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TTHHÈÈSSEE
En vue de l'obtention du
DDOOCCTTOORRAATT DDEE LL’’UUNNIIVVEERRSSIITTÉÉ DDEE TTOOUULLOOUUSSEE
Délivré par l'Université Toulouse III - Paul Sabatier
Discipline ou spécialité : Microbiologie et Génétique moléculaire
JURY
Dr Arturo MUGA, Professeur, Universidad del Pais Vasco, Espagne
Dr Alessandra POLISSI, Professeur, Università degli Studi di Milano-Bicocca, Italie
Dr Peter FALLER, Professeur, Laboratoire de Chimie de Coordination, Toulouse
Dr Jan Willem DE GIER, Professeur associé, Stockholm University, Suède
Dr Marie-Pierre CASTANIE-CORNET, Maître de Conférence, Université de Toulouse
Dr Pierre GENEVAUX, Directeur de Recherche, CNRS, Toulouse
Ecole doctorale : Biologie, Santé, Biotechnologie (BSB)
Unité de recherche : Laboratoire de Microbiologie et Génétique Moléculaire (LMGM)
Directeur(s) de Thèse : Dr Pierre GENEVAUX (Directeur), Dr Marie-Pierre CASTANIE-CORNET (co-
Directeur)
Membre invité: Dr Joen LUIRINK (co-Superviseur)
Rapporteurs : Dr Arturo MUGA, Dr Alessandra POLISSI
Présentée et soutenue par Nicolas Bruel Le Mardi 2 avril 2013
Titre : Hsp33 controls Elongation Factor-Tu stability and allows Escherichia coli growth in
the absence of the major DnaK and TriggerFactor chaperones
2
Contents
ABBREVIATIONS .............................................................................................................................. 5
RÉSUMÉ .............................................................................................................................................. 6
SUMMARY ......................................................................................................................................... 7
INTRODUCTION ................................................................................................................................ 9
Part I: Protein folding and molecular chaperone functions .............................................................. 9
1- Protein folding within the cell ............................................................................................. 9
2- Role of the mRNA and the translation machinery ......................................................... 10
a- Rare codons, translational pausing and protein folding ................................................. 10
b- tRNA concentration modulates nascent chain protein folding ........................................ 11
c- Roles of EF-Tu ................................................................................................................. 12
d- Chaperone function of the ribosome ................................................................................ 13
e- Molecular chaperones, concept, functions and universality............................................ 14
Part II: Chaperone-assisted de novo protein folding ...................................................................... 15
1- The ribosome-bound Trigger Factor ............................................................................... 15
a- Structure / interaction with the ribosome ........................................................................ 15
b- TF substrate interaction and cellular functions ............................................................... 19
2- DnaKJE cycle and interactors .......................................................................................... 21
a- DnaKJE structure and cycle ............................................................................................ 21
b- Functions of DnaK ........................................................................................................... 24
c- Interactors in the cell ....................................................................................................... 24
3- The chaperonin GroESL ................................................................................................... 25
a- Structure of the GroESL complex and chaperonin cycle ................................................. 25
b- Interactors in the cell ....................................................................................................... 26
4- Interplay between TF/DnaKJE/GroESL during de novo protein folding .................... 27
Part III: Interplays between TF, DnaKJE and GroESL and other chaperones during protein
export .............................................................................................................................................. 29
1- Sec Translocation system .................................................................................................. 30
2- Twin Arginin translocation system .................................................................................. 31
Part IV: Chaperone-mediated response to protein misfolding and aggregation ............................. 33
1- Molecular chaperone networks and protein disaggregation ......................................... 33
a- Presentation of other chaperones involved in protein disaggregation ............................ 33
b- Chaperone networks involved in protein disaggregation ................................................ 34
3
2- Hsp33 as a member of the chaperone network ............................................................... 37
a- Hsp33 is involved in the oxidative stress response .......................................................... 37
b- Structure of Hsp33 ........................................................................................................... 38
c- Activation cycle of Hsp33 ................................................................................................ 39
d- Hsp33 deletion and functions in the cell .......................................................................... 42
MATERIALS AND METHODS ....................................................................................................... 44
1- Strains and plasmids ......................................................................................................... 44
a- Bacterial Strains, phages, and culture conditions ........................................................... 44
b- Plasmid construction ....................................................................................................... 45
2- In vivo experiments ............................................................................................................ 46
a- Bacterial viability assay and genetic experiments ........................................................... 46
b- Isolation of protein aggregates and cell fractionation .................................................... 46
c- In vivo pull-down assay ................................................................................................... 47
3- In vitro experiments ........................................................................................................... 47
a- Western blot analysis ....................................................................................................... 47
b- In vitro translation and cross-linking experiments .......................................................... 48
c- Pulse-chase and immunoprecipitation analyses .............................................................. 48
RESULTS ........................................................................................................................................... 50
1- Hsp33 overproduction supports bacterial growth and prevents protein
aggregation in the absence of TF and DnaK .......................................................................... 50
2- Hsp33 function is critical in the absence of both TF and DnaK .................................... 54
3- Hsp33 specifically interacts with EF-Tu .......................................................................... 58
4- Hsp33 triggers EF-Tu degradation by the stress protease Lon ..................................... 63
5- EF-Tu inhibition helps bacterial growth in the absence of TF and DnaK ................... 65
DISCUSSION .................................................................................................................................... 69
REFERENCES ................................................................................................................................... 74
REMERCIEMENTS .......................................................................................................................... 93
4
5
ABBREVIATIONS
AAA+: ATPases Associated with a variety of cellular Activities
DNA: DesoxyriboNucleic Acid
EF-Tu: Elongation Factor-Tu
EF-Ts: Elongation Factor-Ts
FRET: Fluorescence resonance energy transfer
HSP: Heat Shock Protein
IPTG: isopropyl β-D-1-thiogalactopyranoside
JDP: J-domain protein
kDa: KiloDalton
NAC: Nascent polypeptide-Associated Complex
NBD: Nucleotide Binding Domain
NEF: Nucleotide Exchange Factor
NMR: Nuclear Magnetic Resonance
OMP: Outer-Membrane β-barrel Protein
PPIase: Peptidyl-Prolyl cis/trans Iisomerase
RAC: Ribosome-Associated Complex
REMP : Redox Enzyme Maturation Protein
RNA: RiboNucleic Acid
SBD: Substrate Binding Domain
sHSP: small Heat Shock Protein
SRP: Sgnal Recognition Particle
Tat: Twin-Arginine Translocon
TF: Trigger Factor
6
RÉSUMÉ
Le repliement intracellulaire des protéines nouvellement synthétisées est assisté par
des réseaux cellulaires de protéines chaperons. Chez Escherichia coli, la coopération entre les
protéines chaperons Trigger Factor (TF) et DnaK est prédominante dans ce processus. En
accord avec ceci, la délétion simultanée des gènes codants pour ces deux protéines chaperons
conduit à une croissance bactérienne très réduite et à l’accumulationd’un grand nombre de
protéines cytoplasmiques sous forme d’agrégats. Au cours de cette étude, nous avons utilisé
ces phénotypes afin de mettre en évidence des interactions potentielles au sein du réseau de
protéines chaperons in vivo. Nous avons montré que la perte des protéines chaperons TF et
DnaK, et donc des voies de repliements dans lesquelles elles sont impliquées, pouvait être
secourue de façon efficace par la surexpression du chaperon Hsp33, connu pour être activable
en réponse à un stress oxydatif sévère. En outre, la délétion du gène hslO, codant pour Hsp33,
n’était plus tolérée en l’absence de TF et DnaK. Cependant, en comparaison avec d’autres
protéines chaperons comme GroESL ou SecB, la suppression de ces phénotypes par Hsp33
n’a pas pu être attribuée à un éventuel chevauchement de fonctions avec DnaK et TF. Au
contraire, nos résultats montraient qu’ Hsp33 surexprimée fixait de façon spécifique le facteur
d’élongation-Tu (EF-Tu) et favorisait sa dégradation par la protéase Lon. Cette action
synergétique entre Hsp33 et Lon était responsable du rétablissement de la croissance
bactérienne en l’absence de TF et DnaK, possiblement via le rétablissement du couplage entre
la vitesse de traduction et les capacités de repliement des protéines nouvellement synthétisées
du double mutant. Afin de soutenir cette hypothèse, nous avons ensuite montré que la
surexpression de la toxine HipA qui inhibe EF-Tu, était aussi capable de supprimer le
phénotype de thermosensibilité et de réduire significativement l’agrégation des protéines en
l’absence de TF et DnaK.
7
SUMMARY
Intracellular de novo protein folding is assisted by cellularnetworks of molecular
chaperones. In Escherichia coli, cooperationbetween the chaperones Trigger Factor (TF) and
DnaK iscentral to this process. Accordingly, the simultaneous deletion of both chaperone-
encoding genes leads to severe growth andprotein folding defects. Herein, we took advantage
of suchdefective phenotypes to further elucidate the interactions ofchaperone networks in
vivo. We show that disruption of theTF/DnaK chaperone pathway is efficiently rescued by
over-expressionof the redox-regulated chaperone Hsp33. Consistentwith this observation, the
deletion of hslO, the Hsp33 structuralgene, is no longer tolerated in the absence of the
TF/DnaK pathway.However, in contrast with other chaperones like GroESL orSecB,
suppression by Hsp33 was not attributed to its potentialoverlapping general chaperone
function(s). Instead, we showthat over-expressed Hsp33 specifically binds to elongation
factor-Tu (EF-Tu) and targets it for degradation by the proteaseLon. This synergistic action of
Hsp33 and Lon was responsiblefor the rescue of bacterial growth in the absence ofTFand
DnaK,by presumably restoring the coupling between translation andthe downstream folding
capacity of the cell. In support of thishypothesis, we show that over-expression of the stress-
responsivetoxin HipA, which inhibits EF-Tu, also rescues bacterialgrowth and protein folding
in the absence of TF and DnaK.
8
INTRODUCTION
9
INTRODUCTION
Part I: Protein folding and molecular chaperone functions
1- Protein folding within the cell
Proteins need to reach their native three-dimensional structure to become active. This
folding process is determined by the primary amino-acid sequence (Anfinsen, 1973). Protein
folding is not a random event but follows a specific energetic landscape through which
protein gets some native-like structure as folding intermediate. These numerous pathways
drive the protein to its native structure at the most stable energetic point (Dobson, 2004;
Onuchic & Wolynes, 2004). Even if small single domain proteins, having less than 100
residues, can reach their native state without such intermediates (Jackson, 1998), folding of
multi-domain proteins generally need parallel routes of “nucleation-condensation” of a small
number of key residues, which allow the protein to condensate up to its native structure
(Radford et al, 1992). Within the cell, such folding events coupled to the crowded cellular
environment (i.e. proteins, nucleic-acids, complex sugars in high concentration ~400g/L (Ellis
& Minton, 2003)) can lead to protein misfolding and aggregation, often associated with
human diseases (Fig. 1) (Barral et al, 2004; Dobson, 2001; Thomas et al, 1995). To counteract
such noxious pathways, intricate quality control systems have been maintained in all
kingdoms of life (Hartl & Hayer-Hartl, 2002). Such quality controls initially take place during
translation of nascent peptide chains by direct action on the ribosome and/or translational
factors such as tRNAs, the elongation factor Tu (EF-Tu) and the mRNA itself (Caldas et al,
1998; Fedyunin et al, 2012; Komar, 2009; Voisset et al, 2008). Other factors known as
“Molecular Chaperones” actively participate in this folding process by interacting directly
with polypeptide chains in a co- and/or post-translational manner to prevent their aggregation
(Fig. 1).
10
2- Role of the mRNA and the translation machinery
a- Rare codons, translational pausing and protein folding
The folding of nascent peptide chain is incorporated in the kinetics of mRNA
translation during its elongation (Komar, 2009; Komar et al, 1999; Krasheninnikov et al,
1991; Wolin & Walter, 1988). In fact, mRNA sequences contain regions composed of rare
codons slowing down the rate of translation and influencing the folding of the nascent peptide
chain during its translation. Presence of these rare codons is due to the distribution of
aminoacyl-tRNA isoacceptor species in a given organism (Chavancy & Garel, 1981; Garel,
1974; Ikemura, 1981). Such a variation in aminoacyl-tRNA isoacceptor concentration creates
a strong codon bias within any given organism (Sharp et al, 1988). Pause sites were identified
Chaperones
Nascent polypeptide Intermediate Native protein
Aggregates
Fibrillar Amyloids
Fig. 1: Folding pathways of a newly synthesized protein. Proteins can pass through folding intermediates
before reaching their native structure. Misfolding events such as aggregation may occur at each folding step.
Molecular chaperones prevent this aggregation and favor the folding of nascent polypeptides until their native
conformation.
11
by micrococcal nuclease protection assay of mRNA during translation (Hollingsworth et al,
1998; Wolin & Walter, 1988). Indeed, a frequently used codon, which corresponds to a high
concentration in its aminoacyl-tRNA isoacceptor, is translated faster than one infrequently
used. So that kind of punctuation into the mRNA during the translation facilitates, to some
extent, the co-translational folding of the nascent polypeptide because of the positioning of
these pause sites at specific domains in the encoded protein. Remarkably, a correlation
between codon abundance and domain boundaries has been identified (Purvis et al, 1987). In
addition, it has been found that α-helices are frequently associated with high-frequency
codons, whereas β-strands and random coils are preferentially coded by rare codon regions
(Thanaraj & Argos, 1996). Moreover, the location of these pause sites is highly conserved in
some protein families including cytochromes C, globins, γ-B crystallins and chloramphenicol
acetyltransferase (Krasheninnikov et al, 1989; Widmann et al, 2008). To study the relevance
of such pause sites, silent mutations of mRNAs coding the Escherichia coli chloramphenicol
acetyltransferase, Saccharomyces cerevisiae anthranilate synthase indole-3-glycerol
phosphate synthase and Echinococcus granulosus fatty acid binding protein, replacing rare
codons by common ones showed a decrease in their specific activities when compared to the
wild type proteins (Cortazzo et al, 2002; Crombie et al, 1994; Komar et al, 1999).
Furthermore, the proteins produced from these silently mutated mRNAs were misfolded.
Although the kinetics of translation appear to play a significant role in the co-translational
folding of nascent peptide chain, but there are additional factors, including tRNA
concentration, Elongation Factor-Tu (EF-Tu), and ribosome, which participate directly or
indirectly to the folding process.
b- tRNA concentration modulates nascent chain protein folding
tRNAs bring amino-acids to their cognate codons. Remarkably, it has been shown
both in vitro and in vivo that the concentration of tRNAs acceptors can drastically modulate
the rate of translation. Indeed, using an in vitro translational experiment, Anderson first
showed that translation of a poly-U by tRNAPhe
was inhibited at high concentration of
tRNAPhe
(tRNAPhe
/70S ribosome ratio of 7) (Anderson, 1969). In addition, a different study
pointed out that in the presence of a constant concentration of exogenous tryptophan, the rate
of incorporation of this amino acid was considerably diminished when tRNATrp
concentration
was too high, following over-expression of tRNATrp
from a plasmid. Such over-expression
changed the ratio of uncharged tRNATrp
/ [Trp-tRNATrp
] and the pool of uncharged tRNATrp
was found to inhibit Trp incorporation into proteins (Rojiani et al, 1990). Remarkably, they
12
also showed that uncharged tRNATrp
could inhibit incorporation of other amino acids, such as
Lys and Cys (Rojiani et al, 1990). A third team studied the effect of low-abundant tRNAs
upregulation on the folding and solubility of E. coli proteins (Fedyunin et al, 2012). In this
case, upregulation of argU which encodes tRNA4Arg
that pairs with arginine codons AGA and
AGG, ileY encoding for tRNA2Ile
that pairs with AUA codon or leuW encoding tRNA3Leu
that
pairs with CUA codon, modifying the rate of translation of these rare condons, was
responsible of the aggregation of many proteins such as molecular chaperones which had been
found to suffer from this misfolding effect (Fedyunin et al, 2012). This approach revealed the
importance of the translational rate during co-translational folding of all the cellular proteome
and the role of tRNA isoacceptors concentration in this translational rate.
c- Roles of EF-Tu
In E. coli, EF-Tu is the most abundant protein in the cell, representing 5 to 10% of the
total proteins (Furano, 1975). EF-Tu has a GTPase activity and brings aminoacyl-tRNA to the
A site of the ribosome in a GTP-dependent manner. GDP is replaced through interaction with
the nucleotide exchange factor EF-Ts (Agirrezabala & Frank, 2009). EF-Tu and its eukaryotic
homolog EF-1α have many other cellular functions. EF-Tu was found to be an essential
subunit of the Qβ phage replicative complex (Blumenthal et al, 1972). EF-Tu can also
stimulate RNA synthesis by interacting with the transcriptional apparatus (Travers et al,
1970). In Bacillus subtilis, EF-Tu plays an additional role in cell shape maintenance,
interacting with the actin-like MreB (Defeu Soufo et al, 2010). This role was also identified
for EF-1α, as it can bind actin filaments and microtubules and influence the assembly and the
stability of these cytoskeletal polymers (Shiina et al, 1994; Yang et al, 1990). In addition,
Caldas and coworkers also found that EF-Tu could reactivate chemically-unfolded citrate
synthase and α-glucosidase with the same efficiency than DnaKJE in vitro, thus suggesting
chaperone function (Caldas et al, 1998). The eukaryotic EF-1α chaperone activity was also
identified using Phenylalanyl-tRNA synthase and seryl-tRNA synthase as substrates. In this
case, it was shown that the mammalian EF-1α (called eEFiA) could refold and restore the
enzymatic activity of these two aminoacyl-tRNA synthase (Lukash et al, 2004). In conclusion,
EF-Tu does not only bring aminoacyl-tRNAs to the ribosome A site but participates in a
plethora of cellular processes, most likely including protein folding. The last compound of the
translational machinery to have a chaperone function protecting and actively participating in
the co-translational folding of the nascent chain protein is the ribosome itself.
13
d- Chaperone function of the ribosome
Ribosomes represent 30% of the total cell mass with up to 105 and 10
6 ribosomes in a
bacteria and a mammalian cell respectively (Bashan & Yonath, 2008). It is a
ribonucleoprotein complex composed of 2 subunits. The small subunit (30S in bacteria; 40S
in eukaryote) decodes the mRNA during translation and the large subunit (50S in bacteria;
60S in eukaryote) contains the peptidyl transferase center and the ribosomal exit tunnel
through which nascent polypeptides will emerge and be released into the cytosol. The length
of the exit tunnel is about 80 Å to 100 Å with a diameter comprised between 10 Å at its
narrowest point and 20 Å at its widest point. (Ban et al, 2000; Harms et al, 2001; Nissen et al,
2000). These dimensions allow the ribosome to contain a nascent chain of 30 amino-acids in
an extended conformation or 60 amino-acids as an α-helix (Malkin & Rich, 1967; Picking et
al, 1992; Voss et al, 2006). During translation, nascent chain interacts with this exit tunnel
composed of 23S rRNA and proteins L4, L22 and L23 in bacteria (Ban et al, 2000; Harms et
al, 2001; Kosolapov & Deutsch, 2009; Lu & Deutsch, 2005; Lu et al, 2007; Nissen et al,
2000). Such interactions can regulate kinetics of translation and induce translational pausing,
thus facilitating co-translational folding of the nascent peptide chain or its interaction with
ribosome associated factors such as the molecular chaperone Trigger Factor in bacteria (TF)
or the signal recognition particle (SRP) involved in the co-translational targeting of secretory
or membrane proteins to the sec translocon (Eisner et al, 2006; Kramer et al, 2009; Marin,
2008). Moreover, the ribosomal exit tunnel directly participates to the folding of nascent
chains by promoting its compaction or its helical conformation in the lower and upper part of
the tunnel. The middle part of the tunnel is constricted by L4 and L22 proteins (Lu &
Deutsch, 2005). This constriction site avoids protein folding in this area and, interacting
directly with the nascent chain, may modulate the docking site shared by TF or SRP and then
allow the recruitment of each of them to the ribosome exit tunnel (Gu et al, 2003; Houben et
al, 2005; Ullers et al, 2003). The exit pore of the tunnel also plays a role in the co-translational
folding of the nascent chain allowing the formation of tertiary structures such as β-hairspins
(Kosolapov & Deutsch, 2009).
The ribosome itself also possesses some chaperone activity. Indeed, the 50S ribosomal
subunit can interact post-translationally with unfolded proteins and efficiently participate to
their folding (Kudlicki et al, 1994; Voisset et al, 2008). The interaction site of the ribosome
with unfolded protein had been identified near from the peptidyl transferase center, in the
domain V of the 23S rRNA (Chattopadhyay et al, 1996; Pal et al, 1997). The ribosome can
14
also participate in the co-translational folding of newly synthesized proteins in a different
way. Indeed, Kaiser and coworkers (2011) studied the active folding activity of the ribosome
exit tunnel following the folding of the T4 lysozyme and found that the ribosome could
prevent misfolding of the nascent chain by modulating its folding rate (Kaiser et al, 2011).
Furthermore, this exit pore is a docking site for many key proteins involved in the enzymatic
processing of the nascent chain such as the methionine aminopeptidase and the peptide
deformylase (Kramer et al, 2009), for targeting factors such as SRP and SecA, ribosome
associated chaperones like Trigger Factor in bacteria or Ssb/Ssz/Zuotin chaperone triad,
ribosome-associated complex (RAC) and nascent polypeptide-associated complex (NAC) for
the eukaryote S. cerevisiae(Eisner et al, 2006; Ferbitz et al, 2004; Preissler & Deuerling,
2012).
e- Molecular chaperones, concept, functions and universality
Molecular chaperones facilitate protein folding and prevent and/or rescue noxious
protein aggregation. They are involved in many cellular pathways such as: de novo protein
folding, protein disaggregation, protein translocation through a biological membrane,
oligomerisation and control of protein-protein interaction, as well as protein turn over
(Hendrick & Hartl, 1993). Molecular chaperones are conserved in all the three kingdoms of
life, represented by different protein families, and are known to function in a sequential
manner on their substrates co-translationally and post-translationally (Frydman et al, 1994;
Langer et al, 1992; Siegers et al, 1999).
A first group of molecular chaperones interacts with the ribosome and/or the nascent
chain to facilitate co-translational folding. Bacteria use the chaperone Trigger Factor,
interacting at the same time with the ribosome and the nascent chain (Hesterkamp et al, 1996).
The Hsp70 chaperone DnaK from E. coli can also interact with nascent polypeptides to assist
them during their co-translational folding (Teter et al, 1999). Although the eukaryotic
chaperone system does not have Trigger Factor, they do have two different kinds of ribosome
associated complexes acting at the ribosome exit pore to support co-translational folding
(Preissler & Deuerling, 2012). Indeed, in Saccharomyces cerevisiae, the ribosome-associated
complex (RAC) composed of the Hsp70/Hsp40 chaperone system Ssz/Zuotin and another
Hsp70 named Ssb and the heterodimeric nascent chain-associated complex (NAC) may
parallel Trigger Factor function in the folding of the nascent polypeptide chains (Preissler &
Deuerling, 2012).Hsp70 may also bind substrates in a post-translational way to mediate their
folding (Deuerling et al, 1999; Teter et al, 1999). The chaperonins represent the second group
15
of molecular chaperones, mostly participating in the post-translational folding of proteins.
They are structured as double-ring complexes forming a central cavity in each ring, binding
and encapsulating unfolded proteins in one ring, which protect the substrate from the crowded
environment of the cell and help it to fold (Mayhew et al, 1996; Weissman et al, 1996). Other
types of chaperones are more specifically involved in disaggregation, as it’s the case for
Hsp90 and Hsp100, HtpG and ClpB in E. colirespectively.
In the following parts of the introduction, the structures, the mechanisms and the
functions of the major E. coli chaperones involved in de novo protein folding, namely TF,
DnaKJE and GroESL, will be presented. In addition, their functional cooperation with other
chaperone systems during protein export, protein disaggregation and in response to severe
oxidative stress will be developed.
Part II: Chaperone-assisted de novo protein folding
1- The ribosome-bound Trigger Factor
a- Structure / interaction with the ribosome
The ribosome-bound TF is the first molecular chaperone to interact with most of the
newly synthesized polypeptides (Valent et al, 1995). It is a 48kDa protein constituted of 3
distinct domains elongated in a Dragon-like shaped structure (Ferbitz et al, 2004; Hesterkamp
& Bukau, 1996; Zarnt et al, 1997) (Fig. 2A). The N-terminal domain of TF, composed of
amino acids 1 to 149, is the ribosome binding domain, which also contributes to TF chaperone
activity. This domain possesses some structural homology with the molecular chaperone
Hsp33 except for the additional loop involved in the binding of TF to the ribosome via the TF
signature motif “GFRxGxxP” (Genevaux et al, 2004; Hoffmann et al, 2010; Kramer et al,
2002; Kristensen & Gajhede, 2003). A long linker (aa 111 to 149) is found between the
ribosome binding domain and the peptidyl-prolyl cis/trans isomerase domain (PPIase) of TF.
The PPIase domain (aa 150 to 245) is related to the FK506 binding protein family of PPIase
(Hesterkamp & Bukau, 1996; Scholz et al, 1997; Stoller et al, 1995). This domain is
dispensable for chaperone function of TF in vivo, but it can enhance chaperone activity of TF
as an auxiliary binding site of substrates (Genevaux et al, 2004; Hoffmann et al, 2006; Kramer
et al, 2004a; Lakshmipathy et al, 2007; Merz et al, 2006). The C-terminal domain (aa 246 to
432), forming the body of the dragon, is found between the N-terminal domain (the tail of the
16
dragon) and the PPIase domain (the head of the dragon). It interacts with the long linker (aa
111 to 149), which stabilize its structure containing 2 protruding helical “arms” (Ferbitz et al,
2004; Martinez-Hackert & Hendrickson, 2007; Merz et al, 2006). This domain forms the main
chaperone module of TF (Merz et al, 2006).
TF is conserved in bacteria and chloroplasts, but its PPIase domain may be absent in
some species (Hoffmann et al, 2010; Kristensen & Gajhede, 2003). Some bacteria, such as
Desulfitobacterium hafniense, even possess several TF and TF-like molecular chaperones
with specific functions (Maillard et al, 2011). Indeed, among the 3 TF homologs present in
this bacterium, the TF-like chaperone PceT, which lacks the TF N-terminal domain, is
dedicated to the folding and the transport of the reductive dehalogenase PceA through the
Twin arginine translocation system (Maillard et al, 2011).
149 245 432360
NC
Ribosome
binding
domainPPIase
domain Arm 1 Arm 2
A B
43 50
GFRxGxxP
Fig. 2: Structure of the E. coli TF and interaction with ribosome exit tunnel. (A) The upper
part represents the three-dimensional structure ribbon diagram of TF (PBD 1W26). The bottom
shows the domain arrangement of TF and the TF signature motif of the ribosome binding site (aa
43 to 50). (B) Interaction of TF with the exit-tunnel of the ribosome. TF binds L23 and L29 and
covers the exit tunnel from the crowded environment. The star represents the exit pore of the
tunnel. Ribosome binding site of TF is represented in red, the PPIase domain is in yellow, the
Arm 1 is in green and the Arm 2 in blue. Figure adapted from (Ferbitz et al, 2004).
17
TF is an abundant cytosolic protein represented in 2- to 3- fold molar excess over
ribosome (50µM in spite of 20µM respectively) (Crooke et al, 1988). It is constitutively
expressed and dispensable under normal growth conditions (Martinez-Hackert &
Hendrickson, 2009). TF cycles on and off the ribosome in 1:1 stoichiometry in an ATP-
independent manner and creates a protective arch over the ribosomal exit tunnel (Fig. 2B)
(Hoffmann et al, 2010; Kaiser et al, 2006; Rutkowska et al, 2008). During the TF
binding/release cycle, it has been found that the presence of a nascent chain increases the
affinity of TF for the ribosome 2 to 30 fold (Raine et al, 2006; Rutkowska et al, 2008). After
release from the ribosome, TF may stay bound to the nascent chain of large multi-domain
proteins allowing another free TF to bind the ribosome and in some cases to facilitate transfer
of its substrate to the downstream chaperones DnaKJE and GroESL (Kaiser et al, 2006;
Lakshmipathy et al, 2010). In addition, it has been found that TF from Thermotoga maritima
TF (tmTF) can dimerize and encapsulate 2 tmS7 ribosomal proteins (Martinez-Hackert &
Hendrickson, 2009). Such a dimerization of tmTF could create an Anfinsen-like cage which
could protect S7 and participate to its incorporation into the ribosome (Martinez-Hackert &
Hendrickson, 2009) (Fig. 3).
The N-terminal of TF interacts with the ribosome via its signature motif to the L23
ribosomal protein located at the exit port, the ribosomal RNA 23S and additional interaction
occurs with the L29 ribosomal protein (Baram et al, 2005; Kramer et al, 2002; Schlunzen et
al, 2005) (Fig. 2B). This TF-ribosome interaction is crucial for TF chaperone function during
co-translational folding of nascent polypeptide chains (Lakshmipathy et al, 2007). In vitro
experiments showed an early interaction with nascent polypeptide chains as short as 40 amino
acids (Merz et al, 2008). Nevertheless, using ribosome profiling of purified ribosome –
nascent chain – TF complexes and deep sequencing of mRNA fragments protected by these
ribosomes, Oh and coworkers (2011) showed that, in vivo, TF seems to preferentially bind to
nascent polypeptide chains of about 100 amino acids (Oh et al, 2011). This late interaction
between TF and the nascent chain potentially allows earlier interaction with other ribosome
associated nascent chain interacting factors, including the peptide deformylase, the
methionine aminopeptidase and SRP (Ball & Kaesberg, 1973; Bingel-Erlenmeyer et al, 2008;
Keenan et al, 2001).
Recent works identified nascent chains that are disliked by TF: a segment of poly-
alanine folded into a helical conformation (Bhushan et al, 2010; Lu & Deutsch, 2005). Indeed,
replacing 9 amino acids in the N-terminal of GFP by a poly-Alanine sequence near the exit
18
tunnel, the center of the tunnel or the peptidyl transferase center place this helical structure in
different parts of the ribosome tunnel. Using this technique, Lin and coworkers showed that
TF binding to the ribosome was disfavored when this helical structure was in an area near
from the exit tunnel and interacted with L23. This interaction can induce conformational
changes in the ribosomal binding site of TF and reduce its recruitment to the exit pore (Lin et
al, 2012). To validate this approach, they used the signal anchor sequence of a cytoplasmic
membrane protein and the signal sequence of the secretory protein pre-β-lactamase
(Bornemann et al, 2008). These sequences are known to fold into the ribosome tunnel in a
helical structure and to enhance the recruitment of SRP (Berndt et al, 2009; Bornemann et al,
2008; Halic et al, 2006; Woolhead et al, 2004). Both of them reduced the recruitment of TF at
the ribosome exit tunnel after interaction with L23 inside the ribosomal tunnel (Lin et al,
2012). Therefore, the recruitment of TF to the exit pore is modulated by the nascent peptide
chain itself, which modifies the ribosome binding site shared by TF and SRP (Lin et al, 2012).
Trigger Factor
50S
L23
50S
L23
Native proteinProtein
complex
Free dimer
of TF
Fig. 3: Chaperone cycle and assembly function of TF. TF interacts with nascent chains by direct
association with the ribosome. More than one TF molecule may bind a nascent polypeptide of a multi-
domain protein. This protein can be released in a native state or TF may stay bound to it in the cytosol to
fulfill its folding. Martinez-Hackert and Hendrickson (2009) also identified a new role of TF in the assembly
of protein complexes. TF could also form dimers and encapsulate two substrates mediating their folding and
their interaction. This role is important for ribosome biogenesis.
19
b- TF substrate interaction and cellular functions
Several studies were developed to identify preferred interactors of TF. A screening of
2.842 membrane coupled peptides of 20 different proteins originating from bacteria and
eukaryotes was used to identify TF binding motifs (Patzelt et al, 2001). The analysis revealed
that TF preferentially interacts with motifs of 8 residues enriched in aromatic residues (Phe,
Tyr, Trp and His) and basic residues (Arg and Lys) and impoverished in acidic residues (Asp
and Glu). Even if TF possesses a PPIase activity, proline residues do not seem to participate
in substrate binding. TF motifs occurs frequently, about every 32 residues in all protein tested.
This frequent motif allows TF to interact virtually with all the proteins during translation
(Patzelt et al, 2001). However, interactome studies pointed out some preferential substrates of
TF. A comparison between co-purified TF substrates and aggregated proteins isolated in a tig
null-mutant identified 178 potential TF substrates, a majority of them being ribosomal
proteins or proteins from homo- or hetero-oligomers (Martinez-Hackert & Hendrickson,
2009). Recently, a quantitative ribosome profiling analysis of ribosomes whose nascent chains
are bound to TF isolated outer-membrane β-barrel proteins (OMPs) as the strongest TF
interactors (Oh et al, 2011).
Even if TF interacts with a large number of substrates, the mode of action may differ.
TF can help peptide domains shorter than 150 residues to fold within its cradle (Hoffmann et
al, 2006; Merz et al, 2008). In vitro experiments also showed that TF promotes folding of
denatured proteins (Huang et al, 2000; Kramer et al, 2004a; Merz et al, 2006). In addition, TF
can delay co-translational folding of multi-domain proteins like firefly luciferase and β-
galactosidase as presented in the cycle of TF (Part II 1-a). This conclusion has been reached
by measuring the time-dependent enzymatic activity of these two proteins with or without TF
(Agashe et al, 2004). Two recent works have further studied such TF function. Indeed,
Hoffmann and colleagues (2012) showed that TF was found to postpone the disulfide bond
formation of ribosome arrested β-lactamase and barnase, to unfold a folded arrested barnase
and to facilitate its degradation. In addition, modeling the co-translational folding of the N-
terminal domain of β-galactosidase (216 residues), O’Brien and coworkers proposed that TF
could act on the nascent peptide via three different mechanisms: TF might (i) decrease the rate
of structural rearrangements, (ii) avoid tertiary structure formation, (iii) increase the effective
length of the exit tunnel and entanglements between the nascent polypeptide chains (O'Brien
et al, 2012). Such functions of TF could significantly help multi-domain proteins to fold
correctly, thus avoiding protein misfolding.
20
Moreover, interaction of the ribosome-bound TF with its substrate can facilitate
substrate degradation by proteases under certain conditions. Indeed, in vitro experiments
showed that, in the absence of DnaK, TF could favor the degradation of the arrested complex
barnase-arrested/ribosome (Hoffmann et al, 2012). In vivo works also revealed that TF favors
the degradation of EF-Tu in the absence of DnaK following Hsp33 over-expression (Bruel et
al, 2012). These results may pin point a new role of TF when DnaK is sequestered on
aggregates during a stress.
TF may participate in OMPs stability as well (Crooke & Wickner, 1987; Oh et al,
2011; Ullers et al, 2007). Indeed, deletion of the tig gene encoding TF provokes a decrease in
the steady state levels of OMPs compared with wild type cells (Oh et al, 2011) and some
OMPs were isolated as strong TF interactors by ribosome profiling analysis (Oh et al, 2011).
However, in vivo experiments revealed that TF deletion does not facilitate export of SecB-
dependent presecretory proteins, including MBP, OmpA and SecA (Guthrie & Wickner,
1990; Lee & Bernstein, 2002; Ullers et al, 2007). Instead, the deletion of tig even accelerated
the export of some presecretory proteins with Sec translocon (Lee & Bernstein, 2002).
Another example of TF involved in protein secretion was identified in Streptococcus
pyogenes(Lyon & Caparon, 2003). TF is essential for the secretion and the maturation of the
cysteine protease SpeB. The PPIase activity of TF modifies SpeB then TF targets this
substrate to the Sec translocon (Lyon & Caparon, 2003). Moreover, TF over-expression
induces the aggregation of OmpF, partially reflecting the toxicity of TF (Genevaux et al,
2004). Finally, over-expression of TF is toxic and induces a strong filamentation phenotype
similar to the filamentation observed after FtsZ depletion (Genevaux et al, 2004; Guthrie &
Wickner, 1990; Ward & Lutkenhaus, 1985). These findings suggest that TF could be involved
in cell division. In some strain background, TF is also known to be essential for viability at
low temperature (10°C), perhaps due to its possible involvement in ribosome biogenesis and
cell division (Kandror & Goldberg, 1997).
In conclusion, TF facilitates the folding of nascent chains by protecting them from the
crowded cellular environment and delaying formation of unproductive folding intermediates.
In addition, it is involved in ribosome biogenesis and in the stabilization of OMPs.
21
2- DnaKJE cycle and interactors
a- DnaKJE structure and cycle
The Hsp70 (Heat shock protein of 70kDa) molecular chaperone family is the most
conserved protein family in all organisms (Gupta, 1998). Thebest characterized Hsp70 family
member is the bacterial protein DnaK from E. coli. DnaK is a heat shock induced protein
present at about 10 000 copies per cell which double after a shift at 42°C (Genevaux et al,
2007). This molecular chaperone of 638 amino-acids in length possesses an N-terminal
nucleotide binding domain (NBD) of 45kDa with ATPase activity (aa 1 to 380) and a C-
terminal domain of 25kDa as substrate binding domain (SBD) subdivided into a β-sandwich
subdomain and a α-helical domain forming a lid (aa 398 to 638) (Flaherty et al, 1994). Both
domains are connected by a linker (aa 381 to 397) which is highly conserved and possesses a
characteristic 388DVLLLD393 hydrophobic segment essential for allosteric communication
between the NBD and the SBD (Mayer & Bukau, 2005; Swain et al, 2007; Vogel et al, 2006a;
Vogel et al, 2006b). In some eukaryotes the SBD contains an additional short motif
interacting with other partner proteins, like Hop/p60/CHIP that modulate chaperone activity
of Hsp70 (Mayer & Bukau, 2005; Young et al, 2004). The structure of a truncated DnaK (aa 1
to 605) bound to ADP and a substrate peptide (NRLLLTG) was revealed by NMR (Nuclear
Magnetic Resonance) presented in Fig. 4A (Bertelsen et al, 2009). This study showed that the
NBD and the SBD, in this conformation, are independent and mobile. This mobility is
restricted in a cone of ± 35° (Bertelsen et al, 2009). A recent work studied the structure of
DnaK bound to ATP. They showed that in this conformation, the α-helical lid and the β-
sandwich substrate pocket of the DnaK SBD were docked to different positions of the NBD
(Kityk et al, 2012). Then allosteric modifications induced by substrate binding and ATP
hydrolysis will unlock the SBD of DnaK (Kityk et al, 2012). DnaK facilitates protein folding
by cycles of binding/release of protein substrates. While ATP-bound DnaK has low affinity
and fast exchange rates for substrates, ADP-bound DnaK possesses a high affinity for the
substrate and a low exchange rate (Genevaux et al, 2007). The ATP hydrolysis by DnaK is
essential for its chaperone activity but this molecular chaperone has a low rate of ATP
hydrolysis (Karzai & McMacken, 1996; Laufen et al, 1999). Therefore, the DnaK chaperone
function relies on its obligate cochaperone partners, namely the J-domain protein Hsp40/DnaJ
and the nucleotide exchange factor GrpE (Schroder et al, 1993).
22
The DnaKJE cycle starts with the interaction of a substrate-bound DnaJ dimer with the
ATP-bound DnaK. Indeed, DnaJ recruits the substrate and efficiently stimulates ATP
hydrolysis by DnaK up to 1000-fold (Karzai & McMacken, 1996; Laufen et al, 1999; Liberek
et al, 1991). ATP hydrolysis will close the SBD and enhance its affinity for the substrate.
Then, a second cochaperone, namely the nucleotide exchange factor GrpE will bind to the
NBD and facilitate ADP release. The subsequent binding of a new ATP opens the lid of the
SBD and stimulates substrate release (Harrison et al, 1997). Once released from DnaK, the
substrate can reach its native form or necessitates further DnaK cycles (Fig. 4B) or be
transferred to other chaperones, like the chaperonin GroESL to complete its folding (see part
II-4).
DnaK interacts with a broad range of substrates present in different conformations:
folded, misfolded or aggregated. DnaK was found to be highly efficient when the substrates
are misfolded but not aggregated. Indeed, a firefly luciferase variant from Photinus pyralis
was unfolded by Freeze-thaw cycles and urea and incubated in the presence of the DnaKJE
machinery (Sharma et al, 2010). Monitoring folding kinetics, the authors found that while
ATP hydrolysis induces substrate unfolding, substrate release induces spontaneous refolding
of the substrate (Sharma et al, 2010). Another study revealed that the lid of DnaK could stay
opened or partially opened even after hydrolysis of ATP (Kityk et al, 2012; Schlecht et al,
2011). This conformation could allow DnaK to interact with aggregated proteins or amyloids
and participate to their dissolution.
23
NBD SBD
Linker
Lid
A
B
ATP
Pi
ADP
DnaK
DnaJ
Substrate
GrpE
ADP
ATP
ADP
+
++
Native
protein
ATP
Fig. 4: The DnaKJE chaperone system. (A) Three dimensional structure of the E. coli DnaK. (aa 1
to 605) truncated structure of DnaK bound to ADP (Bertelsen et al, 2009) (PDB 2KHO). The N-
terminal NBD (dark green) and the C-terminal SBD (green) are disjointed by a linker (gray). The Lid
of the SBD is in a closed conformation. (B)DnaKJE chaperone cycle. The ATP bound DnaK has a
low-affinity for substrate. The co-chaperone DnaJ mediates the recruitment of substrate to DnaK and
stimulates its ATPase activity. Hydrolysis of ATP, leading to the ADP bound DnaK, induces
allosteric modifications of DnaK allowing the sequestration of the substrate. Then, the Nucleotide
Exchange Factor GrpE facilitates ADP/ATP exchange opening the lid of DnaK and the substrate
release (Genevaux et al,2007).
24
b- Functions of DnaK
It is believed that DnaK interacts and assists the co- and/or post-translational folding
of about 15% of the nascent peptides or newly synthesized proteins (Deuerling et al, 1999;
Teter et al, 1999). DnaK also participates in the refolding of misfolded and aggregated
proteins, translocation through biological membrane and oligomeric complex
assembly/disassembly (Genevaux et al, 2007; Mayer & Bukau, 2005). DnaK acts on protein
aggregates cooperating with other molecular chaperones: ClpB (Hsp100), Hsp31, or the small
heat shock proteins (sHSPs) IbpA IbpB (Genevaux et al, 2007). DnaK is also involved in the
oxidative stress response and controls the heat shock response by stimulating the σ32
degradation (Hoffmann et al, 2004; Straus et al, 1990). The central role of DnaK is revealed
by deletion of its gene. Indeed, a dnaK null mutant in E. coli exhibits multiple phenotypes
including cryosensitivity beyond 20°C, thermosensitivity above 35°C, filamentation, slow
growth at the permissive temperature of 30°C, resistance to bacteriophages λ P1 and P2
infection, loss of motility, sensitivity to nutrient starvation and defective in plasmid
maintenance (Genevaux et al, 2007). DnaK deletion induces a ribosome biogenesis defect
indicating a role for DnaK in ribosome assembly (Al Refaii & Alix, 2009; Maki et al, 2002).
c- Interactors in the cell
Using a screening of 37 different proteins split in more than 4500 peptides and bound
to a cellulose membrane, the recognition motif of DnaK has been defined as an extended five
residue segment composed of hydrophobic amino acids, Leu, Ile, Val Phe and Tyr
preferentially, framed by positively charged residues (Rudiger et al, 1997). These motifs
occur often in proteins, every 50 to 100 amino acids (Rousseau et al, 2006). Such a variety of
cellular functions and high frequency of binding motifs suggest that DnaK interacts with a
large number of newly synthesized proteins. Indeed, a recent analysis of the DnaK
interactome in E. coli revealed that DnaK interacts with more than six hundred proteins under
physiological condition. 80% of these interactors are cytosolic proteins, but DnaK also
interacts with inner membrane proteins (~11%), outer membrane proteins (~3%) and
periplasmic proteins (~3%) (Calloni et al, 2012). The interaction with exported proteins is in
agreement with the role of DnaK in protein export (Randall & Hardy, 2002). Analysis of the
relative enrichment of substrates on DnaK pointed out that 40% of the total mass of DnaK
substrates was highly enriched on DnaK. These enriched proteins were identified to be below
average cellular abundance and with a low solubility property. These proteins are known to be
25
aggregation-prone (Tartaglia et al, 2010; Tartaglia et al, 2007). Interestingly, essential
proteins are under the low enriched protein group of DnaK interactors. This is probably due to
the fact that folding of these substrates is supported by the chaperonin GroESL.
3- The chaperonin GroESL
a- Structure of the GroESL complex and chaperonin cycle
The chaperonin GroESL is the only known chaperone system essential in E. coli(Fayet
et al, 1989). GroESL is an asymmetric complex composed by two heptameric rings of 57kDa
monomer of GroEL (Xu et al, 1997). A GroEL barrel is subdivided in three different domains:
the ATPase equatorial domain, the intermediate domain responsible of GroEL rearrangement
after substrate binding, and the apical domain presenting hydrophobic residues involved in
substrate binding and forming the interaction site with the GroES cochaperone (Bukau &
Horwich, 1998). GroES is structured in heptamer of 10kDa subunits and forms a dome
closing one ring of GroEL. Rings are termed as follow: the cis side is composed of a GroESL
complex encapsulating a substrate and the trans side is made by a free and open GroEL
cylinder (Fig. 5A) (Hartl & Hayer-Hartl, 2009).
At the beginning of the cycle, 7 ATP molecules bind to the trans-ring of GroEL. Then
the substrate is recruited to this GroEL ring. ATP molecules are known to induce
rearrangements of the apical domain of GroEL mediating the capture and the compaction of
the substrate to facilitate its encapsulation (Clare et al, 2012; Lin et al, 2008). Then a GroES
dome encloses the substrate into the cage and creates the new cis-ring of the complex. This
cis-ring interaction between GroES and the apical domains of GroEL will enhance the
hydrophobicity of the cavity, inducing the release of the substrate into the cage formed by the
GroEL cylinder. Then, rearrangements of the apical and the intermediate domains in the cis-
ring will bury the hydrophobic residues and change the environment of the cage. The volume
of the cage will also be enhanced and will be able to accommodate polypeptides up to 60kDa
(Hartl & Hayer-Hartl, 2009). The new substrate enclosed into GroESL complex now folds
into a secure and isolated space during 10s to 15s, time required for ATP hydrolysis. Then
release of ADP and GroES will free the substrate. The cycle will repeat from the new trans-
ring and the substrate will be free folded or will necessitate other cycles of folding into
GroESL (Fig. 5B). This GroESL dependent folding may occur in a passive “Anfinsen cage”
(Horwich et al, 2009; Motojima et al, 2012), with an active role via negatively charged
residues in the cavity (Chakraborty et al, 2010; Lin et al, 2008; Tang et al, 2008; Tang et al,
26
2006) or by forced unfolding (Lin et al, 2008). Action of GroESL mediated folding may differ
in relation to the substrate bound.
b- Interactors in the cell
In vivo, GroESL is known to interact with approximately 10% to 15% of the newly
synthesized cytosolic proteins (Houry et al, 1999). 250 GroESL substrates were identified by
mass spectrometry grouped into 3 different classes using their necessity to be helped by
chaperon systems. The Class I proteins have a relatively low propensity to aggregate and are
chaperone independent. Proteins of Class II need DnaKJE or GroESL to fold properly. Class
III proteins are obligate GroESL substrates These proteins range between 20kDa to 50kDa
and are enriched in (αβ)8 TIM barrel (Kerner et al, 2005). 85 of GroESL substrates are strictly
GroES
GroEL
cis-ring
GroEL
trans-ring
A
B
ADP
7 ATP
+
ATP
GroEL
ATP
GroES
ADP
7 ADP
7 Pi (10-15s)
ADP
7 ATP+
+
7 ADP
Fig. 5: The GroESL chaperonin system. (A) Three dimensional architecture of the GroESL-
7ADP complex (Xu et al, 1997) (PDB 1AON). The GroES heptamer (dark orange) encapsulates
the cavity of the cis-ring of GroEL (orange). The cis-ring of GroEL is in an extended
conformation in comparison to the trans-ring (gray). (B) GroESL chaperonin cycle. Details
concerning the cycle are given in the text.
27
GroESL-dependent for their folding from which 13 are essential proteins. These 85 substrates
occupy 75% to 80% of GroESL capacity in the cell (Kerner et al, 2005).
Recently, Class III proteins were over-expressed and tested for their dependence to
GroESL in GroESL-depleted cells (Fujiwara et al, 2010). They found that 60% of Class III
proteins are in fact GroESL-obligated substrates, the other 40% can use other chaperone
machineries for their folding under over-expressing conditions. These obligate GroESL-
substrates were grouped as Class IV. Note that these proteins possess a positive bias in
alanine and glycine content, making the proteins more aggregation-prone. Moreover, this
group contains six essential proteins for E. coli: DapA, ASD, MetK, FtsE, HemB and KdsA
(Fujiwara et al, 2010).
Interestingly, in the absence of both TF and DnaKJE, GroESL interacts with about 150
additional substrates (Kerner et al, 2005), thus reflecting interplay among these chaperone
systems.
4- Interplay between TF/DnaKJE/GroESL during de novo protein folding
The molecular chaperones TF, DnaKJE and GroESL are known to work as a network
participating together in the folding of nascent chains and newly synthesized proteins (Fig. 6).
Both TF and DnaKJE most likely share more than 300 proteins as common substrates
(Deuerling et al, 2003). Cooperation between these two chaperones had been shown in vitro
using the multi-domain protein firefly luciferase, with TF and DnaKJE acting co-and post-
translationally, respectively (Agashe et al, 2004). Analysis of the effect of TF, DnaKJE and
GroESL on about 800 cytosolic proteins was carried out using a reconstituted cell-free
translation system (Niwa et al, 2012). In this work, the authors showed that either DnaKJE or
GroESL efficiently increase the solubility of more than 66% of the proteins tested. In this
case, TF only had a minor effect on prevention of aggregation of these substrates. These
results strongly support previous in vitro translation experiment (Agashe et al, 2004) (Fig. 6).
To go further inside the chaperone network, Calloni et al (2012) recently characterized
interactors of DnaK following TF deletion or GroESL depletion. In the absence of TF, DnaK
was found to interact with a higher number of substrates: 998, including 95% of the 674
DnaK interactors already characterized in the presence of TF. This extended substrate
interaction of DnaK in the absence of TF correlates with the overlapping of function already
observed between these two molecular chaperones (Deuerling et al, 1999; Teter et al, 1999),
28
i.e.in the absence of TF, DnaK interacts with 2 to 3 times more nascent chains than in the
presence of TF (Teter et al, 1999). Note that in this case, DnaK could also interact with
shorter nascent polypeptides (<30kDa) (Deuerling et al, 1999; Teter et al, 1999).
As state above, E. coli tolerates the single deletion of either tig or dnaK(Deuerling et
al, 1999; Genevaux et al, 2004; Teter et al, 1999) and upregulation of DnaKJE is observed in
tig null-mutant (Deuerling et al, 2003). Simultaneous deletions of both tig and dnaK genes
present synthetic lethality at temperatures ≥30°C with respect to the strain background used
(Deuerling et al, 1999; Genevaux et al, 2004; Teter et al, 1999; Vorderwulbecke et al, 2004).
This double mutant possessed a narrow temperature range of growth from 20°C to 30°C,
presented a filamentous phenotype and accumulated aggregated proteins (Deuerling et al,
1999; Genevaux et al, 2004). When only 15 and 474 proteins aggregated in ∆tig and ∆dnaK
single mutants respectively, this amount of aggregated proteins reached 1087 in the absence
of both molecular chaperones (Calloni et al, 2012). A high number of ribosomal proteins were
also found into these aggregates reflecting the strong defect in ribosomal biogenesis of the
∆tig ∆dnaK double mutant (Calloni et al, 2012). This is in agreement with the role of both TF
and DnaKJE in ribosome biogenesis presented above.
GroESL is upregulated in the absence of DnaK and TF and correspondingly, GroESL
depletion induces DnaKJE system and modifies DnaK interactors (Calloni et al, 2012).
Indeed, in this case, 92 proteins had an increased interaction with DnaK; including 38
GroESL substrates of which 19 Class III obligated GroESL substrates. In contrast, 54 proteins
had a reduced interaction with DnaK, including 11 Class II GroESL substrates (Calloni et al,
2012). This suggests that DnaK could substitute its substrates with GroESL-obligated ones,
preventing their aggregation or mediating their degradation.
Remarkably, when both DnaK and TF are absent, GroESL interacts with 150
additional interactors (Kerner et al, 2005). Under these conditions, GroESL can substitute its
substrates for DnaKJE and TF ones. This partial switch of substrates is most likely
responsible of the partial aggregation of several Class III GroESL-obligatory substrates
(Kerner et al, 2005). Supporting such interactome data, GroESL over-expression suppresses
the growth defect of the ∆tig ∆dnaK double mutant (Genevaux et al, 2004; Vorderwulbecke et
al, 2004).
These results indicate that TF, DnaKJE and GroESL are the main players of the
intricate network of molecular chaperones (Fig. 6), in which DnaKJE appears to be a central
29
node, acting co- and/or post-transnationally downstream of TF and upstream of GroESL
(Calloni et al, 2012).
Part III:Interplays between TF, DnaKJE and GroESL and other
chaperones during protein export
TF
Native
~65-80%
GroESL
~10-15 %~10-20 %
DnaKJE
Fig. 6: Interplay between TF, DnaKJE and GroESL during de novo protein
folding: Nascent polypeptides interact with TF in a co-translational way. A
majority of them (65% to 80%) may reach their native structures without further
assistance. A subset of proteins may need the assistance of other molecular
chaperones to get their native conformation. The DnaKJE system interacts with
polypeptides both in a co- and a post-translational way, mediating the folding of
about 10% to 20% of cytosolic proteins. This chaperone system cooperates with
the chaperonin GroESL by substrate exchange. GroES mediates the folding of
approximately 10% to 15% of cytosolic proteins (Hoffmann et al, 2010).
30
The major TF, DnaKJE and GroESL chaperones participate in the export of proteins
by acting directly on substrates of the Sec and Tat (Twin-arginine translocation) general
secretion pathways (Graubner et al, 2007; Li et al, 2010; Perez-Rodriguez et al, 2007; Phillips
& Silhavy, 1990; Wild et al, 1992; Wild et al, 1996).
1- Sec Translocation system
The Sec translocon, through which substrates will cross the membrane, is composed of
a hetero-oligomeric complex mainly formed by SecYEG located in the inner membrane (Van
den Berg et al, 2004). SecA is a cytosolic protein promoting the translocation of the substrate
through the Sec translocon in an ATP-dependent manner (Schiebel et al, 1991). In E. coli this
system is supported by the molecular chaperone SecB. SecB facilitates export of OMPs and
some periplasmic proteins which have to be kept unfolded to be exported via the Sec-
translocon (Kumamoto & Beckwith, 1985). In agreement with such specific needs, the
tetrameric SecB chaperone possesses an anti-folding activity (Collier et al, 1988; Randall &
Hardy, 2002). It has been shown that TF, DnaKJE and GroESL play a role during protein
export as well (Altman et al, 1991; Calloni et al, 2012; Lecker et al, 1989; Oh et al, 2011;
Phillips & Silhavy, 1990; Qi et al, 2002; Ullers et al, 2007; Wild et al, 1992; Wild et al, 1996)
(Fig. 7). In addition, functional overlaps and complementarities had been observed between
SecB and theses chaperones during protein export. Indeed, both DnaKJE and GroESL
facilitate the export of several SecB-dependent substrates and their over-expression efficiently
rescue the cold-sensitive phenotype of a secB null mutant (Altman et al, 1991; Phillips &
Silhavy, 1990; Qi et al, 2002; Ullers et al, 2007; Wild et al, 1992; Wild et al, 1996).
Furthermore, the export of several OMPs following SecA depletion is abolished when DnaK
functions are altered (Qi et al, 2002), suggesting that DnaK is able to maintain Sec-dependent
OMPs in a translocation-competent state (Qi et al, 2002). In support of such overlaps, both
secB and dnaK mutations are synthetic lethal and a direct interaction between SecB and DnaK
has been recently identified in vivo(Calloni et al, 2012). Remarkably, SecB over-expression
also suppresses the growth defect and the severe protein aggregation observed in the combine
absence of both TF and DnaK (Ullers et al, 2004). This suppression was independent of SecB
export function as judged by the used of the variant E77K impaired in SecA interaction
(Fekkes et al, 1998; Kimsey et al, 1995).This indicates that in addition to its role in protein
export, SecB can perform generic chaperone functions and facilitate cytosolic protein folding
in the absence of TF and DnaK. Finally, such interplays are further supported by the fact that
31
SecB endogenous levels are significantly increased upon depletion of DnaKJE and GroESL,
and reciprocally (Muller, 1996).
Although TF binds some OMPs and most likely facilitates their translocation (see Part
II), it has been shown that TF can also antagonize the action of both SecB and DnaKJE during
protein export, as judged by its ability to severely delay export of some Sec-dependent
proteinswhen SecB or DnaKJE are absent (Lee & Bernstein, 2002; Ullers et al, 2007). A
model presenting the different chaperone pathways involved in the Sec-dependent protein
secretion is show in Fig. 7.
2- Twin Arginin translocation system
The second general secretion pathway in E. coli is the Tat system. The translocon is
composed by three membrane proteins of which encoding genes are organized in an operon:
genes encoding for TatA, TatB and TatC; another one, isolated, encodes for TatE and is a
functional TatA duplication (Sargent et al, 1998). This translocon translocates proteins having
the consensus N-terminal motif (S/T)-R-R-X-F-L-K previously folded into the cytoplasm
(Berks, 1996). Majority of Tat substrates have their own specific chaperones called REMPs
(Redox Enzyme Maturation Proteins) like DmsD, NarJ, NarD or TorD (Turner et al, 2004).
They interact specifically with the twin-arginine leader peptide protecting the substrate from
degradation, maintaining it in a translocation competent state and targeting it to the Tat
translocon (Sargent, 2007; Turner et al, 2004). REMPs are not acting alone and participate in
a chaperone cascade also orchestrated by TF, DnaKJE and GroESL (Perez-Rodriguez et al,
2007). Interactions between DmsD and these molecular chaperones were pointed out by
several in vitro experiments (Li et al, 2010). TF, DnaKJE and GroESL were shown to interact
directly with Tat substrates and to efficiently participate in their translocation (Fig. 7). TF was
shown to interact with signal peptides of TorA and SufI until late translation but did not
participate actively in their translocation (Jong et al, 2004). DnaK is essential for the
translocation of over-expressed CueO, and efficiently binds to the leader peptide of DmsA
(Graubner et al, 2007; Oresnik et al, 2001). Moreover, several Tat substrates appear to be in
an unstable conformation in the absence of DnaK (Perez-Rodriguez et al, 2007). These data
suggest that DnaK may work in cooperation with REMPs to stabilize and protect substrates
from degradation. The GroESL interactome revealed the presence of the Tat-dependent
amidase AmiA as a GroESL substrate, which required GroESL for its folding (Kerner et al,
2005; Rodrigue et al, 1996).
32
SecB
TatABC
Folded
REMP
+
Inner membrane
+
SecA
SecYEG
Cytoplasm
Periplasm
DnaKJEGroESL
TF
Fig. 7: Chaperone networks involved in protein export. Molecular chaperones play different roles in both
systems. In Sec translocation TF, DnaKJE, GroESL, and SecB generally favor secretion by keeping substrates
in an unfolded competent state for translocation. The N-terminal Signal sequence of the substrate is represented
by a green rectangle. In Tat system, the three major chaperones work with REMPs to fold the substrate and
target it to the Tat translocon. The N-terminal Signal sequence of the substrate is represented by a cyan
rectangle.
33
Part IV: Chaperone-mediated response to protein misfolding and
aggregation
The TF, DnaKJE and GroESL chaperone network is not sufficient to rescue protein
aggregation during heat-stress or oxidative protein unfolding. Other heat shock proteins
(Hsps) are induced in these conditions to facilitate the action of DnaKJE and most likely
GroESL. These chaperones include the Clp superfamily proteins (Hsp100), the small Hsps
(sHsps) IbpA and IbpB, HtpG (the Hsp90 homolog) and the redox-regulated chaperone Hsp33
(Hartl, 1996; Hoffmann et al, 2004; Squires & Squires, 1992).
1- Molecular chaperone networks and protein disaggregation
a- Presentation of other chaperones involved in protein disaggregation
Members of the Clp family include protein complexes functioning as disaggregases
and/or proteolytic machines with AAA+ ATPases activities (ATPases Associated with a
variety of cellular Activities). They can be gathered into two groups depending on the number
of Nucleotide Binding Domains (NBD) they possess. Class I AAA+ proteins comprising
ClpA, ClpB, ClpC and ClpE monomers have two different NBDs separated by a coiled coil
middle domain; Class II proteins with ClpX and ClpY monomers have only one NBD
(Schirmer et al, 1996). Some AAA+ proteins possess a protease domain like FtsH and Lon.
ClpB (Hsp100) possesses a disaggregase activity forming a barrel-shaped hexamer with an
axial channel when it is bound to ATP (Akoev et al, 2004). This channel is surrounded by the
two NBDs of ClpB (NBD1 and NBD2), the middle domain (M domain) of the monomer
being outside of the cylinder (del Castillo et al, 2011; Lee et al, 2003). The N-terminal domain
of each monomer forms a crown involved in the interaction with aggregated proteins (Barnett
et al, 2005). Then ATP hydrolysis will mediate disaggregation by ClpB via translocation of
the substrate through the channel (Weibezahn et al, 2004). The M domain, which is structured
as a coiled coil with two wings, is essential for the disaggregase activity of ClpB (Kedzierska
et al, 2003; Lee et al, 2003; Mogk et al, 2003b). It is involved in the stabilization of the
hexameric structure of ClpB (del Castillo et al, 2011). Seyffer and co-workers (2012) showed
that DnaK could bind the M domain and induce allosteric modifications in ClpB, which,
together with substrate interaction, stimulate ClpB ATPase activity (Seyffer et al, 2012). The
E. coliclpB gene possesses an internal translation initiation site expressing two isoforms of
ClpB in vivo: the full length 95kDa ClpB and a truncated isoform of 80kDa without the N-
terminal domain (Squires et al, 1991). It has been shown that these two isoforms cooperate to
34
favor solubilization of aggregated proteins and that each isoform alone is less effective in this
task (Guenther et al, 2012; Zhang et al, 2012). The N-terminal domain is involved in the
interaction with aggregates; this combination of isoforms could give more flexibility at this
domain and then facilitate the interaction with substrates (Nagy et al, 2010). The deletion of
clpB in E. coli is known to present a growth defect at 44°C and is also more sensitive to heat
shock at 50°C than the wild type cell (Squires et al, 1991).
The E. coli IbpA and IbpB proteins (Inclusion body protein) are conserved small heat-
shock proteins of 16 kDa that share about 50% amino acid sequence identity and that are
known to associate with thermally aggregated proteins and inclusion bodies (Allen et al,
1992; Laskowska et al, 1996). These two sHsps are known to form molecular oligomeric
structures (Kitagawa et al, 2002; Shearstone & Baneyx, 1999) and cooperate to disaggregate
aggregated polypeptides. The ibpAB operon can be deleted with no effect on cell during
normal growth conditions but causes a decreased viability during prolonged growth at 50°C
(Kuczynska-Wisnik et al, 2002) and as a sensitivity phenotype to copper-induced stress under
aerobic conditions (Matuszewska et al, 2008).
A third chaperone known to participate in disaggregation is the E. coli Hsp90
chaperone member named HtpG. It is an ATP-dependent chaperone functioning as a
homodimer. HtpG monomer is made up of three domains: an N-terminal domain with ATPase
activity, a middle domain and a C-terminal domain involved in Hsp90 dimerization
(Krukenberg et al, 2011; Mayer, 2010; Pearl & Prodromou, 2006; Wandinger et al, 2008).
HtpG is essential for the activity of the clustered regularly interspaced short palindromic
repeats (CRISPR) system(Yosef et al, 2011) involved in the detection and the degradation of
exogenous DNA (Barrangou et al, 2007; Brouns et al, 2008; Marraffini & Sontheimer, 2008)
and RNA (Hale et al, 2009) in prokaryotes. As for clpB, htpG deletion only shows a growth
defect above 44°C (Bardwell & Craig, 1988).
b- Chaperone networks involved in protein disaggregation
In vitro experiments showed that efficient interaction of ClpB with aggregates was
DnaKJE-dependentand thatClpB and DnaKJE systems are strong partners for protein
disaggregation, acting as a bi-chaperone system by which the substrate is transferred from
DnaKJE to ClpB (Acebron et al, 2009; Goloubinoff et al, 1999; Motohashi et al, 1999;
Zolkiewski, 1999). This substrate transfer is mediated by a direct interaction between DnaK
and the M-domain of ClpB (Seyffer et al, 2012). In addition it has been shown that DnaJ
35
efficiently bindsprotein aggregates independently of DnaK and recruits DnaK to the
aggregates with or without ClpB (Acebron et al, 2008; Winkler et al, 2012). In contrast, ClpB
cannot efficiently bind aggregates in a dnaKmutant (Spence et al, 1990), indicating that the
DnaKJE system first interacts with aggregates and presents the substrates to ClpB. Deletion of
clpB in a dnaK mutant straininduces a strong deleterious effect on E. coli growth when
compared to their respective single mutant controls. Similar results were obtained in a
groES30 mutant allele showing their importance during heat shock (Thomas & Baneyx,
1998). An in vivoFRET (Fluorescence resonance energy transfer) experiment confirmed the
chaperone interaction on aggregates. Indeed, they showed that the DnaJ interaction with
aggregates was followed by recruitment of DnaK. Then DnaK transferred substrates to
ClpB(Kumar & Sourjik, 2012).
The sHsps proteins are strongly recruited to aggregates: 50% of IbpA and IbpB
proteins were bound to aggregates after heat shock at 45°C. This percentage could even reach
80% in ΔclpB and ΔdnaK mutant cells (Mogk et al, 1999), thus revealing the importance of
the chaperone network in the dissolution of aggregates and the reactivation of proteins. How
do IbpA and IbpB favor the dissolution of aggregates mediated by the bi-chaperone
DnaKJE/ClpB? Ratajczak and co-workers (2008) showed that IbpA alone could bind heat-
induced aggregates and reduce their size. However, it could not accelerate the disaggregase
effect of DnaKJE and ClpB. In this case, solitary IbpA even inhibited the downstream action
of the DnaKJE/ClpB bi-chaperone on aggregates, suggesting that it may block the
translocation through the central pore of ClpB. However, IbpB binds to the aggregates only
after IbpA and both IbpA and IbpB favor the action of DnaKJE/ClpB (Kuczynska-Wisnik et
al, 2002). IbpB can thus modify the interaction between IbpA and the aggregates and allow
the action of the bi-chaperone DnaKJE/ClpB (Ratajczak et al, 2009). This cooperation
between the sHsps, ClpB and DnaKJE was additionally tested in vivo using combination of
mutations in these molecular chaperones. In this case the solubilization of protein aggregates
was delayed but not abolished in a ΔibpAB strain when compared with wild type, indicating
IbpA and IbpB are not essential to the solubilization of aggregates but may facilitate the
action of ClpB and DnaKJE (Mogk et al, 2003a). Yet, as observed for clpB mutations (see
above), the combined deletion of ibpABand dnaK exhibitsa strong growth defect when
compared to the respective single mutants(Thomas & Baneyx, 1998). Furthermore, Kumar
and Sourjik (2012) showed that, during severe denaturation conditions, when the DnaKJE
chaperone system is strongly titrated to aggregated or unfolded proteins, both IbpA and IbpB
36
localize with aggregated proteins. ClpB is then recruited to aggregates but DnaKJE is needed
for efficient disaggregation (Fig. 8).
As for ClpB and IbpA/B, HtpG cannot reactivate denatured luciferase in vitro alone.
But its presence significantly stimulates refolding by DnaKJE, up to 1,6-fold when compared
to DnaKJE alone (Genest et al, 2011). This mode of action takes place in a sequential manner,
by first binding of DnaK and the subsequent recruitment of HtpG to the substrate (Genest et
al, 2011). Although the effect of HtpG on aggregates in not well defined yet,the deletion of
htpGeither in a dnaK or a groES30 mutant further affects bacterial growth, thusfurther
arguing for a role for HtpG in the disaggregation pathway (Thomas & Baneyx, 1998).
Together these data reveal the importance of the cooperation between ClpB, IbpA
IbpB and HtpG with the DnaKJE chaperone systems in protein disaggregation (Thomas &
Baneyx, 1998).
IbpAB
oligomersAggregates
ClpB
Native
Proteases
+
DnaKJEGroESL
Unfolded protein
DnaKJ
Fig. 8: Mainmolecular chaperones functions in disaggregation. IbpA (in green) and IbpB (in violet)
oligomers break aggregates into smaller structures. DnaKJ starts to free proteins and cooperate with
the disaggregase ClpB to solubilize them. Then, disaggregated proteins may reach their native
structure with or without the action of DnaKJE and GroESL. If their folding is not possible, because of
a titration of molecular chaperones or an irreversible modification, they will be degraded by proteases.
37
2- Hsp33 as a member of the chaperone network
As stated above, the simultaneous deletion of both dnaK and tig leads to a severe
growth defect and protein aggregation (Deuerling et al, 1999; Genevaux et al, 2004; Teter et
al, 1999; Vorderwulbecke et al, 2004). This strain gives an advantage to study chaperone
networks and to further elucidate interactions between molecular chaperones in vivo. Both
SecB and GroESL have been previously identified as suppressors of these phenotypes
(Genevaux et al, 2004; Ullers et al, 2004; Vorderwulbecke et al, 2004). In this thesis work,
another protein suppressing these sensitive phenotypes has been identified: the molecular
chaperone Hsp33.
a- Hsp33 is involved in the oxidative stress response
Oxidative stress is concomitant with life in aerobic conditions, favoring production of
reactive oxygen species O2- or H2O2. This stress can be induced under culture conditions by
adding H2O2 or HOCl in the medium or after heat shock (Imlay, 2003; Storz & Imlay, 1999;
Winter et al, 2008). Oxidative stress is responsible of the oxidation of all macromolecules in
the cell: DNA, lipids and proteins (Imlay, 2003). This stress even paralyzes the chaperone
network functions inactivating the DnaK molecular chaperone system (Winter et al, 2005).
Proteins oxidize mostly on sulfur group of cysteines and methionines, increasing their
susceptibility to aggregation and degradation (Storz & Imlay, 1999). Some proteins involved
in the oxidative stress response become active through disulfide bound formation (Aslund &
Beckwith, 1999). This is the case of the transcriptional regulators OxyR and SoxR inducing
the oxidative stress response (Ding & Demple, 1998; Zheng et al, 1998). OxyR upregulates 28
genes involved in the protection against oxidative stress such as glutathione oxidoreductase,
glutaredoxin I, Catalase/hydroperoxidase I, Thioredoxin reductase and Thioredoxin 2 (Chiang
& Schellhorn, 2012). SoxR oxidative stress response is involved in superoxide radical
response. SoxR activated by an oxidative stress induces the expression of the transcriptional
regulator SoxS known to regulate the expression of more than 100 genes including the
manganese-superoxide dismutase sodA, the endonuclease IV involved in DNA repair and
yggX protecting iron-sulfur proteins against oxidative stress (Blanchard et al, 2007). A
molecular chaperone was also found to be activated by oxidative stress and participate to this
stress response, the heat shock induced Hsp33 (Jakob et al, 1999).
38
b- Structure of Hsp33
Hsp33 is a very well conserved protein possessing homologues in many prokaryotes
species and also in some eukaryotic unicellular organisms like Trypanosomatidae(Jakob et al,
1999). Synthesis and activity of this protein is regulated by a two level mechanism: the heat
shock response for the transcriptional regulation and an oxidative stress response for its
activation via disulfide bound formation. This modification is reversible (Jakob et al, 1999).
Hsp33 is a 32,9kDa protein structured in three different domains (Fig. 9). The N-
terminal domain of Hsp33 (aa 1-178) is folded as a core domain, very compact, with two anti-
parallel β-sheets of five and four β-strands (Graf & Jakob, 2002). This domain presents some
structural homology with the TF ribosome binding domain except the absence of the loop
region responsible for the ribosome binding (Kristensen & Gajhede, 2003). The linker region
(aa 179-232) is folded in the reduced form of Hsp33 and interacts with the N-terminal domain
(Janda et al, 2004; Vijayalakshmi et al, 2001). The C-terminal domain (aa 232-294) contains
four cysteines absolutely conserved in all Hsp33 homologues (organized as follow: C232
-X-
C234
-(27-32)-C265
-X-Y-C268
) stabilizing a zinc ion in a tetrahedral conformation. This domain
is named the zinc center (Graf et al, 2004; Jakob et al, 2000). This domain is known to be the
oxidative sensor domain of Hsp33 (Jakob et al, 1999).
39
c- Activation cycle of Hsp33
Hsp33 is an ATP-independent molecular chaperone acting as a holdase to prevent
aggregation of oxidized and denatured proteins rather than efficiently participate in their
folding (Jakob et al, 1999). The redox potential of a cell under normal growth conditions is
ranged between -250mV and -280mV keeping Hsp33 inactive (redox potential of -170mV ±
10mV) (Jakob et al, 1999). When an oxidative stress occurs, the cellular redox potential is
about to reach >-150mV which allows the activation of Hsp33 (Gilbert, 1990).
Zn2+
178
N
232 294
C
N-terminal
domain
Linker
zinc center
A
B
Fig. 9: Hsp33 structure. (A)
Three dimensional structure of
reduced Hsp33 ribbon diagram
with a Zn2+
ion bound to the zinc
center (PDB 1VZY). (B) Domain
arrangement of Hsp33 showing
the 4 strictly conserved cysteins
in the zinc center. The N-
terminal is shown in blue, the
linker region in green and the
Zinc center in orange (Kumsta &
Jakob, 2009).
40
The activation of Hsp33 as a molecular chaperone is a two-step process. At the
beginning, reduced Hsp33 is in a monomeric state and the zinc center is folded. When an
oxidative stress occurs, disulfide bound formation is formed between Cys265
and Cys268
releasing the zinc ion and unfolding the zinc center domain (Ilbert et al, 2007; Won et al,
2004). Hsp33 will stay in this oxidized non-active conformation until a denaturing stress
happens. Heat stress favors the unfolding of the linker region and facilitates the disulfide
bound formation between Cys232
and Cys234
unfolding the linker region (Leichert et al, 2008).
Then, activated Hsp33 interacts with its substrates, between the linker region and the N-
terminal domain (Reichmann et al, 2012), as a monomer (Barbirz et al, 2000), a dimer
(Vijayalakshmi et al, 2001) or an oligomeric structures (Akhtar et al, 2004) depending the
oxidized Hsp33 concentration and the temperature (Akhtar et al, 2004; Graf & Jakob, 2002).
The dimerization of activated Hsp33 is favored by high temperature (Akhtar et al, 2004;
Graumann et al, 2001). An in vitro experiment revealed that a small population of Hsp33
oligomers (until octamer) may occur with higher temperature and in late time of oxidative
stress (Akhtar et al, 2004). These authors showed that oligomers of Hsp33 were more efficient
than the dimeric conformation for the protection of thermally unfolded citrate synthase.
Presence of this oligomeric structure is not yet proved in vivo. But during an oxidative stress,
Hsp33 is present in high concentration, which could favor apparition of this structure.
The reversible inactivation of Hsp33 necessitates at the same time reducing and
refolding conditions (Ilbert et al, 2006). Inactivation of the Hsp33 dimer-substrate complex by
glutaredoxin and thioredoxin doesn’t release the substrate into the cytosol. Release of the
substrate from the reduced-Hsp33 requires a functional DnaKJE system (Hoffmann et al,
2004). This substrate exchange mediates the folding of denatured proteins when stress
conditions are over and when sufficient amount of DnaKJE system is reactivated (The Hsp33
chaperone cycle as a dimer is presented in Fig. 10). Oxidative stress is known to drop the ATP
concentration in the cell (Osorio et al, 2003). This ATP depletion combined to oxidative and
heat stresses inactivate DnaK by unfolding of its NBD. This inactivation is reversible when
reducing conditions are restored (Winter et al, 2005). Hsp33 protect unfolded proteins against
aggregation until DnaK’s activity is restored at reduced conditions.
41
Oxidized Hsp33 molecular chaperone activity has been tested in vitro on several
substrates. It can prevent from aggregation denatured citrate synthase, luciferase or the E. coli
protein FtsJ and the eye-lens δ-crystallin (Jakob et al, 1999; Raman et al, 2001). In vivo,
Hsp33 had been found to protect about 80% of aggregated-prone proteins aggressed by
oxidative stress, including metabolic pathways enzymes, fatty acid biosynthesis and proteins
involved in transcription and cell division. Membrane proteins were not protected by Hsp33
(Winter et al, 2005). More recently, binding specificity of Hsp33 was studied by Reichmann
and coworkers (2012). They found that Hsp33 does not seem to have binding sequence
specificity but prefers interact with secondary structured regions, mostly alpha helices. This
secondary structure interaction preference of Hsp33 may avoid the unproductive interaction
between activated Hsp33 and its unfolded C-terminal domain. Although, there is some
overlapping of substrates between DnaKJE and Hsp33; but DnaKJE only interacts with
unfolded proteins whereas Hsp33 binds structured regions (Jakob et al, 1999; Reichmann et
al, 2012). The reduction of Hsp33 induces conformational changes in the molecular
Heat
stress
Hsp33 Dimerization and
interaction with the substrate
Zn2+SS
SS
Reduced
Hsp33
Reducing and physiological
temperature conditionsSS
SS
SS
SS
H2O2 or
HOCL Zn2+
Oxidative
stress
DnaKJE
Native
Activated
Hsp33
Zn2+
Fig. 10: Hsp33 cycle. An oxidative stress is sensed by the Zinc center of Hsp33 via disulfide
bound formation between Cys 265 and 268 removing the Zn2+
ion from the zinc center. Then,
Hsp33 is fully activated by an heat shock inducing the disulfide bound formation of Cys 232 and
234 which unfolds the linker region and the zinc center. This activation mediates Hsp33
dimerization and interaction with unfolded substrates with secondary structures. When reducing
and physiological temperature conditions are recovered, all cysteines of Hsp33 are reduced;
binding of the Zn2+
ion refolds the linker region and the zinc center. This conformational
modification of Hsp33 dimer mediates the unfolding of its secondary structured substrate
allowing its transfer to the DnaKJE chaperone system. Then Hsp33 returns to its monomeric
conformation. Colors are identical to Fig. 9.
42
chaperone refolding the zinc center. These conformational modifications trigger unfolding of
its substrate, loss of stability of the Hsp33-substrate complex and finally allow interaction
with DnaK (Reichmann et al, 2012) or with proteases as Lon (Bruel et al, 2012).
d- Hsp33 deletion and functions in the cell
The Hsp33 encoding gene hslO is located in an operon (Graf & Jakob, 2002). Two
other genes are co-translated, hslR and hslP. hslR encodes for the Hsp15 protein, known to
bind and recycle the complex formed by the 50S ribosomal subunit, the nascent chain and the
tRNA after aborted protein synthesis and hslP encodes for a putative phosphatase (Korber et
al, 2000). hslO can be deleted in E. coli and confers an oxidative sensitivity; this effect is
exacerbated in a cell which is always under oxidative stress after deletion of trxB encoding the
thioredoxin. This in vivo result pin points a key role of Hsp33 during oxidative stress (Winter
et al, 2005). Interestingly, hslO deletion in Vibrio cholerae conduces to thermosensitivity
phenotype and a higher sensitivity to oxidative stress. An in vivo experiment showed that
these phenotypes were suppressed by the E. coli EF-Tu over-expression (Wholey & Jakob,
2012). EF-Tu is known to be subject to thiol modification during oxidative stress and partial
depletion of EF-Tu increases the bleach sensitivity of E. coli(Leichert et al, 2008). In addition,
Hsp33 deletion favored the degradation of the V. Cholerae EF-Tu because of a higher
sensitivity to oxidative stress than E. coli EF-Tu (Wholey & Jakob, 2012).
In this thesis work, we showed that Hsp33 efficiently suppressed the bacterial growth
and protein folding defects observed in the absence of both TF and DnaK, thus revealing a
role for Hsp33 as a major participant of the E. coli chaperone network. However, in contrast
with previous results concerning GroESL and SecB, we now showed that Hsp33 acts
indirectly, by specifically interacting with the essential elongation factor Tu (EF-Tu) and
targeting it for degradation by the protease Lon.
43
MATERIALS
AND
METHODS
MATERIALS AND METHODS
1- Strains and plasmids
a- Bacterial Strains, phages, and culture conditions
Genetic experiments were carried out in the E. coli MC4100 (Casadaban, 1976) or W3110
genetic background strains (Bachmann, 1972). The MC4100 mutant derivatives Δtig::CmR,
ΔdnaKdnaJ::KanR; Δtig::Cm
R ΔdnaKdnaJ::Kan
R, dnaJ::Tn10-42, Δtig dnaJ::Tn10-42 (Genevaux
et al, 2004), Δtig::CmS (Ullers et al, 2007), Δlon::Kan
R(Sakr et al, 2010), ΔclpP::Cm
R (Maurizi et
al, 1990), ΔclpQ::TetR(Yamaguchi et al, 2003) and W3110 Δtig ΔdnaKdnaJ::Kan
R(Genevaux et al,
2004)have been previously described. The ΔhslO::KanR allele was obtained from strain JWK5692
(Keio collection). All mutations described in this study were moved to the appropriate genetic
background by bacteriophage P1-mediated transduction. Bacteria were routinely grown in LB
medium supplemented when necessary with either chloramphenicol (10 μg/ml), kanamycin (50
μg/ml), ampicillin (100 μg/ml), or tetracycline (15 μg/ml).
The construction of the dnaK-protease and dnaK hslO double mutants was performed as
follows. The KanR cassettes from MC4100 ΔhslO::Kan
R and Δlon::Kan
R were first removed using
plasmid pCP20 as described (Datsenko & Wanner, 2000). The ΔdnaKdnaJ::KanR
mutant allele was
then introduced into the hslO and the various protease mutants, thus leading to the construction of
strains MC4100 ΔhslO ΔdnaKdnaJ::KanR, Δlon ΔdnaKdnaJ::Kan
R, ΔclpP::Cm
R
ΔdnaKdnaJ::KanR, and ΔclpQ::Tet
R ΔdnaKdnaJ::Kan
R. To construct strain MC4100 ΔhslO
Δtig::CmRdnaJ::Tn10-42, the Δtig::Cm
R mutant allele was first introduced intoMC4100 ΔhslO,
followed by introduction of dnaJ::Tn10-42(TetR). In this case, selection for transductants was
carried out at 30°C.
The mutant strains used for the Lon-dependent suppression experiments were constructed as
follows. The ΔhslO::KanR mutant allele was first introduced into MC4100 Δlon. Next, the
ΔdnaK52::CmR mutant allele (Bukau & Walker, 1989) was introduced into both MC4100
ΔhslO::KanR and MC4100 Δlon ΔhslO::Kan
R. MC4100 Δtig::Cm
R Δlon::Kan
RdnaJ::Tn10-42 was
constructed by first moving the Δtig::CmR mutant allele into MC4100 Δlon::Kan
R. Note that the lon
and tig genes are 90% linked by P1 transduction on the E. coli chromosome. The dnaJ::Tn10-42
allele was then introduced into MC4100 Δtig::CmR Δlon::Kan
R and selection was done at 30°C. All
DNA cloning and transformations experiments were carried out in strain DH5α (Invitrogen).
45
b- Plasmid construction
Plasmids pSE380ΔNcoI, p29SEN (Genevaux et al, 2004) and pBAD22 (Guzman et al,
1995) have been previously described. To construct the high-copy number plasmid pSE-Hsp33
(pSE380 ΔNcoI-Hsp33) and the low-copy number plasmid p29-Hsp33 (p29SEN-Hsp33), the 879
bp hslO gene was PCR-amplified using primers Hsp33-for (5'-
CGGAATTCTCATGATTATGCCGCAACATGACC-3’) and Hsp33-rev (5’-
CGAAGCTTGGATCCTTAATGAACTTGCGGATCTGC-3’) using MG1655 genomic DNA as
template. The PCR fragment was digested with EcoRI and HindIII and cloned into either
pSE380ΔNcoI or p29SEN previously digested with the same enzymes. A similar cloning procedure
was used to construct the pSE-Hsp33(Y12E), pSE-Hsp33(M172S) mutant derivatives, except that
in this case, plasmids pET11a-hslO(Y12E) or pET11a-hslO(M172S) DNA was used as a template
(Cremers et al, 2010). Plasmid pSE-Hsp33-FLAG (pSE380ΔNcoI-Hsp33-flag tagged) containing
Hsp33 with the C-terminal FLAG tag “GSDYKDDDDKSA” was constructed using primers Hsp33-
for and 33FT-rev (5’-
GCAAGCTTGGATCCTTAGGCGCTTTTATCGTCGTCATCTTTGTAGTCGCTGCCATGAAC
TTGCGGATCTGC -3’) and pSE-Hsp33 as DNA template. The PCR fragment was cloned as an
EcoRI-HindIII digested PCR fragment in pSE380ΔNcoI digested with the same enzymes. Plasmid
pSE-Hsp33(Q151E) was constructed by quick change mutagenesis using the appropriate primers.
Plasmid pSE-Hsp33(1-235) was constructed using primers Hsp33-for and Hsp33(1-235)-rev (5’-
CAAGCTTGGATCCttaCGAGCAGGTGCATTTGAACTC -3’) and pSE-Hsp33 as DNA template.
The PCR fragment was cloned as an EcoRI-HindIII digested fragment into pSE380ΔNcoI digested
with the same enzymes.
The p29SEN-derivatives containing either the yrfG hslR hslO or the hslR hslO operon were
constructed using yrfG-For (5’- GAGAATTC CATATGCATATCAACATTGCCTG -3’) or hslR-
For (5’-GAGAATTC CATATGAAAGAGAAACCTGCTGTTG -3’) forward primers, respectively.
In this case, the primer Hsp33-rev was used as reverse primer.
To construct plasmid p29-EF-Tu, the 1185bp long tufA gene, encoding EF-Tu, was PCR-
amplified using primers tufA-for (5’- GAGAATTCATGTCTAAAGAAAAATTTGAAC-3’) and
tufA-rev (5’- GAAAGCTTTTAGCCCAGAACTTTAGCAAC-3’) and MG1655 genomic DNA as
template. The PCR fragment was digested with EcoRI and HindIII and cloned into p29SEN
digested with the same enzymes.
The toxin gene hipA (1323bp) was PCR amplified using primers HipA-For (5’-
GCGAATCCATGCCTAAACTTGTCACTTG-3’) and HipA-Rev (5’-
46
GCGCATGCTCACTTACTACCGTATTCTC-3’) and MG1655 genomic DNA as template. The
hipA gene was then cloned under the control of an arabinose-inducible promoter as an EcoRI-SphI
fragment into plasmid pBAD22 digested with the same enzymes. The resulting plasmid was named
pBAD-HipA. Note that hipA was cloned under the tightly regulated araBADp promoter to minimize
background expression level of HipA in the absence of inducer, thus avoiding killing by the toxin.
All constructs generated by PCR were sequenced verified.
2- In vivo experiments
a- Bacterial viability assay and genetic experiments
In vivo complementation of the temperature sensitive phenotype of the MC4100 ∆tig::CmR
∆dnaKdnaJ::KanR, MC4100 Δtig dnaJ::Tn10-42, MC4100 Δtig ΔlondnaJ::Tn10-42 and MC4100
Δtig ΔhslOdnaJ::Tn10-42, was performed as follows. Cultures of fresh transformants were first
grown overnight in LB ampicillin glucose 0.4% at the permissive temperature (see Figure legends),
diluted 1/50 into the same medium, further grown to mid-log phase, serially diluted ten-fold and
spotted on LB ampicillin agar plates with or without IPTG (at 5, 50 or 500 µM) and incubated at the
indicated temperatures. Complementation of the W3110 ∆dnaKdnaJ::KanR ∆tig::Cm
R temperature-
sensitive phenotype by HipA was carried out as described above, except that L-arabinose inducer
(0.1, 0.5, 1, or 2%) was used in this case. To monitor Hsp33 toxicity of the MC4100 and its mutant
derivatives, mid-log phase cultures of fresh transformants were grown at the permissive growth
temperature (22°C or 30°C; see Figure legends) in LB ampicillin glucose 0.4%, were serially
diluted ten-fold and spotted on LB ampicillin agar plates with or without IPTG (100 or 500 µM),
and incubated at the indicated temperatures.
The co-transduction frequencies between ΔdnaKdnaJ::KanR and the linked thr::Tet
R alleles
were analyzed using bacteriophage P1-mediated transduction as described previously (Teter et al,
1999), except that MC4100 Δtig and MC4100 Δtig ΔhslO were used as the recipient strains.
b- Isolation of protein aggregates and cell fractionation
Cultures of fresh transformants of MC4100 ∆dnaKdnaJ::KanR ∆tig::Cm
R containing
plasmids pSE380∆NcoI, or pSE-Hsp33 wild type or its mutant derivatives were grown at 22°C,
diluted 1/50 and grown to an OD600 of 0.3 in LBampicillin (100 µg/ml) glucose (0.4%). IPTG
inducer (0.5 mM) was added and cultures were further incubated for 2 h at 22°C. Next, the cultures
were transferred to a 37°C water bath, and shaken (180 rpm) for 1 h and protein aggregates were
isolated as described by Tomoyasu and colleagues (Tomoyasu et al, 2001b). Protein aggregates
47
were prepared and analyzed by SDS-PAGE using 4–15% Mini-Protean TGX gels (Bio-Rad). For
HipA-mediated suppression of protein aggregates, cultures of fresh transformants of W3110
∆dnaKdnaJ::KanR ∆tig::Cm
S carrying plasmid pBAD22, or pBAD-HipA at 22°C were diluted 1/50
and grown to an OD600 of 0.3 in LBampicillin (100 µg/ml). L-arabinose inducer (0.5%) was then
added and cell cultures were further incubated for 4 h at 22°C. Cultures were then transferred to
37°C in a water bath, shaken (180 rpm) for 1 h and protein aggregates were prepared as described
(Tomoyasu et al, 2001b).
c- In vivo pull-down assay
E. coli MC4100, MC4100 ∆tig::CmR, MC4100 ∆dnaKdnaJ::Kan
R, and MC4100 ∆tig::Cm
R
∆dnaKdnaJ::KanR transformed with pSE-Hsp33FLAG were first selected at 22°C on LB agar plates
supplemented with glucose (0.4%) and ampicillin. Overnight cultures of fresh transformants, grown
at 22°C, were diluted to an OD600 of 0.1 and incubated until mid-log phase at 22°C in LB
supplemented with glucose 0.4% and ampicillin. Expression of Hsp33FLAG and
Hsp33(Y12E)FLAG was induced by the addition of 0.5 mM IPTG for 30 min at 22°C and
transferred to 30°C for 1 h and 30 min. Cells were collected by centrifugation (5000g for 5 min at
4°C) and resuspended in 500 µL of cold PD buffer (150 mM NaCl, 50 mM Tris pH 7.5, 1 mg/mL
lysozyme, and protease inhibitors from Roche) supplemented with Benzonase (3 units/mL culture
volume; Invitrogen). Crude cell extracts were obtained following sonication and centrifugation
(14000g for 30 min). Pull-down assays were performed with 500 µL aliquots of cell extract and 50
µL of Anti-Flag M2-Agarose resin (Sigma) in TBS buffer (150 mM NaCl, 50 mM Tris pH 7.5).
Suspensions were gently rocked at 4°C for 2 h, and beads were pelleted by centrifuging for 1 min at
5000g. After washing the beads 3 times with 1 mL of TBS buffer, proteins were eluted in TBS
buffer containing 100µg/mL of peptide Flag (DYKDDDDK). Samples were separated by SDS-
PAGE on 4–20% Mini-Protean TGX gels (Bio-Rad) and stained with Coomassie instant blue
(expedeon).
3- In vitro experiments
a- Western blot analysis
Proteins were separated by SDS-PAGE and transferred to a polyvinylidene difluoride
membrane (Hybond-P, GE Healthcare) using a semidry transfer system (Trans-Blot SD, Bio-Rad)
for 30 min at 10 V.Membranes were blocked for 1 h at RT or overnight at 4°C with 3% nonfat
milk-Tris-buffered saline with Tween 20 (TBS-T; 0.5 M Tris, 1.5 M NaCl, pH 7.4, plus 0.05%
48
Tween 20). Rabbit polyclonal antibody against Hsp33 (1:2000 dilution; a kind gift of Ursula Jakob,
University of Michigan), a mouse monoclonal antibody against EF-Tu (1:10000 dilution; Hycult
Biotech), a rabbit antibody against DnaJ (1:3000 dilution; (Genevaux et al, 2002)) and a rabbit
antibody against TF (1:8000 dilution; (Ullers et al, 2007)) were used as primary antibodies, and
HRP-conjugated rabbit IgG (1:5000; Sigma) or mouse IgG (1:2,000; Sigma) were used as
secondary antibodies. Membranes were developed by chemiluminescence using an ECL Plus
(Prime) immunoblotting detection kit following the manufacturer's protocol (GE Healthcare) with a
luminescence analyser (LAS4000, Fuji).
b- In vitro translation and cross-linking experiments
Strains MC4100 ∆dnaKdnaJ::KanR and ∆tig::Cm
R ∆dnaKdnaJ::Kan
R were first transformed
with pSE-Hsp33FLAG and grown in LB ampicillin medium at 30° and 25°C, respectively, were
used to obtain Hsp33-FLAG enriched translation lysates, as previously described for the SecB
chaperone (Ullers et al, 2004). RpoB150 mRNA was prepared as previously described (Ullers et al,
2004). In vitro translation in the E. coli cell- and membrane-free S-135 extract was carried out using
RpoB150 mRNA as previously described (Ullers et al, 2004; Urbanus et al, 2001). Bifunctional
cross-linking was induced by the addition of 1mM bis(sulfosuccinimidyl) suberate (BS3) for 10 min
at 26°C and quenched at 4°C by adding 1/10 volume of quench buffer (1 M glycine, 100 mM
NaHCO3, pH 8.5). Ribosome-nascent chain complexes were collected as described (Ullers et al,
2003) and analyzed either directly following SDS-PAGE (15%) separation or after
immunoprecipitation (Luirink et al, 1992) with a mouse antibody anti-FLAG (Sigma Aldrich).
c- Pulse-chase and immunoprecipitation analyses
Overnight cultures of MC4100 ∆hslO ∆dnaKdnaJ::KanR transformed with either pSE380 or
pSE-Hsp33 were diluted 1/50 in LB ampicillin, in the presence of 100 µCi/ml 35
S-methionine and
grown until the OD600 reached 0.4. Unlabeled methionine was then added at a final concentration of
15 mM and IPTG at 500 µM to induce expression of Hsp33. Samples were taken at time 0, 60, 120
min and kept overnight on ice cold TCA (10%), and analyzed by immunoprecipitation with anti-
EF-Tu antibodies as described (Jong et al, 2004). The gels were scanned using Fuji FLA-3000
phosphorimaging technology.
49
RESULTS
50
RESULTS
1- Hsp33 overproduction supports bacterial growth and prevents protein aggregation in
the absence of TF and DnaK
The ATP-independent and redox-regulated molecular chaperone Hsp33 (heat-shock protein
of 33 kDa), which is specifically activated by oxidative protein unfolding, appears to be a key
component of the network of heat-induced stress chaperones and proteases in E. coli(Chuang &
Blattner, 1993; Jakob et al, 1999). Specifically, in the presence of stressors such as hypochlorite or
hydrogen peroxide combined with high temperature, sensitive pairs of cysteine residues in Hsp33
are sequentially oxidized, leading to Hsp33 dimerization and activation of its chaperone function.
The resulting Hsp33 dimers bind unfolded substrate proteins and protect them from aggregation
(Jakob et al, 1999). Upon return to reducing conditions, Hsp33-bound substrates are most likely
delivered to DnaK for their subsequent refolding (Hoffmann et al, 2004).
To highlight a possible role of Hsp33 generic chaperone function in the absence of either
DnaK or oxidative stress, we asked whether Hsp33, as part of the chaperone network in E. coli,
could also restore bacterial growth of the severely compromised Δtig ΔdnaKdnaJ chaperone mutant
at higher temperatures (Genevaux et al, 2004). This strain lacks the TF/DnaK folding pathway and
thus represents a valuable tool for the identification of in vivo chaperone function. The bacterial
colony forming experiments presented in Fig. 1A show that indeed, over-expression of Hsp33
(encoded by the hslO gene, the last member of the yrfG-hslR-hslO operon) enables the MC4100
Δtig ΔdnaKdnaJ chaperone mutant to survive at the otherwise non-permissive temperatures of
33°C. In addition, we noticed that Hsp33 over-expression interferes somewhat with overall bacterial
growth, as judged by smaller colony size formation at the permissive temperatures of growth (Fig.
1A; see below). A western blot analysis showing Hsp33 expression levels in the presence of IPTG
inducer at the permissive temperature of 22°C is provided in Fig. 2. Further analysis of the extent of
assistance by Hsp33 in the absence of TF and DnaK showed that suppression by Hsp33 is not strain
background specific and is not further improved by the expression of the full yrfG-hslR-hslO operon
(Fig. 2).
51
As stated above, de novo protein folding is severely impaired in the absence of both TF and
DnaK chaperones. As a consequence, proteins massively accumulate as cytoplasmic aggregates in
the Δtig ΔdnaKdnaJ mutant strain. We took advantage of this observation to ask whether Hsp33
over-expression can also prevent the in vivo protein aggregation that occurs in the absence of TF
50 µM IPTG
37ºC
B
250 kDa -
130 -
100 -
70 -
55 -
35 -
25 -
17 -
11 -
WCE Aggregates
22ºC
33ºC
no IPTG
A
dilution
10-6
10-4
10-2
Hsp33
Fig. 1: Hsp33 over-expression rescues bacterial growth and prevents protein aggregation
in the absence of DnaK and TF. (A) Fresh transformants of strain MC4100 ∆tig::CmR
∆dnaKdnaJ::KanR
containing the plasmid pSE380∆NcoI parental vector, or pSE-Hsp33, or
pSE-Hsp33(Y12E), or pSE-Hsp33(M172S) or pSE-Hsp33(Q151E) were grown at 22°C,
serially diluted ten-fold and spotted on LB ampicillin agar plates with or without IPTG
inducer. Plates were incubated for 1 day at 33°C or 2 days at 22°C. (B) Protein aggregates and
their corresponding whole cell extracts (WCE) were isolated from strain MC4100 ∆tig::CmR
∆dnaKdnaJ::KanR
expressing the pSE-based Hsp33 derivatives, grown at 22°C and transferred
for 1 h at 37°C in the presence of IPTG to induce Hsp33 over-expression. The position of the
protein bands corresponding to Hsp33 or its mutant derivatives is indicated.
52
and DnaK, as previously shown for the overproduction of SecB or GroESL (Ullers et al, 2007).
Hsp33 was over-expressed in the MC4100 Δtig ΔdnaKdnaJ strain and aggregated proteins were
detected following 1 h incubation at the non-permissive temperature of 37°C, as described
(Tomoyasu et al, 2001a). As anticipated from the bacterial viability results, Hsp33 over-expression
efficiently prevented the accumulation of protein aggregates at 37°C (Fig. 1B).
We next asked whether mutations in the hslO gene, which were previously shown to
differentially affect activation of its Hsp33 chaperone function in vitro, could also rescue the defects
of the Δtig ΔdnaKdnaJ mutant in the absence of oxidizing conditions. Specifically, we chose the
following hslO mutations: Hsp33(Y12E), which is constitutively active in vitro but mainly
insoluble in vivo, Hsp33(M172S), activated by high temperature in the absence of oxidizing
conditions (Cremers et al, 2010), Hsp33(Q151E), constitutively monomeric and redox-regulated
with preference for slowly unfolding protein substrates (Chi et al, 2011), and Hsp33(1-235),
33ºC
22ºC
CA B
33ºC
22ºC- Hsp33
Fig. 2: Hsp33 promotes bacterial growth in the absence of DnaK and TF. (A) Western blot
analysis showing steady state expression levels of chromosomally- (vector) and plasmid- (pSE-
Hsp33) encoded Hsp33 in strain MC4100 ∆tig::CmR
∆dnaKdnaJ::KanR
after 6 h induction at
22°C in the presence of 50 µM IPTG and revelation using a rabbit polyclonal anti-Hsp33
antibody.(B) In vivo complementation of W3110 ∆tig::CmR
∆dnaKdnaJ::KanR
temperature-
sensitive phenotype by p29-Hsp33 in the presence of 500 µM IPTG, using the procedure
described in Fig. 1. (C) In vivo complementation in the presence of 500 µM IPTG, of MC4100
∆tig::CmR
∆dnaKdnaJ::KanR
temperature-sensitive phenotype by p29-Hsp33 (hslO) alone, with
Hsp15 (hslR-hslO) or in the context of the whole yrfG-hslR-hslO operon.
53
constitutively active as a dimer in vitro(Kim et al, 2001). The corresponding mutant genes were
cloned in a multicopy plasmid under the control of an IPTG-inducible promoter and tested for their
ability to suppress the growth defects exhibited by the MC4100 Δtig ΔdnaKdnaJ mutant strain. We
found that, in sharp contrast to the Hsp33 wild-type, over-expression of either the Hsp33(Y12E), or
Hsp33(M172S), or Hsp33(1-235) mutant protein did not rescue bacterial growth at 33°C (Fig. 1A;
Fig. 3). These results suggest that the integrity of the redox switch mechanism is critical for the
Hsp33-mediated suppression of the Δtig ΔdnaKdnaJ phenotype. Consistently, neither the
Hsp33(Y12E) nor the Hsp33(M172S) mutant was capable of preventing protein aggregation at
37°C in vivo (Fig. 1B). Instead, these two mutant Hsp33 proteins co-purified with the protein
aggregates. Although it is known that Hsp33(Y12E) is mainly insoluble in vivo(Cremers et al,
2010), it is possible that the presence of the Hsp33(M172S) in protein aggregates represents a bona
fide chaperone-substrate interaction.
33ºC
22ºC
A
70kDa -
55 -
35 -
25 -
17 -
11 -
- Hsp33(1-235)
B
Fig. 3: Hsp33(1-235) mutant does not support bacterial growth in the absence of DnaK
and TF. (A) In vivo complementation of MC4100 ∆tig::CmR
∆dnaKdnaJ::KanR
temperature-sensitive phenotype by pSE-Hsp33 and pSE-Hsp33(1-235) in the presence of
50 µM IPTG, using the procedure described in Fig. 1. (B) Coomassie-stained SDS-PAGE
showing the steady state expression of Hsp33(1-235) at 22°C in strain MC4100 ∆tig::CmR
∆dnaKdnaJ::KanR
after 6 h induction at 22°C in the presence of 50 µM IPTG.
54
The monomeric and redox-regulated Hsp33(Q151E) mutant displayed a sharply different
behavior. It turned out that even at the permissive temperature of growth Hsp33(Q151E) was highly
toxic to bacterial growth at all temperatures tested in the MC4100 Δtig ΔdnaKdnaJ mutant
background when induced(Fig. 1A). In sharp contrast to its deleterious effect on bacterial growth,
monomeric Hsp33(Q151E) fully prevented the accumulation of protein aggregates at the non-
permissive temperature of 37°C, as well as, or even better than Hsp33 wild-type (Fig. 1B). These
findings raise the possibility that the monomeric Hsp33 species, which is believed to be transiently
populated during the Hsp33 activation cascade process and which shows strong preference for
slowly unfolding substrates in vitro(Graf et al, 2004; Jakob et al, 2000), may exhibit robust
substrate binding properties in the absence of the TF/DnaK pathway.
2- Hsp33 function is critical in the absence of both TF and DnaK
The suppression of both the Δtig ΔdnaKdnaJ temperature-sensitive growth phenotype and
the intracellular protein folding defects following Hsp33 overproduction reveal an important
functional cooperation among these three major chaperones. To get additional biological clues for
such interplay we examined the phenotypes of various mutational combinations among these
chaperone genes in the MC4100 strain background. A phenotypic analysis of the double Δtig ΔhslO
and ΔdnaKdnaJ ΔhslO mutants revealed only minor growth differences compared to their
respective single mutant parents, i.e., the double ΔdnaKdnaJ ΔhslO mutant was more temperature-
sensitive for growth at 40°C than its isogenic single ΔdnaKdnaJ mutant (Fig. 4).
55
30ºC
37ºC
40ºC
43ºC
IPTG
50µM
Fig. 4: Comparative growth of the MC4100 Δtig, Δtig ΔhslO,ΔdnaKdnaJ,ΔdnaKdnaJ ΔhslO
chaperone mutants. Mid-log phase cultures of (i) MC4100 ∆tig::CmR
, ∆tig::CmR
ΔhslO,∆dnaKdnaJ::KanR
,∆dnaKdnaJ::KanR
ΔhslO wereserial diluted and spotted on LB agar
plates and (ii)MC4100 ∆dnaKdnaJ::KanR
ΔhslO complemented with p29SEN vector or with
p29-Hsp33 were serial diluted and spotted on LB ampicillin agar plates supplemented with
50µM IPTG at different temperatures.
56
Next, we tested the viability of the triple Δtig ΔdnaKdnaJ ΔhslO mutant compared to
itsisogenic Δtig ΔdnaKdnaJ mutant parent. To do so, we used the same methodology as we
previously described for the construction of the Δtig ΔdnaKdnaJ chaperone mutant (Genevaux et al,
2004; Teter et al, 1999). Specifically, we first investigated the co-transduction frequency between
the ΔdnaKdnaJ::KanR mutant allele and itsnearby ~40% linked thr::Tet
R resistance marker using
either MC4100 Δtig or MC4100 Δtig ΔhslO asrecipient strains (Fig. 5A). As anticipated from our
previous work, the ΔdnaKdnaJ mutation was efficiently transduced into the Δtig single mutant with
the expected 40% co-transduction frequency. In contrast, the occasional ΔdnaKdnaJ::KanRthr::Tet
R
co-transductants formed extremely small colonies and, in addition, appeared at a ten-fold lower
frequency (4% instead compared to the expected ~40%) when the Δtig ΔhslO strain was used as the
recipient. Most likely, the few, slowly growing transductants are due to the continuous
accumulation of unknown extragenic suppressors, which enable the occasional survival of the Δtig
ΔdnaKdnaJ ΔhslO triple mutant.
To further demonstrate such a synergy among the various chaperones, we took advantage of
the phenotype exhibited by null mutations in the dnaJ gene (the main DnaK co-chaperone), which
exhibit a significantly less severe growth phenotype defect due to the presence of the djlA and cbpA
genes, known to code for two additional DnaJ-like family members, capable of partially replacing
DnaJ function (Genevaux et al, 2007; Genevaux et al, 2004). We previously showed that the double
tigdnaJ mutant is temperature-sensitive for growth above 34°C in a strictly DnaK-dependent
manner (Genevaux et al, 2004). Based on these earlier results, we introduced the dnaJ::Tn10-42
mutation into either the MC4100 Δtig or MC4100 Δtig ΔhslO mutant backgroundsat 30°C and
subsequently compared their bacterial growth phenotypes at various temperatures (Fig. 5B). In
agreement with the synthetic lethality observed in the absence of DnaK (Fig. 5A), the triple Δtig
ΔhslO dnaJ mutant growth was shown to be significantly more temperature-sensitive than its
isogenic Δtig ΔdnaJ double mutant parent. As expected, the colony-forming ability of this triple
mutant was fully restored following Hsp33 expression from a plasmid (Fig. 5B). The absence of TF,
Hsp33 or DnaJ protein antigens in the corresponding null mutant strains was confirmed by western
blot analysis (Fig. 5C). Taken together, our above data highlight for the first time the essential
functional interplay between the TF, DnaKJE and Hsp33 chaperones in vivo.
57
Transductants
Recipient strains TetR KanR
MC4100 ∆tig 283 113 (39%)
MC4100 ∆tig ∆hslO 277 13* (4%)
A
B
33ºC
30ºC
37ºC
30ºC
-TF
C
-DnaJ
-Hsp33
Fig. 5: Hsp33 is essential for bacterial viability in the absence of DnaK and TF. (A)
Synthetic lethality among the dnaK, tig and hslO genes was analyzed by P1-mediated
transduction experiments. Transductants were first selected on tetracycline plates following a
two days incubation at 22°C (the permissive temperature for the tig dnaK doublemutant) and
subsequently tested for possession of the kanamycin resistance carried by the linked
ΔdnaKdnaJ::KanR
mutant allele. The co-transduction frequencies of the KanR
marker are
given as percentage of the total number of TetR
transductants. The asterisk indicates the
formation of very small colonies. (B) Synergistic effect of the tig,dnaJ and hslO mutations on
bacterial growth. Mid-log phase cultures of either (i) MC4100 ΔtigdnaJ::Tn10-42 and
MC4100 ΔtigΔhslOdnaJ::Tn10-42, or (ii) MC4100 dnaJ::Tn10-42ΔtigΔhslO containing
p29SEN-Hsp33 were serially diluted ten-fold and spotted on LB agar plates supplemented
with 500 µM IPTG. (C) Whole-cell extracts of MC4100, MC4100 ΔtigdnaJ::Tn10-42 and
MC4100 ΔtigΔhslOdnaJ::Tn10-42 were separated by SDS-PAGE and analyzed by western
blot using anti-TF, -DnaJ and -Hsp33 antibodies.
58
3- Hsp33 specifically interacts with EF-Tu
The genetic suppression results suggested that, as observed for SecB and GroESL
(Genevaux et al, 2004; Ullers et al, 2004), Hsp33 could bind de novo substrates of TF and DnaKJE
and efficiently prevent their aggregation. However, in contrast to SecB, in vitro crosslinking
experiments did not reveal any co-translational interaction with our model substrate RpoB150,
previously shown to interact with TF, DnaK and SecB, and to aggregate in the absence of TF and
DnaK (Fig. 6;(Ullers et al, 2004)). We next proceeded to search for Hsp33 interacting proteins in
four isogenic strains, namely, MC4100 wild type and its Δtig, ΔdnaKdnaJ and Δtig ΔdnaKdnaJ
mutant derivatives. In these strains, in vivo pull-down experiments were carried out using over-
expressed Flag-tagged Hsp33 as bait (Fig. 7A). An analogous pull-down experiment with a lysate
expressing untagged Hsp33 is shown as a control in Fig. 6. We found that one major Hsp33
interacting partner could be detected in all four genetic backgrounds tested, which was identified by
mass spectrometry analysis to be the essential elongation factor EF-Tu, a key regulator of the
polypeptide chain elongation cycle during the translation process (Fig. 7A; (Thompson et al, 1986)).
This finding suggested that the specific interaction between EF-Tu and Hsp33 might be relevant for
the suppression of the Δtig ΔdnaKdnaJ mutant phenotypes by Hsp33.
59
X-link IP
250 kDa -
130 -
100 -
70 -
55 -
35 -
25 -
17 -
Hsp33Flag
EF-Tu
**
*
BAA
30-
46-
69-
97-
14-
RpoB150
Fig. 6: Hsp33 interactor. (A) Search for co-translational interaction between ribosome-
associated nascent chain of RpoB150 and Hsp33-FLAG using in vitro translation of RpoB150
transcripts in a Hsp33-FLAG enriched lysate of MC4100 ∆tig::CmR
∆dnaKdnaJ::KanR
.
Crosslinking was performed with 1mM of BS3 cross-linker or with water (H
2O) as a
control.Complexes were purified and separated directly or immunoprecipitated using an
antibody anti-FLAG and migrated on SDS-PAGE as described by Ullers et al, (2004). The
bracket indicates the expected area for a RpoB150-Hsp33-FLAG crosslinking product. (B)Pull-
down control with untagged (Hsp33) or tagged Hsp33-Flag or Hsp33(Q151E)-Flag expressed in
MC4100, as described in Fig. 7A. Asterisks represent nonspecific interactions with the Anti-Flag
M2-Agarose resin observed in the absence of tagged Hsp33.
60
Next, we monitored the endogenous EF-Tu levels in response to Hsp33 over-expression in
the Δtig ΔdnaKdnaJ mutant. In this case, the Hsp33(Y12E) and Hsp33(Q151E) were used as
inactive and hyper-active controls respectively, as deduced from the results shown in Fig. 1. We
found that the endogenous EF-Tu levels rapidly diminished following Hsp33 over-expression,
especially at the non-permissive temperature of 37°C (Fig. 7B). A prolonged over-expression of
Hsp33 at 22°C also led to a decrease of EF-Tu (Fig. 8). In addition, the effect on EF-Tu was
exacerbated in the presence of the constitutively monomeric and redox-regulated Hsp33(Q151E)
mutant (Fig. 7B), which also efficiently binds EF-Tu as shown in Fig. 6. This last result suggests
that distinct intermediates populating the multistep activation cycle of Hsp33 could possess unique
functions in vivo(Akhtar et al, 2004; Chi et al, 2011; Graf et al, 2004). In contrast, and as expected,
the Hsp33(Y12E) mutant was completely non-functional, behaving like the plasmid vector control
(Fig. 7B).
B
250 -
130 -
100 -
70 -
55 -
35 -
25 -
17 -
Hsp33
EF-Tu
-EF-Tu
250 -
130 -
100 -
70 -
55 -
35 -
25 -
17 -
Hsp33Flag
EF-Tu
A
Fig. 7: Hsp33 specifically interacts with EF-Tu. (A) Identification of the Hsp33 interacting substrates by
pull-down analysis using an Hsp33-FLAG version expressed in either MC4100, or MC4100 ∆tig::CmR
, or
MC4100 ΔdnaKdnaJ::KanR
or MC4100 ∆tig::CmR
∆dnaKdnaJ::KanR
at 30°C. The main co-precipitated
protein obtained in all four strains was EF-Tu as identified by a mass spectrometric analysis. (B)
Endogenous levels of EF-Tu in MC4100 ∆tig::CmR
∆dnaKdnaJ::KanR
following over-expression of Hsp33
wild type or its mutant derivatives Hsp33(Y12E) and Hsp33(Q151E). Cultures of MC4100 ∆tig::CmR
∆dnaKdnaJ::KanR
transformed with the various pSE380-based constructs were grown in LB ampicillin,
supplemented with 0.4% glucose at 22°C to an OD600
of 0.3. IPTG was then added at 500 µM for 30 min at
22°C for pre-induction and cultures were transferred to either 22°, or 30° or 37°C for 1 h and 30 min. Whole
cell extracts were prepared, separated by SDS-PAGE and stained with Coomassie blue. The characterization
of the EF-Tu protein was confirmed by the western blot analysis shown at the bottom of the panel.
61
Although at this stage we cannot completely rule out the possibility that Hsp33, as a
chaperone, directly prevents protein aggregation in the absence of TF and DnaK, our results
strongly suggest that modulation of the rate of polypeptide chain elongation by Hsp33 might be
responsible for bacterial survival. Our in vivo pull-down experiments described above never
revealed a major difference in the qualitative nature of the Hsp33 interactor pattern. Nevertheless,
we reproducibly noticed that the EF-Tu/Hsp33 ratio was higher in the wild type and the Δtig
mutantthan for the ΔdnaKdnaJ or Δtig ΔdnaKdnaJ isogenic mutant strains (Fig. 7A). Consequently,
we tested whether the effect of Hsp33 on EF-Tu was different in the absence of TF, DnaK or both.
To do so, we over-expressed Hsp33 in the four isogenic strains and monitored the EF-Tu levels in
whole cell extracts. The results presented in Fig. 9A show that upon Hsp33 over-expression, the EF-
Tu levels are not significantly affected in the wild-type strain, besides the slight decrease in EF-Tu
observed for the single Δtig mutant at 30°C and 37°C. In contrast, the EF-Tu intracellular levels
250 kDa -
130 -
100 -
70 -
55 -
35 -
25 -
17 -
MC4100 Δtig ΔdnaKdnaJ
vector Hsp33
Hsp33
EF-Tu
time (min)
Fig. 8: Steady state levels of EF-Tu following a prolonged expression of Hsp33 in the
absence of DnaK and TF. Overnight cultures of MC4100 ∆tig::CmR
∆dnaKdnaJ::KanR
transformed with pSE380 vector or with pSE-Hsp33 grown in LB ampicillin glucose
0.4% at 22°C were diluted 1/50 and grown until OD600
0.3. IPTG 500µM was then added
for 30 min at 22°C and strains were transferred at 30°C for 0, 90 min, 120 min, 150 min,
180 min. Whole cell extracts were prepared and separated on SDS-PAGE, and proteins
were stained with Coomassie Blue.
62
dropped very rapidly when Hsp33 was over-expressed in the ΔdnaKdnaJ mutant, even at 30°C. The
expression of Hsp33 in the Δtig ΔdnaKdnaJ mutant resulted in an intermediate behavior. Note that
the Hsp33 over-expression levels were comparable in all instances (Fig. 9A). The results presented
in Fig. 9B suggest that the diminution of the endogenous EF-Tu directly correlates with bacterial
growth inhibition at the otherwise permissive temperature of 22°C, coupled with a marked toxicity
of Hsp33 in the ΔdnaKdnaJ mutant.
-RpoB150
C
*
100µM
IPTG
22ºC
B
-
- EF-Tu
A
130 -
100 -
70 -
55 -
35 -
25 -
17 -
Hsp33
Fig. 9: Hsp33-mediated control of EF-
Tu in the absence of DnaK. (A)
Endogenous EF-Tu levels in MC4100,
MC4100 ∆tig::CmR
, MC4100
ΔdnaKdnaJ::KanR
or MC4100
∆tig::CmR
∆dnaKdnaJ::KanR
following
Hsp33 over-expression were analyzed as
described in the legend to Fig. 7B. Arrows
indicate the position of EF-Tu in whole
cell extracts. (B) Over-expression of
Hsp33 (pSE-Hsp33) in MC4100 and its
mutant derivatives. Bacterial cultures
grown at 22°C were serially diluted ten-
fold and spotted on LB ampicillin agar
plates at 22°C with or without IPTG 100
µM. The letter v stands for the vector and
the number 33 stands for the Hsp33-
carrying plasmid. (C) In vitro translation
of the first 150 amino acids of RpoB using
cell- and membrane-free extracts of
MC4100 ΔdnaKdnaJ::KanR
and MC4100
∆tig::CmR
∆dnaKdnaJ::KanR
mutants over-
expressing Hsp33-FLAG. After in vitro
translation, radiolabeled samples were
resuspended in loading buffer and
separated by a SDS/15% PAGE. The
asterisk indicates the position of the
truncated RpoB150 polypeptide (left lane).
The position of the full length RpoB150 is
indicated as RpoB150 (right lane).
63
These findings suggest that in the absence of TF and DnaK, over-expressed Hsp33 may
target EF-Tu for degradation by proteases. Moreover, the effect on EF-Tu was exacerbated in the
ΔdnaKdnaJ mutant, thus suggesting that TF may also facilitate interaction of EF-Tu with Hsp33,
leading to its subsequent degradation. We first examined whether Hsp33 over-expression
differentially affects polypeptide chain elongation in vitro. To do so, we prepared translation lysates
from both ΔdnaKdnaJ and Δtig ΔdnaKdnaJ mutant strains over-expressing Hsp33, and
subsequently compared their ability to support translation of the RpoB150 mRNA in vitro(Ullers et
al, 2004). In agreement with our previous work, we found that cell lysates obtained from the Δtig
ΔdnaKdnaJ mutant strain indeed efficiently translated RpoB150 mRNA, as judged by the presence
of labeled, full length RpoB150 nascent chains (Fig. 9C;(Ullers et al, 2004)). In sharp contrast,
lysates prepared from the ΔdnaKdnaJ mutant strain expressing Hsp33 were not capable of
translating the RpoB150 mRNA to completion, as judged by the absence of the full length RpoB150
nascent chain, with concomitant appearance of a putative short truncated RpoB150 translation
product (marked with an asterisk; left lane of Fig. 9C). These results are in agreement with the
observed dramatic decrease in the levels of endogenous EF-Tu in the absence of DnaK with the
simultaneous overproduction of Hsp33 and the resulting strong toxicity on bacterial growth(Fig. 9A
and B).
4- Hsp33 triggers EF-Tu degradation by the stress protease Lon
Our results suggest that in the absence of DnaK, and to lesser extent in the absence of both
DnaK and TF, the chaperone Hsp33 specifically binds and delivers EF-Tu to yet unknown
protease(s). In order to identify such protease(s), we took advantage of the strong DnaK-dependent
toxicity of Hsp33 observed in Fig. 9B. We reasoned that mutation in the specific protease might
abolish Hsp33-dependent decrease of endogenous EF-Tu and the resulting toxicity. Therefore, we
combined single deletion of either Δlon, ΔclpP or ΔclpQ major cytoplasmic AAA+ proteases(Sauer
& Baker, 2011) with the ΔdnaKdnaJ mutation at 30°C, over-expressed Hsp33 and monitored its
effect on bacterial growth and EF-Tu. The results presented in Fig. 10A clearly show that in
contrast to ΔclpP and ΔclpQ, the Δlon mutation efficiently suppresses Hsp33 toxicity and restores
endogenous EF-Tu close to wild-type level. Yet, a low amount of EF-Tu was also recovered in the
absence of clpP, suggesting some overlap between Lon and ClpP for this substrate (Fig. 10A,
bottom).
To ensure that the decreased EF-Tu levels were due to EF-Tu degradation and not to an
effect on transcription, the stimulation of EF-Tu degradation by over-expressed Hsp33 was further
demonstrated using pulse chase experiments followed by immunoprecipitation of EF-Tu in the
64
ΔdnaKdnaJ strain expressing Hsp33. The results presented in Fig. 10B show that indeed, Hsp33
significantly accelerates EF-Tu degradation in the absence of DnaK, suggesting that Hsp33
maintains EF-Tu in a conformation competent for degradation.
-EF-Tu
ΔdnaKdnaJ
30ºC
500µM
IPTG
-
-
A
B
ΔhslO ΔdnaKdnaJ
+Hsp33
-
Chase(min) 0 60 120
EF-Tu
Fig. 10: Hsp33 stimulates Lon protease-
mediated degradation of EF-Tu. (A)
Cultures of MC4100 ΔdnaKdnaJ::KanR
,
MC4100 Δlon ΔdnaKdnaJ::KanR
, MC4100
ΔclpP::CmR
ΔdnaKdnaJ::KanR
and MC4100
ΔclpQ::TetR
ΔdnaKdnaJ::KanR
transformed
with pSE380 vector (indicated as v) or pSE-
Hsp33 (indicated as 33), were serially diluted
ten-fold and spotted on LB ampicillin agar
plates with or without 500 µM IPTG inducer
at 30°C. Steady state levels of EF-Tu in the
corresponding strains were analyzed by
western blot analysis using liquid cultures
following a 2 h incubation in the presence of
500 µM IPTG inducer at 30°C. (B) The
kinetics of EF-Tu degradation in vivo in
MC4100 ∆hslO ∆dnaKdnaJ::KanR
bacteria
transformed with either the pSE380 vector or
pSE-Hsp33 was analyzed by first labeling
with 35
S-methionine, followed by a 0, or 60
or 120 min chase with excess, unlabeled
methionine and co-immunoprecipitation.
65
5- EF-Tu inhibition helps bacterial growth in the absence of TF and DnaK
The proposed control of polypeptide chain elongation by a network of chaperones and
proteases suggests that the suppression by Hsp33 in the absence of TF and DnaK is linked to EF-Tu
degradation and not to the previously observed in vitro chaperone activity of EF-Tu(Caldas et al,
1998). In agreement with such a hypothesis, EF-Tu over-expression does not rescue the growth
defect of the MC4100 Δtig ΔdnaKdnaJ mutant strain (Fig. 11A). As a control, the steady state level
of plasmid-encoded EF-Tu is shown in Fig. 12. We next examined whether Hsp33-mediated
degradation of EF-Tu by Lon was responsible for the suppression in the absence of TF and DnaKJ.
We expressed Hsp33 both in the Δtig dnaJ and in the Δtig Δlon dnaJ mutants, and compared
suppression of the temperature-sensitive phenotype by Hsp33. Note that as observed for MC4100
Δtig ΔdnaKdnaJ, over-expression of Hsp33 in the less sensitive MC4100 ΔtigdnaJ mutant strain
similarly stimulated EF-Tu degradation by Lon at high temperature of growth (Fig. 12).
Remarkably, the results presented in Fig. 11B clearly show that in contrast with the expected
suppression observed for the Δtig dnaJ (lon+) double mutant, Hsp33 was not capable of rescuing
bacterial growth in the absence of Lon (Fig. 11B). These results strongly suggest that control of the
endogenous EF-Tu levels is critical when the TF/DnaKJE pathway is compromised.
Next, we took advantage of the HipA toxin of the HipAB toxin-antitoxin module from E.
coli, known to bind to and inhibit EF-Tu by phosphorylation at residue Thr382, thus compromising
polypeptide chain elongation(Hansen et al, 2012; Schumacher et al, 2009). We first asked whether
HipA over-expression could mimic Hsp33-mediated control of EF-Tu in the absence of both TF and
DnaK. To do so, gene encoding HipA was cloned in a plasmid under the control of the tightly
regulated araBADp promoter (to avoid growth inhibition by the toxin) and expressed in the
sensitive Δtig ΔdnaKdnaJ mutant. As expected for a toxin affecting translation, HipA expression
significantly slowed down bacterial growth at the permissive temperature of 22°C (Fig. 11C). In
contrast, at the non-permissive temperature of 37°C, a mild over-expression of HipA efficiently
rescued bacterial growth and prevented protein aggregation in the absence of both TF and DnaK
(Fig. 11C and D). These results indicate that a partial inactivation of EF-Tu function may be
beneficial to bacterial survival in the absence of the major TF and DnaK chaperones and highlights
a possible role for the toxin-antitoxin system in controlling the rate of protein synthesis in response
to protein aggregation (see Discussion below).
66
C
HipA
22°C 33°C
Δtig ΔdnaKdnaJ
- +
37°C
- + - +
30°C
37°C
A
Hsp33-
B
D
WCE Agg.
130 -
100 -
70 -
55 -
35 -
25 -
17 -
37°C
50050-IPTG
22°C
33°C
Fig. 11: EF-Tu inactivation facilitates growth and protein folding in the absence of TF and DnaK
chaperones. (A) EF-Tu over-expression in MC4100 ∆tig::CmR
∆dnaKdnaJ::KanR
. Cultures of
MC4100 ∆tig::CmR
∆dnaKdnaJ::KanR
transformed with p29SEN or with p29-EF-Tu were grown at
22°C, serially diluted ten-fold and spotted on LB ampicillin agar plates with or without IPTG and
incubated at 22° or 33°C. (B) Cultures of MC4100 Δtig::CmR
dnaJ::Tn10-42 or MC4100
Δtig::CmR
dnaJ::Tn10-42 Δlon::KanR
transformed with either p29SEN (indicated as v) or with p29-
Hsp33 (indicated as 33) were grown in LB ampicillin, supplemented with glucose 0.4% at 30°C,
serially diluted ten-fold and spotted on LB ampicillin agar plates with or without IPTG 50 µM at 30°
or 37°C. Protein levels were analyzed by western blot analysis of the extracts following a 2 h
induction with IPTG 50 µM at 30°C. The genetic background of each strain used is shown on top of
each panel. (C) TheHipA toxin efficiently rescues bacterial growth in the absence of TF and DnaK.
Strain W3110 Δtig::CmR
ΔdnaKdnaJ::KanR
pBAD-HipA was grown without (-) or with (+) 0.5% of
L-arabinose inducer at the indicated temperatures. (D) Prevention of protein aggregation by HipA
toxin expressed from the pBAD-HipA plasmid in strain W3110 Δtig::CmR
ΔdnaKdnaJ::KanR
. HipA
expression was induced with 0.5% L-arabinose for 6 h at 22°C followed by 1 h 30 min at 33°C. Whole
cell extracts and aggregates were separated by SDS-PAGE and stained with Coomassie blue.
67
EF-Tu EF-Tu
BA
Fig. 12: EF-Tu expression. (A) EF-Tu over-expression from plasmid p29-EF-Tu in
MC4100 ∆tig::CmR
∆dnaKdnaJ::KanR
strain was induced overnight with 500µM IPTG at
22°C. EF-Tu was revealed by western blot using anti-EF-Tu antibody. (B) EF-Tu
endogenous levels following Hsp33 over-expression in the absence of either TF/DnaJ, or
TF/DnaJ/Lon. Overnight cultures of MC4100 ∆tig dnaJ (lon+
) and MC4100 ∆tig dnaJ
∆lon transformed with pSE-Hsp33 grown in LB ampicillin glucose 0.4% at 30°C were
diluted 1/50 and grown until OD600
0.3. IPTG 500µM was then added for 30 min at 30°C
and strains were transferred at 43°C for 3 h. Whole cell extracts were prepared and
separated on SDS-PAGE, and EF-Tu was revealed by western blot using anti-EF-Tu
antibody.
68
DISCUSSION
69
DISCUSSION
The heat shock protein Hsp33 is a highly specialized redox-regulated molecular chaperone, which is
activated by a combination of both oxidative stress and protein unfolding or aggregation (Kumsta &
Jakob, 2009). In this work, we propose that Hsp33 can also perform key, house-keeping cellular
functions in the absence of exogenous stressors as part of the network of stress chaperones and
proteases in E. coli. This conclusion was reached by demonstrating that over-expression of Hsp33
can efficiently support bacterial growth and prevent accumulation of protein aggregates in the
absence of TF and DnaK. However, in contrast with other chaperones, such as SecB and GroESL
(Genevaux et al, 2004; Ullers et al, 2004; Vorderwulbecke et al, 2004), suppression by Hsp33 does
not seem to rely on its general chaperone function. Instead, it appears that Hsp33 efficiently binds to
the essential elongation factor EF-Tu and specifically targets it for degradation by the Lon protease.
The subsequent decrease in endogenous EF-Tu level may result in a decrease of the intracellular
rate of nascent polypeptide synthesis, thus minimizing the need for action by downstream
chaperones TF and DnaKJE.
Although we do not know yet about the relevance of such pathway under physiological
conditions where all the chaperones are present, our results point toward the existence of an
intricate network of stress chaperones and proteases that result in an inhibition of the rate of
translation in response to severe protein aggregation. It is known that under stress conditions that
affect protein folding, several major, stress-induced chaperones, including DnaK, are recruited to
the accumulating protein aggregates (Winkler et al, 2010). Importantly, such a recruitment of DnaK
to protein aggregates results in the rapid stabilization of the heat shock sigma factor σ32
, thus
leading to a prolonged induction of heat shock proteins, including Hsp33 and Lon. Note that in our
case, both the induction of heat shock proteins and the aggregation-prone conditions are, at least in
part, artificially recreated by the introduction of the tig dnaKdnaJ mutations (Genevaux et al, 2004).
Yet, in the absence of these dominant chaperones, we cannot exclude that induction of the stress
response or accumulation of extragenic suppressors that help bacterial growth without TF and
DnaK could force the bacteria to use pathways that are not utilized under normal conditions (Bukau
& Walker, 1990; Kramer et al, 2004b). In our working model, the increased recruitment of the
major molecular chaperones to preexisting aggregates and newly-synthesized proteins rapidly
overpowers the cellular chaperone capacity available for de novo protein folding. This, in turn,
results in a signal that triggers a transient inhibition of the rate of translation and a subsequent
reduction in the intracellular levels of newly-synthesized polypeptides that normally necessitate
molecular chaperones for their proper folding (Hartl et al, 2011). Such a possible mechanism is
supported by previous studies showing that slowing down translation facilitates de novo protein
70
folding in E. coli(Agashe et al, 2004; Siller et al, 2010) and in mammalian cells, as shown recently
(Meriin et al, 2012).
If such an intricate control of translation rate by the availability of stress chaperones and
proteases indeed exists, then the results presented in this work clearly point to the elongation factor
EF-Tu as one of the major targets of such an inhibition. Interestingly, in addition to the Hsp33/EF-
Tu complex demonstrated in this study, an interaction between EF-Tu and Lon has also been found
in vivo(Butland et al, 2005), further supporting the proposed interplay among chaperones and
proteases. However, it remains to be determined whether EF-Tu is the sole protein directly targeted
for degradation by the Hsp33/Lon pathway and whether other minor interactions that our
experimental procedures are missing could also participate.
It is known that EF-Tu adopts several conformations and populates various intermediate
stages during its functional cycle in polypeptide chain elongation (Agirrezabala & Frank, 2009). At
this time, the actual conformational state(s) of EF-Tu that is recognized by Hsp33 and Lon remains
unknown. Intriguingly, in contrast to our finding that in E. coli Hsp33 stimulates EF-Tu degradation
by Lon, it was recently shown in V. cholerae that Hsp33 was instead required to maintain EF-Tu at
cellular levels sufficient to allow bacterial growth at high temperature under aerobic conditions
(Wholey & Jakob, 2012). In this case, Hsp33 stabilized the highly sensitive V. cholerae EF-Tu
protein from oxidative protein degradation in vivo. Interestingly, the E. coli EF-Tu protein was not
affected under the same conditions (Wholey & Jakob, 2012). The differential results seen with
different bacteria strongly suggest that although the proposed network controlling EF-Tu
intracellular levels is conserved, the overall outcome may differ depending on the particular
organism and/or stress conditions employed.
The fact that degradation of EF-Tu by Hsp33 and Lon predominantly occurs when DnaK is
absent suggests that under normal growth conditions DnaK prevents EF-Tu degradation, either by
directly interacting with EF-Tu or indirectly by inhibiting the σ32
-dependent synthesis of Lon. In
contrast with such a protective effect of DnaK, we found that the presence of TF significantly
stimulates EF-Tu degradation, indicating that TF directly or indirectly facilitates the interaction of
EF-Tu with Hsp33 and/or its subsequent recognition by Lon. Such a hypothesis is strongly
supported (i) by a recent study demonstrating that in contrast with DnaK, the ribosome-bound TF
chaperone can unfold substrates and facilitate their degradation in vitro(Hoffmann et al, 2012), and
(ii) by the synthetic lethality observed between the tig, dnaKdnaJ and hslO mutations (Fig. 5A).
How does Hsp33 contribute to such a network in the absence of oxidative stressors? One
possibility is that Hsp33 is only activated by increasing levels of reactive oxygen species (ROS)
stimulated by the severe protein homeostasis collapse induced by the lack of both TF and DnaK
71
major chaperones (Calloni et al, 2012). However, this seems unlikely because (i) Hsp33 efficiently
binds EF-Tu in the wild type E. coli strain as well, and (ii) EF-Tu is much more efficiently degraded
in the dnaKdnaJ mutant than in the triple tig dnaKdnaJ mutant where proteostasis breakdown is the
most severe (Deuerling et al, 1999; Genevaux et al, 2004; Teter et al, 1999). Therefore, we propose
that a significant fraction of endogenous Hsp33 may be active and physiologically relevant even in
the absence of oxidative stress. This is supported by the synergistic effects observed among the tig,
dnaK (or dnaJ)and hslO mutations. However, the redox and the oligomeric status (monomeric,
dimeric or oligomeric) of cellular Hsp33 that is capable of performing such a function in vivo
remains to be determined (Akhtar et al, 2004; Chi et al, 2011; Graf et al, 2004). In addition, it is not
yet established whether Hsp33 induces EF-Tu degradation under physiological conditions where all
the chaperones are present. As stated above, Hsp33 is activated by oxidative protein unfolding
(Chuang & Blattner, 1993; Jakob et al, 1999), a condition that is known to impede DnaK functions
(Hoffmann et al, 2004). Under such extreme stress condition, quality control of EF-Tu by activated
Hsp33 and Lon could thus become a key to bacterial survival (Fig. 13). Further studies are needed
to elucidate such possible interplay under severe oxidative conditions.
Analogous to the newly identified cooperation between Hsp33 and Lon in controlling of EF-
Tu, over-expression of the HipA toxin of the chromosomally-encoded type-II HipAB toxin-
antitoxin system can also suppress both the growth defect and the accumulation of protein
aggregates in the absence of TF and DnaK. Toxin-antitoxin modules are two-component systems
composed of a stable toxin that forms an inactive complex with its less stable cognate antitoxin. In
response to specific stress conditions the unstable antitoxin is generally degraded by stress proteases
and the thusly free active toxin subsequently targets important cellular processes such as DNA
replication or protein synthesis, resulting in growth inhibition and eventual cell death (Gerdes et al,
2005; Moll & Engelberg-Kulka, 2012; Van Melderen & De Bast, 2009; Yamaguchi et al, 2011) .
Recent work has shown that the E. coli toxin HipA is a protein kinase belonging to the Tor family,
which specifically inhibits EF-Tu and blocks translation (Correia et al, 2006; Schumacher et al,
2009). In agreement with such an activity, over-expression of HipA slows down E. coli growth and
increases antibiotic tolerance and persistence (reviewed in (Lewis, 2010)). Although it is well
established that many stress-responsive toxin-antitoxin systems affect translation in bacteria, the
finding that an active toxin can partially bypass major molecular chaperone requirements in vivo is
novel. An attractive hypothesis is that severe protein aggregation may trigger the activation of
stress-responsive toxin-antitoxin systems in order to control the level of protein synthesis, thus
relieving the overwhelmed downstream network of molecular chaperones until normal conditions
resume. In agreement with this hypothesis, four other well-characterized antitoxins present in E.
72
coli, namely MazE, RelB, MqsA and DinJ, whose respective cognate toxins MazF, RelE MqsR, and
YafQ are known to affect protein synthesis, have been recently shown to be bona fide DnaK
substrates in vivo(Calloni et al, 2012). Clearly, further work is needed to elucidate such a possible
role for chromosomally-encoded toxin-antitoxin systems in bacteria.
EF-Tu degradation
Heat Shock ResponseSynthesis of HSPs
(incl. Lon and Hsp33)
Protein aggregation
Recruitment ofDnaKJE and GroESL
Slowing down of translation (decreased protein load on
chaperones)
STRESS
Adaptation to stress
Fig. 13: Hypothetical model for Hsp33/Lon-mediated degradation of EF-Tu in response to
severe protein aggregation. (1), Denaturing stress induces protein aggregation; (2), DnaKJE and
GroESL are rapidly recruited to protein aggregates; (3), titration of both chaperone machines by
aggregates stabilizes the heat-shock sigma factor σ32
, which subsequently induces the synthesis of
heat-shock proteins (HSPs), including Hsp33 and Lon; (4), Hsp33 and Lon stimulate EF-Tu
degradation, slowing down de novo protein synthesis; (5), the reduced load of newly synthesized
proteins on the downstream chaperones contributes to bacterial survival until normal conditions
resume.
73
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93
REMERCIEMENTS
Je remercie sincèrement les membres du jury d'être venu d'aussi loin afin d’évaluer ce travail.
Je remercie mon directeur de thèse, Pierre ainsi que Marie-Pierre (co-directeur) et Joen (co-
superviseur), pour leurs conseils avisés et leur aide inestimable tout au long de ces années. J’ai
appris énormément de choses à la fois au niveau scientifique et logistique grâce à eux.
Je remercie tous les membres de l’équipe Genevaux, passés et présents: Patricia, Anne-Marie, Elsa,
Ambre, Frédéric, et Sam ainsi que mes deux stagiaires Gaëlle et Thomas. J’y ai rencontré des
personnes formidables qui m’ont apporté un réel soutien scientifique et humain et qui sont devenus
pour moi de très bons amis.
Je remercie les membres de l’équipe de Joen Luirink de l’Université Libre d’Amsterdam: Joen,
Ana, Zora, Sadeeq, Wouter et Gregory de m’avoir accueilli pendant plus de quatre mois afin
d’apprendre et d’appliquer une expérience de traduction in vitro dans des conditions optimales et
pour leur bonne humeur et leur soutien sans failles.
Merci à mes collègues du LMGM avec qui j'ai passé d'excellents moments: Laure, Cécile, Anne,
Brigitte, Véronique en particulier.
Mes travaux ont été financés par une bourse CNRS/Région Midi-Pyrénées. Je remercie donc le
CNRS ainsi que la région Midi-Pyrénées pour leur soutien financier.
Merci à mes parents et à ma petite sœur sans qui je n’aurais jamais pu aller aussi loin dans la vie,
sachant que la route n’est pas encore terminée. Merci de m’avoir laissé accomplir mes rêves.
Merci à Ludo, mon coloc préféré, qui a fait de cette thèse une expérience unique. Nos soirées à
deux vont me manquer.
Et merci à Alexane, un papillon qui s’est posé sur la route de ma vie.
