Thiol-based redox switches evolved as efficient post-translational regulatory mechanisms that enable individual proteins to rapidly respond to sudden environmental changes. While some protein functions need to be switched off to save resources and avoid potentially error-prone processes, protective functions become essential and need to be switched on. In this review, we focus on thiol-based activation mechanisms of stress-sensing chaperones. Upon stress exposure, these chaperones convert into high affinity binding platforms for unfolding proteins and protect cells against the accumulation of potentially toxic protein aggregates. Their chaperone activity is independent of ATP, a feature that becomes especially important under oxidative stress conditions, where cellular ATP levels drop and canonical ATP-dependent chaperones no longer operate. Vice versa, reductive inactivation and substrate release require the restoration of ATP levels, which ensures refolding of client proteins by ATP-dependent foldases. We will give an overview over the different strategies that cells evolved to rapidly increase the pool of ATP-independent chaperones upon oxidative stress and provide mechanistic insights into how stress conditions are used to convert abundant cellular proteins into ATP-independent holding chaperones.
Reactive oxygen species, including superoxide anion and hydrogen peroxide are constantly generated in cells living under aerobic conditions. Effective antioxidant enzymes have evolved to protect the cellular redox homeostasis. Yet, organisms can encounter conditions in which oxidant levels exceed the threshold that can be balanced by cellular detoxification systems, causing a damaging and potentially lethal condition, termed oxidative stress (Imlay 2013; Sies 2015). Reactive oxygen and nitrogen species can modify and damage proteins, lipids, and DNA, thereby challenging the integrity of the entire cell. In proteins, the thiol group of cysteines belongs to the most reactive targets and can undergo different reversible and irreversible modifications. The formation of protein disulfide bonds and mixed disulfides with low-molecular weight (LMW) thiols, like glutathione (GSH), are reversed in most cells by the thioredoxin (Trx) and glutaredoxin (Grx) system. Overoxidation of cysteines to sulfinic or sulfonic acids in contrast causes irreversible damage, leading to protein unfolding and potential aggregation (for recent reviews see Goemans and Collet 2019; Reichmann et al. 2018; Ulrich and Jakob 2019). Oxidants significantly vary in their reactivity with protein thiols and their effects on protein integrity and stability. Hydrogen peroxide, for instance, is a rather slow-acting oxidant—except for proteins with exquisitely peroxide-sensitive thiols—and does not cause substantial protein unfolding or aggregation (Winter et al. 2008; Winterbourn 2008; Winterbourn and Peskin 2016). In comparison, hypochlorous acid, which is produced as a highly potent antimicrobial during mammalian host defense from H2O2 and chloride ions, rapidly modifies a wide range of amino acid side chains, causing widespread protein unfolding and aggregation (Sultana et al. 2020; Winter et al. 2008; Winterbourn et al. 2016).
Excessive levels of H2O2 or HOCl also induce a substantial drop of cellular ATP (Colussi et al. 2000; Kumsta et al. 2011; Winter et al. 2005). The decrease is attributed to the reversible oxidative inactivation of glycolytic enzymes like GAPDH, which results in the redirection of glucose from glycolysis to the pentose phosphate pathway (Hildebrandt et al. 2015; Ralser et al. 2007). Thus, ATP-generating pathways are inactivated in order to produce NADPH, which delivers the electrons for Trx reductase and glutathione reductase to ultimately restore the cellular redox homeostasis (Holmgren et al. 2005). Moreover, cellular ATP is actively converted into long chains of polyphosphate, which not only function as redox inert, high energy storage compounds, but stabilize proteins and have been shown to decrease aggregation in bacteria exposed to oxidative or heat stress (Gray et al. 2014; Yoo et al. 2018).
Depleting cellular ATP levels downregulates all ATP-dependent pathways, including gene expression which reduces the amount of nascent, aggregation-prone polypeptides (Morano et al. 2012; Shenton et al. 2006). Keeping cells in a dormant-like state appears to increase the organisms’ chance of surviving toxic stress conditions. However, decreasing cellular ATP also comes with several downsides. One critical drawback is the inactivation of ATP-dependent canonical chaperones, such as heat shock protein (Hsp) 60, Hsp70 and Hsp90 as well as proteases, like the 26S proteasome (Hartl et al. 2011). How do cells then deal with protein aggregation under oxidative stress conditions? Research over the past two decades revealed that cells respond to oxidative protein unfolding stress by post-translationally activating a distinct set of ATP-independent chaperones, which act as protein holdases that bind unfolding protein intermediates and maintain them in a folding-competent state (Goemans et al. 2018; Jakob et al. 1999; Muller et al. 2014; Reichmann et al. 2012; Wyatt et al. 2014). Once non-stress conditions have been restored, these chaperones get inactivated again and transfer client proteins to ATP-dependent foldases.
Thiol-based redox switches evolved as efficient regulatory mechanisms that allow proteins to immediately respond to changes in the cellular redox state (Leichert and Dick 2015). Since the discovery of bacterial Hsp33 as first redox-regulated chaperone (Jakob et al. 1999), stress-induced activation of ATP-independent molecular chaperones has been shown to protect cellular proteins against irreversible oxidative damage across all kingdoms. In this review, we will focus on thiol-based switches of redox-regulated chaperones, explore the mechanisms by which these proteins sense oxidative stress, and discuss how redox-dependent conformational changes contribute to their function as molecular chaperones.
Oxidative activation of the bacterial chaperone Hsp33
The stress-sensing 33 kDa heat shock protein Hsp33 is chaperone-inactive in its reduced state. Four strictly conserved cysteines arranged in a CXC and CXXC motif coordinate a zinc ion in the C-terminal redox-sensing domain and maintain Hsp33 in a chaperone-inactive conformation (Ilbert et al. 2007; Janda et al. 2004; Vijayalakshmi et al. 2001). Upon exposure of Hsp33 to oxidizing conditions, the four cysteines form two disulfide bonds which cause the release of zinc and induce conformational changes that convert the monomeric chaperone-inactive Hsp33 into highly chaperone-active dimers (Graf et al. 2004; Ilbert et al. 2007; Winter et al. 2008). Importantly, the oxidative activation of Hsp33’s chaperone function is fully reversible. Upon restoring non-stress conditions, Hsp33 dimers are reduced by the Grx and Trx system (Hoffmann et al. 2004). The release of client proteins requires in addition the restoration of cellular ATP levels in order to ensure refolding of clients by the E. coli DnaK (Hsp70) chaperone system (Hoffmann et al. 2004; Reichmann et al. 2012; Winter et al. 2005). Thus, client proteins are only released from Hsp33 when both the cellular redox balance and protein homeostasis have been restored. Recent single molecule studies revealed that activated Hsp33 not only prevents protein aggregation but guides also the folding process by promoting the formation of partially folded conformations in the client proteins (Moayed et al. 2020).
Mechanism of Hsp33 activation
Oxidative disulfide bond formation triggers the conformational change of fully folded to partially disordered, chaperone-active Hsp33 (Bardwell and Jakob 2012; Reichmann et al. 2012). Two intramolecular disulfide bonds are formed between the two cysteines of the CXXC- and CXC-motif, respectively (Figure 1). Since all four cysteines of the tetrahedral zinc center are arranged in an equal distance to each other (Janda et al. 2004; Vijayalakshmi et al. 2001), the question arises how the formation of other disulfide bonds is avoided? Structural and functional analyses revealed that the highly charged, metastable linker region (∼50 aa), which connects the C-terminal redox-sensing domain with the compactly folded, mainly hydrophobic N-terminal domain, plays a crucial role in the oxidative activation of Hsp33. Exposure to H2O2 leads to the formation of only one disulfide bond, connecting the two cysteines of the distal CXXC-motif. Once this disulfide bond has formed, zinc is released and the C-terminal domain of Hsp33 partially unfolds. In this conformation, Hsp33 remains chaperone-inactive, but is primed to become activated upon unfolding conditions. The linker has turned into a thermolabile protein folding sensor that readily unfolds when Hsp33 is exposed to increased temperatures or bile salts (Table 1) (Cremers et al. 2014; Ilbert et al. 2007). Only then, the proximal CXC motif can form the second disulfide bond, which locks the linker region in an unfolded state and constitutively activates Hsp33 as a molecular chaperone (Figure 1) (Cremers et al. 2010; Graf et al. 2004). Kinetically fast oxidants, like hypochlorous acid which cause widespread protein unfolding, induce the formation of both disulfide bonds and activate Hsp33’s chaperone function within seconds at any temperature (Winter et al. 2008). Hsp33 from Chlamydomonas reinhardtii, which lacks the CXC-motif and thus only loosely coordinates zinc, has lost its redox-regulated activation mechanism and can be activated by elevated temperatures even in its reduced form (Segal and Shapira 2015).
|Protein family||Protein||Subcellular localization||Function under non-stress conditions||Activating conditions in vitro||Thiol-based switching mechanism||Chaperone-active conformation||Phenotypes linked to ATP-independent chaperone function||References|
|Hsp33||E. coli Hsp33||Cytoplasm||Unfoldase, potential role in proteostasis||H2O2 + Temp >43 °C, H2O2 + bile salt, HOCl||Disulfide bond formation, zinc release||Partially unfolded, non-covalent dimers||Resistance towards oxidative stress and heat, bile salts or bleach||Cremers et al. 2014; Ilbert et al. 2007; Jo et al. 2019; Rimon et al. 2017; Winter et al. 2008|
|Ruc||M. tuberculosis Rv0991c||Cytoplasm||–||H2O2 and Cu2+||Disulfide bond formation||–||–||Becker et al. 2020|
|Get3/TRC40||Yeast Get3||Cytosol||ATP-dependent TA protein targeting factor||H2O2 and Cu2+||Disulfide bond formation, zinc release||Partially unfolded, tetramers and oligomerics||Resistance towards oxidative stress, heat and metal stress||Powis et al. 2012; Shen et al. 2003; Voth et al. 2014|
|2-Cys Prx||Yeast Tsa1 and 2 (Prx1 and 2)||Cytosol||Peroxidase||H2O2 and/or Temp >43 °C||Hyperoxidation of the peroxidatic Cys||Non-covalent HMW complexes||Resistance towards heat stress, growing under zinc depletion||Jang et al. 2004 Cell; MacDiarmid et al. 2013|
|Human Prx2||Resistance towards H2O2 stress in HeLa cells||Moon et al. 2005|
|A. thaliana Prx||Chloroplast stroma||H2O2, CHP||Decamer and various oligomeric forms||–||Koenig et al. 2013|
|H. pylori AhpC||Cytoplasm||–||Non-covalent HMW complexes||–||Chuang et al. 2006|
|S. mansoni Prx1||Cytosol||H2O2 and/or pH 4.2||Two stacked decamers||–||Saccoccia et al. 2012|
|Anabaena Alr4641||Cytoplasm||H2O2||–||Non-covalent HMW complexes||–||Banerjee et al. 2015|
|P. aeruginosa Prx||Cytoplasm||H2O2||–||HMW complexes||–||An et al. 2010|
|L. infantum mTXNPx||Mitochondrial matrix||Temp >40 °C||Disulfide bond formation (inactivation)||Decamer||Heat tolerance, crucial for parasite infectivity||Castro et al. 2011; Morais et al. 2017; Teixeira et al. 2015|
|Trx||T. brucei Trx2||Mitochondrial matrix||Potential role in mitochondrial protein import||Temp >40 °C||Disulfide bond formation (inactivation)||Monomer||Heat tolerance, increased parasite infectivity||Currier et al. 2019|
|Hsp70||Yeast BiP||ER||ATP-dependent foldase||GSSG, GSH + diamide or HOCl or CHP||Sulfenylation and S-glutathionylation of the single Cys||Monomer||–||Wang et al. 2014; Wang et al. 2016|
|Human BiP (Grp78)||ER||H2O2||Disulfide bond formation||Resistance towards oxidative ER stress||Wei et al. 2012|
|Mammalian Hsc70||Cytosol||GSSG, Temp >43 °C||S-glutathionylation||–||Hoppe et al. 2004|
AhpC, alkylhydroperoxide reductase; BiP, Binding immunoglobulin protein; CHP, cumene hydroperoxide; GET, Guided entry of tail-anchored proteins; Hsc, heat shock cognate protein; Hsp, heat shock protein; Prx, peroxiredoxin; TA, tail-anchored; Trx, thioredoxin.
The C-terminal redox-sensitive domain of Hsp33 mediates the stress-dependent switch from an inactivate to an active chaperone, whereas the highly charged linker region functions as a gatekeeper for Hsp33’s oxidative activation. Reversing the order of the amino acids in the linker region or replacing it by a non-native sequence of similar length results in a constitutively active chaperone (Rimon et al. 2017). All mutant variants showed an increased sensitivity towards oxidation, suggesting that the linker region affects the reactivity of at least some of the cysteines involved. Replacement of the native linker region also affected the transfer of substrates to the DnaK/J system, emphasizing the function of the polar linker domain in client binding and release (Groitl et al. 2016; Rimon et al. 2017).
Hsp33 function in bacteria
Bacteria lacking Hsp33 are significantly more sensitive towards oxidative stress conditions that lead to protein unfolding (Table 1) (Cremers et al. 2014; Jakob et al. 1999; Sultana et al. 2020; Winter et al. 2008). The protective function of Hsp33 has been shown to serve as a first line of defense in E. coli upon oxidative stress and might explain the high abundancy of Hsp33 even under basal conditions (Dahl et al. 2015). Little, however, is known about the in vivo role of Hsp33 under non-stress conditions. Rimon et al. (2017) recently proposed that reduced Hsp33 might serve as a house-keeping factor preserving a “healthy proteome” since Hsp33 interacts with members of the proteostasis network under normal conditions in a linker-independent manner (Reichmann et al. 2018; Rimon et al. 2017). A similar role was suggested by an earlier study showing Hsp33-dependent sequestration of the elongation factor Tu (EF-Tu) for degradation by Lon protease in order to slow down de novo protein biosynthesis in strains lacking the canonical chaperones DnaK and trigger factor (Bruel et al. 2012). Although in vitro studies confirmed that reduced, chaperone-inactive Hsp33 binds to EF-Tu, induces EF-Tu aggregation and thus susceptibility to proteolytic degradation (Jo et al. 2019), in vivo evidence for the specific role of the reduced form of Hsp33 in preserving protein homeostasis under physiological conditions still needs to be provided.
Mycobacterium tuberculosis lacks any Hsp33 homologs. However, very recently the protein Rv0991c has been identified to function analogously to Hsp33 (Becker et al. 2020). The redox-regulated protein with unstructured C-terminus (Ruc) acts as a molecular chaperone upon oxidation, and enables refolding of client proteins by the M. tuberculosis Hsp70 system. Ruc is required for full virulence of M. tuberculosis in mice. Ruc homologs are found throughout Actinobacteria which, except for a single species, lack Hsp33. Thus, it might be reasonably assumed that Ruc fulfills the role of Hsp33 in Actinobacteria in order to prevent irreversible aggregation of unfolding protein intermediates during oxidative stress (Becker et al. 2020).
Dual function proteins with thiol-dependent chaperone activity
Get3: tail-anchored protein targeting factor and molecular chaperone
Hsp33 is highly conserved in bacteria but absent in eukaryotes except for some unicellular flagellates like Chlamydomonas and members of the Trypanosomatidae family. In yeast, Get3 has been identified to function as a Hsp33-like, redox-regulated chaperone activated under ATP-depleting oxidative stress conditions (Powis et al. 2013; Voth et al. 2014). Although structurally unrelated, Get3 shares several features of Hsp33, including the conserved CXC- and CXXC-motif and the ability to coordinate zinc. However, reduced Get3 forms non-covalent homodimers and the cysteines of the respective CXXC-motifs coordinating the zinc ion in the dimer-dimer interface (Stefer et al. 2011) (Figure 2). Like for Hsp33, oxidative disulfide bond formation results in the release of zinc accompanied by partial unfolding of Get3 (Voth et al. 2014). The conformational changes induce the formation of chaperone-active, higher oligomeric species, which bind unfolding protein intermediates and prevent their aggregation. The activation of Get3’s chaperone function is fully reversible and depends also on the restoration of ATP levels (Voth et al. 2014).
In contrast to Hsp33, Get3 has a well-known function under physiological conditions (Table 1). Reduced Get3 acts as an ATPase in the “guided entry of tail-anchored (TA) proteins” (GET) pathway, where it mediates the post-translational insertion of TA proteins into the membrane of the endoplasmic reticulum (ER) (Mateja et al. 2009; Schuldiner et al. 2008; Stefanovic and Hegde 2007). Get3 receives TA proteins from the cochaperone Sgt2, which is bound to the pre-targeting complex (Get4/5) (Figure 2) (for a recent review see Borgese et al. 2019). Sgt2 takes over proteins from cytosolic Hsp70 (Ssa1), which efficiently binds nascent TA proteins. This TA protein relay appears to ensure the gradual loading of Get3 while maintaining the solubility and targeting competence of TA proteins (Cho and Shan 2018). A flexible α-helix lining the substrate-binding groove of Get3 facilitates substrate loading and has been proposed to function as a lid that prevents the interaction of TA proteins with other chaperones (Chio et al. 2019; Mateja et al. 2015). Cargo-loaded Get3 dissociates from the pre-targeting complex and shuttles TA proteins to the GET receptor complex (Get1/2) at the ER (Borgese et al. 2019). ATP hydrolysis seems to be required for Get3 to interact with the receptors. The interaction with Get1 induces the release of ADP and the insertion of the TA proteins into the ER membrane. ATP binding and the interaction with Get4/5 mediate the dissociation of Get3 from the ER and allows Get3 to enter another cycle (Borgese et al. 2019; Hu et al. 2009; Mateja et al., 2015, 2009).
Mechanism of Get3’s chaperone activation
Exposure of Get3 to Cu2+, H2O2 or H2O2/Cu2+ results in oxidative disulfide bond formation and the release of the zinc ion (Voth et al. 2014). Get3 undergoes major conformational changes leading to its partial unfolding and the exposure of hydrophobic surfaces, which allow binding of unfolding protein intermediates. Size exclusion chromatography connected to multi-angle light scattering (SEC-MALS) identified homo-tetramers as the minimal chaperone-active form and subsequent transmission electron microscopy provided a low-resolution structure of the oxidized Get3 tetramer. Importantly, as Get3’s chaperone function is activated, its ATPase activity is switched off. Moreover, the TA binding site appears to be buried, thereby preventing the interaction with TA proteins. These structural changes are fully reversible upon Get3 reduction in the presence of Zn2+ and ATP (Voth et al. 2014). The precise mode of activation and the kinetics of disulfide bond formation remain under investigation. Disulfide mapping revealed the formation of two disulfide bonds in activated Get3, connecting the next adjacent cysteines of the CXC and CXXC motif, respectively. Whereas in vitro studies also revealed the formation of inter-subunit disulfide bonds, only intramolecular disulfide bonds were observed in vivo (Voth et al. 2014).
Get3 function in yeast
Get3 and any of the other components of the GET pathway can be deleted in yeast without causing any obvious growth defects under non-stress conditions. This finding is in stark contrast to the fact that some TA proteins like Sed5 are essential in yeast and suggests that alternative targeting pathways for TA proteins must exist (Powis et al. 2013; Schuldiner et al. 2008). In comparison to non-stress conditions, deletion of get3 leads to growth defects under a number of different stress conditions, including copper and heat stress (Metz et al. 2006; Powis et al. 2013; Shen et al. 2003; Voth et al. 2014). The observation that a Get3 mutant variant, which is deficient in TA protein targeting rescues the copper-sensitive phenotype of a get3 deletion strain (Mateja et al. 2009) was an early indication for another, targeting-independent role of Get3. Since the targeting-deficient mutant was found to have a wild type-like, redox-regulated chaperone function in vitro, we concluded that the copper sensitive phenotype of a get3 deletion strain is likely caused by loss of the Get3 chaperone activity and not by impaired TA protein targeting (Voth et al. 2014). A recent phylogenetic analysis suggested that handling hydrophobic substrates is a fundamental and likely the more ancient property of Get3, irrespective of its role in TA protein targeting. The chaperone function appears to be a common feature among Get3 homologues across phyla and has been later on adapted to serve in TA protein targeting in eukaryotes (Farkas et al. 2019).
TRC40 function in higher eukaryotes
TRC40 is the mammalian homolog of Get3, which post-translationally binds to hydrophobic C-terminal domains of TA proteins and shuttles them to the GET receptors CAML and WRD in the ER membrane (Schuldiner et al. 2008; Stefanovic and Hegde 2007). The knockout of TRC40 is embryonically lethal in mice (Mukhopadhyay et al. 2006). Yet, the conditional, tissue-specific deletion of the WRB receptor subunit affected only a small number of TA proteins (Rivera-Monroy et al. 2016) which is in line with the notion that losing TRC40 may affect the cell independently from TA targeting. Studies in mammalian cell culture systems revealed that even in the absence of a functional TRC40 pathway, most known TRC40 clients are properly targeted to membranes (Coy-Vergara et al. 2019). The finding agrees with previous in vivo studies showing that TA protein targeting is facilitated by several pathways, including the signal recognition particle (SRP) system (Abell et al. 2004; Casson et al. 2017) and the SRP-independent targeting (SND) pathway (Aviram et al. 2016; Hassdenteufel et al. 2017). Therefore, the TRC40 pathway is not essential for TA protein targeting. Considering the function of Get3 as a redox-sensitive molecular chaperone in yeast, it is tempting to speculate that TRC40 also serves as stress-specific chaperone in mammalian cells. Indeed, conditional TRC40 knockout studies in mice suggested that the dependence of syntaxin 5 on TRC40 is not conferred by its transmembrane domain but depends on interactions of TRC40 with its N-terminal cytosolic domain (Rivera-Monroy et al. 2016). Future studies will investigate whether TRC40 indeed functions as a molecular chaperone like its yeast homolog. These studies are, however, not trivial, particularly since purification of TRC40 is very challenging. Nonetheless, obtaining detailed mechanistic knowledge about the two functions of TRC40 is necessary in order to dissect the roles they play in vivo.
2-Cys peroxiredoxins: peroxidase and molecular chaperone
Proteins containing a thioredoxin (Trx)-fold motif are particularly sensitive to changes in the redox milieu and several members of the Trx superfamily have been shown to function as redox-regulated chaperones, such as E. coli Trx reductase and Trx (Kern et al. 2003). Human protein disulfide isomerase (PDI) also operates as oxidoreductase and molecular chaperone in the ER. Thiol-disulfide exchange reactions promote the switching between a “closed” (reduced) and an “open” (oxidized) conformation, which facilitates client binding and release (Karamzadeh et al. 2017; Wang et al. 2012a). Among Trx-fold proteins, peroxiredoxins (Prxs) represent a large and highly conserved family of thiol-based peroxidases showing an exquisitely high reactivity towards H2O2 with catalytic rates of 105–108 M−1s−1 (Chae et al. 1994; Winterbourn and Peskin 2016). Within this family, 2-Cys Prxs belong to the most abundant cellular proteins, which primarily function as efficient scavenger of low levels of H2O2 and catalyze its reduction to water (Nelson et al. 2011). Given their high reactivity with H2O2, 2-Cys Prxs have been shown to act in addition as receptor and transducer of the H2O2 signal (Sobotta et al. 2015; Stocker et al. 2018). In the nucleus, binding of Prx1 to telomeric DNA has been shown to regulate telomerase activity (Aeby et al. 2016; Lu et al. 2013) and Prx-dependent regulations play a crucial role in disease development such as cancer (Forshaw et al. 2019). Under specific stress conditions, 2-Cys-Prxs from various organisms have been shown to become peroxidase-inactive and function as potent ATP-independent chaperone (Table 1) (Jang et al. 2004; Konig et al. 2013; Moon et al. 2005; Rhee and Woo 2020; Toledano and Huang 2016).
2-Cys Prxs form non-covalent head-to-tail dimers (Figure 3). The N-terminal peroxidatic cysteine of one subunit reduces H2O2 under the release of water and formation of a cysteine sulfenic acid. The sulfenic acid intermediate condenses with the C-terminal resolving cysteine of the second subunit, thereby forming an inter-subunit disulfide under the release of a second H2O molecule (Figure 3). The disulfide is subsequently reduced by Trx, which completes the peroxidatic cycle. Eukaryotic 2-Cys Prxs have been shown to be highly sensitive towards oxidative inactivation at elevated H2O2 levels. Oxidation of the cysteine sulfenic acid to a sulfinic acid by another H2O2 molecule results in the loss of peroxidase activity (Wood et al. 2003; Yang et al. 2002). In contrast to most cysteine residues, sulfinylation of the peroxidatic cysteine is reversible. Sulfiredoxin (Srx) catalyzes the ATP-dependent reduction and allows Prx to reenter the peroxidatic cycle (Biteau et al. 2003; Woo et al. 2003). Further oxidation of the sulfinic acid to a sulfonic acid is, however, irreversible.
Oxidation and reduction of typical 2-Cys Prxs are accompanied by large conformational rearrangements (Figure 3). Reduced 2-Cys Prxs favor the formation of (do)decameric, ring-like structures, which dissociate into dimers upon disulfide formation. For Prx1 and Prx2, it has been shown that hyperoxidation induces the formation of high molecular weight (HMW) multi-stacked rings which confer ATP-independent chaperone activity and prevent protein aggregation during oxidative stress (Jang et al. 2004; Moon et al. 2005; Noichri et al. 2015; Wood et al. 2002). Although in Prx1 from Saccharomyces cerevisiae (Tsa1) some chaperone activity can be thiol-independently induced upon heat shock (Jang et al. 2004), the peroxidase-to-chaperone switch is primarily guided by hyperoxidation of the peroxidatic cysteine. A similar functional switch has been described for several other 2-Cys Prxs in different organisms and subcellular localizations as summarized in Table 1.
Mechanism of Prx chaperone activation
In human Prx2, hyperoxidation of the peroxidatic cysteine and switching from a peroxidase to a molecular chaperone has been shown to require the presence of a YF (Tyr-Phe) motif in the C-terminal domain (Moon et al. 2005). In a C-terminally truncated Prx2 mutant, H2O2 exposure failed to trigger the functional and conformational switch (Moon et al. 2005). It appears that the peroxidatic cysteine acts as efficient H2O2 sensor while the YF motif promotes the stress-induced structural rearrangements. Comparing 3D crystal structures of the chaperone-inactive LMW decamer and the chaperone-active HMW double-decamer of Schistosoma mansoni Prx1 provided further insights into the mechanism (Saccoccia et al. 2012). Hyperoxidation of the peroxidatic cysteine which is located at the beginning of the α2 helix results in unwinding of the first turn of the helix concerted by unfolding of the C-terminal tail. Unwinding of the first turn of helix α2 has been previously demonstrated for human Prx1 bound to Srx (Jonsson et al. 2008) and represents most likely a general structural change upon Prx hyperoxidation. Unfolding of the C-terminus changes the surface of the decameric ring and allows interacting with another Prx decamer to form a double decamer which is chaperone-active and binds unfolded proteins (Saccoccia et al. 2012).
The formation of hyperoxidized HMW complexes is reversible in the presence of Srx, which catalyzes the ATP-dependent reduction of the sulfinylated peroxidatic cysteine and restores the peroxidase-active conformation (Jang et al. 2004; Moon et al. 2013). Structural analyses showed that the interaction of hyperoxidized Prx with Srx also relies on the unfolding of its C-terminus (Jonsson et al. 2008; Saccoccia et al. 2012). In some organisms, like H. pylori and trypanosomadis, the mechanism for reducing hyperoxidized 2-Cys Prxs remains to be uncovered since these cells lack known Srx homologs (Toledano and Huang 2016).
In contrast to most known Prxs, the Leishmania infantum mitochondrial 2-Cys Prx (mTXNPx) works as an effective chaperone only in its fully reduced form (Teixeira et al. 2015). Activation of the chaperone function is induced by temperature-mediated restructuring of the decamer, which exposes a previously buried hydrophobic patch and allows binding of unfolded protein clients in the center of the decametric ring (Teixeira et al. 2015; Teixeira et al. 2019). Upon restoration of non-stress conditions, client proteins are released and transferred to ATP-dependent chaperones for refolding. Replacing the oxidation-sensitive peroxidatic cysteine by a serine residue maintains the protein in a decameric and/or higher oligomeric conformation even under non-reducing conditions and results in a constitutively chaperone-active mutant variant (Teixeira et al. 2015). Vice versa, mutant variants that are unable to form decamers no longer protect parasite proteins against temperature-induced aggregation (Morais et al. 2017). L. infantum mTXNPx is essential for host virulence and thermotolerance in the insect form of the parasite (Castro et al. 2011; Teixeira et al. 2015). The fact that these phenotypes can be rescued by a mutant variant that lacks the peroxidatic cysteine indicates that not the peroxidase function is essential under these conditions, but the ability of mTXNPx to protect clients from temperature-induced aggregation.
We recently described a similar switch for mitochondrial Trx2 in Trypanosoma brucei (Table 1). Like Leishmania mTXPNx, the chaperone function of reduced Trx2 is activated by temperature-mediated structural changes and protects other proteins against thermal aggregation by maintaining their folding-competent state. Oxidative disulfide bond formation inhibits activation of the chaperone function. Replacing all five cysteines by serine residues generates a constitutively chaperone-active mutant variant, which complements for the essential function of the wild type protein and confers parasite infectivity in the mouse model (Currier et al. 2019). Both examples show that structural remodeling appears to be necessary for chaperone activation, which can be induced by elevated temperatures as well as possibly by other protein-unfolding stresses (Teixeira et al. 2015). What remains to be investigated is whether this mode of activation only applies to mitochondrial proteins or extends also to cytosolic members of the Trx family.
Thiol-based switching of ATP-dependent foldases to ATP-independent chaperones
Hsp70 proteins are well known for their crucial roles in maintaining cellular proteostasis (Hartl et al. 2011; Rosenzweig et al. 2019). In addition to their ATP-dependent function as foldases, several members of this family have been demonstrated to act as ATP-independent holding chaperones in response to oxidative stress. In yeast, the ER-localized Hsp70 chaperone BiP is sensitive towards changes in the redox milieu of the ER. The single cysteine of BiP becomes oxidized to a sulfenic acid as peroxide levels rise in the ER (Wang et al. 2014). This intermediate can further react with GSH under the formation of a mixed disulfide (Wang and Sevier 2016). Treating recombinant BiP with glutathione disulfide (GSSG) also resulted in the S-glutathionylation, a mechanism that might be of physiological relevance considering the much lower GSH:GSSG ratio in the ER compared to the cytosol (Birk et al. 2013; Schwarzlander et al. 2016). Both, sulfenylation and S-glutathionylation result in inactivation of the ATPase activity and activation of the chaperone holding function in vitro (Wang et al. 2014; Wang and Sevier 2016). The functional switch is fully reversible and the BiP nucleotide exchange factor (NEF) Sil1 has been recently identified to catalyze the reduction of BiP, which restores its ATPase activity and reverses the enhanced holdase activity (Siegenthaler et al. 2017). A Sil1 mutant that lacks its catalytic cysteines is unable to reverse BiP oxidation and restore its ATPase activity. Interestingly, upon reduction, the ATPase activity is further stimulated by Sil1 acting as NEF. Lhs1, another ER-resident NEF of BiP, was unable to restore ATPase activity of oxidized BiP, presumably because Lhs1 does not as a reductant. However, in the presence of both, Sil1 and Lhs1, an enhanced steady-state ATPase activity of BiP was measured. This effect is likely the result of the Sil1-mediated reduction of BiP followed by an enhanced rate of ATP-turnover promoted by Lhs1 and might explain the advantage of having both NEFs in the ER (Siegenthaler et al. 2017).
The functional switch of BiP appears to not only help preventing protein aggregation, but avoids also ATP-consuming and potentially futile cycles of substrate binding and release under oxidative stress conditions. A similar mechanism has been proposed for the mammalian homolog Grp78, where disulfide bond formation was found to increase its ATP-independent chaperone activity (Table 1) (Wei et al. 2012). Interestingly, the ER-localized non-selenocysteine containing glutathione peroxidase NPGPx has been shown to transfer H2O2-derived oxidative equivalents to Grp78 via the formation of a transient intermolecular disulfide bond.
The redox regulation of Hsp70’s chaperone function appears not to be restricted to ER-resident homologs. Mammalian cytosolic heat shock cognate protein Hsc70 is also susceptible to protein S-glutathionylation, which induces an ATP-independent chaperone function and can be reversed by Grx1 (Hoppe et al. 2004). For other cytosolic members of the Hsp70 family, like bacterial DnaK and yeast Ssa1, S-glutathionylation and disulfide bond formation, respectively, have been shown to impair ATP-dependent chaperone function, causing the release from their respective heat shock factors and initiation of the heat shock response (Wang et al. 2012b; Winter et al. 2005; Zhang et al. 2016). The finding emphasizes once more that thiol switches not only act to initiate a specific function, but can also serve to inactivate functions that might be futile or even harmful under stress conditions. In this respect, it is also of particular interest to unravel the role of interaction partners such as J-proteins, which potentially play a crucial role in regulating the functional switch.
The stress-specific activation of ATP-independent chaperones serves as first line of defense to protect cells from the formation and accumulation of potentially toxic protein aggregates under severe stress conditions. As high-affinity binding platforms for unfolding proteins, these chaperones do not require energy, which is particularly important under oxidative stress conditions, where cellular ATP levels drop (Colussi et al. 2000; Winter et al. 2005). In turn, reductive inactivation generally relies on the restoration of ATP levels to ensure that client proteins are only released when ATP-dependent foldases are again functional. It remains to be seen which additional cellular factors contribute to the (in)activation of respective chaperones. In the case of Get3/TRC40, the interaction with the GET receptors triggers nucleotide release, whereas ATP-binding of Get3 is necessary to interact with Get4/5 in order to enter another cycle of TA protein targeting (Figure 2) (Borgese et al. 2019). Since the inactivation of Get3’s chaperone function depends in vitro on the presence of ATP (Voth et al. 2014), it is tempting to speculate that in the cell, GET pathway components contribute to fine-tuning Get3’s functional switch.
Many stress-inducible chaperones are dual function proteins. Whereas in some cases the functions are mutually exclusive and proteins have to lose their non-stress function to turn into chaperones, it is possible that the chaperone function of some stress-inducible proteins also plays a role for the interaction with specific proteins under physiological conditions. However, investigating those aspects is not trivial and reveals one major caveat when studying dual function proteins. In order to assign in vivo phenotypes to a specific protein function, one has to find a way to manipulate one of the activities without affecting the other. Therefore, structural features need to be identified first that allow designing mutations which target only one of the two functions.
The fact that stress-sensing chaperones often are dual function proteins also suggests that more cellular proteins might have the ability to switch to effective chaperones under stress conditions. Systemic approaches are required to globally identify redox-regulated chaperones and investigate their functions under both reducing and oxidizing conditions. The Reichmann lab recently presented a toolbox of diverse techniques for studying redox-regulated chaperone activity, including guidelines for preparing fully reduced and oxidized proteins and mapping of conformational changes via hydrogen-deuterium exchange mass spectrometry (Fassler et al. 2018). In addition to thiol-based switching mechanisms discussed here, more and more thiol-independent mechanisms have been identified to convert proteins into active chaperones during stress as recently outlined in Goemans and Collet (2019), Reichmann et al. (2018) and Sultana et al. (2020).
Equally intriguing is the potential role of redox-regulated chaperones in signaling pathways. It is well-established that oxidants like nitric oxide, peroxynitrite and H2O2 act as second messengers and oxidative thiol modifications serve to transduce molecular signals (D’Autreaux and Toledano 2007; Forman et al. 2014; Sies 2017). 2-Cys Prxs have been shown to transduce H2O2-derived oxidizing equivalents to specific target proteins (Bersweiler et al. 2017; Rhee et al. 2018; Sobotta et al. 2015; Stocker et al. 2018). It will be interesting to see in how far oxidation-induced conformational changes and the ability of 2-Cys Prx to form high-affinity binding platforms affect interactions with specific target proteins and thus, potentially promote the redox relay.
Thiol-based switching mechanisms allow the immediate activation of stress-sensitive chaperones in response to cellular changes. Given the effort of organisms to maintain a healthy proteome, the role of stress-inducible chaperones in protecting cells during disease- or aging-related changes of the redox homeostasis will be of overall interest. With our increasing understanding of tissue-specific redox protein networks (Xiao et al. 2020) that underlie distinct physiologic and metabolic changes, it will be exciting to learn more about tissue-specific functions of redox-regulated chaperones and their role in redox-modified disease signatures.
Funding source: National Institute of Health (NIH)
Award Identifier / Grant number: GM122506
Funding source: Deutsche Forschungsgemeinschaft
Award Identifier / Grant number: SPP 1710, Schw823
This work was supported by grants of the DFG Priority Program SPP 1710 (Schw823) and the NIH (GM122506).
Author contribution: All the authors have accepted responsibility for the entire content of this submitted manuscript and approved submission.
Research funding: This work was supported by grants of the DFG Priority Program SPP 1710 (Schw823) and the NIH (GM122506).
Conflict of interest statement: The authors declare no conflicts of interest regarding this article.
Abell, B.M., Pool, M.R., Schlenker, O., Sinning, I., and High, S. (2004). Signal recognition particle mediates post-translational targeting in eukaryotes. EMBO J. 23: 2755–2764, https://doi.org/10.1038/sj.emboj.7600281. Search in Google Scholar
Aeby, E., Ahmed, W., Redon, S., Simanis, V., and Lingner, J. (2016). Peroxiredoxin 1 protects telomeres from oxidative damage and preserves telomeric DNA for extension by telomerase. Cell Rep. 17: 3107–3114, https://doi.org/10.1016/j.celrep.2016.11.071. Search in Google Scholar
Aviram, N., Ast, T., Costa, E.A., Arakel, E.C., Chuartzman, S.G., Jan, C.H., Hassdenteufel, S., Dudek, J., Jung, M., Schorr, S., et al. (2016). The SND proteins constitute an alternative targeting route to the endoplasmic reticulum. Nature 540: 134–138, https://doi.org/10.1038/nature20169. Search in Google Scholar
Becker, S.H., Ulrich, K., Dhabaria, A., Ueberheide, B., Beavers, W., Skaar, E.P., Iyer, L.M., Aravind, L., Jakob, U., and Darwin, K.H. (2020). Mycobacterium tuberculosis Rv0991c is a redox-regulated molecular chaperone. mBio 11, https://doi.org/10.1128/mbio.01545-20. Search in Google Scholar
Bersweiler, A., D’Autreaux, B., Mazon, H., Kriznik, A., Belli, G., Delaunay-Moisan, A., Toledano, M.B., and Rahuel-Clermont, S. (2017). A scaffold protein that chaperones a cysteine-sulfenic acid in H2O2 signaling. Nat. Chem. Biol. 13: 909–915, https://doi.org/10.1038/nchembio.2412. Search in Google Scholar
Birk, J., Meyer, M., Aller, I., Hansen, H.G., Odermatt, A., Dick, T.P., Meyer, A.J., and Appenzeller-Herzog, C. (2013). Endoplasmic reticulum: reduced and oxidized glutathione revisited. J. Cell Sci. 126: 1604–1617, https://doi.org/10.1242/jcs.117218. Search in Google Scholar
Biteau, B., Labarre, J., and Toledano, M.B. (2003). ATP-dependent reduction of cysteine-sulphinic acid by S. cerevisiae sulphiredoxin. Nature 425: 980–984, https://doi.org/10.1038/nature02075. Search in Google Scholar
Borgese, N., Coy-Vergara, J., Colombo, S.F., and Schwappach, B. (2019). The ways of tails: the GET pathway and more. Protein J. 38: 289–305, https://doi.org/10.1007/s10930-019-09845-4. Search in Google Scholar
Bruel, N., Castanie-Cornet, M.P., Cirinesi, A.M., Koningstein, G., Georgopoulos, C., Luirink, J., and Genevaux, P. (2012). Hsp33 controls elongation factor-Tu stability and allows Escherichia coli growth in the absence of the major DnaK and trigger factor chaperones. J. Biol. Chem. 287: 44435–44446, https://doi.org/10.1074/jbc.m112.418525. Search in Google Scholar
Casson, J., McKenna, M., Hassdenteufel, S., Aviram, N., Zimmerman, R., and High, S. (2017). Multiple pathways facilitate the biogenesis of mammalian tail-anchored proteins. J. Cell Sci. 130: 3851–3861, https://doi.org/10.1242/jcs.207829. Search in Google Scholar
Castro, H., Teixeira, F., Romao, S., Santos, M., Cruz, T., Florido, M., Appelberg, R., Oliveira, P., Ferreira-da-Silva, F., and Tomas, A.M. (2011). Leishmania mitochondrial peroxiredoxin plays a crucial peroxidase-unrelated role during infection: insight into its novel chaperone activity. PLoS Pathog. 7: e1002325, https://doi.org/10.1371/journal.ppat.1002325. Search in Google Scholar
Chae, H.Z., Chung, S.J., and Rhee, S.G. (1994). Thioredoxin-dependent peroxide reductase from yeast. J. Biol. Chem. 269: 27670–27678. Search in Google Scholar
Chio, U.S., Chung, S., Weiss, S., and Shan, S.O. (2019). A chaperone lid ensures efficient and privileged client transfer during tail-anchored protein targeting. Cell Rep. 26: 37–44 e37, https://doi.org/10.1016/j.celrep.2018.12.035. Search in Google Scholar
Cho, H. and Shan, S.O. (2018). Substrate relay in an Hsp70-cochaperone cascade safeguards tail-anchored membrane protein targeting. EMBO J. 37, https://doi.org/10.15252/embj.201899264. Search in Google Scholar
Colussi, C., Albertini, M.C., Coppola, S., Rovidati, S., Galli, F., and Ghibelli, L. (2000). H2O2-induced block of glycolysis as an active ADP-ribosylation reaction protecting cells from apoptosis. Faseb. J. 14: 2266–2276, https://doi.org/10.1096/fj.00-0074com. Search in Google Scholar
Coy-Vergara, J., Rivera-Monroy, J., Urlaub, H., Lenz, C., and Schwappach, B. (2019). A trap mutant reveals the physiological client spectrum of TRC40. J. Cell Sci. 132, https://doi.org/10.1242/jcs.230094. Search in Google Scholar
Cremers, C.M., Knoefler, D., Vitvitsky, V., Banerjee, R., and Jakob, U. (2014). Bile salts act as effective protein-unfolding agents and instigators of disulfide stress in vivo. Proc. Natl. Acad. Sci. U.S.A. 111: E1610–1619, https://doi.org/10.1073/pnas.1401941111. Search in Google Scholar
Cremers, C.M., Reichmann, D., Hausmann, J., Ilbert, M., and Jakob, U. (2010). Unfolding of metastable linker region is at the core of Hsp33 activation as a redox-regulated chaperone. J. Biol. Chem. 285: 11243–11251, https://doi.org/10.1074/jbc.m109.084350. Search in Google Scholar
Currier, R.B., Ulrich, K., Leroux, A.E., Dirdjaja, N., Deambrosi, M., Bonilla, M., Ahmed, Y.L., Adrian, L., Antelmann, H., Jakob, U., et al. (2019). An essential thioredoxin-type protein of Trypanosoma brucei acts as redox-regulated mitochondrial chaperone. PLoS Pathog. 15: e1008065, https://doi.org/10.1371/journal.ppat.1008065. Search in Google Scholar
D’Autreaux, B. and Toledano, M.B. (2007). ROS as signalling molecules: mechanisms that generate specificity in ROS homeostasis. Nat. Rev. Mol. Cell Biol. 8: 813–824, https://doi.org/10.1038/nrm2256. Search in Google Scholar
Dahl, J.U., Gray, M.J., and Jakob, U. (2015). Protein quality control under oxidative stress conditions. J. Mol. Biol. 427: 1549–1563, https://doi.org/10.1016/j.jmb.2015.02.014. Search in Google Scholar
Fassler, R., Edinger, N., Rimon, O., and Reichmann, D. (2018). Defining Hsp33’s redox-regulated chaperone activity and mapping conformational changes on Hsp33 using hydrogen-deuterium exchange mass spectrometry. J. Vis. Exp. 136: 57806, https://doi.org/10.3791/57806. Search in Google Scholar
Forshaw, T.E., Holmila, R., Nelson, K.J., Lewis, J.E., Kemp, M.L., Tsang, A.W., Poole, L.B., Lowther, W.T., and Furdui, C.M. (2019). Peroxiredoxins in cancer and response to radiation therapies. Antioxidants 8, https://doi.org/10.3390/antiox8010011. Search in Google Scholar
Goemans, C.V., Vertommen, D., Agrebi, R., and Collet, J.F. (2018). CnoX is a chaperedoxin: a holdase that protects its substrates from irreversible oxidation. Mol Cell 70: 614–627, https://doi.org/10.1016/j.molcel.2018.04.002, e617. Search in Google Scholar
Graf, P.C., Martinez-Yamout, M., VanHaerents, S., Lilie, H., Dyson, H.J., and Jakob, U. (2004). Activation of the redox-regulated chaperone Hsp33 by domain unfolding. J. Biol. Chem. 279: 20529–20538, https://doi.org/10.1074/jbc.m401764200. Search in Google Scholar
Gray, M.J., Wholey, W.Y., Wagner, N.O., Cremers, C.M., Mueller-Schickert, A., Hock, N.T., Krieger, A.G., Smith, E.M., Bender, R.A., Bardwell, J.C., et al. (2014). Polyphosphate is a primordial chaperone. Mol Cell 53: 689–699, https://doi.org/10.1016/j.molcel.2014.01.012. Search in Google Scholar
Groitl, B., Horowitz, S., Makepeace, K.A.T., Petrotchenko, E.V., Borchers, C.H., Reichmann, D., Bardwell, J.C.A., and Jakob, U. (2016). Protein unfolding as a switch from self-recognition to high-affinity client binding. Nat. Commun. 7: 10357, https://doi.org/10.1038/ncomms10357. Search in Google Scholar
Hassdenteufel, S., Sicking, M., Schorr, S., Aviram, N., Fecher-Trost, C., Schuldiner, M., Jung, M., Zimmermann, R., and Lang, S. (2017). hSnd2 protein represents an alternative targeting factor to the endoplasmic reticulum in human cells. FEBS Lett. 591: 3211–3224, https://doi.org/10.1002/1873-3468.12831. Search in Google Scholar
Hildebrandt, T., Knuesting, J., Berndt, C., Morgan, B., and Scheibe, R. (2015). Cytosolic thiol switches regulating basic cellular functions: GAPDH as an information hub?. Biol. Chem. 396: 523–537, https://doi.org/10.1515/hsz-2014-0295. Search in Google Scholar
Hoffmann, J.H., Linke, K., Graf, P.C., Lilie, H., and Jakob, U. (2004). Identification of a redox-regulated chaperone network. EMBO J. 23: 160–168, https://doi.org/10.1038/sj.emboj.7600016. Search in Google Scholar
Holmgren, A., Johansson, C., Berndt, C., Lonn, M.E., Hudemann, C., and Lillig, C.H. (2005). Thiol redox control via thioredoxin and glutaredoxin systems. Biochem. Soc. Trans. 33: 1375–1377, https://doi.org/10.1042/bst0331375. Search in Google Scholar
Hoppe, G., Chai, Y.C., Crabb, J.W., and Sears, J. (2004). Protein s-glutathionylation in retinal pigment epithelium converts heat shock protein 70 to an active chaperone. Exp. Eye Res. 78: 1085–1092, https://doi.org/10.1016/j.exer.2004.02.001. Search in Google Scholar
Hu, J., Li, J., Qian, X., Denic, V., and Sha, B. (2009). The crystal structures of yeast Get3 suggest a mechanism for tail-anchored protein membrane insertion. PloS One 4: e8061, https://doi.org/10.1371/journal.pone.0008061. Search in Google Scholar
Ilbert, M., Horst, J., Ahrens, S., Winter, J., Graf, P.C., Lilie, H., and Jakob, U. (2007). The redox-switch domain of Hsp33 functions as dual stress sensor. Nat. Struct. Mol. Biol. 14: 556–563, https://doi.org/10.1038/nsmb1244. Search in Google Scholar
Imlay, J.A. (2013). The molecular mechanisms and physiological consequences of oxidative stress: lessons from a model bacterium. Nat. Rev. Microbiol. 11: 443–454, https://doi.org/10.1038/nrmicro3032. Search in Google Scholar
Janda, I., Devedjiev, Y., Derewenda, U., Dauter, Z., Bielnicki, J., Cooper, D.R., Graf, P.C., Joachimiak, A., Jakob, U., and Derewenda, Z.S. (2004). The crystal structure of the reduced, Zn2+-bound form of the B. subtilis Hsp33 chaperone and its implications for the activation mechanism. Structure 12: 1901–1907, https://doi.org/10.1016/j.str.2004.08.003. Search in Google Scholar
Jang, H.H., Lee, K.O., Chi, Y.H., Jung, B.G., Park, S.K., Park, J.H., Lee, J.R., Lee, S.S., Moon, J.C., Yun, J.W., et al. (2004). Two enzymes in one; two yeast peroxiredoxins display oxidative stress-dependent switching from a peroxidase to a molecular chaperone function. Cell 117: 625–635, https://doi.org/10.1016/j.cell.2004.05.002. Search in Google Scholar
Jo, K.S., Kim, J.H., Ryu, K.S., Kang, J.S., Wang, C.Y., Lee, Y.S., Seo, M.D., Lee, Y.H., and Won, H.S. (2019). Unique unfoldase/aggregase activity of a molecular chaperone Hsp33 in its holding-inactive State. J. Mol. Biol. 431: 1468–1480, https://doi.org/10.1016/j.jmb.2019.02.022. Search in Google Scholar
Jonsson, T.J., Johnson, L.C., and Lowther, W.T. (2008). Structure of the sulphiredoxin-peroxiredoxin complex reveals an essential repair embrace. Nature 451: 98–101, https://doi.org/10.1038/nature06415. Search in Google Scholar
Karamzadeh, R., Karimi-Jafari, M.H., Saboury, A.A., Salekdeh, G.H., and Moosavi-Movahedi, A.A. (2017). Red/ox states of human protein disulfide isomerase regulate binding affinity of 17 β-estradiol. Arch. Biochem. Biophys. 619: 35–44, https://doi.org/10.1016/j.abb.2017.02.010. Search in Google Scholar
Kern, R., Malki, A., Holmgren, A., and Richarme, G. (2003). Chaperone properties of Escherichia coli thioredoxin and thioredoxin reductase. Biochem. J. 371: 965–972, https://doi.org/10.1042/bj20030093. Search in Google Scholar
Konig, J., Galliardt, H., Jutte, P., Schaper, S., Dittmann, L., and Dietz, K.J. (2013). The conformational bases for the two functionalities of 2-cysteine peroxiredoxins as peroxidase and chaperone. J. Exp. Bot. 64: 3483–3497, https://doi.org/10.1093/jxb/ert184. Search in Google Scholar
Kumsta, C., Thamsen, M., and Jakob, U. (2011). Effects of oxidative stress on behavior, physiology, and the redox thiol proteome of Caenorhabditis elegans. Antioxid Redox Signal 14: 1023–1037, https://doi.org/10.1089/ars.2010.3203. Search in Google Scholar
Lu, J., Vallabhaneni, H., Yin, J., and Liu, Y. (2013). Deletion of the major peroxiredoxin Tsa1 alters telomere length homeostasis. Aging Cell 12: 635–644, https://doi.org/10.1111/acel.12085. Search in Google Scholar
Mateja, A., Paduch, M., Chang, H.Y., Szydlowska, A., Kossiakoff, A.A., Hegde, R.S., and Keenan, R.J. (2015). Protein targeting. Structure of the Get3 targeting factor in complex with its membrane protein cargo. Science 347: 1152–1155, https://doi.org/10.1126/science.1261671. Search in Google Scholar
Mateja, A., Szlachcic, A., Downing, M.E., Dobosz, M., Mariappan, M., Hegde, R.S., and Keenan, R.J. (2009). The structural basis of tail-anchored membrane protein recognition by Get3. Nature 461: 361–366, https://doi.org/10.1038/nature08319. Search in Google Scholar
Metz, J., Wachter, A., Schmidt, B., Bujnicki, J. M., and Schwappach, B. (2006). The yeast Arr4p ATPase binds the chloride transporter Gef1p when copper is available in the cytosol. J. Biol. Chem. 281: 410–417, https://doi.org/10.1074/jbc.m507481200. Search in Google Scholar
Moayed, F., Bezrukavnikov, S., Naqvi, M.M., Groitl, B., Cremers, C.M., Kramer, G., Ghosh, K., Jakob, U., and Tans, S.J. (2020). The anti-aggregation holdase Hsp33 promotes the formation of folded protein structures. Biophys. J. 118: 85–95, https://doi.org/10.1016/j.bpj.2019.10.040. Search in Google Scholar
Moon, J.C., Hah, Y.S., Kim, W.Y., Jung, B.G., Jang, H.H., Lee, J.R., Kim, S.Y., Lee, Y.M., Jeon, M.G., Kim, C.W., et al. (2005). Oxidative stress-dependent structural and functional switching of a human 2-Cys peroxiredoxin isotype II that enhances HeLa cell resistance to H2O2-induced cell death. J. Biol. Chem. 280: 28775–28784, https://doi.org/10.1074/jbc.m505362200. Search in Google Scholar
Moon, J.C., Kim, G.M., Kim, E.K., Lee, H.N., Ha, B., Lee, S.Y., and Jang, H.H. (2013). Reversal of 2-Cys peroxiredoxin oligomerization by sulfiredoxin. Biochem. Biophys. Res. Commun. 432: 291–295, https://doi.org/10.1016/j.bbrc.2013.01.114. Search in Google Scholar
Morais, M.A.B., Giuseppe, P.O., Souza, T., Castro, H., Honorato, R.V., Oliveira, P.S.L., Netto, L.E.S., Tomas, A.M., and Murakami, M.T. (2017). Calcium and magnesium ions modulate the oligomeric state and function of mitochondrial 2-Cys peroxiredoxins in Leishmania parasites. J. Biol. Chem. 292: 7023–7039, https://doi.org/10.1074/jbc.m116.762039. Search in Google Scholar
Morano, K.A., Grant, C.M., and Moye-Rowley, W.S. (2012). The response to heat shock and oxidative stress in Saccharomyces cerevisiae. Genetics 190: 1157–1195, https://doi.org/10.1534/genetics.111.128033. Search in Google Scholar
Mukhopadhyay, R., Ho, Y.S., Swiatek, P.J., Rosen, B.P., and Bhattacharjee, H. (2006). Targeted disruption of the mouse Asna1 gene results in embryonic lethality. FEBS Lett. 580: 3889–3894, https://doi.org/10.1016/j.febslet.2006.06.017. Search in Google Scholar
Muller, A., Langklotz, S., Lupilova, N., Kuhlmann, K., Bandow, J.E., and Leichert, L.I. (2014). Activation of RidA chaperone function by N-chlorination. Nat. Commun. 5: 5804, https://doi.org/10.1038/ncomms6804. Search in Google Scholar
Nelson, K.J., Knutson, S.T., Soito, L., Klomsiri, C., Poole, L.B., and Fetrow, J.S. (2011). Analysis of the peroxiredoxin family: using active-site structure and sequence information for global classification and residue analysis. Proteins 79: 947–964, https://doi.org/10.1002/prot.22936. Search in Google Scholar
Noichri, Y., Palais, G., Ruby, V., D’Autreaux, B., Delaunay-Moisan, A., Nystrom, T., Molin, M., and Toledano, M.B. (2015). In vivo parameters influencing 2-Cys Prx oligomerization: the role of enzyme sulfinylation. Redox Biol 6: 326–333, https://doi.org/10.1016/j.redox.2015.08.011. Search in Google Scholar
Powis, K., Schrul, B., Tienson, H., Gostimskaya, I., Breker, M., High, S., Schuldiner, M., Jakob, U., and Schwappach, B. (2013). Get3 is a holdase chaperone and moves to deposition sites for aggregated proteins when membrane targeting is blocked. J. Cell Sci. 126: 473–483, https://doi.org/10.1242/jcs.112151. Search in Google Scholar
Ralser, M., Wamelink, M.M., Kowald, A., Gerisch, B., Heeren, G., Struys, E. A., Klipp, E., Jakobs, C., Breitenbach, M., Lehrach, H., et al. (2007). Dynamic rerouting of the carbohydrate flux is key to counteracting oxidative stress. J Biol 6: 10, https://doi.org/10.1186/jbiol61. Search in Google Scholar
Reichmann, D., Xu, Y., Cremers, C.M., Ilbert, M., Mittelman, R., Fitzgerald, M.C., and Jakob, U. (2012). Order out of disorder: working cycle of an intrinsically unfolded chaperone. Cell 148: 947–957, https://doi.org/10.1016/j.cell.2012.01.045. Search in Google Scholar
Rhee, S.G. and Woo, H.A. (2020). Multiple functions of 2-Cys peroxiredoxins, I and II, and their regulations via post-translational modifications. Free Radic. Biol. Med. 152: 107–115, https://doi.org/10.1016/j.freeradbiomed.2020.02.028. Search in Google Scholar
Rhee, S.G., Woo, H.A., and Kang, D. (2018). The role of peroxiredoxins in the transduction of H2O2 Signals. Antioxid Redox Signal 28: 537–557, https://doi.org/10.1089/ars.2017.7167. Search in Google Scholar
Rimon, O., Suss, O., Goldenberg, M., Fassler, R., Yogev, O., Amartely, H., Propper, G., Friedler, A., and Reichmann, D. (2017). A role of metastable regions and their connectivity in the inactivation of a redox-regulated chaperone and its inter-chaperone crosstalk. Antioxid Redox Signal 27: 1252–1267, https://doi.org/10.1089/ars.2016.6900. Search in Google Scholar
Rivera-Monroy, J., Musiol, L., Unthan-Fechner, K., Farkas, A., Clancy, A., Coy-Vergara, J., Weill, U., Gockel, S., Lin, S.Y., Corey, D.P., et al. (2016). Mice lacking WRB reveal differential biogenesis requirements of tail-anchored proteins in vivo. Sci. Rep. 6: 39464, https://doi.org/10.1038/srep39464. Search in Google Scholar
Rosenzweig, R., Nillegoda, N.B., Mayer, M.P., and Bukau, B. (2019). The Hsp70 chaperone network. Nat. Rev. Mol. Cell Biol. 20: 665–680, https://doi.org/10.1038/s41580-019-0133-3. Search in Google Scholar
Saccoccia, F., Di Micco, P., Boumis, G., Brunori, M., Koutris, I., Miele, A.E., Morea, V., Sriratana, P., Williams, D.L., Bellelli, A., et al. (2012). Moonlighting by different stressors: crystal structure of the chaperone species of a 2-Cys peroxiredoxin. Structure 20: 429–439, https://doi.org/10.1016/j.str.2012.01.004. Search in Google Scholar
Schuldiner, M., Metz, J., Schmid, V., Denic, V., Rakwalska, M., Schmitt, H.D., Schwappach, B., and Weissman, J.S. (2008). The GET complex mediates insertion of tail-anchored proteins into the ER membrane. Cell 134: 634–645, https://doi.org/10.1016/j.cell.2008.06.025. Search in Google Scholar
Schwarzlander, M., Dick, T.P., Meyer, A.J., and Morgan, B. (2016). Dissecting redox biology using fluorescent protein sensors. Antioxid Redox Signal 24: 680–712, https://doi.org/10.1089/ars.2015.6266. Search in Google Scholar
Segal, N., and Shapira, M. (2015). HSP33 in eukaryotes - an evolutionary tale of a chaperone adapted to photosynthetic organisms. Plant J. 82: 850–860, https://doi.org/10.1111/tpj.12855. Search in Google Scholar
Shen, J., Hsu, C.M., Kang, B.K., Rosen, B.P., and Bhattacharjee, H. (2003). The Saccharomyces cerevisiae Arr4p is involved in metal and heat tolerance. Biometals 16: 369–378, https://doi.org/10.1023/a:1022504311669. Search in Google Scholar
Shenton, D., Smirnova, J.B., Selley, J.N., Carroll, K., Hubbard, S.J., Pavitt, G.D., Ashe, M.P., and Grant, C.M. (2006). Global translational responses to oxidative stress impact upon multiple levels of protein synthesis. J. Biol. Chem. 281: 29011–29021, https://doi.org/10.1074/jbc.m601545200. Search in Google Scholar
Siegenthaler, K.D., Pareja, K.A., Wang, J., and Sevier, C.S. (2017). An unexpected role for the yeast nucleotide exchange factor Sil1 as a reductant acting on the molecular chaperone BiP. Elife 6, https://doi.org/10.7554/elife.24141. Search in Google Scholar
Sies, H. (2017). Hydrogen peroxide as a central redox signaling molecule in physiological oxidative stress: oxidative eustress. Redox Biol. 11: 613–619, https://doi.org/10.1016/j.redox.2016.12.035. Search in Google Scholar
Sobotta, M.C., Liou, W., Stocker, S., Talwar, D., Oehler, M., Ruppert, T., Scharf, A.N., and Dick, T.P. (2015). Peroxiredoxin-2 and STAT3 form a redox relay for H2O2 signaling. Nat. Chem. Biol. 11: 64–70, https://doi.org/10.1038/nchembio.1695. Search in Google Scholar
Stefanovic, S. and Hegde, R.S. (2007). Identification of a targeting factor for posttranslational membrane protein insertion into the ER. Cell 128: 1147–1159, https://doi.org/10.1016/j.cell.2007.01.036. Search in Google Scholar
Stefer, S., Reitz, S., Wang, F., Wild, K., Pang, Y.Y., Schwarz, D., Bomke, J., Hein, C., Lohr, F., Bernhard, F., et al. (2011). Structural basis for tail-anchored membrane protein biogenesis by the Get3-receptor complex. Science 333: 758–762, https://doi.org/10.1126/science.1207125. Search in Google Scholar
Stocker, S., Maurer, M., Ruppert, T., and Dick, T.P. (2018). A role for 2-Cys peroxiredoxins in facilitating cytosolic protein thiol oxidation. Nat. Chem. Biol. 14: 148–155, https://doi.org/10.1038/nchembio.2536. Search in Google Scholar
Sultana, S., Foti, A., and Dahl, J.U. (2020). Bacterial defense systems against the neutrophilic oxidant hypochlorous acid. Infect. Immun. 88: e00964–19, https://doi.org/10.1145/3334480.3381819. Search in Google Scholar
Teixeira, F., Castro, H., Cruz, T., Tse, E., Koldewey, P., Southworth, D.R., Tomas, A.M., and Jakob, U. (2015). Mitochondrial peroxiredoxin functions as crucial chaperone reservoir in Leishmania infantum. Proc. Natl. Acad. Sci. U.S.A. 112: E616–624, https://doi.org/10.1073/pnas.1419682112. Search in Google Scholar
Teixeira, F., Tse, E., Castro, H., Makepeace, K.A.T., Meinen, B.A., Borchers, C.H., Poole, L.B., Bardwell, J.C., Tomas, A.M., Southworth, D.R., et al. (2019). Chaperone activation and client binding of a 2-cysteine peroxiredoxin. Nat. Commun. 10: 659, https://doi.org/10.1038/s41467-019-08565-8. Search in Google Scholar
Toledano, M.B. and Huang, B. (2016). Microbial 2-Cys peroxiredoxins: insights into their complex physiological roles. Mol Cells 39: 31–39, https://doi.org/10.14348/molcells.2016.2326. Search in Google Scholar
Ulrich, K. and Jakob, U. (2019). The role of thiols in antioxidant systems. Free Radic. Biol. Med. 140: 14–27, https://doi.org/10.1016/j.freeradbiomed.2019.05.035. Search in Google Scholar
Vijayalakshmi, J., Mukhergee, M.K., Graumann, J., Jakob, U., and Saper, M.A. (2001). The 2.2 Å crystal structure of Hsp33: a heat shock protein with redox-regulated chaperone activity. Structure 9: 367–375, https://doi.org/10.1016/s0969-2126(01)00597-4. Search in Google Scholar
Voth, W., Schick, M., Gates, S., Li, S., Vilardi, F., Gostimskaya, I., Southworth, D.R., Schwappach, B., and Jakob, U. (2014). The protein targeting factor Get3 functions as ATP-independent chaperone under oxidative stress conditions. Mol Cell 56: 116–127, https://doi.org/10.1016/j.molcel.2014.08.017. Search in Google Scholar
Wang, C., Yu, J., Huo, L., Wang, L., Feng, W., and Wang, C.C. (2012a). Human protein-disulfide isomerase is a redox-regulated chaperone activated by oxidation of domain a’. J. Biol. Chem. 287: 1139–1149, https://doi.org/10.1074/jbc.m111.303149. Search in Google Scholar
Wang, J., Pareja, K.A., Kaiser, C.A., and Sevier, C.S. (2014). Redox signaling via the molecular chaperone BiP protects cells against endoplasmic reticulum-derived oxidative stress. eLife 3: e03496, https://doi.org/10.7554/elife.03496. Search in Google Scholar
Wang, J. and Sevier, C.S. (2016). Formation and reversibility of BiP protein cysteine oxidation facilitate cell survival during and post oxidative stress. J. Biol. Chem. 291: 7541–7557, https://doi.org/10.1074/jbc.m115.694810. Search in Google Scholar
Wang, Y., Gibney, P.A., West, J.D., and Morano, K.A. (2012b). The yeast Hsp70 Ssa1 is a sensor for activation of the heat shock response by thiol-reactive compounds. Mol. Biol. Cell 23: 3290–3298, https://doi.org/10.1091/mbc.e12-06-0447. Search in Google Scholar
Wei, P.C., Hsieh, Y.H., Su, M.I., Jiang, X., Hsu, P.H., Lo, W.T., Weng, J.Y., Jeng, Y.M., Wang, J.M., Chen, P.L., et al. (2012). Loss of the oxidative stress sensor NPGPx compromises GRP78 chaperone activity and induces systemic disease. Mol Cell 48: 747–759, https://doi.org/10.1016/j.molcel.2012.10.007. Search in Google Scholar
Winter, J., Ilbert, M., Graf, P.C., Ozcelik, D., and Jakob, U. (2008). Bleach activates a redox-regulated chaperone by oxidative protein unfolding. Cell 135: 691–701, https://doi.org/10.1016/j.cell.2008.09.024. Search in Google Scholar
Winter, J., Linke, K., Jatzek, A., and Jakob, U. (2005). Severe oxidative stress causes inactivation of DnaK and activation of the redox-regulated chaperone Hsp33. Mol Cell 17: 381–392, https://doi.org/10.1016/j.molcel.2004.12.027. Search in Google Scholar
Winterbourn, C.C., Kettle, A.J., and Hampton, M.B. (2016). Reactive oxygen species and neutrophil function. Annu. Rev. Biochem. 85: 765–792, https://doi.org/10.1146/annurev-biochem-060815-014442. Search in Google Scholar
Woo, H.A., Chae, H.Z., Hwang, S.C., Yang, K.S., Kang, S.W., Kim, K., and Rhee, S.G. (2003). Reversing the inactivation of peroxiredoxins caused by cysteine sulfinic acid formation. Science 300: 653–656, https://doi.org/10.1126/science.1080273. Search in Google Scholar
Wood, Z.A., Poole, L.B., Hantgan, R.R., and Karplus, P.A. (2002). Dimers to doughnuts: redox-sensitive oligomerization of 2-cysteine peroxiredoxins. Biochemistry 41: 5493–5504, https://doi.org/10.1021/bi012173m. Search in Google Scholar
Wood, Z.A., Poole, L.B., and Karplus, P.A. (2003). Peroxiredoxin evolution and the regulation of hydrogen peroxide signaling. Science 300: 650–653, https://doi.org/10.1126/science.1080405. Search in Google Scholar
Wyatt, A.R., Kumita, J.R., Mifsud, R.W., Gooden, C.A., Wilson, M.R., and Dobson, C.M. (2014). Hypochlorite-induced structural modifications enhance the chaperone activity of human alpha2-macroglobulin. Proc. Natl. Acad. Sci. U.S.A. 111: E2081–2090, https://doi.org/10.1073/pnas.1403379111. Search in Google Scholar
Xiao, H., Jedrychowski, M.P., Schweppe, D.K., Huttlin, E.L., Yu, Q., Heppner, D.E., Li, J., Long, J., Mills, E.L., Szpyt, J., et al. (2020). A quantitative tissue-specific landscape of protein redox regulation during aging. Cell 180: 968–983, https://doi.org/10.1016/j.cell.2020.02.012, e924. Search in Google Scholar
Yang, K.S., Kang, S.W., Woo, H.A., Hwang, S.C., Chae, H.Z., Kim, K., and Rhee, S.G. (2002). Inactivation of human peroxiredoxin I during catalysis as the result of the oxidation of the catalytic site cysteine to cysteine-sulfinic acid. J. Biol. Chem. 277: 38029–38036, https://doi.org/10.1074/jbc.m206626200. Search in Google Scholar
Yoo, N.G., Dogra, S., Meinen, B.A., Tse, E., Haefliger, J., Southworth, D.R., Gray, M.J., Dahl, J.U., and Jakob, U. (2018). Polyphosphate stabilizes protein unfolding intermediates as soluble amyloid-like oligomers. J. Mol. Biol. 430: 4195–4208, https://doi.org/10.1016/j.jmb.2018.08.016. Search in Google Scholar
Zhang, H., Yang, J., Wu, S., Gong, W., Chen, C., and Perrett, S. (2016). Glutathionylation of the bacterial Hsp70 chaperone DnaK provides a link between oxidative stress and the heat shock response. J. Biol. Chem. 291: 6967–6981, https://doi.org/10.1074/jbc.m115.673608. Search in Google Scholar
© 2020 Kathrin Ulrich et al., published by De Gruyter, Berlin/Boston
This work is licensed under the Creative Commons Attribution 4.0 International License.