Lon-Dependent Proteolysis of Ffh, the Protein Component of the Signal Recognition Particle in Escherichia Coli

Lon-dependent proteolysis of Ffh, the protein component of the signal recognition particle in Escherichia coli

Dissertation to obtain the degree Doctor Rerum Naturalium (Dr. rer. nat.) at the Faculty of Biology and Biotechnology Ruhr University Bochum

International Graduate School of Biosciences Ruhr University Bochum Chair for Microbial Biology

submitted by Beate Sauerbrei

from Naumburg, Germany

Bochum August, 2019

First supervisor: Prof. Dr. Franz Narberhaus Second supervisor: Prof. Dr. Danja Schünemann

Lon-abhängige Proteolyse von Ffh, die Proteinkomponente des Signalerkennungspartikels in Escherichia coli

Dissertation zur Erlangung des Grades eines Doktors der Naturwissenschaften der Fakultät für Biologie und Biotechnologie der Ruhr-Universität Bochum

Internationale Graduiertenschule Biowissenschaften Ruhr-Universität Bochum LS Biologie der Mikroorganismen

vorgelegt von Beate Sauerbrei

aus Naumburg

Bochum August, 2019

Referent: Prof. Dr. Franz Narberhaus Korreferent: Prof. Dr. Danja Schünemann Danksagung

Ich bedanke mich herzlichst bei meinem Doktorvater Prof. Dr. Franz Narberhaus für die Ermöglichung dieser Doktorarbeit an seinem Lehrstuhl. Besonders möchte ich mich dafür bedanken, dass ich an dem interessanten Thema meiner Masterarbeit weiterarbeiten durfte. Zu dem bedanke ich für die Unterstützung und die konstruktiven Diskussion die zum Erfolg dieser Arbeit beigetragen haben.

Bei Frau Prof. Dr. Danja Schünemann bedanke ich mich für die freundliche Übernahme des Korreferats und das stetige Interesse an dem Thema meiner Arbeit. Insbesondere möchte ich mich für die bereichernde Diskussion während der gesamten Arbeit bedanken. Außerdem bedanke ich mich für die Bereitstellung der cpSRP54-Konstrukte, die einen bedeutenden Teil zu dieser Arbeit beigetragen haben.

Ein ganz besonderer Dank geht an alle ehemaligen und jetzigen Mitarbeiter der Arbeitsgruppe für „Reguliert Proteolyse“. Dazu gehören Lisa-Marie Bittner, Jan Arends, Blanka Kutscher, Fabian Müller, Alexander Kraus, Simon Brückner und Anna-Maria Möller. Ein ganz großer Dank geht an Fabian Müller und Alexander Kraus für eine bereichernde Zusammenarbeit, den hilfreichen Diskussionen, dem Spaß im Labor und Büro und den unterhaltsamen Stunden während unserer Mittagspausen. Für die engagierte Mitarbeit an diesem Thema möchte ich bei meinen Studenten Fitore Morina, Simon Brückner und Hendrik Strotmeier danken.

Bei dem gesamten Lehrstuhl Biologie der Mikroorganismen möchte ich mich für eine angenehme Arbeitsatmosphäre bedanken. Ich möchte mich bei Dr. Bernd Masepohl für wertvolle Tipps und Diskussion aller Art bedanken. Ein besonders Dank gilt Petra Krämer für die Unterstützung in organisatorischen Fragen und Hanno Böddinghaus für die Unterstützung im IT-Bereich.

Auch möchte ich mich von ganzem Herzen bei meinen Freunden bedanken. Ein ganz großer Dank gebührt dabei Kerstin und Lisa die neben dem Studium und der Doktorarbeit immer für mich da waren und mein Leben neben der Uni sehr bereichert haben.

Ein ganz persönlicher Dank geht an meinen Eltern Gaby und Volker für Ihre großartige Unterstützung während meines gesamten Studiums und ohne die das alles nicht möglich geworden wäre. Ich möchte einfach nur DANKE sagen! Ein ganz besonderer Dank geht an meine Schwester Ellen für die andauernde Unterstützung in allen Lebenslagen und die zahlreichen Kurztrips, die wir während der gesamten Zeit unternommen haben.

Table of contents

Abbreviations ...... I List of figures ...... IV List of tables ...... V A Introduction ...... 1 1. Regulated proteolysis by AAA+ proteases ...... 2 2. Structure and function of the Lon protease ...... 3 2.1 Structure of the Lon proteases...... 3 2.2 Substrate recognition by the Lon protease ...... 6 2.3 Biological relevance of the Lon protease ...... 8 2.3.1 Involvement of the Lon protease in various stress responses ...... 8 2.3.2 Lon protease in motility and biofilm formation ...... 10 2.3.3 Lon proteolysis in cell cycle, TA systems and persister cell formation .. 11 2.3.4 Lon involvement in biosynthetic pathways and metal homoeostasis ...... 12 2.3.5 Lon in protein quality control ...... 13 3. Principles of protein targeting in bacteria ...... 14 3.1 Post-translational protein targeting (Tat and Sec pathway) ...... 14 3.2 Co-translational protein targeting (SRP pathway) ...... 16 4. The universally conserved signal recognition particle ...... 18 4.1 SRP complex in different species ...... 18 4.2 Structure of the minimal SRP complex and its receptor ...... 20 4.3 Structur and function of the chloroplast SRP complex in higher plants ...... 22 5. Objectives of this work...... 26 B Materials and methods ...... 27 1. Materials ...... 28 1.1 Bacterial strains ...... 28 1.2 Plasmids ...... 28 1.3 Oligonucleotides ...... 30 1.3 Laboratory equipment ...... 31 1.4 Chemicals ...... 32 1.5 Enzymes ...... 34 1.6 Kits ...... 34 A

Table of contents

1.7 Antibodies ...... 35 1.8 Media and media supplements ...... 35 2. Software...... 36 3. Methods ...... 36 3.1 Microbiological methods ...... 36 3.1.1 E. coli cell culture ...... 36 3.1.2 Determination of bacteria count ...... 36 3.1.3 Preparation of competent E. coli cells ...... 36 3.1.4 Transformation of E. coli ...... 37 3.1.5 Transformation efficiency ...... 37 3.1.6 Microscopy studies ...... 38 3.2 Molecular biological methods ...... 38 3.2.1 Polymerase chain reaction ...... 38 3.2.2 DNA restriction ...... 39 3.2.3 DNA ligation ...... 39 3.2.4 Preparation of plasmid DNA from E. coli ...... 40 3.2.5 Plasmid preparation from E. coli by commercial kits ...... 41 3.2.6 Agarose gel electrophoresis ...... 41 3.2.7 Extraction of DNA fragments from agarose gel ...... 41 3.2.8 Cloning strategies ...... 42 3.2.8.1 Cloning by restriction and ligation ...... 42 3.2.8.2 Site-directed mutagenesis ...... 42 3.2.9 DNA Sequencing ...... 42 3.3 Protein biochemical methods ...... 43 3.3.1 Expression and solubility studies ...... 43 3.3.2 In vivo degradation experiments ...... 44 3.3.3 Purification of poly-histidine fusion proteins ...... 44 3.3.3.1 Protein overproduction ...... 44 3.3.3.2 Cell disruption by FrenchPress ...... 45 3.3.3.3 Nickel affinity chromatography ...... 45 3.3.4 Protein concentration quantification by Bradford assay ...... 46 3.3.5 In vitro degradation experiments ...... 47

Table of contents

3.3.6 Preparation of protein extracts ...... 47 3.3.7 Denaturing SDS polyacrylamide gel electrophoresis ...... 48 3.3.8 Coomassie staining of polyacrylamide gels ...... 48 3.3.9 Western transfer and immunological detection of proteins ...... 49 3.3.10 Calculation of half-lives ...... 50 C Results ...... 51 1. Investigations on the degradation mechanism of Ffh ...... 52 1.1 Growth phase-dependent proteolysis of plasmid-derived Ffh ...... 52 1.2 Lon is the major protease responsible for Ffh degradation ...... 55 1.3 Cellular Ffh is a stable protein under different stress conditions ...... 56 1.4 The Ffh amount is modulated during bacterial growth ...... 58 1.5 Impact of RpoH on Ffh proteolysis ...... 60 2. Investigations on the recognition mechanism of Ffh by the Lon protease ...... 63 2.1 The Ffh M domain is recognized by the Lon protease...... 63 2.2 Plasmid-encoded M domain induces a lethal phenotype in E. coli...... 64 2.3 Impact of point mutations on stability of Ffh and cpSRP54 from A. thaliana .... 67 D Discussion ...... 71 1. Modulation of the Ffh degradation mechanism in E. coli ...... 72 1.1 Growth phase-dependent Ffh proteolysis is related to its cellular amount ...... 72 1.2 Ffh proteolysis is affected by the sigma factor RpoH ...... 78 2. Recognition mechanism of Ffh by the Lon protease ...... 82 2.1 Lon protease recognizes Ffh via its M domain ...... 82 2.2 Ffh recognition by Lon requires an intact 4.5S RNA-binding motif ...... 84 3. ATP-dependent proteolysis in protein translocation ...... 87 E Summary ...... 90 F Zusammenfassung ...... 93 G References ...... 96 H Appendixes ...... 128 I Publications ...... 141 J Lebenslauf ...... 143 K Erklärung ...... 145

Abbreviations

Abbreviations (p)ppGpp guanosine (penta) or tetraphosphate

a. dest. aqua destilla ta AAA+ ATPase associated with a variety of cellular activities

AC adenylate cyclase

ADP adenosine diphosphate

AHT anhydrotetracycline

Amp ampicillin

Ank ankyrin

approx. approximately

APS ammonium persulfate

ATP adenosine triphosphate

B. th. bachelor thesis

BSA bovine serum albumin

CD chromodomain

Cm chloramphenicol

cp chloroplast

C-terminus carboxy terminus

dATP deoxyadenosine triphosphate

dCTP deoxycytidine triphosphate

dGTP deoxyguanosine triphosphate

DMF dimethylformamide DMSO dimethylsulfoxide

DNA deoxyribonucleic acid

dNTP deoxynucleoside triphosphate

DTT dithiothreitol

dTTP deoxythymidine triphosphate

Abbreviations

EDTA ethylenediaminetetraacetic acid

EtBr ethyl bromide

EtOH ethanol

EV empty vector

Fig. figure

GDP guanosine diphosphate

GTP guanosine triphosphate

HIV human immunodeficiency viruses

HRP horseradish peroxidase

HTH helix-turn-helix

IBD insertion-box domain

IMP inner membrane protein

IPTG isopropyl β-D-1-thiogalactopyranoside

KAc potassium acetate

Km kanamycin

LB lysogeny broth

LHCP light-harvesting chlorophyll a/b-binding protein

M. th. master thesis

MeOH methanol mRNA messenger RNA

MTS membrane targeting sequence n.d. not detectable nt nucleotide

N-terminus amino terminus

OD optical density

PCR polymerase chain reaction

Pi inorganic phosphate

Abbreviations

PMF proton motive force

PMSF phenylmethylsulfonyl fluoride polyP inorganic polyphosphates

PPP protein sample buffer

RNA ribonucleic acid

RNC ribosome-nascent chain rRNA ribosomal RNA

RT room temperature

SAS signal-anchored sequences

SDS sodium dodecyl sulfate

SDS-PAGE sodium dodecyl sulfate–polyacrylamide gel electrophoresis

SIMIBI signal recognition particle, MinD, and BioD

Sp spectinomycin

SR signal recognition particle receptor

SRP signal recognition particle

SSD sensor- and substrate discrimination

T½ half-life

TA toxin-antitoxin

Tab. table

TEMED tetramethylethylenediamine

TIC translocon on the inner chloroplast membrane

TM transmembrane tmRNA transfer messenger RNA

TOC translocon on the outer chloroplast membrane unpubd. unpublished

UV ultraviolet

WT wild-type

III

List of figures

List of figures Fig. A-1: Operating principle of AAA+ proteases...... 3 Fig. A-2: Domain arrangement of the LonA and LonB protease...... 4 Fig. A-3: Biological role of the Lon protease in various bacteria...... 9 Fig. A-4: Post-translational protein targeting in bacteria...... 15 Fig. A-5: Co-translational protein targeting in bacteria...... 17 Fig. A-6: Schematic representation of the SRP complex in different species...... 19 Fig. A-7: Structural overview of Ffh and its receptor FtsY...... 21 Fig. A-8: Post- and co-translational protein targeting in chloroplasts of higher plants...... 23 Fig. C-1: Degradation of cellular and plasmid-derived Ffh at different growth phases...... 53 Fig. C-2: Half-lives of plasmid-derived Ffh in AAA+ protease deficient E. coli strains...... 55 Fig. C-3: Degradation of cellular Ffh after heat stress...... 57 Fig. C-4: Relative protein amounts of cellular Lon, Ffh and plasmid-derived Ffh...... 59 Fig. C-5: Impact of the sigma factor RpoH on Ffh proteolysis ...... 61 Fig. C-6: Degradation of the Ffh domains (NG and M domain)...... 64 Fig. C-7: Transformation efficiencies of plasmid-encoded Ffh and its functional domains. ... 66 Fig. C-8: SRP structure and localization of the evolutionary point mutations...... 68 Fig. C-9: Transformation efficiencies and degradation of cpSRP54 in E. coli...... 69 Fig. D-1: Cellular Ffh amount is modulated by the cytosolic Lon protease...... 77 Fig. D-2: Modulation of Ffh stability in heat shock response...... 81 Fig. D-3: ATP-dependent proteolysis in protein translocation...... 88 Fig. H-1: Half-lives of the corresponding parental strains for AAA+ deficient strains...... 131

Fig. H-2: Purification of His6-Ffh and Lon-His6 and in vitro stability of His6-Ffh...... 132

Fig. H-3: Degradation of cellular Ffh after plasmid-derived His6-Lon overproduction...... 133 Fig. H-4: Degradation of cellular Ffh during oxidative stress conditions...... 134 Fig. H-5: Degradation of cellular Ffh after rifampicin treatment...... 135 Fig. H-6: Degradation of cellular Ffh in ΔrpoH...... 136

Fig. H-7: Light microscopy of MC4100 carrying His6-tag or One hybrid constructs...... 137 Fig. H-8: Deleted region within the M domain after transformation in Δlon...... 138 Fig. H-9: Half-lives of Ffh variants with evolutionary point mutations...... 139

Fig. H-10: Degradation of His6-FusA in MC4100...... 140

List of tables

Tab. B-1: E. coli strains used in this study...... 28 Tab. B-2: Vectors used in this study...... 29 Tab. B-3: Plasmids used in this study...... 29 Tab. B-4: Oligonucleotides used in this study...... 30 Tab. B-5: Laboratory equipment used in this study...... 31 Tab. B-6: Chemicals used in this study...... 32 Tab. B-7: Enzymes used in this study...... 34 Tab. B-8: Kits used in this study...... 34 Tab. B-9: Antibodies used in this study...... 35 Tab. B-10: Media supplements used in this study...... 35 Tab. B-11: Software used in this study...... 36 Tab. B-12: Growth condition for protein overproduction...... 44 Tab. H-1: List of references of Lon substrates in Fig. A-3...... 129

Introduction

A Introduction

A Introduction 1. Regulated proteolysis by AAA+ proteases Bacteria have developed various strategies for adaptation to changing environmental conditions such as temperature, nutrient availability, osmolarity or pH value. They can cope with various stress conditions and colonize a variety of ecological niches. This involves the regulation of gene expression on transcriptional, translational and posttranslational level, whereby each of these strategies has a direct or indirect effect on the protein homeostasis in the cell. On the posttranslational level, the principal of proteolysis is a rapid and effective strategy to adapt the cellular proteome to certain conditions. Due to its irreversibility, this process needs to be controlled closely. On the one hand, general proteolysis includes the degradation of aberrant and damaged proteins and is responsible for the quality control of proteins. On the other hand, regulated proteolysis degrades native folded and functional proteins in order to adapt the proteome to the environment under certain conditions. Proteolysis relies on ATP-independent as well as ATP-dependent proteases, but 90 % of protein degradation in the cytoplasm occur energy-dependently [228]. In the Gram-negative enterobacterium Escherichia coli, regulated proteolysis can occur by the five ATP-dependent AAA+ (ATPase associated with a variety of cellular activities) proteases ClpXP, ClpAP, HslUV, Lon and FtsH. FtsH is the only essential and membrane-anchored protease in E. coli while all others are located in the cytosol and being non-essential. AAA+ proteases share structural and functional similarities. Figure (Fig.) A-1 shows their basic structure and mechanism of proteolysis. In general, AAA+ proteases form a homooligomeric barrel-shaped complex with a central pore. Each monomer consists of an unfoldase including the classical AAA+ module and a proteolytically active protease domain. AAA+ proteases degrade specific substrates, whereby ATP is the driving force (reviewed in [15, 263, 316]). The proteolysis mechanism includes four steps: (I) substrate recognition, (II) unfolding, (III) translocation into the proteolytic chamber and finally (IV) degradation. The initial recognition step comprises an interaction using the unfoldase domain of the AAA+ protease and its substrate. Substrate binding occurs within the axial pore of the oligomeric AAA+ ring and is ATP-independent. Substrate specificity is mediated by recognition motifs, so-called degrons. The localization of the degron within the substrate is highly variable. Recognition motifs are located either at the N- or C-terminus or at internal regions. They can be structured or unstructured. Also, substrates can be recognized indirectly via additional domains or adaptor proteins.

A Introduction

Fig. A-1: Operating principle of AAA+ proteases. AAA+ proteases exhibit an unfoldase (blue) for substrate recognition and unfolding as well as a protease domain (grey) for degradation of the protein into small peptides. Initial recognition of AAA+ protease substrates is mediated by the unfoldase domain and is an ATP-independent step. Repetitive ATP binding and hydrolysis induces conformational changes of the AAA+ module within the unfoldase and results in unfolding and translocation of the substrate into the proteolytic chamber of the protease domain. Finally, substrates are degraded ATP- independently into smaller peptide fragments by sequestration of the proteolytically active sites. Revised from: [38, 316]

Once the interaction of AAA+ protease and substrate has been initiated, ATP-dependent unfolding and translocation of the substrate into the proteolytic chamber follows. The substrate unfolding results from conformational changes within the AAA+ module. This process is driven by repetitive ATP binding and hydrolysis converting chemical energy into mechanical force. These events induce pulling movements, whereby the substrate moves into the central pore. ATP consumption for substrate unfolding and translocation can range from 20 to 500 molecules of ATP [15]. Finally, the unfolded protein is degraded ATP-independently into small peptides by sequestration of the proteolytically active sites within the protease domain [133, 134, 220]. Substrates are cleaved into peptides of about 5 to 25 amino acids in length. Since the degradation mechanism of a Lon substrate was investigated in this dissertation, the structure and biological relevance based on its substrate diversity will be described in more detail below.

2. Structure and function of the Lon protease 2.1 Structure of the Lon proteases The first discovered ATP-dependent protease was the Lon protease, which is highly conserved in all three domains of life. Lon is named according to the observed phenotype of an E. coli lon mutant which forms long undivided filaments, contains multiple nucleoids, and is sensitive to

A Introduction

UV irradiation [3, 155]. The cytosolic Lon protease assembles into a homohexameric ring- shaped structure with a central pore. Like the FtsH protease, a single gene encodes Lon and consequently, a single polypeptide constitutes both domains (unfoldase and protease domain). For E. coli Lon, each polypeptide comprises 784 amino acids with a mass of about 87 kDa. Oligomerization of six monomers is ATP-independent and requires the presence of Mg2+ ions [132]. Among Lon proteases, two classes, LonA and LonB, were defined by sequence analysis [304]. Both classes differ from each other in their structure, as shown in Fig. A-2. LonA is found predominantly in bacteria (including E. coli) and eukaryotes, whereas LonB was identified mainly in archaea and a few bacterial species.

Fig. A-2: Domain arrangement of the LonA and LonB protease. Lon proteases represent the classical AAA+ protease consisting of the unfoldase (blue) and protease domain (grey). The AAA+ module can be divided in two domains, a large α/β domain and a small α domain. Conserved motifs of the AAA+ module are the Walker A (yellow), Walker B (orange), Sensor-1 (light blue), arginine finger (white) and Sensor-2 (green). The protease domain harbors the proteolytically active dyad consisting of serine (S) and lysine (K). Lon B contains an additional transmembrane (TM) domain within its unfoldase. The schematic representation of LonA and LonB is based on the summarized Lon structures described in [303].

LonA consists of three domains: the LonA-specific N-terminal N domain, the central ATPase domain containing the classical AAA+ module and the C-terminal protease domain. The variable N domain is probably responsible for substrate discrimination [92]. Structure analysis suggests a highly charged region (amino acid 211 to 271) forming a coiled-coil domain and mediating a potential substrate discriminatory activity within the N-domain. Furthermore, the N-domain plays a crucial role in assembly and control of the Lon protease because ATPase and protease activity were shown to be altered by deleting parts of the N domain [63, 307]. More recent studies have shown that Lon is available in the cell as a hexamer and as a dodecamer

A Introduction

[368]. A head-to-head formation of two hexamers at their respective N-domains mediates the dodecamer assembly. The N-domains form gates, which mediate substrate selectivity. Compared to a hexamer, a Lon dodecamer is less active in the degradation of large substrates but more active on small substrates. Thus, the balance of hexamer and dodecamer ratio is an important physiological factor for the control of Lon activity. More than 20% of all known Lon proteases are LonB proteases that consist of the conserved AAA+ module and the protease domain but lack the LonA-like N domain [159, 304, 379]. Characteristic for the LonB subfamily is a transmembrane domain, which is located between the nucleotide-binding motifs Walker A and Walker B within the α/β domain of the unfoldase. One or two transmembrane helices are responsible for anchoring the LonB protease in the membrane [304]. LonB often can be found in organisms that do not possess the membrane- anchored FtsH and it is assumed that LonB compensates for the function of FtsH [379]. Beside archaea, LonB is found in other bacterial species, e.g. Pseudomonas aeruginosa, Thermotoga maritima and Bacillus subtilis that possess both types of the Lon protease. The AAA+ module comprises a large α/β Rossmann fold (α/β domain) and a small α-helical domain (α domain) [286, 330, 385]. The α/β domain contains conserved motifs for nucleotide binding called Walker A (GxxxxxGKT/S; x= any amino acid) and Walker B (hhhhDE; h = any hydrophobic amino acid) [372]. The Walker A motif is critical for ATP binding and hydrolysis and the Walker B motif activates a water molecule for nucleophilic attack on the γ-phosphate during ATP hydrolysis [1, 140, 159, 335]. Consequently, mutation of the conserved and catalytically active glutamate of the Walker B motif still allows ATP binding but prevents ATP hydrolysis [13, 75, 191, 380]. The Sensor-1 motif in the AAA+ module contains a polar residue and an arginine finger located within the α/β domain. The polar residue of Sensor-1 plays an important role in the coordination of a water molecule towards a nucleotide by forming a hydrogen-bonding network [208, 399]. Sensor-1 and the glutamate of the Walker B motif coordinate the orientation of the nucleophilic water molecule which is attacked by the negatively charged γ-phosphate of the bound ATP to induce the ATPase activity [266]. Arginine fingers are a common feature in AAA+ proteins. After protease assembly, the arginine of a single subunit reaches into the adjacent subunit [140]. Thus the arginine-finger is involved in oligomerization and ATP sensing and hydrolysis. The small α domain contains at least the conserved Sensor-2. This motif contains an arginine, which further is involved in ATP hydrolysis and substrate remodeling [159, 253]. It also participates in nucleotide binding because it directly interacts with the γ-phosphate of ATP [266].

A Introduction

The protease domain contains a proteolytically active Ser679-Lys722 dyad within the consensus sequence PKDGPSAG and (K/R)(E/D)KXU(A/S), respectively [305]. Mutational analysis revealed serine at amino acid position 679 as catalytically active and further sequence comparisons provided evidence for the catalytically active Ser679-Lys722 dyad [8, 35, 305, 331]. Lon belongs to the class of serine proteases. However, due to its proteolytic center, Lon differs from classical serine proteases, which usually contain a catalytically active triad of Ser-His- Asp [8, 118, 304].

2.2 Substrate recognition by the Lon protease Since there is no Lon-specific recognition motif within the substrates, the recognition is highly diverse. In the following, well-studied substrate recognition motifs for quality control and regulated proteolysis are presented. Lon is involved in the quality control of aberrant proteins [135]. For this purpose, Lon can distinguish aggregated proteins from natively folded proteins by their unstructured regions upon denaturation. In native proteins, hydrophobic amino acids often are hidden inside a protein by their tertiary structure and are inaccessible to proteases [135]. Lon recognizes misfolded proteins, whose tertiary structure is partially disrupted leading to easily accessible hydrophobic and aromatic residues. A well-known example of substrate recognition by Lon for quality control is the so-called SsrA-tag. This tag consists of the amino acids AANDENYALAA and is rich in non-polar amino acids. It is attached to the C-terminus of prematurely terminated proteins during translation at the stalled ribosome. The formation and function of the SsrA-tag, as well as its interaction with Lon will be explained in more detail below (see section A-2.3.5). As part of regulated proteolysis, Lon and other AAA+ proteases recognize fully folded proteins by their structured accessible N- or C-terminus or unstructured regions within the folded protein. Often, naturally unstructured regions are recognized by hydrophobic amino acids, e.g. λN and the antitoxin CcdA (TA: CcdA/CcdB) [227, 366]. N- or C-terminal degradation signals are another possibility for substrate recognition by Lon. A well-studied example is the cell division inhibitor SulA, which is recognized by a C-terminal degradation signal consisting of 20 amino acids (ASSHATRQLSGLKIHSNLYH) [158]. In general, the C-terminal SulA- degron has a moderate hydrophobicity and the underlined amino acids are the core recognition motif. The aromatic histidines within the C-terminal end and the tyrosine at the penultimate position are critical for Lon recognition as shown by site-directed mutagenesis [136, 157]. Examples for Lon substrates recognized by an N-terminal degradation signal are UmuD and SoxS in E. coli as well as HemA from Salmonella typhimurium [119, 326, 376]. Lon recognizes 6

A Introduction

UmuD by the amino acid 12 to 31 (IVTFPLFSDLVQCGFPSPAA) at the N-terminal site of the protein [119]. The primary and auxiliary recognition site contains hydrophobic and aromatic amino acids and is located between residue 15 to 19 (FPLF) and 26 to 29 (FPSP). Furthermore, the prolines acting as helix breakers, make these regions suitable for Lon recognition. Fusion of the N-terminal protease-sensitive amino acid region to an otherwise stable protein results in Lon-dependent proteolysis of the fused product. Thus, the defined region is sufficient for Lon proteolysis. For HemA recognition, the N-terminal 18 amino acids (MTLLALGINHKTAPVSLR) are sufficient for Lon recognition and subsequent proteolysis [376]. It is proposed that HemA proteolysis only occurs when a sufficient heme concentration is present in the cell. Thus, it is a heme-dependent mechanism. In contrast, the recognition motif of SoxS (MSHQKIIQDLIAWIDEHIDQP), the transcription activator of the superoxide stress regulon SoxRS, is rich in polar amino acids playing a critical role in Lon proteolysis [7, 326]. Besides, degradation of SoxS by the Lon protease in vitro is partially prevented by DNA binding to Sox box DNA or RNA polymerase [325]. In contrast, the Lon substrates TrfA and RepE require Lon-DNA interaction for degradation [177]. Moreover, substrate interaction with a ligand or adapter protein may cause protection against proteolysis. The transcriptional regulator ZntR is a substrate of Lon and ClpXP proteolysis [283]. Binding of DNA and Zn2+ protect ZntR towards degradation. Furthermore, this mechanism is common to toxin-antitoxin systems because an antitoxin can be stabilized upon binding to the toxin and vice versa (e.g., CcdB/CcdA). A further mechanism for Lon recognition is mediated by a bipartite degron as shown for the replication initiator DnaA in Caulobacter cresentus [215]. The AAA+ domain of DnaA seems to be sufficient for Lon binding, but proteolysis only occurs also if further regions of the N-terminus are recognized by the protease. Another possibility for Lon recognition is based on the temperature-sensitivity of substrates. For instance, the first enzyme of the methionine biosynthesis HTS (homoserine trans-succinylase) was shown to be more unstable at higher temperatures [34]. In Yersinia pseudotuberculosis the transcriptional virulence regulator RovA becomes accessible for Lon and ClpP degradation upon a temperature-shift from 25 °C to 37 °C. This effect can be attributed to RovA’s loss of DNA-binding ability due to conformational changes [145]. These examples illustrate the high variability in Lon substrate recognition mechanisms. Based on the substrates already mentioned, Lon has been shown to play a role in various cellular processes. The following section focuses on the biological role of Lon by presenting further substrates of different cellular processes in various organisms.

A Introduction

2.3 Biological relevance of the Lon protease Lon is non-essential in E. coli but a lon mutant exhibits a pleiotropic phenotype. A lon-deficient E. coli strain is more sensitive against UV radiation, DNA damaging agents and has a defect in phage development. Colonies of a lon mutant exhibit a mucoid phenotype. These diverse phenotypes demonstrate that Lon has a strong impact on a variety of cellular processes. The different phenotypes can be explained by different Lon substrates. Besides its regulatory function in biological processes, Lon also has a major role in the quality control of proteins, since aberrant proteins accumulate in a lon mutant. In the following section, the biological role will be described based on individual Lon substrates and the role of general proteins in quality control. An overview of known Lon substrates is given in Fig. A-3.

2.3.1 Involvement of the Lon protease in various stress responses A lon mutant is more sensitive to DNA-damaging agents and UV radiation since Lon controls various regulators of the SOS response at the posttranslational level. After DNA damage, the SOS response induces cell cycle arrest and subsequently DNA repair and mutagenesis [285]. This system is under the control of RecA and the transcriptional repressor LexA. RecA controls LexA by autocatalytic cleavage, which induces the SOS response. Lon and ClpXP are responsible for LexA proteolysis. While the N-terminal fragment is subjected to ClpXP and the C-terminal fragment to Lon and ClpXP cleavage [251]. For DNA damage repair, >50 genes are upregulated and several proteins of the SOS response are subject to ATP-dependent proteolysis by Lon, ClpXP and HslUV (e.g., SulA, UmuC/UmuD, RuvB and RecA) [103, 119, 252, 322]. The cell division inhibitor SulA is expressed concomitantly upon DNA damage. Septum formation is blocked by binding of FtsZ to SulA, thus cell cycle is arrested. To resume cell division after DNA repair, SulA is efficiently degraded by Lon and HslUV (reviewed in [364]). The umuCD operon is expressed in late stages of SOS response in order to bypass translesion of error-prone DNA polymerases because DNA-polymerases III can be blocked by DNA lesions (reviewed in [364]). UmuC and UmuD assemble into the DNA polymerase V allowing the bypass of DNA lesions. Both proteins are under the posttranslational control of Lon. Further Lon substrates of the SOS response system are RuvB, RecA and SymE [178, 252]. Lon plays also a role in the superoxide stress response (SoxRS regulon) triggered by reactive oxygen species. The 2Fe-2S cluster of constitutively expressed SoxR becomes oxidized converting the regulator into a transcriptional activator. This initiates expression of soxS, coding for another regulator [80, 84, 113, 114, 149–151, 262, 282, 393]. SoxS induces transcription of the superoxide regulon [56, 80, 111, 112, 214]. 8

A Introduction

Fig. A-3: Biological role of the Lon protease in various bacteria. The figure presents an overview of several Lon substrates and illustrates that the Lon protease has an impact on various cellular processes. In regulated proteolysis, Lon is involved in various stress responses, virulence, pathogenicity, biosynthesis pathways, phage development, cell cycle, toxin-antitoxin system, motility, biofilm formation, metal homeostasis and quorum sensing. In addition, Lon plays an important role in the degradation of proteins as part of the cellular quality control system. The respective references of the Lon substrates are listed in the appendix (Tab. H-1).

After the removal of reactive oxygen species transcription is shut down by Lon- and FtsH- dependent proteolysis of SoxS [126]. A similar system is known for the Lon substrate MarA, the regulator of the Mar regulon [126]. Hence, Lon is part of the regulation of antibiotic resistance. Lon regulates peroxide resistance in B. subtilis by PerR proteolysis. PerR is a transcriptional repressor and a peroxide sensor. If the hydrogen peroxide concentration increases, iron or manganese at the metal-binding site is oxidized. Oxidized PerR has a more open conformation and is subjected to Lon-dependent degradation [4]. Lon is involved in the regulation of the glutamate-dependent acid resistance by degrading the transcription regulator GadE, which positively regulates gadA, gadB and gadC expression. Degradation of GadE shuts down acid resistance response at a neutral pH to prevent expression of acid resistance genes in stationary growth phase [148]. The protein turnover in stationary growth phase is increased, whereas proteins are more stable in fast-growing cells [224]. During stationary growth phase

A Introduction various stress conditions can occur like amino acid starvation, nitrogen limitation, fatty acid limitation and heat shock, which trigger the synthesis of the alarmon ppGpp [22, 108, 142, 369]. This process is called stringent response. During this process, inorganic polyphosphates (polyP; polymer of hundreds of phosphate residues) accumulate in the cell and participate in Lon- dependent proteolysis of free ribosomal protein (e.g S2, L9 and L13) [195, 196]. The ATPase domain of Lon is able to bind polyP and the phosphate polymer acts as an adaptor molecule to stimulate degradation. Binding of polyP to Lon inhibits the formation of a Lon-DNA complex and Lon binds with higher affinity to polyP than to DNA [256]. Interaction between Lon and DNA is also suggested to stimulate ATPase activity [61, 70].

2.3.2 Lon protease in motility and biofilm formation Bacteria are constantly striving to adapt to new environmental conditions in order to settle in new ecological niches. Besides a general modulation of the gene expression profile, the regulation of motility and biofilm formation is essential. It is known that some Lon substrates are involved in these processes. Swimming bacteria such as E. coli or related species move on using flagella. Lon is involved in the regulation of flagella synthesis by degrading certain master regulators, which results in a distinct phenotype. For example, the Gram-negative enterobacterium Proteus mirabilis developed a hyperswarming phenotype by insertion of a mini-Tn5 transposon in the lon gene [72]. The flagellar regulon comprises about 60 genes and can be divided into three classes based on their expression mode. The transcriptional regulator FlhDC is a master activator that positively regulates class II genes of the flagellar regulon. Among these genes is the sigma factor FliA (σ28), which in turn regulates the expression of class III genes. The stability of both regulators, FlhDC and FliA, depends on proteolysis by ATP-driven degradation. FlhDC in P. mirabilis and S. typhimurium is degraded by Lon and ClpXP, respectively [71, 351]. Thus for E. coli, the same principle is assumed. FliA is subject to Lon-dependent proteolysis [19]. FliA binds the negative regulator of flagellin synthesis FlgM preventing interaction of the sigma factor with the RNA polymerase. Thus, FliA-dependent transcription of class II and III genes of the flagellar regulon is repressed. In addition, this interaction of FlgM-FliA prevents Lon-specific proteolysis of free FliA. Since Lon degrades non-bound FliA to FlgM, it is also involved in the repression of flagellar gene expression. Recently, the flagellar hook protein FlgE was found as a Lon substrate [9]. Bacteria change their physiology when they contact a solid surface. Recently, it has been shown that the flagellar activator protein SwrA in B. subtilis is subject to Lon-dependent proteolysis [244]. Furthermore, the proteolysis of SwrA is dependent on the swimming motility inhibitor 10

A Introduction

A protein, SmiA. Therefore, this is the first known example of adaptor-mediated Lon proteolysis. For P. aeruginosa and Vibrio cholerae it was shown that a lon-deficient strain is impaired in biofilm formation. In P. aeruginosa lon mutants biofilm formation is completely defective [225]. In contrast, V. cholerae produces biofilms but the structure is affected in a lon mutant [297]. Surprisingly, the absence of the Lon protease increases the swimming motility and hence possibly allowing colonization of new ecological niches. However, a Lon substrate causing the defect in biofilm formation is not known yet.

2.3.3 Lon proteolysis in cell cycle, TA systems and persister cell formation Besides the presented cellular processes, Lon is also involved in the control of cell cycle, toxin- antitoxin systems and persister cell formation. C. cresentus is a model organism for cell cycle, cell division and cellular differentiation because it exists as both, motile swarming as well as sessile cells. To terminate cell division in stalked cells, the DNA is fully methylated by the DNA methyltransferase CcrM. Subsequently, excess CcrM is continuously degraded by the Lon protease to reach a low CcrM level in the cell [392]. Moreover, CcrM is under positive control of CtrA, which in turn is subject to ClpXP-dependent proteolysis [167]. Further Lon substrates in C. cresentus are DnaA and SciP [120, 171]. Cellular processes like DNA replication or cell division can be controlled by toxin-antitoxin systems (TA systems), which consist of a bacteriostatic toxin and a toxin-neutralizing antitoxin [384, 396]. Lon protease is involved in the control of TA systems through specific degradation of either toxin or antitoxin. This includes the following TA systems: CcdA/CcdB, RelE/RelB, RnlA/RnlB, YefM/YoeB, MazE/MazF, HipA/HipB, DinJ/YfaQ and MqsR/MqsA (underlined protein correspond to Lon substrate) [67, 68, 139, 164, 184, 188, 284, 355, 365, 378]. The bacteriostatic effect of some toxins is directly related to the formation of dormant persister cells [181, 324]. Persister cells are dormant, non-dividing and multidrug-tolerant cells of a bacterial population, which are capable to survive several stress conditions such as nutrient deficiency. The MqsR/MqsA TA system is directly linked to persisters because disruption of mqsA results in increased persister cell formation [339]. In turn the MqsR/MqsA TA system regulates the toxin CspD [184]. The cold shock-like protein CspD acts as a DNA replication inhibitor, which is induced in stationary growth phase [397]. It has been shown that CspD is subjected to Lon proteolysis and dependent on growth phase and growth rate, while degradation of CspD occurs slowly in slow growing cells and is rapid in fast growing cells [200]. CspD degradation is assumed to reverse the toxic effect in stationary growth phase and possibly in persister cells to revive dormant cells.

A Introduction

2.3.4 Lon involvement in biosynthetic pathways and metal homoeostasis A lon mutant exhibits a mucoid phenotype as mentioned above. The mucoid cells are formed by stabilization of the Lon substrate RcsA, a transcription regulator of the capsular polysaccharide (cps) operon [122, 123, 352]. In the presence of Lon, RcsA is unstable and transcription of the cps operon is remarkably low. In contrast, RcsA stabilization in the absence of Lon leads to a high transcription of the cps operon and thus to an increase of colanic acid capsular polysaccharides, which causes the mucoid phenotype. Assembly of a RcsA-RcsB heterodimer protects RcsA against proteolysis [336]. Overexpression of the hslUV protease in a lon mutant suppresses the mucoid phenotype suggesting that RcsA proteolysis depends on the HslUV protease [59, 194]. Moreover, Lon plays a role in regulation of amino acid biosynthesis. The homoserine trans- succinylase (HTS), encoded by metA, is the first enzyme of the methionine biosynthesis pathway and is subject to Lon-dependent proteolysis [34]. HTS is degraded under heat-shock conditions and is more stable at moderate temperatures. Gene expression of metA is repressed by the methionine response repressor MetJ and is activated by the methionine regulon activator MetR. A comparative proteomic approach revealed MetR as a Lon substrate [9]. Furthermore, Lon degrades CysB, a positive regulator of the cysteine biosynthesis pathway [9]. Besides its role in various biosynthetic pathways, Lon is involved in metal homeostasis of zinc and copper. Substrates of ATP-dependent proteolysis are the MerR-like regulator ZntR (zinc homeostasis) and CueR (copper homeostasis). Both are degraded by Lon/ClpXP and Lon/ClpXP/ClpAP, respectively [39, 283]. ZntR binds excess zinc and activates the transcription of the zinc exporter ZntA, resulting in increased efflux of zinc [50, 60, 268]. Proteolysis of ZntR is ligand-controlled because the binding of zinc and DNA stabilizes the protein. The second Lon substrate involved in metal homeostasis is CueR, the key regulator of the Cue system. It activates gene expression of copper tolerance genes (copA and cueO) [269, 337]. CopA represents an efflux pump for monovalent copper (Cu+) and CueO prevents the passage of copper into the cell by converting Cu+ into Cu2+ in the periplasm, which is unable to pass the cell membrane [125, 275, 290, 291]. CueR proteolysis is copper-independent and for protease recognition, the C-terminus needs to be accessible [39]. Hence, ATP-dependent degradation of ZntR and CueR might be responsible for maintaining optimal cellular concentrations of zinc and copper, respectively [39, 283].

A Introduction

2.3.5 Lon in protein quality control In addition to regulated proteolysis, Lon is involved in the quality control of misfolded proteins [228, 300]. Lon has been shown to degrade >50% of abnormal proteins whose translation was terminated prematurely or those having base analogs [116, 229]. The role of Lon and other AAA+ proteases in quality control is important for preventing toxic protein accumulation in the cell. The ribosome rescue system mediated by trans-translation involves the Lon protease in the degradation of incorrectly elongated and terminated translating proteins stalled at the ribosome. The small protein B (SmpB) and the transfer messenger RNA (tmRNA) are required for association with the stalled ribosome [287, 358]. The bifunctional tmRNA (encoded by ssrA) can charge alanine to the A-site of the ribosome like a transfer RNA (tRNA) and codes for a small peptide (the so-called SsrA-tag) like a messenger RNA (mRNA) [189, 358, 361]. The SsrA-tag is fused C-terminally to the translating incomplete protein. Subsequently, SsrA-tagged proteins become suitable for proteolysis by ClpXP, ClpAP, FstH and Lon [66, 124, 146]. ClpXP is the major protease for proteolysis of SsrA-tagged proteins because it degrades >90 % of these proteins whereas Lon (only 2 %) plays a minor role [212]. This system prevents the cell against accumulation of stalled ribosomes. Lon protease is also important for quality control of pre-secretory proteins to protect the cell against toxic accumulation of pre-secretory proteins [308]. A secB lon double mutant accumulates aggregated proteins targeted for export. For example, Lon degrades the precursor molecules proOmpF and proOmpC in the absence of the chaperone SecB. Usually, SecB binds proOmpF and proOmpC to keep it in a suitable state for translocation to the inner membrane. In this way, SecB protects its substrate against Lon-dependent proteolysis. Another Lon substrate for quality control is the molybdenum enzyme TorA (trimethylamine N- oxide reductase 1) [288]. Lon was shown to degrade apo-TorA in vivo and in vitro. In contrast, TorA is protected against Lon proteolysis by binding to its chaperone TorD. Sakr et al. hypothized that a similar mode of action applies to mislocalized proteins of the co-translational protein transport, which is mediated by the signal recognition particle (SRP) complex [308]. The bacterial growth in SRP-depleted cells is facilitated if the proteases Lon and HslUV are present in the cell [30]. Interestingly, both AAA+ proteases become essential in SRP-depleted cells. Further studies identified Ffh, the protein component of the SRP complex, as Lon substrate (Arends, unpub.; [23]). All these findings provide evidence that the Lon protease is involved in the control of protein targeting pathways. Since this work focusses on the

A Introduction proteolysis of Ffh by the Lon protease, the following section presents the different protein targeting pathways, and in particular, the co-translational protein transport.

3. Principles of protein targeting in bacteria In bacteria, several pathways exist to ensure protein targeting and correct localization of newly synthesized membrane proteins or secretory proteins within the cell to sustain cellular homeostasis and membrane integrity. Two main pathways for post-translational and co- translational protein targeting have evolved. In general, both pathways can be divided into three steps: (I) recognition of the target protein in the cytoplasm, (II) targeting to the membrane and (III) integration into or transfer across the inner membrane. The main difference between the two pathways is the synthesis state of the target protein. For post-transitional protein targeting (Tat or Sec pathway; Fig. A-4), the protein is completely synthesized and released from the ribosome. On the other hand, co-translational protein targeting (SRP pathway; Fig. A-5) takes place while the synthesis of the nascent protein at the ribosome proceeds. In the following section, bacterial protein targeting pathways are described.

3.1 Post-translational protein targeting (Tat and Sec pathway) The Tat (twin-arginine translocation) pathway in E. coli consists of three integral membrane proteins (one of each TatA, TatB and TatC), which secrete fully folded proteins across the inner membrane [43, 314, 315, 383]. Signal sequences for Tat substrates comprise 26-58 amino acids and include a conserved sequence motif within the N-terminus of the secreted protein ((S/T)-R- R-x-F-L-K), which has two invariant arginine residues and two hydrophobic amino acid residues following a variable residue [28, 29]. More recent studies show that a simpler twin arginine motif Z-R-R-x-Φ-Φ (Z=any polar residue; Φ=hydrophobic residue) is valid [250]. Tat substrates often require redox cofactors for their function. They obtain the cofactors in the cytoplasm where they reach a folded conformation [311]. This is shown for the periplasmic TorA protein, which has molybdenum as cofactor. Deficiency of intracellular molybdate results in the accumulation of inactive TorA in the cytoplasm and recovery of the molybdenum concentration allows activation, oligomerization and translocation of TorA. For Tat substrate translocation, TatB/TatC provide a Tat substrate binding site and TatA forms a channel in the membrane (for review [245]). It is assumed that Tat substrates are transported across the membrane driven by the transmembrane proton motive force but this mechanism is not well understood.

A Introduction

Fig. A-4: Post-translational protein targeting in bacteria. Proteins completely synthesized and released from the ribosomes are targeted post-translationally across the inner membrane of bacteria. Newly synthesized proteins are recognized in the cytosol by pathway-specific signal sequences (SS) and afterward translocated either in a folded state via the Tat (twin-arginine translocation) pathway or in an unfolded state via the Sec pathway. Both pathways include chaperone binding in the cytosol to maintain the target proteins in a translocation-competent state. The membrane-integral Tat translocon requires three proteins (TatABC) and proton motive force (PMF) for protein secretion. For the Sec pathway, the ATPase SecA binds the unfolded protein in the cytosol or in complex with the SecYEG translocon at the membrane. Sec-dependent translocation is mediated by ATP-driven conformational changes by SecA supporting translocation via the SecYEG/SecDF. After substrate translocation, the signal sequence is cleaved off by a signal peptidase (SPase) and the target protein is released into the periplasm.

The second post-translational protein targeting pathway for secretory or outer membrane protein is Sec-dependent. The main difference to the Tat pathway is the folding state of transported protein because target proteins are completely synthesized but unfolded. Signal sequences contain approximately 20 to 30 amino acids, with a N-terminal positively charged n- region, a central hydrophobic h-region and a C-terminal polar c-region [143]. After release of the unfolded Sec substrate from the ribosome, it is able to interact with the chaperone SecB,

A Introduction other general chaperons (DnaK, GroEL) or trigger factor to maintain them in a translocation- competent state ([82, 97, 154, 345, 359], for review [250]). An additional route of Sec substrate targeting to the translocation system at the membrane involves cytoplasmic SecA. It binds at the ribosome co-translationally to the signal peptide sequence of a substrate, later it binds to the chaperone SecB, which targets the whole complex to the SecYEG translocation system [176]. In both cases, SecB is released before translocation across the inner membrane. Finally, the unfolded substrate is translocated across the inner membrane via ATP-driven conformational changes by SecA, which supports the translocation via the SecYEG. It is proposed, that SecDF has chaperone activity, which is powered by the proton motive force for promoting the final translocation step across the inner membrane [356].

3.2 Co-translational protein targeting (SRP pathway) Co-translational protein targeting of inner membrane proteins is facilitated by the signal recognition particle (SRP) complex and its membrane-anchored receptor FtsY (Fig. A-5). The SRP complex is stoichiometrically underrepresented in comparison to translating ribosomes. In E. coli, a ration of 110 Ffh molecules per 10,000 ribosomes is assumed [168]. Therefore, it is crucial that free SRP complexes rapidly recognize nascent proteins at ribosomes. The SRP complex consists of the highly conserved protein Ffh and the 4.5S RNA. Ffh contains two functional domains, i. e. the M and NG domain. N-terminal hydrophobic signal-anchored sequences (SAS) of nascent proteins emerging from the ribosome (ribosome nascent chain complex = RNC complex) initiate the first step in co-translational protein targeting. SAS differ from signal sequences of secretory proteins in their lack of a defined sequence motif. SAS are sequences of hydrophobic and aromatic amino acids, which are flanked by basic (N-terminal) and polar (C-terminal) amino acids [144]. The main differences between SAS and signal sequences are the absence of helix-breaking amino acids, the absence of a cleavage site for signal peptidases, and often it represents the first transmembrane domain for insertion of the membrane protein [2, 203, 276, 404]. Close contact of the SRP complex to the ribosome is mediated by the interaction of the NG domain and the M domain. The N-terminal GTPase- harboring NG domain interacts via binding the ribosomal proteins uL23 and uL29, which was shown by cryo-electron microscopy and single-particle analysis [137, 233, 413]. A more closer contact of the SRP complex to the ribosome is proposed by the interaction of the M domain to ribosomal proteins L22 and L24 and several rRNA helices [137]. The methionine-rich M domain binds the 4.5S RNA via a helix-turn-helix motif and forms a hydrophobic groove, which is needed for SAS binding [180, 413]. It is assumed that SAS’s are recognized by the 16

A Introduction

SRP complex, when they are exposed at the ribosome [259, 260]. However, recent studies hypothesize that the C-terminal helix of Ffh can reach into the ribosomal tunnel, which raises the possibility for substrate recognition in an early state of translation because the nascent protein is hidden inside the ribosome exit tunnel [81, 169]. Based on the conflicting data it is assumed, that interaction of the nascent protein inside the ribosome exit tunnel takes place for an early state of protein translocation and the nascent protein gets stabilized after the release of the ribosome [190]. After ribosome and substrate binding, the SRP complex interacts with the membrane-anchored SRP receptor (SR) FtsY. Via interaction with FtsY, the SRP-RNC complex is translocated to the inner membrane. FtsY has structural similarities to Ffh because it contains a Ffh homologous NG domain at the C-terminus [238]. Both NG domains represent the interaction site between Ffh and FtsY. Additionally, FtsY has an A domain at the N- terminus [381]. The A domain forms two helices, helix 1 (1-14 aa) with positively charged residues and helix 2 (188-207 aa) and both constitute the membrane anchor [46, 272, 334]. The RNC-SRP complex is recruited to the membrane by a GTP-dependent heterodimerization between Ffh and FtsY [93, 102]. The order of events is still being controversially discussed. One model suggests that the SRP-RNC complex is associated with FtsY in the cytosol and then targeted to the membrane [312].

Fig. A-5: Co-translational protein targeting in bacteria. The signal recognition particle (SRP) complex consisting of Ffh and the 4.5S RNA recognizes nascent polypeptides of the ribosome-nascent chain (RNC) complex by hydrophobic signal-anchored sequences (SAS) in the cytosol. The SRP-RNC complex is targeted to the membrane, where it interacts with the membrane-associated SRP receptor FtsY. GTP hydrolysis initiates dissociation of SRP complex and FtsY. SAS is transferred to the SecYEG/YidC translocation system and the co-translationally targeted protein is integrated into the membrane.

A Introduction

Cell fractionation studies found FtsY in both, the membrane as well as in cytosol fractions [218]. This hypothesis can be supported by structural analysis, as an interaction of FtsY with the SRP-RNA complex in solution has been shown [11, 96, 162, 216, 217]. Contrary data suggest that the interaction takes place at the membrane [235]. High affinity binding of the SRP-RNC complex to its receptor requires the binding of FtsY to lipids and the SecYEG translocation system [47, 199, 334]. Formation of a quaternary SecYEG-FtsY-SRP-RNC complex is the next step in the targeting process [170, 193, 312]. Conformational changes of the Ffh and FtsY NG domain stabilize the SRP-RNC-SR complex [93, 328]. FtsY is partially removed from SecY by its own movements and thus exposes the ribosome-binding site within SecY. On the other hand, the movement of the SRP complex leads to the exposure of the SecY binding site at the RNC [138, 279]. These structural changes promote a stable contact of the RNC to SecYEG. The RNC complex (by ribosomal proteins L23 and 29) binds to the cytosolic loops c4 and c5 of SecYEG [64, 230]. SRP and SecYEG bind related regions of the RNC complex, which supports the hypothesis of prior SRP complex dissociation followed by RNC translocation the SecYEG [25, 27, 105, 236]. The SAS is translocated to the translocation system by binding to the lateral gate of SecY, which is formed by two transmembrane helices [86, 363]. Transmembranes of SecE subunit stabilize the SecY proteins [222, 318] and the function of SecG is more relevant for SecA-dependent protein translocation of secretory proteins [90, 187]. Simultaneous GTP hydrolysis results in the dissociation of the SRP-SR complex [5, 17, 197, 370]. The SRP complex dissociates into the cytosol and FtsY remains bound in the membrane closely to SecYEG or lipids [193, 231]. Finally, the nascent protein is inserted into the inner membrane. Insertion of some SRP-delivered nascent proteins is also possible via the integral membrane insertase YidC as it was shown for the MscL protein or the

F0C protein [99, 362].

4. The universally conserved signal recognition particle 4.1 SRP complex in different species The SRP complex is highly conserved in all three domains of life (reviewed in [278]). It is a ribonucleoprotein, consisting of one or more protein components and an RNA component (SRP RNA) and its composition has changed during evolution. SRP complexes of different species are shown in Fig. A-6. The mammalian SRP complex, which is the best studied eukaryotic SRP complex, contains six proteins (SRP9, SRP14, SRP19, SRP54, SRP68, SRP72; named according to their molecular mass) and the 7S RNA (approx. 300 nt) [360, 374, 382]. The mammalian SRP RNA can be divided into the S- and the Alu-domain [131, 360]. The S-domain 18

A Introduction is composed of the central SRP RNA region (helix 8) and four SRP proteins (SRP19, SRP54, SRP68, SRP72) and is responsible for signal peptide binding and receptor interaction [382]. The Alu-domain is formed by the 5’ and 3’ terminal SRP RNA regions (helices 1 to 5) and interacts with the SRP9/SRP14 heterodimer. Since the mammalian Alu-domain has a tRNA- like structure, it is suggested that it plays a direct role in the translation arrest by blocking incoming tRNA molecules by occupying the ribosomal A-site with the SRP RNA [329, 375]. The mammalian signal recognition particle receptor (SR) consists of two GTPases, the SRα and SRβ [115, 232, 341]. SRα is a peripheral membrane protein and belongs to the class of SRP GTPases. In contrast the SRβ is an integral membrane protein being a member of the Arf/Sar family of small G proteins [234]. The archaeal SRP complex has similarities with the eukaryotic one because it contains the proteins SRP54 and SRP19 as well as the 7S RNA (reviewed in [415]). Likewise, it can divided into the S- and Alu-domain. The Alu-domain has an additional helix, which is involved in stabilization of the SRP RNA. FtsY represents the SR, which is homologous to the eukaryotic SRα.

Fig. A-6: Schematic representation of the SRP complex in different species. Evolutionarily conserved components of the SRP complex are the protein component SRP54/Ffh and the SRP RNA. Complexity of the SRP complex increases from prokaryotes to eukaryotes. In chloroplast, a new type of SRP complex has developed during evolution. In A. thaliana the cpSRP complex contains the additional chloroplast-specific protein cpSRP43. The cpSRP complex in P. patens represents an intermediate form between prokaryotes and higher plants, because it contains cpSRP43 and the SRP- RNA. Schematic representation of Ffh and FtsY is revised from [128].

The bacterial SRP complex exhibits the minimal machinery containing Ffh (protein component) and 4.5S RNA (RNA component) for co-translational protein targeting [31, 280, 292, 298]. Ffh (fifty-four homolog) is homologous to the eukaryotic SRP54, which is highly conserved and is present in all SRP complexes. In bacteria, SRP RNAs vary according to the species, for example 19

A Introduction the 4.5S RNA in Gram-negative bacteria (e.g. E. coli) and the 6S RNA in Gram-positive bacteria (e.g. B. subtilis) [338]. The 4.5S RNA forms a hairpin structure similar to the S-domain of the eukaryotic 7S RNA. In contrast, the 6S RNA comprises the S- and Alu-domain similar to the mammalian SRP RNA. In B. subtilis an additional protein, the histone-like protein HBsu, is able to bind the Alu-domain of the 6S RNA and forms a stable complex with the 6S RNA (helices 1-4 and partly helix 5) and Ffh [249, 280]. Interestingly, the evolution of chloroplast SRP complexes resulted in several variants. The chloroplast SRP complex (cpSRP complex) in higher plants (e.g. Arabidopsis thaliana (A. thaliana)) consists of the conserved cpSRP54 and another unique protein, cpSRP43 [104, 186, 323]. However, the universally conserved SRP RNA got lost in higher plants and thereby the ability to bind a SRP RNA [293, 301]. Further details on the structure and functions of the cpSRP complex in higher plants are described below (see A-4.3). Interestingly, there are transitional variants of the cpSRP complex in lower plants. For instance, the moss Physcomitrella patens contains the universally conserved cpSRP54 and SRP RNA as well as the chloroplast-specific cpSRP43 [353]. In vitro binding studies revealed binding of cpSRP54 and the cpSRP RNA (and 4.5S RNA from E. coli) in P. patens [353]. But binding affinity of cpSRP54 and cpSRP43 is lower than SRP RNA-binding (cpSRP54/cpSRP RNA, Kd: 1.3 μM; cpSRP54/cpSRP43, Kd: 9 μM) [412]. So it is proposed, that cpSRP54 exists in two forms: one bound to SRP-RNA and another one bound to cpSRP43. The SRP RNA in P. patens contains an elongated apical loop instead of the classic tetraloop formed in E. coli [302, 353]. In conclusion, the SRP complex in particular SRP54/Ffh and the SRP RNA is evolutionary conserved and is a fundamental key player for protein translocation in eukaryotic and prokaryotic cells. This is supported by the fact that mammalian SRP54 binds the 4.5S RNA from E. coli [280]. The well-studied cytosolic SRP complex from E. coli represents the minimal core and is described in more detail in the following section.

4.2 Structure of the minimal SRP complex and its receptor Ffh, the protein component of the SRP complex, and its receptor FtsY share structural and functional similarities (Fig. A-7). Both have in common their NG domain, which mediates the GTPase activity of both proteins. Each protein has an additional domain. Ffh exhibits the C- terminal M domain and FtsY the N-terminal A domain. In the following section, first, the individual domains are described and afterwards the NG domains of both proteins are compared. The methionine-rich M domain of Ffh recognizes hydrophobic SASs of nascent proteins at the ribosome with a picomolar affinity and binds the 4.5S RNA [180, 221, 299, 413]. 20

A Introduction

Fig. A-7: Structural overview of Ffh and its receptor FtsY. Both proteins can be divided in three domains while the N and G domain are structurally similar in Ffh and FtsY. They contain four conserved GTPase binding motifs (I, II, III, IV) and the insertion-box domain (IBD) within the G domain. Each protein has an additional domain. Ffh contains the C-terminal M domain, which is responsible for 4.5S RNA-binding by the helix-turn-helix motif (HTH) and signal-anchored sequence binding. FtsY contains the additional N-terminal A domain for membrane localization by the membrane targeting sequence (MTS). Schematic representation of Ffh and FtsY is revised from [313].

The binding site of Ffh and 4.5S RNA was well studied in the year 2000 by Batey et al. [20]. The structure is composed of five helices (helix 1, 2a, 2b, 3, 4) within the M domain and a disordered region of 33 amino acids between helix 1 and 2a. It is speculated that this region (so-called finger loop) is involved in conformational changes during signal sequence binding due to its flexibility [179]. The signal peptide binding is expected in a hydrophobic curve formed by α helix 1, 2 and 4 [180]. Four helices (helix 2a, 2b, 3, 4) form the helix-turn-helix (HTH) motif for SRP RNA-binding, which is distinct from classical HTH-motifs for DNA binding. SRP RNA-binding involves two α-helices (helix 3 and 4), which bind to the minor groove of the 4.5S RNA, instead of a single helix typical for DNA binding proteins [20, 180, 390]. Of particular importance for SRP RNA-binding is a positively charged sequence motif consisting of arginine, serine and glycine residues [6]. The 4.5S RNA (114 nt) folds into a single hairpin structure and contains an apical tetraloop and five internal loops, which are connected by helix forming regions [201, 202]. The tetraloop (-GGAA-) of the 4.5S RNA plays an important role in binding of the SRP complex to its receptor [161]. Gel retardation and membrane targeting experiments revealed an abolished SRP-FtsY interaction, if the tetraloop is mutated into -UUCG-. Other mutations (-UUUU- and –UUCG-) within the tetraloop are able to form a complex with Ffh and FtsY, but GTP hydrolysis was impaired. Further studies confirmed that SRP-SR complex formation and GTP activity on the heterodimer are enhanced by the 4.5S RNA [273, 274]. Ffh binding on the 4.5S RNA is full-filled by a minimal core region of approx. 50 nt, which is necessary and sufficient for binding [20, 207, 319–321]. This region includes an internal symmetric (exhibits non-canonical nucleotides) and asymmetric (exhibits unpaired nucleotides) loop within domain IV of the 4.5S RNA. Domain IV undergoes profound conformational changes, shown by the comparison of the unbounded and bounded state the M domain [20]. 21

A Introduction

The negatively charged A domain of FtsY is responsible for membrane localization [46, 204, 381, 402]. This intrinsically disordered domain binds anionic phospholipids [198, 334]. Binding to the membrane occurs via two sequences flanking the A domain [334]. N-terminal amino acids 2-14 and C-terminal amino acids 188-207 are extended to the first α-helix of the N domain. The second sequence is the so-called membrane targeting sequence (MTS) [272]. The NG domain mediates GTPase activity and interacts in a GTP-controlled manner with the homologous NG domain of the SRP receptor FtsY [93, 102, 169, 197, 327, 328, 405]. They interact in their GTP-bound state with high affinity. In contrast, in their GDP-bound state or non-nucleotide bound state, they bind with low affinity to each other. Both, Ffh as well as FtsY, belong to the GTPase superfamily SIMIBI (signal recognition particle, MinD and BioD) and share structural similarities [206]. The N domain consists of four α-helices with an open end that allows association of the G domain [179]. A small linker of 10 amino acids, which is closely associated with the protein surface, connects both domains and protects the NG domain against proteolysis [221, 414]. Crosslinking experiments have shown that the N domain of Ffh is associated with the ribosomal protein L23 at the exit of the ribosomal tunnel via amino acids 17 and 25 [129]. The G domain of Ffh is folded by five β-sheets and a single α-helix and represents a classical GTPase. Keenan et al. described four GTP binding motifs for Ffh and FtsY, which form a symmetric site for binding two GTPs [179]. A special characteristic of GTPases of the SIMIBI family is the unique insertion-box domain (IBD) within the G domain. It is speculated that this region has a particular role in the SRP-SR interaction [179]. It is presumed that residues of motif II are kept away from the active site of the complex by the IBD allowing SRP-SR formation. Thus it is proposed that the conformational changes in motif II of Ffh and FtsY result in reciprocal stimulation of GTP hydrolysis and finally dissociation of both [162].

4.3 Structur and function of the chloroplast SRP complex in higher plants As briefly described in a previous section (see section A-4.1), the composition of the chloroplast SRP complex has changed during evolution consisting of the highly conserved cpSRP54 and the unique protein cpSRP43 [104, 186, 323]. Hence, the cpSRP complex consists of two proteins instead of a protein component (one or more proteins) and an RNA component (SRP RNA) as in all other known SRP complexes. In higher plants, the SRP RNA is lost and cpSRP54 has no SRP RNA-binding ability due to substitution of two amino acids within the RNA- binding motif [293, 301]. The cpSRP complex has been described to function in both ways, post- and co-translationally (reviewed in [411, 412]) (Fig. A-8). 22

A Introduction

Fig. A-8: Post- and co-translational protein targeting in chloroplasts of higher plants. In higher plants, the cpSRP54 mediates post- or co-translational protein targeting. For post-translational protein targeting of light-harvesting chlorophyll a/b-binding protein (LHCP) a transit complex is formed, which consists of cpSRP54, cpSRP43 and LHCP. Afterwards the transit complex is recruited to the membrane- anchored cpFtsY and forms the docking complex. Finally, LHCPs are integrated into the thylakoid membrane by a complex of the insertase Alb3/cpSecYE/cpFtsY. In addition, plastid-encoded D1 protein is targeted co-translationally to the thylakoid membrane. The cpSRP54 interacts with the ribosome- nascent chain (RNC) complex. Insertion occurs via cpFtsY/cpSecYE/Alb3 and is stimulated by Vipp1. Revised from [410].

The post-translational protein targeting of light-harvesting chlorophyll a/b-binding proteins (LHCPs) in chloroplasts of higher plants mediated by the cpSRP complex is well studied. LHCPs are integral thylakoid membrane proteins for harvesting and transfer of energy to photosystem I and II. LHCP are nucleus-encoded and must be imported into the chloroplast via the translocon on the outer and inner chloroplast membrane (TOC/TIC) complex [44, 166, 270]. The hydrophobic LHCP forms a transit complex with cpSRP54 and cpSRP43 in order to remain 23

A Introduction soluble in the stroma and in a translocation-competent state [117, 323, 400]. The conserved cpSRP54 protein shows structural similarities to other SRP complexes as described above. As usual, it consists of the N-terminal N domain, the central G domain and the C-terminal M domain [104]. The M domain binds cpSRP43 and more precisely, a 10 amino acid residue (RRKRp10) is involved in this interaction [89, 107, 127, 172]. The unique cpSRP43 protein contains three chromodomains (CD1, CD2, CD3) and four ankyrin repeats (Ank1-Ank4) between CD1 and CD2 [117, 186, 332]. The cpSRP54 binding site of cpSRP43 is located in CD2 by two aromatic cages [153]. Moreover, cpSRP43 is crucial for LHCPs binding because it binds the L18 segment (VDPLYPGGSFDPLGLADD) of LHCPs [79, 357]. An interaction of LHCPs to cpSRP54 was not observed. After formation, the transit complex is translocated to the thylakoid membrane. Insertion of the LHCP into the thylakoid membrane requires further components such as the cpSRP receptor cpFtsY, SecYE and Alb3, a homolog of the bacterial insertase YidC.The structure of cpFtsY is similar to other SR since crystal structures showed the four helix bundle of the N domain and five GTP binding motifs within the G domain [57, 333, 353]. It binds to the thylakoid membrane via an amphipathic helix of the N-terminus [226, 333]. In plants, both proteins (cpSRP54 and cpFtsY) interact also via their homologous NG domains as described for the bacterial Ffh-FtsY interaction. Additionally it is suggested that the M domain stabilizes this interaction [58, 165]. LHCP insertion is conducted mainly by the insertase Alb3 [239, 240]. Before insertion of LHCP into the thylakoid membrane, a docking complex is recruited. It is assumed that cpSRP43 and Alb3 directly interact with each other. Interaction of Alb3 and cpSRP43 was shown by several studies, which revealed that positively charged binding motifs of Alb3 are crucial for cpSRP43 binding [16, 88, 100, 209, 210]. Details of the mode of action for alternative LHCP targeting are still being discussed (reviewed in [411]). Alb3 is associated with the cpSecYE translocase, however there is no evidence for an impact of cpSecYE on protein targeting [239, 241]. Furthermore, it was found that the Alb3/cpSecYE complex forms a complex with cpFtsY and the membrane-associated Vipp1 [373]. Thus LHCP are inserted cpSecYE-independently into the thylakoid membrane. It is suspected that the transit complex is delivered to a predefined complex consisting of cpFtsY/Alb3/cpSecYE [411]. In contrast to post-translational protein targeting, a second pool of cpSRP54 is associated with the 70S ribosome in chloroplasts and is assumed to act co-translationally. Experimental data demonstrate cross-linking of cpSRP54 to the nascent chain of the plastid-encoded D1 protein (encoded by psbA) [254, 255]. As part of the photosystem II, the D1 protein forms with the structural similar D2 protein and two chlorophyll molecules a reaction center for the

A Introduction photosynthetic electron transport chain [348]. After interaction of cpSRP54 with the RNC complex, the new complex (cpSRP54-RNC complex) is recruited to cpFtsY. An in vitro reconstitution system of D1 insertion provided a direct evidence for the interaction of D1 and cpFtsY [373]. D1 insertion into the thylakoid membrane is taken over by the cpSecYE/Alb3 translocon [373, 403]. The Vipp1 protein may stimulate the interaction. Whether there is a direct interaction of cpSRP54 and cpFtsY and a dissociation of both by GTP hydrolysis can only be speculated. Since, the mechanism of co-translational protein targeting by cpSRP54 is not well understood.

A Introduction

5. Objectives of this work ATP-dependent proteolysis is a strictly controlled process that allows bacterial cells adaptation to various stress conditions by maintaining protein homeostasis. In a previous master thesis it was demonstrated that Ffh, the protein component of the essential SRP complex, is a substrate for ATP-dependent proteolysis by the cytosolic AAA+ protease Lon. Thus, it was revealed that Lon protease is involved in the control of the co-translational protein targeting pathway. Ffh is degraded especially in the transition from the exponential to the stationary phase, which raises the possibility of a regulated mechanism. A putative recognition motif was suspected within the M domain of Ffh, since it was degraded, whereas the NG domain was stable. Based on these results, this work focused on the modulation of the growth phase-dependent degradation, and on the recognition mechanism of Ffh by the Lon protease. In this work the following questions should be addressed:

1. Modulation of the growth phase-dependent Ffh proteolysis Is Ffh degraded by further AAA+ proteases? Does the cellular Ffh concentration modulate proteolysis? Do other stress conditions, such as heat stress influence Ffh proteolysis? Is Ffh proteolysis dependent on the sigma factor RpoH?

2. Investigations on the recognition mechanism of Ffh by the Lon protease Does the RNA binding motif play a role in recognition of Ffh by Lon? Is the recognition of Ffh by Lon protease influenced by evolutionary point mutations?

3. Is the ATP-dependent proteolysis of Ffh a conserved mechanism? Is the Ffh homologue from chloroplasts of higher plants subject to ATP-dependent proteolysis in E. coli?

Materials and methods

B Materials and methods

B Materials and methods 1. Materials 1.1 Bacterial strains All bacterial strains used in this study are listed in Tab. B-1.

Tab. B-1: E. coli strains used in this study. Strain Relevant genotype References supE44,ΔlacU169 (Ψ80lacZΔM15), hsdR17, recA1, gyrA96, thi1, DH5α [309] relA1

BL21 [DE3] F-, ompT, gal (dcm) (lon), hsdSB (rB-mB-), λ[DE3] [388]

CH1019 X90ssrA:cat[DE3] ΔyefM-yoeB::kan R. T. Sauer

W3110 F‐, IN(rrnD–rrnE)1 [14]

ΔftsH W3110, zad220::Tn10 sfhC21ΔftsH3::kan [344]

F-, Δ(araD-araB)567, ΔlacZ4787(::rrnB-3), λ-, rph-1, Δ(rhaD- BW25113 [12] rhaB)568, hsdR514 (CGSC #7636)

ΔclpP BW25113, ΔclpP723::kan (CGSC #8590) [12]

K12 wild-type (WT) [14]

MC4100 K12, Δara, Δleu [55]

RH166 MC4100, Δara, Δleu, lac- [24]

ΔhslUV MC4100 ΔhslUV::kan [19]

Δlon RH166, Δlon:Tn10 [19]

MG1655 F-, λ-, rph-1 [14]

ΔlonΔclpPΔhslUV MG1655, Δ(clpPX-lon)1196::cat ΔhslVU1172::tet ΔsulA2981 [175] (KY2981)

SKP1101 MC4100, ara+, ffh-1::kan [pSKPP10] [271]

ΔrpoH MC4100, ΔrpoH::kan [408]

1.2 Plasmids All vectors (Tab. B-2) and plasmids (Tab. B-3) used in this study are listed below. To validate the sequence correctness of plasmids sequencing was carried out by Eurofins Genomics or Microsynth SEQLAB. 28

B Materials and methods

Tab. B-2: Vectors used in this study. Vector Relevant characteristics References

R pASK-IBA5(+) Amp , Ptet/Otet, tetR; encodes for N-terminal Strep-tag fusions IBA GmbH

KmR, lacZ, encoding full catalytic domain of AC from B. pertussis pKAC [78] (codons 1-384 of cyaA)

R pET-DuetTM-1 Amp , PT7/Olac, encoding N-terminal His6-tag fusion Novagen®

R pSS Amp , PSP6, PT7 (in vitro transcription plasmid) [104]

R pCA24N Cm , PT5/lac, lacI, gfp [185] +

R pET21b Amp , PT7 R. T. Sauer

Tab. B-3: Plasmids used in this study. Plasmid Vector Relevant characteristics References ffh-10(Ts) cloned into the RSF1030-derived cloning pSKPP10 pLCC29 [271] vector pLCC29; CmR pASK-IBA5(+) derivative, coding region for the pBO1199 pASK-IBA5(+) [200] Strep tag was replaced by his6-cspD pBO1199 derivative, cspD was replaced by ffh Arends, pBO3630 pASK-IBA5(+) (codons 2-453) unpubd. pBO3630 derivative, ffh was replaced by the M pBO3688 pASK-IBA5(+) [23] domain of ffh (codons 296-453) pBO3630 derivative, ffh was replaced by the NG pBO3689 pASK-IBA5(+) [23] domain of ffh (codons 2-295) Arends, pBO4805 pASK-IBA5(+) pBO3630 derivative, encoding FfhS382V unpubd. Arends, pBO4806 pASK-IBA5(+) pBO3630 derivative, encoding FfhC406S unpubd.

pBO4810 pASK-IBA5(+) pBO3630 derivative, encoding FfhG405D This study

pBO4813 pASK-IBA5(+) pBO3630 derivative, encoding FfhG405D C406S [101]

pBO4814 pASK-IBA5(+) pBO3630 derivative, encoding FfhS382V G405D C406S [101]

pBO4816 pASK-IBA5(+) pBO3630 derivative, encoding FfhS382V C406S [101]

cpSRP54 aus A. thaliana (codons: 81-564) with N- AtcpSRP54 pET-DuetTM-1 [293] terminal His6-tag cpSRP54V455S D480G from A. thaliana pBO4864 pSS [293] (codons: 81-564) pBO3630 derivative, Bsp1407I restriction enzyme pBO4817 pASK-IBA5(+) This study site changed to SalI pBO4817 derivative, ffh was replaced by cpSRP54 pBO4818 pASK-IBA5(+) This study from A. thaliana (codons 81-564)

B Materials and methods

Plasmid Vector Relevant characteristics References pBO4818 derivative, truncated cpSRP54, encodes pBO4862 pASK-IBA5(+) This study for the M domain of cpSRP54 from A. thaliana pBO4817 derivative, ffh was replaced by pBO4863 pASK-IBA5(+) cpSRP54V455S D480G from A. thaliana This study (codons 81-564) pKAC derivative, carried HIV protease cleavage pKACp5 pKAC site p5, between codons 224 and 225 of AC from B. [78] pertussis pEC5352 pKAC pKACp5 derivative, p5 replaced by rpoH [265]

Kutscher, pBO4807 pKAC pEC5352 derivative, p5 replaced by ffh unpubd. pEC5352 derivative, p5 replaced by NG domain of Kutscher, pBO4808 pKAC ffh (codons 296-453) unpubd. pEC5352 derivative, p5 replaced by M domain of Kutscher, pBO4809 pKAC ffh (codons 2-295) unpubd.

Lon-His6-pET21b pET21b lon with C-terminal His6-tag R. T. Sauer

- pLon-ASKA pCA24N pCA24N derivative, encoding His6-Lon, gfp [185]

- pFusA-ASKA pCA24N pCA24N derivative, encoding His6-FusA, gfp [185]

1.3 Oligonucleotides Oligonucleotides (Tab. B-4) used in this study were synthesized by Eurofins Genomics and dissolved in a. dest. at a concentration of 100 pmol. Underlined sequences corresponds to restriction sites of type II endonucleases.

Tab. B-4: Oligonucleotides used in this study. Primer Sequence (5’  3’) References Arends, Ffh.fw AAAATGTACATTTGATAATTTAACCGATCGTTTGTCG unpubd. Arends, Ffh.rv AAAAAAGCTTTTAGCGACCAGGGAAGCCT unpubd. Arends, Ffh_NG.rv AAAAAAGTCTTTAGAGAATACGCGACGCGA unpubd. Arends, Ffh_M.fw AAAATGTACAGGCATGGGCGACGTACTGT unpubd. Arends, FfhS382V.fw GGAAGCCATCATCAACGTCATGACGATGAA unpubd. Arends, FfhS382V.rv CGCGCTCTTTCATCGTCATGACGTTGATGA unpubd. Arends, FfhG405D.fw AAACGCCGTATTGCTGCAGATTGCGGTATG unpubd. Arends, FfhG405D.rv TGCACCTGCATACCGCAATCTGCAGCAATA unpubd.

B Materials and methods

Primer Sequence (5’  3’) References

Ffh_G405A.fw GTATTGCTGCAGCTTGCGGTATGCAGGTGC This study

Ffh_G405A.rv CCGCAAGCTGCAGCAATACGGCGTTTAC This study

Ffh_G405N.fw CGTATTGCTGCCAATTGCGGTATGCAGGTG This study

Ffh_G405N.rv CGCAATTGGCAGCAATACGGCGTTTACG This study

Arends, FfhC406S.fw CCGTATTGCTGCCGGATCCGGTATGCAGGT unpubd. Arends, FfhC406S.rv GTCCTGCACCTGCATACCGGATCCGGCAGC unpubd.

Ffh_triple.fw GCTGCAGATTCCGGTATGCAGGTGCAGGACG This study

Ffh_triple.rv GCATACCGGAATCTGCAGCAATACGGCGTTTACGC This study

pBO3630_SalI.fw CATCATCATGTCGACTTTGATAATTTAACCGATCGTTTGTCG This study

pBO3630_SalI.rv AAAAAGTCGACCAGTTGACTGGTGGCC This study

SRP-At-Hind.rv AAAAAAAGCTTTTAGTTACCAGAGCCGAAGC This study

SRP_At_fw2 AAAAAAGGATCCGCAGTTGACTGGTGGCC This study

SRP54_Mut_fw2 AAAAAGTCGACATGGCCATGGAGTTGACTGG This study

SRP54_Mut_rv AAAAAAAGCTTTTAGTTACCAGAGCCGAAGCC This study

Kutscher, ffH_NdeI_for2 AAAACATATGATGTTTGATAATTTAACCGATCGTTTGTCG unpubd. Kutscher, Ffh_XhoI_rev2 AAAACTCGAGGCGACCAGGGAAGC unpubd. Kutscher, ffH_NdeI_MD AAAACATATGGGCATGGGCGACGTACTGTCG unpubd. Kutscher, ffH_XhoI_NGD AAAACTCGAGGAGAATACGCGACGCGATG unpubd.

1.3 Laboratory equipment Laboratory equipment used in this study is listed in Tab. B-5. All other laboratory equipment that is not listed corresponded to the laboratory standard.

Tab. B-5: Laboratory equipment used in this study. Laboratory device Company analytical balance Mettler ToledoTM

autoclave Systec GmbH

B Materials and methods

Laboratory device Company chemiluminescence camera FluorChem SP MultiImage II Alpha Innotech

AlphaImager systems Gel documentation Alpha Innotech

incubator shaker New BrunswickTM Innova® 44 Eppendorf

pH/Ion Benchtop Meter S220 SevenCompact Mettler ToledoTM

VIS-Spectrophotometer Genesys 20 Thermo Scientific

shaking water bath THERMOLAB 1092 GFL

Elektrophoresis Power Supply PowerPac 1000 Biorad

Elektrophoresis Power Supply Power Pack P25 T Biometra GmbH

Thermocycler Mastercylcler nexus GX2 Eppendorf

water purification system Purelab Ultra Elga LabWater

centrufuge Sorvall RC-5C Plus Thermo Scientific

micro cooling centrifuge5415 R Eppendorf

micro centrifuge 5415 D Eppendorf

Heraeus Megafuge 1.0R Heraeus

French Press FA-078 SLM AMINCO

1.4 Chemicals Chemicals used in this study are listed in Tab. B-6.

Tab. B-6: Chemicals used in this study. Chemical Company acetic acid VWR International, LLC

acrylamide/bisacrylamide (37.5:1) Carl Roth GmbH + Co. KG

agar-agar Carl Roth GmbH + Co. KG

agarose Carl Roth GmbH + Co. KG

anhydrotetracycline (AHT) IBA Lifescience

ammonium persulfate (APS) Mallinckrodt Baker

ampicillin (Amp) Sigma-Aldrich

bromophenol blue AppliChem GmbH

bovine serum albumin (BSA) AppliChem GmbH

) calcium chloride (CaCl2 VWR International, LLC

chloramphenicol SERVA Electrophoresis GmbH

cOmpleteTM protease inhibitor coktail Roche

Coomassie Brilliant Blue G250 Merck KGaA

dimethylformamide (DMF) J.T. Baker 32

B Materials and methods

Chemical Company dimethylsulfoxide (DMSO) J.T. Baker dithiothreitol (DTT) Carl Roth GmbH + Co. KG ethylenediaminetetraacetic acid (EDTA) Carl Roth GmbH + Co. KG ethyl bromide (EtBr) Carl Roth GmbH + Co. KG ethanol (EtOH) VWR International, LLC formic acid J.T. Baker glycerol J.T. Baker glycine VWR International, LLC

) hydrogen peroxide (H2O2 J.T. Baker hydrochloric acid (HCl) Fisher Scientific UK Ltd 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid Carl Roth GmbH + Co. KG (HEPES) imidazole Acros Organics isopropyl β-D-1-thiogalactopyranoside (IPTG) AppliChem GmbH isopropanol VWR International, LLC potassium acetate (KAc) VWR International, LLC kanamycin (Km) Carl Roth GmbH + Co. KG potassium chloride (KCl) VWR International, LLC potassium hydroxide (KOH) J.T. Baker methanol (MeOH) VWR International, LLC magnesium acetate tetrahydrate (Mg(OAc)2 x 4 H2O) J.T. Baker magnesium chloride (MgCl2) J.T. Baker magnesium chloride hexahydrate (MgCl2 x 6 H2O) Carl Roth GmbH + Co. KG magnesium sulfate heptahydrate (MgSO4 x 7 H2O) VWR International, LLC sodium carbonate (Na2CO3) VWR International, LLC sodium acetate (NaAc) Sigma-Aldrich sodium chloride (NaCl) Carl Roth GmbH + Co. KG sodium hydrogen carbonate (NaHCO3) Fisher Scientific UK Ltd sodium hydroxide (NaOH) VWR International, LLC nonfat dried milk powder AppliChem GmbH phenylmethylsulfonyl fluoride (PMSF) SERVA Electrophoresis GmbH rifampicin SERVA Electrophoresis GmbH sodium dodecyl sulfate (SDS) AppliChem GmbH spectinomycin Sigma-Aldrich

B Materials and methods

Chemical Company sucrose Fisher Scientific UK Ltd

tetramethylethylenediamine (TEMED) Carl Roth GmbH + Co. KG

Tris Sigma-Alderich

tryptone Becton, Dickinson & Co

tween 20 AppliChem GmbH

Triton X-100 Sigma-Aldrich

yeast extract Becton, Dickinson & Co

β-mercaptoethanol AppliChem GmbH

1.5 Enzymes Commercially available enzymes used in this study are listed in Tab. B-7.

Tab. B-7: Enzymes used in this study. Name Use Company

Pfu-Polymerase PCR Lab stock

PRECISOR High-Fidelity DNA PCR, especially direct mutagenesis BioCat GmbH Polymerase Thermo Fisher T4 DNA ligase DNA ligation SCIENTIFIC Thermo Fisher restriction enzymes type II DNA restriction SCIENTIFIC Thermo Fisher FastDigest restriction enzymes type II DNA restriction SCIENTIFIC

Rnase A Plasmid preparation AppliChem GmbH

Dnase I cell disruption La Roche Ltd

lysozyme cell disruption Sigma-Aldrich

1.6 Kits Commercially available Kits used in this study are listed in Tab. B-8.

Tab. B-8: Kits used in this study. Name Use Company MACHERY-NAGEL NucleoSpin® Plasmid QuickPure plasmid preparation GmbH & Co. KG NucleoSpin® Gel and PCR Clean- DNA purification from agarose gels and MACHERY-NAGEL up PCR product clean up GmbH & Co. KG GE Healthcare Life PD-10 Desalting Columns buffer exchange for recombinant protein Sciences

B Materials and methods

immunological detection of recombinant Penta·His HRP Conjugate Kit QIAGEN His-tagged proteins

1.7 Antibodies Antibodies used in this study are listed in Tab. B-9.

Tab. B-9: Antibodies used in this study. Name Use Dilution Company/Contributor Penta·His HRP immunological detection of 1:4,000 QIAGEN Conjugate recombinant His-tagged proteins immunological detection of a gift from Hans-Georg α-Ffh 1:10,000 cellular Ffh – primary antibody Koch immunological detection of α-Ffh 1:10,000 a gift from Axel Mogk cellular Lon – primary antibody Goat Anti-Rabbit IgG immunological detection of 1:3,000 Bio-Rad Laboratories, Inc (H+L)-HRP Conjugate cellular Ffh – secondary antibody

1.8 Media and media supplements

Before use, culture media were autoclaved for 20 min, at 121 °C and 2 bar. For the preparation of solid medium 1.8 % (w/v) agar-agar was added to the media. Tab. B-10 lists all media supplements used in this study. Aqueous solutions of media supplements were sterile filtered before use.

LB medium [309] NaCl 1 % (w/v) tryptone 1 % (w/v) yeast extract 0.5 % (w/v)

2YT medium [309] NaCl 0.5 % (w/v) tryptone 1.6 % (w/v) yeast extract 0.5 % (w/v)

Tab. B-10: Media supplements used in this study. Media supplement Stock solution Working solution ampicillin (in a. dest.) 100 mg/ml 100 µg/ml

kanamycin (in a. dest.) 50 mg/ml 50 µg/ml

chloramphenicol (in 70% EtOH) 25 mg/ml 25 µg/ml

spectinomycin (in a. dest.) (Sp) 10 mg/ml 300 µg/ml

rifampicin (in 100% MeOH) 50 mg/ml 250 µg/ml

IPTG (in a. dest.) 100 mM varied

AHT (in DMF) 25 µg/ml varied

B Materials and methods

2. Software Software used in this study are listed in Tab. B-11.

Tab. B-11: Software used in this study. Software Use Company CloneManager 7.04 planning cloning Sci-Ed Software

L align alignment of DNA sequences SIB

AlphaEaseFCTM 4.0 densitometric evaluation of western blots Alpha Innotech

Clustal Omega alignment of protein sequences EMBL-EBI

Pymol protein structure simulation Schrödinger Visitron Systems VisiView miroscpoy GmbH Microsoft office 2013 data analysis Microsoft

3. Methods 3.1 Microbiological methods 3.1.1 E. coli cell culture E. coli cultures were grown at 37 °C, 30 °C or 25 °C either on solid LB agar plates or in liquid LB medium on a breeding roller or a shaker at 120 rpm. When required, antibiotics were added for selective cultivation (Tab. B-10.)

3.1.2 Determination of bacteria count The number of bacterial cells was determined spectrophotometrically at an optical density (OD) 8 at 580 nm. An OD580 of 1.0 corresponded to approx. 2 x 10 bacterial cells.

3.1.3 Preparation of competent E. coli cells For the preparation of competent E. coli cells, 100 ml LB medium was mixed with 2 ml Mg2+ solution and 2 ml of the corresponding overnight culture. The main culture was incubated at 37 °C and 120 rpm on a shaker until an OD580 of 0.5 to 0.8 was reached. Afterwards, cell suspension was divided in 2 x 50 ml and cells were harvested by centrifugation (10 min;

4,000 rpm; 4 °C). Cell pellets were resuspended in 25 ml ice-cold CaCl2 buffer, incubated for 60 min on ice and centrifuged again (10 min; 4,000 rpm; 4 °C). Subsequently, cell pellets were resuspended in 5 ml CaCl2 buffer and 1.5 ml 87 % glycerol. Competent cells were divided into 260 µl aliquots and stored at -80 °C until further use.

B Materials and methods

Mg2+ solution MgCl2 x 6 H2O 500 mM MgSO4 x 7 H2O 500 mM

CaCl2 buffer CaCl2 100 mM

3.1.4 Transformation of E. coli For the transformation of E. coli cells, an aliquot of the chemically competent cells (see B-3.1.3) and the plasmid DNA were thawed on ice. Plasmid DNA (100 ng) was added to the competent cells and incubated for 30 min on ice. Transformation was conducted by heat shock at 42 °C for 2 min. To regenerate the cells, 700 µl liquid LB medium was added to the transformation sample and incubated for 45 min at 37 °C on the breeding roller. Finally, 100 µl and the concentrated rest of the bacterial suspension were plated on solid LB agar plates and incubated overnight at 37 °C.

3.1.5 Transformation efficiency Transformation efficiency was determined in E. coli MC4100 and Δlon for plasmid-encoded Ffh, Ffh NG domain, Ffh M domain, cpSRP54 from A. thaliana, cpSRP54 M domain from

A. thaliana and the corresponding empty vectors for His6-tagged constructs and One hybrid constructs. For the transformation 100 ng plasmid DNA and 100 µl of the chemically competent cells were used per construct. Transformation was performed as described under section B-3.1.4 with the following deviations. For cell regeneration 900 µl liquid LB medium was added after heat shock and incubated for 60 min at 37 °C. Cell suspensions were diluted from 100 to 10-4 and 100 µl per dilution and plated on solid LB agar plates. After incubation at 37 °C for 24 h colonies were counted. To calculate transformation efficiency, it was necessary to determine the bacterial titer of the competent cells used in this approach. For this purpose, a transformation sample without plasmid DNA was used, which was treated as described above. Bacterial suspensions were diluted from 100 to 10-8 and 100 µl dilution steps, each aliquot was plated on solid agar and incubated as described above. To determine the bacterial titer, colony number of the control samples (MC4100 and Δlon without plasmid DNA) were calculated for 1 ml bacterial suspension as follows:

Bacterial titer = colonies dilution step plated volume o bacterial suspension

× × f

B Materials and methods

Transformation efficiency was determined using the bacterial titer, the number of transformed cells of 1 ml bacterial suspension and the amount of plasmid DNA as follows:

transformants [transformants/ml] plasmid carried bacteria bacteria titer [bacteria/ml] Transformation efficiency = ng DNA plasmid DNA [ng] 3.1.6 Microscopy studies

For light microscopy of E. coli cells, a main culture was inoculated to an OD580 of 0.05 and was grown at 37 °C until exponential and stationary growth phase, respectively. Before light microscopy, 800 µl 1,5 % agarose solution (dissolved in a. dest.) were applied on the slides.

Cell cultures were diluted to an OD580 of about 0.5 and 10 µl were applied to the prepared slide and covered by a coverslip. Microscope Olympus BX51 was used for microscopy studies with 100-fold magnification. A scale of 5 µm was used for size determination of cells.

3.2 Molecular biological methods 3.2.1 Polymerase chain reaction Polymerase chain reaction (PCR) was used to amplify DNA fragments of plasmid DNA and introduce amino acid substitutions by site-direct mutagenesis [48, 246]. For amplification of DNA fragments, synthetic oligonucleotides containing restriction enzyme sites for type II restriction endonucleases were used as primers. The DNA polymerase used was Pfu polymerase, which has 3'-5'-exonuclease activity and produces blunt ends on DNA fragments. Composition of a PCR approach and the PCR program used are listed below:

PCR approach Pfu polymerase 0.5 U 10x Pfu buffer 5 µl dNTPs (each) 0.2 mM DMSO 0-10 % (v/v) DNA template 100 ng oligonucleotide fw 5-50 pmol oligonucleotide rv 5-50 pmol a. dest. ad 50 µl

B Materials and methods

PCR program denaturation 95 °C 5 min

denaturation 95 °C 1 min annealing varied 1 min 5x elongation 72 °C 2 min

denaturation 95 °C 1 min annealing varied 1 min 25x elongation 72 °C 2 min

finale elongation 72 °C 10 min

10x Pfu buffer KCl 100 mM (NH4)2SO4 100 mM Tris/HCl (pH 8.8) 200 mM MgSO4 20 mM Triton X-100 1 % (v/v) BSA (nuclease free) 1 mg/ml

10x dNTPs dATP,dTTP,dGTP, dCTP each 2 mM

3.2.2 DNA restriction Hydrolytic cleavage of plasmid DNA or DNA fragments was performed with restriction endonucleases type II. The composition of a restriction mix is shown below. The restriction approach was incubated for 60 min at 37 °C or 30 °C depending on the optimal working temperature of the used restriction endonuclease type II. Successful test restriction was verified by agarose gel electrophoresis (see B-3.2.6). A restriction mix was inactivated for 20 min at 70 °C if DNA before ligation.

Restriction mix plasmid DNA 100 ng restriction endonuclease type II 1 U 10 x reaction buffer 1 µl a. dest. ad 10 µl

3.2.3 DNA ligation DNA fragments were ligated using T4 DNA ligase, which forms a covalent phosphodiester bond between the 5'-phosphate end and the 3'-hydroxy end of DNA. Vector/insert ratios were chosen ranging from 1:1to 1:5 for ligation. The composition of a ligation mixture is shown below. Ligation mixtures were incubated for 2 h at room temperature (RT) and afterwards inactivated at 80 °C. The complete ligation mix was transformed in E. coli DH5α (see B-3.1.4). 39

B Materials and methods

Ligation mix plasmid backbone 1 µl insert 1-5 µl T4 DNA ligase 1 µl 10 x T4 DNA ligase buffer 2 µl a. dest. ad 20 µl

3.2.4 Preparation of plasmid DNA from E. coli Plasmid DNA from E. coli was prepared by alkaline lysis [36]. Cells from 2 ml liquid overnight cell culture were pelleted by centrifugation (1 min; 13,200 rpm; RT). Cell pellets were resuspended in 300 µl mix I and incubated for 1 min at RT and afterwards 300 µl mix II was added for cell lysis. Samples were inverted 6 to 8 times and incubated for 5 min at RT. Samples were neutralized by adding 300 µl mix III, inverting 6 to 8 times and incubating samples for 10 min on ice. After centrifugation (15-60 min; 13,200 rpm; 4 °C) the DNA-rich supernatant was transferred to a new reaction tube and DNA was precipitated by adding 630 µl isopropanol. Precipitated DNA was pelleted by centrifugation (30-60 min; 13,200 rpm; 4 °C) and the supernatant was discarded. The DNA pellet was washed with 500 µl EtOH and centrifuged again (5 min; 13,200 rpm; 4 °C). After centrifugation, the supernatant was discarded and the DNA pellet was dried in a heating block at 50 °C. Finally, the dried DNA pellet was dissolved in 20-40 µl a. dest. and stored at -20 °C until further use.

Mix I Tris/HCl (pH 8.0) 50 mM EDTA 10 mM RNase A 0.2 mg/ml RNase A was directly added before use.

Mix II NaOH 200 mM SDS 1 % (v/v)

Mix III KAc 3 M formic acid 1.8 M

B Materials and methods

3.2.5 Plasmid preparation from E. coli by commercial kits For plasmid DNA preparation using a commercial kit, an overnight cell culture of putative positive clones was prepared as described above (see B-3.1.1). NucleoSpin® Plasmid QuickPure kit from MACHERY-NAGEL GmbH & Co. KG (see Tab. B-8) was used for plasmid DNA preparation and was performed according to the enclosed instructions.

3.2.6 Agarose gel electrophoresis DNA separation according to length was performed by agarose gel electrophoresis. DNA was separated in the gel matrix by an electrical field (300 mA, 100-150 V), whereby the migration speed of the DNA fragments depends on their length. For the production of the agarose gel, 1 % agarose were dissolved in 1xTAE buffer, which also was used as electrophoresis buffer. Before loading the agarose gel, the DNA samples were mixed with one-fifth 5x DNA sample buffer. The samples were placed in the gel pockets and the gel was allowed to run for about 1.5 h at 100-150 V. The 1 kb DNA standard (Invitrogen) was used to estimate the DNA fragment length. The DNA was stained with 0,075 % EtBr in the agarose gel and was visualized under UV radiation.

1x TAE buffer Tris/acetic acid pH 7.8 0.8 mM NaAc 0.2 mM EDTA 0.02 mM

5x DNA sample buffer bromphenolblue 0.5 % glycerol 43 % EDTA pH 8.0 100 mM

3.2.7 Extraction of DNA fragments from agarose gel For extraction of PCR products and DNA fragments from agarose gels, DNA was separated by agarose gel electrophoresis by size and DNA was visualized by EtBr staining. The corresponding DNA bands were cut out of the agarose gel under UV radiation. DNA was extracted from agarose gel using the NucleoSpin® Gel and PCR Clean-up kit from MACHERY-NAGEL GmbH & Co. KG. (see Tab. B-8) according to instructions.

B Materials and methods

3.2.8 Cloning strategies 3.2.8.1 Cloning by restriction and ligation Cloning of PCR products was performed by restriction of the PCR insert and the plasmid backbone and subsequent ligation. The PCR products and the plasmid backbone were restricted (see B-3.2.2) via the same singular restriction endonucleases type II sites and then the enzymes were inactivated according to the manufacturer's instructions. The restricted insert and backbone were then separated by agarose gel electrophoresis (see B-3.2.6) to remove restriction fragments. The corresponding fragments were eluted from the agarose gel using a kit (see B-3.2.7). Subsequently, both fragments were ligated using T4 DNA ligase (see B-3.2.3). The complete ligation approach was used for a transformation into E. coli DH5α (see 3.1.4). Appropriate antibiotics were used for the selective cultivation of putative transformants (see B-3.1.1). Positive clones were confirmed by sequencing.

3.2.8.2 Site-directed mutagenesis Site-directed mutagenesis was used to exchange DNA bases in order to generate an amino acid exchange in the protein sequence. PRECISOR High-fidelety DNA Polymerases was used. Complementary overlapping primers were designed containing the desired mutation. During the PCR cycle, the primers anneal to the template DNA and replicate the complete plasmid DNA. The mutated plasmid DNA has a DNA strand break and thus differs from the parental DNA strand. The methylated parental strand is separated from newly synthesized unmethylated mutant DNA by digestion with methylation-sensitive enzyme DpnI (2x 1,5 h; 1,5 µl DpnI each 50 µl PCR product). After DpnI inactivation (80 °C, 10 min), the mutated DNA was transformed into E. coli DH5α, where the strand break was ligated by the host repair system. Seletive cultivation of putative transformants (see B-3.1.1) required the use of suitable antibiotics. Positive clones were confirmed by sequencing.

3.2.9 DNA Sequencing DNA samples were sequenced by Eurofins Genomics or Microsynth SEQLAB. 15 µl of Plasmid-DNA at a concentration between 50 and 100 ng/µl were sequenced. If required, sequencing oligonucleotides diluted in 15 µl a. dest. to a concentration of 10 pmol/µl were supplied. Sequencing results were analyzed for their correctness with the available online tool L align by pairwise sequence alignment.

B Materials and methods

3.3 Protein biochemical methods 3.3.1 Expression and solubility studies To analyze expression of overproduced recombinant proteins, a main culture was inoculated from an overnight culture to an OD580 of 0.05 in 15 ml LB medium with appropriate antibiotics.

Cells were grown at 37 °C to an OD580 of 0.5-0.8. By adding 50 ng/ml AHT, protein overproduction was induced for 30 min at 37 °C. After 0, 15, and 30 min, 1 ml of the culture was removed and the cells were pelleted by centrifugation (10 min; 13,200 rpm, 4 °C) and prepared for SDS-PAGE. The remaining culture was used for solubility studies. Cells were harvested by centrifugation (10 min; 4,000 rpm; 4 °C), washed in 10 ml buffer A and centrifuged (10 min; 4,000 rpm; 4 °C) again. Cell pellets were stored at -20°C. For cell disruption, cell pellets were thawed on ice, resuspended in 3.5 ml lysis buffer and disrupted under pressure (2x 900 psi) via FrenchPress. Subsequently, 2 ml cell lysate were centrifuged (30 min; 13,200 rpm; 4 °C) to separate cell debris and denatured proteins. The supernatant fraction was transferred to a new reaction tube and was stored on ice until sample preparation. The pellet fraction of 2 ml cell lysate was resuspended in 2 ml TE buffer. The pellet and supernatant fractions were prepared for SDS- PAGE, Western transfer and immunodetection.

Buffer A Tris/HCl pH 7.5 20 mM NaCl 200 mM

Lysis buffer Tri/HCl pH 7.5 50 mM NaCl 300 mM imidazole 10 mM glycerol 20 % lysozyme 1 mg/ml DNase I Spade point RNase A Spade point PMSF 0.5 mM

TE buffer Tris/HCl pH 8.0 10 mM EDTA 1 mM

B Materials and methods

3.3.2 In vivo degradation experiments

Stability of recombinant His6-tagged proteins from AHT-inducible expression plasmids and cellular Ffh were measured using in vivo degradation experiments at different growth phases (Fig. C-1 A). For this purpose, an E. coli main culture was inoculated from an overnight culture at OD580 of 0.05 and grown in liquid LB medium at 37 °C (or 30 °C for ΔftsH and SKP1101 + pSKPP10, or 25 °C for ΔrpoH) in a water bath shaker until the corresponding growth phase.

Degradation cultures were taken and expression of plasmid-derived His6-tagged proteins was induced by adding 50 ng/ml AHT for 30 min. For cellular Ffh degradation, cultures were taken and translation was directly blocked or after induction of different stress conditions (e.g. heat stress, oxidative stress). More details about stress conditions used in this study are described in the results section (seeC-1.3). Translation was stopped by addition of 300 µg/ml spectinomycin, samples were taken at different time points, frozen in liquid nitrogen and prepared for SDS- PAGE, Western transfer and immunodetection as described below.

3.3.3 Purification of poly-histidine fusion proteins 3.3.3.1 Protein overproduction For the overproduction of poly-histidine fusion proteins, the corresponding expression plasmid was transformed in E. coli BL21. A main culture was inoculated from an overnight culture to an OD580 of 0.05 in 800 ml 2YT or LB medium. The main culture was incubated to an OD580 of 0.5 to 0.8 at 37°C under constant shaking at 120 rpm. Once the desired OD580 was reached, 1 ml of the culture was transferred to a 1.5 ml reaction tube and the cells were pelleted by centrifugation (10 min; 13,200 rpm; 4 °C). Subsequently, protein overproduction was induced either with 1 mM IPTG (T7-expression system) or 25 ng/ml AHT (Tet-expression system). The recombinant proteins were produced at 30 °C for 3 and 4 h, before 1 ml of the culture was removed and pelleted by centrifugation (10 min; 13,200 rpm; 4 °C). The remaining culture was divided into 400 ml each and pelleted by centrifugation (10 min; 4,000 rpm; 4 °C). Each cell pellet was resuspended in 50 ml 1x buffer A and pelleted again by further centrifugation (10 min; 4,000 rpm; 4 °C). Cell pellets were stored at -20°C until cell disruption. As the conditions for cultivation and protein overexpression for recombinant Lon-His6 and His6-Ffh varied, Tab. B-12 summarize all conditions for cultivation and protein overproduction.

Tab. B-12: Growth condition for protein overproduction. protein plasmid strain growth conditions inducer over production

His6-Ffh pBO3630 BL21 37 °C, LB 25 ng/ml AHT 3 h, 30 °C

Lon-His6 pET21b-Lon-His6 CH1019 37 °C, 2YT 1 mM IPTG 4 h, 30 °C

B Materials and methods

3.3.3.2 Cell disruption by FrenchPress A cell pellet (cells from 400 ml) of each protein to be purified was thawed on ice for cell disruption. Cell pellets were resuspended in 5 ml lysis buffer, which varied (see below). Cells were disrupted under pressure using French-Press. Cells were disrupted two times at 900 psi and the resulting cell lysate was centrifuged (30 min; 13,200 rpm; 4 °C) to remove cell debris and denatured proteins from the cell lysate. Finally, the supernatants were combined and stored on ice until nickel affinity chromatography.

Lysis buffer

His6-Ffh (revised from [26]) Lon-His6 [200] HEPES/KOH pH 7.5 25 mM Tris/HCl pH 7.5 20 mM KCl 500 mM NaCl 200 mM Mg(OAc)2 10 mM lysozyme 1 mg/ml imidazole 10 mM DNase I 10 µg/ml glycerol 10 % DTT 1 mM cOmpleteTM protease inhibitor coktail 4 mM lysozyme 1 mg/ml DNase I 10 µg/ml

3.3.3.3 Nickel affinity chromatography Nickel affinity chromatography was used for purification of native poly-histidine fusion proteins. His6-Ffh and Lon-His6 were purified to determine in vitro stability of His6-Ffh in the presence of Lon-His6. All purification steps were carried out at 4 °C. A column volume of 0.75 ml Ni-NTA Agarose (QIAGEN) was washed with 10 ml a. dest. and equilibrated with 10 ml binding buffer. The cell lysate was pre-incubated with the column matrix for 30-45 min under constant rotation. The saturated column matrix was placed into a column device and allowed to settle at the bottom. Afterwards, the column was opened and the flow through was completely collected. The column matrix was washed four times with 5 ml wash buffer each and the poly histidine fusion proteins were eluted five times with 1 ml elution buffer each. After purification, the column material was cleaned with 10 ml 1 M imidazole (in an appropriate cleaning buffer), 10 ml a. dest. and 10 ml 20 % EtOH. Purified His6-Ffh and Lon-His6 was stored overnight on ice at 4 °C, as longer storage was not possible due to precipitation of the proteins.

B Materials and methods

Binding buffer

His6-Ffh [26] Lon-His6 [200] HEPES/KOH pH 25 mM Tris/HCl pH 7.5 20 mM 7.5 KCl 500 mM NaCl 200 mM Mg(OAc)2 10 mM imidazole 5 mM imidazole 10 mM DTT 1 mM glycerol 10 %

Wash buffer

His6-Ffh [26] Lon-His6 [200] HEPES/KOH pH 25 mM Tris/HCl pH 7.5 20 mM 7.5 KCl 1000 mM NaCl 200 mM Mg(OAc)2 10 mM imidazole 50 mM imidazole 25 mM DTT 1 mM glycerol 10 %

Elution buffer

His6-Ffh [26] Lon-His6 [200] HEPES/KOH pH 25 mM Tris/HCl pH 7.5 20 mM 7.5 KCl 200 mM NaCl 200 mM Mg(OAc)2 10 mM imidazole 500 mM imidazole 150 mM DTT 1 mM glycerol 10 %

3.3.4 Protein concentration quantification by Bradford assay Protein concentration was determined using the Bradford assay [45]. A calibration curve was established with a standard series of 0, 1, 3, 5, and 10 µg BSA. Each calibration sample was filled up to 800 µl with a. dest. For the protein samples, 5 µl of the corresponding purification fraction were mixed with 795 µl a. dest.. Reaction was started by adding 200 µl 5x Bradford reagent (Roti®-Quant from CARL ROTH), incubated for exactly 10 min at RT and measured at OD595 afterwards. Based on the standard series, a standard curve was calculated to determine the protein concentration of the samples as follows:

µg OD595 protein concentration = 1 µl gradient of the standard curve µg × sample volume [µl]

B Materials and methods

3.3.5 In vitro degradation experiments For in vitro degradation experiments, 15 µg of the purified putative Lon substrate and 600 nM

Lon-His6 were pre-incubated with in vitro degradation buffer for 2 min at 37 °C. As a control for Lon activity, 15 µM of the known Lon substrate His6-CspD were used in an in vitro degradation assay [200]. The reaction was started by the addition of 20 mM ATP and/or a. dest. and samples were incubated for 120 min at 37 °C. At defined time points, 20 µl sample were taken and mixed with 5 µl 5x protein sample buffer (5x PPP), boiled at 95 °C for 5 min and stored at -20 °C until further use. Later, samples were separated by denaturing SDS-PAGE and subjected to Coomassie staining or Western transfer.

In vitro degradation buffer [37] Tris/HCl pH 8.0 50 mM DTT 1 mM MgCl2 15mM DMSO 2 % KCl 5 mM HEPES/KOH pH 8.0 25 mM β-mercaptoethanol 0.5 mM sucrose 10 % a. dest. ad 100/200 µl

3.3.6 Preparation of protein extracts For protein extract preparation cells from 1 ml culture were harvested by centrifugation and resuspended in TE buffer volume according to their OD580 (OD580 of 1.0 correspondes to 100 µl TE buffer). Cell pellets were resuspended with TE buffer and a fifth of 5x PPP. Purification fractions and in vitro degradation samples were mixed with a fifth 5x PPP according to their volume. All samples were boiled at 95 °C for 10-15 min and stored at -20 °C until further use for SDS-PAGE.

5x protein sample buffer (5x PPP) Tris/HCl pH 6.8 250 mM SDS 10 % bromophenol blue 0.5 % β-mercaptoethanol 5 % glycerol 50 %

B Materials and methods

3.3.7 Denaturing SDS polyacrylamide gel electrophoresis Protein samples were separated by denaturing SDS-PAGE [309]. Separation was carried out depending on the molecular mass via 12% (protein> 20 kDa) or 15% (protein <20 kDa) separating gels and 5% stacking gels at a voltage of 80-120 V in a electrophoresis chamber. 5 µl protein standard was loaded along for molecular mass determination. For Coomassie stained SDS polyacrylamide gels, BenchMarkTM Protein Ladder from ThermoFisher SCIENTIFIC was used as protein standard. For SDS polyacrylamide gels subjected to Western transfer the PageRulerTM Plus Prestained Protein Ladder from ThermoFisher SCIENTIFIC were used.

1x SDS running buffer Tris 24,76 mM glycine 156,37 mM SDS 3,47 mM

separating gel 12 % 15 % acrylamide/bisacrylamide 12 % (v/v) 15 % (v/v) (37.5:1) Tris/HCl pH 8.8 0.5 M 0.5 M SDS 0.1 % (w/v) 0.1 % (w/v) APS 0.1 % (v/v) 0.1 % (v/v) TEMED 0.04 % (v/v) 0.04 % (v/v)

stacking gel acrylamide/bisacrylamide 5 % (v/v) (37.5:1) Tris/HCl pH 8.8 0.5 M SDS 0.4 % (w/v) APS 0.1 % (v/v) TEMED 0.06 % (v/v)

3.3.8 Coomassie staining of polyacrylamide gels For Coomassie staining, SDS-polyacrylamide gels were incubated for 20 min at RT on the swivel table with staining solution. Subsequently, stepwise washing with destaining solution decolorized the SDS-polyacrylamide gels.

staining solution MeOH 50 % acetic acid 10 % Coomassie billant blue G250 0.15 %

B Materials and methods

destaining solution MeOH 25 % acetic acid 10 %

3.3.9 Western transfer and immunological detection of proteins For immunological detection of poly-histidine fusion proteins and cellular Ffh, protein extracts were transferred to a nitrocellulose membrane (Amersham Protran 0,45 µm nitrocellulose Westernblotting membrane from GE Healthcare Life Science) by Western transfer, which was performed by using ice cold 1x DC buffer at 500 mA for 60 min [87]. After Western transfer, the membrane was blocked with blocking reagent for 60 min. For specific detection of poly- histidine fusion proteins, the membrane was incubated with Penta-His HRP conjugate at a dilution of 1:4,000 overnight. For immunological detection of cellular Ffh/Lon, the membrane was incubated for 1 h with monoclonal α-Ffh (1:10,000)/α-Lon (1:10,000) antibody. After primary antibody treatment, the membrane was washed 4 times for 10 min at RT with 1x TBST and further incubated for 1 h with Goat Anti-Rabbit IgG (H+L)-HRP Conjugate as secondary antibody. Finally the membrane was washed 4 times for 10 min at RT with 1x TBST. The chemiluminescence-based detection was carried out with Immobilon forte™ Western HRP substrate (Merck KGaA) and signals were detected with a chemiluminescence camera.

1x DC buffer [87] NaHCO3 10 mM Na2CO3 3 mM MeOH 20 % (v/v)

blocking reagent non fat dried milk powder 5 % Dissolved in 1x TBST.

1x TBS buffer Tris/HCl (pH 7.5) 50 mM NaCl 150 mM

1x TBST buffer Tris/HCl (pH 7.5) 50 mM NaCl 150 mM Tween 20 0.1 %

B Materials and methods

3.3.10 Calculation of half-lives The relative protein amount was determined by pixel counting of the protein signals to calculate the half-lives of proteins indicating at which time point 50 % of the initial protein amount (at t=0) was still present after the start of degradation. The signal of the initial sample at time 0 (t=0) was considered as 100 %. For calculation, the background was subtracted from the value for each sample. Relative protein amount in % of the following samples was calculated relative to the initial value (t=0). The relative protein amounts (y-axis) were plotted against time (x- axis) and half-lives were calculated in accordance with a linear equation y = mx +b (x=time after degradation start; y= protein amount in % of value t=0). Half-lives were calculated as follows: 1 - 2 b T1 [min]= 2 m

Results

C Results

C Results Membrane protein transport is a crucial process to sustain cellular homoeostasis and membrane integrity. In prokaryotes, one mechanism for membrane protein trafficking is the cotranslational transport by the essential SRP complex, which consists of the protein component Ffh and the regulatory 4.5S RNA [219, 280, 281, 292]. A previous FtsH trapping approach revealed Ffh as substrate of proteolysis in exponential and stationary growth phase [10]. Interestingly, the FtsH protease was not responsible for Ffh degradation. Instead degradation was dependent on the Lon protease ([23]; Arends, unpubd.). While Ffh is rapidly degraded in late exponential and early stationary growth phase, it is a more stable protein in earlier exponential and later stationary growth phases. Growth phase-dependent proteolysis is common for AAA+ proteases. For example, the FtsH substrate YfgM and the Lon substrate CspD are differently degraded [40, 200]. These observations raise the possibility of a regulatory process on Ffh proteolysis and motivated us to further investigate the degradation and recognition mechanism of the essential Ffh protein.

1. Investigations on the degradation mechanism of Ffh 1.1 Growth phase-dependent proteolysis of plasmid-derived Ffh To monitor protein stability during different growth phases, we performed in vivo degradation experiments over the entire bacterial growth with different Ffh variants (Fig. C-1 A). Protein L338P M347T stability of cellular Ffh, His6-Ffh and Ffh was studied in E. coli MC4100, which repressents WT strain of several mutant strains (e.g., Δlon, SKP1101) that were used as reference for different experiments in this work. For in vivo degradation experiments, an appropriate E. coli main culture was grown at 37°C or 30°C and degradation cultures were taken at defined growth phases. Fig. C-1 A shows an example for the bacterial growth of E. coli

MC4100 at 37°C, which was determined photometrically at OD580 from three independent experiments. Degradation cultures were taken in early (I), mid (II) and late (III) exponential growth phase as well as in early (IV) and late (V) stationary growth phases. To analyze the protein stability of His6-Ffh, gene expression from a AHT-inducible high copy plasmid was induced by adding 50 ng/µl AHT to the culture for 30 min. For cellular Ffh and the plasmid- derived FfhL338P M347T variant, an inducible protein overproduction was not required. Translation was stopped by the addition of 300 µg/ml Sp, samples were taken and prepared for SDS-PAGE and Western transfer. Immunodetection was performed using Penta•His HRP Conjugate (for L338P M347T His6-Ffh) and a monoclonal α-Ffh antibody (for cellular Ffh and Ffh ). 52

C Results

Fig. C-1: Degradation of cellular Ffh and plasmid-derived Ffh at different growth phases. (A) Protein stability was analyzed over the entire bacterial growth with in vivo degradation experiments typically at 37°C. Therefore degradation cultures were taken at indicated growth phases (I to V). The shown growth curve and standard deviations were calculated from three biological replicates. Protein stability was analyzed for plasmid-encoded His6-Ffh (pBO3630) in E. coli MC4100 at 37 °C (B), whereby gene expression was induced by adding 50 ng/µl AHT for 30 min. Degradation experiments were conducted for plasmid-encoded FfhL338P M347T (pSKPP10) in E. coli SKP1101 at 30°C (C) and cellular Ffh (D) at 37°C. Afterwards, translation was stopped by the addition of spectinomycin (Sp), samples were taken and prepared for SDS-PAGE. After SDS-PAGE, Western transfer and immunodetection half-lives (T½) and standard deviations were calculated from three biological replicates.

C Results

To examine growth phase-dependent degradation of overproduced His6-Ffh, we performed in vivo degradation experiments in MC4100 at 37 °C (Fig. C-1 B). His6-Ffh was degraded rapidly in late exponential (T½ = 15 min) and early stationary growth phase (T½ = 14 min) and was a more stable protein in early exponential (T½ = 111 min), mid-exponential (T½ = 49 min) and late stationary phase (T½ = 57 min). These results confirmed our previous observations and were consistent with a previous publication showing half-lives of approximately 20 min if high copy plasmid-derived Ffh is not bound to the 4.5S RNA [168]. To analyze whether the copy number of the plasmid or the RNA-binding ability influence the growth phase-dependent proteolysis of Ffh, we performed in vivo degradation experiments at 30 °C with a temperature-sensitive ffh mutant SKP1101 + pSKPP10 (Fig. C-1 C). To reduce the intracellular Ffh level, the chromosomal ffh was disrupted by ffh1::kan and to sustain viability of the ffh mutant, it was complemented with a constitutive medium-copy plasmid encoding the FfhL338P M347T variant. The FfhL338P M347T variant has an inefficient 4.5S RNA- binding ability resulting from two amino acid changes within the C-terminal M domain (L338P and M347T) [271]. Our in vivo degradation experiments showed that the FfhL338P M347T variant is rapidly degraded with similar half-lives in early (T½ = 24 min), mid (T½ = 21 min) and late

(T½ = 20 min) exponential growth phase. The protein was more stable in later growth phases

(IV: T½ = 41 min; V: T½ = 108 min). Compared to His6-Ffh, the growth-phase-dependent degradation was slightly altered, since in earlier growth phases (I to III) the FfhL338P M347T variant was more unstable. These results are comparable to previous studies, where the FfhL338P M347T variant was degraded with a half-life of 10 min (at 30°C) [271]. Moreover, growth phase-dependent degradation of FfhL338P M347T variant, which has no additional tag neither at the N- nor C-terminus suggest that N-terminal His6-tag of His6-Ffh does not influence proteolysis. Previous studies on cellular Ffh degradation showed a stable protein in mid-exponential growth phase in MC4100 for 24h (79 % Ffh have remained) and a more unstable protein in a ΔmsrAΔmrsB deletion mutant, which was defective in the repair of oxidative protein damages [98]. In general, oxidative protein damage was increased during stationary growth phase [264]. Based on these results we assayed whether Ffh is subject to proteolysis when oxidative stress is highest in stationary growth phase. The stability of cellular Ffh in E. coli MC4100 was analyzed at 37 °C over the entire growth (Fig. C-1 D). Our results showed, that cellular Ffh was barely degraded in early (T½ = 103 min) and mid-exponential growth phases (T½ = 83 min) and fully stable with half-lives of >120 min in later growth phases (III to V). In conclusion, cellular

C Results

Ffh was a stable protein, but plasmid-derived Ffh variants were subject to growth phase- dependent proteolysis.

1.2 Lon is the major protease responsible for Ffh degradation To identify the AAA+ protease/s responsible for Ffh degradation, we performed in vivo + degradation experiments with plasmid-derived His6-Ffh utilizing different AAA protease- deficient E. coli strains (Fig. C-2) and their corresponding parental strains (Fig. H-1) during different bacterial growth phases. The growth phase-dependent degradation pattern of His6-Ffh was indistinguishable in the parental E. coli strains (MG1655, W3110, BW25113). In the single knock out mutant strains for the membrane-anchored FtsH protease (ΔftsH) and the cytosolic

HslUV protease (ΔhslUV), His6-Ffh was rapidly degraded in late exponential (III) and/or early stationary (IV) growth phase and was more stable in early exponential (I) and late stationary growth phase (V), a pattern similar to the WT strains.

Fig. C-2: Half-lives of plasmid-derived Ffh in AAA+ protease deficient E. coli strains. A summary of half-lives for degradation of plasmid-derived His6-Ffh in WT (MC4100), ΔftsH, ΔhslUV, ΔclpP, Δlon and ΔlonΔhslUVΔclpP is given. Half-lives and standard deviations for plasmid-derived His6-Ffh degradation were calculated from three biological replicates.

For a strain lacking the cytosolic ClpAP/XP proteases (ΔclpP), His6-Ffh degradation was growth phase-dependent in general, but slightly shifted. In early stationary growth phase (IV) the half-life was approx. fourfold increased (T½ = 56 min) compared to the corresponding WT 55

C Results

BW25113 (T½ = 15 min). Considerable differences where observed in the lon deletion strain

(Δlon), where His6-Ffh was a more stable protein at every tested growth phase (T½ = from

54 min to >120 min) in comparison to the parental strain MC4100. For example, His6-Ffh stability (T½ = 74 min) was increased fivefold at the early stationary growth phase (IV) in Δlon, whereas His6-Ffh was degraded efficiently in MC4100 (T½ = 14 min). Since Ffh was not fully stabilized in a strain lacking the Lon protease, we analyzed Ffh stability in a strain lacking the non-essential and cytosolic proteases Lon, ClpAP/XP and HslUV. It is known that various AAA+ proteases share some of their substrates. For example the Lon substrate CueR is degraded by Lon, ClpAP and ClpXP [39]. In the AAA+ protease triple knock out mutant

ΔlonΔhslUVΔclpP, His6-Ffh was a much more stable protein (T½ = from 97 min to >120 min), than in a Δlon strain showing that Lon is the major protease for Ffh degradation and that ClpAP/XP and/or HslUV play a minor role as backup system. To see whether the presence of Lon and ATP is sufficient for Ffh proteolysis, in vitro degradation experiments with purified Lon and Ffh were attempted (Fig. H-2). Poly-histidine affinity-tagged His6-Ffh and Lon-His6 proteins were successfully purified by Ni-NTA affinity chromatography, and the stability of Ffh was investigated by in vitro degradation experiments over 120 min with or without the Lon protease being present. The reaction was started by addition of ATP, samples were taken at defined time points, prepared for SDS-PAGE and detected by Coomassie staining or immunoblot. Our results showed an unstable His6-Ffh protein under any tested condition in vitro, even in the absence of Lon protease. Therefore, it was not possible to show His6-Ffh degradation in vitro.

To summarize, we found that Lon is the major protease for His6-Ffh proteolysis. To get insights into the physiological relevance of the growth phase-dependent degradation of Ffh, we analyzed protein stability of cellular Ffh under different stress conditions, e.g. heat stress.

1.3 Cellular Ffh is a stable protein under different stress conditions Cellular Ffh was a stable protein under normal growth conditions (LB medium, 37°C). To analyze if cellular Ffh is degraded under other growth conditions, we performed in vivo degradation experiments at diverse stress conditions. First, we determined Ffh stability under heat shock conditions, because Lon is part of the heat shock regulon, and the protease amount is increased upon shift to heat stress on both transcript and protein level [141, 277]. Transcriptome analysis demonstrated that the lon transcript is significantly increased after 15 min of heat stress while ffh transcript was not [141]. Based on these findings, we also assumed an increased Lon amount upon heat stress induction in our experiments and thus a 56

C Results more unstable Ffh protein. In vivo degradation experiments were performed exclusively in early exponential growth phase, because cellular Ffh degraded to some extend at this growth phase (Fig. C-1 D). Additionally, we expected a greater increase of Lon because cellular Lon amount should be lower in earlier growth phases than in later growth phases. Thus, we might expect a more rapid degradation in earlier growth phase in MC4100. For in vivo degradation experiments, an E. coli culture was grown to an OD580 of 0.5 and divided into five degradation cultures (Fig. C-3 A). To induce a heat shock response, cells were shifted to 42 °C for 0, 5, 10, 20, or 30 min.

Fig. C-3: Degradation of cellular Ffh after heat stress. (A) Stability of cellular Ffh was analyzed under heat stress conditions with in vivo degradation experiments. A main culture was grown at 30 °C up to an OD580 0.5 and was divided into five degradation cultures. Each degradation culture was exposed to 42 °C for various incubation times (0-30 min) to induce heat stress. Stability of cellular Ffh was analyzed in E. coli MC4100 (B) and Δlon (C). Translation was stopped by the addition of Sp, samples were taken and prepared for SDS-PAGE. After SDS-PAGE, Western transfer and immunodetection, half-lives were calculated.

The degradation culture without temperature shift (0 min) served as control. In vivo degradation experiments in MC4100 (Fig. C-3 B) and Δlon (Fig. C-3 C) revealed a fully stabilized cellular Ffh protein with half-lives from 103 min to >120 min in E. coli MC4100 and Δlon after heat stress induction. In order to find conditions affecting the stability of cellular Ffh, we performed further in vivo degradation experiments under different stress conditions. We investigated the stability after a directed overproduction of His6-Lon from a high copy plasmid in MC4100 and

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Δlon (Fig. H-3). At different growth phases, Lon overproduction was selectively started by the addition of 1 mM IPTG and subsequently the stability of Ffh was studied. Successful Lon overproduction was confirmed by immunoblot (data not shown). However, cellular Ffh was stable in MC4100 and Δlon in all growth phases. Moreover, we analyzed the stability of Ffh after a shift to oxidative stress conditions (Fig. H-4) because oxidized Ffh is unable to bind 4.5S RNA and is an unstable protein in a ΔmsrAΔmrsB deletion mutant [98]. Additionally, Ffh is also a destabilized protein if not bound to the 4.5S RNA [168, 271]. E. coli MC4100 or Δlon were grown to an OD580 of 0.5, oxidative stress was induced by adding 0-2 mM H2O2 to the degradation culture for 60 min and Ffh stability was monitored. Despite the induction of oxidative stress, Ffh was a stable protein. Finally, we examined Ffh stability in MC4100 after treatment with rifampicin and as control after spectinomycin and spectinomycin/rifampicin treatment (Fig. H-5). We investigated the impact of the transcription inhibitor rifampicin, as we suspected that the 4.5S RNA was no longer transcribed after the addition of rifampicin. Thus, we assumed an increased amount of free Ffh (not bound to the 4.5S RNA) in the cell. However, cellular Ffh was a stable protein over >120 min after rifampicin treatment during the bacterial growth. In conclusion, chromosomally evolved Ffh was a stable protein under all conditions tested.

1.4 The Ffh amount is modulated during bacterial growth Since Jensen et al. demonstrated that Ffh is a stable protein in complex with 4.5S RNA, they suggested that excess of free Ffh was rapidly degraded [168]. Based on these findings, we investigated whether an excess of His6-Ffh was responsible for the growth phase-dependent degradation or whether Lon was able to modulate the Ffh amount in the cell. First, the cellular amounts of Lon and Ffh in MC4100 at different growth phases were compared (Fig. C-4). L338P M347T Subsequently, the cellular amount of His6-Ffh in MC4100 and Δlon and the Ffh variant of the temperature-sensitive ffh mutant (SKP1101 + pSKPP10) were determined. For these experiments, bacterial cultures of MC4100/Δlon or SKP1101+pSKPP10 were grown at 37°C and 30°C, respectively. Samples were taken at t = 0 in growth phases I to V as described for the in vivo degradation experiments (Fig. C-4 A). Samples for plasmid-derived His6-Ffh were withdrawn after an overproduction induced by 50 ng/µl AHT for 30 min. Subsequently, the samples were subjected to SDS-PAGE, Western transfer and immunodetection with either α-Lon or α-Ffh. For quantification of the Lon- or Ffh amount, a sample from early exponential growth phase (I) was assumed to be 100 % and further protein amounts were calculated based

C Results on this value. The total protein extracts were used again as an internal loading control for an SDS-PAGE with subsequent Coomassie staining.

Fig. C-4: Relative protein amounts of cellular Lon, Ffh and plasmid-derived Ffh. The relative protein amounts were analyzed for cellular Lon in MC4100 (A) and cellular Ffh in MC4100 (B) and

Δlon (C). Furthermore, the relative Ffh amount for plasmid-derived His6-Ffh in MC4100 (D) and Δlon (E) as well as the FfhL338P M347T variant in SKP1101 (F) was determined. Samples were taken at defined time points of bacterial growth (growth phase I to V) and prepared for an SDS-PAGE. For plasmid- derived His6-Ffh, samples were taken after His6-Ffh overproduction, which was induced by the addition of 50 ng/µl AHT. After SDS-PAGE and Western transfer, cellular Lon was immunologically detected with a α-Lon antibody and Ffh with a monoclonal α-Ffh antibody. Cellular amounts of Lon or Ffh in MC4100 at growth phase I were defined as 100% for quantification of the relative protein amounts in the following growth phases. The results are based on three biological replicates. As a internal loading control, a section of a Coomassie stained SDS gel at about 70 kDa is shown.

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The experiments showed an almost constant cellular amount of the Lon protease at any tested bacterial growth phase (Fig. C-4 A). In contrast, the cellular Ffh amount in MC4100 was highest in the early exponential growth phase and was gradually decreased to about 31 % of the initial value in the following growth phases (Fig. C-4 B). In a Δlon mutant, the cellular Ffh amount (remaining 64 % Ffh) also decreased to some extent during bacterial growth. This is in agreement with an expected contribution of other protease(s) (Fig. C-4 C). The Lon-mediated degradation of Ffh was most evident when His6-Ffh was produced from an AHT-inducible high copy plasmid, Ffh levels were 23 times higher in MC4100 and 31 times higher in Δlon compared to cellular Ffh in growth phase I (Fig. C-4 D+E). During bacterial growth, His6-Ffh amount was rapidly removed in MC4100 to about 3-fold higher levels than endogenous Ffh in late stationary growth phase (V). In contrast, the removal of excess Ffh in Δlon was muchless efficient and left a 15-fold excess of His6-Ffh over cellular Ffh. For the temperature-sensitive ffh mutant SKP1101 + FfhL338P M347T we observed a comparable Ffh amount in early exponential growth phase as for cellular Ffh in MC4100 (Fig. C-4 F). The amount of the FfhL338P M347T variant gradually decreased during bacterial growth. These results above show that the Ffh amount in the presence of Lon was successively decreased during bacterial growth, while the amount of Lon remained constant. Ffh degradation by Lon or other AAA+ proteases (ClpAP/XP or HslUV) in earlier growth phases reduces excess Ffh. After transition to stationary growth phase, when the Ffh amount was lowest, it was a stable protein probably to maintain a certain minimal level in the cell. To obtain further insights into the degradation mechanism of Ffh, the impact of the sigma factor RpoH (σ32) on Ffh degradation was investigated. RpoH was chosen because interaction between RpoH and Ffh has been demonstrated [213, 237]. It is suggested that this interaction is a negative feedback mechanism in RpoH regulation in which the SRP complex targets RpoH to the inner membrane for FtsH-mediated proteolysis.

1.5 Impact of RpoH on Ffh proteolysis Lon proteolysis can be mediated by substrate-specific adaptor proteins. For instance, in B. subtilis Lon degrades the master flagellar activator protein SwrA only in the presence of the swarming motility inhibitor A (SmiA). The alternative heat-shock sigma factor RpoH was considered as putative factor or adaptor protein to mediate Ffh proteolysis. Previous studies revealed a direct interaction of RpoH and the SRP complex (Fig. C-5 A) [213, 237]. Ffh was found to bind RpoH via its signal peptide binding site within the M domain. The SRP complex targets RpoH to the SecYEG translocation system through the SRP complex-SR interaction and 60

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RpoH is degraded by the membrane-anchored AAA+ protease FtsH under non-heat shock conditions (Fig. C-5 A). Our finding that Ffh is a substrate of growth phase-dependent proteolysis by Lon motivated us to analyze the stability of His6-Ffh in a rpoH-deficient mutant.

Fig. C-5: Impact of the sigma factor RpoH on Ffh proteolysis. (A) Previous studies demonstrated an interaction of the SRP complex with RpoH. Ffh binds RpoH via its signal peptide binding site, targets RpoH to the SecYEG translocase system through the SRP complex-SR interaction and finally, RpoH is degraded by the membrane-anchored AAA+ protease FtsH under non-heat shock conditions [213, 237]. On the other hand, Ffh, the protein component of the signal recognition particle is subject to Lon- dependent proteolysis. (B) Degradation experiments for MC4100 (C) and ΔrpoH (D) were performed to defined growth phases (I to IV) at 25 °C. Translation was stopped by the addition of spectinomycin (Sp), samples were taken and prepared for SDS-PAGE. After SDS-PAGE, Western transfer and immunodetection, half-lives were calculated from two biological replicates. (E) In addition, relative Lon amount of the time point “0 min” of the degradation culture was determined in MC4100 and ΔrpoH. After SDS-PAGE and Western transfer, Lon was immunologically detected with α-Lon antibody. Relative Lon amount was calculated based on growth phase I in MC4100, because this sample was assumed to be 100%.

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Therefore, we performed in vivo degradation experiments of plasmid-derived His6-Ffh (Fig. C-5 C+D) and cellular Ffh (Fig. H-6) in MC4100 and ΔrpoH at 25 °C. We conducted degradation experiments exclusively in growth phase I to IV according to growth phases of MC4100 at 25 °C because it was difficult to distinguish between growth phases in the ΔrpoH mutant (Fig. C-5 B). His6-Ffh was degraded in MC4100 at 25 °C with half-lives from 29 to 59 min in every tested growth phase with a moderate trend of growth phase-dependence

(Fig. C-5 C). His6-Ffh was a more stable protein with half-lives from 60 to 102 min in a rpoH- deficient background indicating a direct or indirect impact of RpoH on His6-Ffh proteolysis (Fig. C-5 D). Additionally, the stability of cellular Ffh was analyzed in rpoH-deficient background. Different from experiments done at 37 °C, cellular Ffh was degraded comparatively fast at 25 °C in growth phase I with T½ = 43 min and was stable in later growth phases (T½ = >120 min) in MC4100 (Fig. H-6 A). In contrast, Ffh was a stable protein with half-lives from 94 min in growth phase I and >120 min in later growth phases in ΔrpoH (Fig. H- 6 B). Compared to MC4100, cellular Ffh was stabilized in ΔrpoH in growth phase I in accordance with the results of plasmid-derived His6-Ffh. To demonstrate whether the amount of Lon was responsible for a more stable Ffh protein in the absence of RpoH, the cellular Lon amounts in MC4100 and ΔrpoH were determined. Samples were taken for each degradation culture at t = 0 after spectinomycin addition. The Lon amount was quantified without (Fig. H-6 C) and after overproduction of His6-Ffh (Fig. C-5 E). After SDS-PAGE and Western transfer, Lon was immunologically detected with α-Lon antibody. The relative Lon amount was determined, whereby growth phase I of MC4100 defined as 100 % and further Lon amounts for MC4100 and ΔrpoH were calculated relative to this value. After

His6-Ffh overexpression in MC4100 the relative Lon amount was constant in growth phases I (100 %) and II (110 %) and approx. 2 times increased in growth phases III In general, the cellular Lon amounts in the ΔrpoH mutant were lower than in MC4100, so a more stable Ffh protein could result from Lon availability. Comparable results were obtained for samples without overproduction of Ffh (Fig. H-6 C). For ΔrpoH a constant Lon amount was shown in all tested growth phases (from 129 % to 160 % relative to its initial value) after overproduction of His6-Ffh. In contrast, the relative Lon amount was increased during bacterial growth without overproduction of His6-Ffh. In this case, it is noticeable that in ΔrpoH the initial Lon amounts in growth phase I and II were lower compared to MC4100 and increased during further bacterial growth. In general, the cellular Lon amounts in the ΔrpoH mutant were lower than in MC4100, so a more stable Ffh protein could result from Lon availability. In conclusion, plasmid-derived

His6-Ffh was rapidly degraded in the presence of RpoH and was more stable in the absence of

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RpoH. This has also been shown for cellular Ffh in the early exponential growth phase. This effect can be caused by the availability of the Lon protease or possibly by the Ffh-RpoH interaction itself.

2. Investigations on the recognition mechanism of Ffh by the Lon protease 2.1 The Ffh M domain is recognized by the Lon protease A previous master thesis had shown that the Lon protease recognizes Ffh within its M domain and this study showed further investigations regarding the Ffh recognition mechanism [23]. The cytoplasmic SRP complex from E. coli represents the simplest variant during evolution and consists of Ffh and the 4.5S RNA (Fig. C-6 A) [31, 280, 292, 298]. Ffh can be divided into two functional domains, the NG and M domain. The methionine-rich M domain binds signal sequences of nascent proteins at the ribosome and interacts with the 4.5S RNA [152, 221, 299, 413]. The GTPase-harboring NG domain interacts with its membrane-anchored SR-receptor FtsY via GTP hydrolysis [74, 83, 233, 310]. To verify the recognition region for Ffh proteolysis we separately analyzed the stability of the two functional Ffh domains (NG and M domain) in MC4100 by in vivo degradation experiments at different growth phases (Fig. C-6). Each domain was overproduced from a high copy plasmid with an AHT-inducible promotor as a N-terminal His6-tagged variant. Plasmid-derived His6-

NG was a stable protein (T½ = >120 min) during bacterial growth in three biological replicates

(Fig. C-6 A). In contrast, His6-M was degraded at each tested growth phases with half-lives from 19 to 49 min (Fig. C-6 B). The degradation profile differs from His6-Ffh full-length protein, since the His6-M was no longer stabilized in late stationary growth phases. Since it was not possible to detect signals for His6-M in late growth phases (IV and V) in this work half- lives for growth phase I to III were calculated from three biological replicates and for growth phases IV and V data were adopted from the M. th. of Beate Sauerbrei [23].

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Fig. C-6: Degradation of the Ffh domains (NG and M domain).(A) The simplest SRP variant exists in E. coli containing the Ffh protein and the 4.5S RNA. To determine a putative motif responsible for Lon recognition we separately monitored stability of the two functional domains, NG (B) and M domain (C). Protein stability was analyzed during different growth phases (growth phase I to IV) by in vivo degradation experiments in MC4100 at 37°C. Overproduction of the two functional domains was induced by the addition of 50 ng/µl AHT for 30 min. Afterwards, translation was stopped by the addition of spectinomycin (Sp), samples were taken to defined time points and prepared for SDS-PAGE. After SDS-PAGE, Western transfer and immunodetection, half-lives and standard deviations were calculated from three biological replicates. Data for the degradation of the M domain were taken from the M. th. of Beate Sauerbrei [23].

Overall, the results show that the M domain is an unstable protein, whereas the NG domain is stable in the cell. Whether the Lon protease is responsible for the degradation of the M domain could not be tested directly by in vivo degradation experiments in a lon-deficient strain because transformation of the expression plasmid for the N terminal His6-tagged M domain let to a synthetic lethal phenotype in a lon-deficient strain [23].

2.2 Plasmid-encoded M domain induces a lethal phenotype in E. coli To investigate the synthetic lethal phenotype of the M domain in Δlon, the transformation efficiencies for N-terminal His6-tagged constructs were determined for His6-Ffh or its functional domains (His6-NG or His6-M) (Fig. C-7 A-C). To analyze whether the M-domain induced lethal phenotype can be reversed by another construct in a lon-deficient strain, the transformation efficiencies of One hybrid constructs were investigated (Fig. C-7 D-F). These 64

C Results plasmids code for the T25 and T18 domains of the adenylate cyclase from Bordetella pertussis and flanking the gene/gene region of interest. The transformation efficiencies were determined in MC4100 and Δlon by counting the plasmid-carrying bacterial colonies after transformation.

All plasmids encoding the His6-tagged variants of Ffh, or the NG or M domains could be transformed into the MC4100. The transformation efficiencies compared to the empty vector

(EV) control were two-fold lower for plasmid-coded His6-Ffh in MC4100. His6-NG showed transformations efficiencies comparable to His6-EV in MC4100. Transformation efficiency of

His6-M was 1,840 times lower than His6-EV. In contrast, only His6-EV, His6-Ffh and the His6-

NG could be transformed into Δlon. Transformation efficiency of His6-Ffh was similar to the

EV control, whereas His6-NG was 2 times increased. After transformation of plasmid-encoded

His6-M in Δlon, no colonies were found confirming the synthetic lethal phenotype of the His6-M construct. One hybrid constructs encoding for T25-Ffh-T18, T25-NG-T18 and T25-M-T18 was transformed into MC4100 and Δlon. In MC4100 transformation efficiencies for T25-Ffh-T18, T25-NG-T18 and T25-M-T18 were slightly decreased in comparison to the T25-EV-18 control, but they showed comparable values to each other. Similar results were observed for Δlon, whereby overall transformation efficiencies were slightly lower than in MC4100. Thus, it was possible to reverse the synthetic lethal phenotype of His6-M by the One-hybrid construct T25- M-T18 in Δlon. The colonies formed after transformation of tested constructs showed various diameters (data not shown). Strains carrying plasmid-encoded His6-Ffh or His6-M formed larger colonies than strains carrying His6-EV or His6-NG. In contrast, strains coding for a One Hybrid construct formed smaller colonies in general. These observations motivated us to analyze the cell morphology of plasmid-carrying MC4100 cells in exponential and stationary growth phase by light microscopy (Fig. H-7). Elongated cells were induced by the presence of plasmid-encoded

His6-M in exponential and stationary growth phase. The cells effect was more pronounced in the stationary than in exponential growth phase. Colonies formed after transformation of other

His6-tag constructs (His6-Ffh, His6-NG), as well as One hybrid constructs (T25-Ffh-T18, T25- NG-T18 and T25-M-T18) did not induce changes in cell morphology in comparison to the respective empty vector control.

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Fig. C-7: Transformation efficiencies of plasmid-encoded Ffh and its functional domains. Transformation efficiencies were determined for Ffh and its functional domains, NG or M domain, as

His6-tag construct or One Hybrid construct. (A) His6-tag constructs encoded an N-terminal His6-tag as well as the corresponding gene sequence (gene X) und the control of a tet promotor (Ptet). Transformation efficiencies were determined for His6-tag constructs in (B) MC4100 and (C) Δlon. (D) Plasmids of the One hybrid constructs encoding the T25 and T18 domains of the adenylate cyclase cyaA from B. pertussis and the corresponding gene sequence was cloned between both domains under the control of a lac promotor (Plac). Transformation efficiencies of the One hybrid constructs were also determined in (E) MC4100 and (F) Δlon. For each construct and strain the corresponding empty vector (EV) was used as a control. Transformation efficiencies were given as “(transformant/ng DNA) x 108” and were calculated from three independent experiments.

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Interestingly, a single clone could be recovered after transformation of the M domain in Δlon and it provided insights into the region within the M domain that triggered the lethal phenotype. The plasmid DNA of the clone was isolated and sequencing revealed a deletion covering the coding region for amino acids 388 to 453, in which the RNA-binding motif of the M domain (aa 382-407) is located. A schematic overview of the deleted region and its localization is shown in Fig. H-8. Thus, we showed that a partly deleted RNA-binding motif was able to reverse the synthetic lethal phenotype of the M domain in Δlon. The availability of an intact or accessible RNA-binding motif seems to mediate the synthetic lethal phenotypes in Δlon. Additionally cell morphology was altered by plasmid-encoded His6-M in E. coli WT.

2.3 Impact of point mutations on stability of Ffh and cpSRP54 from A. thaliana SRP complex is conserved in all three domains of life and its complexity increased during evolution. The prokaryotic SRP complex is the simplest variant consisting of the protein component Ffh and the SRP RNA, e.g. 4.5S RNA in E. coli [31, 280, 292, 298]. Interestingly, chloroplasts of higher plants (e.g. A. thaliana) contain the highly conserved cpSRP54 and the additional protein cpSRP43 [104, 186, 323]. The SRP RNA absent in chloroplasts is, and cpSRP54 carries a mutation of the conserved RNA-binding motif within its M domain [293,

301]. Combined with our result that (A) His6-M is an unstable protein, (B) plasmid-encoded

His6-M induces a synthetic lethal phenotype in Δlon and (C) replacing this effect by deletion of the RNA-binding motif, raises the possibility of a putative recognition sequence within the RNA-binding motif. We investigated whether the stability of Ffh (and cpSRP54) could be altered by point mutations within the RNA-binding motif or whether Ffh degradation is a conserved mechanism. Fig. C-8 A shows a protein model of the whole SRP complex interacting with its SR FtsY, and the exact localization of S382, C406 and G405. All mentioned amino acids are involved in direct intermolecular contact between Ffh and the 4.5S RNA [20]. Binding of the SRP RNA occurs via a helix-turn-helix motif consisting of four helices (Fig. C-8 B). Helix 2b and helix 3 of the RNA-binding motif is mainly involved in the protein-RNA interaction. The conserved SRP RNA-binding motifs SM and GXG from E. coli are mutated to VM and DXG in higher plants (e.g. A. thaliana), which results in the loss of SRP RNA-binding ability [293, 301]. The analyzed amino acids of the SRP RNA-binding motif were named according to the position of Ffh from E. coli (S382M383 and G405X406G407) in the following. First, we investigated the impact of the evolutionary point mutations on Ffh degradation in E. coli by creating N-terminally His6-tagged Ffh variants with single or multiple point mutations. 67

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Fig. C-8: SRP structure and localization of the evolutionary point mutations. (A) Shown is the structure of the SRP complex (Ffh: green; 4.5S RNA: grey) interacting with its membrane-anchored receptor FtsY (FtsY: blue). The protein model was generated based on the structural analysis (PDB ID: 2XXA) from Ataide et al. 2011 [11]. (B) Schematic overview of the organization of the M domain (aa 296 to 453). The helix-turn-helix motif, which is relevant for SRP RNA-binding, consists of four helices. The RNA-binding motif is located from amino acid 382 to 407. The indicated point mutations S382V, G405D and C406S (red-colored amino acids) lead to the loss of SRP RNA-binding ability in cpSRP54 from A. thaliana. Asterisks mark conserved amino acids.

The stability and solubility of the Ffh variants were investigated after overproduction from a AHT-inducible high-copy plasmid (Fig. H-9). If one of the tested point mutations had an effect on the recognition by the Lon protease, the Ffh variant should be more stable in in vivo degradation experiments. These experiments were done as part of a bachelor thesis and of this work [101]. The point mutations S382V or C406S as single or double point mutations did not alter the growth phase-dependent degradation of Ffh in MC4100. In contrast, stability was increased for all Ffh variants (single or multiple point mutations) carrying a point mutation at position G405, but all these Ffh variants were insoluble. However, it remains unclear whether these Ffh variants were really insoluble in vivo or whether they resulted from the experimental procedure during the solubility studies. Taken with care, these results suggest that a point

C Results mutation at position G405 has an effect on the degradation of Ffh preventing recognition by the Lon protease. Since the exchange of the neutral glycine to the acidic aspartic acid was drastic and this could be a reason for the possible insolubility, milder point mutations at position 405 were introduced. The point mutations G405A and G405N also led to a more stable protein in general. However, these Ffh variants were degraded during exponential growth phase with half- lives of 42 and 74 min, respectively. Unfortunately, the G405A exchange also let to an insoluble protein.

Fig. C-9: Transformation efficiencies and degradation of cpSRP54 in E. coli. During evolution, the composition of the SRP complex has changed. (A) The chloroplast SRP complex from A. thaliana consists of two protein components, the highly conserved cpSRP54 and the additional cpSRP43. (B)

Transformation efficiencies were determined for plasmid-encoded N-terminal His6-tagged full-length cpSRP54 (His6-cpSRP54_At) or only M domain (His6-M_At) of A. thaliana. The empty vector (His6- EV) was used as control. The results are based on three independent experiments. Protein stability of cpSRP54 (C) and cpSRP54V455S D480G (D) from A. thaliana was determined by in vivo degradation experiments in E. coli MC4100 at defined growth phases at 37 °C. Degradation cultures were taken at indicated growth phases (I to V) and heterologous protein overproduction was induced for 30 min by adding 50 ng/µl AHT. Afterward, translation was stopped by the addition of Sp, samples were taken and prepared for SDS-PAGE. After SDS-PAGE, Western transfer and immunodetection half-lives (T½ ) were calculated. Standard deviations were determined from three biological replicas.

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In conclusion, the results the Ffh variants with point mutations on amino acid G405 must be interpreted carefully due to their insolubility. It cannot be clearly concluded whether an exchange of amino acid G405 also in natively folded Ffh leads to an altered recognition by Lon. Finally, we investigated whether cpSRP54 from A. thaliana can be recognized and degraded by Lon from E. coli. These results should provide an indication of whether this is a conserved degradation mechanism or whether the missing SRP RNA-binding ability influences the degradation of cpSRP54. Protein stability of cpSRP54 in E. coli was analyzed with in vivo degradation experiments of cpSRP54 from A. thaliana in MC4100 over the entire growth curve

(Fig. C-9 C). The results showed for His6-cpSRP54 a slow degradation in growth phase I

(T½ = 77 min) and II (T½ = 71 min). In later growth phases (III to IV) His6-cpSRP54 was a stable protein (T½ = 112 min and > 120 min). Thus, the growth phase-dependent degradation as described for E. coli Ffh does not apply to cpSRP54, although some degradation was observed. To check the impact of the evolutionary point mutations and the SRP RNA-binding V455S D480G ability on the stability of cpSRP54, we analyzed the mutated His6-cpSRP54 , which had a recovered SRP RNA-binding ability [293]. These point mutations corresponded to the point mutations S382V (V455S in A. thaliana) and G405D (D480G in A. thaliana) in E. coli.

The mutated protein was slightly more unstable in growth phase I (T½ = 59 min) and II

(T½ = 67 min) in comparison to wild-type His6-cpSRP54 and it was stable in later growth V455S D480G phases (III to IV) (Fig. C-9 D). Both proteins, His6-cpSRP54 and His6-cpSRP54 were stable (T½ = > 120 min) in growth phase I and II in Δlon in a single experiment (data not shown) suggesting Lon-mediated proteolysis. It seems that an intact RNA-binding motif is needed for recognition by Lon.

Above it was shown that plasmid-encoded His6-M leads to a synthetic lethal phenotype in a lon-deficient E. coli strain, which was thought to be also induced by an intact SRP RNA- binding motif. To support this hypothesis, we determined the transformation efficiencies of plasmid-encoded His6-cpSRP54_At (full-length protein) and His6-M_At (M domain only) in

MC4100 and Δlon (Fig. C-9 B). We found that the transformation of His6-cpSRP54 and His6- cpSRP54_M into MC4100 and Δlon was possible with comparable transformation efficiencies although in both cases it was lower in comparison to the empty vector. Thus, plasmid-encoded cpSRP54 from A. thaliana or its M domain do not induce a synthetic lethal phenotype in E. coli Δlon, and this supports the above-formulated hypothesis. In summary, an intact SRP RNA- binding motif induced the synthetic lethal phenotype of a lon deletion mutant. Furthermore, it seems to play a role in the recognition of Ffh and cpSRP54 by the cytosolic Lon protease.

Discussion

D Discussion

D Discussion ATP-dependent AAA+ proteases play a crucial role in the regulation of cellular protein homeostasis by degrading certain proteins at defined conditions. On the one hand, misfolded and aggregated proteins are degraded during protein quality control and on the other hand, natively folded functional proteins are degraded during regulated proteolysis. Current research is focused on the identification of novel AAA+ protease substrates using global in vivo trapping approaches (reviewed in [289]). Prerequisite for such an approach is a protease variant with a functional ATPase domain ensuring substrate binding and translocation and a proteolytically inactive protease domain preventing degradation by the AAA+ protease. An in vivo trapping approach of the membrane-bound FtsH protease revealed Ffh, the protein component of the essential SRP complex mediating the co-translational protein transport of inner membrane proteins, as a potential AAA+ protease substrate [10]. Ffh was unstable in the exponential as well as the stationary growth phase. Surprisingly, further investigations have shown that FtsH is not responsible for Ffh degradation but the cytosolic Lon protease. Possible reasons why Ffh was found via an FtsH trapping approach and not via a Lon trapping approach will be discussed later. First investigations on Ffh proteolysis demonstrated a growth phase-dependent degradation mechanism by the cytosolic Lon protease, in which Ffh became rapidly degraded in the transition from exponential to stationary growth phase and was stable in earlier and later growth phases (Arends, unpubd.; [23]). This transition phase is critical for survival because the whole cellular proteome is reprogrammed both on a pre-translational and post-translational level in response to several limitations. In this stringent cellular environment, the proteolytic degradation of Ffh could be a regulated process. Thus, we were interested first in factors or conditions that caused growth phase-dependent degradation of Ffh, and second in the substrate recognition mechanism by the cytosolic Lon protease.

1. Modulation of the Ffh degradation mechanism in E. coli 1.1 Growth phase-dependent Ffh proteolysis is related to its cellular amount Ffh is the protein component of the essential SRP complex, which mediates co-translational protein targeting in E. coli. In addition to Ffh, the bacterial SRP complex contains an SRP RNA (4.5S RNA in E. coli) and is the simplest variant in comparison to the highly developed counterpart in mammalians, which consists of six proteins and a complex SRP RNA (7S RNA) [31, 280, 292, 298, 360, 374, 382]. Already in the 1990s, it was shown that plasmid-derived Ffh from a high-copy plasmid is an unstable protein in E. coli and that its stability is increased 72

D Discussion by 4.5S RNA co-expression [168]. It has been assumed that excess Ffh is subjected to proteolysis when it is not bound to the 4.5S RNA. However, it was not investigated, which protease is responsible for Ffh degradation. Another group has shown that the AAA+ proteases Lon and HslUV become essential by reduced SRP complex levels that do not impair cell growth [30]. These findings suggest a possible link between the SRP complex and AAA+ proteases. Previously, we discovered a direct connection between the SRP complex and the Lon protease since Ffh is degraded Lon-dependently at defined growth phases (Arends, unpubd.; [23]). Growth phase-dependent proteolysis is a common mechanism for the degradation by AAA+ proteases. For instance, YfgM predicted as an ancillary SecYEG translocon subunit and a negative regulator of the RcsB-dependent stress response was rapidly degraded in stationary phase by the essential FtsH protease while it was more stable in earlier growth phases [40, 386]. Stability of YfgM was altered in cells that completely lack the alarmone (p)ppGpp, which induces the stringent response under starvation conditions [40, 76, 163, 354, 394]. Also, the Lon protease participates in growth-regulated proteolysis of the DNA replication inhibitor CspD [200]. It is rapidly degraded in fast-growing cells (mid-exponential phase) and proteolysis is slowed down in slowly growing cells (lag- and stationary phase). The growth phase- dependent Ffh proteolysis and the essential role of Ffh in viability and efficient protein targeting suggested a strictly regulated process and motivated us to investigate the degradation mechanism more closely. At the beginning of this project, in vivo degradation experiments comparing E. coli WT MC4100 and Δlon verified that Lon is the major protease in growth phase-dependent Ffh proteolysis. In accordance with our previous results (Arends, unpubd., [23]), overproduced Ffh

(His6-Ffh) from a high-copy plasmid was rapidly degraded in the late exponential and early stationary growth phases in WT (Fig. C-1 B). However, Ffh was more stable during the earlier and later growth phases. Compared to E. coli WT, the protein was more stable over the entire bacterial growth in a Δlon strain (Fig. C-2). Often AAA+ proteases can share substrates in regulated proteolysis. For example, the HTH-type transcription regulators CueR and ZntR are degraded in a synergistic manner by Lon/ClpAP/ClpXP and Lon/ClpXP, respectively [39, 283]. CueR proteolysis is mainly controlled by the Lon protease while Lon and ClpXP are the major proteases of ZntR degradation. Often quality control of proteins is mediated by more than one AAA+ protease ensuring the efficient degradation of aberrant and mislocalized proteins in the cell. In this context, SsrA-tagged proteins represent a well-studied example of degradation by different AAA+ proteases. The trans-translation system mediates ribosome rescue after ribosome stalling by incorrectly elongated and terminated translating proteins. The incomplete

D Discussion proteins are fused at their C-terminus with the SsrA tag, which acts as a degradation signal for ClpXP, ClpAP, FtsH and Lon [66, 124, 146]. The proteolysis of SsrA-tagged proteins prevents the accumulation of stalled ribosomes in the cell. The involvement of four ATP-dependent proteases in this essential process shows that substrate sharing could be essential and provides a backup system in the cell to ensure viability. We propose substrate sharing for Ffh proteolysis since Ffh is not fully stabilized in the absence of Lon. Analysis of Ffh stability in different AAA+ protease deficient strains indeed demonstrated a completely stable Ffh protein in the triple deletion mutant ΔlonΔhslUVΔclpP (Fig. C-2). Thus, HslUV and/or ClpAP/ClpXP seem to be involved in Ffh degradation but Lon is the major AAA+ protease for Ffh proteolysis. Ezraty et al. have demonstrated by pulse-chase experiments that cellular Ffh was stable over 24 h in E. coli MC4100, while 79 % of the initial protein amount remained [98]. Nevertheless, we studied whether the growth phase-dependent degradation of overproduced Ffh also applies to cellular Ffh, since Ezraty et al. demonstrated a stable Ffh exclusively in the early exponential growth phase. Slightly deviating from the results by Ezraty et al., we found a slow protein degradation (T½ = 103 min and 83 min) for cellular Ffh in the E. coli WT in the early and mid- exponential growth phase whereas in later growth phases it was stable for at least 120 min

(Fig. C-1 D). However, an overproduction of recombinant Lon-His6 in E. coli WT and Δlon did not accelerate the degradation of cellular Ffh (Fig. H-3). We demonstrate that cellular Ffh is barely degraded in E. coli, but this was different to the specific degradation pattern observed for plasmid-derived Ffh. This observation motivated us to investigate whether the degradation of cellular Ffh may be altered by other factors or stress conditions. A possible reason for the growth phase-dependent degradation of overproduced Ffh might be oxidative stress since the highest accumulation of reactive oxygen species (ROS) is assumed during the transition from exponential to stationary growth phase [258]. Furthermore, several pieces of evidence suggest a possible link between oxidative stress and Ffh stability. Cellular Ffh was subjected to proteolysis in a ΔmsrAΔmsrB mutant, which had a defect in the repair of oxidative protein damages [98]. In addition, oxidized Ffh was more unstable as it was not able to bind 4.5S RNA. It is also known that oxidative protein damage can promote the degradation of Lon substrates. For example, oxidized PerR, which changes its conformation as a result of oxidative stress, is thereby subjected to Lon-dependent proteolysis [4]. However, oxidative damage does not necessarily induce growth phase-dependent Ffh degradation, as cellular Ffh was more stable in the transition to the stationary phase than in earlier growth phases in E. coli WT. This result is supported by the unaltered degradation of cellular Ffh after induction of oxidative stress conditions (Fig. H-4).

D Discussion

Thus, there has to be another reason for the growth phase-dependent degradation of plasmid- derived Ffh. Our results provide evidence that the growth phase-dependent degradation of overproduced Ffh is associated with the amount of available Ffh and 4.5S RNA in the cell. We investigated the relative protein amounts of cellular Ffh and recombinant His6-Ffh (after overproduction) in E.coli WT and Δlon (Fig. C-4). The amount of cellular Ffh was gradually decreased during bacterial growth while Lon levels remained unaltered. Thus, the amount of cellular Ffh was high in early growth phases (early and mid-exponential phase), when cell division was fast. The cellular Ffh concentration is lowest in stationary phase, but its stability increases. We conclude for the first time that Ffh proteolysis is stopped when a certain threshold is reached maintaining a certain Ffh level in the cell to ensure co-translational protein targeting of inner membrane proteins. In a Δlon deletion mutant, the Ffh amount was also reduced, but not as drastically as in the E. coli WT (Fig. C-4 C). These results demonstrate that Lon is capable of specifically degrading cellular Ffh, and the degradation is not an effect caused by the overproduction of Ffh and altered protein homeostasis. In addition, this confirms that other proteases are likely to be involved in the degradation of cellular Ffh, as its concentration in slow-growing cells (e.g. stationary phase) was reduced even in the absence of Lon. In conclusion, at a high Ffh concentration, both cellular and plasmid-derived Ffh, appear to be degraded. As mentioned above, Jensen et al. hypothesized that excess Ffh is degraded in the cell if it is not bound to 4.5S RNA [168]. They demonstrated that plasmid-derived Ffh from a high-copy plasmid was unstable and Ffh stability could be restored by concomitant overproduction of 4.5S RNA. This also seems to be an explanation for the growth phase- dependent degradation of overproduced Ffh and the stabilization of cellular Ffh in later growth phases (lower Ffh level) because the 4.5S RNA is represented in the cell in excess over Ffh. Since the literature provided conflicting data on the proportion of Ffh and 4.5S RNA in the cell, Jensen et al. reassessed the concentration of both molecules and measured 400 molecules of the 4.5S RNA per 10000 ribosomes and only 100 Ffh molecules [168]. In the case of overproduced Ffh (high-copy plasmid-derived), we can assume that excess Ffh that has no 4.5S RNA bound will be degraded by Lon. Probably the amount of Ffh was reduced until all molecules were present in a complex with 4.5S RNA protecting Ffh against proteolysis. In stationary growth phase, about three times more molecules of overproduced Ffh remained compared to cellular Ffh, which roughly corresponds to three-quarters of excess 4.5S RNA in the cell described previously. Accordingly, overproduced Ffh could bind the excess (300 molecules) of free 4.5S RNA so that the protein is no longer degraded, which would explain the higher levels of overproduced Ffh in stationary phase. Hence, the amount of 4.5S RNA 75

D Discussion seems to be a limiting factor for Ffh degradation, which might explain the stabilization of plasmid-derived Ffh in the stationary growth phase. Dong et al. proposed that the amount of 4.5S RNA is growth rate-dependent [85]. Whereby at higher growth rates, the 4.5S RNA level was increased in comparison to slower growth rates. Therefore, it might be assumed that the degradation of overproduced Ffh in early growth phases is even more important to prevent the accumulation of free Ffh in the cell. Since free Ffh alone would not be able to carry out its cellular function, probably the cell is attempting to maintain an excess of 4.5S RNA in the cell. It was shown that the M domain can occlude the binding site of the SRP receptor FtsY when Ffh was not bound to the 4.5S RNA and this effect can be reversed by binding the 4.5S RNA [53, 54, 137, 138, 223, 317]. Thus the Lon protease seems to play a critical role in the quality control of the SRP complex by degrading free, non-functional Ffh maintaining excess 4.5S RNA in the cell. Since 4.5S RNA affects the stability of Ffh by protecting overproduced Ffh against degradation by simultaneous overproduction of 4.5S RNA [168], we investigated the effect of an altered RNA-binding ability on Ffh proteolysis. Therefore, we used a temperature-sensitive ffh mutant in which the chromosomal ffh was disrupted by ffh1::kan and complemented with a constitutive medium-copy plasmid encoding the FfhL338P M347T variant to ensure strain viability [271]. These two point mutations (L338P and M347T) are located within the disordered region (between helix 1 and helix 2) of the M domain and result in decreased RNA-binding ability. Crystal structure analysis determined amino acid residues 338 to 370 as a disordered region in the presence of the 4.5S RNA [20]. This region contains the proposed signal peptide recognition site, which is subjected to flexible conformational changes in the absence of a signal peptide [406]. However, binding of the 4.5S RNA to the FfhL338P M347T variant is possible to a limited degree. Park et al. provided the first evidence on the stability of the FfhL338P M347T variant, which was degraded at 30 °C in the early exponential growth phase with a half-life of 10 min. Our results have shown that the FfhL338P M347T variant exhibits a growth phase-dependent degradation pattern distinct from overproduced Ffh (Fig. C-1 C). Rapid degradation was observed from early to late exponential growth phase. In early stationary phase, when overproduced Ffh was rapidly degraded, the FfhL338P M347T variant became more stable. The amount of the FfhL338P M347T variant in early exponential growth phase was comparable to the cellular Ffh in E. coli WT. In this case, the growth phase-dependent degradation of the FfhL338P M347T variant appears to be independent of the Ffh amount. FfhL338P M347T is also degraded in a growth phase-dependent manner but the timing of rapid degradation slightly differs the overproduced Ffh. Since the protein is still degraded in a regulated manner, its integrity appears

D Discussion to be preserved and the different degradation pattern might be attributed to the limited RNA- binding capacity. In summary, the importance of the different Ffh variants are shown in the following figure (Fig. D-1).

Fig. D-1: Cellular Ffh amount is modulated by the cytosolic Lon protease. Cellular Ffh, recombinant L338P M347T His6-Ffh and Ffh can be degraded in E. coli until a certain cellular concentration is reached. Two different pools of Ffh exist in the cell including free Ffh (not bound to the 4.5S RNA) and Ffh in complex with the 4.5S RNA mediating the co-translational protein transport. In each case free Ffh is degraded by the cytosolic Lon protease and in contrast Ffh bound to the 4.5S RNA is maintained to ensure the cellular function of the SRP complex. Thus, the Lon protease appears to be involved in the quality control of the SRP complex in which free non-functional Ffh is degraded.

L338P M347T All variants have in common that cellular Ffh, recombinant His6-Ffh and Ffh can be degraded in E. coli until a certain cellular concentration is reached. Two different pools of Ffh probably exist in the cell including on the one hand free Ffh (not bound to the 4.5S RNA) and on the other hand Ffh in complex with the 4.5S RNA representing the functional SRP complex. In all investigated scenarios we can assume that free Ffh is degraded by Lon and in contrast, Ffh bound to the 4.5S RNA is not degraded to ensure the cellular function of the SRP complex. Thus, Lon protease appears to be involved in the quality control of the SRP complex in which free non-functional Ffh is degraded. However, this process seems to be subject to certain regulation by the Lon protease as Ffh proteolysis is stopped to maintain a certain Ffh level in the cell to sustain the co-translational protein targeting pathway.

D Discussion

Due to the excess of 4.5S RNA, it is assumed that the 4.5S RNA has a dual function in the cell. The 4.5S RNA forms a stable complex with Ffh as well as the elongation factor G (encoded by fusA) [340]. Thereby the 4.5S RNA binds the elongation factor G (FusA) independently of Ffh and is proposed to modulate the interaction of FusA with the ribosome [51, 156, 248, 296, 340]. FusA catalyzes ribosome translocation by one codon along the mRNA during protein biosynthesis whereby GTP hydrolysis is required [389]. Together with our findings, that the 4.5S RNA is present in excess and that the Ffh amount is growth phase-dependently reduced to a certain level, raises the possibility of an opposite degradation pattern of FusA in comparison to Ffh. Thus, we suspected if Ffh was not degraded anymore, FusA might be degraded. With these experiments, we wanted to investigate whether 4.5S RNA-binding to either Ffh or FusA at different growth phases influences the stability of both proteins. However, in vivo degradation experiments showed that FusA is a stable protein over the entire bacterial growth (Fig. H-10). Thus, Ffh proteolysis appears to be independent of the dual role of 4.5S RNA. A further interesting factor for the modulation of the growth phase-dependent Ffh degradation represents the alternative sigma factor σ32 since a direct interaction of both proteins was shown [213, 237]. For that reason, we investigated the influence of RpoH on the degradation of Ffh.

1.2 Ffh proteolysis is affected by the sigma factor RpoH The heat shock sigma factor σ32 (encoded by rpoH) interacts with the RNA polymerase due to rising temperature, thus initiating the transcription of genes (heat shock genes) which are under the control of a σ32 promoter. The cellular RpoH level is strictly controlled at the transcriptional, translational and posttranslational levels and during normal growth conditions, the cellular amount of RpoH is very low. Transcription of rpoH is regulated by two constitutive σ70 promotors and one σ24 promotor as well as it is independent of σ32 [94, 95, 106, 347, 377]. During normal growth conditions, the Shine-Dalgarno sequence of rpoH-mRNA is present in a secondary structure and therefore inaccessible for ribosomes. However, when the ambient temperature rises, this secondary structure melts and translation starts [173, 242, 243, 247, 401]. On post-translational level, RpoH activity is controlled by the chaperone systems DnaK/DnaJ/GrpE and GroEL/ES. RpoH binding to chaperone systems depends on the cellular concentration of misfolded proteins [109, 110, 130, 211, 344, 350]. Thus, RpoH binds to the chaperone systems only at low concentrations of aggregated proteins. RpoH stability + (T½ = ~ 1 min) is mainly controlled by FtsH-dependent proteolysis however, other AAA proteases like Lon, HslUV and ClpXP play a minor role in RpoH degradation [147, 174, 349]. The sigma factor is recognized by its homeostatic control region 2.1 and point mutations within 78

D Discussion this region result in stabilization of the protein [265]. It is suggested that the complex of DnaK/DnaJ chaperone system makes RpoH accessible for FtsH-dependent proteolysis implying a chaperone-mediated RpoH targeting to the membrane [41]. Current studies show that RpoH can be directed to FtsH via alternative targeting pathways. Recently, it was discovered that RpoH was degraded in E. coli by a post-translational modification of the ubiquitin-like protein ThiS [395]. In eukaryotes, the ubiquitin-mediated protein degradation mechanism by the 26S proteasome is a common mechanism for the removal of aggregated protein (reviewed in [18]). Thereby a ubiquitin tag is attached to damaged proteins, which severs a degradation signal for the 26S proteasome. Similar to this system, RpoH is covalently bound to the ubiquitin-like protein ThiS, which leads this complex to ClpXP-dependent proteolysis [395]. Moreover, it was shown that the SRP complex directs RpoH it to the inner membrane for FtsH-dependent proteolysis [213]. RpoH is delivered to the FtsH protease by the interaction of the SRP complex with its receptor FtsY. In vivo cross-linking studies revealed a direct interaction between the protein component of the SRP complex Ffh and RpoH. The sigma factor RpoH interacts via its homoeostatic control region 2.1 with the signal-peptide binding site in the M domain of Ffh. Amino acid residues of RpoH involved in the Ffh-RpoH interaction are L41, E48, K51, T52 and I54 [237]. Amino acid residue I54 is also critical for RpoH recognition by FtsH, whereby additional amino acid residues are involved in recognition [265]. The homeostatic control region is capable to form an amphipathic helix, which might provide a hydrophobic surface for the Ffh-RpoH interaction [237]. With this discovery, it could be shown for the first time that the SRP complex in E. coli can target proteins post-translationally to the membrane. At the beginning of this section, it was mentioned that Ffh was found via an in vivo trapping approach of the FtsH protease. This can probably be explained by the described interaction between Ffh and RpoH. It can be speculated, that the RpoH-SRP-complex might have interacted with FtsH thus Ffh was trapped concomitantly to RpoH. This scenario is plausible since approx. 50 % of cellular RpoH is associated with the inner membrane [213]. In order to investigate a possible influence of RpoH on the stability of Ffh, we did in vivo degradation experiments at 25 °C in E. coli WT and ΔrpoH. Our experiments demonstrated that overproduced Ffh in WT at 25 °C was degraded over the entire bacterial growth. In contrast, Ffh was stabilized in the rpoH deletion mutant in each analyzed growth phase. Additionally, cellular Ffh is degraded more rapidly in the early exponential phase in the WT than in the ΔrpoH mutant. In both cases, Ffh stability depends probably on the available Lon amount in the cell. In the presence of RpoH, the cellular Lon amount was increased in late exponential and early stationary growth phases, whereas it was almost constant in a ΔrpoH mutant. That is

D Discussion presumably the result of an increased lon gene expression depending on the sigma factor RpoH. The rapid proteolysis of Ffh was accompanied by an increased amount of Lon, which can be explained by the induced heat shock response in later growth phases. Lon is part of the heat shock regulon and its cellular concentration is increased on transcript and protein level upon shift to heat stress [141, 277]. Transcriptome analysis revealed significantly increased lon transcript levels after induction of 15 min heat stress while the ffh transcript remained constant [141]. Contrary, another transcriptome analysis measured 3-fold increased ffh transcript levels after heat shock induction [257]. Thus, Ffh and Lon are components of the heat shock regulon controlled by the sigma factor RpoH. At high temperatures or in the transition to the stationary growth phase, aggregated proteins accumulate in the cell and the amount of RpoH molecules is increased to induce gene expression of heat shock genes including AAA+ proteases and chaperones [371]. Under heat shock conditions, AAA+ proteases prevent the toxic accumulation of aggregated proteins by efficiently degrading these proteins and thus ensuring cell integrity. To study the effect of the heat shock response on Ffh proteolysis, we analyzed the stability of cellular Ffh after the induction of a heat shock response, which was generated by a temperature shift from 30 °C to 42 °C. We investigated cellular Ffh since we suspected a more pronounced difference when heat shock conditions affected the proteolysis of Ffh. Our results demonstrated that cellular Ffh was stable after heat shock induction. Therefore, it can be assumed that an intracellular basal level of Ffh is present to maintain co-translational protein transport under heat stress conditions. Depletion of Ffh induced a heat shock response in E. coli preventing a toxic accumulation of aggregated and mislocalized proteins [30, 271]. In a recent study, it was shown that the intracellular amount of aggregated proteins in a Ffh depletion strain was relatively low (only 1% of the proteome) [387]. They suggest that mislocalized proteins in the cytosol are efficiently degraded by e.g. the AAA+ proteases Lon and HslUV. These two proteases become essential after Ffh depletion because they potentially prevent the toxic effect by degrading aggregated proteins [30]. Moreover, the heat shock-induced chaperones GroEL and DnaK are enriched in Ffh-depleted cells at the inner membrane [387]. The authors speculated that both chaperones could be bypass the limited co-translational protein targeting in Ffh-depleted cells. Since GroEL and DnaK interact with auxiliary protein of the SecYEG translocation component SecA [42, 62].

D Discussion

Fig. D-2: Modulation of Ffh stability in heat shock response. Under normal growth conditions (no heat stress) the concentration of the sigma factor RpoH is maintained low by FtsH-dependent proteolysis. One possibility to target cytosolic RpoH to the membrane-anchored FtsH protease occurs via the SRP complex. The main function of the SRP complex is the co-translational protein transport of inner membrane proteins. Within the quality control of the SRP complex free Ffh not bound to 4.5S RNA is degraded by the AAA+ protease Lon. Under heat shock conditions, FtsH no longer degrades RpoH, it accumulates in the cell and induces gene expression of heat shock genes such as chaperones and AAA+ proteases to prevent toxic accumulation of aggregated proteins. In contrast, Ffh concentration decreases under heat stress and Ffh is no longer degraded by Lon to ensure co-translational protein transport.

In conclusion, our results indicate that the increased stability of Ffh in the absence of RpoH represents a correlation between RpoH-Ffh interaction and Ffh stability (Fig. D-2). Under non- heat shock conditions (e.g. early growth phases), RpoH is directed via the SRP complex to FtsH-mediated degradation. Lon can also degrade Ffh under these conditions. However, as described above, faster Ffh degradation is observed in the transition from the exponential to the stationary phase. In contrast, a significant stabilization of Ffh was observed in the later stationary phase. In addition, RpoH could affect growth phase-dependent Ffh degradation. The increased Ffh stability might be due to the availability and activity of RpoH. In the stationary phase or after heat shock induction, aggregated proteins accumulate and the RpoH amount increases leading to a heat shock response. Since RpoH is no longer degraded by FtsH in this 81

D Discussion growth phase, it is not translocated to the membrane via the SRP complex. However, the increased expression of Lon in this phase does not simultaneously lead to increased Ffh degradation. Rather Lon is primarily involved in the degradation of aggregated proteins to prevent a toxic effect. Furthermore, it can be assumed that Ffh is retained in the cell under heat shock conditions to maintain co-translational protein transport in the cell and thus ensure membrane protein integrity. Thus, the growth phase-dependent Ffh proteolysis by the cytosolic Lon protease seems to be modulated by the available Ffh amount in the cell and by the sigma factor RpoH.

2. Recognition mechanism of Ffh by the Lon protease Lon is not essential in E. coli and is involved in the quality control of proteins and the degradation of regulatory proteins controlling cellular processes. Hydrophobic or aromatic amino acids are often relevant for the recognition of substrates by Lon [135, 227, 366]. Lon recognizes substrates by various recognition signals including unstructured accessible N-, C- terminal or internal protein regions. Moreover, structured regions within the C- or N-terminus of a protein serve as recognition sequence. For instance, the cell division inhibitor SulA is recognized by a C-terminal and the DNA polymerase protein V UmuD by a N-terminal degradation signal [119, 158]. In addition, adaptor-mediated and ligand-controlled recognition is a known mechanism for the Lon protease. A detailed overview about recognition principles of the Lon protease is given in section A-2.2.

2.1 Lon protease recognizes Ffh via its M domain This work should give a more profound insight into the recognition of Ffh by the Lon protease. A previous master thesis had shown that Lon recognizes Ffh within its M domain [23]. Therefore, the stability of the functional Ffh domains, NG and M domain was separately analyzed by in vivo degradation experiments. The N-terminal N domain and the central G domain represent a functional unit, which mediates GTP hydrolysis [197, 310]. Interaction between Ffh and its receptor FtsY occurs via their structurally similar NG domains [93, 102]. The C-terminal methionine-rich M domain of Ffh binds the 4.5S RNA and signal-anchored sequences of membrane proteins at translating ribosomes [180, 221, 299, 413]. Both functional domains, NG and M domain, are connected via a flexible linker. In this work, in vivo degradation experiments confirmed that the NG domain was stable during the entire bacterial growth curve (Fig. C-6 B). In contrast, the protein stability of the M-domain was lower at any time point of bacterial growth. The M domain was most rapidly degraded in late stationary 82

D Discussion phase and thus the growth phase-dependent degradation pattern of full-length Ffh was not observed (Fig. C-6 C). We conclude that the Lon protease recognizes Ffh by its RNA-binding M domain but not via the NG domain. We were unable to verify in vivo whether the degradation of the M domain is directly linked to Lon-dependent proteolysis because plasmid-encoded M domain induced a synthetic lethal phenotype after transformation in a lon-deficient strain [23]. This specific phenotype was demonstrated by determining the transformation efficiencies of plasmid-encoded Ffh, NG domain and M domain (Fig. C-7). The respective genes/gene regions were examined as His6-tag (encoding an N-terminal His6-tag) or One hybrid construct (flanked by T25 and T18 domain of the adenylate cyclase of B. pertussis). It was possible to transform MC4100 WT and Δlon with plasmid-encoded Ffh or the NG domain, while only the WT could be transformed with plasmid-encoded M domain. The deleterious phenotype could be avoided by using the One-hybrid constructs for Ffh, the NG and M domain. Both strains, WT and Δlon, could be transformed with all of these constructs. In addition, plasmid-encoded cpSRP54 from A. thaliana and its M domain did not cause a lethal phenotype in the absence of Lon. The synthetic lethal phenotype might be triggered by a basal expression of the recombinant

His6-tagged M domain. The plasmid-encoded M domain is controlled by an AHT-inducible promoter and should usually not allow expression of the recombinant M domain. During in vivo degradation experiments, it was shown that a certain amount of the recombinant protein was already present in the cell before protein expression was induced (data not shown). The leaky expression system could be the reason for a low expression in the absence of inductor (AHT), which was probably sufficient to induce a toxic effect in a lon-deficient strain. Various observations in the literature support the M domain-induced phenotype. It is described that a depletion of the SRP complex, which does not influence bacterial growth, increases the stability of mislocalized inner membrane proteins (IMPs) in a ΔlonΔhslUV strain [30]. As a consequence, mislocalized IMPs accumulate in the cytosol and lead to a lethal phenotype. Under normal growth conditions, Lon and HslUV rapidly degrade mislocalized IMPs preventing the accumulation of these proteins. In vitro studies showed that the M domain alone can bind 4.5S RNA as well as signal peptides. Recombinant M domain binds the 4.5S RNA with the same affinity as full-length Ffh [73]. For the mammalian SRP54-M domain it was shown that it binds signal sequences with weaker affinity than full-length cpSRP54 [414]. Yosef et al. revealed that an overexpression of the SRP complex, Ffh or M domain leads to selective inhibition of membrane proteins [398]. Since the effect of the M domain is the same as for the SRP complex or full-length Ffh, indicates that the M domain can no longer bind its receptor FtsY. Based on this information, it can be speculated for our experiments on the transformation

D Discussion efficiencies that the recombinant M domain binds signal sequences of translating IMPs stalling the nascent ribosome. Due to the absence of the NG domain, a subsequent interaction with the membrane-associated SRP receptor FtsY does not occur and translocation of the membrane protein cannot proceed. This leads to the accumulation of mislocalized IMPs. In a WT strain, the M domain could be recognized by Lon which would rescue ribosomes to maintain viability. in a lon-deficient strain Lon protease is not available to release the stalled ribosome resulting in an accumulation of stalled ribosomes and cytosolic IMPs, which is ultimately toxic to the cell. This scenario is conceivable because ribosome stalling by the M domain could be prevented when the protein is flanked by the T25 and T18 domains at the N- and C-terminus, respectively. It can be assumed that the signal peptide-binding site and the RNA-binding motif are not accessible for binding nascent ribosomes and an interaction did not occur. The constant degradation of the M domain over the entire growth curve in E. coli WT could be explained by a toxic effect of the M domain. In later growth phases, when the M domain is degraded most rapidly, it is even more important to prevent toxic accumulation of aggregated proteins in the cell. Furthermore, plasmid-encoded M domain also leads to an altered phenotype in E. coli WT since we demonstrated elongated cells after transformation. Therefore, it can be speculated that this phenotype is caused by a disturbed cell division resulting from inefficient targeting of SRP substrates. A possible candidate could be the SRP substrate FtsQ, which is required for septum formation during cell division. It was shown that FtsQ-depleted cells also exhibit elongated cells caused by cell division defects (reviewed in [306]). The phenotype induced by the M domain can be affected by many other SRP substrates. However, Lon seems to be essential for the viability of the cell during overproduction of the M domain. To get insights into the recognition mechanism in more detail, point mutations within the RNA- binding motif were inserted assuming that the M domain is recognized by the Lon protease and that the 4.5S RNA increases the protein stability of Ffh.

2.2 Ffh recognition by Lon requires an intact 4.5S RNA-binding motif The SRP complex is conserved in all three domains of life and its complexity has increased during evolution. The prokaryotic SRP complex is the simplest variant, consisting of the protein component Ffh and an SRP RNA, e.g. 4.5S RNA of E. coli [31, 280, 292, 298]. The 4.5S RNA is recognized by the helix-turn-helix motif consisting of four α-helices (helix 2, 2b, 3 and 4) within the M domain of Ffh [20]. The helix-turn-helix-motif facilitates contact between the symmetrical and asymmetrical loop of the 4.5S RNA [21]. The determination of the minimal 4.5S RNA-binding domain of the M domain was performed by limited proteolysis 84

D Discussion

(proteolytical digestion with trypsin and V8 protease) in the presence of the 4.5S RNA followed by MALDI-TOF mass spectrometry [21]. The analysis revealed a core RNA-binding domain consisting of amino acid residues 328 to 432. This fragment was able to bind the 4.5S RNA with the same affinity as full-length Ffh. Surprisingly, our results revealed that deletion of amino acid residues 388 to 453 restore the synthetic lethal phenotype of plasmid-encoded M domain in a lon deletion mutant. This provides a more clear evidence of the importance of the RNA binding motif for Lon recognition. Interestingly, chloroplasts of higher plants (e.g. A. thaliana) contain the highly conserved cpSRP54 and the additional protein cpSRP43 [104, 186, 323]. In chloroplasts the SRP-RNA is lost and cpSRP54 contains a mutated variation of the conserved RNA-binding motif within its M domain [293, 301]. The conserved SRP RNA- binding motifs SM and GXG of E. coli are replaced by VM and DXG in higher plants (e.g. A. thaliana), which leads to the loss of RNA-binding [293, 301]. RNA-binding ability is restored in A. thaliana by homologous mutations (V455S and D480G) towards the E. coli sequence [293]. As mentioned above, 4.5S RNA is involved in Lon-dependent proteolysis of Ffh by protecting the SRP complex against degradation. We investigated whether the stability of Ffh could be altered by evolutionary point mutations (S382V, G405D, C406S) within the RNA-binding motif leading to a loss of RNA-binding in A. thaliana or whether Ffh degradation is a conserved mechanism. In vivo degradation experiments in E. coli WT demonstrated that Ffh variants containing point mutation S382V or C406S were degraded growth-phase- dependently comparable to wild-type Ffh (Fig. H-9) [101]. In contrast, Ffh variants containing point mutations G405D [101], G405A or G405N were stable over the entire bacterial growth. However, it should be noted that Ffh variants carrying a point mutation at position G405 were insoluble. Since the described amino acid residues are involved in an interaction with the 4.5S RNA, the degradation of the Ffh variants might be associated with their function in the RNA- protein interaction. In particular, the altered degradation of the FfhG405D variant and its insolubility might be caused by its impaired RNA-binding ability. In the following, the function of the amino acids S382, G405 and C406 in RNA-protein interaction will be described in more detail. All three amino acid residues are located within the minimal core RNA-binding domain (aa 328 to 432) and are part of the intermolecular contact between Ffh and 4.5S RNA [21]. Direct RNA-protein interaction involves the symmetric and asymmetric loop of 4.5S RNA as well as amino acid residues (A378, N381, S382, M383, K386, S397, R398, R401, G405 and C406) of the M domain located within the minimal core RNA-binding domain. The symmetric loop consists of five non-canonical base pairs (U45-G64, C46-A63, A47-C62, G45-G61, G49- A50) and all nucleotides are involved in the RNA-protein interaction. Amino acid residues

D Discussion

S382, G405 and C406 are located in helix 2b and 3 of the helix-turn-helix motif (Fig. C-8), which interact with nucleotides of the symmetric loop at the minor groove. A hydrogen bond network represents the central feature of the 4.5S RNA-Ffh interaction. Hydrogen bonds are generated between the non-canonical A47-C62 nucleotide pair, A39 of the asymmetric loop and a conserved salt bridge between E387 and R401. The amino acid residues S382 and G405 are involved in the interaction with the nucleotide pair A47-C62. The interaction with C62 is essential as the mutation C62G leads to a lethal phenotype [391]. Moreover, the RNA-binding ability is limited by removal of functional groups of the nucleotide C62. Furthermore, the non- canonical G48-G61 pair belongs to the principle recognition elements. The RNA-protein interaction requires a hydrogen bond network and the binding of two potassium ions. One potassium ion is coordinated by the C46-A63 and U45-G64 base pairs of the 4.5S RNA [20]. The second potassium ion involves the G48-G61 pair and the amino acid residue G405 in its coordination. The conservation of this ion-binding pocket demonstrates that the potassium ion is a key element in the RNA-protein interaction. In addition, the potassium ions appear to stabilize the interaction between Ffh and 4.5S RNA. Thus, it was shown that all three amino acids described have a strong impact on the binding of 4.5S RNA. There is no information in the literature whether a mutation at position G405 affects RNA-binding ability. However, it might be assumed that due to the drastic mutation from a polar glycine to an acidic aspartic acid the coordination of the potassium ion does not occur anymore and thus there is no interaction with the 4.5S RNA. Based on our previously described results, it might be expect that the FfhG405D variant is degraded more rapidly because it is no longer bind the 4.5S RNA. In contrast, the FfhG405D variant was more stable. The tertiary structure of the FfhG405D variant may be drastically altered leading to its insolubility. Thus, aggregated FfhG405D accumulates in the cell while Lon is unable to recognize this variant. This hypothesis can be supported by the fact that the growth of E. coli was markedly inhibited when carrying plasmid-encoded FfhG405D (data not shown). The Ffh variants containing a S382V or C406S point mutation can bind the 4.5S RNA. It has been shown that the point mutation C406S does not affect 4.5S RNA-binding since the M domainC406S retained full RNA-binding ability [21]. An exchange of S382T is suggested to impair RNA-binding ability resulting in fewer Ffh-4.5S RNA complexes [346]. An increased 4.5S RNA concentration promotes an increased Ffh-4.5S RNA complex formation indicating that FfhS382T has a functional SRP receptor interaction [346]. We have shown that the point mutations S382V and C406S, which are unlikely to affect RNA-binding ability, does not affect Ffh recognition by Lon. In combination with the fact that (A) the M domain is an unstable protein, (B) plasmid-derived M domain leads to a synthetic lethal

D Discussion phenotype in a lon-deficient E. coli strain, which can be reversed by a partially deleted RNA- binding motif, suggest an intact RNA-binding motif for Ffh recognition. To investigate whether this hypothesis is valid, the stability of cpSRP54 from A. thaliana in E. coli was determined. We showed that cpSRP54 is degraded slowly in E. coli, and the Lon protease is responsible. Therefore, recognition of cpSRP54 by Lon seems to be impeded. In contrast, the cpSRP54V455S D480G variant showed a slightly accelerated degradation. The point mutations V455S and D480G (restoring the E. coli sequence) in cpSRP54 can restore SRP RNA-binding. Thus, an intact RNA-binding motif is necessary for the recognition of Ffh by Lon. With these results, it was confirmed that an intact RNA-binding motif is important for efficient recognition of Ffh by the Lon protease. Since the wild-type cpSRP54 could also be recognized, the degradation of the protein component of the SRP complex seems to be a partially conserved mechanism. However, the recognition motif in A. thaliana appears to be degenerated. Possibly a degradation of cpSRP54 by Lon can also occurs in chloroplasts, since both components are exist. There is little knowledge about Lon proteases and their substrates in chloroplasts. A. thaliana contains four Lon protease (Lon1, Lon2, Lon3, Lon4) whereby Lon1 and Lon4 belonging to the LonA subfamily and localized in chloroplasts and mitochondria [77, 267, 295]. Both Lon proteases are highly conserved and have a structurally similar ATPase domain, where the SSD-domain (sensor- and substrate-discrimination domain) can vary, suggesting that different substrates can be targeted for ATP-dependent proteolysis [294]. A degradation of the Ffh homologue cpSRP54 by Lon in A. thaliana might be conceivable in chloroplasts, since both components are available.

3. ATP-dependent proteolysis in protein translocation ATP-dependent proteolysis plays a crucial role in the transport of proteins into the inner and outer membrane (Fig. D-3). A key target for AAA+ proteases is the Sec translocon and its ancillary proteins. The core Sec translocon is formed by the membrane-spanning proteins SecYEG and mediates the insertion of inner membrane proteins and the transport of proteins across the inner membrane. Protein targeting to the Sec translocon occurs post- or co- translationally. The post-translational protein targeting pathway requires the assistance of chaperons (e.g SecB) delivering fully folded proteins to the SecYEG translocon. In the co- translational protein targeting pathway, nascent inner membrane proteins are recognized at the translating ribosome by the SRP complex directing them to SecYEG via the membrane-bound SRP receptor FtsY. More detailed information on post- and co-translational protein targeting are given in section A-3 of this work. 87

D Discussion

Fig. D-3: ATP-dependent proteolysis in protein translocation. The Sec translocon consists of the core proteins SecYEG and the ancillary proteins SecD, SecF, YajC and YidC mediating protein translocation in or across the inner membrane. As part of protein quality control, the membrane proteins SecY, SecE and SecD (highlighted in bold) are subjected to FtsH-dependent proteolysis. We have found that the Lon protease controls the co-translational protein targeting pathway by degrading Ffh, the protein component of the signal recognition particle (SRP complex).

Post-translational protein translocation across the inner membrane by SecYEG is associated with the cytoplasmic ATPase SecA, which forms the translocase in this pathway [52]. The ancillary proteins SecD, SecF and YajC are involved in the secretion process of proteins [91]. The insertase YidC assists SecYEG during protein insertion of co-translationally targeted inner membrane proteins. Some of these proteins are degraded by AAA+ proteases in an ATP- dependent manner. Two proteins of the core Sec translocon, SecY and SecE, are subjected to FtsH-mediated proteolysis [182, 367]. SecY is degraded when it is unassembled to SecE translocon since overproduction of SecY in a ftsH mutant caused a deleterious effect that impeded cell growth and protein export [182]. Recently the accessory proteins SecD was identified as a further FtsH substrate, while the insertase YidC was stable [10]. It is assumed that a complex of SecD/SecF/YajC/YidC stabilizes the SecYEG translocon [261]. These substrates are subjected to proteolysis by the essential membrane-anchored FtsH protease, which emphasizes that the Sec translocon as part of its quality control is strictly controlled by ATP-dependent proteolysis. Until now, these processes seemed to be exclusively controlled by FtsH. With this work, we provide new insight into the impact of the AAA+ protease Lon in protein targeting of inner membrane proteins. Since the cytosolic Lon protease is responsible for the degradation of Ffh, the protein components of the essential SRP complex. 88

D Discussion

In this work, it was shown that the protein component, Ffh of the essential SRP complex is predominantly degraded by the cytosolic AAA+ protease Lon depending on the growth phase. Furthermore, the proteases ClpXP, ClpAP and HslUV are involved in the degradation of Ffh but play a minor role. In addition, it was demonstrated that the growth phase-dependent degradation mechanism is modulated by the amount of Ffh available in the cell and by the heat shock sigma factor RpoH. For the recognition of Ffh, it is necessary that Ffh is not bound to the 4.5S RNA in the cell and that it has an intact RNA-binding motif. Furthermore, this degradation mechanism seems to be a partially conserved degradation mechanism, since Lon from E. coli can in principle recognize cpSRP54 from A. thaliana.

Summary

E Summary

E Summary Membrane protein targeting is an essential process for maintaining cellular homeostasis and membrane integrity. One mechanism that allows efficient protein targeting to the inner membrane is the co-translational pathway mediated by the essential signal recognition particle (SRP complex). In bacteria, the highly conserved SRP complex represents a ribonucleoprotein consisting of the protein component Ffh (fifty-four homologue) and a regulatory SRP-RNA (4.5S RNA in E. coli). Ffh can be divided into two functional domains, whereby the M domain binds the 4.5S RNA as well as signal peptides of the nascent membrane proteins and the NG domain mediates a GTPase activity, which has a particular role in the interaction of the SRP complex and its membrane-associated receptor FtsY. The GTP-dependent interaction of the SRP complex and FtsY enables the release of nascent membrane proteins to the SecYEG translocation system. In this work, it was verified that in E. coli the co-translational protein transport on the post- translational level is controlled via ATP-dependent proteolysis. This mechanism is ensured by degrading the protein component of the SRP complex by cytosolic AAA+ proteases, whereas Lon is the major protease for Ffh proteolysis. Since Ffh was more stable in a ΔlonΔhslUVΔclpP deletion mutant, it can be assumed that other AAA+ proteases such as ClpXP, ClpAP and HslUV are involved in Ffh degradation, however they plays a minor role. Ffh is degraded especially during the transition phase from the exponential to the stationary growth phase. The modulation of this growth phase-dependent degradation occurs in dependence of the available Ffh amount in the cell and is possibly coupled to the heat shock response. Ffh was degraded under non-heat shock conditions and stabilized during the heat shock response. Both scenarios provide evidence for a degradation mechanism, whereby excess Ffh that does not bind to the 4.5S RNA is subjected to proteolysis until a minimal cellular Ffh concentration was reached. Thus, the stabilization of Ffh at low concentrations seems to be important in order to ensure co- translational protein transport and membrane integrity in critical growth phases. In addition to the investigation of growth phase-dependent degradation, the recognition mechanism of Ffh by Lon protease was studied. This work confirms that Lon recognizes the RNA-binding M domain of Ffh since recombinant M domain was degraded in vivo, whereas the NG domain was stable. Plasmid-encoded M domain induces a synthetic lethal phenotype in a lon-deficient strain, which could be restored by a RNA binding motif inaccessible to Lon. In vivo degradation experiments of chloroplast SRP54 (cpSRP54) from A. thaliana demonstrated that Lon from E. coli slightly recognizes cpSRP54 , and point mutations within

E Summary the degenerated RNA binding motif of cpSRP54 resulted in an accelerated proteolysis. Thus, the investigated degradation mechanism of Ffh seems to be conserved and the recognition of Ffh by Lon occurs via an intact and for Lon accessible RNA-binding motif.

Zusammenfassung

F Zusammenfassung

F Zusammenfassung Der Transport von Membranproteinen ist ein essentieller Prozess zur Aufrechterhaltung der zellulären Proteinhomöostase sowie Membranintegrität. Ein Mechanismus für den effizienten Transport von Proteinen zu der inneren Membran stellt der co-translationale Proteintransport dar, der durch den Signalerkennungspartikel (SRP-Komplex) vermittelt wird. Der hoch konservierte SRP-Komplex ist ein Ribonukleoprotein, der in Bakterien aus der Proteinkomponente Ffh (fifty-four homologue) und einer regulatorischen SRP-RNA (4.5S RNA in E. coli) besteht. Ffh kann in zwei funktionelle Domänen unterteilt werden, wobei die M- Domäne für die Bindung der 4.5S RNA als auch für die Bindung von Signalpeptiden der naszierenden Membranproteinen verantwortlich ist und die NG-Domäne eine GTPase-Aktivität vermittelt, die für eine Interaktion des SRP-Komplexes mit seinem membran-assoziierten Rezeptor FtsY von Bedeutung ist. Die GTP-abhängige Interaktion von SRP und FtsY ermöglicht die Freisetzung von naszierenden Membranproteinen an das SecYEG- Translokationssystem. In der vorliegenden Arbeit wurde bestätigt, dass der co-translationale Proteintransport auf post- translationaler Ebene durch ATP-abhängige Proteolyse kontrolliert wird, in dem die Proteinkomponente des SRP-Komplexes durch zytosolische AAA+-Proteasen abgebaut wird. Ffh wird vorrangig durch die Lon-Protease abgebaut. Da Ffh in einer ΔlonΔhslUVΔclpP Deletionsmutante am stabilsten war, lässt vermuten, dass weitere AAA+-Proteasen wie ClpXP, ClpAP und HslUV an dem Ffh-Abbau beteiligt sind, diese aber eher eine untergeordnete Rolle spielen. Ffh wird insbesondere in der Übergangsphase von der exponentiellen zur stationären Wachstumsphase abgebaut. Die Modulation dieses wuchsphasenabhängigen Abbaus erfolgt über die zellulär verfügbare Ffh-Menge und ist möglicherweise an eine Hitzeschockantwort gekoppelt. Unter normalen Wachstumsbedingungen wurde Ffh abgebaut und im Rahmen der Hitzeschockantwort stabilisiert. Beide Szenarien liefern Hinweise für einen Abbau von überschüssigem Ffh bis zu einer minimalen Ffh-Konzentration, wobei vermutlich freies Ffh abgebaut wird, welches nicht an die 4.5S RNA gebunden vorliegt. Somit scheint eine Stabilisierung von Ffh bei einer geringen Konzentration umso wichtiger zu sein, damit der co- translationalen Proteintransport und die damit einhergehende Membranintegrität unter kritischen Wachstumsbedingungen gewährleistet werden kann. Neben der Untersuchung des wachstumsphasenabhängigen Abbaus wurde der Erkennungsmechanismus von Ffh durch die Lon-Protease untersucht. In der vorliegenden Arbeit wurde bestätigt, dass Lon die RNA-bindende M-Domäne von Ffh erkennt, da in vivo

F Zusammenfassung rekombinante M-Domäne abgebaut wurde, wohingegen die NG-Domäne stabil war. Plasmid- kodierte M-Domäne führte in einem lon-defizienten Stamm zu einem synthetisch letalen Phänotyp, welcher durch ein für Lon unzugängliches RNA-Bindemotiv aufgehoben wurde. Mit in vivo Degradationsexperimente des chloroplastidären SRP54 aus A. thaliana wurde gezeigt, dass cpSRP54 durch Lon in E. coli degradiert werden konnte, wobei der Abbau langsamer war. Dahingegen war der Abbau für cpSRP54, welches Punktmutationen innerhalb des degenerierten RNA-Bindungsmotivs kodierte, beschleunigt. Somit scheint der Abbaumechanismus von Ffh konserviert zu sein und die Erkennung von Ffh erfolgt über ein intaktes und für Lon zugängliches RNA-bindendes Motiv.

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127

Appendixes

128

H Appendixes

Tab. H-1: List of references of Lon substrates in Fig. A-3. Lon substrate Reference

quality control denatured proteins [135] SsrA-tagged proteins [66] proOmpC/proOmpF [308] apoTorA [288]

stress response LexA [251] SulA [157, 158] UmuC/UmuD [119] RecA [251] RuvB [252] SymE [178] SoxS [126, 325, 326] PerR [4] MarA [33, 126] GadE [148] IbpA/IbpB [37] ribosomal proteins [196]

virulence RovA [145] YomA [160] HilC/HilD [342] HrpR/HrpG [49, 407]

biosynthesis pathways HTS [34] MetR [9] CysB [9] HemA [376] RcsA [352]

phage development λN [121] λXis [205]

cell-cycle, toxin-antitoxin systems, persister cell formation CspD [200] DnaA [215] TrfA [192] CcrM [392] SciP [120] CbpM [65] CcdA/CcdB [365, 366] RelE/RelB [68] RnlA/RnlB [188] YefM/YoeB [67] MazE/MazF [69] HipA/HipB [139] DinJ/YafQ [284] MqsR/MqsA [184, 378]

motility and biofilm formation FliA [19] FhlDC [71] 129

H Appendixes

Lon substrate Reference

FlgE [9] SwrA [244] metal homeostasis ZntR [283] CueR [39] FeoC [183] quorum sensing LasI/RhlI [343] PupR [32] TraR [409]

130

H Appendixes

Fig. H-1: Half-lives of the corresponding parental strains for AAA+ deficient strains. Half-lives for

His6-Ffh stanility in E. coli MG1655, W3110 and BW25113 were summarized. After in vivo degradation experiments, half-lives and standard deviations were calculated from three biological replicas.

131

H Appendixes

Fig. H-2: Purification of His6-Ffh and Lon-His6 and in vitro stability of His6-Ffh. In this figure, an example of purification profiles after SDS-PAGE and Coomassie staining is given for (A) His6-Ffh

(approx. 50 kDa) and (B) Lon-His6 (approx. 90 kDa). Both recombinant proteins were successfully purified via nickel affinity chromatography. Shown are the purification samples: pellet (P), cell lysate (C), samples of the washing steps (W1 and W4) and samples of the elution steps (E1 to E5). Both purified proteins were used for in vitro degradation experiments to analyze the Ffh stability in vitro (C) in the absence and (D) in the presence of Lon either with or without ATP over 120 min. In vitro degradation profiles are shown after Coomassie staining and immunodetection with a monoclonal α-Ffh antibody. Below the relative Ffh amount of each sample is given in % as a bar chart.

132

H Appendixes

Fig. H-3: Degradation of cellular Ffh after plasmid-derived His6-Lon overproduction. Stability of cellular Ffh was analyzed after His6-Lon overproduction with in vivo degradation experiments at various growth phases. A main culture was grown at 37 °C, to defined growth phases degradation cultures were taken and plasmid-derived His6-Lon overproduction (pLon-ASKA) was induced by adding 1 mM IPTG (A). Stability of cellular Ffh was analyzed in E. coli MC4100 (B) and Δlon (C). Translation was stopped by the addition of spectinomycin (Sp), samples were taken and prepared for SDS-PAGE. After SDS- PAGE, Western transfer and immunodetection, half-lives were calculated.

133

H Appendixes

Fig. H-4: Degradation of cellular Ffh during oxidative stress conditions. Stability of cellular Ffh was analyzed under oxidative stress conditions with in vivo degradation experiments. A main culture was grown at 37°C up to an OD580 0.5 and was divided in five degradation cultures (A). Each degradation culture was exposed to 0-2 mM H2O2 for 60 min to induce oxidative stress. Stability of cellular Ffh was analyzed in E. coli MC4100 (B) and Δlon (C). Translation was stopped by the addition of spectinomycin (Sp), samples were taken and prepared for SDS-PAGE. After SDS-PAGE, Western transfer and immunodetection half-lives were calculated.

134

H Appendixes

Fig. H-5: Degradation of cellular Ffh after rifampicin treatment. Stability of cellular Ffh was determined in E. coli MC4100 at defined growth phases with in vivo degradation experiments. A main culture was grown at 37 °C, at defined growth phases degradation cultures were taken and translation was stopped with spectinomycin (Sp) (A), rifampicin (Rif) (B) or Sp/Rif. Afterwards, samples were taken, prepared for SDS-PAGE, Western transfer and immunodetection and half-lives were calculated.

135

H Appendixes

Fig. H-6: Degradation of cellular Ffh in ΔrpoH. Degradation experiments for cellular Ffh MC4100 (A) and ΔrpoH (B) were performed at 25 °C at defined growth phases (I to IV). Translation was stopped by the addition of spectinomycin (Sp), samples were taken and prepared for SDS-PAGE. After SDS- PAGE, Western transfer and immunodetection, half-lives were calculated from two biological replicas. (C) Relative Lon amount was determined in MC4100 and ΔrpoH. After SDS-PAGE and Western transfer, Lon was immunologically detected with α-Lon antibody. Relative Lon amount was calculated relative to Lon amount in growth phase I in MC4100.

136

H Appendixes

Fig. H-7: Light microscopy of MC4100 carrying either His6-tag or One hybrid constructs. Shown are light microscopy images of MC4100 cells carrying either a His6-tag construct or a One Hybrid construct in the exponential or stationary growth phase. Before light microscopy, MC4100 was transformed with the empty vector (EV), plasmid-encoded Ffh, plasmid-encoded NG domain, or plasmid-encoded M domain of the respective construct. The main culture was inoculated and allowed to grow overnight. At an OD580 of 0.5-0.8, cell culture was taken for microscopy in the exponential phase. After overnight incubation, cell culture was used for microscopy of the stationary cells. Microscopy was performed at 100x magnification. The indicated scale corresponds to 5 µm.

137

H Appendixes

Fig. H-8: Deleted region within the M domain after transformation in Δlon. (A) Structure of the SRP complex consisting of Ffh (green) and the 4.5S RNA (grey) and its receptor FtsY (blue). The red- colored region shows the deleted part of Ffh after transformation in Δlon. (B) A schematic overview of the M domain of amino acid (aa) 296 to 453 is shown. The M domain consists of four helices (H1 to H4), where h2 to h4 represent a helix turn helix motif involved in the binding of 4.5S RNA. The conserved RNA-binding motif is located within the helix turn helix motif (h2 b to h3). The red region was deleted and the green region was retained after transformation into Δlon.

138

H Appendixes

>120 18 18 43 >120

>120 15 10 8 >120

98 60 38 20 >120

75 >120 66 96 >120

Fig. H-9: Half-lives of Ffh variants with evolutionary point mutations. The stability of the plasmid- derived His6-tagged Ffh variants during bacterial growth at 37 °C in MC4100 was determined by in vivo degradation experiments. After SDS-PAGE, Western transfer and immunological detection, half-lives were calculated. Half-lives for each Ffh variant are illustrated using a representative example. Half-lives for S382V_C406S, G405D, G405D_C406S and S382V_G405D_C406S were taken from a B. th. of Fitore Morina [101] (B) Solubility studies of the tested Ffh variants. The respective Ffh variant was overproduced by the addition of 50 ng/µl AHT for 30 min. Cells were disrupted by French press and the obtained cell lysate was separated into supernatant and pellet fraction by centrifugation. The supernatant contains the soluble proteins, whereas the pellet contains insoluble proteins and membrane components. Data for the solubility studies (for S382V, C406S, S382V_C406S, G405D, G405D_C406S and S382V_G405D_C406S) were taken from the B. th. of Fitore Morina [101]. n.d. = not detectable

139

H Appendixes

Fig. H-10: Degradation of His6-FusA in MC4100. Protein stability was analyzed over the entire bacterial growth with in vivo degradation experiments typically at 37°C. Therefore, degradation cultures were taken at defined growth phases (I to V). His6-FusA overproduction was induced for 30 min by the addition of 1 mM IPTG. Translation was stopped by the addition of spectinomycin (Sp), samples were taken and prepared for SDS-PAGE. After SDS-PAGE, Western transfer and immunodetection half-lives were calculated. Standard deviations were determined from three biological replicas.

140

Publications

141

I Publications

Research articles:

Möller, P., Busch, P., Sauerbrei, B., Kraus, A., Förstner, K. U., Wen, T.-N., Overlöper, A., Lai, E.-M., and Narberhaus, F. 2019. The RNase YbeY is vital for ribosome maturation, stress resistance, and virulence of the natural genetic engineer Agrobacterium tumefaciens. Journal of Bacteriology 201, 11, e00730-18.

Conference contributions:

B. Sauerbrei, J. Arends, F. Narberhaus (2017): The protein component of the signal recognition particle Ffh is subject to Lon-dependent proteolysis in Escherichia coli. Poster, VAAM Annual Meeting 2017, Jena, Germany, book of abstract poster 565/MTP, p. 132.

B. Sauerbrei, J. Arends, F. Narberhaus (2018): Lon-dependent proteolysis of the signal recognition particle component Ffh in Escherichia coli. ePoster, Annual Conference of the Association for General and Applied Microbiology 2018 , Wolfsburg, Germany, book of abstract poster MTP343, p. 77.

142

Lebenslauf

143

J Lebenslauf

Persönliche Daten

Name Beate Sauerbrei Geboren: 04.04.1988 in Naumburg Staatsangehörigkeit: deutsch Familienstand: ledig

Schulische Ausbildung

08/1998 – 06/2006 Hennebergisches Gymnasium „Georg Ernst“ in Schleusingen Abschluss: Abitur 08/1994 – 06/1998 Hörselberggrundschule in Wutha-Farnroda

Berufsausbildung/Studium/Promotion

seit 10/2015 Anfertigung der Doktorarbeit an der Ruhr-Universität Bochum am Lehrstuhl für Biologie der Mikroorganismen 10/2013 – 09/2015 Studium der Biologie an der Ruhr-Universität Bochum Abschluss: Master of Science (M. Sc.), Gesamtnote: 1,4 10/2010 – 08/2013 Studium der Biologie an der Ruhr-Universität Bochum Abschluss: Bachelor of Science (B. Sc.), Gesamtnote: 1,8 08/2006 – 07/2008 Berufsausbildung zur Biologisch-technischen Assistentin an der Akademie Göttingen Abschluss: Staatlich geprüfte Biologisch-technische Assistentin

Berufserfahrung

10/2015 – 09/2019 Wissenschaftliche Mitarbeiterin am Lehrstuhl für Biologie der Mikroorganismen der Ruhr-Universität Bochum 04/2015 – 08/2015 Studentische Hilfskraft am Lehrstuhl für Biologie der Mikroorganismen der Ruhr-Universität Bochum 08/2008 – 08/2010 Angestellt als Biologisch-technische Assistentin am Institut für Pathologie der Universitätsmedizin Mainz

Stipendien/Graduiertenschulen/Vereingungen

seit 04/2016 Mitglied der Internationalen Graduiertenschule Biowissenschaften (IGB) der Ruhr-Universität Bochum seit 11/2015 Mitglied der Vereinigung für Allgemeine und Angewandte Mikrobiologie 10/2015 – 12/2015 Mitglied der Research School der Ruhr-Universität Bochum Stipendium im Rahmen des Graduiertenkollegs SFB 642

144

Erklärung

145

K Erklärung

ERKLÄRUNG

Ich versichere an Eides statt, dass ich die eingereichte Dissertation selbstständig und ohne unzulässige fremde Hilfe verfasst, andere als die in ihr angegebene Literatur nicht benutzt und dass ich alle ganz oder annähernd übernommenen Textstellen sowie verwendete Grafiken und Tabellen kenntlich gemacht habe. Weiterhin erkläre ich, dass digitale Abbildungen nur die originalen Daten enthalten oder eine eindeutige Dokumentation von Art und Umfang der inhaltsverändernden Bildbearbeitung vorliegt. Außerdem versichere ich, dass es sich bei der von mir vorgelegten Dissertation (elektronische und gedruckte Version) um völlig übereinstimmende Exemplare handelt und die Dissertation in dieser oder ähnlicher Form noch nicht anderweitig als Promotionsleistung vorgelegt und bewertet wurde.

Es wurden keine anderen als die angegebenen Hilfsmittel verwendet.

Die Dissertation wurde gemäß der Promotionsordnung und der Betreuungsvereinbarung angefertigt.

Bochum, den

______(Unterschrift)

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