Vol. 121, No. 1 Printed in U.S.A.
JOURNAL OF BACTERIOLOGY, Jan. 1975, P. 115-120 Copyright 0 1975 American Society for Microbiology
Oxygen-Dependent Inactivation of Glutamine Phosphoribosylpyrophosphate Amidotransferase In Vitro: Model for In Vivo Inactivation CHARLES L. TURNBOUGH, JR., AND R. L. SWITZER* Department of Biochemistry, University of Illinois, Urbana, Illinois 61801 Received for publication 21 October 1974
The oxygen-dependent inactivation of glutamine phosphoribosylpyrophosphate amidotransferase (ATase) is demonstrated in cell extracts of Bacillus subtilis. The rate of inactivation of ATase in vitro is apparently first order with respect to oxygen concentration and ATase activity. ATase inactivated in vitro (or in vivo) cannot be reactivated by a variety of reductants. ATase is significantly stabilized to oxygen-dependent inactivation in vitro in the presence of tetrasodium phosphoribosylpyrophosphate and glutamine together. The effects of the end product inhibitors, adenosine 5'-monophosphate (AMP) and guanosine 5'-monophosphate (GMP), on the stability of ATase are antagonistic. AMP stabilizes ATase, whereas GMP destabilizes the enzyme. The stability of ATase can be manipulated over wide ranges by variations in the AMP/GMP ratio. The effects of AMP and GMP on the inactivation of ATase in vitro are very specific. ATase is partially inhibited by 1,10-phenanthroline, suggesting that the enzyme contains iron (or some other chelatable metal ion). The inactivation of ATase in vitro is proposed to present a model for the reconstruction of the inactivation of ATase in stationary-phase cells of B. subtilis.
In the accompanying communication (10) we reported a rapid, oxygen-dependent inactivation of glutamine phosphoribosylpyrophosphate amidotransferase (ATase, EC 2.4.2.14) in stationary-phase cells of Bacillus subtilis. In an attempt to determine the chemical nature and means of regulation of this inactivation, we have attempted to reconstruct the process in cell extracts. In this paper we demonstrate the oxygen-dependent inactivation of ATase in vitro. Preliminary experiments on the chemical nature of the oxygen-sensitive sites of ATase are presented. Examination of the effects of substrates, end product inhibitors, and ATase concentration on the rate of enzyme inactivation has provided results that enable us to put forward a tentative model for the inactivation of ATase in stationary-phase cells of B. subtilis. (This work was presented in part at the Biochemistry/Biophysics 1974 Meeting, 2-7 June 1974, Minneapolis, Minn.) MATERIALS AND METHODS Chemicals. Chemicals and enzymes were obtained from the following sources: purine and pyrimidine bases, nucleosides and nucleotides and catalase, Sigma Chemical Co.; 1,10-phenanthroline, J. T. Baker Chemical Co.; 1,7-phenanthroline, G. Fred-
erick Smith Chemical Co.; [14Cjadenosine 5'-monophosphate (AMP), New England Nuclear Corp. All other chemicals, enzymes, and buffers were described previously (10) or were reagent grade and commercially available. Bacterial strains. All studies in this paper were conducted with B. subtilis strain 168, which requires tryptophan for growth. Media and culture methods. A buffered minimal medium containing 0.1% glucose described by Anagnostopoulos and Spizizen (1) was used for growth of cells. Culture methods were described in the accompanying paper (10). Preparation of cell extracts. In most experiments, 40-ml samples were withdrawn from a 1-liter culture at determined times during growth. The samples were harvested and stored as described previously (10). The pelleted cells were resuspended in 5 ml of cold buffer A (2) without 2-mercaptoethanol and 100 MAliters of a 2 mg/ml lysozyme solution. The cells were extracted as previously described (10). Enzyme assay. ATase activity was measured by assay I described in the accompanying paper (10). In most experiments 150-,uliter volumes of extract were assayed. It was confirmed that inhibitors of ATase used in this study did not inhibit the assay of glutamate. Incubation of extracts. When extracts were incubated anaerobically, all extraction procedures were performed in a dry box under argon. Extracts were incubated under argon in sealed tubes at 37 C, and samples were assayed at determined times. The tubes 115
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were flushed with argon during sampling and immediately resealed to eliminate oxygen. When extracts were incubated under 1.0 atm of oxygen, 4 ml of extract was placed in a 10-ml Erlenmeyer flask and the flask was capped with a serum stopper. The flask was flushed for 3 min with oxygen, sealed, and incubated at 37 C. In experiments where several samples were assayed at different times, the flask was flushed with oxygen after each sampling. Hydrolysis of AMP in extracts. [14C ]AMP (1 mM) was added to extracts of exponential- and stationary-phase cells and incubated at 37 C for 1.5 h. At determined times, samples were removed and applied to cellulose thin layers (Eastman Chromagram with fluorescent indicator). Carrier adenine and adenosine were spotted on the samples and the chromatograms were developed with 1 M ammonium acetate (4). "4C-labeled adenine, adenosine, and AMP were measured.
RESULTS Inactivation of ATase in vitro. In experiments described in the accompanying communication (10), we observed that ATase activity declined in cell extracts when they were incubated at 37 C, but that this loss of activity was prevented when the extracts were incubated under argon. This finding suggested that the oxygen-dependent inactivation of ATase occurs in vitro as well as in intact stationary-phase cells. The oxygen dependence of ATase inactivation in cell extracts was demonstrated by measuring the effect of oxygen concentration on the rate of inactivation at 37 C. The experiments presented in Fig. 1 show that ATase is stable in the absence of oxygen, and that the rate of inactivation is apparently first order with respect to oxygen concentration. The inactivation also appears to be first order with respect to ATase activity. The results are consistent with the idea that inactivation results from a simple reaction of ATase with molecular oxygen. ATase that was inactivated in vitro (or in vivo) could not be reactivated by incubation with 2-mercaptoethanol, cysteine, dithiothreitol, sodium borohydride, sodium dithionite, or a combination of ferrous chloride, sodium sulfide, and dithiothreitol. The experiments shown in Fig. 1 were conducted with extracts prepared from stationaryphase cells. To ascertain whether the ATase from exponentially growing cells, where no inactivation has taken place, was also susceptible to oxygen-dependent inactivation in vitro, extracts were prepared from cells harvested during exponential growth (48 units of ATase per ml) and incubated as in Fig. 1. The results were very similar to those shown in Fig. 1, except
_-
0.
1 2 Hours of Incubation (37C)
3
FIG. 1. Oxygen dependence of ATase inactivation in cell extracts. Three identical pellets of early stationary-phase cells were extracted as in Materials and Methods. The extracts (30 units of ATase per ml) were incubated under 0 (0), 0.2 (0), and 1.0 (A) atm of oxygen and 150-Aliter samples were assayed at indicated times.
that the rates of inactivation were slightly slower in the extracts of exponential-phase cells (e.g., t½ under air at 37 C was 90 min as opposed to 70 min in the stationary-phase cell extracts). This difference in rate of inactivation might reflect a difference in the concentration of some metabolite in the extracts that affects the inactivation process. It is also possible that the rate of ATase inactivation is dependent on the ATase concentration in the extract. Since the concentration of ATase in the exponentialphase cell extracts (48 units/ml) was higher than that of stationary-phase cell extracts (30 units/ml), one would predict that ATase inactivation would proceed more slowly at higher ATase concentrations if this hypothesis is correct. To examine these possibilities, two 10-ml extracts containing approximately 300 units of ATase per ml were prepared by extracting cells harvested during exponential growth in buffer A without 2-mercaptoethanol. One extract was diluted to several concentrations with buffer A without 2-mercaptoethanol. The second extract was dialyzed for 12 h at 4 C against two 2-liter changes of buffer A without 2-mercaptoethanol through which argon was bubbled continuously. The extract was then dialyzed for one additional hour against 2 liters of buffer A without 2-mercaptoethanol to remove argon. The
INACTIVATION OF ATase IN VITRO
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dialyzed extract was diluted to several concentrations with buffer A without 2-mercaptoethanol. These extracts containing different concentrations of ATase were incubated under air at 37 C, and the rates of inactivation were measured as in Fig. 1 (Table 1). Dialysis of extracts containing 30 to 60 units of ATase per ml did not alter the rates of inactivation. Dialysis of more concentrated extracts, however, significantly altered the half-life of ATase, indicating that small molecules capable of stabilizing ATase were removed. The possibility that these results reflect the instability of a stabilizing element rather than its removal was eliminated by incubating a sample of the 294 units per ml of extract under argon at 4 C for 13 h. When this sample was incubated as above, only 8% (t½ 12 h) of the initial ATase activity was lost. The results also show that the inactivation of ATase proceeds more rapidly when the enzyme is more dilute in the extract. The addition of bovine serum albumin to extracts did not alter the rate of the inactivation. In subsequent experiments, therefore, only dilute extracts were used to minimize interferences by endogenous small molecules. We have also reported initial ATase activities and compared inactivation rates only in extracts with similar initial ATase activities. Effects of substrates and end product inhibitors on the inactivation of ATase in vitro. Studies of ATase from pigeon liver (6) and human placenta (5) have established that substrates and end product inhibitors alter the activity and physical conformation of the enzymes from these sources. These results prompted an investigation of the effects of these substances on the inactivation of B. subtilis ATase in vitro. Samples of late exponentialTABLE 1. Effect of ATase concentration on the irnctivation of ATase in vitro Initial ATase activity
(units/ml) 294 139 64 32 255 124 55 31
Protein concn (mg/ml)
(mg/mi) 5.9 3.0 1.5 0.7 6.4
3.2 1.6 0.8
t, (min)a without dialysis
te(mdi)al-
and early stationary-phase cells were harvested and extracted as described in Materials and Methods. Exponential-phase extracts contained 42 + 4 units of ATase per ml and stationary-phase extracts contained 29 ± 4 units of ATase per ml. After the addition of substrate(s) or purine nucleotide inhibitor(s), the extracts were incubated at 37 C under air or 1.0 atm of oxygen, and the rate of ATase inactivation was determined. It was demonstrated that the hydrolysis of tetrasodium phosphoribosylpyrophosphate (PRPP) and AMP in extracts of exponential- or stationary-phase cells was small during incubation (AMP hydrolysis < 7% of the initial concentration). The addition of 1 mM PRPP or 6 mM glutamine alone to extracts of either exponential- or stationary-phase cells had no significant effect on the rate of inactivation of ATase (Table 2). The effect of both substrates on the inactivation was measured by adding 2.5 mM PRPP and 6 mM glutamine together to an extract of exponential-phase cells and incubating this extract at 37 C for 30 min under 1.0 atm of oxygen. After 30 min, the concentration of PRPP had declined to about 1 mM because of product formation. The substrates and products were removed by dialysis, and the residual ATase activity was assayed. The stability of ATase was increased by nearly fourfold by the presence of both substrates (Table 2). The purine nucleotides AMP and ADP have been shown to be the most potent end product inhibitors of ATase from B. subtilis (8). The effect of AMP on the inactivation of ATase is shown in Fig. 2. Low concentrations of AMP greatly stabilize ATase activity. For comparison, the inhibition of ATase by AMP was also determined and plotted in Fig. 2. The stability of ATase was very substantially increased by concentrations of AMP that bring about relatively little inhibition. The inhibition of ATase by AMP followed a sigmoidal concentration
after dialysis
TABLE 2. Effect of substrates on the inactivation of ATase in vitroa
Stablec 207 90 78 139 104 92 82
Half-lives were measured at 37 C under air. bHalf-lives were measured after 12 h of dialysis under argon. c Corresponds to a 1 to 2% loss of ATase activity after 1.5 h of incubation. a
117
Addition
tA (min)b
None .......................... 1.0 mM PRPP .......................... 6.0 mM glutamine ........................ 2.5 mM PRPP plus 6.0 mM glutamine ......
17 4 2C 13
19 60 X 6d
ATase activity was 38 units/ml. bHalf-lives were measured at 37 C under 1.0 atm of oxygen. c Average of five determinations. d Average of two determinations. a Initial
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.ECY
.)
N
-4 a
CN
0.5
AMP (mM)
FIG. 2. Stabilization and inhibition of ATase by AMP. Stabilization of ATase to oxygen-dependent inactivation (0) was measured by adding the indicated amount of AMP to identical extracts of stationary-phase cells (32 units of ATase per ml) and incubating the extracts at 37 C under 1.0 atm of oxygen. Inhibition of ATase (0) was measured by adding the indicated amount of AMP to the assay mixture described in Materials and Methods.
dependence (Hill coefficient of 3.7 to 4.0), whereas the stabilization of ATase by AMP followed a hyperbolic dependence. An AMP concentration of 0.27 mM gave 50% inhibition under the conditions of assay I. Similar results were obtained when AMP was added to extracts of exponential-phase cells. Guanosine 5'-monophosphate (GMP) is a much weaker inhibitor of ATase from B. subtilis than AMP (8). Inhibition by GMP followed a hyperbolic concentration dependence with 50% inhibition at 2 to 3 mM under the conditions of assay I. The effect of GMP on the inactivation of ATase in vitro is shown in Fig. 3. In contrast to AMP, GMP decreased the stability of ATase to oxygen-dependent inactivation. The effect of GMP on stability was half-maximal at approximately 0.2 mM. Similar results were obtained when GMP was added to extracts of stationaryphase cells. Table 3 summarizes the results of an experiment in which both GMP and AMP were added to identical extracts of stationary-phase cells containing 32 units of ATase per ml. The effects of AMP and GMP clearly antagonize one another. The stability of ATase to oxygen-dependent inactivation in vitro can be manipulated over wide ranges by variations in the AMP/
1.0
1.5
GMP (mM)
FIG. 3. Destabilization of ATase by GMP. After the addition of the indicated amount of GMP to identical extracts of exponential-phase cells (38 units of ATase per ml), the extracts were incubated at 37 C under air and the half-lives were determined.
TABLE 3. Inactivation of ATase in the presence of both AMP and GMP ts (min)b GMP (mM) AMP (mM) 0.48 0.48 0.48 0.48
0
0 0.24 0.48 0.96 0.96
84 57 40 25 6
aInitial ATase activity was 32 units/ml. measured at 37 C under 1.0 atm of
b Half-lives were
oxygen.
GMP ratio in the extract. In separate experiments, it was shown that the addition of 0.5 or 1.0 mM GMP to extensively dialyzed extracts decreased the stability of ATase. Thus, GMP effects a true destabilization rather than acting to displace stabilizing nucleotides in the crude extract. The specificity of the effects of the nucleotide inhibitors of ATase on the inactivation of the enzyme was examined (Table 4}. It is evident that the nucleotide effects are quite specific; only 2'-deoxy AMP (and possibly cytidine 5'monophosphate) was able to significantly mimic the action of AMP. The effect of guanosine 5'-triphosphate may be due to formation of GMP in the crude extracts. Other nucleoside di- and triphosphates were not tested because of their possible breakdown in the crude extracts. Similar results were obtained with ex-
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INACTIVATION OF ATase IN VITRO
119
TABLE 4. Specificity of nucleotide effects on inactivation of ATase in vitroa Additions"
t,,(min)c
None ........... 17d 5'-AMP ........... 95 ....... 34 2'-deoxy 5'-AMP ... 1,10~~i, 0~~~~~~~~~~ 3-AMP ........... 16 ....... 18 3':5'-cyclic AMP ... 40 40_ 17 Adenosine .......... 19 Adenine .......... 5'-GMP ........... 7 20 10 5'-GTP .......... 3':5'-cyclic GMP .18 17 5'-XMP .......... 1.5 2.0 2.5 0.5 1.0 17 5'-IMP .......... 5'-UMP ........... 15 Phenanthroline (mM) 5'-CMP .......... 26 FIG. 4. Inhibition of ATase by phenanthroline. Inhibition was measured by adding the indicated a Initial ATase activity was 38 units/ml. of 1,7 phenanthroline (0) or 1,10phenanthroAll additions were 1.0 mM. Abbreviations not amount to the assay mixture described in Materials line (0) given in text: GTP, guanosine 5'-triphosphate; XMP, xanthine 5'-monophosphate; IMP, inosine 5'-mono- and Methods. phosphate; UMP, uridine 5'-monophosphate; CMP, trations, it was difficult to make a conclusive cytidine 5'-monophosphate. interpretation. c Half-lives were measured at 37 C under 1.0 atm of Recently it was reported by Trotta et al. (9) oxygen. d Average of five determinations. that carbamyl phosphate synthetase from Esch-
phenanthroline
erichia coli was inactivated by hydrogen peroxide generated in enzyme preparations containing low concentrations of dithiothreitol. To cells. Possible chemical nature of the oxygen- determine if inactivation of ATase in extracts of sensitive site of ATase. ATase purified from B. subtilis proceeds by such a mechanism, avian liver (3, 7) contains nonheme iron, which catalase (3,500 units/ml) was added to extracts appears to be involved in both the catalytic and the extracts were incubated at 37 C under reaction and structural integrity of the enzyme. air for 3 h. The rate of inactivation of ATase was Purified pigeon liver ATase is inhibited and the same in extracts with or without catalase. inactivated by 1,10-phenanthroline (6). To in- Catalase activity was measured after incubavestigate the possibility that ATase from B. tion and was > 80% of the initial activity. To subtilis contains nonheme iron, the inhibition of test whether the inactivation of ATase in vitro B. subtilis ATase by 1,10-phenanthroline was was due to photooxidation of a sensitive site on measured (Fig. 4). Inhibition of ATase by the enzyme, extracts were incubated at 37 C 1, 10-phenanthroline appears to be biphasic. under air in the dark. The inactivation of ATase Significant inhibition occurs at low 1,10-phen- was the same in the absence or presence of light. anthroline concentrations; however, the effect DISCUSSION of higher concentrations is appreciably less than InhiThe reconstruction of an oxygen-dependent would be expected with simple inhibition. bition of ATase by 1,7-phenanthroline was also inactivation of glutamine PRPP amidotransfermeasured. This isomer of phenanthroline binds ase represents a first step in elucidating the metals poorly and is often used as a control for chemical mechanism and physiological regula1,10-phenanthroline inhibition. At 0.25 mM tion of the inactivation of this enzyme in 1,7-phenanthroline, ATase activity was inhib- stationary-phase B. subtilis cells in vivo. The ited only 8%, but at higher concentrations the simple kinetics of the inactivation process suginhibition paralleled that observed with gest that the inactivation results from a direct 1,10-phenanthroline. These experiments sug- oxidation of some group on the enzyme, but the gest the presence of a chelatable metal ion; involvement of an inactivating enzyme can only however, because of the fractional inhibition at be conclusively ruled out by purification of the low 1,10-phenanthroline concentrations and ap- ATase. In either case, it is presumed that parent nonspecific inhibition at higher concen- inactivation results from oxidation of a group or tracts from exponential- and stationary-phase
120
TURNBOUGH AND SWITZER
groups that are essential for activity. In the case of a direct oxidation two groups are likely: sulfhydryl groups and ferrous iron. The observation that a variety of reducing agents failed to reactivate inactivated ATase indicates that the inactivation is not a simple oxidation of sulfhydryl groups to disulfides. ATases purified from avian liver (3, 7) have been shown to contain nonheme iron, which is essential for enzymatic activity. Inhibition of B. subtilis ATase by 1, 10-phenanthroline suggested that this enzyme also contains ferrous iron that is essential for activity. The dependence of the rate of oxygendependent inactivation on ATase concentration, together with the observations in the accompanying paper (10) that the activity of ATase is not linearly dependent on concentration, suggests that the enzyme may exist in an equilibrium between aggregated and disaggregated states. A similar equilibrium between aggregated and disaggregated forms has been demonstrated for the ATase of human placenta (5). The ATase purified from pigeon liver also appears to exist in several conformational states (6), and the existence of aggregated forms has been shown (7). It is clear that most of the remaining questions about the chemical nature of the oxygen-dependent inactivation of ATase in vivo could be answered by a study of the inactivation of the pure enzyme. Purification of the enzyme is currently under way in our
laboratory. The results of the study of the inactivation of ATase in vitro also suggest means by which the inactivation of the enzyme could be regulated in vivo. Neither PRPP nor glutamine alone affected the stability of ATase to oxygen, but the two substrates together stabilized the enzyme substantially. It is very likely that PRPP pools decline when cells are starved for carbon substrates and it is virtually certain that glutamine levels are depleted by nitrogen starvation. Thus, a marked drop in either substrate would signal starvation to the cell and would account for the destabilization of ATase in starved cells. The effects of the purine nucleotides, AMP and GMP, on the stability of ATase to oxygen were also specific and effective at relatively low concentrations. AMP stabilized ATase, whereas GMP destabilized the enzyme. The two nucleotides were able to antagonize the action of each other, so that the stability of ATase could be manipulated over wide ranges by variations in the AMP/GMP ratio. We would like to put forward the hypothesis that some combination
J. BACTERIOL.
of these effects of substrates and end product inhibitors could account for the stability of ATase to oxidation in growing cells and the oxygen-dependent inactivation in starved cells. It is possible that these ligands exert their effects on the stability of the enzyme by altering an equilibrium between disaggregated and aggregated forms of the enzyme. Effects of substrates and allosteric inhibitors on the physical state of ATase have been documented for the pigeon liver (6) and human placeta (5) enzymes. The validity of the hypothesis that substrates and/or end products regulate the stability of ATase in vivo can be most readily tested by determining the intracellular levels of the substrates and inhibitors of ATase under a variety of conditions where the enzyme is stable and where it is inactivated in vivo. LITERATURE CITED 1. Anagnostopoulos, C., and J. Spizizen. 1961. Requirements for transformation in Bacillus subtilis. J. Bacteriol. 81:741-746. 2. Deutscher, M. P., and A. Kornberg. 1968. Biochemical studies of bacterial sporulation and germination. VIII. Patterns of enzyme development during growth and sporulation of Bacillus subtilis. J. Biol. Chem.
243:4653-4660. 3. Hartman, S. C. 1963. Phosphoribosyl pyrophosphate amidotransferase. Purification and general catalytic properties. J. Biol. Chem. 238:3024-3035. 4. Hochstadt-Ozer, J., and E. R. Stadtman. 1971. The regulation of purine utilization in bacteria. I. Purification of adenine phosphoribosyltransferase from Escherichia coli K,, and control of activity by nucleotides. J.
Biol. Chem. 246:5294-5303. 5. Holmes, E. W., J. B. Wyngaarden, and W. N. Kelley. 1973. Human glutamine phosphoribosylpyrophosphate amidotransferase. Two molecular forms interconvertible by purine ribonucleotides and phosphoribosylpyrophosphate. J. Biol. Chem. 248:6035-6040. 6. Rowe, P. B., M. D. Coleman, and J. B. Wyngaarden. 1970. Glutamine phosphoribosylpyrophosphate amidotransferase. Catalytic and conformational heterogeneity of the pigeon liver enzyme. Biochemistry 9: 1498-1505. 7. Rowe, P. B., and J. B. Wyngaarden. 1968. Glutamine phosphoribosylpyrophosphate amidotransferase. Purification, substructure, amino acid composition, and absorption spectra. J. Biol. Chem. 243:6373-6383. 8. Shiio, I., and K. Ishii, 1969. Regulation of purine ribonucleotide synthesis by end product inhibition. II. Effect of purine nucleotides on phosphoribosylpyrophosphate amidotransferase of Bacillus subtilis. J. Biochem. 66:175-181. 9. Trotta, P. P., L. M. Pinkus, and A. Meister. 1974. Inhibition by dithiotreitol of the utilization of glutamine by carbamyl phosphate synthetase. Evidence for formation of hydrogen peroxide. J. Biol. Chem. 249:1915-1921. 10. Tumbough, C. L. Jr., and R. L. Switzer. 1975. Oxygendependent inactivation of glutamine phosphoribosylpyrophosphate amidotransferase in stationary-phase cultures of Bacillus subtilis. J. Bacteriol. 121:108-114.
JOURNAL OF BACTERIOLOGY, Jan. 1975, P. 115-120 Copyright 0 1975 American Society for Microbiology
Oxygen-Dependent Inactivation of Glutamine Phosphoribosylpyrophosphate Amidotransferase In Vitro: Model for In Vivo Inactivation CHARLES L. TURNBOUGH, JR., AND R. L. SWITZER* Department of Biochemistry, University of Illinois, Urbana, Illinois 61801 Received for publication 21 October 1974
The oxygen-dependent inactivation of glutamine phosphoribosylpyrophosphate amidotransferase (ATase) is demonstrated in cell extracts of Bacillus subtilis. The rate of inactivation of ATase in vitro is apparently first order with respect to oxygen concentration and ATase activity. ATase inactivated in vitro (or in vivo) cannot be reactivated by a variety of reductants. ATase is significantly stabilized to oxygen-dependent inactivation in vitro in the presence of tetrasodium phosphoribosylpyrophosphate and glutamine together. The effects of the end product inhibitors, adenosine 5'-monophosphate (AMP) and guanosine 5'-monophosphate (GMP), on the stability of ATase are antagonistic. AMP stabilizes ATase, whereas GMP destabilizes the enzyme. The stability of ATase can be manipulated over wide ranges by variations in the AMP/GMP ratio. The effects of AMP and GMP on the inactivation of ATase in vitro are very specific. ATase is partially inhibited by 1,10-phenanthroline, suggesting that the enzyme contains iron (or some other chelatable metal ion). The inactivation of ATase in vitro is proposed to present a model for the reconstruction of the inactivation of ATase in stationary-phase cells of B. subtilis.
In the accompanying communication (10) we reported a rapid, oxygen-dependent inactivation of glutamine phosphoribosylpyrophosphate amidotransferase (ATase, EC 2.4.2.14) in stationary-phase cells of Bacillus subtilis. In an attempt to determine the chemical nature and means of regulation of this inactivation, we have attempted to reconstruct the process in cell extracts. In this paper we demonstrate the oxygen-dependent inactivation of ATase in vitro. Preliminary experiments on the chemical nature of the oxygen-sensitive sites of ATase are presented. Examination of the effects of substrates, end product inhibitors, and ATase concentration on the rate of enzyme inactivation has provided results that enable us to put forward a tentative model for the inactivation of ATase in stationary-phase cells of B. subtilis. (This work was presented in part at the Biochemistry/Biophysics 1974 Meeting, 2-7 June 1974, Minneapolis, Minn.) MATERIALS AND METHODS Chemicals. Chemicals and enzymes were obtained from the following sources: purine and pyrimidine bases, nucleosides and nucleotides and catalase, Sigma Chemical Co.; 1,10-phenanthroline, J. T. Baker Chemical Co.; 1,7-phenanthroline, G. Fred-
erick Smith Chemical Co.; [14Cjadenosine 5'-monophosphate (AMP), New England Nuclear Corp. All other chemicals, enzymes, and buffers were described previously (10) or were reagent grade and commercially available. Bacterial strains. All studies in this paper were conducted with B. subtilis strain 168, which requires tryptophan for growth. Media and culture methods. A buffered minimal medium containing 0.1% glucose described by Anagnostopoulos and Spizizen (1) was used for growth of cells. Culture methods were described in the accompanying paper (10). Preparation of cell extracts. In most experiments, 40-ml samples were withdrawn from a 1-liter culture at determined times during growth. The samples were harvested and stored as described previously (10). The pelleted cells were resuspended in 5 ml of cold buffer A (2) without 2-mercaptoethanol and 100 MAliters of a 2 mg/ml lysozyme solution. The cells were extracted as previously described (10). Enzyme assay. ATase activity was measured by assay I described in the accompanying paper (10). In most experiments 150-,uliter volumes of extract were assayed. It was confirmed that inhibitors of ATase used in this study did not inhibit the assay of glutamate. Incubation of extracts. When extracts were incubated anaerobically, all extraction procedures were performed in a dry box under argon. Extracts were incubated under argon in sealed tubes at 37 C, and samples were assayed at determined times. The tubes 115
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were flushed with argon during sampling and immediately resealed to eliminate oxygen. When extracts were incubated under 1.0 atm of oxygen, 4 ml of extract was placed in a 10-ml Erlenmeyer flask and the flask was capped with a serum stopper. The flask was flushed for 3 min with oxygen, sealed, and incubated at 37 C. In experiments where several samples were assayed at different times, the flask was flushed with oxygen after each sampling. Hydrolysis of AMP in extracts. [14C ]AMP (1 mM) was added to extracts of exponential- and stationary-phase cells and incubated at 37 C for 1.5 h. At determined times, samples were removed and applied to cellulose thin layers (Eastman Chromagram with fluorescent indicator). Carrier adenine and adenosine were spotted on the samples and the chromatograms were developed with 1 M ammonium acetate (4). "4C-labeled adenine, adenosine, and AMP were measured.
RESULTS Inactivation of ATase in vitro. In experiments described in the accompanying communication (10), we observed that ATase activity declined in cell extracts when they were incubated at 37 C, but that this loss of activity was prevented when the extracts were incubated under argon. This finding suggested that the oxygen-dependent inactivation of ATase occurs in vitro as well as in intact stationary-phase cells. The oxygen dependence of ATase inactivation in cell extracts was demonstrated by measuring the effect of oxygen concentration on the rate of inactivation at 37 C. The experiments presented in Fig. 1 show that ATase is stable in the absence of oxygen, and that the rate of inactivation is apparently first order with respect to oxygen concentration. The inactivation also appears to be first order with respect to ATase activity. The results are consistent with the idea that inactivation results from a simple reaction of ATase with molecular oxygen. ATase that was inactivated in vitro (or in vivo) could not be reactivated by incubation with 2-mercaptoethanol, cysteine, dithiothreitol, sodium borohydride, sodium dithionite, or a combination of ferrous chloride, sodium sulfide, and dithiothreitol. The experiments shown in Fig. 1 were conducted with extracts prepared from stationaryphase cells. To ascertain whether the ATase from exponentially growing cells, where no inactivation has taken place, was also susceptible to oxygen-dependent inactivation in vitro, extracts were prepared from cells harvested during exponential growth (48 units of ATase per ml) and incubated as in Fig. 1. The results were very similar to those shown in Fig. 1, except
_-
0.
1 2 Hours of Incubation (37C)
3
FIG. 1. Oxygen dependence of ATase inactivation in cell extracts. Three identical pellets of early stationary-phase cells were extracted as in Materials and Methods. The extracts (30 units of ATase per ml) were incubated under 0 (0), 0.2 (0), and 1.0 (A) atm of oxygen and 150-Aliter samples were assayed at indicated times.
that the rates of inactivation were slightly slower in the extracts of exponential-phase cells (e.g., t½ under air at 37 C was 90 min as opposed to 70 min in the stationary-phase cell extracts). This difference in rate of inactivation might reflect a difference in the concentration of some metabolite in the extracts that affects the inactivation process. It is also possible that the rate of ATase inactivation is dependent on the ATase concentration in the extract. Since the concentration of ATase in the exponentialphase cell extracts (48 units/ml) was higher than that of stationary-phase cell extracts (30 units/ml), one would predict that ATase inactivation would proceed more slowly at higher ATase concentrations if this hypothesis is correct. To examine these possibilities, two 10-ml extracts containing approximately 300 units of ATase per ml were prepared by extracting cells harvested during exponential growth in buffer A without 2-mercaptoethanol. One extract was diluted to several concentrations with buffer A without 2-mercaptoethanol. The second extract was dialyzed for 12 h at 4 C against two 2-liter changes of buffer A without 2-mercaptoethanol through which argon was bubbled continuously. The extract was then dialyzed for one additional hour against 2 liters of buffer A without 2-mercaptoethanol to remove argon. The
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dialyzed extract was diluted to several concentrations with buffer A without 2-mercaptoethanol. These extracts containing different concentrations of ATase were incubated under air at 37 C, and the rates of inactivation were measured as in Fig. 1 (Table 1). Dialysis of extracts containing 30 to 60 units of ATase per ml did not alter the rates of inactivation. Dialysis of more concentrated extracts, however, significantly altered the half-life of ATase, indicating that small molecules capable of stabilizing ATase were removed. The possibility that these results reflect the instability of a stabilizing element rather than its removal was eliminated by incubating a sample of the 294 units per ml of extract under argon at 4 C for 13 h. When this sample was incubated as above, only 8% (t½ 12 h) of the initial ATase activity was lost. The results also show that the inactivation of ATase proceeds more rapidly when the enzyme is more dilute in the extract. The addition of bovine serum albumin to extracts did not alter the rate of the inactivation. In subsequent experiments, therefore, only dilute extracts were used to minimize interferences by endogenous small molecules. We have also reported initial ATase activities and compared inactivation rates only in extracts with similar initial ATase activities. Effects of substrates and end product inhibitors on the inactivation of ATase in vitro. Studies of ATase from pigeon liver (6) and human placenta (5) have established that substrates and end product inhibitors alter the activity and physical conformation of the enzymes from these sources. These results prompted an investigation of the effects of these substances on the inactivation of B. subtilis ATase in vitro. Samples of late exponentialTABLE 1. Effect of ATase concentration on the irnctivation of ATase in vitro Initial ATase activity
(units/ml) 294 139 64 32 255 124 55 31
Protein concn (mg/ml)
(mg/mi) 5.9 3.0 1.5 0.7 6.4
3.2 1.6 0.8
t, (min)a without dialysis
te(mdi)al-
and early stationary-phase cells were harvested and extracted as described in Materials and Methods. Exponential-phase extracts contained 42 + 4 units of ATase per ml and stationary-phase extracts contained 29 ± 4 units of ATase per ml. After the addition of substrate(s) or purine nucleotide inhibitor(s), the extracts were incubated at 37 C under air or 1.0 atm of oxygen, and the rate of ATase inactivation was determined. It was demonstrated that the hydrolysis of tetrasodium phosphoribosylpyrophosphate (PRPP) and AMP in extracts of exponential- or stationary-phase cells was small during incubation (AMP hydrolysis < 7% of the initial concentration). The addition of 1 mM PRPP or 6 mM glutamine alone to extracts of either exponential- or stationary-phase cells had no significant effect on the rate of inactivation of ATase (Table 2). The effect of both substrates on the inactivation was measured by adding 2.5 mM PRPP and 6 mM glutamine together to an extract of exponential-phase cells and incubating this extract at 37 C for 30 min under 1.0 atm of oxygen. After 30 min, the concentration of PRPP had declined to about 1 mM because of product formation. The substrates and products were removed by dialysis, and the residual ATase activity was assayed. The stability of ATase was increased by nearly fourfold by the presence of both substrates (Table 2). The purine nucleotides AMP and ADP have been shown to be the most potent end product inhibitors of ATase from B. subtilis (8). The effect of AMP on the inactivation of ATase is shown in Fig. 2. Low concentrations of AMP greatly stabilize ATase activity. For comparison, the inhibition of ATase by AMP was also determined and plotted in Fig. 2. The stability of ATase was very substantially increased by concentrations of AMP that bring about relatively little inhibition. The inhibition of ATase by AMP followed a sigmoidal concentration
after dialysis
TABLE 2. Effect of substrates on the inactivation of ATase in vitroa
Stablec 207 90 78 139 104 92 82
Half-lives were measured at 37 C under air. bHalf-lives were measured after 12 h of dialysis under argon. c Corresponds to a 1 to 2% loss of ATase activity after 1.5 h of incubation. a
117
Addition
tA (min)b
None .......................... 1.0 mM PRPP .......................... 6.0 mM glutamine ........................ 2.5 mM PRPP plus 6.0 mM glutamine ......
17 4 2C 13
19 60 X 6d
ATase activity was 38 units/ml. bHalf-lives were measured at 37 C under 1.0 atm of oxygen. c Average of five determinations. d Average of two determinations. a Initial
118
J. BACTERIOL.
TURNBOUGH AND SWITZER
.ECY
.)
N
-4 a
CN
0.5
AMP (mM)
FIG. 2. Stabilization and inhibition of ATase by AMP. Stabilization of ATase to oxygen-dependent inactivation (0) was measured by adding the indicated amount of AMP to identical extracts of stationary-phase cells (32 units of ATase per ml) and incubating the extracts at 37 C under 1.0 atm of oxygen. Inhibition of ATase (0) was measured by adding the indicated amount of AMP to the assay mixture described in Materials and Methods.
dependence (Hill coefficient of 3.7 to 4.0), whereas the stabilization of ATase by AMP followed a hyperbolic dependence. An AMP concentration of 0.27 mM gave 50% inhibition under the conditions of assay I. Similar results were obtained when AMP was added to extracts of exponential-phase cells. Guanosine 5'-monophosphate (GMP) is a much weaker inhibitor of ATase from B. subtilis than AMP (8). Inhibition by GMP followed a hyperbolic concentration dependence with 50% inhibition at 2 to 3 mM under the conditions of assay I. The effect of GMP on the inactivation of ATase in vitro is shown in Fig. 3. In contrast to AMP, GMP decreased the stability of ATase to oxygen-dependent inactivation. The effect of GMP on stability was half-maximal at approximately 0.2 mM. Similar results were obtained when GMP was added to extracts of stationaryphase cells. Table 3 summarizes the results of an experiment in which both GMP and AMP were added to identical extracts of stationary-phase cells containing 32 units of ATase per ml. The effects of AMP and GMP clearly antagonize one another. The stability of ATase to oxygen-dependent inactivation in vitro can be manipulated over wide ranges by variations in the AMP/
1.0
1.5
GMP (mM)
FIG. 3. Destabilization of ATase by GMP. After the addition of the indicated amount of GMP to identical extracts of exponential-phase cells (38 units of ATase per ml), the extracts were incubated at 37 C under air and the half-lives were determined.
TABLE 3. Inactivation of ATase in the presence of both AMP and GMP ts (min)b GMP (mM) AMP (mM) 0.48 0.48 0.48 0.48
0
0 0.24 0.48 0.96 0.96
84 57 40 25 6
aInitial ATase activity was 32 units/ml. measured at 37 C under 1.0 atm of
b Half-lives were
oxygen.
GMP ratio in the extract. In separate experiments, it was shown that the addition of 0.5 or 1.0 mM GMP to extensively dialyzed extracts decreased the stability of ATase. Thus, GMP effects a true destabilization rather than acting to displace stabilizing nucleotides in the crude extract. The specificity of the effects of the nucleotide inhibitors of ATase on the inactivation of the enzyme was examined (Table 4}. It is evident that the nucleotide effects are quite specific; only 2'-deoxy AMP (and possibly cytidine 5'monophosphate) was able to significantly mimic the action of AMP. The effect of guanosine 5'-triphosphate may be due to formation of GMP in the crude extracts. Other nucleoside di- and triphosphates were not tested because of their possible breakdown in the crude extracts. Similar results were obtained with ex-
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INACTIVATION OF ATase IN VITRO
119
TABLE 4. Specificity of nucleotide effects on inactivation of ATase in vitroa Additions"
t,,(min)c
None ........... 17d 5'-AMP ........... 95 ....... 34 2'-deoxy 5'-AMP ... 1,10~~i, 0~~~~~~~~~~ 3-AMP ........... 16 ....... 18 3':5'-cyclic AMP ... 40 40_ 17 Adenosine .......... 19 Adenine .......... 5'-GMP ........... 7 20 10 5'-GTP .......... 3':5'-cyclic GMP .18 17 5'-XMP .......... 1.5 2.0 2.5 0.5 1.0 17 5'-IMP .......... 5'-UMP ........... 15 Phenanthroline (mM) 5'-CMP .......... 26 FIG. 4. Inhibition of ATase by phenanthroline. Inhibition was measured by adding the indicated a Initial ATase activity was 38 units/ml. of 1,7 phenanthroline (0) or 1,10phenanthroAll additions were 1.0 mM. Abbreviations not amount to the assay mixture described in Materials line (0) given in text: GTP, guanosine 5'-triphosphate; XMP, xanthine 5'-monophosphate; IMP, inosine 5'-mono- and Methods. phosphate; UMP, uridine 5'-monophosphate; CMP, trations, it was difficult to make a conclusive cytidine 5'-monophosphate. interpretation. c Half-lives were measured at 37 C under 1.0 atm of Recently it was reported by Trotta et al. (9) oxygen. d Average of five determinations. that carbamyl phosphate synthetase from Esch-
phenanthroline
erichia coli was inactivated by hydrogen peroxide generated in enzyme preparations containing low concentrations of dithiothreitol. To cells. Possible chemical nature of the oxygen- determine if inactivation of ATase in extracts of sensitive site of ATase. ATase purified from B. subtilis proceeds by such a mechanism, avian liver (3, 7) contains nonheme iron, which catalase (3,500 units/ml) was added to extracts appears to be involved in both the catalytic and the extracts were incubated at 37 C under reaction and structural integrity of the enzyme. air for 3 h. The rate of inactivation of ATase was Purified pigeon liver ATase is inhibited and the same in extracts with or without catalase. inactivated by 1,10-phenanthroline (6). To in- Catalase activity was measured after incubavestigate the possibility that ATase from B. tion and was > 80% of the initial activity. To subtilis contains nonheme iron, the inhibition of test whether the inactivation of ATase in vitro B. subtilis ATase by 1,10-phenanthroline was was due to photooxidation of a sensitive site on measured (Fig. 4). Inhibition of ATase by the enzyme, extracts were incubated at 37 C 1, 10-phenanthroline appears to be biphasic. under air in the dark. The inactivation of ATase Significant inhibition occurs at low 1,10-phen- was the same in the absence or presence of light. anthroline concentrations; however, the effect DISCUSSION of higher concentrations is appreciably less than InhiThe reconstruction of an oxygen-dependent would be expected with simple inhibition. bition of ATase by 1,7-phenanthroline was also inactivation of glutamine PRPP amidotransfermeasured. This isomer of phenanthroline binds ase represents a first step in elucidating the metals poorly and is often used as a control for chemical mechanism and physiological regula1,10-phenanthroline inhibition. At 0.25 mM tion of the inactivation of this enzyme in 1,7-phenanthroline, ATase activity was inhib- stationary-phase B. subtilis cells in vivo. The ited only 8%, but at higher concentrations the simple kinetics of the inactivation process suginhibition paralleled that observed with gest that the inactivation results from a direct 1,10-phenanthroline. These experiments sug- oxidation of some group on the enzyme, but the gest the presence of a chelatable metal ion; involvement of an inactivating enzyme can only however, because of the fractional inhibition at be conclusively ruled out by purification of the low 1,10-phenanthroline concentrations and ap- ATase. In either case, it is presumed that parent nonspecific inhibition at higher concen- inactivation results from oxidation of a group or tracts from exponential- and stationary-phase
120
TURNBOUGH AND SWITZER
groups that are essential for activity. In the case of a direct oxidation two groups are likely: sulfhydryl groups and ferrous iron. The observation that a variety of reducing agents failed to reactivate inactivated ATase indicates that the inactivation is not a simple oxidation of sulfhydryl groups to disulfides. ATases purified from avian liver (3, 7) have been shown to contain nonheme iron, which is essential for enzymatic activity. Inhibition of B. subtilis ATase by 1, 10-phenanthroline suggested that this enzyme also contains ferrous iron that is essential for activity. The dependence of the rate of oxygendependent inactivation on ATase concentration, together with the observations in the accompanying paper (10) that the activity of ATase is not linearly dependent on concentration, suggests that the enzyme may exist in an equilibrium between aggregated and disaggregated states. A similar equilibrium between aggregated and disaggregated forms has been demonstrated for the ATase of human placenta (5). The ATase purified from pigeon liver also appears to exist in several conformational states (6), and the existence of aggregated forms has been shown (7). It is clear that most of the remaining questions about the chemical nature of the oxygen-dependent inactivation of ATase in vivo could be answered by a study of the inactivation of the pure enzyme. Purification of the enzyme is currently under way in our
laboratory. The results of the study of the inactivation of ATase in vitro also suggest means by which the inactivation of the enzyme could be regulated in vivo. Neither PRPP nor glutamine alone affected the stability of ATase to oxygen, but the two substrates together stabilized the enzyme substantially. It is very likely that PRPP pools decline when cells are starved for carbon substrates and it is virtually certain that glutamine levels are depleted by nitrogen starvation. Thus, a marked drop in either substrate would signal starvation to the cell and would account for the destabilization of ATase in starved cells. The effects of the purine nucleotides, AMP and GMP, on the stability of ATase to oxygen were also specific and effective at relatively low concentrations. AMP stabilized ATase, whereas GMP destabilized the enzyme. The two nucleotides were able to antagonize the action of each other, so that the stability of ATase could be manipulated over wide ranges by variations in the AMP/GMP ratio. We would like to put forward the hypothesis that some combination
J. BACTERIOL.
of these effects of substrates and end product inhibitors could account for the stability of ATase to oxidation in growing cells and the oxygen-dependent inactivation in starved cells. It is possible that these ligands exert their effects on the stability of the enzyme by altering an equilibrium between disaggregated and aggregated forms of the enzyme. Effects of substrates and allosteric inhibitors on the physical state of ATase have been documented for the pigeon liver (6) and human placeta (5) enzymes. The validity of the hypothesis that substrates and/or end products regulate the stability of ATase in vivo can be most readily tested by determining the intracellular levels of the substrates and inhibitors of ATase under a variety of conditions where the enzyme is stable and where it is inactivated in vivo. LITERATURE CITED 1. Anagnostopoulos, C., and J. Spizizen. 1961. Requirements for transformation in Bacillus subtilis. J. Bacteriol. 81:741-746. 2. Deutscher, M. P., and A. Kornberg. 1968. Biochemical studies of bacterial sporulation and germination. VIII. Patterns of enzyme development during growth and sporulation of Bacillus subtilis. J. Biol. Chem.
243:4653-4660. 3. Hartman, S. C. 1963. Phosphoribosyl pyrophosphate amidotransferase. Purification and general catalytic properties. J. Biol. Chem. 238:3024-3035. 4. Hochstadt-Ozer, J., and E. R. Stadtman. 1971. The regulation of purine utilization in bacteria. I. Purification of adenine phosphoribosyltransferase from Escherichia coli K,, and control of activity by nucleotides. J.
Biol. Chem. 246:5294-5303. 5. Holmes, E. W., J. B. Wyngaarden, and W. N. Kelley. 1973. Human glutamine phosphoribosylpyrophosphate amidotransferase. Two molecular forms interconvertible by purine ribonucleotides and phosphoribosylpyrophosphate. J. Biol. Chem. 248:6035-6040. 6. Rowe, P. B., M. D. Coleman, and J. B. Wyngaarden. 1970. Glutamine phosphoribosylpyrophosphate amidotransferase. Catalytic and conformational heterogeneity of the pigeon liver enzyme. Biochemistry 9: 1498-1505. 7. Rowe, P. B., and J. B. Wyngaarden. 1968. Glutamine phosphoribosylpyrophosphate amidotransferase. Purification, substructure, amino acid composition, and absorption spectra. J. Biol. Chem. 243:6373-6383. 8. Shiio, I., and K. Ishii, 1969. Regulation of purine ribonucleotide synthesis by end product inhibition. II. Effect of purine nucleotides on phosphoribosylpyrophosphate amidotransferase of Bacillus subtilis. J. Biochem. 66:175-181. 9. Trotta, P. P., L. M. Pinkus, and A. Meister. 1974. Inhibition by dithiotreitol of the utilization of glutamine by carbamyl phosphate synthetase. Evidence for formation of hydrogen peroxide. J. Biol. Chem. 249:1915-1921. 10. Tumbough, C. L. Jr., and R. L. Switzer. 1975. Oxygendependent inactivation of glutamine phosphoribosylpyrophosphate amidotransferase in stationary-phase cultures of Bacillus subtilis. J. Bacteriol. 121:108-114.
