Chemical Modification of Catalytically Essential Amino Acid Residues in Enzymes Manuel G. Roig Departamento de Fisico-Quimica Aplicada y Tecnicas lnstrumentales, Facultad de Farmacia, Universidad de Salamanca, Salamanca, Spain
A large number of enzymes work by general acid-base catalysis mechanisms. Many of them possess, in their active sites, one or two amino acid residues involved in the catalytic mechanism and function. Among the residues most frequently associated with enzyme activity are Tyr, His, Cys, Ser, Lys, Arg, Asp, and Glu. Different approaches have been taken to identify the catalytically essential amino acid residues in enzymes. Kinetic measurements of t he pH dependence of enzyme activity !,ave been used to determine the pK values for amino acids ,;wolved in catalysis. Substrate specificity studies can also provide general information about the essent ial amino acid residues in the active site. X-ray structure analysis and chemical modification techniques have been employed to identify residues essential to catalysis. In cases where t he amount of enzyme is limited and the amino acid sequence is not available, the chemical protein modification approach seems to be the most appropriate method. When modification of a group in a protein affects the loss of biological activity, it is usually considered essential to that activity. However , interpretations of data such as these must be made with care. Chemical modification of residues that abolish enzymatic activity and that can be protected by a competitive inhibitor, substrate, or coenzyme could also result from at least two other phenomena. First, the modificat ion could occur at a site other than the active site in such a way that they would be of a different conformation and hence be inactivated. Binding of a competitive inhibitor, substrate, or coenzyme could alternatively induce a conformational change in the protein that would make t he group to be modified inaccessible to the reagent. Second, the group modified could be in the active site but might function neit her in substrate binding nor in catalysis. Chemical modification of the residue could induce a conformat ional change in t he active site that could absolish substrate binding, catalysis, or both. Partial loss of enzymatic activity by means of chemical modification can be explained in terms of either of two models, partial inactivation of all the enzyme molecules or complete inactivation of part of the molecules. The two situations a re usually difficult to distinguish. Partial inactivation of t he entire population is normally accompanied by changes in kinetic parameters, pH profiles, and the like, whereas a decrease in the turnover number without a corresponding change in Michaelis constant (KM) is frequently associated with the existen ce of a fraction of molecules whose activities have not been cha nged. Several amino acid residues have been shown to be involved in the catalytic process in numerous enzymes. Arginyl residues serve as complementary, positively charged, recognition sites for a large number of enzymes that act on negatively charged substrates or require anionic cofactors (1). As an example, a single arginyl residue participates in binding NADH to alcohol dehydrogenase (1) . Ser, Cys, His, and Lys residues have been reported to be involved catalytically in many enzymes by forming a covalent enzyme- substrate intermediate. For any modifying agent to be used successfully it should
meet at least two conditions: (1) it must have specificity for a given amino acid residue, and (2) it must react with the am ino acid residue under relatively mild conditions (aq ueous solvent, pH range 5 to 10). Of the several nucleophilic groups commonly found in proteins that a re known to react with haloacetates, sulfhydryl groups of Cys a re intrinsically the most reactive (see below). Reactivity increases with pH since t he anion is t he reactive species. Most often, a pH near neutrality under conditions compatible with physiological activity is best as a first try to determine t he effects of haloacetate modification of sulfhydryl groups. Under denaturing condit ions (urea, SDS, etc.), reactivities of a protein's sulfhydryl groups are similar to t hose of low-molecular-weight sulfhydryl compounds. In the absence of such conditions, considerable variations in reactivity a re observed, reflecting the varied environments of the reacting groups. Neighboring groups can eit her enhance or suppress the reactivity of a group. Greatly enhanced reactivity is often observed for groups in or near catalytic centers of an enzyme. Reactivit ies of Cys residues of proteins can sometimes be either enhanced or suppressed by substrates, allosteric effectors, and other specifically interacting substances. Thus, the coenzyme nicotinamide adenine dinucleotide protects t he two reactive Cys residues of h orse liver alcohol dehydrogenase (ADH) from iodoacetate and, in so doing, prevents a loss of catalytic activity (2). Mercurials also react rapidly and sp ecifically with the sulfhydryl groups of proteins (3) (see below). No other groups are usually affected under normal condit ions. The optim um rate of reaction of mercurials with proteins is usually near pH 5. While higher pH's increase the concentration of the more reactive anionic form Protein-S-, greater reactivity may not result, one reason being the increased compet it ion of OH- ligands for t he mercuria l. Reaction rates of sulfhydryl groups in different proteins and of diffe rent sulfhydryl groups within any one protein can vary greatly but a pproach those of low-molecular-weight thiols in urea or other denaturants. Because of such differences in reactivity, it is often possible to modify selectively a fraction of the total sulfhydryl groups in a protein. Mercurials combine very strongly with sulfhydryl groups. Dissociation constants of thiol- mercurial complexes under conditions such as those normally used for the modification of proteins a re usually 10-zo or less. Removal of mercurials from t heir complexes wit h protein sulfhydryl groups can be accomplished by treatment with competing ligands, particularly with low-molecular-weight t hiols. Where physiological activity has been altered upon reaction with t he mercurial, it can very often be recovered at least in part by such a t reatment. T he above-mentioned chemical agents have been used successfully by many researchers with the aim of probing by chemical modification the identity of the main functional groups involved in the catalytic mechanism of horse liver ADH. In such an enzyme it has been well established t hat Arg47, Cys-46, and Cys-174 are involved in t he active site (1, 4). Volume 64
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Lysil residues have also been implicated in the action of this ADH since activity increases on picolinimidylation and acetimidylation of Lys-228, modifications which in the presence of cofactors and inhibitors prevent both labeling and increase in activity (4, 5). We therefore consider that this enzyme, ADH from horse liver, may be taken as a good working model on which to carry out a series of experiments concerning the chemical modification of proteins wit h the aim of familiarizing the student of biological chemistry with the field of the chemical reactivity of amino acid residues of proteins and with research into the nature of t he active sites in enzymes. In order to obtain a better understanding of the general methodology of t he work in th is ki nd of study of the chemical modificat ion of enzymes, we shall foc us our attention on a single chemical modification, explicitly the modification of the Cys residues in horse liver ADH , considering that it will ill ustrate to a sufficient extent the methods and concepts involved in determ ining wh ich amino acids are implicated in e nzymatic reactions. It should be noted that in t he kind of experiments described he re, the methodology does not involve t he use of sophisticated equipme nt nor expensive reagents, though they do imply careful a ttention on the part of the student in cont rolli ng the condit ions of each chemical mod ification so that it will be selective and in order to be able to follow the protocols of each chemical mod ification and analysis. Experimental
Materials and Methods H orse liver alcohol dehydrogenase (ADH ), coenzymes (,8-NAD+, ,8- NAD H), Sephadex G-25 (fine), iodoacetic acid, iod oacetamide, 5,5'-dithiob is(2-nitrobenzoic acid), p-mercuribenzoate, EDTA, Pipes (Sigma Chemical Co.), and ethanol (Merck) were purchased from t he designated sources. All other chemicals were of reagent grade. Distilled , d eionized water was used throughout t his work. The solutions of the oxidized and reduced forms of ,8-diphosphopyridi ne nucleotide were prepared in the reaction buffer fresh daily and kept on ice. Exclusion of light while working up the iodoacetate solutions is important in ord er to prevent the formation of iodine, which may react with tyrosine and histidine residues. All the buffers and in particular the buffers used in preparation of t he enzyme were deoxygenated by passing a stream of N 2 t hrough t hem for a few minutes. Failure to ta ke this precaution could lead to an underestimation of t he number of Cys res idues in the enzyme due to possible oxidation of some Cys to form S- S bridges which cannot be evaluated by the methods described. The handling of chemicals such as iodoacetic acid , iod oacetamide, p -mercuribenzoate, 5,5'-dithiobis(2-nitrobenzoic acid) must be performed with ca ut ion, avoiding contact wit h the s kin and breath. Enzyme Assay Horse liver ADH (form EE) was obtained as a lyophilized powd er from Sigma. Stock solu tions were prepared in advance by dissolving the protein in 10 mM phosphate buffer (pH 7.5) at 4 °C and dialyzing for th ree days against several changes of a 1000-fold volume excess of the same buffer at 4 °C; cent rifugation at 5000 rpm fo r 30 min, 4 °C, removed insoluble material. The protein concentration was determined using a molar absorptivity at 280 nm of 3.57 X 104 M- 1 cm- 1 (J). The molecular weight of ADH is 83.000 (6). Concentrations of NA D+ and NADH were determined from the absorbance at 260 nm (~ = 18 mM- 1 cm- 1) and 340 nm (~ = 6.22 X 10~ M- 1 cm- 1), respectively (7). The 3-mL reaction mixture consisted of 50 mM pyrophosphate buffer (p H 8.8), I mM (low concentration assay) or 33 mM (high concentration assay) of ethanol, 30 11M (low concentration assay) or 2.5 mM (high concentration assay) of NAD+, and the corresponding concentration (about 0.2 11M) of the ADH or modified ADH (i n the presence of the corresponding modifying reagent concent ration). The presence in the optica l cuvette of a concentration of about lQ-5_ 10- 4 M of the mod ifying reagent (iodoacetic acid or acetamide) does not interfere with the enzymatic assay. A greater or lesser partial loss of enzymatic activity could be checked according to whether the kinetic assay is carried out at high or low concentration of cofactor 78
Journa l of Chemical Education
and substrate; this cou ld reflect changes in the K M val ues for substrate or coenzyme. T he rate of NADH fo rmation was followed at 340 nm using a Sh imadzu UV -120-02 spectrophotometer eq uipped with a t hermostat-circu lator set at 25.0 ± 0.1 °C. Initial velocities were expressed in 11mol of NADH formed per minute. The initial velocities for the high concentrations assay are maximum velocities of the reaction catalyzed by t he enzyme. The kinetic parameters for native horse liver ADH (form EE) are: Michaelis constants for NAD+ and ethanol, 36 11M and 800 11M, respectively, and maximum velocity for the ethanol oxidation reaction of 5 s- 1 (4).
Modification of Horse Liver ADH Modification experiments were performed at 25.0 oc by the addit ion of t he reagent (iod oacetic acid , acetamide, in 50 mM P ipes pH 7.1) to the enzyme solution in t he same buffer. Immediately after the addition, a sample was withd rawn from the reaction mixture and assayed for the zero-time ADH activity (V0 ) (see condit ions of the assay). The progress of t he chemica l modification reaction was monitored by assaying the enzymatic activity ( V) on small a liquots withdrawn at d esignated t ime intervals. In a parallel experiment, the enzyme was incubated ident ically with t he buffer only and the stability of t he enzyme in such conditions was controlled during the incubation time. In order to establis h t he correlation between the loss of ADH activity and the number of Cys resid ues being modified, besides enzymatic activity, the number of Cys was analyzed in only one of t hese aliquots (see below). The (modifying agent)/(enzyme) ratio in t he incubation mixture was around 1()3, chemical about 1-5 mM , enzyme about 1-5 1'M. Protection Studies Protection experiments were perfor med essentially as described above except that t he enzyme was preincubated for 10 min with the coenzymes NAD+, NADH , before adding the modify ing reagent. NAD+ and NADH at the working pH 's will protect t he anion bind ing sites against chemical modification, thereby revealing t he nature of the amino acid residues essential for such binding and for the catalytic process itself. The (cofactor)/(enzyme) ratio was about 200-1000, NAD+ or NA DH 1 mM, enzyme 1-51' M. Each set of enzyme chemical modification experiments can be carried out simultaneously; that is, incubation of enzyme wit h buffer (blank), enzyme p lus reagent, enzyme plus NAD+ plus reagent, and enzyme p lus NADH plus reagent may be in itiated consecutively with 5-10-min intervals between each other. During the interim periods t he student can assay the zero-time ADH activity (Vel of t he preceding incubation mixture. Accordingly, each set of experiments for each reagent (iodoacetic acid or iodoacetamide) (see t he figu re) may be carried out in one 3- hou r laboratory session. Reactions for the Modification of Functional Cys In Proteins (P)
The following is a schematic description of the d ifferent modificati on reactions of the th iol group of Cys t hat are ut ilized in this work. The description includes the kind of react ion involved and the modifying reagent. (1) Carboxymethylation
Reagent: Iodoacetic acid.
p ll
>~
(2) Carboxyamidation
Reagent: Iodoacetamide.
®-s- +
ICH2CONH2
~
®- --cH -CONH
(3) Formation of mercaptide bonds
Reagent: p-mercuribenzoate.
2
2
+
I- (2)
®-s- + Hg~coo- ~ ®-sHg----@-coo- (;lJ (4) Reduction
Reagent: 5,5'-dithiobis(2-nitrobenzoic acid) (Ellman's reagent).
@-sH + NO,--
-s-s-q-NO, coo-
pH
> ll.~
coo(4)
Number of Modified Amino Acid Residues
The correlation between the course of the change in ADH activity and the course of the number of amino acid residues being modified can be established by means of the following method : the aliquots withdrawn at different times from the incubation samples (in the first 3-hour laboratory session) should be dialyzed overnight or passed through a Sephadex G-25 (fine) column against or equilibrated in the incubation buffer (50 mM Pipes pH 7.1) to remove the excess of reagents and maintaining the irreversibility of the modification (check the specific ADH activity of the collected fractions). These specific activities must be similar to the specific activities of the incubation mixtures immediately after modification. The aliquots should be analyzed quantitatively for Cys content (second 3-hour laboratory session) and only the Cys residues unmodified by iodoacetic acid or iodoacetamide are titrated. If at the same time the number of Cys residues of the native enzyme is determined (third 3-hour laboratory session), it will be possible to calculate by differences the number of modified Cys, that is, those Cys protected against titration. For Cys titration of the enzyme, the concentrations of the enzyme itself and of the titration reagents should be known exactly. In view of the laboriousness of the methods to be described below for the determination of the number of Cys residues, the study of the correlation between the loss of ADH activity and the number of modified Cys is carried out with a single aliquot taken from the incubation mixture at a time sufficient for the modification to be important ( V !Vc X 100 = 20 or 30), that is, about 60- 90 min after the modification has begun. Accordingly, the titration of the number of Cys will only be carried out in this aliquot. Ellman's method for the quantitative determination of the number of Cys residues in proteins makes use of the reduction of such groups by the reagent 5,5' -dithiobis(2nitrobenzoic acid) (see reaction 4). The thionitrobenzoate anion is strongly colored and can be determined from its absorbance at 412 nm (~ = 1.36 X 104 M- 1 cm- 1 at pH 8.0). The enzyme was incubated (4-10 1-LM) with 1.5 mM EDTA, 1 mM Ellman's reagent in 10 mM phosphate buffer (pH 8.0), and we measured continuously the absorbance increase at 412 nm against time. In the blank cell were placed all the reagents, except the enzyme, and the readings were also recorded in order to determine the proportional absorbance of the control.
The (absorbance at 412 nm) / 1.36 X 104 (M) ratio is correlated to the thionitrobenzoate concentration, i.e., to the concentration of Cys residues titrated (see reaction 4). This concentration divided by enzyme concentration must then provide the number of Cys residues that are being titrated. The course of this titration can also be followed by means of enzymatic assays at different time intervals, withdrawing aliquots from the optical cuvette. The end point of this titration is reached when the A41 2 nm no longer increases (in about 7-9 h, depending whether the (Ellman's reagent)/ (enzyme) ratio is greater or smaller). The Cys spectrophotometric titration of horse liver ADH was also followed according to the method of Boyer (8, 9), on the basis of the ~ = 7.6 X 103 M- 1 cm- 1 at 255 nm that is produced as a consequence of mercaptide formation on incubation of the enzyme with p-mercuribenzoate (see reaction 3). Successive additions of enzyme to a fixed excess quantity of p-mercuribenzoate cause an increase in the A255 nm until a saturation value is reached when the reagent is no longer in excess with respect to the ADH. A change in the slope indicates the endpoint at the equivalence of p-mercuribenzoate and sulfhydryl groups. The p-mercuribenzoate is prepared about (60 1-LM) by dilution at alkaline pH and addition to 0.33 M acetate buffer (pH 4.6) (the molar absorptivity~ of pmercuribenzoate at 233 nm is 1.69 X 104 M- 1 cm- 1 at pH 4.6) . Insoluble material, if present, can be removed by filtration or centrifugation. One milliliter of this p-mercuribenzoate 60 1-LM is placed in the cuvette; successive 50 JLL aliquots of enzyme (5-9 1-LM) are added, and absorbance increases linearly until a change of slope, endpoint of titration. The (p-mercuribenzoate)/(enzyme) ratio at this point will indicate the number of Cys titrated by p-mercuribenzoate. In the case of our wanting to follow how much the chemical modification would affect the values of the kinetic parameters KM or Vor the pH profile of the native enzyme, we should stop the modification reaction after 2-3 hours by passing the incubation sample through a Sephadex G-25 (fine) column equilibrated in the buffer for removing the excess of reagents and collecting the fractions with modified ADH activity. With these fractions we can plan protocols for determining the kinetic parameters or pH profile. Results and Discussion
Modification of Cys residues
Modification of cysteine residues is known to abolish the activity of horse liver ADH (2, 10). In a model experiment we incubated 7 1-LM enzyme with 4 mM iodoacetate in 50 mM Pipes (pH 7.1) at 25 °C. This 600-fold excess of iodoacetate decreases the activity to 20% of the control in approximately 30- 40 min (see the figure). After this time an average of 1.0 SH group per subunit is carboxymethylated as measured by titration following Boyer's and Ellman 's methods of the native (14 Cys per subunit) and modified enzyme (13 Cys per subunit), after removing the reagents by gel filtration through Sephadex G-25 (fine) equilibrated in 50 mM Pipes (pH 7.1). In the same experiment we checked the protection that the NAD+ and NADH cofactors exert against the enzyme inactivation by iodoacetate (see the figure), indicating that this Cys residue is involved in the active site of the enzyme (anion binding site). This protection by cofactors must be checked by titration of the number of Cys residues in the enzyme before and after incubation with the modifying reagent. The same number of Cys before and after incubation implies no modification of Cys, that is, protection by cofactors against such a modification. Iodoacetamide also inhibits ADH. However, more than a 1500-fold excess of the reagent is required in order to obtain complete inactivation within 3 hours of incubation. This slower and nonspecific (non protected by cofactors) enzyme Volume 64
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January 1987
79
aspect. Thus, linear kinetics (Michaelian) could become nonlinear (non -Michaelian) through the effect of the chemical modification of som e of its essential amino acid residues. With respect to alcohol dehydrogenase, such studies, proposed above but not carried out in the present work, on t he kinetics of chemically modified enzymes could be of prime importance in the control of alcoholism. Thus, modifying agents may be found which increase t he turnover numbers and Michaelis constants for ethanol but not greatly change the Michaelis constants for NAD+ (15 ); such a modified enzyme could be more active in vivo (16). In these circumstances, modified ADH would become saturated at more highly intoxicating concentrations of ethanol than is nat ive enzyme (16).
0
~ X
TIME (min) Changes in the activity of horse liver alcohol dehydrogenase against time o! incubation with the modifying agents (M) iodoacetic acid and iodoacetamide and in the presence of coenzymes (NAD +, NADH). The experimental conditions for each set of experiments .are described in the text.
inactivation by iodoacetamide suggests a role for the carboxyl group of iodoacetate in directing carboxymethylation. The speed and selectivity of carboxymethylation with iodoacetate compared to iodoacetamide led to the suggestion that the carboxyl group of the alkylating agent binds to a positive charge in the vicinity of Cys-46, possibly Arg-47 (11).
The incubation buffer was Pipes because its affinity toward t he enzyme is 6.5 times lower than that of phosphate; t his latter could be a partial protector against chemical modification (12). As previously described (1 3, 14), iodoacetic acid inactivated horse liver ADH in a reaction which was first order with respect to enzyme. Thus a semilogarithmic plot of relative enzymatic activity (V! Vc) against t ime in the first stages of t he modification is linear with the slope being t he apparent k of t he inactivation process (k = 0.06 min- 1). Another interesting possibility of t his kind of study of t he chemical modification of enzymes would be to check if after modification the kinetic parameters KM and V have altered in value compared with those values established for t he native enzyme without modification (see above). If the modification has been directed at the active site, it will logically be reflected in a variation in the maximum velocity V, and, furthermore, the KM for the substrate or coenzyme is likely to change. The influence of each chemical modification on the kind of kinetics followed by the enzyme is another interesting
Summary
The study of the functional amino acid residues involved in the anion binding or in the catalytic activity of horse liver alcohol dehydrogenase by coenzyme protection against t he chemical modification of such residues agrees well with the picture of the active site and anion binding site of the enzyme (1, 3). In this study in particular , we focused our attent ion on the chemical modification of Cys residues. In this way and in agreement with other workers, we found that Cys must be involved in the anion binding site of ADH. The kind of study described here is a first approach, before going into detail with differential labeling, affinity labeling, and other sophisticated technqiues (3) on t he nat ure of essential amino acid residues in enzymes. The experiments described in this work may be carried out in four to five 3-h laboratory sessions. Acknowledgment
An award from t he U.S.- Spanish Joint Committee for Scientific and Technological Cooperation is acknowledged. Literature Cited I. Lange, L. G.; Riordan LJ. F.; Vallee, B. L. Biochemistry 1974, 13, 4361. 2. Li, T. K.; Vallee, B. L. Biochem. Biophys. Res. Commun. 1963, 12, 44. :1. Mean s. C. E.; Feeney, R. E. Chemical Modificat ion of Proteins; Holden· Day: San Francisco, 1972. 4. Plapp, B. V.J. Bioi. Chern. 1910,245, 1727. 5. Dworschack, R.; Torr, C.; Plapp, B. V. Biochemistry 1975, 14, 200.
6. Drum, D. E.; l.i, T. K.; Vallee, B. L. Biochemistry 1969, 8,3783. 7. Abdallah. M. A.; Biellman, J . F.; Nordstrom, B.; Bran den, C.·l. Eur. J. Biochem. t975, 50,475. 8. Boyer, P . D. J. Am. Ch ern. Soc. 1954, 76,433 1. 9. Boyer, P. D.; Segal. H . L. I n A Symposium on the Mechanism of £"nzyme Action, 1st ed.; McElroy, W. D.; Glass, H. B., Eds.; Johns Ho1>ki ns: Baltimore, 1954; p 520. 10. Li, T. K.; Vallee, B. L. Biochemistry 1965, 4, 1195. II. Lange, L. G.; Riordan, J . F.; Vallee, B. L.; Branden, C.-I. Biochemistry t975, /4,3497. 12. Dahl, K. H.; McKinley- McKee, J. S. Eur. J. Biochem. 1980, /03,47. 13. Reynolds, C. H.; McKinley-McKee, J. S. Eur. J. Biochem. 1969, 10,474. 14. Dahl, K. H.: McKi nley-McKee, J . S. Eur. J. Biochem. 1977, 81, 223. 15. Zoltobrocki, M.; Kim, J . C.; P lap1>. B. V. Biochemistry 1974, 13,899. 16. Plapp, B. V. In Alcohol and Aldehyde Metabolizing Systems, 1st ed.; Thurman, R. G.; Yonetani , T .; Williamson, J. R.; Chance, B., Eds., Academic: New York, 1974; pp 91 -
100.
. Applications Sought for Senior and Postdoctoral Research Associateships The National Research Council announces the 1987 Resident, Cooperative, and P ostdoctoral Research Associateships Programs for research in the sciences and engineering to be conducted on behalf of 26 federal agencies or research institutions whose laboratories are located throughout t he United States. T he programs provide PhD scientists and engineers of unusual promise and ability with opportunities to p erform research on problems largely of their own choosing yet compatible with t he research interests of the supporting laboratory. Applications to the National Research Council must be postmarked no later t han April15 and August 15, 1987. Initial awards will be announced in July and November, followed by awards to alternates later. Information on specific research opportunities and federal laboratories, as well as application materials, may be obtained from t he Associateship Programs, Office of Scientific and Engineering Personnel, JH 608-03, National Research Council, 2101 Constitution Avenue, N.W. , Washingt on, DC 20418, (202)334-2760.
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Journal of Chemical Education
