothesis is that TRIM11 is not responsible for mediating ubiquitin or FAT10 ligation to the substrate JunB and has other functional consequences. Generally, overexpression of TRIM11 led to a decreased protein level of ubiquitin, FAT10 and JunB, which could argue for accelerated protein degradation when TRIM11 is present in an excessive amount. This was also observed for Pax6 (Tuoc and Stoykova, 2008), p62 (F. Kundl, Bachelor thesis, 2010 BITg) and Ambra1 (this work). The role of TRIM11 as a FAT10 specific E3 ligase was investigated in in vitro experiments and will be discussed later (see 6.2.13).

Proteasome inhibition augments conjugate formation between JunB and FAT10 Our results show that JunB becomes covalently modified with FAT10 and it would be expected that this modification serves as a signal for proteasomal degradation of the conjugate. Previously, it has been reported that fusion of FAT10 to otherwise long-lived proteins strongly enhanced their degradation rate, suggesting that mono-FAT10ylation is sufficient to target proteins for proteasomal degradation (Hipp et al., 2005). To determine, if the conjugate is degraded through the proteasome, the experiment was repeated with and without the addition of the proteasome inhibitor MG132 (see 6.2.4). As a further proof for the covalent JunB-FAT10 conjugate formation, JunB was co-expressed with a HA-FAT10ΔGG mutant, which is not capable to form isopeptide linkages to target proteins. The amount of JunB-FAT10 conjugates could be augmented significantly, when cells were pre-treated with MG132 for 6 hours before cell lysis, suggesting an involvement of the proteasome for conjugate degradation. As this modification did not occur when the FAT10ΔGG mutant was co-expressed, it additionally confirms the previous result that FAT10 becomes isopeptide- linked to JunB. Besides, co-expression with TRIM11 almost completely abolished the JunB- FAT10 conjugate and no effect after MG132 treatment was visible, which argue in the first instance against the hypothesis, that TRIM11 mediates FAT10 ligation to JunB. A dual role of TRIM11 has been previously described for the transcription factor Pax6 (Tuoc and Stoykova, 2008). TRIM11 mediates on one hand Pax6 degradation via the ubiquitin-proteasome system and TRIM11 overexpression decreased endogenous Pax6 protein levels and repressed Pax6 functions, whereas abrogation of endogenous TRIM11 increased the level of insoluble forms of Pax6 and enhanced apoptosis.

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Discussion

Here the B30.2 domain of TRIM11 was essential for the clearance of insoluble cell proteins. On the other hand, TRIM11 expression is directly regulated by Pax6 which indicate that an autoregulatory feedback loop between TRIM11 and Pax6 maintained a balance between the levels of Pax6 and TRIM11 proteins. In case of JunB, we suspect a role for TRIM11 in controlling JunB and JunB conjugate stability rather than being an E3 ligase that FAT10ylates JunB.

The JunB-FAT10 conjugate becomes proteasomally degraded and FAT10ylation accelerates JunB turnover We could provide evidence that co-expression of JunB does not change the turnover rate of FAT10, since FAT10 turnover remained constant revealing a half life of about 1 h (see 6.2.5). To assure that FAT10ylation is a means to target JunB for proteasomal degradation we next determined the protein half life of unconjugated JunB in the presence and absence of FAT10 expression (see 6.2.6). The degradation rate of monomeric JunB in the absence of FAT10 was only very slightly decreased compares to the JunB turnover in the presence of FAT10, concerning monomeric JunB on protein level. Here we cannot exclude the possibility that the decrease of JunB level occurred somehow due to a covalent interaction with FAT10, which was not detectable on the Western blot. However, pre-treating the cells with MG132 could rescue turnover of monomeric, unconjugated JunB in the presence and absence of FAT10 co-expression and led to a JunB accumulation of about 125 % in both cases. These results indicate that the turnover rate of monomeric unconjugated JunB in the presence of FAT10 was hardly affected. We next aimed to determine the turnover rate of the JunB-FAT10 conjugate in cycloheximide experiments (see 6.2.7). Since FAT10 is degraded by the proteasome and targets FAT10-linked proteins for proteasomal degradation (Hipp et al., 2004; Schmidtke et al., 2009), we investigated in cycloheximide-chase experiments, if the turnover rate of the JunB-FAT10 conjugate and the turnover rate of non-covalently interacting and immunoprecipitated JunB changes over time and was caused by proteasomal degradation. The cycloheximide data indicated that either covalently modified or non- covalently interacting co-immunoprecipitated JunB show a similar protein turnover of about 2.5-3 h, which is much faster, than the turnover rate of monomeric unconjugated JunB in the presence of FAT10. MG132 treatment led to a rescue of the JunB-FAT10 conjugate and the non-covalently with FAT10 co-immunoprecipitated JunB, revealing an involvement of the proteasome in degradation. Moreover, compared to the much shorter protein half-life of 1 h for monomeric FAT10, these data clearly indicate that FAT10 indeed interacts non-covalently or covalently with JunB and degradation is mediated through FAT10 conjugation.

155

Discussion

In further experiments it could also be tested, if co-expression of the interferon-inducible protein NUB1L, which is a non-covalent binding partner of FAT10, can enhance the turnover of JunB as it has been described for FAT10 itself (Hipp et al., 2004). The turnover of JunB in the presence of FAT10 or FAT10 and NUB1L should be investigated in pulse-chase or cycloheximide chase experiments.

Endogenous and ectopically expressed TRIM11 becomes degraded through the proteasome Considering the previous experiments it still remains unclear, if TRIM11 is a FAT10 or ubiquitin specific E3 ligase with JunB as substrate or rather exerts another function as a putative FAT10 substrate, for instance. In this case it would be expected that FAT10 becomes isopeptide linked to TRIM11 with the help of another E3 ligase. So far, no TRIM11- FAT10 conjugate could be detected. To further characterize TRIM11, the protein half life of either endogenous or over-expressed TRIM11-FLAG was determined in a cycloheximide experiment in HEK293 cells (see 6.2.8). Moreover, to evaluate, if TRIM11 is assigned to proteasomal degradation, cells were additionally treated with MG132 for 6 h. In HEK293 cells, a continuous decrease of endogenous TRIM11 could be observed revealing a protein half life of 7-8 hours, in contrast to ectopically expressed TRIM11-FLAG with a much shorter half-life of 5 h. Moreover, treatment with MG132 completely blocked TRIM11-FLAG degradation and increased accumulation, which clearly substantiate an involvement of the proteasome for TRIM11-FLAG turnover, whereas only a slight rescue of endogenous TRIM11 could be measured when MG132 was added. The faster degradation of TRIM11- FLAG occurs maybe due to an excess of protein.

TRIM11 turnover in the presence of FAT10 is slightly accelerated Many RING E3s are known to be ubiquitinated, often by an autocatalytic process. Autocatalytic ubiquitination can be a simple consequence of E3 activity with no functional impact. Alternatively, it could lead to down-regulation of E3 activity owing to degradation by the proteasome, as has been seen for the multi-subunit SCF complex (Galan and Peter, 1999). To investigate the role of TRIM11 as a putative FAT10 substrate, a cycloheximide chase with ectopically expressed TRIM11 and FAT10 was conducted (see 6.2.9). The half-life of HIS- TRIM11 in the presence of FAT10 was about 4 h and therefore slightly accelerated in comparison to 5 h for TRIM11-FLAG (see 6.2.8). These results initially indicated that FAT10 co-expression shortens the protein half life of HIS-TRIM11, but faster degradation can also be caused through the usage of differently tagged TRIM11 constructs.

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Co-expression of TRIM11 does not change the protein turnover rates of JunB and FAT10 Quantitative analysis resulted in a quite similar degradation rate for either FAT10 or JunB, in the presence or absence of over-expressed HIS-TRIM11 (see 6.2.10). The JunB-FAT10 conjugate turnover rate between JunB and FAT10 in the presence of TRIM11 could not be calculated, because co-expression of HIS-TRIM11 almost completely abolished conjugate formation. As we could previously show, co-expression of TRIM11 resulted in severely decreased protein levels either for JunB or FAT10 and TRIM11 itself, without changing the velocity of degradation, which supported a role in regulating and controlling protein stability and abundance and might be involved in protein quality control as has been shown for Pax6, where an autoregulatory feedback loop between TRIM11 and Pax6 maintains a balance between the levels of Pax6 and TRIM11 proteins in cortical progenitors (Tuoc and Stoykova, 2008). On the other hand there remains the possibility that endogenous TRIM11 could have contributed to FAT10 conjugate formation and that overexpression of the protein could not further enhance this process, or leads to depletion of abundant proteins. Knockdown experiments of endogenous TRIM11 via siRNA or shRNA will shed light on the question, if TRIM11 is an essential FAT10 specific E3 ligase with JunB as substrate. If this is indeed the case, we would expect that JunB-FAT10 conjugate formation should be abrogated. This would also be true for further TRIM11 interaction partners and putative substrate proteins.

Conjugate formation of endogenous JunB and FAT10 A general problem in protein-protein interaction studies is that overexpression studies are prone to artefacts, because over-expressed proteins behave in a different way, than under normal, physiological protein concentrations. To test whether a JunB-FAT10 conjugate is formed in the absence of overexpression, endogenous FAT10 was upregulated with TNF-α and IFN-γ in HEK293 cells and co-immunoprecipitation studies were performed with a FAT10-specific monoclonal antibody (mAb) (designated 4F1), followed by Western blot analysis with either a FAT10-specific polyclonal antibody (pAB) or a polyclonal JunB antibody under reducing conditions (see 6.2.11). Non-covalently FAT10 interacting JunB was apparent after immunoblotting with the polyclonal JunB antibody, but only a very weak JunB- FAT10 conjugate band could be detected. Moreover, treatment with the proteasome inhibitor MG132 did not enhanced conjugate formation (data not shown). These data indicate that JunB-FAT10 conjugate formation can indeed occur under completely endogenous conditions but needs to be repeated and confirmed.

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Discussion

Reasons for the weak conjugate band could be the low JunB protein amount in HEK293 cells. Besides, a possible explanation could be that the polyclonal JunB antibody is not sensitive enough to detect the endogenous conjugate band. The reciprocal approach, to immunoprecipitate endogenous JunB and immunoblot with a FAT10 specific antibody however, failed and no conjugate band was detectable, neither in HEK293 cells, nor in HELA cells, where JunB expression has been described to be higher. Subsequent experiments in other cell lines and other antibodies should therefore further investigate, if conjugate formation increases under these conditions.

Conjugate formation of JunB and FAT10 under semi-endogenous conditions Since conjugate formation between JunB and FAT10 was hardly detectable under completely endogenous conditions, co-immunoprecipitation assays were performed under semi-endogenous conditions with ectopically expressed JunB-MYC-FLAG and endogenous FAT10, upregulated via the pro-inflammatory cytokines TNF-α and IFN-γ (see 6.2.12). After immunoprecipitation of JunB and immunoblotting with a polyclonal FAT10 antibody, conjugate formation between JunB and FAT10 could be detected under reducing conditions. Interestingly, JunB expression was apparently increased in INF-γ/TNF-α treated cells, whereas the total protein amount remained stable. JunB, belonging to the AP-1 family of transcription factors, has been described to be induced by a plethora of physiological stimuli and environmental insults (see 3.5.2). For instance, JunB is an immediate early transcription factor induced in response to stimulation of cell surface receptors (Angel and Karin, 1991), growth factors, phorbol esters, stress signals, (Hess et al., 2004; Perez-Albuerne et al., 1993), NF-κB mediated transcriptional induction of JunB in response to oxygen deprivation (Schmidt et al., 2007) and also proinflammatory cytokines like IL-6 or TNF-α (Gomard et al., 2010; Yin et al., 2009) or INF-γ (Gurzov et al., 2011) and therefore might play a role in regulating immune response genes. Recently, it could be demonstrated that JunB can counteract cytokine-induced ER stress and apoptosis and might therefore be part of a defense mechanism triggered by B-cells during exposure to pro-inflammatory cytokines (Gurzov et al., 2011). Further experiments with Real-time PCR analysis upon cytokine treatment would be interesting to further investigate this issue and JunB-upregulation during cytokine induction might build a link to FAT10 induction mediated through the proinflammatory cytokines TNF-α and INF-γ during conditions of inflammation in the cell.

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Discussion

In vitro auto-FAT10ylation assay Previous reports state that JNK regulates the turnover of JunB through its accelerated degradation induced by JNK-mediated phosphorylation of the HECT class E3 ubiquitin ligase Itch (Gao et al., 2004). Moreover, biochemical experiments demonstrated that the HECT domain containing E3 ligase Smurf1 which primarily catalyzes mono- or poly-ubiquitination of protein targets which primarily function in TGF-β signaling pathways, interacts with JunB through the PY motif and targets JunB protein for ubiquitination and proteasomal degradation (Zhao et al., 2010). To test, whether the RING finger containing E3 ligase TRIM11 ligase can mediate FAT10 ligation to JunB, an in vitro FAT10ylation assay with recombinant proteins was performed (see 6.2.13). Moreover, the B-box domain of the TRIM subfamily of RING E3s is structurally related to the RING domain and functions in mediating ubiquitination, while no function in recruiting E2 enzymes has been described (Tao et al., 2008). The coupled immunoprecipitation of JunB and in vitro FAT10ylation assay in the presence of recombinant FLAG-UBA6, His-USE1, FAT10 and GST-TRIM11, however, did not provide evidence that JunB becomes FAT10ylated in presence of an E1, E2 and a putative E3 enzyme under conditions containing a rich source of renewable energy such as ATP, creatine phosphate, creatine phosphokinase and inorganic pyrophosphatase, which could serve as a cell free system for the FAT10ylation reaction in vitro. The efficiency of in vitro assays highly depends on the quality of the enzymes involved. To argue why the in vitro FAT10ylation assay with GST-TRIM11 failed seems to be very troublesome. TRIM11 belongs to the class of single-polypeptide (subunit) E3s that contain a motif known as RING which is associated with another unique type of cysteine-rich, zinc-binding domain, the B- box. The RING and two B-boxes are followed by a leucine coiled-coil forming the tripartite motif. The RING motif is maintained by the arrangement of eight cysteines and histidines in a cross-brace manner that chelate two zinc ions, which is distinct from other zinc finger domains (Borden, 1998). It is proposed that the RING-type E3 ligases serve as scaffolds to bring together the activated ubiquitin-E2 intermediate and the substrate protein to promote the transfer of the ubiquitin molecule to the substrate protein (Lorick et al., 1999). Zinc ligation is required for folding of the domain and for subsequent biological action. One reason why the in vitro FAT10ylation assay failed could be that remaining reduced glutathione concentration after dialysis could have lead to the abstraction of all three zinc atoms and therefore to a three dimension conformational change of TRIM11. Another problem could be the poor solubility and impurity of recombinant GST-TRIM11 protein, due to many degradation products. During in vivo experiments, we could observe that JunB becomes not only monoFAT10ylated, but also mono-ubiquitinated. In addition, an ubiquitin smear indicated that JunB becomes even poly-ubiquitinated.

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Discussion

In order to address the question, if TRIM11 is a further ubiquitin E3 ligase beside Itch (Gao et al., 2004) and Smurf1 (Zhao et al., 2010) for JunB, an in vitro activation assay with ubiquitin and recombinant proteins should be carried out. Moreover, to have a positive control that the in vitro system used in this work is reliable, an in vitro ubiquitination assay with ubiquitin and with recombinant JunB and one of the already published ubiquitin E3 ligases, either Itch (Gao et al., 2004) or Smurf1 (Zhao et al., 2010), should be carried out to have a positive control. To date, USE1 has been described to be the major E2 enzyme in the FAT10ylation pathway (Aichem et al., 2010), which moreover can interact in vivo with TRIM11. It cannot be excluded that further E2 enzymes exists, which mediate FAT10 conjugation and binding to FAT10 specific E3 ligases. To further characterize the function of TRIM11 as a putative FAT10 E3 ligase, the in vitro activation assay should be repeated with the putative FAT10 E2 enzyme BRUCE, which was identified in a yeast two-hybrid screen with UBA6 as bait (this work).

JunB is conjugated with FAT10 on lysine 237 The in vivo experiments provided strong evidence that FAT10 becomes isopeptide linked to JunB. To answer the question which lysine of JunB is involved in FAT10 conjugate formation, co-immunoprecipitation assays with JunB wt and JunB lysine mutants together with FAT10 in HEK293 cells where performed, where JunB lysines (K) are mutated individually (JunB- K237R, -K267R, and -K301R) or in combination (JunB-K237R/K267R/K301R, termed JunB- K3R) to arginines (R) (constructs kindly provided by M. Piechaczyk). All these lysines are embedded in a SUMOylation consensus sequence (Rodriguez et al., 2001), and Garaude et al. could illustrate that lysine 237 is the primary SUMOylation site of JunB (Garaude et al., 2008). Although it is clear that modification of most lysines takes place within the SUMO modification consensus motif, certain substrates are modified on lysine residues where the surrounding sequence does not conform to this consensus. Given that no FAT10 consensus sequence has been identified yet, we investigated the interesting possibility, that JunB may become FAT10ylated on the same lysine residue where JunB becomes conjugated to SUMO. Although binding domains for UBLs other than ubiquitin and SUMO have yet to be identified, proteins containing sequences that match the consensus for UBDs but do not appear to bind ubiquitin have been described (Raasi et al., 2005) which might bind other UBLs instead. The binding of FAT10 to the UBA domains of NUB1L (Schmidtke et al., 2006) is a good example for this concept.

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Discussion

Co-immunoprecipitation assays clearly showed that lysine 237 in JunB is probably the major acceptor site not only for the ULM SUMO, but also for FAT10, since co-immunoprecipitation with FAT10 and either JunB-K237R or JunB-K3R completely abolished the appearance of the lower JunB-FAT10 conjugate band. This raises the interesting question if SUMO and FAT10 generally recognize the same consensus motif and compete for binding. Crucial in this context is to investigate, which functional consequences ensue for the fate of JunB. In contrast to FAT10ylation, modification with the ULMs SUMO and ubiquitin has been described to be reversible. The cycloheximide experiments depicted in Figure 29 clearly show that the JunB-FAT10 conjugate is continuously subjected to degradation by the proteasome which correlates with previous data that fusion of FAT10 to otherwise long-lived proteins strongly enhanced their degradation rate, suggesting that mono-FAT10ylation is sufficient to target proteins for proteasomal degradation (Hipp et al., 2005). Posttranslational modification by SUMO has not been generally associated with increased protein degradation. Rather, similar to nonproteolytic roles of ubiquitin, SUMO modification has been shown to covalently modify a large number of proteins with important roles in many cellular processes including gene expression, chromatin structure, signal transduction, and maintenance of the genome and to regulate transcription, protein localization and activity, for instance, although recent studies indicate that SUMO can also act as a signal for the recruitment of E3 ubiquitin ligases, which leads to the ubiquitination and degradation of the modified protein (Geoffroy and Hay, 2009; Melchior, 2000; Miteva et al., 2010; Verger et al., 2003). For JunB it has been shown, that SUMOylation regulates its ability to induce cytokine gene transcription, like IL-2 and IL-4 in T lymphocytes and likely plays a critical role in T cell activation (Garaude et al., 2008). To clarify if FAT10 and SUMO compete for binding to JunB, endogenous or semi- endogenous co-immunoprecipitation assays and immunoblotting with specific antibodies should be performed to determine, if one modification impairs the other. The upregulation of FAT10 through IFN-γ and TNF-α could thereby also reveal, if these signals somehow influence the interaction between the two proteins. Several studies provide hints about the significance of FAT10 in immune responses, including the synergistic induction of FAT10 in the presence of the proinflammatory cytokines TNF-α and IFN-γ (Liu et al., 1999; Raasi et al., 1999). JunB can also be induced in response to proinflammatory cytokines like IL-6 or TNF-α (Gomard et al., 2010; Yin et al., 2009) and therefore might play a role together with FAT10 during conditions of inflammation, to regulate immune response genes. To prove, if SUMOylation influences FAT10ylation and vice versa, it is necessary to establish a reliable in vitro system to examine, if titration of increasing amounts of either recombinant SUMO or FAT10 modulates the binding pattern.

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Discussion

Since JunB contains 19 lysines and the lower conjugate band is still weakly detectable with the JunB-K3R mutant it is possible that JunB becomes FAT10ylated also on other lysine residues. Unfortunately, neither the JunB nor the FAT10 crystal structure has been solved, which makes it difficult to predict which lysine of JunB is exposed on the surface of the protein. However, database analysis of the JunB amino acid sequence with dbPTM2.0 at (http://dbPTM.mbc.nctu.edu.tw/) predicted that K240 (correlating to K237 in JunB mutant construct, used in this work), K287, K289, K301 are presumably surface exposed. To assure that FAT10ylation occurs only at the lysine residue K237 it is necessary to mutate every lysine in the JunB protein, individually, or in combination and to repeat the co- immunoprecipitation assays with FAT10 and the mutant constructs. The disappearance of the upper but the consistency of the lower conjugate band poses the interesting question, if JunB is coincidentally modified with FAT10 on additional lysine residues (multiple mono- FAT10ylation), which is unlikely because of size reasons, or whether modified forms of JunB become attached to FAT10. These modifications can include other posttranslational modifications, for instance, ubiquitination, acetylation, glycosylation or phophorylation (see 6.2.16). The in vivo data further revealed evidence that JunB interacts not only non- covalently with ubiquitin but also gets mono-ubiquitinated and a protein ladder indicates that most likely poly-ubiquitination takes place as well (see 6.2.3). It has been described in some cases that SUMO and ubiquitin may directly compete for modification of target lysines. Both SUMOylation and ubiquitination are reversible, and, in some cases, dynamic cycles of conjugation/deconjugation may be required for regulated activity (Praefcke et al., 2011). Interesting in this context would be, if ubiquitin becomes attached to the same lysine residue K237 as SUMO and FAT10 and would lead to another level of regulation. Here, the interaction should be investigated in co-immunoprecipitation assays with JunB wt and JunB mutants and ubiquitin. A new method to globally characterize the human ubiquitin-modified proteome (ubiquitinome) was recently developed by Kim et al. (Kim et al., 2011). The combination of using a monoclonal antibody that recognizes di-glycine (diGly)-containing isopeptides followed by trypsin digestion and quantitative proteomics, enables to monitor temporal changes in diGly site abundance in response to both proteasomal and translational inhibition. Moreover, this method can be used to identify substrates for cullin-RING ubiquitin ligases (https://gygi.med.harvard.edu/ggbase/). This analysis additionally confirmed our results that lysine 237 becomes FAT10ylated. Reversible posttranslational modifications are widely used to dynamically regulate protein activity. Proteins can be modified by small chemical groups, sugars, lipids, and even by covalent attachment of other polypeptides.

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The most well-known example of a polypeptide modifier is ubiquitin and posttranslational modification by ubiquitin plays a central role in targeting proteins for proteolytic degradation by the proteasome, although covalent attachment of ubiquitin to proteins can also regulate localization and/or activity independent of proteolysis (Aguilar and Wendland, 2003; Kerscher et al., 2006; Laney and Hochstrasser, 1999). To gain certainty if ubiquitin attachment assigns the conjugate for proteasomal degradation, cycloheximide experiments with or without MG132 are warranted.

Identification of JunB in mass spectrometric analysis Protein-protein interactions can be analyzed directly by precipitation of a tagged bait followed by mass spectrometric identification of its binding partners. To gain absolute certainty about the identity of the JunB-FAT10 conjugate and in order to identify, if FAT10ylated JunB contains post-translational modifications (PTMs), mass spectrometry analysis was performed (see 6.2.15). Moreover, we were interested if lysine K237 involved in conjugate formation could be confirmed. A MYC-FLAG-tagged JunB construct was co-transfected with a HA- tagged FAT10 construct in HEK293 cells and immunoprecipitated with FLAG-M2-agarose beads. JunB could be detected in analyzed bands which size correlated to JunB-FAT10 conjugates, but unfortunately no FAT10 could be identified. One possible explanation could be that the mass-to-charge ratio of charged fragments is disturbed in presence of the HA-tag of FAT10. Moreover, results can also depend heavily on how the sample was prepared. Hence, it will be essential to confirm the conjugate formation with JunB and endogenous FAT10 to avoid the risk of disturbing tags. Another possibility to identify interaction partners is the method of cross-linking. Through the use of appropriate reagents, it is possible to link interacting proteins through cysteine or lysine residues under experimental conditions, including in vivo scenarios. In a subsequent mass spectrometric analysis it is then feasible to unambiguously identify the direct interaction partners of a given protein (Gingras et al., 2007).

Post-translational modification: Phosphorylation of JunB on Serine 259? More than 300 post translational modifications are currently known, including addition of chemical groups, such as phosphate or acetate, addition of complex molecules, such as carbohydrates or lipids, the covalent linkage of small proteins, such as ubiquitin and ubiquitin-like proteins (UBLs), cleavage and modification of side chain residues of specific amino acids. Due to the diversity of post-translational modifications, bioinformatics tools like expasy (http://www.expasy.ch/proteomics) or mass spectrometric analysis are useful to predict putative modifications. For predicted post-translational modification sites by profile Hidden Markov Model (HMM), see chapter 10, addendum (http://dbptm.mbc.nctu.edu.tw/).

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Discussion

In addition to the diversity in dimer-formation, modulating DNA-binding specificity and affinity, JunB protein levels and activity are regulated at the post-transcriptional level. The mechanism of post-translational control is most extensively documented in the case of mitogen- and cellular stress- induced hyper-phosphorylation and, in particular, through the Jun N-terminal kinase (JNK) cascade (Karin et al., 1997; Wagner, 2001). Activated by variant stress stimuli through a MAPK cascade, JNK can translocate to the nucleus, phosphorylate serine and threonine residues within its N-terminal transactivation domain and thereby modulate its transactivation potential (Hess et al., 2004; Karin et al., 1997; Wagner, 2001) . Li et al. reported that JunB in synergy with c-Maf to control IL-4 expression is mediated by the phosphorylation of JunB at Thr102 and -104 by JNK MAP kinase during T-helper cell differentiation (Li et al., 1999). Besides, phosphorylation of JunB has been reported to regulate JunB protein levels in different cellular states. For instance, phosphorylation of JunB by the p34cdc2–cyclin B kinase is associated with lower JunB protein levels in mitotic and early G1 cells (Bakiri et al., 2000) and JunB decrease in late G2 phase is due to accelerated phosphorylation-dependent degradation by the proteasome (Farras et al., 2008). Phosphorylation is one of the most common post translational modifications (PTMs) of proteins and consists in the reversible attachment of a phosphate group to a specific residue of a target protein. Phosphorylation dependent activation and alteration of transcription has been described for JunB residues S79, T102 and T104 (http://www.phosphosite.org). However, fifteen putative phosphorylation sites exist, where eleven have been predicted with quantitative phospho-proteomic analysis (Olsen et al., 2010). We next aimed to determine the possibility of phosphorylation as PTM with phospho-specific JunB-antibodies (see 6.2.16). Immunoblotting with a polyclonal JunB antibody (ab31421), or a serine 259 (S259) phospho-specific JunB antibody (ab30628) revealed (Beausoleil et al., 2004; Olsen et al., 2006) that highest and coincidentally the most prominent JunB-FAT10 conjugate double band is recognized by both, the polyclonal JunB and the Ser259 phospho-specific JunB antibody, suggesting that these band present a phosphorylated form of JunB. Interestingly, this double band was not detectable, when samples were analyzed by immunoblotting with a Ser79 phospho-specific antibody (data not shown), indicating a specific interaction of FAT10 with a Ser259 phosphorylated form of JunB. So far, our findings point to a Ser259 JunB phosphorylation as post-translational modification of the conjugate double band. With this experiment, we could not exclude that a further PTM modification occurs, since the upper as well as the lower conjugate band are recognized with the Ser259 phospho-specific antibody. Further experiments are required to substantiate that only a phosphorylated form of JunB becomes modified with FAT10 and that this phosphorylation is specific.

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Discussion

In future investigations it would therefore also be interesting to immunoprecipitate JunB and JunB-FAT10 conjugates with an antibody, which recognizes the most frequent phosphorylation of the hydroxyl group of serine or threonine or tyrosine residues with whole phospho-tyrosines (4G10), phospho-threonine or phospho-serines antibody and to probe immunoblots with an anti-FAT10 antibody, to confirm previous results. A reciprocal approach would be the removal of attached phosphate residues with phosphatases and to test if FAT10ylation can still occur. Post-translational modification of proteins is a widespread mechanism used by both prokaryotic and eukaryotic cells to modify the activity of key factors that play fundamental roles in cellular physiology.

FAT10 and JunB co-localize at the nuclear membrane and cytosol Alterations of the intracellular distribution of target proteins have been already linked to modification with the small ubiquitin-like modifier (SUMO). Many SUMOylated nuclear proteins localize in nuclear subdomains, such as promyelocytic leukemia nuclear bodies, which may serve to sequester them and thereby suppress their activity. Examination of the PML nuclear bodies, whose principal components are SUMO-modified, has revealed this modification to be essential for their structural and functional integrity. Since most known SUMO substrates are nuclear proteins, it has been suggested that SUMOylation is a nuclear process (Rodriguez et al., 2001). Nuclear localization of SP100, for example, is a requirement for SUMOylation since a cytoplasmic mutant of this protein remains unmodified (Sternsdorf et al., 1999). Endogenous unmodified JunB is nuclear and equally distributed between soluble and nonsoluble nuclear chromatin fractions in primary T cells and Jurkat cells, but showed no concentration in a subdomain. However, SUMOylated JunB does show a striking localization, as it was found exclusively in the insoluble nuclear fraction of human primary T cells, associated with a detergent-resistant nuclear matrix and/or chromatin structure (Garaude et al., 2008). Our previous experiments clearly indicated that JunB is covalently modified with FAT10, whereby proteasome inhibition led to a considerable accumulation of the conjugate. To further investigate the localization and fate of either JunB or FAT10 when co-expressed and to analyze the role of FAT10ylation in this process, we performed confocal laser-scanning microscopy (see 6.2.17). FAT10 in the nucleus was found co-localized with JunB and shifted its subcellular localization from the nucleus towards the nuclear membrane. A previous report underlined this observation, where FAT10ylation of the transcription factor p53 altered its subcellular compartmentalization. Furthermore, Overexpression of FAT10 led to a reduction in the size of promyelocytic leukemia nuclear bodies (PML-NBs) and altered their distribution in the nucleus (Li et al., 2011).

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Discussion

Treatment with the proteasome inhibitor MG132 resulted in almost complete re-localization of JunB to the cytosol, with the functional consequence, that the transcriptional activation capacities of JunB could be impaired. Our results substantiate the precedent data that JunB and FAT10 interact and we strongly suggest that modification with FAT10 regulates JunB localization and function. Moreover, we assume that interaction of JunB and FAT10 occurs preferentially at the nuclear membrane, which requires the translocation of JunB away from the nucleus towards the nuclear membrane. The expression of JunB and FAT10 are cell-cycle regulated and the degradation of JunB at the M phase by the proteasome is absolutely required for proper cell-cycle progression, since constitutive JunB expression results in cell-cycle arrest at late G2/early M phase (Piechaczyk and Farras, 2008). An interesting experiment in this context would be, to perform tracking experiments with endogenous proteins to determine, if protein-protein interaction takes place during certain stages of the cell cycle. Moreover, to confirm the interaction between JunB and FAT10 on the one hand and to further determine their localization within a cell on the other hand, we aim to perform co-localization studies via confocal microscopy in different cellular stages. Moreover, JunB has been described to play a role as an autophagy inhibitor induced by starvation (Yogev et al., 2010; Yogev and Shaulian, 2010). After the aforementioned analysis we want to analyze if JunB FAT10ylation is implicated in autophagy and autophagic degradation which would provide a further role for FAT10 in a different degradation mechanism.

FAT10ylation controls JunB transcriptional activities on minimal AP-1 driven reporter genes JunB is a member of the AP-1 family of transcription factors, which is a dimeric complex composed of members of the Fos (c-Fos, FosB, Fra1, and Fra2) and Jun (c-Jun, JunB, and JunD) bZIP protein families (Foletta et al., 1998). JunB has been first characterized as an inhibitor of Jun function following the observations that an excess of JunB over c-Jun is sufficient to inhibit transactivation of AP-1 reporter genes by c-Jun. Previous in vitro and in vivo experiments clearly documented that JunB is not only a repressor of Jun activity but is also required for the transcriptional activation of key target genes involved in cell cycle regulation, proliferation, differentiation, angiogenesis as well as skin and hematopoietic functions (Florin et al., 2006; Licht et al., 2006; Meixner et al., 2008; Schmidt et al., 2007; Szabowski et al., 2000).

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Moreover, inconsistent with a reputation for being a growth-inhibiting protein, fed by antagonistic effects on c-Jun-dependent transcription and on inhibition of c-Jun-dependent transformation in vitro, JunB has been found to possess growth-promoting activities which contribute to neoplastic transformation in certain tissues (Mathas et al., 2002; Rassidakis et al., 2005; Sreeramaneni et al., 2005). While the molecular mechanisms regulating AP-1 transcription activation were intensively studied, the mechanisms underlying transcriptional repression are so far poorly understood. In general, different mechanisms of gene repression have been proposed, involving inhibition of transcription initiation, inhibition or competition for activating factors as well as epigenetic mechanisms. For JunB, it has been proposed that it acts as a repressor on its own by forming heterodimers with other AP-1 subunits that, as a result, exhibit a much weaker transactivation potential. FAT10ylation of the transcription factor p53 has been recently described to alter its subcellular compartmentalization and to up-regulate its transcriptional activity (Li et al., 2011). The homo-/hetero dimers mediate AP-1 regulated gene expression through binding to the palindromic TPA Responsive Element (TRE), composed of the 7 bp DNA consensus sequence 5’-TGA G/C TCA-3’ (Angel et al., 1987), which are present in promoters of many cellular and viral genes. Thereby, JunB appears to be as effective as Jun in trans-activating reporter genes containing multiple AP-1 binding sites which suggest, that trans-activation by JunB may require synergistic interactions between multiple homodimers bound to adjacent sites (Angel and Karin, 1991; Chiu et al., 1989). Also repressive functions of JunB can depend on the formation of inactive heterodimers with c-Jun or competition with c-Jun homodimers for an AP-1 binding site within the promoter region of the target gene. To evaluate the effect of FAT10 conjugation on transactivation by JunB, we performed co- transfection experiments in JURKAT and HEK293 cells with an AP-1 reporter gene, namely the TRE reporter plasmid that is driven by four minimal canonical binding sites upstream of the TATA box and conducted a luciferase reporter assay (see 6.2.18, 6.2.19). Our results showed that K237 was the primary site for FAT10ylation; when this site was mutated to arginine, we detected a minor level of FAT10ylated JunB. We used the JunB wt and triple mutant, JunB-K3R that was barely FAT10ylated to investigate the role of FAT10ylation on the transcriptional activity of JunB. This JunB-K3R mutant activated the TRE reporter to the same extent as JunB wt in Jurkat and HEK293 cells, indicating that the mutation did not affect basal function. Surprisingly, post-translational modification of JunB with FAT10 led to a 2 fold decreased transcriptional activity of JunB on a reporter gene (AP-1) controlled by the 4xTRE binding sites, in contrast to JunB SUMOylation, which did not affect the trans- activating capacities of JunB for minimal AP-1-driven reporter genes containing 4xTRE binding sites (Garaude et al., 2008).

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Discussion

Moreover, similar lowered trans-activating capacities of FAT10ylated JunB in JURKAT and HEK293 cells were obtained for the cytokine promoters IL-2 and IL-4 (data not shown), containing multiple AP-1 sites, in contrast to SUMOylated JunB, which led to increased reporter-activity in JURKAT cells. Co-expression of either mutant JunB-K3R with FAT10 or FAT10ΔGG, or JunB wt and FAT10ΔGG (negative controls) resulted in a similar transactivation of AP-1 (4xTRE), as JunB wt alone. Taken all the experiments together it seems that JunB-FAT10 conjugation has a direct consequence on trans-activating activities of JunB under these conditions. Why the FAT10-JunB interaction regulates reporter activity of AP-1 (4xTRE) can be due to several reasons. Our in vivo data already indicated that either non-covalent or covalent attachment of FAT10 led to accelerated proteasomal degradation of JunB or JunB-FAT10 conjugate. Thus, we suggest that JunB FAT10ylation assigns the conjugate for rapid degradation via the proteasome, which result in decreased JunB content capable to trans- activate reporter genes. To investigate, if FAT10 mediated degradation of JunB is the major cause for decreased transcriptional activity, the reporterassays should be repeated under conditions of proteasome inhibition. Hitherto, neither the JunB nor the FAT10 structure has been crystallized which makes it difficult to predict, which impact their binding on structural conformation of the complex may have. Attachment of a UBL or UBL chain to a protein creates a protein surface topography in the conjugate that is changed substantially from that of the unmodified protein, at least locally. One consequence caused through FAT10 conjugation could be that dimerization of JunB and hence trans-activating capacities of JunB are impeded. The identified lysine 237 (K237) in JunB, which seems to be the primary site for FAT10ylation directly abuts on the C-terminal basic leucine zipper (bZIP) domain, which allows dimerization to form AP-1 transcriptional complexes and to bind to the DNA backbone. FAT10 ligation could impair the formation of either homo- or heterodimers to members of the AP-1 family and thereby inhibit transactivation capacities of JunB. Moreover, attachment of FAT10 could lead to a change in the JunB-FAT10 complex structure, which additionally could disturb the binding of JunB to the TRE binding sites. A further possibility is that UBL conjugation either links together a set of receptor binding sites in the UBL and target protein, to enhance binding to another macromolecule or masks a receptor-binding site in the target protein to inhibit binding to another macromolecule. By this reasoning, enhanced binding of a UBL-conjugated protein, relative to its unconjugated form, to another “receptor” protein implies the existence of a binding site for the UBL in that other protein (Kerscher et al., 2006). Further work will be required to test this hypothesis and characterize the mechanism by which FAT10 changes JunB activity on cytokine promoters.

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Discussion

Ambra1 interacts with TRIM11 in a yeast two-hybrid screen In a yeast two-hybrid screen with TRIM11 as bait, activating molecule in beclin-1-regulated autophagy (Ambra1) was identified as a putative interaction partner of FAT10 (see 6.2.18). Eleven of 96 positive interacting clones encoded for full length Ambra1. In addition Ambra1 was identified in a Mass spectrometric analysis of FAT10 interacting proteins (A. Aichem et al., manuscript submitted). Ambra1 has been recently identified as a large WD40 domain containing protein that acts as a positive regulator of Becn1 dependent program of autophagy with a crucial role in embryogenesis (Fimia et al., 2007). Ambra1 functional deficiency in mouse embryos led to severe neural tube defects associated with autophagy impairment, accumulation of ubiquitinated proteins, unbalanced cell proliferation and excessive apoptotic cell death, which implies a role for autophagy-dependent protein turnover in the control of neural development (Fimia et al., 2007). The identification of a vertebrate-specific autophagy regulator that is active in an evolutionarily conserved pathway opens up a new scenario involving autophagy in specialized roles during the development of higher eukaryotes. Moreover, dynamic interaction of Ambra1 with the dynein motor complex has been described to regulate autophagosome nucleation, after autophagy induction (Di Bartolomeo et al., 2010). Ambra1 was recently described to be recruited and to interact with the ubiquitin E3 ligase Parkin mediated by the linker domain of Parkin and the N-terminal region of Ambra1, but no evidence for Parkin-mediated ubiquitination of Ambra1 has been found. A model describes, that Parkin ubiquitinates outer mitochondrial membrane proteins and recruits Ambra1, which induces perimitochondrial nucleation of new phagophores through its effect on class III phosphatidylinositol 3-kinase (PtdIns(3)K). The expanding phagophore then associates with LC3, which tethers the phagophore to the ubiquitinated mitochondria via p62 or other receptor proteins (Van Humbeeck et al., 2011). However, interaction of Ambra1 with Parkin serves as a key mechanism for induction of the final clearance step of Parkin- mediated mitophagy (Van Humbeeck et al., 2011), but the question, if Parkin and Ambra1 are degraded along with the mitochondria, or if they dissociate from mitochondria after setting the mitophagic process in motion, still has to be solved. Due to the clear interaction between Ambra1 and the E3 ligase TRIM11 in the screen the connection between these two proteins was further investigated, especially the possibility, if TRIM11 was able to act as a FAT10 specific E3 ligase, which mediates ligation of FAT10 to the putative substrate Ambra1.

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Discussion

Ambra1 interacts with TRIM11 in HEK293 cells The interaction between Ambra1 and TRIM11 was further investigated in HEK293 cells and verified using a co-immunoprecipitation approach (see 6.2.19). To prevent proteasomal degradation of either HIS-TRIM11 or Ambra1-MYC-FLAG, cells were treated additionally with or without MG132 for 6 h before cell lysis. A weak interaction between TRIM11 and Ambra1 could be detected after immunoprecipitation against Ambra1 and Western blot analysis against the HIS-tag of TRIM11, which could be slightly enhanced, when cells were pre- treated with MG132. Moreover, expression of Ambra1 as well as TRIM11 decreased, when co-transfected together, which could be rescued under proteasome inhibition, indicating that Ambra1 and TRIM11 become stabilized after proteasome inhibition. To map the region of TRIM11 which interacts with Ambra1, it is essential to repeat the co- immunoprecipitation assays with TRIM11 deletion constructs. Evidence has been presented that a fragment containing the B-Box and the N-terminal coiled-coil domain is responsible for interactions of many other members of the Trim protein family with their interacting partners (Meroni and Diez-Roux, 2005). The reciprocal approach would be to accomplish the experiments with Ambra1 deletion constructs and full length TRIM11, since Ambra1 contains a WD40 domain, which has been described as a protein-protein interaction motif. Ambra1 is an essential Beclin1 (Becn1) interacting protein which positively regulates autophagy. The Becn1 1-VPS34 complex plays a crucial role in the induction of the autophagic process by generating phosphatidylinositol 3-phosphate (PtdIns(3)P)-rich membranes, which act as platforms for ATG protein recruitment and autophagosome nucleation and several cofactors, such as Ambra1, ATG14 and UVRAG, are necessary for Becn1 complex formation and activity (Fimia et al., 2007; Liang et al., 1999; Sun et al., 2009). Autophagosome formation in mammalian cells is primed by Ambra1 release from the dynein motor complex. In the absence of autophagy, hVps34 and Becn1 are bound in a complex with Ambra1 and p150 (ortholog of Vps15). This complex is bound to cellular microtubules via Ambra1-dynein interactions. When autophagy is induced, Ambra1-DLC1 are released from the dynein complex in an ULK1-dependent manner, and relocalize to the perinuclear region and the endoplasmic reticulum, thus enabling autophagosome nucleation (Di Bartolomeo et al., 2010). Ambra1 downregulation or its overexpression resulted in a significant increase or decrease of cell proliferation rate, respectively (Fimia et al., 2007). These data illustrate that Ambra1 is an indispensible regulator of autophagy initiation and thereby involved in the process of cellular degradation, but hitherto nothing is known about how Ambra1 itself becomes degraded. Since it was identified as an interacting partner of TRIM11, it was further investigated, if Ambra1 could be modified with ubiquitin or FAT10 via TRIM11

170

Discussion

Ambra1 interacts non-covalently with ubiquitin and FAT10 Apart from being an interaction partner of Ambra1, we were hence further interested in the role of TRIM11 as a RING E3 ligase, which might mediate the covalent linkage of either ubiquitin or FAT10 to Ambra1, which potentially would label the protein for degradation through the proteasome pathway (see 6.2.20). We could show that Ambra1 became co-immunoprecipitated with either ubiquitin or FAT10 in HEK293 cells, whereupon MG132 treatment increased the amount of co-immunoprecipitated Ambra1. However, no conjugate formation of Ambra1 was visible neither with ubiquitin nor FAT10. In addition no modification with either ubiquitin or FAT10 was detectable when TRIM11 was over-expressed, suggesting, that Ambra1 becomes non-covalently linked to ubiquitin or FAT10. Moreover, co-expression of TRIM11 even decreased the protein amount of either Ambra1, ubiquitin or FAT10, similar to the case, when JunB, ubiquitin and FAT10 were co-expressed together with TRIM11. These data indicate that non-covalent interaction of Ambra1 with ubiquitin, FAT10 and TRIM11 presumably modulates the function of Ambra1 but not necessarily assigns it for proteasomal degradation.

The proteasome is involved in Ambra1 degradation The previous experiments indicated that Ambra1 becomes stabilized when the proteasome was inhibited. Ambra1 has been described to be an indispensible regulator of autophagy initiation and is thereby involved in the process of cellular degradation (Di Bartolomeo et al., 2010; Fimia et al., 2007; Liang et al., 1999; Sun et al., 2009), but hitherto nothing is known about how Ambra1 itself become degraded and no data were available about the protein half life of Ambra1. Cycloheximide experiments in HEK293 cells indeed provided evidence that Ambra1 becomes degraded via the proteasome pathway, since treatment with MG132 led to a substantial accumulation of Ambra1 to 160 % (see 6.2.21). Though, Ambra1 protein level slowly decreased to ~80 % after 5 h of cycloheximide treatment, indicating that Ambra1 is a long-lived protein. We next asked the question, if FAT10 co-expression and thereby non-covalent interaction with Ambra1 has an impact on the degradation rate of Ambra1. Strikingly, co-expression of FAT10 led to considerably accelerated Ambra1 degradation rate with a protein half-life of about 2.5 h, whereas the turnover rate of FAT10 remained unvaried (~1 h), compared to the half life of single transfected monomeric FAT10 (~1 h). Besides, MG132 treatment did not rescue the Ambra1 degradation, indicating that co-expression of FAT10 alters the degradation rate of Ambra1 and interaction leads presumably to another degradation machinery than the proteasome.

171

Discussion

Two major clearance mechanisms, autophagy and the ubiquitin (Ub)–dependent proteolytic system (UPS), have been described to govern the regulation of intracellular proteolysis. Although proteasomal and autophagy mediated lysosomal degradation use distinct components, several lines of evidence suggest that these pathways may be linked by specific mechanisms. Recent elucidation of selective autophagy has challenged the traditional notion that the two clearance mechanisms are mutually exclusive (Kim et al., 2008; Kraft et al., 2010). The ubiquitin ligase Parkin for instance, was reported to target substrates for both, proteasomal and autophagic degradation, raising questions about the signals and recognition mechanisms that distinguish these degradation routes (Moore, 2006; Narendra et al., 2008; Olzmann et al., 2007). Moreover, several proteins involved in the recognition of autophagy targets are ubiquitin-binding proteins, such as p62/SQSTM1, which contains a carboxy-terminal ubiquitin-associated (UBA) domain, whereby p62 seems to act as an adaptor between ubiquitinated protein aggregates and the autophagic machinery (Bjorkoy et al., 2005; Kim et al., 2008; Pankiv et al., 2007). Moreover, p62 and NBR1, contain a zinc-finger domain, raising the possibility that adaptor components of the UPS that contain zinc-finger domains, such as E3 ligases, might also play a role in selective autophagy (Tasaki and Kwon, 2007). Interestingly, p62 has also been found to be covalently as well as non-covalently linked to FAT10 (A. Aichem et al., manuscript submitted).

In a second cycloheximide approach, we studied the effects of bafilomycin A1, a potent and specific inhibitor of vacuolar H+ ATPase (V-ATPase), on the process of Ambra1 degradation. Cells treated for 20 h with bafilomycin and additionally for 5 h with cycloheximide increased the Ambra1 protein level to ~475 %, whereas co-expression with FAT10 led to a decrease of ~30 % (data not shown). These data strongly suggest the existence of a parallel degradation mechanism for Ambra1, either the proteasomal or autophagic pathway, where a non- covalent interaction with FAT10 seems to play a crucial role to assign Ambra1 for degradation. An interesting experiment would be to repeat co-immunoprecipitation experiments with conditions of autophagy induction, like amino-acid starvation, for instance, which was further tested in immunofluorescence microscopy (see 6.2.22). Further work will shed light on the question, which pathway and which particular conditions are required for Ambra1 degradation. Identification of protein substrates and other components of selective autophagy will aid in the understanding of autophagy interaction network in different cell types and under different physiological conditions (Behrends et al., 2010).

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Discussion

Ambra1 co-localizes with FAT10 in punctuated structures We could already show in co-immunoprecipitation assays that Ambra1 interacted with either ubiquitin or FAT10 in a non-covalent manner. Recently, a link between FAT10 involvement in autophagic processes has been discovered, given that p62/SQSTM, a autophagosomal receptor and signaling adapter was found to be covalently as well as non-covalently linked to FAT10 (A. Aichem et al., manuscript submitted). Moreover, some studies indicate the link of FAT10 to aggregates. FAT10 interacts and co-localizes with histone deacetylase 6 (HDAC6) in aggresomes under conditions of proteasome inhibition in a microtubule dependent manner, suggesting that HDAC6 functions as a linker between the dynein motor complex and FAT10 and FAT10ylated protein cargos (Kalveram et al., 2008). In light of these results, we decided to visualize the non-covalent interaction of Ambra1 and FAT10 with confocal laser-scanning microscopy and to analyze the role of non-covalent interaction in this process (see 6.2.22). Moreover, we wanted to monitor, if the interaction was accompanied by a change in Ambra1 or FAT10 localization. Co-expression of Ambra1 and FAT10 resulted in a clear co-localization in cytosolic punctuated structures, which were not apparent, when cells were singly transfected with either Ambra1 or FAT10, suggesting that Ambra1 and FAT10 indeed interact and further led to trans-location into punctuated structures. Firmia et al. (Fimia et al., 2011) could recently show that autophagy induction resulted in an increased localization of Ambra1 in the perinuclear region. Co-localization studies revealed that a high percentage of Ambra1 relocated to the endoplasmic reticulum, where it partially co-localized with the omegasomes, the sites of autophagosome production. In starvation- induced autophagy, the ER, the outer mitochondrial membrane, the Golgi and even the plasma membrane have all been implicated as sources of autophagosomal membranes. We further tested, if under autophagy conditions, in this case amino acid starvation, the localization of either Ambra1 or FAT10 would be altered. We could observe the formation of punctuated structures, presumably autophagosomes mainly in the cytosol in Ambra1 transfected cells. Besides, Ambra1 and FAT10 co-localized together in punctuated structures in double-transfected cells. Unfortunately, we cannot affirm that punctuated structures are autophagosomes, because no staining with specific autophagosome recognizing antibodies (e.g. LC3) could be performed, due to time limitations.

Besides, the dynamic interaction of Ambra1 with the dynein motor complex has been described to regulate the mammalian autophagy, where ULK1 dependent phosphorylation of Ambra1 leads to a release of the autophagy core complex from dynein (Di Bartolomeo et al., 2010). At this point the Ambra1/Vps34/Beclin1 complex is relocated to the ER or other phagophore assembly sites (PAS) (Fimia et al., 2011).

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Discussion

Therefore, Ambra1 constitutes a direct regulatory link between ULK1 and Becn1-VPS34, which is required for core complex positioning and activity within the cell and there exist a strict and regulatory relationship between cytoskeleton dynamics and autophagosome formation. An interesting experiment regarding Ambra1 and FAT10 localization after inhibiting the activity of ULK1 would give a hint, if FAT10 and Ambra1 co-localize in autophagosomes or if interaction is impaired. Co-immunoprecipitation assays and cycloheximide experiments under proteasome inhibition indicated, that the proteasome is involved in the degradation of Ambra1 and FAT10. To further investigate, if proteasome inhibition has an impact on either FAT10 or Ambra1 localization or distribution in the cell, HEK293 transfected cells were treated with MG132 (10 μM) for 6 h. Interestingly, co-transfection of Ambra1 and FAT10 led to a clear co- localization in punctuated structures widely distributed in the cytosol as well as in the nucleus. Moreover, cells which were singly transfected with Ambra1 showed formation of punctuated structures in the cytosol but considerably reduced in size, compared to amino- acid starved cells. To investigate the sub-cellular distribution of either FAT10 or Ambra1 under proteasomal inhibition and additional autophagy induction, Ambra1 and FAT10 transfected cells were treated for 6 with MG132 and subsequently starved in amino acid and serum-free medium for 4 h. Similar to the case when autophagy was induced, in Ambra1-transfected cells an increased number of punctuated cytosolic structures were apparent which were considerably augmented in size. Due to time limitations no size quantification of punctuated structures was performed, hence these observations are only estimated values. Further experiments are pertinent to analyze Ambra1 and FAT10 localization in different subcellular compartments, with specific antibodies, especially, subcellular markers for ER, autophagosomes, mitochondria, Golgi cisternae, early endosomes, and lysosomes. Especially, staining with LC3 would be helpful to analyze, if appearing punctae, where a clear co-localization of FAT10 and Ambra1 was detectable, are components of formed autophagosomes, and thereby provide a possible link of Ambra1 and FAT10 complexes to autophagy.

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References

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201

Abbreviations

9 Abbreviations 3-AT 3-Amino-1,2,4-triazole 3-MA 3-Methyladenine aa amino acid Ab antibody ALCL anaplastic large cell lymphoma Ambra1 activating molecule in Beclin1-regulated autophagy1 AMP adenosine monophosphate APC anaphase promoting complex AP-1 activating protein-1 APAF-1 apoptotic protease activating factor 1 ARC105 activator-recruited cofactor 105 kDa component ATF activating transcription factor Atg Autophagy related genes ATP adenosine triphosphate Becn1 Beclin1 BIR baculoviral IAP repeat BIRP BIR containing protein BSA bovine serum albumin bp base pair bZIP basic leucine zipper BIRC6 Baculoviral IAP repeat-containing protein 6 Bif-1 Endophilin B1 BRUCE BIR repeat containing ubiquitin-conjugating enzyme cAMP cyclic adenosine monophosphate cDNA complementary DNA CBP CREB binding protein CMA chaperone mediated autophagy CML chronic myelogenous leukemia CNS central nervous system C-terminal carboxy-terminal CRE cAMP response element CREB cAMP response element binding protein CUL cullin dATP Deoxyadenosine triphosphate dCTP Deoxycytinde triphosphate dGTP Deoxyguanosine triphosphate dNTP Deoxynucleotide triphosphate dTTP Deoxythymidine DBD DNA binding domain DMEM Dulbecco’s Minimal Essential Medium DMSO dimethyl sulfoxide DNA desoxyribonucleic acid DTT dithiothreitol DUB deubiquitinating enzyme E. coli Eschericha. coli E1 ubiquitin-activating enzyme E2 ubiquitin-conjugating enzyme E3 ubiquitin protein ligase E6-AP E6-associated protein EGFR epidermal growth factor receptor ERAD endoplasmic reticulum associated degradation ER endoplasmic reticulum ERK extracellular signal-regulated kinase 202

Abbreviations

FAT10 HLA-F adjacent transcript 10 FCS fetal calf serum FLRF fetal liver ring finger Fos Finkel-Biskis-Jinkins osteosarcoma virus oncogene Fra Fos-related antigen G-CSF granulocyte colony-stimulating factor GFP green fluorescent protein GMP1 GTPase-activating protein (GAP)-modifying protein 1 GR Glucocorticoid Receptors Grp 78 Glucose related protein 78 GSH Glutathione GST glutathione S-transferase h hour hMTIIa human metallothionein IIa HA hemagglutinin HCC hepatocellular carcinoma HDAC histone deacetylase HEK human embryonic kidney cells HECT homologous to E6-AP carboxyl terminus HIF hypoxia-inducible factor HIV Human immunodeficiency virus HL Hodgkin lymphoma HRP horseradish peroxidase IAP Inhibitor of apoptosis protein IFN interferon IκB inhibitor of NF-κB IP immunoprecipitation IR inverted repeat ISG15 interferon stimulated gene 15 iTreg induced regulatory T cell JNK Jun N-terminal kinase kDa kilo Dalton LPS lipopolysaccharide MAD2 mitotic arrest deficient 2 Maf musculoaponeurotic fibrosarcoma MAPK mitogen activated protein kinase MDM2 murine double minute 2 MEF mouse embryonic fibroblast MetOH methanol MHC major histocompatibility complex min minute mM millimolar mRNA messenger ribonucleic acid mTOR mammalian target of rapamycin NAD nicotinamide adenine dinucleotide NEDD8 neural precursor cell-expressed developmentally down regulated 8 NEMO NF-κB essential modulator NF-ΚB nuclear factor 'kappa-light-chain-enhancer' of activated B-cells NFAT Nuclear factor of activated T-cells NK cells Natural killer cells Nrdp1 neuregulin degrading protein 1 N-terminal amino-terminal NUB1 NEDD8 ultimate buster 1 NUB1L NEDD8 ultimate buster 1-Long

203

Abbreviations

OD optical density PAGE polyacrylamide gel electrophoresis Pax6 Paired box gene 6 PBS phosphate buffered saline PCNA proliferating cell nuclear antigen PCR polymerase chain reaction PE phosphatidylethanolamine PG prostaglandin Phox2b paired-like homeobox 2b PI3K Phosphoinositide 3-kinase PIC1 PML-interacting protein 1 PINK-1 PTEN-induced kinase I PKA protein kinase A PML promyelocytic leukemia tumor suppressor PTM post-translational modification RBCC RING/B-box/Coiled Coil RFP RET finger protein RING really interesting new gene RNA ribonucleic acid RNF41 ring finger protein 41 RPMI Roswell Park Memorial Institute RT Room temperature RTEC renal tubular epithelial cell SAM signaling lymphocyte-activation molecule SAPK stress activated protein kinase SCF sodium dodecyl sulfate SDS Skp1-Cullin-F-box protein SEM Standard error of the mean siRNA small interfering RNA SLE Systemic lupus erythematosus Smac second mitochondria-derived activator of caspase smARF short form of ARF Smt3p suppressor of MIF2 mutations-3 protein Smurf1 Smad Ubiquitination Regulatory Factor-1 SUMO small ubiquitin-like modifier SV40 simian virus 40 TEMED N,N,N’,N’- Tetramethylethylenediamine TGF-β transforming growth factor β TGN trans-golgi network TIF1 transcriptional Intermediary Factor 1 TNF tumor necrosis factor TPA 12-O-Tetradecanoyl-phorbole-13-acetate TRE TPA responsive element TRIM tripartite motif Trx Thioredoxin Ub ubiquitin UBA ubiquitin-associated UBC ubiquitin-conjugating catalytic domain Ubc9 ubiquitin carrier 9 UBD ubiquitin binding domain UBE1 ubiquitin-like modifier activating enzyme 1 UBL ubiquitin-like protein UDP ubiquitin-domain protein UFM1 ubiquitin-fold modifier 1 UIM ubiquitin-interacting motif

204

Abbreviations

ULM ubiquitin-like modifier URM1 ubiquitin-related modifier 1 USE1 UBA6-specific E2 enzyme 1 USP8/UBPY ubiquitin-specific protease 8 / Y UVRAG UV irradiation resistance-associated tumor suppressor gene VEGF vascular endothelial growth factor VMP1 vacuole membrane protein 1 WB Western blot wt wild type μg microgram μl microliter μM micromolar

Amino Acids Ile I Isoleucine Leu L Leucine Lys K Lysine Met M Methionine Phe F Phenylalanine Pro P Proline Ser S Serine Thr T Threonine Trp W Tryptophan Tyr Y Tyrosine Val V Valline Asx B Aspartic acid / Asparagine Gx Z Glutamic acid / Glutamine

205

Addendum

10 Addendum

Mass spectrometry results 1.1 )

# # # MW calc Accession Coverage Score Description PSMs Peptides AAs [kDa] . pI Heat shock 70 kDa protein 1A/1B P08107 30,89 22 12 641 70,0 5,66 762,14 OS=Homo sapiens GN=HSPA1A PE=1 SV=5 - [HSP71_HUMAN] Protein arginine N-methyltransferase 5 O14744 24,49 17 11 637 72,6 6,29 449,95 OS=Homo sapiens GN=PRMT5 PE=1 SV=4 - [ANM5_HUMAN] Calcium-binding mitochondrial carrier protein Aralar2 OS=Homo sapiens Q9UJS0 23,11 12 11 675 74,1 8,62 378,31 GN=SLC25A13 PE=1 SV=2 - [CMC2_HUMAN] Calcium-binding mitochondrial carrier protein Aralar1 OS=Homo sapiens O75746 7,96 5 4 678 74,7 8,38 171,82 GN=SLC25A12 PE=1 SV=2 - [CMC1_HUMAN] Signal recognition particle 68 kDa protein Q9UHB9 11,80 6 5 627 70,7 8,56 150,02 OS=Homo sapiens GN=SRP68 PE=1 SV=2 - [SRP68_HUMAN] Lamin-B2 OS=Homo sapiens GN=LMNB2 Q03252 4,00 4 2 600 67,6 5,35 122,74 PE=1 SV=3 - [LMNB2_HUMAN] Pescadillo homolog OS=Homo sapiens O00541 5,78 3 3 588 68,0 7,33 106,53 GN=PES1 PE=1 SV=1 - [PESC_HUMAN]

1.2)

# # # MW calc. Accession Coverage Score Description PSMs Peptides AAs [kDa] pI Heat shock cognate 71 kDa protein (Heat P11142 18,42 10 7 646 70,9 5,52 304,16 shock 70 kDa protein 8) - Homo sapiens (Human) - [HSP7C_HUMAN] Stress-70 protein, mitochondrial precursor (75 kDa glucose-regulated protein) (GRP P38646 14,87 8 7 679 73,6 6,16 244,45 75) (Peptide-binding protein 74) (PBP74) (Mortalin) (MOT) - Homo sapiens (Human) - [GRP75_HUMAN] 78 kDa glucose-regulated protein precursor (GRP 78) (Heat shock 70 kDa protein 5) (Immunoglobulin heavy chain- P11021 12,08 8 6 654 72,3 5,16 229,33 binding protein) (BiP) (Endoplasmic reticulum lumenal Ca(2+)-binding protein grp78) - Homo sapiens (Human) - [GRP78_HUMAN] ATP-dependent RNA helicase DDX3X O00571 7,85 4 4 662 73,2 7,18 121,66 OS=Homo sapiens GN=DDX3X PE=1 SV=3 - [DDX3X_HUMAN]

206

Addendum

# # # MW calc. Accession Coverage Score Description PSMs Peptides AAs [kDa] pI Heat shock 70 kDa protein 1A/1B P08107 58,19 100 25 641 70,0 5,66 3338,33 OS=Homo sapiens GN=HSPA1A PE=1 SV=5 - [HSP71_HUMAN] Heat shock 70 kDa protein 1L (Heat shock 70 kDa protein 1-like) (Heat shock P34931 19,34 45 8 641 70,3 6,02 1667,14 70 kDa protein 1-Hom) (HSP70-Hom) - Homo sapiens (Human) - [HS70L_HUMAN] X-ray repair cross-complementing protein 6 OS=Homo sapiens P12956 15,27 9 8 609 69,8 6,64 293,84 GN=XRCC6 PE=1 SV=2 - [XRCC6_HUMAN] Desmoglein-1 OS=Homo sapiens Q02413 3,34 5 2 1049 113,7 5,03 190,77 GN=DSG1 PE=1 SV=2 - [DSG1_HUMAN] Calcium-binding mitochondrial carrier protein Aralar2 OS=Homo sapiens Q9UJS0 14,81 7 7 675 74,1 8,62 190,39 GN=SLC25A13 PE=1 SV=2 - [CMC2_HUMAN] Interferon-induced, double-stranded RNA-activated protein kinase OS=Homo P19525 10,89 5 5 551 62,1 8,40 185,99 sapiens GN=EIF2AK2 PE=1 SV=2 - [E2AK2_HUMAN] Calcium-binding mitochondrial carrier protein Aralar1 OS=Homo sapiens O75746 8,26 6 5 678 74,7 8,38 168,88 GN=SLC25A12 PE=1 SV=2 - [CMC1_HUMAN] Desmoplakin (DP) (250/210 kDa paraneoplastic pemphigus antigen) - P15924 1,81 7 5 2871 331,6 6,81 168,05 Homo sapiens (Human) - [DESP_HUMAN] Pescadillo homolog OS=Homo sapiens O00541 8,33 4 4 588 68,0 7,33 151,73 GN=PES1 PE=1 SV=1 - [PESC_HUMAN] Junction plakoglobin OS=Homo sapiens P14923 7,65 7 5 745 81,7 6,14 141,58 GN=JUP PE=1 SV=3 - [PLAK_HUMAN] Transcription factor jun-B OS=Homo P17275 11,24 3 2 347 35,9 9,22 103,88 sapiens GN=JUNB PE=1 SV=1 - [JUNB_HUMAN]

207

Addendum

# # # MW calc. Accession Coverage Score Description PSMs Peptides AAs [kDa] pI Heat shock cognate 71 kDa protein (Heat P11142 26,47 22 12 646 70,9 5,52 729,26 shock 70 kDa protein 8) - Homo sapiens (Human) - [HSP7C_HUMAN] Stress-70 protein, mitochondrial precursor (75 kDa glucose-regulated protein) (GRP P38646 34,02 24 16 679 73,6 6,16 716,16 75) (Peptide-binding protein 74) (PBP74) (Mortalin) (MOT) - Homo sapiens (Human) - [GRP75_HUMAN] 78 kDa glucose-regulated protein precursor (GRP 78) (Heat shock 70 kDa protein 5) (Immunoglobulin heavy chain- P11021 23,55 15 11 654 72,3 5,16 451,99 binding protein) (BiP) (Endoplasmic reticulum lumenal Ca(2+)-binding protein grp78) - Homo sapiens (Human) - [GRP78_HUMAN] Heat shock 70 kDa protein 1A/1B P08107 18,88 10 8 641 70,0 5,66 350,69 OS=Homo sapiens GN=HSPA1A PE=1 SV=5 - [HSP71_HUMAN] Heat shock 70 kDa protein 1L (Heat shock 70 kDa protein 1-like) (Heat shock 70 kDa -P34931 10,30 6 5 641 70,3 6,02 233,99 protein 1-Hom) (HSP70-Hom) - Homo sapiens (Human) - [HS70L_HUMAN] Heat shock 70 kDa protein 6 (Heat shock P17066 6,38 5 3 643 71,0 6,14 195,41 70 kDa protein B') - Homo sapiens (Human) - [HSP76_HUMAN] Melanoma-associated antigen D2 Q9UNF1 8,42 3 3 606 64,9 9,32 146,32 OS=Homo sapiens GN=MAGED2 PE=1 SV=2 - [MAGD2_HUMAN] X-ray repair cross-complementing protein P12956 6,08 3 3 609 69,8 6,64 136,84 6 OS=Homo sapiens GN=XRCC6 PE=1 SV=2 - [XRCC6_HUMAN] ATP-dependent RNA helicase DDX3X O00571 7,25 5 4 662 73,2 7,18 133,77 OS=Homo sapiens GN=DDX3X PE=1 SV=3 - [DDX3X_HUMAN] Transcription factor jun-B OS=Homo P17275 16,71 3 3 347 35,9 9,22 109,93 sapiens GN=JUNB PE=1 SV=1 - [JUNB_HUMAN]

208

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Predicted Post-Translational Modification Sites (JunB) by profile Hidden Markov Model (HMM) (http://dbptm.mbc.nctu.edu.tw/)

Accessible Locations Substrate Secondary Modification Surface Resource (AA) Sites Structure Area (%)

70 Phosphoserine(CDK) GSYFSGQGS CCCCCCCCC 23.21 HMM Pred.

101 O-linked_Threonine_Man GVITTTPTP CEEECCCCC 21.47 HMM Pred.

102 O-linked_Threonine_Man VITTTPTPP EEECCCCCC 19.26 HMM Pred.

104 Phosphothreonine(CDK) TTTPTPPGQ ECCCCCCCC 39.77 HMM Pred.

113 Methylarginine YFYPRGGGS EECCCCCCC 45.04 HMM Pred.

153 Phosphothreonine(MAPK) MNHVTPPNV HCCCCCCCC 26.02 HMM Pred.

158 O-linked_Serine_GalNAc PPNVSLGAT CCCCCCCCC 27.94 HMM Pred.

158 O-linked_Serine_GlcNAc PPNVSLGAT CCCCCCCCC 27.94 HMM Pred.

173 Phosphotyrosine(Abl) PGGVYAGPE CCCCCCCCC 14.76 HMM Pred.

184 N-linked_Asparagine PVYTNLSSY CCCCCCCCC 21.90 HMM Pred.

186 O-linked_Serine_GalNAc YTNLSSYSP CCCCCCCCC 29.33 HMM Pred.

192 O-linked_Serine_Man YSPASASSG CCCCCCCCC 31.87 HMM Pred.

195 O-linked_Serine_GalNAc ASASSGGAG CCCCCCCCC 40.75 HMM Pred.

206 O-linked_Serine_GalNAc VGTGSSYPT CCCCCCCCC 26.91 HMM Pred.

206 O-linked_Serine_Man VGTGSSYPT CCCCCCCCC 26.91 HMM Pred.

207 O-linked_Serine_GalNAc GTGSSYPTT CCCCCCCCC 36.15 HMM Pred.

207 O-linked_Serine_Man GTGSSYPTT CCCCCCCCC 36.15 HMM Pred.

210 O-linked_Threonine_Man SSYPTTTIS CCCCCCCCC 28.48 HMM Pred. 212 O-linked_Threonine_Man YPTTTISYL CCCCCCCCC 15.48 HMM Pred. 215 Phosphotyrosine(Jak) TTISYLPHA CCCCCCCCC 22.02 HMM Pred.

237 Phosphoserine(PKB) GRGASTFKE CCCCCHHCC 36.88 HMM Pred.

245 Phosphothreonine(PKC) EEPQTVPEA CCCCCCCCC 49.06 HMM Pred.

302 Phosphothreonine(PKC) DKVKTLKAE HHHHHHHCC 36.16 HMM Pred.

Pred. Indicates Predicted

209