Carboxymethyl horse-liver alcohol dehydrogenase

Carboxymethyl horse-liver alcohol dehydrogenase

ARCHIVES OF BIOCHEMISTRY AND 168, 145-162 (19751 BIOPHYSICS Carboxymethyl Horse-Liver Alcohol Ligand-Binding and Kinetic Cysteine-46-Modified ...

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ARCHIVES

OF BIOCHEMISTRY

AND

168, 145-162 (19751

BIOPHYSICS

Carboxymethyl

Horse-Liver

Alcohol

Ligand-Binding and Kinetic Cysteine-46-Modified C. HUGH Department

REYNOLDS’

of Biochemistry,

University

AND

JOHN

of Edinburgh

Properties Enzyme

of the

S. MCKINLEY-McKEE2

Medical United Kingdom

Received October

Dehydrogenase

School, Edinburgh

EH8 9AG Scot/and,

17, 1974

Horse-liver alcohol dehydrogenase was carboxymethylated with iodoacetate, which is known to selectively alkylate cysteine-46 in the polypeptide sequence. Carboxymethyl and native enzyme had the same electrophoretic mobility on starch or polyacrylamide gel, but some separation was achieved when isobutyramide and a low concentration of NADH were present (under these conditions NADH was bound by native enzyme but not by carboxymethyl enzyme). The carboxymethyl enzyme formed ternary complexes with NAD+ and pyrazole or decanoate. The fluorescence emission of NADH was enhanced 7- to B-fold (at 410 nm), and a dissociation-constant of 1.7 PM was calculated at pH 7.4; but, in contrast to native enzyme, neither the affinity nor fluorescence were increased by amides (acetamide or isobutyramide). Carboxymethyl alcohol dehydrogenase possesses catalytic activity. Higher alcohols gave maximum velocities up to 7-fold higher than ethanol (reaching nearly 20% of the activity of native enzyme) while [2H]ethanol showed an isotope-rate effect of 3.3. Although the affinity for aldehydes was considerably increased, the maximum velocity of aldehydereduction was always at least 20% of that shown by native enzyme, and at pH 9.9 it was almost 2-fold greater than with native enzyme. The rate-limiting step in alcohol-oxidation is likely to be the interconversion of ternary complexes (possibly the hydride-transfer step), while in aldehyde-reduction it could still be the dissociation of the enzyme/NAD+ complex. This is also indicated by inhibition experiments with decanoate, pyrazole, and isobutyramide. These results suggest that a major effect of carboxymethylation is upon ternary complexes of enzyme and NADH, which become much more reluctant to form, either by combination of NADH and ligand with the modified enzyme, or by catalytic conversion of the enzyme/NAD+/alcohol complex.

Horse-liver alcohol dehydrogenase (EC 1.1.1.1) has 14 thiol groups per subunit, of which one is reactive towards iodoacetate (l-4); this has been found to be cysteine-46 in the amino-acid sequence (5). Kinetics of inactivation of the enzyme by iodoacetate have shown that iodoacetate can bind ’ Present madu Bello Nigeria. * Present sity of Oslo,

reversibly to the enzyme (6); other ligands such as substrates or coenzymes or their analogues, can affect both the reversible binding and the rate of alkylation (4, 6, 7). Studies on the cysteine-46-carboxymethyl enzyme, described here, complement the above investigations, and show that the most prominent result of this alkylation is to weaken ternary complexes involving NADH, either with aldehyde or with inhibitors. Interpretation of these results is aided very considerably by knowledge of

address: Department of Chemistry, AhUniversity, Zaria, North-Central State, address: Biochemical Oslo, Norway.

Institute,

Univer145

Copyright 0 1975 by Academic Press, Inc. All rights of reproduction in any form reserved.

146

REYNOLDS

AND MCKINLEY-MCKEE

the high-resolution structure of the enzyme determined by Br;indBn and co-workers (8, 9), showing that cysteine-46 is one of the ligands to the active-center zinc atom. Preliminary reports of this work have appeared (1, 10). MATERIALS

AND

METHODS

Horse-liver alcohol dehydrogenase was purchased from Boehringer Mannheim GmbH (Mannheim). It consisted very largely of the EE isoenzyme (11). NADH was obtained from Sigma Chemical Co. (St. Louis, MO), and had a residual absorbance of 2-2.5% at 340 nm, after enzymic oxidation with native liver alcohol dehydrogenase and excess acetaldehyde or butyraldehyde. NAD+ was supplied by Boehringer Mannheim GmbH or P.-L. Biochemicals (Milwaukee, WI). Iodoacetic acid and imidazole were recrystallized as described previously (7). [2H]ethanol (ethanold6) was obtained from Koch-Light (Colnbrook, Bucks, England). Acetaldehyde and n-butyraldehyde were distilled before use. Isopropanol (T. Schuckardt, Miinchen) was guaranteed 99.9% pure. 4-Iodopyrazole was a gift from Prof. R. Hiittel. Other reagents were the purest commercially available. Details of enzyme assay and purity have been given previously (6. 7). Carboxymethylation of horse-liver alcohol dehydrogenase was carried out by allowing iodoacetate (4-12 mM) to react with a dialyzed sample of the native enzyme (1 ml, 50-110 PM, in 40 mM sodium phosphate buffer, pH 7.4) at room temperature (approximately 20°C). Imidazole has been shown to enhance the reactivity of liver alcohol dehydrogenase with iodoacetate (4, 7), and therefore carboxymethylation was routinely performed in the presence of this compound (l-14 mM). It is probable that only the thiol roup (cysteine-46) becomes more “reactive” reactive in the presence of imidazole, thus minimizing possible side-reactions with other residues. In the presence as well as the absence of imidazole, one iodide ion was liberated per inactivated subunit (4). Carboxymethyl enzyme prepared in the absence of imidazole had identical properties regarding residual activity, complex-formation with NAD’ and pyrazole, binding of NADH and effect of amides on this. Therefore it seems unlikely that imidazole is labilizing a residue other than cysteine-46. After l-2 h, when the remaining activity was constant at 2-2.5% (with ethanol as substrate), the protein was separated from imidazole, iodoacetate, and iodide by passage through a column (1.4 x 12 cm) of Sephadex G-25, Medium grade (Pharmacia, Uppsala. Sweden). Spectrophotometric measurements, including enzyme assays, were made using a Gilford Model 2000 recording spectrophotometer, thermostatted at 23.5”C. Fluorimetric measurements for ligand-bind-

ing studies of NADH were performed using a recording spectrofluorimeter (Farrand Optical Co. Inc., NY), with a 150-W xenon-arc lamp: silica cells (1 cm square) were housed in a sample-chamber thermostatted at 23.5”C. Each monochromator had a slit before and after it: for scanning spectra these were (in order in the light-path) 10. 5, 10. and 5 nm. For dissociation-constant determinations these were 20. 20, 10. and 10 nm. Fluorescence measurements of initial-rate kinetics were performed using a sensitive filter fluorimeter with a mercury lamp, developed in this laboratory. Its sample-chamber was thermostatted at 23.5”C. For fluorescence titrations of carboxymethyl enzyme with NADH, excitation was at 325-335 nm. Emission was usually measured at 410 nm. Aliquots of NADH solution were added on a glass stirring rod, while the fluorescence was continuously monitored by the recorder. Concentration increments of NADH were usually 10 x 0.06 pM, 7 x 0.19 KM, and 5 Y 1.2 FM. The fluorescence of a perspex standard was measured before and after each titration; when discrepancies were noted (due to the arc of the lamp jumping) the results were discarded. The standard was only placed in the light-path for short periods of time, as its fluorescence tended to decay. For calculation of dissociation constants of NADH from the results of fluorescence titrations, it was decided to use a graphical method. It was possible that the two subunits in the liver alcohol dehydrogenase molecule might not have behaved independently and identically with respect to either their affinity for NADH or the fluorescence-enhancement upon binding. Native liver alcohol dehydrogenase shows identical and independent sites for reversible NADH-binding (12. 13) but not for pre-steady-state catalysis under some conditions (14). After removal of the enzyme’s zinc atoms, the inactive enzyme can bind NADH, but the sites now appear to be nonequivalent (15), and it was conceivable that carboxymethylation might also render the binding sites nonequivalent. The method chosen for the present investigation was to plot the ratio of concentrations of bound coenzyme to free coenzyme against the concentration of bound coenzyme. This method (which has been used by McKay and Kaplan (16)) is the same as a Scatchard plot (li), but with the axes inverted. When linear, the apparent dissociation constant is given by the negative of the gradient. The fluorescence values with and without enzyme were divided by the value of the perspex fluorescent standard and subtracted to give the enhancement of fluorescence due to some coenzyme becoming enzyme-bound. In order to calculate the concentrations of bound coenzyme it was necessary to know both the concentration of enzyme binding sites and the fluorescence-enhancement that would have been produced when the enzyme was saturated with NADH. The concentration of binding

CARBOXYMETHYL

LIVER

sites was determined by titration with 4-iodopyrazole in the presence of excess NAD ’ (18). while a value for the saturated fluorescence-enhancement was estimated by inspection. If the estimate was inaccurate, this resulted in the graph not extrapolating to the known concentration of binding sites: recalculation with a revised estimate was straightforward. Calculation was greatly facilitated by the use of an Olivetti Programma PlOl desktop computer. At very low concentrations of NADH (below 1 pM) the fluorescence of NADH was anomalously high. with or without enzyme present. This effect was subsequently found to be largely due to aspiration of air through the micropipette used to add aliquots of NADH after cleaning. It may have been due to traces of fluorescent material picked up from the atmosphere. Although this effect should be self-cancelling, it may have contributed to the imprecision seen at low coenzyme concentrations in some of the experiments reported here. Starch-gel electrophoresis was performed in horizontal blocks (19). Vertical disc polyacrylamide-gel electrophoresis was performed in apparatus manufactured by Shandon Scientific Instruments Co. (London). with gels according to Cruft (20). Amino-acid analysis was performed by the accelerated method of Spackman, Moore, and Stein (21) in a Locarte amino-acid analyzer. The content of thiol groups in the enzyme was measured by reaction with 5.5’-dithiobis-(2.nitrobenzoic acid) in 8 M urea (22). Initial-rate measurements of enzyme activity were routinely made in duplicate. with enzyme being added last on a glass stirring rod. However, glycine reacts slowly with aldehydes, and therefore for measurements of aldehyde-reduction in glycine buffer (pH 9.9). aldehyde was added last to start the reaction. RESULTS

General properties of carboxymethyl alcohol dehydrogenase. Carboxymethylation rendered the enzyme somewhat less stable. On keeping, the concentration of binding sites for NAD+ and pyrazole decreased (by about 10% after passing through Sephadex G-25, and reaching 20-30s after 24 h or more at 0-4”C, or after dialysis). This was accompanied by a similar loss of thiol groups: in native horse-liver alcohol dehydrogenase 28 per molecule were found, or 14 per polypeptide chain, in accordance with previous reports (5, 23, 24). The residual catalytic activity (25) also decreased proportionally. Carboxymethyl alcohol dehydrogenase reacted faster than native enzyme with Ellman’s reagent (22)

ALCOHOL

DEHYDROGENASE

147

but nevertheless much slower than nonprotein thiols. Amino-acid analysis (after 24-h hydrolysis in 5.5 N HCl at 1lO’C) of carboxymethyl enzyme showed 1.0-1.2 residues of S-carboxymethyl-cysteine per subunit; no carboxymethyl-histidine (N-l or N-3) was detected. Native enzyme showed no methionine sulphone, which if present could have interfered with analysis of S-carboxymethyl-cysteine. These results are in good agreement with those found from incorporation of [‘%]iodoacetate into the enzyme (2). Polyacrylamide and starch-gel electrophoresis at pH 7.4, pH 9.3, and (with starch-gel) pH 8.4 revealed identical mobilities for the main component of carboxymethyl enzyme and the main (EE) component of native enzyme, by protein staining (with naphthalene black) or activity stain (with butanol as substrate). In more concentrated preparations of carboxymethyl enzyme, a component equivalent to the ES isoenzyme was also seen (11). In an effort to induce electrophoretic separation of native and carboxymethyl enzyme, use was made of an earlier observation (19) that anionic ligands. including NADH, reduced the cathodal mobility of native alcohol dehydrogenase. From the binding data (presented below) it was anticipated that at low concentrations of NADH (0.01-l PM), in the presence of isobutyramide, native enzyme would be largely in the form of the enzyme/NADH/ isobutyramide ternary complex (13) while the carboxymethyl enzyme would be largely uncomplexed. This device was successful in producing some separation of native and carboxymethyl enzymes (Fig. 1); as expected, the native enzyme was retarded. The reverse side of the gel was stained for protein, and it was found that the main protein band for native enzyme and for carboxymethyl enzyme coincided with the respective activity band: The induced electrophoretic separation of native from carboxymethyl enzyme had not separated the catalytic activity in the carboxymethyl enzyme from the bulk of the protein in carboxymethyl enzyme. This is good confirmatory evidence that the ob-

148

REYNOLDS

AND MCKINLEY-MCKEE

03,” 2

4

3

FIG. 1. Starch-gel electrophoresis of native and carboxymethyl liver alcohol dehydrogenases, in the presence of NADH (0.5 PM) and isobutyramide (10 mM). In Tris-chloride buffer, 10 ItIM, pH 8.4, for 15 h at room temperature, and stained for activity with n-butanol as substrate. 1, native enzyme alone (150 kg); 2, native and carboxymethyl enzymes (150 wg each); 3, native enzyme (75 rg) and carboxymethyl enzyme (150 pg); 4, carboxymethyl enzyme alone (150 rd.

25 II-lodoplrazdel

50

75 IpMl

FIG. 2. Titrations of native and carboxymethyl liver alcohol dehydrogenases with 4-iodopyrazole. In 32 mM sodium phosphate buffer, pH 7.4, at 23.5%. NAD+, 136 FM. 0, Native enzyme (2.45 PM); A, Carboxymethyl enzyme (2.6 PM).

served catalytic activity is indeed a genuine property of the cysteine-46-carboxymethyl EE enzyme.

(0.1 PM), which is rather higher than that obtained by kinetics (26). This was the method used for estimating the concentration of binding-sites in samples of carboxymethyl enzyme. Fatty acids form strong ternary complexes with the native enzyme and NAD+ (27). Figure 3 shows that decanoate appears to displace pyrazole from the ternary complex of carboxymethyl enzyme, NAD+, and pyrazole. Native enzyme showed similar competition between pyrazole and decanoate. The dissociation constant for decanoate dissociating from its ternary complex with NAD+ and carboxymethyl enzyme was estimated approximately, the results varying between 1 and 9 PM. The value for native enzyme was 2 PM (13). Equilibrium enhancement

binding: NADH.

fluorescence-

When NADH binds to native liver alcohol dehydrogenase, its fluorescence emission is enhanced considerably, and the peak undergoes a blue-shift (28, 13). With carboxymethyl enzyme, the peak-height is enhanced much less (Fig. 4), but the considerable broadening of the shorter-wavelength part of the spectrum does give an enhancement of 7to 8-fold at 410 nm. This was used to estimate the dissociation constant of NADH from the carboxymethyl enzyme. Both native and carboxymethyl enzymes were found to contain NADH-oxidizing material, presumably aldehyde. Theorell and Winer (12) also found this and corrected for it by calculation. A simpler method was tested, namely the inclusion of low concentrations of ethanol (2-8 mM) in the cuvette. Titrations of native enzyme of

Equilibrium binding: pyrazole complexes. Titrations of carboxymethyl and na-

tive enzyme with 4-iodopyrazole (in the presence of excess NAD+) are shown in Fig. 2. The identical gradients at low concentrations of 4-iodopyrazole indicate that the complex formed with each enzyme has the same extinction coefficient at 290 nm. The curvatures as the end-points are reached are similar; the estimated dissociation constants of 4-iodopyrazole from the (assumed) complex of enzyme, NAD+ and pyrazole derivative were also similar

IFyrazck 1

IpMl

FIG. 3. Titrations of carboxymethyl liver alcohol dehydrogenase with pyrazole: effect of decanoate. In 32 mM sodium phosphate buffer, pH 7.4, at 23.5”C. NAD+, 136 PM. 0, Without decanoate, except as indicated (0); A, + Decanoate (33 PM).

CARBOXYMETHYL

L

LIVER

500

400 h hml

FIG. 4. Fluorescence-emission spectra (uncorrected) of NADH and carboxymethyl liver alcohol dehydrogenase. In 40 mM sodium phosphate buffer, pH 7.4, at 23.5”C. Excitation was at 330 nm. NADH, 0.8 PM. Carboxymethyl enzyme concentrations were: a. zero; b, 0.7 PM; C, 2.1 /AM.

gave good results, the dissociation constant for NADH being 0.296 FM at pH 7.4 (in 40 mM sodium phosphate buffer) compared with 0.31 PM (12, 13) and 0.29 pM (29) found in earlier work (without ethanol) at pH 7.0-7.4. Titrations of carboxymethyl enzyme gave the same values for NADHaffinity and fluorescence-enhancement when titrated in either the presence of ethanol, or in its absence and correcting by calculation (12) for NADH-oxidation. Thereafter, ethanol was routinely included for fluorescence-titrations. The results of titrations of carboxymethyl enzyme with NADH are shown in Figs. 5 and 6. The lines are straight within experimental error (which is largest at low concentrations of NADH), indicating that the two binding-sites of the enzyme probably function identically and independently, both with respect to their dissociation constants for NADH, and their enhancement of fluorescence of bound NADH. The mean dissociation constant for NADH was 1.7 pM, which is about 6-fold higher than for native enzyme (see above). Previously, Li and Vallee (3), using spectrophotometric titration, obtained a value of 40 PM for carboxymethyl enzyme. The reason for this discrepancy is not known. Competition with NADH is shown by AMP (Fig. 5) and decanoate (Fig. 6). Imidazole affects the binding of NADH (Fig. 7), but decreases rather than increases the fluorescence enhancement (Table I). Imidazole strengthens the bind-

ALCOHOL

DEHYDROGENASE

149

ing of NADH to carboxymethyl enzyme at pH 9.8, while at this pH it weakens its binding to the native enzyme (30). The slightly stronger binding of NADH at pH 6.1 compared to pH 7.4, and the weaker binding at higher pH values, is also shown by native liver alcohol dehydrogenase (12, 13). Acetamide (Fig. 7) makes very little difference to the binding of NADH, or to its fluorescence enhancement. Isobutyramide also was virtually without effect. This is in

FIG. 5. Binding of NADH to carboxymethyl liver alcohol dehydrogenase. from fluorescence enhancement, and effect of AMP. In 40 mM sodium phosphate buffer, pH 7.4, at 23.5”C. 0, No other ligand; A, + AMP (122 PM). R represents NADH, and ER the enzymeeNADH complex (one active site).

FIG. 6. Binding of NADH to carboxymethyl liver alcohol dehydrogenase, from fluorescence enhancement, and effect of decanoate. In 40 mM sodium phosphate buffer, pH 7.4, at 23.5”C. 0. No other ligand; A, + decanoate (50 PM); V, + decanoate (100 MM).

150

REYNOLDS

AND MCKINLEY-MCKEE

sharp contrast to native enzyme (27), where a considerable strengthening of NADH-binding is produced by amides, usually accompanied by an enhancement of fluorescence. Ethanol in high concentrations (31, 32)

0

02

04

and dimethyl sulphoxide (33) form ternary complexes with native alcohol dehydrogenase and NADH, giving tighter coenzyme binding, and a decrease and increase, respectively, in fluorescence enhancement. with carboxymethyl enzyme, However, ethanol (3%) and dimethyl sulphoxide (0.3%) had only small effects on NADHbinding and fluorescence-enhancement. Table I summarizes the results obtained from the fluorescence-titration experiments. Kinetics with different alcohols as substrates. When we reported that carboxymethyl liver alcohol dehydrogenase possesses residual catalytic activity (25), ethanol was used as substrate. The activity was 2.2-2.3% of that found for native enzyme. However, as shown in Table II, the maximum velocities for larger primary alcohols, and for cyclohexanol, are considerably greater than for ethanol; the maximum velocity with n-butanol is 6.7-fold higher than with ethanol, reaching 16.5% of the expected value for native enzyme. This implies that the rate-limiting step in alcohol oxidation is no longer the dissociation of the enzyme/NADH complex, as it is in the native enzyme which approximates closely to the Theorell-Chance mechanism

06

FIG. 7. Effects of imidazole, acetamide and altered pH on binding of NADH to carboxymethyl liver alcohol dehydrogenase at 23.5”C. Broken line without points, NADH alone at pH 7.4 (see Fig. 6.). 0, + Acetamide (0.37 Ml at pH 7.4 (40 mM sodium phosphate buffer); 0, + imidazole (base form 29 mM) at pH 7.4 (40 mM sodium phosphate + imidazole buffer); v, at pH 6.1 (80 mM sodium phosphate buffer) A, + imidazole (97 mM1 at pH 9.8 (glycine/ NaOH buffer + sodium phosphate, ionic strength 0.10, 19 mM in glycinate anion). TABLE APPARENT

I

DISSOCIATION CONSTANTS OF NADH FROM CARBOXYMETHYL LIVER ALCOHOL DEHYDROGENASE, DETERMINED BY FLUORESCENCE ENHANCEMENT. (EVALUATED FROM FIGS. 557).

Added ligand

PH

Q

DD (PM)

K,,LC

CM-ADHd (PM) None

7.4

7.2-7.6

1.70

AMP (122 FM) Decanoate (50 PM) Decanoate (100 FM)

7.4 7.4 7.4

8.2 7.7 8.0

3.50 3.16 4.59

Acetamide (0.4 M) Imidazole (base

1.4 7.4

8.8 5.5

2.3 2.9

None

6.1

6.8

None Imidazole

9.8 9.8

3.5

form

-

Native ADHd h) -

115 58.5 57.1 1

32-140 (7) 39-230 (7, 13)

-

29 tTIM)

(97 mM)

1.38 (no binding of NADH detected) 4.42

n Q = factor of fluorescence-enhancement, when NADH binds to carboxymethyl DD = apparent dissociation constant of NADH from carboxymethyl enzyme. c K,, L = calculated dissociation constant of the binary enzyme-ligand complex, NADH. d CM- = carboxymethyl-; ADH = alcohol dehydrogenase.

-

enzyme. assuming competition

with

CARBOXYMETHYL

LIVER

(34). The deuterium-isotope effect of 3.3 is much larger than that found with native enzyme under the same conditions (1.20). These results indicate that the rate-limiting step involves a complex which contains substrate, and could well be the hydride transfer step itself. The increased rates with larger substrates may be due to altered electronic effects at the site of hydride transfer, or to altered orientation of bound substrate; or they may be due to an altered enzyme conformation, i.e., “induced fit” as proposed by Koshland (35). The rate of hybrid-transfer catalyzed by native enzyme has been shown by presteady-state kinetics to be considerably greater for n-propanol than for ethanol (36), which was suggested as being largely due to steric effects. Aliphatic secondary alcohols such as isopropanol gave maximum velocities considerably lower than primary alcohols, with native enzyme (37); the interconversion of ternary complexes appears to be rate limiting. Therefore when secondary alcohols are substrates, the conformation or substrate orientation is presumably rather unfavorable for hydride transfer (36). The low catalytic rates found with isopropanol as substrate for the carboxymethyl enzyme (Table II) imply that the inhibition of hydride-transfer caused by using a secondary alcohol, and by carboxymethylation, are largely independent. The apparent Michaelis constants for isopropanol were similar for native and carboxymethyl enzymes at pH (13.7 and 12.1 mM, respectively, 10.0; 12.6 (Ref. 37) and 6.0 mM at pH 7.0-7.4). This implies that neither carboxymethylation nor change of pH has much effect on the binding of isopropanol to the enzyme/NAD+ complex. Binding of primary alcohols cannot be directly compared because native and carboxymethyl enzymes have different rate-limiting steps. However, pH has a large effect on the affinity of carboxymethyl-enzyme for primary alcohols (see below). it being greater at higher pH values. Primary alcohols give high-substrate inhibition with native liver alcohol dehydrogenase (32, 37-39), while cyclohexanol gives high-substrate activation (39). These

ALCOHOL

151

DEHYDROGENASE TABLE

II

KINETICS OF CARBOXYMETHYL LIVER ALCOHOL DEHYDROGENASE WITH DIFFERENT ALCOHOLS AS SUBSTRATES”

Substrate

Ethanol [*HIEthanol (ethanol-d61 n-Butanol Cyclohexanol n-Octanol Isopropanol

Apparent

K, (mM)

Apparent

(s- I)

V

c/cof Native”

11.6

0.145

2.5

11.6 0.34 2.13 0.041 12.1

0.044 0.97 0.87 0.70 0.011

(0.9) 16.5 14.5 11.9 (17.5)

izAt pH 10.0, with 460 PM NAD’, in glycine-NaOH rates were measured spectrobuffer (42 mM): photometrically for a series of substrate concentrations, and the constants evaluated from double-reciprocal plots. “Compared to ethanol as substrate for native enzyme (for which several different primary alcohols give similar maximum velocities) except that for values in parentheses the substrate shown is compared with each enzyme form.

effects were attributed to a decrease or an increase, respectively, in the rate of dissociation of NADH from a ternary abortive enzyme/NADH/alcohol complex. With carboxymethyl enzyme, no high-substrate inhibition by ethanol was found, even up to 0.5 M. Cyclohexanol and n-octanol, at the highest concentrations used (20 mM and 0.2 mM, respectively) gave a small amount of inhibition (20 and 10%’ approximately). n-Butanol gave quite considerable inhibition (Fig. 8). The rather erratic results at high concentrations may be due to the limit of n-butanol’s solubility in water being approached. However, it seems clear that inhibition is not total, but levels off at around 30% of the maximum activity, which is still over twice that observed for ethanol. It is possible that the inhibition is due to an abortive enzyme/NADH/butanol ternary complex, as with native enzyme; another possibility is that a quaternary enzyme/NAD+/(butanol), complex might form. The calculated K, in the early part of the inhibition is 29 mM. The considerable differences found for the effects of high substrate concentrations on the native and

152

REYNOLDS

AND MCKINLEY-MCKEE

strates, at two pH values (7.4 and 9.9). The results were plotted as double-reciprocal graphs (primary plots), and the slopes and intercepts were replotted against the reciprocal of the concentration of substrate which had been held constant for the primary plot (43). Examples of a primary plot and a secondary plot for each direction (alcohol oxidation or aldehyde reduction) at each pH are shown in Figs. 9-16. Pri-

i 200

400 in-Butonol!

imMi

FIG. 8. High-substrate inhibition of carboxymethyl liver alcohol dehydrogenase by n-hutanol. At 235°C pH 10.0, in 42 mM glycine/NaOH buffer, determined spectrophotometrically. NAD+, 465 pM.

modified enzymes may be because the enzyme/NADH complex is not kinetically significant for carboxymethyl enzyme, or because complexes of the type enzyme/ NADHlligand are not stable with carboxymethyl enzyme. Kinetics of aldehyde reduction. Preliminary studies indicated that the affinity for aldehydes was very much lower for the carboxymethyl enzyme than for native enzyme. To determine a value for the Michaelis constant with acetaldehyde, it was necessary to go up to concentrations of around 1 M, when “high-substrate inhibition” was observed, possibly due to a rapid reaction of acetaldehyde with protein amino groups, or an organic-solvent effect (but see below). As with native enzyme, n-butyraldehyde gave much lower apparent Michaelis constants than acetaldehyde (40), although it too did give some highsubstrate inhibition. This inhibition may well have been due to formation of a ternary complex between enzyme, NAD+ and hydrated aldehyde, the active complex for the aldehyde mutase reaction (41, 42). Maximum velocities with acetaldehyde or n-butyraldehyde were similar at pH 6.1, 7.4, and 9.9, and were always at least 30% of that determined for native enzyme under the same conditions. Steady-state kinetic analyses. Because of the very low affinity of carboxymethyl enzyme for acetaldehyde, steady-state kinetic analyses were performed with nbutanol and n-butyraldehyde as sub-

A ‘5. /

FIG. 9. Kinetics of reduction of n-butyraldebyde by NADH and carboxymethyl liver alcohol dehydrogenase. At pH 7.4, 23.5”C, in 20 mM sodium phosphate buffer, measured spectrophotometrically except at the lowest concentration of NADH, which was measured fluorimetrically. NADH concentrations were as follows: 0, 1.93 pM; A, 9.67 pM; x, 19.3 pM; 0, 29.0 /AM; A, 58.0 PM; v, 193 /AM.

FIG. 10. Secondary plots, of intercepts, from Fig. 9 and from a graph of the same data but with NADH as varied substrate.

CARBOXYMETHYL

LIVER

/ /

l

/iii /

Y’

ALCOHOL

ships between coefficients which in assessing possible mechanisms shown in Table IV. All primary plots appear to within the concentration ranges none of the kinetic coefficients Possible mechanisms are either

IO! I FIG. 11. Kinetics of oxidation of NAD+ and carboxymethyl liver alcohol ase. At pH 7.4 and 23.5”C, in 20 mM phate buffer, measured fluorimetrically. tions of n-butanol were: 0, 0.274 mM; v, 1.37 mM; 0, 2.74 mM; A, 16.4 mM.

n-butanol by dehydrogensodium phosConcentra0.548 mM; x,

153

DEHYDROGENASE

are useful (43) are be linear used, and is zero. a rapid-

/*”

Y

i;i

2

x’

/

FIG. 13. Kinetics of reduction of n-butyraldehyde by NADH and carboxymethyl liver alcohol dehydrogenase. At pH 9.9 and 23.5”C, in glycine/NaOH + sodium phosphate buffer, 10 mM in glycinate anion, ionic strength 0.053, with the rate measured spectrophotometrically. Concentrations of NADH were: v, 9.33 MM; X, 18.66 /.tM; 0, 56.0 KM; A, 373 fiM.

FIG. 12. Secondary plots, of slopes, from Fig. 11 and from a graph of the same data, but with nbutanol as the varied substrate.

mary plots with each substrate varied in turn enabled (from their respective secondary plots) two estimates for each of the four kinetic coefficients to be made: the mean values are shown in Table III. The maximum velocity of aldehyde reduction at pH 9.9 appears to be increased 1.9-fold by carboxymethylation. Some of the relation-

3 i 2 3 I’

/

FIG. 14. Secondary plots of intercepts, from Fig. 13 and from a graph of the same data but with NADH as the varied substrate.

154

REYNOLDS “xy

AND MCKINLEY-MCKEE .

I I

CG!

// /r

/’/ /I ,I,’

.. ..I_

dom-order, rapid-equilibrium mechanism has no other characteristic relationships, and hence cannot be excluded from steadystate analysis alone. Ordered mechanisms (if operating) can be characterized in greater detail. and are considered below. The native enzyme has been shown to approximate closely to the TheorellChance mechanism (34) with primary aliphatic substrates (37, 39, 40, 44, 45), where the rate of reaction in each direction is limited by dissociation of the product coenzyme/enzyme binary complex. There are two characteristic relationships of kinetic constants which may be used to test this (43): 4,.$2/@,2 = &’ (and its converse for the reverse reaction).

15. Kinetics of oxidation of n-butanol by NAD+ and carboxymethyl liver alcohol dehydrogenase, at pH 9.9. Conditions as in Fig. 13, except that measurements were fluorimetric. Concentrations of NAD’ were: A, 1.72 pM; 0, 3.45 PM; x, 6.90 PM; V. 17.2 F~; 0. 51.7 pi; A, 207 Irk. FIG.

FIG. 16. Secondary plots, of slopes, from Fig. 15 and from a graph of the same data but with NAD* as the varied substrate.

equilibrium, random-order mechanism with interconversion of ternary complexes being rate limiting [Dalziel (43) Type I] or an ordered addition of substrates [Dalziel (43) Type II]. Both are characterized by the Haldane relationship 4 ,2’/~12 = K,, , which is obeyed moderately at pH 7.4 and excellently at pH 9.9. However, the ran-

~o’~ch’~~~‘/~-,.~~.~~

= K,.

(i) (ii)

Of these, (i) is perhaps more useful because $,.~J@,2 represents (for an ordered mechanism) the reciprocal of the rate constant for the dissociation of the enzyme/coenzyme complex (43). and therefore 4,. &,/$ ,2 must be I bO’. Applied to the results summarized in Table IV, at pH 7.4, 41.@2/$12 <& (factor of 171, which is compatible with an ordered mechanism for alcohol oxidation, with a kinetically significant ternary complex. f~5~‘.&‘/$,~’ > do (factorof 141, which is apparently not compatible with a simple ordered mechanism for aldehyde reduction (possible reasons for this are considered in the Discussion). The similarity of (bO’‘4,’ &‘/&, .$,. & with Keq appears to be coincidental. At pH 9.9, G,.@$@,~ < &’ (factor of 3): this is compatible with an ordered mechanism for alcohol oxidation with a kinetically significant ternary complex. d,,’ 42’/4,2’ I $0 (factor of 1.141, which is compatible with an ordered mechanism for aldehyde reduction where the rate is limited primarily by the dissociation of the enzyme/NAD+ complex, and ternary complexes are kinetically insignificant. The nonequivalence of Keg and 40’. Q,I’ $,‘I &. $1.@2 (by a factor of 2.7) is to be expected, because it should only hold when the reaction in both directions is limited by

CARBOXYMETHYL

LIVER

ALCOHOL

TABLE

155

DEHYDROGENASE

III

KINETIC CONSTANTS FOH CARBOXVMETHVL LIVER ALCOHOL DEHYDRO~ENASE, CALCULATED FROM STEADY-STATE ANAI.YSES (FIGS. 99161”

Aldehyde reduction Carboxymethyl Native (401’ Carboxymethyl Native’. / Alcohol oxidation Carboxymethyl Native (40)’ Carboxymethyl Native’- 1

7.4 7.1 9.9 9.9

0.058 0.0075 0.261 0.59

7.4

2.17

7.1 9.9 9.9

0.35 0.94

0.19

0.907 0.100

17.2 0.17

2.61

43.2

1.0

-

44.1

5.100

1.1

7.2 234 -

49.6 2.3

127 0.04 3,690 270.500 :360 50,800 -

15.7 13.3 100

297

1.7

-

20.3 3.14 52.7

2.360 20.6 249

12

7.35 0.235 85..5 5.2

22.7 165

-

17.3 133 3.83

1.7

52.9

0.46 2.86 1.06 5.2

50 217 3.9

n &values are defined by the equation: e/c = 4” + $,/tcoenzymel + &/(substrate) + ~,,/(coenzymel(substratel. The lower half of the table (alcohol oxidation) contains @‘-values. (’ Michaelis constant for coenzyme tat infinite substrate concentration). ’ Michaelis constant for substrate (at infinite coenzyme concentration). i( Dissociation constant of the binary enzyme-coenzyme complex. ” From (44). with acetaldehyde and ethanol as substrates assuming a Theorell-Chance mechanism. ’ &values in these rows should be multiplied by 0.83 (and hence V divided by 0.83) for direct comparison with carboxymethyl enzyme, because different methods were used to calculate the enzyme concentration. TABLE

IV

COMBINATIONS OF KINETIC CONSTANTS FOH CARBOXYMETHVL ALCOHOL DEHYDROGENASE,AND THE EQUILIBRIUM CONSTANT FOR THE OVERALL REACTION, FOR TESTING THE KINETIC MECHANISMS PH 7.4 9.9

$0 (s) 0.058 0.261

40’ (s) 2.17 0.94

!e(s) $12 0.123 0.306

@I’@*’ -is) @I2

0.83 0.228

$12’

$J”‘dJl’b’

dJ12

&41@2

2130 13.8

K, 2860 37.1

,435o 14.0

” The equilibrium constant for: NAD ( + n-Butanol = NADH + n-Butyraldehyde + H’ is taken as 0.9 x’ 10 ‘I (401. The enzyme reaction is treated in the reverse direction, and so K,,, the pH-dependent equilibrium constant, is (H-/0.9 x 10 II).

the rate of dissociation of the product enzymelcoenzyme binary complex. Kinetics with inhibitors. The kinetic effects of certain potential inhibitors (isobutyramide, imidazole, pyrazole, and decanoate) were investigated for two reasons: firstly, to extend and complement the equilibrium ligand-binding studies described above; and secondly, to gain further information on the kinetic mechanism of the carboxymethyl enzyme. Isobutyramide and imidazole inhibit alcohol oxidation, as shown in Fig. 17. Both are competitive with ethanol, and give apparent inhibition constants of 3.5 mM for

imidazole and 175 mM for isobutyramide. These may approximate to the dissociation constants of these ligands from ternary complexes with enzyme and NAD+. Native enzyme and isobutyramide form strong ternary complexes with NADH (13) and weaker ones with NAD+ (46). These result in predominantly uncompetitive inhibition with alcohol, because of combination with the kinetically significant enzyme/NADH binary complex (38). Imidazole forms ternary complexes between native enzyme and NAD+ or NADH (13); NADH dissociates more rapidly from its ternary complex than it does from the

156

REYNOLDS

AND MCKINLEY-MCKEE

imidazole from its ternary complex with native enzyme and NAD+ at this pH (13). Imidazole forms a binary complex with carboxymethyl enzyme, shown by protection of its remaining sulphydryl groups from chemical modification (46a). If alcohol oxidation is via a random-order or a preferred-order mechanism, then the competitive inhibition by imidazole could be due to its binary complex, as well as its ternary complex with NAD+. Pyrazole and decanoate are strong inhibitors of butanol oxidation by carboxymethyl alcohol dehydrogenase, competing with substrate (Fig. 18). Calculated inhibiI 02 tion constants are 0.72 PM and 3.7 PM, '/IEthonoll “’ ” O6 respectively, which are similar to values FIG. 17. Kinetics of carboxymethyl liver alcohol given by native enzyme (18, 38). The dehydrogenase at pH 10. In 42 mM glycine/NaOH equilibrium binding experiments described buffer, at 23.5”C, determined spectrophotometrically. above demonstrated that carboxymethyl NAD+, 465 PM. 0, No further additions; A, + enzyme and NAD+ were still able to form Imidazole (4 mM);v, + Isobutyramide (26.7 mM). strong ternary complexes with pyrazole or decanoate. binary complex with enzyme, resulting in The effects of pyrazole and decanoate on activation at high concentrations of alcohol the reduction of n-butyraldehyde are (38). These effects were confirmed at pH 10 shown in Fig. 19. In each case the inhibiby experiments using the conditions of Fig. tion is almost completely uncompetitive. 17, with 10 mM isobutyramide or 10 mru imidazole; ethanol (alone) gave an apparent Michaelis constant of 7.25 mM, with pronounced high-substrate inhibition above 10 mM ethanol (32, 38). Isobutyramide gave almost pure uncompetitive inhibition, with an inhibition constant (evaluated from the intercept on the double-reciprocal plot) of 170 PM. Imidazole approximately doubled both the maximum velocity and the apparent Michaelis constant. It appears that kinetically carboxymethyl enzyme only reveals ternary complexes with these ligands and NAD+, not NADH. The equilibrium binding experiments described above indicated no ternary complex with enzyme, NADH, and isobutyramide, but there probably was one with imidazole. The lack of kinetic evidence for a ternary complex with NADH and imidazole is a further indication of the probable kinetic insignificance of the enzyme/NADH biFIG. 18. Kinetics of carboxymethyl liver alcohol nary complex in alcohol oxidation. The dehydrogenase at pH 7.4. In 20 mM sodium phosphate observed inhibition constant for imidazole buffer, at 23.5”C, determined fluorimetrically. NAD+, (3.5 IIIM) is rather lower than would be 52.4 pM. 0, No inhibitor; A, + Pyrazole (0.6 PM); V, expected for the dissociation constant of + Decanoate (16.7 PM).

CARBOXYMETHYL

LIVER

151

IO.

20

FIG. 19. Kinetics of carboxymethyl liver alcohol dehydrogenase at pH 7.4. In 20 mM sodium phosphate buffer, at 23.5”. determined spectrophotometrically. NADH, 67.5 FM. 0, No inhibitor; 0, + Pyrazole (1.0 PM); A, + Decanoate (16.7 pMj.

Inhibition constants evaluated from the intercepts are 0.47 PM for pyrazole and 4.3 PM for decanoate, which agree well with the values given in the preceding paragraph for alcohol oxidation. With carboxymethyl as well as native alcohol dehydrogenase (18), these compounds appear to combine specifically at low concentrations with the enzymelNAD+ binary complex. The quantitative agreement with values from competition with alcohol suggests that, at high aldehyde and NADH concentrations at pH 7.4, the enzyme is predominantly in the form of the enzyme/NAD+ binary complex, and therefore the dissociation of this complex is rate limiting. The results of kinetic steady-state analysis at pH 7.4 did not point unequivocally to this, although at pH 9.9 this mechanism was indicated. DISCUSSION

Authenticity of the catalytic activity in the carboxymethyl enzyme. When we first reported residual catalytic activity in cysteine-46-carboxymethylated alcohol dehydrogenase (25), it was pointed out that two

ALCOHOL

DEHYDROGENASE

157

other possible sources of catalytic activity might be (a) a minor isoenzyme component which is resistant to alkylation, or (b) the production of a minor enzyme species which is alkylated at a position other than cysteine-46, which possesses catalytic activity and an increased resistance to alkylation at cysteine-46. Some evidence was produced against each of these possibilities, including the demonstration that the carboxymethyl enzyme’s activity and protein showed the same electrophoretic mobility as the main (EE) component of native enzyme. However, the results presented in this work provide confirmation that the catalytic activity is very likely to be a genuine property of the cysteine-46modified enzyme. The evidence may be summarized thus: (i) In the previous report, where ethanol was used as substrate, activity was low, approximately 2.25% of the native enzyme’s activity. The activity for higher alcohols given here is lo-17% of that for native enzyme, and for aldehydes 20-190%. A minor component, from any source, cannot acccunt for these results unless its catalytic activity is considerably activated by carboxymethylation. (ii) The demonstration that carboxymethylation increases the catalytic activity under certain conditions completely rules out the possibility that activity could be due to an unmodified component which was present in native enzyme. (iii) The equilibrium ligand-binding studies must be due to the bulk of the protein present. The effects of the same ligands on catalytic activity were very much as predicted by ligand-binding studies (strong complexes formed by pyrazole compounds and decanoate, but only slight effects of isobutyramide and imidazole). (iv) The electrophoresis experiments described here show that the mobility of both the activity and the protein of the carboxymethyl enzyme migrate together, and (relative to native enzyme) were altered by added ligands as predicted by the known ligand-binding properties of the enzyme. The coincidences of activity properties on the one hand, and “bulk” properties (ligand-binding, and electrophoresis) on

158

REYNOLDS

AND MCKINLEY-MCKEE

the other, render it highly unlikely that the residual activity is due to a component different from isoenzyme EE carboxymethylated at cysteine-46. Effect of carboxymethylation upon ligand-binding properties. The most clearcut effect of carboxymethylation is upon ternary complexes involving NADH. Neither imidazole or amides (acetamide and isobutyramide) cause a significant increase in the fluorescence of enzyme-bound NADH, nor (for amides) a strengthening of the binding of NADH, thus contrasting with native enzyme (13, 27). Yet ternary complexes with NAD+ and pyrazole or decanoate still form readily, as with native enzyme (13, 18). Crystallographic studies have indicated that liver alcohol dehydrogenase exists in different conformations, one typical of free enzyme, and the other of enzymelcoenzyme binary and ternary complexes (47, 48); adenosine diphosphate ribose could bind to the former of these (47), and hence the binding of the nicotinamide portion was implicated in initiating the conformation change. The results reported here imply that there is scme basic difference, however, between ternary complexes involving NADH and NAD+. Those involving NADH appear to be destabilized, perhaps by preventing a necessary conformation change from taking

E&

k k-1

E-NADH

2

k

E-NADH-Ald

+

k-2

place before or during the binding of aldehyde or aldehyde-analog. It is possible that the binding of the dihydronicotinamide of NADH might be disturbed, or become such that it only occupies the “native” or correct orientation for substrate-binding or catalytic reaction for part of the time: this would be consistent with the six- (or more)fold weaker binding of NADH, the smaller fluorescence-enhancement, and reluctance

to bind ligands. The lack of a corresponding effect of NAD’ may be rationalized as follows. NAD+ has a dissociation constant approximately 500 times greater than that of NADH from their respective binary complexes (49) at pH 7 with native enzyme, and so it seems that the nicotinamide of NAD+ may be actually a hindrance to binding (38), and may not itself be bound by the enzyme at all: NAD+ binds considerably less tightly than adenosine diphosphate ribose, at neutral pH. (50). Kinetic properties and mechanism of carboxymethyl alcohol dehydrogenase. For oxidation of alcohols, the interconversion of ternary complexes appears to be rate limiting. The widely differing maximum rates with different alcohols, steady-state analyses at two pH-values (7.4 and 9.9) and the isotope-effect of 3.3 all point to this. The isotope effect suggests that the hydride-transfer step itself is at least partially rate limiting (51). The addition of substrates may be sequential, as for native enzyme (44) although the mechanism is probably preferred order rather than compulsory order (39, 42). The greatly decreased rate of interconversion of ternary complexes for carboxymethyl enzyme would allow much more time for a randomorder binding process to take place. In Scheme 1, k’ would be smaller than both k!, and k-*, if this were the case. E-NAD+-Ale

h’\\’

-+

E-NAD+ +2

,j\ ‘Gil\,! - , k +1\\‘\l, G, ‘>\” E-Ale

a -

E3 k’ + I,,,’,‘/’ ,‘/’.i’ k - Q~,‘ ,,‘;’ k +3 ,;, /;,

The reduction of aldehydes, on the other hand, may still be sequential (i.e., kr, >> k-J, with the dissociation of the enzyme/ NAD+ complex (i.e., k!,) being rate limiting. This was indicated by the comparable maximum velocities in preliminary experiments with acetaldehyde and n-butyraldehyde as substrates, by the uncompetitive inhibition by pyrazole and decanoate, and ’ Ald represents

aldehyde;

Ale represents

alcohol.

CARBOXYMETHYL

LIVER

by steady-state analyses at pH 9.9. The generally high maximum velocities are also compatible with this formulation. The results of the steady-state analysis at pH 7.4, however, are in serious disagreement with this, because the apparent velocity of the last step (i.e., k!,, given by @,Y4,‘.$,‘) was slower (by a factor of 14) than the overall rate (l/4,,). This is unlikely to be due to error in measuring @‘-values, since the results for butanol-oxidation were reasonably precise. An impurity in the (commercial) NAD’ preparation used could have contributed to this effect (44), but it is unlikely that it would have been its sole cause. Another possibility is that there might be a rate-limiting isomerization of the enzyme/NAD+ complex (52). This latter possibility was also proposed recently by Plapp et al. (53) for liver alcohol dehydrogenase which had been activated by chemical modification with picolinimidate. If an isomerisation of the enzyme/ NAD+ binary complex is occurring with carboxymethyl enzyme, then both isomers must combine with pyrazole or decanoate, with similar affinities. The estimates for the dissociation constant of the binary enzyme/NADH complex by kinetic and equilibrium methods are not in good agreement (7.35 PM and 1.7 pM, respectively). This may be partly due to imprecise results for aldehyde-reduction at pH 7.4, to some impurities in the NADH preparation, or to a rather slow isomerization of the enzyme/NADH complex (see above) similar to that observed for lactate dehydrogenase (54). However the latter two suggestions are not consistent with the inhibition experiments showing that the rate-limiting complex binds pyrazole or decanoate, unless a coenzyme impurity is postulated with the NAD+-like property of forming ternary complexes with enzyme and pyrazole or decanoate. The catalytic constant for alcohol oxidation, k’, is decreased by carboxymethylation from a value probably greater than 450 s-’ for n-butanol [see (36)] to 0.46 s1 (Table III) at pH 7.4, and becomes rate limiting. The Michaelis constant for nbutanol is considerably increased, but this

ALCOHOL

159

DEHYDROGENASE

is probably due to the change in kinetic mechanism; for native enzyme, much higher alcohol concentrations were required for maximal rates in pre-steadystate kinetics than under steady-state conditions (55). An increased K, for alcohol was observed for picolinimidylated alcohol which also has ternarydehydrogenase, complex interconversion as the rate-limiting step (53). The quite considerable increase in Michaelis constant for n-butyraldehyde (and, in preliminary experiments, for acetaldehyde) does require explanation, because of the apparent similarity in kinetic mechanism. For an ordered mechanism (43), k_,.k’ m Therefore the increase in $Z could be due to a decrease in k+2, an increase in k_ 2 or a decrease in k. The decrease found in k’ would actually contribute to a decrease in & not an increase, which must be overcome by changes in the other constants. It is perhaps unlikely that a considerable decrease in k is responsible, because this step is still apparently non-rate-limiting in the carboxymethyl enzyme. A decrease in k,, or an increase in k-, (or both) would indicate a decreased stability of the enzyme/NADH/aldehyde ternary complex. This is also indicated by the decrease in k’; in this case the complex is being formed via catalysis. Further information could be obtained by rapid-reaction techniques, which have already been applied to native (14, 34, 36. 55) and lysine-modified (53) alcohol dehydrogenases. Product inhibition might also prove useful for further characterising the mechanism, and this has been successfully applied to both native (56) and lysinemodified alcohol dehydrogenase (57). Carboxymethylation and enzyme structure. Branden and co-workers (8, 9) have determined the structure of native horseliver alcohol dehydrogenase (EE isoenzyme) at 2.4 A resolution; in conjunction with knowledge of the amino-acid sequence (5), each residue has been located (9). Cysteine-46 is one of the ligands to the

160

REYNOLDS AND MCKINLEY-MCKEE

active-center zinc atom, which is the binding site of orthophenanthroline (58, 59, 8), of decanoate and imidazole (38), of pyrazole (18), and possibly of substrates (38, 9); however other anions can bind elsewhere. The neighboring residue, arginine-47, is probably involved in binding the phosphate groups of the coenzyme by electrostatic attraction (9); and serine-48 forms a hydrogen bond with a zinc-bound water molecule and possibly also with histidine51 (9). Carboxymethylation of the sulfur of cysteine-46 would be expected to prevent it binding to the zinc, although it is possible that the introduced carboxyl group might bind instead. In either case the immediate environment of the zinc would be changed, and also the conformation of the side-chain of residue 46 would be affected which might in turn affect the main chain and neighboring side-chains. Optical rotatory dispersion indicates that no large structural change takes place on carboxymethylation (3). Coenzymes and AMP are bound well by the carboxymethyl enzyme, suggesting that no large movement of arginine-47 has taken place. Probably most of the effects of carboxymethylation can be attributed to an alteration of the properties of the active-center zinc atom. The binding of coenzymes and other ligands to native liver alcohol dehydrogenase is pH-dependent; many of the effects were attributed to the ionization of a zincbound water molecule (13, 49) which were abolished when imidazole was added. The pH-dependence of the carboxymethyl enzyme is in some respects different from that of native enzyme: the binding of NAD+ to native enzyme is markedly increased as the pH is raised (13, 49), but the kinetic results reported above indicate that the binding of NAD+ to carboxymethyl enzyme is no stronger at pH 9.9 than at 7.4. The loss of a proton from the zinc-bound water-molecule and consequent change of charge was suggested as being responsible (13, 49), and may be very different in the carboxymethyl enzyme. On the other hand, the pH-dependence of binding of NADH to carboxymethyl enzyme is similar to that of NADH or ADP-ribose to native enzyme (7, 12, 13, 50); it becomes weaker at high pH

even in the presence of imidazole (7, 30) which is probably due to an ionizing group other than the zinc-bound water molecule. The zinc-bound water molecule, when deprotonated, may play an important role in allowing the binding of alcohol as an alcoholate ion (9); this is consistent with the observation that at higher pH-values the carboxymethyl enzyme has a greater affinity for alcohol, and also that the rate of alcohol oxidation is greater. Equivalent observations for native enzyme would require pre-steady-state kinetics, and have not yet been reported. Structural considerations also enable an explanation to be offered as to why cysteine-46 is uniquely reactive among the 14 thiol groups in the polypeptide chain, and among the six thiol groups which are ligands to one or other of the two zinc atoms in the subunit (9). Iodoacetate can form a reversible complex with the enzyme (6, 7). This may well involve binding to arginine-47, which has been suggested to be a general anion-binding site (60,9), although it would normally function in binding the pyrophosphate moiety of the coenzyme molecule: the reversibly bound iodoacetate molecule would then be well placed for alkylating cysteine-46 (Ref. 9). The rather puzzling earlier results (7) showing that orthophenanthroline did not prevent the reversible binding of iodoacetate but did prevent alkylation, and that imidazole stimulated the rate of alkylation, are considerably clarified by the discovery that cysteine-46 is a ligand to the zinc to which these reagents bind. Yeast alcohol dehydrogenase also possessesone thiol group per subunit which is reactive with iodoacetate; the surrounding amino-acid sequence shows homologies with the liver enzyme (61), and its reactivity is greater. Sloan and Mildvan (62) have alkylated the yeast enzyme with a paramagnetic iodoacetamide derivative, and using magnetic resonance they measured the distance from the spin label to a number of protons on enzyme-bound NADH. They deduced from this the conformation of the coenzyme molecule and the relative position of the thiol group to which the spin label was attached, thus providing elegant

CARBOXYMETHYL

LIVER

and independent information on the enzyme structure which agreed very well with the crystallographic results for the liver enzyme. Sloan and Mildvan (62) also detected a change in the conformation of the bound NADH when isobutyramide was added. It would be attractive to equate this with a change which carboxymethylation prevents in the liver enzyme, but this is probably not valid because derivatization itself, with the spin-labeled compound, involved alkylation. Dickinson (63) found that although iodoacetamide-inactivated yeast alcohol dehydrogenase could still bind NADH with the same affinity and fluorescence-enhancement as native enzyme, acetamide now did not increase the fluorescence or the strength of binding of NADH, although it might still be able to bind. He also found (63) that oxidized coenzyme and hydroxylamine would not form a ternary complex, unlike carboxymethyl liver enzyme which readily forms ternary complexes with NAD+. Thus alkylated yeast and liver alcohol dehydrogenases show some similarities and some differences, but the enzymes themselves are rather dissimilar, and different alkylating agents were used. The situation is further complicated by the recent demonstration that the yeast enzyme has two reactive thiol groups, probably adjacent, either of which may be modified (but not both) depending on the reagent and the pH (64, 65). ACKNOWLEDGMENTS The authors thank Prof. R. Htittel (Miinchen) for providing the 4-iodopyrazole; Mr. J. McGowan, Dr. J. Kay and Dr. A. P. Ryle for help with the amino-acid analyses; Dr. I. A. Nimmo for help with the Olivetti Programma PlOl computer; and Dr. D. L. Morris for helpful discussion. REFERENCES 1. MCKINLEY-MCKEE, J. S., MORRIS, D. L., AND REYNOLDS, C. H. (1972). in Structure and Function of Oxidation-Reduction Enzyme (Akeson, A., and Ehrenberg, A., eds.), pp. 613-618, Pergammon, New York. 2. LI, T.-K., AND VALLEE, B. L. (1963). Biochem. Biophys. Res. Commun. 12, 44-49. 3. LI, T.-K., AND VALLEE, B. L. (1965). Biochemistry 4, 1195-1202.

ALCOHOL

DEHYDROGENASE

161

4. EVANS, N., AND RABIN, B. R. (1968). Eur. J. Biothem. 4,548-554. 5. J~RNVALL, H. (1970). Eur. i Biochem. 14, 521-534. 6. REYNOLDS, C. H.. AND MCKINLEFMCKEE, J. S. (1969). Eur. J. Biochem. 10, 474-478. 7 REYNOLDS, C. H., MORRIS, D. L., AND MCKINLEYMCKEE, J. S. (1970). Eur. J. Biochem. 14. 14-26. 8. BR#NDBN, C.-I., EKLUND, H., NORDSTROM. B., BOIWE, T., S~DERLUND, G., ZEPPEZAUER, E.. OHLSSON, I.. AND AKESON, A. (1973). Proc. Nat. Acad. Sci. USA 70, 2439-2442. 9. EKLUND, H., NORDSTROM, B., ZEPPEZACER, E., S~DERLUND, G., OHLSSON, I., BOIWE, T.. AND BRXND~N, C.-I. (1974). Fed. Eur. Eiochem. Sot. Lett. 44, 200-204. 10. REYNOLDS, C. H., AND MCKINLEY-MCKEE, J. S. (1973) 9th International Congress of Biochemistry, Stockholm, Abstracts, p. 52. 11. PIETRUSZKO. R., AND THEORELL, H. (1969). Arch. Biochem. Eiophys. 131, 288-298. 12. THEORELL, H.. AND WINER, A. D. (1959). Arch. Biochem. Biophys. 83,291-308. J. S. 13. THEORELL, H.. AND MCKINLEY-MCKEE. (1961). Acta Chem. Stand. 15, 1811-1833. 14. BERNHARD, S. A., DUNN, M. F., LUISI. P. L.. AND SCHACK, P. (1970). Biochemistry 9, 185192. 15. HOAGSTROM, C. W., IWEIBO, I., AXD WEINER, H. (1969). J. Biol. Chem. 244, 5967-5971. 16. MCKAY, R. H.. AND KAPLAN, N. 0. (1964). Biochim. Biophys. Acta 79, 273-283. 17. SCATCHARD, G. (1949). Ann. N.Y. Acad. Sci. 51, 660-672. 18. THEORELL, H., AND YONETANI, T. (1963). Biochem. 2. 338, 537-553. 19. MCKINLEY-MCKEE, J. S., AND Moss, D. W. (1965). Biochem. J. 96, 583-587. 20 CRUFT, H. J. (1962). Biochem. J. 84, 47P-48P. 21. SPACKMAN, D. H., MOORE, S., AND STEIN, W. H. (1958). Anal. Chem. 30, 118551189. 22. ELLMAN, G. (1959). Arch. Biochem. Eiophys. 82, 70-77. 23. WITTER, A. (1960). Acta Chcm. &and. 14, 1717-1728. 24. J~RNVALL, H., AND HARRIS, J. I. (1970). Eur. J. Biochem. 13, 565-576. 25. REYNOLDS, C. H., AND MCKINLEY-MCKEE, J. S. (1970). Biochem. J. 119, 801-802. 26. THEORELL, H., YONETANI, T., AND &I&ERG, B. (1969). Acta Chem. Stand. 23, 255-260. 27. WINER, A. D., AND THEORELL, H. 11960). Acta Chem. Stand. 14, 1729-1742. 28. BOYER, P. D., AND THEORELL, H. (1956). Acta Chem. Stand. 10, 447-450. 29. ANDERSON, S. R., AND WEBER, G. (1965). Biochemistry 4, 1948-1957. 30. REYNOLDS, C. H., AND MCKINLEY-MCKEE, J. S.

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(1972). Fed. Eur. Biochem. Sot. Lett. 21, 297-299. 31. SHORE, J. D.. AND THEORELL, H. (19661. Arch. Biochem. Biophvs. 116, 255-260. 32. SHORE, J. D., AND THEORELL, H. (1966). Arch. Biochem. Bioph.ys. 117, 375-380. 33. PERLMAN, R. L., AND WOLFF, J. (1968). Science 160, 317-319. 34. THEORELL, H., AND CHANCE, B. (1951). Acta Chem. &and. 5, 1127-1144. 35. KOSHLAND, D. E. (1958). Proc. Nat. Acad. Sci. USA 44, 98-104. 36. BROOKS, R. L., AND SHORE, J. D. (1971). Biochemistry 10, 3855-3858. 37. DALZIEL, K., AND DICKINSON, F. M. (1966). Biohem. J. 100, 34-46. H., AND MCKINLEY-MCKEE, J. S. 38. THEORELL, (1961). Acta Chem. Rand. 15, 1834-1865. 39. DALZIEL, K., AND DICKINSON, F. M. (1966). Biohem. J. 100, 491-500. 40. DALZIEL, K. (1962). Biochem. J. 84, 244-254. 41. ABELES, R. H.. AND LEE, H. A. (1960). J. Biol. Chem. 235, 1499-1503. 42. DALZIEL, K., AND DICKINSON, F. M. (1965). Nature (London) 206, 255-257. 43. DALZIEL, K. (1957). Acta Chem. Stand. 11, 1706-1723. 44. DALZIEL, K. (1963). J. Biol. Chem. 238,2850-2858. 45. THEORELL, H., AND MCKINLEY-MCKEE, J. S. (1961). Acta Chem. Stand. 15, 1797-1810. 46. SIGMAN, D. S., AND WINER, A. D. (1970). Biochim. Biophys. Acta 206, 183-186. 46a. REYNOLDS, C. H.. AND MCKINLEY-MCKEE, J. S. (1974) Fed. Eur. &o&em. Sot. Lett. 46, 8386. 47. ZEPPEZAUER, E., SLIDERBERG, B.-O., BRKND~N,

48. 49. 50. 51.

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