In vivo phosphorylation of liver glycogen synthase. Effect of glucose and glucagon treatment of liver cells on the distribution of the [32~]phosphate

AGNES W. H. TAN AND FRANK Q. NUTTALL ' Metabolic-Endocrine Section, Veterans Affairs Medical Center, Minneapolis, Minn., U.S.A. 55417

and Departments of Biochemistry and Medicine, University of Minnesota, Minneapolis, Minn., U.S.A.

Received June 5, 1992

TAN, A. W. H., and NUTTALL, F. Q. 1993. In vivo phosphorylation of liver glycogen synthase. Effect of glucose and glucagon treatment of liver cells on the distribution of the [32~]phosphate. Biochem. Cell Biol. 71: 90-96.

Glycogen synthase was phosphorylated in vivo by perfusing rat liver or incubating liver cells with [32~]phosphate. It was then isolated by immunoprecipitation and subjected to exhaustive tryptic proteolysis. The trypsin-derived [32~]phosphopeptides were separated by high pressure liquid chromatography (HPLC). Incubation of in vivo phosphorylated synthase with endogenous synthase phosphatase to convert synthase D to synthase R resulted in removal of phosphate from all of the labeled phosphopeptides. In prelabeled liver cells treated with glucagon or glucose, the activities of synthase and phosphorylase changed in the direction expected. The total labeling in the immunoprecipitated synthase was found to be increased to 126% and decreased to 67% of the control with glucagon and glucose treatment, respectively. When the HPLC [32~]phosphopeptide profile of synthase from glucagon-treated animals was compared with that of controls, there were only minor differences in the two profiles. All the peaks were present and the proportion of labeling in each remained similar. There also was only a modest change in the [32~]phosphopeptide profile with glucose treatment when compared with that of controls. These results indicate that regulation of synthase activity in the hepatocyte involves changes in phosphorylation at multiple sites. Indeed, in 32~-labeled liver cells, all of the labeled sites appeared to be involved.

Key words: glycogen synthase, liver, phosphorylation state, glucose treatment, glucagon treatment.

TAN, A. W. H., et NUTTALL, F. Q. 1993. In vivo phosphorylation of liver glycogen synthase. Effect of glucose and glucagon treatment of liver cells on the distribution of the [32~]phosphate. Biochem. Cell Biol. 71 : 90-96.

La glycogtne synthase est phosphorylte in vivo par perfusion du foie de rat ou par incubation des cellules hepatiques avec le [32~]phosphate. Elle est ensuite isolCe par immunoprecipitation et soumise a une prottolyse trypsique exhaustive. Les [32~]phosphopeptides dbivCs de la trypsine sont sCparCs par chromatographie liquide ?I haute pression (HPLC). L'incubation de la synthase phosphorylte in vivo avec une synthase phosphatase endogene pour transformer la syn- thase D en synthase R enleve le phosphate de tous les phosphopeptides marques. Dans les cellules hepatiques prkmar- quees et traitees avec le glucagon ou le glucose, l'activite de la synthase et de la phosphorylase change dans la direction attendue. Le marquage total dans la synthase immunoprecipitee augmente de 126% par traitement avec le glucagon et il diminue de 67% par traitement avec le glucose. I1 n'existe que des differences mineures entre le profil des [32~]phosphopeptides (skpares par HPLC) de la synthase des animaux traites au glucagon et celui des contrales. Tous les pics sont presents et la proportion du marquage dans chacun demeure semblable. Les changements entre le profil des [32~]phosphopeptides des animaux trait& au glucose et celui des contrales sont Cgalement modestes. Ces rksultats montrent que la regulation de l'activitt de la synthase dans les hkpatocytes implique des changements dans la phosphoryla- tion plusieurs sites. En fait, dans les cellules hepatiques marquees avec "P, tous les sites marques semblent impliqub.

Mots cl&s : glycogkne synthase, foie, etat de la phosphorylation, traitement au glucose, traitement au glucagon. [Traduit par la redaction]

Introduction Glycogen synthase, the rate-limiting enzyme in glycogen

synthesis, is under metabolic and endocrine control. Regula- tion of activity occurs via the interconversion of active and inactive enzyme forms. This is brought about by phosphorylation-dephosphorylation of serine residues on the enzyme protein, catalyzed by specific kinases and phosphatases. Using purified kinases, at least 10 different phosphorylation sites have been identified in muscle synthase (for review, see Cohen 1986). Less is known about the liver enzyme. However, recently a cDNA for the rat liver enzyme has been isolated and cloned. From this, the primary sequence has been deduced. The cDNA encoded a protein

ABBREVIATIONS: HPLC, high pressure liquid chromatography; kDa, kilodalton(s); cpm, counts per minute; SDS, sodium dodecyl sulfate; TCA, trichloroacetic acid.

' ~ u t h o r to whom all correspondence should be sent and reprints should be requested, at the following address: Veterans Affairs Medical Center, Metabolic-Endocrine Section 11 lG, One Veterans Drive, Minneapolis, Minn. 55417, U.S.A. Printed in Canada / Imprime au Canada

composed of 703 amino acids with a molecular mass of 80.5 kDa (Bai et al. 1990). Compared with the deduced amino acid sequence of human skeletal muscle synthase (Browner et al. 1989), the C-terminal region was truncated by 33 amino acids and phosphorylation sites la and 16 present in skeletal muscle were absent. Other potential phosphorylation sites also were identified (Bai et al. 1990).

It has been determined previously that there are 6 mol alkali-labile phosphates/mol subunit synthase D. These phosphates have been shown to turnover rapidly and to be involved in the regulation of synthase activity (Tan and Nuttall 1983). However, the distribution of phosphate in the putative phosphorylation sites is unknown.

In a liver cell preparation in which synthase was labeled with [32~]pho~phate and isolated by immunoprecipitation, we have shown a decrease in total 3 2 ~ labeling of synthase when the cells were treated with glucose. There also was an increase in total 3 2 ~ labeling when they were treated with glucagon, compared with untreated cells used as a control (Tan and Nuttall 1985).

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Since there are multiple sites of phosphorylation o n syn- thase (Tan and Nuttall 1983) and many possible physiolog- ical regulators, we were interested in determining whether there a re ma jo r changes in specific 3 2 ~ - l a b e l e d phosphorylation sites after addition of glucagon or glucose to the isolated hepatocyte preparations.

In this paper, we describe the profile of 32~-labeled peptides obtained after exhaustive tryptic digestion of the enzyme protein. The peptides were separated by HPLC. Synthase phosphorylated in vivo, b y perfusing liver o r incubating liver cells with [32~]phosphate, was studied. The phosphorylation profiles of 32~-labeled synthase obtained after treatment of liver cells with glucagon or glucose were compared with those obtained from controls. In addition, we studied the profile of [32~]phosphate-labeled peptide fragments when the in vivo labeled enzyme was activated in vitro by endogenous synthase phosphatase. Our results indicate that the regulation of liver synthase activity occurs through a change in phosphorylation a t essentially all sites labeled over a 60-min period.

Materials and methods Materials

Male Sprague-Dawley rats weighing 200-300 g were used. Trypsin (di henylcarbamyl chloride treated), ['4~]glucose, and

3Y inorganic [ Plphosphate (carrier-free) were from ICN. Glucagon was from Lilly, DE-52 was from Whatman, Nonidet P-40 was from Shell Chemicals, Pansorbin (staphylococcal protein A) was from Calbiochem-Behring, and acetonitrile (HPLC grade) was from Fisher. All other chemicals and biochemicals were of the highest grade available from commercial sources.

Rabbit liver glycogen (type 111, Sigma) used in the enzyme assays was further purified through a mixed-bed ion-exchange resin (Amberlite MB-3). Trifluoroacetic acid, obtained from Sigma, was redistilled before use. UDP-[ '~C]~IUCOS~ was prepared by a method developed in this laboratory (Tan 1979). Preparation of antibodies to synthase and treatment of serum were as described (Tan and Nuttall 1985).

In vitro phosphorylation of liver synthase I by CAMP-dependent protein kinase

Partially purified rat liver synthase I was obtained in a glycogen pellet. A liver extract was prepared in 50 mM glycylglycine - 5 mM Na2S0, (pH 7.2) in the absence of KF and was centrifuged at 45 000 rpm (45 Ti rotor) for 40 min at 25°C. The second pellet was suspended in 50 mM glycylglycine - 5 mM Na2S04 (pH 7.2) and contained synthase I at a concentration of 0.5 U/mL. The phosphorylation reaction mixture contained 0.25 U partially purified rat liver synthase, 0.004 mM CAMP, 2 mM MgCI,, 0.1 mM [Y-,~P]ATP (1000-4000 cpm/pmol), and 50 pg CAMP- dependent protein kinase in a total volume of 500 pL. Phosphoryla- tion was at 30°C for 60 min. The reaction was terminated with the addition of 100 pg of CAMP protein kinase inhibitor.

In vivo phosphorylation of rat liver synthase Two preparations were used to study the in vivo incorporation

of [32~]phosphate into synthase: perfused rat liver and incubated rat liver cells. The perfused liver was used initially, because it was easier to get a pure imrnunoprecipitate starting with a liver glycogen pellet than with a glycogen-free liver extract as obtained in a cell preparation. Secondly, we have much information concerning the interconversion of synthase enzyme forms using the pellet prepa- ration Tan 1982). However, synthase obtained from liver perfused 31 with [ Plphosphate had a much lower specific activity than that which could be obtained from liver cells incubated with [32~]phosphate. Also, prelabeled cells from the same liver could be used for testing the effect of different physiological treatments,

whereas a different perfused liver had to be used for each type of treatment.

Incubation of rat liver cells andperfusion of rat liver with inorganic [32~]phosphate

Liver cells from 24-h fasted rats, prepared as by Berry and Friend (1968), were used. Glucose, at a concentration of 5 mM, was added to the incubation buffer. The phosphate concentration of the incu- bation buffer was reduced to 0.10 mM to increase the specific activity of ATP. Cells with viability greater than 90%, as deter- mined by trypan blue exclusion, were used. The average yield per liver was 5 x lo8 cells. These cells, at a concentration of 1.25 x lo7 cells/mL, were labeled by incubation with about 10 mCi (1 Ci = 37 GBq) of inorganic [32~]phosphate in a shaking water bath at 37°C for 60 min.

For liver perfusion, a rat which had been fasted for 24 h and then refed for 24 h was used. The perfusion buffer was the same as the incubation buffer for Liver cells, except the glucose concen- tration was increased to 10 mM and the buffer was further sup- plemented with 10 mM glutamine, 13.3 mM pyruvate, 1.7 mM lac- tate, and 2 x M insulin (Katz et al. 1979). The first 160 mL of perfusion buffer was allowed to flow through the liver and a recirculation system was set up with the remaining 40 mL. About 8 mCi inorganic [32~]phosphate was added and the liver was perfused for 120 min.

Preparation of liver extract for immunoprecipitation Since a time-dependent nonspecific precipitation of proteins

always occurred in liver extracts, samples had to be partially purified before they could be used for immunoprecipitation. The method has been described elsewhere (Tan and Nuttall 1985). Briefly, a glycogen pellet was prepared from the perfused liver of the fasted and refed rat. This contained 50-75% of the total synthase. a-Amylase, at a concentration of 50 U/mL, was added and the digested sample was used for immunoprecipitation. If an extract of liver cells was used, a-amylase (50 U/mL) was f is t added and a high-speed (105 000 x g, 30 min) supernatant was obtained. This was then treated with ammonium sulfate to 33% saturation. The precipitate, containing about 50% of the original synthase activity was a suitable preparation for immunoprecipitation.

Isolation of 3Z~-labeled synthase by irnmunoprecipitation For a sample containing about 0.7 U synthase, the following

methods were used. To start, two aliquots of Pansorbin (100 and 200 pL of 10% (w/v) suspension of inactivated Staphylococcus aureus of Cowan strain I) were washed twice with a 500-pL solution containing 100 mM KF, 5 mM EDTA, 50 mM Tris-HC1 (pH 7.4), 0.02% NaN,, 1% Nonidet P-40, 0.5% sodium deoxycholate, and 0.1% SDS (buffer A) by centrifugation at 5000 x g for 5 min. The sample containing synthase was added to the 100-pL aliquot of the washed Pansorbin pellet. After resuspension, the tube was rotated, end on end, at room temperature for 1 h. The Pansorbin pellet containing nonspecific proteins was removed by centrifuga- tion at 7000 x g for 5 min. An appropriate amount of antibody was added and the sample was incubated at 30°C for 30 min and at 40°C for overnight. Then the sample was transferred to the tube containing the 200-pL aliquot of washed Pansorbin pellet. After resuspension, the tube was rotated, end on end, at room tempera- ture for 3 h. After centrifugation at 7000 x g for 5 min, the super- natant was discarded. The Pansorbin pellet containing the immunoprecipitate was washed three times with 500 pL of buffer A and then two times with 500 pL of 10 mM phosphate - 0.15 mM NaCl (pH 7.2). The washed pellet was vigorously resuspended in 200 pL of 6 M urea - 2070 SDS, - 5% mercaptoethanol. After centrifugation at 7000 x g for 20 min, the previously immunoprecipitated synthase was in the supernatant.

Tryptic digestion of the irnrnunoprecipitated 32~-labeled synthase and preparation of tryptic phosphopeptides for HPLC

The procedure was essentially that described by Juhl et al. (1983). Samples containing the immunoprecipitated "P-labeled synthase

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92 BIOCHEM. CELL BIOL. VOL. 71, 1993

(A) I4O0

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0 1 J . . . . . . . . lo 0 10 20 30 40 SO 60 70 80 90

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FIG. 1. (A) HPLC profile of tryptic [32~]phosphopeptides of s nthase D labeled in vivo by perfusing rat liver with h [ Plphosphate. Liver was perfused and a glycogen ellet was obtained in the presence of KF as described. The '%-labeled synthase was isolated by immunoprecipitation. The sample was digested with trypsin and the phosphopeptides were analyzed by HPLC using the standard system. (B) Loss of [3z~]phosphate from in vivo phosphorylated sites on synthase during the conversion of synthase D to synthase R in a glycogen pellet. A glycogen pellet was prepared from a rat liver perfused with [32~]phosphate, but in the absence of KF and EDTA. This was incubated at 2S°C for 10 min at which time the activity ratio increased from 0.19 to 0.54 and the radioactivity of the synthase had decreased to 34% of the nonincubated sample. The phosphatase reaction was stopped with the addition of KF and EDTA and the synthase was isolated by immunoprecipitation. The sample was digested with trypsin and the phosphopeptides were analyzed by HPLC using the standard system.

were precipitated with 10% TCA. The TCA precipitate was extracted three times with ether and the powder was suspended in 200 pL of 0.1 M ammonium bicarbonate. The recovery of 32~-labeled synthase after TCA precipitation was 68 + 3% of the immunoprecipitate from control cells, 58 * 4% from the glucose- treated cells; and 73 * 3% from the glucagon cells (n = 4). Trypsin (20 pg) was added and the sample was incubated at 37OC for 16 h. Proteolysis was stopped by the addition of an equal volume of 20% TCA. The TCA supernatant was extracted three times with ether. This sample was used for HPLC analysis of 32~-labeled peptides. We did not attempt to apply a standardized

Separation and measurement of tryptic phosphopeptides of 32~-labeled synthase on HPLC

Reverse-phase chromatography was performed as described by Sheorain et al. (1984), using a Spectra Physics 8700 instrument and a Waters pBondapak (C18) column (0.39 x 30 cm). After a 10-min wash with water, the peptides were eluted over 90 min using a 0-45% acetonitrile gradient. Both the water and acetonitrile contained 0.1% trifluoroacetic acid. Absorption at 206 nm was followed with a LKB Uvicord S. One-millilitre fractions were collected. They were counted for the 3 2 ~ isotope by Cerenkov radiation (Kellogg 1983) at an efficiency of 45%.

Measurement of enzyme activity Glycogen synthase and phosphorylase were measured as

described previously (Tang 1982). In the standard assay, the radioactive glycogen formed was measured. However, when "P-labeled cellular components were present, they interfered with the measurement of the '4~-labeled glycogen formed. The enzymic activity was determined spectrophotometrically by quan- titation of the amount of UDP (Hornbrook et al. 1966) and inorganic phosphate (Fiske and Subbarow 1925) formed, respectively.

Results Loss of in vivo incorporated [32~]phosphate from synthase

during the in vitro activation by synthase phosphatase The phosphopeptide profile of an in vivo phosphorylated

synthase from liver perfused with [32~]phosphate is shown in Fig. 1A. Twelve peaks were identified. In addition, shoulders were present at elution times of 50 min, 59, and 82 min. These have been numbered sequentially. There was extensive labeling of peaks 6 , 7, 9 and 12 in vivo. The most highly labeled peptide of the in vivo profile was peak 12.

In our previous study when synthase D was activated to synthase I in vitro by the synthase phosphatase(s) present in a glycogen pellet preparation, it progressed through an intermediate form (synthase R) with kinetic properties inter- mediate between those of synthase D and synthase I (Tan 1982). All the [32~]phosphates incorporated in vivo were lost during the conversion of synthase D to synthase I in this system (Tan and Nuttall 1985). In the formation of the intermediate synthase R, we were interested in determining if the synthase phosphatase(s) removed phosphate from a specific site or nonspecifically from all the sites. In this experiment, the glycogen pellet isolated from a 32~-labeled perfused rat liver initially had an activity ratio of 0.19 (mostly D form). Incubation of the pellet at 25OC for 10 min resulted in an increase in activity ratio to 0.54 (mostly R form). A longer incubation time would have converted all of the enzyme to the I form. During the conversion of the synthase D to the synthase R from there was a 66% loss of [32~]phosphate from the synthase protein. When the HPLC profile of the tryptic [32~]phosphopeptides of the sample containing mostly the D form (Fig. 1A) is compared with that containing mostly the R form (Fig. lB), it can be seen that all the radioactive peaks were still present. How- ever, the amount of [32~]phosphate labeling had decreased in all of the peaks.

- - .. .

amount of sample to the HPLC column. We were primarily inter- changes in the ~ p ~ ~ ~ ~ ~ f i l ~ of tlyptic [~~p]phosphopeptide ested in comparing the relative peak heights for individual peaks of synthase in response to glucose andglucagon treatment rather than quantitating the entire profile. However, since the samples were small, the majority was injected for analysis and of liver cells resulted in differenes in total cpm used for analyses. The mean It has been established that glucose administration to intact cpm injected were 75 960 + 20 900 from the control cells, animals activates liver synthase by promoting an increased 44 790 + 19 192 for the glucose-treated cells, and 95 213 + 27 133 conversion of the inactive to the active form. Glucagon inac- for the glucagon-treated cells. tivates synthase by the reverse process. Using a liver cell

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NOTES 93

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"0° [ (C)

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FIG. 2. The distribution of [32~]phos hate in phosphopeptides from synthase in hepatocytes. The 3'P-labeled synthase was isolated by immunoprecipitation. A small aliquot was used to analyze the synthase subunit M, by SDS electrophoresis. The rest of the sample was digested with trypsin and the phosphopeptides were analyzed by HPLC using the standard system. The same preparation of cells was used without (A) or with glucagon (B) or glucose (C) addition.

preparation where synthase was labeled and subsequently isolated by immunoprecipitation, we previously reported a decrease in [32~]phosphate content in synthase when prelabeled liver cells were treated with glucose. It was increased with glucagon (Tan and Nuttall 1985). In the present study, synthase in intact liver cells was labeled to high specific activity and the enzyme from the variously treated samples was isolated by immunoprecipitation. Changes in synthase and phosphorylase activities and the extent of labeling were as follows: control cells, 22 + 1.5% synthase I and 18 k 1.2% phosphorylase a; glucagon-

TABLE 1. 3 2 ~ peak heights in trypsin-derived peptides from glycogen synthase in hepatocytes

Peak No. Control Glucose Glucagon

1 226.0k 106.0 250.0k 65.9 216.0-1 98.1 2 232.0-1 88.8 255.0k60.3 282.0 k 164.0 3 246.0k 105.0 289.0+ 56.3 293.05 175.0 4 443.3 k236.6 563.8k299.6 437.0k 190.0 5 604.0 5 204.0 703.0 k 234.0* 603.0 k 168.0 6 592.05 168.0 694.05260.0 610.05 135.0 7 466.0k 143.0 467.0k 147.0 513.0k 147.0 8 470.0k 146.0 443.0k 135.0 416.05 127.0 9 401.05 125.0 456.05 164.0 382.0k 125.0

10 434.0-1 176.0 366.0k 111.0 395.0k 135.0 11 506.0-1 76.6 383.3 k40.8 473.0 k 64.9 12 464.0 * 106.0 282.0k 106.0* 462.0 5 134.0 13 220.0k 97.0 145.0k68.8 240.0k 128.0

NOTE: The data were normalized to a constant 100 000 cpm applied to the column for each preparation to compare individual relative peak heights in the glucose- and glucagon-treated cells to those in the controls. Values are means + SEM (n = 4).

'Statistically different from controls.

treated cells, 12 + 0.6% synthase I, 59 f 1.6% phosphorylase a, 126 k 3.8% of control synthase 3 2 ~

labeling; glucose-treated cells, 55 + 1.3% synthase I, 13 k 0.9'70 phosphorylase a, and 67 k 1.6% of control synthase 3 2 ~ labeling.

The immunoprecipitates were digested with trypsin and the phosphopeptides were analyzed by HPLC. The profile from a typical experiment using control cells is shown in Fig. 2A. The profile from glucagon-treated cells from the same liver also is shown in Fig. 2B. When compared with that of the control cells, there was little difference. When the same cell preparation was incubated with glucose, all of the peaks were greatly reduced, but all were still present. The decreases in peaks 12 and 13 were particularly striking (Fig. 2C). In this experiment, the radioactivity applied to the column was only 37% of that in the control cells.

These experiments have been repeated using four different cell preparations. In each preparation, decreases in all peaks following glucose addition and increases in all peaks fol- lowing glucagon addition were observed. The individual peak heights in each preparation varied even though the labeling pattern was internally quite consistent for each preparation. The variation in relative heights of peaks 5-8 and peaks 11 and 13 were particularly pronounced. Because of these variations, we were not able to accurately quan- titate overall changes in individual peak heights. We have attempted to compare potential differences in various peaks by normalizing, mathematically, the counts in each prepa- ration to a constant cpm applied to the column (Table 1). This analysis suggests that there was little difference in the distribution of [32~]phosphate after glucagon administra- tion. In the glucose-treated cells, peaks 1, 2, 3, and 5 were higher. Peak 11, and particularly peak 12, were lower, suggesting glucose did affect some sites more than others. Overall, however, all of the peaks were reduced in glucose- treaed cells, except for the minor peaks 1, 2, and 3.

Radioautographs of the immunoprecipitates after SDS gel electrophoresis indicated that each of the synthase prepa- rations was essentially pure (data not shown).

Sites on rat liver synthase phosphorylated by CAMP- dependent protein kinase

Our in vivo phosphorylated synthase was obtained from

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94 BIOCHEM. CELL BIOL. VOL. 71, 1993

FIG. 3. HPLC profile of tryptic [32~]phosphopeptides of rat liver synthase I , phosphorylated with CAMP-dependent protein kinase.

perfused rat liver or rat liver cells. Therefore, we wished to determine the HPLC profile of rat liver synthase I phospho- rylated in vitro with CAMP-dependent protein kinase for comparative purposes. A glycogen pellet was used and the conditions were similar to those described by Akatsuka et al. (1985). The labeling pattern was much less complex and clearly different from the in vivo labeling pattern (Fig. 3). Three major peaks were observed. The first and largest eluting at 33 min did not correspond to any of the peaks observed in in vivo labeled preparations. The second cor- responded to peak 7, which was one of the major peaks found in vivo preparations. The third corresponded to peak 11. Peak 12 or peaks 12 and 13, which were major peaks in vivo, were essentially absent. Peaks 6, 9, and 10 also were minor peaks compared with those observed in in vivo preparations.

Using purified rat liver synthase, Imazu et al. (1984) also found three major labeled peaks following phosphorylation by CAMP-dependent protein kinase. They eluted at 15, 20-25, and 29% acetonitrile concentrations. These corre- spond rather closely with those observed in the present study. However, the relative proportion of label in each peak was somewhat different. They reported the late eluting peak to be the major peak. In our study the differences in peaks were not large, but the early eluting peak was the largest.

Discussion Synthase regulation by glucagon

Recent evidence suggests that there are two receptors for glucagon in liver (Wakelam et al. 1986). Binding to one activates adenyl cyclase, elevates the intracellular CAMP concentration, and increases CAMP-dependent protein kinase activity through dissociation of the catalytic and regulatory subunits. Binding to the second receptor results in hydrolysis of phosphatidylinositol with release of inositol triphosphate, which in turn stimulates the release of calcium into the cytosol and activates the calcium-dependent protein kinases. The relative importance of the two mechanisms in glycogen metabolism is unclear. Nevertheless, our data suggest that whether the effects on synthase are due to stimulation of one or both kinase pathways, regulation by glucagon in vivo occurs through a mechanism where the

kinases and phosphatases function in an interactive and coordinated manner and affect all of the sites labeled. Thus, the tryptic profile was little changed although total phosphate labeling was increased (Fig. 2).

In the present study, the 32~-labeled profile of tryptic digests of liver synthase labeled in vivo after glucagon administration clearly was different from that of synthase labeled in vitro with CAMP-dependent protein kinase (Fig. 3). Only three major peaks were identified. The first of these was not observed in the in vivo labeled preparation and peptides corresponding to peaks 12 and 13, which were major peaks in vivo, were essentially absent in the in vitro labeled preparation. Thus, the labeling pattern was different and considerably more complex in vivo. Nevertheless, this does not rule out adenyl cyclase activation by glucagon as a major or only mechanism for increasing the phosphoryla- tion state of synthase in vivo if it is acting in concert with other endogenous kinases.

Akatsuka et al. (1985) also studied the site specificity of CAMP-dependent protein kinase in vitro and of glucagon in vivo on rat liver synthase. Under the in vitro phosphorylating conditions used, they could find only one phosphopeptide by isoelectric focusing. Many peptides were found to be phosphorylated in vivo. Glucagon administra- tion affected the phosphorylation of all these sites. Lawrence et al. (1986) studied the phosphorylation profile of rat adipocytes and found all of the labeled peaks were affected by isoproterenol treatment. Thus, these results are similar to those observed in the present study.

Synthase regulation by glucose via the phosphorylation state The mechanism by which regulators affect increases in

synthase activity is less clear, as the sites dephosphorylated by specific phosphatases have not been well studied. Overall, glucose caused a 33% decrease in the labeling of synthase. If it is acting via a phosphatase with one or more specific sites of action, we would expect the profile after glucose treatment to be considerably different from that of control. However, we found that all the peaks were present and all were lowered, although the extent of lowering may vary. This lack of site specificity also was seen when the in vivo phosphorylated synthase was dephosphorylated in vitro by synthase phosphatase. The HPLC profile of the interme- diate synthase R was found to be the similar to that of synthase D; only the extent of labeling was less. This again suggests that the phosphatases and the kinases function in a coordinated manner and that several sites are involved in regulation in vivo. It should be pointed out that with the method used, minor but consistent changes in radioactive phosphate content in a peptide fragment due to glucose or glycogen treatment would not be detectable.

In a protein as extensively phosphorylated as liver syn- thase, there is another potential problem in interpreting trypsinized peptide profiles. Variations in phosphorylation of specific serine residues may affect the facility with which trypsin may cleave adjacent basic amino acid sites. This could result in some heterogeneity in the 32~-labeled pep- tides, if a peptide has the potential in vivo for phosphoryla- tion at more than one site (Wang et al. 1986). Also, a peptide having three phosphorylation sites and present in vivo as a mixture of mono-, di-, or tri-phosphate forms will elute as either a broad single peak or three closely related peaks from a reverse-phase HPLC column (Poulter et al. 1988). These are inherent limitations of the method.

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NOTES 95

Molecular basis for the phosphorylation-state specific mechanism

Cohen and Antoniw (1973) advanced the concept of the "second site phosphorylation" based o n the study of phosphorylase kinase. Dephosphorylation of the 0-subunit which leads to enzyme inactivation had to be triggered first by phosphorylation of the a-subunit. This idea was extended to glycogen synthase, when two sites of phosphorylation were discovered for the enzyme (Proud et al. 1977). Huang et al. (1975) found that CAMP-dependent protein kinase and phosvitin kinase had to work in a specific order and in a synergistic manner to affect synthase activity and phosphate incorporation. This is good evidence for site-site interaction. After various phosphorylation sites in muscle synthase were identified, the phosphorylation of site 5 by glycogen syn- thase kinase 5 was found to be a prerequisite for phospho- rylation of sites 3a, 3b, and 3c by glycogen synthase kinase 3 (Picton et al. 1982). Using a synthetic peptide containing sequences identical to sites 3a, 3b, 3c, 4, and 5, Fiol et al. (1987) showed that the serines at sites 3a-3c were not phosphorylated by glycogen synthase kinase 3 unless the serine at site 5 was phosphorylated by casein kinase 11.

Thus, for a multiphosphorylated enzyme like glycogen synthase, some interaction between the various phosphoryla- tion sites is to be expected. Through this site-site interaction, activation of a single specific kinase or phosphatase could influence the ability of other kinases and phosphatases present in the cell to act o n synthase and bring about an overall change in the phosphorylation state of the enzyme. This would result in either increases or decreases in the enzyme activity or altered affinity for its allosteric effectors.

Alternatively, a single kinase and phosphatase may be regulating the action of synthase through phosphorylation and dephosphorylation a t several sites. The other identified kinases that have been shown t o phosphorylate synthase in vitro may not phosphorylate synthase in vivo or may provide a steady-state phosphorylation of sites, such as site 5 in muscle, which could keep synthase in a conformation or maintain a recognition site that allows phosphorylation at site 3 or its equivalent in the liver synthase enzyme. Phosphorylation at such sites also could modulate the effi- ciency with which hormonal signals are translated into phosphorylation or dephosphorylation at the primary regulatory sites. In any regard, our data indicate that inactivation of liver synthase by glucagon or activation by glucose administration results in changes in the phosphoryla- tion state of essentially all of the sites identified with regula- tion of the catalytic activity of the enzyme in vivo (Tan and Nuttall 1983). More precise quantitation of the phosphate occupancy of these various sites remains to be determined.

Acknowledgements This work was supported by Merit Review research funds

provided to A.W .H.T. and F.Q.N. by the Veterans Adrnin- istration. We gratefully acknowledge the excellent technical assistance of Faye H. Fang.

Akatsuka, A., Singh, T.J., Nakabayashi. H., Lin, M.C., and Huang, K.P. 1985. Glucagon-stimulated phosphorylation of rat liver glycogen synthase in isolated hepatocytes. J. Biol. Chem. 260: 3239-3242.

Bai, G., Zhang, Z., Werner, R., Nuttall, F.Q., Tan, A.W.H., and Lee, E.Y.C. 1990. The primary structure of rat liver glycogen

synthase deduced by cDNA cloning. J. Biol. Chem. 265(14): 7843-7848.

Berry, M.N., and Friend, D.S. 1968. High-yield preparation of isolated rat liver parenchymal cells. A biochemical and fine struc- tural study. J. Cell Biol. 43: 506-520.

Browner, M.F., Nakano, K., Bang, A.G., and Fletterick, R.J. 1989. Human muscle glycogen synthase cDNA sequence: a negatively charged protein with an asymmetric charge distribu- tion. Proc. Natl. Acad. Sci., U.S.A. 86: 1443-1447.

Cohen, P. 1986. Muscle glycogen synthase. In The enzymes. Vol. 17. Part A. 3rd ed. Edited by P.D. Boyer and E.G. Krebs. Academic Press, New York. pp. 461-497.

Cohen, P., and Antoniw, J.F. 1973. The control of phosphorylase kinase phosphatase by "second site phosphorylation"; a new form of enzyme regulation. FEBS Lett. 34: 43-47.

Fiol, C.J., Mahrenholz, A.M., Wang, Y., Roeske, R.W., and Roach, P.J. 1987. Formation of protein kinase recognition sites by covalent modification of the substrate. J. Biol. Chem. 262: 14 042 - 14 048.

Fiske, C.H., and Subbarow, Y. 1925. The colorimetric determina- tion of phosphorus. J. Biol. Chem. 66(2): 375-400.

Hornbrook, K.R., Burch, H.B., and Lowry, O.H. 1966. The effect of adrenalectomy and hydrocortisone on rat liver metabolites and glycogen synthase activity. Mol. Pharmacol. 2: 106-116.

Huang, K.P., Huang, F.L., Glinsmann, W.H., and Robinson, J.C. 1975. Regulation of glycogen synthetase activity by 2 kinases. Biochem. Biophys. Res. Commun. 65: 1163-1 169.

Imazu, M., Strickland, W.G., and Exton, J.H. 1984. Multiple phosphorylation of rat-liver glycogen synthase by protein kinases. Biochim. Biophys. Acta. 789: 285-293.

Juhl, H., Sheorain, V.S., Schworer, C.M., Jett, M.F., and Soderling, T.R. 1983. Phosphorylation site specificities of glycogen synthase kinases: determination by peptide mapping using high-performance liquid chromatography. Arch. Biochem. Biophys. 222: 518-526.

Katz, J., Golden, S., and Wals, P.A. 1979. Glycogen synthesis by rat hepatocytes. Biochem. J. 180: 389-402.

Kellogg, T.F. 1983. The effect of sample composition and vial type on Cerenkov counting in a liquid scintillation counter. Anal. Biochem. 134: 137-143.

Lawrence, J.C., Jr., James, C., and Hiken, J.F. 1986. Control of glycogen synthase by insulin and isoproterenol in rat adipocytes. J. Biol. Chem. 261: 669-677.

Picton, C., Woodgett, J., Hemmings, B., and Cohen, P. 1982. Multisite phosphorylation of glycogen synthase from rabbit skeletal muscle. FEBS Lett. 150: 191-196.

Poulter, L., Ang, S.-G., Gibson, B.W., Williams, D.H., Holmes. D.F.B., Caudwell, F.B., Pitcher, J., and Cohen, P. 1988. Analysis of the in-vivo phosphorylation state of rabbit skeletal muscle glycogen synthase by fast-atom-bombardment mass spectrometry. Eur. J. Biochem. 175: 497-510.

Proud, C.G., Rylatt, D.B., Yeaman, S.J., and Cohen, P. 1977. Amino acid sequences at the two sites on glycogen synthetase phosphorylated by cyclic AMP-dependent protein kinase and their dephosphorylation by protein phosphatase-111. FEBS Lett. 80: 435-442.

Sheorain, V.S., Juhl, H., Bass, M., and Soderling, T.R. 1984. Effects of epinephrine, diabetes, and insulin on rabbit skeletal muscle glycogen synthase. J. Biol. Chem. 259: 7024-7030.

Tan, A.W.H. 1979. A simplified method for the preparation of pure UDP-[ '~C]~~UCOS~. Biochim. Biophys. Acta, 582: 543-547.

Tan, A.W.H. 1982. Glycogen synthase R in liver of starved rats and starved rats given glucose. J. Biol. Chem. 257: 5004-5007.

Tan, A. W.H., and Nuttall, F.Q. 1983. Endogenous phosphates on liver glycogen synthase D and synthase I. Studies on the number and location. J. Biol. Chem. 258: 9624-9630.

Tan, A.W.H., and Nuttall, F.Q. 1985. In-vivo phosphorylation of liver glycogen synthase. Isolation of the 32~-labeled enzyme and studies on the nature of the bound [32~]phosphates. J. Biol. Chem. 260: 4751-4757.

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96 BIOCHEM. CELL BIOL. VOL. 71, 1993

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Activation parameters for the reconstitution of apotyrosinase by copper

DONALD C. WIGFIELD AND DOUGLAS M. GOLTZ Ottawa-Carleton Chemistry Institute, Department of Chemistry, Carleton University. Ottawa, Ont., Canada KIS 5B6

Received November 16, 1992

WIGFIELD, D. C., and GOLTZ, D. M. 1993. Activation parameters for the reconstitution of apotyrosinase by copper. Biochem. Cell Biol. 71: 96-98.

The reaction of apotyrosinase with divalent copper to give enzymatically active tyrosinase has been studied at pH 8.2 and temperatures from 278 to 303 K. At a 10-fold excess of Cu(I1) over enzyme, the pseudo-first order rate constants range from 1.32 x 10 - 3 s -' to 2.93 x lo-' s -' and yield activation parameters of A H * = 85 + 3 kJ . mol - ' and AS* = 5 k 20 J . mol-' . K- '. The near zero value for the entropy of activation is discussed.

Key words: tyrosinase, copper.

WIGFIELD, D. C., et GOLTZ, D. M. 1993. Activation parameters for the reconstitution of apotyrosinase by copper. Biochem. Cell Biol. 71 : 96-98.

La rtaction de l'apotyrosinase avec le cuivre divalent qui produit l'enzyme active, la tyrosinase, a CtC Ctudite a pH 8,2 et B des tempkratures allant de 278 a 303 K. En presence de 10 fois plus de Cu(I1) que d'enzyme, les constantes de vitesse de rCaction de pseudo-premier ordre vont de 1,32 x s- ' a 2,93 x s - ' et permettent de calculer les param6tres d'activation, AH' = 85 + 3 kJ . mol - ' et AS* = 5 + 20 J . mol - ' .K - I . Nous discutons du fait que la valeur de I'entropie d'activation est presque nulle.

Mots clPs : tyrosinase, cuivre. [Traduit par la rkdaction]

Introduction Tyrosinase catalyzes the rate-determining step in the

biosynthetic pathway leading to melanin. The enzyme requires two atoms of copper and, from the point of view of achieving a full mechanistic understanding of this enzyme, there are two aspects to be considered. These are (a) the mechanism of tyrosine oxidation by the active enzyme and (b) the mechanism by which the apoenzyme acquires copper to become activated. The first aspect appears to be quite complex and mechanisms have been proposed by various groups (Hayaishi 1974; Spiro 1981; Sigel1981; Lerch 1987). Our work has been directed at the less extensively studied second aspect, a reaction known as the reconstitution reaction. In previous publications we have reported the kinetics of the reconstitution reaction and their mechanistic implications (Wigfield and Goltz 1990a), and the pH dependence of the reaction, together with the question of whether Cu(1) or Cu(I1) is the preferred activator of the enzyme (Wigfield and Goltz 1990b). From the mechanistic standpoint, knowledge of the activation parameters is an integral part of the understanding of the reaction, but for this reaction the data are currently incomplete. Two reports are in the literature, the first of which reported the reaction as having an activation energy of about 20 kcal-mol-' (1 kcal = 4.1855 kJ), but gave no supporting data (Kertesz et al. 1972). The second report also concluded that the acti- vation energy was 20 kcal smol- ' (Martinez and Solano). Neither set of investigators reported an Arrhenius A factor or an entropy of activation. In this report we present the Prinled in Canada / Imprime au Canada

temperature dependence of the reconstitution reaction at pH 8.2, together with the A H f and AS' values that fol- low from these data.

Materials and methods Mushroom tyrosinase (Sigma) was used in this work. The

apoenzyme was prepared by incubation of the native tyrosinase preparation in cyanide and was shown to be inactive by use of the usual assay for catalytic activity and to be copper-free using graphite furnace atomic absorption spectrometry. Details of the apoenzyme preparation, its reconstitution, and assay have all been previously documented (Wigfield and Goltz 1990a).

The apoenzyme preparation was kept refrigerated at all times. For the reconstitution reaction at different temperatures, the apotyrosinase preparation was equilibrated at the appropriate tem- perature for 5 min. The temperature range used was 278-303 K, at intervals of 5 K.

After temperature equilibration, 10 PL of copper (11) sulfate solu- tion was added to give a final copper concentration of 1.25 x

M (i.e., a 10-fold excess of copper over enzyme). Mixing was achieved by inversion and the reconstitution was allowed to proceed for various periods of time. The mixture was then added to the assay solution, which effectively quenched the reconstitu- tion. Further reconstitution of apotyrosinase was negligible.

The assay procedure used was the method of dopachrome forma- tion monitored at 475 nm. All assays were performed using a Beckman DU-7 spectrophotometer. Copper determinations were performed using a Perkin-Elmer 5000 atomic absorption spectro- photometer at 324.5 nm.

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