The chromatin architectural protein DEK: assessing its chromatin binding properties
Von der Fakultät für Mathematik, Informatik und Naturwissenschaften der RWTH Aachen University zur Erlangung des akademischen Grades eines Doktors der Naturwissenschaften genehmigte Dissertation
Vorgelegt von
Master of Science Haihong Guo
aus Shanxi, China
Berichter: Universitätsprofessor Dr. rer. nat. Bernhard Lüscher Universitätsprofessor Dr. rer. nat. Ralph Panstruga
Tag der mündlichen Prüfung: 08.03.2016
Dieses Dissertation is auf den Internetseiten der Hochschulbibliothek online verfügbar
Publications:
Parts of this thesis will be submitted for publication:
1) Guo, H., Prell, M., Königs, H., Waldmann, T., Meister, M., Becker, C., Hermans-Sachweh, B. & Kappes, F. “Interrogating protein function via bacterial growth”. Manuscript in preparation and attached to this thesis (see 7.3 in Appendix).
Additional results, not explicitly mentioned in this work but obtained during my doctoral studies, will be also submitted for publication:
2) Smith, E. A., Krumelbeck, E. F., Gole, B., Willis, N., Adams, A., Matrka, M. C. Jegga, A. G., Guo, H., Meetei, A. R., Andreassen P. R., Kappes, F., Scully R., Wiesmüller, L. and Wells, S. I. “Loss of the DEK oncogene disrupts homologous recombination by destabilizing RAD51 filament formation”. Manuscript in preparation.
3) Prell, M., Preisinger, C., Markovitz, D., Guo, H., Schilling, N., and Kappes, F. "Novel chromatin modulatory functions for the TREX component THOC 4 (ALY/Ref)". Manuscript in preparation.
4) Prell, M., Guo, H., Preisinger, C., Ostareck, D., Ostareck-Lederer, A., Naarmann-de Vries, I., and Kappes, F. "Proteomic Interrogation reveals wide-spread functions of the DEK oncogene". Manuscript in preparation.
5) Prell, M., Preisinger, C., Guo, H., Jonak, C., Waidmann, S., and Kappes, F. "The DEK oncogene regulates alternative splicing through selective interaction with SRPK1 and SRPK2". Manuscript in preparation.
Poster presentations:
Guo, H., and Kappes, F. (2013). A rapid technique to identify and mutate DNA binding domains in proteins using bacterial toxicity as read-out. 44th Annual Conference of the German Genetics Society (GfG): Genetics 2013 (Braunschweig, Germany).
Guo, H., Prell, M., Königs, H., Hermanners-Sachweh, B., and Kappes, F. (2014). Studying DNA-binding features of chromatin architectural factors via bacterial growth. Danube Scientific Conference on Epigenetics (2014) (Budapest, Hungary).
Acknowledgements
I would like to thank my supervisor, Ferdinand Kappes, for giving me the opportunity to work in his group and to work on this amazing project. You have taught me a lot during the last four years and your guidance helped me to develop skill not only for scientific research but also for living in Germany. This was my best experience to be your student. It would not have been possible to complete this work without you.
My great gratitude is also to Prof. Dr. Bernhard Lüscher for being my official supervisor and providing a lot of excellent and critical comments.
I would like to thank Prof. Dr. Ralph Panstruga for being my “Zweitgutachter”.
Many thanks to my colleagues in Lab 15! Thank you, Malte, for being a good partner in the
DEK group and always helping and encouraging me. I would like to thank Dr. Jörg Hartkamp for providing a lot of precious advice on my project. Christiane Becker, thank you for managing the excellent scientific environment in the lab, and especially thanks for taking the immunofluorescence photos in my thesis. Many thanks go to every other member in Lab 15, including Fabian, Mathias, Nahleen, Tim, Irina, Sandra, Elke, Jacky and Tobias. I am so delighted and privileged to have worked with you every day.
Special thanks to Dr. Juliana Lüscher-Firzlaff and Prof. Dr. Gerhard Mueller-Newen for providing me with a lot of vectors for my work and Hildegard Schmitz-van-de-Leur for helping with cloning problems. And also to my colleagues in the Lüscher group, including Barbara,
Carolina, Jörg, Marc, Mareike, Weili, Laura and Patricia, thank you for providing a scientific environment in the lab and the help in the lab and for helping me in the hot lab. Thanks to
Hiltrud Königs for taking the EM photos in this thesis. Thanks to Jinyu for helping to edit the photos of protein secondary structure.
I also would like to thank other members in the institute. Thanks to Allison for correcting my thesis and always encouraging me. And also Carolin, Susana, Thomas, Willy, Natalie and
Marcel, thank you everybody for giving me a lot of help during the last four years and helping
me feel happy and welcome in this big team.
I am grateful to the China Scholarship Council (CSC) and to START for supporting me the last four years.
最后，我特别感谢我的父母。你们永远是我坚实的后盾，在背后默默地支持我，在我疲
惫，迷茫的时候给我靠岸的港湾。焉得谖草，言树之背，养育之恩，无以回报，你们永
远健康快乐是我最大的心愿。感谢我的姐姐，妹妹还有弟弟，祝你们天天开心，工作顺
利。另外，我也要感谢男朋友黄旭, 在认识的两年中，你给予我很多工作和生活上的关 心和鼓励，我非常的感激。在以后相伴的日子里，希望我们能够互相鼓励互相扶持，也
祝愿你能早日完成课题，顺利毕业。还有我最亲爱的朋友王晓宇同学，从硕士到博士，
从北京到德国，这六年，我们一起努力奋斗，相互鼓励,相互安慰。你就像是我的亲人， 希望你身体健康，早日博士毕业，也祝我们的友情地久天长。感谢杨建花同学，很开心
认识你并且成为你的室友，非常感谢你对我研究课题提出的很宝贵的建议。在论文完成
之际，也特别感谢所有亚琛的朋友们，非常开心能够在德国认识你们，让我在异国他乡
感受到亲人和朋友的温暖，和你们的相处丰富了我的生活，让我不再感到孤单和害怕，
你们的鼓励和支持让我变的更加勇敢，在这里请接受我最真挚的谢意！
Table of Contents I
Table of Contents
Table of Contents ...... I
List of Figures ...... IV
List of Tables...... VI
Abstract ...... VII
Zusammenfassung ...... IX
1. Introduction ...... 1 1.1 Chromatin ...... 1 1.2 Modifications of chromatin ...... 3 1.2.1 DNA methylation ...... 4 1.2.2 Histone modifications ...... 4 1.2.3 Histone variants ...... 5 1.2.4 ATP-dependent chromatin remodelers...... 6 1.2.5 Non-histone chromatin architectural proteins ...... 7 1.3 Protein motifs for DNA binding ...... 12 1.3.1 Helix-turn-helix (HTH) motifs ...... 13 1.3.2 Zinc finger motifs ...... 13 1.3.3 Leucine zipper ...... 13 1.3.4 Helix-loop-helix (HLH) ...... 14 1.3.5 HMG-box motifs ...... 14 1.3.6 SAP motif ...... 15 1.4 The DEK protein ...... 17 1.4.1 Identification of DEK and its association with human diseases ...... 17 1.4.2 Structure and molecular functions of DEK ...... 18 1.4.3 Cellular functions of DEK ...... 20
2. Aim of this Thesis ...... 25
3. Materials and Methods ...... 26 3.1 Materials ...... 26 3.1.1 Chemicals ...... 26 3.1.2 General buffers ...... 28 3.1.3 Oligonucleotides ...... 29 3.1.4 Plasmids ...... 31 3.1.5 Vectors ...... 34 3.1.6 Enzymes ...... 34 3.1.7 Antibodies ...... 35 3.1.8 E. coli strains ...... 35 3.2 Methods ...... 36 3.2.1 General DNA-related methods ...... 36
Table of Contents II
3.2.2 Protein-related methods ...... 39 3.2.3 Cell culture ...... 43 3.2.4 RNA isolation and cDNA synthesis ...... 44 3.2.5 Real-time PCR (q-PCR) ...... 44 3.2.6 QuickChange site-directed mutagenesis ...... 45 3.2.7 Random mutagenesis by error-prone PCR ...... 45 3.2.8 Bacterial assays ...... 46 3.2.9 Electrophoretic Mobility Shift Assay (DNA-EMSA) ...... 48 3.2.10 Chromatin assembly assay in vitro ...... 48 3.2.11 Creation of HeLa S3 DEK knockout cells ...... 49 3.2.12 Expression of DEK fragments in DEK knockout cells and HeLa S3 cells using lentiviral delivery ...... 49 3.2.13 Cell fractionation ...... 50 3.2.14 Analysis of chromatin changes with MNase digestion ...... 50 3.2.15 RNA pull down assay ...... 51 3.2.16 In situ hybridization for total poly(A)-mRNA localization ...... 52 3.2.17 Northwestern assay ...... 53 3.2.18 Preparation of Pentaprobe RNA for RNA band shift assay ...... 54 3.2.19 RNA electrophoretic mobility shift assay (RNA-EMSA) ...... 54
4. Results ...... 56 4.1 DEK changes chromatin structure at the nucleosomal level in vitro and in cells ...... 57 4.2 Hypothesis-driven design for the creation of “loss-of-DNA-binding” DEK mutants ...... 59 4.2.1 Analysis of DNA binding of the pseudo-SAP/SAP-box of DEK carrying mutations of conserved aromatic amino acids ...... 61 4.2.2 Analysis of DNA binding of the pseudo-SAP/SAP-box of DEK carrying mutations of arginines ...... 64 4.3 The DNA binding activities of DEK interfere with bacterial growth ...... 67 4.3.1 Heterologous expression of DEK in bacteria inhibits growth in liquid cultures ...... 67 4.3.2 DNA binding activities of DEK appear to be responsible for bacterial growth inhibition 69 4.3.3 Activities of the pseudo-SAP/SAP-box domain of DEK are responsible for bacterial growth inhibition ...... 71 4.3.4 Bacterial growth inhibition is due to abundant and aberrant compaction of bacterial DNA by DEK ...... 72 4.4 Establishment of a screening procedure to select “DNA-binding-dead” DEK mutants: Bacterial Growth Inhibition Screen (BGIS) ...... 76 4.5 Detailed analyses of obtained “DNA-binding-deficient” DEK mutants ...... 81 4.6 Analysis of nucleosomal accessibility with a DEK mutant in vitro ...... 85 4.7 Investigation of the relevance of the DNA binding function of DEK in cells ...... 87 4.7.1 Creation of HeLa S3 DEK knockout cells ...... 87 4.7.2 Creation of an inducible, lentivirally-delivered expression system based on the pTRIPZ vector ...... 89 4.7.3 Assessment of the subnuclear distribution of DEK fragment 1-187 ...... 90 4.7.4 The DNA binding and folding activities of DEK are important for global chromatin
Table of Contents III
organization ...... 91 4.8 Identification of the RNA-binding domain of DEK via the BGIS ...... 93 4.9 Applications of the BGIS ...... 99 4.9.1 DNA- and RNA-associated folding activities in the bacterial growth inhibition assay .... 99 4.9.2 Enzymatic activities inhibit bacterial growth ...... 107
5. Discussion ...... 111 5.1 DNA binding and folding activities of DEK and their functions in nucleosomal DNA accessibility ...... 112 5.1.1 Creation of “DNA-binding-dead” DEK mutants ...... 113 5.1.2 Molecular analysis of the “DNA-binding-dead” DEK mutant #11 ...... 115 5.1.3 Analysis of nucleosomal DNA accessibility with DEK mutant #11 ...... 117 5.1.4 Other mutants selected via BGIS ...... 119 5.2 Identification of the RNA-binding domain of DEK ...... 120 5.2.1 RNA binding proteins (RBPs) ...... 120 5.2.2 DEK187-270 is the RNA-binding domain ...... 121 5.3 The functions of DEK within chromatin ...... 123 5.4 The bacterial growth inhibition screen (BGIS) and its applications ...... 127 5.4.1 DNA-/RNA- binding/folding domains in the BGIS ...... 128 5.4.2 Enzymatic activities in the BGIS ...... 130 5.4.3 The BGIS—a useful novel tool for investigating protein functions ...... 131
6. References ...... 132
7. Appendix ...... 148 7.1 Abbreviations ...... 148 7.2 Supplementary figures ...... 151 7.3 Manuscript of publications ...... 156 7.4 Curriculum vitae ...... 194 7.5 Eidesstattliche Erklärung ...... 195
List of Figures IV
List of Figures
Figure 1.1: Structure of a eukaryotic cell and nucleus...... 1 Figure 1.2: Chromatin organization in the eukaryotic cell (modified from (Tonna et al., 2010))...... 2 Figure 1.3: Overview of a selection of important mechanisms in the regulation of chromatin organization...... 3 Figure 1.4: Histone modifications (Bhaumik et al., 2007; Latham and Dent, 2007)...... 4 Figure 1.5: Classification of ATP-dependent chromatin remodelers and their functions in chromatin...... 7 Figure 1.6: Schematic depiction of the structure of HP1 protein...... 9 Figure 1.7: Schematic depiction of the structure of HMG proteins...... 10 Figure 1.8: Schematic depiction of the structure of HMGB1...... 11 Figure 1.9: Schematic depiction of the structure of SAF-A (modified from (Schwander, 2004))...... 12 Figure 1.10: Schematic depiction of the known functional domains and known structural elements of DEK...... 18 Figure 4.1: Current models for DEK functions within the regulation of chromatin structure...... 57 Figure 4.2: DEK alters chromatin structure in general and at the nucleosomal level...... 58 Figure 4.3: Proposed workflow for creation and analyses of “loss-of-function” DEK mutants...... 60 Figure 4.4: Schematic depiction of point mutations in DEK created by Devany et al...... 61 Figure 4.5: DEK-DNA docking model for the identification of amino acids involved in DNA binding...... 62 Figure 4.6: Analyses of recombinant GST-DEK carrying aromatic amino acid mutations via EMSAs...... 63 Figure 4.7: Analyses of recombinant GST-DEK carrying the triple mutation F96/97/182A via EMSA...... 64 Figure 4.8: Depictions of the positions of arginines in the pseudo-SAP/SAP-box using the DEK-DNA docking model...... 65 Figure 4.9: Analyses of recombinant GST-DEK carrying arginine mutations via EMSAs...... 66 Figure 4.10: Summary of all mutations created and tested so far either by Devany et al. (top) or within this thesis (bottom)...... 66 Figure 4.11: DEK expression in E. coli inhibits bacterial growth...... 68 Figure 4.12: Expression of human DEK or Drosophila Dek in E. coli inhibits bacterial colony growth...... 70 Figure 4.13: The pseudo-SAP/SAP domain in DEK is responsible for bacterial growth inhibition...... 71 Figure 4.14: Analysis of DEK-DNA interactions in bacteria using confocal microscopy...... 73 Figure 4.15: Analysis of bacterial DNA via electron microscopy...... 75 Figure 4.16: DEK expression in bacteria results in reduced expression of the16S rRNA gene...... 76 Figure 4.17: The Bacterial Growth Inhibition Screen—BGIS...... 78 Figure 4.18: Confirmation of DNA-binding-dead mutants in the re-transformation assay...... 79 Figure 4.19: Schematic depiction of mutants obtained from the bacterial growth inhibition screen...... 80 Figure 4.20: DEK85-187 mutants #8, #11, and #18 show no signs of bacterial growth inhibition...... 82 Figure 4.21: Electron microscopy assay with DEK mutants...... 82 Figure 4.22: Analysis of DEK85-187 mutants #8, #18 and #11 via EMSA...... 84 Figure 4.23: EMSA analysis of individual mutations of the DEK mutant #11 reverted back to the wt condition. . 85 Figure 4.24: Analysis of nucleosomal accessibility with the DNA-binding-dead DEK1-187 mutant #11...... 86 Figure 4.25: HeLa S3 DEK knockout cells created via a TALEN approach...... 88 Figure 4.26: Cloning of an inducible, lentivirally-delivered expression system (pTRIPZ) and generation of stable cell lines via lentiviral transduction...... 90 Figure 4.27: Assessment of subcellular distribution of DEK fragments...... 91
List of Figures V
Figure 4.28: DEK mutant #11 fails to reduce the nuclear size of DEK knockout cells...... 92 Figure 4.29: Expression of DEK187-270 induces bacterial growth inhibition...... 94 Figure 4.30: DEK187-260 is the potential RNA-binding domain of DEK...... 96 Figure 4.31: Analysis of DEK fragments through a north-western approach...... 97 Figure 4.32: Binding of GST-DEKwt and DEK truncations to ssRNA Pentaprobe sequences...... 99 Figure 4.33: Expression of SAF-A and HMGB1 inhibit bacterial growth...... 100 Figure 4.34: Expression of ASH2L does not induce bacterial growth inhibition...... 102 Figure 4.35: Both DNA- and RNA-associated activities of SAF-A are capable of inducing bacterial growth inhibition...... 103 Figure 4.36: Identification of DNA-binding domains of ALY/REF by the bacterial inhibition screen...... 104 Figure 4.37: EMSAs with ALY/REF full-length and truncations...... 105 Figure 4.38: Proteins containing no DNA or RNA-binding activities exhibit no effect on bacterial growth...... 107 Figure 4.39: Bacterial inhibition assay with CK2 enzyme subunits...... 108 Figure 4.40: Bacterial inhibition assay with PRMT1 and PRMT3...... 108 Figure 4.41: Bacterial inhibition assay with ARTD10 truncations...... 109 Figure 5.1: Depiction of the identified mutations in the DEK mutant #11 in DEK-DNA docking model...... 116 Figure 5.2: Overview of the potential RNA-binding residues in DEK187-270 predicted by BindN program. .... 122 Figure 5.3: Summary of functional domains in DEK with their identified functions...... 123 Figure 5.4: Current models for DEK functions within chromatin...... 126 Supplementary Figure S1: Selection for DEK mutants lacking bacterial growth inhibition by replica stamping the colonies after transformation to LB-Amp plates plus IPTG...... 151 Supplementary Figure S2: Immunoblotting with bacterial colonies transformed pGEX-DEK1-187 and selected from bacterial inhibition screen...... 152 Supplementary Figure S3: Immunoblotting with bacterial colonies transformed pGEX-DEK85-187 and selected from bacterial inhibition screen...... 154 Supplementary Figure S4: Binding of GST-DEKwt and DEK truncations to RNA Pentaprobe sequences...... 155
List of Tables VI
List of Tables
Table 1: Classes of modifications found on histones and their functional relevance (adapted from (Kouzarides, 2007)) ...... 4 Table 2: Histone variants and their functions (modified from (Ramaswamy and Ioshikhes, 2013; Talbert and Henikoff, 2010)) ...... 6 Table 3: Nomenclature for the HMG chromosomal proteins (modified from (Bustin, 2001b)) ...... 10 Table 4: List of mutations in the pseudo-SAP/SAP-box domain (created by Devany et al.) ...... 61 Table 5: List of conserved aromatic amino acids in the pseudo-SAP/SAP-box domain ...... 62 Table 6: List of arginine to alanine mutations in the pseudo-SAP/SAP-box domain ...... 65 Table 7: List of mutations obtained in DEK1-187 and DEK85-187 via BGIS ...... 81 Table 8: Comparison of protein functions between DEK, SAF-A, HMGB1 and HP1 ...... 125
Abstract VII
Abstract
Chromatin is a highly-ordered, yet dynamic structure that is essential in the regulation of all
DNA-dependent processes, with the coordinated interplay of non-histone chromosomal factors playing important roles in its regulation. One such factor is the unique oncoprotein DEK, which has been implicated in the maintenance of heterochromatin via protein-protein-mediated mechanisms. However, given its distinct DNA binding and folding activities, it is highly possible that DEK is directly involved in the regulation of chromatin organization. Our results presented here demonstrate that DEK indeed binds to the entry/exit site of nucleosomal DNA in vitro, and that knocking down DEK expression in cells leads to general alterations in the nucleosomal repeat length of in cellular chromatin. As this aspect has not been studied before, multiple hypothesis-driven strategies were chosen to create
“DNA-binding-dead” DEK mutants by site-directed mutagenesis. Though these initial approaches proved unsuccessful we found that expression of DEK in bacteria resulted in massive compaction of the bacterial genome due to DEK’s DNA-folding activity which resulted in a complete halt in bacterial growth. This fact in combination with random mutagenesis led to the development of a novel screening procedure, now termed bacterial growth inhibition screen (BGIS) that allowed for the selection of loss-of-function mutants, which were subsequently verified by biochemical analyses. By re-expression of one
DNA-binding-dead mutant in newly created DEK knockout cells, first evidence was provided that the DNA binding and folding activities of DEK indeed are involved in the regulation of global chromatin structure.
By subjecting other DEK domains with previously unidentified functions to the BGIS, we were able to identify and verify a hitherto unrecognized RNA-binding domain in DEK.
Prompted by these findings, we next tested whether artificial interference with the biology of bacteria as a tool can also be exploited for investigation of other protein classes. Through these initial studies, we found that DNA- and RNA-binding/folding activities as well as enzymatic activities from several proteins indeed resulted in complete growth inhibition, suggesting a
Abstract VIII rather broad applicability of the BGIS. Despite some limitations to this screen, we believe it could aid in the discovery of functional domains in proteins with yet unknown functions.
Together with the finding that enzyme-dead mutants did not affect bacterial growth, this opens up possibilities for establishing simple, quick and economical ways for high-throughput approaches for the purpose of drug screening using compound libraries.
In conclusion, we established and validated a unique and surprisingly simple screen based on undirected interference with bacterial growth, which allowed us to gain deeper insights into the biology of DEK’s DNA binding activities in the cellular environment. Additionally, due to the seemingly wide applicability and easy implementation in the molecular biology laboratory, we believe that the BGIS may be a valuable addition to the toolbox of methods for the study of protein function.
Zusammenfassung IX
Zusammenfassung
Chromatin ist eine hoch organisierte, jedoch dynamische Struktur, welche essentiell für die
Regulation aller DNA-abhängigen Funktionen ist. Hierbei spielt das koordinierte
Zusammenspiel von nicht-Histone-chromosomalen-Proteinen eine wichtige Rolle. Das
Onkoprotein DEK ist ein solches Protein. Es wurde schon mit der Regulation von
Heterochromatin über Protein-Protein-vermittelte Funktionen in Zusammenhang gebracht.
Basierend auf seinen DNA-bindenden und -faltenden Funktionen könnte DEK auch eine direkte Rolle in der Regulation der Chromatinorganisation spielen. Neue Ergebnisse zeigen, dass DEK in vitro an die Ein- und Austrittsstelle der nukleosomalen DNA binden kann und dass ein Knock-down von DEK in Zellen zu globalen Veränderungen der nukleosomalen
DNA-Länge im zellulären Chromatin führt. Da dieser Aspekt bisher noch nicht untersucht worden ist, wurden, basierend auf diversen Hypothesen, mehrere Strategien angewandt, um nicht mehr DNA-bindende „loss-of-function“-Mutanten von DEK mittels gerichteter
Mutagenese zu generieren. Dieser Ansatz war jedoch nicht zielführend. Es wurde allerdings gefunden, dass bei der Expression in Bakterien die DNA-bindenden Eigenschaften von DEK zu einer massiven Verdichtung des bakteriellen Genoms führten, was schlussendlich einen
Wachstumsstop zur Folge hatte. Basierend auf diesem Befund und in Kombination mit ungerichteter Mutagenese, wurde eine neue Screening-prozedur etabliert, der sogenannte
„Bacterial Growth Inhibition Screen“ (BGIS). Dieser erlaubte die Identifikation von
„loss-of-function“-Mutanten, die durch nachfolgende biochemische Untersuchungen verifiziert wurden. Durch Re-expression einer DNA-Bindungs-defizienten Mutante in neu etablierten DEK Knock-out Zellen, konnten erste Hinweise erbracht werden, dass die
DNA-Faltungsfunktion von DEK in der Tat eine wichtige Rolle in der Regulation der allgemeinen Chromatinstruktur hat.
Weitere BGIS-Untersuchungen von DEK-Bereichen mit unbekannten Funktionen, führten zur
Identifizierung einer bisher unbekannten RNA-Bindedomäne in DEK. Als nächstes wurde untersucht, ob die durch die Expression von Proteinen hervorgerufene Interferenz mit der
Zusammenfassung X
Biologie der Bakterien auch als Möglichkeit für die Identifikation von anderen Proteinklassen herangezogen werden kann. Es konnte in der Tat gezeigt werden, dass diverse DNA- und
RNA-interagierende als auch enzymatische Aktivitäten zu einem Wachstumsstop in Bakterien führten, was auf eine recht breite Anwendbarkeit des BGIS hindeutet. Daher glauben wir, dass dieser Screen mit gewissen Einschränkungen bei der Identifikation von Domänen mit bisher unbekannter Funktion in Proteinen hilfreich sein kann. Da weiterhin Proteinbereiche mit inaktivierter enzymatischer Funktion keinen Einfluss auf das bakterielle Wachstum hatten, könnte BGIS für einfache, schnelle und ökonomische Anwendungen zur Selektion von bioaktiven Substanzen angewendet werden.
Zusammengefasst wurde eine neue Screening-Prozedur (BGIS) etabliert und verifiziert, welche tiefere Einblicke in die Biologie der DNA-Bindung des DEK-Proteins erlaubte. Durch die mögliche breite Anwendbarkeit dieses Screens sowie die einfache Etablierung in jedem molekularbiologischen Labor, glauben wir, dass der BGIS eine hilfreiche neue Methode für das Studium von Proteinfunktionen darstellen könnte.
1 Introduction 1
1. Introduction
1.1 Chromatin
The compartmentalization of discrete biological processes into membrane-enclosed organelles is a distinguishing feature of eukaryotic cells, with the nucleus being the most prevalent compartment (Figure 1.1). The nucleus contains the majority of genetic information, with the large DNA molecules being organized together with histones and a number of non-histone proteins to form the physiological template of DNA, called chromatin.
Figure 1.1: Structure of a eukaryotic cell and nucleus. Shown is a simplified schematic picture of a eukaryotic cell including subcellular structures like mitochondria, endoplasmatic reticulum, cytoskeleton and membrane-enclosed vesicles (bottom). On the top a very simplified schematic overview of the nucleus is given, showing its principle subnuclear organization (from http://www.shmoop.com/biology-cells/all-eukaryotic-cells.html).
On the one hand, the organization of chromatin structure and functions must be rather stable to allow for the establishment of distinct cell- and development-specific functional states. On the other hand, this highly complex and dynamic structure regulates virtually all DNA-dependent processes and therefore plays a vital role in all cellular functions, and therefore must be rather flexible as it is crucial for the response to environmental change (Croston and Kadonaga, 1993;
Kamakaka and Kadonaga, 1993; Paranjape et al., 1994; Wasylyk and Chambon, 1979). The fundamental repeating unit of chromatin is the nucleosome, comprising 146 base pairs of DNA wrapped in 1.67 left-handed turns around a core histone octamer consisting of a central tetramer of histones H3 and H4 and two dimers of histones H2A and H2B, respectively (Luger et al.,
1997). The nucleosome core particles, together with the connecting “linker” DNA, form a
“beads-on-a-string” fiber with a diameter of 11 nm, representing the first level of higher-order
1 Introduction 2 chromatin organization. The linker histone H1, a variant of non-histone proteins, connects two nucleosomes and facilitates the further compaction of chromatin (~50 packing ratio) into the 30 nm fiber which is the second structural level of chromatin organization (Robinson et al., 2006).
While the folding of the chromatin fiber to the next higher level remains much more obscure
(Ausio, 2015) and is currently the subject of intense research (Robinson and Rhodes, 2006;
Tremethick, 2007), it is traditionally suggested that the 30 nm fiber is arranged into higher-order loops with the help of scaffold proteins, among others. Even further compaction leads to a highly condensed chromatin structure as seen in the mitotic chromosomes, which is crucial in ensuring faithful transmission of the complete genetic information to daughter cells (Figure
1.2).
Figure 1.2: Chromatin organization in the eukaryotic cell (modified from (Tonna et al., 2010)). 146 bp of genomic DNA (2 nm in diameter) is wrapped around a histone octamer to form the fundamental core particles termed nucleosomes. With the stabilization of histone H1, the nucleosomes are further compacted leading to the formation of the “10 nm fiber”. Further compaction leads to the appearance of chromatin loops and to distinct and varying densities of chromatin structure (30 nm fiber). The maximal compaction is represented by the mitotic chromosomes (approximately 1,400 nm in width).
Based on cytological staining patterns, unique chromatin compaction states in cellular interphase chromatin can be distinguished, which were historically termed eu- and heterochromatin. Euchromatin comprises a lightly stained, and therefore loose, more accessible structure where the biggest portion of the actively transcribed genome is located, while heterochromatin, which is characterized by intense staining, is highly condensed and consists of mostly genetically inactive satellite sequences. However, using state-of-the-art techniques,
1 Introduction 3 more than two distinct chromatin forms with defined functional features have been identified
(Roudier et al., 2011; van Steensel, 2011) and recent evidence shows that even heterochromatic areas show transcriptional activity (Saksouk et al., 2015).
1.2 Modifications of chromatin
A large number of studies have indicated that dynamic changes between distinct chromatin states influence gene expression (Apostolou and Hochedlinger, 2013; Boskovic et al., 2014;
Schneider and Grosschedl, 2007). Furthermore, cell type- and development-specific chromatin homeostasis is orchestrated by distinct mechanisms, collectively termed epigenetics, which regulate gene activity and expression without alterations to the underlying DNA sequence. So far, several mechanisms and concepts have been proposed (Figure 1.3), with the most important ones being DNA methylation (Bannister and Kouzarides, 2011; Jaenisch and Bird,
2003), histone modifications (Bannister and Kouzarides, 2011; Gillette and Hill, 2015), histone variants (Henikoff and Smith, 2015; Li and Fang, 2015), chromatin remodeling (Bowman,
2010; Clapier and Cairns, 2009), and the more diffuse functions of non-histone chromatin architectural proteins (Zlatanova et al., 2008).
Figure 1.3: Overview of a selection of important mechanisms in the regulation of chromatin organization. Modified from www.abcam.cn/epigenetics/dna-methylation-a-guide, http://www.integratedhealthcare.eu/1/en/histones_and_chromatin/1497/ and (Clapier and Cairns, 2009; Zlatanova and van Holde, 1996). Details are found in section 1.2.1 to 1.2.5.
1 Introduction 4 1.2.1 DNA methylation
DNA methylation usually takes place at the 5’position of cytosines within CpG dinucleotides
(so-called CpG islands), and it is catalyzed by a group of enzymes termed DNA methyltransferases (DNMTs). Studies have shown that DNA methylation contributes to the control of gene expression and has roles in the regulation of nuleosomal structure and genomic integrity (Bird, 2002; Choy et al., 2010).
1.2.2 Histone modifications
The core histones consist of a global domain and a more flexible N-terminus (the unstructured tail). With the help of modification-specific antibodies, mass spectrometry, and a series of other techniques, a number of distinct histone post-translational (PTM) and chemical modifications have been identified (Figure 1.4) (Kouzarides, 2007), which typically coincide with a specific transcriptional activity and a distinct higher-order chromatin organization (Table 1).
Figure 1.4: Histone modifications (Bhaumik et al., 2007; Latham and Dent, 2007). (A) Amino acids with their possible post-translational (PTM) and chemical modifications. (B) Overview of well-characterized post-translational modifications occurring on histones. Most PTMs are found on the N-terminal tails; however the histone fold domain (core) and the C-terminal tails can also be modified. See Table 1 for an overview of their functional relevance.
Table 1: Classes of modifications found on histones and their functional relevance (adapted from (Kouzarides, 2007)) PTM Modifications Functions regulated Acetylation K-ac Transcription, Repair, Replication, Condensation Methylation (lysines) K-me1, K-me2, K-me3 Transcription, Repair Methylation (arginines) R-me1, R-me2a, R-me2s Transcription Phosphorylation S-ph, T-ph Transcription, Repair, Condensation Ubiquitylation K-ub Transcription, Repair Sumoylation K-su Transcription ADP ribosylation E-ar Transcription Deimination R > Cit Transcription Proline isomerization P-cis > P-trans Transcription
1 Introduction 5
In the following sections, only histone lysine methylation will be highlighted as an example for both, activating and inactivating consequences to chromatin.
1.2.2.1 Histone lysine methylation
Histone lysine methylation is catalyzed by histone lysine methyltransferases (HKMTs) which can mono-, di-, or tri-methylate lysine residues at specific positions on histones, resulting in a
“mark” that in concert with downstream effectors (“readers”) can either lead to the “repression” or “activation” of specific genes or genomic regions.
Mono-, di- or trimethylation at lysine (K) 9 and 27 on histone H3 (H3K9Me and H3K27Me) and methylation at K20 on histone H4 (H4K20Me) have been shown to be related to gene repression, with one prominent example being the silencing of homeotic genes. H3K9Me3 is implicated in forming heterochromatin, resulting in silencing of transcription via a conserved mechanism involving the binding of Heterochromatin Protein 1 (HP1), which is a classic example of a “reader” (Boros et al., 2014; Lachner et al., 2001). In turn, binding of HP1 results in the recruitment of the H3K9Me-specific methyltransferase SUV39H1/H2 (KMT1 A/B), a
“writer”, which further sets repressive marks, resulting in a so-called “silencing loop” (Hediger and Gasser, 2006; Kappes et al., 2011b).
In contrast, methylation at K4, K36 and K79 on histone H3 (H3K4Me, H3K36Me and
H3K79Me) has been shown to be generally associated with gene activation. As the most prominent example, both di- and tri-methylated H3K4 are ubiquitously enriched at actively transcribed genes (Inoue et al., 2015; Shen et al., 2014). Little is known about the function of
K3K79 methylation, however, other than its involvement in preventing spreading of heterochromatin and in the maintenance of open chromatin (Schneider and Grosschedl, 2007).
1.2.3 Histone variants
Despite a highly conserved role for the canonical histones, it is now clear that these proteins exist in many variants (Table 2). These function-specific variants can be incorporated into nucleosomes, sometimes independently of replication, and function together with histone modifications in the regulation of transcription, repair, chromosome assembly and segregation
1 Introduction 6
(Shaytan et al., 2015; Talbert and Henikoff, 2010).
Table 2: Histone variants and their functions (modified from (Ramaswamy and Ioshikhes, 2013; Talbert and Henikoff, 2010)) Histone Variant Occurrence Function Canonical Universal Genome packaging H4 H4V Trypanosomes Transcription termination Canonical Widespread Genome packaging H3.1 Universal Transcription, DNA damage response CenH3 (CENP-A, others) Universal Centromere identity H3 H3.3 (H3.2 in plants) Universal Replacement H3t Mammals Testes-specific H3V Trypanosomes Transcription termination Canonical Widespread Genome packaging H2A.X Universal DNA damage response H2A.Z Universal Promoter insulation H2A MacroH2A Animals X chromosome inactivation H2A.Bbd Mammals Transcription activation H2AL1, L2 Mammals Testes-specific Canonical Universal Genome packaging TH2B Mammals Testes-specific H2B H2BFWT Mammals Testes-specific H2BV Trypanosomes Transcription termination
Histone 4 is the most conserved histone amongst all species. Only one single variant, H4V, has been found in Trypanosoma brucei, which was enriched at transcription sites (Siegel et al.,
2009). The centromere-specific histone H3 variant, CenH3 (CENP-A in mammals), is found in most higher eukaryotes and serves as a centromere-specific marker (Yoda et al., 2000). H3.3, another H3 histone variant, only differs in 4 amino acids in comparison to canonical H3, but can replace its position. Unlike the canonical H3, which is deposited strictly at replication forks during S phase of the cell cycle, the H3.3 variant is incorporated throughout the cell cycle with preferential integration both up- and downstream of transcribed genes, suggesting a role as an epigenetic mark for transcriptionally active chromatin (Ahmad and Henikoff, 2002; Chen et al.,
2013; Lund et al., 2015; Snyers et al., 2014). Like H3, several H2A variants have been described, whereas only a few H2B variants have been found, and some are testes-specific (Table 2).
1.2.4 ATP-dependent chromatin remodelers
The usually very large chromatin remodeling complexes catalyze nucleosome rearrangements by either the sliding of nucleosomes along the DNA, the alteration of nucleosomal spacing, the replacing of core histones, or the eviction of entire nucleosomes (Figure 1.5 B). These activities
1 Introduction 7 are strictly ATP-dependent. Based on the sequence homology of the specific ATPase subunit, chromatin remodelers have been classified into four families (Figure 1.5 A): SWI/SNF
(switching defective/sucrose non-fermenting) (Masliah-Planchon et al., 2015; Mohrmann and
Verrijzer, 2005; Wang, 2003), ISWI (imitation switch) (Corona et al., 2007; Corona and
Tamkun, 2004; Mellor, 2006), CHD (chromo domain, helicase, DNA binding) (Marfella and
Imbalzano, 2007; Murawska and Brehm, 2011) and INO80/SWR1 (inositol requiring 80) (Bao and Shen, 2007; Chen et al., 2011; Conaway and Conaway, 2009). Besides the similar ATPase domain, each remodeler shares unique domains reflecting different functions (Figure 1.5 A).
These remodelers promote nucleosome sliding or histone eviction, which leads to an open chromatin structure (Figure 1.5 B) and is essential for DNA replication (Au et al., 2011; Falbo and Shen, 2012; Mermoud et al., 2011), DNA repair (Aydin et al., 2014; Erdel and Rippe, 2011) and gene transcription (Park et al., 2014; Yadon et al., 2013; Zentner et al., 2013).
Figure 1.5: Classification of ATP-dependent chromatin remodelers and their functions in chromatin. (A) The groups of ATP-dependent chromatin remodelers (Clapier and Cairns, 2009). All remodeler families contain a common ATPase subunit characterized by two split parts: DExx (red) and HELICc (orange). The unique domains within the ATPase domain distinguish each family: either short insertion (gray) in SWI/SNF, ISWI, and CHD families, or long insertion (yellow) in INO80 family. Furthermore, each family contains distinct flanking domains: Bromodomain (light green) and HSA (helicase-SANT) domain (dark green) for SWI/SNF family, SANT-SLIDE module (blue) for ISWI family, tandem chromodomains (magenta) for CHD family, and HSA domain (dark green) for INO80 family. (B) Schematic depiction of the molecular mode of chromatin remodelers (modified from (Clapier and Cairns, 2009)).
1.2.5 Non-histone chromatin architectural proteins
Besides the distinct and rather well-studied mechanisms described so far, chromatin structure and accessibility can be additionally or separately regulated by the so-called “linker-protein network” (Zlatanova et al., 2008). Like linker histones, which bind to DNA between nucleosomes and control chromatin condensation and DNA accessibility, multiple other proteins have been identified that may function in a similar fashion through either cooperation with linker histones or as replacements. In the next section, the functions of some prominent
1 Introduction 8 members of the so-called “linker-protein” family, such as HP1, HMG proteins or SAF-A are discussed.
1.2.5.1 Linker histones
Linker histones play a critical role in the stabilization of the 30 nm chromatin fiber (Zlatanova and van Holde, 1996). They consist of a central globular domain and two less-structured long
C- and short N-terminal tails. The crystal structure of the globular domain exhibits a winged-helix motif which allows binding to nucleosomal DNA (Ramakrishnan et al., 1993).
The C-terminal tail plays the most important role in chromatin compaction as it bridges entering and exiting nucleosomal DNA into a four-stranded stem (Hamiche et al., 1996).
Compared to core histones, the linker histones are less conserved. The sequence of the
C-terminal tails are extremely variable, both in length and amino acid composition (Kasinsky et al., 2001). Furthermore, some histone isoforms, such as H1º and H5, are only expressed in a tissue- or development-specific fashion (Zlatanova and van Holde, 1996). Despite significant sequence homology, H5 interacts with linker DNA with much higher affinity as compared to other linker histones, and thereby irreversibly compacts chromatin and inactivates transcription (Zlatanova and van Holde, 1996). Contrarily, other linker histones including H1º compact chromatin and modulate the accessibility of regulatory proteins reversibly (Happel and Doenecke, 2009). This reversible interaction of linker histones (excluding H5) to nucleosomes results in the formation of stem structures by closing the nucleosomal gate (entry and exit site of nucleosomal DNA) and regulating the nucleosomal DNA accessibility. As proposed by Zlatanova et al., H5 “locks” the nucleosomal gate, whereas other linker histones only “close” the gate, or even keep it “ajar”, with the overall gate accessibility depending on the individual dynamic of the linker histone-nucleosome interaction (Zlatanova et al., 2008).
1.2.5.2 Heterochromatin protein 1 (HP1)
Heterochromatin protein 1 (HP1) was originally identified in Drosophila melanogaster as a dominant suppressor of position effect variegation (PEV) (Eissenberg et al., 1990; James and
Elgin, 1986). The HP1 protein family is highly conserved in evolution from fungi to humans
1 Introduction 9 and usually several isoforms exist within the same species (Lomberk et al., 2006; Zeng et al.,
2010). For instance, three main HP1 variants have been identified in mammalian cells, namely
HP1, HP1 and HP1, which all possess 50% amino-acid sequence identity to Drosophila
Hp1 (Li et al., 2002). Despite this sequence similarity, HP1, HP1 and HP1 localize to distinct genomic regions: HP1 and HP1 are mainly localized in heterochromatin, whereas
HP1 is observed in both heterochromatin and euchromatin (Minc et al., 2000).
In mammals, each HP1 paralogue has three functional regions: a conserved N-terminal chromo domain (CD), a conserved C-terminal region (chromo shadow domain (CSD)), and a less-structured and more-variable hinge region that contains the nuclear localization sequence
(Figure 1.6). Through the chromo domain (CD), HP1 binds directly and specifically to either
H3K9Me2 or Me3, which are both markers of heterochromatic gene silencing. Moreover, it was demonstrated that HP1 specifically interacts with the C-terminal globular domain of histone H3 via the CD (Nielsen et al., 2001). In contrast to the CD, the CSD is generally considered a dimerization domain (Brasher et al., 2000). Several studies have demonstrated that the hinge domain binds to DNA or RNA without any sequence specificity and can also bind to linker histones (Keller et al., 2012; Nielsen et al., 2001). Together, the CD interacts with H3K9Me and the hinge domain of HP1 stabilizes this interaction and additionally associates with histones and linker histones, leading to nucleosomal DNA inaccessibility and heterochromatin maintenance. Therefore, it has been proposed that HP1 might function as a
“lock” of the nucleosomal gate, similar to histone H5 (Zlatanova et al., 2008).
Figure 1.6: Schematic depiction of the structure of HP1 protein. The three functional domains of HP1: chromo domain (CD), hinge domain and chromo shadow domain (CSD) were indicated with dark gray, white and light gray rectangles, respectively (Modified from (Schwendemann et al., 2008)).
1.2.5.3 High mobility group (HMG) family
High mobility group (HMG) proteins are the most abundant and ubiquitous non-histone chromatin-associated proteins. HMGs modify chromatin structure by recognizing specific DNA
1 Introduction 10 structures, as opposed to nucleotide sequences via their DNA-binding motifs (Grosschedl et al.,
1994; Murphy et al., 1999). Based on their DNA-binding domains, HMG proteins are subdivided into three families: HMGA (formerly HMG-I/Y, the HMG-AT-hook family),
HMGB (formerly HMG-1/2, the HMG-box family) and HMGN (formerly HMG-14/17, the
HMG-nucleosome binding family) (Bustin, 2001b; Gerlitz et al., 2009) (Figure 1.7 and Table
3).
Figure 1.7: Schematic depiction of the structure of HMG proteins. The HMG proteins are classified into three families based on their different DNA-binding domains: AT-hook region (red) for HMGA family, HMG-boxs (blue) for HMGB family and NBD (pink) for HMGN family. Besides DNA-binding domains, each family further contains distinct domains: H1-like (dark green) domain for HMGA family, a positive-charged N-terminal and a negative-charged C-terminal domain for HMGB family and two split NLS and a CHUD domain for HMGN family, modified from (http://www.biologie.uni-regensburg.de/Zellbiologie/Grasser/Research/Architectural/and (Bustin, 2001a)).
Table 3: Nomenclature for the HMG chromosomal proteins (modified from (Bustin, 2001b)) New name Old name Subfamilies Functional motif (canonical HMGs) (canonical HMGs) HMGA AT-hook HMGA1,2,…n HMG-I/-Y/-C HMGB HMG-box HMGB1,2,…n HMG-1/-2 HMGN NBD HMGN1,2,…n HMG-14/-17 Abbreviations: HMG, high mobility group; NBD, nucleosome binding domain.
The AT-hook motif of HMGA has little, if any, secondary structure while free in solution but transitions to a highly-ordered structure upon DNA binding (Reeves, 2001). This transition promotes the alteration of DNA structure by bending, unwinding, or inducing of looping in linear DNA (Chen et al., 2010; Ozturk et al., 2014). HMGA1 alters DNA structure into a more open state and orchestrates the assembly of transcriptional complexes to regulate gene expression (Bouallaga et al., 2000; Bouallaga et al., 2003; Catez et al., 2004).
HMGB proteins are highly expressed in the nucleus and contain a homologous domain
(HMG-box) (Figure 1.7). This domain consists of around 75 amino acids with an L-shaped fold formed by three -helices (Murphy et al., 1999; Read et al., 1993; Weir et al., 1993). The most abundant non-histone protein in the nucleus, HMGB1, contains two conserved HMG-boxes (A and B) and a long acidic C-terminal tail (Figure 1.8). A characteristic DNA binding feature of
1 Introduction 11
HMGB1 is its preference for distorted DNA (Stros, 2010; Stros et al., 2007), suggesting a role in
DNA damage repair (Ito et al., 2015; Lange and Vasquez, 2009). The C-terminal acidic tail regulates the functions of HMGB1 and is involved in destabilization of nucleosomes by replacement of linker histone H1 which results in a switch of chromatin states (Cato et al., 2008;
Thomas and Stott, 2012). Based on the interaction with and protection of linker DNA (An et al., 1998; Nightingale et al., 1996; Zlatanova and van Holde, 1998), a mechanism was postulated that HMGB proteins bind to the entry and exit site of nucleosomal DNA and directly bind to or compete with linker histones for these common binding sites (Travers,
2003). This binding loosens the nucleosomal gate (“ajar”) and enhances accessibility for chromatin remodeling complexes and also for transcription factors (Travers, 2003; Zlatanova et al., 2008)
Figure 1.8: Schematic depiction of the structure of HMGB1. The two functional HMG-boxes (A and B boxes) and the acidic tail are indicated with green, pink and black rectangles, respectively (modified from (Harris et al., 2012)).
The third family of HMG proteins, HMGN proteins, contains a unique and conserved nucleosomal binding domain (NBD), a bipartite nuclear localization sequence (NLS) and a chromatin-unfolding domain (CHUD) (Figure 1.7). HMGN proteins are present in the nuclei of all mammals and function as architectural factors that modify chromatin structure (Bustin,
2001a; Kugler et al., 2012; Rochman et al., 2010). The interactions between HMGN proteins and nucleosomes via their nucleosomal binding domain can decrease the chromatin condensation and increase the DNA accessibility by interfering with the binding of H1, which could subsequently enhance replication, transcription and DNA repair from chromatin templates (Furusawa et al., 2015; Gerlitz, 2010; Rochman et al., 2009).
1.2.5.4 SAF-A
The nuclear protein SAF-A (hnRNP-U), which is a component of heterogeneous nuclear ribonucleoprotein complexes, was initially shown to specifically bind to scaffold/matrix attachment region DNA elements (S/MARs) and may have functions in the organization of the
1 Introduction 12 controversially discussed nuclear matrix (Fackelmayer et al., 1994; Romig et al., 1992). It contains two functional domains: the N-terminus conveys DNA binding via a SAP-box and the
C-terminal RGG domain containing RNA-binding affinity (Figure 1.9) (Kipp et al., 2000;
Lobov et al., 2001; Schwander, 2004). The RGG domain is responsible for interaction of SAF-A with XIST RNA, which contributes to the silencing of X-linked genes (Helbig and Fackelmayer,
2003; Pullirsch et al., 2010). The DNA-binding domain, namely the SAP-box, conveys
DNA-looping in addition of other functions (Fackelmayer et al., 1994; Gohring and
Fackelmayer, 1997).
Figure 1.9: Schematic depiction of the structure of SAF-A (modified from (Schwander, 2004)). Shown the functional domains of SAF-A: a N-terminal SAP-box (also called SAP-box, yellow) conveys SAR/MAR DNA binding, a C-terminal RGG-box (red) that located between amino acids 714-738 has RNA-binding activity, and two NLS (black). The N-terminal is rich in acidic amino acids such as aspartic acid, glutamic acid and glutamine; with the C-terminal RGG-box containing 129 amino acids and being rich in glycine.
Taken together, higher-ordered chromatin structure is regulated on many different levels by a magnitude of distinct mechanisms: from the DNA level to histone modifications and histone variants, from histones to chromatin remodelers, and further to the conformation of loop domains in the 30 nm chromatin fiber governed by the action of non-histone architectural proteins. All these modulations of chromatin can be regulated by different mechanisms, with the possibility of crosstalk generating complicated and highly complex regulatory circuits in order to enable, maintain, or change the plasticity and dynamics of chromatin.
1.3 Protein motifs for DNA binding
Most proteins affecting the structure of chromatin mediate their functions via protein-DNA interactions. Surprisingly, only one third of these factors bind in a DNA sequence-specific manner, whereas the other protein-DNA interactions occur in a sequence-independent manner.
In the case of DNA sequence-specific DNA-protein interactions, binding is mostly mediated by hydrogen bonds, ionic interactions and/or van der Waals interactions occurring mostly in the major groove of the DNA helix. In contrast, non-sequence-specific interactions with DNA
1 Introduction 13 usually occur between protein functional groups and the DNA sugar-phosphate backbones, thus these interactions are much weaker than sequence-specific interactions, as will be discussed below.
1.3.1 Helix-turn-helix (HTH) motifs
The helix-turn-helix (HTH) motif was the first DNA binding module originally identified in bacterial proteins (Matthews, 2001). It contains two -helices, formed by approximately 20 amino acids, and a short extended amino acid sequence termed “turn”, which connects the two
-helices. In a typical protein-DNA interaction, the second helix (called the recognition helix), which contains distinct amino acids recognizes specific DNA sequences and inserts into the major groove of DNA (Anderson et al., 1981; McKay and Steitz, 1981). In eukaryotes, the HTH motif exists as a modified version termed the homeodomain. This modified HTH folds into three helices, where helix 2 and 3 function in a similar fashion as the secondary helix in bacterial proteins, and are involved in recognizing specific DNA sequences.
1.3.2 Zinc finger motifs
The second most important DNA binding motif, the zinc finger, was first discovered in the transcription factor IIIA (TFIIIA) from Xenopus laevis (Miller et al., 1985). As the name implies, zinc finger proteins incorporate one or two Zn2+ ions in their structure. Both X-ray and nuclear magnetic resonance (NMR) studies revealed that the global structure of the zinc finger motif consists of a two-stranded antiparallel -sheet (containing two cysteines), an -helix
(containing two histidines) and with the zinc ion located in the center and stabilized by the former two parts (Lee et al., 1989; Pavletich and Pabo, 1991). Similar to the HTH motif, the
-helix is responsible for interacting with the major groove of DNA and many zinc finger repeats are required to ensure DNA binding specificity.
1.3.3 Leucine zipper
The leucine zipper was identified in eukaryotic transcription factors such as the GCN4 protein in yeast (Landschulz et al., 1988). These transcriptional factors consist of two parallel, left-handed -helix monomers, which allow for dimerization in a region of approximately 30
1 Introduction 14 residues. Each -helix reveals a periodic repetition of leucine residues at every seventh position.
Since these leucine residues surface at approximately in two-turn intervals and on the same side of helix, the term “leucine zipper” was proposed. Through interactions mediated by the leucine-rich region, two monomers dimerize via the C-termini, whereas the N-termini allow for sequence-specific DNA interactions in the major groove (Krylov and Vinson, 2001; Vinson et al., 1989; Xu and Morrical, 2001).
1.3.4 Helix-loop-helix (HLH)
Another important DNA binding motif that is related to the leucine zipper is the helix-loop-helix
(HLH) motif. The HLH motif similarly contains two left-handed -helices, however, these helices differ in their length and are connected by a variable loop. This structure allows for a specific interaction with DNA and also for protein-protein interactions with other
HLH-containing factors. In a given DNA binding complex, the HLH proteins exist as dimers with the basic amino acids in the -helices in each monomer bridging the specific contacts with the major groove of the DNA (Xu and Morrical, 2001).
1.3.5 HMG-box motifs
As mentioned above, HMG proteins are recognized as the most abundant group of non-histone chromosomal proteins and are subdivided according to their DNA-binding domains (Figure 1.7 and Table 3).
1.3.5.1 AT-hook motif
The AT-hook, the DNA binding motif in HMGA proteins (Figure 1.7 and Table 3), has a consensus sequence of Pro-Arg-Gly-Arg-Pro (with Arg-Gly-Arg-Pro being invariant). It is highly conserved through evolution and one or more AT hooks can be found in proteins that are able to bend, straighten, unwind or induce looping in linear DNA molecules (Reeves, 2001).
NMR studies revealed that the AT-hook forms a C-shaped structure which interacts with the minor groove of adenine-thymine (AT)-rich DNA (Huth et al., 1997). Different from the sequence-specific DNA binding proteins, HMGA proteins can recognize specific DNA structures independent of the nucleotide sequence, and have a preference for binding to
1 Introduction 15 distorted DNA, such as supercoiled DNA and four-way junction DNA.
1.3.5.2 HMG-box motif
All the HMGB proteins share a highly conserved DNA binding motif (HMG-box) that was originally identified through sequence comparisons between the human transcription factor
UBF (hUBF) and that of mammalian HMG1 (Jantzen et al., 1990). Since then, numerous other
HMG-box-containing proteins have been identified. In mammals, these proteins are generally classified into two major groups depending on the motif abundance, function, and DNA binding specificity. The first group consists of proteins with two HMG-boxes and a long highly acidic
C-terminal tail, which is commonly found in canonical HMGB proteins. The second group consists of proteins with mostly a single HMG-box and no acidic C-terminal tails (Stros et al.,
2007). By NMR spectroscopy, the structures of HMG-boxes in HMGB1 (HMG-A box and
HMG-B box) have been determined (Bianchi et al., 1992). The HMG-box consists of approximately 75 amino acids and exhibits an L-shape formed by two arms at an angle of ~80°.
The shorter arm consists of helices I and II, while the longer arm is composed of the extended
N-terminal region and helix III. The overall structure of the motif is maintained by a cluster of conserved residues (Bianchi et al., 1992). In addition to the difference in HMG-box number between the two groups, the other characterizing feature is DNA binding specificity. Generally, two or more HMG-boxes, like those of the proteins in the first group, recognize distorted DNA structure with weak or no sequence specificity, whereas proteins with only one HMG-box, as found mostly in transcription factors, bind to DNA in both a structure- and a sequence-specific manner (Stros, 2010). Furthermore, the DNA-interaction of the HMG-box is accompanied by intercalation of hydrophobic amino acid residues into the minor groove, inducing structural changes to the major groove (Thomas and Travers, 2001).
1.3.6 SAP motif
The SAP or SAF motif (named after SAF-A/B, Acinus and PIAS), subsequently referred to as
SAP-box, was identified as a putative DNA-binding motif by Aravind and colleagues (Aravind and Koonin, 2000) and was independently found in the same year in the SAF-A protein directly
1 Introduction 16
(Kipp et al., 2000). Like the HMG-box motif, this motif also recognizes non-B-form DNA structures and its binding to DNA results in alterations to DNA topology. SAP motifs are found in a number of chromatin-associated proteins, such as the scaffold attachment factor SAF-A and
SAF-B, the DNA repair proteins Ku70, human Acinus, the DNA repair protein Rad18, and the proto-oncogene protein DEK (Aravind and Koonin, 2000; Bohm et al., 2005). Sequence alignments revealed that the SAP-box is a 35-residue motif that forms a helix-extended region containing a number of conserved hydrophobic and charged amino acids as well as an invariant glycine in the extended loop region (Aravind and Koonin, 2000). In general, most of the
SAP-motifs found in various proteins localize to the N- or C-terminus, as seen in SAF-A/B
(N-terminus) and Ku70 (C-terminus). Direct evidence showing that the SAP motif is important for DNA binding was initially gathered from studies involving SAF-A and Ku70. The SAP motif in SAF-A recognizes and interacts with the minor groove of DNA elements called scaffold/matrix attachment regions (Kipp et al., 2000). However, another molecular modeling study has revealed that the SAP-motif in Ku70 is suitable to form electrostatic and hydrophobic interactions with either the major or the minor groove of DNA (Hu et al., 2012), suggesting that
SAP-box-containing proteins may bind to DNA through distinct mechanisms.
Taken together, protein-DNA interaction motifs provide a platform to recognize and to bind to
DNA either sequence-specifically or non-sequence-specifically. In the case of DNA sequence-specific interactions, most of the proteins interact with the major groove of DNA.
However, several proteins can also specifically establish contacts with the minor grooves of
DNA, as observed in the AT-hook motif. In addition, approximately two-thirds of proteins bind to DNA without sequence-specificity, and their interactions with DNA also play many essential roles in cells, especially in chromatin organization. The most prominent examples are histones, which bind to the DNA backbone to generate nucleosomes. Other non-sequence-specific DNA-binding proteins, such as HMG-box proteins and SAP-box proteins are also reported to be involved in regulation of chromatin structure. These proteins typically recognize the DNA structures without DNA-sequence-specificity and induce the alteration of chromatin structure, thus regulating chromatin organization. One such
1 Introduction 17 non-histone chromosomal factor that contains an SAP-box and is known to be involved in shaping chromatin and DNA structure is the DEK oncogene.
1.4 The DEK protein
1.4.1 Identification of DEK and its association with human diseases
The nuclear protein DEK was initially isolated in a subform of acute myelogenous leukemia
(AML), which harbored the rare chromosomal translocation t6:9 that results in the expression of the DEK-NUP214 fusion protein (previously termed DEK-CAN) (Soekarman et al., 1992; von
Lindern et al., 1992). Although the precise role of the DEK-NUP214 fusion still remains largely unclear, it has been shown to interfere with translation initiation and may control cell proliferation, differentiation, and survival of blood cells. Therefore, DEK-NUP214 is considered a prognostic marker for survival and relapse in leukemia (Ageberg et al., 2008;
Broxmeyer et al., 2012; Crans and Sakamoto, 2001; Libura et al., 2003). Due to these discoveries, DEK itself has been classified as a putative proto-oncogene.
Rather quickly after its initial cloning, the DEK protein and DEK-specific auto-antibodies were found in a variety of autoimmune diseases, including juvenile idiopathic arthritis (JIA)
(Mor-Vaknin et al., 2011), systemic lupus erythematosus (SLE) (Wichmann et al., 2000), and sarcoidosis, among others (Dong et al., 1998; Dong et al., 2000). Interestingly, though it is usually found in the nucleus, DEK can be secreted by macrophages either in a free form in exosomes or through passive release in a highly poly(ADP-ribosyl)ated (PARylated) state during apoptosis. Once in the extracellular space, DEK functions as a chemotactic factor and is implicated in triggering various autoimmune events (Hua et al., 2009; Mor-Vaknin et al., 2006).
DEK is also being discussed as a potential urine biomarker, as the DEK protein has been found in the urine of over 80% of bladder cancer patients (Datta et al., 2011; Sanchez-Carbayo et al.,
2003).
DEK has been found to be abundantly expressed in most higher eukaryotes (not in yeast or C. elegans), and its expression varies depending on cellular growth: it is generally upregulated in proliferating cells and shows mostly reduced expression in terminally differentiated cells
1 Introduction 18
(Ageberg et al., 2006; Khodadoust et al., 2009; Waldmann et al., 2004). In healthy tissue DEK shows low expression, and DEK overexpression is found in various tumor types including colon cancer (Carro et al., 2006; Lin et al., 2013), larynx cancer (Carro et al., 2006), bladder cancer
(Carro et al., 2006; Datta et al., 2011; Duggan and Williamson, 2004; Evans et al., 2004;
Sanchez-Carbayo et al., 2003; Wu et al., 2005), hepatocellular carcinoma (Kondoh et al., 1999;
Lin and Chen, 2013; Lu et al., 2005; Yi et al., 2015), melanoma (Kappes et al., 2011a;
Khodadoust et al., 2009), breast cancer (Liu et al., 2012; Privette Vinnedge et al., 2015; Privette
Vinnedge et al., 2012; Privette Vinnedge et al., 2011), and ovarian cancer (Han et al., 2009), among others (Adams et al., 2015; Lin et al., 2015). In summary, DEK is tightly associated with two classes of human diseases, namely disorders relating to both auto-immunity and cancer.
1.4.2 Structure and molecular functions of DEK
DEK is 43 kDa in size and contains 375 amino acids. Lacking any enzymatic activity, it is the only representative of its own protein family as no paralogs have been found thus far (Figure
1.10). Additionally, some of its functional domains show a high interspecies conservation through evolution.
Figure 1.10: Schematic depiction of the known functional domains and known structural elements of DEK. Shown is the linear sequence of DEK with known domains and their functions (yellow: pseudo-SAP-box; red: SAP-box; blue: nuclear localization sequence (NLS); green: C-terminal DNA-binding domain (DEK-C); wavy lines: position of -helices, as revealed by NMR) (Bohm et al., 2005; Devany et al., 2008; Devany et al., 2004; Kappes et al., 2004b)
The only recognizable homology to other known proteins is found within the SAP-box region, thus suggesting that DEK belongs to the family of SAP-box containing proteins (Aravind and
Koonin, 2000; Kipp et al., 2000). However, two distinguishing features set DEK apart, its
SAP-box (amino acids 137-187) is located centrally within the molecule and is accessorized by a so-called pseudo-SAP-box, located N-terminally of the SAP-box (amino acid 87-149)
(Devany et al., 2008). This special domain architecture (pseudo-SAP/SAP-box domain) makes
DEK unique and suggests distinct functions potentially not seen in other proteins. However
1 Introduction 19 there are some similarities to other SAP-box proteins. As seen in other SAP-containing proteins,
DEK’s SAP-box alone shows weak DNA binding in solution. On the other hand, many immobilized DEK SAP-boxes lead to high-affinity DNA binding with subsequent aggregation of the nucleo-protein complexes (Bohm et al., 2005; Kappes et al., 2004b). The NMR structure of the unique pseudo-SAP/SAP domain of DEK revealed a globular five-helix bundle consisting of three -helices (-helices 1-3) in the pseudo-SAP-box and two -helices (-helices 4 and 5) in the SAP-box (Devany et al., 2008). Interestingly, the pseudo-SAP-box exhibits a striking structural similarity to the SAP-box, while lacking any sequence similarity. Chemical shift mapping experiments showed that the -helices (-helices 1-3 and 5) in both domains are responsible for DNA binding, with the -helix 5 also being potentially involved in protein-protein interactions (Devany et al., 2008). This clearly shows that both the pseudo-SAP and SAP-box domains are responsible for DEK’s DNA sequence-unspecific binding activities, as would be expected from a SAP-box-containing protein. Interestingly, DEK has been independently isolated from nuclear extracts based on its DNA/chromatin supercoiling activity
(Alexiadis et al., 2000; Waldmann et al., 2002), which, together with its SAP-box, suggests that it may belong to the large and diversified class of non-histone chromosomal factors. Indeed, similar to other aforementioned non-histone chromosomal factors, DEK preferentially binds to unusual DNA structures (supercoiled DNA, four-way junction DNA, distorted DNA) in a sequence-unspecific manner and its binding also results in structural changes to DNA (Bohm et al., 2005; Waldmann et al., 2003). This DNA-folding activity is conveyed by the pseudo-SAP/SAP-box and also stimulates intermolecular catenation of circular DNA molecules, promotes DNA loop formation and DNA bending, introduces positive supercoils into protein-free DNA and leads to a strong compaction of chromatin templates (Bohm et al.,
2005; Waldmann et al., 2003; Waldmann et al., 2002).
By using two-hybrid screening, a second functional domain in DEK (amino acids 270-350) was identified as both a DNA binding and multimerization domain (Kappes et al., 2004b). As determined by NMR, this domain contains a three-helix-bundle which shows structural similarity to that of the E2F/DP transcription factor family (Devany et al., 2004). Through
1 Introduction 20
-helix 2, DEK309-375 interacts with dsDNA (Devany et al., 2004). Experimental studies have consistently shown that this domain (DEK270-350) displays DNA-binding activity without inducing DNA aggregation or preference for distorted DNA or subsequent alteration of
DNA topology (Kappes et al., 2004b). Additionally, this domain contains multiple CK2 phosphorylation sites which are involved in regulating the DNA binding and multimerization of DEK. Dephosphorylated DEK lacks protein multimerization activity and promotes
DNA-protein interactions as seen in EMSA assays; in contrast, phosphorylation of DEK decreases the DNA binding affinity and results in protein multimerization (Kappes et al., 2004a;
Kappes et al., 2004b).
These observations may suggest that DEK functions as a “linker-protein-like” factor and may be involved in the regulation of higher-order chromatin structure. However, as these functions have been studied exclusively in vitro, the relevance of DEKs DNA and chromatin folding activities in the cellular setting remains largely elusive.
1.4.3 Cellular functions of DEK
1.4.3.1 Functions in the regulation of chromatin structure
In cells, the majority of DEK (90%) localizes to chromatin. Although several studies have reported a re-distribution of DEK within chromatin during the cell cycle, DEK is, at least partly, associated with chromatin during mitosis (Kappes et al., 2001; Matrka et al., 2015). Given its
DNA binding and folding activities seen in vitro, DEK is believed to be involved in chromatin architecture. Indeed, DEK associates with chromatin in vivo and in vitro (Kappes et al., 2001).
Several ChIP studies have demonstrated that DEK localizes to euchromatin and heterochromatin where it can be found associated with active histone modifications, such as
H3K4Me3, and also repressive marks, such as H3K9Me3. However, the precise distribution of
DEK on chromatin appears to be cell type- and growth-specific. Whereas one study showed that
DEK appears to be excluded from the heterochromatic compartment and is enriched at the promoter or enhancer sites of human genes (Hu et al., 2007), another study showed that DEK can directly interact with HP1 and markedly enhance the binding of HP1to H3K9Me3 to
1 Introduction 21 maintain heterochromatin territories in human cells (Kappes et al., 2011b). Consistent with this,
DEK knockdown results in a nuclease-sensitive chromatin structure, as revealed by micrococcal nuclease (MNase) treatment and a substantial reduction of constitutive heterochromatic areas
(Kappes et al., 2011b). Furthermore, DEK appears to have histone chaperone activity under certain conditions: it interacts with histones H2A and H2B (Alexiadis et al., 2000), associates with DAXX in a complex with histone deacetylase 2 (Hollenbach et al., 2002), localizes with a subfraction of chromatin bearing acetylated histone H4 (Hu et al., 2007) and controls H3.3 deposition on chromatin dependent on phosphorylation by CK2 (Ivanauskiene et al., 2014;
Kappes et al., 2011b; Sawatsubashi et al., 2010). DEK also associates with the chromatin remodeling complex B-WICH, which is involved in the replication of heterochromatin
(Cavellan et al., 2006). Therefore, DEK shows functions of an architectural chromatin protein responsible for genomic stability, though with it’s regulation remaining poorly understood.
1.4.3.2 DEK is involved in gene transcription
DEK has furthermore been shown to be involved in the regulation of gene transcription, though the effects and the underlying mechanisms remain paradoxical and somewhat contradictory, which may be due to its multi-functionality and its various functions depending on localization on chromatin (Kappes et al., 2001). Therefore, DEK has been shown to play a dual role in both activation and repression of transcription.
DEK shows high affinity binding at transcription start sites (TSS), and appears to mark active enhancers and promoters (Sanden et al., 2014). It also interacts with the transcriptional activator
AP-2, C/EBP and MLLT 3, and thus is involved in the enhancement of transcription (Campillos et al., 2003; Hu et al., 2005; Sanden et al., 2014). In Drosophila, DEK acts as a transcriptional EcR coactivator leading to the incorporation of the histone variant H3.3
(Sawatsubashi et al., 2010). However, DEK also interacts with HP1 thereby repressing transcription and has been shown to counteract histone acetylation (Lee et al., 2008). DEK is also a substrate of the acetyltransferases CBP (CREB-binding protein), p300, and P/CAF
(p300/CBP-associated factor) (Cleary et al., 2005; Ko et al., 2006), with acetylation of DEK decreasing its affinity for DNA elements within the promoters and therefore altering its
1 Introduction 22 localization (Cleary et al., 2005). In addition, repression of transcription through DEK can also occur by its blocking of the p65 subunit of NF-B (Kim et al., 2010; Sammons et al., 2006).
1.4.3.3 DEK has functions in DNA repair
Given that DEK has a preference for distorted DNA structures, which are intermediate products in DNA damage repair pathways; it is not surprising that DEK has been associated with various aspects of DNA damage repair. First, the C-terminal portion of DEK was reported to reverse the characteristic abnormal DNA-mutagen sensitivity in fibroblasts from ataxia-telangiectasia
(A-T) patients, which indicates a role in the regulation of cellular responses to DNA damage
(Meyn et al., 1993). Furthermore, DEK was shown to be poly(ADP-ribosyl)ated, which is traditionally associated with DNA damage (Kappes et al., 2008) and DEK knockdown in HeLa cells leads to cellular hypersensitivity to genotoxic stress (Kappes et al., 2008). Consistent with this, Deutzmann et al. demonstrated that DEK promotes cell proliferation on replication fork progression via replication stress and counteracts DNA damage to protect mitotic cells and resulting daughter cells from DNA damage induced by replication stress (Deutzmann et al.,
2014).
1.4.3.4 DEK—an oncogene
Based on its DNA associated function within chromatin, DEK is involved in many areas of cell development and death, including cell proliferation, differentiation, apoptosis and senescence.
In proliferating cells, DEK is generally highly expressed, and DEK knockdown reduces cell growth and promotes differentiation (Ageberg et al., 2006; Privette Vinnedge et al., 2011).
Consistently, overexpressed DEK in keratinocytes was shown to repress differentiation and resulted in a more proliferative state (Adams et al., 2015; Wise-Draper et al., 2009).
DEK has also been shown to be an inhibitor of apoptosis and senescence (Wise-Draper et al.,
2006; Wise-Draper et al., 2005), as down-regulation of DEK promotes senescence and apoptosis in papillomavirus-positive cervical cancer cells (Wise-Draper et al., 2006;
Wise-Draper et al., 2005). Similarly, DEK knockdown in canine Transitional cell carcinoma
(TCC) also enhances apoptosis and chemo-sensitivity (Yamazaki et al., 2015); this was also
1 Introduction 23 observed in metastatic melanoma cell lines (Khodadoust et al., 2009). DEK-related inhibition of apoptosis can occur through different mechanisms, either via the destabilization of p53 and decreasing the expression of p53-target genes, or by the activation of the anti-apoptotic protein
MCL-1 (Khodadoust et al., 2009; Wise-Draper et al., 2006). Given all these observations, DEK has been classified as a true oncogene.
1.4.3.5 DEK is involved in RNA-dependent processes
On average, approximately 10% of a cellular DEK population is associated with RNA (Kappes et al., 2001), in line with its implication in multiple RNA-dependent processes. DEK was shown to interact with splicing complexes, and this association was mediated by specific interaction with SR proteins (Le Hir et al., 2000; McGarvey et al., 2000). Subsequently, DEK was found to be a component of the exon-exon junction complex (EJC) which is responsible for linking pre-mRNA splicing and post-splicing events including mRNA export and mRNA decay (Kim and Dreyfuss, 2001; Le Hir et al., 2001). However, follow-up studies have questioned these findings due to cross-reactivity of the antibodies used (Reichert et al., 2002). Soares et al. reported that DEK plays a key role in splice site recognition (Mendes Soares and Valcarcel,
2006), as phosphorylated DEK was required for proofreading of 3’ splicing sites (Kress and
Guthrie, 2006; Mendes Soares and Valcarcel, 2006). Moreover, by using novel “interactome capture” approaches, DEK was identified as an mRNA-binding protein (RBPs) in proliferating
HeLa cells (Castello et al., 2012). Thus, DEK is involved in RNA-related processes, though at this point no RNA-binding domain has been identified.
Taken together, the functions of the unique DEK protein can be divided into three major domains: 1) functions relating to DNA/chromatin structure and function; 2) functions relating to RNA biology; and 3) extracellular functions. As DEK appears to be a multifunctional factor with roles on multiple cellular levels, and only limited information can be deduced from the studies of other proteins, it is rather complex to study and understand its somewhat paradoxical functions. Therefore, in this thesis the goal is to detail the role of its
DNA/chromatin folding activities in global or local chromatin organization. The outcome may lead to further elucidation of its role in tumorgenesis and may even provide a rationale for
1 Introduction 24 targeting DEK in tumor intervention.
2 Aim of this Thesis 25
2. Aim of this Thesis
The DEK protein, a unique chromatin-associated factor, displays multiple functions in cellular processes, though the underlying detailed mechanisms remaining largely elusive. Based on its multiple DNA and chromatin altering activities, DEK has been classified as a non-histone chromosomal factor. Even though the DNA binding activities of DEK are rather well investigated in vitro, the relevance of these activities in the cellular environment is unknown.
Therefore, this thesis aims to elucidate the impact of DEK’s DNA and chromatin binding and folding activities in cells. In order to do so, a series of individual point mutations within the major DNA-binding domain of DEK, the pseudo-SAP/SAP-box, will be created and subsequently tested for their remaining DNA-binding activity in vitro. After identification of
“DNA-binding-dead” mutants, the cellular impact of DEK on chromatin structure can be assessed. Finally, DEK knockout cells will be created and re-expression of DEK and DEK deletion and mutants will be achieved by lentiviral delivery.
3 Materials and Methods 26
3. Materials and Methods
All methods were performed without or with modifications according to standard procedures available in the Institute of Biochemistry and Molecular Biology, RWTH Aachen University.
3.1 Materials
3.1.1 Chemicals
Ordering Chemical Company Lot number number 1,4-Dithiothreitol (DTT) Roth 6908.2 373204255 2-mercapto ethanol, β-Mercaptoethanol Merck S5123240 849 8057300250 2-Propanol Roth 6752.4 364219192 3-methyl-1-butanol (isoamyl alcohol) Merck 818969 820886 3-Morpholinopropanesulfonic acid (MOPS) Serva 29836 11084 Acetic acid (glacial) 100% Merck 1.00063.2511 K45886263 432 Acrylamide 4K - solution (30%) Mix 29:1 Applichem 616-003-00-0 4R014006 Acrylamide 4K - solution (40%) Mix 19:1 Applichem A1640,055 7E001513 Agar bacterial grade MP Biomedicals, LLC 150178 MR29280 Agarose MP biomedicals, LLC 800668 9889J
Ammonium acetate, CH3COONH4 Merck 1116 CC585416
Ammonium chloride, NH4Cl Merck 1.01145.0500 A320145 137 Ammonium peroxydisulfate (APS) Roth 9592.2 303201798
Ammonium sulfate, (NH4)2SO4 Fluka 09980 367647/1 53697 Ampicilin sodium salt Roth K029.1 324213295 Biozym LE agarose Biozym 840004 120212C274967L Blasticidin Invitrogen Ant-bl-1 BLL-36-01A Boric acid molecular biology grade Applichem A2940,1000 0A005929 Bovine serum albumin fraction V (BSA) Serva 11930 17363 Bromphenol blue Roth T116.1 372180743
Calcium chloride dehydrate, CaCl2 Merck 1.02382.1000 TA268982 718 CASY ton OMNI Life Sciences
Chloroform ultrapure Applichem A3633,0500 1E003775 Ciprofloxacin Hydrochloride MP Biomedicals, LLC 199020 4913F Complete tablets (EDTA free, EASY pack) Roche 04 693 132 001 17325400 Coomassie brilliant blue G250 Serva 17524 07975 Cy5-Oligo dT21 MOLBIOL Custom-ordered
DEPC Applichem 1609-47-8 R8938 Dimethyl sulfoxide (DMSO) Merck 1.16743.1000 K25358243 836 Di-potassium hydrogen phosphate trihydrate Merck 1.05099.1000 A911199 625 GR (K2HPO4) Di-sodium hydrogen phosphate dihydrate Merck 1.06580.1000 K44861480 334 (Na2HPO4) dNTP Set, PCR Grade 100 mM 4x 250 µL Qiagen 201913 133219837 Dodecyl sulfate-Na-salt in pellets, NaDS Serva 151-21-3 140489 Doxycycline hydrochloride ICN Biomedical 195044 7654A
3 Materials and Methods 27
Ethanol 70% (V/V) Fischar 603-002-00-5 707 4081 Ethanol absolute for analysis Merck 1.00983.2511 K345982283 447 Ethidium bromide Roth 7870
Ethylenediaminetetraacetic acid (EDTA) Sigma E6758-500G SLBH0908V Fetal Bovine Serum (FBS) Gibco 10270-106 41G3933K Ficoll 400, for molecular biology Sigma 26873-85-8 038K60342 Formaldehyde solution min 37% GR Merck 4003.2500 K17586903 GFP-trap_A Chromo tek Gta-20 140613001A Glutathione sepharose 4B GE healthcare 17-0756-01 71-0248-CO-EF Glycine for molecular biology Applichem A1067,5000 4J017326 Glycogen for molecular biology Roche 10 901 393 001 14267332 Glycerol 85% Merck 1.04091.2500 Z267391 228 HEPES Roth 9105.4 27184596 Hydrochloric acid Roth 4625.2 400162892
Hydrogen peroxide (30%), H2O2 Roth 9681 008-003-01-6 IGEPAL CA-630 (NP-40) MP Biomedicals, LLC 198596 5917J Imidazole, ACS reagent Sigma-aldrich I2399-500G 026K00761 Isopropyl-beta-D-thiogalactopyranoside MP biomedicals, LLC 102101 8305K (IPTG) LB broth (Luria/Miller) Roth X968.1 283216388 L-glutathione (reduced form) Serva 23150 6483 Lipofectamine 2000 Invitrogen 11668-019 1649687 Lithium chloride Merck 5679.0250 A675579 Luminol Aldrich 123072-5G SHBD0162V Magnesium chloride hexayhydrate, KMF Laborchemie Handels GMBH 7791-18-6 6F005518 MgCl26H2O Magnesium acetate tetrahydrate, Merck 5819 630A87219 Mg(CH3COO)24H2O Manganese (II) chloride tetrahydrate, Merck 1.05927.1000 A317127 Mncl24H2O Premier International Foods (UK) Marvel Original Dried Skimmed Milk 92963 Ltd Methanol VWR chemicals 20847295 14I150501 N,N,N’,N’-Tetramethyl ethylene diamine Serva 35925 80935 (TEMED) Orange G Sigma MKBF6993V
Paraformaldehyde pure Serva 31628.01 101205 -Coumaric acid Sigma C9008-1G BCBN0412V Phenol:Chloroform:Isoamyl Alcohol 25:24:1 saturated with 10mm Tris, pH 8.0, 1mM Sigma P2069-100ML 103K0649 EDTA Phenylmethylsulfonylfluorid (PMSF) Roth 6367.2 02145189 Phosphate buffered saline (10x) Sigma P5493-1L SLBH9016 Ponceau S Sigma P-3504 30H3653
Potassium acetate, CH3COOK Merck 1.04820.1000 K32042020 321 Potassium chloride, KCl Sigma P-5405 34H03025
Potassium dihydrogen phosphate, KH2PO4 Merck 1.04873.1000 A377073 218 Potassium hydroxide pellets GR, KOH Merck 5033.0500 C233933 Ribomax Large Scale RNA Production Promega P1300 88386 System - T7
Sodium acetate water free, CH3COONa KMF Laborchemie Handels GMBH KMF.08-319 67776
Sodium azide, NaN3 Merck 1.06688.0100 K40831788 039
3 Materials and Methods 28
Sodium butyrate, Na(C3H7COO) Aldrich 303410-100G MKBG2813V Sodium chloride, NaCl VWR Chemicals 27810.295 13F130010 Sodium dihydrogen phosphate monohydrate, Merck 1.06346.1000 A287446 148 NaH2Po4H2O Sodium fluoride, NaF Merck 1.06449.0250 B440749 515
Sodium hydrogen carbonate, NaHCO3 Merck 6329.1000 K15322429 Sodium hydroxide, NaOH Merck 1.06498.1000 B0847698 241
Sodium orthovanadate, Na3VO4 Sigma 13721-39-6 28K0117 Tris ultrapure Applichem A1085,5000 4G014099 Triton X-100 Aldrich 23472-9 0841EL TRIzol reagent Ambion RNA by life technologies 15596018 13548001 Tween 20 Sigma P2287-500ML MKBH8325V Urea Serva 24524 18938 Vanadyl Ribonucleoside Complex NEB #S1402S 91405 Water Merck 1.15333.2500 Z0294333 320 Xylene cyanol FF Serva 38505 43535 [-32P]-UTP Hartmann Analytic
[-32P]-ATP Hartmann Analytic
3.1.2 General buffers
All buffers were prepared with Millipore water (Merck Millipore).
Transfer buffer: 500 ml 10x Tris/Glycine 50 ml Methanol 50 ml H2O to 500 ml SDS-running buffer (10x): 2 L Tris 60.56 g Glycine 288.26 g SDS (20%) 100 ml H2O to 2 L Coomassie Brilliant Blue G250 staining buffer: 1 L Coomassie Brilliant Blue G250 2 g Methanol 400 ml Acetic acid 100 ml H2O to 1L 10x Tris/Glycine: 2 L Tris 60.2 g Glycine 376 g H2O to 2 L 10x PBS: 2 L NaCl 234.09 g KCl 3.72 g Na2HPO4·2H2O 28.4 g KH2PO4 4.08 g H2O to 2 L
3 Materials and Methods 29
PBS-Tween(PBS-T): Add 0.1% Tween 20 to 1x PBS buffer
De-staining Buffer: 1 L Methanol 400 ml Glacial acetic acid 100 ml H2O to 1 L 10x Laemmli buffer: 10 ml Glycerol 5 ml ß-ME(mercaptoethanol) 4.5 ml 1M Tris, pH 6.8 0.5 ml H2O to 10 ml Add a little orange G 5x Laemmli buffer: Dilute 10x Laemmli buffer 1:2 with 20% SDS
1x Laemmli buffer: Dilute 5x Laemmli buffer 1:5 with water
Ca-ECl: 500 ml Luminol solution (250 mM in DMSO) 5 ml Ca-solution (90 mM in DMSO) 1.1 ml 100 mM Tris-HCl, pH 8.8 to 500 ml 50x TAE: 1 L Tris base 242 g Glacial acetic acid 57.1 ml 0.5 M EDTA 100 ml H2O to 1 L 1x LB medium: 1 L LB brotch (Luria/Miller) 25 g H2O To 1 L LB-agar plate: Add 15 g agar per liter of LB medium 6x DNA loading buffer: 10 ml 0.25% Bromophenol blue 25 mg 0.25% Xylene cyanol 25 mg 15% Ficoll 400 1.5 g 60 mM EDTA 1.2 ml (0.5 M EDTA)
3.1.3 Oligonucleotides
All oligonucleotides were custom-ordered from MWG.
Oligonucleotides used for mutation of aromatic amino acids in the DEK fragment 1-187: Oligo name Sequence (5’---3’) F9697A s 5’-GAA ATT GAG AGG ATA CAT GCT GCT CTA AGT AAG AAG AAA AC-3’
3 Materials and Methods 30
F9697A as 5’-GTT TTC TTC TTA CTT AGA GCA GCA TGT ATC CTC TCA ATT TC-3’ Y114A s 5’-CTA CAC AAA CTG CTT GCC AAC AGG CCA GGC ACT G-3’ Y114A as 5’-CAG TGC CTG GCC TGT TGG CAA GCA GTT TGT GTA G-3’ F130A s 5’-GAA GAA TGT GGG TCA GGC CAG TGG CTT TCC ATT TG-3’ F130A as 5’-CAA ATG GAA AGC CAC TGG CCT GAC CCA CAT TCT TC-3’ F133135A s 5’-GTG GGT CAG TTC AGT GGC GCT CCA GCT GAA AAA GGA AGT GTC C-3’ F133135A as 5’-GGA CAC TTC CTT TTT CAG CTG GAG CGC CAC TGA ACT GAC CCA C-3’ Y142A s 5’-GAA AAA GGA AGT GTC CAA GCT AAA AAG AAG GAA GAA ATG-3’ Y142A as 5’-CAT TTC TTC CTT CTT TTT AGC TTG GAC ACT TCC TTT TTC-3’ F152A s 5’-GAA GAA ATG TTG AAA AAA GCT AGA AAT GCC ATG TTA AAG-3’ F152A as 5’-CTT TAA CAT GGC ATT TCT AGC TTT TTT CAA CAT TTC TTC-3’ F182A s 5’-GTG AAG AGG ATC TTG AAT GCC TTA ATG CAT CCA AAG CC-3’ F182A as 5’-GGC TTT GGA TGC ATT AAG GCA TTC AAG ATC CTC TTC AC-3’
Oligonucleotides used for mutations of arginines in the DEK fragment 1-187: Oligo name Sequence (5’---3’) R93A s 5’-CAG AAA CTT TGT GAA ATT GAG GCG ATA CAT TTT TTT CTA AG-3’ R93A as 5’-CTT AGA AAA AAA TGT ATC GCC TCA ATT TCA CAA AGT TTC TG-3’ R107A s 5’-GAA GAA AAC CGA TGA ACT TGC AAA TCT ACA CAA ACT G-3’ R107A as 5’-CAG TTT GTG TAG ATT TGC AAG TTC ATC GGT TTT CTT C-3’ R116A s 5’-CAC AAA CTG CTT TAC AAC GCG CCA GGC ACT GTG TCC-3’ R116A as 5’-GGA CAC AGT GCC TGG CGC GTT GTA AAG CAG TTT GTG-3’ R153A s 5’-GTT GAA AAA ATT TGC AAA TGC CAT GTT AAA GAG CAT C-3’ R153A as 5’-GAT GCT CTT TAA CAT GGC ATT TGC AAA TTT TTT CAA C-3’ R168A s 5’-GAG GTT CTT GAT TTG GAG GCA TCA GGT GTA AAT AGT GAA C-3’ R168A as 5’-GTT CAC TAT TTA CAC CTG ATG CCT CCA AAT CAA GAA CCT C-3’ R178A s 5’-GTG TAA ATA GTG AAC TAG TGA AGG CGA TCT TGA ATT TCT TAA TGC-3’ R178A as 5’-GCA TTA AGA AAT TCA AGA TCG CCT TCA CTA GTT CAC TAT TTA CAC-3’
Oligonucleotides used for the generation of DEK truncations in pGEX4T1: Oligo name Sequence (5’---3’) EcoR_DEK1-87-sp 5’-CCA TGG AAT TCA TGT CCG CCT C-3’ DEK1-87_Xho_rp 5’-GAG CTC GAG TTA TTT CTG CCC C-3’ EcoR-DEK187-270 5’-CAT GGA ATT CAA GCC TTC TGG CAA AC-3’ DEK187-270_Xho 5’-CTT CCT CGA GTT AAG TAG CTT TCT GTT TAG-3’
Oligonucleotides used for error-prone PCR (template: pGEX4T1-DEK1-187 or pGEX4T1-DEK85-187): Oligo name Sequence (5’---3’) EP_pGEX_sp 5’-GAT CTG GTT CCG CGT GGA TCC CCG GAA TTC-3’ EP_pGEX_rp 5’-CAG TCA GTC ACG ATG CGG CCG CTC GAG-3’
Oligonucleotides used for the generation of SAF-A truncations in pGEX4T1: Oligo name Sequence (5’---3’) SAF-A FLsp 5’-GCC AAG AAT TCC TCA CCA TGA GTT CCT CGC CTG TTA ATG-3’ SAF-A FLRp 5’-CCG TTG CGG CCG CTT ATT GCT CGA GGT CGA CAT AAT ATC CTT GGT G-3’ SAF-A deltaN sp 5’-GCC AAG AAT TCC TCA CCA TGA GTG ACG ACG AGG AG-3’ SAF-A trunc sp 5’-GCC AAG AAT TCA AAA AGC GCA AGG TGA AGC TTC AAA TTC TG-3’ SAF-A trunc rp 5’-CCG TTG CGG CCG CTT ACG TGG CGA CCG GTG GAT CCG-3’
Oligonucleotides used for the generation of DEK truncations in pTRIPZ:
3 Materials and Methods 31
Oligo name Sequence (5’---3’) NLSfor_cut_BsrEc 5’-GAG CTG TAC AAG GCC ACC ATG GTG CCC CCT AAG AAA AAG CGC AAG GTG ACC GGT GAA TTC AAC-3’ NLSrev_cut_BsrEc 5’-GTT GAA TTC ACC GGT CAC CTT GCG CTT TTT CTT AGG GGG CAC CAT GGT GGC CTT GTA CAG CTC-3’ EP_pGEX_sp 5’-GAT CTG GTT CCG CGT GGA TCC CCG GAA TTC-3’ DEK87-HindIIIrp 5’-GCA CAT ACG TAA AGC TTC TGC CCC TTT CCT TGT GCA ATT GTA AAT GGC-3’ DEK87-hindIIIsp 5’-GAT CCC CGG AAT TCG GGC AGA AGC TTT GTG C-3’ DEKx-187rev_Sna 5’-CTT TTA CGT AGC GGC CGC TCG AGA TTA CTT TGG ATG C-3’ DEK186-xfor_BEA CAA GGA TCC GAA TTC AGG CGC GCC CCA AAG CCT TCT GGC AAA CCA TTG CCG-3’ DEKx-375rev_XhS 5’-CTT TAC GTA CAC GAT GCG GCC GCT CGA GAT TAA G-3’ DEKx-260rev_sto 5’-CTT TAC GTA CTC GAG TTA GGC TGT CTT TTT TGG TGG CTC CTC TTC AC-3’ DEKx-87rev_AscI 5’-CTT GGC GCG CCG TTT CTG CCC CTT TCC TTG TGC AAT TGT AAA TGG C-3’
Oligonucleotides used to mutate single amino acids mutations in pGEX4T1-DEK85-187#11 back to wild type condition: Oligo name Sequence (5’---3’) r95h-Sp DEK 5’-CTT TGT GAA ATT GAG AGG ATA CAT TTT TTT CTA AGT AAG-3’ r95h-rp DEK 5’-CTT ACT TAG AAA AAA ATG TAT CCT CTC AAT TTC ACA AAG-3’ m101k-sp DEK 5’-TTC TAA GTA AGA AGA AAA CCG ATG AAC TTA GAA ATC TAC-3’ m101k-rp DEK 5’-GTA GAT TTC TAA GTT CAT CGG TTT TCT TCT TAC TTA GAA-3’ i111k-sp DEK 5’-CTT AGA AAT CTA CAC AAA CTG CTT TAC AAC AGG CC-3’ i111k-rp DEK 5’-GGC CTG TTG TAA AGC AGT TTG TGT AGA TTT CTA AG-3’ e137k-sp DEK 5’-GGC TTT CCA TTT GAA AAA GGA AGA GTC CAA TAT AAA AAG-3’ e137k-rp DEK 5’-CTT TTT ATA TTG GAC TCT TCC TTT TTC AAA TGG AAA GCC-3’ r139s-sp DEK 5’-GGC TTT CCA TTT GAA GAA GGA AGT GTC CAA TAT AAA AAG-3’ r139s-rp DEK 5’-CTT TTT ATA TTG GAC ACT TCC TTC TTC AAA TGG AAA GCC-3’ e150k-sp DEK 5’-GAA GGA AGA AAT GTT GAA AAA ATT TAG AAA TGC CAT G-3’ e150k-rp DEK 5’-CAT GGC ATT TCT AAA TTT TTT CAA CAT TTC TTC CTT C-3’ s166l-sp DEK 5’-GTG AGG TTC TTG ATT TGG AGA GAT CAG GTG TAA ATA G-3’ s166l-rp DEK 5’-CTA TTT ACA CCT GAT CTC TCC AAA TCA AGA ACC TCA C-3’ d181n-sp DEK 5’-CTA GTG AAG AGG ATC TTG AAT TTC TTA ATG CAT CC-3’ d181n-rp DEK 5’-GGA TGC ATT AAG AAA TTC AAG ATC CTC TTC ACT AG-3’
3.1.4 Plasmids
Aromatic amino acid mutants in pGEX4T1(for bacterial expression): Name Template Backbone Primers Restriction sites pGEX4T1- pGEX4T1-DEK1-187 pGEX4T1 F96/97A s; F96/97A as EcoR I/XhoI DEK1-187F96/97A pGEX4T1- pGEX4T1-DEK1-187 pGEX4T1 Y114A s; Y114A as EcoR I/ XhoI DEK1-187 Y114A pGEX4T1- pGEX4T1-DEK1-187 pGEX4T1 F130A s; F130A as EcoR I/XhoI DEK1-187 F130A pGEX4T1- pGEX4T1-DEK1-187 pGEX4T1 F133/135A s;F133/135A as EcoR I/XhoI DEK1-187 F133/135A pGEX4T1- pGEX4T1-DEK1-187 pGEX4T1 Y142A s; Y142A as EcoR I/XhoI DEK1-187 Y142A pGEX4T1- pGEX4T1-DEK1-187 pGEX4T1 F152A s; F152A as EcoR I/XhoI DEK1-187 F152A
3 Materials and Methods 32 pGEX4T1- pGEX4T1-DEK1-187 pGEX4T1 F182A s; F182A as EcoR I/XhoI DEK1-187 F182A pGEX4T1-DEK1-187 pGEX4T1-DEK1-187 pGEX4T1 F182A s; F182A as EcoR I/XhoI F96/97/182A F96/97A
Arginine mutants in pGEX4T1(for bacterial expression): Name Template Backbone Primers Restriction sites pGEX4T1-DEK1-187 R93A pGEX4T1-DEK1-187 pGEX4T1 R93A s;R93A as EcoR I/XhoI pGEX4T1-DEK1-187 R107A pGEX4T1-DEK1-187 pGEX4T1 R107A s; R107A as EcoR I/XhoI pGEX4T1-DEK1-187 R116A pGEX4T1-DEK1-187 pGEX4T1 R116A s; R116A as EcoR I/XhoI pGEX4T1-DEK1-187 R153A pGEX4T1-DEK1-187 pGEX4T1 R153A s; R153A as EcoR I/XhoI pGEX4T1-DEK1-187 R168A pGEX4T1-DEK1-187 pGEX4T1 R168A s; R168A as EcoR I/XhoI pGEX4T1-DEK1-187 R178A pGEX4T1-DEK1-187 pGEX4T1 R178A s; R178A as EcoR I/XhoI
DEK truncations in pGEX4T1(for bacterial expression): Name Backbone Source pGEX4T1-DEKwt pGEX4T1 F. Kappes pGEX4T1-DEK1-187 pGEX4T1 F. Kappes pGEX4T1-DEK187-375 pGEX4T1 F. Kappes pGEX4T1-DEK251-375 pGEX4T1 F. Kappes Name Template Backbone Primers Restriction sites pGEX4T1- pGEX4T1- pGEX4T1 EcoR_DEK1-87-sp; EcoR I/XhoI DEK1-87 DEK1-187 DEK1-87_Xho_rp pGEX4T1- pGEX4T1- pGEX4T1 DEK85_for_BamE; EcoR I/XhoI DEK85-187 DEK1-187 DEKx-187rev_Sna pGEX4T1- pGEX4T1- pGEX4T1 EcoR-DEK187-270; EcoR I/XhoI DEK187-270 DEK187-375 DEK187-270_Xho pGEX4T1-DEK1-187(or pGEX4T1-DEK pGEX4T1 EP_pGEX_sp; EcoR I/XhoI 85-187) random mutations 1-187(or 85-187) EP_pGEX_rp
DEK truncations in pTRIPZ (for re-expression in cells via lentiviral delivery): Name Backbone Source pTRIPZ-spezial-eGFP-DEKwt pTRIPZ-spezial-eGFP F. Kappes Name Template Backbone Primers Restriction sites pTRIPZ-spezial- - pTRIPZ-spezial-eGFP NLSfor_cut_BsrEc; BsrG1/EcoRI NLS-eGFP NLSrev_cut_BsrE pTRIPZ-spezial- pTRIPZ-spezial- pTRIPZ-spezial-NLS-eGFP EP_pGEX_sp; EcoRI/SnaBI NLS-eGFP-DEK eGFP-DEKwt DEKx-187rev_Sna 1-187 pTRIPZ-spezial- pTRIPZ-spezial- pTRIPZ-spezial-NLS-eGFP EP_pGEX_sp; EcoRI/HindIII/SnaBI NLS-eGFP-DEK eGFP-DEKwt; DEK87-HindIIIrp, 187 mutations pGEX4T1-DEK DEK87-hindIIIsp; 85-187 #11 DEKx-187rev_Sna pTRIPZ-spezial- pTRIPZ-spezial- pTRIPZ-spezial-NLS-eGFP DEK186-xfor_BEA; EcoRI/SnaBI NLS-eGFP-DEK eGFP-DEKwt DEKx-375rev_XhS 186-375 pTRIPZ-spezial- pTRIPZ-spezial- pTRIPZ-spezial-NLS-eGFP DEK186-xfor_BEA; EcoRI/SnaBI NLS-eGFP-DEK eGFP-DEKwt DEKx-260rev_sto 186-260 pTRIPZ-spezial- pTRIPZ-spezial- pTRIPZ-spezial-NLS-eGFP DEK186-xfor_BEA; EcoRI/SnaBI NLS-eGFP-DEK eGFP-DEKwt DEKx-87rev_AscI delta SAF
3 Materials and Methods 33 pTRIPZ-spezial- pTRIPZ-spezial- pTRIPZ-spezial-NLS-eGFP EP_pGEX_sp; EcoRI/SnaBI NLS-eGFP-DEK eGFP-DEKwt DEKx-260rev_sto delta C pTRIPZ-spezial- pTRIPZ-spezial- pTRIPZ-spezial-NLS-eGFP DEK186-xfor_BEA; EcoRI/SnaBI NLS-eGFP-DEK eGFP-DEK delta C DEKx-87rev_AscI delta delta
SAF-A in pGEX4T1 (cloned by Nahleen Schilling): Name Template Backbone Primers Restriction sites pGEX4T1- SAF-A FL pEGFP-N1-SAF-A FL pGEX4T1 SAF-A FLsp; SAF-A FLRp EcoRI/NotI pGEX4T1- SAF-N280 pEGFP-N1-SAF-A N280 pGEX4T1 SAF-A trunc sp; SAF-A trunc rp EcoRI/NotI pGEX4T1- SAF-I280 pEGFP-N1-SAF-A I280 pGEX4T1 SAF-A trunc sp; SAF-A trunc rp EcoRI/NotI pGEX4T1- SAF-C280 pEGFP-N1-SAF-A C280 pGEX4T1 SAF-A trunc sp; SAF-A trunc rp EcoRI/NotI
Mutate single mutations of #11 mutant back to wild type (for bacterial expression): Name Template Backbone Primers Restriction sites pGEX4T1-DEK85-187 pGEX4T1-DEK85-187#11 pGEX4T1 r95h-Sp DEK; r95h-rp DEK EcoR I/XhoI #11 R95H pGEX4T1-DEK85-187 pGEX4T1-DEK85-187#11 pGEX4T1 m101k-sp DEK; m101k-rp EcoR I/XhoI #11 M101K DEK pGEX4T1-DEK85-187 pGEX4T1-DEK85-187#11 pGEX4T1 i111k-sp DEK; i111k-rp DEK EcoR I/XhoI #11 I111K pGEX4T1-DEK85-187 pGEX4T1-DEK85-187#11 pGEX4T1 e137k-sp DEK; e137k-rp DEK EcoR I/XhoI #11 E137K pGEX4T1-DEK85-187 pGEX4T1-DEK85-187#11 pGEX4T1 r139s-sp DEK; r139s-rp DEK EcoR I/XhoI #11 R139S pGEX4T1-DEK85-187 pGEX4T1-DEK85-187#11 pGEX4T1 e150k-sp DEK; e150k-rp DEK EcoR I/XhoI #11 E150K pGEX4T1-DEK85-187 pGEX4T1-DEK85-187#11 pGEX4T1 s166l-sp DEK; s166l-rp DEK EcoR I/XhoI #11 S166L pGEX4T1-DEK85-187 pGEX4T1-DEK85-187#11 pGEX4T1 d181n-sp DEK; d181n-rp DEK EcoR I/XhoI #11 D181N
Other plasmids: Name Backbone Source Importin pGST From Müller-Newen lab Importin pGST From Müller-Newen lab Importin pGST From Müller-Newen lab ASH2 FL pGEX4T3 From B. Lüscher lab ASH2 1-121 pGEX4T3 From B. Lüscher lab ASH2 1-279 pGEX4T3 From B. Lüscher lab ASH2 1-279 pGEX4T3 From B. Lüscher lab ASH2 1-394 pGEX4T3 From B. Lüscher lab ASH2 393-628 pGEX4T3 From B. Lüscher lab ASH2 542-628 pGEX4T3 From B. Lüscher lab HMGB1 pGEX2T1 From M. Bianchi SAF-A FL pEGFP-N1 From F. O. Fackelmayer CK2 pGEX-3X From B. Lüscher lab CK2’ pGEX-3X From B. Lüscher lab PRMT1 pGEX From B. Lüscher lab PRMT3 pGEX From B. Lüscher lab ARTD10 1-907 pGEX4T1 From B. Lüscher lab
3 Materials and Methods 34
ARTD10 588-1021 pGEX4T1 From B. Lüscher lab ARTD10 700-1021 pGEX4T1 From B. Lüscher lab ARTD10 818-1025 G888W pGEX4T1 From B. Lüscher lab Ran pGEX4T1 From B. Lüscher lab Grb2 pGEX-3X From B. Lüscher lab Src-SH3 pGEX From B. Lüscher lab PP1-PP12 pCDNA3.1 From K. Bendak ALY/REF FL pGEX-6p From M. Ohno
3.1.5 Vectors
Vectors Features Resistance pGEX4T-1 GST-tag Ampicillin pTRIPZ spezial GFP-tag Ampicillin
3.1.6 Enzymes
3.1.6.1 Restriction Endonucleases
Name Manufacture Catalog number AgeI-HF NEB R3552S ApaI NEB R0114S BamHI-HF NEB R3136S DpnI NEB R0176S EcoRI-HF NEB R3101S HindIII-HF NEB R3104S NheI-HF NEB R3131S NotI-HF NEB R3189S SnaBI NEB R0130S XbaI NEB R0145S XhoI NEB R0146S
3.1.6.2 Polymerases
Name Manufacture Catalog number DNA Polymerase (Klenow) NEB M0210S M-MLV reverse transcriptase Promega M170A pfuUltra DNA Polymerase AD Agilent 600385-51 Q5 HiFi DNA Polymerase NEB M0491S Taq DNA Polymerase NEB M0267L
3.1.6.3 Other enzymes
Name Manufacture Catalog number Antarctic phosphatase NEB B0289S DNase I, RNase-free Roche 10776 785 001 Micrococal Nuclease Thermo Scientific EN0181 Proteinase K Thermo Scientific #EO0491 RNase Inhibitor, Murine NEB M0314S RNase, DNase-free Roche 11 119 915 001
3 Materials and Methods 35
T4 DNA Ligase (1 U/µL ) NEB M0202S T4 DNA Ligase (5 U/µL) Thermo Scientific #EL0017 T4 polynucleotide kinase NEB M0201S
3.1.7 Antibodies
3.1.7.1 Primary antibodies
Dilutions Catalog Antibodies species Manufacture Immunoblot IF number -DEK polyclonal K877 rabbit 1:10,000 1:1000 Kappes, et al.2004 - -DEK polyclonal K878 rabbit 1:10,000 1:1000 Kappes, et al.2004 - -DEK monoclonal mouse 1:1000 1:200 BD Biosciences #610948 -GST monoclonal goat 1:1000 - Rockland #100-101-200 -GFP monoclonal goat 1:1000 - Rockland 600-101-215 - GFP monoclonal mouse - 1:15,000 Rockland 600-301-215 --actin monoclonal mouse 1:100 in PBS-T -
3.1.7.2 Secondary antibodies
For Immunoblotting: Antibodies Species Dilutions Manufacture Catalog number anti-rabbit IgG/HRP goat 1:10,000 Dianova #111-035-003 anti-goat IgG/HRP donkey 1:10,000 Dianova #705-035-003 anti-mouse IgG/HRP goat 1:10,000 Dianova #115-035-003
For immunofluorescence: Antibodies Species Dilutions Manufacture Catalog number anti-rabbit IgG(H+L), donkey 1:1,000 Invitrogen A-21206 Alexa Fluor○R 488 anti-mouse IgG(H+L), donkey 1:1,000 Invitrogen A-31570 Alexa Fluor○R 555
All antibodies were diluted in PBS-T containing 5% milk powder for immunoblotting. When multiple primary antibodies were used, 0.05% sodium azide was added for 30 min prior to the next antibody to kill the HRP signal.
3.1.8 E. coli strains
Chemically competent E. coli DH5, BL21 (DE3) and Top 10 competent cells were produced using standard protocols of the Institute of Biochemistry and Molecular Biology, RWTH
Aachen University (see chapter 3.2.1.6).
3 Materials and Methods 36
3.2 Methods
3.2.1 General DNA-related methods
3.2.1.1 Polymerase Chain Reaction (PCR)
PCR (Polymerase Chain Reaction) is used for amplifying specific DNA sequences based on the ability of DNA polymerase to synthesize a complementary sequence to the template strand. This method relies on thermal cycling, consisting of cycles of repeated heating and cooling of the reaction for DNA melting and enzymatic replication of the DNA. To increase the fidelity of
PCR, the high fidelity DNA polymerase is used for long amplification. In this thesis, Q5 DNA polymerase (NEB) was used in 50 µl PCR reactions: 5 ng DNA template, 0.5 µl 5x Q5 reaction buffer, 0.5 µl forward primer (100 pmol/µl), 0.5 µl reverse primer (100 pmol/µl), 3.5 µl dNTPs
(each 2.5 mM) and 0.5 µl Q5 polymerase (2 U/µl). The PCR program was performed as follows:
98°C 20s as initial denaturing step; 98°C 10 s, (Tm-5)°C 20 s, 72°C 1 min/kb, 30 cycles for elongation; and 5 min at 72°C for final extension. Tm: Melting temperature.
3.2.1.2 Gel electrophoresis
The PCR products were adjusted to 1x DNA loading buffer and loaded onto agarose gels of proper percentage (0.1 µg/ml Ethidium bromide) based on expected DNA size. The run was carried out in 1x TAE buffer at 110 V for 30 min. For size standards the DNA ladder 1 kb plus ladder (Invitrogen, Paisley, UK) was used. DNA was visualized under UV light and photographed by an Intas imaging device.
3.2.1.3 Restriction digestion
Restriction enzymes also referred to restriction endonucleases recognize short, specific (often palindromic) DNA sequences. They cleave double-stranded DNA (dsDNA) at specific sites within or adjacent to their recognition sequences. Restriction digestion is commonly used to insert specific DNA fragments into plasmid vectors, which can be efficiently “cloned” by transformation into bacterial cells. 0.5-1.5 µg DNA was digested with 5-fold excess of enzyme in a total volume of 20-50 µl. After 1 hour digestion at 37°C, the digested DNA was analyzed on
3 Materials and Methods 37
1% agarose gels.
The following formula was used to calculate the needed enzyme volumes (µl ): bp of reference DNA x sites in vector x DNA in µg x 5 times x bp of vector x sites in reference- DNA x enzyme concentration in U/µl
3.2.1.4 Gel extraction
After agarose gel electrophoresis, the DNA was extracted and purified using the QIAquick Gel
Extraction Kit (Qiagen). The DNA was eluted in chromatography water (Merck) and the concentration was measured using the Nanodrop.
3.2.1.5 Ligation
T4 ligase catalyzes the conformation of covalent bonds between the 5’-phosphate of one nucleotide and the 3’-hydroxyl of another in either a cohesive-ended or blunt-ended fashion.
This is used to insert DNA fragment/s into vectors.
For cohesive (sticky) ends, the ligation reaction (20 µl) was set up as follows: T4 ligase buffer (10x) 2 µl digested backbone vector 30 ng digested insert DNA 3-10 fold* T4 ligase (1 U/µl) 1 µl
H2O to 20 µl
*The following formula was used to calculate the amount of insert DNA (ng): bp of insert DNA x 30 ng vector x 3 (to 10) times x bp of vector
The ligation mixture was incubated at RT for 1 hour. For blunt ends, polyethylene glycol (PEG)
4000 (50%, w/v) was added to increase the ligation efficiency. The final concentration of PEG
4000 was 5%. Higher concentrations of T4 ligase (5 U/µl) or overnight incubation was used for difficult-to-ligate samples. After 1 hour incubation, the ligation mixture was directly transformed into chemically competent E. coli cells.
3.2.1.6 Preparation of E. coli competent cells
Glycerol stocks of the E. coli bacteria were streaked on LB plates (without any antibiotic) and
3 Materials and Methods 38 incubated at 37°C overnight. The next day, a single colony was picked and incubated in 2 ml
LB medium at 37°C for 8 hours with 225 rpm shaking. The bacterial culture was mixed with 50 ml fresh LB and cultured at 37°C overnight. The next day, 5 ml of the pre-cultured bacteria was added into 250 ml LB medium and cultured to OD600=0.6. After 10 min incubation on ice, the bacteria were collect by centrifugation (4°C, 2000 x g, 10 min). Next the pellet was gently resuspended with 80 ml TB buffer (15 mM CaCl2, 250 mM KCl, 10 mM Pipes, 55 mM MnCl2, pH 6.7), and incubated on ice (10 min). After one additional centrifugation step, the bacterial pellet was resuspended in 20 ml TB buffer (pH 6.7) and incubated on ice (10 min). DMSO was added to a final concentration of 7% and mixed with the bacteria, and aliquots of 200 µl /each were snap-frozen in liquid nitrogen and stored at -80°C until use.
3.2.1.7 Transformation
Transformation is the process by which foreign DNA is introduced into a cell. Chemical or electrical based transformation is commonly applied. Here we used standard heat-shock transformation of chemically competent bacteria. 1 to 5 ng DNA was gently mixed with 50 μl of thawed DH5 competent cells in a tube and incubated on ice (20-30 min). Next, bacteria were heat shocked at 42°C for 90 seconds (45 seconds for BL21) without shaking, and then placed directly on ice for 2 min. 50 µl of the transformation reaction were spread onto LB-agar plates with the appropriate antibiotics (Ampicillin 100 ng/ml) and plates were incubated at 37°C overnight. When BL21 competent cells were used in transformation, additional culture (37°C,
40 min with shaking) with 450 μl fresh LB medium (without antibiotic) was needed after 2 min incubation on ice.
3.2.1.8 Colony PCR
To quickly identify proper transformants, individual colonies from LB-Agar plates were picked and placed on a master plate. After overnight incubation at 37°C, the colony PCR was performed to determine the presence or absence of insert DNA in plasmid constructs. For multiple colonies, a master mix was prepared and 15 µl were used per PCR reaction. After the
PCR, the amplified DNA was analyzed with 1% agarose gel (1x TAE).
3 Materials and Methods 39 1x colony PCR reaction mix (15 µl): 10x Thermo polymerase buffer 2.5 µl forward primer (10 pmol/µl) 1.5 µl reverse primer (10 pmol/µl) 1.5 µl dNTP mix (2.5 mM each) 2 µl Taq DNA polymerase (NEB) 0.25 µl
H2O to 15 µl PCR program: 1 94°C 60 s initial denaturation 94°C 60 s 2 50°C 60 s 30 cycles 72°C 60 s/kb 3 72°C 5 min final extension 4 4°C pause hold
3.2.1.9 DNA preparation
Bacteria were cultured in 5 ml LB (plus proper antibiotic) overnight and harvested by centrifugation (17,000 x g, RT, 10 min) the next day. The plasmid DNA was purified using
QIAprep Spin Miniprep Kit (Qiagen) following the manufacturer’s instructions. The QIAprep miniprep procedure is based on alkaline lysis of bacterial cells followed by adsorption of DNA onto silica in the presence of high salt. DNA concentration was determined by Nanodrop.
3.2.1.10 DNA extraction (phenol/chloroform) and precipitation (ethanol)
1x volume of phenol:chloroform:isoamyl alcohol (25:24:1) solution was mixed with the DNA sample and centrifuged for 5 min (17,000 x g, RT). The aqueous phase (DNA) was transferred into a new tube. 1/10 volume of sodium acetate and 2.5 volumes of 100% ethanol were added to precipitate the DNA (-20°C, 30 min). The DNA was collected by centrifugation (13,800 x g,
4°C, 15 min) and the pellet was washed with 70% pre-cooled ethanol (80% ethanol was used for DNA smaller than 500 bp). The pellet was either air-dried at RT or at 37°C, and then re-dissolved in chromatography water (Merck).
3.2.2 Protein-related methods
3.2.2.1 Expression of GST-tagged proteins
1-10 ng of pGEX4T1 plasmids carrying the gene of interest (GOI) were transformed into 50 µl
3 Materials and Methods 40
BL21 (DE3) cells and plated onto LB plates/Amp. After overnight incubation (37°C) a single colony was picked and cultured in 5 ml LB/Amp medium overnight with shaking (37°C, 225 rpm). On the third day, the pre-cultured bacteria were diluted 1:100 to 500 ml LB medium in presence of ampicillin (100 ng/ml), and cultured at 37°C with 225 rpm shaking. IPTG (final concentration=0.5 mM) was added to induce protein expression when OD600 reached 0.6. After 1.5 hours induction, the bacteria were collected by centrifugation (6198 x g, 4°C, 15 min).
3.2.2.2 Purification of GST-tagged proteins
After expression of the respective GST-tagged proteins, bacteria were collected by centrifugation and the pellet was resuspended in 10 ml resuspension buffer supplemented with complete protease inhibitor cocktail (Complete) and DTT (1 mM), and then snap frozen and stored at -80°C. Next day, the bacterial solution was thawed at 37°C in a water bath for a few minutes (~15 min) and sonicated (on ice) 5x 10 seconds or more until the cells were completely disrupted (lysis is complete when cell suspension turns from cloudy to translucent). The soluble expressed proteins (supernatant) were separated from insoluble material (pellet) by centrifugation (20,000 x g, 4°C, 15 min). During centrifugation, the Glutathione-sepharose 4B beads were washed three times with resuspension buffer in a 15 ml tube and 500 µl of 1:1 bead slurry was added to the cleared supernatant and rotated for 2 hours at 4°C. The beads were collected (900 x g, 4°C, 10 min) and washed twice with 10 ml Wash buffer I, twice with 10 ml
Wash buffer II, and one time with Wash buffer III. Elution of the GST-tagged proteins was carried out with 200 µl fresh Elution buffer (4°C, 1 hour, with rotation). The first elution was collected (17,000 x g, 4°C, 5 min) and two subsequent elution steps were performed with either half hour or overnight rotation. Eluates were aliquoted (20 µl /each), snap-frozen and stored at -80°C. Resuspension buffer: Wash buffer I: 20 mM Tris, pH 8.0 20 mM Tris, pH 8.0 1 M NaCl 500 mM NaCl 0.5 mM EDTA 0.5 mM EDTA 1 mM DTT (add before use) 0.1% NP40 1 mM DTT (add before use)
3 Materials and Methods 41 Wash buffer II: Wash buffer III: 20 mM Tris, pH 8.0 20 mM Tris, pH 8.0 300 mM NaCl 200 mM NaCl 0.5 mM EDTA 0.5 mM EDTA 0.1% NP40 1 mM DTT (add before use) 1 mM DTT (add before use) Elution buffer: (fresh) 200 mM Tris, pH 8.0 200 mM NaCl 40 mM reduced Glutathione 10% Glycerol
3.2.2.3 SDS-polyacrylamide gel electrophoresis (SDS-PAGE)
SDS-polyacrylamide gel electrophoresis (SDS-PAGE) is the most commonly used technique to separate proteins for the purposes of analysis and purification.
First, polyacrylamide gels were prepared according to the table below. 5x Laemmli buffer was added to protein samples and adjusted to 1x. The samples were boiled for 5 min (95C) and loaded on the gels together with a protein size marker (PageRuler Prestained Protein Ladder).
Separation was carried out by applying 20 mA/per gel until samples reached the resolving gel and then 40 mA/per gel until the dye front reached the bottom of the gel. resolving gel stacking gel 10% 12.5% 4 ml H2O 5.9 ml 4.9 ml 635 μl Acrylamide 4.8 ml 6 ml - 1.5 M Tris, pH 8.8 3.8 ml 3.8 ml 2 M Tris, pH 6.8 - - 313 μl 20% SDS 75 μl 75 μl 25 μl TEMED 15 μl 15 μl 5 μl 20% APS 75 μl 75 μl 40 μl
3.2.2.4 Coomassie staining
Coomassie Brilliant blue dye is commonly used to stain proteins on SDS-PAGE gels. This treatment allows visualizing the protein bands. After SDS-PAGE, the gel was immersed in coomassie staining buffer with gentle shaking (RT). After 15 min, the gel was washed with water and destained with destain buffer until the protein bands were clearly visible.
3.2.2.5 Immunoblot
After SDS-PAGE, the gel was carefully removed from the plate and immersed in transfer
3 Materials and Methods 42 buffer. The PVDF membrane was activated (with methanol for a few seconds) and also immersed in transfer buffer. Next, the transfer sandwich was prepared: paper/gel/membrane/paper, which is wetted in transfer buffer and placed directly between positive and negative electrodes (anode and cathode, respectively) in semi-dry transfer. After transferring the proteins to the membrane (1.5 hours, 0.8 mA/cm2), the PVDF membrane was removed and washed with water, and stained by Ponceau S (0.2% Ponceau Red in 5% acetic acid) for 5 minutes to monitor the quality of transfer. Before blocking, the Ponceau S was washed away with PBS-T buffer and then the blot was blocked with 5% milk (dissolved in
TBST buffer) for 1 hour at RT with gentle shaking. Next, the blot was incubated with the diluted primary antibody in blocking buffer (5% milk in PBS-T, 4C, overnight) after 3 times washing with PBS-T (5 min each time). The next day, the membrane was washed with PBS-T
(3 times, 5 min each, with gently shaking) and incubated with diluted secondary antibody
1:10000 in blocking buffer (5% milk in PBS-T) for 1 hour with agitation (RT). After three wash steps with PBS-T, the blots were developed in chemiluminescent substrate (Ca-ECL) and micrographs were taken by using a LAS-3000 Imaging System from Fuji.
3.2.2.6 Immunofluorescence (IF)
Immunofluorescence (IF) is a technique commonly used to access the sub-cellular location of proteins. The cells were seeded on cover slips treated with 0.01% poly-L-Lysine for 30 min in
12-well plates and cultured overnight. The next day, the cells were rinsed three times with 1x
PBS and then fixed with 4% paraformaldehyde (PFA)/1% sucrose/PBS for 15 min at RT. After washing (3 times) with 1x PBS, the cells were incubated with 50 mM ammonium chloride for
5 min and permeablized with 0.6% Triton X-100/PBS for 5 min. The cells were blocked in
0.1% Triton X-100/4% BSA/PBS for 1 hour after three washes with 1x PBS. The blocking buffer was removed and the primary antibody dilution in 0.1% Tween 20/1%BSA/PBS was added and incubated with the cells in the dark in a humidified chamber. One hour later, the cells were rinsed (3 times with 1x PBS) and incubated with the secondary antibody dilution
(1:1,000) in 0.1% Tween 20/1%BSA/PBS and Hoechst (1:10,000, 1 mg/ml in stock) for 1 hour at RT. After three wash steps with 1x PBS, cover slips were rinsed in water and were mounted
3 Materials and Methods 43 on a slide with a drop of mounting medium and stored overnight in the dark at 4C. The following day, nail polish was used to seal the coverslips to the slides. Images were taken with a Zeiss LSM 710 confocal microscope and edited by ZEN 2012 software (Zeiss).
3.2.3 Cell culture
3.2.3.1 Cells
Hela S3 is the third clone isolated and propagated from the parent heterogeneous HeLa cell culture which was derived from cervical cancer cells. It is a useful cell line in scientific research because it is immortal and easy to culture. Importantly, DEK is overexpressed in
HeLa S3 cells and a substantial amount of work addressing DEK biology has been performed using this cell line. Therefore, in this thesis, HeLa S3 cells were used to create DEK knockout cells.
3.2.3.2 Medium and buffers
Medium/buffers Company Ordering number Lot number Dulbecco's Modified Eagle Medium 1669883 (DMEM) + GlutaMAX (+ 4.5 g/L Gibco 31966-021 following D-Glucose; + pyruvate) Fetal Bovine Serum (FBS) Gibco 10270-106 41G3933K Penicillin-Streptomycin Sigma P0781-100ML 054M4786V 0.05% Trypsin-EDTA (1x) Gibco 25300-054 1667389 Puromycin InvivoGen ant-pr-1 QLL-35-33A Doxycycline hydrochloride ICN Biomedical 195044 7654A
3.2.3.3 Cell culture
Frozen cells were quickly thawed at 37°C in a water bath and transferred drop wise into pre-warmed complete medium (DMEM, 10% FBS and 1% Pen/Strep). After a centrifugation step (5 min, 100 x g), cells were resuspended in complete medium and were incubated in culture dishes at 37°C in an incubator with humidified atmosphere and 5% CO2 overnight. The next day, the medium was changed with fresh complete medium. Before the cells reached confluence, they were passaged. First, old medium was aspirated and cells were washed once with 1x PBS. The cells were detached from the surface of culture plate by adding trypsin for a few minutes. When ≥ 90% of cells detached, the cells were mixed with fresh complete DMEM medium and dispersed by pipetting up and down. 1/6 of the cell suspension was seeded into a
3 Materials and Methods 44 new culture plate containing proper complete DMEM and incubated in an incubator as above.
To freeze the cells as seed stock, the cells were collected by centrifugation (200 x g, 5 min) and resuspended in freeze medium (90% FBS, 10% DMSO). Aliquots were frozen in a freezing apparatus decreasing the temperature approximately 1°C per minute and stored at -150°C. All solutions and equipment in contact with the cells had to be sterile.
3.2.4 RNA isolation and cDNA synthesis
The GST-DEK and GST proteins were expressed in bacteria (chapter 3.2.2.1), and then the total
RNA of bacteria was extracted with TRIzol reagent. 1 µl total RNA was mixed with 0.5 µg random primers in a total volume of 15 µl nuclease-free water, and incubated at 70°C for 5 min to melt the secondary structures within the template. After 5 min of incubation on ice (which prevents secondary structure from reforming), 5 µl 5x M-MLV reaction buffer, 5 µl dNTP mix
(2.5 mM each) and 200 units (1 µl) M-MLV Reverse Transcriptase (Promega) were added and incubated for 1 hour (37°C) in a thermo cycler. The resulting cDNA was stored at -20°C.
3.2.5 Real-time PCR (q-PCR)
Real-time PCR, also called quantitative-PCR, is used for the determination of copy numbers of
PCR templates such as DNA or cDNA in a PCR reaction using SYBR Green, which incorporates into double-stranded DNA and emits a green signal after excitation at a wavelength of λ=494 nm. The green light is detected after each elongation step within a PCR cycle in real time, while the fluorescence increases proportionally to the amount of the PCR product. All experiments were run on a Rotor-Gene 6000/Q (Corbett/Qiagen) in 10 µl reactions using the
SensiMix™ SYBR No-ROX Kit (Bioline). 2 µl cDNA were mixed with a pair of primers for specific genes (16S rRNA, ihfB, cysG), SYBR Green Master Mix (2x), and including dNTPs and DNA polymerase were also added. PCR was performed according to the manufacturer’s recommendations. Relative mRNA expression was calculated by the comparative delta delta Ct method.
3 Materials and Methods 45 Temperate Time Cycles Polymerase activation 95°C 10 min 1 Denaturation 95°C 15 s 40 Annealing 60°C 15 s cycles Elongation 72°C 15 s Melting curve 60-95°C in 0.5°C steps 5 s each step
3.2.6 QuickChange site-directed mutagenesis
The QuickChange site-directed mutagenesis is used to create point mutations, to switch amino acids, or to delete or insert single or multiple amino acids. The basic procedure requires a double-stranded DNA vector with a pair of synthetic complementary oligonucleotide primers containing the desired mutation. During the temperature cycling with PfuUltra High-Fidelity
DNA polymerase AD (Agilent Technologies), a mutated plasmid containing staggered nicks was synthesized. After digestion of the parental DNA template with DpnI, the synthesized site-mutated DNA was selected. Quick-change PCR: 10x PfluUltra HF reaction buffer 2.5 µl DNA template 25 ng forward primer (10 pmol/µl) 1.1 µl reverse primer (10 pmol/µl) 1.1 µl dNTP mix (2.5 mM each) 2 µl PfuUltra-HF DNA polymerase 1 µl
H2O to 50 µl PCR program: 1 95°C 60 s Initial denaturation 2 95°C 30 s 55°C 30 s 17 cycles (Tm-5)°C 60 s/kb 3 68°C 5 min Final Extension 4 4°C pause Hold
After the PCR run, 1 µl DpnI was added to digest the parental template DNA at 37°C for 1 hour.
Next, 1 µl digested PCR product was transformed into 50 µl Top10 competent bacteria.
3.2.7 Random mutagenesis by error-prone PCR
Random mutagenesis is used to introduce mutations into a gene sequence to create a “library” containing thousands of variants of the gene. The variants of the gene in the library would
3 Materials and Methods 46 contain different mutations resulting in slightly altered amino acids. The most commonly used random mutagenesis method is error-prone PCR, which introduces random imposing mutations during PCR by adding manganese ions or by biasing the dNTP concentration. In this study, altered MnCl2 and dGTP concentration were used to modulate the mutation rate.
Error-prone PCR reaction (50 µl ): Calculated mutation rate (per 100 bp) 0.3% 0.4% 0.5% 0.8% 10x Thermo polymerase buffer 5 µl 5 µl 5 µl 5 µl forward primer (100 pmol/µl) 0.5 µl 0.5 µl 0.5 µl 0.5 µl reverse primer (100 pmol/µl) 0.5 µl 0.5 µl 0.5 µl 0.5 µl Template (1 ng/µl) 5 µl 5 µl 5 µl 5 µl MnCl2 (8 mM) 2 µl 2 µl 4 µl 4 µl dGTP (2 mM) 1 µl 2 µl 2 µl 5 µl dNTP mix (2.5 mM each) 3.5 µl 3.5 µl 3.5 µl 3.5 µl Taq DNA polymerase (NEB) 0.5 µl 0.5 µl 0.5 µl 0.5 µl H2O to 50 µl to 50 µl to 50 µl to 50 µl PCR program: 1 94°C 60 s initial denaturation 2 94°C 60 s (Tm-5)°C 60 s 30 cycles 72°C 60 s/kb 3 72°C 5 min final extension 4 4°C pause hold
3.2.8 Bacterial assays
3.2.8.1 Bacterial growth measurement
To monitor the bacterial growth after expression of GST-tagged proteins, 10 ng empty pGEX4T1 vector (GST alone) or pGEX4T1 vector carrying the GOI were transformed into E. coli BL21 competent cells. The colonies were picked the next day from the LB-Agar plate, and cultured in 5 ml LB medium overnight. On the third day, 1:100 of the overnight cultured bacteria were seeded into 50 ml fresh LB medium and incubated at 37°C with shaking. When
OD600 of 0.6 reached, 0.5 mM IPTG was added to induce protein expression. At the same time, the parallel samples without IPTG induction were prepared as control. OD600 readings were taken every half hour.
3.2.8.2 Staining of bacterial DNA with DAPI and electron microscopy
During the bacterial growth measurement, 1 ml bacterial culture was removed and prepared
3 Materials and Methods 47
(centrifugation, 5000 x g, 10 min) for DAPI staining and electron microscopy.
For Hoechst staining, the bacteria were washed once with PBS and collected by centrifugation
(17,000 x g, RT, 5 min). 100 μl paraformaldehyde (PFA, made in 1x PBS, pH 7.4) were added to the bacterial pellet to fix the bacteria (5 min, RT). After one wash with 1 ml PBS, 100 μl lysozyme (2 mg/ml in GTE buffer) was added to lyse the bacterial membrane (RT). After another wash with 1 ml PBS, 100 μl 2% BSA in PBS was added for blocking (20 min, RT), and finally the bacteria were stained by adding 100 μl Hoechst (1 mg/ml) in 2% BSA in PBS for 1 hour at RT. After three wash steps with 1 ml PBS each, the bacteria were resuspended in mounting medium, transferred to a slide and were covered with coverslips. GTE buffer: 50 mM Glucose 10 mM EDTA 25 mM Tris, pH 8.0
For analyses via electron microscopy the bacteria were fixed in 3% glutaraldehyde (Agar
Scientific, Wetzlar, Germany) for at least 4 h, washed in 0.1 M Soerensen’s phosphate buffer
(Merck, Darmstadt, Germany), and post-fixed in 1% OsO4 in 17% sucrose buffer. After fixation, the bacteria were embedded in 2.5% agarose (Sigma, Steinheim, Germany) and then rinsed in 17% sucrose buffer and deionized water and dehydrated by an ethanol series (30, 50,
70, 90 and 100%) for 10 min each with the last step carried out three times. The dehydrated specimens were incubated in propylene oxide (Serva, Heidelberg, Germany) for 30 min, and then in a mixture of Epon resin (Sigma) and propylene oxide (1:1) for 1 h and finally in pure
Epon for 1 h. Samples were embedded in pure Epon. Epon polymerization was performed at
37°C for 12 h and then 80°C for 48 h. Ultrathin sections (70-100 nm) were cut with a diamond knife and picked up on Cu/Rh grids. Negative staining by uranyl acetate and lead citrate (both
EMS, Munich, Germany) was performed to enhance TEM contrast. Samples were viewed at an acceleration voltage of 60 kV using a PhilipsEM400T electron microscope. Samples were kindly prepared by Hiltrud Königs (Institute for Pathology, RWTH Aachen University)
3.2.8.3 Bacterial growth inhibition assay
Plasmid DNA was transformed into E. coli BL21 competent cells (see chapter 3.2.1.7). After
3 Materials and Methods 48
40 min shaking at 37°C, 100 μl transformation was spread onto LB-agar plate either with (0.1 mM) or without IPTG and incubated overnight at 37°C.
3.2.9 Electrophoretic Mobility Shift Assay (DNA-EMSA)
The Electrophoretic Mobility Shift Assay (EMSA), also referred to as band shift assay, is a common technique used to characterize protein DNA/RNA interactions. DNA-EMSA gel assay is based on the observation that complexes of protein and DNA migrate through an agarose gel more slowly than free DNA fragments. Therefore, the binding of protein results in a characteristic upward shift of the DNA on a gel, as monitored under UV light.
Purified proteins were dialyzed on Millipore filters (type: VSWP, pore size: 0.025 μm) against nE100 buffer (20 mM HEPES-KOH, pH 7.6, 100 mM NaCl, 10 mM NaHSO3, 1 mM EDTA ) in the presence of 1 μg/μl of bovine serum albumin (BSA) for 90 min at 4C. 10 ng of supercoiled
DNA was incubated with increasing amount of proteins at 37C for an hour in a total volume of
30 μl nE100 buffer. Prior to loading the samples were adjusted to 10% glycerol, and loaded on
0.6% agarose gels in 1x TBE buffer (50 mM Tris base, 80 mM boric acid, 1 mM EDTA, pH 8) and run at 2 V/cm for 16 hours. The DNA/protein complexes were stained by SYBR Gold
(Invitrogen) and monitored by an Intas image device.
3.2.10 Chromatin assembly assay in vitro
To closely mimic the chromatin environment in vitro, chromatin was assembled by using a commercially available chromatin assembly kit (Active Motif). This kit is based on an
ATP-dependent assembly method that uses purified HeLa core histones, recombinant human chromatin assembly complex ACF and the histone chaperone NAP-1 (h-NAP-1) for assembly of regularly ordered, periodic arrays of nucleosomes. Assembly was carried out by following the manufacturer’s instructions.
GST and GST-tagged proteins were incubated with assembled chromatin at 37C for 30 min.
Then chromatin alterations were analyzed by partial digestion. Each reaction was partially digested with the Enzymatic Shearing Cocktail for 0, 1, 2 or 4 min, respectively. Samples were subsequently deproteinized with Proteinase K, phenol/chloroform extracted and analyzed by
3 Materials and Methods 49
1% agarose gel electrophoresis following the instructions of kit. The purified digested nucleosomal DNA was visualized by SYBR Gold staining under UV light.
3.2.11 Creation of HeLa S3 DEK knockout cells
For the generation of DEK knockout cells (created by Malte Prell), HeLa S3 cells were transfected with Lipofectamin and 5 µg DNA of each of two TALEN plasmids, which target the transcription site of the human DEK gene. After 6 h, the medium was replaced with the normal culture medium (DMEM). Two days after transfection, the cells were seeded in six
96-well plates at a calculated rate of 0.8 cells per well to avoid multiple colonies in one well.
The medium was replaced every two days with 50% fresh medium and 50% conditioned medium. 10 days after seeding the cells were checked for wells containing single colonies and only positive wells were cultured for further experiments. Six weeks after transfection the cells were screened by immunofluorescence with DEK-specific monoclonal antibodies. 7 weeks after transfection the cells were tested using immunoblotting for DEK-negative cells with
DEK-specific monoclonal antibodies. 1,200 cells were seeded in 96 wells and 4 different
HeLa S3 clones were selected in total.
3.2.12 Expression of DEK fragments in DEK knockout cells and HeLa S3 cells using lentiviral delivery
For re-expression of DEK or DEK fragments a lentiviral system based on the pTRIPZ backbone (Open Biosystems) was used. Production of virus-containing supernatant was carried out according to a standard protocol developed by the RNAi consortium at the Broad
Institute (http://www.broadinstitute.org/rnai/public/resources/protocols). A 4-day protocol was used. On day one, HeLa S3 packaging cells were seeded at 1.3-1.5x105 cells/mL (6 mL per plate) in low-antibiotic growth media (DMEM + 10% FBS + 0.1x Pen/Strep) in 6 cm tissue culture plates and the cells were incubated for 24 hours (37 °C, 5% CO2), or until the following afternoon. After the cells reached ~70% confluence, the transfection was carried. To start, 900 ng of packaging plasmid (psPAX2), 100 ng of envelope plasmid (pMD2.G) and 1 μg of the individual pTRIPZ plasmids were combined to a final volume of 30 μL OPTI-MEM. To
3 Materials and Methods 50
90 l OPTI-MEM, 6 l of Lipofectamine 2000 were added and incubated for 5 min. Then the plasmid mix was added dropwise to the transfection reagent after being incubated for 20-30 min at RT prior to adding it to the cells. The cells were then transferred to a biosafety level 2 laboratory and were cultured for 18 hours. The next morning, the medium was removed and was replaced with 6 ml of high-serum medium and the cells were cultured for another 24 hours. The next day the virus-containing supernatant was harvested and filtered through 0.45
m filters prior to transduction of the target cells for 6 hours. After two rounds of transduction, target cells were cultured for at least 24 hours prior to the addition of 1 g/ml puromycin for selection.
3.2.13 Cell fractionation
After expression of GFP, GFP-tagged DEKwt, DEK1-187, or DEK1-187 #11 for two days in the presence of doxycycline, cells were washed three times on the plate with ice cold hypotonic buffer (20 mM HEPES, pH 7.6, 20 mM NaCl, 5 mM MgCl2, 1 mM NaV3O4, 10 mM NaF) and lysed in hypotonic buffer plus protease inhibitor cocktail (Complete tablets EDTA-free, EASY pack, Roche) by Dounce homogenization. After 15 min on ice, the cytosolic supernatant was separated from the nuclear pellet by centrifugation (500 x g, 4C, 5 min). The nuclei were resuspended in hypotonic buffer supplemented with 0.5% Nonident P40 (NP-40) and kept on ice for 15 min to lyse the nuclear envelope. The free nucleosolic proteins were separated by centrifugation (500 x g, 4C, 5 min). The pellet containing the chromatin fraction was extracted for 15 min on ice in extraction buffer (20 mM HEPES, pH 7.6, 1 mM NaV3O4, 10 mM NaF) plus NaCl in concentrations from 50 mM to 450 mM to release chromatin-bound proteins. The final pellet was extracted in 2% SDS. Proteins from each fraction were analyzed by
SDS-PAGE and immuoblotting using GFP-specific antibodies.
3.2.14 Analysis of chromatin changes with MNase digestion
Micrococcal nuclease (MNase) is an important reagent for characterizing chromatin structure because MNase cleaves preferentially in the linker DNA that lies between individual nucleosomes and can digest the chromatin to mononuleosomes. MNase digestion is frequently
3 Materials and Methods 51 applied to assess whether the nucleosomes in a chromosomal domain are covalently modified, and to determine binding of proteins to chromatin.
Cells were washed three times on the plate with pre-cooled hypotonic buffer, and harvested by scraping into 15 ml tubes. Cells were lysed in 5 ml hypotonic buffer plus protease inhibitor
Complete (Complete tablets EDTA-free, EASY pack, Roche) on ice for 7 min. Subsequently, the cells were dounced 10 times using a dounce homogenizer. NP-40 was added dropwise to a final concentration of 0.5% and the cells were incubated on ice for 15 min to lyse the nuclear envelope (vortex 2-3 times). After incubation, the supernatant was removed by centrifugation
(500 x g, 4C, 5 min) and the pellet was washed once with 10 ml hypotonic buffer and twice with 10 ml extraction buffer to remove the residual NP-40. The pellet (nuclei) was resuspended with an appropriate volume of extraction buffer depending on the pellet size. The nuclei suspension was diluted 1:500 in extraction buffer and the DNA concentration was measured by
OD260. The nuclei suspension was incubated with 20 units micrococcal nuclease/50 μg DNA
(300 U/μl, Thermo Scientific) in the presence of 2 mM CaCl2 at 18C. The reaction was stopped after 2, 4, 8, 16 and 32 min with supplement 8 mM EDTA (on ice). The soluble fraction (active chromatin) was separated by centrifugation (16,000 x g, 4C, 10 min). The pellet (inactive chromatin) was resuspended with extraction buffer supplemented with 8 mM EDTA. 5x
Laemmli loading buffer was added to the samples, the digested chromatin was separated on
1% agarose gel in 1x TAE buffer (110 V, 2.5 h), and the DNA was visualized by ethidium bromide staining under UV light. Hypotonic buffer: Extraction buffer: 20 mM HEPES, pH7.6 20 mM HEPES, pH7.6 20 mM NaCl 300 mM NaCl
5 mM MgCl2 1 mM NaV3O4
1 mM NaV3O4 10 mM NaF 10 mM NaF
3.2.15 RNA pull down assay
The Glutathione Sepharose 4 beads were washed three times with RNA structure buffer (10 mM
Tris, pH 7.0, 0.1 M KCl, 10 mM MgCl2). After centrifugation (100 x g, 5 min), the beads were incubated with the same amount of purified GST-tagged proteins in 500 μl RIP buffer (150 mM
3 Materials and Methods 52
KCl, 25 mM Tris, pH 7.4, 0.5 mM DTT, 0.5%NP-40, 1 mM PMSF, protease inhibitor cocktail) at 4C with rotation to bind the proteins to beads. After 1 hour, the unbound proteins
(supernatant) were discarded by centrifugation (100 x g, 5 min). Additionally, total RNA was prepared from HeLa cells with TRIZol Reagent (Life technologies) following the manufacturer’s instructions and quantified by Nanodrop. 1 μg total RNA was added to the beads and incubated with 500 μl RIP buffer for 20 min RT (or 1 h at 4C) with a rotor. The beads
(protein with bound RNA) were separated from supernatant (unbound RNA) and washed with
RIP buffer (3 times). The bound RNA was extracted from beads with TRIzol reagent and measured by Nanodrop.
3.2.16 In situ hybridization for total poly(A)-mRNA localization
Cells were seeded into 12-well plates with cover slips on the bottom that were treated with
0.01% poly-L-Lysine for 30 min one day before. The next day, cells were rinsed with 1x PBS three times and fixed with 4% paraformaldehyde (PFA)/PBS for 10 min at RT. Subsequently, the cells were permeabilized with 100% cold methanol (10 min) and rehydrated with 70% ethanol for at least 10 min. After aspirating the ethanol, the samples were incubated with 1 M
Tris, pH 8.0 for 5 min. Next, the cells were hybridized with 1 ng/μl Cy5-labbeled Oligo-dT
(Molbiol) in hybridization buffer at 37C (in a dark humidified chamber). After one wash with
4x SSC and two washes with 2x SSC, the samples were incubated with the primary antibody dilution (GFP mouse antibody, 1:1,500, Rockland) in 2x SSC/0.1% Triton X-100 at RT for 1 hour in the dark. The following secondary antibody dilution (anti-mouse 488, 1:1,000,
Rockland) and Hoechst (1:1,000, 1 mg/ml in stock) in 2x SSC/0.1% Triton X-100 (in a dark humidified chamber) were added after two rinse steps with 2x SSC. After 1 hour (RT), rinsed cells (twice with 2x SSC) were mounted onto slide with a drop of mounting medium. The samples were kept at 4C in the dark and sealed the next day by using nail polish. Images were taken with a Zeiss LSM 710 confocal microscope and edited by ZEN 2012 software (Zeiss).
3 Materials and Methods 53 20x SSC: Hybridization buffer: 3 M NaCl 1 mg/ml Yeast tRNA 300 mM sodium citrate-HCl, pH 7.0 0.005% BSA 10% Dextran sulfate 4x SSC, 2x SSC: 25% Formamide, deionized dilute 20x SSC with DEPC-treated water 2x SSC buffer
3.2.17 Northwestern assay
The northwestern assay, also known as northwestern blot, is a technique used to detect the interactions between proteins and RNA. Total RNA was isolated from cells with TRIzol
Reagent (Life technologies) following the manufacturer’s instructions. The total RNA was first dephosphorylated at the 5’ end in a 20 μl reaction including 2 U calf intestine phosphate (CIP,
NEB), 10x buffer for CIP, 10 μg RNA substrate and 40 U/μl RNase inhibitor (murine, NEB) at
37°C for 30 min. Subsequently, the total RNA was end-labeled with [-32P]-ATP in a 20 μl reaction including 20 U T4 Polynucleotide kinase (NEB), 10x buffer for T4 PNK forward reaction, dephosphorylated RNA, [-32P]-ATP (3000 Ci/mmol) and 40 U RNase inhibitor at
37°C for 30 min. GST-tagged fusion proteins were expressed in E. coli and purified by
Glutathione Sepharose 4B (GE Health) (see chapter 3.2.2.1 and 3.2.2.2). Under denaturing conditions, the proteins were separated on 12.5% SDS-PAGE, and transferred to PVDF membrane (see chapter 3.2.2.5). After the transfer, the proteins were renatured in renaturation buffer (0.1 M Tris-HCl, pH 7.5 and 0.1 %( v/v) NP-40) (RT,1 hour) and blocked in blocking buffer (0.1 M Tris-HCl, pH 7.5, 5 mM Mg(OAC)2, 2 mM DTT, 5 %( w/v) BSA and 0.01 %( v/v) Triton X-100) at RT for 5 min with shaking. Next the radiolabeled RNA was added to the renatured blot and hybridized in hybridization buffer (0.1 M Tris-HCl, pH 7.5, 5 mM
Mg(OAC)2, 2 mM DTT and 0.01 %( v/v) Triton X-100) for 30 min RT. After 4 wash steps with washing buffer (0.1 M Tris-HCl, pH7.5, 5 mM Mg(OAC)2, 2 mM DTT) (5 min/each), the radioactive signal was obtained by autoradiography. After this, the blot was rinsed with PBS-T and analyzed by immunoblot (see chapter 3.2.2.5). The primary anti-GST antibody (1:1000) and secondary HRP-conjugated antibody (1:10,000, donkey anti-goat HRP, Dianova) were used to detect the protein levels.
3 Materials and Methods 54
3.2.18 Preparation of Pentaprobe RNA for RNA band shift assay
Pentaprobes were cloned into the pcDNA3 plasmid under T7 promoter and kindly supplied from K. Bendak et al. (Bendak et al., 2012). The Pentabrobes (2-3 µg) were linearized with
ApaI (NEB), which locates at 3’ of the Pentaprobe insertion site. The 3’ overhang was filled with
Klenow (Thermo Scientific) in order to allow for optimal activity of the T7 polymerase. After purification of the linearized DNA, the Pentaprobe RNA was transcribed by RiboMAXTM Large
Scale RNA Production System-T7 kit (Promega). The reaction was performed in a 50 µl volume, with 1 µl T7 polymerase, 10 µl T7 buffer, 1 µg linearized DNA, 1.5 µl rGTP/rCTP/rATP (2.5 mM each), 0.5 µl rUTP (1 mM), 8 µl DTT (100 mM), 2 µl RNase inhibitor Murine (40,000 U/ml, NEB) and 1 µl [-32P]-UTP (10µCi/µl) at 37°C for 3 hours.
The formamide loading dye (50 ml: 49 ml formamide, 1 ml 0.5 M EDTA, 0.013 g bromophenol blue, 0.013 g Xylene cyanol) was added to the transcription mixture and then the samples were boiled at 95°C (45 seconds) and kept on ice until loading onto the gel. The RNA was purified via
8% denaturing polyacrylamide gels (14.5 ml 20% acrylamide/7 M urea in DEPC H2O, 21.4 ml 7 M urea, 4.1 ml 10x TBE, 45 µl 20% APS and 45 µl TEMED). Based on the radioactive signal, the gel containing the Pentaprobe RNA was cut out. The gel pieces were crushed and soaked in
H2O at 37°C overnight. The next day, the Pentaprobe RNA was extracted by addition of equal volumes of phenol/chloroform/isoamyl alcohol (25:24:1) and precipitated by 1/10 volume of 3
M NaOAC and 1 volume of 100% ethanol. The labeled RNA was heated at 95°C for 45 seconds and put straight on ice before use in RNA-EMSA.
3.2.19 RNA electrophoretic mobility shift assay (RNA-EMSA)
RNA electrophoretic mobility shift assay (RNA-EMSA) is used to detect the interactions between RNA and proteins. Similarly to DNA-EMSA, the binding of protein also results in a characteristic upward shift of the RNA on the polyacrylamide gel, as monitored using radiolabeled RNA.
In a 30 µl reaction, 2, 4, or 6 pmol purified DEK proteins were incubated with 1 µl
32 [- P]-labeled RNA in EMSA buffer (10 mM MOPS, pH 7.0, 50 mM KCl, 5 mM MgCl2, 1 mM
3 Materials and Methods 55
DTT, 10% glycerol) at 4°C for 30 min. The total RNA-protein solution was loaded to pre-run polyacrylamide gels (4 ml 40% polyacrylamide 19:1, 25 ml EMSA running buffer, 200 µl 20%
APS, 50 µl TEMED) and separated by 200 V for 1 hour. After drying the gel, the labeled RNA was detected by autoradiography.
4 Results 56
4. Results
Based on previous work, two distinct modes of action of DEK within the regulation of chromatin organization have been either determined (Figure 4.1, left side) or proposed (Figure
4.1, right side), which may or may not be subject to crosstalk. On the one hand, DEK can either bind directly to the hinge domain of HP1, or indirectly to its chromoshadow domain via a yet unexplored RNA-mediated mechanism, thereby playing a vital role in maintaining the binding of HP1 to the heterochromatic histone mark H3K9Me3. These protein-protein- and/or protein-RNA-mediated interactions of DEK affect heterochromatin integrity and are important for the maintenance of a proper cell type-specific balance between cellular eu- and heterochromatin (Kappes et al., 2011b). On the other hand, DEK has been identified as a candidate protein during a search for factors that influence chromatin structure and replication efficiency of chromatin templates. Indeed, subsequent work established DEK as a non-histone chromosomal factor with potential chromatin architectural functions based on its specific
DNA binding features, which to date have been identified and studied only in vitro.
Specifically, DEK binds in a sequence-unspecific manner to either protein-free or nucleosomal
DNA in vitro and introduces positive supercoils into DNA resulting in a change of topology
(Alexiadis et al., 2000; Waldmann et al., 2002). These activities suggest that DEK may be involved in the regulation of local or global chromatin organization through directly transmitted effects mediated by its DNA- or chromatin-folding activities. Therefore, DEK’s distinct DNA binding and folding features may influence the nucleosomal structure through direct protein-DNA binding, thereby affecting nucleosomal accessibility or structure with potential effects on all downstream cellular processes (Figure 4.1, right panel).
4 Results 57
Figure 4.1: Current models for DEK functions within the regulation of chromatin structure. Left (verified in cell culture and in the PEV model in Drosophila): DEK modulates local or global heterochromatin structure by augmenting the binding of HP1 to the heterochromatic mark H3K9Me3 (Kappes et al., 2011b). Right (prediction based on biochemical work performed in vitro): DEK may be directly involved in local or global chromatin organization through direct binding to DNA or chromatin. Crosstalk between these two models may exist.
4.1 DEK changes chromatin structure at the nucleosomal level in
vitro and in cells
To investigate whether DEK modulates nucleosomal accessibility, and therefore might have a direct role in chromatin structure, Simian Virus 40 (SV40) minichromosomes were incubated alone or with increasing amounts of recombinant His-DEK and then digested with micrococcal nuclease (MNase) for short periods of time (0, 0.5, 1, or 2 min). Without DEK, SV40 minichromsomes were rapidly digested into mono-, di-, tri-, or tetra-nucleosomes, as revealed by the length of the resulting nucleosomal DNA (Figure 4.2 A). However, incubation of the chromatin templates with DEK resulted in a high molecular weight DNA smear on the gels at short time points (Figure 4.2 A, asterisks). Additionally, the typical nucleosomal “ladder” was not produced, as only the position of the mononucleosome is clearly visible after two minutes of digestion. This clearly indicates the capability of DEK to condense chromatin, at least in vitro, and is in agreement with prior electron microscope studies (Waldmann et al., 2002). In addition, a substantial increase in nucleosomal repeat length (NRL) of approximately 50 bp at the
4 Results 58
mono-nucleosomal level was observed (Figure 4.2 A, red arrows). This elongated nucleosomal
DNA indicates that DEK might bind to the entry/exit site of nucleosomal DNA, which supports
a function for DEK at the nucleosomal level (Figure 4.2 A was generated in the laboratory of Dr.
Tanja Waldmann at the University of Konstanz, Germany and is displayed here with her
permission).
A
Figure 4.2: DEK alters chromatin structure in general and at the nucleosomal level. (A) DEK compacts chromatin in vitro. SV40 minichromosomes were incubated alone or with the indicated amounts of recombinant His-DEK produced in the baculovirus system for 1 h at 37°C, then digested by MNase for 0, 0.5, 1 and 2 min, deproteinized, and analyzed by 0.6 % agarose gel electrophoresis followed by ethidium bromide staining. (B) Stable HeLa S3 control or DEK knockdown cells: A1 (H1-LV vector system expressing two non-redundant shRNAs) or B1 (pLKO.1 vector system expressing 2 individual shRNAs): cells expressing one specific and non-redundant DEK-targeting shRNA; A or B: cells expressing a scrambled shRNA in the respective vector backbone as control. Equal amounts of chromatin, as determined by spectrometry, were analyzed by MNase digestion and the digested chromosomal DNA was analyzed by 0.6% agarose gel electrophoresis with subsequent ethidium bromide staining. The inlet shows the knockdown efficiency as revealed by immonoblotting with DEK-specific antibodies. -actin served as a loading control.
In order to investigate if this general chromatin-altering effect of DEK, which is particularly
pronounced at the nucleosomal level, is also true at the cellular level, the NRL of HeLa S3
cells was analyzed. HeLa S3 cells either stably expressing one or two distinct non-redundant
DEK-specific shRNAs (shDEK A1 and B1), or control cells carrying the corresponding vector
(control A and B) were subjected to MNase digestion. After digestion of equal amounts of
extracted bulk chromatin from these cells, the digested and deproteinized chromosomal DNA
was resolved by agarose gel electrophoresis and visualized by ethidium bromide staining
(Figure 4.2 B). In both DEK knockdown cell lines, a clear shortening of the NRL was observed
4 Results 59 in comparison to the matched control cells (Figure 4.2 B, left panel: one representative gel is shown). To avoid any spurious results from improper digestion due to differing amounts of chromatin between the cell lines, the experiment was repeated three times, and these showed little variation (Figure 4.2 B, right panel: the graph displays the average values of three independent experiments). Even though the effects are in the range of 5-10 bp, this is still remarkable given that bulk chromatin was used. This implies that DEK may also be involved in the regulation of nucleosomal accessibility in cells, and is in support of the hypothesis depicted in Figure 4.1.
4.2 Hypothesis-driven design for the creation of
“loss-of-DNA-binding” DEK mutants
Having determined that DEK appears to have a role in chromatin structure at the nucleosomal level in vitro and in cells, the next goal was to investigate the relevance of these functions in chromatin in greater detail. One possibility to achieve this is to simply delete the pseudo-SAP/SAP-box region of DEK, as it conveys DNA binding and folding activities, and then re-express this DNA binding- and folding-deficient deletion in cells. However, as this domain may also have additional functions and spans a considerable portion of the protein, the major focus was on creating “DNA-binding-dead” point mutations in order to gain a deeper understanding into the mechanism(s) of DEK’s function(s) in chromatin. As multiple strategies were planned for this purpose (presented below), we first lined up a general workflow for testing these mutants in vitro prior to assessing their role in cells. This workflow included the creation of specific point mutations via site-directed mutagenesis, followed by expression and purification of these mutants as GST fusions in bacteria, and subsequent assessment of DNA binding and folding activities using EMSA, topology, and chromatin assembly assays. Mutants showing substantial reduction or even loss of DNA binding in these assays were ideally subjected to circular dichroism (CD) spectroscopy or even NMR analysis in order to record and understand potential changes to the secondary structure. Finally, mutants that passed all tests were analyzed in comparison to wtDEK in cellular systems (Figure 4.3).
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Figure 4.3: Proposed workflow for creation and analyses of “loss-of-function” DEK mutants. See text above for details.
The NMR structure of the DEK fragment aa 68-226 was published a number of years ago
(Devany et al., 2008), and, importantly, Devany and colleagues discovered a novel
DNA-binding motif in DEK, namely the pseudo-SAP domain, which locates N-terminally of the canonical SAP-box and displays a similar structure to the SAP motif despite lacking any sequence homology. Based on this NMR structure and associated chemical shift perturbation analyses, a series of 24 single and 1 double alanine substitution of positively charged, surface-exposed amino acids were created (Figure 4.4 and Table 4). Devany and co-workers tested these mutants for their remaining DNA binding and folding activities via EMSA and topology assays (Devany et al., 2008). Surprisingly, no effect on either the DNA-binding activity or the DNA-supercoiling properties of those single or the double mutants was observed, suggesting a rather complex biology underlying the DNA-binding activity of DEK. DEK, at this point, is the only known protein exhibiting the DNA-binding domain architecture of a combined pseudo-SAP/SAP-box motif. Owing to this, the DNA binding activities of DEK may be largely dependent on the cooperation of its two N-terminal dsDNA-binding domains rather than a single amino acid.
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Figure 4.4: Schematic depiction of point mutations in DEK created by Devany et al.. Shown is the schematic structure of DEK with known functional domains (yellow: pseudo-SAP-box; red: SAP-box; blue: NLS; green: C-terminal DNA-binding domain). Position and nature of mutations created and tested by Devany et al. are depicted as color-coded lollipops (top).
Table 4: List of mutations in the pseudo-SAP/SAP-box domain (created by Devany et al.) Single mutations K84A, K87A, K100A, K101A, K102A, K111A, K124A, K125A, K137A, K143A, K144A, K145A, K150A, Lysine (K): K151A, K158A, K177A and K187A Arginine (R): R93A, R107A, R116A, R168A and R178A Tyrosine (Y): Y114A Double mutant K177A/R178A
As the efforts of Devany et al. yielded no loss-of-DNA-binding mutants, we next chose multiple different strategies to create such mutants in the pseudo-SAP/SAP-box domain of
DEK. The selection of candidate amino acids was guided by interspecies comparison, a newly generated DEK-DNA docking model, literature search, and other experimental evidence available to us, as is outlined below.
4.2.1 Analysis of DNA binding of the pseudo-SAP/SAP-box of DEK carrying mutations of conserved aromatic amino acids
As interspecies conservation is particularly high in the pseudo-SAP/SAP-box region of DEK, the sequences of multiple species were aligned in order to identify conserved amino acids which may indicate an involvement in DNA binding (Table 5). Indeed, at nine positions between alpha helices one and five in the pseudo-SAP/SAP-box region, aromatic amino acids were found to be conserved. In particular phenylalanines (F) or tyrosines (Y) were identified, or if absent, tryptophan (W) as a conservative substitution was present. These particular amino acids were furthermore displayed on a newly generated DEK-DNA docking model, which supported a feasible location for DNA interaction (Figure 4.5), as numerous studies have identified that aromatic amino acids play an important role in protein-DNA recognition. Although phenylalanine has been associated less commonly with the interactions with DNA (Ahmad et al., 2004; Mandel-Gutfreund and Margalit, 1998), it has been implicated in the formation of
4 Results 62 specific aromatic hydrogen bonds that are responsible for DNA binding (Parkinson et al.,
1996). Additionally, conserved phenylalanine (F) residues are more likely to be involved in
DNA binding, as they are often found in TATA box-interacting factors in which they mediate interactions with DNA (Baker and Grant, 2007; Luscombe and Thornton, 2002). Furthermore, experimental work has demonstrated that benzene and indole bind more strongly to DNA bases than other aromatic groups (Guckian et al., 2000). Therefore, it is tempting to speculate that the highly conserved aromatic amino acids are responsible for DNA binding and folding by DEK.
Table 5: List of conserved aromatic amino acids in the pseudo-SAP/SAP-box domain Helix 1 1 2 3 Adj. to 3 Adj.to3 4 4 5 Human F96 F97 Y114 F130 F133 F135 Y142 F152 F182 Bovine F F Y F F F Y F F Rat F F Y F F F Y Y F Mouse F F Y F F F Y F F Chicken F F Y F F F Y F F - - - Drosophila F F F F Y F (E) (N) (V) Xenopus Y F F F F F Y W F - - Rice F Y F F W F F (K) (Q) - - - Zebrafish Y F F F Y F (H) (S) (L) - - - - Arabidopsis F F F W F (H) (K) (K) (C) Phenylalanine (F) and Tyrosine (Y) show high conservation through species. Amino acids in red denote species-specific substitutions with similar chemical propensities.
Figure 4.5: DEK-DNA docking model for the identification of amino acids involved in DNA binding. Left: The DEK fragment DEK78-187 was used in this in silico docking model which was generated using http://cssb.biology.gatech.edu/skolnick/webservice/DP-dock/index.html. The helices and extended loops in the pseudo-SAP/SAP-box domain were labeled with different colors. Light blue: -helix 1, dark blue: -helix 2, cyan: -helix 3, green: loop connecting pseudo-SAP and SAP-box, light pink: -helix 4, dark pink: -helix 5. Right: All conserved aromatic amino acids (F and Y) were labeled with sticks and spheres.
All conserved aromatic amino acids (F and Y), as shown in Table 5, Figure 4.5 and Figure 4.6
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A, were subsequently mutated to alanine (A) using QuickChange site-directed mutagenesis. In order to facilitate expression and to avoid interference from DNA binding activities present in the C-terminal DNA-binding domain, the DEK fragment 1-187 was used for cloning. After verification of the individual mutations via sequencing and alignment, the individual mutants
(GST-DEK1-187 plus mutations as shown in Fig. 4.6 A) were expressed and purified from E. coli and analyzed in comparison to GST-DEK1-187 (control) by EMSA (Figure 4.6 B). As expected, GST-DEK1-187 displayed high affinity to supercoiled (form I) and also relaxed
(form II) DNA, and formed the characteristic large nucleo-protein complexes that remained in the pockets of the gels (Figure 4.6 B, control). However, despite of some mutants showing reduced DNA binding affinity (F96/97A, Y114A, Y142A, F152A and F182A), none of the mutants showed loss of DNA binding.
Figure 4.6: Analyses of recombinant GST-DEK carrying aromatic amino acid mutations via EMSAs. (A) Schematic depiction of created aromatic acid mutations within DEK. The positions of all conserved aromatic amino acids (F and Y) within the pseudo-SAP/SAP-box domain were labeled with color-coded lollipops. (B) EMSAs. 10 ng of supercoiled DNA (I: superoiled DNA; II: relaxed DNA) was incubated without protein (minus, lane 0) or with increasing amounts (4, 8, 16, 32, 64 and 128 pmol) of GST-DEK1-187 (control, lane 1-6, upper and lower panel) or with individual GST-DEK1-187 mutants: F96/97A (upper panel, lane 7-12), Y114A (upper panel, lane 13-18), F130A (upper panel, lane 19-24), F133/135A (upper panel, lane 25-30), Y142A (lower panel, lane 7-12), F152 (lower panel, lane 13-18) and F182A (lower panel, lane 19-24) at 37°C for 1 hour. Then the DNA-protein complexes were separated on 0.6% agarose gels in 0.5x TBE and visualized by SYBR GOLD staining. Utilized proteins were further analyzed by SDS-PAGE and coomassie staining (lower, right panel), showing that similar concentrations of individual protein preparations were used. Note that BSA, which is added prior to filter dialysis and does not affect the DNA-binding activity, varies in concentration.
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As the double mutant F96/97A and the single mutant F182A showed less DNA-binding activity, we thought that combining these three mutations into one triple mutant protein
GST-DEK1-187 F96/97/182A, would results in substantially reduced DNA binding affinity. As shown in Figure 4.7, the triple mutant protein indeed exhibited less DNA-binding activity as compared to the control protein. However, the residual DNA-binding activity was still substantial, and therefore not adequate for downstream studies of this triple mutant. In addition, proteins from multiple independent purifications showed variations in their
DNA-binding activity (data not shown).
Figure 4.7: Analyses of recombinant GST-DEK carrying the triple mutation F96/97/182A via EMSA. EMSA was performed without protein (minus, lane 0), or with increasing amounts (4, 8, 16, 32, 64 and 128 pmol) of GST-DEK1-187 (control, lane 1-6), mutant proteins GST-DEK1-187 F96/97A (lane7-12), GST-DEK1-187 F182A (lane13-18), and triple mutant GST-DEK1-187 F96/97/182A (lane 19-24) and analyzed as described in Figure 4.6. Utilized proteins were further analyzed by SDS-PAGE and coomassie staining (right panel).
4.2.2 Analysis of DNA binding of the pseudo-SAP/SAP-box of DEK carrying mutations of arginines
As all aromatic amino acid mutants created still showed substantial DNA-binding activity, we focused next on arginine, an amino acid known to potentially intercalate into DNA. Indeed, six arginines were found in the pseudo-SAP/SAP-box of DEK (Figure 4.8). It was attractive to mutate arginines as previous data showed that incubation of linear DNA with DEK resulted in an increase in length of approximately 10% as seen in electron microscopy studies (Waldmann et al., 2002). Additionally, the DNA binding and folding activities of DEK were shown to be competitively reduced by the addition of ethidium bromide (Tanja Waldmann, personal communication), which is a strong intercalator into the minor groove of DNA. Together, this
4 Results 65 evidence may indicate that DEK can intercalate into the minor groove of DNA via arginines.
Furthermore, statistical analysis of atomic interactions revealed that arginine has a higher propensity to interact with base edges than with the rest of the nucleotide (Lejeune et al., 2005).
Rohs et al. proposed a new mode for protein-DNA recognition via the recognition of local sequence-dependent minor-groove shapes. Among all amino acid residues that contact the minor groove, 28% are arginines, which are particularly enriched in narrow minor groove-binding proteins (a groove width of <5.0 Å) (Rohs et al., 2009).
Figure 4.8: Depictions of the positions of arginines in the pseudo-SAP/SAP-box using the DEK-DNA docking model. Similar to Figure 4.5, all arginines (R) in pseudo-SAP/SAP-box domain were displayed by sticks and spheres (right panel) within docking model (left).
To examine whether arginine residues are responsible for DEK’s binding to DNA, arginines located in the pseudo-SAP/SAP-box domain were mutated to alanine using again the
QuickChange approach (Table 6). After verification of the respective new mutants through sequencing, they were expressed, purified, and tested in EMSA in comparison to the control protein GST-DEK1-187 (Figure 4.9). Intriguingly, none of the new mutants showed loss of
DNA-binding activity. Some mutants showed slightly reduced DNA-binding activity (R93A,
R107A, R116A and R168A); however, some showed even stronger DNA-binding activity
(R153A and R178A).
Table 6: List of arginine to alanine mutations in the pseudo-SAP/SAP-box domain Arginine mutants (R) R93A, R107A, R116A, R153A, R168A and R178A
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Figure 4.9: Analyses of recombinant GST-DEK carrying arginine mutations via EMSAs. (A) Schematic depiction of created arginine mutations within DEK. The positions of all arginines within the pseudo-SAP/SAP-box domain were labeled with color-coded lollipops. (B) EMSAs were performed without protein (lane 0) or with increasing amounts (4, 8, 16, 32, 64 and 128 pmol) of GST-DEK1-187 (control, lane 1-6), GST-DEK1-187 mutant proteins: R93A (upper panel, lane7-12), R107A (upper panel, lane13-18), R116A (upper panel, lane 19-24), R153A (upper panel, lane 25-30), R168A (lower panel, lane 7-12) and R178A (lower panel, lane 13-18) and analyzed as described in Figure 4.6. Utilized proteins were further analyzed by SDS-PAGE and coomassie staining (right panel).
In summary, a series of mutations in aromatic amino acids (F and Y) and arginines (R), in addition to the once created by Devany et al., were tested for their DNA-binding activity and are schematically summarized in Figure 4.10. Even though some mutants showed reduced
DNA-binding activity, it appears that the DNA-binding activity of DEK is neither based solely on aromatic amino acids nor on arginines, nor did the guidance by the DNA-DEK docking model and the NMR structure analysis yield single key amino acids responsible for DNA binding. This again suggests that the biology underlying the DNA-binding activity of DEK may be rather complex, as its unique DNA-binding domain architecture suggests.
Figure 4.10: Summary of all mutations created and tested so far either by Devany et al. (top) or within this thesis (bottom).
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4.3 The DNA binding activities of DEK interfere with bacterial
growth
4.3.1 Heterologous expression of DEK in bacteria inhibits growth in liquid cultures
Given that all DEK mutants created by rational and hypothesis-driven design did not yield the desired “DNA-binding-dead” mutants, we next chose to implement a random mutagenesis approach in order to obtain such mutants. In order to do so, an appropriate pre-screening procedure that has the capability to select for “loss-of-function” mutants has to be developed first, as sequencing and subsequent manual testing of a large number of mutants typically created by error-prone PCR was not feasible.
Based on this criterion, we evaluated our available DEK expression systems in the laboratory, namely bacteria, insect and yeast cell systems for potential use as pre-screening tool.
Interestingly, heterologous expression of DEK in all three systems appeared to induce substantial negative biological effects in the host cells. Expression in the yeast system
Klyveromyces Lactis produced hyperglycosylated DEK species with a very low yield, whereas expression of His-DEK in the baculovirus system produced hyperphosphorylated DEK (Malte
Prell and Ferdinand Kappes, personal communication). In bacteria, expression yield depended on the chosen tag. Whereas His-fusions were only marginally expressed, GST-DEK-fusions were produced. The resulting proteins, however, can be purified in rather low quantities and show instant and substantial protein degradation, pointing to a certain level of toxicity in bacteria (Figure 4.11 B). These observations furthermore suggest that expression of DEKwt may negatively interfere with the biology of bacteria, as was also seen in the other expression systems. In comparison to yeast or insect cells, bacteria were by far the most practical and most economical choice for a potential pre-screening system. Therefore, we explored whether expression of GST-DEK in bacteria may be a suitable system for screening of DEK mutants produced by random mutagenesis. First, bacterial growth after expression of GST (Figure 4.11
A, red dashed line) or GST-DEK (Figure 4.11 A, black dashed line) after induction with IPTG
4 Results 68 was observed over time by measurement of OD600. In parallel, growth of both bacterial strains without induction was also recorded as control (Figure 4.11 A, solid lines). Without induction both bacterial strains grew equally well, and the induction of GST expression slowed down the growth, as is expected by the immense production of GST (Figure 4.11 A, red lines). However, after a brief gap phase the induction of GST-DEK expression resulted in a complete stop of bacterial growth (Figure 4.11 A, black line). Successful expression of either GST or GST-DEK was further verified by lysing the respective bacteria and analysis via immunoblotting (Figure
4.11 B).
Figure 4.11: DEK expression in E. coli inhibits bacterial growth. (A) Bacterial growth curves. Cultured bacteria transformed with either GST (red lines) or GST-DEKwt (black lines), were induced with IPTG
(0.5 mM; dashed lines) or left uninduced (solid lines), and OD600 values were measured at the indicated time points (error bars: SD of six independent experiments). (B) Analysis of protein expression. The bacteria from A were lysed after 3 hours and protein expression was analyzed by immunoblotting using GST- and DEK-specific antibodies. Note that substantial protein degradation is visible with DEK-specific antibodies. (C) Growth inhibitory effect of DEK is transient. Bacteria as in A were kept under culture conditions without further addition of
IPTG and OD600 values were measured after 4, 6 or 12 hours.
We next wondered if the observed growth inhibition is due to cytostatic or cytotoxic effects of
DEK. This was assessed by monitoring growth after continued culture without further addition of IPTG. Interestingly, bacteria previously expressing GST-DEK started to re-grow after 12
4 Results 69 hours and reached OD600 values comparable to the other strains. This indicates that DEK-induced growth inhibition is due to transient cytostatic effects rather than cytotoxic effects
(Figure 4.11 C). Taken together, these results clearly show that the expression of GST-DEKwt in bacteria exhibits a growth inhibitory effect.
4.3.2 DNA binding activities of DEK appear to be responsible for bacterial growth inhibition
As DEK expression in liquid bacterial cultures inhibited growth, we were next interested if this effect could be also observed in classical transformation experiments. Therefore, easy-to-perform transformation assays, from now on referred to as “bacterial inhibition assay” were used (Figure 4.12 A). After transformation of the indicated plasmids to bacteria via heat shock, half of the transformants were plated on regular LB-Amp plates, and the other half were plated on LB-Amp plates supplemented with 0.1 mM IPTG. In this assay, immediate induction of GST expression showed no obvious effect on bacterial growth and therefore had no effect on colony formation. However, expression of either human GST-DEK or Drosophila GST-Dek resulted in empty plates, indicating complete inhibition of bacterial colony formation (Figure
4.12 B). As this halt in colony growth was observed with the expression of both human and
Drosophila DEK/Dek, this may indicate that conserved regions in these molecules are responsible for the inhibition. Therefore, the protein sequences of the two species were aligned via NCBI protein BLAST
(http://blast.ncbi.nlm.nih.gov/Blast.cgi?PROGRAM=blastp&PAGE_TYPE=BlastSearch&LIN
K_LOC=blasthome), and revealed that only two domains display high similarity. One localizes to the N-terminus with 44% identity, and overlaps with the pseudo-SAP/SAP-box domain, and the other domain is found on the C-terminal part with 38% identity and overlapping with the
C-terminal DNA-binding domain (Figure 4.12 C). This hints to DNA binding activities of
DEK as a potential cause for the observed bacterial growth inhibition.
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Figure 4.12: Expression of human DEK or Drosophila Dek in E. coli inhibits bacterial colony growth. (A) Overview of the transformation assay. (B) pGEX 4T1 plasmids carrying either the cDNA of GST (control), full-length wt human DEK (bottom left), or full-length wt Drosophila Dek (bottom right) were transformed into BL21, and grown overnight on LB/AMP plates either in the absence or presence of IPTG. Micrographs of the plates were taken the next day. (C) BLAST alignment of human DEK (NP_003463) and Drosophila Dek (CAD30679). Overlapping regions are shown in blue and red, whereas gaps in the alignment are indicated by letters in black and dashes. Amino acids shown in red indicate high interspecies conservation, whereas blue letters indicate less conservation. The positions of the DNA-binding domains of human DEK are indicated by black solid lines.
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4.3.3 Activities of the pseudo-SAP/SAP-box domain of DEK are responsible for bacterial growth inhibition
The key biochemical activities of DEK are the introduction of positive supercoils into closed circular DNA and preferential binding to DNA of unusual structure, such as cruciform DNA
(Waldmann et al., 2003; Waldmann et al., 2002). Importantly, these activities are transmitted by the pseudo-SAP/SAP-box domain (Bohm et al., 2005). To determine whether the observed inhibition of bacterial growth is due to the activities of this domain, we examined DEK truncations in the bacterial growth inhibition assay.
Figure 4.13: The pseudo-SAP/SAP domain in DEK is responsible for bacterial growth inhibition. (A) Transformation assay as in Figure 4.12 A with pGEX4T1 plasmids expressing the GST-tagged proteins full-length DEK, the N-terminal fragment DEK1-187 and the N-terminal truncations DEK1-87, DEK85-187. Top: Schematic domain structure of full-length DEK, DEK1-187, DEK1-87 and DEK85-187. (B) Bacteria grown in culture media either transformed with pGEX4T1-GST (red line), pGEX4T1-DEK1-87
(GST-DEK1-87, black lines) and pGEX-DEK85-187 (GST-DEK85-187, blue lines) were induced at OD600 0.6 with IPTG (0.5 mM; dashed lines) or left untreated (solid lines), and OD600 values were measured at the indicated time points.
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As shown in Figure 4.13 A, full-length GST-DEK and GST-DEK1-187 both suppress colony formation on agar plates in the presence of IPTG, as did the pseudo-SAP/SAP-box-containing fragment DEK85-187, indeed highlighting DNA binding activities as cause of growth inhibition. In support of this notion, expression of the far N-terminal part GST-DEK1-87, with no known DNA binding and folding activities, showed no inhibitory effect. This finding was furthermore evaluated in liquid cultures, and confirmed that expression of DEK85-187, but not
DEK1-87, inhibited bacterial growth (Figure 4.13 B). Because the truncation DEK85-187 possesses the pseudo-SAP/SAP-box functional domain of DEK, bacterial growth inhibition is very likely due to the DNA binding and folding activities of this domain.
4.3.4 Bacterial growth inhibition is due to abundant and aberrant compaction of bacterial DNA by DEK
Having identified that the pseudo-SAP/SAP-box domain of DEK was responsible for the observed bacterial growth inhibition, we next wished to investigate this phenomena in greater detail. We therefore monitored the higher-order organization of the bacterial genome, the nucleoid, using confocal and electron microscopy. First, bacterial DNA was stained with DAPI after either a brief induction of GST-DEKwt expression or without induction as a control.
Whereas bacterial DNA was found more or less evenly distributed in the bacteria prior to induction (Figure 4.14 A), expression of GST-DEKwt for only 30 min resulted in a markedly different appearance of the DNA as a dotted pattern. This may indicate that the DNA binding and folding activities of DEK induce substantial structural changes including substantial compaction of the bacterial genome. To further investigate if recombinantly-expressed
GST-DEKwt indeed binds to bacterial DNA, immunofluorescence was applied (Figure 4.14 B).
After 30 min of induced expression of GST-DEKwt, a clear co-localization between DEK and
DNA was observed in both the GST- and the DEK-specific antibody-treated cells (Figure 4.14
B). However, after pro-longed expression of GST-DEKwt (1.5 and 3 hours), only partial co-localization of the DEK signal with the DAPI signal was observed, with the bacterial genome undergoing strong compaction (Figure 4.14 B and C (left panel)).
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Figure 4.14: Analysis of DEK-DNA interactions in bacteria using confocal microscopy. (A) E. coli BL21 transformed with either the empty pGEX4T1 vector or pGEX4T1-DEKwt were cultured for 30 min in the absence or presence of IPTG (0.5 mM), fixed, stained with DAPI, and analyzed by confocal microscopy. Scale bar: 10 m. (B) Bacteria as in A were cultured for 0.5, 1.5 or 3 hours in the presence of IPTG, fixed, stained with GST- and DEK-specific antibodies and DAPI, and analyzed by confocal microscopy. Scale bar: 5 m. (C) Overlay of signals as in B between DEK, DAPI or GST were analyzed using Zeiss Zen software with the immunofluorescence intensities being measured along the indicated arrows. (D) A schematic model of proteolytic cleavage occurring to GST-DEK in bacteria as possible explanation for the diffusion and disconnection of the GST- and DEK-specific signals.
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Interestingly, a similar dislocation between GST- and DEK-specific signals was observed at later time points (Figure 4.14 B and C (right panel)). After 30 min of GST-DEKwt expression,
DEK and GST signals display a substantial overlay, but after 1.5 hours this overlay was strongly reduced. This loss of co-localization could be due to GST-DEK degradations as seen before in immunoblotting (Figure 4.11 B and Figure 4.14 D) and might represent a bacterial mechanism to ensure survival by inactivating the factor possessing the aberrant DNA binding activities.
Taken together, GST-DEK expressed in bacteria appears to interact with bacterial DNA and induces substantial compaction of the bacterial genome.
To detail this finding using a more suitable methodology, the structure of the bacterial nucleoid was further analyzed by electron microscopy (Figure 4.15). Without induction of protein expression, the bacterial DNA (visible in grey) was evenly distributed in the bacterial cells with no particular patterns visible (Figure 4.15, red arrows). Upon the expression of GST, a slight compaction of DNA was observed, which can be attributed to sterical effects due to high expression levels of GST in these cells. However, expression of DEK, even for only 30 min, induced a massive compaction of the bacterial genome (Figure 4.15 A, orange arrow). In order to quantify these compaction events, we counted bacterial cross sections either containing DNA
(Figure 4.15 A and B, blue arrow) or cross sections without any DNA staining visible (Figure
4.15 A and B, green arrow) in multiple micrographs from different replicates. Whereas the expression of GST alone only marginally increased the number of cross sections without DNA, expression of GST-DEK led to a three-fold increase in cross sections lacking DNA (Figure 4.15
C). Upon prolonged expression of GST-DEK, the grade of compaction increased to an even greater extent (data not shown). These findings clearly indicate that the DNA binding and folding activities of DEK induce substantial compaction of the bacterial genome. This furthermore suggests that, at least in bacteria, DEK has functions in higher-order genome organization.
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Figure 4.15: Analysis of bacterial DNA via electron microscopy. (A) Bacteria as in Figure 4.14 were analyzed by electron microscopy. Red arrows indicate selected bacterial nucleoids before protein induction, the orange arrow indicates a representative substantially compacted nucleoid (longitudinal section), and green and blue arrows indicate representative bacterial cross sections without or with DNA staining visible. Shown are representative micrographs with similar effects seen in three independent experiments. Scale bar: 2m. (B) Magnification of images of bacterial cross section. The cross section of bacteria in A (right panel) with or without DNA visible is shown with blue arrows and green arrows respectively. (C) The percent of bacterial cross sections with or without DNA visible was quantified and is displayed as bar diagram. At least 5000 cross sections per condition were counted in total.
This substantial compaction of bacterial DNA may severely interfere with the biology of bacteria and may be responsible for the observed growth inhibition. To further investigate this, we assessed the expression of three bacterial genes by qPCR upon expression of DEK (Figure
4.16). The expression of 16S rRNA (rrsA), which is commonly used as a reference gene in
4 Results 76 bacteria, increased to 1.8 fold upon induction of GST, while the expression of GST-DEK resulted in an 80% decrease in expression (Figure 4.16). As this effect was also seen for the genes ihfB and cysG (data not shown), this observation suggests that the expression of DEK in bacteria interferes with the gene expression program due to severe compaction of the genome, which would result in a transient growth inhibition as expression of vital genes would be either greatly reduced or abolished completely.
Figure 4.16: DEK expression in bacteria results in reduced expression of the16S rRNA gene. E. coli BL21 transformed with either the empty pGEX4T1 vector (expressing GST alone) or pGEX4T1-DEKw were cultured for 3 hours in the presence (gray bar) or in the absence (black bar) of IPTG. Total RNA was extracted and the expression of 16S rRtNA (rrsA) was monitored by RT-qPCR. Error bars indicate SD from three individual experiments, measured in technical duplicates.
Taken together, DEK expression in bacteria suppresses the expression of vital bacterial genes via robust and aberrant compaction of the bacterial genome, which results in the inhibition of bacterial growth.
4.4 Establishment of a screening procedure to select
“DNA-binding-dead” DEK mutants: Bacterial Growth
Inhibition Screen (BGIS)
The results presented so far indicate that the observed bacterial growth inhibition is most likely caused by the aberrant DNA binding and folding activities of DEK upon expression. As our goal was to identify mutants lacking precisely these activities, we hypothesized that combining a random mutagenesis approach with monitoring bacterial growth could provide a means to
4 Results 77 screen for mutants lacking DNA binding and folding activities, as these mutants are expected to lack any bacterial growth inhibitory effect. Based on this assumption we lined up a novel screening procedure which consisted of the following steps (Figure 4.17): a) mutagenesis and creation of mutant libraries via error-prone PCR; b) transformation of plasmid libraries to E. coli
BL21 and plating on regular LB/AMP plates; c) subsequent replica stamping of obtained colonies on LB-AMP plates supplemented with IPTG for four consecutive days; d) collection of surviving colonies on master plates; e) small-scale expression and selection of mutants with proper molecular weight (to avoid unwanted STOP codons, non-sense codons, etc.); f) sequencing of obtained plasmids and analysis of the mutations; and finally g) expression of mutants and assessment of their DNA-binding activity via EMSA followed by further downstream in vitro or cellular assays. Specifically, mutant libraries using either the cDNA of the DEK fragment 1-187 or solely the cDNA of DEK85-187, which spans the entire pseudo-SAP/SAP-box, were generated by error-prone PCR. This was achieved by altering manganese and dGTP concentrations during the PCR and was performed ultimately at calculated mutational rates of 0.3-0.9% mutations per 100 bp PCR product. The mutated PCR products were ligated to a pGEX4T1 vector backbone and transformed into E. coli BL21 and plated initially on LB-AMP plates without IPTG (Figure 4.17 A and B, I). The next day, the obtained bacterial colonies were replica-stamped onto LB-AMP-IPTG (0.1 mM) plates and cultured overnight, with repetition of this step for several rounds (Figure 4.17 A and B, II-V).
During this time most colonies stopped growing and vanished, yet the surviving colonies, which tended to overgrow, were collected on master plates in the presence of IPTG (Figure 4.17 A and
B, VI). Next, the individual colonies were picked and cultured in 96-well plates and the expression of the respective mutant proteins was analyzed by immunoblotting of the lysates with GST-specific antibodies (Figure 4.17 A, C, Supplementary Figure S2 and Supplementary
Figure S3). As expected, a significant number of colonies either produced no detectable protein, or appeared to have acquired STOP codons, as mostly products with the molecular weight of
GST were found (Figure 4.17 C). By sequencing some of these “GST-only” expressers we were able to indeed verify the introduction of STOP codons (data not shown). Importantly, we also found mutants that appeared to have the correct corresponding molecular weight in which either
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DEK1-187 or DEK85-187 was used for mutagenesis. From these colonies, the respective plasmid DNA was then extracted via mini-prep.
Figure 4.17: The Bacterial Growth Inhibition Screen—BGIS. (A) Schematic depiction of the individual steps of the screening procedure. (B) After transformation of mutants generated by error-prone PCR, bacteria were initially plated onto LB-Amp plates without IPTG (I). Subsequent rounds of replica stamping of the colonies to LB-Amp plates plus IPTG selected for mutants that lack bacterial growth inhibition (II-V). Surviving colonies were eventually collected on master plates (VI). Shown is one representative example of multiple replicates (see also in Supplementary Figure S1 for more examples). (C) Bacterial colonies as in B (B, V) were induced with IPTG (0.5 mM), lysed and analyzed by immunoblotting with GST-specific antibodies, and using GST and GST-DEK1-187 (or GST-DEK85-187) as positive controls. Shown are representative blots from multiple replicates (see Supplementary Figure S2 and Supplementary Figure S3 for the complete set).
In order to avoid any false-positive clones, we first re-transformed all obtained individual plasmids to our transformation assay. Indeed, some mutants selected in this way, still showed bacterial growth inhibition, and were therefore not investigated further but rather discarded.
However, the majority of mutants who come out of the BGIS showed no detectable growth
4 Results 79 inhibition (Figure 4.18).
Figure 4.18: Confirmation of DNA-binding-dead mutants in the re-transformation assay. All obtained plasmids from the BGIS (Figure 4.17, Supplementary Figure S2 and Supplementary Figure S3) were re-transformed into bacteria and bacterial growth was assessed in the presence or absence of IPTG. Shown are a few representative clones. Plasmids with remaining bacterial growth inhibition were discarded as false-positives.
Mutants showing no detectable growth inhibitory effect after re-transformation were finally sent for sequencing with the appropriate primers, and mutations in the individual plasmids were analyzed (Figure 4.19). As we initially performed our mutational analysis in the DEK fragment
1-187, we were surprised to identify several mutations that were located in the region DEK1-87
(Figure 4.19 A and Table 7); however these mutants clearly lacked the ability to inhibit bacterial growth in transformation assays. This may indicate that these particular amino acids may be involved in the regulation of folding of this DEK domain itself, and thus may result in an overall unfavorable protein structure (see chapter 5.1.4). Though these observations were indeed intriguing, we chose to exclusively use the pseudo-SAP/SAP-box domain (DEK85-187) for mutational analyses, which yielded a number of interesting mutational patterns (Figure 4.19 B and Table 7). Altogether, six rounds of the BGIS were carried out (Supplementary Figure S3).
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Figure 4.19: Schematic depiction of mutants obtained from the bacterial growth inhibition screen. (A) Mutational patterns obtained in the DEK fragment 1-187 after sequencing and alignment. Shown are the positions of mutated amino acids, with the wt condition being shown on the top and the obtained mutations depicted below. (B) Mutational patterns obtained in the DEK fragment 85-187 after sequencing and alignment. Mutants were numbered sequentially as obtained from the BGIS. Obtained mutations in A and B are further summarized in Table 7. (C) Distribution of identified mutations from A and B are summarized and schematically depicted in the DEK structure 1-187.
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Table 7: List of mutations obtained in DEK1-187 and DEK85-187 via BGIS DEK1-187 mutants colony number mutations #7 R65G, N126K, N181S #11 E30V, K158M #23 M1T E64K #27 V70A #38 E38K DEK85-187 mutants colony number mutations #1 K87E, E105D, L113I,R116T, K150G #2 L109Q, F130L, Q141L, K144M, K145E, N172S #5 K125R, S139G, K145E, N172D, K187E #6 L106P, G138E, C161R, L179 stop #8 F153S, S159N, V176M, R178G #11 H95R, K101M, K111I, K137E, S139R, K150E, L166S, N181D #14 K101E, E105G, K124E, V127G, E136G, N154D, M156T, L157I, I160T, E162G, K187R #18 K137E, M156L, K158E, S169P #19 E92D, T103T, K124R, S131G, K137E, N172D #28 E136G, K143E, G170R, K177 frame shift #36 K87R, E92G, F97S, N115D, V140A, E147G, N180S For clarity the mutations shown in Figure 4.19 were summarized in this table.
Through this novel screening procedure, more than fifty individual mutations were identified in total, which are summarized in Figure 4.19 C and Table 7. It appears that there were no obvious mutational hotspots. Importantly, no single or double mutations were selected through this screen, as most mutants carry at least four individual mutations. Thus it appears that particular combinations of amino acids are important in conveying DNA binding of DEK. Additionally, we noticed that mutations either cluster in the pseudo-SAP-box (not shown) or, more frequently, in the SAP-box domain, or in both domains.
4.5 Detailed analyses of obtained “DNA-binding-deficient” DEK
mutants
Based on the individual mutational landscapes in the obtained mutants, we selected three distinct mutants for detailed downstream analysis (Figure 4.20 C). Mutant #11 showed a total of
8 mutations spread throughout both DNA-binding domains. Mutants #8 and #18 showed a total of four mutations, all of which were localized in the SAP-box region. Prior to expression and purification of these mutants, we verified that no signs of growth inhibition were observed as assessed in transformation assays or in liquid cultures upon expression (Figure 4.20 A and B)
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Figure 4.20: DEK85-187 mutants #8, #11, and #18 show no signs of bacterial growth inhibition. (A-B) Plasmids of mutants, as shown in C, were re-transformed into bacteria and bacterial growth was assessed in the presence or absence of IPTG. A: transformation assay; B: growth in liquid culture (similar results were obtained with mutant #8 and #18 (data not shown)). (C) Schematic depiction of DEK85-187 mutants #8, #11 and #18.
Figure 4.21: Electron microscopy assay with DEK mutants. Bacteria expressing GST, or the GST-DEK85-187 mutants #11 and # 18 were cultured and induced with 0.5 mM IPTG at 37 °C for 1 hour, then analyzed by electron microscopy. Red arrows indicate the bacterial DNA before protein induction, and the blue arrows indicate the bacterial DNA after protein expression. Scale bar: 2 m. Similar results were obtained with mutant #8 (data not shown).
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Next we re-expressed these mutants in bacteria and subjected the resulting specimens to analysis via electron microscopy. As would have been expected, expression of these mutants in bacteria showed no sign of abundant compaction of the bacterial genome, thus additionally confirming the loss of DNA-compaction activities in these mutants (Figure 4.21).
To finally test these mutants for their DNA-binding activity, all mutant proteins were expressed in bacteria, purified and subjected to analysis via EMSA (Figure 4.22 A and B), as performed before (Figure 4.6, Figure 4.7 and Figure 4.9). Whereas GST-DEK85-187 bound preferentially to supercoiled (I) DNA before binding to relaxed DNA (II), mutant #11 exhibited no signs of
DNA-binding activity (Figure 4.22 A). Thus, we successfully identified a “DNA-binding-dead”
DEK mutant. Mutants #18 and #8 showed residual, albeit strongly reduced DNA-binding activity to both DNA forms. GST alone showed no DNA-binding activity. Interestingly mutant
#18 and #8 showed no activity in topology assays (data not shown), suggesting that both the pseudo-SAP as well as the SAP-box are needed for this activity. The mutants were furthermore subjected to EMSA with linear DNA (III) alone, showing identical behavior (Figure 4.22 B).
Prior to proceeding with testing of the “DNA-binding-dead” mutant #11 in the cellular environment, we wanted to investigate whether all of the eight mutations were in fact biologically relevant for DNA binding, or if some of them represented irrelevant “passenger” mutations that were simply co-selected during the BGIS. Therefore, all eight mutations were reverted back individually to the original wt condition and tested for their DNA-binding activity in EMSAs (Figure 4.23). To our surprise, it appeared that indeed all eight mutations were biologically relevant, as residual DNA-binding activity was found in every individual reverted mutant, though the R139S mutant only slightly rescued the DNA-binding activity. This suggests that our newly established bacterial screen truly has the capacity to select for biologically relevant mutations, and furthermore underscores that the DNA-binding activity of DEK is in fact indeed rather complex. Furthermore, bacterial growth inhibition appears to be dependent on the DNA-folding activity of DEK rather than simply on its DNA-binding activity, as mutants
#18 and #8 were selected as positive hits in the BGIS, despite exhibiting remaining
DNA-binding activity via the pseudo-SAP-box domain.
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Figure 4.22: Analysis of DEK85-187 mutants #8, #18 and #11 via EMSA. (A-B) 10 ng of supercoiled DNA (form I, A) or linearized DNA (form III, B) were incubated with increasing amounts (10, 40 and 300 pmol) of GST (lane 2-4), GST-DEK85-187 (lane 5-7), GST-DEK85-187 mutants #11 (lane 8-10), #18 (lane 11-13) and #8 (lane 14-16) or without protein (lane 1) at 37°C for 1 hour. The nucleoprotein complexes were analyzed on 0.6% agarose gels electrophoresis in 0.5x TBE and visualized by SYBR GOLD staining. Utilized proteins were further analyzed by SDS-PAGE and coomassie staining (A, B right panel). (C) Schematic depiction of DEK85-187 mutants #11, #18 and #8 indicating their DNA-binding activity on the right.
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Figure 4.23: EMSA analysis of individual mutations of the DEK mutant #11 reverted back to the wt condition. The identified mutations of DEK mutant #11 were individually mutated back to wt, and then the expressed, purified and tested by EMSAs. 10 ng supercoiled DNA were incubated with increasing amounts (10, 40 and 300 pmol) of GST-DEK85-187 (lane 2-4), GST-DEK85-187 mutants #11 (lane 5-7), #11 mutations reverted back to wt: R95H (lane 8-10), M101K (lane 11-13), I111K (lane 14-16), E130K (lane 17-19), R139S (lane 20-22), E150K (lane 23-25), S166L (lane 26-28), D181N (lane 29-31) or without protein (lane 1) at 37°C for 1 hour. The nucleoprotein complexes were analyzed on 0.6% agarose gels electrophoresis in 0.5x TBE and visualized by SYBR GOLD staining. Utilized proteins were further analyzed by SDS-PAGE and coomassie staining (lower panel).
4.6 Analysis of nucleosomal accessibility with a DEK mutant in
vitro
Having identified a true “DNA-binding-dead” DEK mutant and confirmed that all of its specific mutations were relevant for the DNA-binding activity, we next tested the activity of this particular mutant on chromatin structure. We only focused on the mutant #11 for this purpose and did not test mutants #18 and #8 in further assays. We first assembled regularly spaced chromatin using a commercially available chromatin assembly kit (Active Motif). The resulting chromatin templates were incubated with either GST, GST-DEK1-187 or
GST-DEK1-187 #11, and nucleosomal accessibility and nucleosomal repeat length were analyzed by treating the samples with the provided enzymatic shearing cocktail for the
4 Results 86 indicated time points (Figure 4.24). This produced a typical nucleosomal ladder (see also
Figure 4.2 A), and showed that incubation with DEK1-187wt resulted, as expected, in a more resistant chromatin structure and in longer nucleosomal repeat length. This effect was less pronounced than in Figure 4.2 A, in which full-length DEK had used, suggesting some influence by the C-terminal DNA-binding domain of DEK on the regulation of its overall
DNA binding. Importantly, the “DNA-binding-dead” mutant #11 did not alter chromatin structure, as neither an increase in nucleosomal repeat length, nor an overall enhanced protection of nucleosomal DNA was observed. This further confirmed that the newly identified “DNA-binding-dead” mutant #11 appeared to lack binding affinity to nucleosomal
DNA, thus we were able to exclude effects via DEK-histone mediated interactions.
Figure 4.24: Analysis of nucleosomal accessibility with the DNA-binding-dead DEK1-187 mutant #11. In a 100 µl reaction 1 µg of circular DNA was assembled into chromatin following the manufacturer’s recommendations. Following chromatin assembly, the reaction was divided in three tubes, and each tube either incubated with GST, GST-DEK1-187 or GST-DEK1-187 #11 at molar ratio of protein/DNA=100 at 37°C for 30 min. After incubation, chromatin was digested with Enzymatic Shearing Cocktail for 0, 1, 2 and 4 minutes, respectively. Samples were subsequently deproteinized with Proteinase K, DNA was extracted with phenol/chloroform and analyzed by 1% agarose gel electrophoresis followed by SYBR GOLD staining. Lane 2-4 represents the input.
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4.7 Investigation of the relevance of the DNA binding function of
DEK in cells
Through the above described screening procedure we were able to select and characterize one specific DEK mutant—DEK85-187 #11, which showed lack of DNA binding in various in vitro analyses. Additionally, we identified other potentially interesting mutants with some remaining
DNA binding affinity via one of the two N-terminal DNA-binding domains. Our next step was to develop a suitable cellular system to comparatively study the importance of the DNA binding and folding activities of DEK within cellular chromatin.
4.7.1 Creation of HeLa S3 DEK knockout cells
We have previously established multiple stable DEK knockdown (KD) cell lines, using constitutive or inducible lentiviral systems that deliver non-redundant shRNAs targeting the
DEK mRNA at non-redundant positions. However, residual DEK levels of 2-10% are present in even the best scenario, and re-expression of DEK in these cells is hampered, as we do not intend to alter the DEK cDNA to allow for shRNA escape (Ferdinand Kappes, personal communication). In order to test DEK functions using forced overexpression, we wanted to generate a “clean” cellular environment that was devoid of any residual DEK molecules, and we therefore designed TALEN (Transcription activator-like effector nuclease) pairs (sequence specific nucleases) in order to create knockout cell lines. The TALEN constructs, which target the region around the transcription start site, were developed together with the company
Cellectis (http://www.cellectis.com/en/gene-editing) and were thoroughly tested for any potential off-target effects and other relevant criteria prior to transfection. In addition, the targeted genomic region from all used cell lines was sequenced in order to identify any SNPs or other mutations present that could have compromised the success of this approach.
Using this technique, DEK knockout clones in multiple cell lines (created by Malte Prell,
Institute for Biochemistry and Molecular Biology, RWTH Aachen) were selected by immunofluorescence and immunoblotting. For the purpose here, only the HeLa S3 1E7 cell clone was used (Figure 4.25). Even though most DEK-specific antibodies available to us
4 Results 88 confirmed successful knockout, one DEK-specific polyclonal antibody (K877) detected low expression of a DEK truncation, which was present in all clones (Figure 4.25 A) and is currently under further investigation. Even though it appears that these cells are not true DEK knockout cells, we referred to them as such for the sake of simplicity throughout this thesis. A more compelling analysis of these cells is part of the work of Malte Prell and will be presented in his thesis. However, the overall morphology of the clone 1E7 was assessed for this work by confocal microscopy (Figure 4.25 B). In comparison to a matched control clone, the DEK knockout clone showed dramatic differences in overall morphology, nuclear size and chromatin structure. In particular, the gain in nuclear size was noteworthy.
Figure 4.25: HeLa S3 DEK knockout cells created via a TALEN approach. (A) DEK expression in HeLa S3 cells (control) and in selected DEK knockout cells was analyzed with DEK-specific mono- or polyclonal antibodies as indicated, -actin served as a loading control. (B) DEK knockout results in increased nuclear size, altered nucleolus distribution, and an overall change in morphology as revealed by confocal microscopy. Scale bar: 5 m.
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4.7.2 Creation of an inducible, lentivirally-delivered expression system based on the pTRIPZ vector
To investigate whether DEK re-expression in the DEK knockout cells could restore these particular nuclear alterations, and to further analyze the effect of “DNA-binding-dead” DEK mutants to chromatin structure, a lentiviral inducible system was chosen to express DEK and
DEK truncations or mutants in human cells. We first cloned the DNA binding regions and a series of other DEK regions or mutations as eGFP fusions into a custom-altered, inducible, lentivirally-delivered over-expression system that is based on the pTRIPZ vector (Open
Biosystems). An overview of all constructs created throughout this thesis is shown in Figure
4.26 A, though only a select few from this list were chosen for overexpression. The lentiviral vectors carrying either GFP alone or DEK fragments were transduced into HeLa S3 DEK control cells (not shown) and knockout cells (in cooperation with Malte Prell) in order to create stable cell lines. This system appeared to be tightly regulated and non-leaky, as no detectable expression of either construct was observed in the absence of doxycycline (Figure 4.26 B, left panel). Expression of some of these constructs was tested by induction with doxycycline, followed by subsequent testing of the behavior of the eGFP-DEK fusions in relation to endogenous DEK. By using cell fractionation approaches, immunofluorescence co-localization studies, cell cycle behavior, and other functional assays, we verified that eGFP-DEK behaves similarly to endogenous DEK (data not shown; Malte Prell and Ferdinand Kappes, personal communication). Thus, few artifacts due to over-expression of these eGFP fusions were expected. Upon the addition of doxycycline we found that most fragments were well expressed with approximately similar intensities. However, DEK fragments 85-187 and 68-226 showed no expression, suggesting some level of toxicity for the cells. Prompted by this finding we decided to re-express the DNA-binding-dead mutant #11 as a DEK1-187 fusion instead of as a
DEK85-187 fusion as originally planned.
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Figure 4.26: Cloning of an inducible, lentivirally-delivered expression system (pTRIPZ) and generation of stable cell lines via lentiviral transduction. (A) Overview of the DEK fragments created in the pTRIPZ system throughout this thesis. (B) Analysis of expression of DEK fragments. HeLa S3 DEK knockout cells (Figure 4.25) were transduced with lentiviral particles carrying pTRIPZ constructs with DEK fragments (from A), and were left untreated (left panel) or were induced with doxycycline (200 ng/ml) for 24 hours (right panel). Expressions were analyzed by immunoblotting with GFP-specific antibodies and -actin-specific antibodies as loading control.
4.7.3 Assessment of the subnuclear distribution of DEK fragment 1-187
After cloning, transduction and selection of control and DEK knockout cell lines expressing either GFP, GFP-DEKwt, the N-terminal fragments GFP-DEK1-187 or the DNA-binding-dead mutant GFP-DEK1-187 #11, we first assessed their nuclear distribution. Cellular DEK is mostly associated with chromatin and is eluted from the nuclei with 250 mM salt (Kappes et al.,
2001). Therefore we analyzed whether the “DNA-binding-dead” DEK mutant localizes differently compared to the wt form. GFP-tagged DEKwt, DEK1-187 and DEK mutant 1-187
#11 were expressed in DEK knockout cells, and then the protein distribution was examined using a cell fractionation assay (Figure 4.27). The majority of GFP was extracted in the cytoplasm, nucleosol and with 50 mM NaCl, as is expected of GFP (Figure 4.27 A). Consistent with previous studies, most of the GFP-tagged DEKwt became dissociated from nuclei with
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150 mM to 250 mM NaCl, and only small amounts were extracted with low or high concentration of salt (50, 100 and 450 mM) (Figure 4.27 B). In comparison to DEKwt, the
GFP-DEK1-187 truncation was extracted at lower salt concentrations (50 and 100 mM NaCl)
(Figure 4.27 C), and the DNA-binding-dead mutant was mostly present in the nucleosol, with some binding to chromatin, as seen in the fractions containing 50 or 100 mM NaCl, or in the pellet (Figure 4.27 D).
Figure 4.27: Assessment of subcellular distribution of DEK fragments. GFP (A) and GFP-tagged DEKwt (B), DEK1-187 (C) and DEK mutant 1-187 #11 (D) were expressed in DEK knockout cells by using the pTRIPZ inducible system. After two days of induction with doxycycline (200 ng/ml), cells were subjected to a cell fractionation approach yielding cytosol, nucleosol, and the nuclear fraction, which was eluted with 50, 100, 200, 250 and 450 mM NaCl. Equal aliquots of the supernatants and the final pellet were analyzed by SDS-PAGE and immunoblotting using GFP-specific antibodies.
4.7.4 The DNA binding and folding activities of DEK are important for global chromatin organization
Having created DEK knockout cell lines with the ability to express various DEK fragments using the inducible pTRIPZ lentiviral system and having evaluated the subnuclear localization of DEK1-187 and the DNA-binding-dead mutant, we next chose to investigate whether the
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Figure 4.28: DEK mutant #11 fails to reduce the nuclear size of DEK knockout cells. (A) HeLa S3 DEK knockout cells (Figure 4.25 B) were transduced with lentiviral particles either carrying pTRIPZ constructs with DEK wild type, DEK N-terminal 1-187, or DEK1-187 DNA-binding-dead mutant #11, and were induced with Doxycycline (200 ng/ml) or left untreated for 24 hours. Cells were fixed and the DNA was stained with DAPI. The images were taken and converted to binary images. (B) The nuclear size from A was measured by the Image J software. The graph shows the median nuclear size represented in arbitrary units. The box plot shows the 5th and 95th percentile. T-test was applied to the results as indicated. (C) The expression of the GFP-tagged DEK and GFP-tagged DEK fragments was monitored by immunoblotting with GFP-specific antibodies, and -actin antibodies as loading control.
DNA binding activities of these DEK molecules play a role in global chromatin organization.
To do this, we initially cultured cells on cover slips and induced the expression of DEKwt,
DEK1-187 or DEK1-187 #11 in the DEK knockout cells for 24 hours. After cell fixation and staining of DNA with DAPI, confocal tiling micrographs were taken (Figure 4.28). In order to allow for unbiased analysis of nuclear size, we used Image J. The Hoechst channel was converted to binary mode (Figure 4.28 A) and the nuclear size was measured by the Image J function “measure size” in at least 1000 cells for each condition (Figure 4.28 B) . Expression of full-length DEKwt significantly reduced the median nuclear size of DEK knockout cells, as was anticipated. Interestingly, the expression of DEK1-187 in these cells also decreased the nuclear
4 Results 93 size to the level observed with full-length DEK, suggesting that the N-terminus was sufficient for reverting the nuclear phenotype in knockout cells.
However, re-expression of the GFP-DEK1-187 #11 mutant did not lead to a reduction in nuclear size, even though the fragment was abundantly expressed as assessed by both immunoblotting (Figure 4.28 C) and IF analyses (data not shown). This provides strong evidence that the DNA binding and folding activities transmitted by the pseudo-SAP/SAP-box of DEK indeed have important functions in the regulation of nuclear size, and therefore may also play a vital role in the regulation of global chromatin structure.
Taken together, we have shown that DEK can directly interact with nucleosomal DNA, which lead to alterations in the structure and accessibility of chromatin in purified systems as well as in cells. We furthermore established a bacterial screen based on the aberrant DNA-compacting activities of DEK when expressed in bacteria, which allowed us to create numerous DEK mutants, including a “DNA-binding-dead” mutant, resulting from random mutagenesis.
Importantly, re-expression of this mutant in DEK knockout cells showed no compaction of cellular chromatin. Thus, the DNA binding and folding activities of DEK appear to be important for global chromatin organization. This is the first evidence suggesting a role for the pseudo-SAP/SAP domain of DEK in higher-order chromatin organization on a cellular level.
As studies have shown that the C-terminal DNA-binding domain may also contribute to the chromatin organization, we were next interested in creating a full-length DEK molecule lacking DNA binding via the N-terminal as well as the C-terminal DNA-binding domains to further investigate the DNA-binding activity of DEK.
4.8 Identification of the RNA-binding domain of DEK via the
BGIS
Having successfully used the BGIS to create DNA-binding-dead mutants in the N-terminal part of DEK, we next chose to use this screen for mutational analysis of the C-terminal
DNA-binding domain in order to create a full-length molecule lacking any DNA-binding
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Figure 4.29: Expression of DEK187-270 induces bacterial growth inhibition. (A) pGEX vectors carrying DEKwt, C-terminal fragment DEK187-375 and the C-terminal truncations: DEK187-270 and DEK251-375 shown in the upper panel were analyzed by the bacterial growth inhibition screen. (B) RNA-pull down assay with GST and GST-tagged proteins from A. Purified GST and GST-tagged DEK proteins: DEKwt, DEK1-187, DEK187-375, DEK1-87, DEK187-270 and DEK251-375 were coupled to Glutathione Sepharose 4 beads. Subsequently, protein-coupled beads were incubated with total RNA extracted from HeLa cells for 20 min at RT and bound RNA was precipitated and measured. activity. To do this, C-terminal truncations were initially analyzed in the transformation assay
(Figure 4.29 A). As anticipated, expression of DEK187-375 showed inhibition of bacterial colony formation to the same extent as the full-length protein. Surprisingly, the expression of
DEK251-375, which includes the C-terminal DNA-binding domain, did not inhibit bacterial colony formation. As this domain possesses DNA-binding activity but lacks DNA-folding activity, this may indicate that DNA folding activities are needed for the inhibition of bacterial
4 Results 95 growth (discussed in more detail in chapter 5.4.1). Instead, the DEK truncation DEK187-270 showed inhibition of bacterial growth, which was as pronounced as that from either
DEK187-375 or full-length DEKwt. Interestingly, besides being a rather unstructured region, this domain has thus far not been associated with any function. As about 10% of the cellular
DEK population is found associated with RNA, and no RNA-binding domain in DEK has been discovered yet, we speculated that this region might harbor an RNA-interacting or RNA-folding motif. This notion was further supported by the fact that DEK has been recently identified as an
RNA-binding protein in proteome-wide interaction studies, in addition to its already described roles in RNA biology (Castello et al., 2012). For our initial study investigating RNA-binding by DEK, RNA pull-down assays were performed with GST-tagged DEK truncations together with total RNA extracted from HeLa cells. After multiple wash steps, bound RNA in the individual samples was extracted, precipitated, measured by Nanodrop, compared to the initial input, and RNA binding affinity was expressed as % input. Our results from this indicated that
DEK187-270 exhibited the highest affinity for RNA (Figure 4.29 B).
However this assay was not very reliable because of low reproducibility, we next used IF to investigate potential co-localization of GFP-DEK fusion variants with protein-coding mRNAs
(Figure 4.30 B) as visualized by hybridization with a fluorescently labeled oligo-dT probe. In order to do so we used our stably transduced wt HeLa S3 cell lines, as created in chapter
3.2.12. The cells were grown on cover slips and were induced with doxycycline to either express GFP-DEKwt or other GFP-DEK truncations (DEK187-375, DEK186-260 and DEK delta delta (lacking all DNA-binding domains) (Figure 4.26 A and Figure 4.30 A). Cells were first fixed and hybridization with Cy5-labeled oligo-dT oligonucleotides was carried out to mark all mRNAs carrying a poly-A tail. Cells were mounted and analyzed using confocal microscopy (Figure 4.30 B). DEKwt and the C-terminal fragment showed substantial overlap with protein coding mRNAs, while the N-terminal truncation showed only minor co-localization. In contrast, the DEK truncations 187-260 and a truncation lacking all DNA binding domains (delta delta) showed the highest overlap with mRNA (Figure 4.30 B). This may indeed indicate that the fragment DEK187-260 could potentially be the long sought after
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RNA-interacting domain of DEK.
Figure 4.30: DEK187-260 is the potential RNA-binding domain of DEK. (A-B) The indicated DEK fragments from A were transduced into HeLa S3 cells to create stable cell lines. Expression of the individual fragments was induced with doxycycline (200 ng/ml) for 2 days. Cells were fixed, hybridized with oligo-dT oligonucleotides and stained using GFP-specific antibodies. The expression of the individual fragments was shown in Figure 4.26 B. Scale bar: 2 m. Right panels: Intensity profile graphs between mRNA (red) and DEK (green). Line scan graphs showing the immunofluorescence intensity along the indicated positioned arrows in the merge pictures. (C) Shown is a representative parent cell expressing GFP-DEKwt that divided into two daughter cells during telophase, underscoring the specificity of RNA hybridization. Scale bar: 2 m. Right panel: Intensity profile graph between mRNA (red) and DEK (green) was measured as described in B.
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These findings so far showed that the DEK fragment 187-270 may exhibit substantial
RNA-binding affinity. To further explore this possibility, northwestern studies with distinct
DEK fragments were performed (Figure 4.31). Purified GST, GST-DEKwt, DEK1-187,
DEK187-375, and DEK187-270 were separated by SDS-PAGE and transferred to PVDF membranes. After a renaturing step, the membranes were incubated with total RNA extracted from HeLa cells that was end-labeled with [-32P]-ATP. After extensive washing steps, bound
RNA was visualized by autoradiography. We found that RNA was not bound by high amounts of
GST, while wtDEK and the C-terminal fragment showed strong RNA binding. In contrast, the
N-terminal part of DEK showed negligible RNA binding, whereas bound RNA was observed with the fragment 187-270. These data furthermore suggest that the DEK fragment 187-270 is an RNA-binding domain in DEK, or at least, plays a large part in RNA-binding of DEK.
Figure 4.31: Analysis of DEK fragments through a north-western approach. 2 pmol of the indicated purified proteins were separated on 12.5% SDS-PAGE and transferred to PVDF membranes. Subsequently the proteins were renatured and incubated with radioactively-labeled RNA. After washing steps the membrane was exposed to an X-ray film (autoradiography). Duplicates of purified GST protein and triplicate of GST-DEK proteins were loaded. Total protein amounts were additionally monitored by immunoblotting using GST-specific antibodies (middle) and ponceau S staining (bottom).
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To further substantiate the finding that the DEK fragment 186-260 may be the RNA binding domain, we chose to carry out RNA-EMSAs. RNA-EMSAs using total RNA extracted from cells are difficult to carry out as the RNAs vary dramatically in size and nature of their secondary structures, which complicates assessment of binding. Recently, Bendak et al. developed a simple technique that allows screening for RNA-binding proteins in simple
RNA-EMSA experiments (Bendak et al., 2012). In an attempt to mimic the most abundant mRNA secondary structures, Bendak et al. designed sequences, all 516 bp in length based on computational algorithms that include every possible 5 nucleotide-long motif as six double stranded 100-bp long overlapping sequences (Pentaprobes, PP, Figure 4.32 A). In total, they cloned 12 sequences into pCDNA 3.1 and each of them allowed for transcription of one of the
12 Pentaprobes sequences as single stranded RNA of defined size. After radioactive labeling of the Pentabrobes via in vitro transcription and subsequent purification of the Pentaprobes, purified GST and GST-DEK fusions were mixed with one of 12 Pentaprobes. The protein/RNA complexes were then analyzed by RNA electrophoresis mobility shift assay (RNA-EMSA)
(Figure 4.32 B). GST alone was used as negative control to exclude any effect of the GST tag.
Incubation of purified full-length GST-DEKwt with different ssRNA Pentaprobes results in a clear shift in EMSAs (Figure 4.32 B and Supplementary Figure S4), implying that DEK can indeed bind to the ssRNA. Consistent with the northwestern assay, DEK1-187 exhibited only weak affinity for the Pentaprobes. However, the C-terminal fragment DEK187-375, as well as
DEK187-270, showed strong binding to ssRNA (Figure 4.32 B and Supplementary Figure S4).
These data strongly suggest that the DEK domain 187-270 indeed exhibits RNA-binding activity.
In summary, the BGIS facilitated us in identifying the RNA binding domain in DEK. These results suggest that this screening tool could also be applied to identify potential RNA-binding or folding domains in proteins and therefore may have an even broader applicability as initially thought.
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Figure 4.32: Binding of GST-DEKwt and DEK truncations to ssRNA Pentaprobe sequences. (A) Secondary structure of twelve RNA pentaprobes used for RNA-EMSAs (from (Bendak et al., 2012)) (B) RNA-EMSAs. 32P-labeled RNA Pentaprobes (PP5: left, PP6: right) were incubated without (minus) or with increasing amounts (0.5, 1 and 2 pmol) of GST or GST-tagged DEK or DEK truncations (DEK1-187, DEK187-375 and DEK187-270). All samples were separated on 8% polyacrylamide gels and exposed to an X-ray film (autoradiography).
4.9 Applications of the BGIS
4.9.1 DNA- and RNA-associated folding activities in the bacterial growth inhibition assay
Thus far, DEK-induced interference with bacterial growth guided us in mutating the DNA binding and folding domain in DEK that resulted in biologically valuable mutants for further downstream studies. Additionally, we identified a previously unknown RNA binding domain in
DEK via monitoring bacterial growth after expression of DEK fragments in bacteria. Based on these observations, we hypothesized that interfering with the biology of bacteria by heterologous expression of other genes of interest may be used more generally as a tool to study
DNA-binding and RNA-binding domains. In order to investigate this notion we subjected
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Figure 4.33: Expression of SAF-A and HMGB1 inhibit bacterial growth. (A) Transformation assay with GST-HMGB1 or GST-SAF-A. (B) Bacterial growth curve. Cultured bacteria transformed with either GST (red lines), GST-HMGB1 (upper panel, black lines) or GST-SAF-A (lower panel, black lines) were induced with IPTG (0.5 mM; dashed lines) or
IPTG was omitted (solid lines), and OD600 values were measured at the indicated time points. (C) Bacteria from B were analyzed by electron microscopy. Red arrows indicate regular bacterial nucleoid before induction of protein expression and the blue arrows indicate the bacterial nucleoid after protein expression. Scale bar: 2 m.
4 Results 101 well-known chromatin architectural factors to the BGIS (Figure 4.33).
As mentioned in chapter 1.2.5, HMGB1 and SAF-A are two of the most abundant non-histone architectural proteins in cells. HMGB1 functions as a linker protein that binds to the entry and exit site of nucleosomal DNA and remodels chromatin accessibility via its DNA-associated activities. As a SAP-box-containing protein, SAF-A specifically binds to scaffold/matrix attachment region (S/MARs) elements and has functions in the nuclear matrix and in global chromatin organization. Considering the similarity of chromatin-associated functions based on
DNA binding activities, both proteins were analyzed by the bacterial growth inhibition screen.
As expected, expression of both proteins led to bacterial growth inhibition similar to what was seen after the induction of DEK expression (Figure 4.33 A and B). Furthermore, expression of both proteins resulted in structural changes to the bacterial genome. HMGB1 led to a similar compaction of the DNA as seen with DEK, while SAF-A induced more subtle changes to the structure of the nucleoid (Figure 4.33 C). Thus, it appears that the effect of bacterial growth inhibition is not exclusive for DEK, as other chromatin architectural factors were capable of inhibiting bacterial growth. This hints at a rather broad applicability of the BGIS in studying
DNA binding activities of other factors.
These three factors, DEK, HMGB1, and SAF-A, all exhibit DNA folding activities combined with DNA binding affinities. Although the bacterial growth inhibition is induced by
DNA-associated activities, it seems that, at least with DEK, DNA folding rather than pure DNA binding is the essential key that leads to bacterial growth inhibition, as no effect was observed with the C-terminal DNA-binding domain of DEK. Additionally, DEK mutant #8 and #18 showed no bacterial growth inhibition, despite remaining DNA-binding activity in EMSAs. In order to better understand the kind of DNA-related activities that can be monitored by the BGIS, we next investigating a chromatin-associated protein with weak DNA binding affinity—the thrithorax protein ASH2L— in the bacterial inhibition assay (Figure 4.34 B).
ASH2L (Absent, Small, or Homeotic-Like) was originally identified based on the analogy with
Drosophila Ash2, which is a known transcriptional regulator (Ikegawa et al., 1999). It is known to be a component of the Set1/Ash2 histone methyltransferase (HMT) complex that specifically
4 Results 102 methylates lysine 4 of histone H3, which is associated with activation of gene transcription (Dou et al., 2006; Steward et al., 2006). As shown in Figure 4.34, five different ASH2L domains were tested for their inhibition of bacterial growth: a PHD finger domain, a helix-winged-helix
(HWH) motif, a central nuclear localization signal (NLS), a SPRY domain and an Sdc1/Dpy-30 interaction (SDI) domain. Upon examination via the BGIS, neither full-length ASH2L nor the truncations including HWH domain (aa 162-273), which has direct DNA-binding activity, showed signs of bacterial growth inhibition (Figure 4.34 B). Thus it appears that DNA folding activities of the proteins tested were responsible for the bacterial growth inhibition observed.
Consequently, the applicability of the BGIS may be limited to the study of DNA folding activities and may not be a useful tool for studying DNA binding activities of transcription factors or other proteins lacking DNA-folding activity.
Figure 4.34: Expression of ASH2L does not induce bacterial growth inhibition. (A) Schematic depiction of the structure of ASH2L (from (Ullius, 2014)). Shown are the identified functional domains of ASH2L: a PHD domain (aa 95-161), a helix-winged-helix domain (aa 162-273), a NLS (aa 299-316), an SP1a and RYanodine receptor domain (SPRY, aa 360-583) and a C-terminal Sdc1/Dpy-30 interaction motif (aa 602-628). (B) GST (control) and GST-tagged ASH2L full-length and indicated-truncations were analyzed in transformation assays.
4 Results 103
As we also identified an RNA-interacting motif in DEK using the bacterial growth inhibition assay, this may indicate that RNA-associated activities of proteins are also capable of inducing bacterial growth inhibition. As SAF-A possesses both DNA- and RNA-binding activities, we next investigated bacterial growth inhibition using specific SAF-A fragments. Based on the location of its two functional regions (SAP-box and RGG-box) (Figure 4.35 A), three SAF-A truncations SAF-A N280, SAF-A I280 and SAF-A C280 were individually cloned into the pGEX vector and examined by the BGIS (Figure 4.35 B). Interestingly, two SAF-A truncations,
SAF-A N280 containing the SAP-box and SAF-A C280 containing the RGG-box, exhibited similar bacterial growth inhibition to that observed from full-length SAF-A. However, the fragment SAF-A I280 did not show such an effect. Therefore, both DNA folding and
RNA-associated activities of this factor were capable of inducing bacterial growth inhibition.
Figure 4.35: Both DNA- and RNA-associated activities of SAF-A are capable of inducing bacterial growth inhibition. (A) Schematic depiction of SAF-A structure. (B) GST-tagged SAF-A full-length and indicated truncations (N280, I280 and C280) were examined by the bacterial growth inhibition screen. GST was used as a control.
To expand on these findings, we subjected another well-known RNA-binding protein,
ALY/REF to this assay (Figure 4.36). ALY/REF is an mRNA export factor and a crucial member of the TREX (Transcription/Export complex), though it was originally described as a transcriptional co-activator (Osinalde et al., 2013). ALY/REF binds to mRNA via a
4 Results 104
Figure 4.36: Identification of DNA-binding domains of ALY/REF by the bacterial inhibition screen. (A) Schematic depiction of the structure of ALY/REF. Via NMR analysis amino acids 77-181 were determined as RNA recognition motif (Perez-Alvarado et al., 2003). (B) Transformation assay with GST-tagged ALY/REF full-length and truncations ALY1-100, ALY77-182 and ALY182-257. (C) Bacterial growth curve. Cultured bacteria transformed with GST, GST-ALY, GST-ALY1-100, GST-ALY77-18 or
GST182-257 were induced with IPTG (0.5 mM; dashed lines) or IPTG was omitted (solid lines), and OD600 values were measured at the indicated time points and displayed with indicated color.
4 Results 105
RNA-binding motif (RRM) and is implicated in multiple processes including splicing, RNA export and nuclear RNA stability (Kohler and Hurt, 2007; Stubbs and Conrad, 2015; Stubbs et al., 2012) (Figure 4.36 A). As expected, expression of full-length ALY/REF displayed strong inhibition of bacterial growth. However, the RNA-binding motif, ALY77-181 showed no growth inhibitory effect. Surprisingly, the two other truncations ALY1-100 and ALY182-257 completely inhibited bacterial growth on plates as well as in liquid culture (Figure 4.36 B and
C). We hypothesized that these two domains in ALY may be DNA-folding domains, and therefore purified the GST-tagged ALY/REF truncations and analyzed them by EMSA (Figure
4.37).
Figure 4.37: EMSAs with ALY/REF full-length and truncations. EMSA was performed without protein (lane 0) or with increasing amounts (3.75, 7.5, 15, 30 and 60 pmol) of GST (control, lane 1-5), GST-ALY full-length (lane 6-10), GST-ALY1-100 (lane 11-15), GST-ALY77-181 (lane 16-20) and GST-ALY182-257 (21-25). The utilized proteins were visualized by coomassie staining.
4 Results 106
To our surprise, full-length GST-ALY showed strong binding to DNA, which to date has not been reported. Additionally, both truncations ALY1-100 and ALY182-257 also exhibited strong affinity to DNA, though slightly weaker than the full-length protein. However, the truncation
ALY77-181, which contains the RNA-binding motif, showed no binding to DNA. Prompted by this finding we tested whether ALY has DNA-supercoiling activity, which was confirmed by our findings (data not shown, Malte Prell and Nahleen Schilling, personal communication).
Therefore, by using this assay we were able to identify two new DNA-folding domains in
ALY/REF, a protein so far exclusively associated with RNA biology. These results further substantiated that DNA folding rather than DNA binding is important for bacterial growth inhibition (Malte Prell, personal communication).
Since DNA- and RNA-associated activities can induce bacterial growth inhibition, we were next interested in whether different protein classes also exhibit bacterial growth inhibition.
Therefore, we chose a set of proteins with no known direct functions in DNA or RNA metabolism. The importin family members and Ran function as transport proteins that control and regulate protein import and export between nucleus and cytoplasm. Growth factor receptor-bound 2 (Grb2) is an intracellular adaptor protein and is required for regulation of receptor tyrosine kinase (RTK) signaling. Src is one member of Src-family kinases (SFKs) that phosphorylate specific tyrosine residues in other proteins. The SH3 domain analyzed in this thesis is one functional region that possesses no catalytic activity and is responsible for interaction with other proteins. Given that no DNA- or RNA-binding or folding activities have been identified for this set of proteins, their expression in bacterial should show no effect on bacterial growth. Indeed, none of the tested proteins induced bacterial growth inhibition, as there was no difference in colony growth in comparison to the control after protein expression
(Figure 4.38).
4 Results 107
Figure 4.38: Proteins containing no DNA or RNA-binding activities exhibit no effect on bacterial growth. pGEX vectors carrying importin 1, 5 and 7, Ran, Grb2 and Src SH3 were transformed into E. coli BL21 and examined by BGIS.
4.9.2 Enzymatic activities inhibit bacterial growth
Having established that both DNA- and RNA-folding activities, but not DNA binding functions, can be identified and screened via the BGIS, we were next interested in whether enzymatic activities of proteins exhibit effects on bacterial growth. Post-translational modification of proteins, which allows cells to react quickly to a changing environment, is a hallmark of signal transduction. Therefore, we sought out to examine the bacterial growth inhibition potential of a kinase (CK2), protein arginine methyltransferases (PRMT 1 and 3), and the mono-ADP-ribosyltransferase ARTD10 using the bacterial growth inhibition assay.
CK2 is a widespread serine/threonine kinase expressed in all eukaryotic organisms and is comprised of catalytic subunits (α or α’) and regulatory subunits (β). It phosphorylates more than 300 proteins that participate in various cellular processes (Meggio and Pinna, 2003; Salvi et al., 2009), and it functions either as free catalytic subunit or as a holoenzyme with two CK2 kinase units and two regulatory CK2 units (Ortega et al., 2015; Ortega et al., 2014).
Surprisingly, in our test both catalytic subunits of CK2 exhibited bacterial growth inhibition, as no bacterial colony appeared after induction of expression (Figure 4.39). This strongly indicates
4 Results 108 that the BGIS may be also applied to kinases to identify their catalytic functional domains.
Figure 4.39: Bacterial inhibition assay with CK2 enzyme subunits. The pGEX vector alone (control) and vectors carrying CK2 subunits CK2 or CK2’ were transformed into E. coli BL21 and examined by the BGIS.
Arginine methylation is an important post-translational modification involved in numerous cellular functions, including DNA repair, transcription, translation, chromatin remodeling, signaling pathways, RNA processing and mRNA splicing. According to methylation symmetry, the PRMT enzymes are classified into 2 group types. Here two members in group I,
PRMTs 1 and 3, which have a common catalytic methyltransferase domain (Figure 4.40 A) were chosen to be examined by the bacterial inhibition screen (Figure 4.40 B). Both PRMT 1 and PRMT3 exhibited bacterial growth inhibitory effects, however, this inhibition was not as dramatic as seen with either DEK or the other chromatin architectural factors, as a few bacterial colonies still survived after protein expression.
Figure 4.40: Bacterial inhibition assay with PRMT1 and PRMT3. (A) Schematic depiction of the structure of PRMT 1 and 3. (B) GST (control), GST-tagged PRMT1 and PRMT3 were examined by BGIS.
ADP-ribosylation is a post-translational modification performed by enzymes that add
ADP-ribose to a substrate protein by using the cofactor NAD+. The mono-ADP-ribosyltransferase ARTD10 (formerly PARP10), which was originally identified as
4 Results 109 a MYC-interacting protein (Yu et al., 2005), contains an RNA recognition motif (RRM) on the
N-terminus and a PARP domain on the C-terminus that is able to transfer one ADP-ribose moiety to itself, to core histones, and to other proteins (Figure 4.41 A). Recent developments hint at the involvement of ARTD10 mono-ADP-ribosylation in transcriptional regulation,
DNA repair, insulin secretion, and immunity. Similar to the RNA recognition by ALY, the
ARTD10 truncation 1-907 containing this domain did not show any bacterial growth inhibition
(Figure 4.41 B). Contrarily, the expression of C-terminal truncations 588-1021 and 700-1021 containing the PARP domain, suppressed bacterial colony formation. Remarkably, the expression of the inactive mutant G888W, which is devoid of any catalytic activity failed to inhibit bacterial growth. These results suggest that expressing enzymatic activities in bacteria can also induce bacterial inhibition, most likely by interfering with bacterial signaling cascades or by introduction of modifications not present in bacteria.
Figure 4.41: Bacterial inhibition assay with ARTD10 truncations. (A) The schematic structure of ARTD10. It consists of an RNA recognition motif (RRM), a glycine-rich region (GRD) and three ubiquitin-interaction motifs (UIM), and its catalytic mono-ADP-ribosyltransferase (mART) domain, modified from (Till et al., 2008). (B) Indicated ARTD10 truncations (1-907, 588-1021 and 700-1021) and the inactive ARTD10 truncation 818-1025 mutant G888W were examined by BGIS.
In summary, using bacterial growth as a read-out for studies could be applied to a variety of protein classes for initial pilot studies as well as identification of DNA- and RNA-folding and enzymatic activities. As it appears that the expression of enzyme-dead mutants of enzymes does not inhibit bacterial growth, as exemplarily shown for ARTD10, this suggests that bacteria might also be used for drug-screening or drug development purposes.
4 Results 110
Taken together, the bacterial inhibition screen allows for the identification of potential DNA- and RNA-functional domains within proteins, especially factors that may induce alterations to chromatin structure. Furthermore, combined with random mutagenesis, this approach can also be employed as a screening tool for the creation of “loss-of-function” mutants of proteins.
More generally, this technique appears to be capable of analyzing the functional catalytic domains of enzymes and could be a useful tool for creating inactive mutants and selecting inhibitors. The efficiency and the potential wide applicability may make this technique a novel, useful tool in the repertoire of analyzing protein functions. Additionally, the BGIS could possibly be employed for screening medical drugs targeting the expressed activities in bacteria.
5 Discussion 111
5. Discussion
A substantial and constantly increasing number of studies have reported that DEK
(over)expression is related to the development and progression of certain auto-immune disorders and a large number of genetically highly diverse and aggressive tumor entities
(Mor-Vaknin et al., 2011; Piao et al., 2014; Privette Vinnedge et al., 2015; Riveiro-Falkenbach and Soengas, 2010). These findings, along with more functional studies, showing that DEK fulfills the criteria typically seen in a bona fide oncogene, have led to DEK being discussed as a novel cancer biomarker and as a potentially attractive target for therapeutic tumor treatment regimens (Datta et al., 2011; Khodadoust et al., 2009; Liu et al., 2012; Martinez-Useros et al.,
2014). This notion is furthermore underscored by an increased interest by industrial companies in identifying inhibitors of DEK functions, and in particular of its DNA binding and folding activities
(http://www.venenumbiodesign.com/wp-content/themes/venenumbiodesignV3/posters/DEK%
20Poster%20for%20SLAS.pdf). However, despite the rather large number of available studies, detailed knowledge about the molecular mode(s) of action of its multifaceted activities, and in particular of its DNA binding activities in cells, is still limited. This in turn had hampered the rational development of strategies to target this abundant and unique oncogene for tumor therapy purposes.
Based on its distinct DNA- and chromatin-binding properties identified in vitro (Bohm et al.,
2005; Kappes et al., 2004b; Waldmann et al., 2003; Waldmann et al., 2002), which led to the classification of DEK as a unique chromatin architectural protein, DEK appears to be able to shape cellular chromatin structure. As local and global chromatin organization affects virtually all DNA-dependent processes, it is crucial to understand the contribution of DEK’s activities in cells and the potential (dis)regulation of these activities in tumor cells in this context. This would not only enrich our knowledge of general chromatin biology but could as well aid in detailing DEK’s contribution to tumorigenesis.
5 Discussion 112
In support of its classification as a chromatin architectural protein, DEK has been shown to affect (hetero)chromatin integrity through a number of protein-protein-mediated interactions
(Kappes et al., 2011b; Waldmann et al., 2002), which most likely are regulated and integrated by the staggering number of post-translational modifications of DEK. On the other hand, new data in the present work show that DEK can modulate nucleosomal accessibility (Figure 4.2), suggesting that directly transmitted effects of its DNA binding and folding activities may play an additional or complementary role in the regulation of chromatin structure. These activities are transmitted by the unique pseudo-SAP/SAP-box domain of DEK (see chapter 1.4). The main goal of this body of work was to better understand the mechanism of DEKs DNA binding and folding activities and to investigate how these influence cellular chromatin structure.
Our initial hypothesis-driven approach based on the creation of single point- or limited combination-mutations in the pseudo-SAP/SAP-box domain of DEK did not result in the identification of DNA-binding-dead mutants (Figure 4.4, Figure 4.6, Figure 4.8, Figure 4.9 and
Figure 4.10). Therefore we developed a new screening procedure to circumvent these limitations (see chapter 4.4). Using interference with biological circuits in bacteria allowed us to obtain mutants that had either lost DNA binding or showed reduced DNA-binding activity
(Figure 4.22). Additionally, the identified mutational landscapes in these mutants may further aid in deciphering the molecular mechanism of DEK’s DNA-related functions. Further downstream studies with the DNA-binding-dead mutant allowed us to gain deeper insights into
DEK’s DNA binding biology. Together with newly created DEK knockout cells and a newly developed, inducible, lentiviral-delivered expression system (Figure 4.25 and Figure 4.26), we created new tools that have the potential to be highly valuable in future, more-detailed studies of
DEKs function in chromatin structure and gene regulation.
5.1 DNA binding and folding activities of DEK and their functions
in nucleosomal DNA accessibility
As mentioned above, DEK has been classified as an architectural protein that may regulate chromatin organization via its DNA-, chromatin- and/or histone-binding affinities, as well as by
5 Discussion 113 its DNA folding activities. DEK transmits its functions through known protein-protein-mediated interactions with chromatin remodeling factors (Cavellan et al.,
2006) or HP1(Kappes et al., 2011b), and these interactions indirectly affect
(hetero)chromatin integrity. Furthermore, DEK has been shown to be involved in interactions with core histones such as H2A, H2B and H4, and their variants (e.g. H3.3), and thereby may modulate chromatin organization (Alexiadis et al., 2000; Ivanauskiene et al., 2014). However, as the three DNA binding modules in DEK cover more than 50% of its structure, it is possible that DEK’s conserved DNA binding activities are additionally important for the regulation of cellular functions. This aspect of DEK biology has been neither studied in detail nor looked at in a cellular context before. We have been able to show in the present work that DEK is indeed involved in directly regulating the accessibility of chromatin in vitro and in cells (Figure 4.2,
Figure 4.24 and Figure 4.28). These effects appear to be solely dependent on the interaction with DNA, as a direct interaction of unmodified GST-DEK1-187 with histones was not observed under the experimental conditions used here (Figure 4.24). Thus, regulation of nucleosomal DNA accessibility by unmodified DEK appears to be histone-independent and is most likely mediated by its DNA binding and folding activities. However, the contribution of the C-terminal DNA-binding domain of DEK to either its overall DNA-binding activity or its role in histone interaction was not studied here. Additionally, DEK is known to undergo numerous post-translational modifications in cells (Kappes et al., 2004a; Kappes et al., 2008;
Tabbert et al., 2006), which may regulate or integrate spatial or temporal interactions with specific histones. Together, DNA- and/or protein-mediated interactions of DEK may modulate the access of other DNA repair-, transcription-, or replication-related proteins to nucleosomal
DNA, leading to further influence on subsequent cellular processes.
5.1.1 Creation of “DNA-binding-dead” DEK mutants
Considering the importance of DNA binding and folding activities to chromatin organization, it is important to understand the mechanisms of these activities. This is not uncomplicated, as the interactions of most chromatin architectural factors with chromatin and/or DNA are rather complex and somewhat diffuse in nature, making the identification of specific effects rather
5 Discussion 114 difficult.
Many in vitro studies have observed that the DNA binding and folding activities of DEK are transmitted by its unique and major DNA-binding module consisting of the pseudo-SAP and
SAP domain (Bohm et al., 2005; Kappes et al., 2004b). Though not fully understood, the functions of SAP domains of other proteins have been studied before, and are somewhat comparable to the activity of the SAP-box in DEK. Interestingly, mutation of the highly conserved invariant glycine in the SAP-box, which usually creates DNA-binding-dead SAP mutants in other proteins (e.g. for SAF-A) (Schwander, 2004), did not abolish the DNA binding in DEK (data not shown). Furthermore, the contribution of the unique pseudo-SAP domain, which is located N-terminally of the canonical SAP-box, to DNA binding is not understood (Bohm et al., 2005). As revealed by NMR structure, both domains share a striking structural similarity to each other (Devany et al., 2008). Therefore, the DNA binding, and in particular the DNA folding activities are most likely transmitted by cooperative efforts of this unique pseudo-SAP/SAP-box domain. Although a series of mutations were created in both domains in this work and in the work of Devany et al., it appears that single or double substitutions in either domain individually are not capable of abolishing the DNA binding and folding activities of DEK, and that combinatorial mutations of multiple amino acids concomitantly in both domains are needed. However, given the rather large number of mutants created so far, creating additional combinatorial mutational patterns was necessary to identify relevant mutants lacking DNA binding and folding activities. As this task would most likely be very time-consuming, we established a screening procedure that allowed for unbiased, undirected identification of DEK mutants. This was made possible due to the rather surprising finding that DEK’s DNA folding activities lead to bacterial growth inhibition (see chapter 4.3).
Based on this observation we were able to establish a screening procedure that yielded the desired DNA-folding-dead DEK mutant from a mutant library created by random mutagenesis
(Figure 4.17) (the general applicability and limitations of this assay are discussed in chapter
5.4). Interestingly, no single substitution mutants were selected by this screen, which indicates that the DNA binding and folding activities are much more complex in nature and suggests
5 Discussion 115 that multiple amino acids are involved in this process (Figure 4.19 B). Importantly, the
“DNA-binding-dead” DEK mutant #11 was selected via this screen and is discussed in detail below.
5.1.2 Molecular analysis of the “DNA-binding-dead” DEK mutant #11
Sequence analysis of the mutant #11 revealed eight distinct amino acid substitutions (Figure
4.20 and Figure 5.1), which initially appeared to be a surprisingly high number of mutations.
However, reverting back each of the identified mutations individually clearly showed that every single mutated amino acid contributes to the overall DNA-binding activity of DEK
(Figure 4.23). This furthermore substantiated that this newly developed screen does indeed allow for the selection of biologically relevant mutations.
Interestingly, four lysines, either present in the pseudo-SAP or SAP-box, were found substituted to alternative amino acids. Although all of these particular lysines were mutated to alanine individually in previous approaches (see Figure 4.4 and Table 4), they were found simultaneously mutated using the bacterial screen. As revealed by the NMR structure of DEK, these lysines are present on the surface of the protein. Due to its positive charge, lysine is generally considered a universal amino acid candidate for the interaction with negatively charged DNA. Indeed, through the analysis of 129 protein-DNA complexes, lysine was the most common amino acid found engaged in hydrogen bonding with the phosphate backbone of
DNA (Luscombe et al., 2001; Luscombe and Thornton, 2002). Interestingly, all lysine substitutions found in the DEK mutant #11 weaken this hydrogen bond-mediated
DNA-interaction either through alteration of the charge (K137E and K150E) replacement of hydrogen bonding with van der Waals contacts (K111I), or potential complete loss of contact
(K101M). Besides lysine substitutions, mutations in histidine, serine, leucine and asparagine were also found in the DEK mutant #11, again dispersed over both SAP-related domains.
Unexpectedly, histidine 95 and serine 139 were found replaced by arginines (H95R, S139R), which are positively charged and in principle would suggest an increased DNA interaction, though this was not the case. One possible explanation of this could be the structural function of histidine and serine. Pseudo-SAP and SAP domain generate a unique topology of two
5 Discussion 116 helix-extended loop-helix (HEH) subdomains (Aravind and Koonin, 2000). The first subdomain contains the N-terminal pseudo-SAP helix 1-2-loop-3 and the second subdomain includes the C-terminal SAP domain 4-loop-5 (Figure 5.1), and a long linker loop is attached between these two subdomains (Devany et al., 2008). Histidine 95 located in the -helix of the pseudo-SAP domain might be involved in stabilization of the -helix due to its inflexibility. Serine 139 is located right next to the -helix of the SAP domain, and therefore could probably be regulated by phosphorylation and may have an influence on helix remodeling and DNA binding (Kirchler et al., 2010). Therefore, arginine replacements of these two residues might partially weaken the protein secondary structure, thereby resulting in reduction or abolishment of DNA interaction. Similarly, the other two mutations (L166S and
N188D) located on the loop domain and -helix 5 in the SAP domain, respectively, could also lead to a failure to rotate the helix or the angle between the two helices. In conclusion,
DEK-DNA interaction is most likely generated by hydrogen bonding (between the protein side chains and DNA’s phosphate backbone) which may be supported by the stabilization of the specific protein domain structure (two HEH motifs). To investigate this, CD/NMR structure of
DEK mutant #11 needs to be determined further, which was not pursued here due to time limitations.
Figure 5.1: Depiction of the identified mutations in the DEK mutant #11 in DEK-DNA docking model. Left: DEK-DNA docking model. Right: All the mutations identified in the DEK mutant #11 were labeled in the DEK-DNA docking model (left) with sticks and spheres (right panel).
5 Discussion 117
5.1.3 Analysis of nucleosomal DNA accessibility with DEK mutant #11
5.1.3.1 Nucleosomal DNA accessibility
Accessibility to nucleosomal DNA is generally considered as a common regulatory mechanism to regulate the access of factors to the template DNA.
So far, several concepts for the regulation of nucleosomal accessibility have been proposed.
The histone code and chromatin remodelers involved in the regulation of the plasticity of higher-order chromatin structure are well established (Bartke and Kouzarides, 2011; Strahl and
Allis, 2000). The combinatorial patterns of histone post-translational modifications along with multiple chromatin remodelers can move nucleosomes along the DNA or make DNA more accessible by eviction of core histones. Another model termed the “chromatosome-like particle
(CLiP)” has been proposed. In this model, proteins being bound at the entry/exit site of the nucleosomal DNA are termed “linker proteins” due to their similar function to linker histones
(Zlatanova et al., 2008). These linker proteins making contact with either the entry or the exit site, or both, can leave the nucleosomal gate “ajar” or lock the nucleosomal gate, whereas linker histones are believed to close the nucleosomal gate. Consequently this results in allowance or inhibition of access to nucleosomal DNA.
5.1.3.2 DEK mutant #11 failed to affect nucleosomal DNA accessibility
Compared to other linker proteins, DEK shares resemblances with HMGB proteins, a class of well-characterized linker proteins, despite having no sequence similarity. They both exhibit
DNA binding affinity and a preference for binding to distorted DNA. Moreover, they can introduce positive supercoils into DNA and bend DNA. In addition, they can both directly interact with nucleosomal DNA. All these similarities suggest that DEK might function, at least partly, as a “linker protein” sitting at the entry/exit site of nucleosomal DNA and thus may regulate chromatin structure similar to HMGB proteins.
The experiments presented in this study show that DEK can change the nucleosomal DNA accessibility in vitro and DEK knockdown in cells leads to a reduction in nucleosomal DNA repeat length (Figure 4.2). Interestingly, a DEK isoform in Arabidopsis thaliana, called DEK3,
5 Discussion 118 that shares two consensus sequences with human DEK (SAP domain and C-terminal
DNA-binding domain), was recently identified as a chromatin-associated protein capable of regulating nucleosomal occupancy and gene expression (Waidmann et al., 2014). This suggests a possible evolutionarily conserved function for the SAP domain, and maybe also the pseudo-SAP-box of DEK in the regulation of nucleosomal DNA accessibility.
In the present work, eight amino acids present in pseudo-SAP/SAP-box were selected via the bacterial growth inhibition screen and identified as mediating DNA binding and folding activities. In vitro, the “DNA-binding-dead” DEK mutant (#11) failed to change nucleosomal
DNA accessibility and nucleosomal repeat length (Figure 4.24), which confirms the essential role of the pseudo-SAP/SAP-box domain in DNA binding and folding activities. To further investigate the biological relevance of the DNA binding and folding activities of DEK, DEK mutant (#11) was analyzed in DEK knockout cells. Surprisingly, a slow cell proliferation (data not shown) and strikingly altered cell morphology (cell shape, nuclear size and nucleus distribution) was observed in DEK knockout cells (Figure 4.25 B). Therefore, we speculated whether the DNA folding activities of DEK were able to rescue this phenotype. Expression of
DEKwt for only a short time led to a substantial reduction in nuclear size, and therefore was able to restore the DEK-deficient phenotype (Figure 4.28 B). Interestingly, re-expression of the N-terminal truncation DEK1-187 showed a similar activity to full-length DEK (Figure
4.28 B), suggesting that DEK1-187 is capable of organizing nuclear compaction. Meanwhile, these results also indicate that the C-terminal DNA-binding domain may not have critical functions in chromatin structure alterations. However, re-expression of the DEK mutant (#11) in DEK knockout cells failed to decrease the nuclear size in cells (Figure 4.28 B). This indicates that the chromatin-associated function of DEK is based on its DNA folding activities that are mediated by the pseudo-SAP/SAP-box domain. Further work is needed to substantiate this finding, however, as this particular mutant also showed a different subnuclear localization compared to full-length DEK (Figure 4.27). These results clearly indicate that DNA binding is crucial for the sub-nuclear localization of DEK and may be important in guiding DEK to its regions of action. This may further imply that post-translational modification of DEK, e.g.
5 Discussion 119 phosphorylation or poly(ADP-ribosyl)ation, which both strongly weaken the DNA binding and folding activities of DEK (Fahrer et al., 2010; Kappes et al., 2004a; Kappes et al., 2008), could mimic the phenotype of the “loss-of-DNA-binding” DEK mutant in the cellular environment. Thus, DNA binding itself and regulation thereof may play a critical role in regulating either activating or inactivating transcriptional effects of DEK, as well as distribution of DEK to either eu- or hetrochromatin regions.
In summary, the DNA binding and folding activities of DEK mediated by pseudo-SAP/SAP-box domain can control the distribution of DEK within chromatin, and indeed play a role in global chromatin organization. Based on its DNA binding and folding activities, DEK may modulate local and global chromatin structure which facilitates or inhibits the recruitment of other factors to nucleosomal DNA and influences subsequent cellular processes.
5.1.4 Other mutants selected via BGIS
Besides mutant #11, two other mutants (#8 and #18) that exhibited strongly reduced, yet definite DNA-binding activity were selected by the BGIS (Figure 4.22). In comparison to mutant #11, the identified mutations in these mutants were only present in the SAP domain
(Figure 4.22 C). As we also identified mutants with only mutations in the pseudo-SAP domain via the BGIS (data not shown), this strongly indicates that cooperative effects of both domains are necessary for the typical DNA folding activities of DEK. As DEK was recently identified as having roles in replication fork progression under conditions of DNA replication stress
(Deutzmann et al., 2014; Kavanaugh et al., 2011), it is tempting to speculate that the
DNA-folding activity may have important functions in this fundamental process. In particular fork reversal is observed under replication stress and DEK may aid in resolving these issues by its supercoiling activity (Deutzmann et al., 2014). Even though further studies are needed to study this, the mutants #8 and #18 may represent suitable tools to investigate this hypothesis. As both mutants show residual DNA-binding activity via the pseudo-SAP domain and thus should localize properly to chromatin, but lack any DNA supercoiling activity (data not shown), the role of DEK’s DNA-folding activities in this scenario could be addressed with
5 Discussion 120 these tools.
Surprisingly, several mutations present in DEK1-87 were also selected by the bacterial growth inhibition screen (Figure 4.19 A and Table 7), though they have not been further tested in
EMSAs. As no DNA binding affinity is associated with this domain, selection of these mutations may imply that this rather flexible domain may be important for proper protein folding, thereby regulating the DEK-DNA interaction via changes to DEK’s secondary structure. This domain is targeted by ample post-translational modifications, which presents the possibility for phospho-mimic mutants to be used to assess their role in the DNA-binding activity via the BGIS. Moreover, one could imagine a more systematic assessment of structural features of other proteins using this bacterial screen.
5.2 Identification of the RNA-binding domain of DEK
5.2.1 RNA binding proteins (RBPs)
The dynamic processes of a living cell are orchestrated by coordinated temporal, spatial, and stoichiometric regulation of gene expression. Even though transcriptional control represents a major regulatory hub for gene expression, subsequent regulatory circuits involving cellular
RNA-dependent processes like splicing, transport, localization, translation or RNA decay play vital roles. All these processes are regulated by the interplay of RNAs with RNA binding proteins (RBPs), mostly in form of ribonucleoprotein complexes (RNPs). Therefore, interactions of RBPs with RNA affect the post-transcriptional fate of transcripts in cells, and defects in the expression or functions of RBPs may be related to numerous diseases.
For many decades, intensive efforts have been undertaken to identify and understand RBPs and their specific interactions with RNA. In vivo ultraviolet (UV) cross-linking has been used to introduce covalent bonds between RNA and RBPs (Brimacombe et al., 1988). Based on this, conventional UV-cross linking (cCL) and related modified methods entailing photoactivatable-ribonucleoside-enhanced cross-linking (PAR-CLIP) have been employed to identify RNA-interacting proteins and their specific RNA targets more broadly (Hafner et al.,
2010). Recently, several hybrid methods combining these two UV cross-linking methods (cCL
5 Discussion 121 and PAR-CL) have been created. For instance, poly(A)-mRNA can be captured by using hybridization with oligo(dT) oligonucleotides, with the associated proteins subsequently being analyzed by mass spectrometry in an unbiased fashion. Immunoprecipitation of known
RNA-interacting proteins can also be introduced to identify RNA binding activities and RNA targets by analysis of associated RNAs using q-PCR, RNA-seq or microarrays (Castello et al.,
2012; Castello et al., 2013; Strein et al., 2014). In vitro, Systematic Evolution of Ligands by
Exponential enrichment (SELEX) and Electrophoretic Mobility Shift Assay (EMSA) are employed to systematically identify RNA-protein interactions. In silico, several computational programs (BindN, RBPmap, RBPDB) have been developed to predict and identify the RBPs and its RNA targets (Cook et al., 2011; Paz et al., 2014; Wang and Brown, 2006). With these techniques, a substantial number of additional proteins that bind to RNA have been identified.
5.2.2 DEK187-270 is the RNA-binding domain
For quite some time, DEK has been reported to be involved in RNA-dependent processes, suggesting a role for DEK in RNA biology. More recently, and as revealed by an “interactome capture” approach, DEK was clearly identified to have mRNA-binding affinity (Castello et al.,
2012). A functional relevance of DEK in the interaction with both SR (serine/arginine-rich) proteins and the exon-exon junction complex (EJC) was identified, and DEK was found to be required for proofreading 3’ splice sites by interaction with the splicing factor U2AF35 (Kress and Guthrie, 2006; Mendes Soares and Valcarcel, 2006). Additionally, evidence in our laboratory suggests an involvement of DEK in alternative splicing and mRNA transport (Malte
Prell, personal communication). However, no RNA-binding domain in DEK had been identified prior to the present work. In order to closely understand DEK’s role in this RNA biology, knowledge about the nature and localization of its RNA-binding domain is crucial.
Initially, multiple in silico methods were performed in this thesis to identify a potential
RNA-binding domain in DEK. As DEK is the only representative in its family, only its different isoforms were blasted using PSI-blast. According to several databases (RBPDB and
RBPmap) that collect most known RNA-binding proteins, no RNA-binding activity was predicted for DEK (data not shown). However, upon examination by the bacterial growth
5 Discussion 122 inhibition approach, the DEK truncation 187-270 showed unexpected effects in bacterial growth inhibition (Figure 4.29 A). Interestingly, when only using this domain and subjecting it to the BindN program (Wang and Brown, 2006) a potential RNA-binding was predicted. As shown in Figure 5.2, the highest putative RNA-binding score of 80% specificity was obtained in this domain. Three other potential RNA binding domains were additionally predicted (data not shown). However, after examining this domain with other RNA-binding domain prediction programs, no clear RNA-binding was predicted for DEK (data not shown).
Figure 5.2: Overview of the potential RNA-binding residues in DEK187-270 predicted by BindN program. Potential RNA-interacting residues are labeled with “+” and in pink, the non-RNA-binding residues are labeled with “-” and in green. The numbers represent the confidence level from 0 (lowest) to 9 (highest).
By subjecting this domain to several experimental approaches, we could indeed confirm that this domain is the RNA-interacting domain in DEK. Confocal microscopy analysis revealed that DEK truncations containing amino acids 187-270 show strong co-localization with endogenous protein-coding mRNAs (Figure 4.30). Northwestern and RNA-EMSAs experiments further confirmed the RNA-binding activity of this domain (Figure 4.31 and
Figure 4.32). However, interaction of DEK with RNAs appears not to be limited to protein coding RNAs, as immunoprecipitation of DEK and analysis of small non-coding RNAs revealed that DEK may interact with a number of microRNAs (Ferdinand Kappes, personal communication). Thus, using the BGIS allowed us to identify the long sought-after
RNA-binding domain in DEK. As performed with the pseudo-SAP/SAP-box domain of DEK, the bacterial growth inhibition screen could be further employed to screen for “RNA-binding dead” mutants, which would provide a means to obtain better and deeper understanding of the
5 Discussion 123 role of DEK in RNA-associated processes, including the intriguing potential interaction with microRNAs.
Therefore, through its RNA-binding affinities of either mRNAs or non-coding RNAs, DEK has possible functions in post-transcriptional processes, especially in mRNA splicing, which could affect the fate of RNAs in subsequent steps including export, location and translation. In addition, RNA-associated functions of DEK in concert with its DNA-associated activities could further affect chromatin organization.
5.3 The functions of DEK within chromatin
As the present work has uncovered new insights into the biology of DEK, we present here an updated scheme to collectively depict DNA-, RNA- and protein-associated activities of DEK within chromatin (Figure 5.3).
Even though DEK is the only representative of its protein class, here we compare its functions to other, better-understood chromatin architectural factors. Based on DNA-, RNA- and protein-associated activities of DEK, several proteins (SAF-A, HMGB1 and HP1 were chosen and their functions compared to DEK. This comparison may aid in revealing common and distinct roles for DEK in the regulation of chromatin organization, and proposes a possible updated framework model for DEK’s role in the regulation of chromatin structure (Table 8 and
Figure 5.4).
Figure 5.3: Summary of functional domains in DEK with their identified functions.
5 Discussion 124
SAF-A is able to bind both DNA and RNA via its N-terminal SAP-box and its C-terminal RGG box (Fackelmayer et al., 1994; Kipp et al., 2000). As a SAP-box-containing protein, it has roles in the regulation of chromatin structure and gene transcription (Gohring and Fackelmayer, 1997;
Romig et al., 1992). Additionally, it associates with pre-mRNAs via its RGG domain and influences pre-mRNA processing and other aspects of mRNA metabolism and transport
(Fackelmayer et al., 1994; Kiledjian and Dreyfuss, 1992). Contrarily, the SAP-box of DEK is located centrally within the molecule and an extra DNA-binding domain (pseudo-SAP-box) allows DEK to exhibit other activities such as DNA-bending, -looping and -folding.
Interestingly, without any other consensus sequence with SAF-A, similar growth inhibitory effects of the RNA and DNA binding and folding activities were identified in the bacterial growth inhibition screen for both proteins.
Despite lacking sequence similarity, HMGB1 shares surprisingly similar DNA binding and folding activities to DEK, especially the preference for binding to distorted DNA, alteration of
DNA supercoiling and DNA bending. Also, heterologous expression of HMGB1 in bacteria resulted in bacterial growth inhibition and compaction of bacterial DNA. Both HMGB1 and
DEK exhibit chromatin-binding affinity with consequent alterations to nucleosomal DNA accessibility. However, unlike DEK, which shows RNA-binding affinity, only a few
RNA-related activities of HMGB1 have been found and only branched RNA structures have been found to bind to it (Bell et al., 2008). Therefore, DEK functions might resemble HMGB1 functions in terms of roles in chromatin organization due to the similar DNA-associated activities, however, DEK may have additional functions based on its RNA-binding and potential RNA-folding activities.
As a regulator of chromatin structure, HP1 possesses both DNA- and RNA-binding activities via its hinge domain, which links two functional distinct globular domains: N-terminal chromo domain (CD) and the C-terminal chromoshadow domain (CSD) (Figure 1.6). Interestingly, neither of these domains showed any growth inhibitory effect in the bacterial growth inhibition screen (data not shown). The CD domain functions as a reader module recognizing H3K9Me3, although the affinity is rather weak (Jacobs and Khorasanizadeh, 2002; Nielsen et al., 2002).
5 Discussion 125
This interaction is stabilized by the CSD domain, which induces protein dimerization to provide an interaction surface for many other chromatin proteins that are collectively involved in the stabilization of heterochromatin structure (Azzaz et al., 2014; Nishibuchi and Nakayama,
2014). Besides the aforementioned CSD domain, the hinge domain also enhances this interaction: its RNA-binding activities may guide HP1 to H3K9Me and the DNA binding activities act to stabilize this interaction (Meehan et al., 2003; Muchardt et al., 2002). Therefore, both the DNA- and RNA-associated activities are involved in the enhancement of its affinity to nucleosomes, thereby regulating the chromatin accessibility.
In conclusion, through comparing DEK’s functions with other proteins (Table 8), similar yet distinctively different modes of actions have been identified. For instance, although DEK’s SAP domain sequence shows homology to the one in SAF-A, its DNA-associated activities are more consistent with the ones seen in HMGB1. On the other hand, its RNA-associated activities display more similarities to SAF-A or HP1.
Table 8: Comparison of protein functions between DEK, SAF-A, HMGB1 and HP1 Functions DEK SAF-A HMGB1 HP1 Yes, via pseudo-SAP/SAP domain Yes, via SAP domain, Yes, via HMG boxes, Yes, via hinge domain, and C-termial DNA-binding domain, DNA-binding without DNA preference for without DNA preference for distorted structure-specificity distorted DNA structure-specificity DNA(pseudo-SAP/SAP) Yes, via HMG boxes, Yes, via pseudo-SAP/SAP domain, Yes, via SAP domain, DNA-folding DNA supercoiling No DNA supercoiling and bending DNA looping and bending RNA-binding Yes, via DEK187-270 Yes, via RGG domain No Yes, via hinge domain RNA-folding Yes, via DEK187-270 Yes, via RGG domain No No Yes, via its DNA regulation of binding and folding Yes, via CD and hinge nucleosomal Yes, keep the nucleosomal gate -- activities, leave the domain, “lock” the DNA inaccessible nucleosomal gate nucleosomal gate accessibility “ajar”
Currently, two models have been proposed to explain DEK’s role within chromatin. Through the protein-protein-mediated interaction of DEK with HP1, DEK indirectly regulates chromatin organization (Kappes et al., 2011b). In this study, we found that DEK can also directly interact with nucleosomal DNA and regulate its accessibility based on its DNA binding and folding activities, which are similar to HMGB1.
Considering that the RGG domain in SAF-A and DEK187-270 both have RNA-binding activities that show inhibition in the bacterial growth screen, one might speculate that these
5 Discussion 126
RNA-associated activities are also indirectly involved in shaping chromatin structure. As discovered here from studying SAF-A and HP1, their RNA-interactions translate into
DNA-related processes such as genome expression and chromatin organization. In other words,
RNA-interacting modules may not only be important in recognizing and transporting an underlying RNA structure, but may also be an important part in guiding DNA-dependent activities to their final position in the genome, thus may also influence DNA-dependent processes and chromatin stability (Naro et al., 2015). Given that DEK can also interact with non-coding RNAs, RNA could play a role in guiding DEK to chromatin.
Figure 5.4: Current models for DEK functions within chromatin. (A) DEK can regulate chromatin organization through indirect interactions (Kappes et al., 2011b). (B) DEK directly interacts with chromatin via DNA interactions and affects chromatin structure. (C) A crosstalk between these two directions may be regulated by the guidance of RNA-interactions.
Taken together, we have achieved a closer insight into DEK’s function within chromatin
(Figure 5.4), and our results add to the work that suggests that DEK is involved in the regulation of chromatin structure via three potential routes. These may entail direct interaction with nucleosomes, as revealed in this thesis (Figure 5.4 B), or indirect interaction (Figure 5.4
A) via protein-protein mediated interactions, or, though pure speculation, via RNA-mediated guidance (Figure 5.4 C). Through direct binding to HP1, DEK can stabilize the binding of
HP1 to H3K9Me3 (Figure 5.4 A). This indirect interaction of DEK then maintains heterochromatin integrity (Kappes et al., 2011b). In addition to indirect interaction, DEK can also directly bind to the entry/exist of nucleosomes via its pseudo-SAP and SAP domain and
5 Discussion 127 then modulate local and global nucleosomal DNA accessibility due to its DNA binding and folding activities (Figure 5.4 B). Both modes of modulation could further affect gene transcription, DNA repair and genomic stability. The C-terminal DNA-binding domain of
DEK seems not to be essential for chromatin organization. However, as seen in chromatin accessibility assay, the N-terminal DEK fragment 1-187 protected nucleosomal DNA from
MNase digestion, though not as potently as full-length DEK. Therefore, the C-terminal
DNA-binding domain may have roles in the regulation of the DNA-associated functions of the full-length molecule. Given that the RNA-binding domain was also identified by the BGIS, we now speculate that RNA-binding of DEK may be also involved in the regulation of chromatin organization. As proposed in Figure 5.4 C, the RNA-binding of DEK, especially to non-coding
RNA species, could indicate roles in guiding DEK to either HP1 or to nucleosomal DNA in particular genomic regions. Therefore, RNA-binding may provide a bridge between these two models (Figure 5.4 C).
5.4 The bacterial growth inhibition screen (BGIS) and its
applications
Proteins take center stage in performing a variety of functions in living cells, with their functions usually mediated by interactions with other molecules including DNA, RNA and other proteins. Therefore, screening procedures for functional domains in proteins are helpful in understanding the roles of proteins in multiple cellular processes. In silico, computational methods are used to predict potential functional domains based on homology-, sequence- and structure-similarity of proteins. In vitro and in vivo, a number of experimental methods are employed to characterize the binding targets of proteins. In addition, bacterial- and yeast-based systems, among others, are also used to identify protein-DNA, -RNA and -protein interactions (Allen et al., 1995; Joung et al., 2000; Phizicky and Fields, 1995; Vidal and
Legrain, 1999). In this body of work, a new bacteria-based system termed bacterial growth inhibition screen (BGIS) was established and validated. This allowed for the identification of functional domains (DNA/RNA binding/folding domains and catalytic domains) of proteins. In
5 Discussion 128 combination with random mutagenesis this screen furthermore allowed for the unbiased identification of “loss-of-function” mutants. Together, the BGIS may provide a useful tool to discover and functionally analyze protein domains.
5.4.1 DNA-/RNA- binding/folding domains in the BGIS
Driven by the pitfalls in designing DNA-binding-deficient DEK mutants, we explored the use of bacteria for this purpose. Bacteria are widely used for cloning and protein expression purposes, however, at least to our knowledge, their use in directly assisting in the characterization of protein function has not been explored yet. Based on our observation that expression of certain protein classes in bacteria results in a general growth attenuation, we hypothesized that interfering with bacterial biological circuits via conventional heterologous expression of genes of interest (GOIs) may provide a means to identify and characterize functional domains in proteins. Using such an undirected and unbiased approach may be particularly useful for the identification of domains in proteins with yet unknown function and for the investigation of activities that are not as thoroughly investigated or have proven difficult to explore via standard approaches.
We first studied the expression of GST-DEK in the bacterial system, which represented the
“proof-of-concept” study. Compared with yeast, baculoviral or other eukaryotic expression systems which are able to place post-translational modifications on proteins, E. coli produces unmodified proteins. Given the abundant number of post-translational modifications identified in DEK, which collectively inhibit or regulate its DNA binding, this approach allowed us to focus purely on its DNA binding affinities. We found that the expression of unmodified
GST-DEK strongly inhibited bacterial growth (see chapter 4.3). This was due to the massive compaction of the bacterial genome upon the expression of DEK (Figure 4.14, Figure 4.15 and
Figure 4.16). Using this system in combination with error-prone PCR allowed us to identify a series of mutants that showed no or reduced DNA-binding activity (Figure 4.22). To a first approximation, DNA-folding activities, but not DNA binding activities, can be studied using this bacterial assay. Only protein domains with DNA-supercoiling activity showed inhibition in growth upon heterologous expression. This was seen for DEK, HMGB1, and SAF-A
5 Discussion 129
(Figure 4.12 and Figure 4.33). Importantly, this assay also allowed us to identify previously unrecognized DNA-folding domains in the mRNA transport adapter ALY/Ref, which had not been associated with DNA binding and folding activities before (Figure 4.36).
In contrast, protein domains with only DNA binding affinities showed no inhibition of bacterial growth. ASH2L and HP1, which both have DNA binding affinity but no
DNA-folding activity, showed no growth inhibition (Figure 4.34). Furthermore, the BGIS selected for the mutants #8 and # 18 (and others), which both show remaining DNA-binding activity, yet lost their DNA-folding activity in vitro (data not shown). In support of this, the
C-terminal DNA-binding domain in DEK also showed no growth inhibition (Figure 4.29 A).
Therefore, the BGIS may be useful in the identification of activities that cause structural changes to DNA (DNA-folding) rather than just pure DNA binding activities.
Additionally, several mutations present in the far N-terminal DEK domain 1-87 were selected by bacterial growth inhibition screen (Figure 4.19 A and Table 7). Even though this domain has no DNA-binding activity, it appears that it may be important in the regulation of the folding of the pseudo-SAP/SAP-box domain of DEK. This further indicates that the BGIS may also be used to investigate the role of protein folding in the activity of given proteins. Additionally, regulation of DNA-folding via post-translational modifications could be quickly assessed via subjecting mimic mutants to the BGIS.
Besides DNA-folding activities, we identified that certain RNA-binding domains can be identified via the BGIS. First, we identified the previously unknown RNA-binding domain in
DEK (see chapter 4.8), which further underscores that the BGIS may be a useful tool for the identification of novel functional domains in proteins. Additionally, the RNA-binding domain of SAF-A as well showed bacterial growth inhibition (Figure 4.35), however this inhibition was observed for neither the RNA-binding domain of HP1 (data not shown) nor the RRM domain of ALY/REF (Figure 4.36), again suggesting that only certain RNA-binding activities can be assessed via the BGIS.
To further explore the applicability of the BGIS, a set of proteins with no known direct
5 Discussion 130 functions in DNA or RNA metabolism were tested, and this resulted in no sign of growth inhibition (Figure 4.38). The tested importin family members, as well as Ran, Grb2 and SH3 domains are believed to not have any functional relevance in bacteria. This suggests that perhaps only functional domains with at least some relation to bacterial biology can be screened using this assay, which also clearly indicates the limitations of this approach.
Taken together, the bacterial growth inhibition screen was capable of identifying DNA-folding and RNA-binding domains in not only DEK but in additional proteins. Furthermore, this screen combined with random mutagenesis was applied to create and successfully select
“loss-of-DNA-binding” DEK mutants, which are helpful for understanding the DNA binding and folding actives of DEK within chromatin more extensively. Similarly, this approach could be used in exploring the RNA-binding/-folding activities of DEK.
5.4.2 Enzymatic activities in the BGIS
We additionally identified that the BGIS may be useful for studying enzymatic activities, thus extending the capabilities of this screening procedure significantly. Several enzymatic activities, including a kinase (CK2), two protein arginine methyltransferases (PRMTs), and the mono-ADP-ribosetransferase (ARTD 10) also showed substantial bacterial growth inhibition
(Figure 4.39, Figure 4.40 and Figure 4.41). This further indicates that enzymatic activities could possibly be tested via the BGIS more broadly. Together with the finding that the enzyme-dead mutant of ARTD10 did not affect bacterial growth, this opens up possibilities for establishing simple, quick and economical ways for functional screening of enzymatic activities. Furthermore, one could envision the establishment of high-throughput approaches for the purpose of drug screening using large, compound libraries. This newly developed technique could provide a fast route to pre-select bioactive compounds and may aid in speeding up drug development procedures. Considering that the BGIS would be easy to implement in every molecular laboratory and is fast to perform, it could be used as a pre-screening tool for the identification of catalytic functional domains within enzymes and for the creation of catalytically dead mutants. In addition it could potentially be implemented in a high-throughput pre-screening approach for inhibitory bioactive compounds for enzymes.
5 Discussion 131
5.4.3 The BGIS—a useful novel tool for investigating protein functions
Though the human genome has been determined, the function of a large portion of the human proteome remains to be revealed. Despite many efforts having been made to identify potential functions of proteins, the functions of a multitude of factors are still barely understood, especially when they possess neither homologous sequences to other proteins nor known functional consensus domains.
In this dissertation, an easy-to-perform screening technique has been established based on bacterial growth inhibition. Unlike classic bacterial inhibition assays which use bacterial colony growth to monitor the abnormal presence of phenylalanine in blood (phenylektonuria
(PKU)) (Rohr et al., 1996), our screen can be used to directly investigate protein function upon heterologous expression in bacteria. In spite of some limitations, we believe that this screen can potentially analyze both DNA/RNA binding/folding activities and catalytic activities of proteins. This could possibly aid in the discovery of functional domains in proteins with yet unknown functions. By further examining the isolated functional domains after random mutagenesis, “loss-of-function” mutants may be also created in a nearly high-throughput fashion. Additionally, pre-screening approaches for bioactive compounds based on bacterial growth inhibition could be established. This technique requires no special equipment andcan be easily implemented in every molecular biology laboratory. Due to its easy implementation and its rather wide applicability, we believe that the BGIS can be a valuable addition to the method repertoire for the identification and functional analysis of protein domains.
6 Reference 132
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7 Appendix 148
7. Appendix
7.1 Abbreviations
°C degree Celsius aa amino acid ACF ATP-utilizing chromatin assembly and remodeling factor Amp ampicillin ARTD ADP-ribosyltransferase diphtheria toxin-like ASH2 absent, small or homeotic 2 ATP adenosine triphosphate -ME -mercaptoethanol bp base pair BSA Bovine serum albumin cDNA complementary DNA Ci curie C-terminal Carboxy-terminal DMEM Dulbecco's modified Eagle medium DMSO dimethylsulfoxid DNA Deoxyribonucleic acid Dnase Deoxyribonucleosidease dNTP Desoxynucleoside triphosphate Dox. Doxycyclline dsDNA Double stranded DNA DTT Dithiothreitol E. coli Escherichia coli EDTA Ethylene diamine tetraacetic acid EMSA Electrophoretic mobility shift assay FCS fetal calf serum g gram or relative centrifugal force GCN5 general control nonderepressible 5, acetyltransferase GFP green fluorescent protein GST Glutathione-S-transferase h hour H3K9Me histone H3 lysine 9 methylation HAT histone acetyltransferase HDAC hisone deacetylase HMG High mobility group HMT histone methyltransferase HP1 Heterochromatin protein 1
7 Appendix 149 HRP Horseradish peroxidase IF Immunofluorescence IgG Immunoglobulin G IPTG Isopropyl -D-1-thiogalactopyranoside K lysine kb kilo bases kDa Kilo daltons LB lysogeny broth micro(10-6) M Molar me1/me2/me3 mono-/di-/trimethylated min minute(s) MNase Micrococcal Nuclease MW Molecular weight NLS nuclear localization sequence NMR nuclear magnetic resonance NP-40 Nocidet P40 NRL Nucleosomal repeat length N-terminal Amino-terminal p300/CBP p300/CREB-binding protein PAGE Polyacylamide gelelectrophoresis PAR poly(ADP-ribose) PARP poly-ADP-ribose polymerase PBS Phosphate-buffered saline PCR Polymerase chain reaction PFA Paraformaldehyde PHD Plant homeo domain PMSF phenylmethylsulphonyl fluoride PP Pentaprobe PRMT protein arginine methyltransferase PTM post-translational modification qPCR quantitative PCR R arginine RAS rat sarcoma viral oncogene RNA Ribonucleic acid rpm rotation per minute RRM RNA-recognition motif RT Room temperature RT-qPCR Reverse transcriptase quantitative polymerase chain reaction s.d. standard deviation SDS Sodium Dodecyl Sulphate sec second SH3 Src-homology-3
7 Appendix 150 shRNA small hairpin RNA ssDNA single stranded DNA SU(VAR)3-9 Suppressor of position effect variegation 3-9 SV40 simian virus 40 TALEN transcription activator-like effector nuclease TBE TRIS/Borate/EDTA TBP TATA-binding protein Tween-20 Polyoxyethy-lene(20)-sorbitan monolaurate U unit UIM Ubiquitin-interaction motif UV Ultraviolet light V Volts v/v volume per volume proportion Vol Volume(s) w/v weight per volume proportion WB western blot wt wild type
7 Appendix 151
7.2 Supplementary figures
Supplementary Figure S1: Selection for DEK mutants lacking bacterial growth inhibition by replica stamping the colonies after transformation to LB-Amp plates plus IPTG. After transformation of mutants generated by error-prone PCR, bacteria were initially plated onto LB-Amp plates without IPTG (I). Subsequent rounds of replica stamping of the colonies to LB-Amp plates plus IPTG selected for mutants that lack bacterial growth inhibition (II-V). Surviving colonies were eventually collected on master plates (VI). Shown are three examples of multiple replicates
7 Appendix 152
Supplementary Figure S2: Immunoblotting with bacterial colonies transformed pGEX-DEK1-187 and selected from bacterial inhibition screen. Randomly-mutagenized pGEX-DEK1-187 plasmids were transformed into E. coli and scanned by the bacterial inhibition screen. After a few rounds thesurviving bacterial colonies were induced with IPTG (0.5 mM), lysed and analyzed by immunoblotting with GST-specific antibodies, and using GST and GST-DEK1-187 as positive controls.
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Supplementary Figure S3: Immunoblotting with bacterial colonies transformed pGEX-DEK85-187 and selected from bacterial inhibition screen. Randomly-mutagenized pGEX-DEK85-187 vectors were transformed into E. coli and scanned by bacterial inhibition screen. After a few rounds thesurviving bacterial colonies were induced with IPTG (0.5 mM), lysed and analyzed by immunoblotting with GST-specific antibodies, and using GST and GST-DEK85-187 as positive controls.
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Supplementary Figure S4: Binding of GST-DEKwt and DEK truncations to RNA Pentaprobe sequences. EMSAs showed indicated 32P-labeled RNA Pentaprobes (PP1-PP8) incubated with increasing amount (2, 4 and 6 pmol for PP1-PP4 and 0.5, 1, 2 pmol for PP5-PP8) of GST, GST-DEKwt, GST-DEK1-187, GST-DEK187-375 and GST-DEK187-270. All complexes were analyzed on polyacrylamide gels and exposed to an X-ray film (autoradiography).
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7.3 Manuscript of publications
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7.4 Curriculum vitae
Personal data
Name: Haihong Guo Nationality: Chinese Date of birth: 02.07.1986 Place of birth: Xiaoyi, Shanxi, China
Education
09/2011-present Graduate student, Institute for Biochemistry and Molecular Biology, RWTH Aachen University, Germany
09/2009-07/2011 Master degree, Institute for Biochemistry and Molecular Biology, School of Science, Beijing Jiaotong University, Beijing, China
09/2005-07/2009 Bachelor degree, Biotechnology, School of Agriculture, Shanxi Agricultural University, Shanxi, China
09/2001-07/2005 Xiaoyi high school, Xiaoyi, Shanxi, China
09/1998-07/2001 Xiaoyi Xincheng middle school, Xiaoyi, Shanxi, China
09/1993-07/1998 Xiaoyi Xincheng primary school, Xiaoyi, Shanxi, China
Participation in academic conferences
11/2014 Danube Scientific Conference on Epigenetics (2014), Budapest, Hungary
09/2014 30th Ernst Klenk Symposium in Molecular Medicine: DNA Damage Response and Repair Mechanisms in Aging and Disease, Cologne, Germany
09/2013 44th Annual Conference of the German Genetics Society (GfG): Genetics 2013, Brauschweig, Germany
03/2013 Third Clinical Epigenetics International Meeting, Solingen, Germany
09/2012 43rd Annual Conference of the German Genetics Society (GfG): Chromatin and Epigenetics, Essen, Germany
06/2012 PhD/Master course: Advanced Microscopy and Vital Imaging 2012, Maastricht, Netherlands
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7.5 Eidesstattliche Erklärung
Ich erkläre eidesstattlich, dass ich die vorliegende Dissertation selbständig verfasst und alle in
Anspruch genommenen Hilfen in der Dissertation angegeben habe. Des Weiteren erkläre ich, dass die vorliegende Dissertation nicht bereits als Diplomarbeit oder vergleichbare
Prüfungsarbeit verwendet worden ist.
Aachen, den
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(Haihong Guo)