©1995 by The American Society for Biochemistry and Molecular Biology, Inc.
Exposure of Hydrophobic Surfaces on the Chaperonin GroEL Oligomer by Protonation or Modification of His-401 (*)

(Received for publication, December 12, 1994; and in revised form, January 27, 1995)

Don L. Gibbons Paul M. Horowitz (§)

From the Department of Biochemistry, University of Texas Health Science Center, San Antonio, Texas 78284-7760

ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
FOOTNOTES
ACKNOWLEDGEMENTS
REFERENCES

ABSTRACT

Hydrophobic exposure on the chaperonin GroEL is increased 6-10-fold after the protein is treated with the His-reactive reagent diethyl pyrocarbonate (DEP), or the solution pH is lowered to 5.5. The induced hydrophobic surfaces have the same 1,1`-bis(4-anilino)naphthalene-5,5`-disulfonic acid (bis-ANS) binding characteristics as unperturbed GroEL: a K 3.5 µM, a maximum intensity at 500 nm, and an average fluorescence lifetime of 8.0 ns. The pK for the pH-induced transition is 6.6, most likely attributable to the only histidine in GroEL, His-401, located in the intermediate domain. The modification of one histidine residue per monomer upon DEP treatment is supported by the correlation between the change in the absorbance at 242 nm for the N-carbethoxyhistidyl derivative and the increase in bis-ANS fluorescence. GroEL at pH 5.5 is tetradecameric and can capture urea-denatured rhodanese and release it as active enzyme. The GroEL-rhodanese complex is more stable to dissociation by 2.25 M urea than the complex formed at pH 7.8. We propose that His-401 is in a conformationally sensitive region such that protonation or modification can lead to increased exposure of hydrophobic surfaces capable of binding folding intermediates.


INTRODUCTION

The proper folding and transport of nascent polypeptide chains in vivo can be facilitated by the group of proteins termed molecular chaperones (1, 2). One class of chaperones, the chaperonins, has been found in all prokaryotes, mitochondria, and chloroplasts (3). The extensively studied chaperonin from Escherichia coli, GroEL, the homologue of the mitochondrial matrix protein hsp60 (heat shock protein with a M(r) = 60,000), has been shown to facilitate in vitro refolding of several chemically denatured proteins, including rhodanese (4, 5), ribulose-bisphosphate carboxylase/oxygenase (Rubisco) (6, 7), and citrate synthase (8). GroEL is a tetradecamer (14-mer) of 57.2-kDa subunits arranged as two stacked seven-membered rings (3, 9, 10). It has been shown that the range of possible substrates for this chaperonin is rather broad, with GroEL capable of binding about half of the soluble proteins from E. coli cell lysate with relatively high affinity (11). Such promiscuity brings into question the specific recognition motif, which now appears to involve in some cases, but is not necessarily restricted to, an amphipathic helix that may be stabilized or induced by binding GroEL (12-16). Despite studies supporting a model of GroEL interacting with substrate through multiple binding sites (17, 18), the mechanism of binding and functional release is still obscure, especially since the differing requirements for release indicate various degrees of interaction between chaperonin and substrate in the binary complexes(16) . Some proteins require only K and MgATP for release of functional enzyme from the binary complex, whereas others also require the co-chaperonin GroES (4-6, 19).

It has been proposed that the chaperonin binds the ``molten globule'' or ``compact intermediate'' form of substrate proteins, a fairly compact state displaying significant secondary structure, increased exposure of hydrophobic surfaces compared with the native protein, and lacking a rigid tertiary structure (4, 20). These hydrophobic surfaces have been suggested to mediate the binding between chaperonin and substrate(2) . However, according to quantitative bis-ANS (^1)binding studies(5) , there are very few available sites on unperturbed GroEL for such binding. Even in the papers discussing molten globule intermediates as substrates, it is assumed that increased binding of a hydrophobic probe by the complex is due entirely to exposure of surfaces on the substrate protein(4) , without any concomitant increase in the exposure of hydrophobic surfaces on GroEL. It has not been shown by direct biochemical evidence that GroEL has the necessary complementary hydrophobic surfaces for interaction with a folding intermediate of substrate.

The recently released crystal structure of a GroEL tetradecamer shows each monomer with an equatorial, intermediate, and apical domain (21). The authors attribute the lack of resolution for residues 222-248 in the apical domain to a region of probable flexibility. As shown by the companion mutagenesis study, single mutations of hydrophobic residues in this region and flanking it (Y199E, Y203E, F204E, L234E, L237E, L259S, V263S, and V264S) abolish peptide binding (22). Interestingly, the putative hydrophobic binding pocket defined by the crystal structure and mutagenesis work is not buried in the structure or apparently blocked by it but rather lines the entrance to the inner channel and should be readily accessible to binding by hydrophobic probes in solution. In fact, several studies visualize via electron microscopy or suggest based on biochemical evidence that GroEL binds substrate proteins in this channel (23-26). The intermediate domain is composed of two stretches of amino acids (134-190 and 377-408) which form three alpha-helices and three beta-strands that not only connect the equatorial and apical domains, but also make inter-monomer contacts between two highly conserved stretches of residues(21) . Mutagenesis of residues at several places in the intermediate domain (I150E, S151V, A152E, A383E, A405E, and A406E) shows significant effects on one or more properties of the chaperonin: folding of substrate, ATPase activity, substrate binding, or GroES binding. Thus, the intermediate and apical domains appear to have dynamic properties, critical to the interaction between GroEL and substrate, that are not revealed by the static crystal structure.

In this study, we show that GroEL displays a pH-dependent exposure of hydrophobic surfaces, as measured by the fluorescence of the hydrophobic probe bis-ANS. At pH 5.5, under conditions where the protein retains its tetradecameric structure, exposure of hydrophobic surfaces on GroEL can be induced to a level of 6-10-fold over that observed at pH 7.6. The pH dependence of the hydrophobic increase shows a pK 6.6. This value, in addition to experiments with the His-reactive reagent diethylpyrocarbonate, leads us to suggest that the single histidine in GroEL monomers, residue 401 in the intermediate domain, is in a region of the protein that has flexibility or is sensitive to structural perturbations that can account for the observed pH-triggered exposure of hydrophobic surfaces. Furthermore, binding of non-native rhodanese to GroEL at pH 5.5 occurs to produce a complex that is more stable to urea dissociation than a comparable complex at pH 7.8 described previously(17) .


MATERIALS AND METHODS

Reagents and Proteins

All reagents used were of analytical grade. bis-ANS and 1-anilinonaphthalene-8-sulfonic acid (1,8-ANS) were obtained from Molecular Probes (Junction City, OR).

Rhodanese was prepared as described previously (27) and stored at -70 °C as a crystalline suspension in 1.8 M ammonium sulfate. Rhodanese concentrations were determined using A = 1.75 (28) and a molecular mass of 33 kDa (29). Rhodanese activity was assayed using a colorimetric method based on the absorbance at 460 nm of the complex formed between the reaction product, thiocyanate, and ferric ion(28) . The chaperonin, GroEL, was purified from lysates of E. coli cells bearing the multicopy plasmid pGroESL which were grown at 37 °C (30). The purification was a modified version of previously published protocols (31), except that the Mono Q and hydroxylapatite columns were excluded, but including the batch treatment with Affi-Gel blue as described in(19) . After purification, GroEL was dialyzed against 50 mM Tris-HCl, pH 7.5, and 1 mM dithiothreitol, then made 10% (v/v) in glycerol, rapidly frozen in liquid nitrogen, and stored at -70 °C. The protomer concentration of GroEL was measured using the bicinchoninic acid protein assay (Pierce) according to the procedure recommended by the manufacturer.

Chaperonin-assisted Refolding

Rhodanese was denatured in 8 M urea, 200 mM sodium phosphate buffer, pH 7.4, and 1 mM beta-mercaptoethanol for 1 h, 25 °C, at a protein concentration of 90 µg/ml. Unfolded rhodanese was diluted (109 nm final concentration) into 50 mM Tris-HCl, pH 7.8, or 5 mM citrate, pH 5.5, 200 mM beta-mercaptoethanol, 50 mM sodium thiosulfate, and GroEL at a concentration of 2.5 µM (monomer). To discharge rhodanese from the binary complex and assess recoverable activity, urea (2.25 M final concentration) was added to samples(17) , which were then incubated 30 min at room temperature and assayed for rhodanese activity as described above. For samples where the complex was formed at pH 5.5, an aliquot of sample was shifted into a urea solution (2.25 M final concentration) containing 200 mM beta-mercaptoethanol, 50 mM sodium thiosulfate, and buffered with Tris-HCl (50 mM final concentration) to hold the pH at 7.8. After 30 min the rhodanese assay was performed on this sample to assay recoverable activity.

Fluorescence Spectroscopy

Fluorescence of bis-ANS or 1,8-ANS was measured using an SLM model 500C fluorometer at an excitation wavelength of 360 nm. For acid titration curves, microliter aliquots of 1 mM HCl were added to a sample containing 1 µM GroEL and 10 µM bis-ANS, with a starting pH as indicated in figures. Upon each addition of HCl, an emission spectrum was taken and the pH measured. For DEP experiments the same procedure was followed, except that GroEL was pretreated with the indicated concentrations of DEP using a 10 mM stock made by freshly diluting concentrated DEP in ethanol. It should be noted here that some experiments (titration curves) were performed using HCl to achieve certain pH values and other experiments (ultracentrifugation and chaperonin-assisted refolding) were performed using citrate buffer. However, no significant difference in maximum fluorescence was seen between the two methods.

Measurement of Fluorescence Lifetimes

Fluorescence lifetimes for bis-ANS were measured in solutions containing 10 µM GroEL, 50 µM bis-ANS, and either 50 mM Tris-HCl, pH 7.8, or HCl to pH 5.5. Lifetimes were derived from phase-modulation measurements (32) on an SLM 48000 fluorometer (SLM Instruments, Inc.). The temperature of the samples was maintained at 25 ± 0.02 °C by using a circulating water bath (RTE-5, Neslab) and a jacketed cell holder. Glycogen solutions were used as zero lifetime standards. Samples were excited at 360 nm, and emission was monitored at 500 nm using an interference filter. For each sample, the phase shift and demodulation were measured as a function of modulation frequency at 2, 4, 8, 16, 32, and 64 MHz. Fluorescence lifetimes were calculated from these data using software supplied by SLM, which employs the modified analyses described by Brent (33) in making the computations.

Sedimentation Analysis

Samples of GroEL (0.44 µM 14-mer) in 50 mM Tris-HCl, pH 7.8, or 50 mM citrate, pH 5.5, were subjected to sedimentation velocity analysis in a Beckman XL-A analytical ultracentrifuge. The runs were performed using 12-mm double sector cells in a four-hole Ti-60 rotor. The temperature was kept constant at 25 °C. The solutions had an A = 0.6-1.0, and the rotor speed for different runs was either 20,000 or 27,000 rpm. The scans were analyzed by the method of van Holde and Weischet (34) using the UltraScan ultracentrifuge data collection and analysis program (B. Demeler, Missoula, MT). Measured sedimentation coefficients were corrected to s(w) values, corresponding to a solvent with a viscosity and density of water at 20 °C.

Absorbance Spectroscopy

Spectrophotometric determinations and recording of difference absorbance spectra were carried out with a Shimadzu UV260 double-beam recording spectrophotometer or a Varian Cary 219 double-beam recording spectrophotometer. Diethylpyrocarbonate was freshly diluted with cold ethanol to 20 mM for each experiment. Protein samples containing 16.6 µM GroEL in 100 mM NaH(2)PO(4), pH 7.0, were treated with DEP (either 0.25 mM or 0.5 mM final concentration) at 25 °C. The modification of GroEL by DEP was initially followed by taking spectra between 220 and 300 nm. Since the most significant change was observed at 242 nm, the rate and extent of the N-carbethoxyhistidine formation were followed by measuring the difference in absorbance at 242 nm between the treated enzyme and a control enzyme treated with an equal volume of ethanol. The final concentration of ethanol in both samples was < 3.0%. The number of histidine residues modified was calculated using a value of 3200 M cm for the molar absorptivity difference for N-carbethoxyhistidine at 242 nm (35) .

Reaction mixtures with diethylpyrocarbonate, displaying maximal absorbance change at 242 nm, were treated with hydroxylamine adjusted to pH 7 (0.5 M final concentration). The extent and rate of the reaction with NH(2)OH were followed spectrophotometrically, either by taking spectra between 220 and 300 nm or by following the absorbance change at 242 nm.


RESULTS

Binding of the Hydrophobic Probe Bis-ANS Increases Dramatically under Conditions where the pH of the Solutions Containing GroEL Is Lowered or the Protein Is Treated with DEP

Fig. 1shows the fluorescence of bis-ANS upon binding to GroEL at different pH values (filled circles). These results are representative of acid titrations that consistently produced a 6-10-fold increase in fluorescence between pH 7.6 and pH 5.5. The same results were seen when the hydrophobic probe 1,8-ANS was used instead of bis-ANS (not shown). The effect was reversible as evidenced by the fact that the curve could be retraced in reverse by starting with GroEL at pH 5.5 and titrating with NaOH (open squares). The approximate pK(a) for this transition is 6.6, which is within the range of pK(a) values for the imidazole side chain of histidine. According to the amino acid sequence for GroEL, there is only one His, located at position 401(3) , which is in the intermediate domain in the crystal structure(21) .


Figure 1: Bis-ANS fluorescence of DEP-treated and untreated GroEL at different pH values. A solution with 1 µM GroEL was treated with 0.25 mM DEP at room temperature for 1 h (filled diamonds) or with 0.50 mM DEP at room temperature for 50 min (open circles). bis-ANS (final concentration, 10 µM) was added to the treated protein, and the fluorescence at 500 nm was measured with excitation at 360 nm. Microliter aliquots of 1 mM HCl were used to adjust the pH of the solution between 7.8 and 4.5. At each pH value the fluorescence intensity was read and the pH, as shown, was measured. Filled circles, a solution with 1 µM GroEL (untreated) and 10 µM bis-ANS was titrated with HCl between pH 7.8 and 4.5. As above, the pH was measured at each point where the fluorescence was measured. Open squares, a solution at pH 5.5, containing the same concentrations of GroEL and bis-ANS used above, was titrated with NaOH to pH 7.6.



To further test the possibility that the observed pH-dependent exposure of hydrophobic surfaces was due to the protonation of the His imidazole group, a His-reactive reagent was used to pretreat GroEL before acid titration. DEP reacts with His side chains to produce an N-carbethoxyhistidyl derivative (35-37). The top curve in Fig. 1shows the titration of GroEL that had been pretreated with 0.25 mM DEP at room temperature for 1 h (filled diamonds). The approximate maximum intensity of bis-ANS fluorescence achievable upon addition of acid (e.g. the intensity at pH 5.5) was not affected, but the modified protein displayed considerably more hydrophobic surfaces at neutral pH. In fact, when GroEL was pretreated with 0.5 mM DEP at room temperature for 50 min, prior to the acid titration analysis, 97.3% of the maximum fluorescence seen at pH 5.5 was seen at neutral pH (Fig. 1, open circles). Thus, modification with DEP abolished the observed pH-dependent change in bis-ANS fluorescence described above.

Reaction of DEP with residues other than histidine, especially lysine or tyrosine, has been demonstrated (35, 38). However, since the reaction requires an unprotonated nucleophile, modification of His is the most likely explanation given the solution conditions used here. To support this, we carried out several experiments to verify that the modification was occurring at His-401. First, the absorbance of the reaction was monitored. As shown in Fig. 2A, the result was an increase in absorbance at 242 nm as has been previously demonstrated for DEP modification of histidine in other proteins(35, 36, 37, 38) , whereas there was no significant change at any other wavelength, including 278 nm (the wavelength at which modified tyrosines absorb). Second, hydroxylamine was used to test for the reversibility of DEP modification. Melchior and Fahrney (39) found that 0.5 M NH(2)OH, adjusted to pH 7, removes N-carbethoxy groups from imidazole. In the present case, absorbance spectroscopy showed that hydroxylamine completely reversed the modification by DEP (data not shown). Since reversal has been shown to be possible with only histidyl and tyrosyl derivatives, but not with other modified residues(35, 38) , and since tyrosine does not appear to account for the modification observed here, we take this as evidence of a His modification. It is also evidence that the His residue is only singly modified and has not formed a derivative that is modified at both imidazole nitrogens, which is not reversible by treatment with NH(2)OH(35) .


Figure 2: A, absorbance spectra of DEP-modified GroEL. A solution of 16.6 µM GroEL in 100 mM NaH(2)PO(4), pH 7, was treated with 0.5 mM DEP at room temperature and the difference absorbance spectra were taken at various times. Representative curves are shown at 40 s, 105 s, 5 min, and 7 min 40 s (starting from bottom). B, relationship between fluorescence of DEP-modified GroEL and the number of histidine residues modified. Filled circles, the absorbance and fluorescence of solutions containing 16.6 or 1 µM GroEL, respectively, 100 mM NaH(2)PO(4), pH 7, and 0.5 mM DEP were followed for 30 min. For various time points, the fluorescence is plotted versus number of His residues modified (based on absorbance, using Delta = 3200 M cm). The solid line is a best fit to the first seven data points, extrapolated to 100% fluorescence. The extrapolated value for the number of His residues modified/monomer GroEL at 100% fluorescence is 1.19.



Since DEP modification produced an increase in bis-ANS fluorescence and in absorbance at 242 nm, the kinetics for the reaction were followed by both absorbance and fluorescence spectroscopy. In each case the data were best fit by exponential curves, displaying pseudo-first order kinetics with a theoretical A(max) = 0.10 and k = 0.004 s; a F(max) = 5.0 and k = 0.009 s. Fig. 2B shows the results after the data was treated as described previously (35, 36, 37) . The curve relating fluorescence to the number of His residues modified (based on absorbance) extrapolates at 100% fluorescence to 1.19 His residues modified per GroEL monomer, which is consistent with the interpretation that one histidine per monomer was modified.

The Nature of Bis-ANS Binding to GroEL Does Not Change Significantly at Low pH

To analyze whether or not the increased bis-ANS fluorescence at low pH was due to a change in the number of available sites on the GroEL tetradecamer or instead due to an increased binding affinity between protein and probe, bis-ANS titrations were performed. Fig. 3shows the two bis-ANS titration curves for GroEL at pH 5.5 and pH 7.8. According to parameters for the fitted curves, the I(max) for GroEL is 0.47 at pH 7.8 and 4.9 at pH 5.5. At pH 7.8 the K(d) = 3.5 µM, and at pH 5.5 the K(d) = 3.9 µM. These numbers are consistent with a large increase (approximately 10 times) in the number or size of hydrophobic sites that are presented at low pH and a small change in the relative binding affinity.


Figure 3: Binding of bis-ANS to GroEL at pH 5.5 and pH 7.8. Separate samples of 1 µM GroEL at pH 5.5 (as adjusted by HCl), with increasing amounts of bis-ANS, were excited at 360 nm, and the fluorescence intensity was read at 500 nm (upper curve, squares). The curves were generated using a nonlinear least squares procedure in the software program, PS Plot (Poly Software International, Salt Lake City, UT). The K and I(max), as taken from the fitted curve, are 3.9 and 4.9, respectively. Separate samples were prepared and fluorescence measured as above, except that a 50 mM Tris-HCl, pH 7.8, buffer was used (lower curve, circles). The K and I(max) for this curve (as taken from the fitted curve) are 3.5 and 0.47, respectively.



Consistent with this is the fact that the wavelength of maximum fluorescence intensity did not change significantly between pH 7.8 and pH 5.5 (data not shown). The wavelength of bis-ANS fluorescence shifted 60 nm upon binding to GroEL from aqueous solution, whereas the maximum shift upon pH change was <5 nm. This suggests that the hydrophobic nature of the binding sites did not change significantly, since it has been shown that the wavelength maximum of bis-ANS fluorescence is sensitive to the hydrophobic character of the binding site (40, 41).

To confirm that these observed changes in fluorescence intensity were in fact due to an increase in the number of bis-ANS molecules bound, and not due to different fluorescence quantum yields at the different pH values, lifetime measurements were made at the two extremes (pH 7.8 and 5.5). The decay at each pH was best fit by three components. At pH 7.8: 1 = 1.78 ns, alpha1 = 0.188; 2 = 6.57 ns, alpha2 = 0.794; and 3 = 130.24 ns, alpha3 = 0.0185. At pH 5.5: 1 = 1.51 ns, alpha1 = 0.230; 2 = 6.93 ns, alpha2 = 0.751; and 3 = 128.45 ns, alpha3 = 0.0191. The reduced ^2 values for the data at pH 7.8 and 5.5 were 2.94 and 0.904, respectively, consistent with the fact that the sampling error decreases as the signal increases, and the reduced ^2 approaches 1. Thus, the average lifetimes were 7.96 ns and 8.00 ns at pH 5.5 and 7.8, respectively. The average lifetimes and the fractional contributions of the individual components indicate that the fluorescence lifetime did not vary with pH. Therefore, there was no significant change in the quantum yield of the bis-ANS as a function of pH so that an increase in the area of hydrophobic surface best accounts for the observed increase in fluorescence.

The GroEL Molecule Remains a Tetradecamer at Low pH

We were concerned that such a large increase in the area of hydrophobic exposure might be due to the GroEL molecule becoming denatured or disassembling into monomers upon the change in pH. To test for this possibility, ultracentrifuge analyses, as described in (17) , were carried out on samples of GroEL at pH 7.8 and 5.5 (data not shown). In both cases the chaperonin population was homogenous and displayed an s(w) value of approximately 23 S, consistent with previously reported values for the intact tetradecamer (17) . Thus, no significant shift in the s(w) value was observed that would indicate either denaturation or disassembly.

At Low pH, GroEL Retains Its Ability to Bind the Non-native State of the Substrate Protein Rhodanese

The simple exposure of more hydrophobic surface area on the GroEL molecule at pH 5.5 does not necessarily mean that the chaperonin is in a state that can productively bind non-native rhodanese and release reactivated protein. We used the colorimetric rhodanese activity assay to test whether GroEL retained its ability to capture non-native rhodanese and then release reactivated protein when incubated at 2.25 M urea(17) . Control experiments showed that urea-unfolded rhodanese, incubated at pH 5.5 for 5 min without GroEL, then shifted to pH 7.8, showed only 2.5% recoverable activity after a 30-min incubation in 2.25 M urea. We were able to take advantage of this loss of recoverable rhodanese activity at pH 5.5 by designing the experiment so that the complex was formed at pH 5.5, but the samples were shifted to pH 7.8 to check for recoverable rhodanese activity (as outlined under ``Materials and Methods''). Any measured activity would be attributable to rhodanese that had bound to GroEL and was therefore protected from inactivation. When unfolded rhodanese was incubated in the presence of GroEL at pH 5.5 for 5 min prior to shifting pH and performing the urea discharge assay, 23.5% recoverable rhodanese activity was obtained. As a further control, unfolded rhodanese was incubated at pH 5.5 for 5 min, without GroEL, then shifted into a pH 7.8 solution containing GroEL and incubated in 2.25 M urea for 30 min. In this case only 8.5% recoverable activity was obtained, suggesting that 5% of the rhodanese inactivated at pH 5.5 could be reactivated after switching into a pH 7.8 buffer containing GroEL. From these experiments it seems clear that GroEL retains its ability to bind the non-native form of rhodanese at low pH in a manner that protects the rhodanese from inactivation and allows for reactivation, in a pH 7.8 solution, upon urea-induced discharge from the complex.

The GroEL-Rho(I) Complex formed at pH 7.8 Is Stable to Dissociation upon Lowering the pH to 5.5

The binding in the binary complexes formed at pH 7.8 and pH 5.5 were compared by again taking advantage of the fact that non-native rhodanese is inactivated at the lower pH. GroEL-Rho(I) complexes formed at pH 7.8 for 5 min and then either maintained at pH 7.8 or switched to pH 5.5 for 20 min had approximately the same recoverable rhodanese activity, 19 and 23%, respectively. The rhodanese was not inactivated in the switch to pH 5.5. Instead, the rhodanese activity was retained as in the experiment above where GroEL was present in the low pH incubation. This indicates that the complex formed at pH 7.8 is maintained when the pH is lowered and therefore that binding at low pH is at least as strong as at pH 7.8.

The Low pH GroEL-Rho(I) Complex Is More Stable to Dissociation by 2.25 M Urea Than the Complex at pH 7.8

To directly compare whether the complex formed at low pH is more stable than the one formed at pH 7.8, the stabilities of the complexes against urea dissociation were compared. We confirmed that the best urea-induced recovery of rhodanese activity from a complex formed at pH 7.8 (38%) was achievable by a 2-h incubation of the complex in 2.25 M urea(17) . Incubations longer than 2 h did not yield any more recoverable activity ( (17) and data not shown). Therefore, if complexes formed at pH 5.5 are equally as stable to 2.25 M urea, then after 2 h of incubation they should show no additional discharge of reactivable rhodanese. As a control, the complex formed at pH 5.5 was incubated for 2 h and showed a yield of 23% recoverable rhodanese activity (comparable with the pH 5.5 complex noted above formed during a 5-min incubation), which was lower than the pH 7.8 maximum recovery, because some background of unbound rhodanese spontaneously refolded at 7.8, but was inactivated at 5.5. In contrast, the complex formed at pH 5.5 and pretreated with 2.25 M urea for 2 h (at pH 5.5) yielded only 11.5% recoverable activity, a 50% reduction. In summary, the results show that after a 2-h incubation in 2.25 M urea, the low pH complex retained 50% of its bound and reactivable rhodanese, conditions where the pH 7.8 complex discharged its maximum amount of recoverable rhodanese. This demonstrates the greater stability of the pH 5.5 binary complex in urea.

As a control to verify that this apparent difference in urea stability was due to the stability of the complex and not due to the difference in urea effectiveness at different pH values, possibly because of differential cyanylation, the stability of the GroEL tetradecamer at the two pH values against urea dissociation was examined as described previously(50) . The results were consistent with the fact that the low pH form shows higher fluorescence, even in 0 M or low concentrations of urea (not shown), and were also consistent with an unfolding transition and decreased fluorescence between approximately 2 and 4 M urea. Based on these results, there was no significant difference between the effectiveness of urea at pH 5.5 and pH 7.8, meaning that the greater stability of the GroEL-Rho(I) complex was not due to a decreased effectiveness of the chaotrope.


DISCUSSION

The mechanism for the capture of folding intermediates by GroEL and the release of functional proteins is not well understood, although pieces of the scheme are coming together from different lines of investigation. There is now good evidence that hydrophobicity is a necessary characteristic for the substrate (42-44) and that in some cases it must be present in an amphiphilic context(12, 13) . Additionally, based on the crystal structure and mutant studies (21, 22) , there is now a better idea of where this complementary hydrophobic binding site might exist on the chaperonin. The challenge is to incorporate this structural information with current models based on the dynamics and biochemistry of the system. To bind substrate proteins in non-native, molten globule states, GroEL must be able to display significantly more hydrophobic surfaces than has previously been demonstrated(5) . Additionally, the generality of this kind of interaction suggests that exposure of these ``sticky'' surfaces would somehow be regulated, with exposure predicated on induced fit of the non-native substrate or a triggered response, such that nonproductive interactions do not interfere with the chaperonin. Such an induced response has been proposed for another chaperone, SecB (45-47), is consistent with previous reports for folding of beta-lactamase by GroEL (48) and is reasonable considering the structural transition in GroEL recently shown by Hansen and Gafni (10, 49).

That GroEL displays the same significant enhancement of bis-ANS fluorescence when the pH is lowered or when treated with DEP suggests very similar structural changes in the tetradecamer under the two conditions. It is important to realize that this large conformational change in the oligomer occurs simply with the protonation or N-carbethoxylation of His-401. The His residue in question falls in the middle of a highly conserved, extended alpha-helix in the intermediate domain which, at its N-terminal end, makes contacts to a highly conserved region of the adjacent monomer(363-378) in the tetradecamer(21) . Based on the amino acid sequence, this alpha-helix has considerable hydrophobic character in the region around 401 (3, 50), and although we cannot say where the side chain lies, until the three-dimensional coordinates are released, it is reasonable that it is buried in a hydrophobic region of the protein such that protonation perturbs the native structure. Introducing a charge into a hydrophobic environment might be expected to have dramatic conformational consequences, explaining the increased exposure of hydrophobic surfaces. Alternatively, the arrangement of side chains may be such that N-carbethoxylation causes a steric clash that distorts the structure of the intermediate domain. We are not suggesting a mechanistically important role for His-401, since published sequences show that the His is not a highly conserved residue (51), and since not all site-directed mutations at this position are deleterious (see below). Rather, our contention is that His-401 is in a structurally sensitive region where small perturbations, especially due to the introduction of charge, can produce large increases in exposure of hydrophobic surfaces. This would be consistent with the idea that the intermediate domain serves as a hinge region within a monomer and/or as a connecting region that allows ``communication'' between adjacent monomers in the tetradecamer(21) . In fact, the mutations made by Fenton et al.(22) in the intermediate domain show results that support a similar conclusion.

It is interesting that neither the His Phe mutant made by Fenton et al.(22) nor a similarly uncharged substitution we have made (His Leu) showed any non-native characteristics, when evaluated according to urea stability, ATP-dependent chaperonin activity, and ability to arrest rhodanese refolding. (^2)This is in contrast to another mutant, where a constitutive negative charge was introduced (His Asp), which yielded little soluble, recoverable GroEL in the supernatant after centrifugation of the E. coli cell lysates (where GroEL is normally found during purification). The protein was instead found aggregated in inclusion bodies.^2 These seemingly disparate results make sense within the confines of the present model. If exposure of hydrophobic surfaces on GroEL is affected by flexibility in the intermediate domain, then mutation to a constitutive negative charge at residue 401 could significantly change the structure in that region. The result would be a protein that aggregates in vivo due to the exposure of large amounts of hydrophobic surface. It is interesting to note here that DEP-modified GroEL, which constitutively displays significant hydrophobic surface, can arrest the spontaneous refolding of denatured rhodanese, but will not release the active enzyme, even upon addition of MgATP. (^3)Similarly, some of the single mutations of residues in the intermediate domain (I150E, S151V, A383E, and A405E) result in a protein that lacks the ability to release substrate or has a decrease in release(22) .

Other recent work (50, 52) shows that various salts and polycations can induce exposure of hydrophobic surfaces on GroEL without dissociation of the tetradecamer or can increase the affinity between GroEL and substrate. Based on that work and the present results, it is tempting to speculate that exposure of functional hydrophobic surfaces on GroEL can be triggered by ionic perturbations. This would be in keeping with the earlier results demonstrating that GroEL preferentially binds amphiphilic alpha-helices that are positively charged (12, 13, 53) and makes sense, given the recent results suggesting that divalent cations cause structural changes in GroEL that increase the rates of cross-linking between monomers (54).


FOOTNOTES

*
This research was supported by Research Grants GM25177 and ES05729 from the National Institutes of Health and Welch Grant AQ 723 (to P. M. H.). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore by hereby marked ``advertisement'' in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

§
To whom correspondence should be addressed: Dept. of Biochemistry, University of Texas Health Science Center, 7703 Floyd Curl Dr., San Antonio, TX 78284-7760. Tel.: 210-567-3737; Fax: 210-567-6595.

(^1)
The abbreviations used are: bis-ANS, 1,1`-bis(4-anilino)naphthalene-5,5`-disulfonic acid; DEP, diethyl pyrocarbonate; 1,8-ANS, 1-anilinonaphthalene-8-sulfonic acid.

(^2)
G.-X., Luo, personal communication.

(^3)
S. Hua, personal communication.


ACKNOWLEDGEMENTS

We thank Dr. Boris Gorovits for assistance with the lifetime measurements.


REFERENCES

  1. Gething, M.-J., and Sambrook, J. (1992) Nature 355, 33-45 [CrossRef][Medline] [Order article via Infotrieve]
  2. Ellis, R. J., van der Vies, S. M., and Hemmingsen, S. M. (1990) Biochem. Soc. Symp. 55, 145-153
  3. Hemmingsen, S. M., Woolford, C., van der Vies, S. M., Tilly, K., Dennis, D. T., Georgopoulos, C. P., Hendrix, R. W., and Ellis, R. J. (1988) Nature 333, 330-334 [CrossRef][Medline] [Order article via Infotrieve]
  4. Martin, J., Langer, T., Boteva, R., Schramel, A., Horwich, A. L., and Hartl, R.-U. (1991) Nature 352, 36-42 [CrossRef][Medline] [Order article via Infotrieve]
  5. Mendoza, J. A., Rogers, E., Lorimer, G. H., and Horowitz, P. M. (1991) J. Biol. Chem. 266, 13044-13049 [Abstract/Free Full Text]
  6. Goloubinoff, P., Christeller, J. T., Gatenby, A. A., and Lorimer, G. H. (1989) Nature 342, 884-889 [CrossRef][Medline] [Order article via Infotrieve]
  7. Viitanen, P. V., Lubben, T. H., Reed, V., Goloubinoff, P., O'Keefe, D. P., and Lorimer, G. H. (1990) Biochemistry 29, 5665-5671 [Medline] [Order article via Infotrieve]
  8. Buchner, J., Schmidt, M., Fuchs, M., Jaenicke, R., Rudolph, R., Schmid, F. X., and Kiefharber, T. (1991) Biochemistry 30, 1586-1591 [Medline] [Order article via Infotrieve]
  9. Hendrix, R. W. (1979) J. Mol. Biol. 129, 375-392 [Medline] [Order article via Infotrieve]
  10. Hansen, J. E., and Gafni, A. (1993) J. Biol. Chem. 268, 21632-21636 [Abstract/Free Full Text]
  11. Viitanen, P. V., Gatenby, A. A, and Lorimer, G. H. (1992) Protein Sci. 1, 363-369 [Abstract/Free Full Text]
  12. Landry, S. J., and Gierasch, L. M. (1991) Biochemistry 30, 7359-7362 [Medline] [Order article via Infotrieve]
  13. Landry, S. J., Jordan, R., McMacken, R., and Gierasch, L. M. (1992) Nature 355, 455-457 [CrossRef][Medline] [Order article via Infotrieve]
  14. Zahn, R., Spitzfaden, C., Ottiger, M., Wuthrich, K., and Pluckthun, A. (1994) Nature 368, 261-265 [CrossRef][Medline] [Order article via Infotrieve]
  15. Schmidt, M., and Buchner, J. (1992) J. Biol. Chem. 267, 16829-16833 [Abstract/Free Full Text]
  16. Schmidt, M., Bucheler, U., Kaluza, B., and Buchner, J. (1994) J. Biol. Chem. 269, 27964-27972 [Abstract/Free Full Text]
  17. Mendoza, J. A., Demeler, B., and Horowitz, P. M. (1994) J. Biol. Chem. 269, 2447-2451 [Abstract/Free Full Text]
  18. Mendoza, J. A., Butler, M. C., and Horowitz, P. M. (1992) J. Biol. Chem. 267, 24648-246-54 [Abstract/Free Full Text]
  19. Schmidt, M., Buchner, J., Todd, M. J., Lorimer, G. H., and Viitanen, P. V. (1994) J. Biol. Chem. 269, 10304-10311 [Abstract/Free Full Text]
  20. Ptitsyn, O. B., Pain, R. H., Semisotnov, G. V., Zerovnik, E., and Razgulyaev, O. I. (1990) FEBS Lett. 262, 20-24 [CrossRef][Medline] [Order article via Infotrieve]
  21. Braig, K., Otwinowski, Z., Hedge, R., Boisvert, D. C., Joachimiak, A., Horwich, A. L., and Sigler, P. B. (1994) Nature 371, 578-586 [CrossRef][Medline] [Order article via Infotrieve]
  22. Fenton, W. A., Kashi, Y., Furtak, K., and Horwich, A. L. (1994) Nature 371, 614-619 [CrossRef][Medline] [Order article via Infotrieve]
  23. Chen, S., Roseman, A. M., Hunter, A. S., Wood, S. P., Burston, S. G., Ranson, N. A., Clarke, A. R., and Saibil, H. R. (1994) Nature 371, 261-264 [CrossRef][Medline] [Order article via Infotrieve]
  24. Ishii, N., Taguchi, H., Sasabe, H., and Yoshida, M. (1994) J. Mol. Biol. 236, 691-696 [CrossRef][Medline] [Order article via Infotrieve]
  25. Langer, T., Pfeifer, G., Martin, J., Baumeister, W., and Hartl, F.-U. (1992) EMBO J. 11, 4757-4765 [Abstract]
  26. Braig, K., Simon, M., Furuya, F., Hainfeld, J. F., and Horwich, A. L. (1993) Proc. Natl. Acad. Sci. U. S. A. 90, 3978-3982 [Abstract]
  27. Miller, D. M., Kurzban, G. P., Mendoza, J. A., Chirgwin, J. M., Hardies, S. C., and Horowitz, P. M. (1992) Biochim. Biophys. Acta 1121, 286-292 [Medline] [Order article via Infotrieve]
  28. Sorbo, B. H. (1953) Acta Chem. Scand. 7, 1129-1136
  29. Ploegman, J. H., Drent, G., Kalk, K. H., Hol, W. G. J., Heinrikson, R. L., Keim, P., Weng, L., and Russel, J. (1978) Nature 273, 124-129 [Medline] [Order article via Infotrieve]
  30. Goloubinoff, P., Gatenby, A. A., and Lorimer, G. H. (1989) Nature 337, 44-47 [CrossRef][Medline] [Order article via Infotrieve]
  31. Todd, M. J., Viitanen, P. V., and Lorimer, G. H. (1993) Biochemistry 32, 8560-8567 [Medline] [Order article via Infotrieve]
  32. Lakowicz, J. R. (1983) Principles of Fluorescence Spectroscopy , Plenum, New York
  33. Brent, R. P. (1973) Algorithms for Minimization without Derivatives , Prentice-Hall, New York
  34. van Holde, K. E., and Weischet, W. O. (1978) Biopolymers 17, 1387-1403
  35. Miles, E. W. (1977) Methods Enzymol. 47, 431-442 [Medline] [Order article via Infotrieve]
  36. Dominici, P., Tancini, B., and Voltattorni, C. B. (1985) J. Biol. Chem. 260, 10583-10589 [Abstract/Free Full Text]
  37. Kumagai, H., Utagawa, T., and Yamada, H. (1975) J. Biol. Chem. 250, 1661-1667 [Abstract]
  38. Burstein, Y., Walsh, K. A., and Neurath, H. (1974) Biochemistry 13, 205-210 [Medline] [Order article via Infotrieve]
  39. Melchior, W. B., Jr., and Fahrney, D. (1970) Biochemistry 9, 251-258 [Medline] [Order article via Infotrieve]
  40. Rosen, C. G., and Weber, G. (1969) Biochemistry 8, 3915-3920 [Medline] [Order article via Infotrieve]
  41. Turner, D. C., and Brand, L. (1968) Biochemistry 7, 3381-3390 [Medline] [Order article via Infotrieve]
  42. Zahn, R., Axmann, S. E., Rucknagel, K.-P., Jaeger, E., Laminet, A. A., and Pluckthun, A. (1994) J. Mol. Biol. 242, 150-164 [CrossRef][Medline] [Order article via Infotrieve]
  43. Zahn, R., and Pluckthun, A. (1994) J. Mol. Biol. 242, 165-174 [CrossRef][Medline] [Order article via Infotrieve]
  44. Staniforth, R. A., Cortes, A., Burston, S. G., Atkinson, T., Holbrook, J. J., and Clarke, A. R. (1994) FEBS Lett. 344, 129-135 [CrossRef][Medline] [Order article via Infotrieve]
  45. Randall, L. L., Topping, T. B., and Hardy, S. J. S. (1990) Science 248, 860-863 [Medline] [Order article via Infotrieve]
  46. Hardy, S. J. S., and Randall, L. L. (1991) Science 251, 439-443 [Medline] [Order article via Infotrieve]
  47. Randall, L. L. (1992) Science 257, 241-245 [Medline] [Order article via Infotrieve]
  48. Zahn, R., and Pluckthun, A. (1992) Biochemistry 31, 3249-3255 [Medline] [Order article via Infotrieve]
  49. Hansen, J. E., and Gafni, A. (1994) J. Biol. Chem. 269, 6286-6289 [Abstract/Free Full Text]
  50. Horowitz, P. M., Hua, S., and Gibbons, D. L. (1995) J. Biol. Chem. 270, 1535-1542 [Abstract/Free Full Text]
  51. Mazodier, P., Guglielmi, G., Davies, J., and Thompson, C. J. (1991) J. Bacteriol. 173, 7382-7386 [Medline] [Order article via Infotrieve]
  52. Okazaki, A., Ikura, T., Nikaido, K., and Kuwajima, K. (1994) Structural Biol. 1, 439-446
  53. Rosenberg, H. F., Ackerman, S. J., and Tenen, D. G. (1993) J. Biol. Chem. 268, 4499-4503 [Abstract/Free Full Text]
  54. Azem, A., Diamant, S., and Goloubinoff, P. (1994) Biochemistry 33, 6671-6675 [Medline] [Order article via Infotrieve]

©1995 by The American Society for Biochemistry and Molecular Biology, Inc.