Purification and properties of fructose 1,6-diphosphatase from Polysphondylium pallidum

ARCHIVES

OF

BIOCHEMISTRY

Puri~cation

and

AND

BIOPHYSICS

Properties

114, 31-37 (1966)

of Fructose

Polysphondyfiom

1 ,&Diphosphatase

from

pallidurn

0. M. ROSEN2

Received December 30, 1965 Fructose 1,6-diphosphatase (FDPase) has been purified 400-fold from the cellular slime mold Polysphondylium pallidz~m. As isolated, the enzyme specifically hydrolyzes the C-l phosphate group of fructose 1,6-diP and sedoheptulose l,?-diP with the production of stoichiometric amounts of fructose 6-P and sedoheptulose 7-P, respectively. The enzyme has an alkaline pH optimum {9X+9.5), requires Mgft or Mn++, and is active with low concentrations of substrate. It exhibits no activity at pH 7.5 unless EDTA is added. In the presence of 0.1 mM EDTA, the activity at pH 7.5 becomes equal to that at pH 9.5. Cysteine, diethyldithiooarbamic acid, and Z-mercaptoethanol were also effective in inducing enzymic activity at pH 7.5, but much higher concentrations were required than with EDTA. The enzyme was not significantly inhibited by AMP, dAMP, or FDP. There was no alteration in the level of Fl)Pase or phosphofructokinase activity during the course of morphologic differentiation.

A specific fructose 1,6-diphosphatase (FDPase)3 that catalyzes the reaction: D-fructose 1 ,6-diP + HZ0 -+ n-fructose 6-P + Pi was originally described by Gomori in 1943 (1) , FDPases, subsequently isolated from a variet’y of sources, have many properties in common (2-9). They have a narrow

range of specificity, require Mg+f or Mn+f, function optimally at alkaline pH, are stimulated by EDTA, and are active with low concentrations of substrate. In addition, they are generally inhibited by AMP, high concentrations of FDP (lo), or both. There is evidence to suggest that FDPase may be involved in the regulation of carbohydrate metabolism and that the relative rates of glu~oneogenes~ and glycolysis are to some degree determined by the net effect of the antagonistic activities of FDPase and phosphofructokinase (PFK) (11-14). The cellular slime molds exist as independent myxamebae for part of their life cycle (15). In the absence of nutrients, on a solid substrate, the myxamebae aggregate and undergo differentiation into a multicellular organism consisting of two cell types: stalk and spore. The process of differentiation is highlighted by tCheeonversion of a ceil undergoing glyeolysis and synth~izing protein into a multicellular organism that synthesizes carbohydrate at the expense of endogenous reserves. During this process, the

1 Supported by grants from the National Institutes of Health (GM 11301) and the Nat#ional Science Foundat*ion (GB-4313). Communication No. 57 from the Joan and Lester Avnet Institute of Molecular Biology. 2 Postdoctoral Fellow of the United Cerebral Palsy Foundation. 3 Abbreviat,ions used: FDPase, fructose 1,6-diphosphatase; SDPase, sedoheptulose 1,7-diphosphatase; SDP, sedoheptulose 1,7-diphosphate; S7P, sedoheptulose 7-phosphate; FDP, fructose 1,6-diphosphate; F6P, fructose 6-phosphate; FlP, fructose 1-phosphale; PFK, phosphofructokinase; TEA, triethanolamine; DDTC, diethyldithiocarbamic acid; Pi, inorganic phosphate; DHAP, dihydroxyacet~one phosphate; CM Sephadex, carboxymethyl Sephadex; CMC, carboxymethyl cellulose; G6P’, glucose 6-phosphate; GlP, glucose l-phosphate. 31

32

ROSEN

content of protein, RNA, and the dry weight per cell decrease, but the carbohydrate content increases and appears to be positively correlated with differentiation (16). In view of the postulated role of FDPase in gluconeogenesis we undertook a study of the properties of this enzyme in the cellular slime mold, Po~~~~h~nd~~~u??2 ~a.l~i~u~n. The present commu~lication report,x the parGal purification of FDPase from the organism and a description of some of its properties, during the process of including its activity differentiation. ~~AT~R~ALS

AND

~fETHODS

Materials. All chemicals were obtained from commercial sources unless otherwise stated. FDP (98y0 pure) and TPN were obtained as t’he sodium salts from Sigma Chemical Co., St. Louis. The enzymes employed in the assays were obtained as crystalline suspensions from Boehringer and Sons, ~~aIlnheirn, Germany. When necessary, barium or calcium salts of organic phosphate esters were converted to the corresponding sodium salts by passage through Dowex-50 in t,he sodium form. SDP was prepared by a modification of the method of Smyrniotis and Horecker (17,18). Transaldolase and DHAP were generously provided by Mr. 0. Tchola and Mr. D. Morse, respectively, Department of Molecular Biology, Albert Einstein College of Medicine. Protamine sulfate was purchased from Eli Lilly and Co., Indianapolis; CM Sephadex (C-50, coarse) from Pharmacia Chemicals, Uppsala, Sweden. Myxamebae of Po~ysp~o~dyl~~~?n. p~~l~~~rn~ were grown axenieally in liquid medium (19). Organisms in the various stages of differentiation were obtained by plating the myxamebae on washed agar plates. For enzyme purification, the myxamebae were harvested by centrifugation, washed, and stored frozen at -20”. ~~e~~ods. Protein was determined by the met,hod of Lowry (20). FFP (Z), FDP (21), FDP aldolase (22), PFK (23), and SDP (24) were assayed Inorganic phosphate spectrophotometrically. (Pi) was measured by the method of Fiske and SubbaRow (25). S7P was assayed spectrophotometrically by measuring the F6P formed when D-glyceraldehyde 3-P was added to the S7P in the presence of purified transaldolase. The reaction mixture (1 ml) contained: tris buffer, pH 7.5 (0.05 M); phosphoglucose isomerase (5 pg); glucose 6-phosphate * Culture generously supplied by Dr. M. Sussman, Brandeis University, Waltham, Massachusetts.

dehydrogenase (1 rg); transaldolase (1 pg); DHAP (0.65 mfif); triose phosphat,e isomerase (10 pg); and TPN (0.5 m&f). The increase in absorbance at 340 rng was measured. All spe~trophotometric measurements were made in a Gilford recording spectrophotometer at, room temperat,ure. De$nition of enzyme unit. Unless otherwise stated, the studies reported in this communication refer to t,he “FDPase” and not to the “SDPase” capacit,y of t,he enzyme preparation (see Resulls: St80ichiometry and substrate specificity). The unit of enzyme was defined as the amolmt required to catalyze the conversion of 1.0 rmole of FDP to F6P in 1 minute at room temperature. Specific activity was expressed as units per milligram protein. Unless ot,herw-ise stat,ed, the FDPase used in these studies had a specific activity of 5 rmoles per minute per miliigram protein and was assayed at pH 9.5 by using the spectrophot,ometric assay. Enzyme assay I (“Xpectrophotomefric Assay”). at FDPase was assayed spectrophotometrically 340 rnp by measuring the rate of reduction of TPN. The incuba,tion mixt,ure (1.0 ml) contained 0.02 1% glycine buffer, pH 9.5; 0.5 mM TPN; 0.2 m&f FDP; 1.0 rn*%’ MgC12; glucose 6-P dehydrogenase (5.0 pg); phosphohexose isomerase (5.0 pg) ; and an appropriate aliquot of FDPase. The final pH measured in the euvette was 9.3. When the reaction was performed at pH 7.5, triethanolamine (TEA) buffer (0.02 iv) was used in place of glycine buffer. Where indicated, EDTA (0.5 mM) was added. The optical density at 340 rnp was recorded between 6 and 10 minutes. II (“Inorganic Phosphate l+nzyme assay -
Purification of the enzyme. All operations were performed at 4” unless otherwise stated. Centrifugation was carried out at 10,000 rpm in the SS No. 34 rotor of a refrigerated Sorvall centrifuge (RC-2). Table I gives a summary of the purification procedure.

FRUCTOSE

PURIFICATION

33

TABLE I OF FDPASE FROM P~~~~pho~d~~~u~ pal~~du~ Total

FW2iOD

1. Cell-free extract 2. Protamine sulfate fraction 3. Ammonium sulfate fraction saturation) 4. Sephadex G-25 eluate 5. CM Sephadex fraction=

FROM P. PALLIDUM

1,6-DIPHOSPHATASE

(ml)

vol.

T”~~~~~

Sp. act. (units/mg)

Over-all recovery (%I

14.8 18.5 20.0

2.68 2.68 2.17

0.013 0.025 0.045

106 81

40.0 10.0

2.14 0.66

0.045 5.50

80 25

(40-60~o

a If CMC is substituted for CM Sephadex, enzyme with a specific activity of 10.6-12.0 units/mg may be obtained. The over-all recovery, however, is only one half of that found when CM Sephadex is employed. TABLE

11

STOICKIOMETRY OF THE REACZIONS CATALYZED BY FDPAsE” Reaction component

FDP F6P

hitial cmc. @moles/ml)

Final cont. bmles/ml)

Nc;yf

Pi

0.710 0.0 0.220

0.395 0.330 0.580

-0.315 -to.330 +0.360

SDP S7P Pi

0.595 0.0 0.180

0.070 0.415 0.630

-0.525 +0.415 $0.450

0 The incubation mixture (1.0 ml) contained 1.0 mM MgC12; 0.62 M glycine buffer, pH 9.5; 0.1 m&f EDTA, purified FDPase (1 pg) and substrate as indicated. A control mixture without enzyme and a tube with enzyme alone were incubated separately. After 1 hour at room temperature, the reaction mixtures were heated to 90” for 5 minutes, cooled, and assayed for FDP, SDP, F6P, S7P, and Pi (see ilfelhods).

Step I : Cell-free extract. Frozen P. pallidum myxamebae (10 gm) were ground for 5 minutes in a chilled mortar with 20 gm of alumina. The enzyme was extracted in 0.05 M tris buffer, pH 7.9, the suspension was centrifuged, and the residue was discarded (cell-free extract). Step 2: Protamine sulfate precipitation. To the supernatant fluid obtained in Step I, 36 volume of 1% protamine sulfate was added with stirring. The precipitate which formed was removed by centrifugation (protamine sulfate fraction). Step 3: Ammonium suljate fractionation. Crystalline ammonium sulfate (24.3 mg/lO ml of solution) was added with stirring and

the res~t~t p~~eipitate was removed by centrifugation. To the supernatant fluid was added additional solid ammonium sulfate (14.3 mg/lO ml), and the precipitate was collected by centrifugation. The pellet was dissolved in 20 ml of 5 m&f malonate buffer, pH 6.0 (A~o~urn Sulfate Fraction). Step 4: Sephadex G-25 eluate, The enzyme solution was passed through a 150 ml Sephadex G-25 column previously equilibrated with 5 mM malonate buffer, pH 6.0. Fractions of the eluate which contained enzymic activity were tested with a conducti~ty meter to confirm separation from ammonium sulfate and pooled (Sephadex G-25 Eluate). Step 5: CM Sephadex &&ion.5 CM Sephadex was prepared by suspending the resin in 0.2 M malonic acid and slowly adjusting the pH of the slurry to pH 5.0 with 10 N NaOH. The slurry was then poured into a 0.5 X 10 cm column and washed exhaustively with 5 mM malonate (pH 6.0) until the effluent reached pH 6.0 (approximately 20 column volumes were required). The enzyme solution was applied to the column at a rate of 1.0 ml per minute. The column was then washed successively with 5 and 10 mlM malonate buffer, pH 6.0. In each instance a protein peak was eluted and the washing was continued until the absorbance of the effluent at 280 rnp was less than 0.010 (10~150 ml was required). The enzyme was then eluted with a solution of 2.0 m&f FDP in 15 nrib’ malonate buffer, pH b A similar procedure for the elution of mammalian FDPase by substrate from B substituted cellulose column was first described by Pogell (3).

34

ROSEN

6.0. Fractions were collected (2 ml) and those with specific activities of 4.5 or above were pooled. The enzyme appeared in a peak between fractions 2 and 6 (CM Sephadex fraction). The enzyme solution could be stored at -20” for several weeks without appreciable loss of activity. ~t~~~h~o~~etryand s~~str~ spe~~e~t~. The purified enzyme catalyzed the hydrolysis of the phosphate group on C-l of fructose 1,6diphosphate and sedoheptulose 1,7-diphosphate to yield equimolar amounts of F6P and S7P, respectively (Table II). The rat’io of “FDPase” to “SDPase” activities was approxi~tely 0.8 and remained unchanged during the 400-fold purification. No hydrolysis of G6P, F6P, FlP, or GlP was observed when these were tested at concentrations of 1.0 mM with the Pi assay. E$ect of @!l and ELWA. In TEA and glycine buffers the purified FDPase of P. pallidurn showed maximal activity between pH 9.0 and pH 9.5 with no activity at pH 7.5. In the presence of EDTA, however, a new peak of hydrolytic activity appeared at pH 7.5-8.0 (Fig. 1). Under these conditions the activity in the neutral pH range became similar to that measured at pH 9.5. EDTA stimulated the pH 9.5 activity 2-fold.

6

7

6 PH

9

IO

FIG. 1. Effect of EDTA on the pH activity curve. The enzyme activity was measured speetrophotometrically using purified FDPase (1.0 pg) (see Methods). TEA buffer was used in the pH range 6.W3.5, and glycine buffer in the pH range M-10.0. The lower curve (X-X) represents the values obtained in the absence of EDTA; the upper curve {.--+) represents the values obtained in the presence of 0.5 mM EDTA.

/

/3

MERCAPTOETHANOL I

I

CONCENTRATION

IQ” OF ACTIVATOR

1

10-z &l)

FIG. 2. Compounds which activate FDPase at pH 7.5. The enzymio activity was measured spectrophotometrically at pH 7.5 with purified FDPase (1 pg) (see Methods). Control tubes containing [‘activator” and F6P were run simultaneously to eliminate any errors due to the assay procedure.

The effect of EDTA on the “SDPase” activity of the enzyme preparation was similar to that described for the “FDPase” activity. There was no hydrolysis of SDP at pH 7.5 unless EDTA was present in the reaction mixture. The effect of EDTA on the pH activity curve was observed with crude as well as purified enzyme preparations. In the presence of 1.0 mM Ylg++ and 0.1 ti FDP, the concentration of EDTA required for maximal effect was approximately 0.1 m&f (Fig. 2) at pH 7.5 and 9.5. There was marked inhibition of enzymic activity when the EDTA concentration was increased to 1.0 mM. Effect of some other compounds on enzymic activity at pH 7.5. A variety of other compounds was tested to see whether they would enhance hydrolytic activity at pH 7.5 in TEA buffer with Mg+f (1.0 mM). The following were tested at concentrations ranging from 0.01 to 1.0 m.M and were found to be ineffective : gluconate, pyruvate, fumarate, maleate, succinate, malate, histidine, hydro~ort~one, imidazole, acetyl-CoA, gly-

PRUCTOSE TABLE

l,&DIPHOSPHATASE

FROM

P. PALLZDUM

III

TABLE

EFFECT 0~ AMP AND FDP ON FDPASE

35

IV

ENZYME A~TXVITIES DURING DIRFERENTII.~TION~-6

ACTIVITY FD+se YP m2””

0.01 0.10 0.50

1.00

g22

y:gnc.

F6PlhOW) 5.5 6.0 5.0 4.8

FDPaseactivity (pm&s F6P/hour) pH 7.5

0.00 0.10 0.20 1.00

3.9 4.2 4.0 4.0b

pH 9.5

5.3 5.3 5.2 5.2

0 Eneymic activity was measured spectrophotometrically at pH 9.5 (to determine the effect of FDP) or at pH 7.5 and 9.5 (to determine the effect of AMP). In all cases, purified FDPwe (15 rg) and EDTA (0.5 mM) were present in the reaction mixture. The reactions were initiated by the addition of FDP (l-10 ~1). Where indicated, AMP (l-10 ~1) was added just prior to the addition of FDP. b Similar results were obtained when dAMP was substituted for AMP.

tine, pyrophosphate, glutathione, a-ketoglutarate, ATP, isocitrate, and citrate. Several compounds tested did induce some activity at pH 7.5; these were diethyl~thiocarb~~ic acid (DDTC), cysteine, and 2-mercaptoethanol (Fig. 2). None was as active as EDTA on a molar basis. When optimal concentrations of these compounds were added together with EDTA (0.5 mM), the activity measured at pH 7.5 was equal to that induced by EDTA alone. ideal r~~uire~~n~. The FDPase from P. pallidurn has an absolute requirement for a divalent cation. This is best satisfied by 1 mM Mg++. I\!In++ is less effective than Mg++ ; at optimal concentrations (0.1 m&f) it is only one third as active as the optimal concentration of Mg++. The “SDPW” activity of P. pal~idum FDPase is likewise dependent upon the presence of added Mg++. Zn++ and Ca++ ions did not replace Mg++. E$ect of substrate concentration. The enzyme was fully active at 0.01 m.M FDP6 and 0.1 mM SDP.’ Higher concentration of B It was dXicult to measure enzymic activity with concentrations of FDP below 0.01 mM. 7 Neither the Pi assay nor the S7P spectrophotometric assay were sufficiently sensitive for detailed studies to be performed with SDP concentrations below 0.1 m&f.

stage:,

&$;

/ “,acm

1 zz$

Fruit

EIWIIW

1. FDPase a. b. 2. PFK 3. FDP Aldolase

Enzyme activity (rmoles/min/ mg protein)

/ 0.014

0.022

0.017

1 0.016 0.052 0.037

0.015 0.075 0.029

0.015 0.079 0.047

0.017 0.019 0.075 0.047

GCells were harvested from the agar plates by

scraping them into 0.05 M tris buffer, pH 7.9. The cell suspensions were washed with buffer, centrifuged, and frozen. The frozen cells were then ground with twice their weight of alumina, extracted with buffer, and assayed immediately. The protein concentration of the extracts was approximately 10 mg/ml. FDPase, PFK, and FDP dolase were assayed spectrophotometrically by using 5-20 pl of extract (see Methods). FDPaee was measured at pH 7.5 (in the presence of EDTA) and at pH 9.5 with identical results. The two series of values outlined above (a and b) represent the activities measured in two separate experiments at pH 9.5. b The specific activities recorded are not corrected for the diminution in cellular protein concentration that occurs during the process of differentiation (16).

FDP (up to 1.0 mM) did not inhibit (Table III). Possible ~n~~bit~r~. Fluoride was an effective inhi~tor at 0.01 M. The enzymic activity was inhibited by 60 %. Unlike most of the FDPases reported in the literature (6, 10, 13, 26, 27) the P. pallidurn enzyme was not significantly inhibited by AlMP or dAMP at contentrations between 0.1 and 1.0 mM (Table III). Enzyme activity during cellular diffwentiation of P. pallidurn. The activity of FDPase was assayed in cell-free extracts at 4 different stages of differentiation.8 The myxamebae, the multicellular aggregate, the precuImin~tion stage, and the differentiated fruiting body (spores and stalks). The enzyme was assayed at pH 7.5 (in the *The hibited tration, enzyme

activity measured in these extracts exthe same response to pH, substrate concencations, and EDTA as reported for the purified from the myxamebae.

36

ROSEN

presence of EDTA) and at pH 9.5. The results (Table IV) reveal no significant alteration of FDPase activity during this process. Phosphofructokinase and FDP aldolase activities also showed no change during differentiation. DISCUSSION

The fructose 1,6-diphosphatase partially purified from the cellular slime mold P. pallidurn resembles that found in other organisms. It is stimulated by EDTA and is active with low concentrations of substrate. In the absence of EDTA the enzyme is active only at alkaline pH. It requires the presence of either Mg++ or Mn*. Like the mammalian liver FDPase (4, 5), SDP may serve as substrate for the FDPase of P. pallidurn. Since the enzyme has been purified only partiahy, it cannot be stated with certainty that the “FDPase” and “SDPase” activities are due to a single protein. However, the ratio of activities with these two substrates remained constant over a 400-fold purification, and the pH optimum, Mg++ requirement, and effect of EDTA were the same for the hydrolysis of both substrates. It is of interest that SDPase activity has been found to be present in every system which has been examined, either as a single enzyme with FDPase and SDPase activity (mammalian liver) or as a specific enzyme distinct from FDPase [Can&da stalks (24, 27), ~~cha~orn~~es cerevisiae (2)]. It has been suggested that SDPase may provide an alternate pathway for the conversion of FDP to F6P (2,28).9 The conversion of FDP to FGP by FDPase serves to provide G6P for glucose and glycogen synthesis. It is therefore a required step in gluconeogensis and in the reversal of 9 With the FDPase isolated from mammalian liver, the capacity to hydrolyze SDP is far less sensitive to AMP inhibition than is the capacity to hydrolyze FDP (0. M. Rosen, unpublished results). In C. z&&s, SDPase activity is not. increased by growth of the organism on 2- and a-carbon intermediates and is not inhibited by AMP (I. Zarembok, S. M. Rosen, and 0. M. Rosen, unpublished results). Thus the SDPase and FDPase activities in these two species appear to be under different types of controls suggesting that their functions in uivo are distinct.

glycolysis. The mechanism(s) of control of this enzyme in v&o is consequently of great interest and, from the reports in the literature, appears to vary from species to species. For example, the activity of mammalian liver FDPase has been reported to increase 2- to 3-fold during gluconeogenic stress (14); the FDPase activity in Cu~d~da utilis increases 4- to 5-fold (29); the enzyme in ~acchar~~~ces cerevisiae (13) can be detected only when these organisms are grown on 2- and S-carbon intermediates. On the other hand, the FDPase activity in Escherichiae coli is not altered during growth on various carbon sources, although the requirement for FDPase during growth on 2- and S-carbon intermediates has been clearly documented (3O).‘O Thus an increase in FDPase activity (as measured in cellfree extracts) may not always parallel the requ~ements of enhanced glueoneogenesis. The cellular slime mold undergoes a morphogenetic sequence which is accompanied by a marked increase in gluconeogenesis and polysaccharide biosynthesis. The enzyme which catalyzes a step in the bios~thesis of one such polysac~h~de in ~~t~osteliurn d~sco~m (31) has been shown to appear late in the developmental sequence shortly before the actual synthesis of the mucopolysaccharide occurs. It could not be detected in the myxamebae. It was of interest, therefore, to determine whether similar changes occurred in the activity of FDPase and phosphofructokinase during the process of differentiation. The results of this investigation revealed no alteration in the specific activities of these enzymes as measured in vitro. Since they are undoubtedly regulated in v&o, modification of their activity by the intracellular “milieu” or by changes in cellular localization may occur. In an extensive review of the biochemical alterations in morphogenesis, Wright (32) has summarized much evidence to suggest that alterations in concentrations of metabolites rather than in enzyme concentration are frequently the rate limiting factors in viva. FDPase inhibition by AMP or excess lo An FDPase-negative mutant of E. Coli isolated by Fraenkel and Horeeker (30) is unable to grow on 2- and 3-carbon intermediates.

FRUCTOSE

1,6-DIPHOSPHATASE

substrate (FDP) or both, could provide one type of control. However, neither AMP nor l?DP sig~ficax~tly inhibit the P. ~~l~~~??z enzyme. The appearance of activity in t’he neutral pH range in the presence of low concentrations of EDTA or higher concentrations of DDTC, cysteine, or mercaptoethanol is very similar to that already reported to occur in the C. utilis enzyme (27). Further investigations of naturally occurring compounds which might produce a similar effect in viva are of great interest since the reversible conversion of an enzyme active only at pH 9.0-95 to one fully active at pH 7.5 may represent, a method for regulat.ing its activity in viva. ACKNOWLEDGMENT

The aut,hor is indebted to Dr. B. L. Horecker, Depart,ment of Molecular Biology, Albert Einstein College of Medicine, N.Y., in whose laboratory most of this work was performed. His constant and helpful discussions were encouragement invaluable. REFERENCES 1. GOMORI, G., J. Viol. Chem. 148, 139 (1943). 2. RACKER, E., AND SCHROEDER, E. A. R., AT&. Biochem.. Biophys. 74, 326 (1958). 3. POGELL, B. M., Biochem. Biophys. Res. Commun. 7, 225 (1962). 4. PONTREMOLI, S., TR~NIELLO, S., LUPPIS, B., AND WOOD, W. A., 2. BiOt. Che?% 2% 3459 (1965). 5. BONSIGNORE, A., MANGIAROTTI, G., MANGIAROT’I’T, M. A., DEFLORA, A., AND PONTREMOLI, S., J. Biol. Chem. 233, 151 (1963). 6. MENDICINO, J., AND VASARHELY, F., J. Riot. 7. 8.

9.

10.

Ch,em. 288, 3528 (1963). APP, A. A., AND JAGENDORF,A. T., Biochem. Biophys. dcta 86, 427 (1964). FOSSITT, D. D., AND BERNSTEIN, I. A., J. Bacleriol. 86, 598 (1963). SMILLIE, R., in “Fructose 1 ,&Diphosphatase (R. W. md Its Role in. Gluconeogenesis McGilvery and B. M. Pogell, eds.), p. 31. American Institute for Biological Sciences, Washington, D.C. (1964). TAKETA, K., AND POGELL, B. M., J. Biol. Chem. 240, 651 (1965).

FROM P. PALLIDUM

37

11. PASSONNEAU, S. V., AND LO%-RY, 0. H., Biothem. Biophys. Res. Commun. 7, 10 (1962). 12. MCGIL~ERY, It. W., in “Fructose 1,6-Diphos-

phatase and It,s Role in Gluconeogenesis” (R. W. McGilvery and B. M. Pogell, eds.), p. 3. American Institute for Biological Sciences, Washington, I). C. (1964). 13. GRANCEDO,C., SALAS, M. L., GINER, A., AND SOLS, A., Biochem. Bioph,ys. Res. Co~~m~~n. 20, 15 (1965). 14. WEBER, G., SINGHAL: R. L., AND SRIVASTA, S. K., Proc. (1965).

Natl.

Acad.

Sci. U. S. 63, 96

15. SIXSMAN, M., dnn. 16. 17. 18. 19. 20. 21. 22. 23. 24. 25. 26. 27. 28. 29.

Rev. Microbial. 10, 21 (1956). WHITE, G. J., AND SUSSMAN, M., Biochim ~~ophy~. Acta 63, 285 (1961). SMYRNIOTIS, I?., AND HORECKER, B. L.,J. Biol. Chem. 218, 745 (1956). KOWAL, J., CREMONA, T., AND HORECKER, B. L., Arch. Biochem. Biophys. 114, 13 (1966). SUSSMAN,M., Science 139, 338 (1963). LOWRY, 0. Hi., ROSEBROUGH, N. .I., FARR, A. L., AND RANDALL, R. J., J. Viol. Chem. 193, 265 (1951). COOPER, J., SRBRE, P. A., TABACHNIK, M., AND RACKER, E., Arch. Biochem. Biophys. 74, 306 (1958). RACKER, E., J. Biot. Chem. 167, 843 (1947). Wu, R., AND RACKER, E., J. Biol. Chem. 234, 1029 (1959). PONTREMOLI, S., AND GRAZI, E., Bull. See. Chim. Biol. 42, 50 (1960). FISKE, C. H., AND SUBBARO~~-, Y., J. Biol. Chem. 66, 375 (1925). FRAENKEL, D. G., PONTREMOLI, S., AND HORECKER, B. L., Arch. Biochem. Biophys. 114, 4 (1966). ROSEN, 0. M., ROSEN, S. M., AND HORECKER, B, L., Arch. Biochem. Biophys. 112, 411 (1965). BONSIGNORE, A., PONTREMOLI, S., AND GRAZI, E., Ital. J. ~~oGhern. 10, 52 (1961). ROSEN, S. M., ROSEN, 0. M., AND HORECKER, B. L., Biochem. Biophys. Res. Commun. 20,

3 (1965). 30. FRAENKEL, D. G., AND HORECKER, B. L., J. Bacterial. 90, 837 (1965). 31. SUSSMAN,M., AND OSBORN,M. J., Proc. niati. Ad. Sei. U. S. 62, 81 (1964). Biochem32. WRIGHT, B. E., in “Comparative istry” (M. Florkin and H. S. Mason, eds.), Vol. 1, p. 1. Academic Press, New York (1964).