Research Collection
Doctoral Thesis
Mechanistic studies of mycobacterial caseinolytic proteases (Clp)
Author(s): Warweg, Jannis
Publication Date: 2014
Permanent Link: https://doi.org/10.3929/ethz-a-010376409
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ETH Library
Diss. ETH No. 21564
Mechanistic studies of mycobacterial caseinolytic proteases (Clp)
A dissertation submitted to
ETH Zurich
for a degree of
Doctor of Science
presented by
Jannis Warweg Dipl. Biologie, Universität Göttingen born on September 26th, 1982
German
accepted on the recommendation of:
Prof. Eilika Weber-Ban Prof. Rudi Glockshuber Prof. Peter Sander
2014
Contents
1 Summary/Zusammenfassung 5
1.1 Summary 5 1.2 Zusammenfassung 7
2 Introduction 9
2.1 Mycobacteria 9 2.2 Energy dependent protein degradation in Mycobacteria 10 2.2.1 Mycobacterial chaperone-proteases 10 2.2.2 Cellular function of chaperone-proteases 12 2.2.3 Substrate recognition by chaperone-proteases 15 2.3 Mycobacterial Clp Protease 19 2.3.1 Regulation of clp genes in Mycobacteria 19 2.3.2 Architecture and mechanisms of Clp proteases 20 2.3.3 The interaction between the ClpP particle and ATPases 24 2.3.4 Molecular details of Clp ATPases 25 2.4 Pathogenesis and drug design 29 2.4.1 Role of chaperone-proteases during pathogenesis 29 2.4.2 Drug design 30 2.4.3 The Clp protease activation molecule ADEP 31
3 Aim of the study 33
4 Material and methods 34
4.1 Plasmids 34 4.2 Expression and purification of proteins 35 4.3 Preparation of ClpP1P2 complexes 37 4.4 Assembly of the complex between protease and chaperone analyzed by analytical gelfiltration 37 4.5 Crystallography 38 4.6 Data collection, structure determination, refinement and alignment 38 4.7 ClpP peptidase activity 39 4.8 Processing of ClpP1ClpP2 39 4.9 ATPase Assays 39 4.10 Analysis of the Mtb H37Rv proteome for ClpX degrons 40 4.11 Degradation of ssrA-substrates, FR-linker proteins and Pup conjugated proteins by Clp chaperone-proteases 40 4.12 Peptide inhibition of ClpXP degradation 40 4.13 Pupylation of ClpP2 and Adk 41
3 Contents
4.14 Degradation of ClpP2-Pup by the proteasome 41
5 Results 42
5.1 ClpP1P2 assembled in a tetradecamer 42 5.2 Crystallization and structural comparison of Mtb ClpP1 and ClpP2 47 5.3 Analysis of the N-terminal processing in the P1P2 complex 50 5.4 Contribution of the individual subunits to the peptidase activity 54 5.5 ClpP function in combination with AAA binding partners 57 5.5.1 Characterization of the two possible binding partners ClpX and ClpC1 57 5.5.2 ClpX stimulates proteolytic sites of ClpP 60 5.5.3 Scan of the Mtb genome for proteins carrying ssrA-like degrons 63 5.5.4 Small peptides inhibit ClpXP1P2 dependent degradation 65 5.6 Post-translational modifications of ClpP1P2 66 5.6.1 Pupylation 66 5.6.2 Phosphorylation of ClpP1 68
6 Discussion 70
7 Bibliography 76
8 Appendix 92
8.1 Abbreviations 92 8.2 Properties and sequences of the protein constructs 95 8.3 Structures of activator and peptide substrates 102
9 Acknowledgment 105
10 Curriculum Vitae 106
4
1 Summary/Zusammenfassung
1.1 Summary
Recently identified drug targets with a potential to combat tuberculosis are the Clp chaperone-proteases of Mycobacterium tuberculosis. The Clp proteases share a common architecture of two stacked rings that form a cavity with 14 internal active centers which are sequestered from the environment. Mycobacterium tuberculosis contains two genes for the caseinolytic-serine-proteases clpP1 and clpP2. The interaction of the Clp particle with a Clp ATPase is a requirement for the chaperone-protease to enable its degradation activity. The available ATPases in Mycobacterium tuberculosis ClpX or ClpC recognize proteins, exhibiting a degradation motif. The ATPases deliver free energy from ATP hydrolysis to unfold the substrate and to translocate the unraveled polypeptide chain into the associated ClpP particle. In this thesis the composition and assembly pathway of the mycobacterial Clp peptidase were investigated by analytical size exclusion chromatography. For the assembly the homo- heptameric rings of ClpP1 and ClpP2 associate into a double-ring consisting of one ClpP1 and one ClpP2 ring. In these complexes truncated N-termini of ClpP1 and ClpP2 were observed. The cleavage product was described as a propeptide. The propeptide of ClpP1 included the first six amino acids (VSQVTD) and for ClpP2 the motif sequence was VNSQNSQIQPQA. A gel-based assay combined with mass spectrometric analysis demonstrated that propeptide cleavage could only occur when both subunits ClpP1 and ClpP2 were present. Tests with active and inactive subunits revealed the restriction of ClpP2 to only auto catalytically cleave itself, but ClpP1 was able to process itself and cross process ClpP2. A requirement for the activities of the Mtb Clp proteases was the presence of an activator (Z-LL-H) which is a dipeptide with a N-terminal protecting group, that had been described in the literature. However, in the presence of the ATPase ClpX, protein degradation could be detected in the absence of an activator peptide. Furthermore, the association with ClpX was sufficient for maturation (propeptide cleavage) of ClpP1P2 in the absence of the activator peptide (Z-LL- H). Finally, the influences of two post-translational modifications on the ClpP particle were investigated. The phosphorylation of ClpP1 at threonine 17 reduced the peptidase activity
5 1 Summary/Zusammenfassung
in complex with ClpP2. However, the modification of ClpP2 by Pup-conjugation of the lysines 174 or 181 showed an increased effect on the activity, beside the fact that the conjugated Pup-tag served as a degradation signal for the mycobacterial proteasome.
6 1 Summary/Zusammenfassung
1.2 Zusammenfassung
In der Entwicklung neuer Medikamente gegen Tuberkulose sind mögliche Ansatzpunkte, die für den Erreger Mycobacterium tuberculosis essentiellen Chaperone-Proteasen. Mycobacterium tuberculosis besitzt zwei Gene für caseinolytische-Serine-Proteasen clpP1 und clpP2. Clp Proteasen haben eine konservierte Architektur, die aus zwei Ringen mit je sieben Untereinheiten besteht, die einen abgeschotteten Innenraum bilden. In diesem befinden sich die 14 aktiven Zentren, die zum Schutz vor unkontrollierter Proteolyse von der Umgebung abgeschirmt sind. Der ClpP Partikel ist nur durch Interaktion mit Clp ATPasen zum Proteinabbau befähigt. In Mycobacterium tuberculosis existieren die ATPasen ClpX und ClpC1, die in der Lage sind ATP zu hydrolysieren. Eine weitere Funktion der Clp ATPasen ist die Erkennung von Proteinen, die ein Degradationsmotiv aufweisen. Sie verwenden die frei werdende Energie zur Entfaltung der gebundenen Substrate und für die Weitergabe der entfalteten Polypeptidkette an die Clp Proteasen. Die Proteasen spalten dann die Aminosäurekette in kleine Fragmente. In dieser Doktorarbeit wurden die Zusammensetzung und der Aufbau von myco- bakteriellem ClpP1 und ClpP2 durch analytische Größenausschlusschromatographie untersucht. Dabei bilden die heptameren Untereinheiten ClpP1 und ClpP2 einen Doppelring, mit jeweils einem ClpP1- und einem ClpP2-Ring. In diesen Komplexen konnte ein Reifungsprozess, in Form N-terminaler Abspaltungen der Propeptide von ClpP1 und ClpP2, beobachtet werden. Das Propeptid von ClpP1 besteht aus sechs Aminosäuren (VSQVTD) und für ClpP2 beinhaltet das Motiv die Sequenz VNSQNSQIQPQA. Eine Gel- basierte Untersuchung mit anschließender Massenanalyse zeigte, dass die Reaktion die Anwesenheit beider Untereinheiten ClpP1 und ClpP2 benötigte. Ein Test mit aktiven und inaktiven Untereinheiten bewies, dass ClpP2 sich ausschließlich selbst autokatalytisch spaltet, während ClpP1 sich selbst und auch ClpP2 prozessiert. Die Propeptid-Abspaltung sowie die Peptidase-Aktivität mit unterschiedlichen fluorogenen Substraten benötigten die Zugabe eines in der Literatur beschriebenen Aktivators (Z-LL-H), ein Dipeptid, das eine N- terminale Schutzgruppe besitzt. Dagegen konnten wir in Gegenwart der ATPase ClpX Proteinabbau ohne den Zusatz weiterer Aktivator Peptide beobachten. Des Weiteren reichte die Interaktion mit ClpX aus, um in der Protease ClpP1P2 den Abspaltungsprozess des N-Terminus in Abwesenheit des Aktivator-Peptids (Z-LL-H) auszulösen.
7 1 Summary/Zusammenfassung
Ferner wurde der Einfluss von post-translationalen Modifikationen auf den ClpP Komplex untersucht. Die Phosphorylierung von ClpP1 am Threonin 17 reduzierte die Peptidaseaktivität im Komplex mit ClpP2. Dagegen konnte für die Pup-Modifikation der Lysine 174 oder 181 von ClpP2 eine Aktivitätszunahme gemessen werden. Außerdem konnte der konjugierte Pup-Tag den Abbau durch das prokaryotische Proteasom induzieren.
8
2 Introduction
2.1 Mycobacteria
Mycobacteria belong to the Actinobacteria. This phylum contains Gram-positive bacteria with high GC content in their genome. Typically, these bacteria exhibit special cell wall characteristics by building mycelium. Members of this group are aerobes and share a few similarities to eukaryotic cells, such as exospores, a eukaryotic-like fatty-acid synthetase (FasI) (Gago et al. 2011), and a sub-group the Mycobacteria produces sterols (Cavalier- Smith 2002). Another analogy is the existence of a proteasome (Striebel et al. 2009). Many members of the Actinobacteria live in close relation with eukaryotic hosts, either as symbionts (nitrogen fixing or gastrointestinal species) (Ventura et al. 2007), or in a few cases as prominent pathogens such as Mycobacterium leprae or Mycobacterium tuberculosis. The genus Mycobacterium contains around 100 members of rod shaped bacteria. The cell wall of Mycobacteria typically consists of a high density of lipids. Because of the thick layer, these organisms are acid fast and bear resistance against most antibiotics. Two subgroups emerge: rapid growers with 1-4 hours doubling time (e.g. M. smegmatis) and slow growers with a generation time of 6-24 hours such as M. tuberculosis (Tortoli 2006).
Mtb is the causative agent of tuberculosis, a disease that kills 1.8 million people every year. Roughly one third of the world's population presumably harbors latent tuberculosis and in 2–10% of all cases M. tuberculosis reactivates during their lifetime (Gedde-Dahl et al. 1952; Marks et al. 2000). Over 0.5 million cases of tuberculosis with multi-drug resistant cell lines (WHO 2012) making the discovery of new drug targets of major global interest. Tuberculosis is a droplet infection of the lungs. In this tissue, the macrophages, the immune cells of the host, phagocyte the bacteria without the ability to eliminate the organism. The survival of Mtb in macrophages is based on the inhibition of phagosome maturation (Russell et al. 2001) and the ability to replicate in this environment. The properties of the phagosome are distinguished by acidic conditions generated by the vacuolar ATPase proton pump. Further reactive oxygen species and nitrogen intermediates (RNIs) are produced and are responsible for irreversible damage of microbial proteins during host invasion by pathogens. Additionally, a variety of hydrolytic enzymes cause degradation of phagosomal contents in macrophages. During the later phase of the immune response, granulomas develop: rings of macrophages
9 2 Introduction
and lymphocytes around the necrotic infection area. Human granulomas are hypoxic and miss airway communication (Haapanen et al. 1959). The hypoxic response is controlled by the two-component system DosR - DosS (DevR – DevS) (Dasgupta et al. 2000). In response to low oxygen or NO exposure, the sensor kinases DosS and DosT autophosphorylate their specific conserved histidine residues (Roberts et al. 2004). The signal cascade proceeds by the phosphoryl transfer to DosR, enhancing its DNA binding affinity and resulting in the upregulation of 48 genes involved in dormancy (Park et al. 2003). The pathogen also reacts to nutrient stress in the granulomas in form of a “starvation response”, including a down- regulation of biosynthetic pathways and low oxygen consumption. In this transient phase Mtb enters the non-replicating state called dormancy, enabling the survival in phagosomes for decades. Active tuberculosis can be triggered by reactivation (Parrish et al. 1998).
2.2 Energy dependent protein degradation in Mycobacteria
2.2.1 Mycobacterial chaperone-proteases
Chaperone-proteases exist in all domains of life. The cytoplasmic degradation in eukaryotes is predominantly executed by the 26S proteasome, while in bacteria a set of chaperone- proteases are responsible, including members of the caseinolyic protease (Clp) family. Exceptional for bacteria, the Actinobacteria (including the genus Mycobacterium) have homologs of the 20S proteasome (Li et al. 2010). It is not clear, if these bacteria developed an ancestral form of the eukaryotic proteasome (Valas et al. 2008), or inserted the proteasome later by horizontal gene transfer (Volker et al. 2002). Bacterial chaperone-proteases, the Clp proteases as well as Lon and FtsH, share several common features. These molecules have cylindrical shapes with layers of protease- and layers of ATPase-rings. Like the eukaryotic homolog, the mycobacterial proteasome contains four protease rings. Each of the two inner rings consist of seven PrcB subunits and both outer layers have the same amount of PrcA subunits. The proteasome is sandwiched by two hexameric rings of the ATPase Mpa. The Clp chaperone-protease complexes also feature a protease core, the ClpP particle.
ClpP comprises two rings, each with seven subunits. The ClpP14 particle is flanked on both sides by hexameric Clp ATPase rings. A mismatch between the hexameric ATPase and the heptameric protease exist for the proteasome and the Clp chaperone-proteases. Further, the ATPase and the protease are separated on two polypeptide chains. On the contrary, for Lon and FtsH the ATPase and the protease are encoded on one polypeptide. They assemble
10 2 Introduction
in hexamers without a mismatch (Figure 1). Separately encoded Clp ATPase and peptidase subunits are widely conserved. ClpP was first identified in E. coli (Katayama et al. 1986) and is also present in the mitochondria of eukaryotes but seems to be absent in archaea. The majority of bacteria possesses one clp gene, the exceptions being Actinobacteria, Chlamydiae and Cyanobacteria with multiple isoforms.
Figure 1: The set of mycobacterial chaperone-proteases exhibit a similar architecture. These macromolecules consist of a sequestered protease particles (grey). The membrane anchored FtsH forms a hexameric protease ring in contrast ClpP1P2 assembles in two heptameric rings, and the proteaseome comprises even four rings of seven subunits. The barrel-shape protease core interacts with a hexameric AAA+ ATPase (orange), being is responsible for substrate recognition and delivery (Laederach et al. manuscript in preparation).
The number and types of Clp ATPases vary even among closely related organisms. Two classes of Clp ATPases exist: ClpX (class 1) with one ATP-binding domain or ClpA and ClpC (class 2) with two ATP-binding domains. In most bacteria ClpX and one version of the class 2 ATPases exist. As a general rule ClpX and ClpC are always present in Gram-positive bacteria including the genus of the Mycobacteria, while ClpA is found in Gram-negatives (Frees et al. 2007). In Bacillus subtilis, the number of Clp molecules per cell was estimated at 1400 ClpX hexamers, 250 ClpC hexamers, and 1200 tetradecameric ClpP, during exponential growth (Gerth et al. 2004). Beside the caseinolytic chaperone-proteases, protein degradation in bacteria has been
11 2 Introduction
extensively reported for two other systems (Lon and FtsH). In contrast to most bacteria, Mycobacteria do not possess the cytoplasmic chaperone-protease Lon, but they do contain FtsH. This membrane associated metalloprotease has a zinc ion in the catalytic site. The metal ion is coordinated by a glutamate and two histidine residues. Furthermore, the Zn2+ Ion coordinates a water molecule that functions as the nucleophile in the proteolytic reaction (Deuerling et al. 1995). FtsH has the function to regulate heat shock, therefore a substrate is σ32 (Tomoyasu et al. 1995) (Table 1).
Table 1: Comparison of chaperone-proteases
Clp Lon Proteasome FtsH HslUV
membrane Cellular location Cytoplasmic cytoplasmic cytoplasmic cytoplasmic bound ClpS, MecA, HflK, HflC, HflD, Additional adapters MscB, SspB, PolyP, PinA SpoVM, MgtR RssB ssrA -tag, ssrA-tag, N- linear pupylated endogenous endogenous Recognition motifs end rule aromatic substrates motifs motifs residues separate encoded on separate encoded on separate subunit: ATPase partner same subunit: same subunit: ClpA, ClpX, polypeptide Mpa polypeptide HslU ClpC serine serine threonine Zn threonine Active center protease protease protease metalloprotease protease
2.2.2 Cellular function of chaperone-proteases
Proteolysis guarantees homeostasis by removing misfolded, mislocalized, aggregated, damaged or unneeded proteins to avoid toxicity for the cell. Protein aggregates are biologically inactive and interfere with vital cellular functions. In clpP knockout mutants 20- 30% of newly synthesized proteins aggregate (Kock et al. 2004). Protein abnormalities can be caused through mutations in DNA, errors in transcription or translation, post-translational damage (arising through heat or oxygen radicals), and spontaneous denaturation. A broad set of proteins can be degraded by ClpAP, ClpCP or ClpXP. ClpXP has more than 100 identified substrates (Baker et al. 2012; Flynn et al. 2003; Neher et al. 2006). In most cases, the cell-associated proteases of E. coli and B. subtilis are non-essential for normal growth (Lipinska et al. 1989). The proteases ClpXP and Lon only become essential
12 2 Introduction
for E. coli during heat shock if DnaK, the major Hsp70 chaperone, shows reduced levels (Tomoyasu et al. 2001). The role of these protease complexes in certain pathogens on the other hand seems very different. There the molecular machines are essential for viability and virulence (Raju et al. 2012).
Table 2: Protease overview FtsH ClpP ClpX ClpC ClpA ClpB Proteasome Lon HslUV Mitochondria + + + - - + + - Chloroplasts + + + + -/+ + + + - Gram- negative Salmonella sp. + + + - + + - + + E. coli + + + - + + - + + Gram-positive B. subtilis + + + + - - - + + S. aureus + + + + - + - - + S. pneumoniae + + + + - + - - - M. tuberculosis + + + + - + + - - M. leprae + + + + - + + - - M. smegmatis + + + + - + + + - C. diphteriae + + + + - + - - - C. glutamicum + + + + - + - - -
The functions of chaperone-proteases can be separated into two types, general versus regulatory proteolysis. During general proteolysis, damaged or misfolded proteins are eliminated, which is part of the protein quality control of the cell. In regulatory proteolysis mainly transcription factors and signal transduction proteins are removed. The proteolysis of such proteins arranges a fast and efficient way to control crucial checkpoints for diverse cellular pathways (Schmidt et al. 2009).
General proteolysis: selection between misfolded and native proteins
Environmental stress conditions, such as extreme pH or heat, trigger the destabilization of protein structures. During protein unfolding, hydrophobic regions normally buried in the inner core of native proteins get exposed at the surface. Unfolded hydrophobic polypeptide stretches get tangled up to irreversible aggregates. These hydrophobic signals are enriched in aromatic residues and selectively target wrong folded proteins for chaperone-protease
13 2 Introduction
recognition (Gur et al. 2008; Rüdiger et al. 1997). Misfolding can also occur under non- stressful conditions, especially during de novo folding after translocation (Kramer et al. 2009). The ATPase ClpC is responsible for recruiting ClpP to degrade non-native proteins in B. subtilis and S. aureus (Krüger et al. 2000). ClpX fulfills a variety of roles: Mutants in B. subtilis are heat sensitive in contrast to S. aureus, where deletion improves heat resistance (Frees et al. 2003). Moreover, the Clp system is involved in different stress responses. Knockout mutants of clp genes exhibit a broad range of phenotypic changes, e.g. enhancing general stress sensitivity. One substrate is SPX (YjbD; suppressor of ClpX and ClpP) (Nakano et al. 2002), the major stress regulator under oxidative stress. HdiR was identified in L. lactis (Savijoki et al. 2003) and resembles the SOS response regulator LexA (DNA damage). HdiR also gets degraded by Clps. In B. subtilis, ClpP is responsible for 50% of the cellular protein turnover (Kock et al. 2004) and the down-regulation of central metabolism in glucose-starved cells (Gerth et al. 2008).
Regulated proteolysis: substrate stability and control by the accessibility of the degradation signal
The controlled elimination of enzymes or signal transduction factors regulates key steps in either metabolism or anabolism, which relies on specific recognition of degrons. Different strategies are known. One possibility is an always accessible tag at the N- or C-terminus such as RecN (Neher et al. 2006). But also post-translational modifications by degron fusion brand proteins for degradation. Another possibility for degradation labeling is the interaction with a factor that remodels the substrate and reveals a shielded degradation tag. A prominent example is the transposase MuA of phage Mu, in which the tetramer exposes a signal recognized by ClpX (Abdelhakim et al. 2008). A further event for tagging a protein to degradation is an endo-proteolytic cleavage which generates a new C or N-terminal degron (e.g. the cleavage of the N-domain of RseA) and makes the protein a substrate for ClpXP (Schmidt et al. 2009). For sporulation σH (SpoOH) is activated by ClpX and after completion of sporulation degraded in a process ClpC is involved in (Liu et al. 2000). A well described substrate of ClpCP in B. subtilis is the master regulator of competence development ComK during vegetative growth (Frees et al. 2007; Turgay et al. 2001), where degradation results in competence development in stationary phase cells (Turgay et al.
14 2 Introduction
1998). Furthermore, aberrant cell morphology like abnormal cell wall structures, filamentation due to degradation, lack of MurAA (catalysis first step in peptidoglycan biosynthesis), or overproduction show the same filamentous phenotype as a ClpP knockout (Kock et al. 2004). Additionally, the knockout is impaired in cell division (FtsZ inhibitor of cell division is degraded by ClpP), separation (Msadek et al. 1998; Nair, et al. 2000), and in initiating developmental programs. An organism in which ClpP is essential for viability is Caulobacter crescentus. There, the peptidase degrades the cell cycle regulator CtrA and the clpX or clpP knockouts arrest in the cell-cycle.
2.2.3 Substrate recognition by chaperone-proteases
The chaperone generally recognizes substrates via C- or N-terminal signals such as the ssrA-tag, the Pup-tag, or the N-degron. Both N- or C-terminal degrons are treated equally. During unraveling, the chaperone-protease starts from the point of attachment and moves along the polypeptide backbone. The rate depends on the structural stability of the substrate protein around the degron site and not of the overall structural stability against chemical denaturation or global unfolding (Kenniston et al. 2003). Structural elements are responsible for the completeness of the degradation. Substrates displaying exposed unstructured loops at their surface and α-helices beside the degradation initiation side facilitate unraveling in contrast to buried β-strands (Lee et al. 2001). The process is highly cooperative when unfolding overcomes the first energy barrier beside the recognin, then the rest of the structure follows fast (Prakash et al. 2004). Although eukaryotes and prokaryotes share the basic strategy to accommodate general protein quality control in sequestered and chambered proteases, the basic mechanism for substrate selection is different. While in eukaryotic cells proteins are branded by ubiquitin for degradation, in most prokaryotes no comparable system exists. Instead bacteria have solved the specific assignment to degradation by engaging diverse adaptor proteins.
TmRNA encoding the ssrA-tag, a degron for Clp Proteases
For Clp proteases, a set of short peptide sequences acts for direct or indirect substrate recognition. One example is the eleven amino acid long C-terminal ssrA-tag (AADSHQRDYALAA in Mtb). This degron is encoded on a small rescue RNA molecule called
15 2 Introduction
tmRNA and was first discovered in Mycobacterium tuberculosis. This rescue mechanism occurs during translation failure, due for example to mRNAs missing their stop codon. The tmRNA binds to stalled ribosomes and reactivates ribosome translation by transfer of the stalled chain to tmRNA and translating its encoded ssrA-tag. Consequently, the ssrA-tag is fused to the polypeptide chain of the unfinished translation product. This rescue mechanism marks the unfinished protein, with possible impaired function or even cell toxicity, for degradation. The tmRNA recycles the ribosome by adding a stop codon behind the degradation tag to the unfinished polypeptide and enables the degradation of the messenger RNA. Most cellular proteins can be targeted for ClpAP or ClpXP dependent degradation by the C-terminal fusion of the ssrA-tag (Dougan et al. 2003; Farrell et al. 2005; Gottesman et al. 1998).
Adaptor proteins: expanding and regulating chaperone-protease function
Not all degrons are directly recognized by the unfoldase. The recognition process becomes much more specific with additional adaptors (Schmidt et al. 2009). Adaptors are defined as protein factors that link recognition of substrate degrons to the chaperone-protease. Adaptor proteins usually dock to the N-terminal domains of the ATPases, the least conserved and therefore most specific part for a certain chaperone. The flexibility of the N-domains of the ATPases plays an important role for the delivery. The N-domains of ClpC and ClpA are highly similar and bacteria usually possess one or the other but not both. The main difference of the N-domains is the electrostatic surface potential which defines the binding specificity for the regulatory molecule (Kojetin et al. 2009). The ~10 kDa C4-zinc binding domain of ClpX is required for the association with the adaptor protein SspB and contributes to binding of certain substrates (Banecki et al. 2001; Dougan et al. 2003; Levchenko et al. 2003; Wojtyra et al. 2003). Although the N-domain of ClpX is redundant for degradation of ssrA-tagged proteins, it can play a crucial role for some substrates. The unfoldase recognizes the C-terminal amino acids LAA and especially the negative charge of the α-carboxylate of the ssrA-tag (Flynn et al. 2001) via the basic RKH loop (Farrell et al. 2007; Martin et al. 2007; Siddiqui et al. 2004). In contrast, the adaptor SspB binds the N-terminal part of the ssrA-motif. Beside the recognition of ssrA-tagged substrates, SspB binds the C-terminus of the processed RseA N-domain and targets the anti-sigma factor to ClpXP dependent degradation (Flynn et al. 2004). The crystal structure of SspB shows that both substrates bind to the same groove, but in the opposite direction (Levchenko et al. 2003). In contrast to its delivered substrate, SspB does not undergo
16 2 Introduction
degradation and serves multiple rounds as an adaptor by interacting with the chaperone- proteases in a manner that prevents it from being recognized as a substrate. The function of SspB is restricted to substrate recognition and hand over. The adaptor protein SspB increases the affinity of ssrA-tagged substrate degradation significantly (~10 fold) (Bolon et al. 2004), ClpX pulls at the ssrA-tag and thereby loses contact to the SspB adapter (Baker et al. 2012) which is released for another round of substrate recruitment. Surprisingly, even though ClpX is part of the Mtb proteome, an sspB gene has not been found. Further adaptors for ClpX are UmuD (targets UmuD’ for degradation and is involved in UV- protection) (Neher et al. 2003), CpdR in C. crescentus which is involved in regulatory proteolysis (Abel et al. 2011) and YjbH (delivers Spx for protein turnover) (Larsson et al. 2007). The regulatory adapter molecules connect the chaperone-proteases to different biological processes in response to environmental and or developmental cues (Table 3). Moreover, the adaptor proteins provide an additional regulatory layer to control protein degradation. The adaptors themselves can undergo modification by anti-adaptors. In E. coli, the regulator of σS (the adapter RssB) is even further regulated by multi anti-adaptor proteins, such as IraP, IraM, and IraD which prevent binding of substrate σS. This is one example of specific substrate inhibition in response to stress (Bougdour et al. 2006; Muffler et al. 1996; Schmidt et al. 2009). An adaptor for the ATPase ClpC is the molecule MecA, which assists in assembling of the chaperone in B. subtilis. Accordingly, MecA-ClpC build complexes, whereby six subunits of MecA are required to form the ClpC hexamer. In Gram-positive bacteria, the master regulator of protein quality control (CtsR) is phosphorylated at various arginines and inactivated by MscB (Fuhrmann et al. 2009).
Table 3: List of adaptor functions Chaperone Adapter Function Organism ClpX RssB degradation of σs E. coli UmuD target UmuD' for degradation SspB enhance degradation of ssrA tagged substrates and ATPase activity ClpC MecA ClpC assembly B. subtilis YjbH substrate delivery (Spx) McsB degradation of CtsR ClpA ClpS degradation of N-end rule substrates E. coli and inhibition of ssrA tagged substrates
17 2 Introduction
The N-end rule pathway
The adapter protein ClpS binds to the N-domains of the chaperones ClpA or ClpC. In E. coli, ClpS is encoded by a gene directly upstream of the clpA gene. The binding of ClpS to the chaperone promotes the degradation of N-end rule substrates and at the same time competitively inhibits the recognition of substrates, which depend on internal binding sites of the chaperone such as the ssrA-tag (De Donatis et al. 2010). Generally, the “N-end rule” refers to the fact that a protein’s half-life depends on the nature of its N-terminal residue. Destabilizing N-terminal residues are phenylalanine, tyrosine, tryptophan, and leucine which serve directly as recognition signals for degradation (referred to as the N-degron) (Dougan et al. 2002; Tobias et al. 1991). The secondary destabilizing residues lysine and arginine are recognized by L/F-tRNA protein transferase which then attaches an N-terminal primary destabilizing phenylalanine or leucine (Ninnis et al. 2009; Shrader et al. 1993). Orthologs of ClpS are also present in Mtb: But as ClpA is missing in Actinobacteria, another chaperone (ClpC or ClpX) must be involved as the binding partner.
Pupylation as a recruitment route to the proteasome
One of the best described degradation tags in M. tuberculosis is the prokaryotic ubiquitin like post-translational modification (Pup) which functions as a recognition sequence for proteasome-dependent degradation (Pearce et al. 2008). The Pup-tag has no sequence or structural homology to eukaryotic ubiquitin. In Actinobacteria, the single enzyme PafA is responsible for the ligation of the substrate and the Pup-tag (Striebel et al. 2009b). The eukaryotic ubiquitination in contrast includes three steps (activation, conjugation, and ligation) (Kerscher et al. 2006). However, both tags carry a diglycine motif and are ligated by an isopeptide bond to an amino group of a lysine on the substrate surface (Striebel et al. 2009a). In eukaryotes, ubiquitin is not only a degradation tag, but also functions as a post- translational modification in protein regulation. A few members of the Actinobacteria contain Pup, without offering a proteasome. It is possible that Pup not only functions as a degradation tag but also contributes to protein regulation. Among of the target protein clients for pupylation in Mtb ClpP2 was identified, opening the possibility that Clp complexes might be regulated by pupylation.
18 2 Introduction
2.3 Mycobacterial Clp Protease
The Mtb genome encodes the chaperone-proteases Clp, FtsH, and the proteasome. The proteasome may handle RNIs during infection of macrophage (Gandotra et al. 2007). The chaperone-proteases FtsH and ClpP appear to be essential cellular components for Mtb (Sassetti et al. 2003). These components are responsible for protein turn-over, quality control, and transcription regulation due to environmental changes.
2.3.1 Regulation of clp genes in Mycobacteria
The survival of Mtb in macrophages is linked to changes in the expression profile of ~600 genes (Schnappinger et al. 2003). A global transcriptome analysis of the log-phase and hypoxic stress identified ~100 genes responsible for reactivation. One of those genes is Rv2745c, encoding the Clp protease gene Regulator (ClgR). This indicates that the Clp proteases are involved in the transition from latent to active tuberculosis (Sherrid et al. 2010). The ClpP protease genes (clpP1, clpP2) form a gene cluster together with the gene coding for the chaperone ClpX which is located three genes upstream (clpX = Rv2457) (Figure 2). In the same locus directly beside clpP1, the trigger factor tig (Rv2462c) is encoded. The other annotated Clp chaperone genes clpC1 (Rv3596c) and clpC2 (Rv2667) are scattered at distant locations. Transposon mutagenesis screens predict clpC1 and clpX as essential genes for Mtb (Sassetti et al. 2003).
Figure 2: Gene locus of Clp genes. The clpC1 gene is located in a single operon, in contrast to ClpX which build a gene cluster with clpP1 and clpP2.
Both ClpP subunit genes (clpP1, clpP2) of Mtb are upregulated during microaerobic and hypoxic growth (Muttucumaru et al. 2004). ClpP1clpP2 (Rv2461c and Rv2460c in H37Rv) are arranged in one single bicistronic operon. They are both required for Mtb viability (Raju et al. 2012). The Mtb ClpPs are upregulated during reactivation from non-replicating phase (Sherrid et al. 2010) and are important during macrophage persistence. A Mtb strain with
19 2 Introduction
deleted clgR gene is deficient in the pH control of the phagosome. Additionally, ClgR regulates the protease genes (PtrB and HtrA like protease) and chaperone genes (Acr2, ClpB, and the chaperone Rv3269), thereby controlling a network which is responsible for homeostasis (Estorninho et al. 2010). The transcription activator ClgR is highly conserved in Gram-positive bacteria with high GC content. For example, ClgR regulates also the clp genes in C. glutamicum (Engels et al. 2004), S. lividans (Bellier et al. 2004), and Bifidobacterium breve (Ventura et al. 2005). The expression of ClgR is generally upregulated during heat stress (Stewart et al. 2002). Accordingly, the transcription of clp genes is also connected to the heat shock response. ClgR binds a palindromic sequence motif upstream of the clpP1P2 gene locus, as well as clpC in C. glutamicum. But transcription can be upregulated during heat stress independently of ClgR (Engels et al. 2004). Furthermore, ClgR is degraded in a ClpCP dependent manner. Upstream of the clpP1P2 operon an additional σECF promoter sequence is arranged, which is also able to activate the transcription of clpP1P2 and clpC in response to heat stress (Engels et al. 2004). A knockdown of clpP1 and clpP2 in Mycobacterium tuberculosis shows reduced growth and lack of replication after infection of macrophages (Carroll et al. 2011). The importance of ClpP1 and ClpP2 was confirmed by promoter replacement, showing that depletion of the genes causes cell death in vitro and during infection in mice (Raju et al. 2012). On the other hand, the over-expression of ClpP2 is toxic, indicating that both inhibition and activation are lethal (Ollinger et al. 2012). Nevertheless, ClpP2 is one of the ten most abundant proteins of Mtb (Schubert et al. 2013).
2.3.2 Architecture and mechanisms of Clp proteases
Clp protease particles function in conjunction with ATPase partners that bind to the ends of the ClpP cylinder. Degradation substrates are recognized by the controlling ATPases. ClpP serves as the proteolytic component of the complex and cleaves any polypeptide which is fed into the ClpP cylinder by the ATPase. The ClpX translocation rate during degradation is considerably slower than the ClpP14 cleavage rate of model substrates for E. coli. The small entry pores of the ClpP rings are closed by the ClpP N-termini (Thompson et al. 1994). These portals exhibit 10 Å restricted diameters and are located at both ends of the particle (Wang et al. 1997). The peptidase alone does not degrade proteins, because the multiple
20 2 Introduction
active sites are located inside the proteolytic chamber and are isolated from the surrounding solution. But sequestering the catalytic sites has an energetic price, due to connected substrate unfolding and translocation in the chamber for peptide hydrolysis. The active sites are located in three dimensions ~25 Å to the neighboring sites. A distance spanned by ~8 amino acids in an extended chain. The ClpP proteases (~200 amino acids) share the same architecture forming a cylinder with a sequestered chamber. This chamber is roughly spherical and can accommodate several hundred residues. The large interior chamber covers ~50 Å. The proteolytic core spans approximately 100 Å in height and diameter (Wang et al. 1997) and consists of two stacked rings each with seven subunits. Each ClpP monomer structure comprises six repeats of α/β-fold (αA/β1/β2, αB/β3/β4, αC/β5/β6, αD/β7/β8, αF/β10, and αG/β11). The ClpP monomers have a hammerhead structure, consisting of an N-terminal axial loop (residues 15-31 in E. coli), a hatchet-shaped subunit with a globular head domain, expanding along residues 32-138 and 172 -207, and a handle region (β strand 6 and α-helix E, residues 139-171) (Wang et al. 1997). The central E-helix bares a glycine-rich loop region and is essential for the inner ring contacts. Surprisingly, truncations in these regions do not result in dissociation of the double-ring, presumably due to a high degree of flexibility in the “handle” area. The two rings dissociate at low temperature and under high salt concentration. The handle region is accountable for the back-to-back interaction of the two heptameric, doughnut like rings. This interaction is particularly hydrophobic in nature. The contacts of the globular head stabilize the main body of the ring structure. The intra-ring interactions are biased by charge to charge contacts. The catalytic site is located in every monomer between the handle and the head domains (Gribun et al. 2005; Wang et al. 1997). In the active sites of the serine proteases ClpP1 and ClpP2, a catalytic triad with a central serine hydroxyl-group functions as a strong nucleophile, due to a histidine serving as proton acceptor and an aspartate positioning the histidine by electrostatic interactions (Dodson et al. 1998). The reactive serine attacks the carbonyl-group of the substrate, resulting in an intermediate which is stabilized by the oxyanion-pocket. The collapse of the intermediate results in the hydrolysis of the peptide bond and the release of the free amino-group. In the final rate-limiting step, the carbonyl group gets released from the enzyme (Ding et al. 1994). The hydrophobic S1 binding pocket in ClpP and the catalytic triad distinguish the favored amino acid characteristics in the P1 cleavage position. For E. coli ClpP, the specificity is not high, but non-polar residues in P1 are preferred. Whereas GFP degradation by ClpXP
21 2 Introduction
results in ~80% of peptides, cleaved after L, G, M, A, and Y (Baker et al. 2012; Thompson et al. 1994). Expressed Mtb ClpP1 and ClpP2 have no peptidase activity against model peptide substrates described for E. coli ClpP in vitro. Only ClpP1 and ClpP2 in combination exhibit peptidase activity and only in the presence of small activator peptides. One of these molecules is Z-LL-H which shows the highest known activation rate (Akopian et al. 2012). In Listeria monocytogenes the activity of the asparagine protease ClpP1 is increased 75- fold in the complex with ClpP2. Both proteases have different cleavage preferences, whereby ClpP1 favors small amino acids in substrate peptide P1 site (Zeiler et al. 2013). During degradation the peptide bonds are hydrolyzed, leading to the release of small peptide products with a typical size distribution of ~900 Da (8-9 peptides). These results favor a proteolysis mechanisms for ClpAP in which allosteric interactions between chaperone and protease only allow alternate translocation and proteolysis (Jennings et al. 2008). If the proteolysis mechanism might also be influenced by size-dependent escape of the products, or on a combination of size-dependent escape and a size dependent cleavage is not yet elucidated. The peptide product release after the cleavage is not entirely clear. Fragments might exit the same way they entered the lumen, namely via the axial pores (Thompson et al. 1994). Alternatively, peptides might escape through transiently formed exit pores of ClpP. It has been suggested that in a transient state the protease rotates into a compressed conformation, thereby opening exit pores in the ring interface (Gribun et al. 2005; Sprangers et al. 2005).
Figure 3: The protease ClpP in an active (pdb 1TYF) and an inactive (pdb 2CE3) conformation. Both structures reveal the overall barrel like tetradecameric double-ring shape. Furthermore, the inactive ClpP complex exhibits an equatorial rotation with pores in this axis.
22 2 Introduction
It has been shown that the ClpP particle can exist in different conformational states. Crystal structures describe two conformations (Figure 3; Table 4). The overall cylindrical structure is similar in both structures. The main difference results from the flexible handle region which is involved in the ring-ring interaction. A different angle between the handle and head region of each monomer generates an equatorial rotation leading to compressed shape. These compressed ClpP complexes are catalytically inactive. The active site which is located between the head and the handle domain is distorted. The distance between the histidine and the serine of the catalytic triad is increased to ~7 Å in contrast to the active conformation with 2.8 Å distance between the active site residues (Ingvarsson et al. 2007). It is not clear if the compact form is an early stage during complex assembly or a functionally relevant conformation during the protease cycle. This compressed conformation reveals 14 pores in the axis between both ClpP rings. The structures of active E. coli ClpP do not feature these pores (Kimber et al. 2010; Lee et al. 2010).
Table 4: Inactive- (in which residues 139-150 are typically disordered (Kimber et al. 2010)) versus active-structures
Compact inactive structures Active extended structures Streptococcus pneumonia E. coli (Gribun et al. 2005) (Wang et al. 1997) Mycobacterium tuberculosis Homo sapiens mitochondria (Ingvarsson et al. 2007) (Kang et al. 2004) Plasmodium falciparum Helicobacter pylori (Vedadi et al. 2007) (Kim et al. 2008)
In E. coli, the ClpP subunits are expressed in a preform (proenzyme) and first have to be auto catalytically cleaved at their N-termini to become active. The cleaved fragment is called propeptide. The removal of a propeptide was also observed for both ClpP versions of M. tuberculosis. For ClpP1, the propeptide is six amino acids long and for ClpP2 it has a length of 12 amino acids (Akopian et al. 2012). The N-terminus is located at the axial gates of the cylinder-shaped ClpP14 particle and can be detected in an “up” or “down” conformation (Alexopoulos et al. 2012). The “up” conformation can be observed in E. coli, human mitochondria, and S. pneumonia enzymes (Gribun et al. 2005; Kang et al. 2004). These studies revealed that the N-terminus forms a loop in which the first ~7 residues line the axial pore, while amino acids ~8-16 form a flexible loop, extending out of the pore. The “down” conformation was discovered in E. coli; this structure shows a less defined N-terminus, but the first 11 residues point into the pore (Bewley et al. 2006). The hydrophobic residues P4,
23 2 Introduction
V6, I19, L24, and F49 help to stabilize the down conformation and close the pore (Lee et al. 2010). Close to open transition accompanies interaction with the chaperone. The interaction influences the motion of these loops. The “up” conformation of loops correlates with an open state. The main part of ClpP 18-193 remains unchanged by this interaction (Effantin et al. 2010). Deletion of the first seven N-terminal residues of ClpP offers increased accessibility for peptides up to 30 amino acids in size through the constricted pore to the protease chamber (Jennings et al. 2008). The normal adjustment of the loops in “down” conformation and the resulting size limitation of the pore form a barrier which only peptides smaller than 5-10 amino acids can pass (Thompson et al. 1994).
2.3.3 The interaction between the ClpP particle and ATPases
The N-terminus of the sevenfold symmetric ClpP particle as well as hydrophobic patches at the surface of ClpP predominantly mediate the interaction with the hexameric chaperone rings (Gribun et al. 2005). The central core particle forms the proteolytic chamber, and the cognate regulatory particle flanks either end of the core, resulting in the symmetry mismatched chaperone-protease. The central interaction between Clp chaperone- and protease-ring is characterized by a 6:7 mismatch (Beuron et al. 1998). On the contrary, the hexameric ClpY/HslU unfoldase binds the two hexameric rings ClpQ/HslV without symmetry mismatch (Kessel et al. 1995; Sousa et al. 2000).
Figure 4: Interaction of ClpA with ClpP. The complex is stabilized by weak flexible interactions. Involved parts are the pore 2 loops and the IGF/IGL loops (blue) of the chaperone (pdb 1KSF) and hydrophobic patches (yellow) and the N-terminus (red) of the protease (pdb 1TYF).
24 2 Introduction
The fully assembled chaperone ring associates via flexible loops, containing the IGL/IGF motif, to hydrophobic patches on the surface of ClpP about 54 Å away from the axial pores (Figure 4). All six loops in the ATPase are required for strong ClpP binding and efficient degradation. Deletion of one IGF/L motif weakens the interaction dramatically around 40- fold and the deletion of two loops results in failure of binding. Certain weak interactions like the IGL binding maintain stable during degradation cycles, while other parts in the machinery undergo frequent conformational changes. For example, the association between the flexible pore-2 loops at the bottom of the chaperone and the N-terminal loops of the peptidase subunits are controlled by the ATP load of the ATPase (Martin et al. 2007). Subsequently, the conformational changes by ATP hydrolysis in the chaperone also enhance the proteolytic activity of the protease. Most information of the interaction between chaperone and protease were reported for the E. coli system. However, for the Mtb chaperone ClpC1, activity (Kar et al. 2008) and interaction with the protease ClpP2 have been reported (Barik et al. 2010).
2.3.4 Molecular details of Clp ATPases
The two Clp ATPases ClpX (426 aa and ~46 kDa) and ClpC1 (848 aa and ~93 kDa) are encoded in Mtb. The Mtb ClpC1 is upregulated after hypoxia along with ClgR, the Clp protease gene regulator (Sherrid et al. 2010). The protein ClpC2 (252 aa and ~26 kDa) shares the same name but exhibits no ATPase domain. The AAA+ (ATPases Associated with diverse cellular Activities) ATPases ClpX and ClpC1 are the motor components of the chaperone-protease complexes and form ring structures of six subunits, with one (ClpX) or two (ClpC1) nucleotide-binding domains (NBD), also named D1 and D2 (Figure 5). The single AAA+ domain of ClpX is homologous to the second AAA+ binding site of Class 2 ATPases. In Class 2 ATPases such as ClpC1, NBD1 and NBD2 form two stacked rings, the D1 and D2 tier. In the ClpP-assembled state the D2 tier caps the protease.
Figure 5: The structural organization of the Mtb Clp ATPases. ClpX possesses one ATPase domain in contrast to ClpC1 which contains two (NBD1 and NBD2; homolog structures in same green color). The IGL/IGF-loops (grey) of ClpX and ClpC1 mediate the interaction with the proteolytic particle. Both ATPases comprise one or two flexible N-terminal domains (red) involved in adaptor and substrate recognition. ClpX bears an N-terminal C4 binding motif which is required for dimerization.
25 2 Introduction
The Clp chaperone-proteases belong to the class of AAA+ ATPases which are an abundant class of nucleotide binding and hydrolysis proteins. The AAA+ ATPases offer energy to direct molecular remodeling including various processes (protein unfolding and degradation, vesicular fusion, peroxisome biogenesis, and the formation of membrane complexes) (Iyer et al. 2004). The P-loop is a conserved nucleotide phosphate binding motif, also referred to as Walker A motif (Saraste et al. 1990; Walker et al. 1982). The catalysis includes Mg2+ for the hydrolysis of the β–γ phosphate bond of the bound nucleotide (ATP or GTP). The energy of the ATP hydrolysis delivers the power for the reaction cycle of the chaperone-protease. The Clp chaperone-protease reaction cycle consists of initial tag recognition of the substrate by the ATPase ring. A fast unfolding process follows, combined with gate opening of the protease, hand in hand with translocation of the substrate into the ClpP protease and degradation (Figure 6) (Maglica et al. 2009).
Figure 6: Model of the bacterial chaperone-protease (grey; pdb 1TYF and 1KSF). Reaction cycle consists of degron recognition by ATPase, unfolding of the substrate (GFP in green (pdb 1B9C) and tag in red), translocation of the unraveled polypeptide chain into the protease chamber and rapid degradation into small peptides.
The degron, e.g. the ssrA-tag binds to different loops in the ATPase pore during the cycle. For Clp ATPases, the loops are located at the top, the middle, and lower parts of the central pore. ATP hydrolysis drives conformational changes within these loops, resulting in translocation by repetitive up and down movements and unraveling force that induces the threading of the denatured chain through the unfoldase pore (Hinnerwisch et al. 2005; Martin
26 2 Introduction
et al. 2008a; Wang et al. 2001). The upper loop discriminates as a filter, while the remaining loops offer binding sites deeper in the channel. The flexible diaphragm loop is located in the D2 ring and contains the GYVG motif. The conserved aromatic-hydrophobic GYVG-motif has a crucial role in translocation. Removal of tyrosine even in a few ClpX subunits results in slippage, frequent disruption of denaturation, and an increased ATP consumption (Martin et al. 2008b). Mutations in all loops disrupt the recognition of the degradation tag (Martin et al. 2007; Siddiqui et al. 2004). The protein sequence of ClpC1 possesses two N-terminal domains (NTD). The distal N- domains in particular feature the highest flexibility in an overall highly dynamic complex (Effantin et al. 2010). A flexible linker connects the N-domain with the NBD1 (Guo et al. 2002), enabling optimal substrate recruitment above the NBD1 ring. The Mtb ClpC1 has a long C-terminus which seems important for hexamerization (Bajaj et al. 2012). Clp chaperones with two ATP-binding domains assemble only when nucleotides are present. For ClpA, the D1 ATPase domain is required for the assembly and the nucleotide binding domain of the D2 tier transforms the energy from ATP hydrolysis into substrate unfolding. Beside the Walker A and B motif another motif is conserved in AAA+ ATPases, the “second region of homology” (SRH) which is involved in ATP hydrolysis in the NBD1 (Gamer et al. 1996). The SRH of ClpA includes the sensor 1 which is a polar residue and the arginine finger (R370) (Karata et al. 1999) of the active site. Both residues contact the ɣ-phosphate of the nucleotide. A ClpA mutation at position (R370K) failed to bind ClpP (Joshi et al. 2004).
ATP hydrolysis rates of the AAA+ motor components depend on the load state with its substrate. High rates of ATP hydrolysis can be observed when the enzyme is degrading an unfolded polypeptide backbone, in contrast to the degradation of a folded substrate, whereby ATP turnover is slower. The speed of ClpA alone along the substrate string ranges between kinetic rates of ~14 amino acids up to ~19 amino acids s-1 under saturated ATP concentrations (Rajendar et al. 2010). In complex with the proteolytic particle ClpP, the chaperone reaches rates of 50 amino acids s-1 (Kress et al. 2009). The steady state kinetic parameters for E. coli ClpX hydrolysis of nucleotide depend on the presence of ClpP and/or protein substrates (Burton et al. 2003; Kim et al. 2001). The hydrolysis rate under saturated ATP concentrations varies between 100-600 min-1 depending on the interaction with the protease (Baker et al. 2012).
27 2 Introduction
Each subunit of the hexameric ClpX ATPase consists of two parts, a small (65-314) and a large one (65-314) which are connected by hinges. These hinges are flexible. Rigid interactions in the ring are between one small domain and the large one of the clockwise neighbor subunit (Glynn et al. 2012). The orientation between these single ClpX blocks can vary significantly in crystal structures and the homo-hexameric ring appears asymmetric. In these structures only four ClpX subunits could bind nucleotides at the same time and two ClpX subunits were nucleotide free in each round of hydrolysis (Figure 7). The conformational motions due to hydrolysis are necessary for translocation, while keeping a sufficient robust architecture to force native protein structures to unfold (Glynn et al. 2009; Hersch et al. 2005; Stinson et al. 2013).
Figure 7: Composition of two homo hexameric chaperone rings of ClpX (single subunits in different colors) in nucleotide free (pdb 3HTE) and bound conformation (pdb 3HWS) which results in an allosteric structural motion of the ring.
ClpX translocates polypeptides of all properties including large, small, charged, or hydrophobic amino acids. The ATPase does not recognize special polypeptide features along the polypeptide chain (Barkow et al. 2009). ClpX even translocates polypeptide chains with internal disulfide bonds simultaneously through the narrow pore (Burton et al. 2001). The unfoldase translocates processively in one direction similar to other molecular motors that move along a linear track such as nucleic acid helicases and kinesin (Rajendar et al. 2010). Presumably, ClpX uses van der Waal’s contacts along the polypeptide chain (Baker et al. 2012). During translocation through the narrow pore, ClpX creates a pulling force that unravels the bound substrate which cannot fit in its native form through the small entry (Martin et al. 2008c). The diameter of the pores of Clp ATPases are even for small native proteins to narrow to pass through. ClpX moves 1-3 nm which are 5-8 amino acids per ATP.
28 2 Introduction
The translocation rate of the unfolded polypeptide chain can reach up to 80 aa/s under ATP saturation (Maillard et al. 2011). The ATPase uses the energy from ATP hydrolysis for pulling the polypeptide chain with a stall force of ~20 pN through the pore to reduce the structural stability of the native protein (Maillard et al. 2011). Each pulling generates ~3 kcal/mol of mechanical work. Most unfolding events are very fast with less than 1 ms between degron binding and unfolded substrate (Aubin-Tam et al. 2011). In fact, the interaction with the protease ClpP accelerates ClpX protein unfolding 2-3 times (Kim et al. 2000). The global thermodynamic substrate stability appears uncorrelated to its degradation properties. Indeed, destabilizing mutations close to the degradation tag facilitate the degradation and result in reduced ATP consumption (titin-127 variants require 20–500 ATP cycles and Arc-ssrA ~150 ATP molecules) (Burton et al. 2001). Findings favor a denaturation model with repeating rounds of a consistent unfolding force. The denaturation of the native substrate and especially the translocation of the polypeptide chain is the rate limiting step of ClpXP proteolysis (Kenniston et al. 2003). Besides a few exceptions, the overall degradation of all energy-dependent proteases is processive.
2.4 Pathogenesis and drug design
Although intracellular chaperone-proteases are usually not virulence or pathogenesis factors in themselves, they can contribute to or support pathogenesis by various mechanisms, like for example by regulating the gene expression of virulence genes. Moreover, the macromolecular chaperone-proteases influence the viability of bacteria (e.g. pathogens) by controlling competence, sporulation, the cell cycle as well as protein quality control in conditions which promote protein denaturation such as the host immune response during infection.
2.4.1 Role of chaperone-proteases during pathogenesis
In Yersinia sp., ClpX and Lon are required for the expression of the Type III secretion system by degradation of expression repressor YmoA (Mota et al. 2005). The Type III secretion system forms structures to inject effector molecules into the eukaryotic host cells, which enables the system to kill macrophages (Losada et al. 2005). YmoA also regulates the adhesion factor Invasin, facilitating the entry of Yersinia enterocolitica into host cells (Ellison et al. 2003; Isberg et al. 1987).
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In E. coli, ClpXP degrades the stress response σS (Hengge-Aronis 2002). In Salmonella enterica serovar Typhimurium, σS is an essential pathogenesis factor (Fang et al. 1992). A study showed that a Listeria monocytogenes clpP mutant lost its ability to infect mice (Gaillot et al. 2000), due to low concentrations of listerolysin O, a virulence factor involved in phagosome lysis. ClpC knockout of the same strain results in reduced expression levels of invasion genes inlA, inlB, and actA (Nair et al. 2000). Moreover, the chaperones ClpL and ClpC are important for mice pathogenesis of Streptococcus pneumonia (Hava et al. 2002; Polissi et al. 1998). A clpP knockout in S. pneumonia is avirulent (Ibrahim et al. 2005; Robertson et al. 2002). The ATPase ClpX is important for spa transcription. This results in different levels of Staphylococcal Protein A, an important surface protein (Frees et al. 2003). Additionally, ClpXP is involved in transcription of pathogenic genes, encoding hemolysin (an extracellular protease) in S. aureus (Butler et al. 2006; Frees et al. 2003).
2.4.2 Drug design
Nearly all bacterial pathogens have developed numerous defense mechanisms against antimicrobial treatment, causing the need for novel drugs to fight antibiotic resistance. The outbreak of S. aureus (MRSA) and totally drug resistant Mtb disfavor earlier results in treatment and control. The nightmare of a post-antibiotic period emerges in which our current antibiotics become ineffective against pathogenic species. But in principle almost all essential bacterial pathways could serve as therapeutic targets. Bacterial proteases are often indispensable for normal growth or virulence and offer diverse contact points for pharmacological agents. Different successful applications in numerous diseases target proteases. Typical drug treatments imply extracellular proteases, involved in blood pressure control or blood glucose level control (dipeptidyl peptidase). Furthermore, the drugs dabigatran (Pradaxa; Boehringer Ingelheim) and rivaroxaban (Xarelto; Bayer) target thrombin and factor Xa both necessary in blood coagulation (Raju et al. 2012). Different compound libraries were screened for inhibitory effects of ClpP, resulting in potential therapeutic compounds (Gersch et al. 2013). Moreover, docking experiments to hydrophobic pockets aim in the same direction (Tiwari et al. 2010). The activation of the protease ClpP by the small natural antibiotic acyldepsipeptide (ADEP) results in uncontrolled degradation of the proteome and causes cell death. Further attempts to identify molecules with the potential to activate ClpP revealed new compounds which are named Activators of Self- Compartmentalizing Proteases 1 to 5 (ACP 1 to 5). The activators exhibit drug-like characteristics with improved bioactivity and bactericidal effects (Leung et al. 2011). The 30 2 Introduction
chemical structures of the ACPs are unrelated to ADEP. But also the Clp ATPases have the potential to become drug targets. The natural compound cyclomarin for example targets ClpC1 of M. tuberculosis and is toxic for the organism (Schmitt et al. 2011).
2.4.3 The Clp protease activation molecule ADEP
It is possible to activate the degradation of the Clp protease in an unregulated way by acyldepsipeptide (ADEP) which results in cellular toxicity. This compound also activates the Mtb protease ClpP1P2.
A
B
Figure 8: Structural diversity of ClpP from B. subtilis as described by Lee et al. 2011. A) ClpP is on the left in a compressed conformation (PDB 3TT6), in the center structure ClpP is bound to ADEP which is highlighted in red (PDB 3KTI), and on the right ClpP is in an expended conformation (PDB 3KTG). The seven ClpP monomers in each ring are colored in a gray spectrum. A structure of the natural ADEP 1 as referred by Hinzen et al. 2006 is shown in B. The acyldepsipeptides differ at R1 (H, 3-F, and 3,5-F2) and R2 (C3H7 or C5H11).
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These small natural antibiotics are produced by Streptomyces hawaiiensis and mimic the binding of the chaperone by interacting with the same hydrophobic pockets on the surface of ClpP which normally serve as binding sites for the ATPase. The antimicrobial effect is caused by the disregulation of ClpP. A library of the isolated acyldepsipeptides (ADEP 1-4) was chemically optimized to increase the effect (Figure 8D) (Hinzen et al. 2006). In the absence of the chaperone, the acyldepsipeptide opens the N-terminal extensions of ClpP in a β-hairpin conformation and generates a 20 Å extend pore which triggers an uncontrolled degradation of proteins, causing cell death (Brötz-Oesterhelt et al. 2005; Lee et al. 2010; Li et al. 2010) (Figure 8B). ADEPs bypass the requirement for an unfoldase. The binding of ADEP1 in vivo redirects the activity away from substrates and toward various nascent polypeptides, such as those emerging from the ribosome, resulting in inhibition of cell division and leading to cell death (Kirstein et al. 2009). Furthermore, ADEP induces the dissociation of the complex between ClpC and ClpP that otherwise forms in the presence of the adaptor protein MecA of B. subtilis (Brötz-Oesterhelt et al. 2005; Kirstein et al. 2009) and blocks ClpAP interaction while activating the protease allosterically like ClpA (Li et al. 2010). The effect of the addition of ADEP to B. subtilis cells appears similar to the previous observed clpP mutant phenotype (Molière et al. 2009). Moreover, ADEP appears to have antimicrobial activity for Mtb (Ollinger et al. 2012).
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3 Aim of the study
Mycobacterial genomes have been found to encode two ClpP protease subunits (clpP1, clpP2) and for Mtb both genes were found to be essential. Both ClpP subunits in Mtb had largely unknown functions and their assembly state had not been described. It was also still unclear why the gene duplication of clpP1P2 had occurred. The first aim of the thesis was the recombinant production of the ClpP1 and ClpP2 subunits either from a coexpression plasmid or by individual expression. Analysis of the composition and assembly of the ClpP peptidase cylinder from the ClpP1 and ClpP2 subunits should provide insight which particles exist, homooligomeric, heterooligomeric, or both. Further, biochemical analysis was planned to investigate the peptidolytic activity of the assembled ClpP particles and the interaction with the potential ATPase partners. Two such AAA+ ATPases are encoded in the Mtb genome, ClpX and ClpC1. A set of potential adapter proteins (ClpS and ClpC2) could be found in the Mtb H37Rv genome, but little information was available on substrates of the ClpP complex.
Another aspect of this thesis was the elucidation of the regulatory influence of post- translational modifications on the ClpP particle composition and activity, since components of the system were identified in proteomic studies cataloguing phosphorylation sites (ClpP1) as well as modification sites for prokaryotic ubiquitin-like protein Pup (ClpP2). The influence of phosphorylation and pupylation on particle assembly and activity was investigated in vitro.
For in vitro reconstitution of the Clp dependent degradation system of Mtb, all components (i.e. the proteases ClpP1, ClpP2, the chaperones ClpX and ClpC1 and the potential adaptors ClpC2 and ClpS as well as a set of substrates) were cloned and heterologously expressed in either E. coli or M. smegmatis. The corresponding proteins were purified and tested for their assembly state by analytical gel filtration. A photometric assay was employed to study the peptidase activity of ClpP1P2 against various substrates. The chaperone partners ClpX and ClpC1 were tested for their ATPase activity and interaction behavior with the ClpP subunits.
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4 Material and methods
4.1 Plasmids
The single protease-coding genes clpP1 and clpP2 and the fusion constructs clpP1P2 and clpP2P1 were originally cloned by Frank Imkamp in pETDuet-1 (Novagen) vectors. These T17E constructs had an uncleavable C-terminal H4-tag. The phosphomimetic mutants (P1 and P1T17A) were generated by site-directed mutagenesis PCR of clpP1 in the pETDuet-1 vector. The clpP2 wild type, clpP1, and clpP2 active site variants were cloned with Nco I and Hind III in pPROEX HTb plasmids (Invitrogen). Therefore, wild type ClpP1 and ClpP2 were mutated by Quick Change PCR according to the Stratagene protocol to ClpP1S98A and S110A ClpP2 (with a TEV cleavable N-terminal H6-tag). Also the N-terminal deletion constructs (Δ7ClpP1, Δ11ClpP2, Δ13ClpP2, and Δ18ClpP2) were generated of full length ClpP1 and ClpP2 which were cloned in pPROEX HTb plasmid. All the clp genes were designed with the start codon ATG encoding methionine instead of the Mtb start codon valine. These constructs had an N-terminal TEV cleavable H6-tag. Wild type clpX and the deletion constructs of the first 62 amino acids were cloned in pET24 via Nde I and Sac I with a C-terminal TEV cleavable GFP H6-tag. In addition, ClpX was cloned with an uncleavable C-terminal Strep-tag in the pET24. An alternative cloning strategy was chosen for clpC1 which included fx-cloning with only one restriction enzyme Sap I. The gene was cloned by Jürg Laederach in pINITIAL. Afterwards the gene clpC1 was transferred in a p7X with a PreScission cleavable C-terminal H10-tag (Geertsma et al. 2011). Furthermore, the gene clpC2 was cloned in a mycobacterial expression vector pMyC with an uncleavable C-terminal H6-tag. The Mtb ClpS adaptor (Rv1331) was cloned in pET16b expression vector (Invitrogen) with an uncleavable N-terminal H10-tag. The Clp substrates malate dehydrogenase (Rv1240) (MDH) was fused to a C-terminal Mtb ssrA-tag (AADSHQRDYALAA) and cloned with an N-terminal TEV cleavable H6-tag in pET20 (modified by Cyrille Delley). The genes Rv2308, Rv2961, and Rv3832c were cloned with BamH I and Hind III into a modified pET20 vector (Novagen) with H6-Thioredoxin-TEV- tag. In addition, a GFPuv with a C-terminal Mtb ssrA-tag was generated using the restriction enzymes Nde I and BamH I for the pET20b expression vector (Invitrogen). The adenylate kinase (Adk) encoding gene RV0733 and the gene RV0543c encoding a ClpS-like protein were cloned in pET24 with a C-terminal TEV cleavable GFP H6-tag.
34 4 Material and methods
4.2 Expression and purification of proteins
The ClpP1 and ClpP2 plasmids were transformed into E. coli BL21(DE3) (Invitrogen) cells.
The cells were grown in LB-medium at 37 °C to OD600 and induced with 0.1 mM isopropyl β-D-1-thiogalactopyranoside (IPTG). Proteins were expressed at 25 °C overnight. The harvested cells were resuspended in lysis buffer containing 50 mM Tris-HCl pH 7.5, 300 mM
NaCl, 10% (v/v) glycerol, 5 mM β-mercaptoethanol, 20 mM imidazole, 3 mM MgCl2, and DNAase (10 U/l of growth culture). The recombinant proteins were purified by Ni2+-affinity chromatography (HiTrap IMAC HP, GE Healthcare) in the same buffer and eluted with a breakage buffer containing 150 mM imidazole. The TEV cleavable N-terminal H6-tag of Δ7ClpP1, ClpP1S98A, Δ11ClpP2, Δ13ClpP2, Δ18ClpP2, and ClpP2S110A were digested by TEV-protease (Invitrogen) overnight at 4°C in a dialysis buffer (50 mM Tris-HCl pH 7.5, 300 mM NaCl, 10% (v/v) glycerol, and 5 mM β-mercaptoethanol) to remove the imidazole. The TEV-protease was added in a molar ratio of 1:50 to the target protein. The TEV-protease and the histidine-tags were removed by a second Ni2+-affinity chromatography.
The expression vectors of the ClpX chaperone variants were transformed in E. coli
BL21(DE3) (Invitrogen) cells. The cells were grown in LB-medium at 37 °C to OD600, induced with 0.1 mM isopropyl β-D-1-thiogalactopyranoside (IPTG). The ClpX variants were expressed at 16 °C for 16 hours. The cells of the ClpX expression with a C-terminal TEV cleavable GFP histidine-tag were dissolved in lysis buffer containing 50 mM Tris-HCl pH 7.5, 300 mM NaCl, 10% (v/v) glycerol, 5 mM β-mercaptoethanol, 20 mM imidazole, 12.5 μM PepstatinA, 1 mM PMSF, protease inhibitor cocktail (20 μl/l of growth culture, Sigma), 3 mM MgCl2, and DNAase (10 U/l of growth culture). ClpX was further purified by Ni2+-affinity chromatography (HiTrap IMAC HP, GE Healthcare) in the same buffer and eluted with a breakage buffer containing 150 mM imidazole. The GFP histidine-tag was cleaved off by the TEV-protease (Invitrogen) (in a molar ratio of 1:50 substrate) overnight at 4°C in dialysis buffer (50 mM Tris-HCl pH 7.5, 300 mM NaCl, 10% (v/v) glycerol, and 5 mM β-mercaptoethanol). A second Ni2+-affinity chromatography was performed to remove the GFP histidine-tag and the TEV protease. The cell pellets of the ClpX expression with a C-terminal fused Strep-tag were resuspended in breakage buffer 50 mM Tris-HCl pH 7.5, 300 mM NaCl, 10% (v/v) glycerol, 5 mM β- mercaptoethanol, 12.5 μM PepstatinA, 1 mM PMSF, protease inhibitor cocktail (20 μl/l of growth culture, Sigma), 3 mM MgCl2, and DNAase (10 U/l of growth culture) and cracked by
35 4 Material and methods
sonification. The recombinant protein was purified by Strepcolumn (StrepTrap HP, GE Healthcare) and eluted with breakage buffer supplemented with 2.5 mM Streptavidin.
The chaperone ClpC1 plasmid was transformed into E. coli BL21(DE3) (Invitrogen) cells and the protein was expressed under the same conditions as the chaperone ClpX at 16 °C. The cells were harvested, centrifuged, resuspended in lysis buffer as described for ClpX, purified by Ni2+-affinity chromatography in the same buffer and eluted with a breakage buffer containing 150 mM imidazole. The PreScission cleavable C-terminal H10-tag was cleaved off by PreScission protease (ratio 1 U/100 μg substrate) overnight at 4 °C in dialysis buffer (50 mM Tris-HCl pH 7.5, 300 mM NaCl, 10% (v/v) glycerol, and 5 mM β-mercaptoethanol). 2+ The H10-tag and the PreScission protease were removed by a second Ni -affinity chromatography.
The plasmids of the adapter proteins ClpS (Rv1331), ClpS-like (Rv0543C), and also ClpC2 were transformed in M. smegmatis MC2155. The cells were grown at 37 °C in Middlebrook 7H9 medium (Difco) and induced with 0.2% (w/v) acetamide. Expression was performed at 25 °C overnight. The harvested cells were dissolved in the previously described lysis buffer. The recombinant proteins were further purified by Ni2+-affinity chromatography (HiTrap IMAC HP, GE Healthcare). The purified proteins were eluted at concentrations of 150 mM imidazole. The ClpS like protein was diluted in dialysis buffer and the TEV cleavable GFP histidine tag was cleaved off and removed as described for ClpX.
The plasmids of the potential ClpX substrates MDH-ssrA, Rv2308, Rv2961 and Rv3832c were transformed into E. coli BL21(DE3) (Invitrogen) cells and the proteins were expressed under the same conditions as the chaperone ClpX. The cells were harvested, centrifuged, resuspended in lysis buffer as described for ClpX. The further purification by Ni2+-affinity chromatography and digestion of the N-terminal TEV cleavable H6-tag of MDH-ssrA as well as for Rv2308 and Rv3832c the H6-Thioredoxin-tags followed the same protocol as described for the ClpP variants. Finally, the TEV protease and the affinity tags were removed by a second Ni2+-affinity chromatography. The ClpX substrate GFP-ssrA was purified as described by Kress et. Al. The heat stable protein was purified by heat denaturation (30 min at 65 °C) and ion exchange chromatography (Q-Sepharose Fast Flow and Source30Q).
36 4 Material and methods
All protein constructs were finally purified by size-exclusion chromatography. Due to the size of the proteins, they were applied on a Superdex 200, Superdex 75, or Superose 6 column (GE Healthcare) and stored in 50 mM Tris-HCl pH 7.5, 150 mM NaCl, 10% (v/v) glycerol, and 1 mM DTT. The protein identities were examined by ESI mass spectrometry. All proteins were finally concentrated by concentrators with 3.5, 10, or 30 kDa cut-off size and concentrations were determined via the specific protein absorbance at 280 nm (the molar extinction coefficients are listed in the appendix).
4.3 Preparation of ClpP1P2 complexes
Equimolar ratios of ClpP1 (60 μM) and ClpP2 (60 μM) were mixed for the formation of the active mycobacterial ClpP complex. The subunits were incubated for 60 minutes in 50 mM Tris-HCl pH 7.5, 150 mM NaCl, 10% (v/v) glycerol, and 1 mM DTT at room temperature (25 °C). The assembly state of the complex was followed by analytical gelfiltration (Superdex 200 24 ml; GE Healthcare) and analysis of the elution fraction on 15% SDS-PAGE. The molecular masses were verified by mass spectroscopy. For the comparison of Mtb and E. coli ClpP. 300 μM ClpP1ClpP2 were compared with 60 μM E. coli ClpP by analytical size exclusion chromatography (Superose 6). 150 μM of the deletion constructs Δ11ClpP2, Δ13ClpP2, and Δ18ClpP2 were incubated with 150 μM Δ7ClpP1 and the assembly states were analyzed on a Superdex 200 column. ClpP1 (15 μM) and ClpP2 (15 μM) as well as the complex of ClpP1 (15 μM) and Δ11ClpP2 (15 μM) were preincubated with 1 mM activator (Z-LL-H) for two hours at 25 °C and compared by size exclusion chromatography (Superose 6) at RT.
4.4 Assembly of the complex between protease and chaperone analyzed by analytical gelfiltration
To monitor complex formation of 2 µM preprocessed ClpP1P214 and 4 µM ClpC16 analytical size exclusion (GE Healthcare) runs on Superose 6PC 2.4 ml were performed at 25 °C and compared with E. coli ClpP (2 µM) and ClpA (4 µM). Therefore, 30 µL samples of the proteins were incubated for one hour and 30 minutes at room temperature. The chaperones ClpX
(1.5 µM) or ClpC1 (1.5 µM) were incubated for one hour with 20 mM MgCl2 and 0.75 mM ATPɣS or 1 mM ATP. Moreover, the column was equilibrated in gelfiltration buffer, containing
37 4 Material and methods
50 mM Tris-HCl pH 7.5, 150 mM NaCl, 10% glycerol, 1 mM DTT, 20 mM MgCl2, and 0.75 mM ATPɣS. The absorbance was measured at 227 nm.
4.5 Crystallography
To achieve the crystallization of ClpP1 and ClpP2, the proteins were purified in a minimal buffer containing 20 mM Tris-HCl pH 7.5, 75 mM NaCl, 10% glycerol, and 5 mM DTT. The initial protein concentrations varied between 7-10 mg/ml. For co-crystallization, traces were supplemented with 3 mM activator Z-LL-H (PeptaNova). The crystallization was performed at 26 °C in sitting-drop vapor diffusion plates. 1 µl protein solution was added at an equal volume of 1:1 to the reservoir solution (0-20% PEG 3500, 100 mM BisTris Propane pH 8-9, 10% glycerol, and 100 mM lithium sulfate or sodium sulfate). First crystals of ClpP2 grew in 0.1 M sodium acetate trihydrate pH 4.6 and 8% PEG 4000. The fine screen contained conditions between 0 and 10% polyethylene glycol (PEG) 4000, 6000, or 8000, and 0.1 M sodium acetate trihydrate pH 5-6. Different cryo-conditions were used, including 20% PEG 500, 30% MPD, or 20% glycerol to protect the crystals during freezing.
4.6 Data collection, structure determination, refinement and alignment
Data sets of ClpP1 and ClpP2 were collected at beamline X06Sa of the Swiss Light Source (Paul Scherrer Institute, Villigen, Switzerland). Data indexing and integration was performed with XDS (Kabsch, 2010). The program CCP4 was used for scaling, merging of diffraction data, and refinement. Structure determination was carried out with Coot (Emsley et al. 2010). Molecular graphics were created in PyMOL (http://www.pymol.org/). For the structural alignment, the structure of ClpC2 was predicted by Phyre2 and aligned in PyMOL with ClpC1 N-domain (PDB 3WDB). Other alignments were performed by UniProt using the Clustal distance matrix and neighbor joining algorithm for pairwise alignment which was visualized with Jalview in ClustalX colors. The protein sequences were taken from the TubercuList database and the sequences of the protein constructs were visualized by Expasy ProtParam tool. The masses and the molar extinction coefficients were also calculated by using the Expasy ProtParam tool.
38 4 Material and methods
4.7 ClpP peptidase activity
The peptidase activity of ClpP1P2 was tested independently of the chaperone ClpX or ClpC1 for small fluorophoric peptide-substrates. The used peptides Ac-DEVD-AMC, Boc-VLK- AMC, Glt-AAF-AMC, Suc-GGL-AMC, and the ClpP1P2 substrate Z-GGL-AMC (Akopian et al. 2012) were all commercially available (Bachem, PeptaNova, and Sigma). The structures of the peptide substrates were visualized by using ChemSketch. The peptides were dissolved in DMSO. The cleavage between the peptide and the AMC entity caused an increase in fluorescence. Therefore, the excitation wavelength was 380 nm and the emission was set to 440 nm. The peptidase activity of ClpP1P2 was calculated by using a calibration curve of AMC. The used ClpP1 and ClpP2 protein constructs were incubated with 0.75 mM activator overnight to complete the processing reaction. The complexes were purified by size exclusion chromatography (Superdex 200) to remove the propeptides and the activator.
For the peptidase assay, 0.5 µM preassembled ClpP1P214 were incubated with 0.1 mM peptides in 50 mM Tris-HCl pH 7.5, 150 mM NaCl, 10% glycerol, 1 mM DTT, and 6% DMSO (in higher concentrations the protein was unfolded, observed by CD spectra). The reaction was measured in addition of 0.75 mM activator Z-LL-H (Akopian et al. 2012).
4.8 Processing of ClpP1ClpP2
In the processing assay equimolar concentrations (3 µM) of ClpP1 and ClpP2 were incubated at 25 °C. The reaction buffer (buffer R) contained 50 mM Tris-HCl pH 7.5, 150 mM NaCl, 10% glycerol, and 1 mM DTT in the addition or the absence of 0.75 mM activator Z-LL-H. The samples were further analyzed on 15% SDS-PAGE. The preprocessed ClpP1P2 which was incubated overnight in the presence of 0.75 mM Z- LL-H, was purified on gelfiltration S200 column or by multiple rounds with centricons (10000 MWCO) to remove the activator.
4.9 ATPase Assays
The ATPase activity of the chaperones ClpX and ClpC1 in combination with the different proteases ClpP1 ClpP2 was measured in a continuous spectroscopic assay. The release of phosphate during the hydrolysis of ATP to ADP was measured by the phosphorylation of 7- methylinosin by purine nucleosidase phosphorylase (PNPase; Sigma). This coupled reaction resulted in hypoxanthine and the depletion of absorbance at 291 nm (Rieger et al.
1997). The production of Pi could be calculated according to a calibration curve with
39 4 Material and methods
KH2PO4. The reaction was carried out in reaction buffer 50 mM Tris-HCl pH 7.5, 150 mM NaCl, 10%
(v/v) glycerol, and 1 mM DTT referred to as buffer R with the ATPase (0.2 µM Mtb ClpXhex,
0.2 µM Mtb ClpC1hex, or 0.2 µM E. coli ClpXhex) at 23 °C. The measurement was initiated by 5 mM ATP, 0.1-1.5 U/mL PNPase, and 1 mM 7-methylinosine. The ATPase assay was recorded in a Cary UV-Vis spectrophotometer (Varian) with an absorbance cuvette (1 cm path length). The measured ATPase reactions were buffer corrected by the following equation: (exp curve - buffer) + 2.16 (the measured absorbance at 291 nm at the start of the reactions).
4.10 Analysis of the Mtb H37Rv proteome for ClpX degrons
The proteome screen of the Mtb strain H37Rv (UniProt database) for known ClpX recognition sites contained the C-terminal tags LAA, VAA, NVA, RRKKAI, RAKKVA, and RVKHPA (Baker et al. 2011). Therefore, we copied the complete mycobacterial proteome in FASTA format into Microsoft Word 2010 and screened the file by using the search function for the C-terminal sequence + Word wrap.
4.11 Degradation of ssrA-substrates, FR-linker proteins and Pup conjugated proteins by Clp chaperone-proteases
1 µM chaperone ClpXhex or ClpC1hex with or without 3 µM of the adaptors ClpC2 and ClpS
(Rv1331) and 1 µM ClpP1P214 were preincubated for one hour at 23 °C in buffer R. For the assay, the preincubated chaperone-proteases were mixed with 3 µM substrate (GFP-ssrA, MDH-ssrA or λR-ssrA (purified according to Kress et al. 2009b), FR-GFP (purified according to Kress et al. 2009b), or (GFP-Pup or Adk-Pup)). Furthermore, the degradation assay included an ATP-regeneration system (Sigma: 40 mM creatine phosphate and 20 U/mL phosphocreatine kinase). Degradation was initiated with 5 mM ATP and samples were taken at various time points during overnight incubation. The aliquots were quenched in SDS sample buffer and heat treated at 95 °C. Equivalent volumes of the degradation reaction were monitored by SDS-PAGE.
4.12 Peptide inhibition of ClpXP degradation
The degradation reaction contained the same reaction preparation described for MDH-ssrA in the presence of 1.5 mM peptide (Z-GGL-AMC, Suc-LLVY-AMC, FR (N-end rule), LGF, RGLGFGA (hepta LGF), or NPSITRD (pore2)). Aliquots were taken at time point 0, after 40 4 Material and methods
one, and two days and the samples were analyzed by SDS-PAGE.
4.13 Pupylation of ClpP2 and Adk
The reaction mixture contained 0.5 μM PafA, 15 μM Pup, 5 μM Pup substrate (ClpP2 or
Adk), 5 mM ATP, 20 mM MgCl2, and ATP regeneration system (40 mM creatine phosphate and 20 U/mL phosphocreatine kinase). The pupylation was performed overnight at 37 °C in reaction buffer (50 mM HEPES-NaOH pH 7.5, 150 mM NaCl, 10% glycerol, and 1 mM DTT). The pupylation completeness of the proteins was analyzed on 15% SDS-PAGE. The conjugated ClpP2-Pup and Adk-Pup were purified by size exclusion chromatography and Superdex 200 peaks were analyzed on 15% SDS-gel. Furthermore, for ClpP2 multiple pupylation reaction rounds were performed. The pupylation of ClpP2 was further analyzed densitometrically by ImageJ to determine the produced ClpP2.
4.14 Degradation of ClpP2-Pup by the proteasome
For the degradation assay of ClpP2-Pup, Pup-Strep was applied to the pupylation reaction and was further purified by Strep affinity chromatography. The elution buffer contained Tris- HCl pH 7.5, 150 mM NaCl, 10% glycerol, and 1 mM DTT supplemented with 2.5 mM Streptavidin. The purified Strep-tagged ClpP2-Pup was degraded by mycobacterial Mpa, PrcA, and PrcB. The reaction mixture also contained the ATP regeneration system with 40 mM creatine phosphate and 20 U/mL phosphocreatine kinase. Samples were quenched with SDS, heat inactivated, and further analyzed by SDS-PAGE and Coomassie staining.
41
5 Results
5.1 ClpP1P2 assembled in a tetradecamer
To study the properties of the proteases ClpP1 and ClpP2 of Mtb, we expressed the single Mtb ClpP1 and ClpP2 subunits separately in E. coli RosettaTM (DE3) cells under the control of an IPTG inducible lacUV5 promoter, in case of ClpP1 fused to a C-terminal H4-tag and in case of ClpP2 with a C-terminal H4-tag or an N-terminal TEV-cleavable H6-tag. Alternatively, we coexpressed ClpP1P2 with one subunit fused to a H4-tag followed by the other untagged. The proteins were purified and yielded high amounts of pure protein (~15-20 mg/L of culture). Protein identity was confirmed with electrospray ionization mass spectrometry (ESI- MS) analysis. The masses agreed within the experimental error (+/- 0.01%) with the theoretical masses according to the amino acid sequences listed in the TubercuList database (for ClpP1 21675.6 Da and for ClpP2 23507.7 Da).
Figure 9: Purification of coexpressed ClpP1 and ClpP2 subunits through Ni-affinity chromatography. SDS-PAGE analysis of purification fractions (1 lysate, 2 wash, and 3 elution) on 15% SDS-PAGE and Coomassie stained. The elution (3) contained on the right additionally 75 mM imidazole beside the 125, 150, and 175 mM imidazole of both purifications. In the gel on the left ClpP2 was purified via its uncleavable C-terminal H4-tag and on the right ClpP1 carried the H4-tag.
From coexpression in E. coli BL21(DE3) (Invitrogen) cells (Figure 9), we could only purify the tagged subunits and not the complex of ClpP1P2. The corresponding counterpart was located in the wash. However, together with the purified ClpP2 two additional bands were visible on 15% SDS-PAGE analysis. N-terminal sequencing of the elution mixture resulted in 90% full length ClpP2 carrying the wild type N-terminus as well as two N-terminally truncated forms, cleaved before position 13 and 18. We did not obtain this processing in the separately expressed subunits.
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Nevertheless, the purified Mtb ClpP1P2 complex did not cleave the model substrates as described for Clp homologs from other organisms. Either as single subunits or in the complex and independent of the N-terminal length, ClpP1P2 possessed no intrinsic peptidase activity (data not shown), until the group of Goldberg discovered by chance a group of peptides which activate the Mtb protease. The data demonstrated that only the addition of activator, such as Z-LL-H to the complex stimulated the peptidase activity of
ClpP1P2 to measurable rates (Akopian et al. 2012).
Figure 10: The ClustalX alignment included six closely related a) ClpP1 with 30 % identity (60 identical positions of 200 amino acids) and b) ClpP2 with 36 % identity (80 identical positions of 210 amino acids) from Actinobacteria. Three Mycobacteria (M. tuberculosis (MTB), M. leprae (Mle), and M. smegmatis (Ms)), two Corynebacteria (C. glutamicum (Cglu) and C. diphtheria (Cdip)), and Nocardia farcinica (Nfa) were compared. Alignment was visualized by Jalview.
The alignment of actinobacterial ClpP1 or ClpP2 homologs revealed a high overall homology (Figure 10) but diverse N-termini. Therefore, even closely related ClpP2 varied in the composition and the length of the N-termini between three and 16 polypeptides. Moreover, M. smegmatis ClpP1 possesses a five amino acid long C-terminal extension.
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Figure 11: Analysis of ClpP assembly state by size exclusion chromatography on a Superdex 200 column. a) The oligomeric state of Clp1 (dashed line), ClpP2 (grey line), and ClpP1ClpP2 (black line) was analyzed using analytical gelfiltration at RT. Fractions of the peaks were applied on a 15% SDS-gel. b) Samples of a) were further used for mass spectrometry. Additionally, the sequences of ClpP2 and the N-terminal deletion constructs of ClpP2 (green) were aligned.
The assembly state of ClpP1 and ClpP2 as well as ClpP1 mixed with ClpP2 was assessed using size exclusion chromatography on a Superdex 200 resin. The single subunits ClpP1 or ClpP2 eluted at positions corresponding to around 150 kDa in mass, which corresponds to about half of the mass of the E. coli ClpP double-ring (Figure 11a). This suggests that ClpP1 and ClpP2 each is assembled into 7-membered homooligomeric single rings. Due to its higher molecular mass ClpP2 (2 kDa/monomer) (11 ml) eluted earlier than ClpP1 (12 ml). In a third run equal amounts of the two subunits were mixed and incubated together for one hour at RT prior to application onto the gelfiltration column to allow for assembly of heterooligomers. The mixed ClpP1P2 formed higher oligomeric states which eluted in a main peak with two residual minor peaks at the positions of the 7-membered single rings. The newly formed complex displayed the same elution position (Figure 12) as E. coli ClpP. Therefore, the elution peak of the protease corresponds to a complex mass of 300 kDa, presumably arranged into a tetradecamer and assembled from the two heptameric single rings. The elution peak yielded a complex with 1:1 ratios of ClpP1 and ClpP2 as shown by SDS-PAGE analysis (see gel slice above elution profile). The elution samples of the three runs were additionally analyzed by mass spectrometry (Figure 11b). The analysis of the complex revealed three mass peaks, one corresponding to full length ClpP2, one
44 5 Results
corresponding to full length ClpP1, and the third peak corresponding to ClpP2 lacking the first 12 residues (22761 Da). Taking into account the mass for the H4-tag at the C-terminus, the TubercuList mass for mature ClpP2 (22212 Da) closely matched processed ClpP1 (22145 Da) for molecular weight.
Figure 12: Comparison of Mtb and E. coli ClpP complexes by analytical size exclusion chromatography. Superose 6 size exclusion chromatography was performed at room temperature to follow the association of mixed ClpP1P2 (black) complexes. For the gelfiltration runs equimolar amounts of ClpP1 and ClpP2 were incubated for one hour at 25 °C and same treated E. coli ClpP (blue) was used as a control as well as the standard protein masses.
Figure 13: The ClpP1P2 complex assembled in 60 minutes at room temperature. In the comparison of complex assembly over the time, at 0 (grey), 15 (black) 30 (light grey), and 60 (dark grey) minutes by analytical size exclusion chromatography Superdex 200, the complex of ClpP1P2 assembled in 60 minutes. 60 μM of the single subunits ClpP1 (dashed line) and ClpP2 (dashed line) and the standard protein masses were used as controls.
The complex assembled in 60 minutes (Figure 13). Even without incubation, assembly occurred during the gelfiltration run. Experiments with concentrations of ClpP1 fivefold higher than ClpP2 or vice versa resulted in the same amount of complex (data not shown), supporting the formation of a particle that is composed of ClpP1 and ClpP2 in a strict 1:1 stoichiometry.
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One possible role for the N-terminal propeptides could be in support of assembly of the double-ring particle. To address the function of the N-terminus in complex formation, we cloned mature versions of ClpP1 and ClpP2 (Δ7P1, Δ11P2) and purified them. Despite the information from the mass spectrometry that 12 N-terminal residues are cleaved off, Δ11P2 was cloned, because Δ12P2 starts with an arginine and exhibited solubility problems. Purified Δ7P1 eluted at a size of ~150 kDa on the analytical size exclusion chromatography, indicating that it also formed a heptameric single ring. When mixed with full length ClpP2 it displayed the identical assembly behavior as full length ClpP1, forming the Δ7P1P2 double- ring. The assembly equilibrium could be shifted toward single rings when incubation and gel filtration took place at 4 °C, which resulted in the individual single rings running separately. These results suggest that hydrophobic interactions between the two rings (data not shown) are the main driving force underlying P1P2 assembly.
Figure 14: ClpP2 deletion constructs failed to bind ClpP1 monitored by analytical size exclusion chromatography Superdex 200 at RT. Complexes of Δ7ClpP1, Δ11ClpP2 (green), Δ13ClpP2 (purple), and Δ18ClpP2 (red) in equal concentrations were preincubated for one hour at 25 °C. The void volume and the standard protein masses are labeled in the figure.
When Δ7P1 was incubated with Δ11P2, the Δ7ClpP1Δ11P2 double-ring particle was not formed (Figure 14). This would suggest that the N-terminus of ClpP2 is important for the interaction with ClpP1. The purified variants of ClpP2 with N-terminal deletions resulted in higher oligomeric states which underscore the second important function of these residues (Figure 14). Two peaks eluted from the gelfiltration column for the mixed complexes of the ClpP1P2
46 5 Results
truncations. The broad peak of the ClpP2 constructs comprised an inhomogeneous complex formation with different oligomeric forms. In the literature, synthetic peptides were described which enhance the peptidolytic activity of Mtb ClpP1P2 in vitro, referred to as “activators”. These molecules should disassemble
P114 and P214 and assist in the assembly of the mixed complex, consisting of a P17 and a
P27 ring (Akopian et al. 2012). However, treatment of the complexes with an activator for two hours did not result in disassembly or a new formation of the processed or mature or N-terminally truncated ClpP1P2 delta complexes, compared to full length ClpP1P2 (Figure 15). Furthermore, we did not obtain a positive effect in complex formation for full length ClpP1P2 in the presence of the small molecule (Z-LL-H) at room temperature.
Figure 15: Assembly comparison of native ClpP1P2 (black) with ClpP1 Δ11ClpP2 (green). For analytical size exclusion chromatography on Superpose 6 at RT, complexes in equimolar concentrations were preincubated for two hours at 25 °C in the presence of 1 mM activator (Z-LL-H). Additionally, the gelfiltration buffer contained 1 mM Z-LL-H. The standard protein masses are labeled in the chromatogram.
5.2 Crystallization and structural comparison of Mtb ClpP1 and ClpP2
While several structures are available for homologs from other bacteria, for the Mtb system only a structure for ClpP1 had been solved. This structure shows some characteristics of an inactive ClpP particle with a compact overall form, including pores in the equatorial axis (supposed to be peptide product exits). No structure of the mixed Mtb Clp protease complex consisting of the two different subunits ClpP1P2 has been described in the literature. To solve the structure of a mixed ClpP1P2 particle we set up sitting drop crystallization screens with ClpP1P2 in equal amounts. As controls we set up drops of the individual subunits ClpP1 and ClpP2 under the same conditions to identify wells that only produced crystals with both subunits present. Optimized conditions of presumed ClpP1P2 crystals contained 0-20%
47 5 Results
PEG 3500, 100 mM BisTris Propane pH 8-9, 10% glycerol, and 100 mM lithium sulfate or sodium sulfate. Crystals under these conditions grew to a length of 150 µm and a width of 100 µm, but further analysis revealed that the complex disassembled under these conditions and only ClpP1 crystallized.
Figure 16: Crystals of ClpP2 and the complex of ClpP1ClpP2 diffracted up to 8 Å.
Co-crystallization of ClpP1 together with the activator resulted in a structure with a determined resolution of 2.6 Å by molecular replacement (Figure 16). Careful examination of difference maps phased with the previously determined ClpP1 structure (PDB 2CBY) revealed no clearly defined new electron density that could be assigned to the activator peptide (Z-LL-H) (Figure 17). Regions with additional electron density were found close to the active site, where flexible loops were stabilized, potentially helping to form an active catalytic triad. However, we did not obtain conformational changes of the active site residues for the 2.6 Å structure of P1 in the presence of the activator. In this structure the distance between the histidine and the serine of the catalytic triad were the same as described in the literature (Ingvarsson et al. 2006). Further structural similarities with the published ClpP1 structure included no electron density for the first 14 residues and an assembly state of two ClpP1 rings, which is in contrast to the gelfiltration results. Moreover, the activator did not influence the compressed conformation of the protease with pores tarned between the two rings.
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Figure 17: Electron density map (Fo-Fc map) of ClpP1 and ClpP1 in addition with activator Z-LL-H. The map reflected coinciding density (blue), new density (green), and clashing density (red). The overall overlay of the polypeptide chain (pink) in the density was congruent.
Crystallization experiments were also carried out with ClpP2 alone. First crystals of ClpP2 grew within one week in 0.1 M sodium acetate trihydrate pH 4.6 and 8% PEG 4000 (Figure 16). The biggest crystals had a dimension of 150 µm x 30 µm. Unfortunately, the crystals diffracted poorly and the best resolution for ClpP2 was around 12 Å. The two Mtb protease homologs ClpP1 and ClpP2 feature a high degree of similarity, with 42.5% identity for ClpP1 (85 identical amino acids of 200 amino acids). The main surface conservation was located at the binding sites for the chaperone ring and at the bottom for the other protease ring. Most residues shared at least similar characteristics. The highest diversity was arranged in the pore and in the cleft between subunits in the ring sphere. A high conservation in symmetric patches around the pore most likely indicated a functionally important structure (Figure 18).
Figure 18: Surface of a single ClpP1 ring, with a resolution of 2.6 Å. The color of the protease ring displayed the conservation between mycobacterial ClpP1 and ClpP2: the scale ranged from high identity (blue) to no conservation (white).
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The S1 pocket determines the required amino acid characteristics of the substrate polypeptide chain (P1 position) after which the protease cleaves. To find substrates whose P1 amino acid was suited for the ClpP1 and ClpP2, we compared the residues of the S1 pocket (Figure 19). The S1 pocket of E. coli ClpP was formed by the amino acids F103, S106, I124, P126, K148, M151, N152, and L155 (Kress et al. 2009). In comparison with ClpP1 and ClpP2, most amino acids were conserved. Therefore, the residues of ClpP1 had higher similarity to E. coli ClpP. Due to the alignment, ClpP2 differed in two residues (position K162R and L168T).
Figure 19: Structure of the S1 pocket of ClpP (PDB 1TYF) in which residues involved in the surface contact are labeled and highlighted in red. The ClustalX alignment of E. coli ClpP and Mtb ClpP1 ClpP2 below the structure was visualized with Jalview, and residues of the S1 pocket were highlighted with red symbols.
5.3 Analysis of the N-terminal processing in the P1P2 complex
As processing of the ClpP N-termini to remove the propeptides appeared to depend on complex formation, processing activity, and its connection to complex formation was investigated. The results in the previous section showed that for the N-terminal processing of ClpP1 or ClpP2, the other subunit has to be present, since no processed subunits could be detected by mass spectrometric analysis of ClpP1 or ClpP2 alone. To analyze this in more detail, processing reactions were performed with either ClpP1 alone, ClpP2 alone, or a mixture of ClpP1 and ClpP2. As the activator peptide described by Goldberg and his group appears to influence peptidolytic activity, propeptide cleavage was investigated both in the absence and in the presence of activator Z-LL-H (Akopian et al. 2012). Samples at the reaction start and after overnight incubation were analyzed on Coomassie stained SDS- PAGE (Figure 20). For ClpP1 and ClpP2 no processing was observed.
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Figure 20: N-terminal processing of 3 μM ClpP1, 3 μM ClpP2, and equimolar concentrations of ClpP1ClpP2 +/- Z-LL-H (activator) at 25 °C. Reactions of different time points (0, 15, 30, 60 minutes, and ON) were analyzed on 15% SDS-PAGE and Coomassie stained. Number below was the time in minutes or hours, after which the reaction was quenched with SDS and heat inactivated at 95 °C.
However, in the complex of ClpP1P2 cleavage at least of the ClpP2 propeptide could be detected. In presence of the activator, the cleavage of the ClpP2 propeptide was fast: on a time scale of one hour the band corresponding to full length ClpP2 disappeared almost completely. For exact determination of the cleavage product, aliquots were analyzed by mass spectrometry. The obtained mass correlated to Δ12ClpP2. While processing of ClpP2 is easily detected by SDS-PAGE via the disappearance of the P2 band, it is an unfortunate coincidence that the truncated P2 runs at the same apparent molecular size in SDS-PAGE as full length P1 band. The mature P1P2 complex formed only one band at 21 kDa on SDS gels (Figure 20; Figure 22c). In absence of the activator, processing of ClpP2 occurred slowly. After overnight incubation, the decrease in the ClpP2 band was barely detectable. However, in the mixed complex and in the presence of the activator, a small incomplete down-shift of the P1 band by the appearance of a thin band underneath the P1 band was observed over the time-course of 60 minutes (Figure19). Purified Δ18ClpP2 with full length ClpP1 resulted in completely processed ClpP1 (Figure 21). The processing for the N- terminus of ClpP1 occurred overnight and resulted in the Δ7ClpP1. Therefore, the shift of the P1 band during processing was smaller compared to the P2 processing (Figure 21). To test, if processed ClpP1 can support ClpP2 propeptide cleavage, the analysis was carried out with the truncated ClpP1 variant. Incubation of purified Δ7ClpP1 and full length ClpP2 generated the Δ12ClpP2 in the same period of time.
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Figure 21: N-terminal cleavage of 3 μM Δ7ClpP1, 3 μM ClpP2, and equimolar concentrations of ClpP1Δ18ClpP2 +/- 0.75 mM Z-LL-H (activator). At indicated time points (zero, 15, 30, 60 minutes, and overnight) aliquots were heat inactivated at 95 °C, mixed with SDS and applied on 15% SDS-gel.
To investigate the contribution of each individual subunit in the cleavage reaction we generated the inactive variants ClpP1S98A and ClpP2S110A with mutated serine to alanine in the catalytic triad and tested the processing reaction with these variants (Figure 22). As expected the single inactive subunits ClpP1S98A or ClpP2S110A were unable to auto catalytically process themselves alone (Figure 22b), ruling out co-purification of E. coli ClpP subunits that might give false activities. The masses of the unprocessed proteins were confirmed by mass spectrometric analysis of the samples (Figure 22a). When ClpP1S98A was incubated with ClpP2, ClpP2 was processed to Δ12ClpP2, but ClpP1 remains unprocessed (Figure 22d). For incubation of active ClpP1 with inactive ClpP2S110A, a thin band appeared under ClpP1 during the reaction, but it was impossible to distinguish if the gel band of the overnight sample corresponded size wise to ClpP1 or to Δ7ClpP1. The mass spectrometry of the sample identified the gel band as Δ7ClpP1 and Δ12ClpP2. The processing of P2 in Figure 22d can be seen by the disappearance of the P2 full length band over the time. However, the half inactive complexes with a serine mutation in ClpP1 or ClpP2 resulted in different processing of the complexes. Therefore, the subunits did not equally contribute to the processing. Nevertheless, these results also reflected that the cross processing belonged to the activity of the active catalytic triad in the proteolytic Mtb ClpP.
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Figure 22: The propeptide cleavage reactions of ClpP1 and ClpP2 were recorded on 15% SDS-PAGE. The processing state of a) the single subunits ClpP1 and ClpP2, b) the active and inactive subunits, c) the fully active and inactive complexes ClpP1P2 and ClpP1S98AP2S110A, and d) the half active complexes ClpP1S98AP2 and ClpP1P2S110A were analyzed by using a gel based assay. All measurements were performed with 0.75 mM Z-LL-H (activator). The samples of different time points were quenched with SDS and heat inactivated at 95 °C. Finally, molecular weight of the single subunits and different active or inactive ClpP1P2 complexes were verified after 18 hours of cross processing by mass spectrometry analysis. The mass of full length ClpP1 corresponded to ClpP1S98A (blue), the N-terminal processed ClpP1 was marked in red. For unprocessed ClpP2S110A, the masses were highlighted in green and the mature ClpP2 was underlined in yellow.
The propeptide of ClpP1 is six amino acids long (VSQVTD) with a mass of 647 Da. For ClpP2, the propeptide sequence contains the motif VNSQNSQIQPQA (1313 Da). The sequence of the ClpP1 and ClpP2 constructs with highlighted propeptide cleavage sites are listed in the appendix. To test the possibility that interaction with AAA+ partner might stimulate processing and thus replace the activating role of the activator peptide, the propeptide cleavage was analyzed in presence of ClpX. As shown in Figure 23, the presence of the ATPase ClpX alone was insufficient to induce the N-terminal cleavage reaction of the protease in absence of the activator molecule (Figure 23a). In addition, the presence of a peptide of the ssrA amino
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acid sequence which simulated an element of the unstructured substrate chain, was ineffectual in triggering the reaction (Figure 23b).
Figure 23: Analysis of the influence of the chaperone ClpX and a peptide with the sequence of the Mtb ssrA-tag on the processing activity of ClpP1P2. a) Cross processing of ClpP1P2 in the presence of ClpX without the activator. Samples were quenched after different time points (30 and 60 minutes, 3 hours, and overnight) and were applied on 15% SDS-gel. b) Overnight incubation of 3 µM
ClpP1P214 with the ssrA peptide (AADSHQRDYALAA) did not result in N-terminal processing which was followed on 15% SDS-PAGE.
5.4 Contribution of the individual subunits to the peptidase activity
In absence of an AAA+ partner, ClpP can act as a peptidase, as has been shown for E. coli ClpP complexes and others (Thompson et al 1994). As Mtb ClpP complexes consist of two subunits, it is of interest to investigate how the individual subunits contribute to the peptidase activity of the assembled particle. To test peptidase activity, a fluorescence-based assay was employed using fluorescently labeled tri- or tetra-peptide model substrates (Ac-DEVD-AMC, Boc-VLK-AMC, Suc-LLVY- AMC, Glt-AAF-AMC, and Suc-GGL-AMC) that differed in the amino acid properties at the P1 position, the amino acid bound to the AMC. The substrates also differed in the protection groups at the N-terminus which might influence the accessibility to the proteolytic core of the ClpP1P2 particle. The structures of the peptide substrates with their specific cleavage sites are listed in the appendix. All substrates carried an AMC fluorophore at their C-terminus which produced a fluorescence signal increase when cleaved off, since bound and free AMC show different fluorescence yields. The measurement of ClpP peptidase activity was started 20 seconds after rapid mixing with the peptide substrate. The recorded fluorescence maximum for AMC is located at 440 nm. The fluorescence change corresponds to the amount of cut peptides resulting from the peptidolytic reaction of the ClpP particle. The peptidolytic activity was determined in the linear portion of the time course between 360 and
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420 seconds to exclude the lag phases in the peptidase reactions that exhibited such a phase (e.g. Suc-LLVY-AMC and Suc-GGL-AMC).
Figure 24: Peptidase assay of active and inactive ClpP1P2 complexes. The activity assay contained the preprocessed complexes with subunits in equal concentrations and 0.1 mM Ac-DEVD-AMC.
In order to characterize the contributions to peptidase activity by ClpP1 and ClpP2 subunits, the fully active Mtb P1P2 peptidase was compared to complexes assembled from one active and one inactive ring. All three complexes ClpP1P2, ClpP1S98AP2, and ClpP1P2S110A had no peptidase activity for the peptide Ac-DEVD-AMC (Figure 24). A very low activity was observed for the substrate Boc-VLK-AMC (Figure 25) and for Suc-LLVY-AMC (Figure 26), as no significant increase in fluorescence intensity could be detected during the measured time trace.
Figure 25: In the peptidase assay of the model substrate 0.1 mM Boc-VLK-AMC the full active peptidase ClpP1P2 had the highest cleavage rate of the tested complexes. The peptidase activity of active and inactive ClpP1P2 complexes was monitored, by using preprocessed complexes which were purified via centricons (10000 MWCO).
For the model peptide Glt-AAF-AMC (Figure 27) peptidase activity could be detected. Interestingly, the highest peptidase activity was achieved for ClpP1P2S110A where only ClpP1
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ring is active. The recorded activity rate was five times higher compared to the wild type. The complex ClpP1S98AP2 had no measurable activity, like the completely inactivated complex ClpP1S98AP2S110A.
Figure 26: Peptidase assay of preprocessed purified active and inactive ClpP1P2 complexes. All complexes contained the subunits in 1:1 ratios. The activity assay was started by addition of 0.1 mM Suc-LLVY-AMC.
Figure 27: The peptidase assay of active and inactive ClpP1P2 complexes of which the fluorogenic peptide Glt-AAF-AMC. For the assay, all complexes were preincubated overnight with 0.75 mM Z-LL-H and subsequently purified via centricons (10000 MWCO). The protease complex P1P2S110A (red) had a turnover number of ~0.03 min-1 and P1P2 (black) a turnover number of ~0.01 min-1. The activity assay contained 0.1 mM Glt-AAF-AMC.
A similar result was obtained with the model peptide Suc-GGL-AMC, where the complex ClpP1P2S110A also achieved a higher peptidase activity than the wild type complex (Figure 28). Surprisingly, the time courses with the peptide showed a pronounced lag phase that could not readily explained. The measured activity rate was 10 times higher than for ClpP1P2. In summary, it can be concluded that ClpP2 activity does not make a strong contribution to overall peptidase activity for the measured peptides. In fact, it appears that ClpP1 was even more active in context of the inactive ClpP2 ring, but the proteolytic activity depended on the interaction with the other Mtb ClpP subunit.
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Figure 28: In the proteolytic reaction of active and inactive ClpP1P2 complexes, ClpP1P2S110A had a turnover number of ~0.11 min- (linear slope between 360 and 420 seconds) All complexes were preincubated overnight with 0.75 mM Z-LL-H (samples were analyzed with mass spectrometry fig. 11). The concentration for the model substrate was 0.1 mM Suc-GGL-AMC.
5.5 ClpP function in combination with AAA binding partners
The peptidolytic activity of ClpP against model fluorogenic peptides provided a possibility to assess the in vitro activity of the E. coli peptidase. Yet, it is unknown if this peptidase activity of ClpP has any in vivo relevance. The only known in vivo activity of the Clp protease is in association with the Clp ATPases in protein degradation. To test the assembly state of the chaperone-proteases, as well as the ATPase and degradation activity we purified the Mtb ATPases ClpX and ClpC1.
5.5.1 Characterization of the two possible binding partners ClpX and ClpC1
One of the prerequisites for a fully active protease particle is the ATPase driven activity of the chaperone component. The ATPase activity of Mtb ClpX was therefore determined and compared to the activity of the E. coli ClpX. The Mtb ClpX has a slightly lower ATPase activity than the one of E. coli (Figure 29) with the calculated turnover rate for Mtb ClpX at ~30/sec. In contrast, the second ClpP1P2 associated chaperone of Mtb ClpC1 did not exhibit ATP hydrolysis activity, suggesting that it might not be assembled, since ATPase activity requires formation of the hexameric ring. The ATP turnover could be followed in the coupled assay by the phosphorylation of 7-methylinosin by PNPase resulting in hypoxanthine accompanied by an absorbance decrease at 291 nm. The highest ATP turnover was determined using the phase between three and four minutes.
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Figure 29: ATPase assay of 0.2 µM Mtb ATPases ClpX (green), ClpC1 (blue), and as a control 0.2 µM ClpXE. coli were measured with 5 mM ATP for six minutes. The measured ATPase reactions are buffer corrected. The turnover numbers were calculated in the linear range between 3 and 4 minutes.
The AAA binding partner only interacts as an assembled hexameric ring with the ClpP protease. To investigate the assembly state of ClpC1 analytical gelfiltration runs were performed (Figure 30). In the comparison, the gelfiltration runs of ClpC1 were performed at RT in the absence (red) and in the presence of different nucleotides. The samples including nucleotides were preincubated with 20 mM MgCl2 and 1 mM ATP or 0.75 mM ATPɣS for one hour at room temperature. The elution volume of ClpC1 shifted from 1.7 ml to 1.6 ml upon addition of ATP (yellow) or ATPɣS (blue). However, the shifts in elution position are too small for hexamerization but rather might correspond to formation of a ClpC1 dimer.
Figure 30: Gelfiltration runs of ClpC1 on a Superose 6 2.4 ml column. The analytical size exclusion chromatography of ClpC1 + ATPɣS
(blue) contained 0.75 mM ATPɣS and 20 mM MgCl2 in the running buffer, additionally the gelfiltration buffer of ClpC1 +ATP (yellow) was supplemented with 1 mM ATP and 20 mM MgCl2. The protein absorption was detected at 227 nm. The standard protein masses are labeled above the size exclusion runs.
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To test whether perhaps the ClpP1P2 particle as binding partner could induce hexamerization of ClpC1 by serving as a template for assembly into the ring, analytical size exclusion chromatography on a Superose 6 column (2.4 ml) was performed in presence of ClpP1P2 and ATPγS. Also with this setup no hexamerization of ClpC1 could be observed and no complex was formed between ClpP1P2 and ClpC1 (Figure 31). Both runs of ClpC1 alone and with added ClpP1P2 eluted at 1.6 ml, in contrast to the control (ClpAP of E. coli) which eluted as the active complex at 1.2 ml. It is possible that an assembly adaptor similar to MecA of B. subtilis might be required for Mtb ClpC1-ring assembly.
Figure 31: A Superose 6 (2.4 ml) column was used for the analytical size exclusion chromatography to compare ClpC1 alone and incubated with complex ClpP1P2. All gelfiltration runs contained 0.75 mM in the running buffer and the absorption was measured at 227 nm. The standard protein masses are labeled in the chromatogram.
A potential adapter molecule encoded in the Mtb genome that might assist in the correct assembly of ClpC1, such as the ClpC assembly factor MecA of B. subtilis, might be the protein ClpC2. ClpC2 features 20% sequence similarity (substitutions of amino acids with similar side chain properties, scoring > 0.5 in the Gonnet PAM 250 matrix) and 11 % identity (32 identical positions of 252 amino acids) to the adaptor molecule MecA (alignment not shown), but mainly came to attention due to its high similarity to the ClpC1 N-domain with 61 identical positions (Figure 32b and c). As ClpC2 is a small protein homolgous to a portion of ClpC1, yet lacks the canonical ATPase domain, we concluded that it might not act as an alternative ClpP-binding partner but rather as an adapter to ClpC1. However, the size exclusion chromatography of a preincubated sample containing ClpC1 and ClpC2 revealed no higher oligomeric assembly state (data not shown). Alignment of ClpC2 with the protein ClpS1 of A. thaliana yielded 20% sequence similarity
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(data not shown) and 10 % identity with 25 identical positions of 252 amino acids. ClpS1 of A. thaliana exhibits a low homology to E. coli ClpS, but also functions in N-degron recognition (Nishimura et al. 2013). I therefore tested its potential role in the degradation of N-end rule model substrates. However, the FR-li-GFP, an N-end rule model substrate was not degraded by the protease ClpP1P2 in presence of ClpC1 and ClpC2 (Figure 32a).
Figure 32: a) The mixture of ClpC1C2P1P2 was unable to degrade the N-end rule substrate FR-linker-GFP. The samples of 0, 4 hours, and overnight were quenched with SDS and heat inactivated. Final analysis was carried out by SDS-PAGE and Coomassie staining. b) Structural alignment of the N-domain of ClpC1 (PDB 3WDB blue) and a structure prediction of ClpC2 by Phyre2 using PyMOL. c) The sequence alignment of ClpC1 and ClpC2 was performed by UniProt alignment, visualized by Jalview, and colored in ClustalX style.
5.5.2 ClpX stimulates proteolytic sites of ClpP
The ATPase dependent activation of ClpP by structural remodelling is presumably a prerequisite to perform degradation by chaperone-proteases (Lee et al. 2010). The degradation activity of the Mtb ClpXP complex was first tested by using the two ssrAMtb- tagged model substrates.
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Figure 33: ClpX activated degradation of λR-ssrA (PDB 1LMB) by the protease ClpP1P2 +/- activator molecules (Z-LL-H). In the assays 1 µM substrate was degraded by 1 µM ClpX and 1 µM (ClpP1ClpP2, ClpP1, or ClpP2). Samples were taken at time point 0, 1, and 3 hour, and overnight and analyzed on 15% SDS gels.
To assay the efficiency of ClpP1P2 in protein quality control we fused an ssrA-tag to an enzyme of the Mtb proteome. We fused a ssrA-tag C-terminally to the Mtb malate dehydrogenase.
Figure 34: Only the complex of ClpP1P2 interacted with the ATPase ClpX to degrade the ssrA-tagged Mtb substrate malate dehydrogenase (PDB 3HHP). For the gel-based assay (15% SDS) 1 µM substrate was degraded by 1 µM ClpX and 1 µM (ClpP1ClpP2, ClpP1, or ClpP2) +/- activator (Z-LL-H). Numbers below the gel indicated the time points (0, 1 hour, 3 hours, and overnight) after which reaction was stopped (heat and SDS).
ClpP1 and ClpP2 alone with an otherwise identical setup were used as controls. The degradation time courses were recorded both in the absence and the presence of the activator peptide and were started by addition of 5 mM ATP. It should be noted here that unprocessed, but preincubated and therefore fully assembled ClpP1 and ClpP2 were used in this experiment. The propeptide processing in the sample containing the activator Z-LL- H, contributed to the overall degradation time courses observed. As can be seen in Figures 33 and 34, the ClpP1P2 particle in combination with ClpX degrades the malate dehydrogenase-ssrA (MDH-ssrA) model substrate as well as the λR-ssrAE. coli (N-terminal domain of λ–repressor fused to the ssrA-tag) independent of the activator. The activator also
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does not appear to have any influence on the rate of the degradation, as the bands in the presence and in the absence of activator disappeared on the same time scale. Degradation of substrates was measured by a decrease in gel band intensity of the corresponding protein. This suggests that peptide activator is not needed in the context of the fully assembled protease complex to support protease activity. As expected, ClpP2 or ClpP1 alone were not able to support this reaction even in presence of ClpX. The degradation is ssrA-dependent, since MDH without the ssrA-fusion is not degraded (Figure 35).
Figure 35: a) Degradation assays of MDH-ssrA and untagged MDH (1 µM) by 1 µM ClpX and 1 µM (ClpP1ClpP2, ClpP1, or ClpP2). Samples of time point 0, after 1, 2, 3, 4, 5 hours, and overnight were loaded on 15% SDS-PAGE and Coomassie stained. b) Degradation assay of MDH-ssrA by ClpP1P2 in the absence of ClpX. The samples were quenched with SDS and heat inactivated at 95 °C and further followed by 15% SDS-PAGE.
As mentioned above, in the degradation experiments the complex ClpP1ClpP2 was not preprocessed by the activator. Therefore, the fact that ClpP1P2 in the presence of ClpX without activator were able to degrade the substrates MDH-ssrA and the λR-ssrAE. coli suggests that ClpX-dependent propeptide processing must have occurred but only in presence of a degradation substrate (Figure 23a and 35a). To test which subunit contributed most to the ClpX associated degradation, we performed degradation of MDH-ssrA with ClpP particles consisting of one active and one inactive ring, ClpP1S98AP2 and ClpP1P2S110A (Figure 36), respectively. This time, preprocessed particles were used.
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Figure 36: In vitro degradation assays of MDH-ssrA at room temperature. In the assays 1 µM substrate was degraded by 1 µM ClpX and 1 µM (ClpP1ClpP2, ClpP1, or ClpP2). Aliquots of time point 0, after 1, 2, 4 hours, and overnight were applied on 15% SDS gels.
The propeptide processed ClpP1P214 was able to degrade the 35 kDa MDH-ssrA on a time scale that required overnight incubation. Despite ClpX present, ClpP1 (data not shown) and ClpP2 alone were not able to degrade the MDH-ssrA completely. Surprisingly, the partially active complexes ClpP1S98AP2 and ClpP1P2S110A accelerated the degradation rate of substrate MDH-ssrA, relative to the complex with intact ClpP1 and ClpP2 triades. The substrate band disappeared completely within a four hours reaction. The overall degradation rates of the partially active ClpXP1P2 chaperone-proteases were similar independent of which subunit was inactivated. However, our results demonstrated that the complex of ClpP1P2S110A was slightly faster than ClpP1S98AP2. In the performed assay, the complex degraded most of the MDH-ssrA in two hours. To investigate ClpP1P2 responsibility for the degradation of MDH-ssrA, as a control an assay without chaperone ClpX was performed (Figure 35b). The missing ClpX in this assay was accountable for the degradation inability and the remaining substrate band obtained in the final reaction sample.
5.5.3 Scan of the Mtb genome for proteins carrying ssrA-like degrons
It has been shown in E. coli and other bacteria that ClpP in association with the ClpX ATPase partner is responsible for degradation of ssrA-tagged substrates. This also appears to be the case for GFP-ssrAE. coli in Mtb in the presence of the E. coli adapter SspB (Kim et al 2011). It was furthermore shown for the E. coli system that some substrates were directly recognized by this degradation complex based on C-terminal amino acid sequences that
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resemble the ssrA-tag, in particular the terminal three residues. A preferred degron appeared to be the presence of the LAA-tripeptide at the very C-terminus. We therefore searched the Mtb proteome for amino acid sequences ending in this tri- peptide or related sequences. The assessment resulted in 9 possible substrates for ClpXP dependent degradation (Table 5). Two of these potential substrates were assigned essential based on Himar-1 transposon mutagenesis screen (Griffin et al 2011). The essential proteins were FbiC, a FO synthase that is involved in cofactor biosynthesis (Darwin et al. 2003) and PapA3, a possible adapter of the polyketide synthase. Furthermore, two substrates were probably involved in virulence as toxin-antitoxin systems (MazE9 and VapC27). Another substrate was annotated as a potential transcription regulator (Rv2308) based on the existence of a helix-turn-helix motif within its structure. The identified substrates bared three different C-terminal degradation tags; five had a LAA-tag, three contained the VAA-tag, and one the motif NVA. Three of these hypothetical substrates (Rv2308, Rv2961, and Rv3832c) were cloned, expressed and purified to test for degradation. However, Rv2961 did not yield soluble protein during expression and the purified Rv3832c was not degradable by ClpXP1P2.
Table 5: Screen of the Mtb proteome against known C-terminal ssrA-like degrons
The ClpX associated degradation system could degrade almost completely the putative transcription regulator Rv2308 during overnight incubation (Figure 37). The results for the degradation assays of MDH-ssrA were similar compared to the degradation of the substrate Rv2308 with the C-terminal LAA degron. In the degradation assays, the absence and presence of the activator displayed again no influence on the degradation rate of the substrate Rv2308.
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Figure 37: a) Degradation attempts of 1 µM Rv2308 by 1 µM ClpX and 1 µM ClpP1ClpP2 in the presence or the absence of the activator were analyzed by SDS-PAGE and Coomassie staining. Samples were taken at time point 0, after 1, 3 hours, and overnight. The Cpk was part of the regeneration system. b) Degradation of Rv2308 by ClpX in complex with the single subunits ClpP1 or ClpP2 were followed on 15% SDS-PAGE.
5.5.4 Small peptides inhibit ClpXP1P2 dependent degradation
The hexameric chaperones ClpX and ClpC bind to hydrophobic patches on the sevenfold symmetric ring surface of ClpP via loops carrying a conserved motif at their tip consisting of a glycine flanked by two hydrophobic residues (LGF). The small antibiotic ADEP also interacts with the same binding sites and its binding opens the N-terminal pores of ClpP leading to unregulated proteolysis. We designed two small peptides of three (LGF) and seven (RGLGFGA) amino acid length imitating the LGF loop of Mtb ClpX and one peptide that simulated a second interaction between chaperone and protease, referred to as pore 2 loop peptide (NPSITRD).
Figure 38: The inhibitory effects of small peptides on the degradation of 1 µM MDH-ssrA by 1 µM ClpX and 1 µM (ClpP1ClpP2, ClpP1, or ClpP2) were tested. Samples of various time points (0, 1, and 2 days) were applied on 15% SDS-gel.
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First, the potential peptidase-stimulating effect analogous to an ADEP mode of action was tested, However, none of the LGF containing peptides lead to an increase in ClpP1P2 peptidase activity in absence of the ATPase partner (data not shown) indicating that they either did not bind or their binding (unlike ADEP) did not lead to opening of the pores. We then tested the possible inhibitory effect of the two IGF peptides on the ClpXP1P2 dependent degradation of MDH-ssrA (Figure 38). Activity was measured over a total period of two days in the presence of hepta and tri LGF peptides. Both peptides lead to a reduction of the degradation rates of MDH-ssrA. Furthermore, the samples of the last time point after two days still contained 2/3 of the substrate. The normal degradation rate required 15 hours for the complete degradation of MDH-ssrA by ClpXP1P2. However, this inhibition did not appear to be that specific, since the presence of another small peptide Z-LLVY-AMC that did not contain the LGF motif also inhibited the degradation of the MDH-ssrA by ClpP1P2X.
5.6 Post-translational modifications of ClpP1P2
5.6.1 Pupylation
ClpP2 was detected in a proteomic study that identified proteins post-translationally modified with prokaryotic ubiquitin-like protein Pup. To confirm that ClpP2 can indeed be pupylated and to characterize the properties of the modified complex, we carried out in vitro pupylation of ClpP2. The recorded pupylation time course (Figure 39b) shows that ClpP2 can be pupylated up to 20% under optimized reaction conditions (50 mM HEPES-NaOH pH7.5, 150 mM NaCl, 1 mM DTT, and 10% glycerol overnight at 37 °C. After three rounds of pupylation, 30% of ClpP2 were ligated to Pup (Figure 40). Mass analysis indicated two possible lysines as pupylation sites (K174 or K181). On the contrary, for ClpP1 no pupylation could be observed under the same conditions (data not shown). The pupylated ClpP2 still assembled into the double-ring complex with ClpP1 (Fig. 40a). Surprisingly, the complex ClpP1P2-Pup eluted later than the unpupylated complex in size exclusion chromatography (Superdex 200 resin), although each conjugated Pup adds a mass of ~7 kDa to ClpP2 (Figure 39).
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Figure 39: ClpP2 could be pupylated and the post-translational modified ClpP2 was still able to assemble with ClpP1 in a higher oligomeric complex. a) Analytical size exclusion chromatography were performed on Superdex 200 (left) of the subunits ClpP1 (dashed line), ClpP2 (dashed line), the complexes of ClpP1P2-Pup (red), and ClpP1P2 (black) at RT. The standard protein masses are labeled in the figure. b) Pupylation of ClpP2 (right) at 37 °C on 15% SDS-gel was further analysed by ImageJ. The numbers above are the time in hours after which the reaction was quenched with SDS and heat inactivated at 95 °C. In the final reaction sample 19% of ClpP2 monomers were pupylated.
The ClpP1P2-Pup had a slightly increased peptidase activity compared to wild type (Figure 40). Moreover, Pup serves in Mtb as a degradation tag that targets modified substrates for proteasomal degradation. Therefore, the Pup-conjugated ClpP2 of the pupylation reaction, was tested as a substrate for proteasome degradation. Degradation in context of the ClpP1P2 particle was not tested.
Figure 40: Purification and peptidolytic activity of ClpP2-Pup. a) Purification of ClpP2-Pup from a Pup conjugation assay with PafA. b) Peptidase activity of substrate Z-GGL-AMC by protease act ClpP1P2-Pup (red), ClpP1P2 (black) in addition with 0.75 mM Z-LL-H, and ClpP1P2 without (green) 25 °C
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About 75% of the ClpP2-Pup disappeared after two hours. In the overnight sample of the reaction, the ClpP2-Pup was completely degraded. But beside the pupylated ClpP2 the associated non pupylated ClpP2 was also removed. In the overnight sample only a tiny band of ClpP2 remained. This is surprising, since it suggests that only one tagged subunit of a ClpP2 heptamer was enough to induce complete degradation by the proteasome, including the untagged subunits (Figure 41 a).
Figure 41: a) ClpP2-Pup was degraded by Mpa with Mtb proteasome. For the performed assay at RT, measured ClpP2-Pup was pupylated by Pup-Strep and purified via Strep column. Samples of six time points were applied on 15% SDS-gel and Coomassie stained. b) Pup was not a degradation signal for ClpXP. The degradation assay of purified adenylate kinase conjugated to Pup was analyzed by SDS-PAGE and Coomassie staining.
Using a pupylated substrate, we tested whether perhaps the Mtb ClpXP complex could also degrade Pup-tagged proteins, since Pup is unstructured and by this nature might be recognized. However, ClpXP1P2 displayed no Pup-dependent degradation of pupylated mycobacterial adenylate kinase (Adk; Rv0733) (Figure 41 b).
5.6.2 Phosphorylation of ClpP1
In a proteomic study that catalogued Mtb proteins that are phosphorylated (Prisic et al. 2010), ClpP1 was found modified at threonine 17. To mimic the phosphorylation at this position and investigate its behaviour in the ClpP1P2 complex formation, we engineered a phosphomimetic variant ClpP1T17E that simulated the charge at threonine 17, when it is phosphorylated. Another variant was generated ClpP1T17A which represented the unphosphorylated protein. Our results indicated that the complex ClpP1T17EP2 possessed a reduced activity in the peptidase assay, but also ClpP1T17AP2 had a lower activity, compared to the wild type (Figure 42).
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Figure 42: a) In the peptidase assay of ClpP1P2 and the two phosphomimetic mutants the substrate Suc-GGL-AMC was cleaved in the presence of 0.75 mM activator.
69
6 Discussion
ClpP protease complexes are universal in the bacterial kingdom, the only exception being Mycoplasma and relatives which feature an exceptionally reduced genome (Yu & Houry 2007). To act on protein substrates, the ClpP double-ring complex must associate with an ATPase chaperone ring that unfolds the substrate and translocates it into the ClpP proteolysis chamber (Weber-Ban et al. 1999). Usually, at least two alternative ATPase partners co-occur in all bacterial organisms, allowing formation of different ClpP-chaperone complexes. The existence of two ClpP subunits (ClpP1 and ClpP2) in M. tuberculosis offers potential for an even larger combinatorial set of protease complexes. Whereas in E. coli and many other Gram-negative bacteria the clpP gene can be disrupted without affecting viability, it is essential in Mycobacteria. The higher modular complexity of the Clp-protease system in Mtb along with its promise as a drug target, make this system particular interesting to study. Understanding how the Mtb ClpP protease complexes are assembled and regulated could give clues on how to design drugs that target this system. The fact that clpP1 and clpP2 are located within the same operon suggested to us that the subunits are coexpressed in the organism rather than expressed differentially under certain conditions. However, whether the two subunits would form two separate homooligomeric particles or might form a heterooligomeric complex was not known. The results presented in this thesis support a heterooligomeric double-ring architecture composed of two homooligomeric heptameric rings, one ClpP17 and one ClpP27. During the preparation of this study, another study was published that demonstrated formation of a ClpP17ClpP27 particle (Akopian et al. 2012). However, while our analytical gel filtration analysis showed that the individual ClpP1 and ClpP2 subunits assembled only to the stage of single rings,
Goldberg and colleagues detected only ClpP114 and ClpP214 double-rings and postulated that the limiting step in formation of the heterooligomer is the dissociation of the homooligomeric double-rings. In fact, they proposed that the double-rings are so stable that a small peptide activator Z-LL-H is required to dissociate the rings quantitatively and allow hetero-complexes to form. In contrast, our data are consistent with a model where assembly into heterooligomeric particles does not require disruption of the homooligomeric particles but occurs spontaneously upon mixing of the two subunits. This assembly appears to require intact propeptides, since we observed that double-ring could not be achieved from ClpP2 subunits that were expressed without their propeptides. It is possible that parameters like
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ionic strength, temperature, and glycerol concentration contribute to the discrepancy in the findings. The formation of single heptameric ClpP rings has been observed for human mitochondrial ClpP particles (Kang et al. 2005). In absence of the ATPase partner ClpX, the ClpP subunits form single, heptameric rings. It was shown that the single rings of human ClpP are inactive due to an inactive conformational arrangement of the catalytic triad, thereby preventing exposure of cytosolic proteins to active protease sites. Upon association with ClpX the ClpP single rings make a double-ring and at the same time the catalytic triad is activated. Measurements of the ClpP2 abundance suggests that some ClpP2 at least are available in free form in the cell (Schubert et al. 2013). The inability of uncomplexed protease subunits
ClpP17 and ClpP27 to degrade randomly accessible cytosolic proteins or peptides is presumably also highly important for Mtb viability (Lee et al. 2010). Our assessment of the peptidase- activity of ClpP17 and ClpP27 single rings showed that they were catalytically inactive towards a range of fluorogenic peptides (Boc-VLK-AMC, Suc-LLVY-AMC, Glt-AAP- AMC, and Suc-GGL-AMC) likely due to a misarranged catalytic triad. Only mixed ClpP1P2 complexes showed peptidase activity for the substrates. To observe peptidase-activity, we also had to make use of the activator peptide confirming the recently published results that ClpP1P2 alone required an activation by Z-LL-H for peptidase activity (Akopian et al. 2012). The activator peptide is a derivate of the proteasome inhibitor Z-LLL- H which was discovered by chance to have the opposite effect on Mtb ClpP, namely activation of peptidase activity. In principle, the small activators share a similar length with the cleavage products (3-8 amino acids) of the Clp proteolysis.
In our peptidase assays we screened a range of substrates with acidic, basic or hydrophobic properties and small amino acids in the P1 position. It should be mentioned that the chemical properties of the associated protection group of the substrate might also influence substrate accessibility and could therefore interfere with the preferred characteristics of ClpP1P2 for the amino acid in the P1 position. Nevertheless, the ClpP1 in the complex with an inactive ClpP2 ring shows a similar preference as E. coli ClpP (Arribas et al. 1993) which prefers large aromatic amino acid in the P1 position (Arribas et al. 1993). Therefore, the highest cleavage rate was achieved for Glt-AAF-AMC with a phenylalanine in position P1 and for Suc-GGL-AMC, carrying a hydrophobic leucine in the P1 position. Surprisingly, ClpP1 in the half inactivated complex had a very low cleavage rate for the peptide Ac-DEVD-AMC. We surmise that this is one reason why the N-terminal cleavage of
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ClpP1 propeptide is significantly slower than that of ClpP2. The main cleavage site for ClpP1 processing was found to be D6-M7. For ClpP2 on the other hand, the cleavage occurred between A12 and R13. The ClpP1P2 complex showed no hydrolysis for the substrates Ac-DEVD-AMC and Boc- VLK-AMC. These peptides might get no access to the active sites or were released before P1P2 could cleave. We did not test the influence of the chaperone ClpX on the hydrolysis rates of P1P2 for these substrates. For E. coli Clp, it was shown that some peptides which did not display high turnover rates required the association with ClpX in presence of nucleotide to promote hydrolysis (Thompson et al. 1994). In contrast to the cleavage rates reported for E. coli ClpP in the literature, the measured Mtb protease ClpP1P2 peptidase activity is ~1000 fold lower. This is most likely due to hindered substrate accessibility. The fluorophoric substrates Suc-GGL-AMC and Suc-LLVY-AMC have a succinyl protection group and showed a lag phase in the first 300 seconds of the peptidase assay. The lag phase may be occured from replacement reactions in which the substrates competed with propeptides from the previous preprocessing reaction to the substrate binding sites. In these reactions also the activator itself may have interfered with the cleavage reaction by interacting to residues involved in substrate binding or directly to the catalytic triad which is required for the peptide cleavage reaction. One somewhat puzzling result was our observation that the half inactivated complex ClpP1P2S110A was able to function as a peptidase five times stronger than the fully active complex. Furthermore, the inactivation of ClpP2 significantly enhanced the cleavage rates for the substrates Glt-AAP-AMC and Suc-GGL-AMC. One possible explanation would be that all peptidase assays have to be carried out under the makeshift help of the activator peptide. It is not clear where this activator binds and how it activates the peptidase activity but regardless of the exact mechanism, it is possible that it binds more strongly to one ClpP subunit than to the other and therefore more strongly enhances one over the other.
The maturation of ClpP was already described for E. coli, where a propeptide of 14 amino acids is cleaved off, to allow better substrate access to the proteolytic core (Maurizi et al. 1990). For the Mtb ClpP subunits, the individual subunits alone (even in the presence of the activator) had no intrinsic N-terminal cleavage ability. The presence of the other subunit was required to induce the cleavage. Interestingly, when double-ring complexes were generated that consisted of one catalytically active and one inactive ring, we found that the subunit ClpP2 exclusively processed itself and was unable to process the inactive ClpP1 within the
72 6 Discussion
same particle. ClpP1 on the other hand not only processed itself but also cross processed ClpP2. One possible explanation for this observation might be the length of the propeptides. The ClpP1 propeptide is short and can therefore likely only reach the active sites in the ring it is part of. The ClpP2 propeptide is almost twice as long and might for that reason be able to reach the catalytic sites of the opposite ring. After the truncation both particles feature a similar length of their N-termini. While analysis of the assembly and processing of the ClpP particle is important and peptidase activity measurements have also been a useful tool in studying ClpP complexes, it needs to be noted in this context that the role of ClpP inside the cell is not as a peptidase but as a protease. Hence, the activity toward fluorogenic peptide substrates is an artificial system and the question of in vivo relevant ClpP activity still needs to be addressed in the context of an AAA+ partner and with protein substrates. We therefore postulated that an activator might not be needed for processing in presence of an AAA+ partner. We tested processing of ClpP1P2 in presence of the chaperone ClpX. We found that appearance of N- terminal processing during degradation of MDH-ssrA by ClpXP1P2 without Z-LL-H caused ClpP1P2 activity independent of activator. Although it appeared that processing is facilitated by ClpX in presence of protease substrate, the results remained somewhat inconclusive because ClpX in absence of a substrate did not generate the same effect. These results support a scenario where substrate translocation into the ClpP protease chamber stimulates conformational changes essential for protease activity and processing of the N-terminus, because we could not obtain processing of ClpP1P2 in association with ClpX alone. The presence of substrate was required, but a peptide of the ssrA-tag sequence alone was not sufficient for ClpP1P2 processing. Our observed set of substrates is based on the C-terminal degradation tag (ssrA) for ClpXP. For a complete set of all interacting proteins, including substrates with unknown recognition motifs and adapters of ClpXP pull downs with ClpX in presence and absence of our trapped protease particle ClpP1S98AP2S110A could result in better insights.
So far the natural targets of Mtb ClpXP dependent proteolysis are not well described: only an artificial in vivo degradation assay of GFPssrA in complex with SspB of E. coli (Kim et al 2011) and the specific degradation of anti-sigma factor (RseA) and FtsH by ClpC1 and ClpP2 (Barik et al. 2010) are reported. Here we report the first in vitro reconstitution of an active ClpP1P2 chaperone complex of Mtb. We obtained activity for ClpXP1P2 and not for one of the single subunits in complex with the chaperone. Our results indicate that only a mixed
73 6 Discussion
protease complex of ClpP1P2 possesses the ability to interact with the chaperone in a functional mode. Furthermore, for the degradation activity in complex with ClpX no additional factors were required and activator was not added in those reactions. Therefore, we conclude that the previously described activator Z-LL-H only influenced the in vitro peptidolytic activity of P1P2 but is not necessary in context of the fully assembled chaperone-protease. The unfolding activity of Mtb ClpX presented in this study is relatively weak and the chaperone ClpX failed to unfold GFP-ssrA. It is also possible that the low degradation rate is due to insufficient assembly with the chaperone. For ClpX we could not detect the interaction with ClpP in analytical size exclusion chromatography, nor in electron microscopy studies (data not shown), suggesting that association is not very stable. Nevertheless, the ATPase activity, the degradation activity and the influence on ClpP1P2 processing were clear hints that association between the protease core and the ATPase is taking place. In degradation assays with the ATPase ClpX we obtained higher degradation rates for both half-inactive complexes than for the fully active ClpP1P2 particle. A possible explanation for the increased degradation activity of the half-inactive complexes ClpP1S110AP2, ClpP1P2S98A and the additional higher rate for peptide hydrolysis in ClpP1P2S98A could be a different complex conformation which raised the substrate accessibility (e.g. active complex and N- termini in “up” conformation) or facilitates the substrate release (compressed conformation with additional exit pores). However, it is at this point unclear what these conformational adjustments could be. It should be pointed out here that there is in vivo relevance to the activity of mixed active and inactive complexes of P1P2. It was reported in the literature that ClpP1 can be phosphorylated in Mtb (Prisic et al. 2010). When we analysed a phosphomimetic variant, we observed an inhibitory effect on the peptidase activity similar to the half-inactive complex ClpP1S100AP2. But for a better comparison of the constructs further degradation- and processing experiments should be performed. Furthermore, for ClpP2 a post-translational modification was also described. It was shown that ClpP2 can be covalently modified with the prokaryotic ubiquitin-like protein Pup (Festa et al. 2010). We could not completely conjugate all of ClpP2 to Pup, so we could not determine if 100% pupylation might stronger influence the peptidase activity, but 30% Pup- conjugation of ClpP2 complexed with ClpP1 slightly increased the peptidase activity in the presence of the activator. Another regulatory effect of Pup we observed for the activity of ClpP2 was in the presence of the proteasome which degraded not only the pupylated ClpP2
74 6 Discussion
but also the associated unpupylated ClpP2 proteases. It might be that pupylation is a means to selectively remove ClpP2 from the ClpP1P2 system. To which end this might be useful is not clear at the moment.
Because ClpP is so well conserved in pathogens and because regulatory proteolysis seemed essential in different pathogens, the design of Clp inhibitors is of central interest. Here we demonstrated the success of small peptide compounds in competition with ClpX binding to ClpP1P2. However, the effectiveness of small chaperone-protease inhibitors is limited by permeability of the compounds through the thick cell wall barrier of M. tuberculosis. Therefore, the treatability of Mtb must be tested in vivo. However, it is not known if drugs aiming at the inhibition of the Clp system of pathogens would not interfere with human mitochondria function, since this organelle also exhibits a Clp version which functions in a very similar way.
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8 Appendix
8.1 Abbreviations
AAA+ ATPase associated with diverse cellular activities GC Guanine-cytosine content of DNA DNA Deoxyribonucleic acid RNA Ribonucleic acid FasI Fatty-acid synthetase HslUV ATP dependent protease ClpYQ Mtb Mycobacterium tuberculosis Clp Caseinolytic protease ssrA Transfer-messenger RNA Pup Prokaryotic ubiquitin-like protein ATP Adenosine triphosphate ADP Adenosine diphosphate ATPyS Adenosine-5'-O-(3-thiotriphosphate) dosR Transcription regulator in Mtb RNIs Reactive nitrogen intermediates iNOS Interferon-ɣ-inducible nitric oxide synthase DnaK 70 kilodalten heat shock protein GroEL 60 kilodalten heat shock protein groES 10 kilodalten heat shock protein CtsR Gram positive master regulator of protein quality control tig Trigger factor σ32 RNA polymerase sigma factor RpoH σB Alternative sigma factor HtrA High temperature requirement protease A = peptidase Do FtSH ATP dependent zinc metalloprotease SulA Cell devision inhibitor La Lon protease Zn2+ Zinc °C Degree Celsius IGL/F Amino acid sequence HflK Modulator of FtsH protease
92 8 Abbreviations
HflC Modulator of FtsH protease HflD High requency lysogenization protein SpoVM Stage V sporulation protein M SpoIIAB Anti-sigma F factor SpoOH sporulation σH MgtR Regulatory peptide in Salmonella typhimurium ClpS ATP-dependent Clp protease adapter protein MecA Adapter protein MscB Arginine Kinase of B. subtilis SspB Stringent starvation protein B rssB Regulator of RpoS PolyP Inorganic polyphosphate PinA Puroindolie-A Mpa Mycobacterium proteasome ATpase Rv Virulent variant of Mycobaterium tuberculosis H37 strain ADEP Acyldepsipeptide Aa Amino acid kDa Kilodalten NBD Nucleotide-binding domains NTD N-terminal domains ClgR Clp gene regulator Z-Leu-Leu-H Benzyloxylcorbaonyl-L-Leucyl-L-Leucinol PtrB Protease II (Oligopeptidase B) Acr2 Chaperone of alpha-crystallin family pH According to Carlsberg “power of hydrogen” RecN DNA repair protein MuA Bacteriophage Mu RseA Anti-sigma-E factor RsiW Anti-sigma-W factor ECF sigma factor σW SPX YjbD suppressor LAA ClpXP degron sequence (leucine, alanine and alanine) SOS Is not a abbreviation, is a morse code FtsZ Cell devision protein LexA Repressor
93 8 Abbreviations
CtrA Cell cycle response regulator YmoA modulating protein RpoS sigma-38 or KatF inlA invasion genes actA Actin, alpha skelet muscle ACP Activators of Self-Compartmentalizing Proteases Å Angstrom S1 oxyanion hole F Phenylalanine S Serine I Isoleucine P Proline K Lysine M Methionine D Aspartate L Leucine V Valine T Threonine pdb Protein Data Bank hClpX human ClpX GFP Green Fluorescent Protein aa/s Amino acid per second
KD Dissociation constant C- or N- Carboxy- or amino- terminal tmRNA Transfer-messenger RNA Mg2+ Magnesium umuD involved in SOS response CpdR two component receiver protein σS Sigma S Ira anti-adaptor proteins ComK Competence transcription factor ComS Regulator of genetic competence 26S proteasome Svedberg unit PafA proteasome asociated factor A
94
8.2 Properties and sequences of the protein constructs
ClpP1 (RV2461c)
-1 -1 MW: 22125 Da ε280: 14565 M cm 10 20 30 40 50 60 MSQVTDMRSN SQGLSLTDSV YERLLSERII FLGSEVNDEI ANRLCAQILL LAAEDASKDI
70 80 90 100 110 120 SLYINSPGGS ISAGMAIYDT MVLAPCDIAT YAMGMAASMG EFLLAAGTKG KRYALPHARI
130 140 150 160 170 180 LMHQPLGGVT GSAADIAIQA EQFAVIKKEM FRLNAEFTGQ PIERIEADSD RDRWFTAAEA
190 200 LEYGFVDHII TRAHVNGEAQ HHHH
Δ7ClpP1 (RV2461c)
-1 -1 MW: 21463 Da ε280: 14565 M cm 10 20 30 40 50 60 SMRSNSQGLS LTDSVYERLL SERIIFLGSE VNDEIANRLC AQILLLAAED ASKDISLYIN
70 80 90 100 110 120 SPGGSISAGM AIYDTMVLAP CDIATYAMGM AASMGEFLLA AGTKGKRYAL PHARILMHQP
130 140 150 160 170 180 LGGVTGSAAD IAIQAEQFAV IKKEMFRLNA EFTGQPIERI EADSDRDRWF TAAEALEYGF
190 VDHIITRAHV NGEAQ
ClpP1S98A (RV2461c)
-1 -1 MW: 22109 Da ε280: 14565 M cm 10 20 30 40 50 60 SMSQVTDMRS NSQGLSLTDS VYERLLSERI IFLGSEVNDE IANRLCAQIL LLAAEDASKD
70 80 90 100 110 120 ISLYINSPGG SISAGMAIYD TMVLAPCDIA TYAMGMAAAM GEFLLAAGTK GKRYALPHAR
130 140 150 160 170 180 ILMHQPLGGV TGSAADIAIQ AEQFAVIKKE MFRLNAEFTG QPIERIEADS DRDRWFTAAE
190 200 ALEYGFVDHI ITRAHVNGEA Q
95 8 Properties and sequences of the protein constructs
ClpP1T17E (RV2461c)
-1 -1 MW: 22153 Da ε280: 14565 M cm 10 20 30 40 50 60 MSQVTDMRSN SQGLSLEDSV YERLLSERII FLGSEVNDEI ANRLCAQILL LAAEDASKDI
70 80 90 100 110 120 SLYINSPGGS ISAGMAIYDT MVLAPCDIAT YAMGMAASMG EFLLAAGTKG KRYALPHARI
130 140 150 160 170 180 LMHQPLGGVT GSAADIAIQA EQFAVIKKEM FRLNAEFTGQ PIERIEADSD RDRWFTAAEA
190 200 LEYGFVDHII TRAHVNGEAQ HHHH
ClpP1T17A (RV2461c)
-1 -1 MW: 22095 Da ε280: 14565 M cm 10 20 30 40 50 60 MSQVTDMRSN SQGLSLADSV YERLLSERII FLGSEVNDEI ANRLCAQILL LAAEDASKDI
70 80 90 100 110 120 SLYINSPGGS ISAGMAIYDT MVLAPCDIAT YAMGMAASMG EFLLAAGTKG KRYALPHARI
130 140 150 160 170 180 LMHQPLGGVT GSAADIAIQA EQFAVIKKEM FRLNAEFTGQ PIERIEADSD RDRWFTAAEA
190 200 LEYGFVDHII TRAHVNGEAQ HHHH
ClpP2 (RV2460c)
-1 -1 MW: 24088 Da ε280: 10430 M cm 10 20 30 40 50 60 MNSQNSQIQP QARYILPSFI EHSSFGVKES NPYNKLFEER IIFLGVQVDD ASANDIMAQL
70 80 90 100 110 120 LVLESLDPDR DITMYINSPG GGFTSLMAIY DTMQYVRADI QTVCLGQAAS AAAVLLAAGT
130 140 150 160 170 180 PGKRMALPNA RVLIHQPSLS GVIQGQFSDL EIQAAEIERM RTLMETTLAR HTGKDAGVIR
190 200 210 KDTDRDKILT AEEAKDYGII DTVLEYRKLS AQTAHHHH
Δ11ClpP2 (RV2460c)
-1 -1 MW: 22283 Da ε280: 10430 M cm 10 20 30 40 50 60 SARYILPSFI EHSSFGVKES NPYNKLFEER IIFLGVQVDD ASANDIMAQL LVLESLDPDR
70 80 90 100 110 120 DITMYINSPG GGFTSLMAIY DTMQYVRADI QTVCLGQAAS AAAVLLAAGT PGKRMALPNA
130 140 150 160 170 180 RVLIHQPSLS GVIQGQFSDL EIQAAEIERM RTLMETTLAR HTGKDAGVIR KDTDRDKILT
190 200 AEEAKDYGII DTVLEYRKLS AQTA
96 8 Properties and sequences of the protein constructs
Δ13ClpP2 (RV2460c)
-1 -1 MW: 22056 Da ε280: 10430 M cm 10 20 30 40 50 60 SYILPSFIEH SSFGVKESNP YNKLFEERII FLGVQVDDAS ANDIMAQLLV LESLDPDRDI
70 80 90 100 110 120 TMYINSPGGG FTSLMAIYDT MQYVRADIQT VCLGQAASAA AVLLAAGTPG KRMALPNARV
130 140 150 160 170 180 LIHQPSLSGV IQGQFSDLEI QAAEIERMRT LMETTLARHT GKDAGVIRKD TDRDKILTAE
190 200 EAKDYGIIDT VLEYRKLSAQ TA
Δ18ClpP2 (RV2460c)
-1 -1 MW: 21482 Da ε280: 8940 M cm 10 20 30 40 50 60 SFIEHSSFGV KESNPYNKLF EERIIFLGVQ VDDASANDIM AQLLVLESLD PDRDITMYIN
70 80 90 100 110 120 SPGGGFTSLM AIYDTMQYVR ADIQTVCLGQ AASAAAVLLA AGTPGKRMAL PNARVLIHQP
130 140 150 160 170 180 SLSGVIQGQF SDLEIQAAEI ERMRTLMETT LARHTGKDAG VIRKDTDRDK ILTAEEAKDY
190 GIIDTVLEYR KLSAQTA
ClpP2S110A (RV2460c)
-1 -1 MW: 23610 Da ε280: 10430 M cm 10 20 30 40 50 60 SMNSQNSQIQ PQARYILPSF IEHSSFGVKE SNPYNKLFEE RIIFLGVQVD DASANDIMAQ
70 80 90 100 110 120 LLVLESLDPD RDITMYINSP GGGFTSLMAI YDTMQYVRAD IQTVCLGQAA AAAAVLLAAG
130 140 150 160 170 180 TPGKRMALPN ARVLIHQPSL SGVIQGQFSD LEIQAAEIER MRTLMETTLA RHTGKDAGVI
190 200 210 RKDTDRDKIL TAEEAKDYGI IDTVLEYRKL SAQTA
ClpS (RV1331)
-1 -1 MW: 12593 Da ε280: 26477 M cm 10 20 30 40 50 60 MAVVSAPAKP GTTWQRESAP VDVTDRAWVT IVWDDPVNLM SYVTYVFQKL FGYSEPHATK
70 80 90 100 110 LMLQVHNEGK AVVSAGSRES MEVDVSKLHA AGLWATMQQD RHHHHHHHHH H
97 8 Properties and sequences of the protein constructs
ClpS like protein (Rv0543c)
-1 -1 MW: 12106 Da ε280: 13980 M cm 10 20 30 40 50 60 MNRFLTSIVA WLRAGYPEGI PPTDSFAVLA LLCRRLSHDE VKAVANELMR LGDFDQIDIG
70 80 90 100 VVITHFTDEL PSPEDVERVR ARLAAQGWPL DDVRDREEHA ENLFQ
ClpC1 (RV3596c)
-1 -1 MW: 94282 Da ε280: 35870 M cm 10 20 30 40 50 60 MFERFTDRAR RVVVLAQEEA RMLNHNYIGT EHILLGLIHE GEGVAAKSLE SLGISLEGVR
70 80 90 100 110 120 SQVEEIIGQG QQAPSGHIPF TPRAKKVLEL SLREALQLGH NYIGTEHILL GLIREGEGVA
130 140 150 160 170 180 AQVLVKLGAE LTRVRQQVIQ LLSGYQGKEA AEAGTGGRGG ESGSPSTSLV LDQFGRNLTA
190 200 210 220 230 240 AAMEGKLDPV IGREKEIERV MQVLSRRTKN NPVLIGEPGV GKTAVVEGLA QAIVHGEVPE
250 260 270 280 290 300 TLKDKQLYTL DLGSLVAGSR YRGDFEERLK KVLKEINTRG DIILFIDELH TLVGAGAAEG
310 320 330 340 350 360 AIDAASILKP KLARGELQTI GATTLDEYRK YIEKDAALER RFQPVQVGEP TVEHTIEILK
370 380 390 400 410 420 GLRDRYEAHH RVSITDAAMV AAATLADRYI NDRFLPDKAI DLIDEAGARM RIRRMTAPPD
430 440 450 460 470 480 LREFDEKIAE ARREKESAID AQDFEKAASL RDREKTLVAQ RAEREKQWRS GDLDVVAEVD
490 500 510 520 530 540 DEQIAEVLGN WTGIPVFKLT EAETTRLLRM EEELHKRIIG QEDAVKAVSK AIRRTRAGLK
550 560 570 580 590 600 DPKRPSGSFI FAGPSGVGKT ELSKALANFL FGDDDALIQI DMGEFHDRFT ASRLFGAPPG
610 620 630 640 650 660 YVGYEEGGQL TEKVRRKPFS VVLFDEIEKA HQEIYNSLLQ VLEDGRLTDG QGRTVDFKNT
670 680 690 700 710 720 VLIFTSNLGT SDISKPVGLG FSKGGGENDY ERMKQKVNDE LKKHFRPEFL NRIDDIIVFH
730 740 750 760 770 780 QLTREEIIRM VDLMISRVAG QLKSKDMALV LTDAAKALLA KRGFDPVLGA RPLRRTIQRE
790 800 810 820 830 840 IEDQLSEKIL FEEVGPGQVV TVDVDNWDGE GPGEDAVFTF TGTRKPPAEP DLAKAGAHSA
850 GGPEPAARLE VLFQ
98 8 Properties and sequences of the protein constructs
ClpC2 (RV2667)
-1 -1 MW: 27449 Da ε280: 8480 M cm 10 20 30 40 50 60 MPEPTPTAYP VRLDELINAI KRVHSDVLDQ LSDAVLAAEH LGEIADHLIG HFVDQARRSG
70 80 90 100 110 120 ASWSDIGKSM GVTKQAAQKR FVPRAEATTL DSNQGFRRFT PRARNAVVAA QNAAHGAASS
130 140 150 160 170 180 EITPDHLLLG VLTDPAALAT ALLQQQEIDI ATLRTAVTLP PAVTEPPQPI PFSGPARKVL
190 200 210 220 230 240 ELTFREALRL GHNYIGTEHL LLALLELEDG DGPLHRSGVD KSRAEADLIT TLASLTGANA
250 AGATDAGATD AGHHHHHH
ClpX (RV2457c)
-1 -1 MW: 47577 Da ε280: 18255 M cm 10 20 30 40 50 60 MARIGDGGDL LKCSFCGKSQ KQVKKLIAGP GVYICDECID LCNEIIEEEL ADADDVKLDE
70 80 90 100 110 120 LPKPAEIREF LEGYVIGQDT AKRTLAVAVY NHYKRIQAGE KGRDSRCEPV ELTKSNILML
130 140 150 160 170 180 GPTGCGKTYL AQTLAKMLNV PFAIADATAL TEAGYVGEDV ENILLKLIQA ADYDVKRAET
190 200 210 220 230 240 GIIYIDEVDK IARKSENPSI TRDVSGEGVQ QALLKILEGT QASVPPQGGR KHPHQEFIQI
250 260 270 280 290 300 DTTNVLFIVA GAFAGLEKII YERVGKRGLG FGAEVRSKAE IDTTDHFADV MPEDLIKFGL
310 320 330 340 350 360 IPEFIGRLPV VASVTNLDKE SLVKILSEPK NALVKQYIRL FEMDGVELEF TDDALEAIAD
370 380 390 400 410 420 QAIHRGTGAR GLRAIMEEVL LPVMYDIPSR DDVAKVVVTK ETVQDNVLPT IVPRKPSRSE
430 RRDKSAENLF Q
Potential ClpX substrate with C terminal VAA motif (Rv2308)
-1 -1 MW: 26775 Da ε280: 35410 M cm 10 20 30 40 50 60 SMRADMSVTS MLDREVYVYA EVDKLIGLPA GTAKRWINGY ERGGKDHPPI LRVTPGATPW
70 80 90 100 110 120 VTWGEFVETR MLAEYRDRRK VPIVRQRAAI EELRARFNLR YPLAHLRPFL STHERDLTMG
130 140 150 160 170 180 GEEIGLPDAE VTIRTGQALL GDARWLASIA TPGRDEVGEA VIVELPVDKA FPEIVINPSR
190 200 210 220 230 YSGQPTFVGR RVSPVTIAQM VDGGEEREDL AADYGLSLKQ IQDAIDYTKK YRLARLVAA
99 8 Properties and sequences of the protein constructs
Potential ClpX substrate with C terminal LAA motif (Rv2961)
-1 -1 MW: 14574 Da ε280: 19480 M cm 10 20 30 40 50 60 SVEHGNPHDA PQLAPAVERI TTRAGRPPGT VTADRGYGEK RVEDDLHDLG VRTVAIPRKG
70 80 90 100 110 120 RPSQARRAEE QRPSFRRTVK WRTGSEGRIS TLKRNYGWNR SCIDGTEGTR IWTRHGILTH
130 NLIKISSLAA
Potential ClpX substrate with C terminal VAA motif (Rv3832c)
-1 -1 MW: 20853 Da ε280: 22585 M cm 10 20 30 40 50 60 SMAMNLLHRR HCSSAGWEKA VANQLLPWAL QHVELGPRTL EIGPGYGATL QALLGLTASL
70 80 90 100 110 120 TAVEVDNSMV ERLNRRYGQR ARIIRGDGTQ TGLPDDHFTS VVCFTMLHHV ASAQLQDQLF
130 140 150 160 170 180 AEAYRVLQPG GVFAGSDGVP SLPFRLIHIA DTYTPIAPAD LPGRLRAVGF TDIHVDVAGA
190 RLRWRATKPV AA
MDH-ssrA (RV1240)
-1 -1 MW: 35779 Da ε280: 33460 M cm 10 20 30 40 50 60 SVSASPLKVA VTGAAGQIGY SLLFRLASGS LLGPDRPIEL RLLEIEPALQ ALEGVVMELD
70 80 90 100 110 120 DCAFPLLSGV EIGSDPQKIF DGVSLALLVG ARPRGAGMER SDLLEANGAI FTAQGKALNA
130 140 150 160 170 180 VAADDVRVGV TGNPANTNAL IAMTNAPDIP RERFSALTRL DHNRAISQLA AKTGAAVTDI
190 200 210 220 230 240 KKMTIWGNHS ATQYPDLFHA EVAGKNAAEV VNDQAWIEDE FIPTVAKRGA AIIDARGASS
250 260 270 280 290 300 AASAASATID AARDWLLGTP ADDWVSMAVV SDGSYGVPEG LISSFPVTTK GGNWTIVSGL
310 320 330 340 EIDEFSRGRI DKSTAELADE RSAVTELGLI AADSHQRDYA LAA
100 8 Properties and sequences of the protein constructs
MDH (RV1240)
-1 -1 MW: 34408 Da ε280: 31970 M cm 10 20 30 40 50 60 SVSASPLKVA VTGAAGQIGY SLLFRLASGS LLGPDRPIEL RLLEIEPALQ ALEGVVMELD
70 80 90 100 110 120 DCAFPLLSGV EIGSDPQKIF DGVSLALLVG ARPRGAGMER SDLLEANGAI FTAQGKALNA
130 140 150 160 170 180 VAADDVRVGV TGNPANTNAL IAMTNAPDIP RERFSALTRL DHNRAISQLA AKTGAAVTDI
190 200 210 220 230 240 KKMTIWGNHS ATQYPDLFHA EVAGKNAAEV VNDQAWIEDE FIPTVAKRGA AIIDARGASS
250 260 270 280 290 300 AASAASATID AARDWLLGTP ADDWVSMAVV SDGSYGVPEG LISSFPVTTK GGNWTIVSGL
310 320 330 EIDEFSRGRI DKSTAELADE RSAVTELGLI
Adenylate kinase (RV0733)
-1 -1 MW: 20919 Da ε280: 8940 M cm 10 20 30 40 50 60 MRVLLLGPPG AGKGTQAVKL AEKLGIPQIS TGELFRRNIE EGTKLGVEAK RYLDAGDLVP
70 80 90 100 110 120 SDLTNELVDD RLNNPDAANG FILDGYPRSV EQAKALHEML ERRGTDIDAV LEFRVSEEVL
130 140 150 160 170 180 LERLKGRGRA DDTDDVILNR MKVYRDETAP LLEYYRDQLK TVDAVGTMDE VFARALRALG
KENLFQ
Propeptide cleavage site SHD Residues of the catalytic triad Amino acid exchange LAA Substrate recognition motif for ClpX
H4 or ENLFQ Sequence stretches that are not part of the encoded sequence but result from addition of affinity tags or protease cleavage of affinity tags
101
8.3 Structures of activator and peptide substrates
Z-Leu-Leu-H CH3
CH CH
HC CH O H2C CH3
HC C O NH C CH H
CH CH2 C CH NH C
O H2C CH3 O
CH
CH3
Suc-LLVY-AMC O
C OH
CH2 CH2 CH3
NH C HC H3C HC O CH2
C O
HN O H3C OH
H3C CH C CH CH3 CH C
CH CH2 NH HC HC CH
H3C C NH C CH
O CH CH2
O C
NH
CH C
HC CH
C C
O C CH3
C CH
O
102 8 Structure of activator and peptide substrates
Suc-GGL-AMC O
C
O CH
O C C
C CH O CH 3 C CH2 HO C CH2 HC CH
C NH
H2C NH
C CH2 NH C O NH C HC O
O CH2
CH CH3
H3C
Glt-AAF-AMC
CH CH H3C
HC CH C CH
C CH CH C C O
H2C NH C C O
O CH C CH CH
C NH O
H3C NH HC
O CH C CH3
C NH O
O CH2 CH2
C CH2
HO
103 8 Structure of activator and peptide substrates
BOC-VLK-AMC CH3
CH CH O O
O H2C CH3 O HC C C
H3C O NH C CH NH C C C CH
C C CH NH C CH NH CH C
H3C CH3 O CH O CH2 CH3
H3C CH3 H2C
CH2
H2C
NH2
Ac-DEVD-AMC CH O 3
NH C C HO HC O CH2
C NH O H3C
O CH C CH CH3
CH2 CH2 NH HC
HO C C NH O
O O CH C CH CH CH3
H2C NH C C HC
C O CH C CH2
HO O C
O
Protease cleavage site
104
9 Acknowledgment
Ohne die Hilfe der folgenden Personen wäre die Arbeit in dieser Weise nicht möglich gewesen.
Eilika Weber-Ban Rudolf Glockshuber Peter Sander Kürşad Turgay
Frank Striebel Wolfgang Kress Namit Ranjan Ethan Guth Frank Imkamp Dennis Öscelik Markus Sutter Jonas Barandun Marcel Bolten Cyrille Delley Julia Leodolter Zeljka Maglica Jürg Laederach Michal Ziemski Michael Burger
Sebastian Klinge Daniel Böhringer
Besonderer Dank gilt meinen Eltern, meiner Partnerin, den Mitbewohner/innen und Freunden.
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10 Curriculum Vitae
Personal data Jannis Warweg 26.09.1982, Duisburg, Germany German citizenship
Scientific career
2010 – 2014 Ph.D. Studies Laboratory of Prof. Weber-Ban Institute of Molecular Biology and Biophysics ETH Zürich, Switzerland
2009 “Diplomarbeit“(similar to Master's thesis) Laboratory of Prof. Ficner Department for Molecular Structural Biology Georg-August-Universität Göttingen, Germany
2006 – 2007 Internship in molecular biology and genetics Laboratory of Prof. Fritz Institute for Microbiology and Genetics Georg-August-Universität Göttingen, Germany
Education
2005 – 2009 Study of biology Georg-August-Universität Göttingen, Germany Major: biochemistry Minor: immunology, microbiology
2003 – 2005 Study of biotechnology Technische Universität Hamburg-Harburg, Germany
2002 – 2003 Civilian service Haus Alstertal/Alten und Pflegepension GmbH Hamburg, Germany
1993 – 2002 Secondary school Stadtgymnasium Detmold, Germany
Selected work experience
2010 – 2011 Organizing committee of the 8th Annual Retreat of the MLS PhD Program Bergün, Switzerland (2011)
2007 Student assistant Laboratory of Prof. Steinfeld Department of Pediatrics and Absolescent Medicine Göttingen, Germany
106