- Email: [email protected]
Reversible inactivation of d -fructose 1,6-diphosphatase by adenosine triphosphate and cyclic 3′, 5′-adenosine monophosphate
ARCHIVES
OF
BIOCHEMISTRY
Reversible
.lND
Inactivation Adenosine
116,
BIOI’HTSICS
Department
(1966)
of D-Fructose Triphosphate
Adenosine JOSEPH
436-445
MENDICINO, RABINDRA of Biochemistry,
1,6-Diphosphatase and
Cyclic
by
3’, 5’-
Monophosphate”’ CHARLES BEAUDREAU, N. BHATTACHARYYA The Ohio
Received
May
Slate
University,
Columbus,
AND
Ohio
19, 1966
An analytical method for the quantitative recovery and purification of fructose l,G-diphosphatase from incubation mixtures containing crude kidney extracts has been described. Utilizing this procedure it has been demonstrated that renal fructose 1,6-diphosphatase was inactivated on incubation with ATP and cyclic-3’,5’-AMP. Fructose 1,6-diphosphatase present. in crude kidney extracts was inactivated by ATP and Mg++ alone, and the rate of this inactivation was increased in the presence of cyclic 3’,5’-AMP. Ptlrified enzyme was inactivat,ed by ATP only in the presence of crude kidney extracts. Fructose 1,6-diphosphatase was also inact,ivated when kidney slices were incubated in the presence of epinephrine. When fructose 1 ,G-diphosphatase was incubated in the presence of an ATa2P generating syst,em in crude extracts, 32P was incorporated into the enzyme protein with a concomitant decrease in enzyme activity. Very little or no a2Pi was incorporated in the absence of adenosine nncleotide. The inactivated enzyme was reactivated on incubation with crude kidney extracts. The possible relationship of these effects to those obtained with three other enzymes involved in the synthesis and degradation of glycogen and the mechanism of action of epinephrine is discussed.
In previous studies with reconstructed gluconeogenic multienzyme systems it was demonstrated that the concentration of fructose 1,6-diphosphate and the activity of fructose 1,6-diphosphatase were ratelimiting in the formation of hexose monophosphate from noncarbohydrate precursors (1, 2). Further studies on fructose 1,6-diphosphatase, isolated from kidney homogenates, showed that the purified enzyme was inhibited by AMP and high concentrations of its substrate, fructose 1,6diphosphate (3). The specific activity of the different fructose 1,6-diphosphatase prep1 Dedicated to Luis F. Leloir on the sixtieth birthday. 2 This invest.igation was supported States Public Health Service grant AM the National Institute of Arthritis and Diseases.
occasion
arations purified during the course of this investigation varied greatly, in spite of the fact that electrophoretic and ultracentrifugal data indicated the same degree of purity for each of these preparations (3). These observations could be explained if the activity of the enzyme was altered “in viva” or during its isolation without a detectable change in the physical properties of the enzyme. This possibility was investigated in kidney extracts. The present communication reports studies on the nature of the enzymic inactivation of fructose 1,6-diphosphatase in kidney extracts in the presence of ATP and cyclic-3’) 5’-AMP. Evidence is presented which indicat#es that fructose 1,6-diphosphatase is inactivated by an ATP- and magnesium-dependent phosphorylation of the enzyme. The inactive enzyme can be
of
his
by United 10341 from Metabolic 436
ItEVERSIBLE
ISACTIVATION
OF
recouvertetl to an active form by incubation wit,h crude kidney ext’racts. It’ appears that the two forms of the enzyme arc interconvertible by the a&ion of a protein kinase and a phospho-protein phosphstase present in kidney extra&s. The enzyme was also inactivated when kidney slices were inc:ubatmed with cpinephrine. The possible relationship of these properties to those rcport,ed for phosphorylase, glycogen synthetJasc, and phosphofructokinase and the mechanism of action of cpinephrine is discussed. EXPERIMENTAL Assam
CJJ’ fructose
PIIOCEI>UI~E: I,&diphosphchse
acli~~ify.
The enzyme was assayed by rltiliaing the methods developed in a previous study (3). The standard proced~lre was based on the determination of Pi formed from fructose-l ,&dip. Except where otherwise indics.ted, the reaction mixtlu+e was incuhated at 30” for 5 minutes and contained in 9 ml: 100 mM Tris-HCl, pH 8.0; 12 mM MgC12; 5 my cysteilie; 0.06 mM fructose-1,6-dip; and an appropriate amount of enzyme. The reaction was stopped with 1 ml of 1057, trichloroacetate acid and an aliquot was removed and assayed for P;. With crttde extracts or preparations containing large amollnts of P, the frllctose l,G-diphosphatase activity was determined by measuring the rate of formation of fructose-6-P from frllctose-l,G-diP (3). The frllct,llse-6-P formed in the reaction u-as assayed spectrophotometrically by measuring the reduction of TPN in the presence of phosphohcxose isomerase and glucose-B-P dehydrogenase (4). One mlit of activity in all cases represents t,he formation of 1 pmole of fructose-6-P or Pi per minute, and specific activity is expressed as rmitr: per milligram of protein. .I ssag 0-f jrztche 1 ,6-diphosphaluxe inac/itding and nctirraling ~eacfions. This assay depends on the ahility of kidney extracts to catalyze the inactivat,ion of frllctose 1 ,Gdiphosphatase when incubated with ilTP and Mg++, or on the ability to catalyze the reactivation of the inactive enzyme when incubated with dialyzed kidney extracts. In order to study the inflrlence of various compounds and incubation conditions on fructose 1,6-diphosphatase in crude extracts, it was necessary to develop methods for the quantitative recovery of t,he enzyme from preincrthation mixtures. It, was not possible to obtain reliahle or consistantlp reproducible results by examining the activity of frl~ct ose 1 ,&diphosphat,ase directly on aliquots of crltde incrtbation mixtures. III previous studies it was ohserved that when either ATP or
U-FRUCTOSE
1 ,G-DIPHOSPHATASE
437
frllct ose-1,6-diP was incubated with crude kidney extracts, very high values for Pi release were obtained (3). The ATP was converted to ADP and AMP hy the act.ion of ATPase and myokinase present in the extracts. Fructose-l,G-diP was converted to triose phosphates in these preparations, and P, was formed by hydrolysis of these compounds hy nonspecific phosphatases or by dephosphorylation of the oxidized products derived from these esters. Moreover, the addition of as lit)tle as 1 X 10e3 M ATP or ATIP to the dialyzed crllde extract increased the nonspecific hydrolysis of fructose-1,6-dip. In addition, the AMP formed diiring preincubation experiments with ATP and magnesillm colrld prodllce a significant error in the subseclllent determination of fructose 1,6-diphosphatase activity, since this nucleotide has heen shown to inhibit the reaction (3). Therefore, it was necessary, especially in experiments involving crude enzyme preparations, to reisolate the fructose 1 ,&diphosphatase before assaying this enzyme. Procedures for the quantitative recovery of fructose l,A-diphosphat,ase from incubation mixtures with an 80-fold purification were developed. The reaction mixtllres from incubat,ion experiments were routinely diluted to 30 ml with 0.05 M Tris-HCl, pH 8.0, at 3” to stop any flu+ther enzymic activity. Experiments in which the enzyme was allowed to remain at 3” for various times showed that the activity of fructose l,A-diphosphatase did not change after this dihltion. The mixtllres were then passed thrcnlgh a cellulose-P column and fructose 1 ,B-diphosphatase was quantitatively elut,ed from the column by the procedure outlined in Table I. The resldts summarized in this Tahle illrlstrate the recovery of the enzyme by this procedure. A dialyzed crrtde kidney extract, was Iised as the source of fructose 1,Gdiphosphatase in t,hese experiments. The total activity of fructose 1,Gdiphosphatase and the amnlmt of protein isolated by this proceclllre were directly proportional t,o the amount of crude extract used, and the specific acbivity of the isolated enzyme was constnnt and independent of the amount of extract added. The crude extract was the only source of fructose 1,6-diphosphatase in experiment,s 1,2, and 3. In order to eliminate the possibilit,y that proportional losses of enzyme had occrlred dllring isolation, varinbls amounts of purified fructose I,& diphosphatase were added to 4 ml of the crude extract, and the enzyme was reisolated hy chromat)ography on cellulose-P. As shown in experiment 4, Table I, a sample containing additional purified enzyme had proportionately more activity, indicating that essentially all of the fruct,ose 1,6diphosphatase activity present in the sample had been recovered in the pllrified fractioll.,
TABLE
I
The crude extract, was prepared by homogenizing fresh swine kidneys it1 ogle vol~une of 0.05 JI TrisHCI, pH 8, and centrifuging the suspension twice at 30,OOOg for 20 minutes. 111 the rspclriments shown in t,he table, the indicated amounts of crude kidney extracts were dilllted to 30 ml with 0.05 RI Tris-WC], pH 8, at 3” and passed into cellulose-P columns (3 X 4 cm). The column was washed with 30 ml of 0.05 M Tris-HCl, pH 8, and then with GO ml of 0.05 M Tris-HCI, pH S-0.1 hr NaCl. These solutions did not contain frllctose 1 ,G-diphosphatnse and were discarded. The enzyme was then qlumtitntivel>removed from the column with 60 ml of 0.05 M Tris-HCl, pH 8-l M NaCl. The resrdts of a typical isolation are summarized below. The purified fructose 1 ,G-diphosphatase added in experiment 4 had an activity of 12 units/ml and a specific activity of 1.8. When incltbation experiments were performed, samples were routinely dilllted to 30 ml wit,h 0.05 M Tris-HCl, pH 8, at 3” and passed into cellulose-P colllmns (3 X 5 cm). Experiment:
Crude extract added (ml) Purified fructose 1,6-diphosphatase Units of fructose 1,6-diphosphat,aee 0.05 M Tris-HCI, pH 8-l .O hf NaCl Mg protein/ml Specific activity (units/mg protein)
added (ml) activity/ml of cellulose-P eluant
PzrriJcalion and concentration of fruclose 1,6diphosphatase. This enzyme was purified as described previously (3). In order to use the purified enzyme as a substrate in the present studies, it was necessary to concentrate the enzyme and store it for long periods of time without, repeated freezing and thawing, since lyophilization or freezing resulted in large losses of activity (3). The following methods were used to concentrate and store the purified enzyme. Solutions were concentrated by dialysis against glycerol at 0”. When the concentration of glycerol inside the dialysis tubing became very high, the sack was placed in a freezer at, -10” with one end in a beaker containing 0.5yc glycerol. Positive pressure was aim-ays maintained on the dialysis tnbiug by placing a heavy weight on it. In this manner, the enzyme was further concentrated and some of the glycerol was removed. The enzyme can be stored in glycerol-water sollltions at -20” for at least a year without appreciable loss of act)ivity. The solution does not freeze under these conditions. Before use, aliquots of the enzyme solution were dialyzed twice for 30 minutes against 3 liters of 0.005 M P-mercaptoethylamine-0.005 M Tris-HCl, pH 8. Preparation of kidney extracfs. Unless otherwise indicated, crude kidney extracts were prepared by homogenizing fresh swine kidney in 1 volume of ice-cold 0.05 111Tris-HCl, pH 8, for 5 minutes in a Waring Blendor. The suspension was centrifuged twice at 30,000 g for 20 minlltes and the super-
1
2
3
4
2.0 -
1.0
G.0
4.0 1 .o
0.101 0.056 1.78
0.195 0.108 1.85
0.295 0.166 1.77
0.390 0.220 1.77
natant fluid was dialyzed for 1 hour against 1 liters of 0.05 M Tris-HCl, pH 8. Kidney cortex tisslle slices were prepared by the procedure of Stadie and Riggs (5). The slices were washed with a 0.151 M NaCl solution at 3”. The amount of tissue was determined by weighing the moist slices. For the study of fructose 1,6-diphosphatase inactivation, incubations were carried out in a Dubnoff metabolic shaker bath under an atmosphere of air at, 37” for 30 minutes. The incubation mixtllre contained, in a final volume of 20 ml: 2 gm of washed kidney slices, 0.154 M NaCl, and the compound being tested. The incltbations were terminated by removing the slices and washing them with cold 0.154 &I NaCl. The slices were immediately homogenized in 15 ml of 0.05 M Tris-HCl, pH 8 at 3”, and the SIISpension was centrifuged at 30,OOOg for 20 minutes. The precipitate was washed with 15 ml of 0.05 M Tris-HCl, pH 8. The combined extracts were then passed into a cellulose-P column and the frllctose 1,6-diphosphatase was isolated and assayed. MaleriaZs. Labeled ATP-+P was prepared by the method of Glynn and Chappell (6). The LiCl which was used to elute the nucleotide from a Dowex-l-Clcolumn was removed from the final preparation by paper chromatography with 85% ethanol, and the specific activity was adjusted to 1 X 10” cpm/per mole. CTP-+P and I-TP-yJ2P were prepared from CDP and UDP, respectively, by incubating these compounds with 10 pmoles of MgCle, 10 pmoles of fructose-l,&diP. z2P;, and a
REVERSIBLE
IXACTWATIOX
INACTIVATION
OF FRITTOSE
OF
D-FRUCTOSE
TABLE
II
1 ,6-DIPHOSPHATASE
l,G-UIPHOSPIIATASE
BY ATP
AND
439
CYCLIC-~',S'-A&~P
The reaciion mixtlxe was incubated at 30” for 5 minutes and contained, in 3 ml: 70 rnhr Tris-HCl, pH 7.5; 5 ml?% 3-P-glycerate; 7 mM cysteine; 8 mM MgCl?; fructose 1,6-diphosphatase, crltde kidney extract,; ATP; and cyclic-3’,5’-AMP as indicated. The reaction was stopped by adding 30 ml of distilled water at 2” alld the dilated solutions were immediately passed into cellulose-P columns. Elution was carried out) as described in the text. The 0.05 M Tris-IICl, pH 8-l M NaCl fraction contained all of the frrrctose 1 ,F-diphosphatase and had a volume of 60 ml. The 0.05 M Tris-HCI, pH SO.1 M NnCl fraction and frnctiolls beyond GO m! of 0.05 >t Tris-HCl, pH 8-l M NaCl contained no activity. The fructose 1, ti-diphosphatase activity was measured by the standard assay procedure. Experiment:
2
1
4
3
3
6
___-
-
Addit ions Prlrified fructose 1,6-diphosphatase Crnde kidney extract (ml) ATP (pmoles) Cyclic-3’,5’-AMP (pmoles) Activity X 102/m1 Protein (mg/ml) Specific act,ivit,y
(ml)
-
1.0 -
1.0 40 -
10.6 O.OG 1.76
dialyzed O-60Tij ammonillm sulfate enzyme fraction from r,zbbit, muscle (7). It was not necessary to add either I>PN or ATP to this system, althollgh the addition of either of these compounds greatly increased the rate of formation of CTP and UTP. The nncleotides were isolated hy ionexchange clhromatography. Inorganic phosphate was estimated by a modificat,ion of the method of Fiske and SuhbaRow (8, 9). In concentrated solutions protein was determined by the method of Lowry el nl. (lo), and in dilllte sollltions protein was assayed by spectrophotometric deiermination (11, 12). RESULTS
I~nactivation of fructose 1 ,6-diphosphatase by L4TP and cyclic-3’,5’-AMP. When kidney extracts were incubated with ATP there was a striking and Cyclic-3 ,5’-ARIP decrease in the activity of fruct’ose 1,6diphosphatase (Table II). Adenosine triphosphate, at a concentration of 14 nmr, was more effect’ive than 0.6 mM cyclic-3’) 5’. Ahll’, and the addition of both compounds resulted in the greatest’ inactivation. The decrease in activity with ATP plus cyclic-3’) 5’-AMP was about 30-fold. However, the amount ‘of protein isolated remained unchanged and therefore the specific: activity of the enzyme isolated in experiments 2, 3, and 4 decreased. It may be that’ the inactivatJed fructose 1,6-diphosphatase is still present in this cellulose-P fraction, since only the
0 .6 O.OG 0.11
1.0 40 1.5 0.4 0.06 0.06
1.0 1.8 2.6 0.05 0.52
0.5 20 1.8 10.5 0.1 1.05
0.5 10.3 0.1 1.03
activity was affected by incubation with ATP, while t’he concentration of protein remained unchanged. In order to exclude the possibility that the chromatographic properties of fructose 1, G-diphosphat’ase had been altered, the other fractions eluted from 6he celIulose-P column were tested for act#ivity. They were found to be inactive. To be certain that A?v:IP, or some other compound capable of inhibiting fructose 1,6-diphosphatase, was not responsible for the effects observed, l-ml aliquots of the 0.05 RI Tris-HCl-1 nr KaCl fraction from experiments 1, 2, and 3 were combined and assayed. The activity of t,he combined sample was 0.122, which was almost the same as the sum of the individual activities in experiments 1, 2, and 3 in Table II. If an inhibitor was present in the inactive fractions from experiments 2 and 3, it would also be expected to inhibit t’he active enzyme from experiment 1. Since little or 110 inhibition was observed when the fract’ions were combined, it was likely that the inactivation obtained with ATP resulted from an alteration of t,he enzyme. Similar results were obtained when purified fruct,ose 1 ,6-diphosphatase was incubated with diluted kidney ext’racts, ATE’, and cyclicHowever, these compounds 3’ ,5’-AMP. had uractically no effect on purified fructose
440
RlENI>lCIPU’O, TABLE
INFLI;EWE
OF
NUCLEOSIDE VATION
ATP
ASI)
III
C~X~ES,PRAUON
TRIPHOSPHATES OF FRL-WOSE
BlSAITl)RJZ.4U,
AKD ON
‘I’HE
OTHER
T,u,zc:~vI-
1 ,(i-I)II’HOSPHA’l’ASE
The assay of fructose l,G-diphosphatssc :uld the inactivation reaction were carried ollt as described in E.rpe,.inlentul Pwcadwe. The reaction mixtrlre was inclthated al. 30” for 5 minrltes and contained, in 1.5 ml: 50 m&r Tris-HCl, pH 7.5; 1 IIIM M&l,; 5 mu cqstrine; nucleoside triphosphate as indicated in the table; and 1 ml of dialyzed kidney extract. The percentage inactivation, in each case, was obtained hv dividing the decrease in activity after preincnbation in the presence of the nucleoside triphosphate by the activity in the absence of added nucleotide. The activities were calcldatetl on the basis of the 1 ml of kidney extract, added. Fructose 1,6-diphosphatase activity (units/ml)
Additions
Inac$;tion -
None CTP, crrP, GTP, ATP, ATP, ATP,
5 5 5 5 3 2
X 1OF M
5.0 -1.5
x
lo-’
M
4.9
X X X X
1O-3 1O-3 1O-3 lop3
M
1.7 1.4 3.5 4.5
M M M
0 10 2 G 72 30 10
I .Miphosphatase in the absence of t’he crude kidney extracts as shown in expcriments .5 and 6 in Table II. Efect of other nucleoside triphosphates and the conceniration of ATP on the inactivation of fructose 1,6-diphosphatase. Other compounds were test’ed to see whether they would replace ATP in the syst,em inactivating fructose 1 ,6-diphosphatase. At concentrations of .5 X 10H3 M, GTP, and UTP had very little or no effect (Table III). Many other nucleotides including AMP, IMP, CMP, GMP, IDP, CDP, GDP, TDP, dCMP, adenosine, cytosine, and guanosine were inactive. High concentrations of ADP caused some inactivation, but this was due to the formation of ATP by the myokinase present in crude kidney extracts. The influence of ATP concentration on the rate of inactivation of fructose 1,6-diphosphatase is shown in Table III. The rate of inactivation decreased very rapidly with decreasing ATP concentration. However, the rate was considerably higher when an ATP regenerating system was present, since
J~IIATTA(:I~AI1~~~,\
ATP is very quic~kly hytlrolyzc~tl to ;\T>P and iZ;\lP in this system. l’wpwties 04’ inactivatin~q s$dfw. The inactivating system slro~d an :lbsolutc rcquiremclnt for 1\Ig++ and the apparent pH optimunl was at, 7.5. L4s shown in Table IV the inactivation of purified fructose 1 ,6-diphosphntast~ required t,hc addition of cruclc kidney cstrart,. Dialysis of ihe crude ext’ract had 110 effect OII t’he inac+tivation of fructose 1 , Miphosphatnse, but boiled dialyzed extract was inactive. JIoreovor, inactivation of purified frurtose 1,6-tliphosphat~ase was found to bc dependent 011 the amount of crude kidney cxt’ract added. InJluence oj” epinephrine on j’wcfose 1,6diphosphatase activity. The possible relationship of the fructose 1,6-diphosphatnse inactivating system in crude kidney extracts to the influence of cpinephrine on glpcolysis in t,he intact cell was investigat’cd. Epinephrine or glucagon would be expected to inactivate fructose 1,6-diphosphatasc, since administrat,ion of these compounds has been shown to result in the formation of cyclic-3 , ;5’-AMP (1.3, 14). This effect should stimulate the inactivat,ion of fructose 1,6-diphosphnt,ase. This possibility was supported by t,he results of experiments in which intracellular fructose 1 ,6-diphosphatase was inachivatcd when kidney cortex slices were incubated with rpinephrine. As shown in Table Y incubation with epinephrine resulted in almoat complete inactivation of fructose 1, Ci-diphosphatnse. There was some inactivation even wit#hout added epinephrinc, and this may represent an enclogenous inactivation of this enzyme. It should be noted that’ the experiment shown in Table V was not typical of all the experiments undertaken. The extent of inactivation of fructose 1,6-diphosphatase was dependent OII the act’ivity of this enzyme in the particular tissue preparation used. In general, tissue from kidneys with higher levels of enzyme showed greater decreaseswhen incubated with epinephrine. Mechanism of inactivation of fiucfose 1,6diphosphataseby ATP. It was possible that ATP inactivated fructose 1,6-diphosphatase by adsorption at a regulatory site, which appears to be distinct from the catalytic
REVERSIBLE
ISACTIVATION
OF
u-FRUCTOSE
TABLE PROPERTIES
OF
THE
SYSTEM
IV
INACTIVATING
FRUCTOSE
1 ,6-DIPHOSPHATASE
The reaction mixture was incubated at 30” for 5 minutes and contained, pH 7.5; 8 IriM cysteine; 8 mM MgClz; and when indicated, 10 mM ATP and boiled extract,. The pllrified fructose 1,6-diphosphatase preparation added activity. The inactivating reaction was assayed as described in the text.
in 1.5 ml: 60 mM Tris-HCI, 1 ml of crude, dialyzed, contained about 2 units
Fructose Incubation
+ Crude extract $ ATP + crude
conditions
extract crude
2.64 0.54
extract
+ fructose 1,6-diphosphatase extract + fructose l,A-diphosphatase
4.2 0.98
$- Boiled dialyzed $- ATP f boiled
extract dialyzed
2.12 2.09
-+ fructose 1,6-diphosphatase extract + fructose 1,6-diphosphatase
TABLE OF
FRUCTOSE
V
1,6-I)IPHOSI?HATASE ~VITH
The cedure. kidney
1,6-diphosphatase activity (units)
$- Crude extract $- ATP + crude
INAc’TIVATION
WHEN
KIDNEY
SLICES
ARE
INCUBATED
EPINEPHRINE
incubation conditions and enzyme assays were carried out as described in Ezperinlenlal Each vessel was incubated for 1 hour at 37” and contained, in 20 ml: 0.154 M NaCl and slices. The control vessel was kept at 3’ and epinephrine was added as indicated. Additions
None (without incubation None (incubated at 37”) Epinephrine (50 @g/ml) Epinephrine (10 fig/ml) Epinephrine (5 pg/ml)
or of
2.67 0.66
extract
+ Dialyzed crude $- ATP + dialyzed
441
1.6.DIPHOSPHATASE
at 3”)
site and is inaccessible to other nucleoside triphosphates (15, 16). Thus, an allosteric type of inhibition might be involved in which the presence of ATP altered the shape of the enzyme. When the enzyme was incubated in the presence of crude kidney extracts with AT32P and ATP S-14C, only 9 emerged from the cellulose-P column with the enzyme peak, indicating that an irreversible reaction of ATP with the enzyme had taken place to form a modified inactive enzyme. The 32P-labeled protein could be precipitated with 50 % ammoniunl sulfate or dialyzed wit’hout any significant 10s~of radioactivity. The best incorporations of 32P into t,he
Pro-
2 gm of
Fructose l,h-diphosphatase activity (umts/gm wet weight of kidney)
Specific activity of isolated fructose l,6-diphosphatase (units/mg protein)
1.71 1.14 0.12 0.34 0.48
1.7 1.1 0.1 0.3 0.5
enzyme were obtained with an AT32P generating system, since as indicated earlier ATP was rapidly hydrolyzed in crude kidney extracts. When the regenerating system was used high concentrations of ATn3P were maintained throughout the incubation period. This system depended on the formation of ATz3P from ADP and 32Pi by the action of aldolase, triosephosphate dehydrogenase, and 1,3 dip-glycerate kinase present in the crude kidney extracts. Fructose 1,6-diphosphate was employed as the substrate for the ATP regenerating system. As shown in Fig. 1 when AT33P was present in the reaction mixture, the decrease in activity of fructose 1,6-diphosphatase was
442
MENDICINO.
REAUDREAU,
0.7
-3
AND
BHATTACIIAII1-YA
7
0.6-
-6
f
, WITH / EXTRACT
ML
OF
ELUANT
FIG. 1. Incorporation
of 32Pi into fructose 1,6diphosphatase in the presence of AT32P and isolation of azP,-labeled fructose 1,6-diphosphatase on cellulose-P. The reaction mixtures were incubated at 30” for 20 minutes and contained, in 13 ml: 25 mM Tris-HCl, pH 7.5; 15 mM cysteine, pH 7.5; 12 mM MgClz; 1 mC z2Pi; and 10 ml of dialyzed kidney extract. The reaction mixture designated “+ATP” also contained 7 mM ADP and 15 mM fructose 1,6-diphosphate. The ot,her incubation mixture designated “-ATP” was incubated in the absence of an AT32P generating system. The elution method described in Ex~e~inLental Procedure was used.
accompanied by a concomitant incorporation of 32P into this enzyme fraction. The radioactivity emerging from the cellulose-P column on elution with 0.05 M Tris-HCl, pH 8-l M NaCl was coincident with the peak of enzymatic activity in each case. The enzyme was not inactivated in the absenceof ATP and very little 32P was incorporated into this enzyme fraction in the absence of adenosinenucleotides as shown in the curves designated “ -ATP”. The figure showsonly the final fractions of the chromatogram cont.aining fructose 1,6-diphosphatase activity. AT32P and 32Pwere completely and rapidly eluted from this cation exchange column in the earlier fractions. When UT32P and CT3”P were incubated
o.70”-pm
L 20
-L-L--J 40 MINUTES
60
60
FIG. 2. Reactivation of inactivated fructose 1,6-diphosphatase by crude kidney extracts. The complete reaction mixt,ure contained, in 10 ml: 60 mM Tris-HCl, pH 7.5; 12 mM MgClz; 0.25 ml of crude kidney extract,, and 2 ml (about 1 unit of activity) of inactivated fructose 1 ,B-diphosphatase. The inactive enzyme was prepared by concentrating and dialyzing the 0.05 M Tris-HCl, pH 8-1 M NaCl fraction obt,ained from a cellulose-P column after applying a reaction mixtlu-e COIItaining crude kidney extract which had been incubated with ATP and Mg++. No react,ivation was observed in the absence of either inactivated fructose 1,6-diphosphatase or crude kidney extract.
with crude kidney extract’s in the presence of 32Pi and a regenerating system (7), fructose 1,6-diphosphat’ase was not inactivated and very little 32P was incorporated into the fructose 1,6-diphosphatase fraction eluted from the cellulose-P column. Reactivation of inactive fructose I ,&diphosphataseby kidney extracts. Experiments were performed to determine whether an enzyme in kidney extracts might convert inactive fructose 1,6-diphosphatase to an active enzyme. The results summarized in Fig. 2 show that when purified inactive fructose 1,6-diphosphatase was reincubated
REVERW3LE
INACTIVATION
OF
with crude kidney extracts a marked reactivation of the enzyme occurred. The increase in the activity of fructose 1 ,Gdiphosphat’ase was accompanied by a decrease in the content of 3flP bound to the protein. Thus it would appear t’hat the inactivation caused by incubation with ATP and iVlg++ can be reversed by other enzymes present in crude kidney homogenates.
Kidney and liver are the main sites of gluconeogenesis in mammals, and the metabolism and regulation of glycogen and glucose Imetabolism in these tissues are closely related to the energy requirements of animal l&sues. Moreover, the rates of glycolysis and gluconeogenesis are sensitive to the needs of tbe organism, and the overall rates of these two processes have been shown to vary widely as a result of hormonal changes, exercise, diet and the presence of pathological conditions. The mechanism by which a .hormone acts to regulate an entire pathway involving many enzymes is not yet clearly understjood, although present evi dcnce da’es indicate that multienzyme systems are regulated at specific rate limiting stages. There are two sites in which t’he synthesis of glycogen is thought to differ from the breakdown of glycogen in liver and kidney, the principal gluconeogenic organs in mammals. The reactions at these two steps are catalyzed by four different enzymes, which are all present in the soluble cytoplasmir fraction Iof the liver and kidney cell. These reactions are illustrated by the following equations. Fructose-Ii-1’
+
Phosphofructokinase
ATI’
(1)
Fructose-l Fruct,ose-1 Fructose
,(i-diP
) Fructose G-P + I’hosphorylase
+
+ ADP
+ Hz0
1, F-diphosphatase
Glycogen,,,+1)
,G-diP
(2) Pi
,
I’,
(3)
Glucose-l-I’ UDP-D-glucose Glyrogen
+ Glycogetq,,
$ Glycogen,,, synt11etnse
(4) ) UDP
+
Glycogenl,+,,
D-FRUCTOSE
1 ,B-DIPHOSPHATASE
443
Fructose 1,6-diphosphatase and glycogen synthetase act in the direction of synthesis of glycogen, and phosphorylase and phosphofructokinase are active in Dhe breakdown of glycogen. It is evident that both sets of enzymes cannot operate simultaneously since the net result would be a substrate-mediated hydrolysis of ATP and UDP-n-glucose to ADP and P 1 and glucose-l-P and UDP, respectively. In t#he present investigation experiments were carried out to elucidate t’he mechanisms involved in the regulation of the activity of fructose 1,6-diphosphatasc in kidney cells. The evidence obtained suggests that those factors which stimulate glycogen breakdown may also function to inactivate fructose 1,6-diphosphatase. The experiments reported clearly demonstrated the marked effect of ATP and cyclic3’ ,5’-AMP upon the activity of renal Dfructose 1,6-diphosphatase in t’he presence of crude kidney extracts. Two distinct effects have been observed, one causing inactivation of the enzyme in the presence of ATP and Mg++ and the other causing a rcactivation of the inactive enzyme by an enzyme present’ in crude kidney extracts. This latter enzyme may function to reactivate fructose 1,6-diphosphatasc after the release of glycolytic stress. A similar response of the inactivating reaction to cpinephrine in its intracellular site was consistent with factors regulating the activity of fructose 1,6diphosphatase in t(he intact cell. Epinephrine exerts it’s glycolytic effect by increasing t#he concentration of active muscle phosphorylase. i2n activating and inactivating enzyme catalyzed the interconversion of 2 forms of phosphorylase. Other studies with liver phosphorylase indicated that a similar mechanism was involved, except that there was no increase in the molecular weight of the active form of phosphorylase (17). The activating enzyme in each case was a kinase which catalyzed the phosphorylation of inact,ive phosphorylase by ATP. Heart and liver protein kinases also acted on muscle phosphorylase. Recent results indicate that epinephrine influences this system by increasing the formation of cyclic-3’,5’-AlIP from ,4TP
444
MENDICINO,
Inactive Active Active
Phosphorylaled glycogen synthetase
BEAUDREAU,
AND
BHATTACHARYYA
enz,ymes
Phosphorylase Phosphofrtlctokinase
Protein
kinase
st,imulat,es
Active Active Inactive Inactive
glycolysis I)IAGIIAM
(13, 14). The inactivating enzyme was a protein phosphat’ase which catalyzed the release of Pi from the active forms of phosphorylase. This enzyme had a broad specificity and dephosphorylsted other phosphoproteins but did not hydrolyze nonprotein phosphate esters. Likewise, more recent studies have indicated that the related enzyme, glycogen synthetase, is inactivated by conditions which activate phosphorylase. Incubation of muscle tissue preparations with epinephrine or homogenates with ATP and cyclic-3’,5’AMP resulted in a decrease in glycogen synthetase activity (1s). Other studies with partly purified enzyme preparations also showed that incubation with ATP could decrease the activity of glycogen synthetase (19). It has been suggested that this enzyme exists in an act’ive and inactive form (20). At the fructose 1,6-diphosphate level similar results have been obtained for phosphofructokinase (21, 22). The evidence thus far indicates that epinephrine may exert its regulatory effects by simultaneously stimulating and inactivating appropriate enzyme reactions involved in the degradation and synthesis of glycogen. In conclusion, it may be tentively postulated that epinephrine stimulates the formation of intracellular cyclic-3’, 5’-AMP at the cell wall. In the presence of this compound an intracellular protein kinase may then catalyze the phosphorylation and activation of phosphorylase and phosphofructokinase and the inactivation of glycogen synthetase and fructose 1 ,6-diphosphatase as shown in the following Diagram A. This would be expected to be an active and very rapid process. It is interesting that such a
Dephosphor~glaied enqpnes glycogeu spn t hetax. fructose 1,Gdiphosphalasc phosphorylaae phosphofrnctclkitl:Iso
A
mechanism would not require the hormone to enter the cell, and indeed the steady state level of hormone in extracellular fluid and blood plasma could be regulated by endocrine tissues which would then control the breakdown of carbohydrate in other organs. A decrease in the concentration of hormone in the extracellular fluid, in conjunction with the destruction of cyclic-3’, 5’-AMP in the cell and activation of phosphoprotein phosphatase, would dephosphorylate the four enzymes and restore a glyconeogenie poise in the liver and kidney cells. The specific and simultaneous action of other enzymes on these four enzymes involved at two rate-limiting steps in carbohydrate metabolism and the possible relationship of this system t’o the formation of cyclic3 ,5’-AMP from ATP by epinephrine at the cell wall remains to be established. One of the difficulties in postulating a plausible role for the phosphoprotein phosphatasc and protein kinase may lie in the fact t,hat the activity and enzymic properties of these enzymes must also be changed as a result of the interaction of epinephrine at t,he cell wall, since if bot,h of these enzymes remained active they would catalyze a protein-dependent hydrolysis of ATP. l
IREVERSIBLE
INACTIVATION
OF
5. STADIE, W. C., AND RIGGS, B. C., J. Biol. Chena.164,687 (1944). G. GLYNN, I. M., AND CHAPPELL, J. B., Biochem. J. 90, 147 (1964). 7. MENDICINO, J., J. Biol. Chem. 237, 165 (1962). 8. FISKE, (C. H., AND SUBBAROIV, Y., J. Biol. Chem. 66, 375 (1925). 9. L~HKINN, K., AND JENR.~SRIK, L., Biochena. J. 178, 419 (1926). 10. Lousy, 0. II., ROSEBROUGH, N. J., FARR, A. L., ARD RANDALL, R. J., J. Biol. Chem. 193, 265 (1951). 11. W.\RBUI~G, O., AND CHRISTIAN, W., Biochem. z. 310, 384 (1941). 12. LXYNE, E., in “Methods in Enzymology” (S. P. Colowick and N. 0. Kaplan, (eds.), Vol. III, p. 447. Academic Press, New York (1957:. 13. RALL, T. W., SUTHERLAND, E. W., AND BERTCIET, J., J. Biol. Chem. 224, 463 (1957).
U-FRUCTOSE
1,6-DIPHOSPHATASE
445
14. SUTHERLAND, E. W., AND RALL, T. W., J. Biol. Chem. 232, 1077 (1958). 15. MENDICINO, J., AND MUNTZ, J. A., J. Biol. Chem. 233, 178 (1958). 16. HURWITZ, J., RAPPEL, L. A., AND HORECKER, B. L., J. Biol. Chem. 226, 525 (1957). 17. RALL, T. W., SUTHERLAND, E. W., AND WOSIL.IIT, W. D., J. Biol. Chem. 218, 483 (1956). 18. BELOCOPITO~, E., Arch. Bio&em. Biophys. 93, 457 (1961). 19. TRAUT, R. R., AND LIPMANN, F., J. Biol. Chem. 238, 1213 (1963). 20. ROSELL-PEREZ, M., VILL.U+PALASI, C., AND LARNER, J., Biochemistry 1, 763 (1962). 21. PASSENNEAU, J. V., AND LOTVRY, 0. H., Biothem. Biophys. Research Common. 7, 10 (1962). 22. MANSOUR, T. E., J. Biol. Chem. 240, 2165 (1965).
OF
BIOCHEMISTRY
Reversible
.lND
Inactivation Adenosine
116,
BIOI’HTSICS
Department
(1966)
of D-Fructose Triphosphate
Adenosine JOSEPH
436-445
MENDICINO, RABINDRA of Biochemistry,
1,6-Diphosphatase and
Cyclic
by
3’, 5’-
Monophosphate”’ CHARLES BEAUDREAU, N. BHATTACHARYYA The Ohio
Received
May
Slate
University,
Columbus,
AND
Ohio
19, 1966
An analytical method for the quantitative recovery and purification of fructose l,G-diphosphatase from incubation mixtures containing crude kidney extracts has been described. Utilizing this procedure it has been demonstrated that renal fructose 1,6-diphosphatase was inactivated on incubation with ATP and cyclic-3’,5’-AMP. Fructose 1,6-diphosphatase present. in crude kidney extracts was inactivated by ATP and Mg++ alone, and the rate of this inactivation was increased in the presence of cyclic 3’,5’-AMP. Ptlrified enzyme was inactivat,ed by ATP only in the presence of crude kidney extracts. Fructose 1,6-diphosphatase was also inact,ivated when kidney slices were incubated in the presence of epinephrine. When fructose 1 ,G-diphosphatase was incubated in the presence of an ATa2P generating syst,em in crude extracts, 32P was incorporated into the enzyme protein with a concomitant decrease in enzyme activity. Very little or no a2Pi was incorporated in the absence of adenosine nncleotide. The inactivated enzyme was reactivated on incubation with crude kidney extracts. The possible relationship of these effects to those obtained with three other enzymes involved in the synthesis and degradation of glycogen and the mechanism of action of epinephrine is discussed.
In previous studies with reconstructed gluconeogenic multienzyme systems it was demonstrated that the concentration of fructose 1,6-diphosphate and the activity of fructose 1,6-diphosphatase were ratelimiting in the formation of hexose monophosphate from noncarbohydrate precursors (1, 2). Further studies on fructose 1,6-diphosphatase, isolated from kidney homogenates, showed that the purified enzyme was inhibited by AMP and high concentrations of its substrate, fructose 1,6diphosphate (3). The specific activity of the different fructose 1,6-diphosphatase prep1 Dedicated to Luis F. Leloir on the sixtieth birthday. 2 This invest.igation was supported States Public Health Service grant AM the National Institute of Arthritis and Diseases.
occasion
arations purified during the course of this investigation varied greatly, in spite of the fact that electrophoretic and ultracentrifugal data indicated the same degree of purity for each of these preparations (3). These observations could be explained if the activity of the enzyme was altered “in viva” or during its isolation without a detectable change in the physical properties of the enzyme. This possibility was investigated in kidney extracts. The present communication reports studies on the nature of the enzymic inactivation of fructose 1,6-diphosphatase in kidney extracts in the presence of ATP and cyclic-3’) 5’-AMP. Evidence is presented which indicat#es that fructose 1,6-diphosphatase is inactivated by an ATP- and magnesium-dependent phosphorylation of the enzyme. The inactive enzyme can be
of
his
by United 10341 from Metabolic 436
ItEVERSIBLE
ISACTIVATION
OF
recouvertetl to an active form by incubation wit,h crude kidney ext’racts. It’ appears that the two forms of the enzyme arc interconvertible by the a&ion of a protein kinase and a phospho-protein phosphstase present in kidney extra&s. The enzyme was also inactivated when kidney slices were inc:ubatmed with cpinephrine. The possible relationship of these properties to those rcport,ed for phosphorylase, glycogen synthetJasc, and phosphofructokinase and the mechanism of action of cpinephrine is discussed. EXPERIMENTAL Assam
CJJ’ fructose
PIIOCEI>UI~E: I,&diphosphchse
acli~~ify.
The enzyme was assayed by rltiliaing the methods developed in a previous study (3). The standard proced~lre was based on the determination of Pi formed from fructose-l ,&dip. Except where otherwise indics.ted, the reaction mixtlu+e was incuhated at 30” for 5 minutes and contained in 9 ml: 100 mM Tris-HCl, pH 8.0; 12 mM MgC12; 5 my cysteilie; 0.06 mM fructose-1,6-dip; and an appropriate amount of enzyme. The reaction was stopped with 1 ml of 1057, trichloroacetate acid and an aliquot was removed and assayed for P;. With crttde extracts or preparations containing large amollnts of P, the frllctose l,G-diphosphatase activity was determined by measuring the rate of formation of fructose-6-P from frllctose-l,G-diP (3). The frllct,llse-6-P formed in the reaction u-as assayed spectrophotometrically by measuring the reduction of TPN in the presence of phosphohcxose isomerase and glucose-B-P dehydrogenase (4). One mlit of activity in all cases represents t,he formation of 1 pmole of fructose-6-P or Pi per minute, and specific activity is expressed as rmitr: per milligram of protein. .I ssag 0-f jrztche 1 ,6-diphosphaluxe inac/itding and nctirraling ~eacfions. This assay depends on the ahility of kidney extracts to catalyze the inactivat,ion of frllctose 1 ,Gdiphosphatase when incubated with ilTP and Mg++, or on the ability to catalyze the reactivation of the inactive enzyme when incubated with dialyzed kidney extracts. In order to study the inflrlence of various compounds and incubation conditions on fructose 1,6-diphosphatase in crude extracts, it was necessary to develop methods for the quantitative recovery of t,he enzyme from preincrthation mixtures. It, was not possible to obtain reliahle or consistantlp reproducible results by examining the activity of frl~ct ose 1 ,&diphosphat,ase directly on aliquots of crltde incrtbation mixtures. III previous studies it was ohserved that when either ATP or
U-FRUCTOSE
1 ,G-DIPHOSPHATASE
437
frllct ose-1,6-diP was incubated with crude kidney extracts, very high values for Pi release were obtained (3). The ATP was converted to ADP and AMP hy the act.ion of ATPase and myokinase present in the extracts. Fructose-l,G-diP was converted to triose phosphates in these preparations, and P, was formed by hydrolysis of these compounds hy nonspecific phosphatases or by dephosphorylation of the oxidized products derived from these esters. Moreover, the addition of as lit)tle as 1 X 10e3 M ATP or ATIP to the dialyzed crllde extract increased the nonspecific hydrolysis of fructose-1,6-dip. In addition, the AMP formed diiring preincubation experiments with ATP and magnesillm colrld prodllce a significant error in the subseclllent determination of fructose 1,6-diphosphatase activity, since this nucleotide has heen shown to inhibit the reaction (3). Therefore, it was necessary, especially in experiments involving crude enzyme preparations, to reisolate the fructose 1 ,&diphosphatase before assaying this enzyme. Procedures for the quantitative recovery of fructose l,A-diphosphat,ase from incubation mixtures with an 80-fold purification were developed. The reaction mixtllres from incubat,ion experiments were routinely diluted to 30 ml with 0.05 M Tris-HCl, pH 8.0, at 3” to stop any flu+ther enzymic activity. Experiments in which the enzyme was allowed to remain at 3” for various times showed that the activity of fructose l,A-diphosphatase did not change after this dihltion. The mixtllres were then passed thrcnlgh a cellulose-P column and fructose 1 ,B-diphosphatase was quantitatively elut,ed from the column by the procedure outlined in Table I. The resldts summarized in this Tahle illrlstrate the recovery of the enzyme by this procedure. A dialyzed crrtde kidney extract, was Iised as the source of fructose 1,Gdiphosphatase in t,hese experiments. The total activity of fructose 1,Gdiphosphatase and the amnlmt of protein isolated by this proceclllre were directly proportional t,o the amount of crude extract used, and the specific acbivity of the isolated enzyme was constnnt and independent of the amount of extract added. The crude extract was the only source of fructose 1,6-diphosphatase in experiment,s 1,2, and 3. In order to eliminate the possibilit,y that proportional losses of enzyme had occrlred dllring isolation, varinbls amounts of purified fructose I,& diphosphatase were added to 4 ml of the crude extract, and the enzyme was reisolated hy chromat)ography on cellulose-P. As shown in experiment 4, Table I, a sample containing additional purified enzyme had proportionately more activity, indicating that essentially all of the fruct,ose 1,6diphosphatase activity present in the sample had been recovered in the pllrified fractioll.,
TABLE
I
The crude extract, was prepared by homogenizing fresh swine kidneys it1 ogle vol~une of 0.05 JI TrisHCI, pH 8, and centrifuging the suspension twice at 30,OOOg for 20 minutes. 111 the rspclriments shown in t,he table, the indicated amounts of crude kidney extracts were dilllted to 30 ml with 0.05 RI Tris-WC], pH 8, at 3” and passed into cellulose-P columns (3 X 4 cm). The column was washed with 30 ml of 0.05 M Tris-HCl, pH 8, and then with GO ml of 0.05 M Tris-HCI, pH S-0.1 hr NaCl. These solutions did not contain frllctose 1 ,G-diphosphatnse and were discarded. The enzyme was then qlumtitntivel>removed from the column with 60 ml of 0.05 M Tris-HCl, pH 8-l M NaCl. The resrdts of a typical isolation are summarized below. The purified fructose 1 ,G-diphosphatase added in experiment 4 had an activity of 12 units/ml and a specific activity of 1.8. When incltbation experiments were performed, samples were routinely dilllted to 30 ml wit,h 0.05 M Tris-HCl, pH 8, at 3” and passed into cellulose-P colllmns (3 X 5 cm). Experiment:
Crude extract added (ml) Purified fructose 1,6-diphosphatase Units of fructose 1,6-diphosphat,aee 0.05 M Tris-HCI, pH 8-l .O hf NaCl Mg protein/ml Specific activity (units/mg protein)
added (ml) activity/ml of cellulose-P eluant
PzrriJcalion and concentration of fruclose 1,6diphosphatase. This enzyme was purified as described previously (3). In order to use the purified enzyme as a substrate in the present studies, it was necessary to concentrate the enzyme and store it for long periods of time without, repeated freezing and thawing, since lyophilization or freezing resulted in large losses of activity (3). The following methods were used to concentrate and store the purified enzyme. Solutions were concentrated by dialysis against glycerol at 0”. When the concentration of glycerol inside the dialysis tubing became very high, the sack was placed in a freezer at, -10” with one end in a beaker containing 0.5yc glycerol. Positive pressure was aim-ays maintained on the dialysis tnbiug by placing a heavy weight on it. In this manner, the enzyme was further concentrated and some of the glycerol was removed. The enzyme can be stored in glycerol-water sollltions at -20” for at least a year without appreciable loss of act)ivity. The solution does not freeze under these conditions. Before use, aliquots of the enzyme solution were dialyzed twice for 30 minutes against 3 liters of 0.005 M P-mercaptoethylamine-0.005 M Tris-HCl, pH 8. Preparation of kidney extracfs. Unless otherwise indicated, crude kidney extracts were prepared by homogenizing fresh swine kidney in 1 volume of ice-cold 0.05 111Tris-HCl, pH 8, for 5 minutes in a Waring Blendor. The suspension was centrifuged twice at 30,000 g for 20 minlltes and the super-
1
2
3
4
2.0 -
1.0
G.0
4.0 1 .o
0.101 0.056 1.78
0.195 0.108 1.85
0.295 0.166 1.77
0.390 0.220 1.77
natant fluid was dialyzed for 1 hour against 1 liters of 0.05 M Tris-HCl, pH 8. Kidney cortex tisslle slices were prepared by the procedure of Stadie and Riggs (5). The slices were washed with a 0.151 M NaCl solution at 3”. The amount of tissue was determined by weighing the moist slices. For the study of fructose 1,6-diphosphatase inactivation, incubations were carried out in a Dubnoff metabolic shaker bath under an atmosphere of air at, 37” for 30 minutes. The incubation mixtllre contained, in a final volume of 20 ml: 2 gm of washed kidney slices, 0.154 M NaCl, and the compound being tested. The incltbations were terminated by removing the slices and washing them with cold 0.154 &I NaCl. The slices were immediately homogenized in 15 ml of 0.05 M Tris-HCl, pH 8 at 3”, and the SIISpension was centrifuged at 30,OOOg for 20 minutes. The precipitate was washed with 15 ml of 0.05 M Tris-HCl, pH 8. The combined extracts were then passed into a cellulose-P column and the frllctose 1,6-diphosphatase was isolated and assayed. MaleriaZs. Labeled ATP-+P was prepared by the method of Glynn and Chappell (6). The LiCl which was used to elute the nucleotide from a Dowex-l-Clcolumn was removed from the final preparation by paper chromatography with 85% ethanol, and the specific activity was adjusted to 1 X 10” cpm/per mole. CTP-+P and I-TP-yJ2P were prepared from CDP and UDP, respectively, by incubating these compounds with 10 pmoles of MgCle, 10 pmoles of fructose-l,&diP. z2P;, and a
REVERSIBLE
IXACTWATIOX
INACTIVATION
OF FRITTOSE
OF
D-FRUCTOSE
TABLE
II
1 ,6-DIPHOSPHATASE
l,G-UIPHOSPIIATASE
BY ATP
AND
439
CYCLIC-~',S'-A&~P
The reaciion mixtlxe was incubated at 30” for 5 minutes and contained, in 3 ml: 70 rnhr Tris-HCl, pH 7.5; 5 ml?% 3-P-glycerate; 7 mM cysteine; 8 mM MgCl?; fructose 1,6-diphosphatase, crltde kidney extract,; ATP; and cyclic-3’,5’-AMP as indicated. The reaction was stopped by adding 30 ml of distilled water at 2” alld the dilated solutions were immediately passed into cellulose-P columns. Elution was carried out) as described in the text. The 0.05 M Tris-IICl, pH 8-l M NaCl fraction contained all of the frrrctose 1 ,F-diphosphatase and had a volume of 60 ml. The 0.05 M Tris-HCI, pH SO.1 M NnCl fraction and frnctiolls beyond GO m! of 0.05 >t Tris-HCl, pH 8-l M NaCl contained no activity. The fructose 1, ti-diphosphatase activity was measured by the standard assay procedure. Experiment:
2
1
4
3
3
6
___-
-
Addit ions Prlrified fructose 1,6-diphosphatase Crnde kidney extract (ml) ATP (pmoles) Cyclic-3’,5’-AMP (pmoles) Activity X 102/m1 Protein (mg/ml) Specific act,ivit,y
(ml)
-
1.0 -
1.0 40 -
10.6 O.OG 1.76
dialyzed O-60Tij ammonillm sulfate enzyme fraction from r,zbbit, muscle (7). It was not necessary to add either I>PN or ATP to this system, althollgh the addition of either of these compounds greatly increased the rate of formation of CTP and UTP. The nncleotides were isolated hy ionexchange clhromatography. Inorganic phosphate was estimated by a modificat,ion of the method of Fiske and SuhbaRow (8, 9). In concentrated solutions protein was determined by the method of Lowry el nl. (lo), and in dilllte sollltions protein was assayed by spectrophotometric deiermination (11, 12). RESULTS
I~nactivation of fructose 1 ,6-diphosphatase by L4TP and cyclic-3’,5’-AMP. When kidney extracts were incubated with ATP there was a striking and Cyclic-3 ,5’-ARIP decrease in the activity of fruct’ose 1,6diphosphatase (Table II). Adenosine triphosphate, at a concentration of 14 nmr, was more effect’ive than 0.6 mM cyclic-3’) 5’. Ahll’, and the addition of both compounds resulted in the greatest’ inactivation. The decrease in activity with ATP plus cyclic-3’) 5’-AMP was about 30-fold. However, the amount ‘of protein isolated remained unchanged and therefore the specific: activity of the enzyme isolated in experiments 2, 3, and 4 decreased. It may be that’ the inactivatJed fructose 1,6-diphosphatase is still present in this cellulose-P fraction, since only the
0 .6 O.OG 0.11
1.0 40 1.5 0.4 0.06 0.06
1.0 1.8 2.6 0.05 0.52
0.5 20 1.8 10.5 0.1 1.05
0.5 10.3 0.1 1.03
activity was affected by incubation with ATP, while t’he concentration of protein remained unchanged. In order to exclude the possibility that the chromatographic properties of fructose 1, G-diphosphat’ase had been altered, the other fractions eluted from 6he celIulose-P column were tested for act#ivity. They were found to be inactive. To be certain that A?v:IP, or some other compound capable of inhibiting fructose 1,6-diphosphatase, was not responsible for the effects observed, l-ml aliquots of the 0.05 RI Tris-HCl-1 nr KaCl fraction from experiments 1, 2, and 3 were combined and assayed. The activity of t,he combined sample was 0.122, which was almost the same as the sum of the individual activities in experiments 1, 2, and 3 in Table II. If an inhibitor was present in the inactive fractions from experiments 2 and 3, it would also be expected to inhibit t’he active enzyme from experiment 1. Since little or 110 inhibition was observed when the fract’ions were combined, it was likely that the inactivation obtained with ATP resulted from an alteration of t,he enzyme. Similar results were obtained when purified fruct,ose 1 ,6-diphosphatase was incubated with diluted kidney ext’racts, ATE’, and cyclicHowever, these compounds 3’ ,5’-AMP. had uractically no effect on purified fructose
440
RlENI>lCIPU’O, TABLE
INFLI;EWE
OF
NUCLEOSIDE VATION
ATP
ASI)
III
C~X~ES,PRAUON
TRIPHOSPHATES OF FRL-WOSE
BlSAITl)RJZ.4U,
AKD ON
‘I’HE
OTHER
T,u,zc:~vI-
1 ,(i-I)II’HOSPHA’l’ASE
The assay of fructose l,G-diphosphatssc :uld the inactivation reaction were carried ollt as described in E.rpe,.inlentul Pwcadwe. The reaction mixtrlre was inclthated al. 30” for 5 minrltes and contained, in 1.5 ml: 50 m&r Tris-HCl, pH 7.5; 1 IIIM M&l,; 5 mu cqstrine; nucleoside triphosphate as indicated in the table; and 1 ml of dialyzed kidney extract. The percentage inactivation, in each case, was obtained hv dividing the decrease in activity after preincnbation in the presence of the nucleoside triphosphate by the activity in the absence of added nucleotide. The activities were calcldatetl on the basis of the 1 ml of kidney extract, added. Fructose 1,6-diphosphatase activity (units/ml)
Additions
Inac$;tion -
None CTP, crrP, GTP, ATP, ATP, ATP,
5 5 5 5 3 2
X 1OF M
5.0 -1.5
x
lo-’
M
4.9
X X X X
1O-3 1O-3 1O-3 lop3
M
1.7 1.4 3.5 4.5
M M M
0 10 2 G 72 30 10
I .Miphosphatase in the absence of t’he crude kidney extracts as shown in expcriments .5 and 6 in Table II. Efect of other nucleoside triphosphates and the conceniration of ATP on the inactivation of fructose 1,6-diphosphatase. Other compounds were test’ed to see whether they would replace ATP in the syst,em inactivating fructose 1 ,6-diphosphatase. At concentrations of .5 X 10H3 M, GTP, and UTP had very little or no effect (Table III). Many other nucleotides including AMP, IMP, CMP, GMP, IDP, CDP, GDP, TDP, dCMP, adenosine, cytosine, and guanosine were inactive. High concentrations of ADP caused some inactivation, but this was due to the formation of ATP by the myokinase present in crude kidney extracts. The influence of ATP concentration on the rate of inactivation of fructose 1,6-diphosphatase is shown in Table III. The rate of inactivation decreased very rapidly with decreasing ATP concentration. However, the rate was considerably higher when an ATP regenerating system was present, since
J~IIATTA(:I~AI1~~~,\
ATP is very quic~kly hytlrolyzc~tl to ;\T>P and iZ;\lP in this system. l’wpwties 04’ inactivatin~q s$dfw. The inactivating system slro~d an :lbsolutc rcquiremclnt for 1\Ig++ and the apparent pH optimunl was at, 7.5. L4s shown in Table IV the inactivation of purified fructose 1 ,6-diphosphntast~ required t,hc addition of cruclc kidney cstrart,. Dialysis of ihe crude ext’ract had 110 effect OII t’he inac+tivation of fructose 1 , Miphosphatnse, but boiled dialyzed extract was inactive. JIoreovor, inactivation of purified frurtose 1,6-tliphosphat~ase was found to bc dependent 011 the amount of crude kidney cxt’ract added. InJluence oj” epinephrine on j’wcfose 1,6diphosphatase activity. The possible relationship of the fructose 1,6-diphosphatnse inactivating system in crude kidney extracts to the influence of cpinephrine on glpcolysis in t,he intact cell was investigat’cd. Epinephrine or glucagon would be expected to inactivate fructose 1,6-diphosphatasc, since administrat,ion of these compounds has been shown to result in the formation of cyclic-3 , ;5’-AMP (1.3, 14). This effect should stimulate the inactivat,ion of fructose 1,6-diphosphnt,ase. This possibility was supported by t,he results of experiments in which intracellular fructose 1 ,6-diphosphatase was inachivatcd when kidney cortex slices were incubated with rpinephrine. As shown in Table Y incubation with epinephrine resulted in almoat complete inactivation of fructose 1, Ci-diphosphatnse. There was some inactivation even wit#hout added epinephrinc, and this may represent an enclogenous inactivation of this enzyme. It should be noted that’ the experiment shown in Table V was not typical of all the experiments undertaken. The extent of inactivation of fructose 1,6-diphosphatase was dependent OII the act’ivity of this enzyme in the particular tissue preparation used. In general, tissue from kidneys with higher levels of enzyme showed greater decreaseswhen incubated with epinephrine. Mechanism of inactivation of fiucfose 1,6diphosphataseby ATP. It was possible that ATP inactivated fructose 1,6-diphosphatase by adsorption at a regulatory site, which appears to be distinct from the catalytic
REVERSIBLE
ISACTIVATION
OF
u-FRUCTOSE
TABLE PROPERTIES
OF
THE
SYSTEM
IV
INACTIVATING
FRUCTOSE
1 ,6-DIPHOSPHATASE
The reaction mixture was incubated at 30” for 5 minutes and contained, pH 7.5; 8 IriM cysteine; 8 mM MgClz; and when indicated, 10 mM ATP and boiled extract,. The pllrified fructose 1,6-diphosphatase preparation added activity. The inactivating reaction was assayed as described in the text.
in 1.5 ml: 60 mM Tris-HCI, 1 ml of crude, dialyzed, contained about 2 units
Fructose Incubation
+ Crude extract $ ATP + crude
conditions
extract crude
2.64 0.54
extract
+ fructose 1,6-diphosphatase extract + fructose l,A-diphosphatase
4.2 0.98
$- Boiled dialyzed $- ATP f boiled
extract dialyzed
2.12 2.09
-+ fructose 1,6-diphosphatase extract + fructose 1,6-diphosphatase
TABLE OF
FRUCTOSE
V
1,6-I)IPHOSI?HATASE ~VITH
The cedure. kidney
1,6-diphosphatase activity (units)
$- Crude extract $- ATP + crude
INAc’TIVATION
WHEN
KIDNEY
SLICES
ARE
INCUBATED
EPINEPHRINE
incubation conditions and enzyme assays were carried out as described in Ezperinlenlal Each vessel was incubated for 1 hour at 37” and contained, in 20 ml: 0.154 M NaCl and slices. The control vessel was kept at 3’ and epinephrine was added as indicated. Additions
None (without incubation None (incubated at 37”) Epinephrine (50 @g/ml) Epinephrine (10 fig/ml) Epinephrine (5 pg/ml)
or of
2.67 0.66
extract
+ Dialyzed crude $- ATP + dialyzed
441
1.6.DIPHOSPHATASE
at 3”)
site and is inaccessible to other nucleoside triphosphates (15, 16). Thus, an allosteric type of inhibition might be involved in which the presence of ATP altered the shape of the enzyme. When the enzyme was incubated in the presence of crude kidney extracts with AT32P and ATP S-14C, only 9 emerged from the cellulose-P column with the enzyme peak, indicating that an irreversible reaction of ATP with the enzyme had taken place to form a modified inactive enzyme. The 32P-labeled protein could be precipitated with 50 % ammoniunl sulfate or dialyzed wit’hout any significant 10s~of radioactivity. The best incorporations of 32P into t,he
Pro-
2 gm of
Fructose l,h-diphosphatase activity (umts/gm wet weight of kidney)
Specific activity of isolated fructose l,6-diphosphatase (units/mg protein)
1.71 1.14 0.12 0.34 0.48
1.7 1.1 0.1 0.3 0.5
enzyme were obtained with an AT32P generating system, since as indicated earlier ATP was rapidly hydrolyzed in crude kidney extracts. When the regenerating system was used high concentrations of ATn3P were maintained throughout the incubation period. This system depended on the formation of ATz3P from ADP and 32Pi by the action of aldolase, triosephosphate dehydrogenase, and 1,3 dip-glycerate kinase present in the crude kidney extracts. Fructose 1,6-diphosphate was employed as the substrate for the ATP regenerating system. As shown in Fig. 1 when AT33P was present in the reaction mixture, the decrease in activity of fructose 1,6-diphosphatase was
442
MENDICINO.
REAUDREAU,
0.7
-3
AND
BHATTACIIAII1-YA
7
0.6-
-6
f
, WITH / EXTRACT
ML
OF
ELUANT
FIG. 1. Incorporation
of 32Pi into fructose 1,6diphosphatase in the presence of AT32P and isolation of azP,-labeled fructose 1,6-diphosphatase on cellulose-P. The reaction mixtures were incubated at 30” for 20 minutes and contained, in 13 ml: 25 mM Tris-HCl, pH 7.5; 15 mM cysteine, pH 7.5; 12 mM MgClz; 1 mC z2Pi; and 10 ml of dialyzed kidney extract. The reaction mixture designated “+ATP” also contained 7 mM ADP and 15 mM fructose 1,6-diphosphate. The ot,her incubation mixture designated “-ATP” was incubated in the absence of an AT32P generating system. The elution method described in Ex~e~inLental Procedure was used.
accompanied by a concomitant incorporation of 32P into this enzyme fraction. The radioactivity emerging from the cellulose-P column on elution with 0.05 M Tris-HCl, pH 8-l M NaCl was coincident with the peak of enzymatic activity in each case. The enzyme was not inactivated in the absenceof ATP and very little 32P was incorporated into this enzyme fraction in the absence of adenosinenucleotides as shown in the curves designated “ -ATP”. The figure showsonly the final fractions of the chromatogram cont.aining fructose 1,6-diphosphatase activity. AT32P and 32Pwere completely and rapidly eluted from this cation exchange column in the earlier fractions. When UT32P and CT3”P were incubated
o.70”-pm
L 20
-L-L--J 40 MINUTES
60
60
FIG. 2. Reactivation of inactivated fructose 1,6-diphosphatase by crude kidney extracts. The complete reaction mixt,ure contained, in 10 ml: 60 mM Tris-HCl, pH 7.5; 12 mM MgClz; 0.25 ml of crude kidney extract,, and 2 ml (about 1 unit of activity) of inactivated fructose 1 ,B-diphosphatase. The inactive enzyme was prepared by concentrating and dialyzing the 0.05 M Tris-HCl, pH 8-1 M NaCl fraction obt,ained from a cellulose-P column after applying a reaction mixtlu-e COIItaining crude kidney extract which had been incubated with ATP and Mg++. No react,ivation was observed in the absence of either inactivated fructose 1,6-diphosphatase or crude kidney extract.
with crude kidney extract’s in the presence of 32Pi and a regenerating system (7), fructose 1,6-diphosphat’ase was not inactivated and very little 32P was incorporated into the fructose 1,6-diphosphatase fraction eluted from the cellulose-P column. Reactivation of inactive fructose I ,&diphosphataseby kidney extracts. Experiments were performed to determine whether an enzyme in kidney extracts might convert inactive fructose 1,6-diphosphatase to an active enzyme. The results summarized in Fig. 2 show that when purified inactive fructose 1,6-diphosphatase was reincubated
REVERW3LE
INACTIVATION
OF
with crude kidney extracts a marked reactivation of the enzyme occurred. The increase in the activity of fructose 1 ,Gdiphosphat’ase was accompanied by a decrease in the content of 3flP bound to the protein. Thus it would appear t’hat the inactivation caused by incubation with ATP and iVlg++ can be reversed by other enzymes present in crude kidney homogenates.
Kidney and liver are the main sites of gluconeogenesis in mammals, and the metabolism and regulation of glycogen and glucose Imetabolism in these tissues are closely related to the energy requirements of animal l&sues. Moreover, the rates of glycolysis and gluconeogenesis are sensitive to the needs of tbe organism, and the overall rates of these two processes have been shown to vary widely as a result of hormonal changes, exercise, diet and the presence of pathological conditions. The mechanism by which a .hormone acts to regulate an entire pathway involving many enzymes is not yet clearly understjood, although present evi dcnce da’es indicate that multienzyme systems are regulated at specific rate limiting stages. There are two sites in which t’he synthesis of glycogen is thought to differ from the breakdown of glycogen in liver and kidney, the principal gluconeogenic organs in mammals. The reactions at these two steps are catalyzed by four different enzymes, which are all present in the soluble cytoplasmir fraction Iof the liver and kidney cell. These reactions are illustrated by the following equations. Fructose-Ii-1’
+
Phosphofructokinase
ATI’
(1)
Fructose-l Fruct,ose-1 Fructose
,(i-diP
) Fructose G-P + I’hosphorylase
+
+ ADP
+ Hz0
1, F-diphosphatase
Glycogen,,,+1)
,G-diP
(2) Pi
,
I’,
(3)
Glucose-l-I’ UDP-D-glucose Glyrogen
+ Glycogetq,,
$ Glycogen,,, synt11etnse
(4) ) UDP
+
Glycogenl,+,,
D-FRUCTOSE
1 ,B-DIPHOSPHATASE
443
Fructose 1,6-diphosphatase and glycogen synthetase act in the direction of synthesis of glycogen, and phosphorylase and phosphofructokinase are active in Dhe breakdown of glycogen. It is evident that both sets of enzymes cannot operate simultaneously since the net result would be a substrate-mediated hydrolysis of ATP and UDP-n-glucose to ADP and P 1 and glucose-l-P and UDP, respectively. In t#he present investigation experiments were carried out to elucidate t’he mechanisms involved in the regulation of the activity of fructose 1,6-diphosphatasc in kidney cells. The evidence obtained suggests that those factors which stimulate glycogen breakdown may also function to inactivate fructose 1,6-diphosphatase. The experiments reported clearly demonstrated the marked effect of ATP and cyclic3’ ,5’-AMP upon the activity of renal Dfructose 1,6-diphosphatase in t’he presence of crude kidney extracts. Two distinct effects have been observed, one causing inactivation of the enzyme in the presence of ATP and Mg++ and the other causing a rcactivation of the inactive enzyme by an enzyme present’ in crude kidney extracts. This latter enzyme may function to reactivate fructose 1,6-diphosphatasc after the release of glycolytic stress. A similar response of the inactivating reaction to cpinephrine in its intracellular site was consistent with factors regulating the activity of fructose 1,6diphosphatase in t(he intact cell. Epinephrine exerts it’s glycolytic effect by increasing t#he concentration of active muscle phosphorylase. i2n activating and inactivating enzyme catalyzed the interconversion of 2 forms of phosphorylase. Other studies with liver phosphorylase indicated that a similar mechanism was involved, except that there was no increase in the molecular weight of the active form of phosphorylase (17). The activating enzyme in each case was a kinase which catalyzed the phosphorylation of inact,ive phosphorylase by ATP. Heart and liver protein kinases also acted on muscle phosphorylase. Recent results indicate that epinephrine influences this system by increasing the formation of cyclic-3’,5’-AlIP from ,4TP
444
MENDICINO,
Inactive Active Active
Phosphorylaled glycogen synthetase
BEAUDREAU,
AND
BHATTACHARYYA
enz,ymes
Phosphorylase Phosphofrtlctokinase
Protein
kinase
st,imulat,es
Active Active Inactive Inactive
glycolysis I)IAGIIAM
(13, 14). The inactivating enzyme was a protein phosphat’ase which catalyzed the release of Pi from the active forms of phosphorylase. This enzyme had a broad specificity and dephosphorylsted other phosphoproteins but did not hydrolyze nonprotein phosphate esters. Likewise, more recent studies have indicated that the related enzyme, glycogen synthetase, is inactivated by conditions which activate phosphorylase. Incubation of muscle tissue preparations with epinephrine or homogenates with ATP and cyclic-3’,5’AMP resulted in a decrease in glycogen synthetase activity (1s). Other studies with partly purified enzyme preparations also showed that incubation with ATP could decrease the activity of glycogen synthetase (19). It has been suggested that this enzyme exists in an act’ive and inactive form (20). At the fructose 1,6-diphosphate level similar results have been obtained for phosphofructokinase (21, 22). The evidence thus far indicates that epinephrine may exert its regulatory effects by simultaneously stimulating and inactivating appropriate enzyme reactions involved in the degradation and synthesis of glycogen. In conclusion, it may be tentively postulated that epinephrine stimulates the formation of intracellular cyclic-3’, 5’-AMP at the cell wall. In the presence of this compound an intracellular protein kinase may then catalyze the phosphorylation and activation of phosphorylase and phosphofructokinase and the inactivation of glycogen synthetase and fructose 1 ,6-diphosphatase as shown in the following Diagram A. This would be expected to be an active and very rapid process. It is interesting that such a
Dephosphor~glaied enqpnes glycogeu spn t hetax. fructose 1,Gdiphosphalasc phosphorylaae phosphofrnctclkitl:Iso
A
mechanism would not require the hormone to enter the cell, and indeed the steady state level of hormone in extracellular fluid and blood plasma could be regulated by endocrine tissues which would then control the breakdown of carbohydrate in other organs. A decrease in the concentration of hormone in the extracellular fluid, in conjunction with the destruction of cyclic-3’, 5’-AMP in the cell and activation of phosphoprotein phosphatase, would dephosphorylate the four enzymes and restore a glyconeogenie poise in the liver and kidney cells. The specific and simultaneous action of other enzymes on these four enzymes involved at two rate-limiting steps in carbohydrate metabolism and the possible relationship of this system t’o the formation of cyclic3 ,5’-AMP from ATP by epinephrine at the cell wall remains to be established. One of the difficulties in postulating a plausible role for the phosphoprotein phosphatasc and protein kinase may lie in the fact t,hat the activity and enzymic properties of these enzymes must also be changed as a result of the interaction of epinephrine at t,he cell wall, since if bot,h of these enzymes remained active they would catalyze a protein-dependent hydrolysis of ATP. l
IREVERSIBLE
INACTIVATION
OF
5. STADIE, W. C., AND RIGGS, B. C., J. Biol. Chena.164,687 (1944). G. GLYNN, I. M., AND CHAPPELL, J. B., Biochem. J. 90, 147 (1964). 7. MENDICINO, J., J. Biol. Chem. 237, 165 (1962). 8. FISKE, (C. H., AND SUBBAROIV, Y., J. Biol. Chem. 66, 375 (1925). 9. L~HKINN, K., AND JENR.~SRIK, L., Biochena. J. 178, 419 (1926). 10. Lousy, 0. II., ROSEBROUGH, N. J., FARR, A. L., ARD RANDALL, R. J., J. Biol. Chem. 193, 265 (1951). 11. W.\RBUI~G, O., AND CHRISTIAN, W., Biochem. z. 310, 384 (1941). 12. LXYNE, E., in “Methods in Enzymology” (S. P. Colowick and N. 0. Kaplan, (eds.), Vol. III, p. 447. Academic Press, New York (1957:. 13. RALL, T. W., SUTHERLAND, E. W., AND BERTCIET, J., J. Biol. Chem. 224, 463 (1957).
U-FRUCTOSE
1,6-DIPHOSPHATASE
445
14. SUTHERLAND, E. W., AND RALL, T. W., J. Biol. Chem. 232, 1077 (1958). 15. MENDICINO, J., AND MUNTZ, J. A., J. Biol. Chem. 233, 178 (1958). 16. HURWITZ, J., RAPPEL, L. A., AND HORECKER, B. L., J. Biol. Chem. 226, 525 (1957). 17. RALL, T. W., SUTHERLAND, E. W., AND WOSIL.IIT, W. D., J. Biol. Chem. 218, 483 (1956). 18. BELOCOPITO~, E., Arch. Bio&em. Biophys. 93, 457 (1961). 19. TRAUT, R. R., AND LIPMANN, F., J. Biol. Chem. 238, 1213 (1963). 20. ROSELL-PEREZ, M., VILL.U+PALASI, C., AND LARNER, J., Biochemistry 1, 763 (1962). 21. PASSENNEAU, J. V., AND LOTVRY, 0. H., Biothem. Biophys. Research Common. 7, 10 (1962). 22. MANSOUR, T. E., J. Biol. Chem. 240, 2165 (1965).