Unfolding of Metastable Linker Region Is at the Core of Hsp33 Activation as a Redox-regulated Chaperone^*

Strains and Plasmids

The Hsp33-M172S and Hsp33-Y12E mutants were generated by introducing single site mutations into the wild-type Hsp33 gene (hslO) using pUJ30 (pET11a-hslO) (3) as a template. The plasmids were transformed into JH13 (BL21, ΔhslO) (12), generating the expression strains JHa21 (JH13, pET11a-hslO-M172S) and CC4 (JH13, pET11a-hslO-Y12E). Additional strains used in this study are UJ83, a BL21 strain expressing wild-type Hsp33 from a pET11a vector, and CC20, a JH13 strain carrying the empty pET11a plasmid.

Purification of Wild-type and Hsp33 Mutant Proteins

Cultivation of the Hsp33-overexpressing strains for protein purification was conducted as previously described (12). For the induction of the Hsp33-M172S mutant, the protocol for wild-type Hsp33 was followed (3). To overexpress large amounts of soluble Hsp33-Y12E protein, cells were grown to an A₆₀₀ of 0.6–0.8 at 37 °C and then shifted to 18 °C. Once the temperature was reached, Hsp33-Y12E expression was induced with 1 mm isopropyl-1-thio-β-d-galactopyranoside (IPTG)⁴ for 24 h. Afterward, the standard protocol for the Hsp33 purification was followed (12). All proteins were stored in 40 mm potassium phosphate buffer (KH₂PO₄) (pH 7.5) at −20 °C. Reduced, zinc reconstituted Hsp33_red was prepared as previously described (12). To prepare oxidized Hsp33, 50 μm Hsp33_red was incubated in 2 mm H₂O₂ at either 30 °C (Hsp33_{ox30 °C}) or 43 °C (Hsp33_{ox43 °C}) for 3 h. Oxidants were removed using NAP-5 columns.

Hsp33 Chaperone Activity Assay

The chaperone activity of wild-type Hsp33 and its variants was measured as previously described (13). In short, 12 μm citrate synthase (CS, Roche) was denatured with 4.5 m Gdn-HCl in 40 mm HEPES-KOH (pH 7.5) overnight at room temperature. To initiate protein aggregation, denatured CS was diluted to a final concentration of 75 nm in 40 mm HEPES-KOH, pH 7.5 at 30 °C under continuous stirring in the absence or presence of a 4× molar excess of wild-type Hsp33 or Hsp33 variants. To assess the activity of reduced wild-type Hsp33 and the variants at heat shock temperatures, CS (final concentration 150 nm) was incubated in the absence or presence of a 4× molar excess of Hsp33 in 40 mm HEPES-KOH, pH 7.5 at 43 °C. Light scattering was monitored at λ_ex/λ_em at 360 nm using a Hitachi F4500 fluorescence spectrophotometer equipped with a temperature-controlled cuvette holder and stirrer.

Far-UV CD Spectroscopy

To determine changes in the secondary structure of wild-type Hsp33 and its variants, far-UV CD spectra were recorded in 20 mm KH₂PO₄, pH 7.5 at 20 °C using a Jasco-J810 spectropolarimeter as previously described (6). To determine the thermostability of Hsp33 and its derivates, the CD signal at either 195 nm or 222 nm was recorded at temperatures from 20 to 50 °C or 20 to 80 °C, respectively. The temperature was increased at a rate of 1 °C per minute. The temperature was controlled with a Jasco Peltier device.

ANS Fluorescence Measurements

The fluorescence probe bis-ANS (Molecular Probes) was used to test for the presence of hydrophobic surfaces in wild-type Hsp33 and the variants as previously described (6).

Thiol Redox State of Hsp33 and Its Variants

To assess whether oxidation of Hsp33 occurred during the chaperone activity assay, thiol trapping with the 490-Da thiol-reactive probe AMS was performed immediately after the activity measurement (3). An aliquot (815 μl) of the activity assay mixture was supplemented with 100% (v/v) trichloroacetic acid to a final concentration of 10% (v/v) trichloroacetic acid and incubated on ice for 30 min. The solution was centrifuged (30 min, 16,000 × g, 4 °C), and the protein pellet was resuspended in 20 μl of AMS-DAB solution (15 mm AMS, 6 m urea, 200 mm Tris-HCl, pH 8.5, 50 mm EDTA, 7.5% w/v SDS). After a 1-h incubation in the dark with shaking (1,300 rpm), 5 μl of 5× non-reducing Laemmli buffer was added, and the sample was loaded onto a non-reducing SDS-PAGE gel. Proteins were visualized by Western blot or Coomassie Blue staining of the gels.

Phenotype, Hsp33 Expression Levels, and Aggregate Formation in Strains Expressing Wild-type Hsp33 and Mutants

Overnight cultures of strains expressing either wild-type Hsp33 (UJ83), the two Hsp33 variants (CC4, JHa21), or no protein (CC20) from a pET11a vector in MOPS minimal medium (10× MOPS modified buffer (TEKnovo), 0.2 (w/v) glucose, 132 mm K₂HPO₄ pH 7, 20 μm thiamine) were prepared. Cell cultures were diluted 1:100 into MOPS minimal medium, supplemented with 50 μm IPTG, and cultivated at 37 °C. Growth was monitored every hour with absorbance measurements at A₆₀₀. To compare soluble and insoluble proteins in these strains, 5 ml of the culture were removed, and cells were harvested by centrifugation (5,000 × g, 4 °C, 10 min) after 5.5 h of cultivation. Cells were resuspended in 650 μl of sucrose buffer (50 mm Tris-HCl pH 8.0, 25% w/v sucrose, 1 mm NaEDTA, 10 mm DTT) and homogenized by sonication (Kontes, Micro Ultrasonic cell disrupter, 2 × 10–15 s with the output control set to 70). 5 μl of lysozyme (50 mg/ml), 12.5 μl of DNase l (2 mg/ml), and 2.5 μl of MgCl₂ (0.5 m) were added, followed by the addition of 650 μl of lysis buffer (50 mm Tris-HCl pH 8.0, 1% Triton X-100, 100 mm NaCl, 0.1% sodium azide, 0.1% sodium deoxycholate, 10 mm DTT). Samples were incubated at room temperature for 15 min, frozen at −80 °C for 20 min, and thawed at 37 °C for 15 min. Then, 2.5 μl of MgCl₂ (0.5 m) were added, and the solution was incubated at room temperature until the viscosity decreased. NaEDTA was added, and the lysate was centrifuged (11,000 × g, 4 °C, 20 min). The supernatant was removed, and the pellet was resuspended in 500 μl of washing buffer A (50 mm Tris-HCl, pH 8.0, 0.5% Triton X-100, 100 mm NaCl, 1 mm NaEDTA, 1 mm DTT). The solution was again homogenized by sonication as before. Protein aggregates were pelleted by centrifugation (11,000 × g, 4 °C, 20 min). The supernatant was discarded, and the pellet was resuspended in 500 μl of washing buffer B (50 mm Tris-HCl pH 8.0, 100 mm NaCl, 1 mm NaEDTA, 1 mm DTT), and again homogenized by sonication. The washing steps were repeated three times. The final pellet was resuspended in 2× reducing Laemmli buffer and boiled for 5 min. Aggregate formation was visualized by SDS-PAGE followed by Coomassie Blue staining of the gels.

Homology Modeling and Calculation of the Connectivity Map

The homology modeling of reduced E. coli Hsp33 was performed using the MODELLER software through an interface of the MODWEB modeling server. The crystal structure of reduced Bacillus subtilis Hsp33 (PDB code:1VZY) was used as a template (7). The E. coli Hsp33 sequence was aligned to the B. subtilis Hsp33 structure revealing a sequence identity of 24%. To validate the model, the highly conserved core domain of Hsp33 (residues 7–178) was aligned (RMSD = 0.3 Å). A connectivity map was derived using the AQUAPROT server, which calculates interatomic interactions between residues in two chains (14). For this purpose, the E. coli Hsp33 sequence was divided into N-terminal (amino acids 7–178) and C-terminal (amino acids 179–288) domains. The results were confirmed by a standalone version of the same algorithm, which computes interactions within one chain. The definition of interactions between residues was based on the “all-atom contact” method (15), calculating the detailed atomic contacts including hydrogen bonds, van der Waals, aromatic and electrostatic interactions (15). The van der Waals interactions also reflect side chain packing between protein residues. The residue interaction network was plotted using the Cytoscape V2.6.3 program (16).

Site-specific Mutations to Alter the Linker Binding Interface

The surface of the Hsp33 N-terminal β-sheet, which is masked by the folded linker in the crystal structure of reduced, inactive Hsp33 (Fig. 1A), has been postulated to serve as a substrate binding site for unfolded proteins when Hsp33 is oxidized and active (8–10). Analysis of the Hsp33 x-ray structure reveals a cluster of highly conserved hydrophobic amino acids whose side chains face toward the linker region under reducing conditions and might interact with unfolded substrate proteins under oxidizing conditions. We focused on two conserved residues that mapped to the hydrophobic platform, Tyr¹² on β-strand 1 and Met¹⁷² on β-strand 9 (Fig. 1B, supplemental Fig. S1). We predicted that alteration of these residues would affect the surface features of the four-stranded β-sheet but not the stability or fold of the Hsp33 N-terminal domain. To efficiently disrupt potential hydrophobic interactions, we individually replaced Tyr¹² with glutamate (Hsp33-Y12E) or Met¹⁷² with the polar residue serine (Hsp33-M172S). We then overexpressed both mutant proteins in BL21 strains lacking the endogenous Hsp33 gene and determined their solubilities. Hsp33-M172S was fully soluble upon IPTG-induced overexpression at 37 °C. In contrast, the Hsp33-Y12E variant was overexpressed but largely insoluble under these conditions. A 24-h induction with IPTG at 18 °C was required to accumulate significant amounts of soluble Hsp33-Y12E. We purified the two variants according to the wild-type Hsp33 protocol and prepared fully reduced, zinc-reconstituted proteins (Hsp33-Y12E_red, Hsp33-M172S_red) for subsequent functional studies.

Mutations in the Hsp33 N-terminal Domain Dramatically Alter Functional Regulation

Hsp33 activation by fast-acting oxidants such as HOCl occurs within the mixing time of the experiment, making evaluation of activation kinetics and determination of oxidation intermediates technically challenging. In contrast, H₂O₂-mediated activation of Hsp33 is significantly slower (t_½ of ∼20 min) and depends on the additional presence of unfolding conditions (43 °C) (Fig. 2A, open circles). In the absence of unfolding conditions (H₂O₂, 30 °C), an oxidation intermediate accumulates, which appears to lack the second disulfide bond (11) and is inactive as a chaperone (Fig. 2A, inset). To determine how the introduced mutations affect the redox regulation and chaperone function of our Hsp33 variants, we monitored their activity upon incubation in 2 mm H₂O₂ at either 30 or 43 °C. As shown in Fig. 2A, both variants displayed chaperone activity very similar to that of activated wild-type Hsp33 when incubated with H₂O₂ at 43 °C for 2 h. All three oxidized variants suppressed the aggregation of chemically denatured CS to the same extent. These results clearly indicate that the introduced mutations did not substantially affect the substrate binding affinity of Hsp33. Similar results were observed when the activity of the proteins was tested using thermally unfolding malate dehydrogenase as an in vitro chaperone substrate (supplemental Fig. S2A).

In contrast to the chaperone function, which seems unchanged, the regulation of Hsp33 chaperone function appears to be dramatically affected by the mutations. Activation of Hsp33-M172S was significantly faster at 43 °C and lacked the lag phase characteristic of wild-type Hsp33 (Fig. 2A, compare Hsp33-M172S, closed squares and wild-type Hsp33, open circles). This lag phase has been shown to correlate to the formation of the critical second disulfide bond in Hsp33 and is thought to represent the rate-limiting step in its activation (11). Moreover, activation of Hsp33-M172S no longer required the simultaneous presence of oxidizing and unfolding conditions. Incubation of Hsp33-M172S in 2 mm H₂O₂ at 30 °C was fully sufficient for its activation (Fig. 2A, inset, closed squares) whereas wild-type Hsp33 (Fig. 2A, inset, open circles) remained inactive. Likewise, simple incubation of reduced, zinc-reconstituted Hsp33-M172S at elevated temperatures (43 °C) caused activation of its chaperone function (Fig. 2B). In contrast to the oxidative activation of Hsp33, however, which is irreversible unless reducing agents are added, thermal activation of Hsp33-M172S was fully reversible and required the activity tests to be performed at 43 °C (Fig. 2B).

Whereas Hsp33-M172S maintained at least some requirement for post-translational regulation, the freshly reduced and zinc-reconstituted Hsp33-Y12E mutant appeared to be fully active even without incubation in H₂O₂ or exposure to elevated temperatures (Fig. 2, A and B). Additional exposure to H₂O₂ at either 30 or 43 °C did not further increase its chaperone activity, suggesting that Hsp33-Y12E is constitutively active (Fig. 2A, open squares). To exclude the possibility that high oxidation sensitivity of the active site cysteines in the two Hsp33 variants leads to their uncontrolled oxidation and subsequent activation during the chaperone assays, we performed thiol trapping experiments on the reduced proteins immediately after the activity assays to assess their cysteine oxidation status (Fig. 2B). We made use of the 490-Da thiol-alkylating reagent AMS, which irreversibly reacts with reduced cysteines and causes a distinct mass increase that is easily discernable on SDS-PAGE. We found that the cysteines in both mutant proteins are reduced and remain reduced even during incubation at 43 °C. This result agrees well with zinc binding studies, which revealed that both mutant proteins remain fully zinc-reconstituted even in the presence of a 20-fold excess of the zinc chelator PAR (K_α = 2 × 10¹² m⁻¹ at pH 7) (17) (supplemental Fig. S2B).

To test the oligomerization status of the reduced Hsp33 variants, we performed analytical ultracentrifugation and confirmed that both variants are monomeric, as is reduced wild-type Hsp33 (data not shown) (12, 18). These results are in contrast to previous studies, which suggested that dimerization of Hsp33 is required for the full activation of its chaperone function (6, 18). Some of these conclusions were based on a single Hsp33 mutation designed to render Hsp33 constitutively monomeric (Hsp33-E150R) (6). Our results with the Hsp33-Y12E mutant suggest that Hsp33 does not require dimerization for its chaperone function in vitro. It is therefore conceivable that the E150R mutation not only interferes with Hsp33 dimerization but, more importantly, with substrate binding. This possibility remains to be investigated. Nevertheless, we conclude from our results that single amino acid mutations at the linker binding interface of the Hsp33 N terminus are sufficient to generate Hsp33 variants that function as highly effective chaperone holdases despite their reduced, zinc-reconstituted, and monomeric conformation.

Hsp33 Variants Show a Decrease in Surface Hydrophobicity

Our results indicated that mutations designed to alter the hydrophobic character of the N-terminal β-sheet platform affect the redox regulation but not the chaperone function of Hsp33. To assess whether the mutations actually caused a detectable change in hydrophobicity, particularly in those surfaces that specifically become exposed upon Hsp33 activation, we recorded the fluorescence spectra of reduced and oxidized wild-type and mutant Hsp33 proteins using the hydrophobic probe bis-ANS. As previously shown, wild-type Hsp33 undergoes substantial conformational rearrangements upon oxidative activation, which lead to the exposure of extensive hydrophobic surfaces that interact with bis-ANS (10) (Fig. 3, compare inactive Hsp33_red, trace a and activated Hsp33_{ox43 °C}, trace a′). When both mutant proteins were exposed to the same oxidizing conditions (2 mm H₂O₂, 43 °C, 3 h), which fully activated wild-type Hsp33, their extent of surface hydrophobicity was significantly reduced (Fig. 3, compare traces b′ and c′ with a′). These results confirmed that the mutations did indeed affect the hydrophobicity of surface areas that become exposed during the activation process. Furthermore, analysis of the surface hydrophobicity of the reduced, constitutively active Hsp33-Y12E mutant revealed that not all of the hydrophobic surface areas exposed upon oxidative activation of Hsp33 are necessary for its chaperone function. This mutant protein is as active as oxidized wild-type Hsp33 when reduced and zinc-coordinated but shows significantly less overall surface hydrophobicity (Fig. 3, trace c).

Close Correlation between Linker Destabilization and Hsp33 Activity

Based on the proposed activation model, Hsp33 chaperone function depends on the formation of the critical disulfide bond between Cys²³² and Cys²³⁴, which is both kinetically and thermodynamically linked to the folding status of the linker region (4). Mutations that destabilize the linker region should therefore enhance the rate of disulfide bond formation and activation. To investigate whether the M172S and Y12E mutations indeed altered the thermostability of the linker region in the Hsp33 variants, we performed far-UV circular dichroism (CD) spectroscopy to monitor changes in Hsp33 secondary structure as a function of temperature. As previously shown, when fully reduced and zinc-reconstituted, wild-type Hsp33 begins to irreversibly unfold at temperatures above ∼50 °C (supplemental Fig. S3A) (1). When fully oxidized, zinc-depleted, and activated (Hsp33_{ox43 °C}), the linker region and zinc binding domain of wild-type Hsp33 remain unfolded at temperatures as low as 20 °C (Fig. 4), and the N-terminal domain begins to unfold at a temperature that is similar to the onset of unfolding of reduced Hsp33 (supplemental Fig. S3A). In stark contrast, the reduced, zinc-reconstituted Hsp33-M172S variant begins to unfold at temperatures as low as 30 °C (Fig. 4). This unfolding (T_m ∼ 40 °C), which is fully reversible up to 50 °C (supplemental Fig. S3B), is highly reminiscent of the oxidized, inactive Hsp33_{ox30 °C} intermediate, which lacks zinc coordination and has a substantially destabilized linker region (T_m < 43 °C) (1). A further increase in the incubation temperature of reduced Hsp33-M172S led to the irreversible unfolding of the remaining protein with the onset of unfolding at a temperature that was again similar to reduced wild-type Hsp33 (supplemental Fig. S3A). These results suggest that the M172S mutation does not globally destabilize Hsp33. Our finding that the linker region of Hsp33-M172S reversibly unfolds upon exposure to heat shock temperatures (supplemental Fig. S3B) agrees well with our previous observation that the reduced Hsp33-M172S acts as a molecular chaperone when tested at 43 °C but not at 30 °C (Fig. 2B). We concluded from these results that unfolding of the linker region is a critical element for the chaperone function of Hsp33.

This conclusion was further supported by the CD analysis of the reduced, constitutively active Hsp33-Y12E mutant, which showed dramatically decreased secondary structure content, suggestive of a constitutively unfolded linker region (Fig. 4). Subsequent incubation with H₂O₂ caused some additional loss of secondary structure as monitored by CD spectroscopy, which likely corresponds to the oxidative zinc release and the unfolding of the zinc binding domain (supplemental Fig. S3C). Unfolding of the remaining protein structure started at temperatures similar to the wild-type Hsp33_red and Hsp33-M172S_red mutant protein, suggesting that also this substitution did not globally destabilize Hsp33 (supplemental Fig. S3A).

These results provide good evidence that destabilization of the linker region by single mutations in the Hsp33 N-terminal linker binding domain is sufficient to eliminate Hsp33 post-translational regulation. They further imply that the stability of the linker region plays a central role in the chaperone function of Hsp33. The Hsp33 redox-sensing zinc binding domain appears to primarily serve as the molecular toggle whose redox status controls the thermostability of the linker region and ultimately the activity of the protein.

Tyrosine 12-a Central Interaction Hub between the Hsp33 N and C Terminus

To better understand how substitution at either Met¹⁷² or Tyr¹² can exert such dramatic effects on Hsp33 functional regulation, we decided to analyze the interatomic connectivity of each of these residues with their surroundings. Unfortunately, the structure of reduced E. coli, Hsp33 is not known, and the only two available crystal structures of E. coli Hsp33 are domain-swapped and truncated protein variants (9, 19). We therefore chose to model the structure of reduced E. coli Hsp33 using the x-ray structure of reduced B. subtilis Hsp33 (1VZY) as a template (sequence identity 24%) (7). The quality of the obtained model, which we calculated using the software MODELLER (20), appeared to be very high, as reflected by the maximum GA341 score of 1 (21). When we analyzed the location of Tyr¹² in this model (Fig. 5A, magenta spheres), we found that this residue makes extensive contacts with a large number of linker residues (Fig. 5A, cyan spheres). To characterize the architecture and connectivity of these interacting residues in more detail, we computed a contact network using AQUAPROT (14). This type of presentation reveals the environmental importance of specific residues and unveils potential cooperative interactions between residues (22). The contact map further confirmed the high connectivity of Tyr¹² with residues located in the Hsp33 linker region (Fig. 5B). Tyr¹² was found to interact with at least five residues in the linker region predominantly by side chain interactions, with one of the strongest contacts formed between Tyr¹² and Glu²¹⁹. Therefore, substitution of Tyr¹² with a glutamate as in our Hsp33-Y12E variant should not only cause a decrease in residue packing but also strong repulsive interactions between the Hsp33 N and C terminus. This would explain how substitution of a single residue in the Hsp33 N terminus could lead to separation of the linker region from the N-terminal β-sheet platform and subsequent linker unfolding. The side chain atoms of Met¹⁷² are involved in interactions with three residues, only one of which, Leu²¹⁴, is located in the linker region (data not shown). This result might explain why the M172S mutation leads to a more moderate destabilization of the linker region.

Lack of Post-translational Regulation of Hsp33 Affects E. coli Growth

Our results suggested that we generated two Hsp33 mutants that lack their proper post-translational redox regulation in vitro. These mutants provided us now with the opportunity to investigate what role the post-translational regulation of Hsp33 plays in the cell. We cultivated BL21ΔhslO strains expressing either wild-type Hsp33 or the variants from an IPTG-inducible pET11a vector in the presence of increasing concentrations of IPTG at 37 °C (supplemental Fig. S4A). We observed a significant growth defect in strains expressing the constitutively active Hsp33-Y12E mutant at IPTG concentrations as low as 50 μm (Fig. 6A, open circles). In contrast, no growth defect was detected in strains expressing wild-type Hsp33 or the temperature-controlled Hsp33-M172S mutant protein under the same growth conditions. Increasing the cultivation temperature to 43 °C, however, made expression of Hsp33-M172S disadvantageous for E. coli as well (supplemental Fig. S4B). These results provided preliminary evidence that expression of Hsp33 variants lacking their tight post-translational redox control might be harmful to bacteria. Analysis of the respective cell lysates using SDS-PAGE revealed that all three Hsp33-expressing strains contained similar steady-state levels of Hsp33 (supplemental Fig. S4C). The Hsp33 expression levels in the presence of 50 μm IPTG were estimated by Western blot analysis to be ∼20-fold higher than the expression level of chromosomally encoded Hsp33 under non-stress conditions (data not shown). No obvious change in the general protein expression pattern (i.e. induction of the heat shock response) was observed (supplemental Fig. 4C). These results argued against the possibility that the growth defect observed in Hsp33-Y12E expressing strains is due to massive overexpression of a partially unfolded protein. When we analyzed and compared the soluble and insoluble protein population in these three strains, however, we detected a significant difference; a substantial proportion of Hsp33-Y12E protein was found to be insoluble and co-aggregated with a large number of different E. coli proteins (Fig. 6B). This co-aggregation led to a small but reproducible decrease in the overall amount soluble E. coli proteins. In contrast, both wild-type Hsp33 or Hsp33-M172S proteins remained mostly soluble at 37 °C and did not cause any protein aggregation. Cultivation of the Hsp33-M172S expressing strain at temperatures that activated the mutant protein in vitro (Fig. 2B) and caused a growth defect in vivo (supplemental Fig. S4B), however, also sequestered a large number of E. coli proteins into the insoluble protein pellet (supplemental Fig. S4D). These results suggest that in the absence of any post-translational regulation, activated Hsp33 binds to either newly synthesized or pre-existing cellular proteins, which may cause Hsp33 to co-aggregate with these proteins. It remains to be determined whether the formation of aggregates per se, or the sequestration of specific E. coli proteins necessary for optimal bacterial growth is responsible for the observed growth defect. In either case, these results provide a first clue as to why chaperone holdases, such as Hsp33, require tight functional regulation.