Does fusion of domains from unrelated proteins affect their folding pathways and the structural changes involved in their function? A case study with the diphtheria toxin T domain

Alexandre Chenal1, Philippe Nizard1, Vincent Forge2, Martine Pugnière3, Marie-Odile Roy3, Jean-Claude Mani3, Florent Guillain2 and Daniel Gillet1,4

1 Département d'Ingénierie et d'Etudes des Protéines, CEA-Saclay, 91191 Gif sur Yvette cedex, 2 Biophysique Moléculaire et Cellulaire, UMR 5090, Département de Biologie Moléculaire et Structurale, CEA-Grenoble,17 rue des Martyrs, 38054 Grenoble cedex 9 and 3 CNRS-UMR 5094, Faculté de Pharmacie, 15 avenue C. Flahault, 34093 Montpellier cedex 5, France


    Abstract
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
We investigated whether the structural and functional behaviors of two unrelated protein domains were modified when fused. The IgG-binding protein ZZ derived from staphylococcal protein A was fused to the N- and/or C-terminus of the diphtheria toxin transmembrane domain (T). T undergoes a conformational change from a soluble native state at neutral pH to a molten globule-like state at acidic pH, leading to its interaction with membranes. We found that this molten globule state was not connected to the GdnHCl-induced unfolding pathway of T. The pH-induced transition of T, and also the unfolding of T and ZZ at neutral and acidic pH, were unchanged whether the domains were isolated or fused. The position of ZZ, however, influenced the solubility of T near its pKi. SPR measurements revealed that T has a high affinity for membranes, isolated or within the fusion proteins (KD< 10-11 M). This work shows that in the case of T and ZZ, the fusion of protein domains with different stabilities does not alter the structural changes involved in folding and function. This supports the use of T as a soluble membrane anchor.

Keywords: diphtheria toxin/fusion protein membrane anchor/protein A/protein folding


    Introduction
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Protein fusion technology has become an essential tool in many fields of basic science and biotechnology. It allows the creation of new proteins, which have the functional properties of the protein domains that are linked together. Recent data suggest that protein domains may accept the insertion of another protein without major consequences for the folding and function of both fusion partners (Collinet et al., 2000Go). In contrast, the addition of a single N-terminal residue may destabilize a protein (Ishikawa et al., 1998Go; Chaudhuri et al., 1999) and it has been shown that adjacent domains in a protein may be more flexible within the protein than alone (Engen et al., 1999Go). Still, few examples have been described in detail. In particular, little is known about the relations between domains within non-natural combinations when one of the fusion partners undergoes important structural changes related to its function.

One of the applications of protein fusion technology is the development of membrane anchors for soluble proteins. The anchoring of soluble proteins to membranes has useful applications, including the attachment of antibodies to cells (Nizard et al., 1998Go, 2001Go; de Kruif et al., 2000Go) to create new interaction sites at their surface and the binding of immunomodulatory molecules to tumor cells to produce novel anti-cancer vaccines (McHugh et al., 1999Go; van Broekhoven et al., 2000Go).

We have shown that the transmembrane (T) domain from diphtheria toxin may function as a membrane anchor for soluble proteins fused at its N- or C-terminus, including cytokines (Liger et al., 1998Go) and the IgG-binding protein ZZ (Nizard et al., 1998Go, 2001Go). ZZ was generated by duplication of a mutated B domain from the staphylococcal protein A (Ljungberg et al., 1993Go; Jansson et al., 1998Go). Diphtheria toxin is a 58 kDa protein organized in three domains: receptor binding (R), transmembrane (T) and catalytic (C) (Choe et al., 1992Go; Bennett and Eisenberg, 1994Go). After binding to its cell surface receptor, the toxin is internalized. The acidic pH in the endosome triggers the insertion of T in the membrane. This assists the translocation of C to the cytoplasm where it blocks protein translation by ADP-ribosylation of elongation factor 2 (Lemichez et al., 1997Go). The structure of the T domain (~20 kDa) is organized in three layers of {alpha}-helices (Choe et al., 1992Go; Bennett and Eisenberg, 1994Go): a central hydrophobic helical hairpin sandwiched and hidden from the solvent by two amphiphilic layers. Acidic pH induces a conformational change leading to a molten globule-like state with exposure of the hydrophobic parts of the central layer, promoting interaction with the membrane (Zhan et al., 1994Go).

In order to progress in the design and applications of these membrane anchors, it is important to investigate in detail whether the presence of a `foreign' protein fused to the N- or C-terminus or at both ends of the T domain would modify its structural transition involved in membrane binding. This transition leads to a partial exposure of the hydrophobic core of T (Zhan et al., 1994Go). The close presence of another protein may lead to alterations of this functional transition, undesired interactions between domains within the fusion protein and/or alterations of the kinetics of binding of T to membranes. Furthermore, in terms of the large-scale production of such proteins for therapeutic use, it is of interest to investigate whether the folding pathways of the fusion partners are altered.

Here we studied these problems using three fusion proteins in which the IgG-binding protein ZZ (Ljungberg et al., 1993Go; Jansson et al., 1998Go) was fused to the N- or C-terminus or to both ends of the T domain (Nizard et al., 1998Go, 2001Go). The structural transitions involved in membrane binding or folding were monitored by circular dichroism (CD) and fluorescence at various pHs and/or increasing concentrations of GdnHCl. The kinetics of binding of the fusion proteins and the isolated T domain to membranes were measured by surface plasmon resonance (SPR).

We show that each domain retains its structural and functional features within the fusion proteins. However, differences in solubility were found depending on the protein constructions. The consequences for the use of T as a membrane anchor are discussed.


    Materials and methods
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Recombinant proteins

The recombinant proteins T, ZZ, ZZ-T, T-ZZ and ZZ-T-ZZ have been described previously (Nizard et al., 1998Go, 2001Go). In the isolated T domain, Cys201 (native diphtheria toxin numbering) has been mutated to Ser. The proteins were further purified on a 1 ml anion-exchange column (HiTrap Q-Sepharose HP, Amersham Pharmacia Biotech, Uppsala, Sweden).

CD spectropolarimetry

CD experiments were performed on a CD6 spectrodichrograph (Jobin-Yvon Instruments, Longjumeau, France) calibrated with d10-camphorsulfonic acid (Hennessey and Johnson, 1982Go). All spectra were recorded at 22°C with constant N2 flushing. The scans were recorded using a bandwidth of 2 nm and an integration time of 1 s at a scan rate of 0.5 nm/s. All measurements were performed 1 h after sample preparation, except those with GdnHCl, which were monitored at least 16 h after sample preparation. The spectra were corrected using a blank and a smoothing algorithm was then applied with the minimum filter in the CD6 software (CDMax, filter 5). The mean residue ellipticity in the far-UV, [{theta}]f, and the molar ellipticity in the near-UV, [{theta}]n, were calculated from the relations and , respectively, where {theta}m is the measured ellipticity in degrees, C is the concentration in moles per liter, l is the pathlength of the cell in centimeters and N is the number of residues. The values of [{theta}]f and [{theta}]n are expressed in degrees square centimeters per decimole residue and degrees square centimeters per decimole, respectively. The normalized ellipticity, en, was calculated from the relation , where em is the ellipticity measured for a given pH and/or GdnHCl concentration, emax is the maximum ellipticity measured in the series and emb and emaxb are the corresponding blanks. For neutralized samples, the normalized ellipticity is given as 2en, according to the dilution factor.

Each near-UV spectrum represents the average of 30 scans. In all cases, the protein concentrations were >30 µM. A 500 µl volume of protein solution in 20 mM sodium phosphate, pH 7.8, were added to either 500 µl of 20 mM sodium phosphate, 1 mM sodium acetate, pH 7.6, or 20 mM sodium phosphate, 50 mM sodium acetate, pH 4.4. In acidic pH experiments, the pH was kept at 4.4 for 2 h before neutralization. In neutralization experiments, the pH of the acidic samples was brought back to 7.6 by dilution in an equal volume of 400 mM sodium phosphate, pH 9.

Each far-UV spectrum represents the average of five scans. The protein concentration was 5 µM. Samples were prepared as described for near-UV experiments, but in a final volume of 250 µl.

Far-UV CD as a function of pH

Sodium phosphate solutions (20 mM) buffered from pH 7.6 to 4 were prepared with addition of increasing amounts of 1 M sodium acetate, pH 3.6, to reach a final acetate concentration from 1 to 100 mM. The samples were then prepared with these buffered solutions as described for far-UV in the previous section and the pH was measured for each sample. Ellipticity measurements were performed for 15 s and in triplicate at 208, 222 and 250 nm. The results are given as normalized ellipticity.

Fluorescence spectroscopy

Fluorescence measurements were performed with an FP-750 spectrofluorimeter (Jasco, Tokyo, Japan) at 22°C in a thermostated cell holder and using a 1 cm pathlength quartz cell. Bandwidths of 5 nm were used for both excitation and emission beams. The excitation wavelength was fixed at 295 nm where the contribution of tyrosine residues is negligible. The emission spectra were recorded from 300 to 500 nm at a scan rate of 60 nm/min. Each maximum emission wavelength ({lambda}max) is the average calculated from five emission spectrum measurements. The protein concentration was 5 µM. First, proteins were mixed for 1 h at 22°C in a final volume of 500 µl of 20 mM sodium phosphate, pH 7.8. Then, samples of varying pH were prepared as described in the previous section. The final volume was 1 ml. Neutralized samples were obtained by adding 500 µl of 400 mM sodium phosphate, pH 9, to 500 µl of each sample. {lambda}max was corrected for blank measurements.

CD and fluorescence spectroscopy as a function of GdnHCl

Two stock solutions of 7 M GdnHCl and 20 mM phosphate were prepared and the pH was adjusted to 7.8 and 4.4 with NaOH and sodium acetate, respectively. A range of solutions of GdnHCl concentrations from 0.18 to 6.3 M at pH 7.8 and 4.4 were prepared by dilution of the stock solutions with 20 mM sodium phosphate, pH 7.8, and 20 mM sodium phosphate and with 50 mM sodium acetate, pH 4.4, respectively. For near-UV CD and fluorescence experiments, protein solutions in 20 mM sodium phosphate, pH 7.8, were mixed with the set of GdnHCl solutions at neutral or acid pH in a final volume of 1 ml. The final concentration of proteins was 5 µM. For far-UV experiments, the preparations were identical, but in a final volume of 250 µl. The refractive index of each sample was measured after the experiments were completed to determine the concentration of GdnHCl.

SPR experiments

The binding studies were performed on L1 sensor chips (Biacore, Uppsala, Sweden) coated with lipids. The coating was obtained by injecting small unilamellar vesicles (SUV) up to complete coverage of the surface. SUV were made from phosphatidylcholine–cholesterol (PC–Chol) (8:2) by sonication and their size (<100 nm) was checked by quasi-elastic light scattering. The surface of the L1 sensor chip was cleaned by a 5 min injection of 20 mM CHAPS at a flow-rate of 10 µl/min. A 20 µl volume of SUV, at a lipid concentration of 1.5 mM in PBS buffer, was injected at 25°C at a flow-rate of 2 µl/min. The lipid layer was then washed at 10 µl/min with 10 mM NaOH. Formation of the lipid layer on the L1 chip led to a signal of the order of 10 000–11 000 Ru (Erb et al., 2000Go). At the end of each lipid surface preparation, a control for non-specific binding was performed using BSA in PBS.

Protein binding kinetics were analyzed by SPR using a BIAcore 2000 apparatus (Biacore). Binding of each protein was performed on a new PC–Chol surface stabilized with running buffer (10 mM citrate, pH 4.2, 150 mM NaCl) to avoid the need for a regeneration step. A negligible baseline drift showed the stability of the lipid layer during the analysis. To determine the kinetic constants given in Table IGo, each protein (T, ZZ, ZZ-T, T-ZZ and ZZ-T-ZZ) was injected at three concentrations (18, 36 and 72 nM) in running buffer for 180 s at a constant flow-rate of 20 µl/min followed by a dissociation phase of 600 s. The dissociation time was increased up to 8 h for the highest concentration. To compare the dissociation rates at pH 4.2 and 7.4, each protein was bound at pH 4.2 in running buffer. Then, a first dissociation step was performed for 600 s in the same buffer, followed by a second dissociation step at pH 7.4 by injection of 300 µl of PBS. Each experiment was repeated three times. Association and dissociation curves at acidic pH were analyzed using BIAevaluation 3.1 software (Biacore) using a global fitting method. The kinetic rates of the second dissociation at pH 7.4 were estimated by separate fitting using a simple 1:1 bimolecular interaction model.


View this table:
[in this window]
[in a new window]
 
Table I. Kinetic constants of the interaction of the proteins with PC–Chol membranes
 

    Results
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
The three fusion proteins used in this study, in which the ZZ protein is fused to the N- or C-terminus or to both extremities of the T domain, have been described previously (Nizard et al., 1998Go, 2001Go). These constructs are named ZZ-T, T-ZZ and ZZ-T-ZZ. At pH >7 the T domain remains soluble and does not interact with lipid bilayers. For convenience, we refer to pH from 7 to 7.8 as neutral pH. Two of the recombinant proteins, the isolated T domain and the fusion protein T-ZZ, precipitated around pH 5.6 and 5.2, respectively, which corresponded to their estimated pKi. The proteins ZZ, ZZ-T and ZZ-T-ZZ remained soluble whatever the pH. Therefore, all the experiments performed at acidic pH were done at pH 4.2 at which all proteins were soluble, unless stated otherwise.

Far-UV CD spectra of the proteins at neutral and acidic pH

We studied the secondary structure of the recombinant proteins at neutral and acidic pH using far-UV CD. The far-UV CD spectra of the isolated T domain and of the ZZ protein indicated a mainly {alpha}-helical content (Figure 1A and BGo), in agreement with their crystal structure (Deisenhofer, 1981Go; Choe et al., 1992Go; Cedergren et al., 1993Go; Bennett and Eisenberg, 1994Go). The fusion proteins also exhibited a rich {alpha}-helical content (Figure 1C–EGo). For each protein, the CD spectra were identical when recorded at acidic pH and after neutralization (Figure 1Go). These results show that the T domain and the ZZ protein retained their helical secondary structure, isolated or within the fusion proteins, at neutral and acidic pH.



View larger version (34K):
[in this window]
[in a new window]
 
Fig. 1. Far-UV CD spectra of the recombinant proteins subjected to a cycle of pH. (A) T, (B) ZZ, (C) ZZ-T, (D) T-ZZ and (E) ZZ-T-ZZ at neutral pH (7.8), acidic pH (4.4) and neutralized pH (7.6). Recordings for all three pHs are shown for each protein. (F) Normalized ellipticity at 222 nm of T ({circ}), ZZ (+), ZZ-T ({square}), T-ZZ ({lozenge}) and ZZ-T-ZZ ({triangleup}) as a function of pH. The decrease in ellipticity at 222 nm observed for T and T-ZZ between pH 6.5 and 5 was due to aggregation at the pKi of these proteins.

 
Near-UV CD spectra of the proteins at neutral and acidic pH

Near-UV CD allows the detection of aromatic residues engaged in rigid chiral environments, revealing their embedding in tertiary structure (Adler et al., 1973Go). Any decrease of a near-UV CD signal reflects reduction in the order of tertiary structure.

At neutral pH, the near-UV CD spectrum of T revealed a major peak at 292 nm, which may be attributed to a Trp constrained in an organized tertiary structure (Figure 2AGo). This Trp signal disappeared at acidic pH (Figure 2A and FGo, inset) and was recovered after neutralization. This indicated that at acidic pH the T domain underwent a reversible structural transition with reduced tertiary organization.



View larger version (38K):
[in this window]
[in a new window]
 
Fig. 2. Near-UV CD spectra of the recombinant proteins subjected to a cycle of pH. (A) T, (B) ZZ, (C) ZZ-T, (D) T-ZZ and (E) ZZ-T-ZZ at neutral pH (7.8), acidic pH (4.4) and neutralized pH (7.6). Recordings for all three pHs are shown for each protein. (F) Near-UV CD spectra of ZZ-T as a function of pH. The inset shows the pH-dependence of the normalized ellipticity at 292 nm. This was monitored using ZZ-T because the isolated T domain aggregated around pH 5.5, its estimated pKi.

 
The near-UV CD spectrum of ZZ (Figure 2BGo) showed two negative fine structures at 262 and 268 nm, which may be attributed to packed Phe residues (Strickland, 1974Go). These signals were unaffected by acidic pH, showing that the tertiary structure of ZZ was unchanged whatever the pH.

The near-UV CD spectra of the three fusion proteins displayed the Trp signal found in T together with the two Phe signals found in ZZ (Figure 2Go). As for the isolated fusion partners, the Trp signal was sensitive to pH whereas the Phe signals were not. These results showed that all the recombinant proteins were folded at neutral pH and that the T domain retains its capacity to relax its tertiary structure reversibly at acidic pH within the fusion proteins.

Far-UV CD spectra of the proteins during denaturation by GdnHCl at neutral and acidic pH

The unfolding of the secondary structure of the proteins in response to the addition of GdnHCl was followed by the decrease in negative ellipticity at 222 nm. Unfolding of the T domain occurred in two stages (Figure 3AGo). The first stage, in which close to 60% of the ellipticity was lost, had a midpoint transition at 0.8 M GdnHCl. This led to an intermediate state predominating between 1.5 and 2 M GdnHCl. Most of the remaining ellipticity was lost in a second process, with a midpoint at 3 M GdnHCl. At acidic pH, at which T maintained its secondary structure but lost its tertiary constraints, the unfolding of the secondary structure was proportional to the concentration of GdnHCl above 1 M and occurred at higher concentrations of GdnHCl as compared with neutral pH.



View larger version (25K):
[in this window]
[in a new window]
 
Fig. 3. Normalized ellipticity at 222 nm of the recombinant proteins subjected to GdnHCl-induced unfolding. (A) T, (B) ZZ, (C) ZZ-T, (D) T-ZZ and (E) ZZ-T-ZZ at pH 7.8 (open symbols) and pH 4.4 (closed symbols).

 
Unfolding of the secondary structure of ZZ was identical at neutral and acidic pH and occurred sharply, starting at 3 M GdnHCl with a midpoint around 3.6 M GdnHCl (Figure 3BGo), as previously described for neutral pH (Cedergren et al., 1993Go; Myers and Oas, 2001Go).

The denaturation of the fusion proteins showed complex patterns, reflecting addition of the denaturation curves of each of the fusion partners. At neutral pH, the first stage of unfolding of T appeared as a loss of ~30% ellipticity for ZZ-T and T-ZZ (Figure 3C and DGo) and 20% ellipticity for ZZ-T-ZZ (Figure 3EGo), in accordance with the presence of one or two ZZ domains, respectively. The unfolding intermediate of T was detectable between 1.5 and 2 M GdnHCl. The second stage of unfolding of T starting above 2.5 M GdnHCl seemed detectable, although it was partly hidden by the denaturation of ZZ. At acidic pH, for each fusion protein, denaturation occurred at higher GdnHCl concentrations than at neutral pH, as for the isolated T domain (Figure 3Go).

Overall, the data showed that unfolding of T was different at neutral and acidic pH, as opposed to ZZ for which denaturation was identical. At neutral pH, the unfolding of T followed a two-step process. At acidic pH, {alpha}-helices were more stable and their denaturation was a continuous process. In addition, T and ZZ retained their respective secondary structure denaturation profiles at each pH studied, when fused together. Thus, the denaturation of the secondary structure of each domain, T and ZZ, seemed to behave independently within the fusion proteins.

Near-UV CD spectra of the proteins during denaturation by GdnHCl at neutral and acidic pH

As shown above (Figure 2AGo), the near-UV CD spectrum of T revealed a major peak at 292 nm, which may be attributed to a Trp constrained in an organized tertiary structure. Increasing concentrations of GdnHCl led to a loss of this Trp signal, and the concomitant appearance of a strong negative Tyr signal at 280 nm, between 0.6 and 1.25 M (Figure 4A and BGo). Subsequently, between 2.45 and 3.5 M GdnHCl, the signal at 280 nm disappeared. This revealed three different states in the folding process of the tertiary structure of the T domain, as was observed above for its secondary structure: folded, intermediate, unfolded. At acidic pH, at which T maintained its secondary structure (Figure 1AGo) but lost its tertiary constraints (Figure 2AGo), none of the peaks observed at neutral pH were present, whatever the concentration of GdnHCl, and the CD spectrum remained flat (not shown).



View larger version (36K):
[in this window]
[in a new window]
 
Fig. 4. Near-UV CD signals of the recombinant proteins subjected to GdnHCl-induced unfolding. (A) From top to bottom, near-UV CD spectra of T at 0, 3.5 and 2.45 M GdnHCl. (B) Near-UV signals of T ({circ}) and ZZ-T-ZZ ({triangleup}) at 280 ± 2 nm (closed symbols) and at 292 ± 2 nm (open symbols) as a function of GdnHCl. (C) From top to bottom, near-UV CD spectra of ZZ-T-ZZ at 0, 4.5, 3.8 and 2.4 M GdnHCl. (D) Normalized signal at 262 and 268 nm of ZZ (+) and ZZ-T-ZZ ({triangleup}) as a function of GdnHCl.

 
The near-UV CD spectrum of ZZ in Figure 2BGo displayed two negative CD fine structures at 262 and 268 nm, corresponding to packed Phe residues. These signals were affected by GdnHCl concentrations above 3 M and remained slightly visible at 5 M (Figure 4DGo). The same result was obtained at acidic pH (not shown).

The near-UV CD spectra of the three fusion proteins were also recorded at increasing concentrations of GdnHCl. The signals at 292 and 280 nm corresponding to T and at 262 and 268 nm corresponding to ZZ followed the same modifications at the same GdnHCl concentrations as for the isolated domains (Figure 4CGo, B and D for ZZ-T-ZZ, not shown for ZZ-T and T-ZZ). The spectrum of ZZ-T-ZZ (Figure 4CGo) at neutral pH at 2.5 M GdnHCl exhibited the negative signal of T at 280 nm and the two fine bands of ZZ around 262 and 268 nm. This strongly suggests that T adopted its intermediate state on its folding reaction, whereas ZZ remained stable. Therefore, both partners appeared to retain their own unfolding process when fused. The ZZ Phe signals were stronger for ZZ-T-ZZ than for the isolated ZZ protein. This was because the triple fusion protein ZZ-T-ZZ contained twice as many ZZ domains as the isolated ZZ protein. At acidic pH, the denaturation spectra of the fusion proteins corresponded to the sum of the spectra of the isolated T and ZZ domains: no signal for T and vanishing signals at 262 and 268 nm for ZZ, above 3 M GdnHCl (not shown).

Overall, these results showed that ZZ was fairly resistant to denaturation whatever the pH and that unfolding of its tertiary structure followed a two-state process (folded–unfolded), insensitive to pH variations. In marked contrast, the denaturation of the tertiary structure of the T domain exhibited a complex profile at neutral pH. The reaction exhibited a folding intermediate state between the native and the unfolded states. At acidic pH, no tertiary structure elements were detected during the whole denaturation process of the T domain. These data indicate that T and ZZ retained their respective tertiary structure denaturation profiles at neutral and acidic pH when fused together. Thus, each domain behaves consistently with a model in which they unfold independently within the fusion proteins.

Fluorescence of the proteins as a function of pH and GdnHCl

The {lambda}max of a Trp reflects the polarity of its environment when excited at 295 nm. In the non-polar core of a globular protein, {lambda}max is ~330 nm, whereas in a hydrophilic environment {lambda}max reaches 355 nm. Thus, any change in {lambda}max reveals conformational changes in a protein. The T domain contains two Trp whereas ZZ is devoid of Trp. Hence, only T contributed to fluorescence when excited at 295 nm.

Figure 5AGo shows the {lambda}max of T and the three related fusion proteins subjected to a range of pHs. The increase in {lambda}max (from 336 to 342 nm) when the pH decreased indicated that one or both Trp became more exposed to the solvent at acidic pH. This is in agreement with previous observations made with the isolated T domain (Wang et al., 1997Go; Malenbaum et al., 1998Go). For all proteins the transitions were identical and occurred between pH 6 and 5, with mid-transition around pH 5.5. After neutralization, {lambda}max returned to those found initially at neutral pH (Figure 5AGo, closed symbols).



View larger version (19K):
[in this window]
[in a new window]
 
Fig. 5. Fluorescence of the recombinant proteins as a function of pH and GdnHCl. {lambda}max of T ({circ}), ZZ-T ({square}), T-ZZ ({lozenge}) and ZZ-T-ZZ ({triangleup}), (A) as a function of pH (open symbols) and after neutralization (closed symbols) and (B) as a function of GdnHCl at pH 7.8 (open symbols) and at pH 4.4 (closed symbols).

 
Figure 5BGo shows the {lambda}max of T and the three related fusion proteins subjected to increasing concentrations of GdnHCl, at neutral pH (open symbols) or acidic pH (closed symbols). At neutral pH, the {lambda}max was shifted from 336 to 355 nm for all proteins when the concentration of GdnHCl increased from 0.6 to 2 M. At acidic pH, the {lambda}max was shifted from 342 to 355 nm for all proteins when the concentration of GdnHCl changed from 1 to 4.25 M.

Changes in the fluorescence of ANS are used to detect the presence of solvent-exposed hydrophobic clusters on proteins. The exposure of hydrophobic clusters by the T domain following exposure at acidic pH has already been documented (Zhan et al., 1994Go). Using ANS fluorescence, we have found that exposure of hydrophobic clusters as a function of pH was identical for the T domain and the three fusion proteins (not shown). In the presence of the protein ZZ, ANS did not show any fluorescence change whatever the pH. After neutralization, the {lambda}max and fluorescence intensity returned to those found at neutral pH (not shown), indicating a similar reburying of the hydrophobic clusters for all the proteins.

Overall, these results are in agreement with the near-UV CD experiments, which allowed monitoring of the pH-induced transition and the GdnHCl-induced denaturation pathway of T at neutral pH, isolated or within the fusion proteins. The increased solvent accessibility of the Trp at acidic pH measured by intrinsic fluorescence and the exposure of hydrophobic clusters are correlated with the loss of tertiary structure constraints observed by near-UV CD. In addition, Trp fluorescence allowed monitoring of the denaturation of T at acidic pH, a process that could not be detected by near-UV CD. The data indicate that the T domain followed different denaturation pathways at neutral and acidic pH. They show also that the interactions that stabilize the T domain are different at neutral and acidic pH. Finally, the ZZ protein, once fused to the T domain, did not influence any of its studied transitions, i.e. pH-induced, GdnHCl-induced denaturation at neutral pH and GdnHCl-induced denaturation at acidic pH.

Binding of the recombinant proteins to membranes studied by SPR

We used SPR to compare the interaction of the three fusion proteins ZZ-T, T-ZZ and ZZ-T-ZZ and the isolated T domain with lipid membranes at acidic pH. The study was performed on PC–Chol (8:2) membranes because Chol increased the stability of the membranes immobilized on the chip of the BIAcore apparatus. At pH 4.2 all proteins underwent binding to the lipid membrane (Figure 6Go) in a concentration-dependent manner (data not shown). In contrast, no binding was observed at pH 7.4 (not shown). The protein ZZ did not bind to the membrane at either acidic or neutral pH.



View larger version (24K):
[in this window]
[in a new window]
 
Fig. 6. SPR analysis of the interactions of the recombinant proteins with a PC–Chol phospholipid bilayer. Sensorgram curves for T ({blacktriangledown}), ZZ (*), ZZ-T (•), T-ZZ ({triangleup}) and ZZ-T-ZZ ({circ}). The association and first dissociation steps were performed at pH 4.2 and the second dissociation at pH 7.4.

 
The sensorgrams agreed best with a two-state binding model with conformational change as shown by the fitting of the curves. In such a model, analyte A binds to ligand B, then the complex AB changes to AB*, which cannot dissociate directly to give A + B according to the following equilibrium:

The kinetic constants calculated from the fitted curves are given Table IGo. For all four proteins, T, ZZ-T, T-ZZ and ZZ-T-ZZ, the two association rates (1.53x105 M-1 s-1 < ka1 < 2.40x105 M-1 s-1; 8.96x10-3 s-1 < ka2 < 6.32x10-3 s-1) and the first dissociation rates (2.32x10-3 s-1 < kd1 < 1.45x10-3 s-1) were similar. The second dissociation rates, kd2, were in the region of 10-6 s-1, except for ZZ-T (kd2 = 3.99x10-5 s-1). The apparent dissociation constants (KD = kd1/ka1xkd2/ka2) were difficult to extract accurately from the data owing to the slow rates of the second dissociation. Indeed, when the dissociation step was performed over 8 h at pH 4.2, no further dissociation was measured for the four proteins (not shown). Estimations gave KD < 10-11 M for all proteins except for ZZ-T for which KD {approx} 7x10-11 M.

We then studied dissociation at neutral pH (pH 7.4), following the association and the first dissociation at acidic pH (Figure 6Go). The dissociation curves fitted correctly with a simple 1:1 bimolecular interaction model, with {chi}2 values <1. For all proteins, a fast dissociation was observed at neutral pH, with kd values in the region of 10-3 s-1 (Table IGo).

The results of this SPR study show that the T domain bound to lipid membranes at acidic pH with a high apparent affinity (KD <10-11 M). These results are consistent with a model in which binding occurs according to a two-step mechanism: a low-affinity binding followed by a conformational change, leading to a high-affinity interaction with a very slow dissociation rate. Neutralizing the pH led to a fast dissociation. Interestingly, the ZZ protein, which does not interact with the membrane, did not seem to alter the membrane binding kinetics of the T domain when fused to its N- or C-terminus or to both extremities. A slight decrease in binding affinity was observed for ZZ-T, but not for ZZ-T-ZZ.

Affinity of an IgG2a for ZZ-T or T-ZZ bound to membranes studied by SPR

ZZ is capable of interacting with the Fc part of IgG from various species (Ljungberg et al., 1993Go). Using SPR, we measured the dissociation constants of the interaction of a mouse monoclonal IgG2a with the ZZ part of the fusion proteins ZZ-T and T-ZZ, after its binding to membranes attached to the L1 chip of the BIAcore apparatus. The fusion proteins were first allowed to bind to the membranes at pH 4.2. The interaction of the IgG2a with ZZ was performed at acidic pH because the T domain dissociated from the membrane at neutral pH. The dissociation constants deduced from the association and dissociation curves (not shown) were similar for the two fusion proteins: 0.7x10-8 M < KD < 10-8 M. These values are 3–5-fold higher than those found for the interaction of ZZ with human IgG at neutral pH (Ljungberg et al., 1993Go; Jansson et al., 1998Go). This small discrepancy may be explained by the acidic pH used here, the IgG species difference and/or a steric effect due to the proximity of the membrane. The IgG2a did not bind to the membrane in the absence of the fusion proteins. Thus, the affinity of an IgG for ZZ bound to membranes by the T domain is similar when ZZ is fused to the N- or C-terminus of T.


    Discussion
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
The structural change of T at acidic pH involved in its function has the characteristics of a molten globule

The structural transition of the T domain at acidic pH has been studied previously by disulfide trapping, fluorescence proximity studies, TNS (Zhan et al., 1994Go) and Trp fluorescence (Wang et al., 1997Go; Malenbaum et al., 1998Go). This transition has been described as an opening of the three helical layers composing T, with an exposure of the hydrophobic core of the domain. The acidic pH transition of the whole diphtheria toxin, probed by spectroscopy (Blewitt et al., 1985Go) and hydrophobic photoactivatable reagent (D'Silva and Lala, 1998Go), has been described as a partial unfolding leading to a molten globule-like structure. Here we have presented new data, which further establish that the T domain adopts a molten globule state at acidic pH, as based on the following classical criteria (Ptitsyn, 1992Go; Kuwajima, 1996Go). CD experiments showed that at acidic pH, T retained more than 90% of its secondary structure (Figure 1A and FGo) but changed its tertiary structure, with a loss of constraint on Trp (Figure 2A and FGo, inset). In agreement with other studies (Zhan et al., 1994Go; Wang et al., 1997Go; Malenbaum et al., 1998Go), the increase in ANS fluorescence (not shown) and the shift of the {lambda}max of Trp fluorescence from 336 to 342 nm (Figure 5AGo) indicated exposure of hydrophobic clusters and a partial exposure of Trp to the solvent. This transition was fully reversible after neutralization, even after 2 h at low pH. Altogether, these structural characteristics at acidic pH are in agreement with the definition of a molten globule (Ptitsyn, 1992Go; Kuwajima, 1996Go). This conclusion was further demonstrated by two additional observations. First, the denaturation of T by GdnHCl is less cooperative at acidic than at basic pH (Figures 5B and 3AGoGo), as observed for other proteins in a molten globule state (Christova et al., 2000Go). This indicates that the protein surface accessible to the solvent is larger at acidic pH (Myers et al., 1995Go). Second, 1D NMR spectra of T at acidic pH were poorly resolved with broader peaks in the region of aliphatic side chains together with a considerable diminution of the chemical shift dispersion as compared with spectra in the native state (unpublished results). This behavior has also been related to the molten globule state (Alexandrescu et al., 1993Go; Arai and Kuwajima, 2000Go).

The molten globule state of T at acidic pH is not connected to its GdnHCl-induced unfolding pathway

The molten globule has been described as a major class of folding intermediate, which occurs on the folding pathway of globular proteins and which can be observed during kinetic refolding or stabilized by very low pH (between 4 and 2) as well as by denaturing agents (Ptitsyn et al., 1990Go; Haynie and Freire, 1993Go; Kuwajima, 1996Go; Forge et al., 1999Go; Fujiwara et al., 1999Go). Interestingly, the T domain has a very different behavior. First, its molten globule transition is completed at pH 5 in solution. Second, the folding intermediate of the T domain that we identified by GdnHCl denaturation is not a molten globule and has a very different signature (Table IIGo). Namely, it displayed (i) <50% of the helical content found in the native conformation (Figure 3AGo), (ii) a loss of tertiary constraint on the Trp as in the molten globule, but with the presence of non-native tertiary constraints on Tyr (Figure 4A and BGo), (iii) fully solvated Trp (Figure 5BGo) and (iv) the absence of hydrophobic clusters detectable by ANS fluorescence (not shown). This unusual dual unfolding pathway could be explained by the fact that the molten globule state of the T domain is involved in its function, i.e. insertion into the membrane of the endosome at a mild acidic pH. If this conformation existed as a folding intermediate, it would be dangerous to the bacteria expressing it. Such a conformation could interact with, and permeabilize, the membrane of the bacteria. Indeed, expression of the T domain in the periplasm of Escherichia coli under acidic conditions blocks bacterial growth (O'Keefe et al., 1992Go; and our observations). This is not the case when T is expressed in the cytoplasm.


View this table:
[in this window]
[in a new window]
 
Table II. Comparison of the structural and functional characteristics of the molten globule and unfolding intermediate states of the T domain
 
A growing number of proteins are found to be able to shift to a molten globule state under physiological conditions (Wright and Dyson, 1999Go; Dunker et al., 2001Go). Basically, they can be divided into two groups. The first group encloses proteins that are in a molten globule state in the absence of their ligand: metal ions (Christova et al., 2000Go), prosthetic groups (Uversky et al., 1997Go; Bychkova et al., 1998Go), nucleic acids (Hornby et al., 1994Go; Carroll et al., 1997Go), other proteins (Kirkitadze et al., 1998Go), etc. The second group encloses soluble proteins (domains) that turn into a molten globule in order to penetrate into, or translocate across, a membrane. These include, for example, cytochrome c (Bychkova et al., 1996Go), the cholesterol transporter StAR (Bose et al., 1999Go), pore-forming and translocating bacterial toxins such as the staphylococcal {alpha}-toxin and colicins (Parker and Pattus, 1993Go; Vecsey-Semjen et al., 1996Go; Zakharov et al., 1998Go). The channel-forming domain of colicins shares structural and functional resemblance with the diphtheria toxin T domain (Parker and Pattus, 1993Go). However, its molten globule state does not occur in solution, but rather in the early steps of interaction with the membrane (Zakharov et al., 1998Go). Finally, members of the pro- or anti-apoptotic proteins of the Bcl-2 family also share close structural resemblance with the T domain (Reed, 1997Go). They acquire a molten globule conformation at a moderately acidic pH, which promotes interaction with membranes and formation of dimers, a sequence of events involved in apoptosis (Xie et al., 1998Go).

The recombinant proteins have different solubility at acidic pH

Few studies have described the influence of a protein domain on its neighbor during folding or functional transitions within fusion proteins. The ZZ protein was shown to act as a solubilizing partner for proteins with a tendency to aggregate during folding, increasing the yield of production (Samuelsson et al., 1994Go). However, it is not known whether the folding pathways of the fusion partners were modified. All the recombinant proteins used in the present study were soluble during unfolding experiments. The situation was different during the pH-induced transition of T. The proteins T and T-ZZ precipitated around their estimated pKi, whereas ZZ-T and ZZ-T-ZZ remained soluble. At this pH (~5.5), hydrophobic clusters are already present at the surface of T (Zhan et al., 1994Go; and data not shown). However, the aggregation of T is not due directly to these clusters because T is soluble below pH 5 (Figure 1FGo). Hence the net charge of the domain plays a role. A foreign protein at the N-terminus of T could mask areas of T involved in aggregation as suggested previously for a ZZ-insulin-like growth factor I fusion protein (Samuelsson et al., 1994Go). If this were the case, these areas would not correspond to the regions of T involved in membrane binding because all fusion proteins have the same capacity to bind to (Figure 6Go) and to permeabilize membranes (Nizard et al., 2001Go). Finally, one cannot exclude that the peptide linker between ZZ and T, which is highly hydrophilic with five Ser residues, may have a solubilizing effect. In summary, fusion of the ZZ protein may be a useful tool for structural studies in conditions favoring aggregation of a given protein.

The pH-induced transition of T and the unfolding of T and ZZ are unaffected by protein fusion

The influence of the fusion of protein domains on their structure and function is not a trivial issue. The addition of a single N-terminal residue may destabilize a protein (Ishikawa et al., 1998Go; Chaudhuri et al., 1999Go). It has been shown that adjacent domains in a protein may be more flexible within the protein than alone (Engen et al., 1999Go). In this case, an interface exists between both domains. Another study showed that a protein domain could tolerate fairly well the insertion of various proteins at various positions (Collinet et al., 2000Go). Both proteins (enzymes) were functional, but their activities and folding transitions were affected to some extent, depending on the site of insertion. The authors suggested that autonomous folding units do not remain autonomous once their polypeptide chains are connected. This was found for inserted proteins with both extremities engaged in the fusion protein. In the present work, we found the opposite. The respective structural and functional behaviors of the T and ZZ domains were identical whether they were isolated or fused. However, compared with the situation described by Collinet et al. (Collinet et al., 2000Go), each domain has more freedom. A flexible linker of 12 residues was added between ZZ and the N-terminus of the T domain (Nizard et al., 2001Go). Similarly, the C-terminal residues of the T domain (from 377 to 386 native diphtheria toxin numbering) plus two residues introduced by the gene construction stick out of the globular structure of the domain and constitute a linker pushing the fused protein away from T. This design was empirically chosen to avoid steric hindrance between the two fusion partners, as the goal is to use the T domain as a soluble membrane anchor for functional proteins.

The pH-induced transition of T was not affected by the fusion of ZZ, even in the triple fusion protein. In the native toxin, the T domain is placed between the catalytic and the receptor-binding domains. At acidic pH, T interacts with the catalytic domain, in addition to other proteins, provided that these other proteins also adopt a molten globule conformation (Ren et al., 1999Go). This property may be involved in the process of translocation of the catalytic domain. In the case described here, the ZZ protein appeared very stable whatever the pH. This stability may prevent interaction with the T domain in the molten globule conformation. Indeed, in cell-binding studies with the ZZ-T protein, the ZZ domain is not translocated across the membrane (Nizard et al., 2001Go). In contrast, translocation of foreign proteins fused to the N-terminus of the T domain has been described, provided that they are highly unstable (Klingenberg and Olsnes, 1996Go). Hence the pH stability of the protein to be anchored to membranes using T should be taken into consideration.

Functional integrity of the T domain and the ZZ protein within the fusion proteins

SPR experiments have shown that the T domain has the same capacity to interact with phospholipid bilayers, isolated or within the three fusion proteins (Figure 6Go). These data gave for the first time quantitative values of the kinetics of binding of T to membranes. In particular, they showed that T has a high apparent affinity for membranes at acidic pH (KD <10-11 M). Strictly, this is an apparent affinity because the analytes interact with a surface instead of discrete binding sites. However, many authors use the term affinity for protein–membrane interactions (Mozsolits et al., 2001Go; Stahelin and Cho, 2001Go). This high affinity is an important feature for the use of T as a membrane anchor. In another study we have shown that all three fusion proteins attach to cell membranes with similar efficacy (Nizard et al., 2001Go), which is consistent with the present results. However, one cannot conclude that the affinity of T for cell membranes is identical. Cell membranes are much more complex than lipid vesicles. Indeed, while we found that binding to PC–Chol membranes (Figure 6Go) and to PC–PA vesicles (unpublished results) is reversible when the pH is neutralized, binding to cell membranes is stable for more than 1 day (Nizard et al., 2001Go). The interaction of the T domain with the membranes agrees with the model where this interaction takes place in two steps: a low-affinity binding stage followed by a change in conformation leading to a stable membrane-associated state. These two steps may reflect a superficial binding of the molten globule state observed in solution, followed by penetration into the membrane (Zhan et al., 1994Go).

In conclusion, this work shows that in the case of the ZZ protein, there is no structural or functional caveat against using the T domain as a soluble membrane anchor. It is of importance to study how such a useful result extends to any soluble proteins. In this respect, we have already shown that T could attach human IL-2 and murine IL-3 fused at its C-terminus to the surface of cells (Liger et al., 1998Go). The fusion proteins described here may also be used to anchor IgG to cell membranes, taking advantage of the IgG-binding property of ZZ (Nizard et al., 1998Go, 2001Go). For new constructs, it is important, however, to look for and avoid pH values that could cause precipitation. The T domain possesses several interesting properties as an anchor. Attachment to membranes is fast and triggered by a simple pH drop. It accepts the fusion of proteins at either of its extremities or at both at the same time. Hence it offers the possibility of choosing the orientation of the soluble protein, its N- or C-terminus towards the membrane, and it allows membrane attachment of two different proteins at the same time in a triple fusion protein. However, it is not suitable for attachment to liposomes because it dissociates after neutralization and it destabilizes lipid vesicles (Nizard et al., 2001Go). Many applications may be found for membrane anchors including the design of cancer vaccines (McHugh et al., 1999Go; van Broekhoven et al., 2000Go), linkage of enzymes to membranes, cell tagging and design of artificial receptors.


    Notes
 
4 To whom correspondence should be addressed. E-mail: daniel.gillet{at}cea.fr Back


    Acknowledgments
 
This work is dedicated to the memory of Jean-Claude Mani. We thank A.Urvoas for her help with the spectrofluorimeter. P.N. and A.C. were supported by the Ministère de l'Education Nationale, de la Recherche et de la Technologie. This work was supported by the Commissariat à l'Energie Atomique, the Association pour la Recherche contre le Cancer (grant 5491) and the Comité Départemental de l'Essonne de la Ligue Nationale Contre le Cancer.


    References
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Adler,A.J., Greenfield,N.J. and Fasman,G.D. (1973) Methods Enzymol., 27, 675–735.[Medline]

Alexandrescu,A.T., Evans,P.A., Pitkeathly,M., Baum,J. and Dobson,C.M. (1993) Biochemistry, 32, 1707–1718.[ISI][Medline]

Arai,M. and Kuwajima,K. (2000) Adv. Protein Chem., 53, 209–282.[ISI][Medline]

Bennett,M.J. and Eisenberg,D. (1994) Protein Sci., 3, 1464–1475.[Abstract/Free Full Text]

Blewitt,M.G., Chung,L.A. and London,E. (1985) Biochemistry, 24, 5458–5464.[ISI][Medline]

Bose,H.S., Whittal,R.M., Baldwin,M.A. and Miller,W.L. (1999) Proc. Natl Acad. Sci. USA, 96, 7250–7255.[Abstract/Free Full Text]

Bychkova,V.E., Dujsekina,A.E., Klenin,S.I., Tiktopulo,E.I., Uversky,V.N. and Ptitsyn,O.B. (1996) Biochemistry, 35, 6058–6063.[CrossRef][ISI][Medline]

Bychkova,V.E., Dujsekina,A.E., Fantuzzi,A., Ptitsyn,O.B. and Rossi,G.L. (1998) Fold. Des., 3, 285–291.[ISI][Medline]

Carroll,A.S., Gilbert,D.E., Liu,X., Cheung,J.W., Michnowicz,J.E., Wagner,G., Ellenberger,T.E. and Blackwell,T.K. (1997) Genes Dev., 11, 2227–2238.[Abstract/Free Full Text]

Cedergren,L., Andersson,R., Jansson,B., Uhlen,M. and Nilsson,B. (1993) Protein Eng., 6, 441–448.[Abstract]

Chaudhuri,T.K. et al. (1999) J. Mol. Biol., 285, 1179–1194.[CrossRef][ISI][Medline]

Choe,S., Bennett,M.J., Fujii,G., Curmi,P.M., Kantardjieff,K.A., Collier,R.J. and Eisenberg,D. (1992) Nature, 357, 216–222.[CrossRef][ISI][Medline]

Christova,P., Cox,J.A. and Craescu,C.T. (2000) Proteins, 40, 177–184.[CrossRef][ISI][Medline]

Collinet,B., Herve,M., Pecorari,F., Minard,P., Eder,O. and Desmadril,M. (2000) J. Biol. Chem., 275, 17428–17433.[Abstract/Free Full Text]

Deisenhofer,J. (1981) Biochemistry, 20, 2361–2370.[ISI][Medline]

de Kruif,J., Tijmensen,M., Goldsein,J. and Logtenberg,T. (2000) Nature Med., 6, 223–227.[CrossRef][ISI][Medline]

D'Silva,P.R. and Lala,A.K. (1998) J. Biol. Chem., 273, 16216–16222.[Abstract/Free Full Text]

Dunker,A.K. et al. (2001) J. Mol. Graph. Model., 19, 26–59.[CrossRef][ISI][Medline]

Engen,J.R., Smithgall,T.E., Gmeiner,W.H. and Smith,D.L. (1999) J. Mol. Biol., 287, 645–656.[CrossRef][ISI][Medline]

Erb,E.M., Chen,X., Allen,S., Roberts,C.J., Tendler,S.J., Davies,M.C. and Forsen,S. (2000) Anal. Biochem., 280, 29–35.[CrossRef][ISI][Medline]

Forge,V., Wijesinha,R.T., Balbach,J., Brew,K., Robinson,C.V., Redfield,C. and Dobson,C.M. (1999) J. Mol. Biol., 288, 673–688.[CrossRef][ISI][Medline]

Fujiwara,K., Arai,M., Shimizu,A., Ikeguchi,M., Kuwajima,K. and Sugai,S. (1999) Biochemistry, 38, 4455–4463.[CrossRef][ISI][Medline]

Haynie,D.T. and Freire,E. (1993) Proteins, 16, 115–140.[ISI][Medline]

Hennessey,J.P.,Jr and Johnson,W.C.,Jr. (1982) Anal. Biochem., 125, 177–188.[ISI][Medline]

Hornby,D.P., Whitmarsh,A., Pinarbasi,H., Kelly,S.M., Price,N.C., Shore,P., Baldwin,G.S. and Waltho,J. (1994) FEBS Lett., 355, 57–60.[CrossRef][ISI][Medline]

Ishikawa,N., Chiba,T., Chen,L.T., Shimizu,A., Ikeguchi,M. and Sugai,S. (1998) Protein Eng., 11, 333–335.[Abstract]

Jansson,B., Uhlen,M. and Nygren,P.A. (1998) FEMS Immunol. Med. Microbiol., 20, 69–78.[CrossRef][ISI][Medline]

Kirkitadze,M.D., Barlow,P.N., Price,N.C., Kelly,S.M., Boutell,C.J., Rixon,F.J. and McClelland,D.A. (1998) J. Virol., 72, 10066–10072.[Abstract/Free Full Text]

Klingenberg,O. and Olsnes,S. (1996) Biochem. J., 313, 647–653.[ISI][Medline]

Kuwajima,K. (1996) FASEB J., 10, 102–109.[Abstract/Free Full Text]

Lemichez,E., Bomsel,M., Devilliers,G., vanderSpek,J., Murphy,J.R., Lukianov,E.V., Olsnes,S. and Boquet,P. (1997) Mol. Microbiol., 23, 445–457.[ISI][Medline]

Liger,D., Nizard,P., Gaillard,C., vanderSpek,J.C., Murphy,J.R., Pitard,B. and Gillet,D. (1998) Protein Eng., 11, 1111–1120.[Abstract]

Ljungberg,U.K., Jansson,B., Niss,U., Nilsson,R., Sandberg,B.E. and Nilsson,B. (1993) Mol. Immunol, 30, 1279–1285.[CrossRef][ISI][Medline]

Malenbaum,S.E., Collier,R.J. and London,E. (1998) Biochemistry, 37, 17915–17922.[CrossRef][ISI][Medline]

McHugh,R.S., Nagarajan,S., Wang,Y.C., Sell,K.W. and Selvaraj,P. (1999) Cancer Res., 59, 2433–2437.[Abstract/Free Full Text]

Mozsolits, H., Wirth, H.J., Werkmeister, J. and Aguilar, M.I. (2001) Biochim. Biophys. Acta, 1512, 64–76.[ISI][Medline]

Myers,J.K. and Oas,T.G. (2001) Nature Struct. Biol., 8, 552–558.[CrossRef][ISI][Medline]

Myers,J.K., Pace,C.N. and Scholtz,J.M. (1995) Protein Sci., 4, 2138–2148.[Abstract/Free Full Text]

Nizard,P., Liger,D., Gaillard,C. and Gillet,D. (1998) FEBS Lett., 433, 83–88.[CrossRef][ISI][Medline]

Nizard,P., Chenal,A., Beaumelle,B., Fourcade,A. and Gillet,D. (2001) Protein Eng., 14, 439–446.[Abstract/Free Full Text]

O'Keefe,D.O., Cabiaux,V., Choe,S., Eisenberg,D. and Collier,R.J. (1992) Proc. Natl Acad. Sci. USA, 89, 6202–6206.[Abstract]

Parker,M.W. and Pattus,F. (1993) Trends Biochem. Sci, 18, 391–395.[CrossRef][ISI][Medline]

Ptitsyn,O.B., Pain,R.H., Semisotnov,G.V., Zerovnik,E. and Razgulyaev,O.I. (1990) FEBS Lett., 262, 20–24.[CrossRef][ISI][Medline]

Ptitsyn,O.B. (1992) In Creighton,T.E. (ed.), The Molten Globule State. Freeman, New York, pp. 243–300.

Reed,J.C. (1997) Nature, 387, 773–776.[CrossRef][ISI][Medline]

Ren,J., Kachel,K., Kim,H., Malenbaum,S.E., Collier,R.J. and London,E. (1999) Science, 284, 955–957.[Abstract/Free Full Text]

Samuelsson,E., Moks,T., Nilsson,B. and Uhlen,M. (1994) Biochemistry, 33, 4207–4211.[ISI][Medline]

Stahelin, R.V. and Cho, W. (2001) Biochemistry, 40, 4672–4678.[CrossRef][ISI][Medline]

Strickland,E.H. (1974) CRC Crit. Rev. Biochem., 2, 113–175.[Medline]

Uversky,V.N., Narizhneva,N.V., Ivanova,T.V., Kirkitadze,M.D. and Tomashevski,A. (1997) FEBS Lett., 410, 280–284.[CrossRef][ISI][Medline]

van Broekhoven,C.L., Parish,C.R., Vassiliou,G. and Altin,J.G. (2000) J. Immunol., 164, 2433–2443.[Abstract/Free Full Text]

Vecsey-Semjen,B., Mollby,R. and van der Goot,F.G. (1996) J. Biol. Chem., 271, 8655–8660.[Abstract/Free Full Text]

Wang,Y., Malenbaum,S.E., Kachel,K., Zhan,H., Collier,R.J. and London,E. (1997) J. Biol. Chem., 272, 25091–25098.[Abstract/Free Full Text]

Wright,P.E. and Dyson,H.J. (1999) J. Mol. Biol., 293, 321–331.[CrossRef][ISI][Medline]

Xie,Z., Schendel,S., Matsuyama,S. and Reed,J.C. (1998) Biochemistry, 37, 6410–6418.[CrossRef][ISI][Medline]

Zakharov,S.D., Lindeberg,M., Griko,Y., Salamon,Z., Tollin,G., Prendergast,F.G. and Cramer,W.A. (1998) Proc. Natl Acad. Sci. USA, 95, 4282–4287.[Abstract/Free Full Text]

Zhan,H., Choe,S., Huynh,P.D., Finkelstein,A., Eisenberg,D. and Collier,R.J. (1994) Biochemistry, 33, 11254–11263.[ISI][Medline]

Received November 14, 2001; revised February 8, 2002; accepted February 19, 2002.