Eur. J. Biochem. 66, 43-47 (1976)

Activation Parameters of the Adenosine Triphosphatase of Micro co ccus lysodeik t icus A Comparison of the Soluble and Membrane-Bound Forms of the Enzyme

Juan AYALA, Manuel NIETO, Josk CARREIRA, and Emilio MUNOZ

Seccion Bioquimica de Membranas, Consejo Superior de Investigaciones Cientificas, Madrid

(Received December 15, 1975/March 19, 1976)

The Arrhenius plots for the membrane-bound ATPase and its soluble form purified from Micro- coccus lysodeikticus, presented discontinuities near 30 “C at pH 7.5. Glycerol-containing lipids were not responsible for these discontinuities. The values of the enthalpies of activation were 12 (soluble) and 22 (membrane-bound) kcal/mol (50.2 and 92.0 kJ/mol) above 30°C and 42 (soluble) and 29 (membrane-bound) kcal/mol (175.7 and 122.3 kJ/mol) below that temperature.

The results suggested that both molecular forms of the ATPase were able to adopt at least two different structures, above and below the critical temperature. Of the two, only the high- temperature structure seemed to be enzymically active. In the case of lipid-dependent ATPases, such as the Escherichia coli enzyme, the transition between both enzyme structures probably occurred with simultaneous “melting” of their lipid microenvironment.

As pointed out by Raison in a recent review [l], “the significance of abrupt changes in activation energy of some enzymic reactions and physiological pro- cesses has been the subject of considerable contro- versy”. These discontinuities or “breaks” in the Arrhenius plots can be justified thermodynamically by assuming that the system undergoes a phase change at the critical temperature [2]. In the case of membrane- bound enzymes these “breaks” have been associated with phase-changes in the lipid microenvironment sur- rounding the enzyme whose activity is being measured. This interpretation has been put forward for the mito- chondrial ATPase [ 11, sarcoplasmic reticulum ATPase [3] and Acholeplusma laidfawii Mg2 + -ATPase [4], based on solid but indirect experimental evidence. On the other hand there are conflicting reports for the ATPase from Escherichia coli [5 - 71.

In all the cases reported the enzymes appeared to be lipid-dependent and no comparison could be made between the membrane-bound form and the truly soluble lipid-free protein. Recently, our laboratory has reported a purification of the membrane ATPase of Micrococcus lysodeikticus that yields homogenous protein, that is either completely free of lipid, or if present, only in minute amounts [8,9]. This afforded a good opportunity to compare the parameters of ac-

EnzymcJs. Adenosine triphosphatase, ATP phosphohydrolase or ATPase (EC 3.6.1.3); trypsin (EC 3.4.21.4).

tivation of the soluble and the membrane-bound forms of the enzyme.

MATERIALS AND METHODS

ATPase Preparations

Soluble, active ATPase from M . lysodeikticus strain PNB, non-stimulated by trypsin, was prepared as previously described [lo, 111. Its specific activity when assayed with saturating concentrations of ATP-Ca2 +

(10 mM) at 40°C and pH 7.5 was 20 pmol . mg-’ . min-’. After subjecting a culture of M . lysodeikticus to pulses of [2-3H]glycerol for periods of time ranging from 1 to 30 generations, the isolated soluble ATPase had less than 1 mol of glycerol containing lipid per 100 mol of enzyme (Muiioz, C., unpublished ob- servations).

We used as the membrane-bound form of the enzyme, the standard membrane preparation of M . lysodeikticus strain PNB [ l l ] washed three times with Tris-HC1 buffer (30 mM, pH 7.5). It had a basal ATPase activity of 0.19 pmol/mg per min that was stimulated by trypsin (3 min preincubation at 37 “C with the protease, 1.0 mg/mg of membrane protein) to a maximum value of 0.4 pmol/mg per min when incubated at 40°C with 10mM ATP-Mg” as in- dicated below.

44 Activation Parameters of ATPase

ATPase Assay

Phosphate liberation during ATP hydrolysis was recorded continuously using the pH-stat technique as described by Nieto et al. [12]. The reaction vessel was fitted with a jacket through which water at the desired temperature % 0.05 "C was circulated from a constant temperature bath (Lauda K2RD, West Germany). A combined Radiometer Semimicroelectrode was used and adequately standardized at each temperature [ 131.

The temperature inside the reaction vessel was continuously monitored by means of a thermocouple connected to a digital thermometer YEW 2509 (Yoko- gawa Elect. Works, Ltd, Japan). The reaction mix- tures, contained in a final volume of 1.5 ml, had the following final concentrations: KCI (30 mM), ATP (10 mM) and CaC12 (soluble enzyme) or MgC12 (membrane-bound) (10 mM). The reaction was started by addition of 7-23 pg/ml of soluble enzyme or 100 pg of membrane protein/100 pg of trypsin per ml previously incubated for 3 min at 37 "C. The pH was maintained constant at a value of 7.5.

Chemicals

ATP disodium salt was purchased from PL Bio- chemicals (Milwaukee, Wisconsin, U.S.A.) and used without further purification. The trypsin was from Calbiochem (San Diego, Calif., U.S.A.). All other chemicals were analytical grade from Merck (Darm- stadt, West Germany).

RESULTS

K, of the Enzymes at 37 and 50 "C

It was desirable to work under conditions of sub- strate saturation (about 10 x K,) at all temperatures. Therefore the K, values for the membrane-bound and soluble form of the ATPase were determined from double-reciprocal plots using seven substrate (ATP- Ca2+ or ATP-Mg2+, molar ratio l / l ) concentrations ranging from 0.2 to 4 mM. The values of K, derived from the intercepts were 0.50 mM at 37 "C and 0.23 mM at 50°C in the case of the soluble form. For the membrane-bound enzyme the corresponding values of K,,, were 0.37 mM (37°C) and 0.63 mM (50°C). These relatively small variations in K, with temper- ature made us confident that a concentration of 20 mM ATP-cation was sufficient to saturate the enzyme in all cases.

Arrhenius Plots and Activation Parameters for the Different Forms of M. lysodeikticus ATPasr

The soluble form of the ATPase underwent a thermal denaturation transition with midpoint near

Table 1. Lags in the time-course of ATPase activity below the critical temperature The incubation mixtures for the assay of ATPase activity were as described in the Methods section. The reaction was initiated by addition of the enzyme, and the period of time during which there was no activity plus the period in which this activity was not linear with time (i.e. when the progress curve was concave looking from the top of the y axis) were measured. This lag period was followed by another 4-5 min of linearity in the plot of activity against time

Temperature Period during which the enzyme was not active or activity was non-linear with time

form BA membranes

"C min

20 25 30

6 4.2 2

1 0 0

49°C at pH 7.5 and ionic strength 0.03 M, in the absence of substrate. The enzyme was stabilized by the presence of substrate (i.e., under the experimental conditions of assay) and the midpoint of the transition then became 54°C (Ayala, J., and Nieto, M., un- published observations). Therefore, the specific ac- tivity of both forms of ATPase, at an ATP-cation concentration of 10 mM, was determined in the range of temperatures from 20°C to 55°C. The ATPase activity was linear with time for 2-4 min at all temperatures above 30 "C. However, below this criti- cal temperature the time-course of activity exhibited time-lags whose duration depended on the tempera- ture of the assay (Table 1). After the time-lag the progress curve was linear for about 4- 5 min. The values of specific activity obtained were represented as a function of temperature in the form of Arrhenius plots as shown in Fig. 1. Discontinuities were present in both plots but were more obvious for the soluble form of the enzyme. Both forms had higher energies of activation below the critical temperature. The value of the critical temperature, i.e. that at which the discontinuity appeared, was 30°C for both the soluble and the membrane-bound forms of the enzyme.

The difference in activation energy below and above the critical temperature was 30 kcal/mol(l25.5 kJ/mol) for the soluble form BA. Therefore, the discontinuity observed was a "real break" in the sense discussed by Belehradek [14]. For the membrane-bound enzyme the difference in energy of activation was only 7 kcal/ mol (29.3 kJ/mol).

The enthalpies and free enthalpies of activation were calculated according to the transition state theory [15,16] and from these values the entropies of activation were derived. The results are shown in Table 2 for both forms of the enzyme above and below

J. Ayala, M. Nieto, J. Carreira, and E. Mufioz 45

the critical temperature. The value of A H# and A S' changed sharply at the critical temperature, whereas d G' was sensibly constant in the interval 20- 55 "C, an example of enthalpy-entropy compensation. The values of AH' and A S f were always positive and, below critical temperature, were comparatively large.

This suggested a wide remodeling of the protein- solvent system as a consequence of (or the condition for) the formation of an active complex with ATP- cation. In the case of the membrane-bound enzyme the activation parameters were calculated assuming that the ATPase represented either 2.5% or 10% of the total membrane protein. As shown in Table 2 the two assumptions led to very small differences in the calculated values of A H' and A Sf.

Comparison with Escherichia coli Membrane-Bound A TPase

The discontinuity present in the Arrhenius plot of the membrane-bound ATPase from M. lysodeikticus occurred at the same temperature as that for the soluble active enzyme. However, since trypsin stimu- lation was necessary due to the extremely low basal activity [9], it could be argued that the membrane protein was degraded, and, therefore, was not repre- sentative of the native membrane protein. This pos- sibility was difficult to test with M. lysodeikticus, but could be examined with the E. coli K 12 ATPase which had a greater basal specific activity [17]. Arrhenius plots for both the trypsin-stimulated and basal ac- tivities of the membrane-bound form of this enzyme are shown in Fig.2. A discontinuity was present at about 19 "C in both cases and the values of the energies of activation obtained above and below 19°C were

I I I I about 3 kcal/mol(l2.5 kJ/mol) higher for the protease- activated ATPase (Table 3). -4 I

3.0 3.1 3.2 3.3 3.4 3.5 103ir (K- ' )

Fig. 1. Arrhenius plots for M. lysodeikticus ATPase. Enzyme assays were carried out as indicated in Methods and V values determined DISCUSSION under the conditions described in the text. The plots correspond to soluble ATPase ( ~ C I ) , and membrane-bound, trypsin- stimulated (A -A) enzyme. Two or three independent measure- ments of V were performed at each temperature. When a single point is plotted this is due to a coincidence of the values determined

The Origin Of the Discontinuities in the Arrhenius Plots Of Bacterial ATPuses

The sharp discontinuity present in the Arrhenius plot of the soluble ATPase occurred at the same

Table 2. Ictivation parameters for the hydrolysis of ATP-Ca2+ catalyzed by the soluble and membrane-bound forms of M. lysodeikticus ATPasr U I p H 7.5 The free enthalpy of activation A C # was calculated from V values expressed in s- ' using a value of 380000 for the molecular weight of the enzyme [ l l ] . In the case of the membrane-bound ATPase the calculations were made assuming that this enzyme represented 10% (w/w) of the total membrane protein [9] or 2.5%. The latter value was a minimal content calculated assuming equal turnover numbers for the soluble and membrane-bound forms of the enzyme. The Qlo values were determined from the experimental points using all the possible temperature intervals below and above the critical temperature. The average f S.D. is shown in the table

ATPase form t A G + A H? A S ? Qio

"C

Soluble 25 40

Membrane-bound 25 40 25 40

10 % of the membrane protein

2.5 % of the membrane protein

kcal/mol (kJ/mol) calxmoI- 'xK- ' (J x mol-' x K-')

10.2 * 2.1 15.7 (65.7) 42.3 (177.0) 89 (372.3) 15.3 (64.0) 12.0 (50.2) 11 (46.0) 1.94 & 0.18

16.6 (69.4) 29.2 (122.2) 42 (175.7) 16.2 (67.8) 22.2 (92.9) 19 (79.5) 15.8 (66.1) 29.2 (122.2) 45 (188.3) 5.2 & 0.7 15.3 (64.0) 22.2 (92.9) 22 (92.0) 3.3 f 0.6

46 Activation Parameters of ATPase

f (“C) 40 35 30 25 20 15 10

‘ t *&

-3 3.2 3.3 3.4 3.5 3.6

103/r (n-’) Fig.2. Arrhenius plots ,fbr L. coli K I2 rncnihranc~-bound ATPuse. The enzyme was assayed as indicated in Table 3 in the absence of protease (O---O) or after 3 min preincubation at 37 ‘ C with trypsin (0-0)

Table 3. Ener,+.s of’ uctivatiun for the I1yU5.olysi.s of ATP-M?’ ccrtulyxd by E. coli mcmhrane-hound ATPuse non-stirnuluted and stirnulotrd hy rrypsin E. c d i K 12 membranes prepared as described by Corao cf ul. [21] were assayed at pH 7.5 using the pH-stat as described in Methods. In a final volume of 1.5 ml the reaction mixtures contained the following final concentrations: ATP, 8 m M ; MgC12, 4 m M ; KCI, 30 mM; membrane protein, 67 pg>ml. The I-caction was initiated by addition of the membrane-bound enzyme alone or after 3 min preincubation at 37 ‘C with trypsin (ratio trypsinimembrane protein, 112, w/w) and was linear for at least 1.5 min. QIO was calculated as described in Table 2

‘C kcal/mol (kJ/mol)

N o trypsiin below 19 19.9 (S3.3) 3.63 0.51 above 19 7.0 (29.3) 1.84 0.35

Trypsin stimulated below 19 23.4 (97.9) 4.09 k 0.41 above 19 10.6 (44.4) 2.25 i 0.56

temperature as that for the membrane-bound enzyme. The absence of lipids in the soluble preparation implies that these discontinuities cannot be interpreted as a consequence of a phase-change in the lipid phase. They can only be attributed to the protein itself and at least two possibilities are open. In the first place it might occur that hydrolysis of ATP by this enzyme proceeds through more than one step after the formation of the enzyme-substrate complex or that complex formation takes place in two different types of active site. Situations of this kind have been discussed by Dixon and Webb [16], i.e. as the tempera- ture increases the rate-determining step may change and with it the activation energy of the process. A second possibility that we think more probable is that of a temperature-dependent change in the struc-

ture of the enzyme itself. Such a rearrangement in the structure of the protein-solvent system at the critical temperature is supported by the large values of A H # and A S+. The same is also suggested by the existence of time-lags in the progress curves at temper- atures below the critical. This change could be a re- arrangement of the positions of the subunits or a conformational change in one or several of them. We do not think subunit dissociation to be involved because, in the absence of substrate, it did not occur in the range from 20 to 50 ‘C (Ayala, J., and Nieto, M., unpublished observations).

The divalent cation-dependent ATPase from h4. lysodeikticus is similar in many respects to those from other microorganisms and to mitochondria1 ATPase [18 - 201. Because the claims of lipid-originated discon- tinuities in the Arrhenius plots appear to be well substantiated one could wonder why the breaks due solely to the protein, as described in this work, were not observed in addition to those due to changes in the lipid environment. It could be argued that neither the soluble forms of M. lysodeeikticus ATPase, nor the membrane-bound enzymes stimulated by trypsin were the complete enzyme present in the membrane. How- ever, the presence of a single discontinuity in the Arrhenius plot of the ATPase from E.coli for both trypsin-stimulated and basal activities, would appear to negate that possibility. Therefore, at least for this type of membrane proteins, these discontinuities in the Arrhenius plots seem to arise as a result of the transition between a low and a high-temperature protein structure. This transition probably occurs simultaneously with the “melting” of their lipid microenvironment. i.e. the phase transitions in both types of molecule, protein and lipid, occur at the same temperature. The question remains as to whether the change of phase in the protein induces the cor- responding change in the lipids; or whether it is a cooperative phenomenon involving a limited domain of the membrane structure. The indirect observations of Eleetr and Inesi [22] and Lee and Gear [23] also point to a similar conclusion.

The value found by us for the critical temperature in E. coli K 12 ATPase (19 “C) was virtually identical to that reported by Sweetman and Griffiths [ 5 ] for the same strain of the microorganism, although the values of the energy of activation were somewhat different [ 5 ] . Our values of E, were in better agree- ment with those reported by Siiieriz and Farias [7] for E. coli L 010, but their critical temperature was near 33 “C, much higher than that found in this work. Differences in bacterial strain and growth conditions might account for the difference.

This work was supported in part by grants from Eli Lilly de Espaiu, S.A. and the Fondo Nucional puru el Desarrollo de la Investigucibn Cientificu. We thank M r A. M. Garcia and Miss P. I’alacios for skilful technical assistance.

J . Ayala, M. Nieto, J . Carreira, and E. Muiioz 41

REFERENCES

1. Raison, J . K. (1973) J . Bioenerg 2. Kumamoto, J., Raison, J . K . & Lyons, J . M. (1971) J. Throrc,r.

3. Madeira, V . M . C., Antunes-Madeira, M. C. & Carvalho, A.

4. Hsung, J. C., Huang, L., Hoy, D. J. & Haug, A. (1974) Con.

5. Sweetman. A. J. & Griffiths, D. E. (1971) Bioc1ic.m. J . 121.

6. Davies, P. L. & Bragg, P. D. (1972) Biochim. Biophjs. Acto,

7. Siiieriz, F. & Farias, R . N. (1973) FEBS Lett. 32, 30-32. 8. Andreu. J . M. & Mufioz, E. (1975) Biochim. Biophj~ . Ac /u ,

9. Muiioz, E., Salton, M. R . J . , Ng, M. H. & Schor, M. T. (1969)

10. Carreira, J.. Muiioz, E.. Andreu, J. M. & Nieto, M. (1976)

11. Carreira, J . , Andreu, J . M., Nieto, M. & Mufioz, h. (1976)

Biol. 31, 47-51.

P. (1974) Biochenz. Biophy,T. Res. Comrnun. 58, 897-904.

J. Biochmni. 52, 974- 980.

1 17- 124.

266, 273-281.

387, 228-233.

Eur. J . Bioclzeni. 7, 490 - 501.

B i ~ ~ h i ~ l . B i ~ p l l ~ , ~ . Actcl, 436, 183- 189.

Mid. Cell. Bioclzem. 10, 67- 76.

12. Nieto, M., Muiioz, E., Carreira, J . & Andreu, J. M. (1975) Biochim. Biophys. Actcr. 413, 394 - 41 4.

13. Bates, R . J . (1973) in Determinotion o f p H . Throri, ond Proc- rice, p. 73, John Wiley & Sons Inc., New York.

14. Belehrlidek, J . (1957) Annu. Rr i ' . Phjsiol. 10. 59 -X2. 15. Johnson, F. H., Eyring, H . & Polissar, M. J . (1954) Thc Kirietic,

Basis ofMoleculur Biology, p. 187, John Wiley & Sons Inc., New Y ork.

16. Dixon, M. & Webb, h. C. (1971) in Enzymc.~, 2nd edn, p . 150, Longmans, London.

17. Carrcira, J. & Muiioz, E. (1975) Mol. Cell. Biocken2. 0, 85-95. 18. Andreu, J. M. , Albendea, J . A. & Mufioz, E. (1973) Ew. J .

19. I'edersen, P. L. (1975) J . Bioenerget 20. Abrams, A. & Smith, J . B. (1974) in T/7e Enz.vmes (Boyer.

P. D.. ed.) vol. 10, pp. 395-429, Academic Press, New York.

21. Corao, M., Serrano, J . A., Leal, J . A, , Puig, J . & Muiioz, E. (1974) Microhiol. E.sp. 27, 283-298.

22. Eleetr, S. & Inesi, G. (1972) Biochim. Biupl?j.s. Acto. 200,

23. Lec, M. P. & Gear, A. R. L. (1974) J . Biol. Clwm. 240,

Biocheni. 37, 505-515.

178 - 185.

7541 -7549.

J . Ayala. M . Nieto. J. C'arreira, and E. Mufioz. SecciOn Bioquirnica de Mcmhranas. Ccntro dc Invcstigaciones Biolbgicas, C.S.I.C., VclLzque7 144, Madrid-6, Spain