- Email: [email protected]
A possible role for oxygen inactivation in the regulation of amidophosphoribosyltransferase activity in mammalian cells
A POSSIBLE ROLE FOR OXYGEN INACTIVATION IN THE REGULATION AMIDOPHOSPHORIBOSYLTRANSFERASE ACTIVITY IN MAMMALIAN CELLS
OF
RICHARD L. LEFF, MITSUO ITAKURA, ALBERT UDOM and EDWARD W. HOLMES Howard Hughes Medical Institute Laboratories, Departments of Medicine and Biochemistry, Duke University Medical Center, Durham, North Carolina
Glutamine phosphoribosylpyrophosphate amidotransferase [(EC 2.4.2.14)] (ATase) catalyzes the first and probable rate-limiting reaction unique to the pathway of purine biosynthesis de novo, and consequently changes in the activity of this enzyme are thought to be important in controlling the rate of purine ribonucleotide synthesis in the cell (I, 2). Prior reports have reviewed the control of mammalian ATase by the ligands PP-ribose-P and purine ribonucleotides (1, 3). Less is known about the mechanisms which control the rate of synthesis and degradation of ATase in eukaryotic cells. In prokaryotes the rate of synthesis of ATase is regulated in part by derepression which is in turn controlled by the availability of exogenous purines (4). Similar mechanisms have not been reported to affect ATase activity in mammalian cell lines (5). Recently Switzer and colleagues have reported an elegant series of studies which demonstrate a novel mechanism for controlling the rate of degradation of ATase in B. subtilis. These investigators have shown that ATase is an oxygen-sensitive enzyme (6, 7), a consequence of the relative ease with which the iron-sulfur center (8) of this protein is oxidized, and that degradation of the enzyme in B. subtilis is preceded by oxygen inactivation (9, 10). Sensitivity of ATase to oxygen inactivation, and consequently the rate of proteolytic degradation of this enzyme, varies depending on the growth cycle of the bacterium (9-11). During log-phase growth the enzyme is relatively resistant to oxygen inactivation, while during stationary-phase the enzyme is relatively sensitive to oxygen inactivation. The factor or factors which control the sensitivity of ATase to oxygen inactivation in the cell have not been identified as yet, but the ligands PP-ribose-P and purine ribonucleotides are thought to be unlikely candidates for this regulatory role (10), prompting these investigators to suggest that covalent modification of ATase may control the sensitivity of this enzyme to oxygen inactivation. Irrespective of the basis for this change in oxygen-sensitivity, the data presented by this group of investigators demonstrate that variability in the sensitivity of ATase to oxygen ~R
ZZ-N
403
404
R I C H A R D L, L E F F , el al.
inactivation is an important mechanism for controlling the rate of ATase degradation in B. subtilis. In this presentation we would like to summarize the data we have accumulated which suggest that a similar mechanism may play a role in controlling the rate of ATase degradation in mammalian cells. We will begin with a description of some of the physical properties of human ATase, since this is the best characterized of the mammalian forms of ATase. We have purified the enzyme from human placenta using the series of steps outlined in Table 1 (12), It was necessary to perform all steps after the initial homogenization in an anaerobic chamber and even then the recovery of catalytic activity was quite low. Under optimal conditions we recovered only 7% of the catalytic activity in the initial homogenate and we achieved a 1778fold purification of the enzyme. Based on this analysis we estimate that ATase accounts for no more than .005% of the soluble protein in human placenta. As shown in Figure 1 this preparation of ATase was over 95% pure as judged by SDS-PAGE. In 7 enzyme preparations we obtained a mean subunit molecular weight of 27,000 + 1500. As illustrated in Figure 1 only one polypeptide chain was evident on SDS-PAGE, We conclude from these studies that the native enzyme, mol. wt. 133,000 (13), is composed of 4-5 identical subunits. The purified enzyme has a yellow-brown color and as shown in Table 2 it contains both iron and sulfide (12, 14). In comparing 7 different enzyme preparations we found the iron and sulfide content (expressed in moles per mole of subunit) to vary, similar to the results reported for different preparations of the B. subtilis enzyme (8). The mean values obtained for all 7 preparations were 3.8 moles of iron and 3.2 moles of sulfide per mole of subunit. This iron-sulfur protein is sensitive to oxygen inactivation and prior studies have demonstrated that molecular oxygen, rather than an oxygen radical, is responsible for inactivation of human ATase (14). Since catalytic activity of the human enzyme can be partly reconstituted by incubation of the inactive TABLE 1. P U R I F I C A T I O N OF H U M A N A M I D O P H O S P H O R I B O S Y L T R A N S F E R A S E *
Step I. II. II1. IV. V. VI. VIL
Ceil Extract DE-52 (NH4)2SO4 plus G-75 Hydroxyapatite Ultrogel Aminohexylagarose Affigel-Blu~
Volume (ml)
Protein (mg)
920 33,235 655 3,013 142 646 250 140 24 43 5 11 . " 11 ~ ' 1.4
Specific Activity (nmol/hr/mg)
-
30 142 502 1,796 4,672 5,290 53,333
Yield 100% 43% 33% 25% 20% 6% 7% .
Fold Purification 1 4.7 17 60 156 176 1778
*All steps after the initial homogenization were carried out at room temperature in an anaerobic chamber (95% N J 5 % H2),
lOOK
68 K
43K
-
-
-
36K
-
29K
-
21K
-
17K
-
-
27K
FIG. 1. SDS gel analysis of the h u m a n ATase preparation obtained from step VII of the purification procedure described in Table 1. The left lane is loaded with molecular weight standards; the right lane is loaded with the purified ATase preparation.
PROBING THE INFRA-STRUCTURE OF THYMIDYLATE SYNTHASE AND DEOXYCYTIDYLATE DEAMINASE FRANK MALEY, MARLENE BELFORT and GLADYS MALEY Center for Laboratories and Research,New York State Department of Health, Empire State Plaza, Albany, New York 12201
INTRODUCTION At the Eighth Symposium on Enzyme Regulation we presented data on regulatory processes affecting both deoxycytidylate (dCMP) deaminase (EC 3.5.4.12) and thymidylate (dTMP) synthase (EC 2.1.1.45) (1). The sequential relationship of these enzymes within the pyrimidine deoxynucleotide pathway, coincident with the marked responsiveness of dCMP deaminase to allosteric end-product regulation (2), provides most animal cells with a particularly sensitive means of regulating dTMP for DNA synthesis. The fact that nature has committed the responsibility for the synthesis of dUMP to 3 allosteric enzymes, dCMP deaminase (2), ribonucleotide reductase (3), and deoxyuridine (thymidine) kinase (4), cannot help but suggest that the formation of this compound plays an important if not rate controlling step in cell division. Our findings at the previous symposium dealt primarily with kinetic responses of the deaminase, from which information regarding the functional groups involved in substrate and effector binding was derived. Responses of the synthase to methotrexate were demonstrated in vivo, and the elevated levels of this enzyme were shown subsequently to be due to an altered steady state resulting from an impairment in enzyme degradation (5). At the time of the earlier meeting an in depth analysis of the structures of the synthase and deaminase was not possible due to the absence of methods which would provide the necessary amounts of both enzymes required for this type of study. These problems have been resolved since by the development of procedures for, 1) isolating methotrexate resistant high dTMP synthase (6, 7) producing strains ofLactobacillus caseL 2) incorporating the dTMP synthase gene from Escherichia coli and phage T4 into amplification plasmids, and 3) the synthesis of a folate analog which has provided an extremely efficient affinity column for the purification of dTMP synthase from various sources (8). In addition a large scale fermentation procedure was developed to isolate dCMP deaminase from T2-phage infectedE, coll. Although they are not from a eucaryotic source, the similarity in properties of these enzymes from 413
414
FRANK MALEY, et al.
bacterial and animal sources suggests that meaningful comparisons can be made, particularly with respect to their phylogenetic evolution. The studies to be presented are but a start in this direction, from which it is hoped that a more complete picture will be available in the near future.
MATERIALS
AND METHODS
Chemicals. Labeled deoxynucleoside mono- and triphosphates were purchased from New England Nuclear Corp., while [214C]5-fluorodeoxyuridylate (FdUMP) and [63H]FdUMP were from Moravek Biochemicals. The unlabeled ribo- and deoxyribonucleotides were obtained from P-L Biochemicals or from Sigma. Hn-dUMP,4-N-hydroxy-dCMP, 5-hydroxymethyl(HM)-dCMP and HM-dCTP were prepared as described previously (911); 5-nitro-dUMP was a generous gift of Dr. Daniel V. Santi. The folylpolyglutamates employed were kindly provided by Dr. Charles M. Baugh. Oligonucleotide linkers were purchased from New England Biolabs and dideoxynucleoside triphosphates and heptadecanucleotide universal primer were from P-L Biochemicals. Enzyme preparation and assay. L. casei dTMP synthase was purified and crystallized as previously described (12). The enzyme was stored routinely at 4°C in 20 mM potassium phosphate, pH 7.0, containing 20 mM 2mercaptoethanol. The E. coli and T-even phage dTMP synthases were purified from crude extracts using primarily the quinazoline affinity column procedure of Rode et al. (8). The affinity column was generously provided by Drs. Wojciech Rode and Joseph R. Bertino. To assay these enzymes, the spectrophotometric procedure of Wahba and Friedkin (13) was employed. Homogeneous T2-phage dCMP deaminase was prepared as described earlier (14), and assayed by a continuous recording method (15). T4-DNA ligase and BAL-31 nuclease were purchased from New England Biolabs. The Klenow fragment ofE. coli DNA polymerase was obtained from Bethesda Research Labs. Restriction enzymes, such as BamHI, ClaI, EcoRv, EcoRI, HindlII, PvulI, PstI, and others were from either of the above suppliers and used according to their instructions. Bacterial strains, phage strains and plasmids. The various recombinant plasmids containing the T4- and E. coli synthase genes were constructed from the plasmid vector pBR322 and then transferred to the high amplifying expression plasmid pKC30 (16). The methodology employed in both cases has been described in detail (17, 18). DNA andprotein sequence analysis. Both strands of the thyA gene from E. coli were sequenced by the dideoxy chain termination method of Sanger (19).
d T M P SYNTHASE A N D d C M P D E A M I N A S E
415
Single-stranded DNA templates of either BAL-31 generated deletions or specific rest riction fragments were cloned into sequencing vectors M 13 mp8 or M 13mp9 (20). Both DNA and protein sequence data were stored and analyzed by programs 1 and 2 of Larson and Messing (21). Protein sequencing. The methods employed for determining the complete amino acid sequences of both L. casei dTMP synthase (22-24) and T2-phage dCMP deaminase (25) have been described, as has the procedure for isolating and analyzing the FdUMP-ternary complex peptide (23, 26). Most of the isolated peptides were sequenced using a Beckman 890B sequencer.
RESULTS
AND
DISCUSSION
Thymidylate Synthase Structural analysis of substrate and inhibitor binding sites. The development of MTX resistant strains ofL. casei (6, 7) that contained a 100-fold enrichment in dTMP synthase, coupled with a modification in the original purification procedure (12), were essential steps in providing adequate amounts of enzyme for the desired structural studies. As a consequence the enzyme was verified to be a 73,000 dalton dimer composed of identical subunits, each consisting of 316 amino acids (22-24). An added dividend was the clarification of the 5fluorodeoxyuridylate (FdUMP) binding region of this protein, first isolated as a nonapeptide (26) and then located more precisely at cysteine residue 198 (Fig. 1) (23). The involvement of a nucleophile at the catalytic site was originally suggested by Santi and Brewer (27) from model studies and subsequently verified by the elegant mechanistic studies of Santi and his group with the synthase from L. casei (28). However, since FdUMP was employed in these studies it was not completely certain that dUMP and FdUMP are bound to the same amino acid, although equilibrium dialysis studies reveal these 2 nucleotides to compete for the same region (29), if not the same amino acid. These studies have, in addition, verified our original kinetic studies with the chick embryo synthase (30), showing that dUMP binds to the enzyme prior to the second substrate, 5,10-methylenetetrahydrofolate (5,10-CH2H4PteGlu). However, there still remains the unexplained anomaly of dUMP binding to a single site, possibly by a half-the-sites mechanism, while FdUMP binds to two sites (one per subunit) (29, 31). To locate the dUMP binding region precisely requires the fixation of dUMP in a manner tight enough to facilitate its isolation as a peptide, similar to that described for the FdUMP ternary complex (26, 32). We have been able to affect this fixation recently by exposing the enzyme to UV light in the presence of dUMP alone (Table 1). The fact that compounds which are mechanism based inhibitors of dUMP, such as FdUMP (28) and 5-nitro-dUMP (33, 34), impair the fixation, with the latter obviously a more tightly bound compound than FdUMP (about 1000× better), suggests
416
FRANK MALEY~ et al.
10
20
30
40
H-MET-LEU-GLU-GLN-PR•-TYR-LEU-A•P-LE•-ALA-LY•-LYs-VAL-LE•-A•P-GL•-GLY-HIS-PHE-LYSPRO-ASP-ARG-THR-HIS-THR-GLY-THR-TYR-SER-ILE-PHE-GLY-HIS-GLN-MET-ARG-PHE-AsP-LEu
-
"6b
SER~LYs-GLY-PHE~PR~-LE~LE~-THR-THR-LY~-LY~-~AL-PR~-PHE-GLY-LEu-~LE-LY~-SER-GL~-~
70 80 LEu-TRP-PHE-LEu-HIS-GLY-AsP-THR-ASN-ILE-ARG-PHE-LEu-LEu-GLN-HIs-ARG-AsN-HIs90 100
ILE-TRP-AsP-GLU-TRP-ALA-PHE-GLu-LY~-TRP-vAL-LYs-SER-AsP-GLu-~YR-H~s-GLY-PR~-AsP110
MET-THR-AsP-PHE-GLY-H IS-ARG-SER-GLN-[YS-AsP-PRO-GLU-PHE-ALA-ALA-VAL-TYR-H
IS-I~-
130 140 GLu-MET-ALA•LY•-PHE-AsP•A•P-ARG••AL-LEu-HIs-A•P-A•P-ALA-PHE-ALA-ALA-LYs-TYR-GLY• 150 160 A•P-LEu-GLY-LE•-VAL-TYR•G•Y-SER•G•N-TR•-ARG-ALA-TRP-HI•-THR-SER-LY•-GLY-A•P-THR170 180 ILE-A~-GLN-LEU-GLY-A~P-~AL-ILE~GLU-GLN-ILE-LYs-THR-HIs-PR~-TYR~SER-ARG-ARG-LEU-
]90 ............ F ................. I LE-VAL-SER-ALA-TRp-AsN-PRo-GLu-AsP'VAL-PRO-THR-MET ~.LA-[Eu~PR~P,~:~ ~_7.T, R~.: 210 220 GLN-PHE-TYR-VAL-AsN-AsP-GLY-LYS-LEu-SER-LEU-GLN-LEu-TYR-GLN-ARG-SER-ALA-
~ ~ 23O 24O A~P~LE~PHE~LE~GLY~VAL~R~E~A~N~ILE~ALA~SER~TYR~ALA~LE~LE~THR~HIs~LE~VAL~ 250 260 ALA-HI~-GLu-CY~-GLY-LE~-~L~-VAL~GLY-GL~PHE-ILE-H~s-T~R-PHE-GLY-A~P-ALA-Hts-LEU270
TYR~AL~A~N~H~-LE~A~P~GLN~LE~LYs-GL~GLN-LE~-~ER-ARG-~HR-PR~-ARG-PR~-ALA-~-
290
300
THR-LE~-GLN-LEu-A~N-PR~-A~P-LY~HIs-AsP~ILE-PHE-Asp-PHE-AsP-MET-LYs-A~P~ILE-LYsLEU-LEu-AsN-TYR-AsP-PRo-TYR-PRO-ALA-I~-LYS-ALA-PRo-VAL-ALA-VAL-OH FIG. 1. The primary sequence ofL. casei thymidylate synthase designating the active site region for FdUMP (area encompassing cysteine-198) and the region labeled by PteGlu7 (area encompassing lysine 50, 51 and lysine 58). Reproduced from (36).
that d U M P is binding to cysteine-198 or to an amino acid in its proximity. If this bond is not labile, we should be able to locate the precise binding site of dUMP. The fact that U M P and 4-N-hydroxy-dCMP inhibit is not surprising in view of the fact that the former can act as a substrate and the latter as an inhibitor of the synthase (30). Since folate exists in nature primarily as a polyglutamate, which can bind to the synthase in the absence of d U M P in contrast to the monoglutamate derivatives of folate (29, 35), the location of this binding site in the synthase was undertaken. For this purpose labeled pteroyl heptaglutamate was used, with the most distal glutamate to the pteroic acid moiety being activated with a water soluble carbodiimide (36). To enhance the specificity of binding, d U M P was included in the reaction, resulting in the fixation of the activated
409
AMIDOPHOSPHORIBOSYLTRANSFERASE AMIDOPHOSPHORIBOSYLT RANSFE RASE
HYPOXANTHINE - GUANINE PHOSPHORIBOSYL TRANSFE RASE
100
I00 9O
90 80
80
70
70
60
6O
~5o
~ so
>
~ 40 o N 30
o 30 N
It~ 20
I0
8
16 24 32 40 HOURS OF CULTURE
48
~sx
10
I 8
L I I 16 24 32 HOURS OF CULTURE
I 40
I 48
FIG. 4. Cells were grown as described in the legend to Figure 3 exceptthe tissue culture medium contained 10/.tg/ml of cycloheximide. activity was proportional to the oxygen concentration in the culture medium. For comparison, hypoxanthine-guanine phosphoribosyltransferase (EC 2.4.2.8) (HPRTase) activity was also quantitated in these cell extracts, and the loss in HPRTase activity was not dependent upon the oxygen concentration in the medium (Fig. 4). We conclude from these studies that the decline in ATase activity in this mammalian cell line when no new protein is being synthesized is largely an oxygen-dependent process. These results are consistent with the in vitro data and support the hypothesis that oxygen inactivation may play a role in regulating ATase activity in eukaryotic cells. Much remains to be done to establish that the rate of ATase degradation in mammalian cells is controlled by variations in the sensitivity of this enzyme to oxygen inactivation, as it is in B subtilis. A specific antibody to ATase is needed to determine the rate of inactivation and degradation of this enzyme in the intact cell. In addition, it will be necessary to demonstrate that the sensitivity of ATase to oxygen inactivation varies urtder different growth conditions which are associated with changes in ATase activity before accepting that this mechanism plays a physiologically important role in regulating ATase degradation and activity in the cell. While the studies reviewed in this presentation do not establish a role for ozygeaa inactivation in the Control of ATaSe actix/~ty in eukaryotic celis, the results are consistent with this hypothesis and will hopefully serve as a stimulus to additional studies in this area. I f this hypothesis were
410
RICHARD L. LEFF, et al.
s u b s t a n t i a t e d , it w o u l d p r o v i d e an e x p l a n a t i o n for the p r e s e n c e o f an i r o n - s u l f u r c e n t e r in this e n z y m e w h i c h d o e s n o t c a t a l y z e a k n o w n o x i d a t i o n r e d u c t i o n r e a c t i o n a n d it w o u l d e x t e n d o u r k n o w l e d g e r e g a r d i n g m e c h a n i s m s w h i c h c o n t r o l p r o t e i n d e g r a d a t i o n in e u k a r y o t i c cells.
SUMMARY H u m a n a n d o t h e r m a m m a l i a n f o r m s o f A T a s e , i n c l u d i n g the C h i n e s e h a m s t e r e n z y m e , are o x y g e n - s e n s i t i v e e n z y m e s a n d h u m a n A T a s e , like the e n z y m e f r o m B. s u b t i l i s , is an i r o n - s u l f u r p r o t e i n . W h e n p r o t e i n s y n t h e s i s is i n h i b i t e d in c u l t u r e d C h i n e s e h a m s t e r cells, A T a s e a c t i v i t y is lost in a n o x y g e n d e p e n d e n t r e a c t i o n . T h e h y p o t h e s i s is d e v e l o p e d t h a t the sensitivity o f A T a s e to o x y g e n i n a c t i v a t i o n c o n t r o l s the rate o f d e g r a d a t i o n o f this e n z y m e in m a m m a l i a n cells, s i m i l a r to the m e c h a n i s m w h i c h has b e e n d e m o n s t r a t e d for r e g u l a t i o n o f A T a s e d e g r a d a t i o n in B. s u b t i l i s .
REFERENCES I. 2. 3. 4. 5. 6. 7. 8.
9. 10. 11. 12.
E.W. HOLMES, Regulation ofpurine biosynthesis de novo, Uric A c i d ( W . N. KELLEY and I. M. WEINER, eds.), Handbook of Experimental Pharmacology 51, 21-42 (1978). M. ITAKURA, R. L. SABINA, P. W. HEALD and E. W. HOLMES, Basis for the control of purine biosynthesis by purine ribonucleotides, J. Clin. Investig. 67, 994-1002 (1981). E. W. HOLMES, Kinetic, physical and regulatory properties of amidophosphoribosyltransferase, Advances in Enzyme Regulation 19, 215-234 (1981). J.S. GOTS, Regulation of purine and pyrimidine metabolism, pp. 225-256 in Metabolic Pathways, 3rd ed., vol. 5 (H. J. VOGEL, ed.), Academic Press, New York (1971). D.W. MARTIN and N. T. OWEN, Repression and depression of purine biosynthesis in mammalian hepatoma cells in culture, J. Biol. Chem. 247, 5477-5485 (1972). C.L. TURNBOUGH and R. L. SWITZER, Oxygen dependent inactivation of glutamine phosphoribosylpyrophosphate amidotransferase in vitro: Model for in vivo inactivation, J. Bact. 121, 115-120 (1975). D. A. BERNLOHR and R. L. SWITZER, Reaction of B. subtilis glutamine phosphoribosylpyrophosphate amidotransferase with oxygen chemistry and regulation by ligands, Biochemistry 20, 5675-5681 (1981). B.A. AVERILL, A. DWIVEDI, P. DEBRUNNER, S. J. VOLLMER, J. Y. WONG and R. L. SWlTZER, Evidence for a tetranuclear iron-sulfur center in glutamine phosphoribosylpyrophosphate amidotransferase from B. subtilis, J. Biol. Chem. 255, 6007-6010 (1980). M. E. RUPPEN and R. L. SWITZER, Degradation of B. subtilis glatamine phosphoribosylpyrophosphate amidotransferase in vivo, J. Biol. Chem. 258, 2843-2851 (1983). D. A. BERNLOHR and R. L. SWITZER, Regulation of B. subtilis glutamine phosphoribosylpyrophosphate amidotransferase inactivation in vivo, J. Bact. 153, 937-949 (1983). C. L. TURNBOUGH and R. L. SWITZER, Oxygen-dependent inactivation of glutamine phosphoribosylpyrophosphate amidotransferase in stationary-phase cultures ofB. subtilis, J. Bact. 121, 108-141 (1975). A. UDOM and E. W. HOLMES, Purification and characterization of the physical properties of amidophosphoribosyltransferase from human placenta, Federation Proc. 42, 2266 (1983).
AMIDOPHOSPHORIBOSYLTRANSFERASE
41 1
13. E. W. HOLMES, J. B. WYNGAARDEN and W. N. KELLEY, Human glutamine phosphoribosylpyrophosphate amidotransferase: Two molecular forms interconvertible by purine ribonucleotides and phosphoribosylpyrophosphate, J. Biol. Chem. 248, 6035-6040 (1973). 14. M. ITAKURA and E. W. HOLMES, Human amidophosphoribosyltransferase: An oxygensensitive iron-sulfur protein, J. Biol. Chem. 254, 333-338 (1979). 15. R.V. LEBO and N. M. KREDICH, Inactivation of human-glutamylcysteine synthetase by cystamine: Demonstration and quantification of enzyme- I igand complexes, J. Biol. Chem. 253, 2615-2623 (1978).
AER 22-N*
OF
RICHARD L. LEFF, MITSUO ITAKURA, ALBERT UDOM and EDWARD W. HOLMES Howard Hughes Medical Institute Laboratories, Departments of Medicine and Biochemistry, Duke University Medical Center, Durham, North Carolina
Glutamine phosphoribosylpyrophosphate amidotransferase [(EC 2.4.2.14)] (ATase) catalyzes the first and probable rate-limiting reaction unique to the pathway of purine biosynthesis de novo, and consequently changes in the activity of this enzyme are thought to be important in controlling the rate of purine ribonucleotide synthesis in the cell (I, 2). Prior reports have reviewed the control of mammalian ATase by the ligands PP-ribose-P and purine ribonucleotides (1, 3). Less is known about the mechanisms which control the rate of synthesis and degradation of ATase in eukaryotic cells. In prokaryotes the rate of synthesis of ATase is regulated in part by derepression which is in turn controlled by the availability of exogenous purines (4). Similar mechanisms have not been reported to affect ATase activity in mammalian cell lines (5). Recently Switzer and colleagues have reported an elegant series of studies which demonstrate a novel mechanism for controlling the rate of degradation of ATase in B. subtilis. These investigators have shown that ATase is an oxygen-sensitive enzyme (6, 7), a consequence of the relative ease with which the iron-sulfur center (8) of this protein is oxidized, and that degradation of the enzyme in B. subtilis is preceded by oxygen inactivation (9, 10). Sensitivity of ATase to oxygen inactivation, and consequently the rate of proteolytic degradation of this enzyme, varies depending on the growth cycle of the bacterium (9-11). During log-phase growth the enzyme is relatively resistant to oxygen inactivation, while during stationary-phase the enzyme is relatively sensitive to oxygen inactivation. The factor or factors which control the sensitivity of ATase to oxygen inactivation in the cell have not been identified as yet, but the ligands PP-ribose-P and purine ribonucleotides are thought to be unlikely candidates for this regulatory role (10), prompting these investigators to suggest that covalent modification of ATase may control the sensitivity of this enzyme to oxygen inactivation. Irrespective of the basis for this change in oxygen-sensitivity, the data presented by this group of investigators demonstrate that variability in the sensitivity of ATase to oxygen ~R
ZZ-N
403
404
R I C H A R D L, L E F F , el al.
inactivation is an important mechanism for controlling the rate of ATase degradation in B. subtilis. In this presentation we would like to summarize the data we have accumulated which suggest that a similar mechanism may play a role in controlling the rate of ATase degradation in mammalian cells. We will begin with a description of some of the physical properties of human ATase, since this is the best characterized of the mammalian forms of ATase. We have purified the enzyme from human placenta using the series of steps outlined in Table 1 (12), It was necessary to perform all steps after the initial homogenization in an anaerobic chamber and even then the recovery of catalytic activity was quite low. Under optimal conditions we recovered only 7% of the catalytic activity in the initial homogenate and we achieved a 1778fold purification of the enzyme. Based on this analysis we estimate that ATase accounts for no more than .005% of the soluble protein in human placenta. As shown in Figure 1 this preparation of ATase was over 95% pure as judged by SDS-PAGE. In 7 enzyme preparations we obtained a mean subunit molecular weight of 27,000 + 1500. As illustrated in Figure 1 only one polypeptide chain was evident on SDS-PAGE, We conclude from these studies that the native enzyme, mol. wt. 133,000 (13), is composed of 4-5 identical subunits. The purified enzyme has a yellow-brown color and as shown in Table 2 it contains both iron and sulfide (12, 14). In comparing 7 different enzyme preparations we found the iron and sulfide content (expressed in moles per mole of subunit) to vary, similar to the results reported for different preparations of the B. subtilis enzyme (8). The mean values obtained for all 7 preparations were 3.8 moles of iron and 3.2 moles of sulfide per mole of subunit. This iron-sulfur protein is sensitive to oxygen inactivation and prior studies have demonstrated that molecular oxygen, rather than an oxygen radical, is responsible for inactivation of human ATase (14). Since catalytic activity of the human enzyme can be partly reconstituted by incubation of the inactive TABLE 1. P U R I F I C A T I O N OF H U M A N A M I D O P H O S P H O R I B O S Y L T R A N S F E R A S E *
Step I. II. II1. IV. V. VI. VIL
Ceil Extract DE-52 (NH4)2SO4 plus G-75 Hydroxyapatite Ultrogel Aminohexylagarose Affigel-Blu~
Volume (ml)
Protein (mg)
920 33,235 655 3,013 142 646 250 140 24 43 5 11 . " 11 ~ ' 1.4
Specific Activity (nmol/hr/mg)
-
30 142 502 1,796 4,672 5,290 53,333
Yield 100% 43% 33% 25% 20% 6% 7% .
Fold Purification 1 4.7 17 60 156 176 1778
*All steps after the initial homogenization were carried out at room temperature in an anaerobic chamber (95% N J 5 % H2),
lOOK
68 K
43K
-
-
-
36K
-
29K
-
21K
-
17K
-
-
27K
FIG. 1. SDS gel analysis of the h u m a n ATase preparation obtained from step VII of the purification procedure described in Table 1. The left lane is loaded with molecular weight standards; the right lane is loaded with the purified ATase preparation.
PROBING THE INFRA-STRUCTURE OF THYMIDYLATE SYNTHASE AND DEOXYCYTIDYLATE DEAMINASE FRANK MALEY, MARLENE BELFORT and GLADYS MALEY Center for Laboratories and Research,New York State Department of Health, Empire State Plaza, Albany, New York 12201
INTRODUCTION At the Eighth Symposium on Enzyme Regulation we presented data on regulatory processes affecting both deoxycytidylate (dCMP) deaminase (EC 3.5.4.12) and thymidylate (dTMP) synthase (EC 2.1.1.45) (1). The sequential relationship of these enzymes within the pyrimidine deoxynucleotide pathway, coincident with the marked responsiveness of dCMP deaminase to allosteric end-product regulation (2), provides most animal cells with a particularly sensitive means of regulating dTMP for DNA synthesis. The fact that nature has committed the responsibility for the synthesis of dUMP to 3 allosteric enzymes, dCMP deaminase (2), ribonucleotide reductase (3), and deoxyuridine (thymidine) kinase (4), cannot help but suggest that the formation of this compound plays an important if not rate controlling step in cell division. Our findings at the previous symposium dealt primarily with kinetic responses of the deaminase, from which information regarding the functional groups involved in substrate and effector binding was derived. Responses of the synthase to methotrexate were demonstrated in vivo, and the elevated levels of this enzyme were shown subsequently to be due to an altered steady state resulting from an impairment in enzyme degradation (5). At the time of the earlier meeting an in depth analysis of the structures of the synthase and deaminase was not possible due to the absence of methods which would provide the necessary amounts of both enzymes required for this type of study. These problems have been resolved since by the development of procedures for, 1) isolating methotrexate resistant high dTMP synthase (6, 7) producing strains ofLactobacillus caseL 2) incorporating the dTMP synthase gene from Escherichia coli and phage T4 into amplification plasmids, and 3) the synthesis of a folate analog which has provided an extremely efficient affinity column for the purification of dTMP synthase from various sources (8). In addition a large scale fermentation procedure was developed to isolate dCMP deaminase from T2-phage infectedE, coll. Although they are not from a eucaryotic source, the similarity in properties of these enzymes from 413
414
FRANK MALEY, et al.
bacterial and animal sources suggests that meaningful comparisons can be made, particularly with respect to their phylogenetic evolution. The studies to be presented are but a start in this direction, from which it is hoped that a more complete picture will be available in the near future.
MATERIALS
AND METHODS
Chemicals. Labeled deoxynucleoside mono- and triphosphates were purchased from New England Nuclear Corp., while [214C]5-fluorodeoxyuridylate (FdUMP) and [63H]FdUMP were from Moravek Biochemicals. The unlabeled ribo- and deoxyribonucleotides were obtained from P-L Biochemicals or from Sigma. Hn-dUMP,4-N-hydroxy-dCMP, 5-hydroxymethyl(HM)-dCMP and HM-dCTP were prepared as described previously (911); 5-nitro-dUMP was a generous gift of Dr. Daniel V. Santi. The folylpolyglutamates employed were kindly provided by Dr. Charles M. Baugh. Oligonucleotide linkers were purchased from New England Biolabs and dideoxynucleoside triphosphates and heptadecanucleotide universal primer were from P-L Biochemicals. Enzyme preparation and assay. L. casei dTMP synthase was purified and crystallized as previously described (12). The enzyme was stored routinely at 4°C in 20 mM potassium phosphate, pH 7.0, containing 20 mM 2mercaptoethanol. The E. coli and T-even phage dTMP synthases were purified from crude extracts using primarily the quinazoline affinity column procedure of Rode et al. (8). The affinity column was generously provided by Drs. Wojciech Rode and Joseph R. Bertino. To assay these enzymes, the spectrophotometric procedure of Wahba and Friedkin (13) was employed. Homogeneous T2-phage dCMP deaminase was prepared as described earlier (14), and assayed by a continuous recording method (15). T4-DNA ligase and BAL-31 nuclease were purchased from New England Biolabs. The Klenow fragment ofE. coli DNA polymerase was obtained from Bethesda Research Labs. Restriction enzymes, such as BamHI, ClaI, EcoRv, EcoRI, HindlII, PvulI, PstI, and others were from either of the above suppliers and used according to their instructions. Bacterial strains, phage strains and plasmids. The various recombinant plasmids containing the T4- and E. coli synthase genes were constructed from the plasmid vector pBR322 and then transferred to the high amplifying expression plasmid pKC30 (16). The methodology employed in both cases has been described in detail (17, 18). DNA andprotein sequence analysis. Both strands of the thyA gene from E. coli were sequenced by the dideoxy chain termination method of Sanger (19).
d T M P SYNTHASE A N D d C M P D E A M I N A S E
415
Single-stranded DNA templates of either BAL-31 generated deletions or specific rest riction fragments were cloned into sequencing vectors M 13 mp8 or M 13mp9 (20). Both DNA and protein sequence data were stored and analyzed by programs 1 and 2 of Larson and Messing (21). Protein sequencing. The methods employed for determining the complete amino acid sequences of both L. casei dTMP synthase (22-24) and T2-phage dCMP deaminase (25) have been described, as has the procedure for isolating and analyzing the FdUMP-ternary complex peptide (23, 26). Most of the isolated peptides were sequenced using a Beckman 890B sequencer.
RESULTS
AND
DISCUSSION
Thymidylate Synthase Structural analysis of substrate and inhibitor binding sites. The development of MTX resistant strains ofL. casei (6, 7) that contained a 100-fold enrichment in dTMP synthase, coupled with a modification in the original purification procedure (12), were essential steps in providing adequate amounts of enzyme for the desired structural studies. As a consequence the enzyme was verified to be a 73,000 dalton dimer composed of identical subunits, each consisting of 316 amino acids (22-24). An added dividend was the clarification of the 5fluorodeoxyuridylate (FdUMP) binding region of this protein, first isolated as a nonapeptide (26) and then located more precisely at cysteine residue 198 (Fig. 1) (23). The involvement of a nucleophile at the catalytic site was originally suggested by Santi and Brewer (27) from model studies and subsequently verified by the elegant mechanistic studies of Santi and his group with the synthase from L. casei (28). However, since FdUMP was employed in these studies it was not completely certain that dUMP and FdUMP are bound to the same amino acid, although equilibrium dialysis studies reveal these 2 nucleotides to compete for the same region (29), if not the same amino acid. These studies have, in addition, verified our original kinetic studies with the chick embryo synthase (30), showing that dUMP binds to the enzyme prior to the second substrate, 5,10-methylenetetrahydrofolate (5,10-CH2H4PteGlu). However, there still remains the unexplained anomaly of dUMP binding to a single site, possibly by a half-the-sites mechanism, while FdUMP binds to two sites (one per subunit) (29, 31). To locate the dUMP binding region precisely requires the fixation of dUMP in a manner tight enough to facilitate its isolation as a peptide, similar to that described for the FdUMP ternary complex (26, 32). We have been able to affect this fixation recently by exposing the enzyme to UV light in the presence of dUMP alone (Table 1). The fact that compounds which are mechanism based inhibitors of dUMP, such as FdUMP (28) and 5-nitro-dUMP (33, 34), impair the fixation, with the latter obviously a more tightly bound compound than FdUMP (about 1000× better), suggests
416
FRANK MALEY~ et al.
10
20
30
40
H-MET-LEU-GLU-GLN-PR•-TYR-LEU-A•P-LE•-ALA-LY•-LYs-VAL-LE•-A•P-GL•-GLY-HIS-PHE-LYSPRO-ASP-ARG-THR-HIS-THR-GLY-THR-TYR-SER-ILE-PHE-GLY-HIS-GLN-MET-ARG-PHE-AsP-LEu
-
"6b
SER~LYs-GLY-PHE~PR~-LE~LE~-THR-THR-LY~-LY~-~AL-PR~-PHE-GLY-LEu-~LE-LY~-SER-GL~-~
70 80 LEu-TRP-PHE-LEu-HIS-GLY-AsP-THR-ASN-ILE-ARG-PHE-LEu-LEu-GLN-HIs-ARG-AsN-HIs90 100
ILE-TRP-AsP-GLU-TRP-ALA-PHE-GLu-LY~-TRP-vAL-LYs-SER-AsP-GLu-~YR-H~s-GLY-PR~-AsP110
MET-THR-AsP-PHE-GLY-H IS-ARG-SER-GLN-[YS-AsP-PRO-GLU-PHE-ALA-ALA-VAL-TYR-H
IS-I~-
130 140 GLu-MET-ALA•LY•-PHE-AsP•A•P-ARG••AL-LEu-HIs-A•P-A•P-ALA-PHE-ALA-ALA-LYs-TYR-GLY• 150 160 A•P-LEu-GLY-LE•-VAL-TYR•G•Y-SER•G•N-TR•-ARG-ALA-TRP-HI•-THR-SER-LY•-GLY-A•P-THR170 180 ILE-A~-GLN-LEU-GLY-A~P-~AL-ILE~GLU-GLN-ILE-LYs-THR-HIs-PR~-TYR~SER-ARG-ARG-LEU-
]90 ............ F ................. I LE-VAL-SER-ALA-TRp-AsN-PRo-GLu-AsP'VAL-PRO-THR-MET ~.LA-[Eu~PR~P,~:~ ~_7.T, R~.: 210 220 GLN-PHE-TYR-VAL-AsN-AsP-GLY-LYS-LEu-SER-LEU-GLN-LEu-TYR-GLN-ARG-SER-ALA-
~ ~ 23O 24O A~P~LE~PHE~LE~GLY~VAL~R~E~A~N~ILE~ALA~SER~TYR~ALA~LE~LE~THR~HIs~LE~VAL~ 250 260 ALA-HI~-GLu-CY~-GLY-LE~-~L~-VAL~GLY-GL~PHE-ILE-H~s-T~R-PHE-GLY-A~P-ALA-Hts-LEU270
TYR~AL~A~N~H~-LE~A~P~GLN~LE~LYs-GL~GLN-LE~-~ER-ARG-~HR-PR~-ARG-PR~-ALA-~-
290
300
THR-LE~-GLN-LEu-A~N-PR~-A~P-LY~HIs-AsP~ILE-PHE-Asp-PHE-AsP-MET-LYs-A~P~ILE-LYsLEU-LEu-AsN-TYR-AsP-PRo-TYR-PRO-ALA-I~-LYS-ALA-PRo-VAL-ALA-VAL-OH FIG. 1. The primary sequence ofL. casei thymidylate synthase designating the active site region for FdUMP (area encompassing cysteine-198) and the region labeled by PteGlu7 (area encompassing lysine 50, 51 and lysine 58). Reproduced from (36).
that d U M P is binding to cysteine-198 or to an amino acid in its proximity. If this bond is not labile, we should be able to locate the precise binding site of dUMP. The fact that U M P and 4-N-hydroxy-dCMP inhibit is not surprising in view of the fact that the former can act as a substrate and the latter as an inhibitor of the synthase (30). Since folate exists in nature primarily as a polyglutamate, which can bind to the synthase in the absence of d U M P in contrast to the monoglutamate derivatives of folate (29, 35), the location of this binding site in the synthase was undertaken. For this purpose labeled pteroyl heptaglutamate was used, with the most distal glutamate to the pteroic acid moiety being activated with a water soluble carbodiimide (36). To enhance the specificity of binding, d U M P was included in the reaction, resulting in the fixation of the activated
409
AMIDOPHOSPHORIBOSYLTRANSFERASE AMIDOPHOSPHORIBOSYLT RANSFE RASE
HYPOXANTHINE - GUANINE PHOSPHORIBOSYL TRANSFE RASE
100
I00 9O
90 80
80
70
70
60
6O
~5o
~ so
>
~ 40 o N 30
o 30 N
It~ 20
I0
8
16 24 32 40 HOURS OF CULTURE
48
~sx
10
I 8
L I I 16 24 32 HOURS OF CULTURE
I 40
I 48
FIG. 4. Cells were grown as described in the legend to Figure 3 exceptthe tissue culture medium contained 10/.tg/ml of cycloheximide. activity was proportional to the oxygen concentration in the culture medium. For comparison, hypoxanthine-guanine phosphoribosyltransferase (EC 2.4.2.8) (HPRTase) activity was also quantitated in these cell extracts, and the loss in HPRTase activity was not dependent upon the oxygen concentration in the medium (Fig. 4). We conclude from these studies that the decline in ATase activity in this mammalian cell line when no new protein is being synthesized is largely an oxygen-dependent process. These results are consistent with the in vitro data and support the hypothesis that oxygen inactivation may play a role in regulating ATase activity in eukaryotic cells. Much remains to be done to establish that the rate of ATase degradation in mammalian cells is controlled by variations in the sensitivity of this enzyme to oxygen inactivation, as it is in B subtilis. A specific antibody to ATase is needed to determine the rate of inactivation and degradation of this enzyme in the intact cell. In addition, it will be necessary to demonstrate that the sensitivity of ATase to oxygen inactivation varies urtder different growth conditions which are associated with changes in ATase activity before accepting that this mechanism plays a physiologically important role in regulating ATase degradation and activity in the cell. While the studies reviewed in this presentation do not establish a role for ozygeaa inactivation in the Control of ATaSe actix/~ty in eukaryotic celis, the results are consistent with this hypothesis and will hopefully serve as a stimulus to additional studies in this area. I f this hypothesis were
410
RICHARD L. LEFF, et al.
s u b s t a n t i a t e d , it w o u l d p r o v i d e an e x p l a n a t i o n for the p r e s e n c e o f an i r o n - s u l f u r c e n t e r in this e n z y m e w h i c h d o e s n o t c a t a l y z e a k n o w n o x i d a t i o n r e d u c t i o n r e a c t i o n a n d it w o u l d e x t e n d o u r k n o w l e d g e r e g a r d i n g m e c h a n i s m s w h i c h c o n t r o l p r o t e i n d e g r a d a t i o n in e u k a r y o t i c cells.
SUMMARY H u m a n a n d o t h e r m a m m a l i a n f o r m s o f A T a s e , i n c l u d i n g the C h i n e s e h a m s t e r e n z y m e , are o x y g e n - s e n s i t i v e e n z y m e s a n d h u m a n A T a s e , like the e n z y m e f r o m B. s u b t i l i s , is an i r o n - s u l f u r p r o t e i n . W h e n p r o t e i n s y n t h e s i s is i n h i b i t e d in c u l t u r e d C h i n e s e h a m s t e r cells, A T a s e a c t i v i t y is lost in a n o x y g e n d e p e n d e n t r e a c t i o n . T h e h y p o t h e s i s is d e v e l o p e d t h a t the sensitivity o f A T a s e to o x y g e n i n a c t i v a t i o n c o n t r o l s the rate o f d e g r a d a t i o n o f this e n z y m e in m a m m a l i a n cells, s i m i l a r to the m e c h a n i s m w h i c h has b e e n d e m o n s t r a t e d for r e g u l a t i o n o f A T a s e d e g r a d a t i o n in B. s u b t i l i s .
REFERENCES I. 2. 3. 4. 5. 6. 7. 8.
9. 10. 11. 12.
E.W. HOLMES, Regulation ofpurine biosynthesis de novo, Uric A c i d ( W . N. KELLEY and I. M. WEINER, eds.), Handbook of Experimental Pharmacology 51, 21-42 (1978). M. ITAKURA, R. L. SABINA, P. W. HEALD and E. W. HOLMES, Basis for the control of purine biosynthesis by purine ribonucleotides, J. Clin. Investig. 67, 994-1002 (1981). E. W. HOLMES, Kinetic, physical and regulatory properties of amidophosphoribosyltransferase, Advances in Enzyme Regulation 19, 215-234 (1981). J.S. GOTS, Regulation of purine and pyrimidine metabolism, pp. 225-256 in Metabolic Pathways, 3rd ed., vol. 5 (H. J. VOGEL, ed.), Academic Press, New York (1971). D.W. MARTIN and N. T. OWEN, Repression and depression of purine biosynthesis in mammalian hepatoma cells in culture, J. Biol. Chem. 247, 5477-5485 (1972). C.L. TURNBOUGH and R. L. SWITZER, Oxygen dependent inactivation of glutamine phosphoribosylpyrophosphate amidotransferase in vitro: Model for in vivo inactivation, J. Bact. 121, 115-120 (1975). D. A. BERNLOHR and R. L. SWITZER, Reaction of B. subtilis glutamine phosphoribosylpyrophosphate amidotransferase with oxygen chemistry and regulation by ligands, Biochemistry 20, 5675-5681 (1981). B.A. AVERILL, A. DWIVEDI, P. DEBRUNNER, S. J. VOLLMER, J. Y. WONG and R. L. SWlTZER, Evidence for a tetranuclear iron-sulfur center in glutamine phosphoribosylpyrophosphate amidotransferase from B. subtilis, J. Biol. Chem. 255, 6007-6010 (1980). M. E. RUPPEN and R. L. SWITZER, Degradation of B. subtilis glatamine phosphoribosylpyrophosphate amidotransferase in vivo, J. Biol. Chem. 258, 2843-2851 (1983). D. A. BERNLOHR and R. L. SWITZER, Regulation of B. subtilis glutamine phosphoribosylpyrophosphate amidotransferase inactivation in vivo, J. Bact. 153, 937-949 (1983). C. L. TURNBOUGH and R. L. SWITZER, Oxygen-dependent inactivation of glutamine phosphoribosylpyrophosphate amidotransferase in stationary-phase cultures ofB. subtilis, J. Bact. 121, 108-141 (1975). A. UDOM and E. W. HOLMES, Purification and characterization of the physical properties of amidophosphoribosyltransferase from human placenta, Federation Proc. 42, 2266 (1983).
AMIDOPHOSPHORIBOSYLTRANSFERASE
41 1
13. E. W. HOLMES, J. B. WYNGAARDEN and W. N. KELLEY, Human glutamine phosphoribosylpyrophosphate amidotransferase: Two molecular forms interconvertible by purine ribonucleotides and phosphoribosylpyrophosphate, J. Biol. Chem. 248, 6035-6040 (1973). 14. M. ITAKURA and E. W. HOLMES, Human amidophosphoribosyltransferase: An oxygensensitive iron-sulfur protein, J. Biol. Chem. 254, 333-338 (1979). 15. R.V. LEBO and N. M. KREDICH, Inactivation of human-glutamylcysteine synthetase by cystamine: Demonstration and quantification of enzyme- I igand complexes, J. Biol. Chem. 253, 2615-2623 (1978).
AER 22-N*