Volume 22, n u m b e r 1

MOLECULAR t~; CELLULAR BIOCHEMISTRY

N o v e m b e r 30, 1978

KINETIC CHARACTERIZATION OF PLASMA MEMBRANE ATPASE FROM SACCHAROMYCES CEREVISIAE Jan A H L E R S , Erika A H R and Angelika S E Y F A R T H

Institut fiir Biochemie und Molekularbiologie, Fachbereich Biologie, Freie Universitiit Berlin EhrenbergstraBe 26-28, 1000 Berlin 33 [Germany]

Summary 1. Plasma m e m b r a n e preparations have been isolated from spheroplasts of Saccharomyces cerevisiae, strain R XII, via lysis and subsequent differential centrifugation. These preparations are almost devoid of mitochondrial contamination. 2. The plasma m e m b r a n e A T P a s e is fairly stable when refrigerated, but loses activity at 8 °C and above. Below p H 5.6 the ATPase is irreversibly inactivated. The enzyme also splits G T P and ITP, although to a lesser extent. 3. Mg2+-ions are essential as part of the reactive substrate, MgATP, and furthermore they activate the ATPase. Optimal conditions depend on substrate concentration. When the concenSYMBOLS AND ABBREVIATIONS E, EH + S P K"

I~mH+~,/H+ KMg K~g

Ki

free enzyme with various degrees of protonation substrate product modified Michaelis constant (in the presence of activator) apparent constants, which depend on H+ ion concentration dissociation constant of the EMg complex dissociation constant of the EMgS complex inhibition constant = dissociation constant of the EMg2 complex N-Tris-(hydroxymetyl)-methyl-glycine morpholino-ethyl-sulfonicacid N,N' dicyclohexylcarbodiimide

Tricin MES DCCD ENZYMES ATPase (3.6.1.3) Chitin synthetase (2.4.1.16) cytochrom-c-oxidase (1.9.3.1)

Dr. W. Junk b.v. Publishers

tration of free Mg 2+ ions exceeds about 0.1 mM, competitive inhibition occurs. 4. In the range of p H 5 . 6 - 9 . 2 two functional groups dissociate. One, with pKb = 8.1 + 0.1 participated in substrate binding and another one with pKb ' = 8.1 + 0.1 is involved in substrate splitting. 5. The experiments with group-specific inhibitors suggest that an a - a m i n o group and a sulfhydryl residue are involved in substrate binding and conversion. Furthermore, imidazole, tryptophan and carboxyl residues may be important for the catalytic process.

INTRODUCTION Whereas mitochondrial ATPase from yeast has been quite well characterized (e.g. ref. 1-4), little is known about the structure and kinetic properties of yeast plasma m e m b r a n e ATPase. Plasma m e m b r a n e ATPase was found to be dependent on Mg 2+ ions, which can be replaced by Mn z+ ions. In contrast to the mitochondrial ATPase, it is not oligomycin-sensitive [5-7]. The plasma m e m b r a n e ATPase has a p H optimum of about 7, which is somewhat lower than that of the mitochondrial ATPase [6-8]. Similar results are reported by DELHEZ et al. [9]. SIGLER and KOTYK [10] find an inhibition of plasma m e m b r a n e ATPase by D C C D but no effect of phyenylmethylsulphonyl-fluoride. The function of yeast plasma membrane ATPase is as little understood as its structure and properties. FotrRY et al. [11] discuss a participation of the ATPase in m e m b r a n e transport via translocation of protons in accordance with the Mitchell hypothesis [12]. In order to approach the question of yeast plasma membrane ATPase, it is necessary to gain more

The Hague, The Netherlands

39

information about the structure and properties of this enzyme. For membrane-bound enzymes a kinetic characterization is very suitable as it is not necessary to purify the enzyme very much, and this enables one to work under conditions which are similar to those found in vivo. However, it is neccessary to obtain a preparation which is not contaminated by ATPases from other cell organelles (e.g. mitochondrial ATPases). In this paper we characterize the ATPase kinetically with the main emphasis on the examination of the mechanism of substrate conversion at the active site and on the mechanism of action of Mg 2÷ ions. The etiects of pH and group-specific inhibitors on the activity were studied to get information about the structure of the active site. This information is also very important for examination of the membrane, as the ATPase is the best known marker enzyme for yeast plasma membrane. Thus to get information about phospholipid-protein interactions, about distribution of individual phospholipids in the membrane, and about possible changes in the structure and function of the membrane with growth phase and/or growth conditions, a comprehensive knowledge of the ATPase is necessary. Materials and Methods

Enzyme preparation Saccharomyces cerevisiae (strain R XII, a kind gift of Dr. Kotyk, Prague) was grown in a medium containing 2% glucose, 1% yeast extract and 0.5% peptone under aerobic conditions. The cells were harvested in the exponential growth phase (3-5 x 107 cells/ml). 108 cells/ml were incubated for 15 min at 30 °C in 0.1 ~lTris-C1, pH 8, 5 m u E D T A and 5 m u dithioerythritol. After washing, 101° cells were incubated in 1 ml snail enzyme, 2 ml 20 mu dithioerythritol and 4 ml 1.2 u KC1 t o digest the cell wall. Formation of spheroplasts was followed under a light microscope. After about 30-40 min more than 95% of the cells had been converted into spheroplasts. The snail enzyme was eliminated by centrifuging at 700 x g for 2 min and washing twice with an osmotic stabilizer (0.6 M mannitol, 2% (w/v) glucose, 5 mu sodium citrate, pH 6). The pellet was then resuspended in the 1 : 2.5 diluted osmotic stabilizer (101° cells in 15 ml) and the cells were lysed in a Potter-Elvehjem homogenizer (10 strokes). The homogenate was centrifuged as described in the following scheme: Ii homogenat.e_] 5 rain centrifugation at 3500 x g ~~

Reagents

cell debris 1 discarded nuclei J

All reagents were analytical reagent grade. supernatant 0 ATP, oligomycin, and cytochrome c were pro5 rain centrifugation ducts of Boehringer (Mannheim, Germany). at 6000 x g Tris(hydroxymethyl)-aminomethane, ~ mitochondria morpholinoethyl-sulfonic acid (MES), NI supernatantll Tris(hydroxymethyl)-methylglycine (Tricin) and ascorbic acid were from Serva (Heidelberg, 10 min centrifugation Germany). Snail enzyme containing per ml at at 7500 x g least 100 000 units fl-glucuronidase (Fishman) ~ mitochondria +plasma membranes and 1 000 000 units sulfatase (Roy) was from discarded Industrie Biologique Francaise, Gennevilliers, supernatant 2 France. All group specific inhibitors except 2, 3-butanedione (Merck, Darmstadt, Germany), 20 rain centri~ugation N-ethyl-N'- (3-di-methylaminopropyl)-carbodiimide, at 31 000xg N-ethyl-5-phenylisoxazolium 3'sulfonate and Nplasma membranes cyclohexyl-N'-[2-(4-morpholinyl)-ethyl]-carbodi[ supernatant discarded imide-methyl-p-toluene-sulfonate (Fluka AG, Neu-Ulm, Germany) were from Serva (Heidelberg, Germany). All other reagents were from The pellets were washed twice and finally suspended in 20 mM Merck, Darmstadt, Germany). Tris-Cl, pH 7.5 (0.5 ml for 101° cells starting material)

3]

40

Enzyme assays ATPase: The ATPase activity was determined by continuously recording the amount of inorganic phosphate released as described recently

[13-16]. The standard assay contained 0.5 mM MgC12, 0.5 mM ATP, 100 mM Tris-C1, pH 7.5 and 10-40/~g protein in a total volume of 5 ml. [MgATP] was calculated using eqn. 1:

[MgATP] = [MgZ+]t+ [ATP]t KM~ + A T P - - 2 ~/([MgZ+]t + [ATP]t + KM~ATP)2 - [Mg2+]t[ATP]'

From the [MgATP] obtained the concentrations of free ATP and free Mg 2+ were calculated using [Mg2+] = [MgZ+]t- [MgATP]

(2)

[ATP] = [ATP]t- [MgATP]

(3)

We determined KMgATP by titrating ATP in the presence and absence of MgZ+-ions. We obtained a value of 0.22 raM, which is in good agreement with KMgATV= 0.215 mM reported by WOLF and ADOLPH[17] for similar assays. To examine the oligomycin sensitivity, the ATPase activity was determined in the presence and absence of 10/zg oligomycin/ml. To detect, whether the pH effects are reversible or not, the enzyme was preincubated for 10 min at 30 °C at various hydrogen-ion concentrations. After readjusting the pH to 7.5 the activity was determined under standard conditions. To test the influence of free Mg 2+ ions on ATPase activity one has to take into account that a part of the total amount of Mgz+ ions is complexed by ATP resulting in the true substrate MgATP. Using equations 1-3 we calculated the total amounts of MgC12 and ATP necessary to obtain distinct concentration of MgATP at various concentrations of free Mg 2+. The measurements were performed in 100 mM Tris-C1, pH 7.5. To examine the pK-values of the functional groups we performed [MgATP] variations at different pH values from 5.6-9.2. The concentration of free Mg z+ was 0.1 raM. The experiments were performed with 40 mM MES or Tricin buffer. To achieve an optimal C1- concentration, 80 mM KC1 was added. The pH-dependence of the dissociation constant have been taken into account as described recently [14].

(1)

Influence of group-specific inhibitors. About 50/xg of plasma membrane ATPase (P3) was preincubated with various inhibitors under the conditions given in Table 4. After preincubation the enzyme was transferred to 100 mM Tris-C1, pH 7.5 and the reaction was immediately initiated by adding 0.5 mM ATP and 0.5 mM MgC12. To follow the time-course of inhibition, the enzyme was preincubated with the inhibitor in presence and absence of 0.5 mM ATP and 0.5 mM MgC12. Immediately and after 2.5, 5 . . . . . . min the activity of an aliquot was determined under standard conditions. Cytochrome-c-oxidase: Cytochrome-c oxidase was determined using a slightly modified procedure of SLATER[18]. Chitin- synthetase : Chitin-synthethase-activity was determined as described by CABIn [19].

Determination of protein concentration Protein was determined by the method of SCHAFFNERand WEISSMANN[20] using bovine serum albumin as a standard.

Estimation of error All values used for kinetic characterization of plasma membrane ATPase are averages of two or more experiments performed in duplicate. To evaluate the error of the determinations of enzyme activity as well as of the determination of the kinetic constants Km and V, the ATPase activity was determined six times in duplicate at each of the following substrate concentrations: 18 ~M 35 /~M, 58 /xM, 80 tXM, 120 /zM and 200/xM. [C1-] was 67 raM. Tris-C1 was used as pH-buffer (0.1 M, pH 7.5). For each series the mean value and the standard deviation was calculated and after -i wards an examination of outliers was performed 41

Experimental and Discussion

(+4s criterion (21)). The Bartlett test [22] was used to determine whether there are significant differences for the standard deviations between each series. Furthermore, the results were tested for normal distribution by plotting the classified relative measurements xiJxi in the probability paper [23] (i stands for the different concentrations of substrates, j for the 12 repeated measurements). To examine the error of K m and V and to find out which graphical procedure is optimal to evaluate the kinetic constants, the values obtained are plotted in the following linearized graphs:

Examination of the purity of the plasma membranes High ATPase activity is not only reported to be associated with plasma membranes from yeast, but also with the mitochondrial membrane [1-4]. Thus it is necessary to separate the mitochondrial from the plasma membranes. We tried to obtain relatively pure membrane pellets by fractional centrifugation. Each fraction obtained was tested for activity of cytochrome-c oxidase (mitochondrial marker) and chitin synthetase, which is reported to be a specific plasma membrane marker [29]. The ATPase activity was also determined in the presence and absence of 10 ~g oligomycin/ml. The results are compiled in Table 1. The distribution of cytochrome-c oxidase and chitin synthetase activity shows that most of the mitochondrial membranes are precipitated in P1, although some plasma membrane contamination is present, whereas the concentration of plasma membranes is quite high in P3, which is only slightly contaminated with mitochondrial membranes. Both fractions are present in P2. These results are in agreement with the determinations of ATPase activity. The mitochondrial ATPase is reported to be oligomycin-sensitive [30] in contrast to the plasma membrane ATPase [5]. However, the different response to oligomycin cannot be taken as an absolute indication for mitochondrial-or plasma membrane, as solubilized mitochondrial ATPase is also oligomycin-insensitive [6], and the plasma

1/v versus 1/[S] [S]/v versus IS] v versus v/[S] and according to the method of Eisenthal and Cornish-Bowden [24]: V / v - Km/[S] = 1 From each of these graphs we obtained 6 results for Km and V by plotting the respective experimental values of the estimation performed in duplicate. The mean value and the standard deviation were calculated.

Derivation of the rate equation for the reaction The derivation of the rate equation was performed on the basis of a rapid equilibrium reaction according to B o t t s and MORALES[25], LAIDLER [26], OHLENBUSCH [27] and CLELAND

[281.

Table 1 Activities of some enzyme markers fraction

homogenate pellet 0 pellet 1 pellet 2 pellet 3 supernatant 3

42

cytochrom-coxidase [m units/mg]

chitin-synthetase [arbitrary units/rag]

ATPase [units/mg]

total ATPase [units per 5.10 x° cells starting material]

% residual ATPase activity after inhibition by oligomycin

6 5 80 7 3 0

1.0 2.7 1.3 5.0 10.6 1.2

0.37 0.57 1.18 1.29 1.42 0.01

146 30 15 14 22 15

66 44 35 52 79 95

membrane ATPase usually shows some oligomycin sensitivity [7, 8]. However, Table 1 indicates that fraction P1 is inhibited much more strongly than P3. The residual activities are in the same range as reported by FUHRMANNet al. [8] as well as by Kuo and LAMPEN [7]. Taken together, the results from Table 1 show that the contamination of plasma membranes by mitochondria is very slight, so that kinetic characterizations of plasma membrane ATPase are not significantly disturbed. Furthermore, storage of the membrane preparations at 18 °C inactivated the mitochondrial ATPase (unpublished results), whereas the plasma membrane ATPase remains stable (see below). Therefore possible mitochondrial contaminations do not disturb the kinetic investigations. Finally the kinetic properties of the ATPase in P1, especially the pH dependence of the kinetic constants (unpublished results), are quite different from those of P3 ATPase (see below). A contamination of the plasma-membrane ATPase by vacuoles cannot be ruled out absolutely. However, if the results of WIEMKEN [28] can be transmitted on our yeast strain, the vacuoles should not be destroyed when the spheroplasts are disrupted by the method employed in this work and thus are pelleted at lower speed. Furthermore the activity of the ATPase from vacuoles is rather low [28]. -

Estimation of error

The examination of the experimental values showed that the data are free from outliers, are

normally distributed and have a constant relative standard deviation. A random error for the individual measurement of 6.1% was found and for the measurement performed in duplicate, 4.3%. The values for Km and V obtained from the 4 graphical procedures are reported in Table 2. In accordance with the results of a theoretical paper from ATKINS and NIMMO [31] it can be seen that with the kind of error mentioned above the 4 procedures do not differ very much in accuracy and precision for the determination of Km and V. Therefore in the present paper the kinetic constants were examined by means of the double reciprocal plots. With duplicate measurements the error for Km is about 10% and for V 5%. ATPase stability To be able to perform kinetic characterizations it is absolutely necessary to avoid changes in enzyme activity and properties during the whole time necessary to perform a series of experiments (e.g. substrate variations at various concentrations of activator). To detect the influence of pH on the kinetic constants it is furthermore important to perform the measurements in a pH range where no irreversible denaturation takes place. In Fig. 1 the relative activity is plotted as a function of the time of storage under various conditions. It can be seen that the plasma membrane ATPase cannot be stabilized completely. However, storage at - 1 8 °C leads only

Table 2 Comparison of four plots for the determination of kinetic constants K~

Sx

[~M]

[/zM]

47.8

4.5

9.4%

1.133

0.048

4.2%

49.7

5.8

11.7%

1.132

0.064

5.6%

v - [S---]

49.0

5.4

11.1%

1.147

0.055

4.8%

according to Eisenthal and Cornish-Bowden [24]

48.0

4.9

10.1%

1.137

0.044

3.9%

plot

1

1

v IS] Is]

---[S] V

V

s , . 100

Km

V

s,

Sx. 100

V

~ moles Pi min mg

43

- ~ i

I

I

I

I

I

I

I

ments are not disturbed by the presence of an unspecific phosphatase.

Influence of [Mg 2÷] on ATPase activity In Fig. 2 the influence of the concentration of free Mg 2÷ ions at various [MgATP] on plasma membrane ATPase activity is plotted. We obtained bell-shaped curves with a wide optimum, similar to results obtained from E, coli ATPase [32], The [Mg 2÷] optimum rises with increasing [MgATP]. For further characterization of the effect of Mg 2+ ions we performed [MgATP] variations at various concentrations of free Mg 2+ in the range where Mg 2+ activates and in the range where it inhibits. In Fig. 3 the 1/v versus 1/[Mg 2+] plots at different concentrations of MgATP are shown for the activating range. We obtained straight lines with a common point of intersection on the abscissa. Plotting 1/v versus 1/[MgATP] for various concentrations of free Mg 2+ (not shown), we also obtained straight lines with a common point of intersection on the abscissa. From the common points of intersection in both plots the values of

700

~C3

.

; 1

o 2

3

I

I

I~

I

/+

5

6

7

t [ days ]

Fig. 1. Stabilityof plasma membrane ATPase. Plasma membrane ATPase (pellet 3) was stored at -18°C (V1), -18 °C in the presence of 5% methanol (O), 8 °C ('O) and 22 °C (I). After the indicated time the activitywas determined under standard conditions. to about 30% loss of activity, probably from the process of refrigerating or thawing, and afterwards the activity remains fairly stable. Under the same conditions mitochondrial preparations lose their ATPase activity nearly completely (unpublished .results). Thus storage of plasma membrane ATPase under these conditions not only supplies one with enzyme of constant activity, but also destroys possible mitochondrial contamination. The examination whether the pH effects on plasma membrane ATPase are reversible or not showed that hydrogen ion concentrations in the range of pH 5.8-9.2 produce only reversible changes. When the pH is readjusted to pH 7.5 the original activity is recovered. Below pH 5.8 an inactivation occurs which cannot be reversed totally. Therefore preparations of plasma membranes described in the literature, using buffer of low pH, are not suitable.

Substrate specificity From table 3 the relative activity of plasma membrane ATPase with various substrates can be seen. The nucleotides GTP and ITP are split at only half the velocity of ATP. UTP, ADP and CTP are hydrolysed much more slowly. /3glycerophosphate, although present at a ten times higher concentration, is only converted to a negligible degree. Thus the kinetic measure44

Km and KMg can be calculated: Km= 50/zM,

KMg = 11/~M

As discussed recently for alkaline phosphatase of pig kidney (13) these results indicate a random mechanism of action of substrate and activator with the enzyme, binding of substrate and Mg 2+ ions occurring independently of each other. Table 3 Substrate specificityof yeast plasmarnembrane ATPase substratea] ATP GTP ITP UTP CTP ADP /3-glyeerophosphate /3-glycerophosphate

relative activityunder standard conditions 100 b] 47 46 13 7 13 3.6 6 (pH 9)

alThe concentrations of nucleotides were 0.5raM and of /3-glycerophosphate 5 mM bacorresponds to a specificactivityof 1.4 units/ragprotein.

I

I

I

I

I

o

1.5

_-~ ~

o S ~ ~I-.

°~ °~o

•

1.0

05

I

I

'SO

&5

I

I

/,.0

3.5

I

30 log [Mg z+]

-

Fig. 2. v versus-log[Mg 2÷] plotted for various constant concentrations of MgATP. Experiments were done at 30 °C, pH 7.5, 100 mM Tris-Cl [MgATP]: 0.02 mM (©), 0.04 mM (11) 0.08 mM (A), 0.12 mM (0)

I

I

I

I

I

0 . 2 0 mM (D)

•

•~

k o

7

~a'~"~'~

I

,

10

,

,

,

25

50

Ilqq

,

I

I

100

10

20

I

Fig. 3. 1/v versus 1/[Mg2+], plotted for various concentrations of MgATP. Experiments were done at 30 °C, pH 7.5 in 100 mM Tris-C1 [MgATP]: O = 10 ~M • = 2 0 tzr~

[] = 40 t ~ • = 60/~

A = 80 t~ra • = 120/ZM

~ ~.~ I

30

40

~ 50

~9/,~Firr~M-'J

Fig. 4. 1/v versus I/[MgATP] plotted for various concentrations of Mg 2+. Values were taken from the descending part of the plot v versus -log[Mg 2+] [Mg2+]: 0.10 mM (O). 0.18 mM (Vq) 0.32 mM (A), 0.50 mra (11) 0.71 mM (0). Inset: Plot of I ~ Mg2+ versus [Mg 2+] The values were taken from the intercepts of the straight lines of Fig. 4 with the abscissa.

45

In Fig. 4 the results from the descending part of the curves in Fig. 2 are replotted in such a way that 1/v versus 1/[MgATP] plots at various concentrations of Mg 2+ resulted. We obtained straight lines with a common point of intersection on the ordinate. To evaluate the inhibition constant Ki, we plotted the apparent Km values obtained from Fig. 4 versus [Mg z+] (inset in Fig. 4). The intercepts of the resultant straight line with the ordinate and the abscissa are Km and Ki respectively. As a mean value of three determinations we obtained Km= 40/ZM± 10/XM and Ki = 0.5 mM+0.15 mM. These results are quite similar to those obtained with E. coli ATPase [32]. From the linearity of the 1/v versus 1/[MgATP] plots at all [Mg z+] employed, we can deduce that MgATP is the true substrate. The occurrence of straight lines in 1/v versus 1/[Mg 2+] plots for activating Mg z+ concentrations and in 1/v versus [Mg z+] plots for inhibitory magnesium concentrations indicate that free ATP, whose concentration is always changed in the opposite direction when the concentration of Mg 2+ is varied at constant [MgATP] (eqns. 1-3), has no influence on ATPase activity. This interpretation is in accordance with plots of 1/v versus 1/[MgATP] at constant [ATP] but variable [Mg2+], where curved lines are obtained (not shown). The results shown in Figs. 3 and 4 are summarized in the following scheme: E + S + Mg 2+

K~',.JI

~"

ES / / + S = MgATP Mg 2+

'Jl

KMg

EMg + S • ~" EMgS +

K;'L EMg + Products

Mg 2+

EMg2 Besides gaining information about the mechanism of reaction it is of interest to deduce a rate equation, which shows how the initial rate depends on the concentrations of E, S and Mg 2+. In analogy to a recent discussion [13] and on the basis of a rapid equilibrium reaction 46

mechanism the following rate equation was calculated for the scheme above: v. IS] V =

K J l + K~g ~(l+[Mg2+]~ \ [Mg2+]] k Ki ] K~K~ + IS](1 + K~[Mg2~+])

(4)

At large concentrations of Mg 2+ equation (4) simplifies to eqn (5), which is a typical equation for a competitive inhibition and represents the results shown in Fig. 4. V. [S] V =

2+

(5)

1

At low concentrations of Mg 2÷ eqn (4) simplifies to eqn. (6), which is in accordance with the random mechanism of activation and represents the values plotted in Fig. 3 V. [S] --~g ] +[S] 1+ K~[Mg2+]] K'm 1 + [Mg2+ Influence of p H on the kinetic constants Between pH 5.6 and pH 9.2, the dependence of the reaction rate of plasma membrane ATPase on the MgATP and H + concentration was investigated at a constant free Mg 2+ concentration of 0.1 raM. At a lower pH the ATPase is no longer stable and at higher pH the activity is too low. Optimal concentrations of free Mg 2+ ions and C1- (80 mM) were chosen. At each pH value, 1/v was plotted against 1/[MgATP] (not shown). The kinetic constants KmH+ and ~x-i+ were derived from the straight lines. In Fig. 5 log ~,rI+, log (Qn+/I(mn+) and -log I ~ n+ are drawn as a function of the pH value. According to the theory given by DIXON [34], and by DIXON and WEan [35], the pK values of the functional groups which participate in binding and conversion of MgATP by the plasma membrane ATPase can be deduced from these graphs. In the plot of log ~n+ versus pH one inflection point at pH 8.1 + 0.1 appears. This means that a functional group with pK~ = 8.1 dissociates in the ES-complex. The

l

I

I

/2

I

I

I

I

I

constant (unpublished results), the difference in behaviour can be used to check the purity of the membrane fraction. The following scheme represents the results obtained with plasma membrane ATPase:

I

0 o

°

08-~:

~

\

\

r'~,.

~°\\~

E+ H

00

K~, E

o

l

I

I

I

I

I

I

I

E+HS . ~ " ES

4.7 o

0 O__

0

0 °

0

O

0

0

0

0

1

E+H+p

.o 0

O O

3.9 I

'

J

I

I

I

I

I

I

I

I

I

I

59, .55' 5.1 o

\

4.7 4.3 I

513

I

I

6~

68

I

72

I

76

I

I

80 ' 84

8~ 92pH

Fig. 5. Plots of log QH+,_ log I~mH+ and log (Vla+/I(mI~+) versus pH. Experiments were done at 30 °C. [Mg 2+] = 0.1 raM, pH buper: 40 mM MES (below pH 7) or 40 mM Tricin. The QI~+ and I(mH+ values were obtained from 1/v versus 1/[MgATP] plots. 0: measured values. Solid lines: theoretical curves, drawn according to the theory described by Dixon and Webb [,35], considering that they miss the intersection point of their asymptotes (broken lines) by a vertical distance of log. 2

protonated form of this group takes part in substrate splitting. From the plot of log (VH+/I(~+) versus pH one can see that another group, which takes part in substrate binding when it is protonated (pKb = 8.1+0.1), dissociates in the same range. This pKb may be due to a functional group in the free enzyme or may represent a dissociation constant of the substrate. As a protonisation of ATP or MgATP does not occur at pH 8 [36, 37] this group must be in the enzyme. In the pI(~ + versus pH-plot the two functional groups produce effects on the curve, thus cancelling each other and producing a straight line. As quite different plots are obtained with mitochondrial ATPase from yeast, e.g. log V increases with pH and above pH 7 remains

The pK values indicate that a sulfhydryl group of cysteine and/or an a-amino group may be involved in the catalytic process. However, as the dissociation constants of the side chains in amino acids are often altered in a protein, further investigations are neccessary to determine the nature of the functional groups which are represented by the above calculated pKvalues. One possible tool to gain such information is the use of group-specific inhibitors. However, one has to keep in mind, that experiments with such inhibitors only provide some further hints about the active site, but cannot prove absolutely, which amino-acid-sidechain is involved.

Influence of group-specific inhibitors As group specific inhibitors are seldom absolutely specific, and sometimes fail to react because of steric hindrance, it is necessary to use a great number of such inhibitors to determine the functional groups of the active and/or regulatory centers of the enzyme. Therefore to get reliable results we used, whenever possible two or more reagents for each functional group which might participate in substrate binding and conversion. In Table 4 the inhibitors employed, the main target group of each inhibitor, the conditions of incubation and the concentration where 50% inhibition is found, are compiled. It can be seen that all amino-and sulfhydryl-specific reagents inhibit the plasma membrane ATPase from Saccharomyces cerevisiae at very low concentrations. Therefore these functional groups are probably involved in the catalytic process. This is in accordance with the pK values obtained from the pH studies. However, as amino-groupspecific reagents may also react with sulfhydryl 47

Table 4 Effect of group specific inhibitors on ATPase activity time of preincubation min

plsoal

0.1 M carbonate pH 9.6

15

4.5

amino (39)

0.1 M carbonate pH 8.8

70

4.5

sulfhydryl (40)

0.05 M Tris-C1pH 7.5

60

5.9

sulfhydryl (41)

0.1 M Tris-C1 pH 7.5

15

4.5

tyrosyl (42) tyrosyl (42) arginyl (43) imidazolium (44) seryl (45)

0.1 M Tris-C1 pH 7.5 0.02 M Tris-C1 pH 8 0.05 M Tris-C1 pH 7.5 0.1 M Tris-Cl pH 6.6 0.1 M Tris-CI pH 7.5

25 60 140 10 30

3.5 1.8 3.2 1.9

tryptophane (46)

0.1 M Tris-C1 pH 7.5

15

3.0

carboxyl (47) carboxyl (47)

0.1 M Tris-Cl pH 7.5 0.02 M MES pH 6.0

60 25

4.9 3.5

carboxyl(47)

0.03 M MES pH 5.8

20

2.8

carboxyl(48)

0.03 M MES, pH 5.8

10

2.4

inhibitor

main target group

2, 4, 6-trinitrobenzenesulfonic acid 1, 2-naphtoquinone4-suffonic acid p-chloro-mercuribenzoate (pCMB) 5, 5'-dithiobis (2-nitrobenzoic acid) N-acetylimidazole tetranitromethane 2, 3-butanedione diethylpyrocarbonate phenylmethane-sulfonylfluoride 2-hydroxy-5-nitrobenzyl bromide DCCD N-ethyl-N'-(3-dimethylaminoprpyl)-carbodiimide N-cyclojexyl-N'-2-(4-morpholinyl)-ethylJ-carodiimidemethyl-p-toluene-sulfonate N-ethyl-5-phenylisoxazolium3' sulfonate

amino (38)

buffer

a] negative logarithm of inhibitor concentration causing 50% inhibition of the ATPase activity groups, one cannot in this case completely rule out the possibility, that the inhibition by these reagents is due to a reaction with SH-groups. Of the reagents for tyrosyl residues, only tetranitromethane, but not N-acetylimidazole inhibits the ATPase. As tetranitromethane may also react with sulfhydryl groups, we suppose that no tyrosyl residue is located within the active site. The reagents which react with imidazole, tryptophan and carboxyl residues inhibit the A T P a s e only at considerably higher concentrations. Thus it is possible that these functional groups are involved in the catalytic process, but one cannot rule out the possibility that unspecific reactions may be the reason for the inactivation. Except for imidazole, these groups posses p K values beyond the range where p H measurements have been performed. Therefore further information about the involvement of these groups cannot be obtained at present. Interestingly, D C C D , which is a strong and 48

specific inhibitor of bacterial m e m b r a n e ATPases (51, 52], also inhibits yeast plasma m e m brane A T P a s e at very low concentrations. Seryl- and arginyl-specific reagents did not inhibit at all. Therefore one can conclude that at least no exposed seryl- and arginyl residues seem to be involved in the catalytic process. With all reagents which inhibit the ATPase, the time-course of inactivation was examined in the presence and absence of MgCI2 and ATP. In no case were activator and substrate able to protect the enzyme from inactivation.

Conclusion These results are only a first indication of a possible structure of the active site and the mechanism of enzyme action. Much m o r e work has to be done to find out about the role of each functional group on substrate binding and conversion, as well as about binding of Mg 2÷ ions. A possible approach would be to investigate which of the kinetic constants (K~, KMg,

Kin, V, Kb, Kb), reported in this paper changes after partial inactivation by the group-specific inhibitors. Furthermore, to get the desired information it is necessary to examine the dissociation constants Ki, KMg under different states of protonisation of the functional groups. These measurements are in progress in our laboratory. Acknowledgement We thank Mrs. Sabine Rade, Mr. Andreas Cremer and Dr. Uwe Reichert for valuable discussion and Mrs. Eva Nehls for skilfull technical assistance. This work was supported by a grant of the Deutsche Forschungsgemeinschaft.

References 1. Takeshige, K., Hess, B., B6hm, M., Zimmer-Telschow, H., 1976. Hoppe-Seyler's Z. Physiol. Chem. 357, 1605-1622. 2. Grifliths, D. E., Houghton, L., 1974. Eur. J. Biochem. 46, 157-167. 3. Schatz, G., Penefsky, H. S., Racker, E. 1967. J. Biol. Chem. 242, 2552-2560. 4. Somlo, M., Reid, R. A., Krupa, M., 1977. Biochem. J. 162, 51-59. 5. Matile, Ph., Moor, H., Miihlthaler, K. 1967. Arch. Microbiol. 58, 201-211. 6. Christensen, M. S. Cirillo, V. P. 1972. J. Bacteriol. 110, 1190-1205. 7. Kuo, S. C., Lampen, J. O. 1975. Biochim. biophys. Acta 389, 145-153. 8. Fuhrmann, G. F., Wherli, E., Boehm, C. 1974. Biochim. biophys. Acta 363, 295-310. 9. Delhez, J., Dufour, J. -P., Thines, D and Goffeau, A. 1977. Eur. J. Biochem. 79,319-328. 10. Sigler, K., Kotyk, A., 1976. Mol. Cell. Biochem. 12, 73-79. 11. Foury, F., Boutry, M., Cotteau, A. 1976. Arch. intern. Biochim. 84, 618-619. 12. Mitchell, P. 1973. Bioenergetics 4, 63-91. 13. Ahlers, J. 1974. Biochem, J. 141, 257-263. 14. Ahlers, J., Kabisch, D., Giinther, Th. 1975. Can. J. Biochem. 53, 658-665. 15. Arnold, A., Wolf, H. U., Ackermann, B. P., Bader, H. 1976. Analyt. Biochem. 71,209-213. 16. Lacy, J. 1965. Analyst 90, 65-75. 17. Wolf, H. U. Adolph, L. 1969. Eur. J. Biochem. 8, 68-74. 18. Slater, E. C. 1949. Biochem. J. 44, 305-318. 19. Cabib, E. (1972) in: Methods in Enzymology (Ginsburg, V., ed.) Vol. 28, pp. 572-580, Academic Press, New York and London. 20. Schattner, W., Weissmann, C. 1973. Anal. Biochem. 56, 502-514.

21. Sachs, L. 1974. in: "Angewandte Statistik", p. 219, Springer Verlag, Berlin, New York. 22. Sachs, L. 1974. in: Angewandte Statistik", p. 384, Springer Verlag, Berlin, New York. 23. Cavalli-Sforza, L. 1974. in: Biometrie, pp. 167-170, G. Fischer Verlag, Stuttgart. 24. Eisenthal, R., Cornish-Bowden, A. 1974. Biochem. J. 139, 715-720. 25. Botts, J., Morales, M. 1953. Trans. Faraday Soc. 49, 696-707. 26. Laidler, K. J. 1956. Trans. Faraday Soc. 52, 13741382. 27. Ohlenbusch, H. D. 1962. Die Kinetik der Wirkung von Effektoren auf station~ire Fermentsysteme, SpringerVerlag, Berlin. 28. Wiemken, A. 1975. Methods in Cell Biology 12, 99-109. 29. Duran, A., Bowers, B., Cabib, E. 1975. Proc. Nat. Acad. Sci. USA 72, 3952-3955. 30. Schatz, G. 1965. Biochim. biophys. Acta 96, 342-345. 31. Atkins, G. L., Nimmo, J. A. 1975. Biochem. J. 149, 775-777. 32. Ahlers, J., Giinther, Th. 1975. Z. Naturforschg. 30c, 412-416. 33. Ahlers, J., Giinther, Th. 1975. Arch. Biochem. Biophys. 171, 163-169. 34. Dixon, M. 1953. Biochem. J. 55, 161-170. 35. Dixon, M., Webb, E. C. 1965. in: Enzymes, Longmanns, Green and Co., Ltd., London. 36. Wolf, H. U. 1972. Biochim. biophys. Acta 266, 361375. 37. Sill6n, L. G., Martell, A. E. 19641 Chem. Soc. Spec. Publ. no. 17. 38. Fields, R. 1972. in: Methods in Enzymology (Hirs, C. H. W. and Timasheff, S. N., eds). Vol. 25, pp. 464-468, Academic Press Inc., New York, London. 39. Matsushima, A., Hachimori, Y., Inada, Y. and Shibata, K. 1967. J. Biochem. 63, 328-336. 40, Riordan, J. F. and Vallee, B. L. 1972. in: Methods in Enzymology, Vol. 25, pp. 451-453. 41. Habeeb, A. F. S. A. 1972. in: Methods in Enzymology, Vol. 25, pp. 457-459. 42. Riordan, J. F., Vallee, B. L. 1972. in: Methods in Enzymology, Vol. 25, pp. 500-506, 515-521. 43. Takahashi, K. 1968. J. Biol. Chem. 243, 6171-6179. 44. Ovadi, J., Libor, S., ElSdi, P. 1967. Acta Biochim. Biophys. Acad, Sci. Hung. 2, 455-458. 45. Gold, A. M. 1972. in: Methods in Enzymology, Vol. 25, pp. 706-711. 46. Horton, H. R., Koshland, D. E. Jr. 1972. in: Methods in Enzymology, Vol. 25, pp. 468-482. 47. Carraway, K. L., Koshland, D. E. Jr. 1972. in: Methods in Enzymology, Vol. 25, pp. 616-623. 48. Phillips, J. L., Azari, D. 1971. Biochem. 10, 1160. 49. Patel, L., Kaback, H. R. 1976. Biochem. 15, 27412746. 50. Roisin, M. P., Kepes, A. 1973. Biochim. biophys. Acta 305, 249-259. 51. Peter, H. W., Ahlers, J. 1975. Arch. Biochem. Biophys. 170, 169-178. 52. Feinstein, D. L., Fisher, R. J. 1977. Biochem. J. 167, 497--499. 49

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