- Email: [email protected]
On the activation of human and rat fructose 1,6-bisphosphatase
Inr. J. Biochem. Vol. 13. pp. 337 lo 341 0 Pergamon Press Lid 1981. Printed in Great Brrtam
0020.71 IX/81/030337-OSSO2.C0/0
ON THE ACTIVATION OF HUMAN AND RAT FRUCTOSE 1,6-BISPHOSPHATASE ANTONIO
From
the Department Anderson
ORENGO* and DOMITILAM. PATENIA
of Biochemistry, The University of Texas, System Cancer Center, Hospital and Tumor Institute, Houston, Texas 77030, U.S.A.
M.D
(Received 7 July 1980) Abstract-l. Human and rat liver Fructose 1,6_bisphosphatase have low specific activities at physiological pHs. 2. Cysteine, threonine, serine and glycine activate the enzyme present in cytosols. A small amount (5%) of the enzyme is bound to microsomes. This fraction does not require any activator for maximal activity. 3. A factor that is capable of activating the cytosol enzyme has been isolated and partially purified from liver microsomes and co-elutes with the microsomal-bound enzyme. A similar factor has been
isolated from the cytosol fraction of 2 hepatomas.
INTRODUCTION Since the first description of fructose 1,6-bisphophatase (FDPase) by Gomori (1943), the alkaline pH optimum of the enzyme has been a disturbing property in the assignment of a physiological role to the enzyme. All the mammalian phosphatases studied were reported to have a pH optimum of 9-9.5. The question then arises of whether the alkaline pH optimum is an intrinsic property of the enzyme or an artifact attributable to the method of purification. Nakashima & Horecker (1971) attributed the lack of activity at neutral pH to limited proteolysis. Traniello et al. (1971) claimed the purification of a native enzyme with an optimal activity at neutral pH. However, this enzyme preparation still required (ethylenedinitrilo) tetraacetic acid (EDTA) as an activator. Pogell et al. ‘(1968), instead, advanced the hypothesis that during the purification procedure an essential activator was lost. In other studies (Baxter et al., 1972) they were able to demonstrate that oleic acid activates FDPase and could fulfill the role of the natural activator. Allen & Blair (1972) presented evidence that certain phospholipids particularly diphosph&nositide were a potent activators of the phosphatase. We present evidence that human and rat FDPases are substantially activated at neutral pH by a few amino acids, in particular by those that yield pyruvate in their oxidative degradation. Moreover, a factor was isolated from the microsomal fraction of the rat liver that proved to be an excellent activator of the phosphatase at pH 7.5.
sulfate were products of Swartz-Mann, Orangeburg, NY; DEAE cellulose was purchased from Whatman, Clifton, NJ. Young female Sprague-Dawley rats (9GllOg) were purchased from Gibco, Madison, WI. The human and rat
phosphatases were purified according to Orengo & Patenia (1980, 1976). Experiments
Purijication
should
was prepared
of an activating
factor from
rat liver microsomes
Frozen livers (150 g) were thawed at room temperature and were forced through a tissue press. The pulp was homogenized in 300 ml of 25 mM Tris-acetate-25OmM sucrose-5 mM MgCI,, pH 7.5, in a Potter-Elvehjem homogenizer which was kept in a water-ice bath. During homogenization I.5 ml of’PMSF (1 M in isopropanol) were added. The homogenate was centrifuged at ICOOg for IOmin. The sediment was discarded, and the supernatant was re-centrifuged at 7500 g for 20 mins. The microsomal fraction was sedimented bv centrifugation at 125,ooOg for 120 min. The supernatantwas discarded and the pellets were dissolved in I50 ml of 25 mM Tris-Cl-l% Triton X-100. pH 8.3. To achieve the maximal solubilization of the sediment the suspension was homogenized in the cold with a Potter-Elvehjem homogenizer. After a centrifugation at 45,OOOy for 40min the supernatant was collected and labeled as Fraction 1.
Fructose 1,6-bisphosphate trisodium salt was purchased from Boehringer Mannheim Biochemicals, New York, NY; glucose 6-phosphate dehydrogenase was from P.L. Biochemicals, Inc., Milwaukee, WI; phosphoglucose isomerase and L-amino acids were from Sigma, St Louis, MO: Tris+trishydroxymethylamino methane) and ammonium all correspondence
liver extract
livers (20g) were thawed at room temperature
MATERIALS AND METHODS
* To whom
with crude extract
as follows. Frozen and forced through a tissue press model I41 from Harvard Apparatus Co., Inc., Millis, MA. Forty milliliters of 0.154 M ice-cold KCI, pH 7.3, were added to the liver pulp, which was homogenized in less than 2 min using a tissumizer, model SDT, from Tekmar Company, Cincinatti, OH. During homogenization. 0.20 ml of phenylmethylsulfonylfluoride (PMSF) (I M in isopropanol) was added. After addition of another 40 ml of 0.154 M KCI, pH 7.3, the homogenate was centrifuged at 125,OOOg for 60 min. The supernatant was collected and dialyzed against 25 mM Naphosphate buffer, pH 7.5. Ten milliliters of liver extract were dialized against 101 of buffer. The buffer was changed 5 times at 12 hr intervals. Human
chromatography at pH 8.3 A DEAE-cellulose column (2.5 x 26 cm) was packed and equilibrated with 800ml of 25 mM Tris-Cl-l% Triton X-100, pH 8.3. Fraction 1 (16mg/ml, 150ml). was applied MAE-cellulose
be addressed. 337
ANTONIO ORENGO and DOMITILA M. PATENIA
338
to the column and washed through with 3OOml of the eauilibratina buffer. A gradient from o-O.5 M NaCl in the equilibrating buffer was-used to elute the column. The total volume of the gradient was 600ml. The flow rate was I5 mlihr. The peak of the activity was eluted at a concentration of 0.15 M NaCI. The volume of the fractions was 2.3 ml. Proteins were measured according to Lowry ef al. (1951). The fractions with the highest activities were pooled. yielding Fraction 2. DEAE-cr//ulo.w
chromatoyruphy
at pH Y.5
A second DEAE column (2.5 x 26cm) was packed and eauilibrated with 800 ml of 5 mM TrisCI, pH 9.5, and applied to the column, The column was then washed with distilled water until the proteins eluted were below 0.02 mg,‘ml. A gradient of NaCl (O-l.0 M) in the equilibrating buffer was then applied to the column. The peak of the active fractions was eluted at 0.34 M NaCI. The total volume of the gradient was 600 ml. the volume of the fractions 4.75 ml. and the flow rate was 15 ml/hr. Proteins were measured according to Lowry cv al. (1951). The fractions with highest activity were pooled, yielding Fraction 3. Fraction 3 was lyophilized and dissolved in 22ml of water. The solution was then shaken with 4 volumes of chloroform. The chloroform phase was discarded, and after a brief centrifugation. the water phase was collected and lyophilized. The lyophilized material was then dissolved in 10ml of distilled water and used in the experiments here described. Prrpuration
oftrscitic
@ids
The various ascitic fluids were brought to pH 8.0 by addition of Tris and centrifuged at 125.000 g for 30 min. The supernatants were collected and used in the activation experiments. Prrparution
of tumor w//s
extract.5
Tumor cells were separated from the ascitic fluid by centrifugation. The cells were suspended in 5 mM Trissglycine buffer. pH 8.3 and broken in the presence of PMSF (0.5 ml of 1 M solution per 100 ml of homogenate) with the aid of a tissuemizer. The extract was centrifuged at 125,000 g for 30 min. and the supernatant was used in the activation experiments.
All the reagents for the enzyme assays were adjusted to pH 7.0 with NaOH when it was necessary. The pH measurements were made at room temperature, The routine assay for neutral activity was carried out at 37-C. The reaction -mixture contained 0.02 M triethanolamineeO.02 M diethanolamineeHCI, pH 7.5; 2 mM MgSO,; 0.3 mM NADP+ ; 0.1 mM fructose 1,6-PZ; 4 pg of glucose 6-phosphate dehydrogenase (140 U/mg); 7 fig-of hexosephosphate isomerase (535 Ujmg), and an appropriate amount of the enzyme. A unit of activity was defined as 1 pmol of fructose 6-phosphate formed per min. A Cary 15 recording spectrophotometer equipped with a Lauda circulating bath was used to monitor the enzymic rate at 1Osec intervals. The activating compounds, EDTA, mercaptoethanol, L-amino acids and microsomal fractions, were added at the concentrations indicated. All the maximal velocities are reported as percentage fractions of the maximal velocity obtained in presence of EDTA. In all cases, the reactions were started by the addition of FDPase.
RESULTS The activation of human liver FDPase by EDTA, 2-mercaptoethanol and amino acids is presented in Table 1. The activation by EDTA gave a sigmoidal response, with an apparent number of 3 binding sites
Table 1. Activation
of human liver fructose tase at pH 7.5 Liver extract
Addition EDTA 2-Mercapto ethanol Cysteine Threonine Serine Glycine Alanine Histidine Lysine
(r% 0.0005vt 0.49 0.14 0.44 I .80 2.00 5.80 0.10 8.40
l,6-biphospha-
Pure enzyme
v,,,*
(r%
VI,,*
100
0.0007t
IO0
90 90 68 89 99 81 72 71
4.25 0.22 13.60 5.30 21.30 22.00 0.90 22.80
94 83 57 90 89 83 75 64
* All maximal velocities are reported as a percent fraction of the maximal velcocity obtained in presence of EDTA. t These are K’.
and a K’ Qf 0.7 PM. Cysteine, serine, glycine and alanine were’found to be the most effective activators of the enzyme among the 20 amino acids that we tested. Three other amino acids, histidine, threonine and lysine, were capable of activating the phosphatase. but to a limited degree. No activation by amino acids could be detected at pH9.5. The phosphatase activated by cysteine is as sensitive to 5’-AMP inhibition as the one activated by EDTA. In comparing the activation of an extract extensively dialyzed with a highly purified preparation of the enzyme, it was apparent that during the purification the sensitivity of the phosphatase to the activation by amino acids is somewhat diminished. The same pattern of activation, however, could be noted. 2-mercaptoethanol is also capable of activating the phosphatase. We have obtained similar results with the rat liver phosphatase although somewhat higher concentrations of amino acids were required. The pattern of activation presented is reproducible, although the K, values may vary from experiment to experiment with different preparations of the phosphatase. FDPase has been found to be an enzyme of the cytosol in all mammals studied. We noticed, however, that approx 5% of FDPase activity was found bound to the microsomal fraction of the rat liver. This phosphatase was sensitive to 5’-AMP and was fully active at pH 7.5 in absence of EDTA. The phosphatase could be easily salubilized by treatment with Triton X-100 and partially purified by two successive chromatographies on DEAE cellulose at pH 8.3 and 9.5. In order to differentiate between the presence of an activating factor or of a phosphatase undegraded by proteases, we added 2 pg of highly purified cytosol FDPase to all the fractions eluted from the DEAEcellulose column at pH9.5 and the assays were carried out at neutral pH. The results of this experiment are presented in Fig. 1. Two large peaks of an activating factor were eluted. The first overlapped the microsomal-bound FDPase. The factor appears to be a protein or a compound associated to a protein. At the stage of purification that we have achieved, the factor is not dialyzable and is not soluble in chloroform.
Activation of fructose i,&bisphosphatase
339
0.32 M NaCl
8
6
100
50
150
200
Fractions Fig. 1. Elution of rat microsomal-bound FDPase and activating factor from a DEAE-cellulose column. Fraction II was diluted 5 times with 0.005 M tris-Cl, pH 9.5, and applied to a DEAE-cellulose column (2.5 x 25 cm) equilibrated with the diluting buffer. The column was washed with 430 ml of water, and fractions were eluted with a linear gradient of NaCl (O-l.0 M. 600 ml) in the same Ttis-Cl buffer. The microsomal-bound FDPase was assayed in the routine reaction mixture at pH 7.5 in the absence of EDTA. In parallel assay, an appropriate amount of rat liver cytosol FDPase was added to the reaction mixtures in order to measure the activating microsomal factor. The fractions that contained both the microsomal phosphatase and the activating factor were corrected for the contribution of the microsomal FDPase before plotting the elution profile of the activating factor. The proteins were measured according to the method Lowry er al. (9).
Activation
of Liver F,t -6 Bisphosphatase
After treatment
of an aqueous
solution
of the activat-
ing factor with several volumes of chloroform, the water phase retained 75% of its activity. paralleled by an equal recovery of its protein content. When the kinetics of activation of an appropriate amount of rat cytosol FDPase by the rat microsomal factor was studied, a sigmoidal response was noted (Fig. 2). The rat microsomal activating factor was also capable of activating human liver cytosol FDPase, although with a lower apparent number of binding sites. The microsomal factor was as active as EDTA in increasing the specific activity of the phosphatase at neutral pHs. Pogell er al. (1968) reported that extracts of Novikoff hepatoma cells were capable of increasing the FDPase activity of dialyzed rabbit liver supematant. This finding is noteworthy since neither Pogell et al. (1968) nor we could detect FDPase activity in Novikoff hepatoma cells extracts, In Table 2, we showed the tests of several tumor cell extracts using a highly purified rat liver FDPase. The activator was present in 2 20 Rat Liver
40 M~crosomal
60 Activator
80
100
(pg of Protein)
Fig. 2. Human liver FDPase (2.1 pg) and rat liver FDPase (2.3~~) were activated by the above increasing concentrations of rat microsomai activator (0-iOO jzg of protein). The activity of the phosphat~e was determined at pH 7.5 as described in Materials and Methods. B.C 13,3--r
hepatoma cell extracts and in their ascitic fluids but not in the other tumor extracts and ascitic fluids tested. DISCUSSION The co-existence of phosphofruetokinase and FDPase in the cytosols of muscle, renal and liver cells
340
ANT~NIOORENG~and DOMITILAM. PATENIA Table 2. Activation of rat liver fructose 1,4-biphosphatase ascitic fluids Final concentration (protein, mg)
Addition
by tumor cell extracts and
Activity Activation (U min-’ pH 7.5) (folds)
0
0.08
1.0
Ascitic fluid from Novikoff tumor
0.70 1.00
0.40 0.48
5.0 6.0
Novikoff tumor cell extract
0.30 0.50
0.55 0.58
6.9 1.3
None
Ascitic fluid from rat tumor AS-30-D (Smith et al., 1970)
1.20
0.17
2.1
2.40
0.24
3.0
Extract from rat tumor AS-30-D
0.40 0.80
0.20 0.34
2.5 4.3
As&tic fluid from-L-1210 Mouse leukemia
0.25
0.10
I.3
Ascitic fluid from mouse Erlich tumor-mustard resistant
0.37
0.09
1.1
Ceil extract of Erlich mustard resistant tumor
0.35
0.18
2.2
0.40 0.60
0.16 0.15
2.0 1.8
Bovine serum albumin
raises the questions of the regulation of these enzymic activities. The simultaneous operation of the 2 enzymes should generate a futile cycle of phosphorylation and dephosphorylation. with a concomitant waste of ATP. Scheme I.
ATP
ADP
Fructose,6-phosphate
I-6 bisphos~ate. Since there is no evidence of separate subcellular localization of the 2 enzymes. one must assume that a fine interplay of inhibition and activation takes place, allowing a modulation of the enzymic activities according to physiological needs. The phosphatase has been found to be of very low activity in cytosot extracts that were assayed at physiological pH. However. the chelating agent EDTA was found to activate the phosphatase. The activation by this artificial compund. although of interest in the study of the mechanism of catalysis. does not shed light on the in ~.ico regulation of the enzymic activity. We present evidence here that the 4 amino acids that exclusively yield pyruvate in their oxidative degradation are capable of activating the enzyme at concentrations which may be compatible with control by those occurring in l.iCO. The regulation of glucose homeostasis and gluconeogenesis is under several types of regulatory mech-
anisms We are limiting, here. our studies to mechanisms that may play a role in the regulation of enzymic activities. Other mechanisms involving increased de noro synthesis of enzymes incited by hormones or effecters are probably of equal or greater importance in determining the rate of gluconeogenesis. Possibly the mechanisms regulating enzymic activities are responsible for the immediate adjustment of rates, while mechanisms involving enzyme synthesis should be considered physiological adaptations to long-lasting situations! The activation by FDPase by amino acids may operate at the initiating steps, where an alternative exists between gluconeogenesis and oxidation through the Krebs cycle. In studying the rate of gluconeogenesis from various precursors in rat kidney cortex, i(rebs (1964) noted that the rate obtained from alanine, serine, valine, isoleucine and threonione were not significantly different from the rate obtained in the absence of added substrate. For example, a rate of 263 pmol g dry weight- ’ hr- ’ was found after addition of L-glutamate, while a value of approx 201.lmol was found for alanine. These differences are not clearly evident in experiments using liver perfusion or liver slices. It should be pointed out, however, that it is quite difficult to measure gluconeogenetic rates in liver. where carbohydrates are rapidly converted to fat, cholesterol, and amino acids. In addition, liver stores carbohydrates and appears to be less permeable to polyvalent anions, i.e. glutamic acid (Krebs, 1964). It is possible. therefore, that amino acids such as cysteine or serine (Table 1) could substantially increase the flow through fructose l,&bisphosphatase in some phases of the physiolo~cal adaptions during which gluconeogenesis is required.
Activation of fructose 1,6-bisphosphatase The finding that a small percentage of FDPase is bound to the microsomal fraction of the liver may be of interest. Allen & Blair (1972) had in fact suggested from their study of activation of the phosphatase by phospholipids that the enzyme may be attached to membranes and therefore influenced by their conformational state. The microsomal fraction also contains an activating factor, a part of which is co-eluted with the phosphatase. The chemical nature of this activating factor is not known at the present time. It appears to be a protein or at least a compound strongly bound to a protein. It was not dialyzable and does not appear to be soluble in chloroform. Further studies are necessary for its characterization. Finally, tumor cells can be viewed as metabolic parasites, since they almost exclusively utilize glucose with a large production of lactate. It is known that gluconeogenesis is enhanced in liver and kidney of tumor-bearing animals. The finding that cytosol extracts of hepatoma and their ascitic fluid are capable of activating FDPase raises the important question of whether some tumor cells have high concentration of the activator in soluble form. The tumor could, by secreting it, activate FDPase in the liver and kidney and, possibly, in muscles. This would generate a futile cycle of phosphorylation and dephosphorylation of fructose 6-phosphate with a waste of high energy equivalents, which could contribute to the profound cachexia that many tumors induce in their host. REFERENCES ALLEN M. B. & BLAIR J. McD. (1972) The regulation
of
rabbit liver fructose 1,6-diphosphatase activity pholipids in vitro. Biochem. J. i30, 1167-1169.
341 by phos-
BAXTER R. C.. CARLWN C. W. & P~GELL B. M. (1972) Stimulation of the neutral activity of rabbit liver fr&zto& 1,6_Diphosphatase by fatty acids. J. biol. Chem. 247, 2969-2971. GOMORIG. (1943) Hexosediphosphatase J. biol. Gem. 148, 139-149. KREBS H. A. (1964) The Croonian lecture, 1963. Gluconeoeenesis. Proc. R. Sot. Lond. 159B. 545-564. LOWRY 0. H., ROSENBROUGHN. J., FARR A. L. & RANDALL R. J. (1951) Protein measurement with the Folin phenol reagent. J. biol. Chem. 193, 265-275. NAKASHIMA K. & HORECKER B. L. (1971) Modification of the catalytic properties of rabbit liver fructose diphosphatase by a particulate fraction from liver. Arch Biothem. Biophys. 146, 153-160. ORENGOA. & PATEN~AD. M. (1976) Exploitable molecular mechanisms in hibernation-I. Liver diphosphofructose phosphatase of the rat and hamster: a comparison. Como. Biochem. Phiol. 55B. 283-291. ORENG~A. & PATEN~AD. M. (1980) Human liver fructose 1,6-biphosphatase. Purification and properties. Int. J. Biochem. 13, 329-335. POGELL B. M., TANAKA A. & S~~DONS R. C. (1968) Natural activators for liver fructose 1.6-diphosphatase and the reversal of adenosine 5’-monoph’osphate. Inhibition by muscle phosphofructokinase. J. hiol. Chem. 243. 1356-1367. SMITH D. F.. WALBORG E. F. & CHANG J. P. (1970) Establishment of a transplantable ascites variant of rat hepatoma induced by 3’-methyl I-4-dimethylaminoazobenzene. Cuncer Res. 30, 2306-2309. TRANIELLO S.. PONTREMOLI S., TASHIMA Y. 8~ HORECKER B. L. (1971) Fructose 1,6_diphosphatase from liver. Isolation of the native form with optimal activity at neutral pH. Archs Biochem. Biophys. 146. 161-166.
0020.71 IX/81/030337-OSSO2.C0/0
ON THE ACTIVATION OF HUMAN AND RAT FRUCTOSE 1,6-BISPHOSPHATASE ANTONIO
From
the Department Anderson
ORENGO* and DOMITILAM. PATENIA
of Biochemistry, The University of Texas, System Cancer Center, Hospital and Tumor Institute, Houston, Texas 77030, U.S.A.
M.D
(Received 7 July 1980) Abstract-l. Human and rat liver Fructose 1,6_bisphosphatase have low specific activities at physiological pHs. 2. Cysteine, threonine, serine and glycine activate the enzyme present in cytosols. A small amount (5%) of the enzyme is bound to microsomes. This fraction does not require any activator for maximal activity. 3. A factor that is capable of activating the cytosol enzyme has been isolated and partially purified from liver microsomes and co-elutes with the microsomal-bound enzyme. A similar factor has been
isolated from the cytosol fraction of 2 hepatomas.
INTRODUCTION Since the first description of fructose 1,6-bisphophatase (FDPase) by Gomori (1943), the alkaline pH optimum of the enzyme has been a disturbing property in the assignment of a physiological role to the enzyme. All the mammalian phosphatases studied were reported to have a pH optimum of 9-9.5. The question then arises of whether the alkaline pH optimum is an intrinsic property of the enzyme or an artifact attributable to the method of purification. Nakashima & Horecker (1971) attributed the lack of activity at neutral pH to limited proteolysis. Traniello et al. (1971) claimed the purification of a native enzyme with an optimal activity at neutral pH. However, this enzyme preparation still required (ethylenedinitrilo) tetraacetic acid (EDTA) as an activator. Pogell et al. ‘(1968), instead, advanced the hypothesis that during the purification procedure an essential activator was lost. In other studies (Baxter et al., 1972) they were able to demonstrate that oleic acid activates FDPase and could fulfill the role of the natural activator. Allen & Blair (1972) presented evidence that certain phospholipids particularly diphosph&nositide were a potent activators of the phosphatase. We present evidence that human and rat FDPases are substantially activated at neutral pH by a few amino acids, in particular by those that yield pyruvate in their oxidative degradation. Moreover, a factor was isolated from the microsomal fraction of the rat liver that proved to be an excellent activator of the phosphatase at pH 7.5.
sulfate were products of Swartz-Mann, Orangeburg, NY; DEAE cellulose was purchased from Whatman, Clifton, NJ. Young female Sprague-Dawley rats (9GllOg) were purchased from Gibco, Madison, WI. The human and rat
phosphatases were purified according to Orengo & Patenia (1980, 1976). Experiments
Purijication
should
was prepared
of an activating
factor from
rat liver microsomes
Frozen livers (150 g) were thawed at room temperature and were forced through a tissue press. The pulp was homogenized in 300 ml of 25 mM Tris-acetate-25OmM sucrose-5 mM MgCI,, pH 7.5, in a Potter-Elvehjem homogenizer which was kept in a water-ice bath. During homogenization I.5 ml of’PMSF (1 M in isopropanol) were added. The homogenate was centrifuged at ICOOg for IOmin. The sediment was discarded, and the supernatant was re-centrifuged at 7500 g for 20 mins. The microsomal fraction was sedimented bv centrifugation at 125,ooOg for 120 min. The supernatantwas discarded and the pellets were dissolved in I50 ml of 25 mM Tris-Cl-l% Triton X-100. pH 8.3. To achieve the maximal solubilization of the sediment the suspension was homogenized in the cold with a Potter-Elvehjem homogenizer. After a centrifugation at 45,OOOy for 40min the supernatant was collected and labeled as Fraction 1.
Fructose 1,6-bisphosphate trisodium salt was purchased from Boehringer Mannheim Biochemicals, New York, NY; glucose 6-phosphate dehydrogenase was from P.L. Biochemicals, Inc., Milwaukee, WI; phosphoglucose isomerase and L-amino acids were from Sigma, St Louis, MO: Tris+trishydroxymethylamino methane) and ammonium all correspondence
liver extract
livers (20g) were thawed at room temperature
MATERIALS AND METHODS
* To whom
with crude extract
as follows. Frozen and forced through a tissue press model I41 from Harvard Apparatus Co., Inc., Millis, MA. Forty milliliters of 0.154 M ice-cold KCI, pH 7.3, were added to the liver pulp, which was homogenized in less than 2 min using a tissumizer, model SDT, from Tekmar Company, Cincinatti, OH. During homogenization. 0.20 ml of phenylmethylsulfonylfluoride (PMSF) (I M in isopropanol) was added. After addition of another 40 ml of 0.154 M KCI, pH 7.3, the homogenate was centrifuged at 125,OOOg for 60 min. The supernatant was collected and dialyzed against 25 mM Naphosphate buffer, pH 7.5. Ten milliliters of liver extract were dialized against 101 of buffer. The buffer was changed 5 times at 12 hr intervals. Human
chromatography at pH 8.3 A DEAE-cellulose column (2.5 x 26 cm) was packed and equilibrated with 800ml of 25 mM Tris-Cl-l% Triton X-100, pH 8.3. Fraction 1 (16mg/ml, 150ml). was applied MAE-cellulose
be addressed. 337
ANTONIO ORENGO and DOMITILA M. PATENIA
338
to the column and washed through with 3OOml of the eauilibratina buffer. A gradient from o-O.5 M NaCl in the equilibrating buffer was-used to elute the column. The total volume of the gradient was 600ml. The flow rate was I5 mlihr. The peak of the activity was eluted at a concentration of 0.15 M NaCI. The volume of the fractions was 2.3 ml. Proteins were measured according to Lowry ef al. (1951). The fractions with the highest activities were pooled. yielding Fraction 2. DEAE-cr//ulo.w
chromatoyruphy
at pH Y.5
A second DEAE column (2.5 x 26cm) was packed and eauilibrated with 800 ml of 5 mM TrisCI, pH 9.5, and applied to the column, The column was then washed with distilled water until the proteins eluted were below 0.02 mg,‘ml. A gradient of NaCl (O-l.0 M) in the equilibrating buffer was then applied to the column. The peak of the active fractions was eluted at 0.34 M NaCI. The total volume of the gradient was 600 ml. the volume of the fractions 4.75 ml. and the flow rate was 15 ml/hr. Proteins were measured according to Lowry cv al. (1951). The fractions with highest activity were pooled, yielding Fraction 3. Fraction 3 was lyophilized and dissolved in 22ml of water. The solution was then shaken with 4 volumes of chloroform. The chloroform phase was discarded, and after a brief centrifugation. the water phase was collected and lyophilized. The lyophilized material was then dissolved in 10ml of distilled water and used in the experiments here described. Prrpuration
oftrscitic
@ids
The various ascitic fluids were brought to pH 8.0 by addition of Tris and centrifuged at 125.000 g for 30 min. The supernatants were collected and used in the activation experiments. Prrparution
of tumor w//s
extract.5
Tumor cells were separated from the ascitic fluid by centrifugation. The cells were suspended in 5 mM Trissglycine buffer. pH 8.3 and broken in the presence of PMSF (0.5 ml of 1 M solution per 100 ml of homogenate) with the aid of a tissuemizer. The extract was centrifuged at 125,000 g for 30 min. and the supernatant was used in the activation experiments.
All the reagents for the enzyme assays were adjusted to pH 7.0 with NaOH when it was necessary. The pH measurements were made at room temperature, The routine assay for neutral activity was carried out at 37-C. The reaction -mixture contained 0.02 M triethanolamineeO.02 M diethanolamineeHCI, pH 7.5; 2 mM MgSO,; 0.3 mM NADP+ ; 0.1 mM fructose 1,6-PZ; 4 pg of glucose 6-phosphate dehydrogenase (140 U/mg); 7 fig-of hexosephosphate isomerase (535 Ujmg), and an appropriate amount of the enzyme. A unit of activity was defined as 1 pmol of fructose 6-phosphate formed per min. A Cary 15 recording spectrophotometer equipped with a Lauda circulating bath was used to monitor the enzymic rate at 1Osec intervals. The activating compounds, EDTA, mercaptoethanol, L-amino acids and microsomal fractions, were added at the concentrations indicated. All the maximal velocities are reported as percentage fractions of the maximal velocity obtained in presence of EDTA. In all cases, the reactions were started by the addition of FDPase.
RESULTS The activation of human liver FDPase by EDTA, 2-mercaptoethanol and amino acids is presented in Table 1. The activation by EDTA gave a sigmoidal response, with an apparent number of 3 binding sites
Table 1. Activation
of human liver fructose tase at pH 7.5 Liver extract
Addition EDTA 2-Mercapto ethanol Cysteine Threonine Serine Glycine Alanine Histidine Lysine
(r% 0.0005vt 0.49 0.14 0.44 I .80 2.00 5.80 0.10 8.40
l,6-biphospha-
Pure enzyme
v,,,*
(r%
VI,,*
100
0.0007t
IO0
90 90 68 89 99 81 72 71
4.25 0.22 13.60 5.30 21.30 22.00 0.90 22.80
94 83 57 90 89 83 75 64
* All maximal velocities are reported as a percent fraction of the maximal velcocity obtained in presence of EDTA. t These are K’.
and a K’ Qf 0.7 PM. Cysteine, serine, glycine and alanine were’found to be the most effective activators of the enzyme among the 20 amino acids that we tested. Three other amino acids, histidine, threonine and lysine, were capable of activating the phosphatase. but to a limited degree. No activation by amino acids could be detected at pH9.5. The phosphatase activated by cysteine is as sensitive to 5’-AMP inhibition as the one activated by EDTA. In comparing the activation of an extract extensively dialyzed with a highly purified preparation of the enzyme, it was apparent that during the purification the sensitivity of the phosphatase to the activation by amino acids is somewhat diminished. The same pattern of activation, however, could be noted. 2-mercaptoethanol is also capable of activating the phosphatase. We have obtained similar results with the rat liver phosphatase although somewhat higher concentrations of amino acids were required. The pattern of activation presented is reproducible, although the K, values may vary from experiment to experiment with different preparations of the phosphatase. FDPase has been found to be an enzyme of the cytosol in all mammals studied. We noticed, however, that approx 5% of FDPase activity was found bound to the microsomal fraction of the rat liver. This phosphatase was sensitive to 5’-AMP and was fully active at pH 7.5 in absence of EDTA. The phosphatase could be easily salubilized by treatment with Triton X-100 and partially purified by two successive chromatographies on DEAE cellulose at pH 8.3 and 9.5. In order to differentiate between the presence of an activating factor or of a phosphatase undegraded by proteases, we added 2 pg of highly purified cytosol FDPase to all the fractions eluted from the DEAEcellulose column at pH9.5 and the assays were carried out at neutral pH. The results of this experiment are presented in Fig. 1. Two large peaks of an activating factor were eluted. The first overlapped the microsomal-bound FDPase. The factor appears to be a protein or a compound associated to a protein. At the stage of purification that we have achieved, the factor is not dialyzable and is not soluble in chloroform.
Activation of fructose i,&bisphosphatase
339
0.32 M NaCl
8
6
100
50
150
200
Fractions Fig. 1. Elution of rat microsomal-bound FDPase and activating factor from a DEAE-cellulose column. Fraction II was diluted 5 times with 0.005 M tris-Cl, pH 9.5, and applied to a DEAE-cellulose column (2.5 x 25 cm) equilibrated with the diluting buffer. The column was washed with 430 ml of water, and fractions were eluted with a linear gradient of NaCl (O-l.0 M. 600 ml) in the same Ttis-Cl buffer. The microsomal-bound FDPase was assayed in the routine reaction mixture at pH 7.5 in the absence of EDTA. In parallel assay, an appropriate amount of rat liver cytosol FDPase was added to the reaction mixtures in order to measure the activating microsomal factor. The fractions that contained both the microsomal phosphatase and the activating factor were corrected for the contribution of the microsomal FDPase before plotting the elution profile of the activating factor. The proteins were measured according to the method Lowry er al. (9).
Activation
of Liver F,t -6 Bisphosphatase
After treatment
of an aqueous
solution
of the activat-
ing factor with several volumes of chloroform, the water phase retained 75% of its activity. paralleled by an equal recovery of its protein content. When the kinetics of activation of an appropriate amount of rat cytosol FDPase by the rat microsomal factor was studied, a sigmoidal response was noted (Fig. 2). The rat microsomal activating factor was also capable of activating human liver cytosol FDPase, although with a lower apparent number of binding sites. The microsomal factor was as active as EDTA in increasing the specific activity of the phosphatase at neutral pHs. Pogell er al. (1968) reported that extracts of Novikoff hepatoma cells were capable of increasing the FDPase activity of dialyzed rabbit liver supematant. This finding is noteworthy since neither Pogell et al. (1968) nor we could detect FDPase activity in Novikoff hepatoma cells extracts, In Table 2, we showed the tests of several tumor cell extracts using a highly purified rat liver FDPase. The activator was present in 2 20 Rat Liver
40 M~crosomal
60 Activator
80
100
(pg of Protein)
Fig. 2. Human liver FDPase (2.1 pg) and rat liver FDPase (2.3~~) were activated by the above increasing concentrations of rat microsomai activator (0-iOO jzg of protein). The activity of the phosphat~e was determined at pH 7.5 as described in Materials and Methods. B.C 13,3--r
hepatoma cell extracts and in their ascitic fluids but not in the other tumor extracts and ascitic fluids tested. DISCUSSION The co-existence of phosphofruetokinase and FDPase in the cytosols of muscle, renal and liver cells
340
ANT~NIOORENG~and DOMITILAM. PATENIA Table 2. Activation of rat liver fructose 1,4-biphosphatase ascitic fluids Final concentration (protein, mg)
Addition
by tumor cell extracts and
Activity Activation (U min-’ pH 7.5) (folds)
0
0.08
1.0
Ascitic fluid from Novikoff tumor
0.70 1.00
0.40 0.48
5.0 6.0
Novikoff tumor cell extract
0.30 0.50
0.55 0.58
6.9 1.3
None
Ascitic fluid from rat tumor AS-30-D (Smith et al., 1970)
1.20
0.17
2.1
2.40
0.24
3.0
Extract from rat tumor AS-30-D
0.40 0.80
0.20 0.34
2.5 4.3
As&tic fluid from-L-1210 Mouse leukemia
0.25
0.10
I.3
Ascitic fluid from mouse Erlich tumor-mustard resistant
0.37
0.09
1.1
Ceil extract of Erlich mustard resistant tumor
0.35
0.18
2.2
0.40 0.60
0.16 0.15
2.0 1.8
Bovine serum albumin
raises the questions of the regulation of these enzymic activities. The simultaneous operation of the 2 enzymes should generate a futile cycle of phosphorylation and dephosphorylation. with a concomitant waste of ATP. Scheme I.
ATP
ADP
Fructose,6-phosphate
I-6 bisphos~ate. Since there is no evidence of separate subcellular localization of the 2 enzymes. one must assume that a fine interplay of inhibition and activation takes place, allowing a modulation of the enzymic activities according to physiological needs. The phosphatase has been found to be of very low activity in cytosot extracts that were assayed at physiological pH. However. the chelating agent EDTA was found to activate the phosphatase. The activation by this artificial compund. although of interest in the study of the mechanism of catalysis. does not shed light on the in ~.ico regulation of the enzymic activity. We present evidence here that the 4 amino acids that exclusively yield pyruvate in their oxidative degradation are capable of activating the enzyme at concentrations which may be compatible with control by those occurring in l.iCO. The regulation of glucose homeostasis and gluconeogenesis is under several types of regulatory mech-
anisms We are limiting, here. our studies to mechanisms that may play a role in the regulation of enzymic activities. Other mechanisms involving increased de noro synthesis of enzymes incited by hormones or effecters are probably of equal or greater importance in determining the rate of gluconeogenesis. Possibly the mechanisms regulating enzymic activities are responsible for the immediate adjustment of rates, while mechanisms involving enzyme synthesis should be considered physiological adaptations to long-lasting situations! The activation by FDPase by amino acids may operate at the initiating steps, where an alternative exists between gluconeogenesis and oxidation through the Krebs cycle. In studying the rate of gluconeogenesis from various precursors in rat kidney cortex, i(rebs (1964) noted that the rate obtained from alanine, serine, valine, isoleucine and threonione were not significantly different from the rate obtained in the absence of added substrate. For example, a rate of 263 pmol g dry weight- ’ hr- ’ was found after addition of L-glutamate, while a value of approx 201.lmol was found for alanine. These differences are not clearly evident in experiments using liver perfusion or liver slices. It should be pointed out, however, that it is quite difficult to measure gluconeogenetic rates in liver. where carbohydrates are rapidly converted to fat, cholesterol, and amino acids. In addition, liver stores carbohydrates and appears to be less permeable to polyvalent anions, i.e. glutamic acid (Krebs, 1964). It is possible. therefore, that amino acids such as cysteine or serine (Table 1) could substantially increase the flow through fructose l,&bisphosphatase in some phases of the physiolo~cal adaptions during which gluconeogenesis is required.
Activation of fructose 1,6-bisphosphatase The finding that a small percentage of FDPase is bound to the microsomal fraction of the liver may be of interest. Allen & Blair (1972) had in fact suggested from their study of activation of the phosphatase by phospholipids that the enzyme may be attached to membranes and therefore influenced by their conformational state. The microsomal fraction also contains an activating factor, a part of which is co-eluted with the phosphatase. The chemical nature of this activating factor is not known at the present time. It appears to be a protein or at least a compound strongly bound to a protein. It was not dialyzable and does not appear to be soluble in chloroform. Further studies are necessary for its characterization. Finally, tumor cells can be viewed as metabolic parasites, since they almost exclusively utilize glucose with a large production of lactate. It is known that gluconeogenesis is enhanced in liver and kidney of tumor-bearing animals. The finding that cytosol extracts of hepatoma and their ascitic fluid are capable of activating FDPase raises the important question of whether some tumor cells have high concentration of the activator in soluble form. The tumor could, by secreting it, activate FDPase in the liver and kidney and, possibly, in muscles. This would generate a futile cycle of phosphorylation and dephosphorylation of fructose 6-phosphate with a waste of high energy equivalents, which could contribute to the profound cachexia that many tumors induce in their host. REFERENCES ALLEN M. B. & BLAIR J. McD. (1972) The regulation
of
rabbit liver fructose 1,6-diphosphatase activity pholipids in vitro. Biochem. J. i30, 1167-1169.
341 by phos-
BAXTER R. C.. CARLWN C. W. & P~GELL B. M. (1972) Stimulation of the neutral activity of rabbit liver fr&zto& 1,6_Diphosphatase by fatty acids. J. biol. Chem. 247, 2969-2971. GOMORIG. (1943) Hexosediphosphatase J. biol. Gem. 148, 139-149. KREBS H. A. (1964) The Croonian lecture, 1963. Gluconeoeenesis. Proc. R. Sot. Lond. 159B. 545-564. LOWRY 0. H., ROSENBROUGHN. J., FARR A. L. & RANDALL R. J. (1951) Protein measurement with the Folin phenol reagent. J. biol. Chem. 193, 265-275. NAKASHIMA K. & HORECKER B. L. (1971) Modification of the catalytic properties of rabbit liver fructose diphosphatase by a particulate fraction from liver. Arch Biothem. Biophys. 146, 153-160. ORENGOA. & PATEN~AD. M. (1976) Exploitable molecular mechanisms in hibernation-I. Liver diphosphofructose phosphatase of the rat and hamster: a comparison. Como. Biochem. Phiol. 55B. 283-291. ORENG~A. & PATEN~AD. M. (1980) Human liver fructose 1,6-biphosphatase. Purification and properties. Int. J. Biochem. 13, 329-335. POGELL B. M., TANAKA A. & S~~DONS R. C. (1968) Natural activators for liver fructose 1.6-diphosphatase and the reversal of adenosine 5’-monoph’osphate. Inhibition by muscle phosphofructokinase. J. hiol. Chem. 243. 1356-1367. SMITH D. F.. WALBORG E. F. & CHANG J. P. (1970) Establishment of a transplantable ascites variant of rat hepatoma induced by 3’-methyl I-4-dimethylaminoazobenzene. Cuncer Res. 30, 2306-2309. TRANIELLO S.. PONTREMOLI S., TASHIMA Y. 8~ HORECKER B. L. (1971) Fructose 1,6_diphosphatase from liver. Isolation of the native form with optimal activity at neutral pH. Archs Biochem. Biophys. 146. 161-166.