Purification and characterization of the inactive Ca2+, Mg2+-activated adenosine triphosphatase of the unc A− mutant Escherichia coli AN120

ARCHIVES

OFBIOCHEMISTRY

Purification Adenosine

AND BIOPHYSICS

178, 486-494

and Characterization of the Inactive Ca2+,Mg2+-Activated Triphosphatase of the uric A- mutant Escherichia co/i AN120 P. D. BRAGG

Department

(1977)

of Biochemistry,

University

of British Received

AND

HOU

c.

Columbia,

Vancouver,

B.C., Canada

V6T 1 W5

June 1, 1976

The inactive Ca2+, Mg2+-activated ATPase of the uric A- mutant Escherichia coli AN120 and the active enzyme of the normal strain AN180 were purified to homogeneity. Both enzymes contained five different subunits (a-e) with the same molecular weights and stoichiometry. Both enzymes gave identical cross-linked products with the bifunctional reagent dithiobis(succinimidy1 propionate), suggesting that the subunits were arranged in the same way in the two enzymes. Treatment of the enzyme of AN120 with Natosylphenylalanine chloromethyl ketone-trypsin did not reveal latent ATPase activity. The ATPase of AN120 bound normally to ATPase-depleted membranes of AN180 or AN120 to reconstitute aerobic-driven energy-dependent transhydrogenase activity. The enzymes from both AN120 and AN180 contained one to two molecules of both ATP and ADP per molecule of ATPase. Each enzyme could also bind one molecule of exogenous F’HIADP per molecule of ATPase. The dissociation constants of the 13HlADP-ATPase complexes formed by the enzymes from AN120 and AN180 were 3.55 and 9.3 pM, respectively. Both enzymes reacted with 7-chloro-4-nitrobenzo-2-oxa-1,3-diazole with the substitution of a tyrosine residue at the active site. It is concluded that the loss of hydrolytic activity in the ATPase of AN120 is due to an alteration at the active site which is reflected in nucleotide binding but may involve a catalytically active amino acid.

determining the role of the Ca2+, Mg2+activated ATPase in energy-linked procoxidative phosphorylation have become of esses such as active transport (11, the muincreasing importance in elucidating the tants containing the inactive ATPase are function of the components of the various potentially of use in studies of the cataenergy-linked systems present in the lytic mechanism of this enzyme. membranes of these organisms (1). The The first mutant of this type, E. coli mutants may be grouped into several AN120, was isolated by Butlin et al. (9). It classes (2-61, in one of which (uric A-) the was subsequently shown by the same activity of the membrane-bound Ca2+, group that the inactive enzyme could be Mg2+-activated ATPase’ is much lower released from the membrane by washing than normal. In these mutants the ATP- at low ionic strength, but the enzyme was ase may be present in an inactive form (7) not further characterized (7). In this paor absent (7, 8). Besides being useful in per we report the purification and preliminary characterization of the ATPase of AN120. ’ Abbreviations used: ATPase, adenosine triphos-

Mutants of Escherichia coli and Salmonella typhimurium which are defective in

phatase; NBD-chloride, 7-chloro-4-nitrobenzo-2-oxa1,3-diazole; Tricine, N-tris(hydroxymethyl)methyl glycine; DCCD, dicyclohexylcarbodiimide; SDS, sodium dodecyl sulfate; DEAE, diethylaminoethyl; TPCK, Nstosylphenylalanine chloromethyl ketone. 486 Copyright All rights

0 1977 by Academic Press, Inc. of reproduction in any form reserved.

MATERIALS

AND

METHODS

Materials. Bovine serum albumin, phosphoenolpyruvate, and rabbit muscle pyruvate kinase were obtained from Calbiochem. ADP and ATP, and fire-

MUTANT

ATPase

487

dex G-25 (2 x 12 cm) equilibrated with the same fly lantern extract and 7-chloro-4-nitrobenzo-2-oxabuffer. The “urea” particles were resedimented from 1,3-diazole (NBD-chloride) were purchased from PL Biochemicals and Sigma Chemical Company, rethe column effluent as above. spectively. [G-3H]ADP was supplied by New Eng Reconstitution of transhydrogenase activity. The land Nuclear Corp. stripped or urea particles were suspended in 50 mM Tris-H,SO, buffer, pH 7.8, containing 10 mM MgCl,, Escherichia coli AM180 (arg E-, thi?) and AN120 and 1% bovine serum albumin, at a concentration of (arg E-, t/z-, uric A401), a generous gift of Professor 20-32 mg of protein/ml. The particles (25 or 50 ~1) F. Gibson (Australian National University, Canwere preincubated for 5 min at 37°C with various berra), were grown at 37°C to the late exponential amounts of purified ATPase in a cuvette with 1 ml of phase with vigorous aeration on a minimal salts medium containing 22 m&i glucose, 12 pM ferric Tris-H,SO, buffer, pH 7.8, containing 10 mM MgCl,, 0.1% bovine serum albumin, 0.1 mM dithiothreitol, citrate, thiamine. HCl(1 mg/liter), arginine (50 mgl preincubation, the liter), and 0.1% (w/v) casein hydrolysate (Difco viand 0.16 M sucrose. Following tamin-free casamino acids). For experiments involvtranshydrogenase assay was carried out directly on the contents of the cuvette. The final volume of the ing transhydrogenase, the cells were grown on the same medium but without casein hydrolysate. complete assay mixture was 1.2 ml. Preparation of membrane particles, coupling facEnzyme assays. Protein, NADH oxidase, Ca”+ATPase, respiration and ATPtor, and purified ATPase. Cells were converted to and Mg2+-activated driven energy-dependent transhydrogenase, and enwashed respiratory particles as previously described ergy-independent transhydrogenase activities were (10). The washed particles were suspended in, and dialyzed overnight against, 1 rnM Tris-HCl buffer, measured exactly as described previously (11) except that the final concentration of sucrose in the transpH 7.5, containing 0.5 mM EDTA, 0.1 mM dithiothreitol, and 10% glycerol (10). The dialyzed particle hydrogenase assays was 0.133 M. ATPase activity suspension was centrifuged (12O,OOOg,2 h) to yield a was determined in the presence of 5 mM CaCl, unpellet of “stripped” particles. The supernatant fluid less indicated otherwise. Enzyme activities were measured at 37°C. was recentrifuged to remove residual particulate material and then concentrated to about one-tenth Gel electrophoresis. Analytical disc gel electroof its volume by ultrafiltration through an XM 100 phoresis was performed either on 5% polyacrylamide membrane (Amicon Corp.) to yield the “coupling buffer, gels prepared and run in 0.05 M Tris-glycine factor” (protein, 8-10 mg/ml). pH 8.7, or on 5 or 7.5% polyacrylamide gels containATPase was purified to homogeneity from the ing 0.1% sodium dodecyl sulfate (SDS). For the latcoupling factor by chromatography on DEAE-celluter system, the protein sample was depolymerized at lose followed by sucrose gradient centrifugation. 37°C for 1 h, and then at 100°C for 3 min in 4 M urea1% SDS-l% mercaptoethanol, and run as described The procedure followed the method used previously (10) except that chromatography on Sepharose 6B previously (12). Protein bands were stained with was omitted since it did not improve the purity of Coomassie blue and the gels were scanned at 500 nm with a Gilford Model 240 spectrophotometer the enzyme and the sucrose gradient was prepared in triethanolamine buffer in place of the usual Tris equipped with a linear transporter. buffers and dithiothreitol was omitted. The same Reaction ofATPase with NBD-chloride. To 0.5 ml procedure was used for preparing the ATPases from of ATPase (l-2 mg of protein/ml) in 0.02 M triethaboth AN180 and AN120. Although the ATPase ac- nolamine buffer, pH 7.5, containing 0.5 mM EDTA tivity could not be measured in fractions from and 22.5% sucrose was added 0.05 ml of 0.5 M triAN120, the fractions equivalent to those for the tine-KOH buffer, pH 9.5. This adjusted the pH of preparation of the ATPase of AN180 were taken at the enzyme solution to pH 8.5. NBD-chloride (5 ~1 of each step. Following sucrose density gradient cen- a 0.1 mM solution in ethanol) was added and the trifugation, the enzyme was located by analysis of solution was kept for 60 min at room temperature. fractions by SDS polyacrylamide gel electrophoreThe reacted ATPase was then separated from unsis. reacted NBD-chloride by chromatography on a “Urea” particles were prepared by suspending Sephadex G-50 column (9 x 135 mm) equilibrated stripped particles (65-100 mg of protein) to 4 ml in 1 with 0.05 M triethanolamine buffer, pH 7.5, containmM Tris-HCl buffer, pH 7.5, containing 0.5 mM ing 4 mM EDTA and 0.2 M sucrose. The reacted EDTA, 0.1 mM dithiothreitol, and 10% glycerol. ATPase appeared in the void volume of the column, Urea (10 M) in the same buffer was added with The entire procedure was carried out in the dark. Estimation ofprotein-bound ATP and ADP. Samstirring to give a final concentration of 2 M. After incubating for 30 min at O”C, the particles were ples (0.1 ml) of the ATPase at about 1 mg of protein/ sedimented by centrifuging at 224,000g for 1 h and ml in 0.02 M triethanolamme buffer, pH 7.5, conthen suspended in 1 ml of the above buffer without taining 0.5 mM EDTA and 22.5% sucrose, were diurea. Contaminating urea was removed from this luted with an equal volume of water. Perchloric acid (0.02 ml; 40%) was added and after incubation for 10 suspension by passage through a column of Sepha-

488

BRAGG AND HOU

min at 0°C the precipitate was removed from the solution by centrifuging. Samples (5 ~1) of the supernatant were analyzed for ATP and ADP without neutralization. The ATP content was determined by the luciferase method using the procedure of Stanley and Williams (13). ADP was determined in the same system following conversion to ATP by preincubating the components of the assay system with the extract and standard amounts of ADP in the scintillation vial with 0.01 ml of 0.01 M phosphoenolpyruvate and 1.5 units of pyruvate kinase for 10 min at room temperature prior to the addition of the firefly lantern extract. The amount of ADP was then calculated by subtracting the previously determined value for ATP from the total amount of ATP. In order to correct for possible quenching, standard solutions of ATP and of ADP in the range of 2.5 .x lo-l2 to 5 x lo-ii M were assayed in the presence of the perchloric acid extract and the light emission observed was corrected by subtraction of the value obtained with the same amount of extract but without added ATP or ADP. Light emission was measured with a Unilux II scintillation spectrophotometer (Nuclear Chicago), with the coincidence gate switched off, for 0.2 min at 2.0 min after the addition of firefly lantern extract. Binding of 13HIADP by ATPase. ATPase (0.07 to 0.125 mg of protein) was incubated for 60 min at room temperature in 0.5 ml 0.05 M Tris-HCl buffer, pH 8.0, containing 4.5% sucrose, 1 mM EDTA, 0.25 nmol of 13H1ADP (5.56 Cilmmol), and 1 to 50 nmol of nonradioactive ADP. Controls were set up in which the enzyme was omitted. The reaction was terminated by adding 0.2 ml of bovine serum albumin (10 mg/ml in Tris-EDTA buffer) and 5.0 ml of 3.5 M ammonium sulfate solution in Tris-EDTA buffer. The mixture was filtered through a Millipore filter (0.45-pm pore size; 25 mm diameter) which was then washed successively with 0.5-, 0.5-, l.O-, 0.5-, and 0.5-ml volumes of the ammonium sulfate solution. The filters were dried under an infrared lamp and dissolved in Bray’s scintillation fluid, and the radioactivity was determined in a Packard Tri-Carb liquid scintillation spectrophotometer, Model 2425.

alfold by certain ions. We have examined the effect of 0.3 M KC1 and 0.3 M sodium acetate on the Ca2+- and Mg2+-activated ATPase activities of both membrane particles and the solubilized enzyme (“coupling factor”) in order to determine if inclusion of these salts in the assay mixture for the enzyme would allow us to detect it during purification. Although little stimulation was obtained with KCl, sodium acetate was more effective particularly with the solubilized enzyme where eight- to tenfold stimulation was observed (Table I). Thus, in our initia\ experiments on purifying the ATPase of A~120 we assayed ATPase activity with and without 0.3 M sodium acetate. Purification of this ATPase to homogeneity was attained by following the procedure used previously to purify the active enzyme from several strains of E. coli and Salmonella typhimurium (5, 10, 12). This method involved chromatography of the coupling factor on DEAE-cellulose followed by sucrose density gradient centrifugation. The location of the ATPase of AN120 during this procedure was followed by SDS-gel electrophoresis since the subTABLE I EFFECT OF SALTS ON ATPase ACTIVITIES OF MEMBRANE PARTICLES AND COUPLING FACTOR OF

pH

Activator

As originally shown by Butlin et al. (9), the Ca2+- or Mg2+-stimulated ATPase activity in membrane particles of the mutant strain AN120 is much lower than in the normal strain AN180. However, Gunther and Mariss (14) have reported that the ATPase activity of membrane particles of the mutant strain can be stimulated sever-

Specific activityb Particles C;;fJi;g

7.6

Mg

None KC1 Na-acetate

12 18 22

15 14 142

Ca

None KC1 Na-acetate

8 8 12

3 0 7

W

None KC1 Na-acetate

15 19 23

15 15 120

Ca

None KC1 Na-acetate

7 8 12

0 0 I

RESULTS

Purification of the Ca2+, MgzC-Activated ATPase of AN120

AN120” Addition

8.5

a ATPase activity was assayed with 2.5 mM MgCl, or CaCl,. The concentration of KC1 and sodium acetate, when present, was 0.3 M. b Nanomoles per minute per milligram of protein.

MUTANT

units of the enzyme could be readily detected. Acetate-stimulated ATPase activity cochromatographed with the enzyme subunits on DEAE-cellulose but was completely separated from them on the sucrose gradient. The fractions containing the acetate-stimulated ATPase activity were shown to contain acetokinase by the method of Lipmann and Tuttle (15). Therefore, it is probable that acetokinase was responsible for the acetate-stimulated ATPase activity. The Ca”+, Mg2+-ATPase of AN120 sedimented in the sucrose gradient to a position identical to that found with the normal strain AN180. These fractions contained no detectable Caz+-activated ATPase activity. The Mg*+-stimulated ATPase activity was less than 0.2% of the normal strain. KC1 (0.3 M) and 0.3 M sodium acetate did not stimulate the ATPase activities of these fractions. Treatment of the ATPase from AN120 with TPCK-trypsin as previously described (12) did not reveal latent enzyme activity. Cross-Linking

of ATPase Subunits

The purified ATPases of AN120 and AN180, both of which gave a single protein band on electrophoresis on polyacrylamide gels in Tris-glycine buffer, pH 8.7, were compared by SDS-polyacrylamide gel electrophoresis (Fig. 1). Both enzymes gave five polypeptides which had identical molecular weights to those previously determined for the (Y-Esubunits of the ATPases from other strains ofE. coli (12). The ratio of the five subunits, as determined by the intensity of staining with Coomassie blue, agreed with the previously found stoichiometry of (Y&$E (12). Treatment of these ATPases with the bifunctional cross-linking reagent dithiobis(succinimidy1 propionate) by the method used before (12, 16) and examination of the products by SDS-polyacrylamide gel electrophoresis gave the results shown in Fig. 1. The ATPases of AN120 and AN180 gave identical results. The y-e subunits were not detected on the gels (results not shown) while the (Y and /3 subunits were much reduced in amount but both to the same extent. Although

489

ATPase

FIG. 1. Electrophoresis on 7.5% (1,2) and 5% (3,4) polyacrylamide gels containing SDS of untreated purified Ca*+, Mg*+-activated ATPase (1, AN180; 2, AN120) and cross-linked ATPase (3, ANHO; 4, AN120). Peaks a-e represent subunits a-~. Peak Y is the a-P cross-linked product.

other, higher molecular weight crosslinked products could be found, the major new band was that of the a-p cross-linked dimer described previously (12). These results indicate that the ATPases from both the mutant and the normal strain have identical subunit composition and arrangement. Coupling

Factor Activity

Washing membrane particles of E. coli with low ionic strength buffer results in the loss of both aerobic- and ATP-driven energy-dependent transhydrogenase activities. This is due to removal of the Ca2+, Mg2+-activated ATPase since addition of the purified enzyme will restore these activities (10). The aerobic-driven transhydrogenase activity only is recovered by adding dicyclohexylcarbodiimide (DCCD). This is probably due to the ability of DCCD to decrease the permeability of stripped membrane particles to protons (17).

490

BRAGG

AND

Membrane particles of AN180 have both aerobic- and ATP-driven transhydrogenase activities, whereas only the former is present in particles from AN120 (Table II). Removal of the ATPase from membrane particles of these strains by dialysis at low ionic strength resulted in complete loss of the ATP-driven transhydrogenase of AN180 and in the reduction of the aerobic-driven activities of both strains by about 80%. The residual aerobic-driven activity was removed by washing the stripped particles with 2 M urea (Table II). Urea treatment removed further amounts of ATPase but had no effect on energyindependent transhydrogenase and NADH oxidase activities of the membrane particles. Addition of the purified ATPase of AN180 restored the aerobic- and ATPdriven transhydrogenase activities of both stripped and urea-treated particles from strains AN180 and AN120. This confirms the observation of Cox et al. (18) that the absence of the ATP-driven activity in ZLW A- mutants was due to modification of the ATPase and not due to alterations in the rest of the transhydrogenase system. Addition of the purified ATPase of AN120 or DCCD restored only the aerobic-driven transhydrogenase in urea-treated particles of AN180 and AN120. If stripped particles of AN180 were used, then the ATPase of AN120 could stimulate the ATPdriven reaction to some extent. It is probable that the stripped particles of AN180

still contain some latent ATPase which is not present in sufficient amounts to retain the impermeability of the membrane to protons (17,19). Thus, this ATPase cannot drive the ATP-dependent transhydrogenase reaction. Addition of the ATPase of AN120, although unable to drive the ATPdependent transhydrogenation, will restore the proton impermeability of the membrane and permit the residual amount of bound active ATPase to carry out this reaction. Half-maximal activation of the aerobicdriven and ATP-driven transhydrogenase activities of urea-treated particles of AN180 was given by 31 and 35 kg of AN180 ATPase protein/mg of particle protein, respectively. Half-maximal activation of the aerobic-driven reaction in the same particles was obtained with 33 pg of AN120 ATPase protein/mg of particle protein. This indicates that the ATPase of AN120 binds as readily to the membrane as that of AN180. Reaction ofATPase

with NBD-Chloride

The ATPases of mitochondria and chloroplasts are inactivated by the binding of 1 mol of 7-chloro-4-nitrobenzo-2-oxa-1,3diazole (NBD-chloridellmol enzyme. A single tyrosine residue at the active site of the enzyme reacts with this reagent as shown by the appearance of a characteristic absorbance maximum at about 385 nm in the spectrum of the substituted enzyme

TABLE EFFECT OF ATPases

HOU

II

OF NORMAL AND MUTANT STRAINS ON ENERGY-DEPENDENT TRANSHYDROGENA~E ACTIVITIES OF STRIPPED PARTICLE@

Addition

Transhydrogenase

activity

of particled

Aerobic AN180 None DCCD ATPase (AN180) ATPase (AN120)

AN180 (4

ATP AN120

AN120 (u)

AN180

AN180 (4

AN120

AN120 (4

4 18 25

5 20 29

10 24 27

13 16 26

0 0 53

0 0 29

0 0 33

0 0 25

22

30

25

23

6

0

0

0

a Membrane particles of stripped AN180 (0.63 mg) or AN120 (0.97 mg) or urea-stripped AN180 (u) (1.1 mg) or AN120 (u) (1.6 mg) were incubated with DCCD (0.1 pmol) or purified ATPase of AN180 (0.06 mg) or AN120 (0.11 mg) for 5 min at 37°C prior to the assay. * Nanomoles per minute.

MUTANT

(20, 21). NBD-chloride has been found to inhibit the ATPase of E. coli also. A group on the P-subunit was covalently modified by the reagent but was not identified (22). Since thiol and amino groups also react with NBD-chloride, we attempted to identify the reactive amino acid. Moreover, since this group appeared to be involved at the active site of the ATPase it was possible that it had been replaced in the mutant enzyme. The ATPases of AN120 and AN180 were reacted with NBD-chloride and the excess reagent removed by gel filtration on Sephadex G-50. The yellow substituted enzyme emerged in the void volume of the column. The spectra of the reacted enzymes are shown in Fig. 2. The spectra of the products from AN120 and AN180 are very similar, both showing an absorption maximum at 390 nm which is due to the NBD derivative of a tyrosine residue. Addition of dithiothreitol (data not shown) results in the loss of this absorption peak as has been observed also with the mitochondrial ATPase (20). These results suggest that the loss of activity produced by the addition of NBDchloride to the ATPases of wild-type strains ofE. coli is due to the substitution of a tyrosine residue on the p-subunit of the enzyme. The lack of enzyme activity in the ATPase of AN120 cannot be due to replacement of this tyrosine residue.

491

ATPase

L

300

I

400

500

Wavelength “Ill

FIG. 2. Absorption spectra of purified Ca”+, Mg2+-activated ATPases of AN180 and AN120 after reaction with NBD-chloride. BL, baseline.

the nucleotides could also function in the regulation of ATPase activity (28, 29). Slater (23) has reported that, in contrast to membranes of wild-type E. coli, the membrane of an unspecified ATPase-negative mutant of this organism did not contain bound ATP or ADP. We measured the bound nucleotides of the purified ATPases of AN180 and AN120. The nucleotides were released from the enzyme by perBound Nucleotide Content of ATPase chloric acid and analyzed by the luciferase Tightly bound nucleotides have been assay for ATP. ADP was converted to ATP found in the energy-transducing mem- with phosphoenolpyruvate and pyruvate kinase prior to analysis. The purified branes of mitochondria, chloroplasts, ATPases of AN180 (six separate analyses) chromatophores of Rhodospirillum ruband AN120 (four separate analyses) conrum, cell membranes ofE. coli, andstreptococcus faecalis (23-25). Some of the nu- tained 1.13 & 0.09 (-t standard deviation) cleotide is associated with the coupling and 1.45 -+ 0.31 molecules of ATP per molecule of ATPase, and 1.79 -+ 0.55 and 1.79 ATPase of these organisms since purified beef-heart ATPase contains three mole-* + 0.39 molecules of ADP per molecule of cules of ATP and two molecules of ADP ATPase, respectively. per molecule of enzyme and one molecule of each nucleotide per molecule of enzyme ADP Binding to the ATPase is found in the purified ATPase of 5’. faeExogenous ADP will bind to the isocalis. A role for these nucleotides in the lated ATPases of microorganisms as well phosphorylation reactions of oxidative as to mammalian mitochondrial and chlophosphorylation or photophosphorylation roplast ATPases (28-30). Smith and has been suggested (23, 26, 27). However, Abrams (31) found that the dissociation

492

BRAGG

AND

HOU

constant for the ATPase-ADP complex formed from the S. faecalis enzyme was 2 x 1O-7 M at pH 7.5 in the presence of EDTA. The stoichiometry of binding was uncertain, but two binding sites may be present. The enzyme from Alcaligenes faecalis reacted with exogenous ADP to form a complex with a dissociation constant of 1.5 x 10-j M at pH 8 in the presence or absence of 1 mM MgCl,. The molar binding ratio varied between 0.4 and 1.8 (32). A molar binding ratio of 1.7 molecules of ADP per molecule of enzyme and a dissociation constant of 8.1 x lo-” M at pH 7.4 in the presence of 5 mM MgCl, was given by the purified ATPase of Mycobacterium phlei

(33).

0

0

a3

0.6

The binding of 13HlADP to the ATPases ii of AN180 and AN120 was measured at pH FIG. 3. Binding of [3H]ADP to purified Ca*+, 8.0 in the presence of 1 mM EDTA. The Mg*+-activated ATPases of AN180 and AN120. The ATPase-ADP complex was precipitated data are plotted as recommended by Scatchard (34) and the lines are drawn by the least-squares by the addition of bovine serum albumin, as a carrier protein, followed by ammo- method. v, molecules of ADP bound per molecule of ATPase; S, micromolar concentration of free ADP. nium sulfate solution and the precipitate collected on a Millipore filter. Control exThese results indicate that exogenous periments showed that bovine serum alADP will form a more stable complex with bumin did not bind ADP. Furthermore, the ATPase of AN120 than with that of Catterall and Pedersen (30) have demonAN180. strated that the ammonium sulfate precipitation step does not result in the DISCUSSION release of ADP from the mitochondrial exogenous ADP-ATPase complex. The reThe purified ATPase from the mutant sults were plotted by the method of Scat- strain AN120 differs from that of the norchard (34) (Fig. 3). The ATPase-ADP mal strain AN180 by the almost complete complexes of AN180 and AN120 had disso- absence of hydrolytic activity with Ca*+ or ciation constants of 9.3 and 3.55 PM, re- Mg*+ as activating ions. This is not due to spectively. The stoichiometries of binding the loss of subunits since the mutant enwere 0.62 and 0.63 molecules of ADP per zyme has identical sedimentation behavmolecule of enzyme, respectively. In other ior to the parent in a sucrose gradient and experiments where the precipitated en- contains five polypeptide subunits which zyme-ADP complex on the Millipore filter appear to be identical both in molecular was washed twice instead of five times weight and stoichiometry to those of the with the ammonium sulfate solution, the ATPase of the normal strain. Moreover, stoichiometry of binding for the ATPases cross-linking experiments suggest that of both strains was 0.8 to 1.1 molecules of the subunits of AN120 and AN180 are arADP per molecule of enzyme and the dis- ranged in a similar manner. sociation constants for the complexes of The E subunit of the ATPase can act as the ATPases of AN180 and AN120 were 15 an inhibitor of the hydrolytic activity of to 25 and 7.7 to 8.6 PM, respectively. These the enzyme (12,35). This inhibition can be values are probably less reliable than relieved by trypsin treatment which dethose given above since there was a stroys this subunit (12, 22). Trypsin treatgreater scatter of points in the Scatchard ment of the purified ATPase of AN120 did plots due to contamination of the enzyme- not reveal latent hydrolytic activity. ADP complex by free [3H]ADP. Thus, the absence of activity in this ATP-

MUTANT

ase is not due to stronger binding of the E subunit to the enzyme. Furthermore, the ability of the mutant ATPase to bind to ATPase-depleted membranes to act as a coupling factor to restore the aerobicdriven energy-dependent transhydrogenase indicates that the 6 subunit, which is involved in the binding (35), is unaltered in the mutant. The amount of enzyme required for half-maximal activity was the same for both mutant and normal enzymes . The tightly bound ADP and ATP found in isolated mitochondrial and other energy transducing ATPases may have a role at the active site or in the control of the enzyme (23, 26-29). However, the ATPases of AN120 and AN180 have similar levels of tightly-bound ADP and ATP (one to two molecules of both nucleotides per molecule of ATPase) so that the difference between the two enzymes is not caused by different levels of bound nucleotides. This may not hold for all mutants of E. coli in which the ATPase activity is absent since Slater (23) has shown that membranes of another mutant, in contrast to those of the parent, had no bound ATP and ADP. Extra ADP can be taken up by the enzyme on incubation with exogenous ADP. The relationship between this and the tightly bound ADP and ATP is not clear. In mitochondrial ATPases two binding sites can be detected. The dissociation constant of the low affinity binding site appears to be similar to the Ki for competitive inhibition of ATP hydrolysis by ADP, whereas the high affinity site has a similar dissociation constant to the K, for ADP in oxidative phosphorylation (30). In the ATPases of AN180 and AN120, only one binding site with a dissociation constant of 9.3 and 3.55 PM, respectively, could be detected. Thus, exogenous ADP is bound more strongly by the mutant enzyme. However, it is probably unlikely that this would be responsible for the complete loss of ATPase activity in the mutant enzyme. The Ki for competitive inhibition of ATP hydrolysis by ADP in normal strains is 0.3 to 0.75 mM (36, 37). This binding site for ADP was not detected in our experiments, probably because the

493

ATPase

bound ADP held weakly at this site would be removed in the washing steps used in our assay procedure. An attempt to use the ultrafiltration technique of Hilborn and Hammes (28) to separate the ADPenzyme complex from the reaction mixture was unsuccessful due to the difficulty of completely removing unbound 13HlADP from the filters. The binding site that we measured in the ATPases of AN180 and AN120 may correspond to the high affinity site found in the mitochondrial enzyme. However, the K, for ADP for oxidative phosphorylation has not yet been determined for E. coli. Tyrosine and arginine residues, and possibly carboxyl and amino groups, appear to be involved at the active site of the E. coli ATPase (38). We have found that a tyrosine residue reacts with NBD-chloride with complete inactivation of the enzyme. However, this reactive amino acid is present in both normal and mutant enzymes and so cannot be responsible for the difference in their hydrolytic activities. We have not yet examined the possible replacement of other potential active site amino acids in the enzyme of AN120. We conclude that the loss of hydrolytic activity in the mutant enzyme is due to an alteration at the active site of the enzyme which is reflected in nucleotide binding but may involve a catalytically active amino acid. It does not seem to involve changes in the stoichiometry of the subunits or in their arrangement in the enzyme molecule. ACKNOWLEDGMENTS We are pleased to thank Professor Frank Gibson (Australian National University) for the generous gift of the strains used in this work and the Medical Research Council of Canada for financial support. REFERENCES 1. SIMONI, R. D., AND POSTMA, P. W. (1975) Ann. Rev. Biochem. 44, 523-554. 2. COX, G. B., AND GIBSON, F. (1974) Biochim. Biophys. Actu 346, l-25. 3. DANIEL, J., ROISIN, M.-P., BURSTEIN, C., AND KEPES, A. (1975) Biochim. Biophys. Acta 376, 195-209. 4. KANNER, B. I., NELSON, N., AND GUTNICK, D. L. (1975) Biochim. Biophys. Acta 396, 347-359.

494

BRAGG

5. KAY, W. W., AND BRAGG, P. D. (1975) Biochem. J. 150, 21-29. 6. THIPAYATHASANA, P. (1975) Biochim. Biophys. Acta 408, 47-57. 7. Cox, G. B., GIBSON, F., AND MCCANN, L. (1974) Biochem. J. 138, 211-215. 8. ROSEN, B. P. (1973) J. Bacterial. 116, 1124-1129. 9. BUTLIN, J. D., Cox, G. B., AND GIBSON, F. (1971) Biochem. J. 124, 75-81. 10. BRAGG, P. D., AND Hou, C. (1972)FEBS Lett. 28, 309-312. 11. BRAGG, P. D., DAVIES, P. L., AND Hou, C. (1972) Biochem. Biophys. Res. Commun. 47, 12481255. 12. BRAGG, P. D., AND Hou, C. (1975) Arch Biothem. Biophys. 167, 311-321. 13. STANLEY, P. E., AND WILLIAMS, S. G. (1969) Anal. Biochem.,29, 381-392. 14. GUNTHER, T., AND MARISS, G. (1974) 2. Naturforsch. 29c, 60-62. 15. LIPMANN, F., AND TUTTLE, L. C. (1945) J. Biol. Chem. 159, 21-28. 16. BRAGG, P. D. (1975)J. SupramolecularStruct. 3, 297-303. 17. ALTENDORF, K., HAROLD, F. M., AND SIMONI, R. D. (1974) J. Biol. Chem. 249, 4587-4593. 18. Cox, G. B., GIBSON, F., AND MCCANN, L. (1973) Biochem. J. 134, 1015-1021. 19. HINKLE, P. C., AND HORSTMAN, L. L. (1971) J. Biol. Chem. 246, 6024-6028. 20. FERGUSON, S. J., LLOYD, W. J., LYONS, M. H., AND RADDA, G. K. (1975) Eur. J. Biochem. 54, 117-126. 21. DETERS, D. W., RACKER, E., NELSON, N., AND NELSON, H. (1975) J. Biol. Chem. 250, 10411047. 22. NELSON, N., KANNER, B. I., AND GUTNICK, D. L. (1974) Proc. Nat. Acad. Sci. USA 71, 2726-

AND

HOU

2724. 23. SLATER, E. C. (1974) Biochem. Sot. Trans. 2, 1149-1163. 24. YAMAMOTO, N., YOSHIMURA, S., HIGUTI, T., NISHIKAWA, K., AND HORIO, T. (1972) J. Biothem. 72, 1397-1406. 25. ABRAMS, A., NOLAN, E. A., JENSEN, C., AND SMITH, J. B. (1973) Biochem. Biophys. Res. Commun. 55, 22-29. 26. BOYER, P. D., CROSS, R. L., AND MOMSEN, W. (1973) Proc. Nat. Acad. Sci. USA 70, 28372839. 27. HARRIS, D. A., AND SLATER, E. C. (1975) Biochim. Biophys. Actu 387, 335-348. 28. HILBORN, D. A., AND HAMMES, G. G. (1973) Biochemistry 12, 983-990. 29. CANTLEY, L. C., AND HAMMES, G. G. (1975) Biochemistry 14, 2968-2975. 30. CATTERALL, W. A., AND PEDERSEN, P. L. (1972) J. Biol. Chem. 247, 7969-7976. 31. SMITH, J. B., AND ABRAMS, A. (1974) Fed. Proc. 33, 1257. 32. ADOLFSEN, R., AND MOUDRIANAKIS, E. N. (1976) Arch. Biochem. Biophys. 172, 425-433. 33. KALRA, V. K., LEE, S.-H., RITZ, C. J., AND BRODIE, A. F. (1975) J. Supramolecdar Struct. 3, 231-241. 34. SCATCHARD, G. (1949)Ann. N.Y. Acad. Sci. 51, 660-672. 35. SMITH, J. B., STERNWEIS, P. C., AND HEPPEL, L. A. (1975) J. Supramolecular Struct. 3, 248255. 36. KOBAYASHI, H., AND ANRAKU, Y. (1972) J. Biothem. 71, 387-399. 37. CARREIRA, J., AND MuAoz, E. (1975) Mol. Cell. Biochem. 9, 85-95. 38. AHLERS, J., KABISCH, D., AND GUNTHER, T. (1975) Canad. J. Biochem. 53, 658-665.