Methanol-stabilized intermediates in the thermal unfolding of ribonuclease A

.I. Mol.

Hiol.

(1982)

160, 87-116

Methanol-stabilized Thermal Unfolding Characterization

Intermediates in the of Ribonuclease A

by ‘H Nuclear Magnetic Resonance

Ro(:E:K G. BIKINGEK. AXI) ASTHOSY I,. FISK? Division,

of Xatural Santa

(Received

24 August

Aciences, Cruz,

University

of California

CA 95064, V.S.A.

1981, and in revised form

13 May 1982)

The thermal unfolding of ribonuclease A has been studied in solutions of 25. 35 and 500;, methanol (v/v)* using 360MHz proton magnetic resonance spectroscopy. Several observations indicate that the native structure of the protein in methanol cryosolvents is very similar to that in aqueous solution. A detailed analysis of the unfolding process has been made using the C-2 protons of the imidazole side-chains of the four histidine residues. As denaturation proceeds new resonances appear. whose chemical shifts correspond to neither native nor fully unfolded species. These have been assigned to particular His residues by selective deuteration studies. The thermal denaturation transitions reveal a multiphasic process in each of the solvents. and become less co-operative with increasing concentrations of methanol. The denaturation is fully reversible with no evidence of hysteresis. The new resonances that appear during the unfolding process are attributed to partially folded species. which are stabilized by the presence of the relative]) hydrophobic methanol. Eased on the temperature dependence of the chemical shifts and the relative areas of the various resonances. a detailed sequence of events has been proposed to describe the unfolding process. Key features include the initial genera1 loosening of the two domains, the subsequent movement of the upper S-peptide region (residues 13 to 25) away from the main body of the protein. followed by partial separation of the sheet structure and full exposure of the Nterminal helix, leading to complete separation of the “winged domains”. and ultimately the loss of the residual sheet and helix structure.

1. Introduction The mechanism of protein folding is currently undergoing much experimental and theoretical scrutiny (Jaenicke, 1980). One of the key questions concerns whether the folding process is nucleationor intermediate-controlled (Baldwin, 1980). If bhe former is the case, the folding process would essentially be totally co-operative. whereas in the latter case partially folded intermediates would exist, although there may not be enough to be detectable. Numerous experimental investigations have demonstrated that under normal conditions of protein denaturation, e.g. aqueous solution and temperatures 225”C, the majority of proteins exhibit a two-state transition : that is. the only populated species are the native (folded) and denatured

88

Ii. G. HIHIN(:ER

ANI)

A. I,. FINK

(fully unfolded) states. At present there are a number of systems where it is apparent that more than a simple two-stat.e situation exists: many of these have been summarized in a recent review (Baldwin & Creighton, 1980). In several of these cases it is clear that the additional species detected are not partially folded intermediates ; e.g. they may represent slow and fast-folding species due to cisltrans isomerization of proline (Brandts et al., 1975: Schmid & Baldwin, 1978), or they may reflect the individual folding of domains in a multi-domain protein (e.g. Tiktopulo & Privalov, 1978: Robson & Pain, 1976). Since normally transient intermediates in enzyme catalysis can be detected, stabilized and characterized using subzero temperatures in conjunction with fluid aqueous/organic solvent systems (Douzou, 1977 ; Fink, 1977 ; Fink & Cartwright, 1981), it occurred to us that the cryo-enzymological approach might also be valuable in the detection of transient partially folded intermediates in protein folding. In fact if the intermediate-controlled theories of folding are correct, the combination of low temperature and cosolvent’ would be expected to be particularly effective in stabilizing partially folded intermediates. The basis for this statement is as follows : a simplified picture of the sequence of events during folding would involve initial formation of nucleation sites leading to regions of secondary structure, linked by random chain segments. Subsequently, the units of secondary alpha and beta structure would pack together due to hydrophobic interactions. The final stage of folding would then involve the essential completion of solvent exclusion and minor rearrangements to yield the maximum favorable intramolecular interactions involving both hydrophobic interactions and hydrogen bonding. Low temperatures would be expected to strengthen hydrogen bonds, and weaken hydrophobic interactions, thus tending to stabilize the initially formed regions of secondary structure. The greater hydrophobicity of organic cosolvents would also be expected to stabilize the initially formed units of secondary structure, with their exposed non-polar groups, relative to the situation in aqueous solution. Since preliminary experiments verified these expectations (Fink & Grey, 1978). we have embarked on an extensive investigation of protein folding in the presence of organic cosolvents, and at subzero temperatures. A number of factors suggested that ribonuclease A would be a suitable candidate for initial studies. These include its well-characterized structure: the lack of significant aggregation when it is unfolded, even in high cosolvent concentrations; the large number of previous studies concerning its folding : evidence for a partially folded intermediate in aqueous solution (Blum et al., 1978 ; Hagerman et al., 1979 : Schmid & Baldwin, 1979: Schmid & Blaschek, 1981); its excellent solubility properties in cryosolvents which make n.m.r.t studies feasible: and finally, observations that indicate that ethanol and methanol cryosolvents have no adverse effects on the structure or catalytic properties of the enzyme (Fink & Grey, 1978: G. A. Petsko, personal communication; Fink, Kar & Kotin, unpublished data). We ascribe the lack of adverse effects of ethanol and methanol on RNAase A to the preferential exclusion of the cosolvent from the vicinity of the protein (Pittz & Timasheff, 1978; Fink & Cartwright, 1981) : i.e. charged groups on the surface result t Abbreviations used : n.m.r., nuclear magnetic resonance: DSS. sodium-Z,B-dimethvl-2-Yila~ntan~. R-sulfonate: p.p.m.. parts per million: u.v.. ultraviolet light.

INTERMEDIATES

IN

RNAase

UNFOLDING

89

in unfavorable interactions with the relatively more hydrophobic cosolvent (compared to water), leading to the exclusion of the cosolvent molecules from such areas. and hence preferential hydration. Previous studies of the thermal denaturation of RNAase A using n.m.r. have provided some evidence for the presence of intermediate species in the reaction, especially at very low pH values. For example, Westmoreland and Matthews (1973) observed evidence for multistate thermal denaturation at pH 1.3. Subsequently, Benz & Roberts (1975a,b) extended this study to include other pH values, as well as unfolding induced by urea and guanidinium chloride. Evidence has also been found hy Blum et ~2. (1978) for an intermediate in refolding, in which a new resonance appears at a chemical shift corresponding to that of the C-2 proton of His12 in the isolated S-peptide. The above studies all utilized proton n.m.r. and the wellresolved C-2 proton signals from the His residues. Recently, Howarth (1979) has examined the 13C n.m.r. spectrum of RNAase A during thermal denaturation. and also finds evidence to support intermediate states. Scheraga and co-workers, on the basis of a variety of different data, have attempted to delineate the pathway of folding in RNAase A (Burgess & Scheraga, 1975; Nemethy & Scheraga, 1979; Matheson & Scheraga, 1979). The present investigation demonstrates the potential of our approach to reveal stabilized, partially folded intermediates, and to allow determination of the pathway of folding.

2. Materials and Methods Methanol-d, was obtained from Stohler Isotope Chemicals Inc., and *H20 from Bio-Rad I,aboratories. Ribonuclease A was purchased either from Calbiochem-Behring Corp. or Worthington Biochemical Corp. The enzyme was further purified, essentially by the method of Taborsky (1959). using a Sephadex CM-50 column with a sodium chloride gradient (001 M t,o 0.25 M) in Tris buffer (pH 8.0). The enzyme eluted around 0.11 M-sodium chloride. After exhaustive dialysis against distilled water at 4°C the sample fraction was lyophilized. S-protein was prepared from RNAase A by the method of Doscher & Hirs (1967). as modific,d by Chavez 8.1Scheraga (1980). The resulting protein fractions were desalted and concentrated using an Amicon UM-2 membrane?, lyophilized, and stored at -20°C. The backbone amide protons of the S-protein were exchanged in *H,O at pH 6.93. 45°C. for 10 min. The peptide backbone protons of RNAase A were exchanged using Markley’s (1!)75) procedure E. By repeating the process 2 or 3 times the residual HO’H peak could be made vrry small. The resultant product was lyophilized and stored at - 20°C. Selective deuteration was accomplished as follows: RNAase A was dissolved in *Hz0 (99.8%) (40 mg/ml) and the pH* (see below) adjusted to 890 with Na02H. The solution was then incubated at 40°C. Samples were removed at 3, 6 and 16.day intervals. Each sample was allowed to cool to ambient temperature, and the pH* adjusted to 3.00 to quench the exchange reaction. A white precipitate formed during the process and was removed by centrifugation. The snpernatant was further purified by ultrafiltration using an Amicon PM-10 membrane, and washed with deionized, distilled water. The solution was then lyophilized. and the backbone protons exchanged as mentioned above. Solutions of deuterium-exchanged ribonuclease A in 25 and 35”/0 methanol-d, were prepared at room temperature as follows: the cryosolvents were mixed on a v/v basis. i.e. 2,>: 75. methanol-d,/*H20 (99.9%) and adjusted to a predetermined ionic strength using KCI and th(sn mixed with lyophilized enzyme. DSS was generally added to the solvent prior to t ~
90

R. (:. BIRINGER

ASI)

A. 1,. FINK

mixing with RNAase. The pH* was then adjusted with ‘HCI or NaO’H as necessary. Solutions of the enzyme in 50% methanol-d, were prepared at 1‘C by mixing the cryosolvent with the dry enzyme. If the enzyme and methanol cryosolvent were mixed at 25°C. where the protein is well into its denaturation transition. a gel was formed. If the enzyme was prepared in 5O”/b methanol at 1°C. where no significant denaturation has yet occurred. gel formation was not observed at any subsequent temperature. In order t,o excslude moisture from the air. the samples were prepared in a dry-box under an atmosphere of nitrogen. The pH* values reported here are the observed pH-meter readings obtained at 25“C using a glass combination electrode. The values of pH* thus represent t,he apparent protonic activity in the deuterocryosolvent (Hui Ron Hoa & Douzou. 1973). Absorbance measurements of the unfolding were performed at 286 nm using a Gary 118 spectrophotometer. Typical concentrations used were 0.1 mg of Rh’Aase A/ml. Reported t, values were obtained by asrertaining the temperature at which half the maximal change in absorbance or area had occurred. (a) S.w.r.

rnrnsl~rrrnents

,411 measurements were performed with a modified Bruker 360 MHz spectrometer. equipped with a Nicolet 1180 data system. unless otherwise noted. All chemical shifts are referred to an internal DSS standard; at each temperature the spectrum was calibrated by setting the DSS peak to 0 p.p.m. The reproducibility in the chemical shifts of resonances is estimated as kOOO3 p.p.m. The data for 35”/; and 500,; methanol were obtained from 3 or 4 separate experiments (samples), each covering the whole temperature range examined. and were completely reproducible. Peak areas were integrated by electronic integration. Areas were generally reproducible within a few per cent and averages of 2 or 3 measurements were used for each data point. All RSAase A spectra were obtained with a standard l-pulse sequence. using the following parameters: 90” pulse (typically 18 ps), acquisition delay (200 ps), pulse delay (minimum 3 s). The long pulse delay allbwed complete equilibration of the His C-2 resonances (see Results) prior to the start of the next ptilse sequence. Gated decoupling was used to eliminate solvent resonances. A maximum of T,,V was applied to the HO’H peak, and not more than 0.5 V to the methanol mak. RampIe concentrations were generally 20 mg/ml. The spectra were artificially line-&oad&ed ,b.y 05 Hz to improve the signal/noise ratio. A temperature c$iliiaiion ciirve: was prepared for each sample. The chemical shift of the H0”H peak was me&ui’k;d&&~e$pec% to DSS as a function of temperature using a Jeol FX100 instrument. Th~~:,t&$&&u& was monitored with a thermocouple probe inserted directly into the n.m.r:I:‘&be:, When the HO’H shift had stabilized at a particular temperature the probe wa8i ‘ins@Gd and its value was recorded. The process was repeated until 2 successive measurements gbvethe same shift and temperature values. respecntively. .4 linear dependence of chemical shift tin temperature over the 0 t,o 70°C range was found for all the solvents examined. Spectra for the S-protein were obtained using a standard Redfield 21412 pulse sequence to improve the signal/noise ratio. as only a small amount of protein was used. All peaks were referenced to DSS from a subsequent standard l-pulse experiment. (I))

Kxpwirnrnts

to detect nggrrgntion

A number of methods were used to assess the state of association of ribonuclease A at the high concentrations used in these n.m.r. experiments. Laser light-scattering was used to determine the diffusion coefficient with a Malvern Scientific Corp. light-scattering photometer, equipped with a Malvern model K7025 correlator and a 15 mW He-Ne laser. ITltracentrifugation (sedimentation equilibrium) experiments were carried out with a Spinco model E instrument. equipped for absorbance monitoring. In each case samples of 20 mg/ml in aqueous and aqueous/methanol solvents. pH* 3.0. were used. Columns (1 cm x 30 cm) of Biogel P60 were used for gel permeation chromatography. The sample was 0.5 ml of a 40 mg/ml protein solution. These experiments were performed at 25°C. pH* 3.0 (@05 Mformate). Pl M-KU. in aqueous and aqueous/methanol solvents.

ISTEKMEI)IATES

9.0

8.5

IN

8.0

RNAase

7.5

I’NFOLI)lS(:

7.0

HI

6.5

p.p.m.

of methanol on the aromatic region of thr proton spectrum of KSAase A at ‘H,O. 22’C. pH* :34X).0.1 M-KCI: R. 2’”o /,>methanol-d,. 12 ‘c’. pH* 280.0~1 M-KC! : 12-C. pH*&81. 0.1 wKC1: I). so”, methanol-d,. -3 C. pH* 3.00. 0.1 M-KCI.

3. Results (a) Environmental

effects on spectra

The effect of increasing the concentration of methanol on the aromatic region of t,he prot,on n.m.r. spectrum of RNAase A is shown in Figure 1. These spectra, with the exception of the 50% methanol system, were taken at temperatures below the denaturation transition temperature, so that only the native state was present. In the ca.se of SOY0 methanol a small amount of non-native material was present. ac.rounting for the resonances between those of the C-2 protons of His48 and HislOh. The similarities between the spectra of the native states in aqueous and methanolic solvents indicate that the presence of methanol does little to perturb t’he native &ructure. The experiment,al renditions chosen for each sample were determined from a

92

R. (:. HIRISGER

ANI) 13..I,. FINli

series of studies in which the ionic strength, pH*. protein and methanol concentrations were varied. In this manner the conditions necessary to produce the best-resolved spectrum of the native protein were ascertained. At low ionic strength (below 0.1 M), in aqueous methanol, the resonances corresponding to the C2 protons of His1 19 and His105 severely overlapped at all methanol concentrations examined, as well as in ‘H,O. Increasing ionic strengt’h (of KU) serves to change the chemical shifts of these resonances so as to improve their resolution, but at the same time lowers the solubility of the protein. Lower values of pH* ( <4) increase the solubility, but decrease the t, (Painter & Fink, unpublished data). The chemical shifts of these protons are also sensitive to pH, and are resolved only over a limited range of pH* values. The particular pH* range is dependent on the cosolvent concentration and ionic strength. Increasing met,hanol concentration decreases the t, (Painter & Fink, unpublished data), and decreases the solubility. In order to eliminate the effect of T1 relaxation on the peak areas, long pulse delays were used. The optimum delay was determined from a series of spectra taken at temperatures where the protein (35y0 methanol) was native (65, 14.4 and 21.O”C), partially unfolded (27.2”C). and fully unfolded (40.2”C). For each temperature 100 acquisitions were obtained with a pulse delay of either 3, 5 or 10 seconds. At temperatures below 27°C the areas of each peak were equivalent ( +50/‘,) regardless of pulse delay. At 27.2”C the areas of all His C-2 resonances exrept that, of His48 (native) were independent of pulse delay. For His48 a pulse delay of 3 s gave a peak area 20% smaller than that at a delay of 5 or 10 seconds. At 40.2”C the peak areas of the unfolded species were independent’ of pulse delay. All experimental data reported below were obtained with a S-second pulse delay in the 20 to 40°C range, and a 3-second delay for all other temperatures. We have previously shown that the denaturation of R,“Aase A in aqueous/methanol solvents is fully reversible as judged by the complete reversal of changes in the near-ultraviolet spectrum and return of catalytic activity (Paint,er & Fink, unpublished data: Fink & Grey. 1978). The unfolding process was also found to be completely reversible as judged by n.m.r.: the same sample could be completely unfolded, partially unfolded, and refolded several times and give essentially identical spectra. (b) Thermal

denaturation

in 25:;

nletharlol

The temperature dependence of the chemical shifts of the signals corresponding to the C-2 protons in the native conformation are shown in Figure 2. The chemical shifts for each of the imidazoles show temperature dependence, consistent with a fast exchange process. Furthermore the temperature-dependent changes in the chemical shift are not uniform. For simplicity we have connected the data points by straight lines, rather than curves. The deviation from linearity for His12 is quite striking. For the other residues the deviations are much smaller but nevertheless clearly beyond the experimental error (f0.003 p.p.m., which lies within the symbols). The reproducibility of the chemical-shift data from sample to sample, coupled with the observation that similar patterns appear in the case of higher methanol concentrations, also indicates that’ these small changes in the slope of the chemical shift versus temperature are real.

INTERMEUIATES

IN

RNAase

ITNFOLI)IN(:

93

FIN:. 2. Temperature dependence of the chemical shifts for the native resonances of the His C-2 protons of RKAase A in 25% methanol-d, (pH*2+30, 0.1 M-KCI). His12 (0); His119 (A); His105 (0); His48 (0).

Starting at 3O”C, new resonances appear in the spectrum (Fig. 3, trace A). These are ascribed to the presence of partially folded species (see below), and have been assigned to particular residues using selective deuteration, as outlined in the next section. We have labeled these resonances Pi, where i corresponds to the residue number of the particular His C-2 proton responsible for that resonance. The temperature dependence of the chemical shifts of the partially folded species (Fig. 4) also shows complex, multiphasic behavior. Breaks in these plots appear at 405 and 51.5”C, and they coalesce into a single peak above 54°C. The latter also exhibits a temperature-dependent chemical shift, suggesting that the unfolded protein is in fast exchange between two or more conformers. Note that the chemical shift of peak PIo5 (Fig. 4) corresponds closely to that of the unfolded state (U), i.e. solvent-exposed. A more detailed analysis is given in the Discussion. The effect of temperature on the areas of the peaks corresponding to both the native and the partially folded states, is shown in Figure 5(a) and (b). The loss of area of the peaks corresponding to the native protein (Fig. 5(a)) reveals a pret,ransition region in which some differences are observed for the different C-2 protons, followed by a co-operative process in which the rate of area loss for each C2 proton is the same within experimental error. (Note that in Figure 5(a), (b) and (c) the areas are represented as the fraction of the total area of peaks in the C-2 imidazole region, i.e. both native, partially and fully unfolded.) This may be compared with the data for unfolding as measured by changes in the ultraviolet

94

Ii. G. KIKIN(:ER

ANI)

A. I,. FINK

FIN:. 3. The spectra of the native and partially folded His C-2 protons. Traw -4. %5’& methanol-d,. pH* 2.80. 0.1 M-Kcl. 442”(‘: R. %‘:,, methanol-d,. pH* 281~ 0.1 M-KU. 35B‘C: C. 50°b rnrthanol-dl. pH* 3GO. 0.1 M-KCI, 33l”CY The numerals at thr top of the Figure refer to the His residues responsible for the resonance. Thr signals in the vicinity of 85 to 8.7 p.p.m. correspond to partially folded species (see text).

light absorbance (which values for the midpoints

reflects Tyr exposure: see Figure of the transition (t,).

(c) Selective deuteration

and saturation

5(d)). Table

tramjw

1 shows the

st,udies

In order to ascertain whether the slow-exchanging resonances (Fig. 3) corresponded to individual imidazole C-2 protons, or to four equivalent imidazoles in four different environments we resorted to both saturation transfer and selective deuteration experiments. In the former, each of the resonances corresponding to the native and partially folded structures was irradiated during the delay period, and decreases in intensity were sought in any or all peaks. No significant decrease

INTERMEI)IATES

IS

KNAase

U’NFOI,I)IS(:

8.65

0.60

40

30

60

50 t PC)

Flc:. 4. Tesmperature dependence of the chemical shifts of the partially mt~thanol-d, (pH* 280. 0.1 M-K(‘I). l’,,9. P,, (0): P,os (0): P,, (A).

folded species in 15”,,

(a) 25 20 15 IO 5 a aC P s 100 z % if

(b)

P

0.2 -

ZS-

a

O.Ok

5C

/

I

4c 3c 2c IC

C IO

20

30

40 f PC)

50

60

IO

20 30 f PC)

40

FIG:. 3 Itelativr populations of various species during the unfolding of KNAasr A in 2ri(& mrt~hanol-d, (pH* PM. 0.1 M-K(Y) as a function of temperature. (a) Area of native His C-2 peaks: His12 (0). -1 I!) (A). -105 ([7). -48 (0); (h) area ofpartially-folded C-2 resonances: 1’,,9. I’,, (0). P,os (a). P,, (A) (see text): (c) sum of thr areas of the native C-2 peaks expressed as a fraction of the total His Cd peak area: (d) ahso~hanc~r at 286 nm reflecting the relative solvent exposure of the huritd tyrosinrs (Mi mg KSAaw A/ml).

96

R. G. HIRINGER

ANI)

A. 1,. FINK

TAHIX 1 Transition

midpoints,

as determined

from n.m.r.

peak area and absorbance changes I, (;(‘f

Methanol

(y<,)

n.m.r.

drg(‘) rl 286

The 1, is defined as the midpoint between the temperature at which all the signal emanated from thr native material. and that at which no signal from thr native state remained. t Hiphasir transition. the t, is given for each phase (see Fig. 13(d)).

was observed, implying either slow exchange between the native state and that (or those) responsible for the partially folded resonances, or no exchange. When incubated in ‘Hz0 at pH* 8 the imidazole (J-2 protons of RNAase A exchange with solvent at different rates, in the order His105 > -119 > -12 > -48 (Markley, 1975). It was thus possible to prepare samples in which the C-2 protons of the imidazole side-chains could be identified unambiguously on the basis of the relative peak areas. Samples that had been exchanged for 3, 6, and 16 days were lyophilized and prepared in 35 and 50% methanol solutions. Some selected spectra illustrating the selective decrease in intensity of some of the partially folded intermediates are shown in Figure 6(a) to (d). The differential changes in the areas of the resonances for the partially folded species, along with the individual magnitudes (as compared to the areas under native conditions), clearly show that each may be assigned to a particular C-2 resonance. Assignments can be made as follows in order of decreasing chemical shift: Hisll9, -48, -105, -12, as indicated at the top of Figure 3. The assignments for both 35 and 500;, methanol are consistent, and quite unambiguous. (d) Thermal denaturation in 35”/;, meth,anol The data for this solvent system are basically similar to those for 25% methanol, but reveal greater detail about the partially folded species. Figure 7 shows the effect of temperature on the chemical-shift values of the resonances corresponding to the native protein. Again, biphasic behavior is observed. the break in the plots occurring near the temperature at which the resonances corresponding to the partially folded species appear (Figs 3, trace B and 8). The temperature dependence of the chemical shift of the partially folded species (Fig. 8), is complex, and multiphasic. Major breaks occur around 25 (the exact value is different for each resonance), 48, 52 and 56°C. The break at 48°C indicates the separation of the lowfield peak into two resonances, Pr,a and P,,. At 56”C, P,a5 and Pi2 overlap, and remain so to the highest temperature examined. Based on the data obtained for 25 and 50% methanol, and examination of the spectrum of free His, we believe that after the coalescence of Pro5 and P,, the chemical shift corresponds to that of a fully solvent-exposed imidazole.

INTERMEDIATES

IN

RNAase

ITNFOLl)ING

FII:. 6. Spectra of the His C-P region of partially folded species with various degrees of selectire deuteration. The incubation times (see text) were, from top to bottom: 16,6 and 0 days, respectively, for (a) and (b). and 16,6.3 and 0 days, respectively. for (c)and (d). (a) 50% methanol. pH* 3~00, (~1 M-KC]. 52°C: (b) as in (a) but. at 40°C: (c) 35% methanol. pH* 281. @l M-KU. 46°C: (d) as in (c) but at 51°C.

R. C:. HIRINGER

98

AND

A. 1,. FINK

a 8.75 t 8.10

8.05

I

0

8.00

48 :_ I/

IO

4(

20 30 f PC)

FIN:. 7. Temperature dependence of the chemical shift of the native methanol-d, (pH* 2.81, 0.1 M-KCI).Symbols as for Fig. 2.

His C-2 resonances in 35”,

8.65

20

30

40

50

60

70

f ("Cl FIG:. 8. Temperaturr dependence of the chemical shifts of the partially folded His C-2 peaks in XV?,, methanol-d, (pH* 281. *1 M-KCI). P,,g (0): P,, (0); FIos (A): PI2 (Cl).

The changes in area in t,he native and partially folded resonances are shown in Figure 9(a), (b) and (c) as a function of temperature, and compared with that monitored by changes in the U.V. absorbance (Fig. 9(d)). The t, values are given in Table 1. The apparent decreased area of the native His48 peak (Fig. 9(a)) probably stems from its slow exchange with 2H+ and 02H-, as noted by Markley (1975), and discussed by Sudmeier et al. (1980). Whereas some of the decreased area of His105

30

20I-

a IC,5 P 6 !2 c,.k= o 5cIis

(b) ,

4c,-

fp-“/

30 2c,IO 0

0

20

IO

30

40

50

60

70

a

PII:. 9. Relative populations of various species in the unfolding of RNAasr .4 in X5’+,, methanol-d, (pH* 2T+1.0.1 M-KU) as a function of temprrature. (a) Area of the native His C-2 peaks. gvmbols as for Fig. 2: (b) areas of the His C-2 peaks from the partially folded species. symbols as for Fig. 7; (c) sum ot the areas of the peaks from the native His C-2. expressed as a fraction of the total His C-2 area; (d) absorbance at 286 nm (WI mg protein/ml).

100

R. (4. BIKINGEH

AND

A. I,. FINK

could arise from overlap errors due to the close chemical shifts of His105 and Hisll9, there is, however, a clearcut decrease in the area of the native His105 resonance prior to the main transition (Fig. 9(a)). The His48 native resonance also indicates a pre-transition decrease in area. This pret,ransition is evident in Figure 9(c), and even more so in the data for 50% methanol (Fig. 13), and also in the absorbance measurements (Fig. 9(d)). Pretransition phenomena have been previously reported for RNAase A in aqueous solution (e.g. see Wang et al., 1980). (e) Thermal

denaturation

in 50% methanol

The temperature-dependence of the native C-2 proton chemical shifts in 50% methanol are shown in Figure 10. The overall shapes of the chemical shift versus temperature plots are similar to those for 35% methanol. New resonances (compared to those from the native state) appeared, corresponding to partially folded intermediates, at temperatures as low as - 10°C (Figs 3, trace C, 11 and 12). The temperature dependence of the chemical shifts (Fig. 12) for the resonances of these partially folded species are again multiphasic, and reveal some interesting features. For example, at 52°C and above the peaks designated PIos and PIz have the same chemical shift as the ultimate fully unfolded species, which is not formed until 75°C. This suggests that the imidazoles of His12 and HislOS are fully exposed to

FM:. IO. Temperature dependence of the chemical shifts of the native His C-2 resonances in 500/, methanol-d, (pH* 390. 0.1 M-KU). Symbols as for Fig. 2.

31.6

62-2

58.8

6-5

-1-l

I

9.0

,

,

,

I

8.5

I

44.7

, 8:O p.p.m.

102

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ANI)

A. I,. FlXK

.

8.721 870

i 8.68 t 8.66 i d ci 8.64 m

:

1

1

1

I

I

1

IO

20

30

40

50

60

I

70

t (“C) FIG:. 12. Temperature dependence of the chemical shifts for the His (‘-2 protons in the partially states in SOY,, methanol-d, (pH*WO, @I wliC1). l’,, (A): P,,, (0); PLO5 (0): P,, (0).

solvent some 25 degC or more below the tem,perature residues become so exposed.

folded

at which the other two imidazole

The chemical shifts of the partially folded intermediates are qualitatively similar to those observed in 35% methanol, at temperatures above 45°C. Below 45°C there are a number of significant differences between the results for the different methanol concentrations (see Figs 8 and 12). Figure 11 shows the imidazole C-2 proton region for a number of temperatures in t’he 50% methanol solvent system. The narrow linewidths throughout the transition region indicate that no significant aggregation can be occurring. Further evidence to support the lack of aggregation comes from the fact that essentially identical spectra are obtained in the transit,ion region with protein concentrations ranging from 10 to 50 mg/ml. The area changes for the native and partially folded peaks are shown as a fimct~ion of temperat,ure in Figure 13(a) and (b). The total area of peaks corresponding to the native state as a function of total area of His (‘-2 prot’ons is shown in Figure 13(c). The transition is clearly biphasic, as is the corresponding transition as determined by tyrosine exposure using changes in the U.V. spectrum (Fig. 13(d)). The t, values are given in Table 1. For Figure 13(c) the t, is taken as the temperature at which the sum of the native-peak areas is equal to the sum of the areas of the partially unfolded species. The transition region, from the temperature ate which the first n.m.r. evidence for non-native material occurs (- 10°C) to that at which all the native state is gone, covers 50 deg(‘! The total transition, from initial evidence of unfolding to complete unfolding covers almost 90 degC. For comparison, the transition region spans about 50 de& when measured by the absorbance change at 286 nm (Fig. 13(d)).

(a)

6 t- 0 @ St-4

0.6 0.4

W

0.2 0.0 E 0020 8 ‘0

0.015

2 x 2

FIN:. 1:s. Relative (pH* 3TM). 0.1 M-KU) Fig. %: (II) awas of PloS (0). P,, (0): His C-Z peak area:

0.010

populations of various species in the unfolding of RNAase A in NV& methanol-d, as a function of temperature. (a) Areas of the native His C-Z peaks. symbols as for the His C-T peaks corresponding to the partially folded sprcirs: I’, ,9 (0). P4s (,I). (r) sum of the arvas of thr native (‘-2 resonances rxpremed as a fracti(m of thr total (d) ahsorbanrr at ~?86nm (MI7 mg/ml).

101

K.

t:.

BIRlS(:ER

ANI)

A.

I,.

FlSli

X number of control experiments were performed to establish that the high concentration of protein used in these experiments did not result in aggregation. Samples of 20 mg prot’ein/ml in aqueous solution, and 25 and 50?, met)hanol, were subjected to laser light-scattering at 25°C’. The calculated diffusion coefficient for the aqueous sample was 11 + 1 x IOF’ cm2 s- ‘, in excellent agreement with the reported value (Rothen. 1940). Af%er correction for the viscosit)y of the met)hanol cryosolvent,s values of IO5 + 1 X IO-’ cm2 s- ’ were obtained for these solvent’s, ($el filtration experiments using Biogel P60 revealed no evidence for species wit)h molecular weights greater t’han that of the monomer. (‘ontrol experiments indicated that, the dimer was readily resolved under our experimental conditions. Sedimentation equilibrium experiments were run with low concentrations of protein and demonstrated the absence of’detectable concentrations of anything but the monomer. Melting curves for t,he thermal denaturation of RNAase A (Ad 286) in methanol cryosolvents are essentially identical regardless of protein c*oncent,ration. The possibilit,y that any of’ t)he observed resonances in the n.m.r. spectra are due to aggregation of the protein may therefore be diswunted.

4. Discussion The present, investigat,ion indicates that) there are advantages in studying protein folding in the presence of organic cosolvents. Clearly the unfolding of RXAase A is more complex in aqueous/methanol than in the absence of rosolvent. Since methanol is more hydrophobic t,han water its presence might be expected to stabilize part,ially folded intermediates during folding or unfolding, if non-polar residues are more exposed in such int’ermediate stat,es than in t)he folded native prot,ein. In other words. the driving force for completion of folding in partially folded species would be decreased in the presence of methanol. This expectation is borne out I)g the observations of partially folded intermediate species, as revealed by n.m.r. (Fig. 3), and the decrease in the co-operativity of the denaturation transition (e.g. Fig. 13(d)). The use of cryosolvents means that the refolding reaction can be followed at subzero temperat,ures. at whic*h the rates will be much slower than at ambient temperatures (Biringer & Fink. unpublished data). Judging by t,he similarity of the spectra in Figure 1 the struct’ure of the native protein (i.e. below the onset of the denaturation transition) in t,he presence of methanol must be very similar to that) of the natire state in aqueous solution. (‘omparison of the chemical shifts. for example, of the peaks from t,he His V-2 protons in aqueous and methanolic solvents indicates that the only significant difference is the chemical shift of His48. This part)iwlar His residue and its chemical shift seem quite sensitive to the protein’s environment. The difference between aqueous and methanol spectra for this residue could reflect small conformat)ional differences in this region of the structure, or perhaps a specific solvent effect. The changes observed in Figure I for t)he chemical shift of the proton of His12 reflect the different pH values at which the spectra were obtained, A detailed analysis of the Tyr and Phe contributions to the

INTERMEDIATES

IN

ItNAase

I’NFOLDINC:

I 0.5

aromatic region of the spectrum using resolution enhancement’(Biringer & Fink, unpublished data) indicates that there are no significant changes in the chemical shifts of these residues on going from aqueous to methanolic solvent systems. The apparent broadening of the resonances from Tyr and Yhe, especially notable in the spectrum from 5Oo/o methanol at -3°C (Fig. l), is due to the close proximity of several resonances, such that a small increase in line-width, due to decreased temperature, increased solvent viscosity, and decreased ring-flipping rates, results in considerable overlap of adjacent resonances. The analysis of the results of the present investigation will be restricted to considerations of the His C-2 protons. The key features involve the interpretation of the observed changes in chemical shifts and peak areas in terms of the underlying structural changes. For the sake of the following discussion, the molecular architecture of RNAase 4 can be considered to be composed of a bent beta-sheet-like structure, effectively forming two wings, with which three alpha-helices are associated (Wlodawer et ab.. 1982). The helix from residues 3 to 13 lies within the cleft of the two wings, whereas those of residues 24 to 33 and 50 to 60 are on the outside of the wings. The two wings with their associated helices form the two domains of the native structure. The hinge region between the wings consists of two antiparallel strands, enc*ompassing residues 75 to 78 and 104 to 106. (a) Analysis

of 25% methanol! data

The chemical-shift behavior, and the peak areas, as the protein is taken through the transition region, reveal a multi-state process. At temperatures below 20°C the chemical shifts of all four resonances corresponding to the native state change in a linear manner with temperature (Fig. 2), and without significant change in peak area (Fig. 5(a)). Such behavior is indicative of an exchange process that is fast on the n.m.r. time scale, and is probably associated with a general loosening of the native structure. We will represent this process by equation (l), in which N’ is similar in structure to the native state, N. The steepness of the slopes for His12 and His48 indicates that significant structural changes are occurring in the vicinity of these residues. NeN’.

(1)

Around 40°C there is a break in the chemical shift versus temperature plots. which is followed by another linear region, of different slope. This observation suggests a new fast-exchange process occurring at temperatures above 4O”C, and is consistent with the scheme shown in equation (2): NL=N’*N”.

(2)

Henceforth, we will represent the three native-like species of equation (2) as [N*]. The difference in chemical shift between N, N’ and N” indicates that they are all similar in structure, with the largest environmental changes in the vicinity of His12 and His48. For all four native resonances, the change in chemical shift as the temperature is raised is toward the chemical shift of the fully unfolded state, 4

106

R. B. HIRINGER

AND

A. 1,. FINK

implying more exposure to solvent as the unfolding progresses, and hence a general loosening of the structure. The partially folded species (Fig. 3, trace A) also exhibit temperature-dependent chemical shifts (Fig. 4), and again are characteristic of a multiphase process. The chemical shifts corresponding to His12 and His105 in the partially folded state show little temperature dependence below 41°C. In fact the chemical shift for His105 is close to that expected for a fully exposed residue. Since the imidazole of His105 is in a very exposed situation to begin with, it is not so surprising that the side-chain is exposed to the solvent at a relatively early stage in unfolding. The changes in the slope of the chemical shift for the other residues in the partially folded intermediates are most consistent with three different species in fast exchange with each other. i.e. equation (3), where P predominates below 4O”C, P’ around 40°C and P” by 50°C: PeP’*P”.

(3)

These species will be referred to collectively as [P*]. At 54°C the chemical shifts all coalesce, presumably reflecting the formation of the unfolded, fully solventexposed state. However, the non-linearity of the chemical shift for U in Figure 4 suggests that there is a fast-exchange process occurring here also. One explanation is that there are at least two forms of unfolded protein present, U’ and U, which we will represent as [U*]. Alternatively (in view of the analysis of the 50% methanol data) it is possible that the fast-exchange process involves at least one partially folded intermediate state, as well as the unfolded state. The changes in area (Fig. 5(a) and (b)) indicate that the slow exchange processes of the system can be represented by : [N*] e [P*J G= [u*].

(4)

All residues exhibit changes in their shift behavior at 41°C. The residue most affected is His12 (native and Pi,). This suggests that the major structural change at this stage involves the N-terminal helix region, whereas the minor effects observed for His119 indicate that major changes do not occur in this region of the active site. Changes in the flexible loop, residues 17 to 25, could explain the effects on His12. Support for this idea is found in the changes observed for His48, since its chemical shift is highly dependent on its relat,ive orientation to Tyr25, and Asp14 (Lenstra et al., 1979). The behavior of the chemical shift of His119 would therefore be an extension of this through a HislB-His119 interaction. Similar behavior for the chemical shifts of both native His12 and Pi2 indicates that the structural difference responsible for their chemical shifts lies within the N-terminal helix itself. At 52°C substantial changes occur about Hisl2, -48 and -119 (native and partially folded) (Fig. 4). In the light of the above results this probably represents the expulsion of the S-peptide region from the cleft, and the subsequent parting of the two domains in the 40 to 50°C range. Further changes probably represent dismantling of the residual beta-sheet in the two domains. The non-coincidence of the area curves of the native resonances below 41°C suggests a pre-transition region (clearly resolved in Fig. 5(c)) in which different

INTERMEDIATES

Izi

RNAase

l’NFOLDIN(~

107

parts of the molecule are undergoing preferential loss of structure. Comparison of the transition curves from the U.V. absorbance measurements to those from the loss of area of the native peaks (Figs 5(c) and (d)) reveals substantial differences in the apparent t, values (39 and 45”C, respectively). In both cases, however, t,he t,ransition covers essentially the same temperature range. In the case of unfolding monitored by U.V. absorbance the experiment measures the exposure to solvent of the four tyrosine residues (25,73,92,97), which are buried or partially buried in the native structure. In the case of the n.m.r. data of Figure 5(c) it is the loss of native environment of His that is responsible for the observed transition. We can therefore conclude that the Tyr residues become exposed to solvent in the early part of the unfolding and are essentially fully exposed when the His C-2 protons are present in the partially folded state. In other words the n.m.r. data reveal that structuw is preseljt after the buried Tyr residues are exposed to solvent, suggesting that the partia.lly folded state(s) reflect(s) secondary structure rather than hydrophobic clusters, or ordered structures involving secondary elements of structure that are packed together through hydrophobic interactions. This line of reasoning will be elaborated subsequently. (b) Analysis of 35% methanol data Since the nature of the analysis and the data (qualitatively) are similar for all t*hree methanol concentrations we will only consider a few of the results in 35O/;, methanol. The chemical shifts of the resonances corresponding to His12 and His1 19 show little temperature dependence below 2O”C, whereas those of His48 and Hisl05, especially the former, exhibit marked temperature dependence (Fig. 7). We interpret these observations to mean that the regions of native structure involving His48 and His105 are in fast exchange with a second conformation (or conformations), while the structure around His12 and His119 is still nat,ive-like. These results are consistent with a general loosening of the structure about the hinge region, or increased movement of the two domains relative to one another, and especially the unravelling of the loop from 17 to 25 linking the S-peptide region t,o the rest of the molecule. The significant change in slope occurring around 2O”C, especially for Hisl2. indicates the beginning of additional structural changes, affecting most of the molecule. but greatest for His12. As in the case of 25% methanol it is the resonances of His12 that show the greatest deviation from those of the native state, implying major changes in the binding of the S-peptide region to the rest of the molecule. The temperature dependence of the chemical shifts of the partially folded species is more complex than in the 25% methanol case. The peaks corresponding to the partially folded species P, r9 and P,s overlap below 48°C (Fig. 8). The changes in slope of the chemical shift versus temperature plots that occur at 48, 52 and 56°C suggest several partially folded species in fast exchange with each other. The residues corresponding to Pi2 and PloS are essentially fully exposed to solvent above 56”C, whereas the residues responsible for Pas and P, i 9 still have residual structure around them, even at temperatures as high as 76°C. The temperature

108

R. (:. BIRINGER

AND

A. 1,. FINK

dependence of the chemical shifts of I’,, and I’,,, at the highest temperatures observed in this study indicate a fast exchange process, probably between partially folded and unfolded states. Examination of the effect of temperature on the areas of the peaks corresponding to partially folded states also reveals complexity in the 45 to 56°C region (Fig. 9). Based on both chemical shift and area effects it is clear that the partially folded species below 48°C ([I?:]) and above 56°C ([I?$]) are significantly different. We can therefore represent the overall unfolding minimally as in equation (5), where the symbol * implies the possibility of more than one similar substate : IN*] G [Iy] T= [P2*] e [Q] t Iv*]. (5) From the chemical shift difference above and below 48°C we can conclude that the major differences between [I’:] and [I’;] involve the regions around residue 12. Similarly, the main differences between [I’?] and [Pt] involve the further solvent exposure of His12 and greater exposure of His48. In [I?$] (above 56°C) His105 and His12 are fully solvent-exposed whereas His48 and His119 still have some residual structure in their vicinity. Thus [P,$] is definitely not a compact, globular structure. Reference to the onset of the unfolding transition as revealed by exposure of the buried tyrosine residues (Fig. 9(d)) indicates that these side-chains become more accessible to solvent during the [N*] to [I’?] transition. This must mean that the structure about these residues becomes significantly dismantled at this stage of the unfolding. The pre-transition loss in area of the peaks corresponding to the native state as a function of temperature (Fig. 9(a)) reveals that the slow-exchange process involving His48 occurs considerably earlier than that for the other residues. In fact the apparent t, for His48 appears 5 degC lower (33°C) than that for the other His residues. Thus the native structure about His48 (Tyr25 and the S-peptide region) is lost considerably in advance of major effects in the vicinity of the other His residues. The data of Figure 9(d) not only indicate that the three buried Tyr residues are fully solvent-exposed at a temperature at which there is still considerable structure remaining, as judged by the n.m.r. His c-2 proton data (Fig. 8), but that the loss of native structure correlates well with that of Tyr exposure (Figs 7, 9(c) and (d)). These observations point out the inherent limitations of using solvent exposure of aromatic residues as a probe for protein structure. Furthermore, it seems likely that the partially folded states present above 48°C ([I’f], [I??]) still have significant structure, probably regions of structure (secondary?) linked by random sections of polypeptide chain. It is interesting to note that at even the highest accessible temperature some residual structure remained in the vicinity of His48 and Hisll9. However, the similarity in chemical shift to that of fully exposed His suggests that the remaining structure in [P3*] is probably minimal. (c) Analysis

of 50% methanol

data

The effect of temperature on the chemical shifts of the native resonances suggests several native-like species in fast exchange on the n.m.r. time scale, which we will represent as [N*]. In general these effects are similar to those noted in 35% methanol. It, is interesting that the signals from the partially folded states apparently first

INTERMEDIATES

IN RNAase I'NFOI,I)IX(:

1O!)

appear at different temperatures. The area of the peak corresponding to native His12 decreases from 25”/0 at - 1°C to 137; at lO”C, and remains essentially at this level until 33°C. Examination of the spectra of Figure 11 indicates that, a resonance is present near X.45 p.p.m. at - 1.1“C, which has not been assigned. This peak was observed in all samples of RNAase A in 50% methanol in the - 10 to 0°C temperature range. Our tentative assignment, based on peak area and chemical shift, is that it is from His48 in a different conformational environment than in the native state and t,he partially folded state in which the His48 C-2 is assigned to I’,, (Fig. 3, trace V). This interpretation implies an additional partially folded intermediate. in slog exchange with the native state, responsible for the resonances at 8.73 and 8.45 p.p.m. On the basis of the chemical shifts we would assume that this intermediate, formed early in the unfolding, has a relatively compact st~rurture. at least compared to the subsequent partially-folded species. A minimum interpretation of the fast-exchange processes indicates the exist)ence of at least four, and most likely nine, different partially folded species : individual species predominate at 0, 10, 22, 28, 34, 38, 45, 52 and 66°C. It is apparent that there are major differences between the conformations of some of these partially folded species, which are in fast exchange with each other. This can be readily seen from a comparison of the spectra in Figure 11, and from the effects of temperature on chemical shift, and peak area. From the chemicalshift data we will represent the species below 20°C as [ PT] ; [I’? J will represent the substate between 20 and 3O”C, [I’31 will represent those bet)ween 30 and 52°C’. and [I’,*] those between 52 and 66°C. Perusal of the area data of Figure 13(c), and the Tyr exposure curve (Fig. 13(d)) suggests that a significant structural change is completed around 20°C. It, is noteworthy that a change in shape of the chemical shift VPTS’U~S temperature data is also observed for all the native resonances at this temperature (Fig. 10). The species above 75°C is assumed to be fully unfolded. and will be represented as U. However, as can be seen from the peak width in the spectrum of Figure 11 there is still evidence for some residual structure a,t 75°C’. The instability of the solvent at high temperature precluded experiments above 75°C: therefore, we cannot make any judgements as to whether I! represents a single population or not. The minimum pathway for unfolding in FiOO’,,met,hanol is shown in equation (6), where * denotes the possibility of several similar substates:

The fact that the transition in this case covers some 90 degC implies drasticall) reduced co-operativity compared wit,h the situation in the presence of 10~ concentrations of methanol, or in its absence, as well as the potential for a detailed evaluation of the pathway for unfolding (see section (d), below). The decreased cooperativity is a reflection of the stabilization of partially folded int,ermediates h> the cosolvent. (d) Pathway

for folding

and unfolding

It is reasonable on both theoretical and experimental grounds to consider that the order in which the protein unfolds from its native state will mirror the pathway

110

K.

(:.

KIKISGER

ANI)

A.

I,.

FINK

in which folding occurs. It also seems reasonable to assume that intermediates formed early in t,he unfolding transition (i.e. native-like conditions), would be likely to occur early in t’he kinetic pathway of unfolding, and those formed under strongly denaturing conditions in t’he unfolding transition would occur late in the unfolding pathway. Thus, if we can determine t’he chronology of unfolding, we can assume that refolding would be the exact reverse. under comparable experimental conditions. Confirmation of this assumption comes from : (1) the observation that t’he spectrum obtained is identicaal at any temperature in the transition region, whether it is approached from below, starting with native enzyme. or from above, starting with unfolded protein : and (2) also from preliminary refolding experiments at subzero temperatures in which part’ially folded species, closely resembling those seen in unfolding (Fig. 11) are observed (Biringer &r Fink, unpublished data). Our present analysis of the folding pathway is limited by t,he fact that, effectively, we have only the four imidazoles as probes, and can therefore only provide details about the environment~s of these residues. Fortunately. these His residues are strategically located. We will now describe t,he picture that emerges from the analysis of the SO’& methanol data for the pathway of unfolding. This analysis may also be applied to the 35?, met’hanol data, as t,he patterns of chemical shift and area changes are similar. The chemical-shift data (both slow and fast) exchange) reveal that the earliest stages of unfolding involve the environment, of His@. The immediate struct,ure about the imidazole of His48 involves Tyr25. and the region around residues 14 to 18. A likely cause of the observed effects would be the unfolding of residues 13 to 25 (cf. blatheson & Scheraga, 1979). One would expect such changes to affect’ His12 as well. The chemical shift of His12 at this point in the unfolding indicat,es that the His side-chain is st’ill in an environment not) very different from that in t,he native st)ruct’ure. and thus we expect the helix (residues 3 to 12) to be still intact. The change in absorbance due to Tyr exposure (Fig. 13(d)) also reveals a minor t)ransition. complete by dOY”, followed by the major transition. It is possible that the minor transition corresponds to the exposure of one of the three buried Tgr residues, t,he major transition being the exposure of the remaining t’wo. If this interpretation is caorrect. the init,iallg exposed Tyr would be residue 25. This in turn means that Tyr73 and Tyr97 become exposed to solvent in the 20°C to 40°C region. The continued change in chemical shift of the native resonances as the t,etnperature rises to 38°C’ indicates that the native structure undergoes a progressive loosening, the trend being in the direct’ion of more solvent exposure for all four (‘-2 protons. The area of the peaks from His105 and His119 in the nativelike state indicates a pre-transition change around 20°C. Both residues are part of the beta-sheet structure involving the C t,erminus of the molecule. and it’ is reasonable to expecat that significant’ structural changes in the vicinity of one of these residues might be coupled wit)h those of the other. It, is possible t,hat movement apart of the tw:o domains (wings) of ribonuclease A would result in such effects. At 40°C’ all the native peaks have disappeared. and an abrupt rhange in slope of the chemical shift for I’,, occurs. Coupled with further changes over the next

IS’I’EKhlEI)IATCS

IN RXAast,

17NFOI,I)IN~:

III

10 deg, these observations probably reflect further separat,ion of the domains and dismantling of the sheet’ structure in the region of residues 42 to 49 and SO to 86. In addition, the absorbance changes reveal that the formerly buried Tyr residues are now all exposed to solvent at this point. It is interesting to note that if the last two Tyr residues to become exposed are Tyr73 and Tyr97. the latter does interact with this region of beta-sheet. The rapid area changes in both native and partially folded peaks near -WC’ indicate that a rather large structural change occurs at this stage. IVe believe that this change corresponds to the movement of the S-peptide region residues (1 to 25) away from the two domains. This would allow the t,wo domains tjo open signific*antlg and thus explain a massive change in all peak areas. We base this on two criteria : (1) agreement with the spectrum of t,he S-protein (Fig. 14) ; and (2) consist’ency with the data reported by Matheson & Scheraga (I 979). The proton t1.rn.r. spectrum of the S-protein was obtained in .?O~, methanol. under identi4 conditions to the RXAase A spectra and is shown in Figure 1-C.The spectrum of the S-protein at 23Y7 is similar to that of RNAase A at 10°C’. Since the S-protein does

11%

R. (:. HIRIS(:EK

AND A. I,. FISK

not have the S-peptide region to interact with it, we conclude that at 40°C the Speptide no longer interacts with the two wings of the native RNAase A structure. To be consistent with this the chemical shift of P,, in the 40°C t,o 50°C (50% methanol) region must only reflect interactions between Phe8, Glnll and HislZ. Thus the small change in the shift behavior of P,, at’ 40°C indicates that the major interaction responsible for t,he chemical shift of P,, is due to intra-helix, rather than peptide-protein, interactions. At 52°C the chemical shifts of P,,, and P,, merge and in fact assume the same chemical shift as fully exposed histidine. Therefore, we conclude that at this point the S-peptide region, containing the N-terminal helix, has completed unfolding, thus exposing His12 to solvent. Brown & Klee (1971) have reported that the helix in a peptide consisting of the first 13 residues of RNAase unfolds in aqueous solution at temperatures above 26°C. The complete exposure of His105 (Pro5) at this stage indicates that the residual structure in the vicinity of Hisl05, and in part’icular the C-terminal region of beta-sheet (residues 105 to 124) becomes ‘unhinged”. Sime His119 is not fully exposed at this stage, the results suggest that there must still be some residual structure in its vicinity. Similarly, the existence of residual st’ructure in the vicinity of His48 indicates that t,here is still some of the beta-sheet from residues 41 to 49 and 80 to 87 remaining. At 66°C t’he remaining structure involving His48 (P4s) and His1 19 (PI r9) begins to disappear. Thus, to a first approximation, one can characterize the unfolding process as beginning with a general loosening of bhe native structure. followed by the separation of residues 12 to 25 from the remainder of the molecule. Subsequently, major structural changes occur throughout the molecule, reflecting the separation of the N-terminal region (residues 1 to 25) from the cleft between the wings of the molecule, and leading to the formation of partially folded substates ([Pf]). These are characterized by t’he essentially full solvent exposure of His105 and Tyr25, but retention of the helix involving residues 3 t’o 12. In ]Pz] further changes have occurred in the vicinity of His48 and His1 19. These changes may signal the partial unravelling of the beta-sheet structure involving residues in both t,he C-terminal and hinge regions. In ]PJ] the His12 side chain becomes fully exposed to solvent, reflecting the unfolding of t)he N-terminal helix, and the remaining structure about His48 and His1 19 begins to disappear. One possible interpretation of the results is that in the partially folded states we have regions of secondary structure linked by random polypeptide chain. Thus secondary strucoture is lost last in unfolding, and by implication, is formed first in refolding. Blum et al. (1978) have previously report,ed n.m.r. evidence from studies in aqueous solution t,o support the early formation of the N-terminal helix during refolding (see below). (e) Comparison

to n.m.r.

studies in aqueous solution

Westmoreland 8r Matthews (1973) investigated the thermal unfolding of RNAase A at pH 1.3. Their key observations were that the loss of area for the resonances of the native His C-2 protons was not simultaneous, implying at least two regions of the molecule unfold at different temperatures; and temperature dependence of t,he chemical shifts, implying fast-exchange processes involving a

INTERMEI)IATES

IN RNAaue I'XFOLI)ING

I I3

minimum of three native-like species. As with the results in the presence of methanol the environments of His12 atid His48 began to change first. The unfolding at pH 1.5, 2.9 and 55 was also studied by Benz & Roberts (197%). Their data are also consistent with different regions of the protein unfolding independently, and with early changes in the regions of Hisl2, -48 and -119. In a related study, using unfolding induced by guanidine . HCl or urea, they report da,ta consistent with local unfolding around His12 (Benz & Roberts, 19756). Further evidence for the existence of partially folded species in the thermal unfolding of RNAase A comes from a recent ’ % n.m.r. study by Howarth (1979) ; the conclusion of this work was that the first region to unfold is residues 17 to 25, which in turn affects the environment of His48, due to the Tyr25-His48 interaction (Lenst’ra et al.. 1979). ThcJ kinetics of refolding of RNAase A at lO”C, pH 2, was studied by Blum et r/l. (1978) by monitoring the His C-2 protons at 360 MHz. Two non-native peaks were observed and attributed to fully unfolded (U) and partially folded (X) states. The latter was assigned to the C-2 proton of His12 in an S-peptide-like environment. On t,his basis they concluded that the S-peptide region refolded more rapidly than the rest of the molecule, since the areas of the U and X peaks decreased simultaneously. and at the same rate at which the native resonances appeared. The relativr positions of U and X are the same as those observed for U and YIZ (the partially folded resonance assigned to Hisl2; see Figs 3, 11 and 1%) in aqueous/methanol. (“lose examination of the resonance U in Figure 2 of Blum et al. (1978) indicates that the peak probably consists of two closely spaced resonances. In view of the results in aqueous/methanol it is quite possible that the resonance U, assigned t)o the fully unfolded state may in fact represent the signal from a partially folded species, and be equivalent to peaks P4s, P,,,, P,,, in Figures 11 and 12. for example. ApJjlication of the T,, off-resonance technique to obtain inforrnation regarding internal mobilit’y of RNAase A during guanidine. HCl-induced unfolding indicates increased mobility about His12 and His105 prior to the beginning of the transition (,James & Sawan, 1979). Sevclral previous investigations have indicated that the unfolded form of KXAase A often studied may not be completely random, but still possess some rcJsidua1 structure (Benz &, Roberts, 1975a; Westmoreland & Matthews. 1975: Howarth. 1979: Lenstra et al., 1979). These observations are also consistent with our results in aqueous/met,hanol. The results in 25% methanol show a single peak with essentially the expected chemical shift for solvent-exposed His, From the temperature dependence of the chemical shift (Fig. 4) it is clear that several unfolded conformations are in fast, exchange. The separation of the signals from the part,ially folded intermediates is much smaller in 25% methanol than with the higher methanol concentrations, and we suspect that, the single peak observed above 55°C is a composite of closely overlapping resonances from partially folded intermediates. In 35% methanol the protein is not fully unfolded even at 79°C’. whereas in riO:i, methanol it is close to fully unfolded at this temperature. This is a consequence of the overall denaturation transition occurring at a lower t,emperat,ure in 500/, methanol compared to 350/o methanol.

114

K. (:.

HIRIN(:ER

AXI)

A. I,. FINli

Comparison of the temperature at which denaturation is complete as judged by exposure of buried Tgr (measured by u.v. absorbance). with t’hat at which unfolding as measured by n.m.r. is complete. indicates t,hat Tgr exposure occurs at experiments at subzero a relatively early stage of unfolding. In wfolding temperatures we find the converse, namely t)he large u.r. absorbance change occurs late in refolding (Lust,ig Pr Fink, unpublished observat’ions). These results in methanol are consistent, with proposals that Tyr92 and ‘I+-25 are exposed in the initial stages of unfolding in aqueous solut,ion (Burgess & Scheraga, 1976). The limited amount of n.m.r. dat,a involving intermediates in RNAase A folding/unfolding in aqueous solution, which can be compared with the results in aqueous/methanol, show not,hing to indicate that the process is sign
(f) Oth‘rr

e~~idpnw for

intcwnedintes

in R1VL4nse A folding

Baldwin and co-workers have found evidence for the existence of discret’e intermediate states in folding and unfolding in aqueous solution. Both kinetic (Hagerman 6 Baldwin. 1976: Sal1 et al.. 1978: Cook et nl., 1979: Schmid. 1980,1981 : Schmid & Blaschek, 1981) and structural (Blum et al., 1978: Schmid & Baldwin. 1979) evidence for the presence of at least two partially folded intermediat’e stat,es has been found. For example. by tritium-labeling the exchangeable amide protons of the unfolded protein and then refolding under conditions where exchange-out, is slower than refolding, Schmid &, Baldwin (1979) showed that a significant1 number of tritium atoms were trapped in this manner. implying the existence of a stabilized, h4’drogen-hondetl structure at’ an early stage of refol&g. Several other approaches hare yielded indirect evidence for the step-wise unfolding of RNAase A. Many of these hare been summarized by Burgess & Scheraga (1975) and by Ma,theson 8 Scheraga (1979). However, the present work is the first in which detailed spectra of such species have been obtained.

(g) ~‘onclrrsion
INTERMEDIATES

IN

RNAasp

I’SFOLI)IS(:

I 1.i

cleft and then disruption of the N-terminal region of beta-sheet. concomitant with exposure of the remaining tyrosine residues and the uncoiling of the N-terminal helix. ultimately followed by loss of the remaining structure. especially in the sheet (il to 74, which is undoubtedly stabilized by the disulfide between (‘y&5 and C’ys7P. (3) The data in 35% and 5O”/b methanol reveal at least three distinct partialI>, folded substates. In fact. comparison of t,he results as t,he concentration of’ methanol is progressively increased reveals progressively more st,abilization of the partially folded states. At this stage it is not possible to debermine whether the!? represent, successive stages in the folding process. or whether, at certain stages. the preponderance of evidencr* multiple parallel pathways occur. However. suggests that a series of discrete st)eps occurs. (1) The information provided by the His (‘-2 probes alone does not unambiguously allow one to determine whethkr in the folding process the initialI> formed structure involves secondary structure or hydrophobic clusters : however. we strongly suspect that it) is the former. It is clear from the Tyr exposure data that the hydrophobic “cores” t,hat exist in the native structure cannot be present in the partially folded specaies. In fact the data are inconsistent with any hydrophobic, clusters involving Tyr in the partially folded states. We therefore conclude t,hat the last structural elements in unfolding are secondary in nature. and hence. in refolding. the first’ structure must be of hydrogen-bonding origin, rather than a hydrophobic cluster. This rrwarch was supported by grants CM20555 from The Xational Itrstitutvs of Health. ilnd I’(‘M79- 13766 from Thtb National Science Foundation. as well as grants SSF (~P43633 and R’IH RR0071 1 to Stanford Magnetic R(~sonance Laboratory. The technical assistanw of I)r I). Kar in preparing the initial samples. and I)r A. Ritrriro in the !WK-\var~ is graW’ull> ~rppr~~(+Lted.\I’e thank Professor Richard Shaf&r. I-.(!. San Franc~isw. for rutlning the lights sc*attwing r~xperimrwts. Thta instrumentation for thrsr t~xpwittwnts wits purvhawtl \\.ith funds from grant no. I’41 (N274!&?, from the Kat,ional Institutes of Htaalth. We also t’xprtw our appreciation to I)r Alan Lohr at Stjanford ITnirrrsity for running thtb rllt,rHc~t,trtrifilg;Itioll t~xprrimcwts.

REFERESCES Eatdwin. R. I,. (1980). In I’rotri~ Po/dir,g (Jaenickr. R.. rd.). pp. 36%386, fSlserier!Xorth Holland. Nrw York. R. I,. & (‘reighton, T. (1980). In I’rotri~ b’oldittg (,Jatwi(akr. R.. wi.). pp. 217 260. Baldwin. E:lstvier/North Holland. X;ew York. Gnz. F. W. bt Roberts. (:. (‘. K. (1975r/). ./. lMo/. Biol. 91. 335-365. Htbnz. P. W. & Roberts. (:. (‘. K. (19750). ./. Mol. Rid. 91, 367--X35. Hlum. A. I).. Smallcomb~. S. H. 8: Baldwin. R. I,. (1978). ./. IVo/. Rio/. 118. 30,Sp31fi. Hrandt.s. *I. F., Halvorwn. H. R. & J3rennan. M. (1975). Riochetttistr,t/. 14. 4!).‘,3-4!)63. Hrown. .J. l-2. & KIw. W. A. (l!J71). Hiochetttistry. 10. 476-479. Hurgrss. A. W. & Schrraga. H. (1975). .1. Thror. Rio/. 53. 403 420. (‘havrz. I,. (Z. Jr 6t Srheraga. H. (1980). Riochemistry. 19. 1003-1012. f‘ook. K. H.. Schmid. F. X. 8: Raldtvin. R. 1,. (1979). I’roc. Sort. .-Irrrd. Sri.. f~.S..-i. 76. 615i 6161. I)ouxou. I’. (1977). (‘ryohiochevttistr!/. Academic Press. R;cw York. I)osc~hvr. .\I. 8r Hirs. (‘. H. W. (1967). Riochettrixtry. 6. SO+ 312. Fink. A. I,. (l!lii). .-1c~ts.Chettt. RPS. 10. 233-339.

116

H. (:. HIKINCRR

AXI)

A. I,. FINli

Fink. A. I,. & Cartwright. S. *J. (1981). CRC CM. Rm. Biocknc.. p. 145. Fink. A. I,. & Grey. H. I,. (1978). In Riornoleculor A’tructure clnd Function (Agris. I’. F.. Sykes. B. & Loeppky. R.. rds). pp. 471-477. Academic Press, New York. Hagerman. P. ?J. & Baldwin. It. I,. (1976). Biochemistry. 15, 1462-1473. Hagerman. I’. J.. Schmid. F. X. & Baldwin. R. I,. (1979). Riochemistryy. 18. 293-296. Howarth, 0. W. (1979). Riochitr~ Riophys. Actn, 576. 163-175. Hui Bon Hoa. (:. & Douzou. I’. (1973). J. Hiol. Chrrrc. 248. 4649-4656. Jarnickr. R.. ed. (1980). Pro/& Fold&g. Elsevier/Korth Holland, New York. James. T. I,. & Sawan. S. P. (1979). J. .-!~wr. Chant. Sot. 101. 70.50-7055. Len&a. CJ.A.. Holscher. H. (:. *J. 11.. Stab. S.. Reintema. J. J. &, Kaptein. R. (1979). E:ur. J. Riochern. 98. 385-397. Matheson. R. R. ,Jr. R: Srheraga, H. A. (1979). Hiochenktry. 18. 2437F2445. blarklry. J. I,. (1975). J%ochrrnistry. 14. 3554-3561. Nail. H. T.. CareI. J. &r IMdwin. R. I,. (1978). J. Mol. Riol. 118. 317-330. Semethy. (:. 8: Schrraga. H. .4. (1979). I’roc. Srrr. dcnd. Sci.. IT.S.A~, 76. 6050-6054. Pitti. E:. 1’. 8: Timasheff. S. R’. (1978). Riochpwistry. 17, 615-625. Roltson. H. & Pain. R. H. (1976). Rio&err/. .J. 155. 331-344. Rothen. A. (1940). J. C:u,i. J’hpiol. 24. 203. Schmid. F. S. (1980). In /‘r&iv Folding (,Jaenickr. R.. rd.). pp. 387-400, Elsevier/North Holland. Il’rw York. Schtnid. F. X. (1981). Eur. .J. Niochern. 114. IO&IO!). Snt. dcnd. Sci.. C.S..-l. 75. 4764-4768. Schtnid. F. S. & Baldwin. R. I,. (1978). l’roc. Schtnid. F. X. 8: Baldwin. R. I,. (1979). .J. Mol. Rio!. 135. 199-215. Schmid. F. X. & Blaschek. H. (1981). Ew. ,/. Riochern. 114. 111-117. Sudtnrier. .I. I,.. Evrlhoch. J. I,. 8: ~Jonsson. K.-H. (1980). .J. Mngnrt. Res. 40. 377-390. Taborsky, G. (1959). ./. Rio/. (‘hem. 234. 2652-2656. Tiktopulo. E. I. P: l’rivalov, 1’. IA. (1978). FII:RS Letters. 91. 57-58. Wang. (‘.-C., Cook. K. H. & Pwora. R. (1980). Riophys. Chem. 11. 439-442. M’rsttnoreland. I). (:. RTMatthews. (‘. R. (1973). I’roc. .Vat. =Icnd. Sci.. U.S.A. 70. 914-918. M’lodawer. .4.. Kott,s. R. & Sjolin. 1,. (1982). .J. Riol. Chern. 257. 13251332. Edited hy S. Rrennu