Phosphorylation- and ligand-induced conformational changes of rat liver fructose-1,6-bisphosphatase

ARCHIVES

OF BIOCHEMISTRY

Vol. 248, No. 2, August

AND

1,

BIOPHYSICS

pp. 604-611,1986

Phosphorylationand Ligand-Induced Conformational of Rat Liver Fructose-l ,6-bisphosphatase H. VIDAL,l INSERM

B. ROUX,*

U197, FacultC de M&&cine Alexis Carrel, de Physicc-chimie biologique, Universiti Received

November

6,1985,

AND

rue G. Paradin, Claude Bernard, and in revised

Changes

J. P. RIOU F-69372 F-69622 form

March

Lyon Ckdex Villeurbanne,

08, and Yaborata’re France

24,1986

The effects of cyclic AMP-dependent phosphorylation on the structural properties of rat liver fructose-1,6-bisphosphatase were investigated by uv difference spectroscopy and circular dichroism. The incorporation of 4 mol of phosphate per mole of fructose1,6-bisphosphatase induces a significant increase in the a-helix content of the enzyme without affecting its spectrophotometric properties. The addition of fructose 1,6-bisphosphate or fructose 2,6-bisphosphate also affects the conformation of the enzyme. However, both the phosphorylated and the nonphosphorylated forms exhibit similar ligand-induced conformational changes. These results show that cyclic AMP-dependent phosphorylation of fructose-1,6-bisphosphatase induces a specific conformational change. They also suggest that this modification does not alter the interaction of the enzyme protein with fructose 1,6-bisphosphate and fructose 2,6-bisphosphate. o 1986 Academic Press, Inc.

PZ) and for AMP (3). On the other hand, Rittenhouse et al. (9) have reported the absence of phosphorylation-induced modifications, while Meek and Nimmo (10) as well as Ekman and Dahlquist-Edberg (11) have described a twofold increase of the apparent Km of the enzyme for the substrate. Recently, Ekdahl and Ekman (12) have reported that nonphosphorylated Fru-1,6P,ase is more readily inhibited by both AMP and fructose 2,6-bisphosphate (Fru 2,6-Pz) than the phosphorylated enzyme. Since these reports had not allowed a precise description of the effects of cyclic AMP-dependent phosphorylation on Fru1,6-Ppase catalytic activity, we investigated the effect of phosphorylation on the conformation of the enzyme protein by circular dichroism and uv difference spectroscopy studies. The experiments were designed to study (i) specific effects of phosphate incorporation in the enzyme protein and (ii) interactions of both forms of Fru-1,6-P2ase with its metabolic effectors.

Rat liver fructose-1,6-bisphosphatase (Fru-1,6-P,aseF EC 3.1.3.11), a key gluconeogenic enzyme (1,2), has been shown to be a substrate for the cyclic AMP-dependent protein kinase (3-5). Phosphorylation occurs both in vitro (3-5) and in vivo (3, 6) and is stimulated by glucagon in isolated rat liver cells (7). These findings support the hypothesis that cyclic AMP-dependent phosphorylation is involved in the regulation of Fru 1,6-P,ase activity. Nevertheless, reports showing that glucagon affects the enzyme’s properties are scarce (8) and there has been some debate concerning the effects of in vitro phosphorylation on the enzyme activity. We have previously reported that phosphorylation increases enzyme activity without modifying the affinity for fructose l,&bisphosphate (Fru 1,61 To whom correspondence should be addressed. ’ Abbreviations used: Fru-1,6-Passe, fructose-1,6hisphosphatase; Fru 1,6-Pa, fructose 1,6-bisphosphate; Fru 2,6-Pa, fructose 2,6-hisphosphate; SDS, sodium dodecyl sulfate. 0003-9861186 Copyright All rights

$3.00

0 1986 by Academic Press, Inc. of reproduction in any form reserved.

604

CONFORMATIONAL

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The data presented herein suggest that cyclic AMP-dependent phosphorylation of rat liver Fru-1,6-PPase induces significant modifications of the enzyme protein’s secondary structure. MATERIALS

AND

METHODS

Materials All chemicals were of reagent grade and were purchased from Merck or Sigma. All enzymes and coenzymes were produced from Boehringer (Meylan, France), [y-32P]ATP was from Amersham (London), and DEAE-trisacryl and hydroxylapatite were from IBF (France). Fru 1,6-PZ was freed of possible contamination by Fru 2,6-P* by a 60-min acid treatment (13). Fru 2,6PZ (Sigma) concentration was determined by conversion to Pi plus fructose 6-P by acid treatment and enzymatic determination of fructose 6-P (14). No Fru 1,6-PZ could be detected in the Fru 2,6-Pa solution.

Methods Purijkation procedures and assay of enzyme activities. Rat liver Fru-1,6-Pzase was purified from male Sprague-Dawley fed rats (IFFA-CREDO, Lyon) as previously described (3). The catalytic subunit of rat liver cyclic AMP-dependent protein kinase was prepared by the method of Sugden et al. (15) except for the replacement of DEAE-cellulose by DEAE-trisacryl and omission of the final gel filtration step. Each purified enzyme protein displayed a single band on polyacrylamide microslab gel electrophoresis in the presence of sodium dodecyl sulfate (SDS) (16). The molecular weight of the Fru-1,6-Pzase subunits was estimated to be 41,000 +_ 1000 (Z f SD, n = 9). Fru-1,6-P,ase was assayed by the spectrophotometric method of Pontremoli et al. (17) as previously described (3). One unit of Fru-1,6-Passe is defined as the amount of enzyme which catalyzes the hydrolysis of 1 pmol of Fru 1,6-P* per minute at 30°C. Purified Fru-1,6-P,ase had a specific activity of 29 + 2 (3c + SD, n = 7) units per milligram protein. The ratio of activity at pH 7.4 to activity at pH 9.4 was always greater than 4. Protein kinase activity was determined using histone as substrate by the method of Corbin and Reimann [18]. The filter paper used in this method was washed according to the procedure of Gill and Walton (19). One unit of protein kinase is the amount of enzyme which catalyzes the transfer of 1 pmol of Pi from [y-32P]ATP to histone per minute at 30°C. Protein concentration was determined by the method of Lowry et al (20) with bovine serum albumin as standard.

LIVER

FRUCTOSE-l,&BISPHOSPHATASE

605

Phmphmylatim of Fru-1,6-Pwe. Fru-1,6-Pzase (23 mg * ml-‘) was incubated at 35°C in a final volume of 0.2 ml that contained 35 mM potassium phosphate, pH 6.8,6 mM MgClz, 0.2 mM dithiothreitol, 0.2 mM [y‘*P]ATP (20-100 cpm . pmoll’), and 20-50 pg. ml-’ of the catalytic subunit of the cyclic AMP-dependent protein kinase (sp act 1 to 2 X 10s units * mg protein-‘). Labeled phosphate incorporation into the enzyme substrate was estimated by the method of Corbin and Reimann (18). When analyzed on polyacrylamide slab gel electrophoresis in the presence of SDS, “Pi was detected in a single band migrating with an apparent molecular weight of 41,000. The number of moles of Pi incorporated per mole of Fru-1,6-P*ase was calculated on the basis of a molecular weight of 164,000 for the rat liver enzyme. The purified Fru-1,6-Pzase was considered to be nonphosphorylated because about 4 mol of Pi per mole of enzyme could be incorporated routinely and because Pilkis et al (21) have reported that rat liver Fru-1,6-P*ase, purified by the same methodology, contained 0.2 to 0.3 mol of Pi per mole of Fru-1,6-Pzase. When a stock solution (lo-15 mg) of phosphorylated rat liver enzyme was prepared, a 4-ml vol was used and the reaction went on until 3.5 to 4.5 mol of Pi per mole of enzyme was incorporated. The phosphorylated Fru-1,6-Pzase was then separated from [y-32P]ATP and from the catalytic subunit of the cyclic AMP-dependent protein kinase by ammonium sulfate precipitation and gel filtration on superfine Sephadex G-100 equilibrated in 50 mM, Tris-HCl, pH 7.4, containing 1 mM dithiothreitol and 0.1 mM EDTA. The labeled enzyme, stored at -20°C in this buffer, was stable for at least 1 month. Ultraviolet differace spectroscopy. Ultraviolet difference spectra were recorded on a Cary Model 210 spectrophotometer. To study the interaction of the enzyme with substrates or effecters, the tandem cell method of Herskowitz (22) was used with a pair of matched rectangular tandem cells (2 X 0.4-cm path length). Difference spectra were recorded at 30°C from 240 to 320 nm on a full scale of 0.02 absorbance unit. Fru-1,6-Pzase (0.2-0.5 mg. ml-‘) was diluted in 50 mM Tris-HCl, pH 7.4, containing 0.1 mM EDTA and 1 mM dithiothreitol. Aliquots (l-10 ~1) of the desired ligand were added sequentially with Hamilton microsyringes as described by O’Brien and Kapoor (23). Circular dichroiam CD spectra were recorded at room temperature on a Jobin-Yvon Mark IV dichrograph using a cylindric cell of 0.02-cm path length between 200 and 250 nm at a rate of 6 nm 9 mini. Protein solutions were diluted to l-2 mg * ml-’ in the storage buffer and cleared by filtration on a 0.25~pm filter (Acrodisk, Gelman) before use. Molar ellipticities were calculated from the mean of at least four spectra recorded on one sample and each experiment was repeated with three different enzyme preparations in order to ensure that the findings were reproducible. Ellipticities were calculated

606

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ROUX,

as previously described (24) on the basis of a molecular weight of 164,000 for the rat liver enzyme. Contributions of the various secondary structures (a-helix, P-sheet, and random coil) to the overall conformation of the enzyme were estimated from the molar ellipticities according to the model of Chen et al (25). Each spectrum was analyzed by using a nonconstrained multilinear least-squares computer program; i.e., the sum of the amounts of secondary structure can equal any value in order to obtain the best fit of the Chen model. Modifications of the molar ellipticity of Fru-1,6Pzase during phosphorylation were studied by recording kinetically the signal at 222 nm using an expanded scale. ATP (0.2 mM), MgClz (6 mM), and the catalytic subunit of the cyclic AMP-dependent protein kinase were mixed with the enzyme protein and introduced together in the cell before the recording was started. To avoid a possible contribution of the catalytic subunit to the recorded spectrum, the amount of catalytic subunit protein never exceeded 10% of the amount of Fru-1,6-Passe. % incorporation in Fru-1,6Pzase was measured in parallel as described above. RESULTS

Phosphorylation-induced conformational changes of rat liver Fru-1,6-Pzase were first studied by circular dichroism. Figure 1 shows the CD spectra of the phosphorylated and nonphosphorylated enzyme. A highly significant difference was observed in the region of 215-225 nm with a 30% decrease in the molar ellipticity at 222 nm. Contributions of the various secondary structures to the overall conformation were calculated using the model of Chen et al. (25). This model allowed the 30% molar ellipticity decrease to be attributed to an increase in the a-helix content of 30 to 36% (Table I). The data concerning ,f3sheet and random coil contents were more scattered because of the relative imprecision of measurement below 210 nm. Therefore, no conclusion concerning the effect of phosphorylation on the P-sheet and random coil contents could be drawn. During spectra recordings (about 60 min) the enzyme activity and the pH 7.4/pH 9.4 activity ratio were stable (data not shown). To obtain more insight into the relationship between the stoichiometry of phosphorylation and the a-helix content variations, these parameters were recorded during the phosphorylation reaction. Fig-

AND

RIOU

I 1 200

I

1 220 WAVELENGTH

240 (nm)

FIG. 1. Circular dichroism spectra of nonphosphorylated (0) and phosphorylated (0) rat liver Fru-1,6P,ase. Each point represents the mean f SD of three different Fru-1,6-Pzase preparations. Conditions were as described under Materials and Methods. The theoretical curve (-) was calculated according to the model of Chen et al. (25).

ure 2 shows that when rat liver Fru-1,6Pzase was incubated with ATP-Mg and the catalytic subunit of the cyclic AMP-dependent protein kinase, a time-related change in the molar ellipticity was observed at 222 nm. When 32P incorporation was measured in parallel, it was found that the molar ellipticity was affected only after the incorporation of 4 mol of Pi per mole of Fru-1,6Pzase. The relatively slow rate of the reaction was related to the low temperature at which the recording was performed (ZO’C) and to the small amount of catalytic subunit added. These kinetic experiments were also performed with regard to the concentrations of the catalytic subunit. Figure 3 shows that the more catalytic subunit that was added, the earlier the changes occurred. In every case, the decrease in the molar ellipticity proceeds at the same rate as shown by the parallelism of the sigmoidal curves. The signal was unaffected by addition of the catalytic subunit

CONFORMATIONAL

CHANGES

OF

RAT

LIVER

TABLE INFLUENCEOFEFFECTORSONTHEPERCENTAGEOFSECONDARY AND PHOSPHORYLATED

I STRUC~UREOFNONPHOSPHORYLATED(NP) (P) Fru-1,6-Passe” % of secondary

o-Helix

0

Fructose

P

30 f 0.6

36 k 0.8

29 zk 0.5 28 + 0.6

35 f 0.6 36 + 0.6

24 + 0.6

27 f 0.4

NP 8f3

Random P 15 f

coil

NP

Sum

P

NP

P 100

3

58 f 3

49f3

96

13 2 2 14 f 2

13 + 2

47 f 3

23f4

89

12 f

2

57 f 3

5724

99

105

12 k 2

18 * 2

45 f 4

52f3

81

97

1,6-bisphosphate

5OpM 250

structure

P-Sheet

NP

Effector

607

FRUCTOSE-1,6-BISPHOSPHATASE

@I

Fructose

71

2,&bisphosphate

25~~

a Estimated by circular dichroism according to the model of Chen et al (25) with a nonconstrained multilinear least-squares program. Conditions were as described under Materials and Methods. The percentages of secondary structure are expressed as values f standard deviation.

alone or by addition of the phosphorylation mixture without the catalytic subunit or without ATP. During the incubation pe-

riod, enzyme activity was stable and the apparent molecular weight of the Fru-1,6Pease subunit, estimated by SDS-slab gel electrophoresis, remained constant (data not shown). These data show that cyclic AMP-dependent phosphorylation of rat

-9x)

I -9.0

I 200

100

TIME

300 (min.)

100

200 TIME

FIG. 2. Time course of the phosphorylation-induced incorporation of “Pi in Fru-l,&Paase (0) and changes in the molar ellipticity (0) (recorded at 222 nm). Fru1,6-Passe (1.5 mgeml-‘) was incubated as described under Materials and Methods with ATP-Mg and 30 pgeml-’ of the catalytic subunit of cyclic AMP-dependent protein kinase in either a cylindrical cell for molar ellipticity measurement or in a test tube with [y-SZPJ4TP-Mg for “Pi incorporation measurement.

300

(min.)

FIG. 3. Phosphorylation-induced changes of Fru-l,BPasse molar ellipticity (recorded at 222 nm) as a function of the amount of catalytic subunit of the cyclic AMP-dependent protein kinase. Nonphosphorylated Fru-l,&P,ase (1.5 mg . ml-‘) was incubated in the cylindrical cell as indicated in the text with various amounts of the catalytic subunit. +, No catalytic subunit; 0, 30 pg. ml-‘; A, 75 pg - ml-‘; 0, 150 fig - ml-‘.

608

VIDAL,

ROUX,

AND

liver Fru-l,&Pzase induces a decrease of the molar ellipticity which can be attributed mainly to an increase in the a-helix content. CD was also used to study the interaction of phosphorylated and nonphosphorylated Fru-1,6-Pzase with Fru 1,6-Pz and Fru 2,6Pz. Table I shows that Fru 1,6-P2 did not affect the calculated percentage of a-helix even at high concentrations. In contrast, Fru 2,6-P2 produces a significant decrease in the percentage of a-helix. Similar changes were observed with both forms of the enzyme: thus, Fru 2,6-Pz produced a decrease in the a-helix content but did not alter phosphorylation-induced differences in the a-helix content. Perturbations of the Fru-1,6-Pzase aromatic residue environment were studied by uv difference spectroscopy. No difference spectrum was observed between the phosphorylated and nonphosphorylated forms of the enzyme. The addition of either Fru 1,6-Pz or Fru 2,6-Pz (Fig. 4) induced a difference spectrum with maximum absorbance peaks at 294 and 250 nm with no discernible wavelength shift of the absorption bands. These absorbance changes were related to ligand concentration and were saturable. When molar absorption difference at 294 nm was plotted as a function

320

300

280 h

FIG. 4. The uv difference spectra recorded on the 0.02 absorbance nm*s-‘) either with (A) Fru 1,6-P* pM (f); or with (B) Fru 2,6-Pa at 1

280 (nm)

RIOU

of effector concentration (Fig. 5), a sigmoidal curve was observed with half-maximal changes occurring at 165 PM Fru 1,6PZ and at 10 PM Fru 2,6-Pz. Similar results were observed with both the nonphosphorylated and the phosphorylated forms of the enzyme. DISCUSSION

The present report shows that phosphorylated rat liver Fru-1,6-Pzase exhibits a secondary structure different from that of the nonphosphorylated enzyme as shown by the increase in the a-helix content as estimated by circular dichroism according to the model of Chen (25). This conformational change can be attributed to the specific phosphorylation of the enzyme by the catalytic subunit of the cyclic AMP-dependent protein kinase since, first, the change occurred in a kinetic manner when 4 mol of phosphate per mole of Fru-1,6-Pzase was incorporated; second, a relationship was shown between the amount of catalytic subunit added and the time at which the change occurred; and third, there was no change when the catalytic subunit or ATP was omitted in the phosphorylated medium. Nevertheless, Fru-1,6-P2ase is known to be very sensitive to proteolysis, and the

240

320

300

280

260

240

A (nm)

of nonphosphorylated Fru-1,6-Passe (0.4 mg. ml-‘). Spectra were scale (period 10 s, spectral bandwidth 1 nm, and scan rate 0.2 at 60 p&! (a), 80 pM (b), 100 JLM (c), 150 @d (d), 245 pM (e), and 340 PM (a), 2 pM (b), 6 FM (c), 10 PM (d), 15 pM (e), 20 pM (f), 36 pM (g).

CONFORMATIONAL

CHANGES

loo FRU. l-6

OF

200 P2

RAT

300 (lchl)

FIG. 5. Changes in the molar extinction coefficient for the nonphosphorylated (0) and phosphorylated

cyclic AMP-dependent phosphorylation of the enzyme might have sensitized the enzyme to proteolytic cleavage (26). Therefore, the observed change could have been related to proteolysis. Such an event in our experiments is unprobable since the pH 7.4/pH 9.4 enzyme activity ratio was stable during the phosphorylation reaction and since at the end of the experiment 32Pwas found to comigrate with nonphosphorylated purified Fru-1,6-Pzase on SDS-slab gel electrophoresis with an apparent subunit molecular weight of 41,000. Therefore, the observed changes in the a-helix content of Fru-1,6-Pzase can be related to the incorporation of phosphate. In addition, these changes proceed at a constant, relatively slow rate (60-90 min) and occurred only after complete phosphorylation of the four Fru-1,6-Pzase subunits. Thus, no relationship could be determined between the rate of the incorporation of phosphate and the increase in the a-helix content. These findings suggest that rat liver Fru-1,6-Pzase exhibits at least two different stable conformations corresponding to the nonphosphorylated and the phosphorylated forms of the enzyme. To our knowledge, this is the first report of an enzyme protein undergoing conformational change after cyclic AMP-dependent phos-

LIVER

400

609

FRUCTOSE-1,6-BISPHOSPHATASE

5 FRU.Z-6

15

10 PZ

at 294 nm as a function (A) Fru-1,6-P,ase.

20

(liiw

of ligand

concentration

phorylation. This result adds new insight concerning the effect of phosphorylation on the secondary structure of enzyme proteins, but the significance of the increased a-helix content is presently unknown. If this change is involved in the biochemical properties of Fru-1,6-Pzase, the two forms of the enzyme might be expected to react differently with specific ligands. Such an event was not observed. Either by CD or uv difference spectroscopy analysis, each form of the enzyme exhibited the same characteristic spectra after addition of Fru 1,6-Pz or Fru 2,6-Pz. Therefore, it appears that phosphorylation modifies the secondary structure of rat liver Fru-1,6-Pzase but that this conformational change does not affect the interactions with ligands. Studies of ligand interactions with Fru1,6-Pzase using the same optical methods were also of interest. Fru 1,6-Pz, the substrate of the enzyme, did not affect the secondary structure of Fru-1,6-P2ase as shown by CD, but induced a characteristic uv difference spectrum. This spectral effect is principally due to perturbations in the environment of the tyrosine residues since Fru-1,6-P2ase does not contain tryptophan (27). The sensitivity of these changes to the addition of ligand was similar to that reported by McGrane et al. (28). The high

610

VIDAL,

ROUX,

concentration of Fru 1,6-Pz required to elicit half-maximal molar absorption change at 294 nm, compared to the apparent Km of the enzyme, could be related to the postulated existence of an inhibitory binding site for the substrate (28) or more probably to the lack of Mgzf ions in the medium. Me ions are indeed needed for catalytic activity but cannot be used in these experiments since Fru 1,6-P2 is rapidly hydrolyzed in its presence. The effect of Fru 2,6-Pz was somewhat different from that of Fru 1,6-PZ. When Fru 2,6-P2 was studied by uv difference spectroscopy, similar perturbations at 293 and 250 nm were observed, suggesting, as previously shown (28, 29), that Fru 2,6-P2 interacted with the active site of the enzyme, but with a much higher affinity than Fru 1,6-P2. In addition, Fru 2,6-Pz produced a significant decrease in the a-helix content of the enzyme as shown by CD. These observations suggest that in addition to its ability to bind to the active site Fru 2,6-Pz induces an overall conformational change. This hypothesis is supported by the findings that Fru 2,6-Pz on the one hand is a competitive inhibitor of the enzyme and on the other hand potentiates AMP-mediated inhibition of Fru-1,6-Pzase activity (29). Moreover, recent NMR studies have shown that Fru 2,6-Pz interacts with the active site of bovine liver Fru-1,6-Passe and induces structural modifications of the enzyme (30). In conclusion, the combined circular dichroism and uv difference spectroscopy data presented in this work demonstrate that cyclic AMP-dependent phosphorylation of rat liver Fru-1,6-Pzase is associated with a significant modification of its secondary structure but that this structural change does not modify the interactions with specific ligands. More studies are needed to determine whether such modifications are involved in regulatory process, requiring definite secondary structures such as protein-protein interactions (31). ACKNOWLEDGMENTS We are indebted to M. Odeon and S. Terfous their invaluable technical assistance and typing

for ex-

AND

RIOU

pertise, respectively. The discerning Rousset and C. Dumontet in correcting is deeply appreciated.

advice of B. the manuscript

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B&him Biophys. Acta 662,265-270. 12. EKDAHL, K. N., AND EKMAN, P. (1985) J. Bid Chem 260,14173-14179. 13. PILKIS, S. J., EL-MAGHRABI, M. R., GUMMING, D. A., PILKIS,

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Methods in Enzymology (Wood, W. A., ed.), Vol. 89, pp. 101-107, Academic Press, New York. 14. LANG, G., AND MICHAL, Enzymatic Analysis Vol. 3, pp. 1238-1242, Beach, Fla. 15. SUGDEN, P. H., HOLLADAY, AND

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422. 16. WEBER, K., AND OSBORN, M. (1975) in The Proteins (Neurath, H., and Hill, R. L., eds.), 3rd ed., Vol. 1, pp. 180-224, Academic Press, New York. 17. PONTREMOLI, S., TRANIELLO, S., LUPPIS, B., AND WOOD, W. A. (1965) J. Biol Chem 240, 34593463. 18. CORBIN, J. D., AND REIMANN, E. M. (1974) in Methods in Enzymology (Hardman, J. C., and O’Malley, B. W., eds.), Vol. 38, pp. 287-289, Academic

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19. GILL, G. N., AND WALTON, G. M. (1979) in Advances in Cyclic Nucleotide Research (Brooker, G., Greengard, P., and Robison, G., eds.), Vol. 10, pp. 93-106, Raven Press, New York. 20. LOWRY, 0. H., ROSEBROUGH, N. .I., FARR, A. L., AND RANDALL, J. J. (1951) J. Biol Chum. 193, 265-271. 21. PILKIS, S. J., EL-MAGHRABI, M. R., COVEN, B., CLAUS, T. H., TAGER, H. S., STEINER, D. F., KEIM, P. J., AND HENRIKSON, R. L. (1980) J. Bid Chem 255,2770-2775. 22. HERSKOWITZ, T. T. (1967) in Methods in Enzymology (Hirs, C. H. W., ed.), Vol. 11, pp. ‘748775, Academic Press, New York. 23. O’BRIEN, M. D., AND KAPOOR, M. (1975) FEBSLett. 58,366-368. 24. Roux, B., FELLOUS, G., AND GODINOT, C. (1984) Biochemistry 23,534-537.

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25. CHEN, Y. H., YANG, J. T., AND MARTINEZ, H. M. (1972) Biochemistry 11,4120-4131. 26. BERGSTROM, G., EKMAN, P., HUMBLE, E., AND ENGSTROM, L. (1978) B&him. Biophys. Acta 532,259-267. 27. BENKOVIC, S. J., AND DE MAINE, M. M. (1982) in Advances in Enzymology (Meister, A., ed.), Vol. 53, pp. 45-82, Wiley, New York. 28. MCGRANE, M. M., EL-MAGHRABI, M. R., AND PILKIS, S. J. (1983) J. BioL Ckm. 258,10445-10454. 29. PILKIS, S. J., EL-MAGHRABI, M. R., PILKIS, J., AND CLAUS, T. H. (1981) J. Biol. Chem. 256, 36193622. 30. GANSON, N. J., AND FROMM, H. J. (1985) J. Biol. Chem. 260,2837-2843. 31. MCGREGOR, J. S., SINGH, V. N., DAVOUST, S., MELLONI, E., PONTREMOLI, S., AND HORECKER, B. L. (1980) Proc. Nat1 Ad Sci. USA 77,3889-3892.