University of Patras, Medicine Department Laboratory of Biological Chemistry

Elongation Factor P

and its Role in Environmental Stress Adaptation

Master Thesis Stavropoulou Maria

Patras, 2012

University of Patras, Medicine Department Laboratory of Biological Chemistry

Elongation Factor P

and its Role in Environmental Stress Adaptation

Master Thesis

Stavropoulou Maria

Patras, 2012 Περίληψη 1

Περίληψη

Ο βακτηριακός παράγοντας επιμήκυνσης EF-P, είναι μια διαλυτή πρωτεΐνη που βοηθά στο σχηματισμό του πρώτου πεπτιδικού δεσμού, αλληλεπιδρώντας με το ριβόσωμα και το εναρκτήριο tRNA. Η κρυσταλλική δομή του EF-P δείχνει ότι μιμείται στη μορφή το tRNA. Ορθόλογες πρωτεΐνες έχουν βρεθεί και στα αρχαία και τα ευκαρυωτικά κύτταρα, γνωστές ως aIF5A και eIF5A, αντίστοιχα. Ο eIF5A, για τον οποίο αποδείχθηκε πρόσφατα ότι συμμετέχει και στο στάδιο της επιμήκυνσης της μετάφρασης, υφίσταται μια μοναδική μετα-μεταφραστική τροποποίηση στη λυσίνη 50 (Κ50), μέσω της προσθήκης σε αυτή ενός σπάνιου αμινοξέος, της υπουσίνης (hypusine). Μια παρόμοια τροποποίηση αποδείχθηκε ότι υφίσταται ωστόσο και ο EF-P της Escherichia coli (E. coli), ο οποίος τροποποιείται μετα-μεταφραστικά στη λυσίνη 34 (Κ34) με τη βοήθεια των ενζύμων YjeA και YjeK. Το YjeA αποτελεί παράλογο της δεύτερης κλάσης των tRNA συνθετασών της λυσίνης (LysRSs: Lysyl-tRNA synthetases), και καταλύει την προσθήκη της λυσίνης επάνω στον EF-P. Το YjeK είναι μια 2,3 αμινομουτάση της λυσινης (LAM: Lysine-2-3-aminomutase) και είναι αρμόδια για τη μετατροπή της α-λυσίνης σε β-λυσίνη. Εντούτοις, πρόσφατες έρευνες έδειξαν ότι ο πλήρως τροποποιημένος EF-P απαιτεί ένα επιπλέον ένζυμο, το YfcM, το οποίο ενεργεί ως υδροξυλάση και υδροξυλιώνει τον C4 ή C5 άνθρακα της K34 του EF-P. Στη συγκεκριμένη μελέτη εστιάσαμε στην εξέταση του μηχανισμού δράσης του EF-P και ειδικότερα στις επιπτώσεις που μπορεί να έχουν τα διαφορετικά στάδια των τροποποιήσεών του σε κύτταρα E. coli. Χρησιμοποιώντας τα knockout E. coli στελέχη της Keio (Δefp, ΔyjeK, ΔyjeA, ΔyfcM), ελέγξαμε την επίδραση διαφόρων περιβαλλοντικών συνθηκών στα στελέχη (διαφορετικές θερμοκρασίες, συνθήκες διατροφής, ευαισθησία σε αντιβιοτικά), δείχνοντας ότι το Δefp στέλεχος έχει μειωμένη αύξηση και ότι τα μεταλλαγμένα στελέχη παρουσιάζουν ευαισθησία σε μη-ριβοσωματικούς αναστολείς, όπως για παράδειγμα την αμπικιλίνη και τη ριφαμπικίνη. Επιπλέον, εξετάσαμε τη δυνατότητα των μεταλλαγμένων στελεχών να ανακάμπτουν στην ανάπτυξή τους παρουσία του κατάλληλου πλασμιδίου και είδαμε ότι ο EF- P είναι σημαντικός για την in vivo ανάπτυξη της E. coli σε στρεσσογόνες καταστάσεις. Όπως αναφέρθηκε πρόσφατα, οι YjeA, YjeK και EF-P πρωτεΐνες συμβάλουν στη μείωσης της τοξικότητας στη Salmonella. Επιπλέον, έχει δειχθεί πως ο EF-P είναι μια από τις πρωτεΐνες, που παίζουν σημαντικό ρόλο στην κινητικότητα των βακτηρίων στο Bacillus subtilis. Ωστόσο, έρευνα των Josenhans και Suerbaum το 2002, έδειξε πως η κινιτηκότητα και η τοξικότητα συχνά συνδέονται μεταξύ τους. Έτσι εξετάσαμε, τα μεταλλαγμένα στελέχη για την ικανότητά τους να κινούνται, δημιουργώντας μαστίγια, σε semi-solid θρεπτικό υλικό. Περαιτέρω έρευνες χρησιμοποιώντας external fluorescence staining και confocal microscopy, αποκαλύψε διαφορές στη μορφολογία των μεταλλαγμένων στελεχών της E. coli. Επίσης, με στόχο τη μελέτη της πρωτεΐνης YfcM, η λειτουργία της οποίας δεν είναι ακόμη γνωστή, απομονώσαμε και καθαρίσαμε την YfcM, με Νi-NTA agarose beads και gel filtration για τη μελλοντική κρυσταλοποίησή της,. Τέλος, μελλοντικός στόχος μας είναι η κλωνοποίηση του ακόλουθου πολυκιστρονικού γονιδίου “- yjeK - yjeA - yfcM - Ηis-efp - “, με σκοπό την υπερέκφραση και την κρυσταλλοποίησή του, ώστε να λάβουμε μια εικόνα του πως μοιάζει ολοκληρωμένο το μονοπάτι της τροποποίησης του EF-P, και επιπλέον, να μελετήσουμε καλύτερα τη λειτουργία του EF-P σε εκχυλίσματα της μετάφρασης από διαφορετικές μεταλλάξεις. Abstract 2

Abstract

Bacterial elongation factor P (EF-P) is a poorly understood soluble protein that has been shown to enhance the first step of peptide bond formation through an interaction with the ribosome and initiator tRNA. The crystal structure of EF‐P shows that EF‐P mimics the tRNA shape. Orthologous proteins have been found in both archaeal and eukaryotic systems, known as aIF5A and eIF5A, respectively. eIF5A, which was recently shown to increase translation elongation rates, is post-translationally modified at a highly conserved lysine residue (K50) through the addition of the rare amino acid hypusine. A similar pathway was recently elucidated for EF-P, in which EF-P is post-translationally modified by the enzymes YjeA and YjeK at lysine 34, corresponding to a homologous site of hypusylation in a/eIF5A. As a paralog of class II LysRS, YjeA catalyzes the addition of lysine onto EF-P, but is incapable of modifying tRNA. YjeK is a 2,3-(β)-lysine aminomutase and is responsible for converting lysine to β-lysine, which YjeA was recently shown to recognize as a preferred substrate for EF-P modification. However, fully modified EF-P requires a third enzyme, YfcM, which acts as a hydroxylase and hydroxylates the C4 or C5 position of K34 of EF-P, but not the added β-lysine. Based on a complete description of the EF-P modification and pathway, in this project we focused on further studies to address the mechanism of action of EF-P and especially to investigate how the different stages of EF-P’s modifications can affect E. coli cells. Using E. coli Keio knockout collection (Δefp, ΔyjeK, ΔyjeA, ΔyfcM) and E. coli Keio parental strain (wild-type) as reference, we checked the effect of the deletion strains on the cells under different environmental stress conditions (varying growth temperatures and nutrition conditions, susceptibility to antibiotics), showing that Δefp strain has growth defects and that E. coli efp mutants show sensitivity to non-ribosomal inhibitors, such as ampicillin and rifampicin, suggesting a possible secondary role of EF-P related to the cell envelope. Moreover, we tested the ability of deletion strains to restore viability in the presence of the appropriate plasmid and showed that EF-P is important for cell viability under certain conditions in E. coli. As reported previously, YjeA and YjeK are important in bacteria virulence. In addition, EF‐P is recognized as one of the proteins important for bacteria motility in Bacillus subtilis. However, motility and virulence are often linked together. Here, we tested deletion strains for their ability to produce flagella. Further, using external fluorescence staining and confocal microscopy we revealed differences in morphology of the E. coli deletion strains, and we performed Histidine tag protein purification with Ni-NTA agarose beads and gel filtration, in order to purify YfcM, an uncharacterized protein, and set initial screens for crystallization. Finally, our future goal is to clone the following polycistronic construct, “- yjeK - yjeA - yfcM - his-efp -“, overexpress and crystallize it, so as to see the crystal structure of the whole modification pathway of EF-P and study better the function of EF-P in translation extracts from different mutants.

Examining Committee 3

Examining Committee

Dinos George Thesis Adviser Assistant Professor of Biological Chemistry School of Medicine, University of Patras, Patras, Greece

Wilson N. Daniel Thesis Adviser Group Leader in Department of Chemistry and Biochemistry, Gene Center, Ludwig-Maximialians University (LMU) Munich, Germany

Kalpaksis Dimitrios Member of Committee Professor of Biological Chemistry School of Medicine, University of Patras, Patras, Greece

Acknowledgements 4

“I was taught that the way of progress was neither swift nor easy”

Marie Curie

Acknowledgements 5

Acknowledgements

This master thesis was held in collaboration with the Gene Center of Ludwig-Maximilian University in Munich. The writing of this thesis has been one of the most significant academic challenges I have ever had to face until now. Without the support, patience and guidance of the following people, this study would not have been completed. In one way or another, each one individually contributed, so it is to them that I owe my deepest gratitude.

First of all, my utmost gratitude goes to my supervisor Assistant Professor George Dinos, who gave me the opportunity to work in his lab. I am heartily thankful for his trust, support and help in every step I was about to take. He was always there smiling and advising me, always for my own good.

I would also like to thank Professor Dimitrios Kalpaksis, for participating in my examining committee. He continually and convincingly conveyed a spirit of adventure in regard to research and he was always there to help or advise me when needed.

Furthermore, this is a great opportunity to express my honor to Dr. Daniel Wilson, who was my thesis adviser in LMU and agreed with pleasure to be a member of my committee. Daniel’s encouragement, supervision and support from the preliminary to the concluding level, enabled me to develop an understanding of the subject I had to work with. His excitement in regard to science kept me motivated during the whole period I was working in his lab.

Special thanks go to all members of Wilson’s and the neighboring Beckmann’s lab for fruitful hints, as well as for humorous private discussions. However, I owe my deepest gratitude to my good friend and supervisor in Munich, Dr. Agata Starosta, for her patience and unselfish, but also unfailing support, as my thesis supervisor. Agata has been an exemplary editor, as constructive with her suggestions as she was meticulous with her corrections.

For help in the preparation of this thesis, I am also grateful to many friends and colleagues. I cannot mention all of them, but they include my co-workers in Greece, who gave me a helpful mixture of criticism and advice, and of course my friends. Their whole-hearted and enthusiastic belief in me was always very encouraging. I am sincerely grateful for the memories we have shared and those we have yet to create.

Above all, I thank my parents, who always supported me and my studies. They believed in me, in all my endeavors and encouraged me to keep going. Their love, support and constant patience have taught me so much about sacrifice, discipline and compromise.

Financial support for this study was provided by IKYDA 2010.

. Maria Stavropoulou Table of Contents 6

Table of Contents

Περίληψη ...... 1

Abstract ...... 2

Examining Committee ...... 3

Acknowledgements ...... 4

Table of Contents ...... 6

List of Figures ...... 10

Lists of Tables ...... 13

Abbreviations ...... 14

Introduction

1. The Ribosome ...... 17 1.1. Overview of the Prokaryotic Ribosome Structure ...... 17 1.2. Structure of the Prokaryotic Ribosomal Subunits ...... 18 1.3. 30S Ribosomal Subunit ...... 18 1.4. 50S Ribosomal Subunit ...... 19 2. Protein Synthesis in Prokaryotic Cells ...... 21 2.1. Initiation of Protein Synthesis ...... 24 2.2. Peptide Elongation and Translocation ...... 25 2.3. Termination of Protein Synthesis and Ribosomal Recycling ...... 27 3. Antibiotics Targeting the Ribosome ...... 28 3.1. Protein Synthesis Inhibitors ...... 30 3.2. Tetracyclines: Inhibitors of Accommodation of the A-tRNA ...... 30 3.3. Aminoglycosides: Inhibitors Inducing Misreading in Translocation ...... 30 Table of Contents 7

3.4. Spectinomycin: Inhibitor of Translocation by Stabilizing an Intermediate State of the Ribosome ...... 32 3.5. Macrolides: Inhibitors of Peptidyltransferase Center ...... 32 3.6. Chloramphenicol: Inhibitor of the A-site of the Peptidyltransferase Center ...... 33 3.7. Lincosamides: Overlap in Binding Site with Macrolides and Chloramphenicols .. 33 3.8. Oxazolidinones: Inhibitors of the A-site Accommodation ...... 33 3.9. Hygromycin A: Inhibitor of the Peptidyltransferase Center ...... 34 3.10. Viomycin: Inhibitor of Translocation by Stabilizing Hybrid State Formation ...... 35 4. Bacterial Resistance towards Antibiotics...... 35-36 5. Elongation factor EF-P ...... 36-50 5.1. Role and Structure of EF-P ...... 36 5.2. Eukaryotic Translation Initiation Factor 5A (eIF5A): The Eukaryotic Orthologue of EF-P ...... 38 5.3. EF-P Modification Pathway in E. coli ...... 41 5.4. YjeK, YjeA and YfcM: Enzymes Modifying E. coli EF-P ...... 43 5.4.1. Lysine-2-3-aminomutase ...... 43

5.4.2. Lysyl-tRNA synthetase (LysRS) ...... 46 5.4.3. YfcM, a Hydroxylase ...... 49

6. Aim of the Project ...... 51

Materials and Methods

7. Materials...... 52-55 7.1. Chemicals ...... 52 7.2. Bacterial Strains ...... 52 7.3. Growth Media ...... 53 7.4. Equipment ...... 54 7.5. Plasmids ...... 54 7.6. Primers ...... 55 8. Methods ...... …………………………………………………………………………………………..56-69 Table of Contents 8

8.1. Microbiological Analysis ...... 56 8.1.1. Growth Analysis of E. coli Keio Collection ...... 56 8.1.2. Growth under Stress Condition ...... 56 8.1.2.1. Growth of Cells in Various Media………………………………………………………….56 8.1.2.2. Temperature-sensitive Growth in Various Media at 37oC and 44oC………56 8.1.2.3. Antibiotic Susceptibility Assay……………………………………………………………….56 8.2. Rescue Experiments ...... 57 8.3. Motility Test ...... 57 8.4. Confocal Microscopy………………………..………………………………………………………………58 8.5. Generating the Templates for Expression and Purification ...... 60 8.5.1. Polymerase Chain Reaction (PCR) ...... 60 8.5.2. PCR Purification and Gel Extraction...... 61 8.5.3. Agarose Gel Electrophoresis ...... 61 8.5.4. Site Directed Mutagenesis ...... 61 8.6. Recombination Cloning in Bacteria ...... 62 8.6.1. Preparation of Competent Cells………………………………………………………………….…….62 8.6.2. Digestion by Restriction Enzymes ………………………………………………………….…..……..62 8.6.3. Ligation and Transformation……………………………………………..………………….…………..62 8.6.4. Colony Polymerase Chain Reaction (PCR)……………………….…………………….…………..63 8.6.5. Miniprep Isolation of Plasmid DNA………………………………………….………………………..64 8.7. Analysis of the Generated Clones………………………………………………………….………….64 8.7.1. Expression in E. coli………………………………….………………………………………………………..64 8.7.2. Sodium Dodecyl Sulphate-Polyacrylamid Gel Electrophoresis (SDS-Page) and Coomasie Staining…………………………………………………………………………………….……….65 8.7.3. 6xHis-tag Protein Purification using Ni-NTA Agarose Beads……………………………….66 8.8. Gel Filtration…………..………………………………………………………………………………….….…68

Results

9. Results ...... 70-84

9.1. Deletion of efp or yjeA lead to growth defects in E. coli ...... 70 9.2. efp is a Nonessential Gene for Cell Viability ...... 72 9.3. E. coli Deletion Strains Show Sensitivity to Nonribosomal Inhibitors ...... 73 9.4. EF-P is Important in Cell Viabillity under Stress Conditions in E. coli ...... 77 9.5. Lysine is the Required Substrate for YjeA in vivo ...... 79 9.6. ΔyfcM strain is might be Virulent for E. coli ...... 80 9.7. Cells Lacking YjeA or YjeK have Phenotypes Similar to those Lacking EF-P………. 81 Table of Contents 9

9.8. YfcM Purification for Crystallization ...... 83 9.9. Cloning of the Polycistronic Construct: -yjeK - yjeA - yfcM - his-efp- ...... 84

10. Discussion and Perspectives ...... 85-88 11. References ...... 89-106 12. Appendix...... 107-118

List of Figures 10

List of Figures

Figure 1.1. Crystal structure of 70S ribosome in complex with termination factors…………………18 Figure 1.2.1.1. Structure of 30S SU…………………………………………………………………………….……….…..19 Figure 1.2.2.1. Structure of 50S SU………………………………………………………………………………………….20 Figure 1.2.2.2. The ribosomal tunnel of bacterial and eukaryotic ribosomes……………….………….21 Figure 2.1. The secondary and tertiary structure of tRNA…………………………….………………………….22 Figure 2.2. The two ribosomal subunits……………………………………………………………………………………23 Figure 2.3. Overview of bacterial translation……………………………………………………………………………24 Figure 2.2.1. Schematic of Peptide Bond Formation on the Ribosome…………………………………….27 Figure 3.1. Antibacterial drug targets………………………………………………………………………………………28 Figure 3.2. The sites of antibiotic action for the different stages of protein synthesis……………..29 Figure 4.1. Antibiotic resistance increases among pathogens………………………………………………….35 Figure 4.2. (A) Between 1962 and 2000, no major classes of antibiotics were introduced. (B) Antibacterial agents approved 1983–2007………………………………………………………………………………36 Figure 5.1.1. Ribbon model of E. coli EF-P.…………………………………………………………………..…………..37 Figure 5.1.2. EF-P is composed of three β-barrel domains……………………………………….………………37 Figure 5.1.3. The structure of EF-P bound to the ribosome……………………………………………………..38 Figure 5.2.1. Structure comparison of EF-P and eIF-5A………………………………………………….…………39 Figure 5.2.2. 3D structure of translation elongation factors…………………………………………….………39 Figure 5.2.3. Ribbon diagram and representative sketch of the structure of aIF5A from M. jannaschii…………………………………………………………………………………………………………………………….…..40 Figure 5.2.4. Model structure of eIF5A (A) and the hypusine synthesis pathway (B)…………….….40 Figure 5.2.5. Amino acid sequence conservation of eIF5A, DHS and DOHH in eukaryotes……….41 Figure 5.3.1. Model for YjeA, YjeK, and EF-P Function ...... 42 Figure 5.3.2. (A) Modification pathway of EF-P. (B) Modification pathway of eIF-5A. (C) Relative positions of K34 of EF-P (green) and the acceptor stem of the P-tRNA (blue) when bound on the ribosome………………………………………………………………………………………………………………………….………43 Figure 5.4.1.1. Reaction catalyzed by LAM……………………………………………………………………….………44 List of Figures 11

Figure 5.4.1.2. Tetrameric packing of LAM subunits…………………………………………………………………44 Figure 5.4.1.3. Structural Analysis of YjeK and LAM…………………………………………………………………45 Figure 5.4.2.1. Structural analysis of YjeA and LysRS2………………………………………………………………47 Figure 5.4.2.2. Alignment of protein sequences of the three isoforms of lysyl-tRNA synthetases………………………………………………………………………………………………………………………………48 Figure 5.4.3.1. Characterization of the hydroxylase YfcM……………………………………………..……...…49 Figure 8.2.1. The Confocal Laser Scanning Microscope Zeiss LSM710 and ZEN navigation software ...... 58 Figure 8.2.2. Light path and image formation in a CLSM ...... 59 Figure 8.2.3. Absorption and fluorescence emission profiles of propidium iodide bound to dsDNA ...... 59 Figure 8.5.3.1. Ni-NTA spin purification procedure ...... 66 Figure 8.6.1. Common terms in gel filtration ...... 68 Figure 8.6.2. Theoretical chromatogram of a high resolution fractionation ...... 69 Figure 8.6.3. The figure shows a schematic of a section through a Superdex particle ...... 69 Figure 9.1.1. Growth curves of wild type E. coli and deletion strains grown in LB media, showing that Δefp strain has growth defects...... 70 Figure 9.1.2. Growth under starving conditions……………………………………………………………………….71 Figure 9.2.1. Effect of temperature shift on the growth of wild-type and mutant cells ...... 73 Figure 9.3.1. The figure shows the graphs for each antibiotic tested in alphabetical order ...... 77 Figure 9.4.1. Δefp, ΔyjeK and ΔyjeA cannot restore wild-type phenotype in E. coli ...... 78 Figure 9.4.2. Δefp and ΔyjeA mutants restored the wild-type phenotype from the first till the last day of the experiment suggesting the important role of EF-P and YjeA in E.coli cells...... 79 Figure 9.6.1. Motility Test ...... 80 Figure 9.7.1. Morphology of wild-type and mutant strain in CLSM ...... 82 Figure 9.8.1. Purification of YfcM ...... 83 Figure 9.8.2. (A). Gel filtration of elution fraction of YfcM. (B) Coomassie stained gel of elution fractions after gel filtration ...... 83 List of Figures 12

Figure 9.9.1. The polycistronic gene cloned in pQE70 vector and digested with SphI, SacI and BamHI restriction enzymes ...... 84 Figure 9.9.2. In the figure the arrow indicates where the polycistronic gene should be, that is around 3.300 bp ...... 84

List of Tables 13

List of Tables

Supplementary Table 1. List of Primers for the Polycistronic Construct ...... 107

Supplementary Table 2. Maps of Vectors used...... 109

Supplementary Table 3. Sequences of Escherichia coli K-12 Proteins ...... 112

Supplementary Table 4. Antibiotics in clinical use and modes of resistance ...... 113

Supplementary Table 5. Chemical Structure of Different Classes of Antibiotics ...... 114

Supplementary Table 6. MICs of antibiotics tested...... 115

Supplementary Table 7. List of Primers for K34 Mutant ...... 118

Abbreviations 14

Abbreviations

30S SU 30S Subunit 50S SU 50S Subunit Å Ångström A. baumannii Acinetobacter baumannii A. baylyi Acinetobacter baylyi A. tumefaciens Agrobacterium tumefaciens aa amino acid aaRS aminoacyl-tRNA synthetase aa-tRNA aminoacyl-tRNA aIF5-A archaeal translation initiation factor 5A Apr Apramycin APS Ammonium Persulfate A-site aminoacyl-tRNA site Asp-tRNA synthetase Aspartyl-tRNA synthetase ATP Adenosine triphosphate B. subtilis Basillus subtilis C. burnetii Coxiella burnetii C. subterminale Clostridium subterminale CAM Chloramphenicol Clin Clindamycin Cryo-EM Cryo-Electron Microscopy D. radiodurans Deinococcus radiodurans Da Dalton Deoxyhypusine Nε-(4-aminobutyl)-lysine DHS Deoxyhypusine synthase DMSO Dimethylsulfoxide DNA Deoxyribonucleic acid dNTPs Deoxynucleotide triphosphates DOHH Deoxyhypusine hydroxylase E. coli Escherichia coli EF-G Elongation factor G EF-P Elongation factor P EF-Tu Elongation factor Tu eIF5-A eukaryotic translation initiation factor 5A E-site exit-site (binds deacylated-tRNA) Fe iron FeS iron (II) sulfide Met fMet-tRNAf N-formylmethionine-tRNA gDNA genomic DNA GDP Guanosine diphosphate GDPCP Guanosine 5’-β,γ-methylnetriphosphate GTP Guanosine triphosphate h44, h7, h34, h31, h16/H17 helix44, 7, 34, 31, helix16/Helix17 HygA Hygromycin A H. influenzae Haemophilus influenzae H. marismortui Haloarcula marismortui Abbreviations 15

Hypusine Nε-(4-amino-2-hydroxybutyl)-lysine IF1, 2, 3 Initiation factor 1, 2, 3 IPTG Isopropyl-β-D-1-thiogalactopyranoside K. pneumoniae Klebsiella pneumoniae K34 Lysine 34 Kan Kanamycin KOW motif Kyrpides, Ouzounis, Woese motif Ksg Kasugamycin L. pneumophila Legionella pneumophila LB media Luria bertani media Lnz Linezolid Lys50 Lysine 50 LysAM Lysine-2,3-aminomutase M. jannaschii Methanococcus jannaschii MDa Mega Dalton MIC Minimum Inhibitory Concentration min minute MITF Microphthalmia-associated transcription factor mQ H2O milli-Q H2O mRNA messenger RNA MS Mass Spectometry NC Nascent Chain OB domain Oligonucleotide-binding domain OD600 Optical density P. aerophilum Pyrobaculum aerophilum P. horikoshii Pyrococcus horikoshii PCR Polymerase Chain Reaction PLP Pyridoxal-5-phosphate post-70SIC 70S post-initiation complex Pre-30SIC 30S pre-initiation complex P-site Peptidyl-tRNA site PTC Peptidyltransferase center P-tRNA Peptidyl-tRNA RF1, 2, 3 Release Factor 1, 2, 3 RNA Ribonucleic acid RPPs Ribosome protection proteins r-proteins Ribosomal proteins RRF Recycling release factor rRNA ribosomal RNA RT Room temperature S. typhimurium Salmonella typhimurium SAM S-adenosylmethionine SD Shine-Dalgarno SDS Sodiumdodecylsulfate SDS-PAGE SDS-Polyacrylamide gel electrophoresis sec seconds Spc Spectinomyci Abbreviations 16

Str Streptomycin T. thermophila Tetrahymena thermophila T. thermophilus Thermus thermophilus TAE Tris-acetate-EDTA TB media Terrific broth media TEMED N, N, N’, N’-tetramethylethylene-1,2-diamine Tet Tetracycline TPF 1,3,5,-triphenylformazan Tris Tris-(hydroxymethyl)-aminomethane tRNA transfer RNA TTC Triphenyltetrazolium chloride V. cholera Vibrio cholera Vio Viomycin Y. pestis Yersinia pestis Introduction 17

1. Introduction

1.1. Overview of the Prokaryotic Ribosome Structure

Ribosomes were first observed in the middle of 1950 by a Romanian biologist George Palade in the electron microscope and classified as dense particles or organelles (Palade, 1955), a study that earned him the Nobel Prize. Though, the term “ribosome” was proposed by the scientist Richard B. Roberts in 1958. They are macromolecular nanomachines of more than 2.3 MD (Mega Dalton) that translate the genetic information into functional proteins (Nierhaus and Wilson, 2004) and they are composed of two-thirds RNA and one-third protein by mass. An Escherichia coli (E. coli) cell contains around 20.000 ribosomes, constituting about 25% of the dry cell mass (Bremer et al, 1987). Moreover, structure of the ribosome has shown that the ribosome is a ribozyme whose active sites consist of RNA and whose assembly and functions are assisted by ribosomal proteins (Moore and Steitz, 2003; Rodnina et al, 2007). The bacterial 70S ribosome is composed of two unequal subunits, a small 30S subunit (30S SU) and a large 50S subunit (50S SU), with molecular weights of approximately 800 and 1.500 kDa (kilo Dalton), respectively, where S stands for the Svedberg unit for sedimentation velocity (Laursen et al, 2005; Wilson and Nierhaus, 2007; Jenner et al, 2010). For almost 30 years crystallographers have sought to solve the structure of the ribosome, the largest and most complicated RNA–protein complex in the cell. The general structure of the ribosome is known since the early 1970s. In the early 2000s though, the structure has been achieved at high resolutions, on the order of a few Å (ångströms). The first papers describing atomic structures of ribosomes derived from high resolution X-ray studies that appeared in 2000 and since then many more have been published. Those first structures include a 2.4 Å resolution structure of the large subunit from Haloarcula marismortui (Ban et al, 2000), a 3.0 Å resolution structure (Wimberly et al, 2000) and a 3.2 Å resolution structure (Pioletti et al, 2001) of the 30S SU from Thermus thermophilus, and a 3.1 Å resolution structure of the 50S SU from Deinococcus radiodurans (Harms et al, 2001). In the recent years, the structure of the 70S ribosome has been determined at atomic resolution (2.8 Å) (Selmer et al, 2006). Additionally, a crystal structure refined to 3.6 Å resolution of the ribosome trapped with elongation factor G (EF-G) in the post-translocational state using the antibiotic fusidic acid was published (Gao et al, 2009). Ramakrishnan and co-workers determined the crystal structure of the ribosome complexed with elongation factor Tu (EF-Tu) and aminoacyl tRNA (aa-tRNA) at 3.6 Å resolution (Schmeig et al, 2009) and recently, they also presented the crystal structure of the 70S ribosome with release factor 3 (RF3) in the presence of a nonhydrolyzable GTP (guanosine- 5’-triphosphate) analogue, GDPCP (guanosine 5′-β,γ-methylenetriphosphate), refined to 3.8 Å resolution (Fig. 1.1C) (Jin et al, 2011). Crystal structures of T. thermophilus 70S ribosome complexes bound with release factor 1 (RF1) in response to UAA or UAG codons and release factor 2 (RF2) in response to UAA or UGA codons were determined at 3.0–3.6 Å resolution, respectively (Fig. 1.1A,B) (Korostelev et al, 2008; 2011; Laurberg et al, 2008; Weixlbaumer et al, 2008), while crystal structures of the bacterial ribosomal recycling factor (RRF) bound to a 70S ribosomal complex (Dunlkle et al, 2011) were also solved. Introduction 18

Further, studies about the eukaryotic ribosome have been also published. Recently, the complete crystal structure of the yeast Saccharomyces cerevisiae 80S ribosome has been shown at a 3.0 Å resolution (Ben-Shem et al, 2011). Moreover, crystal structure of Tetrahymena thermophila 40S subunit in complex with eukaryotic translation initiation factor 1 (eIF-1) at 3.9 resolution (Rabl et al, 2011) and T. thermophila structure of 60S subunit in complex with eukaryotic translation initiation factor 6 (eIF-6), co-crystallized with the antibiotic cycloheximide at 3.5 Å resolution (Klinge et al, 2011), were published.

Figure1.1. Crystal Structure of 70S Ribosome in complex with termination factors. (A, B) Crystal structures of the 70S translation termination complexes bound with RF1 (3.2 Å resolution) and RF2 (3.0 Å resolution) (Korostelev et al, 2011). (C) Structure of RF3●GDPCP bound to 70S ribosome refined to 3.8 Å resolution (Jin et al, 2011).

1.2. Structure of the Prokaryotic Ribosomal Subunits

1.2.1. 30S Ribosomal Subunit

The 30S SU contains a single RNA roughly 1500 nucleotides in length, 16S rRNA, and 21 ribosomal proteins (r-proteins) (S1-S34) (Ramakrishnan and Moore, 2001). According to Yonath and coworkers, 30S SU’s traditional subvisions are the “head,” “neck,” and “body”; the body has a “shoulder,” a “platform,” and a “foot” with a “toe”. The head has a “nose” with a further protuberance, the “beak” (Fig. 1.2.1.1A). Three long helices, h44, h7, and h16/H17 run parallel to the long axis of the subunit (Schluenzen et al, 2000). The 30S SU provides (i) the mRNA- binding machinery, (ii) the path along which the mRNA progresses, (iii) the decoding center (Fig. 1.2.1.1B) and (iv) most of the components that control translation fidelity. The main function of 30S SU though is decoding. The decoding center organizes mRNA and tRNA translocation and controls fidelity in codon–anticodon interactions by monitoring base pairing between them (Green and Noller, 1997; Ramakrishnan, 2002) and it is located at the upper part of the body and the lower part of the head (Schluenzen et al, 2000). Introduction 19

Figure 1.2.1.1. Structure of 30S SU. (A) RNA is shown in gold, using a ribbon backbone and simple lines for base pairs. The differently colored helical segments and loops are the proteins. The major subdivisions are labeled: H, head; B, body; S, shoulder; P, platform; N, nose; F, foot. (B) A surface representation of the subunit, viewed from the side of the 50S SU. The latch is circled in cyan and the decoding center in red (Schluenzen et al, 2000).

1.2.2. 50S Ribosomal Subunit

In E. coli, the 50S SU contains a 2900-nucleotide RNA, 23S rRNA, an RNA of about 120 nucleotides, 5S rRNA, and 36 r-proteins (L1-L36) (Fig. 1.2.2.1) (Ramakrishnan and Moore, 2001). The secondary structure of 23S is divided into six large domains, within which domain V is crucial for its peptidyl transferase activity (Ban et al, 2000). Each domain has secondary structure, is highly symmetric in tertiary structure and is protrude by proteins between their helices. Proteins are mostly on the surface and largely absent from the active site inside. The main function of the proteins is to stabilize the tertiary structure and orientation of the rRNA (Ban et al, 2000). 50S SU includes the activity that catalyzes peptide bond formation (peptidyl transfer reaction) (Ban et al, 2000; Moore And Steitz, 2011), prevents premature polypeptide hydrolysis (Ban et al, 2000), provides a binding site for the GTPase dependent translational Introduction 20

factors – G-protein (assists initiation, elongation and termination) (Ban et al, 2000), and helps protein folding after synthesis (Ban et al, 2000; Nissen et al, 2000). The active site for peptide bond formation, the peptidyl-transferase center (PTC), is located in a cleft on the intersubunit side of the 50S SU (Simonovic and Steitz, 2009). As the nascent polypeptide chain (NC) is being synthesized, it passes through a tunnel within the large subunit and emerges at the solvent side where protein folding occurs (Wilson and Beckmann, 2011). X-ray structures of bacterial and archaeal ribosomal particles have revealed that the ribosomal tunnel is predominantly composed of rRNA (Nissen et al, 2000; Harms et al, 2001; Schuwirth et al, 2005; Selmer et al, 2006), consistent with an overall electronegative potential (Lu et al, 2008; Lu et al, 2007). Additionally, r-proteins, like L4 and L22, contribute to the formation of the tunnel wall, and form a ‘constriction’ where the tunnel narrows (Nissen et al, 2000). Near the tunnel exit, in bacteria-specific extension of L23 (L25 in eukaryotes) occupies a similar position to the r-protein L39e of eukaryotic and archaeal ribosomes (Harms et al, 2001; Schuwirth et al, 2005; Selmer et al, 2006) (Fig. 1.2.2.2).

Figure 1.2.2.1. Structure of 50S SU. (A) Secondary structure of 23S RNA. (B) Tertiary structure of 50S SU. Each domain has a similar color in the secondary and tertiary structures (Ramakrishnan and Moore, 2001). Introduction 21

Recently, studies by Beckmann and co-workers, showed that cryo-EM enabled NCs to be directly visualized within the ribosomal tunnel, extending from the PTC to the exit site on the back of the 50S SU (Becker et al, 2009). Nevertheless, the functional role of the ribosomal tunnel is not clear yet, as for many years, it was thought to be just a channel for the NCs, but lately there are evidences that for some NCs the tunnel plays a more active role (Deutsch, 2003) in nascent polypeptide folding and translational stalling (Wilson and Beckmann, 2011).

Figure 1.2.2.2. The ribosomal tunnel of bacterial and eukaryotic ribosomes. Transverse sections through the (A) bacterial and (B) eukaryotic large ribosomal subunit to reveal the tunnel components (Wilson and Beckmann, 2011).

2. Protein Synthesis in Prokaryotic Cells

Protein synthesis occurs in the 70S ribosome and it’s the process by which messenger- RNA (mRNA), that carries the genetic code to the ribosome, is translated into proteins. For protein synthesis it is necessary to select transfer-RNAs (tRNAs) that bind to the required amino acids, according to the genetic code, so that the amino acids are accurately polymerized (Yonath, 2005). tRNAs are actually non-ribosomal entities, L-shaped in their tertiary structure (Fig. 2.1), that connect the RNA and protein worlds since at one end of the molecule they have an anticodon that is complementary to a codon specifying a particular amino acid, and at the other end, the specific amino acid is linked to the 3’ terminal CCA-end of the tRNA by an ester bond (Wilson, 2009). On the 70S ribosome there are three main binding sites for tRNA: (i) The aminoacyl-site (A-site), where the decoding takes place. Here, the correct aa-tRNA is selected based on its codon-anticodon interaction, respective to the codon of the mRNA currently displayed. (ii) The peptidyl-site (P-site), where peptidyl tRNAs (tRNA bearing the nascent Introduction 22

polypeptide chain) is located, carrying the elongating polypeptide chain. (iii) The exit-site (E- site) through which deacylated or uncharged tRNAs pass as they are released from the ribosome (Fig. 2.2) (Ramakrishnan et al, 2002; Zaher and Green, 2009a; Jenner et al, 2010; Malys and McCarthy, 2011).

Figure 2.1. The secondary (left) and tertiary (right) structure of tRNA (Cooper, 2000).

During protein synthesis a single ribosome can incorporate 10-20 amino acids (aa) per second (Bremer and Dennis, 1996). The aa incorporation is very accurate with only one misincorporation per 3000 aa (Bouadloun et al, 1983). Translational fidelity relies on the combined accuracy of two basic processes: (i) the aminoacylation of tRNAs with their cognate amino acid by the aminoacyl-tRNA synthetases and (ii) the selection of cognate aminoacyl- tRNAs by the ribosome in cooperation with the GTPase EF-Tu. These two processes, which together ensure the specific acceptance of a correctly charged cognate tRNA into the A-site, operate prior to peptide-bond formation. However, recent studies reported the identification of an additional mechanism that contributes to high fidelity protein synthesis following peptidyl transfer, where RF3 plays a major role (Zaher and Green, 2011). RF3 is a GTPase integrally involved in the removal of the class I RF following peptide release (Freistroffer et al, 1997; Grentzmann et al, 1998; Zavialov et al, 2001), but has no effect on the rate constants for peptide release on authentic termination complexes (Freistroffer et al, 1997). In this retrospective quality control step, incorporation of an aa from a non-cognate tRNA into the growing polypeptide chain leads to a general loss of specificity in the A-site of the ribosome, and thus to a propagation of errors that results in abortive termination of protein synthesis. Interestingly, it was shown recently that when RF2 and RF3 were added together to a variety of P-site-mismatched complexes, release activity was substantially accelerated (Zaher and Green, 2009b). So, it was reported that the resulting rate constants for the release reaction can be remarkably fast for some complexes, in a range where this promiscuity could influence the fidelity of protein synthesis in vivo (Zaher and Green, 2009b; Zaher and Green, 2011). Introduction 23

Interestingly, Ehrenberg and co-workers found that RF3 could stimulate release on certain ribosome complexes containing a near-cognate stop codon in the A-site (Freistroffer et al, 2000).

Figure 2.2. The two ribosomal subunits. Left: The small ribosomal subunit from T. thermophilus showing positions of codon–anticodon interactions of A-, P- and E-tRNAs (Schluenzen et al, 2000). Right: The large ribosomal subunit from D. radiodurans (Ban et al, 2000). The figure presents the two stalks controlling the A-site tRNA entrance (L7/L12) and the E-site tRNA exit (L1), which are known to undergo a coordinated lateral movement during elongation. Insert: a tRNA molecule on which its two functional domains (the anticodon loop and CCA 3’ end. The brown oval indicates the portion of the tRNA molecule interacting with the 30S subunit, and the blue oval shows the portion bound to the 50S subunit (Bashan and Yonath, 2008).

Furthermore, protein synthesis can be divided into three distinct phases: initiation, elongation, and termination/recycling (Fig. 2.3) (Schmeing and Ramakrishnan, 2009). Each of these phases has a specific set of translation factors that regulate the process (Wilson and Nierhaus, 2007). In vitro translation systems have demonstrated that three initiation factors (IF1, IF2, and IF3) (Simonetti et al, 2009), three elongation factors (EF-G, EF-Tu, and EF-Ts), and three of the four termination factors (RF1 or RF2, RF3, and RRF) are necessary and sufficient for synthesis (Shimizu et al, 2001), although RF3 does not appear to be essential in vivo and is missing from 72 of the 191 available bacterial genomes as shown by Tenson and co-workers (Margus et al, 2007). However, many additional factors may be required in vivo, as for example EF-P, which was shown to stimulate the first peptide bond during initiation translation (Ganoza and Aoki, 2000; Swaney et al, 2006) and LepA (EF4), which has a role in counteracting mistranslocated ribosomes (Qin et al, 2006). Introduction 24

Figure 2.3. Overview of bacterial translation (Schmeing and Ramakrishnan, 2009).

2.1. Initiation of Protein Synthesis

Translation initiation (Fig. 2.3) is a process in which mRNA, initiator tRNA, initiation factors, and small and large ribosomal subunits associate with each other to form a complex. Initiation of translation in bacteria begins with the formation of a 30S pre-initiation complex (pre-30SIC) (Schmeing and Ramakrishnan, 2009). First, initiation factor 3 (IF3) binds rapidly to the 30S SU and blocks premature subunit docking (Subramanian and Davis, 1970). Then, mRNA binds to the 30S SU near its 3’ end in a process enabled by IF3. The binding of mRNA to the 30S SU is facilitated by a Shine-Dalgarno (SD) sequence in the mRNA, which is complementary to the 16S rRNA (anti-SD sequence) (Kaminishi et al, 2007; Schmeing and Ramakrishnan, 2009). Additionally, presence of initiation factor 1 (IF1) prevents the binding of the initiator tRNA to the 30S SU A-site (Ramakrishnan, 2002; Simonetti et al, 2009). IF1 interacts with the decoding center and may therefore indirectly regulate the association of the ribosomal subunits by influencing the conformation of the 30S SU. Thus IF1 may have a role in ensuring the accuracy of translation initiation (Ramakrishnan, 2002). IF1 is also known to increase the affinity of IF2 and IF3 for the 30S complex and in this way enhance their activities (Antoun et al, 2006; Pavlov Introduction 25

et al, 2008). Once the 30S SU·IF3·mRNA – complex is formed, initiation factor 2 (IF2), in a GTP- dependent manner, binds to the complex, and promotes the binding of the initiator N-formyl Met methionine-tRNA (fMet-tRNAf ) to the pre-30SIC (Gualerzi and Pon, 1990; Yusopuva et al, 2006; Kaminishi et al, 2007). The formation of the pre-30SIC is followed by the binding of the 50S SU and the eventual formation of 70S post-initiation complex (post-70SIC). However, there are indications that the newly formed 70SIC has to undergo a series of conformation changes (Pon and Gualerzi, 1984; Yusopuva et al, 2006; Kaminishi et al, 2007). Those conformational changes include tightening of the intersubunit contacts (Hennelly et al, 2005), a ratchet-like Met movement of the 30S relative to the 50S SU and relocation of fMet-tRNAf into the P-site upon GTP-hydrolysis catalyzed by IF2, bound to 30S P-site, and the release of IF1 and IF3 (Allen et al, 2005; Grigoriadou et al, 2007a). After the formation of 70SIC IF2 is also released (Marzi et Met al, 2003). The importance of the positioning of fMet-tRNAf in the 70SIC is highlighted by the presence of elongation factor P (EF-P) in E. coli. In the 70SIC EF-P binds to the region overlapping with the E-site and helps to correctly position fMet-tRNA for the first peptide-bond formation (Glick and Ganoza, 1975; Blaha et al, 2009). The 70S ribosome is now ready to enter the elongation cycle. Later, when translation is completed, IF3 is needed for the dissociation of the complex (Benne et al, 1973; Antoun et al, 2003; 2006).

2.2. Peptide Elongation and Translocation

The elongation of the nascent peptide by one aa according to the corresponding codon on the mRNA is performed by the ribosome in a cyclic manner (Vesper and Nierhaus, 2004). The polypeptide chain grows in the direction of the N-terminus to the C-terminus (Dintzis, 1961). Two main elongation factors, EF-Tu and EF-G are involved in the elongation cycle (Fig. 2.3) (Noble and Song, 2008; Schmeig and Ramakrishnan, 2009), and the cycle can be divided into the following three basic reactions: (i) occupation of the A-site by the incoming tRNA, (ii) peptide-bond formation, and (iii) translocation (Vesper and Nierhaus, 2004; Beringer and Rodnina, 2007; Frank et al, 2007; Schmeig and Ramakrishnan, 2009). The A-site occupation can be subsequently divided into two reactions. In the first reaction, decoding, EF-Tu associated with GTP and the aa-tRNA (ternary complex, aatRNA•EF- Tu•GTP), enters the A-site based on the codon-anti-codon interaction. Following the delivery of the aa-tRNA to the A-site, the GTPase center of EF-Tu is activated leading to the GTP hydrolysis and release of the EF-Tu•GDP complex from the ribosome (Schmeig and Ramakrishnan, 2009). The second reaction is known as the accommodation step, wherein the release of the EF- Tu•GDP from the ribosome allows the aa-tRNA to be bound to the A-site by docking the aminoacyl residue of the aa-tRNA to the PTC of the 50S SU. The A- and P-site occupied aa-tRNAs are now ready for the formation of the peptide bond (Vesper and Nierhaus, 2004). This state of the ribosome is called the ‘pre-translocational’ state (PRE) (Vesper and Nierhaus, 2004; Schmeig and Ramakrishnan, 2009). Introduction 26

The second step of the elongation cycle is a peptide-bond formation, which leads to the subsequent formation of a peptidyl-tRNA at the A-site and a deacylated tRNA at the P-site (Vesper and Nierhaus, 2004). The reaction catalyzed by the PTC of the ribosome is the aminolysis of an ester bond, with the nucleophilic α-amino group of A-site aa-tRNA attacking the carbonyl carbon of the ester bond linking the peptide moiety of P-site peptidyl-tRNA (Fig. 2.2.1) (Beringer and Rodnina, 2007; Frank et al, 2007; Schmeig and Ramakrishnan, 2009; Simonovic and Steitz, 2009). The third and final step of the elongation cycle is the translocation. During translocation, A-site bound peptidyl-tRNA and P-site deacylated tRNA move to the P- and E-sites, respectively. This state of the ribosome is called the ‘post-translocational state’ (POST) (Vesper and Nierhaus, 2004). The P- and E-sites of the ribosome are occupied and the A-site is vacant and waits to receive the new aa-tRNA•EF-Tu•GTP ternary complex based on the codon on the mRNA, thus the elongation cycle is repeated. However, during the next A-site occupation, the E-site tRNA has to be released (Vesper and Nierhaus, 2004; Beringer and Rodnina, 2007; Frank et al, 2007). This movement of the tRNA•mRNA complex within the ribosome by one codon is catalyzed by elongation factor G (EF-G) (Spahn and Nierhaus, 1998; Wilson and Noller, 1998). Finally, the ribosome continues to translate the remaining codons on the mRNA as more aa-tRNA bind to the A-site, until the ribosome reaches a stop codon on mRNA (UAA, UGA, or UAG) (Petry et al, 2008). Translocation involves multiple large conformational changes in the ribosome (Horan et al, 2007; Schuwirth et al, 2005; Spahn et al, 2004; Valle et al, 2003; Savelsbergh et al, 2003; Peske et al, 2004). In particular, it is thought to begin with a ratchet-like motion of the small 30S SU relative to the 50S SU (Valle et al, 2003; Horan et al, 2007), followed by swiveling of the head of the small subunit (Schuwirth et al, 2005; Spahn et al, 2004) and “unlocking” of the tRNA binding groove to allow peptidyl-tRNA in the P-site to pass into the E-site (Schuwirth et al, 2005; Pan et al, 2007). These conformational changes lead to discrete movements of bound tRNAs, where the tRNAs move first with respect to the 50S SU into hybrid binding states and then move with respect to the 30S SU (Moazed and Noller, 1989). An interesting turn was given to the translocation research in the last decade, when it was shown that in addition to the spontaneous forward translocation the ribosome is able to undergo translocation in the reverse direction (back- or retrotranslocation) (Shoji et al, 2006; Konevega et al, 2007). At about the same time Qi and co-workers identified a previously known protein LepA (Dibb and Wolfe, 1986; Bijlsma et al, 2000) as the factor that catalyzes translocation in the reverse direction (Qin et al, 2006). Although structurally similar to EF-G (Evans et al, 2008), the LepA-catalyzed back-translocation proceeds through different intermediate states (Connell et al, 2008; Liu et al, 2010).

Introduction 27

Figure 2.2.1. Schematic of peptide bond formation on the ribosome. The α-amino group of aminoacyl-tRNA in the A-site (red) attacks the carbonyl carbon of the pept-tRNA in the P-site (blue) to produce a new, one amino acid longer pept-tRNA in the A-site and a deacylated tRNA in the P-site (Beringer and Rodnina, 2007).

2.3. Termination of Protein Synthesis and Ribosomal Recycling

During termination (Fig. 2.3), a stop codon in the A-site is recognized by class-I release factors (RFs), RF1 (recognizes codons UAA, UAG) or RF2 (recognizes codons UAA, UGA) in prokaryotes (Capecchi, 1967; Scolnick et al, 1968). These factors induce the hydrolysis of peptidyl-tRNA, resulting in the release of the newly synthesized protein from the ribosome (Kisselev and Buckingham, 2000). The class II release factor RF3 accelerates the dissociation of class I factors from the ribosome after peptide release in a GTP-dependent manner as mentioned before (Freistroffer et al, 1997; Grentzmann et al, 1998; Zavialov et al, 2001), leaving the ribosome leaving the POST complex. The binding of RF3 to the ribosome–RF1/2 complex in the GDP form is thought to induce RF3 to exchange GDP for GTP (Zavialov et al, 2001). The crystal structure of RF3–GDP resembles EF-Tu in the GTP form (Gao et al, 2007). The same study showed that the binding of RF3 in the GTP form to the ribosome induces conformational changes likely to destabilize the binding of class I RFs, thus leading to their dissociation from the ribosome (Gao et al, 2007). Ribosome recycling (Fig. 2.3) is regarded as the fourth step in protein synthesis. Upon termination of protein synthesis the ribosome recycling factor (RRF) -in conjunction with EF-G and IF3- promotes the dissociation of the ribosome into its subunits (Karimi et al, 1999; Hirokawa et al, 2006). There are two models over the action of RRF and EF-G (Karimi et al, 1999; Peske et al, 2005; Zavialov et al, 2005; Hirokawa et al, 2006; Gao et al, 2005). In model 1 (Hirokawa et al, 2006), RRF and EF-G not only catalyze the dissociation (splitting) of 70S ribosomes into subunits but they also catalyze the release of mRNA and tRNA. By contrast, in model 2 (Karimi et al, 1999; Peske et al, 2005; Zavialov et al, 2005; Gao et al, 2005), RRF and EF- G catalyze only the dissociation of the 70S ribosome into subunits. In this model, IF3 is required for the release of tRNA, and then mRNA is released spontaneously (Hirokawa et al, 2006) and reassociation is prevented (Peske et al, 2005; Zavialov et al, 2005; Vesper and Wilson, 2006).

Introduction 28

3. Antibiotics Targeting the Ribosome

Throughout history, there has been a continual battle between humans and the multitude of microorganisms that cause infection and disease (Tenover, 2006). An antibiotic is a substance or compound that inhibits the growth of or destroys microorganisms (Waksman, 1947). Antibiotics are derived from three sources: moulds or fungi; bacteria; and synthetic or semi-synthetic compounds. They can be used either internally or topically, and their function is either to inhibit the growth of pathogens or to kill them. Current antimicrobial therapies, which cover a wide array of targets (Walsh, 2003), fall into two general categories: bactericidal drugs, which kill bacteria with an efficiency of >99.9%, and bacteriostatic drugs, which merely inhibit growth (Pankey and Sabath, 2004). Antibacterial drugs target interactions are well studied and predominantly fall into three classes: (i) inhibition of DNA replication and repair, (ii) inhibition of cell-wall turnover, and (iii) inhibition of protein synthesis (Fig. 3.1) (Walsh, 2000).

Figure 3.1. Antibacterial drug targets. There are five main antibacterial drug targets in bacteria: cell-wall synthesis, DNA gyrase, metabolic enzymes, DNA-directed RNA polymerase and protein synthesis. The figure shows the antimicrobial agents that are directed against each of these targets. In the case of protein synthesis, aminoglycosides and tetracyclines inhibit 30S RNA, and macrolides, chloramphenicol and clindamycin inhibit 50S RNA (Coates et al, 2002).

As illustrated in figure 3.2, protein synthesis due to high complexity of the ribosome represents a main target for clinically relevant antibiotics (Gale et al, 1981). In fact, antibiotics have been identified that inhibit almost every step of translation, although with differing degrees of specificity (Wilson, 2009). Structures of antibiotics in complex with ribosomal particles have revealed that antibiotics target mainly (i) the tRNA-mRNA pathway on the 30S SU (Brodersen et al, 2000), (ii) the PTC (Schlunzen et al, 2001; Hansen et al, 2003; Harms et al, 2004; Schlunzen et al, 2004; Bashan et al, 2003) and (iii) the exit tunnel on the 50S SU Introduction 29

(Schluenzen et al, 2001; Hansen et al, 2003; Harms et al, 2004; Schlunzen et al, 2004). However, there are exceptions, such as the orthosomycins (evernimicin and avilamycin) and thiopeptides (thiostrepton and micrococcin) which bind to distinct sites on the 50S SU and interfere with translation factor binding (Mikolajka et al, 2011). In terms of the ribosome, target specificity appears to be conferred by the regions surrounding the drug binding sites. This outer layer of less conserved components may also contribute to differential binding and inhibition of antibiotics across distinct bacterial species. Generally though, antibiotics targeting the ribosome interact mostly, if not exclusively, with rRNA, which reflects the fact that the functional centers of the ribosome are rich in RNA. Exceptions include the thiopeptides, streptomycins and spectinomycins, where r-proteins L11, S12 and S5 contribute to their respective binding sites (Wilson, 2009). Further, as reviewed by Champney and co-workers, there are a large number of antibiotics that act as inhibitors of ribosomal subunit assembly in addition to their well characterized role as protein synthesis inhibitors. Recently, there has been evidence that the assembly defects are secondary effects that result from inhibition of translation instead of direct effects on assembly (Siibak et al, 2009). Particularly, when translation is inhibited then production of ribosomal proteins is prevented whereas transcription continues, leading to an excess of rRNA over r-proteins. This non-physiological imbalance between rRNA and r-proteins has been proposed to lead to the observed defects in ribosomal assembly, at least in the presence of the antibiotics erythromycin and chloramphenicol (Siibak et al, 2011).

Figure 3.2. The sites of antibiotic action for the different stages of protein synthesis (reviewed by Wilson, 2009). Introduction 30

3.1. Protein Synthesis Inhibitors

3.1.1. Tetracyclines: Inhibitors of Accommodation of the A-tRNA

Tetracycline (Tet) is produced from Streptomyces aureofaciens (Duggar, 1953) and has been used to treat a variety of bacterial infections since its introduction into medicine. Several binding sites of tetracycline on the 30S SU have been identified by crystallographic studies (Pioletti et al, 2001; Brodersen et al, 2000). The primary binding site of tetracycline (Tet1) is located on the head of the 30S SU, where the drug interacts with the irregular minor groove of h34 and the loop of h31 (Brodersen et al, 2000; Pioletti et al, 2001). Tet prevents the stable binding of the ternary complex aa-tRNA•EF-Tu•GTP to the A-site of the ribosome (Blanchard et al, 2004). Additionally, there are a variety of mechanisms that give rise to Tet resistance (Chopra and Roberts, 2001). The most important and unique mechanism though is the use of the so-called ribosome protection proteins (RPPs), such as Tet(O) and Tet(M) (Connell et al, 2003).

3.1.2. Aminoglycosides: Inhibitors Inducing Misreading in Translocation

Most antibiotics that target the ribosome are bacteriostatic (Kohanski et al, 2007). Aminoglycosides induce cell death and are therefore considered as a bactericidal class of antibiotics (Davis, 1987). However, aminoglycosides induce cell death at dramatically lower concentrations and incubation times than other translation inhibitors (Bakker-Woudenberg et al, 2005). The bactericidal effect of aminoglycosides is thought to result from misreading and misfolding of membrane proteins, which leads to oxidative stress and cell death (Kohanski et al, 2007; 2008). Aminoglycoside antibiotics are generally composed of a sugar moiety and an amino group. Those that are derived from bacteria of the Streptomyces genus are named with the suffix –mycin (such as streptomycin, spectinomycin, kanamycin, kasugamycin, apramycin), whereas those that are derived from Micromonosporaa (Kroppenstedt et al, 2005) are named with the suffix –micin (such as gentamicin (Dewick, 2009)). Aminoglycosides mainly target the 30S SU, where they usually bind to the 16S rRNA (Shakil et al, 2008). They specifically bind to the A-site internal loop within the deep groove of helix 44 (h44) of the 16S rRNA in the highly conserved region of the decoding center (Carter et al, 2000; Ogle et al, 2003), where they interfere with two flexible adenine residues (A1492, A1493; Ogle et al, 2001). These residues are involved in the selection of the cognate aa-tRNA during elongation. Consequently, these antibiotics affect the fidelity of protein translation (Moazed and Noller, 1987; Fourmy et al, 1996; Carter et al, 2000; Pape et al, 2000). They also seem to act as rigid molecular braces that prevent conformational changes important for ribosome function. An additional binding site for aminoglycosides, such as gentamicin and neomycin, has been observed in helix 69 (H69) of the 23S rRNA, where it has been proposed to inhibit the action of the ribosome recycling factor (RRF) during the ribosome recycling phase of translation (Borovinskaya et al, 2007a). However, it remains to be determined whether this secondary binding site plays a role in the inhibitory activity of aminoglycosides. Some of the known aminoglycosides and their mechanism of action are analyzed below: Introduction 31

 Apramycin (Apr): Apr is produced by Streptomyces tenebrarius (Ryden and Moore, 1977) and contains a bicyclic sugar moiety and a monosubstituted deoxystreptamine (O'Connor et al, 1976). Apr stands out among aminoglycosides for its mechanism of action which is based on blocking translocation and its ability to bind also to the eukaryotic decoding site despite differences in key residues required for Apr recognition by the bacterial target. The drug binds in the deep groove of the RNA which forms a continuously stacked helix comprising non-canonical C-A and G-A base pairs and a bulged-out adenine. The binding mode of Apr at the human decoding-site RNA is distinct from aminoglycoside recognition of the bacterial target, suggesting a molecular basis for the actions of Apr in eukaryotes and bacteria (Ryden and Moore, 1977).

 Kasugamycin (Ksg): Ksg is an antibiotic produced by Streptomyces kasugiensis that exhibits activity against a wide variety of microorganisms, but has low toxicity against plants, humans, fish and animals. Ksg was shown to prevent the formation of the pre- Met 30SIC by interfering with fMet-tRNAf , which binds to the ribosomal P-site in E. coli ribosomes (Schlünzen et al, 2006). In addition, recent studies have shown that treatment of the E. coli cells with Ksg in vivo leads to the formation and accumulation of stable 61S ribosomal particles. These particles comprise an intact 50S SU. However, many proteins of the small subunit were absent (Kaberdina et al, 2009). Furthermore, Ksg specifically inhibits translation initiation of canonical but not of leaderless mRNA. For initiation on leaderless mRNA, the overlap between mRNA and Ksg is reduced and the binding of tRNA is further stabilized by the presence of the 50S SU, minimizing Ksg efficacy (Kaberdina et al, 2009). The primary binding site of kasugamycin (Ksg1) is at the top of h44 (Schlünzen et al, 2006; Schuwirth et al, 2006). A secondary binding site (Ksg2) was also found in T. thermophilus located in the E-site (Vila-Sanjurjo et al, 1999).

 Streptomycin (Stp): Stp is structurally related to the aminoglycoside family of antibiotics and exhibits the same classical hallmark, i.e. Stp induces translational misreading (Kurland et al, 1996). Stp binds to a distinct site on the ribosome and therefore mediates its inhibitory and misreading effects by an unrelated mechanism to aminoglycosides. Unlike aminoglycosides that bind in h44, Stp has a single binding site on the 30S SU that connects helices from all four different domains of the 16S rRNA, namely h1, h18, h27 and h44, and makes interactions with r-protein S12 (Sharma et al, 2007). Selection of the correct or cognate tRNA by the ribosome results in domain closure of the 30S SU, where the head and platform close in on the A-tRNA (Ogle et al, 2002). The domain closure induced by Stp binding (Ogle et al, 2002) has been proposed to reduce the rate of GTPase activation for cognate tRNAs, while slightly enhancing the rate for near- cognate tRNAs, such that the overall rates are similar and rate-limiting (Gromadski and Rodnina, 2004), thus explaining the high translational misreading that Stp induces.

Introduction 32

3.1.3. Spectinomycin: Inhibitor of Translocation by Stabilizing an Intermediate State of the Ribosome

Spectinomycin (Spc) is an aminocyclitol antibiotic closely related to the aminoglycosides, produced by the bacterium Streptomyces spectabilis (Lewis and Clapp, 1961; Mason et al, 1961; Hoeksema and Knight, 1975). Spc is important in clinical and veterinary use, as it induces only low levels of bacterial resistance (Brocklehurst, 2002; Wisselink et al, 2006). Spc is constituted from two glucose moieties and like aminoglycosides, inhibits translocation of tRNA and mRNA in the ribosome (Bilgin et al, 1990; Fredrick et al, 2003; Peske et al, 2004), but the mechanism of inhibition differs since Spc appears to stabilize an intermediate state of the ribosome that occurs during translocation (Pan et al, 2007). Spc binds to the neck region of the 30S SU, where it interacts predominantly with the minor groove of h34 (Carter et al, 2000; Borovinskaya et al, 2007b). Biochemical data (Peske et al, 2004) and the crystal structure of Spc bound to the E. coli 70S reveals that the presence of the drug rotates and locks the head in a distinct position relative to the body, suggesting that Spc traps the ribosome in a translocation intermediate by preventing the rotation of the head necessary for movement of the tRNAs and mRNA into the P- and E-sites (Borovinskaya et al, 2007b).

3.1.4. Macrolides: Inhibitors of Peptidyltransferase Center

Macrolides represent a large class of polyketide compounds synthesized by actinomycetes that inhibit eubacterial, but not archaeal nor eukaryotic, protein synthesis (Gaynor and Mankin, 2003; Poehlsgaard and Douthwaite, 2003; Takashima, 2003; Katz and Ashley, 2005; Mankin, 2008). Macrolide antibiotics bind in a position adjacent to PTC, located within the exit tunnel of the 50S SU, through which the polypeptide chain passes during translation. Clinically used macrolides have a 14- (erythromycin, telithromycin), 15- (azithromycin), or 16-membered (tylosin, spiramycin and carbomycin) lactone ring, to which amino sugars are attached at various positions. Multiple structures of a variety of macrolides bound to bacterial and archaeal large subunits reveal a common binding mode, such that the general orientation and conformation of the lactone ring and C5-sugar is similarly placed to establish interactions with A2058 and A2059 of the 23S rRNA (Schlünzen et al, 2001; Hansen et al, 2002; Tu et al, 2005; Wilson et al, 2005). The emergence of bacterial resistance to macrolide antibiotics has led to the development of ketolides, semi-synthetic derivatives of macrolides where the C3-sugar is replaced with a keto group. The broader spectrum of activity of the ketolides seems to be related to the presence of their additional sidechains and modifications, such as the alkyl-aryl sidechain of telithromycin. Translation inhibition by macrolide antibiotics restricts the synthesis to short oligopeptides that eventually fall off the ribosome as oligopeptidyl-tRNAs, in a poorly defined process called “drop-off” (Otaka and Kaji, 1975; Menninger and Otto, 1982; Tenson et al, 2003). Moreover, the size of the oligopeptides synthesized appears to be related to the extent to which the macrolides occlude the tunnel, such that macrolides like spiramycin with a C5-disaccharide allow the synthesis of 2–4 amino acids, whereas smaller macrolides, such as erythromycin with a C5-monosaccharide, permit 6–8 aa to be polymerized before drop-off (Tenson et al, 2003). Indeed, macrolides with extensive side chains extending from the C5 position even inhibit peptidyltransferase activity to varying Introduction 33

extents, such as carbomycin (100% inhibition), spiramycin (85%) and tylosin (~60%), whereas those with shorter side chains, such as erythromycin, do not (Poulsen et al, 2000).

3.1.5. Chloramphenicol: Inhibitor of the A-site of the PTC

Chloramphenicol (CAM) was originally isolated from Streptomyces venezuele and displays a broad-spectrum activity, inhibiting a wide range of Gram-positive and Gram-negative bacteria, but not translation in eukaryotes. CAM binds in the A-site of the PTC of a bacterial ribosome (Schlunzen et al, 2001), in a position overlapping the methylated-tyrosyl-moiety of puromycin (Puro) (Schmeing et al, 2002) or more generally the aminoacyl moiety of an aa-tRNA. This is consistent with biochemistry showing that CAM interferes with the puromycin reaction as well as the binding of small tRNA fragments to the A-site of the PTC (Celma et al, 1971), but stimulates the binding of tRNA fragments to the P-site (Ulbrich et al, 1978). Cam is a classic elongation inhibitor, in the sense that addition of the drug to growing bacterial cells blocks the ribosomes on the mRNA and protects the peptidyl-tRNA from hydrolysis, enabling the visualization of polysomes on sucrose gradients. It has also been shown to influence translational accuracy, promoting stop codon readthrough and framshifting, but unlike the aminoglycosides, does not induce misincorporation (Thompson et al, 2002; 2004). CAM alters the protection pattern of multiple nucleotides within the PTC, including A2451, G2505 and U2506, but also nucleotides that are located in the ribosomal tunnel, such as A2058 and A2059 (Moazed and Noller, 1987; Rodriguez-Fonseca et al, 1995). Finally it has been reported to enhance the premature release of short oligopeptidyl tRNAs in vitro (Rheinberger and Nierhaus, 1990), reminiscent of the macrolide antibiotics.

3.1.6. Lincosamides: Overlap in Binding Site with Macrolides and Chloramphenicols

The lincosamides are excellent inhibitors of Gram-positive bacteria, but not of Gram- negative, with clindamycin also affecting protozoa, such as Pneumocystis carinii, but not inhibiting archaeal, or eukaryotic systems, such as rabbit reticulocyte (Spizek et al, 2004). Lincosamides bind within the A-site of the PTC (Schlunzen et al, 2001; Tu et al, 2005).

 Clindamycin (Clin): Clin is a semi-synthetic derivative of lincomycin (Spizek et al, 2004). Specifically, the prolyl-moiety of Clin overlaps the aminoacyl-moiety of A-tRNA and the binding site of CAM, whereas the sugar moiety extents into the ribosomal tunnel and overlaps the binding position of the macrolide antibiotics, such as erythromycin (Schlunzen et al, 2001; Tu et al, 2005). The majority of the interactions involve hydrogen bonds from hydroxyl groups on the sugar-moiety of Clin with nucleotides of the PTC, such as the nucleobases of A2058 and A2062, as well as the backbones of G2505 and A2503 (Douthwaite et al, 1992).

3.1.7. Oxazolidinones: Inhibitors of the A-site Accommodation

The oxazolidinones are a class of synthetic antibiotics that act against a wide spectrum of Gram-positive and anaerobic bacteria as well as exhibiting activity against multi-drug Introduction 34

resistant Gram-positive bacteria, such as MRSA (methicillin-resistant Streptococcous aureus). The first oxazolidinone to enter into the market was linezolid, launched in 2000 under the tradename Zyvox by Amersham Pharmacia (now Pfizer). Novel oxazolidinones are ranbezolid (Das et al, 2005; Kalia et al, 2009) and Rx-01 (Franceschi and Duffy, 2006; Skripkin et al, 2008), which display improved inhibitory activities compared to linezolid (Das et al, 2005; Lawrence et al, 2008), and even act against linezolid-resistant strains (Lawrence et al, 2008; Skripkin et al, 2008). In particular, oxazolidinones are used to treat skin and respiratory tract infections caused by S. aureus and Streptococci strains, as well as being active against vancomycin- resistant Enterococcus faecium (Prasad, 2007; Hutchiinson, 2004; Bozdogan and Appelbaum, 2004). Oxazolidinones have been suggested to inhibit translational events ranging from initiation and the formation of the first peptide bond, EF-G dependent translocation during elongation and frameshifting, to non-sense suppression during termination (Swaney et al, 1998; Thompson et al, 2002; Aoki et al, 2002).

 Linezolid (Lnz): Crystal structures of Lnz bound the bacterial (Wilson et al, 2008) and archaeal large subunits (Ippolito et al, 2008) reveal that it binds to the A-site of the PTC, in a position overlapping the binding sites of anisomycin, CAM as well as the aminoacyl moiety of an A-site bound tRNA. Binding of linezolid stabilizes the nucleobase of U2585 (E. coli numbering used throughout) in an orientation that is distinctly different from when A and P-site tRNA ligands are bound, suggesting that Lnz induces a nonproductive conformation of the PTC (Wilson et al, 2008). In these structures, the aromatic ring of lnz, anisomycin and CAM are similarly located, in good agreement with the observation that oxazolidinones compete with CAMs, lincosamides and puromycins for ribosome binding (Lin et al, 1997; Skripkin et al, 2008). An elegant series of in vivo crosslinking experiments (Colca et al, 2003) led to model for the binding position of Lnz at the A-site of PTC (Leach et al, 2007), which is in excellent agreement with the subsequent crystal structures.

3.1.8. Hygromycin A: Inhibitor of the Peptidyltransferase Center

Hygromycin A (HygA) is a secondary metabolite produced by Streptomyces hygroscopicus (Pittenger et al, 1953). Nevertheless, it was not until 30 years later that HygA was shown to target the ribosome by inhibiting the peptidyltransferase reaction as well as blocking the binding of CAM to the ribosome, suggesting that the binding sites of the two antibiotics are closely related (Guerrero and Modolell, 1980). Moreover, chemical footprinting studies have shown that HygA competes for binding with 16-membered macrolide antibiotics bearing disaccharides attached the C5 position of the lactone ring (Poulsen et al, 2000). Structure- activity relationships have indicated that the aminocyclitol moiety at one end of the molecule is essential for biological activity, whereas the 5-dehydrofucofuranose can be chemically substituted by an allyl group (Hayashi et al, 1997).

Introduction 35

3.1.9. Viomycin: Inhibitor of Translocation by Stabilizing Hybrid State Formation

Viomycin (Vio), a cyclic peptide antibiotic produced by Streptomyces sp. (Kitagawa et al, 1972), is a member of the tuberactinomycin family that is active against Mycobacterium tuberculosis. Vio was the first tuberactinomycin used to treat tuberculosis (Bartz et al, 1951) until it was replaced by the structurally related capreomycin, which is less toxic but is now only used as a second-line drug (Jain and Dixit, 2008). Vio may have two binding sites on the ribosome, one on the small subunit and one on the large, and also protects nucleotides 912– 915 and A1408 in the 16S rRNA as well as U913 and G914 in the 23S rRNA from chemical attack (Moazed and Noller, 1987). In particular, Vio stabilizes the peptidyl-tRNA at the A-site and the deacylated tRNA in a hybrid P/E state (Peske et al, 2004; Shoji et al, 2006; Ermolenko et al, 2007; Pan et al, 2007). Thus, like Spc, Vio inhibits translocation (Liou and Tanaka, 1976; Modolell and Vazquez, 1977) by trapping the ribosome in an intermediate state on the translocation pathway (Ermolenko et al, 2007), and additionally induces back-translocation (Shoji et al, 2006) and translational misreading (Wurmbach and Nierhaus, 1983), analogous to the aminoglycosides.

4. Bacterial Resistance towards Antibiotics

Soon after the first use of antibiotics in medicine, resistant organisms were seen to arise during therapy (Abraham and Chain, 1940). As antimicrobial usage increased, so did the level and complexity of the resistance mechanisms exhibited by bacterial pathogens (Tenover, 2006). More strains of pathogens have become antibiotic resistant, and some have become resistant to many antibiotics and chemotherapeutic agents, a phenomenon called multidrug resistance (Nikaido, 2009). Resistance has developed to every main class of antibiotic, both natural and synthetic, over the course of 1 year to more than a decade after the first clinical use. Therefore, resistance, has proven not to be a matter of if, but a matter of when (Walsh, 2003) and it is increasing even among pathogens (Fig. 4.1) (Livemore, 2004). Selection of resistance, though its frequency varies with: (i) the regimens and extent of use, (ii) the effectiveness of infection control, and (iii) random factors, such as the initial escape of resistance genes to mobile DNA and into biologically ‘fit’ strains (Livemore, 2004).

Figure 4.1. Antibiotic resistance increases among pathogens (Livemore, 2004). Introduction 36

Nevertheless, between 1962 and 2000, no major classes of antibiotics were introduced (Fischbach and Walsh, 2009) (Fig. 4.2A) and the last years there has been a decreased number of new drugs in the market (Fig. 4.2B) (Fox, 2006). So, the subsequent decades have seen a cycle of emergence of resistant microbes, followed by the subsequent development of new antibiotics. These have included brand new classes of drugs as well as medicinal chemical elaboration of established classes to avoid resistance. This cyclical approach has proven highly successful, and we are now employing fourth, fifth, and sixth generation antibiotics in many cases. Consequently, the continued evolution and selection of new resistance genes, along with the emergence of multi-drug resistant organisms, are driving the need for new drugs (Morar and Wright, 2010).

Figure 4.2. (A) Between 1962 and 2000, no major classes of antibiotics were introduced (Fischbach and Walsh, 2009). (B) Antibacterial agents approved 1983–2007. Decreased number of new drugs in the market (Fox, 2006).

5. Elongation Factor EF-P

5.1. Role and Structure of EF-P

EF-P was originally identified as a soluble protein that binds to the ribosome and stimulates peptide bond formation in protein synthesis on 70S ribosomes (Glick and Ganoza, 1975; Aoki et al, 1991; Ganoza and Aoki, 2000; Swaney et al, 2006). The EF-P protein is 20.6 kDa (Aoki et al, 1991) and there are approximately 0.1 – 0.2 copies of it per ribosome in E. coli, a ratio similar to that of the initiation factors (An et al, 1980; Cole et al, 1987). The efp gene encoding it has been cloned and sequenced (Aoki et al, 1997a) and it occurs at a unique site at 94.3 min on the E. coli chromosome (Aoki et al, 1991). EF-P, apart from its role in initiation translation, was detected in the monosome and polysome fractions, suggesting that EF-P may also be involved in elongation step of translation (Aoki et al, 2008). As shown by Ganoza and Aoki, EF-P under certain conditions enhances the translation of natural and synthetic mRNAs in vitro (Ganoza and Aoki, 2000). Another possible function that researchers focused on was whether EF-P can alter indirectly the affinity of the ribosome for aa-tRNAs, thus increasing their reactivity as acceptors for peptidyl transferase (Aoki et al, 1997b; Glick and Ganoza, 1975) and the role of EF-P as an assistance in accommodation of the side chains of amino acids of the growing polypeptide chain (Aoki et al, 1991; Swaney et al, 2006). Deletion of the efp gene leads Introduction 37

to defects in (i) growth of E. coli (Yanagisawa et al, 2010) and Acinetobacter baylyi (de Grecy et al, 2007), (ii) swarming (Kearns et al, 2004) and sporulation (Ohashi et al, 2003) of Bacillus subtilis, and (iii) virulence of Agrobacterium tumefaciens (Peng et al, 2001). In 2004, Yokoyama and collaborators, reported the crystal structure of EF-P from T. thermophilus HB8 at a 1.65 Å resolution (Hanawa-Suetsugu et al, 2004), while recently, Choi and Choe published the crystal structure of EF-P from Pseudomonas aeruginosa at 1.75 Å resolution (Choi and Choe, 2011). The structure revealed that the protein is composed of three β-barrel domains (domains I, II, and III) (Fig. 5.1.1) (Yanagisawa et al, 2010; Hanawa-Suetsugu et al, 2004; Blaha et al, 2009; Choi and Choe, 2011) and has an overall L-shape, reminiscent of a tRNA (Hanawa-Suetsugu et al, 2004; Blaha et al, 2009; Choi and Choe, 2011). Domain I is an N- terminal KOW-like domain (Fig. 5.1.2A). The KOW motif is found in a variety of ribosomal proteins and the bacterial transcription antitermination proteins NusG (Kyrpides et al, 1996). Domain II is the central OB domain, which forms an oligonucleotide-binding (OB) fold, but it is not clear yet if this region is involved in binding nucleic acids (Fig. 5.1.2B) and domain III is C- terminal domain which adopts an OB-fold, with five β-strands forming a β-barrel in a Greek-key topology (Fig. 5.1.2C) (Hanawa-Suetsugu et al, 2004). EF-P binds to both the 30S and 50S ribosomal subunits (Hanawa-Suetsugu et al, 2004). Moreover, the crystal structure of EF-P bound to the 70S T. thermophilus ribosome at 3.5 Å resolution published by Steitz and coworkers (Blaha et al, 2009), shows that EF-P binds at a distinct position that is adjacent to the P-site tRNA, between the P- and E-sites (Fig. 5.1.3). So, EF-P spans both ribosomal subunits and contacts the initiator tRNA near the anticodon stem-loop on the 30S SU, the D-loop, and the acceptor stem on the 50S SU (Blaha et al, 2009).

Figure 13. 5.1.1 Ribbon model of E. coli EF-P. EF-P domain 1 (residues 3–64), domain 2 (residues 65–128) and domain 3 (residues 129–187) are shown (Yanagisawa et al, 2010).

Figure 5.1.2. EF-P is composed of three β-barrel domains. (A) Crystal structure of translation initiation factor 5A from Pyrococcus horikoshii. (B) Crystal structure of translation EF-P from T. thermophilus HB8. (C) Crystal structure of translation EF-P from T. thermophilus HB8. Introduction 38

EF-P bears no obvious amino acid sequence similarity to any other prokaryotic translation factor (Aoki et al, 1991). Nevertheless, EF-P is conserved in all bacteria (Bailly et al, 2010) and orthologous to archaeal and eukaryotic initiation factor 5A (a/eIF-5A) (Kyrpides and Woese, 1998). As reviewed by Valentini and co-workers, similarly to EF-P, eIF-5A was also identified based on its ability to stimulate Met-puromycin formation (Park et al, 2010). Further, domains II and III of EF-P share a very similar topology, while the structures of domains I and II of EF-P are superposable on the structures of the M. jannaschii, P. aerophilum and P. horikoshii eIF-5A proteins (Hanawa-Suetsugu et al, 2004).

Figure 5.1.3. The structure of EF-P bound to the ribosome. (A) Overview of the E- and P-site tRNAs bound to the 70S ribosome. (B) Overview of EF-P and P-site tRNA–binding in the 70S ribosome. EF-P is shown as a surface representation in shades of magenta to indicate the different domains (I, II, and III) of the protein (Blaha et al, 2009).

5.2. Eukaryotic Translation Initiation Factor 5A: The Eukaryotic Orthologue of EF-P

The putative eIF5A is a highly conserved and essential protein, present in all organisms from archaea to mammals, but not in eubacteria (Chen and Liu, 1997). Despite being highly conserved and essential for eukaryotic cell proliferation (Lee et al, 2009; Nishimura et al, 2011), the critical cellular role of eIF5A remains unclear. eIF5A was initially purified from ribosomes and it was described as being involved in the formation of the first peptide bond (Benne et al, 1978; Smit-McBride et al, 1989; Park, 1989). Recently, studies in yeast, demonstrated that eIF5A promotes translation elongation rather than translation initiation (Zanelli et al, 2006; Gregio et al, 2009; Saini et al, 2009). eIF5A stimulates translation directly and functions as a general translation elongation factor in a manner determined by its hypusine modification (Saini et al, 2009). According to Zanelli and Valentini, eIF5A function is essential for polarized growth, a process necessary for the G1/S transition in yeast (Zanelli and Valentini, 2005). Introduction 39

Synthetic lethality was furthermore revealed between mutants of eIF5A (Frigieri et al, 2008), but the mechanism triggered by eIF5A to accomplish this cell cycle function is still unknown.

Figure 5.2.1. Structure comparison of EF-P and eIF-5A. (A) Superimposition of the ribbon diagrams of T. thermophilus EF-P (blue) and M. jannaschii eIF-5A (yellow). (B) Amino acid residues conserved in EF-Ps and eIF-5As color-coded on the surface of T. thermophilus EF-P (Hanawa- Suetsugu et al, 2004).

eIF5A shares sequence and structural similarity with the first two domains of EF-P as shown by Yokoyama and co-workers (Fig. 5.2.1) (Hanawa-Suetsugu et al, 2004) and together with aIF5A they lack a carboxy-terminal domain III found in bacterial EF-P (Fig. 5.2.2.) (Blaha et al, 2009). Additionally, the crystal structure of the M. jannaschii eIF5A shows that the protein is made of two β-sheet domains arranged in an elongated structure that is about 63 Å long and 26 Å wide (Fig. 5.2.3) (Kim et al, 1998). Figure 5.2.3 shows the arrangement of domain I, which has the β1 to β6 strands and the helix. Domain II resembles the OB fold (Murzin, 1993) and harbors strands β7 to β11, while the hypusine site of modification occurs in a long loop between strands β3 and β4 (Kim et al, 1998).

Figure 5.2.2. 3D structure of translation elongation factors. The 3D structure of representative examples of e(a)IF5A ⁄ EF-P proteins is drawn to demonstrate the structural similarity between eukarya, archaea and bacteria. The position of the unique modifications hypusine attached to conserved amino acids (numbered) is indicated by arrows (Greganova et al, 2011). Introduction 40

eIF5A is the only protein known to contain the unusual amino acid residue hypusine [Nε- (4-amino-2-hydroxybutyl)-lysine] (Chen and Liu, 1997; Wolff et al, 2007), which was first isolated by Kakimoto and co-workers (Shiba et al, 1971). After determination of its chemical structure, this aa was named hypusine, based on its two structural components, hydroxyputrescine and lysine (Shiba et al, 1971). Hypusylation is formed exclusively involving two enzymatic steps (Fig. 5.2.4) catalysed by deoxyhypusine synthase (DHS) and deoxyhypusine hydroxylase (DOHH) enzymes (Park et al, 2010). Of great interest is the high sequence conservation of each of the three proteins (eIF5A, DHS and DOHH) shown in figure 5.2.5. In the first step of the hypusine-reaction, DHS catalyzes the transfer of the 4-aminobutyl moiety of spermidine to the ε-amino group of lysine residue of eIF5A precursor to form an intermediate, deoxyhypusine [Nε-(4-aminobutyl)-lysine] residue. This intermediate is subsequently hydroxylated by DOHH (Abbruzzese et al, 1986) to complete hypusine synthesis and eIF5A activation (Zanelli et al, 2007). eIF5A and its deoxyhypusine/hypusine modification are vital for eukaryotic cell proliferation (Gerner et al, 1986; Chen and Liu, 1997; Park, 2006).

Figure 5.2.3. Ribbon diagram and representative sketch of the structure of aIF5A from M. jannaschii. (a) The sketch shows the eleven β- sheets in two domains of the molecule joined by flexible links that contain the Lys-54 site of hypusine modification. (b) Ribbon structure of the eIF5A, showing the two domains of the molecule linked by the flexible linker that harbors the hypusine residue. The molecule bears opposite charges in the C- and N-terminal ends (Kim et al, 1998).

Figure 5.2.4. The hypusine synthesis pathway (Park et al, 2006). Introduction 41

Mutations at the hypusine attachment site Lys50 in human eIF5A completely blocked deoxyhypusine synthesis whereas substitutions in its vicinity resulted in reduced efficiency of deoxyhypusine synthesis or inhibition of the hydroxylation reaction catalyzed by DOHH (Cano et al, 2008). A truncated peptide consisting of 80 residues of human eIF5A was nearly as good a substrate as the full-length protein for hypusine attachment (Kang et al, 2007).

Figure 5.2.5. Amino acid sequence conservation of eIF5A, DHS and DOHH in eukaryotes. In each case (eIF5A-1, DHS and DOHH), the human sequence is shown. The degree of conservation is indicated by color coding: red, 100% identity; dark orange to yellow, conservative replacements (e.g. F, Y; D, E; K, R, L, V, I, M; T, S; A, G, C, S) with >80 to >50% sequence identity; white, no significant sequence identity. The diagram is based on eukaryotic sequences (40 for eIF5A, 36 for DHS and 14 for DOHH) chosen from a range of eukaryotic phyla and species. Critical amino acids for binding of substrates are shown by symbols under the residues (Wolff et al, 2007).

5.3. EF-P Modification Pathway in E. coli

The unique hypusine modification (Shiba, 1971) is a strictly conserved aminobutyl moiety of spermidine attached to lysine K50 of domain I of eIF5A and has been found in all eukaryotes examined so far (Park et al, 2010). In addition, it also occurs in certain archaea (Bartig et al, 1992), but has not been detected in bacteria. Interestingly, the equivalent lysine 34 (K34) of endogenous E. coli EF-P has been reported to carry a modification of ~144 Da (Aoki et al, 2008). Bound to the ribosome, K34 of EF-P, and by analogy the hypusine of eIF-5A, would interact with the CCA-end of the P-tRNA, suggesting that EF-P and eIF-5A stimulate peptide bond formation through stabilization and positioning of the acceptor end of the P-tRNA (Blaha et al, 2009). Introduction 42

In E. coli two unrelated enzymes, YjeK and YjeA (also termed PoxA/GenX), was proved to be implicated in the EF-P modification pathway (Bailly et al, 2010; Ambrogelly et al, 2010; Navarre et al, 2010; Yanagisawa et al, 2010; Roy et al, 2011; Park et al, 2012). YjeK is a lysine 2,3-aminomutase that converts (S)-α-lysine to (R)-β-lysine (Behshad et al, 2006), whereas YjeA has homology to class II lysine-tRNA synthetases that lacks the tRNA anticodon recognition domain (Bailly et al, 2010). Based on published data, deletion of yjeA or yjeK in Salmonella strains leads to growth defects, antibiotic sensitivity and attenuation of virulence (Kaniga et al, 1998; Bearson et al, 2006; Navarre et al, 2010; Bearson et al, 2011; Zou et al, 2012). Furthermore, in vitro biochemical assays indicate that YjeA cannot aminoacylate tRNALys (Ambrogelly et al, 2010), but can activate lysine and transfer it to K34 of EF-P (Ambrogelly et al, 2010; Navarre et al, 2010; Yanagisawa et al, 2010; Roy et al, 2011; Park et al, 2012). Recently, (R)-β-lysine was shown to be 100-fold more efficient as a substrate for lysinylation of EF-P by YjeA than either (S)-β-lysine or α-lysine (Roy et al, 2011). Curiously, mass spectrometry (MS) analyses from recently published data indicate that in vitro lysinylation of K34 of EF-P by YjeA results in a ~128 Da modification (Navarre et al, 2010; Yanagisawa et al, 2010; Roy et al, 2011; Park et al, 2012) leaving a difference of ~16 Da unexplained, when compared with the ~144 Da reported for endogenous EF-P (Aoki et al, 2008).

Figure 5.3.1. Model for YjeA, YjeK, and EF-P Function. YjeA catalyzes aminoacylation (with lysine) of the same conserved lysyl residue of EF-P. YjeK is shown here modifying the Lys-Lys side group to generate a fully active form of EF-P. Alternatively, YjeK may act on the lysine substrate prior to its ligation to EF-P (Navarre et al, 2010).

With the fact that the currently accepted EF-P modification pathway (Navarre et al, 2010) (Fig. 5.3.1) is incomplete, this ~16 Da unexplained mass difference was studied by Peil et al. recently, showing that in vivo endogenous EF-P carries a unique ε(R)-β-lysylhydroxylysine modification of ~144 Da, thus, leading to the existence of a new, unidentified protein, which was the third missing enzyme resulting in the fully modified EF-P in vivo (Peil et al, in press). For this reason, based on immuno-precipitation methods, coupled with high resolution MS, they demonstrate firstly, that lysinylation of EF-P by YjeK and YjeA results in the addition of β-lysine to the ε-amino group of K34 (Peil et al, in press). Additionally, they reported that fully modified EF-P requires the presence of YfcM, a hydroxylase, as the last enzyme in the EF-P modification pathway and proposed the potential mechanism for the post-translational modification of EF-P (Fig. 5.3.2A) (Peil et al, in press). For this experiment chymotrypsin proteolysis of EF-P from the E. coli AT713ΔyfcM strain (auxotrophic to lysine and arginine) produced a fragment containing K34 with a mass increase of +128.09 Da compared to the unmodified EF-P (Peil et al, in press). Introduction 43

When yfcM was exogenously overexpressed from a plasmid (pYfcM) in the AT713ΔyfcM strain, the mass of the chymotrypsin fragment was +144.09 Da larger than the unmodified fragment and identical to the mass increase observed for endogenous EF-P (Peil et al, in press). These experiments were also repeated in the presence of LysC fragments, leading to the same mass differences compared to unmodified EF-P (Peil et al, in press). Based on this study, YfcM seems to be dependent on the action of YjeK and YjeA, since the absence of either of these genes leads to a predominance of completely unmodified EF-P in vivo. Particularly, the lack of +16 Da modified EF-P peptides in any of the ΔyjeK or ΔyjeA MS spectra suggested that YfcM cannot hydroxylate unmodified EF-P and consequently acts after YjeK and YjeA (Peil et al, in press). It should also be mentioned that the +144 Da modified peptide was never detected in the ΔyjeK strain (Peil et al, in press). This proves that α-lysinylated EF-P is a poor substrate for YfcM and β- lysinylated EF-P is preferred, a situation that shares similarities to the eukaryotic situation where deoxyhypusinylation of eIF-5A by DHS is a prerequisite for the subsequent hydroxylation by DOHH (Park et al, 2010) (Fig. 5.3.2B). In contrast to eIF-5A where the added spermidine moiety is hydroxylated by DOHH, YfcM was proved to hydroxylate the C4 or C5 position of K34 of EF-P, but not the added β-lysine (Peil et al, in press). Further, based on T. thermophilus structure (Blaha et al, 2009) K34 and the lysinylation moiety extend towards the PTC and stabilize the CCA-end of the tRNA (Fig. 5.3.2C) (Peil et al, in press).

Figure 5.3.2. (A) Modification pathway of EF-P. The conserved K34 of endogenous E. coli EF-P bears a 144.09 Da modification, which results from two distinct reactions, namely a 128.09 Da lysinylation performed by YjeK and YjeA, followed by an additional 16 Da hydroxylation by YfcM. (B) Modification pathway of eIF-5A. (C) Relative positions of K34 of EF-P (green) and the acceptor stem of the P-tRNA (blue) when bound on the ribosome. The γ, δ and ε carbon atoms of K34 and the C74, C75 and A76 positions of the tRNA are indicated (Peil et al, in press).

5.4. YjeK, YjeA and YfcM: Enzymes Modifying E. coli EF-P

5.4.1. Lysine-2-3-aminomutase

The enzyme lysine-2-3-aminomutase (LAM), encoded by the gene kamA, was first identified in Clostridium subterminale SB4, has a molecular weight of about 285.000, and is composed of six subunits with a molecular weight of about 47.000 (Song and Frey, 1991). LAM contains an iron-sulphide cluster and is extremely oxygen sensitive, according to Frey and co- workers (Chen et al, 2000). The enzyme catalyzes the interconversion of (S)-α-lysine to (S)-β- Introduction 44

lysine (Behshad et al, 2006), which is the initial step in the metabolism of lysine as a source of carbon and nitrogen in Clostridia (Chirpich et al, 1970). In other organisms, β-lysine is used in cellular defense mechanisms, such as the production of antibiotics (Grammel et al, 2002). The amino group is transferred from the α- to the β-carbon atom, as shown in figure 5.4.1.1. The reaction is typical of adenosylcobalamin-dependent enzymes; however, unlike B12-dependent enzymes, LAM does not contain adenosylcobalamin and is not activated by cobamide coenzymes (Petrovich et al, 1992; Chen et al, 2000).

Figure 5.4.1.1. Reaction catalyzed by LAM (Petrovich et al, 1992).

Barker and co-workers purified the enzyme to near homogeneity and examined the molecular properties and cofactor requirements (Chirpich et al, 1970). LAM is a member of the “radical SAM” superfamily, a highly diverse family of enzymes, known members of which use S- adenosylmethionine (SAM) as a radical initiator in catalysis (Sofia et al, 2001). The enzyme is activated by SAM, which is necessary for the transfer of the hydrogen between the a- and β- carbons of lysine and β-lysine that is mediated by a radical mechanism (Moss and Frey, 1987; Baraniak et al, 1989). The LAM consists of an iron-sulfur cluster [4Fe – 4S], which keeps the structural integrity of the protein and is important for the function of the aminomutase (Petrovich et al, 1992). For the activity of the protein also pyridoxal 5’-phosphate (PLP) is required (Ruzicka et al, 2007). The crystal structure of LAM of C. subterminale SB4 has been solved to 2.1 Å resolution by Ringe and co-workers (Lepore et al, 2005). Regarding to this, LAM crystallizes as a homotetramer in the asymmetric unit. The homotetramer consists of a dimer of tightly associated dimers (Fig. 5.4.1.2) (Lepore et al, 2005).

Figure 5.4.1.2. Tetrameric packing of LAM subunits. Two views of the tetramer at 0° and rotated by 90°. Each subunit is represented as a separate surface and color. Thus, one domain-swapped, zinc-coordinated dimer corresponds to the red/green or yellow/blue subunit combinations (Lepore et al, 2005). Introduction 45

Currently, there are 53 prokaryotic proteins with aa sequences homologous to that of LAM from C. subterminale SB4. The aligned amino acid sequences of the E. coli protein encoded by the gene yjeK and of clostridial LAM shows 33 % identities, and the E. coli protein is one of the least similar to clostridial LAM (Behshad et al, 2006). Unlike kamA in C. subterminale, the gene yjeK does not appear in a gene cluster associated with lysine metabolism. In E. coli, yjeK is flanked by genes annotated as elongation factor protein, GroEL/ES chaperone proteins, membrane proteins and others that seem to be expressed under stress or in the stationary phase (Behshad et al, 2006). The product of yjeK is a LAM with low activity relative to clostridial LAM, and it functions by an analogous but stereochemically variant mechanism. YjeK catalyzes the conversion of (S)- α-lysine to (R)-β-lysine (Behshad et al, 2006). The low activity of the E. coli YjeK compared to the clostridial LAM may indicate that (S)-β-lysine might not be the real in vivo substrate, or that β-lysine is required in low amounts for the production of a specialized molecule (Behshad et al, 2006). Because the functions of variant forms of the same enzyme in different organisms are often different, they frequently display different activities. So, classic case is the alcohol dehydrogenases from horse liver and yeast, which differ by 100-fold in catalytic activity, yet they are structurally and apparently mechanistically similar (Behshad et al, 2006). A similar difference in activity has been also noted for LAM from C. subterminale and B. subtilis (Chen et al, 2000), and LAM from E. coli which is even less active than the Bacillus enzyme. Moreover, sequence alignment analysis revealed that the YjeK that clusters with the efp gene can be separated from the canonical LAM involved in Lys degradation pathway. The major difference between the two families is that the YjeK proteins lack the C-terminal multimerization domain present in the LAM family of proteins (Fig. 5.4.1.3) (Lepore et al, 2005). Additionally it was shown recently, that YjeK plays an important role in the pathway for the modification of a lysine residue in protein chain elongation factor EF-P (Bailly and de Crécy-Lagard, 2010).

Figure 5.4.1.3. Structural analysis of YjeK and LAM. (Right) 3D structure of LAM from Clostridium subterminale SB4 (Lepore et al, 2005) in blue with the C-terminal multimerization domain in pink. (Left) 3D-model of YjeK from Acinetobacter baylyi based on C. subterminale SB4 (Bailly and de Crécy-Lagard, 2010).

5.4.2. Lysyl-tRNA synthetase (LysRS) Introduction 46

Aminoacyl-tRNA synthetases (aaRS) play a major role in translation during protein synthesis, since they are a class of enzymes that charge tRNAs with their cognate amino acids (Hausmann and Ibba, 2008). AaRSs catalyze a two-step reaction. In the first step they activate their amino acid by forming an aminoacyl-adenylate, in which the carboxyl of the amino acid is linked in to the α-phosphate of ATP, by displacing pyrophosphate. Second, the enzyme-bound reaction intermediates are transferred to the 3’ acceptor end of the tRNAs docking onto their active sites (Hausmann and Ibba, 2008). Because tRNAs cannot distinguish amino acids conjugated to their ends, the correct recognition of aa and tRNAs by these enzymes is a crucial determinant to maintain the fidelity of protein synthesis. The energy required for the formation of the ester bond between the amino acid carboxylate group and the tRNA acceptor stem is supplied by coupling the reaction to the hydrolysis of ATP (Onesti et al, 2000). There are two classes of aminoacyl-tRNA synthetase depending on the amino acid they are responsible for.

 Class I lysyl-tRNA synthetase (LysRS1) has two highly conserved sequence motifs. Class I enzymes are generally (though not always) monomeric, and attach the carboxyl of their target amino acid to the 2' OH of adenosine 76 in the tRNA molecule (Perona et al, 1993; Delarue, 1995).

 Class II lysyl-tRNA synthetase (LysRS2) has three highly conserved sequence motifs. Class II enzymes are generally dimeric or tetrameric, and attach their amino acid to the 3' OH of their tRNA, except for phenylalanine-tRNA synthetase which uses the 2' OH (Perona et al, 1993; Delarue, 1995).

Except for their regular function, it was shown, for many aaRSs, that they have diverse function, as an elaborate network of protein–protein interactions is required for efficient translation in all domains of life. AaRSs must correctly bind a diverse array of molecules to perform their essential function of aminoacylation, including ATP, aa, tRNAs, and in some cases other aaRSs. Also, several aaRSs associate with additional or auxiliary protein factors that are primarily used in cellular tasks beyond translation (Hausmann and Ibba, 2008). Although many of these associations were first described in eukaryotic cells, numerous multienzyme complexes containing aaRSs have recently been identified in both bacteria and archaeal. Bacterial aaRSs are generally thought to function as stand-alone proteins to carry out the fundamental task of aminoacylation. However, a handful of binary complexes comprised of one aaRS and a second protein factor have recently been discovered in bacteria, like for example the complex formation between YbaK and prolyl-tRNA synthetase (ProRS), which is essential for efficient hydrolysis of mischarged Cys-tRNAPro (Hausmann and Ibba, 2008), the aspartyl-tRNA synthetase, which stably associates with amidotransferase in the presence of tRNAAsn, forming the transamidosome complex (Hausmann and Ibba, 2008) and the tryptophanyl-tRNA synthetase, which has been shown to associate with nitric oxide synthase (NOS) (Buddha et al, 2004a, b). Although binary complexes containing one aaRS are generally the norm in bacteria, archaea and eukaryotes have been found to harbor higher order complexes composed of Introduction 47

multiple aaRS activities in association with a variety of cellular factors. In archea ProRS has been found to associate in M. jannaschii with a metabolic protein, Mj1338 (Lipman et al, 2003), while lysyl-tRNA synthetase, leucyl-tRNA synthetase and prolyl-tRNA synthetase associate in order to form a stable complex for enhanced aminoacylation of LeuRS (Praetorius-Ibba et al, 2007). In eukaryotes these complexes tend to be larger than those discovered in bacteria and archaea and also perform a wider range of functions from aminoacylation to noncanonical functions beyond translation. In S. cerevisiae glutamyl- and methionyl-tRNA synthetases (GluRS and MetRS, respectively) stably interact with aminoacyl-tRNA synthetase cofactor I (Arc1p), a nonsynthetase accessory protein (Simos et al, 1996, 1998; Galani et al, 2001; Karanasios et al, 2007). In addition to the Arc1p : GluRS : MetRS complex, S. cerevisiae also harbors a complex between seryl-tRNA synthetase (SerRS) and Pex21p, a protein involved in peroxisome biosynthesis (Rocak et al, 2002). A connection between tyrosyl-tRNA synthetase (TyrRS) and a protein involved in cell wall biosynthesis (Knr4) has also been identified in S. cerevisiae from DNA microarray, yeast two hybrid, and pull-down experiments (Ivakhno and Kornelyuk, 2005).

Figure 5.4.2.1. Structural analysis of YjeA and LysRS2. (Right) Class II lysyl-tRNA synthetase from E. coli (Onesti et al, 2000) in purple. (Left) 3D structure of YjeA from Salmonella typhimurium in pink and (Bailly and de Crécy-Lagard, 2010).

In E. coli there were found 2 distinct genes, which encode LysRS. lysS, which is expressed constitutively and lysU, whose expression can be induced (Emmerlich and Hirshfield, 1987) and is expressed under certain physiological stress conditions, like heat shock (Onesti et al, 1995). YjeA from E. coli, a protein that specifically aminoacylates EF-P with lysine (Zou et al, 2012; Yanagisawa et al, 2010), has homology to LysRS2 family of enzymes that catalyze the addition of lysine to its cognate tRNALys (Bailly and de Crécy-Lagard, 2010; Zou et al, 2012), but a number of studies have failed to show that YjeA can aminoacylate a tRNA (Kong et al, 1991; Kaniga et al, 1998; Ambrogelly et al, 2010). The analysis of the three dimensional structure of E. coli YjeA (Fig. 5.4.2.1) revealed that YjeA lacks the tRNA anticodon recognition domain and contains only the catalytic core of LysRS2 deprived of the anticodon binding domain (ABD) Introduction 48

(Bailly and de Crécy-Lagard, 2010). This absence of the ABD, points to a function different from tRNA aminoacylation as already observed for other aaRSs catalytic core homologs (Roy et al, 2003). Alignments of the LysU, LysS and YjeA also display that the YjeA is shorter and lacks one domain, as shown in figure 5.4.2.2. In addition, it was shown recently that in agreement with its well-conserved lysine binding site, YjeA can activate in vitro L-lysine and lysine analogs (Ambrogelly et al, 2010). Moreover, it was reported that YjeA can catalyze the ATP-dependent ligation of lysine to EF-P by the addition of β-lysine to the ε-amino group of K34 (Roy et al, 2011; Peil et al, in press). The shape of the protein substrate mimics that of the L-shaped tRNA (Yanagisawa et al, 2010). Furthermore, in vivo analyses revealed that EF-P lysinylation by YjeA is enhanced by YjeK (Yanagisawa et al, 2010) and Navarre and co-workers also suggested an interaction between yjeA and yjeK gene that is critical for virulence and stress resistance in Salmonella enterica (Navarre et al, 2010; Zou et al, 2011). Salmonella poxA (or yjeA) and yjeK mutants display nearly identical phenotypes, including increased resistance to S-nitrosoglutathione (GSNO) and hypersusceptibility to a large number of unrelated antimicrobial compounds (Navarre et al, 2010; Bearson et al, 2011; Zou et al, 2012). These mutant strains also display markedly reduced virulence in a mouse model of infection (Kaniga et al, 1998; Navarre et al, 2010). The mechanisms underlying the pleiotropic phenotypes of poxA mutants have remained unresolved for over 2 decades (Zou et al, 2012).

Figure 5.4.2.2. Alignment of protein sequences of the three isoforms of lysyl-tRNA synthetases. Protein sequences (from NCBI) were used for alignment. Alignment was done using ClustalW2 software (http://www.ebi.ac.uk/Tools/clustalw2/index.html). Stars show amino acids which are common in all three proteins. Dots display similarity of the properties of amino acids. 5.4.3. YfcM, a Hydroxylase

Introduction 49

In order to identify the missing enzyme in the modification pathway of EF-P, Peil et al. searched proteins associated with YjeA, YjeK and EF-P, using the STRING (Search Tool for the Retrieval of Interacting Genes/Proteins) database (http://string.embl.de). A query using E. coli PoxA (YjeA), revealed an association of EF-P, YjeK and three additional proteins: YfcM, HflD and ECs0957. From literature it was known that HflD is involved in the λ lysis-lysogeny switch and that ECs0957 has homology with pyruvate dehydrogenases. In contrast, BLAST searches revealed that YfcM has no reasonable homology with any protein of known structure or function (Peil et al, in press). To investigate the potential involvement of YfcM in the modification of EF-P in vivo, MS experiments were carried out revealing YfcM as the third enzyme in EF-P modification, and as a potential hydroxylase (mono-oxygenase) that modifies K34 of lysinylated EF-P (Peil et al, in press). Moreover, it is suggested that YfcM is more likely to use a NAD- or FAD-based cofactor similar to other bacterial lysine hydroxylases, rather than Fe2+ or α-ketoglutarate/ascorbic acid co-factors as used by eukaryotic lysine hydroxylases (Peil et al, in press). Additionally, fragmentation data show that YfcM hydroxylates either the C4 (γ) or C5 (δ) position, which is the first report of C4/C5-hydroxylase activity in bacteria (Peil et al, in press).

Figure 5.4.3.1. Characterization of the hydroxylase YfcM. Phylogenetic tree (left) and genomic neighborhood (right) of representative YfcM proteins in bacteria. Only bootstrap support values >50% are shown (Peil et al, in press).

Previous bioinformatic analysis of 725 bacterial genomes revealed the presence of efp, yjeA and yjeK genes in 200 genomes (28%) (Bailly and de Crecy-Lagard, 2010). Recent further Introduction 50

extended analysis by Peil et al. detected the presence of YfcM protein in 179 species clustering within the γ-proteobacteria phyla out of 1268 completely sequenced bacterial genomes (Fig. 5.4.3.1) (Peil et al, in press). All bacterial species containing yfcM gene have also efp, yjeA and, with one exception (Haemophilus influenzae strain PittGG), yjeK (Peil et al, in press). Moreover, a lysine, equivalent to position 34 in E. coli EF-P, is strictly conserved in organisms that contain YfcM and yfcM gene is always located in a distinct region of the chromosome and has a prmC- aroC-mepA-yfcA-yfcM-yfcL operon structure, which is highly conserved within the Enterobacteriaceae family (Fig. 5.4.3.1) (Peil et al, in press). Except for that, yfcM seems to neighbor other modification enzymes, such as the methyltransferases prmB and prmC, which modify ribosomal protein L3 and release factor RF1/RF2, respectively, as well as mnmC and yrdC (rimN) which are involved in tRNA modification (Bujnicki et al, 2004; Harris et al, 2011) (Fig. 5.4.3.1) (Peil et al, in press). Finally, the taxonomic distribution shows that yfcM is present with the yjeK and yjeA genes in most common Gram-negative pathogenic bacteria, such as Acinetobacter baumannii, Escherichia coli O157:H7, Haemophilus influenzae, Klebsiella pneumoniae, Legionella pneumophila, Salmonella typhimurium, Vibrio cholerae and Yesinia pestis (Fig. 5.4.3.1) (Peil et al, in press).

Aim of the Project 51

6. Aim of the Project

Based on a complete description of the EF-P modification and pathway (submitted Peil et al, 2011), in this project we focused on further studies to address the mechanism of action of EF-P and especially to investigate how the different stages of EF-P’s modifications can affect E. coli cells.

1. Using E. coli Keio Knockout Collection (Δefp, ΔyjeK, ΔyjeA, ΔyfcM) and E. coli Keio parental strain (wild-type) as control, we are investigating the effect of the deletion strains on the cells under different environmental stress conditions:

 varying growth temperatures,  varying nutrition conditions, and  susceptibility to antibiotics, showing that those mutations affect the in vivo activity of EF-P and that E. coli efp mutants show sensitivity to non-ribosomal inhibitors, suggesting a secondary effect of EF-P on the cell-membrane integrity under stress conditions or stationery phase.

2. Moreover, by a rescue experiment, we transformed Δefp, ΔyjeK, ΔyjeA, ΔyfcM with an efp-, yjeK-, yjeA- and yfcM-containing plasmid, respectively, in order to test the ability of deletion strains to recover in presence of the appropriate plasmid.

3. In order to restore the wild-type phenotype, different K34 mutants, prepared by site directed mutagenesis were used.

4. Motility and virulence are often linked together (Josenhans and Suerbaum, 2002). EF-P was previously reported to be important for bacteria motility in B. subtilis (Kearns et al, 2004). Here we are performing motility tests with triphenyltetrazolium chloride (TTC) for the deletion strains, in order to determine if they are motile or not.

5. Using Confocal Laser Scanning Microscopy (CLSM) and fluorescence staining we monitored differences in morphology of the E. coli deletion strains.

6. Further, so as to identify the uncharacterized protein, YfcM, a pilot experiment was performed, where YfcM was cloned into pET21b, purified using Ni-NTA agarose beads, and purified proteins were further purified using the gel filtration column Superdex G75.

7. We cloned half of the polycistronic “- yjeK - yjeA - yfcM - His-efp –“, which was desinged in order to overexpress fully modified EF-P in BL21 expression strain, crystallize it, see the crystal structure of the whole modification pathway of EF-P and study better the function of EF-P in translation extracts from different mutants.

Materials 52

7. Materials

7.1. Chemicals

Chemicals which are not listed came from Roth. Buffers used are listed in the methods part (see Methods).

Chemicals Supplier Acetic acid Roth, Karlsruhe APS Sigma-Aldrich, München Bromophenol Blue Merck, Darmstadt DMSO Merck, Darmstadt Ethanol Merck, Darmstadt Glycerin Merck, Darmstadt Glycerol Merck, Darmstadt Isopropanol Merck, Darmstadt Kalioum Chlorid Merck, Darmstadt NaCl Merck, Darmstadt NaH2PO4 Merck, Darmstadt Propidium iodide (Pi) Invitrogen Rotiphorese Gel 30 Roth, Karlsruhe SDS SERVA, Heidelberg SDS-Page Standard New England BioLabs, Frankfurt/Main Spermine Fluca, München SYTO9 Invitrogen TEMED Roth, Karlsruhe Triphenyltetrazolium chloride (TTC) Sigma- Aldrich, München Tris Roth, Karlsruhe β-Mercaptoethanol Sigma- Aldrich, München

7.2. Bacterial Strains

 In all experiments E. coli Keio parental strain BW25113 (wild-type) was used as reference (control).  Experiments were conducted using the E. coli Keio Knockout Collection, which is a set of precisely defined, single gene deletion mutants of all nonessential genes in E. coli K12 (Baba et al, 2006). Δefp (JW4147), ΔyjeK (JW4106), ΔyjeA (JW4116) and ΔyfcM (JW5381) strains were obtained from the National BioResource Project (NBRP) at the National Institute of Genetics (NIG), Japan (Baba et al, 2006)) and contained one antibiotic resistance marker, kanamycin. The ΔyjeK strain was taken from plate 38, since the isolate on plate 37 is not a bone fide yjeK knock-out strain (Yamamoto et al, 2009).  The cloning strain E. coli NovaBlue (Genotype: endAI hsdR17 (rK12-mK12+) supE44 thi-1 recA gyrA96 re1Al lac F’[proA+B+lac1qZDM15::Tn10] (TetR)) was obtained from Novagen. Materials 53

 The expressing strain E. coli BL21 (Genotype: F-; ompT; gal [dcm] [lon]; hsdSB (rB-, mB-); λ (DE3) +/- pLysE (CAMR)) was obtained from Novagen.

7.3. Growth Media

Media Components 10 gr Peptone

Luria Bertani Media (LB) (V = 1 lt) 5 gr Yeast Extract 10 gr NaCl 16 gr Trypton 2xTY Media (V = 1 lt) 10 gr Yeast Extract 5 gr NaCl 12 gr Trypton

Terrific Broth Media (V = 1 lt) 24 gr Yeast Extract 9.4 gr Dipotassioum phosphate 2.2 gr Monopotassioum phosphate 3 gr Meat Extract

Swimming Media M1 (V = 1 lt) 5 gr Tryptone 5 gr NaCl 0.3%Agar 15 mM (NH4)2SO4 Minimal Media I 8 mM MgSO4 27 mM KCl 7 mM Na-citrate 50 mM Tris/HCl, pH 7.4

0.6 mM KH2PO4 autoclave 2 mM CaCl2 10 μM MnSO4 1 μM FeSO4 added after autoclaving 0.2 % carbon source (autoclaved, glucose or glycerol) M9 salts:

Minimal Media M9 (5x, 250 ml) 16 gr Na2HPO4 3.75 gr KH2PO4 0.625 gr NaCl

1.25 gr NH4Cl autoclave 0.5 ml of 1M MgSO4 25 μl of 1M CaCl2 added after autoclaving 5 ml of 0.2 % carbon source (autoclaved glucose or glycerol)

Materials 54

7.4. Equipment

Equipment Company

96-well flat bottom Microtiter Plates Corning Incorporated

Centrifuge 5415 D Eppendorf, Hamburg

Centrifuge 5417 R Eppendorf, Hamburg

Confocal Laser Scanning Microscope LSM 710, Zeiss Zeiss

Electrophoresis Power Supply: EPS 301 Amersham Pharmacia Biotech

Gradient Stationip Biocomp

Heidolph REAX 2 Heidolph Instruments GmbH & co.KG

Incubator Shaker Series InnonaR 44 New Brunswick Scientific GmbH

Microfluidizer Microfluidics International Corporation, USA

MJ Research PTC – 200 Peltier Thermal Cycler BioZym Diagnostic GmbH

NanoDrop (ND-1000) Spectrophotometer Peqlab Biotechnologie GmbH, Erlangen

Petri plates Sarstedt AG & Co, Germany

SF-1-V Thomtech Labortechnik und Industrie

Sorvall EvolutionRc Thermo Scientific

TECAN Infinite M1000 Tecan Group Ltd., Switzerland

Thermomixer Comfort Eppendorf, Hamburg

Unichromat 1500 UniEQuiP GmbH

7.5. Plasmids

The efp, yjeK, yjeA, yfcM genes were PCR amplified from gDNA of E. coli K-12 strain MC4100 using the primers listed in Supplementary Table 1. For all experiments either the pET- system (Novagen), the pQE-system (Qiagen) or the pRSFDuet-system (Novagen) were used. The Materials 55

RSFDuet vector has a kanamycin resistance gene (grown in 30 μg/ml Kan), whereas the pET21b and pQE70 vectors contain an ampicillin resistance gen (grown in 100 μg/ml Amp). pET-system was used for the overexpression of YfcM, while all three vector-systems were used in order to clone the polycistronic construct (see Methods below). All the vector maps and sequences of the proteins can be found in Supplementary Table 2.

7.6. Primers

All primers are listed Supplementary Table 1 in appendix and were used in a concentration of 10 pmol/μl for PCR reactions. For sequencing the concentration of the primers was 2 pmol/μl.

Methods 56

8. Methods

8.1. Microbiological Analysis

In the following experiments E. coli Keio parental strain was used as control in the same conditions as the E. coli Keio knockout clones, but was grown without the presence of any resistance marker. Overnight cultures were grown in falcons (15 ml), while fresh cultures were prepared in 96-well microtiter plates, containing 250 μl LB media, 10 μl of the overnight cultures and the respective antibiotic when needed. Growth was monitored by measuring the optical density (OD) in TECAN Infinite M1000 at a wavelength of 600 nm (OD600) and the final graphs were prepared in SigmaPlot 11.0. The sequences of YjeK, YjeA, YfcM and EF-P proteins from NCBI are listed in Supplementary Table 3.

8.1.1. Growth Analysis of E. coli Keio Collection. E. coli Keio knockout clones (Δefp, ΔyjeK, ΔyjeA, ΔyfcM) were grown overnight in 4 ml LB media and 25 μg/ml kanamycin (kan). o Fresh cultures were grown in 96-well microtiter plate and incubated at 37 C. OD600 was measured almost every hour and when stationary-phase was reached, incubation stopped, and a growth curve of OD600 vs. time was prepared.

8.1.2. Growth under Stress Conditions

8.1.2.1. Growth of Cells in Various Media. E. coli Keio knockout clones were grown under varying nutrition conditions till stationary-phase. The cultures were grown in rich media, and also in minimal media with glucose and glycerol as carbon source. The OD600 was measured periodically and graphs of OD600 vs. time for each media were prepared.

8.1.2.2. Temperature-sensitive Growth in Various Media at 37oC and 44oC. In order to test the rate of growth in various temperatures E. coli Keio knockout clones were grown in different media (LB, 2xTY, M9 glucose, M9 glycerol, MM glucose and MM glycerol) and the cells were incubated at the beginning at 37 oC. After about 2 hrs the temperature o was shifted to 44 C and the cultures were grown another 2 hrs. OD600 was measured and a growth curve of OD600 vs. time for each strain was prepared.

8.1.2.3. Antibiotic Susceptibility Assay. In order to test the susceptibility of various antibiotics targeting the ribosome on the E. coli Keio knockout clones, the minimum inhibitory concentration (MIC) was determined in a 96-well microtiter plate. MIC in microbiology is the lowest concentration of an antimicrobial agent that will inhibit the visible growth of a microorganism after incubation. The assay was carried out in standard, sterile 96- well microtiter plates. The method is based on dilutions where, bacteria are inoculated into a liquid growth media and they are tested for their ability to produce visible growth in microplate wells of broth containing different dilutions of an antimicrobial agent. For this reason a stock solution of each antimicrobial under test was prepared. Methods 57

From the stock solution, eleven ‘principle’ dilutions were made (range between 0.5 – 1024 μgr/ml) using either sterile mQ H2O or 0.75% NaCl as a solvent. The assay was started by adding 250 μl LB media, 10 μl of overnight bacterial suspension incubated at 37 oC and 10x fold of each antibiotic’s dilution into the wells. The microplate was incubated in Thermomixer comfort (Eppendorf, Hamburg) shaking at 600 rpm at 37 oC. Growth was assessed after incubation for a defined period of time (16–20 hrs) and the antimicrobial activity of the compounds was determined by following the bacterial growth as optical density of the suspension on the plate reader at wavelength of 600 nm. Between the measurements, the incubation was continued in room temperature and final graphs were presented with SigmaPlot 11.0.

8.1.3. Rescue Experiments

The gene expressed from a recombinant construct can restore the wild-type phenotype. This can provide us conclusive evidence that knockout of a targeted gene, and not an off-target effect, is responsible for an observed phenotype. In this study, we used the E. coli Keio knockout clones and E. coli Keio parental strain as references and transformed the following knocked out strains: Δefp, ΔyjeK, ΔyjeA and ΔyfcM with an efp-, yjeK-, yjeA- and yfcM- containing plasmid, respectively. Cultures from the above strains were prepared in 15 ml falcons containing 5 ml LB media plus 25 μg/ml Kan and incubated overnight at 37 oC. Next morning dilutions (1:50) of the overnight cultures were prepared in fresh falcons and 20 μl of each dilution was added in 250 μl LB media in the 96-well microplate. The cells grew in o Thermomixer comfort shaking at 600 rpm at 37 C. After 4.15 hrs OD600 was measured in plate reader, the overnight cultures of the first day were left shaking in the incubator and the assay was repeated the same way as above for 4 days in a row.

8.1.4. Motility Test

Motility Test in a semi-solid agar is used to demonstrate if an organism is motile or non- motile and to determine if organisms are motile by means of flagella. The location of the flagella is determined by the bacterial species. Non-motile bacteria do not possess flagella. The production of flagella is also subject to culture conditions; for example, some bacteria are motile at different temperatures from those at which they are normally incubated. Motility Test is a modification of the formula of Tittsler and Sandhoizer. The medium contains M1 media, agar and TTC. TTC (2,3,5-triphenyltetrazolium chloride) is a redox indicator used to differentiate between metabolically active and inactive tissues. It is a soluble compound which is taken up by the bacterial cells. The white compound is enzymatically reduced to red TPF (1,3,5- triphenylformazan) in living bacteria and tissues due to the activity of various dehydrogenases (enzymes important in oxidation of organic compounds and thus cellular metabolism). Once the substance has been absorbed by the bacteria, it is reduced, releasing the acid formazan, a highly pigmented red, insoluble compound. Motility is indicated by the lateral spread of a pink color (formazan) throughout the media. The semi-solid agar was composed of M1 media, 0.37% agar and 0.01% TTC. Prior to inoculation, the media should be brought to room temperature. Here we are using a multipoint inoculation system in a square petri plate. Each plate contains Methods 58

20 ml semi-solid agar and a drop of liquid bacterial overnight culture. Inoculation was done using a straight sterile needle. The plates were examined for evidence of motility after overnight incubation at 37 oC.

8.2. Confocal Laser Scanning Microscopy

To determine the morphology of E. coli Keio knockout clones we used Confocal Laser Scanning Microscopy (CLSM). Confocal microscopy is a technique for obtaining high- resolution optical images with depth selectivity (Pawley, 2006). The principle of confocal imaging was originally patented by Marvin Minsky in 1961, but it took another thirty years and the development of lasers for CLSM to become a standard technique toward the end of the 1980s (Pawley, 2006). This CLSM design combined the laser scanning method with the 3D detection of biological objects labeled with fluorescent markers. The key feature of confocal microscopy is its ability to acquire in-focus images from selected depths, a process known as optical sectioning. Images are acquired point-by-point and reconstructed with a computer, allowing three-dimensional reconstructions of topologically complex objects. A conventional microscope "sees" as far into the specimen as the light can penetrate, while a confocal microscope only images one depth level at a time. In effect, the CLSM achieves a controlled and highly limited depth of focus. CLSM is actually a conventional microscope which is equipped with a laser light source, the laser scanning head and an automatic focusing stage connected to a monitor and PC (Fig. 8.2.1). Operation is possible in two modes: in reflected light or in fluorescence mode. The confocal arrangement of an illumination pinhole and a conjugated detector pinhole ensures that only information from the focal plane reaches the detector. Using this arrangement, images of objects with a resolution up to 1.4x higher than in a non-confocal system can be obtained.

Figure 8.2.1. The Confocal Laser Scanning Microscope Zeiss LSM710 and ZEN navigation software.

The advantage of confocal light microscopy is the capturing the light emitted by a single plane of a sample. A laser beam scans the specimen pixel by pixel and line by line. A pinhole conjugated to the focal plane obstructs the light emerging from objects outside that plane so that only light from objects that are in focus can reach the detector (Fig. 8.2.2). The pixel data gathered using this method are then assembled to form an image that represents an optical Methods 59

section of the specimen and is distinguished by high contrast and high resolution in the X, Y and Z planes. Several images generated by means of shifting the focal plane can be combined into a 3D image stack.

Figure 8.2.2. Light path and image formation in a CLSM. Beam path of the excitation and emission light in a confocal laser scanning microscope. The confocal pinhole allows the creation of optical sections by hiding fluorescence signals of non-focal levels.

Having our cells in a 96-well microtiter plate we wanted to see their three-dimensional structure. The cells were stained with SYTO9 and Propidium iodide (Pi) dyes in order to visualize the ratio of the live/dead cells. SYTO9 is a green-fluorescent nucleic acid stain, it has been shown to stain live and dead Gram-positive and Gram-negative bacteria, and it is a component of the LIVE/DEAD BacLight Bacterial Viability Kits (L-7007, L-7012, L-13152) working together with Pi. SYTO9, a 5 mM solution in 100 μl DMSO, allows cell permeability and when bound to nucleic acids, the maximum absorption is 485 nm and the maximum fluorescence emission is 498 nm. Pi dye binds to DNA by intercalating between the bases with little or no sequence preference and with a stoichiometry of one dye per 4–5 base pairs of DNA. It is membrane impermeant and generally excluded from viable cells, commonly used for identifying dead cells in a population and as a counterstain in multicolor fluorescent techniques. When bound to nucleic acids, the maximum absorption for PI is 535 nm and the maximum fluorescence emission is 617 nm (Fig. 8.2.3). Stacks of optical serial sections (optical thickness 1 µm) were recorded using a confocal laser scanning microscope (LSM 710, Zeiss) with a 40× PlanNeofluar (NA 1.3) oil immersion objective. The confocal images were analyzed with Zeiss Efficient Navigation (ZEN) 2009 light edition software.

Figure 8.2.3. Absorption and fluorescence emission profiles of propidium iodide bound to dsDNA. Methods 60

8.3. Generating the Templates for Expression and Purification

8.3.1. Polymerase Chain Reaction (PCR)

The polymerase chain reaction (PCR; Erlich, 1989) is a technique that amplifies specific DNA fragments from minute quantities of source DNA material, even when that source DNA is of relatively poor quality. All PCR reactions were set up using KOD Hot Start Polymerase (Novagen). 1 μl of templates was used (except for the Colony PCR). The reactions were prepared in the following scheme in a total volume of 50 μl for PCR.

PCR

mQ H2O 30 μl DMSO 3 μl Reaction Buffer 4 5 μl

25 mM MgSO4 3 μl 2 mM dNTPs 5 μl Forward primer 0.4 μl Reversed primer 0.4 μl KOD DNA polymerase 0.4 μl Template 2 μl

The constructs were amplified using the following program.

PCR Temperature Time 95 oC 30’’ 95 oC 15’’ 60 oC 15’’ 72 oC 60’’ x5 95 oC 15’’ 65 oC 15’’ 72 oC 60’’ x20 72 oC 5’ 12 oC forever

Depending on the size of the PCR bands produced and the discrimination needed, band visualization was accomplished by 1.2% agarose gel electrophoresis and analyzed under UV light.

Methods 61

8.3.2. PCR Purification and Gel Extraction

For PCR purifications and gel extractions the kits of Qiagen were used according to the protocol given in the handbooks of the kit. The DNA was always eluted in 35 μl of mQ H2O.

8.3.3. Agarose Gel Electrophoresis

Agarose gel electrophoresis is a powerful separation method frequently used to analyze DNA fragments generated by restriction enzymes, and it is a convenient analytical method for determining the size of DNA molecules in the range of 500 to 30,000 base pairs. It can also be used to separate other charged biomolecules such as dyes, RNA and proteins. The separation occurs because smaller molecules pass through the pores of the gel more easily than larger ones, i.e., the gel is sensitive to the physical size of the molecule. DNA samples were analyzed by 1.2% agarose gel electrophoresis. Samples were mixed with SYBRsafe for DNA and 6x loading buffer. For separation 1.2% agarose in 1x TAE (Tris-acetate-EDTA) buffer was used. The gel was run at 120 V/200 mA for 30 min. For size determination 2-loq DNA marker (NEB) was applied as well to the gel.

8.3.4. Site Directed Mutagenesis

By site directed mutagenesis, mutations were created at a defined site in a DNA molecule. First step in site directed mutagenesis was the amplification of the mutant DNA by typical PCR. The reactions were prepared according to KOD polymerase protocol. The constructs were amplified using the following program, while elongation time was extended to 5 min. Site Directed Mutagenesis Temperature Time 95 oC 2’ 95 oC 20’’ 60 oC 20’’ 70 oC 3:30’ x5 95 oC 20’’ 60 oC 20’’ 70 oC 3:30’ X7 72 oC 5’ 12 oC forever

Following the reaction, the product was digested with DpnI. DpnI only cleaves at methylated sites, so it chews up the template plasmid but not the PCR product. 1 μl DpnI and 5 μl Buffer 4 (New England BioLabs, NEB) were added to the reaction and were incubated for 1 hour at 37 oC. After digestion step, E. coli Nova Blue competent cells were transformed with 10 μl of the PCR reaction and LB+Kan containing petri plates were plated and incubated at 37 oC overnight. Grown colonies were sent for sequencing (EUROFINS MWG). Methods 62

8.4. Recombination Cloning in Bacteria

8.4.1. Preparation of Competent Cells

Generating competent cells is a laboratory procedure in which cells are passively made permeable to DNA, using conditions that do not normally occur in nature.

Solutions: 100 mM MgCl2 100 mM CaCl2 in 15% Glycerol

The main culture was inoculated using 2 ml of overnight culture in 100 ml 2xTY media o plus the respective antibiotic. The bacteria were grown at 37 C until they reached an OD600 of 0.4 – 0.6. The cells were collected by centrifugation for 10 min (3000 rpm, 4 oC). The pellet (cells) were resuspended in ice–cold 50 ml MgCl2 and incubated on ice for 30 min, followed by centrifugation (10 min/3000 rpm/4 oC). The pellet was resuspended in 10 ml Ca+2 solution and aliquots of 200 μl were frozen in liquid nitrogen and stored at -80 oC.

8.4.2. Digestion by Restriction Enzymes

To ligate a DNA fragment (also called insert) into a plasmid vector, first of all is need to prepare the insert and the vector in a way that the fragment can be inserted into the vector. For this, fragment and vector, must have compatible ends after digestion. In order to design a polycistronic gene, consisting of 4 genes, the 4 genes and the vector pQE70 were digested with the same restriction enzymes producing complementary single-stranded tails. All restriction digestions were done overnight at 37 oC. All used enzymes were purchased from NEB. The pQE70 vector was digested with SphI and BamHI, while the polycistronic gene was digested as shown in the following scheme.

All reactions took place in presence of 1 μl of the respective restriction enzyme, 4 μl Buffer 4 (NEB) and 1 μl BSA. The inserts were purified by PCR purification kit (Qiagen) and the plasmids were purified by gel extraction kit (Qiagen) according to manufacturer’s instructions.

8.4.3. Ligation and Transformation

The digested plasmids were dephosphorylated with the Antarctic phosphatase (NEB) for 30 min at 37 oC. Antarctic Phosphatase catalyzes the removal of 5´ phosphate groups from DNA and RNA. Since phosphatase-treated fragments lack the 5´ phosphoryl termini required by Methods 63

ligases, they cannot self-ligate. After heat inactivation of the enzyme for 10 min at 65 oC, the plasmids were ligated with the inserts (using a ratio 1:3 and 1:6) in the total volume of 10 μl, containing 1 μl ligation buffer and 1 μl T4 ligase (Fermentas). The ligation was carried out at 22 oC at least for one hour using T4 DNA ligase under Standard conditions (Sambrook et al, 1989). Following transformation, competent cells were thawed on ice and the ligation mix was added on the top of them and mixed gently. After 30 min of incubation on ice, the cells were heat shocked for 45 sec at 42 oC. 100 μl of 2xTY media were added per mixture and cells were shaken at 37 oC for at least 45 min (650 rpm). After transformation the cells were plated on LB agar containing the appropriate antibiotics and incubated overnight at 37 oC.

8.4.4. Colony Polymerase Chain Reaction (PCR)

Colony PCR was used after ligation and transformation to screen colonies and determine if the cloning worked, by checking to see if the insert is present, and estimating its size. The reactions were prepared in the following scheme in a total volume of 10 μl.

Colony PCR DMSO 0.8 μl Reaction Buffer 4 1 μl

25 mM MgSO4 0.6 μl 2 mM dNTPs 1 μl forward primer 0.3 μl reversed primer 0.3 μl KOD DNA polymerase 1 μl

Selected colonies of bacteria were labeled and picked with a pipette tip from overnight incubated petri plate (37 oC) and added directly to the PCR mix. PCR is then conducted to determine if the colony contains the DNA fragment or plasmid of interest. The constructs were amplified using the following program.

Colony PCR Temperature Time 95 oC 2’’ 95 oC 20’’ 60 oC 20’’ 70 oC 40’’ x5 95 oC 20’’ 60 oC 20’’ 70 oC 40’’ x20 72 oC 5’ 12 oC forever

After the PCR reactions were completed, the PCR products were run in 1% agarose gel (120 V/200 mA/30 min) in order to analyze the results of the amplification. The gel was checked Methods 64

under UV light. Any colonies which give rise to an amplification product of the expected size are likely to contain the correct DNA sequence. For backing-up the picked colonies a new petri plate with LB agar and the appropriate antibiotic was marked on the bottom and left for overnight incubation at 37 oC. The next day according to PCR’s results, the promising colonies were selected, inoculated in 5 ml of LB media (supplemented with the appropriate antibiotic) and incubated overnight at 37 oC in order to prepare minipreps according to Qiagen Kit.

8.4.5. Miniprep Isolation of Plasmid DNA

After bacteria have been transformed either with plasmid vector DNA or with plasmids containing inserts, the plasmid (or recombinant) DNA needs to be isolated from the bacterial chromosome. In this study DNA isolation of the generated clones was purified using the DNA miniprep kit (Qiagen). The isolation was done according to the manual of the kit. 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 (Vogelstein and Gillespie, 1979). The unique silica-gel membrane used in QIAprep Miniprep Kits completely replaces glass or silica slurries for plasmid minipreps. The procedure consists of three basic steps: (i) preparation and clearing of a bacterial lysate, (ii) adsorption of DNA onto the QIAprep membrane, and (iii) washing and elution of plasmid DNA. The DNA was always eluted in 50 μl mQ H2O. Then 18 μl of isolated DNA were sent to EUROFINS MWG for sequencing. The sequences were analyzed by using the Clone Manager Suite software (version 1.3.0.0.).

8.5. Analysis of the Generated Clones

8.5. 1. Expression in E. coli

Expression of recombinant proteins can be approached in general by starting with a plasmid that encodes the desired protein, introducing the plasmid into the required host cell, growing the host cells and inducing expression, and ending with cell lysis and SDS-PAGE analysis to verify the presence of the protein. E. coli Novablue cells transformed with generated constructs was cultured overnight at 37 oC (130 rpm). The main culture (terrific broth media plus respective antibiotic) was inoculated with overnight culture (diluted 1:100) and grown until an OD600 of 0.6, followed by isopropyl-β-D-thiogalactoside (IPTG) induction (final concentration 1 mM). After 3 hours of incubation at 30 oC, cells were centrifuged at 3000 rpm/15 min/4 oC and the pellet was stored at -20 oC until processing (purification).

Methods 65

8.5.2. Sodium Dodecyl Sulphate-Polyacrylamid Gel Electrophoresis (SDS-Page) and Coomasie Staining

The theory behind the SDS-Page is to separate proteins based on size through the use of a stacking gel and a resolving gel. SDS is a strong anionic detergent which disrupts the secondary, tertiary and quaternary structures of a protein and leads to the formation of a linear polypeptide chain. Therefore, if a cell is incubated with SDS, the membranes will be dissolved and the proteins will be solubilized by the detergent, plus all the proteins will be covered with many negative charges. Under the influence of the applied electric field, the negatively charged protein migrates from cathode (-) to anode (+) in accordance to their size. In SDS-Page the proteins have to be put into an environment that will allow different sized proteins to move at different rates. The environment of choice is polyacrylamide, which is a polymer of acrylamide monomers. Polyacrylamide gel consists of chains of polymerized acrylamide which are cross- linked by a bi-functional agent, i.e. N, N’-methylene-bis- acrylamide. The gel acquires rigidity and tensile strength from the cross-links formed by bisacrylamide. These cross-links forms pores in the gel through which SDS-protein complexes pass.

SDS running buffer (10x) 0.25 M Tris 1.918 M Glycerin 1% [w/v] SDS add 1000 ml H2O

SDS sample buffer (3x) 62.5 mM Tris-HCl, pH 6.8 10% Glycerol 20% SDS 0.7 M β-Mercaptoethanol 1% bromphenol blue

APS (ammonium persulfate) 10% [w/v] APS add H2O

Coomasie Blue Staining Solution 0.25% Coomasie Blue R 250 40% Methanol 10% acetic acid add H2O

Destaining Solution 40% Ethanol 10% acetic acid add H2O

Molecular Weight Protein Marker New England BioLabs (NEB) (Broad Range)

Methods 66

The proteins were separated according to their sizes by SDS-page electrophoresis (Laemmli, 1970). The composition of the gels is given in Table 8.5.2.1. Samples were mixed with samples buffer and denaturated for 5 min at 95 oC. After loading the proteins on the gel, they were run at 190-200V until the sample reached the end of the separating gel (at least 1 hr). The gels were stained by Coomasie Blue staining solution for at least 30 min (Merril, 1990). After destaining the gels were analyzed.

Table 8.5.2.1. Composition of SDS gels. The volumes are given for 6 gels. The Tris solution has a different pH for the stacking and separation gel.

Stacking gel 4% Separating gel 15% Rotiphorese R Gel 30 2 ml 15 ml

H2O 10,8 ml 3,4 ml 1M Tris 1,9 ml (pH 6.8) 11,2 ml (pH 8.5) Bis-Acrylamid 2 ml 15 ml 10% SDS 150 μl 300 μl TEMED 15 μl 15 μl APS 150 μl 300 μl 0.1% PyroninY 46 μl ------

8.5.3. 6xHis-tag Protein Purification using Ni-NTA Agarose Beads

Figure 8.5.3.1. Ni-NTA spin purification procedure. Methods 67

Recombinant proteins containing one or more 6xHis affinity tags, located at either the amino and/or carboxyl terminus of the protein, can bind to the Ni-NTA groups on the matrix with an affinity far greater than that of antibody-antigen or enzyme-substrate of the protein. Binding of the 6xHis tag does not depend on the three-dimensional structure of the protein. The high affinity of the Ni-NTA resins for 6xHis-tagged proteins is due to the specificity of the interaction between histidine residues and immobilized nickel ions and to the strength with which these ions are held to the NTA resin. The purification procedure can be divided into three stages: preparation of the cell lysate and binding of the 6xHis-tagged protein to Ni-NTA silica, washing, and elution of the 6xHis-tagged protein (Fig. 8.5.3.1). After centrifugation most of the non-tagged proteins flow through. Residual contaminants and non-tagged proteins are removed by washing with buffers of slightly reduced pH or with buffers containing a low concentration of imidazole. Monomers and multimers elute at concentrations of imidazole greater than 100 mM and 200 mM, respectively. Purified protein is subsequently eluted in aliquots of 100-200 μl.

Components of the Buffers:

Lysis Buffer 50 mM Phosphate pH 8.0 300 mM NaCl 5 mM Imidazole pH 8.0

Wash Buffer 50 mM Phosphate pH 8.0 300 mM NaCl 10 mM Imidazole pH 8.0

Elution Buffer 50 mM Phosphate pH 8.0 300 mM NaCl 250 mM Imidazole pH 8.0

E. coli cells were grown in 50 ml LB media at 37 oC, containing the respective antibiotics. The cells were lysed in 15 ml lysis buffer. 5 μl of 10 mg/ml DNAseI were added to the suspension and then cells were incubated on ice for 15 min. After clearing the lysate by centrifugation at 17000 rpm/20 min/4 oC, the supernatant was incubated with the Ni-NTA agarose beads for an hour at 4 oC while rotating. The beads were washed twice with lysis buffer before applying the supernatant. Flow-through was collected in a falcons and pre-incubated with the supernatant beads were washed in 10 ml wash buffer twice. The protein was eluted in 750 μl elution buffer in six steps. Aliquots of each fraction were loaded on SDS-gel for analysis. Aliquots from each step, including supernatant and flow-through, were collected and loaded on SDS-gel for analysis.

Methods 68

8.6. Gel Filtration

Gel filtration is the simplest and mildest of all chromatography techniques and separates molecules on the basis of differences in size as they pass through a gel filtration medium packed into a column. Unlike ion exchange or affinity chromatography, molecules do not bind to the chromatography medium so buffer composition does not directly affect resolution (the degree of separation between peaks). Separations can be performed in the presence of essential ions or cofactors, detergents, urea, guanidine hydrochloride, at high or low ionic strength, at 37 oC or in the cold room according to the requirements of the experiments. Purified proteins can be collected in buffer containing: 50 mM NaH2PO4 (pH 8.0), 150 mM NaCl and 2 mM β-mercaptoethanol. The technique can be applied in two distinct ways:  Group separations, and  High resolution fractionation of biomolecules.

The medium (i.e. Superdex, Sephadex, Sephacryl or Superose) is a porous matrix in the form of spherical particles that have been chosen for their chemical and physical stability, and inertness (lack of reactivity and absorptive properties). The pores of the matrix and the space in between the particles are equilibrated with buffer. The liquid inside the pores is usually referred to as the stationary phase and this liquid is in equilibrium with the liquid outside the particles, referred to as the mobile phase. Figure 8.6.1 shows the most common terms used to describe the separation process of gel filtration.

Figure 8.6.1. Common terms in gel filtration.

Results from gel filtration are usually expressed as an elution profile or chromatogram that shows the variation in concentration (typically in terms of UV absorbance at A280nm) of sample components as they elute the column in order of their molecular size (Fig. 8.6.2). Molecules that do not enter the matrix are eluted in the void volume, Vo as they pass directly through the column at the same time speed as the flow of buffer. Molecules with partial access to the pores of the matrix elute from the column in order of decreasing size. Molecules with full access to the pores move down the column, but do not separate from each other. These molecules usually elute just before one total column volume, Vt, of buffer has passed through the column. Methods 69

Figure 8.6.2. Theoretical chromatogram of a high resolution fractionation. Large molecules leave the column first followed by smaller molecules in order of their size. The entire separation process takes place as one total column volume of buffer passes through the gel filtration medium.

Here, proteins after Ni-NTA purification were further purified using the gel filtration column Superdex G75 (Amersham biosciences). Superdex is a composite medium based on highly cross-linked porous agarose particles to which dextran has been covalently bonded. The result is media with high physical and chemical stability, due mainly to the highly cross-linked agarose matrix, and excellent gel filtration properties determined mainly by the dextran chains (Fig. 8.6.3). The filtration was done at 4 oC. Fractions of 1.5 ml volume were collected and loaded in SDS-gel for analysis.

Figure 8.6.3. The figure shows a schematic of a section through a Superdex particle.

Results 70

9. Results

9.1. Deletion of efp or yjeA lead to growth defects in E. coli

To examine whether the in vivo activity of EF-P is affected by its single-residue substitutions that abolish the lysinylation by YjeA, we used the Keio collection of E. coli K-12 single-gene deletion mutants (Baba et al, 2006): the Δefp strain, the ΔyjeK strain, the ΔyjeA strain and the ΔyfcM strain. In the experiment, the above strains were grown for 4.5 hrs in LB o media at 37 C till they reached stationary-phase. The OD600 was measured periodically. The following graph represents the growth of each strain over time, having wild-type E. coli as control. The Δefp and ΔyjeA mutant strains grew slower than the wild-type strain (Fig. 9.1.1), while the ΔyfcM strain grows the same way as wild-type. The ΔyjeK strain grew also slower than the wild-type, but faster than Δefp and ΔyjeA.

Growth Curves of Deletion Strains

1.0

0.8

0.6

OD (600 nm) 0.4 wt efp yjeK

0.2 yjeA yfcM

0.0 0 50 100 150 200 250 300 Time (min)

Figure 9.1.1. Growth curves of wild-type E. coli and deletion strains grown in LB media, showing that Δefp strain has growth defects.

Additionally, it was examined whether the phenotype of E. coli mutant strains can be more severe under stress conditions in the absence of efp gene and/or if EF-P is partially modified or not. For that reason Δefp, ΔyjeK, ΔyjeA and ΔyfcM strains were grown under varying stress conditions, namely starving conditions, influence of temperature and susceptibility to antibiotics. So, as shown in figure 9.1.2, all starved cultures, containing glycerol or glucose as carbon source were grown much slower compared to the 2xTY or LB media. In the lag-phase the cultures growing in glucose media grew faster compared to the glycerol cultures. Moreover, the growth rates of Δefp and ΔyjeA strains’in either 0.2% glucose or glycerol containing media, was approximately half that of wild-type E. coli, while ΔyfcM grows almost as fast as wild-type. According to these data, we conclude that the modification is important for active EF-P under starving conditions, but the viability of the cells does not depend on the modification, and Results 71

further, Δefp and ΔyjeA strains in vivo lead to growth defects in E. coli. The present in vivo study of the EF-P mutants clearly showed the importance of the lysyl modification at K34 in EF-P in vivo. In future studies, therefore, it would be interesting to determine how the putative aminoacyl–EF-P functions in the peptidyl transferase active site.

2xTY Media LB Media

1.0 1.0

0.8 0.8

0.6 0.6

0.4 0.4 (600 OD nm)

OD (600 nm) wt wt efp efp yjeK yjeK 0.2 yjeA 0.2 yjeA yfcM yfcM

0.0 0.0 0 50 100 150 200 250 300 0 50 100 150 200 250 300 Time (min) Time (min)

M9 Glucose Media M9 Glycerol Media

0.35 0.28

0.26 0.30 0.24

0.25 0.22

0.20 0.20 0.18 0.15 0.16

OD (600 nm) (600 OD

OD (600 nm) (600 OD wt wt 0.10 efp 0.14 efp yjeK yjeK 0.12 yjeA yjeA 0.05 yfcM yfcM 0.10

0.00 0.08 0 50 100 150 200 250 300 0 50 100 150 200 250 300 Time (min) Time (min) MM Glucose Media MM Glycerol Media 0.5

0.30

0.4 0.25

0.3 0.20

0.15 0.2

OD (600 OD nm)

OD (600 nm) (600 OD wt wt efp 0.10 efp yjeK yjeK 0.1 yjeA yjeA yfcM 0.05 yfcM

0.0 0.00 0 50 100 150 200 250 300 0 50 100 150 200 250 300 Time (min) Time (min)

Figure 9.1.2. Growth under starving conditions. Δefp and ΔyjeA strains in vivo and under starving conditions lead to growth defects in E. coli. Results 72

9.2. efp is a Nonessential, but Important Gene for Cell Viability

E. coli Keio knockouts and E. coli parental strain were tested for temperature-sensitive growth at 37 and 44 °C in different growth media containing 25 μgr/ml Kan (except the wild- type strain). According to previous studies by Ganoza and co-workers, when the temperature was shifted to 44 °C, the growth of wild-type cells was unaffected however Δefp strain growth rate begins to slow almost immediately. (Aoki et al, 1997b). Here we show that when the temperature was shifted at 44 °C, after the cells had entered the logarithmic phase of growth, mutant strains slowed a growth for approximately 20-50 min, re-adapted to the new environmental conditions and resumed growing (Fig. 9.2.1). Given that yjeA and yjeK are not essential for the viability of either E. coli or Salmonella, we suggest that efp is a nonessential gene and may instead be required for the synthesis of a subset of proteins necessary for stress tolerance, a fact that comes in agreement with recent published data about Salmonella strains (Zou et al, 2012).

Results 73

Figure 9.2.1. Effect of temperature shift on the growth of wild-type and mutant cells. Cells were grown first at 37 oC. After 160 min of growth, the temperature was shifted to 44 oC. All strains re-adapted after 20-50 min to the new environmental conditions and kept growing. In case of M9 glucose/glycerol and MM glucose/glycerol media, cells are growing much slower, but again after the temperature shift they re-adapt and keep growing.

9.3. E. coli Deletion Strains Show Sensitivity to Nonribosomal Inhibitors

Wild-type E. coli and mutant strains were also tested for their ability to produce visible growth measuring MICs in 96-well microplate wells containing 2xTY media and dilutions of antibiotics from different pharmacological classes and with different mechanisms of action. Visible growth was measured by turbidity at each concentration following 16-20 hrs of incubation at 37°C. Wild-type E. coli and non-ribosomal inhibitors were used as controls and the prepared graphs are shown in figure 9.3.1. The different classes and chemical structures of the antibiotics tested are listed in appendix in Supplementary Tables 4 and 5, respectively. If the posttranslational modification catalyzed by YjeA, YjeK and YfcM is essential for EF- P activity, the phenotypes of the yjeK, yjeA, yfcM and efp mutants would be expected to be similar. Nevertheless, E. coli yjeA, yjeK and efp mutants are sensitive to a variety of antibiotics, in contrast to wild-type and yfcM mutant. So, the examined strains could be separated in a wider range of two groups: (group 1) wild-type and ΔyfcM strains, and (group 2) Δefp, ΔyjeK and ΔyjeA strains. Based on our data, group 1 shows usually resistance to the majority of antibiotics and group 2 is often more sensitive to varying classes of antibiotics (Fig. 9.3.1). Antibiotics targeting Gram-positive bacteria, like bacitracin and vancomycin, had as expected no effect on E. coli (Gram-negative) cells. In particular, bacitracin interferes with the dephosphorylation of the C55-isoprenyl pyrophosphate, a molecule that carries the building- blocks of the peptidoglycan bacterial cell-wall outside of the inner membrane (Stone and Strominger) and vancomycin acts by inhibiting proper cell-wall synthesis (Gold, 2001). Additionally, antibiotics like ampicillin, rifampicin and norfloxacin were used as controls, as they are not protein synthesis inhibitors and it was expected that they wouldn’t have any effect on the strains. Specifically, ampicillin is a β-lactam antibiotic inhibiting the cell-wall synthesis in bacteria (Rolinson, 1980). Rifampicin inhibits bacterial RNA synthesis by binding to the β- subunit of DNA-dependent RNA polymerase, thus blocking RNA transcription (Ehrlich et al, Results 74

1973). Norfloxacin functions by inhibiting DNA gyrase, a type II topoisomerase, and topoisomerase IV (Drlica and Zhao, 1997), enzymes necessary to separate bacterial DNA, thereby inhibiting cell division. Interestingly though, ampicillin, rifampicin and norfloxacin, in low concentrations (4 μgr/ml), had a different from the expected effect on our strains (Fig. 4), which showed sensitivity to those antibiotics. Moreover polymyxins, which disrupt the plasma membrane, causing leakage (Neu and Gootz, 1996) also inhibited the growth of the mutants in extremely low concentrations (data not shown). Consequently, as the mutant strains had increased susceptibility to those antibiotics, we hypothesize that E. coli EF-P does not only play a role in initiation and/or elongation of translation (Glick and Ganoza, 1975; Aoki et al, 2008), but also it might participate in molecular pathways that deal with the intergrity of cell membrane of E. coli cells, indicative of a perturbation in the cell envelope. Based on previous findings in Salmonella strains that yjeA and yjeK mutants were more susceptible to the aminoglycoside gentamicin, the susceptibility of deletion strains to this class of antibiotics was also assessed in E. coli (Navarre et al, 2010). Regarding the aminoglycosides spectinomycin and tetracycline, Δefp, ΔyjeK and ΔyjeA strains were more susceptible than either ΔyfcM and wild-type strains, as expected, as they all affect initiation and/or elongation of translation, starting from 16 μg/ml and 2 μg/ml, respectively, but always Δefp strain was slightly more susceptible than ΔyjeK and ΔyjeA (Fig. 9.3.1). Kasugamycin had a milder effect on the cells, starting growth inhibition at 128 μg/ml. Under the effect of clindamycin, erythromycin, H2O2, hygromycin A and linezolid the strains were resistant and sensitivity was obvious only in very high concentrations of the antibiotics, which was expected as they interfere mainly with the PTC center, something that did not happen under the effect of chloramphenicol, to which cells were susceptible in concentrations starting from 16 μg/ml. MICs from the antibiotics tested are listed in Supplementary Table 6.

1.2 1.2

wt 1.0 1.0 efp yjeK yjeA 0.8 yfcM 0.8

0.6 0.6

OD (600 OD nm) (600 OD nm) 0.4 0.4 wt efp yjeK 0.2 0.2 yjeA yfcM Col 1 vs yfcM: 32

0.0 0.0 0.1 1 10 100 0.1 1 10 100 Ampicillin (g/ml) Apramycin [g/mL]

Results 75

1.4 1.2

wt 1.2 1.0 efp yjeK 1.0 yjeA 0.8 yfcM

0.8 0.6 0.6

OD (600 OD nm)

OD (600 OD nm) wt 0.4 0.4 efp yjeK yjeA 0.2 0.2 yfcM

0.0 0.0 0.1 1 10 100 1000 0.1 1 10 100 Bacitracin [g/ml] Chloramphenicol [g/ml]

1.2 1.2

wt 1.0 1.0 efp yjeK yjeA 0.8 0.8 yfcM

0.6 0.6

OD (600 OD nm) (600 OD nm) 0.4 wt 0.4 efp yjeK 0.2 yjeA 0.2 yfcM

0.0 0.0 0.1 1 10 100 1000 1 10 100 1000 Clindamycin [g/ml] Erythromycin [g/ml]

1.2 1.2

wt wt 1.0 1.0 efp efp yjeK yjeK yjeA yjeA 0.8 0.8 yfcM yfcM

0.6 0.6

OD (600 OD nm)

OD (600 OD nm) 0.4 0.4

0.2 0.2

0.0 0.0 0.1 1 10 100 0.1 1 10 100 H O [g/ml] Gentamicin [g/ml] 2 2

Results 76

1.2 1.0

1.0

0.8

0.8

0.6 0.6

OD (600 OD nm) 0.4 (600 nm) OD wt wt 0.4 efp efp yjeK yjeK 0.2  yjeA yjeA 0.2 yfcM yfcM

0.0 0.0 0.1 1 10 100 1000 0.1 1 10 100 1000 Hygromycin A [g/ml] Kasugamycin [g/ml]

0.7 1.0

0.6 wt efp 0.8 yjeK 0.5 yjeA yfcM 0.6 0.4

0.3

OD (600 OD nm) 0.4 wt (600 OD nm)  efp 0.2 yjeK 0.2 yjeA yfcM 0.1

0.0 0.0 0.1 1 10 100 1000 0.1 1 10 100 Linezolid [g/ml] Norfloxacin [g/ml]

1.2 1.0

1.0

0.8 0.8

0.6 0.6

OD (600 OD nm)

OD (600 OD nm) wt 0.4 wt 0.4 efp efp yjeK yjeK yjeA 0.2 yjeA 0.2 yfcM yfcM

0.0 0.0 0.1 1 10 100 0.1 1 10 100 Rifampicin [g/ml] Spectinomycin [g/ml] Results 77

1.2 1.0 wt 1.0 efp yjeK 0.8 yjeA 0.8 yfcM

0.6 0.6

OD (600 OD nm) OD (600 OD nm) 0.4 0.4 wt efp yjeK  0.2 0.2 yjeA yfcM

0.0 0.0 0.1 1 10 100 0.1 1 10 Streptomycin [g/ml] Tetracycline [g/ml]

1.2 1.0

1.0

0.8

0.8

0.6 0.6

OD (600 OD nm) OD (600 nm) OD 0.4 wt 0.4 wt efp efp yjeK yjeK  0.2 yjeA 0.2 yjeA yfcM yfcM

0.0 0.0 0.1 1 10 100 1000 0.1 1 10 100 1000 Vancomycin [g/ml] Viomycin [g/ml]

Figure 9.3.1. The figure shows the graphs for each antibiotic tested in alphabetical order. Ampicillin. Apramycin. Bacitracin. Chloramphenicol.

Clindamycin. Erythromycin. Gentamicin. H2O2. Hygromycin A. Kasugamycin. Linezolid. Norfloxacin. Rifampicin. Spectinomycin. Streptomycin. Tetracycline. Vancomycin. Viomycin.

9.4. EF-P is Important in Cell Viabillity under Stress Conditions in E. coli

In this experiment we are testing whether the transformed mutant strains containing a plasmid with a normal copy of the mutated genes have the ability to restore viability compared to the Keio collection of E. coli K-12 single-gene deletion mutants. When a gene is knocked-out and something abnormal is observed, then re-introduction of that gene should "rescue" the original state of the cell, hence the term. As shown in figure 9.4.1 wild-type recovers directly, and ΔyfcM strain recovers relatively quickly, even at the fourth day, showing that the mutation is not lethal for E. coli, that yfcM gene is not important for the cell viability, and that even in its absence YjeK and YjeA can modify EF-P, but not completely. The growth rate of Δefp, ΔyjeK and ΔyjeA strains on the other hand, is half the wild-type, suggesting the negative effect of those mutations on E. coli viability (Fig. 9.4.1) and showing that the absence of either of these genes (yjeK or yjeA) leads to a predominance of completely unmodified EF-P in vivo. Results 78

Knock-out strains

1.0

0.8

0.6

(600 nm)

OD OD 0.4

wt efp 0.2 yjeK yjeA yfcM 0.0 0 1 2 3 4 Time (days)

Figure 9.4.1. Δefp, ΔyjeK and ΔyjeA cannot restore wild-type phenotype in E. coli.

The pQE70 plasmid, containing the efp, yjeA or yfcM genes, restored the cell growth of the deletion mutants; expect the ΔyjeK strain (Fig. 9.4.2). The Δefp and ΔyjeA cells grew approximately as fast as the wild-type cells. The fact that the introduction of the yjeA gene into the respective mutant restored up to the level observed for wild-type the mutant phenotype, indicates that yjeA gene is required for the in vivo growth of E. coli and for the modification pathway of EF-P. In contrast, under the growth conditions tested, the pQE70 plasmid, con- taining the yjeK gene, failed to restore the cell growth of the ΔyjeK mutant (Fig. 9.4.2) all four days, suggesting that yjeK gene might have other functions in E. coli, for example YjeK participates in the degradation pathway of lysine, and that’s why secondary effects are observed. Further, the overexpression of YjeK from the plasmid could have made the cells toxic and for that reason there is no recovery of growth. ΔyfcM strain recovers slightly slower than Δefp and ΔyjeA strains, however, shows a relative influence on the E. coli viability, as even after the fourth day is rescued. Above all though, in this experiment we should take into concideration the fact that the proteins are expressed under the control of lac promoter, resulting in general in secondary effects for the cells. Nevertheless, the direct recovery of pEF- P- and pYjeA-containing strains, suggests that both enzymes are important in vivo for cell viability of E. coli under certain stress conditions. Results 79

Knock-out strains plus recovery plasmid

1.2

1.0

0.8

0.6

(600 nm)

OD

0.4 wt efp (pEF-P) 0.2 yjeK (pYjeK) yjeA (pYjeA) yfcM (pYfcM) 0.0 0 1 2 3 4 Time (days)

Figure 9.4.2. Δefp and ΔyjeA mutants restored the wild-type phenotype from the first till the last day of the experiment suggesting the important role of EF-P and YjeA in E.coli cells.

9.5. Lysine is the Required Substrate for YjeA in vivo

It has been reported that EF-P is post-translationally modified on Lys34 by a moiety with a mass consistent with lysine (Aoki et al, 2008). In order to identify whether wild-type phenotype can be restored by substitution of lysine with other amino acids, we repeated rescue experiments and growth under starving conditions, by replacing lysine 34 (K34) either with alanine, arginine, glutamine, asparagine, aspartic acid or glutamic acid (as shown below).

1. K143A 2. K143R 3. K34Q 4. K34N 5. K34D 6. K34E

The mutations were done by site directed mutagenesis. The primers used are listed in the appendix in Supplementary Table 7. After amplification of the mutants with PCR, the products were digested with DpnI and then transformed in Δefp competent cells. The positive colonies were incubated overnight for mini-preparation. According to the data (not shown) when the Lys34 residue was substituted by another amino acid, none of the amino acids tested restored wild-type phenotype, suggesting that lysine residue is essential for the in vivo modification of EF-P. Subsequently, despite the general role EF-P is proposed to play in translation, it is clear that mutations in yjeA and yjeK affect the expression of a relatively small Results 80

subset of proteins under the conditions tested. It is likely that the effects of YjeA and YjeK on the expression of some of these proteins are indirect.

9.6. Δefp strain does not Posses Flagella

Figure 9.6.1. Motility Test. Δefp and ΔyjeK strains do not possess flagella suggesting that they might cause perturbation to the cell envelope. ΔyjeA and ΔyfcM strain are attenuated for virulence in the E. coli.

In literature it was reported that EF-P and the lysinylation pathway are critical for attenuation of virulence in bacteria (Kaniga et al, 1998; Peng et al, 2001; Bearson et al, 2006; Navarre et al, 2010; Bearson et al, 2011) and that EF-P is recognized as one of the proteins important for bacteria motility, that it is not swarming (Kearns et al, 2004). Additionally, as reviewed by Josenhans and Suerbaum (Josenhans and Suerbaum, 2002), motility and virulence are often linked together. Based on our hypothesis that E. coli EF-P might be involved in cell- wall synthesis pathways, in this experiment we tested if the mutant strains are motile by means of flagella. Cells were grown in semi-solid agar overnight at 30 oC. Growth is indicated by the presence of a red color into the soft medium around from the point of inoculation, and as motility occurs, small to very large regions of color can be observed. Theoretically, as efp is a Results 81

non-motile gene, we are expecting that Δefp strain is going to be motile. However, as shown in figure 9.6.1, motility of Δefp strain is impaired, as it does not possess flagella compared to the wild-type, confirming our hypothesis that efp might be related to proteins causing perturbation to the integrity of cell membrane. ΔyjeK is also less motile than wild-type, confirming the fact that YjeK is known to interact with membrane proteins and other proteins that seem to be expressed under stress or in the stationary phase (Behshad et al, 2006). In ΔyjeA strain, where there is no lysinylation of EF-P, we are expecting that the strain is going to be motile, as expected for Δefp strain. As shown in figure 9.6.1 the strain is motile, proposing that the mutation might cause virulence to the cells, if motility and virulence are linked together (Josenhans and Suerbaum, 2002), and that yjeA might not participate in the same pathways like efp and yjeK regarding the cell-wall synthesis, as it is also known from Bailly and de Crécy- Lagard that in E. coli only efp/yjeK genes are clustered together (Bailly and de Crécy-Lagard, 2010). In order though to confirm such a suggestion experiments in aminals, for example mice or flies, should be carried out. These data are in agreement with reported studies suggesting that yjeA mutant was highly attenuated for virulence in mouse models of infection (Navarre et al, 2010). Regarding ΔyfcM strain, as it is a non-characterized protein yet, we interestingly observe that the strain is the most motile compared to wild-type and the rest of the mutants. Based on published data and our hypothesis however, ΔyfcM strain should have a phenotype similar to ΔyjeK and ΔyjeA, as YfcM is dependent on YjeK and YjeA for the post-translationally modification of EF-P (Peil et al, in press). The fact that in this experiment ΔyfcM strain is a “super swimmer” suggests that the strain might also be attenuated for virulence in E. coli cells. For sure though, there are a lot to be done still in order to find out the exact function of YfcM, except its role in the modification pathway of EF-P.

9.7. Cells Lacking YjeA or YjeK have Phenotypes Similar to those Lacking EF-P

Based on our data from motility tests and the fact that mutant strains are susceptible to antibiotics targeting cell-wall, except initiation/elongation of translation, we investigated further the morphology of the mutant strains. First attempt to see the morphology of the strains was done by Gram-staining and editing the slides under light microscope (data not shown). However, in order to have 3D and of higher resolution images, we confirmed light microscopy data using CLSM and the ZEN 2009 light edition software for the analysis of the images. Figure 9.7.1 shows the morphology of each strain. Compared to wild-type, where we can see typical rod-shaped bacteria, Δefp, ΔyjeK and ΔyjeA strains appear smaller and more compact, while they also have phenotypes similar to each other. Additionally, they possess fibers (black arrows in Fig. 9.7.1), which in microbiology is indicative of cell membrane “sickness” of the E. coli cells. On the other side, cells in ΔyfcM strain have different morphology, as they are more elongated compared to the other mutants, and they look more alike to wild- type. This phenotype also seems to have morphological defects that we cannot explain precisely yet, because our knowledge about YfcM, and its structure and function, is insufficient. Results 82

Wild-type Δefp

ΔyjeK ΔyjeA

c

ΔyfcM

Figure 9.7.1. Morphology of wild-type and mutant strains in CLSM. Δefp, ΔyjeK and ΔyjeA strains show similar morphology, in contrast to ΔyfcM which appears more elongated compared to wild-type. All mutants indicate growth defects and “sickness” of the cells. Results 83

9.8. YfcM Purification for Crystallization

The goal of crystallization is usually to produce a well-ordered crystal that is lacking in contaminants and large enough to provide a diffraction pattern when hit with X-ray. This diffraction pattern can then be analyzed to discern the protein’s three-dimensional structure. Proteins, like many molecules, can be prompted to form crystals when placed in the appropriate conditions. However, prior to starting protein crystallization, we first need to purify the protein in relatively large quantities (few milligrams). High purity and homogeneity of the sample are crucial for the crystallization to be successful. In this study YfcM [His-tag (C-terminal)], an uncharacterized protein, cloned into pET21b vector, was purified using Ni-NTA agarose beads as described in Methods part. After purification the fractions were loaded to a SDS gel (Fig. 9.8.1). 1/10 of each washing and elution fraction was loaded and 1/500 of supernatant and flow through were applied to the gel. The SDS gel was stained with Coomassie Brilliant Blue and destained for 30 min. Proteins after Ni- NTA purification were further purified using gel filtration column (Fig. 9.8.2A) and the corresponding elution fractions were applied to SDS gel (Fig. 9.8.2B).

YfcM

Figure 9.8.1. Purification of YfcM. YfcM was successfully purified using Ni-NTA agarose beads.

YfcM

Figure 9.8.2. (A). Gel filtration of elution fraction of YfcM. Peaks indicated by arrows were applied to SDS gel and after gel electrophoresis stained with Coomassie Brilliant Blue dye and analyzed. (B) Coomassie stained gel of elution fractions after gel filtration. Arrows indicate YfcM. Results 84

9.9. Cloning of the Polycistronic Construct: - yjeK - yjeA - yfcM - his-efp -

In order to study better the function of EF-P in translation extracts from different mutants and to see the crystal structure of the whole modification pathway of EF-P, we are trying to clone the polycistronic gene shown in figure 9.9.1. The process is still in initial stage, nevertheless the future goal is to clone the polycistronic gene in pET- or pRSFDuet-system, express it in BL21 expression strain and set-up screening for crystallization in robot.

Figure 9.9.1. The polycistronic gene cloned in pQE70 vector and digested with SphI, SacI and BamHI restriction enzymes.

After trying lots of primers we now have cloned half of the polycistronic gene. Additionally, until now the most successful ligation was in pQE70 vector. So, yjeK-yjeA (A-R) gene was PCR amplified using “for SphI yjeK new #1” and “rev #2 yjeA SacI” primers, while yfcM-his-efp (M-E) gene using “for pQE ME SphI SacI” and “rev efp BamHI” primers. The PCR products were purified (QIAGEN kit), digested with SphI/SacI (AR) and SacI/BamHI (ME) restriction enzymes and ligated in pQE70 , which was also digested with SphI/BamHI, in order to create the polycistronic ARME (Fig. 9.9.2). Figure 9.9.2 shows an agarose gel and the kilobases of each single gene in order to clone the polycistronic. We can see further where the ARME is expected to be.

yjeK – yjeA – yfcM – His-efp (ARME)

̴ 3.300 bp

Figure 9.9.2. In the figure the arrow indicates where the polycistronic gene should be, that is around 3.300 bp. Discussion and Perspectives 85

10. Discussion and Perspectives

Since its identification in 1975, the role of EF-P in protein synthesis has remained enigmatic. While EF-P is not required to reconstitute protein synthesis in vitro, some studies reported a mild stimulation of peptide synthesis upon the addition of EF-P (Glick and Ganoza, 1975; Swaney et al, 2006). Moreover, EF-P was previously reported to be essential for viability in E. coli (Aoki et al, 1997b; Gerdes et al, 2003; Henderson and Hershey, 2011), a hypothesis that was consistent with the observation that all bacteria, including those with a highly reduced genome, encode at least one efp homolog (Bailly et al, 2010; Gil et al, 2004; Glass et al, 2006; Navarre et al, 2006). However, a systematic high-throughput attempt to disrupt every nonessential gene in E. coli found that at least one strain of E. coli could tolerate a disruption of efp and remain viable (Baba et al, 2006; Yamamoto et al, 2009). The hypersusceptibility of efp, yjeA and yjeK E. coli mutants to a wide variety of antimicrobials, their poor growth under stress conditions (varying temperature and growth media), and attenuated virulence are additionally indicative of a perturbation in the cell envelope. Recently it was also reported that Salmonella yjeA and yjeK mutants are hypersensitive to a variety of growth inhibitors, including dyes, detergents, and sulfometuron methyl, an inhibitor of acetolactate synthase (Zou et al, 2012). So, current findings indicate that efp is a nonessential gene for viability and that loss of EF-P might affect the production of relatively few proteins (Zou et al, 2012), many of which are more highly expressed in the absence of EF-P and may be required for stress tolerance. The basis for a previous report (Aoki et al, 1997) that efp is an essential gene and is required for protein synthesis in E. coli remains still unclear. It is possible though, that a difference in strain background or growth conditions, such as high temperature or low- osmolarity medium (Zou et al, 2012), led to the earlier conclusion that this factor is essential. In addition to E. coli, EF-P has been successfully knocked out in Salmonella, Agrobacterium and Acinetobacter (Baba et al, 2006; de Crécy et al, 2007; Peng et al, 2001; Yamamoto et al, 2009). In the case of the Acinetobacter efp mutant, suppressor mutations conveying enhanced growth were obtained after a few generations (de Crécy et al, 2007). Taken together, these studies suggest that EF-P may be nonessential in many bacterial species. The fact that YjeA catalyzes the lysyl modification of a specific lysine residue of EF-P is a previously unknown example of a post-translational protein modification. Thus, there have been reported various protein modifications, such as glycosylation, acetylation, methylation, phosphorylation, biotinylation, lipoylation, phosphopantheteinylation, hypusination, ubiquitina- tion and others (Rucker and Wold, 1988). The present lysylation is analogous to biotinylation and lipoylation (Reche and Perham, 1999), which are catalyzed by biotinyl protein ligase (for example, BirA) and lipoyl protein ligase (for example, LplA), respectively. The catalytic core structure of BirA resembles those of the class II aaRSs (Safro and Mosyak, 1995), whereas the LplA structure does not resemble those of the class II aaRSs, and the LplA sequence shares no clear sequence homology with the aaRSs. Therefore, YjeA is the first aaRS paralog that has been shown to modify a protein with an amino acid (Yanagisawa et al, 2010). In eukaryotes, the function of EF-P is mediated by eIF5A (Bartig et al, 1992). Recently, eIF5A was shown to be an essential translation elongation factor (Saini et al, 2009), which Discussion and Perspectives 86

strictly conserves a unique post-translational residue modification of a specific lysine residue to hypusine, in a two-step reaction catalyzed by deoxyhypusine synthase (DHS) and deoxyhypusine hydroxylase (DOHH). However, DHS and DOHH are completely unrelated to the aaRSs. The hypusine modification is essential for eIF5A function and cell viability (Park et al, 2006; Wolf et al, 2007). Furthermore, it has been consistently observed that disruption of yjeK leads to milder phenotypes than disruption of yjeA does (Navarre et al, 2010). These findings are extended by showing that the most severe phenotypes are obtained by deletion of efp. With the structure of fully modified EF-P now known (Peil et al, in press) the focus can shift to understanding the means by which YjeA has adapted substrate specificities and functions distinct from that of the well-known members of the aaRS super family. Recent data (Roy et al, 2011; Peil et al, in press) place YjeK upstream of YjeA in the modification pathway. In particular, according to Peil et al, for the fully modified modification pathway of EF-P, YjeK first converts (S)-α-lysine to (R)-β- lysine, then YjeA uses ATP to lysinylate EF-P by the addition of β-lysine to the ε-amino group of K34 (Behshad et al, 2006), and afterwards YfcM uses molecular oxygen to hydroxylate the C4 (γ) or C5 (δ) position of K34 of EF-P (Peil et al, in press). Several other studies have also demonstrated that YjeA is able to use lysine as a substrate for the modification of EF-P in vitro, albeit at a greatly reduced rate (Ambrogelly et al, 2010; Navarre et al, 2010; Roy et al, 2011; Yanagisawa et al, 2010). One plausible model for the relative severity of phenotypes is that in the absence of YjeK, YjeA attaches a lysine instead of a β-lysine to EF-P to generate a partially active protein. However, in the absence of YjeA, no such modification can occur, resulting in EF- P with an unmodified lysyl side chain that has little to no activity. Mutation of the lysyl side chain to an alanine could reduce the activity of EF-P even further, thereby creating a dominant negative effect and further exacerbating the phenotypes of the efp mutant. The role of EF-P in bacterial physiology also remains unclear. The novel finding from Zou et al. that EF-P plays a role in membrane integrity in Salmonella, and particularly in the translational regulation of a limited number of proteins that, when perturbed, renders the cell susceptible to stress by the adventitious overexpression of an outer membrane porin (Zou et al, 2012) is interesting in light of two reports, which link eIF5A to cell wall integrity in yeast and may also be true for the distantly related genus Agrobacterium (Chatterjee et al, 2006; Valentini et al, 2002). Several eukaryotic species encode multiple isoforms of eIF5A. It has been reported that eIF5A can selectively bind certain RNA molecules, supporting the notion that these factors may also modulate the translation of specific transcripts instead of acting as a general translation factor (Clement et al, 2003; Clement et al, 2006; Xu et al, 2004). aIF5A and eIF5A are generally considered to be essential for cellular growth, and some reports indicate that they play a role in peptide chain elongation beyond the formation of the first peptide bond, although these points remain controversial (Gregio et al, 2009; Henderson et al, 2011; Saini et al, 2009). While the mechanism by which EF-P, aIF5A, and eIF5A affect peptide bond synthesis at the peptidyl transferase center is likely similar, there are also considerable differences among the three proteins (Zou et al, 2011) regarding their structure as both aIF5A and eIF5A lack the third beta-barrel domain present in EF-P (Blaha et al, 2009). Future investigation could focus on how EF-P specifically targets its cognate transcripts and the role of this factor in bacterial physiology. Discussion and Perspectives 87

A logical next step to gain insight into the interworkings of the EF-P-YjeK-YjeA-YfcM modification pathway is to further investigate the divergent evolution of substrate recognition for YjeA. To define the basis for YjeA’s substrate recognition, crystallographic studies should be carried out with several forms of YjeA and varying substrates. Results of crystallographic studies will then be used to perform kinetic analyses of YjeA variants to further define the molecular basis for β-lysine specificity. Crystallographic studies should also be performed in order to visualize YfcM structure, as yfcM is still a non-characterized gene, with unknown function except its role in the modification pathway of EF-P as shown by Peil et al. (Peil et al, in press). Additionally, our future goal is to design the polycistronic gene and crystallize it, so that the studying of the function of EF-P in translation extracts from different mutants will be achieved. Another means of understanding the evolution of EF-P as a tRNA mimic is to investigate the role of its homologue, EF-P2. EF-P2 also exists in some bacterial organisms, either in addition to EF-P or in place of it. Because EF-P2 contains an arginine in place of a lysine at residue 34, the site of EF-P post-translational modification, it is possible that EF-P2 acts in a uniquely modified form or even as an unmodified protein. Based on these differences, it will be beneficial to gain information on the functional state of EF-P2, as well as to investigate the potential role of EF-P2 in translational pathways. This can be accomplished through testing for phenotypes produced in conditions similar to those evaluated with EF-P (such as hypoosmolarity and antibiotic stress), as well as to observe effects of EF-P2 on virulence in mouse models of infection. This information will aid researchers in evaluating the role of EF- P/EF-P2 in organisms lacking the YjeK/YjeA/YfcM pathway. As a second means of assessing the role of EF-P in translation systems, future work could focus on determining the action of modified EF-P on the ribosome. Although previous reports have detailed the structure of EF-P bound to the 70S ribosome (Blaha et al, 2009), this work was limited in that it did not include information on how modified EF-P associates with the different ribosomal sites. By judging the affinity of β-lysine-EF-P for the ribosome, it will be possible to detect a mode of ribosomal binding for β-lysine-EF-P. By comparing how modified and unmodified EF-P form initiation complexes with other ribosomal proteins, as well as determining what components are necessary for optimal EF-P activity, it will be possible to gain insight into the role of EF-P in peptide bond formation. Based on findings from this proposed work, researchers can then work to further define EF-P in its role in translation initiation and/or elongation. It is still unclear as to whether the homologous protein in eukaryotic systems, eIF5A, operates to enhance formation of the first peptide bond or aids in the incorporation of amino acids during translation elongation. Likewise, the role for EF-P is equally unknown. Measuring polysome/monosome ratios as well as monitoring 70S initiation complex formation in the presence of modified and unmodified EF- P will pinpoint the effect of EF-P on initiation. Additionally, the role of EF-P in elongation will be studied by employing poly(Phe) synthesis assays. Furthermore, polysome profiling and in vivo reporter assays will show if the YjeK-YjeA-YfcM-EF-P pathway favors processing of a subset of particular mRNA elements as opposed to acting on global protein synthesis. Overall, future work will be directed to provide knowledge of the possible mechanism by which EF-P influences translation, and largely contribute to a better understanding of EF-P across many species of bacteria. Ultimately we will gain a greater grasp on roles for aaRSs outside of amino-acylating Discussion and Perspectives 88

tRNA for protein synthesis, antibiotic resistance and virulence models, as well as post- translational gene regulation in bacteria. Finally, in order to determine whether yjeA, yjeK, yfcM and efp mutants share similar virulence phenotypes, each mutant should be assayed for its ability to cause disease in mice following intraperitoneal inoculation or flies. Pilot virulence experiments have been already performed in our lab in flies for wild-type and efp mutant. Wild-type strain caused lethality within 5 days of infection, in contrast to efp mutant, which was attenuated for virulence (data not shown).

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Appendix 107

Supplementary Table 1. List of Primers for the Polycistronic Construct.

Oligoname (Metabion International AG) Sequence

#1 yjeK FOR 5’-CCA GGT ATA ACA TAT GGC GCA TAT TGT AAC CCT AAA T-3’

#4 efp REV 5’-CGA TTT ACG CGA GCT CTT ACT TCA CGC GAG AGA CGT A-3’ for Bluescript 5’-GCC GCT CTA GAA CTA GTG GAT CC-3’ rev Bluescript 5’-CTC GAG GTC GAC GGT ATC GAT AAG-3’ for AmRs Polycis 5’-CTC CAG CTA CGC CAG CAG TAA GAA GGA GAT ATA CCA T-3’ rev AmRs Polycis 5’-CAA ATT TTA CGC AGG AAT AAC ACC ATG GTA TAT CTC C-3’ for yfcM-efp 5’-GTG GCC GGA AGC GCT CAA CTA AGA AGG AGA TAT ATC T-3’ rev yfcM-efp 5’-CGT GAT GAT GAT GAT GAT GCA TTC TAG ATA TAT CTC C-3’ for yjeA-yfcM 5’-CTT TAG CGT TGA CCG GGC ATA AGA AGG AGA TAT AGG A-3’ 5’-CTT TAG CGT TGA CCG GGC ATA AGA AGG AGA TAT AGG ATC CAT GAA CAG TAC ACA for yjeA-yfcM new CCA CTA CG-3’ rev yjeA-yfcM 5’-CGT AGT GGT GTG TAC TGT TCA TGG ATC CTA TAT CTC C-3’ for yfcM out polycis 5’-CTT TAG CGT TGA CCG GGC ATA AGA AGG AGA TAT AGG A-3’ rev yfcM out polycis 5’-GTA CGT TGC TCC GTG ATG ATG ATG ATG ATG CAT GGA T-3’

SphI yjeK FOR polyciston 5’-CCA GGT ATA AGC ATG CAT GGC GCA TAT TGT AAC CCT A-3’

BglII efp REV polycistron 5’-CGA TTT ACG CAG ATC TTT ACT TCA CGC GAG AGA CGT A-3’ 5’- CGA TTT ACG CGA GCT CTA TAT CTC CTT CTT ATG CCC GGT CAA CGC TAA AGG CG-3’ #2 yjeA SacI new rev 5’-CAT TAA AGA GGA GAA ATT AAG CAT GCC GAG ATT TAC GCG AGC TCA TGA ACA GTA for pQE ME SphI SacI CAC ACC ACT ACG-3’ 5’-CGT AGT GGT GTG TAC TGT TCA TGA GCT CGC GTA AAT ATA GGC ATG CTT AAT TTC rev pQE ME SphI SacI TCC TCT TTA ATG-3’ for gfp fix pQE 5’-CTC ATC ATC ATC ATC ATC ATT AAG CTT AAT TAG CTG AGC TTG G-3’ rev gfp fix pQE 5’-CCA AGC TCA GCT AAT TAA GCT TAA TGA TGA TGA TGA TGA TGA G-3’ for efp Nde #1 5’-GAT TTA TTA CAT ATG GGC AGC AGC CAT CAT CAT CAT CAT CA-3’

Appendix 108

Table Continued.

for efp Nde #2 5’-GAT TTA TTA CAT ATG CAT CAT CAT CAT CAT CAC-3’ for GFP pQE70 SphI 5’-CCA GGT ATA AGC ATG CTT ACT AAA GGT GAA GAA CTT TTC-3’ rev GFP pQE70 BamHI 5’-CGA TTT ACG CGG ATC CAT GAT GAT GAT GAT GAT GAG-3’

Duet efp NdeI for 5’-GAT TTA TTA CAT ATG CAT CAT CAT CAT CAT CAC GCG TG-3’

Duet efp XhoI rev 5’-CAG GTA TAA CTC GAG TTA CTT CAC GCG AGA GAC GTA TTC-3’

Duet yfcM NcoI for 5’-GAT TTA CGC CCA TGG GTA ACA GTA CAC ACC ACT ACG AG-3’

Duet yfcM SacI rev 5’-CGA TTT ACG CGA GCT CTT AGT TGA GCG CTT CCG GCC ACG-3’ for Sac IF5A SphI pQE70 5’-CCA GGT ATA AGC ATG CTT TCT GAC GAA GAA CAC ACC TTT G-3’ rev Sac IF5A BamHI pQE70 5’-CGA TTT ACG CGG ATC CTT AAT CAG ATC TTG GAG CTT CCT TGA A-3’ for Tkod IF5A SphI pQE70 5’-CCA GGT ATA AGC ATG CTT GGA GAC AAG ACT AAG GTT CAG-3’ rev Tkod IF5A BamHI pQE70 5’-CGA TTT ACG CGG ATC CTT ACT CGC CCC TGA TCT TCT TTA TC-3’ for Sac IF5A SphI pQE70 5’-CCA GGT ATA AGC ATG CTT TCT GAC GAA GAA CAC ACC TTT G-3’ rev Sac IF5A BamHI pQE70 5’-CGA TTT ACG CGG ATC CTT AAT CAG ATC TTG GAG CTT CCT TGA A-3’ for SphI yjeK new #1 5’-CCA GGT ATA AGC ATG CAT GGC GCA TAT TGT AAC CCT AAA TAC C-3’ 5’-CGA TTT ACG CGA GCT CCG ATT TAC GCG GAT CCT ATA TCT CCT TCT TAT GCC CGG rev #2 yjeA SacI TCA ACG CTA AAG GCG-3’ 5’-CCA GGT ATA AGG ATC CAT GCA CCA CCA CCA CCA CCA CCA CGG AGC AAC GTA rev efp BamHI TAG CAA CG-3’

Appendix 109

Supplementary Table 2. Maps of Vectors used.

Appendix 110

Table continued.

Appendix 111

Table continued.

Appendix 112

Supplementary Table 3. Sequences of Escherichia coli K-12 Proteins.

EF-P 188 aa protein Accession: AAC77107.1 GI: 1790590

MATYYSNDFRAGLKIMLDGEPYAVEASEFVKPGKGQAFARVKLRRLLTGTRVEKTFKSTDSAEGADVVDMNLTYLYND GEFWHFMNNETFEQLSADAKAIGDNAKWLLDQAECIVTLWNGQPISVTPPNFVELEIVDTDPGLKGDTAGTGGKPAT LSTGAVVKVPLFVQIGEVIKVDTRSGEYVSRVK

YjeK 342 aa protein Accession: AAC77106.1 GI: 1790589

MAHIVTLNTPSREDWLTQLADVVTDPDELLRLLNIDAEEKLLAGRSAKKLFALRVPRSFIDRMEKGNPDDPLLRQVLTSQ DEFVIAPGFSTDPLEEQHSVVPGLLHKYHNRALLLVKGGCAVNCRYCFRRHFPYAENQGNKRNWQTALEYVAAHPELD EMIFSGGDPLMAKDHELDWLLTQLEAIPHIKRLRIHSRLPIVIPARITEALVECFARSTLQILLVNHINHANEVDETFRQAM AKLRRVGVTLLNQSVLLRDVNDNAQTLANLSNALFDAGVMPYYLHVLDKVQGAAHFMVSDDEARQIMRELLTLVSGY LVPKLAREIGGEPSKTPLDLQLRQQ

YjeA 325 aa protein Accession: AAC77115.2 GI: 87082379

MPLSKTGDLTMSETASWQPSASIPNLLKRAAIMAEIRRFFADRGVLEVETPCMSQATVTDIHLVPFETRFVGPGHSQG MNLWLMTSPEYHMKRLLVAGCGPVFQLCRSFRNEEMGRYHNPEFTMLEWYRPHYDMYRLMNEVDDLLQQVLDCP AAESLSYQQAFLRYLEIDPLSADKTQLREVAAKLDLSNVADTEEDRDTLLQLLFTFGVEPNIGKEKPTFVYHFPASQASLA QISTEDHRVAERFEVYYKGIELANGFHELTDAREQQQRFEQDNRKRAARGLPQHPIDQNLIEALKVGMPDCSGVALGV DRLVMLALGAETLAEVIAFSVDRA

YfcM 182 aa protein Accession: AAC75386.1 GI: 1788666

MNSTHHYEQLIEIFNSCFADDFNTRLIKGDDEPIYLPADAEVPYNRIVFAHGFYASAIHEISHWCIAGKARRELVDFGYWY CPDGRDAQTQSQFEDVEVKPQALDWLFCVAAGYPFNVSCDNLEGDFEPDRVVFQRRVHAQVMDYLTNGIPERPARFI KALQNYYHTPELTAEQFPWPEALN

Reference http://www.ncbi.nlm.nih.gov/protein Appendix 113

Supplementary Table 4. Antibiotics in clinical use and modes of resistance (Morar and Wright, 2010).

Antibiotic Class Examples Targets Mode of Resistance β-lactams Penicillins (ampicillin) Peptidoglycan Hydrolysis Cephalosporins biosynthesis Efflux (cephamycin) Altered target Aminoglycosides Gentamicin Translation Phosphorylation Streptomycin Acetylation Spectinomycin Nucleotidylation Efflux Altered target Glycopeptides Vancomycin Peptidoglycan Reprogramming of peptidoglycan biosynthesis biosynthesis Tetracyclines Tigecycline Translation Monooxygenation Efflux Altered target Macrolides Erythromycin Translation Hydrolysis Azithromycin Glycosylation Phosphorylation Efflux Altered target Lincosamides Clindamycin Translation Efflux Altered target Streptogramins Synercid Translation C-O lyase (type B streptogramins) Acetylation (type A streptogramins) Efflux Altered target Oxazolidinones Linezolid Translation Efflux Altered target Phenicols Chloramphenicol Translation Acetylation Efflux Altered target Quinolones Ciprofloxacin DNA replication Acetylation Efflux Altered target Pyrimidines Trimethoprim Cl metabolism Efflux Altered target Suflonamides Sulfamethoxazole Cl metabolism Efflux Altered target Rifamycins Rifampin Transcription ADP-riboxylation Efflux Altered target Cationic peptides Colistin Cell membrane Efflux Altered target

Appendix 114

Supplementary Table 5. Chemical Structure of Different Classes of Antibiotics. (A) Tetracycline (reviewed by Wilson, 2009). (B) Apramycin (Ryden and Moore, 1977). (C) Kasugamycin. (D) Streptomycin. (E) Spectinomycin. (G) Erythromycin. (F) Chloramphenicol. (H) Clindamycin (reviewed by Wilson, 2009). (I) Linezolid. (J) Hygromycin A (Donohoe et al, 2009). (K) Viomycin (Barkei et al, 2009). (L) Gentamycin (reviewed by Wilson, 2009).

Appendix 115

Supplementary Table 6. MICs of antibiotics tested.

Name of Antibiotic MIC dilutions (μg/ml)

Ampicillin ≥0 ≥0.125 ≥0.25 ≥0.5 ≥1 ≥2 ≥4 ≥8 ≥16 wild-type + + + + + + + − − Δefp + + + + + + + − − ΔyjeK + + + + + + + − − ΔyjeA + + + + + + + − − ΔyfcM + + + + + + + − −

Apramycin ≥0 ≥1 ≥2 ≥4 ≥8 ≥16 ≥32 ≥64 wild-type + + + + + − − − Δefp + + + + − − − − ΔyjeK + + + + + − − − ΔyjeA + + + + + − − − ΔyfcM + + + + + + + +

Bacitracin ≥0 ≥ 0.5 ≥ 1 ≥ 2 ≥ 4 ≥ 8 ≥ 16 ≥32 ≥64 ≥128 ≥256 ≥512 wild-type + + + + + + + + + + + + Δefp + + + + + + + + + + + + ΔyjeK + + + + + + + + + + + + ΔyjeA + + + + + + + + + + + + ΔyfcM + + + + + + + + + + + +

Chloramphenicol ≥0 ≥0.125 ≥0.25 ≥0.5 ≥ 1 ≥ 2 ≥ 4 ≥ 8 ≥16 ≥ 32 ≥ 64 ≥128 wild-type + + + + + + + + + + + − Δefp + + + + + + + + − − − − ΔyjeK + + + + + + + + + − − − ΔyjeA + + + + + + + + + + + − ΔyfcM + + + + + + + + + + − −

Clindamycin ≥0 ≥ 16 ≥ 32 ≥ 64 ≥128 ≥256 ≥512 wild-type + + + + + + + Δefp + + + + + + + ΔyjeK + + + + + + + ΔyjeA + + + + + + + ΔyfcM + + + + + + +

Erythromycin ≥0 ≥4 ≥8 ≥16 ≥32 ≥64 ≥128 ≥256 ≥512 ≥1024 wild-type + + + + + + + + − − Δefp + + + + + + + − − − ΔyjeK + + + + + + − − − − ΔyjeA + + + + + + + − − − ΔyfcM + + + + + + + + − − ‘+’ Indicates growth of bacteria, ‘−‘ indicates inhibition of growth of bacteria. Table continued. Appendix 116

Name of Antibiotic MIC dilutions (μg/ml)

Gentamicin ≥ 0 ≥0.125 ≥0.25 ≥0.5 ≥1 ≥2 ≥ 4 ≥ 8 ≥16 ≥ 32 ≥ 64 ≥128 wild-type + + + + + + + − − − − − Δefp + + + + + − − − − − − − ΔyjeK + + + + + + − − − − − − ΔyjeA + + + + + + + + − − − − ΔyfcM + + + + + + + + + + − −

H2O2 ≥0 ≥8 ≥16 ≥32 ≥64 ≥128 wild-type + + + + + − Δefp + + + + − − ΔyjeK + + + + − − ΔyjeA + + + + − − ΔyfcM + + + + + −

Hygromycin A ≥ 0 ≥16 ≥ 32 ≥64 ≥128 ≥256 ≥512 ≥1024 wild-type + + + + + + − − Δefp + + + + + − − − ΔyjeK + + + + + + − − ΔyjeA + + + + + + + − ΔyfcM + + + + + + − −

Kasugamycin ≥0 ≥1 ≥2 ≥4 ≥8 ≥16 ≥32 ≥64 ≥128 ≥256 wild-type + + + + + + + + − − Δefp + + + + + + + + − − ΔyjeK + + + + + + + + − − ΔyjeA + + + + + + + + − − ΔyfcM + + + + + + + + − −

Linezolid ≥ 0 ≥32 ≥64 ≥128 ≥256 ≥512 ≥1024 wild-type + + + + + + − Δefp + + + + + − − ΔyjeK + + + + + − − ΔyjeA + + + + + + − ΔyfcM + + + + + + −

Norfloxacin ≥0 ≥0.125 ≥0.25 ≥0.5 ≥1 ≥2 ≥4 wild-type + + + + + − − Δefp + + + + + − − ΔyjeK + + + + − − − ΔyjeA + + + + − − − ΔyfcM + + + + + − − ‘+’ Indicates growth of bacteria, ‘−‘ indicates inhibition of growth of bacteria.

Table continued. Appendix 117

Name of Antibiotic MIC dilutions (μg/ml)

Rifampicin ≥0 ≥0.5 ≥1 ≥2 ≥4 ≥8 ≥16 ≥32 ≥64 wild-type + + + + + + + − − Δefp + + + + + + − − − ΔyjeK + + + + + + + − − ΔyjeA + + + + + + + − − ΔyfcM + + + + + + + − −

Spectinomycin ≥ 0 ≥1 ≥2 ≥4 ≥8 ≥16 ≥32 ≥64 wild-type + + + + + + − − Δefp + + + + + + − − ΔyjeK + + + + + − − − ΔyjeA + + + + + + − − ΔyfcM + + + + + + − −

Streptomycin ≥ 0 ≥0.125 ≥0.25 ≥0.5 ≥1 ≥2 ≥ 4 ≥ 8 ≥16 wild-type + + + + + + + − − Δefp + + + + + + + − − ΔyjeK + + + + + + + + − ΔyjeA + + + + + + + + − ΔyfcM + + + + + + + + +

Tetracycline ≥ 0 ≥0.25 ≥0.5 ≥1 ≥2 ≥ 4 ≥ 8 wild-type + + + + + − − Δefp + + + + − − − ΔyjeK + + + + − − − ΔyjeA + + + + − − − ΔyfcM + + + + + − −

Vancomycin ≥ 0 ≥0.5 ≥1 ≥2 ≥ 4 ≥ 8 ≥16 ≥32 ≥64 ≥128 ≥256 ≥512 wild-type + + + + + + + + + + + + Δefp + + + + + + + + + + + + ΔyjeK + + + + + + + + + + + + ΔyjeA + + + + + + + + + + + + ΔyfcM + + + + + + + + + + + + + + + + + + + + + + + +

Viomycin ≥ 0 ≥ 16 ≥ 32 ≥ 64 ≥128 ≥256 ≥512 wild-type + + + + + + + Δefp + + + + + + − ΔyjeK + + + + + + − ΔyjeA + + + + + + − ΔyfcM + + + + + + + ‘+’ Indicates growth of bacteria, ‘−‘ indicates inhibition of growth of bacteria.

Appendix 118

Supplementary Table 7. List of Primers for K34 Mutants.

Oligoname (Metabion International AG) Sequence for K34Q 5’-GAA TTC GTA AAA CCG GGT CAG GGC CAG GCA TTT GCT CGC-3’ rev K34Q 5’-GCG AGC AAA TGC CTG GCC CTG ACC CGG TTT TAC GAA TTC-3’ for K34N 5’-GAA TTC GTA AAA CCG GGT AAC GGC CAG GCA TTT GCT CGC-3’ rev K34N 5’-GCG AGC AAA TGC CTG GCC GTT ACC CGG TTT TAC GAA TTC-3’ for K34D 5’-GAA TTC GTA AAA CCG GGT GAC GGC CAG GCA TTT GCT CGC-3’ rev K34D 5’-GCG AGC AAA TGC CTG GCC GTC ACC CGG TTT TAC GAA TTC-3’ for K34E 5’-GAA TTC GTA AAA CCG GGT GAG GGC CAG GCA TTT GCT CGC-3’ rev K34E 5’-GCG AGC AAA TGC CTG GCC CTC ACC CGG TTT TAC GAA TTC-3’ for K143A 5’-GAT ACC GAT CCG GGC CTG GCA CGT GAT ACC GCA GGT ACT G-3’ rev K143A 5’-CAG TAC CTG CGG TAT CAC CTG CCA GGC CCG GAT CGG TAT C-3’ for K143R 5’-GAT ACC GAT CCG GGC CTG CGT GGT GAT ACC GCA GGT ACT G-3’ rev K143R 5’-CAG TAC CTG CGG TAT CAC CAC GCA GGC CCG GAT CGG TAT C-3’ for K34Q 5’-GAA TTC GTA AAA CCG GGT CAG GGC CAG GCA TTT GCT CGC-3’ rev K34Q 5’-GCG AGC AAA TGC CTG GCC CTG ACC CGG TTT TAC GAA TTC-3’