Vol. 28, No. 3 Printed in U.S.A.
JOURNAL OF BACTERIOLOGY, Dec. 1976, p. 766-775 Copyright © 1976 American Society for Microbiology
Unique Aspects of the Regulation of the Aspartate Transcarbamylase of Serratia marcescens JAMES R. WILD,' WILLIAM L. BELSER, AND GERARD A. O'DONOVAN* Department of Biology, University of California, Riverside, California 92502, and Department of Biochemistry and Biophysics, Texas A&M University, College Station, Texas 77843* Received for publication 7 July 1976
Aspartate transcarbamylase (ATCase; EC 2.1.3.2) from Serratia marcescens HY has been purified 134-fold. Its properties are unique. Unlike the ATCase from Escherichia coli and Salmonella typhimurium, the S. marcescens HY enzyme activity is not feedback inhibited by any purine or pyrimidine nucleotide effectors; instead, the enzyme is activated by both cytidine 5'-triphosphate and adenosine 5'-triphosphate. Like the ATCase from E. coli and S. typhimurium, adenosine 5'-triphosphate alters the [S],.5 of the enzyme and, in contrast, cytidine 5'-triphosphate does not alter the [S],.5 but, instead, alters the Vma.. As has been shown for both E. coli and S. typhimurium, effector sensitivity may be selectively dissociated from catalytic activity by treatment with heat, parachloromercuribenzoate, or neohydrin. The dissociated enzyme possesses threefold higher specific activity than the native enzyme. The sedimentation coefficient of the native enzyme is approximately 11.4S, whereas the dissociated enzyme has a value of 6.0S. Whereas it has been possible to reconstitute the E. coli and the S. marcescens ATCase enzymes from their own homologous subunits, it has not been possible to make hybrid enzymes of catalytic and regulatory heterologous subunits from each other. It was not possible to detect repression of ATCase formation after growth of prototrophic strains of S. marcescens HY supplemented with 200 ,tg of uracil per ml, but eightfold derepression was observed after uracil withdrawal in pyrimidine auxotrophs. Aspartate transcarbamylase (ATCase; EC study. Bethell and Jones have classified such 2.1.3.2) is the first enzyme unique to the path- ATCases as "B-type" enzymes (2); these allosway of pyrimidine biosynthesis and therefore terically regulated ATCases are subject to seoccupies an important position in the regula- lective dissociation of effector response and tion of pyrimidine nucleotide formation. Al- characterize all genera of the Enterobacteriathough ATCase is common to all microorga- ceae (2, 25, 27). Serratia marcescens belongs to the Enteronisms possessing a pathway for de novo pyrimidine biosynthesis, the allosteric control of the bacteriaceae (7), but differs from other memenzyme is effected by many nucleotides; the bers of the family in several respects. S. marnegative effectors that have been identified for cescens and Enterobacter (Aerobacter) aerodifferent systems include uridine 5'-triphos- genes represent an extreme in guanine + cytophate (UTP), cytidine 5'-triphosphate (CTP), sine content (58%) among the enterobacteria uridine 5'-monophosphate (UMP), and cytidine (23). The majority of the nucleotide sequences 5'-monophosphate (CMP) (2, 9, 14, 25, 27). AT- of strains of S. marcescens are not related to Case from Escherichia coli has been studied other members of the family by any of the extensively. It contains two nonidentical pro- criteria used by Brenner et al. (4), and thus tein subunits: one, the catalytic subunit, pro- Serratia appears to be the most divergent gevides the active sites of the enzyme, whereas nus of the family. These differences have been the other, the regulatory subunit, provides the supported by the determination of immunologibinding sites for nucleotide effectors (6, 12). cal indexes of dissimilarity of the tryptophan Table 1 summarizes some characteristics of the synthetase a subunit (24) and ,32 subunits (28) ATCase from E. coli that are pertinent to this in comparative studies. The extensive studies in the tryptophan pathway have been recently I Present address: Genetics Section, Texas A&M Uni- reviewed by Crawford (10). In this article we versity, College Station, TX 77843. describe the ATCase from S. marcescens HY 766
VOL. 128, 1976
S. MARCESCENS ATCase
767
TABLE 1. Properties of native ATCase and its catalytic and regulatory subunits from E. colia Property
(2C3) (3R2)
Native enzyme
Mercurials
M o Mercaptoethanol
2C3
Catalytic
Subunit structure ......... ...... (2C3) (3R2) 2(C3) Molecular weight ................ 310,000 2 (100,000) Sedimenation coefficient ......... 11.7S 5.8S Molecular weight of each polypeptide chain 34,000 Shape of saturation curve ........ Sigmoidal Hyperbolic Catalytic activity (binding of substrates) ....................... + + Binding of effectors ........ ...... + a Determined according to O'Donovan and Neuhard, with modifications (27).
and examine some of its unique regulatory properties. MATERIALS AND METHODS
+
3R2
Regulatory
3(R2) 3 (34,000) 2.8S
17,000
+
termination described by Gerhart and Pardee (14). Standard assay mixtures contained: 5 to 10 mM aspartate, pH 7.0; 0.4 mM carbamyl phosphate, pH 7.0; 40 mM potassium phosphate, pH 7.0; and 25 to Bacterial strains. S. marcescens HY wild type 50 ,ul of extract. The volume was adjusted to 0.50 ml and pyrimidine-requiring auxotrophs derived from with distilled water. A final nucleotide concentrathis strain were used in these studies: HYpyrF3 was tion of 2 mM was utilized for all effector studies confirmed by enzyme assay and shown to lack oroti- unless otherwise specified. The reaction mix was dine 5'-phosphate decarboxylase activity (EC incubated at 30°C for 15 min at an extract concentra4.1.1.23); HY pyrEl lacks orotate phosphoribosyl tion such that the final absorbance developed by the transferase (EC 2.4.2.10). colorimetric assay at 560 nm was 0.400 to 0.800. The Growth conditions. The minimal medium used amount of carbamyl aspartate produced was deterwas Serratia minimal medium (SM) defined by La- mined three times during the 15-min reaction to brum and Bunting (21) and contained, per liter of ensure linearity. distilled water: K2HPO4, 8.0 g; ammonium citrate Enzyme hybridization. ATCase was partially dibasic, 5.0 g; MgSO4- 7H2O, 0.5 g; FeCl3, 0.02 g. The purified from a pyrimidine auxotrophic strain wild-type strain HY was grown overnight in 1-liter (pyrF3) of S. marcescens HY by the method of quantities of SM at 30°C. The cells were harvested Gerhart and Holoubek (13), with one modification: by centrifugation during the exponential phase of in the heat step we used a temperature of 58°C ingrowth (optical density at 660 nm = 0.400). The wild stead of 72°C for E. coli. An overall purification of type was used to prepare enzymes for effector stud- 134-fold was achieved, thus giving a protein prepaies. ration homogeneous by gel electrophoresis. Bacterial auxotrophs were grown in SM + uracil ATCase of E. coli K-12 was prepared from a leaky (25 ug/ml). Derepression was achieved by allowing pyrF strain by the method of Gerhart and Holoubek the cells to deplete the available uracil supply at an without modification (13). In both cases the native aggregates were dissooptical density at 660 nm of about 0.250 and continuing starvation for 90 to 120 min in the absence of ciated by p-chloromercuribenzoate (p-CMB) or neouracil. Pyrimidine auxotrophs of S. marcescens re- hydrin treatment (26; M. W. Kirschner, Ph.D. thequired uniquely high levels (50 ,ug/ml) of uracil for sis, University of California, Berkeley, 1971), and maximal yield. the subunits were separated by zone centrifugation Preparation of cell extracts. Cell extracts were on a 5 to 25% sucrose gradient and diethylaminoprepared by sonication of washed bacterial suspen- ethyl-cellulose anion-exchange chromatography, as sions as described by Hutchinson and Belser (19). previously described (16). All extracts were passed over Sephadex G-25 or diChemicals. The chemicals used in these studies alyzed overnight against a 1,000-fold excess of 50 were commercially obtained. 2-Mercaptoethanol, mM potassium phosphate buffer, pH 7.5, containing ethylenediaminetetraacetic acid, dithiothreitol, L2 ,uM ZnSO4. During sonication the extracts were aspartate, and carbamyl phosphate (dilithium salt) maintained below 8°C. The extract was clarified by were purchased from Sigma. p-CMB (sodium salt) centrifugation at 39,000 x g for 1 h. The enzyme and the following nucleotides and nucleosides were preparation was stable for several weeks when obtained from Calbiochem: CTP, cytidine 5'-diphosphate, cytidine 5'-monophosphate (CMP), cytidine, stored at 40C. Analytical methods. Protein was estimated by UTP, uridine 5'-diphosphate, UMP, uridine, guanothe method of Lowry et al. (22), with crystalline sine 5'-triphosphate, guanosine 5'-monophosphate, bovine serum albumin as the standard. The enzy- adenosine 5'-triphosphate (ATP), and adenosine 5'matic activity of ATCase was measured by the de- monophosphate. Catalase was obtained from Wortermination of carbamyl aspartate formed. Carba- thington Biochemicals. Neohydrin was a gift from J. myl aspartate was assayed by the colorimetric de- C. Gerhart.
768
J. BACTERIOL.
WILD, BELSER, AND O'DONOVAN
RESULTS and reassociation. dissociation Enzyme The reconstitution of ATCase from catalytic and regulatory subunits prepared by dissociation withp-CMB is summarized in Table 2. The data are presented as relative enzyme activity (1 unit = 1.0 ,umol of carbamyl aspartic acid produced/min per 25 ,ul of initial sample). Thus, the same concentration of ATCase was assayed for each purified extract before and after dissociation. Separation of the subunits produced a threefold increase in catalytic activity for E. coli (0.220 to 0.661 units) and for S. marcescens (0.440 to 1.240 units). After dissociation from the 3.OS regulatory subunit, the 6.0S catalytic subunit did not possess an effector response to 2 mM CTP for E. coli (0.661 versus 0.665 units) and S. marcescens (1.240 versus 1.257 units). It was possible to reconstitute both the E. coli and S. marcescens native enzymes from their respective homologous subunits and regain the original effector response to CTP, accompanied by a decrease in relative enzyme activity in the absence of effector. The ratio of the percentage of activity in the presence and absence of CTP provides a valuable index for evaluating the reconstitution. The inhibition of ATCase from E. coli and the activation of the ATCase from S. marcescens were virtually identical before dissociation and after reconstitution. We have been unable to reconstitute the heterologous (hybrid) subunits and form the hybrid enzyme that possessed either positive or negative effector response; i.e., the catalytic subunit from S. marcescens mixed with the regulatory subunit from E. coli did not possess any effector response to CTP.
Physical characterization of ATCase. (i) Estimation of sedimentation coefficients. Sedimentation coefficients were determined for the ATCase of S. marcescens HY by sucrose gradient sedimentation. A sample of 0.2 ml of fresh enzyme extract (20 mg/ml) was layered onto a 5-ml, linear 5 to 25% sucrose gradient. The native ATCase was centrifuged in a Beckman Spinco L-2 ultracentrifuge for 13 h at 39,000 rpm at 2°C in an SW39 rotor. The desensitized enzyme (treated with 1 mMp-CMB) was centrifuged for 17 h under the same conditions. Three-drop samples were collected from the bottom of the gradients and diluted with 0.25 ml of 50 mM potassium phosphate buffer, pH 7.5. Each sample was assayed for ATCase activity, and sedimentation profiles are shown in Fig. 1 and 2. Using a value of 11.4S for the catalase marker, sedimentation coefficients were estimated for the enzymes under both conditions (19). The native enzyme has a value of 11.4S and the dissociated enzyme has a value of 6.OS. (ii) Response to nucleotide effectors. The effects of various nucleotides, nucleosides, and
I-
E
5; I-
0.10
TABLE 2. Reconstitution ofATCase from E. coli and S. marcescens HY Relative activity % ActivType of en(+tPI Source - CTP + CTP (+CTP/ zyme 6.0a
E. coli E. coli
0.220 0.661
0.106 0.665
48.1 100.6
3.0b
E. coli
0.233 0.440 1.240
0.096 0.970 1.257
41.1 220 107
0.176
0.334
190
0.350
0.360
102.8
11.7
11.4 6.0a 6.0b
3.0b 6.0b
3.0b
S. marcescens S. marcescens S. marcescens S. marcescens S. marcescens E. coli I
Effector response selectively dissociated by treatment with p-CMB. b Reconstitution of subunits separated by treatment with p-CMB. a
N
I 0
z
z
LUJ
co
005 m
0
5
10 15 20 FRACTION NUMBER
25
FIG. 1. Sucrose gradient sedimentation of native ATCase from S. marcescens HY. A sample of0.4 mg of crude extract was layered onto a 5-ml, linear 5 to 25% sucrose gradient and centrifuged at 39,000 rpm for 13 h in an SW39 rotor at 2°C. Symbols: ATCase (O); catalase (0), used as a marker. The sloping line represents the relative sedimentation values developed by the gradient.
S. MARCESCENS ATCase
VOL. 128, 1976 2.0
E
0.10
§
1.0 w
~~~~~~~z
z
~ ~ ~
0
5
~
10
~
~
~
15
~
20
-0.05
25
FRACTION NUMBER
FIG. 2. Sucrose gradient sedimentation ofp-CMB (2 mM)-treated ATCase from S. marcescens HY. A sample of 0.4 mg oftreated extract was layered onto a 5-ml, linear 5 to 25% sucrose gradient and centrifuged at 39,000 rpm forl3 h in an SW39 rotor at2°C. Symbols: ATCase (a); catalase (0) was used as a marker, measured by absorbance at 405 nm.
free bases on the activity of ATCase in S. marHY wild type were studied. Stimulation by various effectors was determined under standard assay conditions, using an aspartate concentration of 5 mM. No inhibition was observed with any of the effectors (Table 3). Cytosine compounds stimulated enzymatic activity in the order: CTP > cytidine 5'-diphosphate > CMP > cytidine. Deoxycytidine compounds were equally effective as activators as the ribonucleotides. Under similar conditions, ATP was the most effective activator and guanosine 5'-triphosphate showed neither inhibition nor activation. The pH of the reaction mix was then varied in an attempt to identify negative effectors; none were discovered. Various concentrations of aspartate and carbamyl phosphate were included in the assay mix with different potential inhibitors, but still no inhibition was found. (iii) Combinations of effectors. Table 4 presents the effect of various combinations of nucleotides upon the activity of ATCase. The reaction conditions were identical to those used for Table 1 studies, except 2 mM concentrations of each effector were used. The total effector con-
cescens
769
centration was 4 mM. Guanosine 5'-triphosphate, UTP, and UMP had no effect on the activation by CTP. The nucleoside monophosphates in different combinations had no effect on enzymatic activity. The combination of ATP and CTP showed an intermediate response when compared with either effector alone. (iv) Saturation of effectors. The saturation of the nucleotide effectors ATP and CTP is summarized in Fig. 3. At concentrations up to 5 mM, UMP or CMP shows no effect on the activity of ATCase. Both CTP and ATP show saturation of effector activation at 2.5 mM. (v) Kinetics of substrate dependence. Figures 4 and 5 summarize the dependence of ATCase activity of S. marcescens HY on the concentrations of carbamyl phosphate and aspartate. In the absence of effectors, the native enzyme shows a sigmoidal response when the aspartate concentration is varied (Fig. 4) and TABLE 3. Effect of various nucleotide and nucleoside effectors on the activity of S. marcescens HY ATCasea Effector
Enzyme activity (carbamyl as partate produced)"
% Activityc
100 25.2 None 189 47.7 CTP 142 34.8 CDP 108 27.2 CMP 105 26.6 Cytidine 100 25.3 UTP 98 24.5 UDP 97 24.4 UMP 102 25.8 Uridine 101 25.6 dTTP 108 27.2 dTMP 101 25.6 Thymidine 187 47.2 dCTP 110 27.9 dCMP 104 26.2 dUTP 97 24.4 dUMP 104 26.3 GTP 101 25.5 GMP 249 62.9 ATP 96 24.2 AMP a ATCase activity was determined in the presence of 2 mM effector, 5 mM i-aspartate, 0.4 mM carbamyl phosphate, at pH 7.0. Abbreviations: CDP, Cytidine 5'-diphosphate; UDP, uridine 5'-diphosphate; dTTP, deoxythymidine 5'-triphosphate; dTMP,
deoxythymidine 5'-monophosphate; GTP, guanosine 5'-triphosphate; GMP, guanosine 5'-monophosphate; AMP, adenosine 5'-monophosphate. b Expressed as nanomoles of carbamyl aspartate produced in 15 min at 300C per milligram of protein. c
The activity of ATCase without effectors is set
at 100%.
WILD, BELSER, AND O'DONOVAN
J. BACTZRIOL.
TABLz 4. Effect of combinations of nucleotide effectors on the activity of ATCase from S. marcescens Hya
ment, activity was not lost, but actually increased 2.7-fold (Table 5). A high ionic strength 1.0 M (NH4)2S04 and high pH (pH 9.2) provide heat stability to effector response ofthe enzyme and were utilized during partial purification of the enzyme. Enzymatic activity and effector response were determined as described for Table 3. Another method utilized to separate effector response from catalytic activity was treatment with the mercurialp-CMB at 2 mM. Enzymatic activity was greatly reduced (90%) by the initial treatment with this concentration of mercurial. Specific recovery of the enzymatic activity, without effector response, was observed after 3 h of dialysis against 100 volumes of potassium phosphate buffer, pH 7.5, containing 50 mM dithiothreitol and 0.1 mM ethylenediaminetetraacetic acid. Consistent with the report of Gerhart and Pardee for E. coli ATCase (14), the loss of effector response by treatment with p-CMB resulted in a threefold increase in the specific activity of ATCase over the untreated enzyme. Nucleotide activation was also separated from catalytic activity by treatment with neohydrin at 3 mM. (vii) pH studies. Effector responses showed
770
Effector
Carbamyl aspartate produced"
% Activity,
100 25.2 189 47.7 249 62.9 100 25.3 104 26.3 97 24.4 108 27.2 101 25.5 221 55.9 190 48.0 GTP+CTP 178 44.9 UTP+CTP 187 47.2 UMP+CTP 98 24.5 UMP+CMP 105 26.6 GMP+CMP a ATCase activity was determined, as in Table 3, in the presence of 2 mM nucleotide. When the effect of two nucleotides present simultaneously was determined, 2 mM concentrations of each potential effector were present. See Table 3 for abbreviations. b Expressed as nanomoles produced in 15 min at 30°C per milligram of protein. c The activity of ATCase without effectors is set at 100%.
None CTP ATP UTP GTP UMP CMP GMP ATP+CTP
an apparently hyperbolic response to variations in carbamyl phosphate concentration (Fig. 5). The sigmoidal response to aspartate is maintained in the presence of effectors. However, CTP elicits an alteration in V., whereas ATP causes a change in the IS .5. The estimated Hill coefficient (11) in the presence of CTP is similar to that in the absence of effector (napp = 1.8 to 1.9). The value is somewhat lower in the presence of ATP (napp = 1.5 to 1.6). Substrate activation by aspartate is observed in the presence and absence of effectors. The [SL.5 of the native enzyme for aspartate is approximately 65 mM. The [SL.5 for aspartate in the presence of ATP is approximately 20 mM. The hyperbolic response of carbamyl phosphate concentration and velocity is maintained in the presence of nucleotide effectors, and the activation appears to be uncompetitive (see Fig. 5, inset). (vi) Selective destruction of effector response. Selective destruction of activation by both CTP and ATP was obtained by three independent methods. When the cell extract (3 ml) was heated at 60°C for 3 min in 50 mM potassium phosphate buffer, pH 7.5, ATCase could not be activated by 5 mM CTP or ATP, nor could it be inhibited by UMP or CMP. 2-Mercaptoethanol, dithiothreitol, or ethylenediaminetetraacetic acid (1 mM) did not provide any protection to the enzyme. After heat treat-
k-C-) 4
5 4 3 EFFECTORS x 10-3 FIG. 3. Saturation of effectors of S. marcescens HY ATCase. Enzyme activity (micromoles of carbamyl aspartate x 10115 min per milligram of protein at30°C) was determined in the presence of increasing concentrations of nuclotides. Symbols: CTP (0); ATP (-); and UMP or CMP (U) in the standard reaction mix containing 5 mM aspartate, 0.4 mM carbamyl phosphate, and 40 mM phosphate buffer, pH 7.0. 2 1 MOLARITY OF
VOL. 128, 1976
S. MARCESCENS ATCase
P 2.0 w
>.
1.5
w
0
20 40 60 80 MOLARITY OF L-ASPARTATE x10-3
100
FIG. 4. Velocity-substratesaturationcurvesinthe of several nucleotides (at 2 mM concentration) at pH 7.0. Enzyme activities are expressed as micromoles of carbamyl aspartate x 100/15 min per milligram of protein at 30°C. The reaction mixes contained 0.5 mg of protein with 0.4 mM carbamyl phosphate and 40 mM, potassium phosphate, and varying concentrations of aspartate were used to initiate the reaction. Symbols: ATP (a); CTP (0); CMP or UMP (U); no effector (O). presence
771
(viii) Effect of 2-thiouracil. 2-Thiouracil has been shown to be a growth inhibitor when added to exponentially growing cultures of E. coli (5). It has been proposed that 2-thiouracil is converted to 2-thiouridine-5'-monophosphate, which functions as a CTP analogue in inhibiting the catalytic activity of ATCase (17). 2Thiouracil (50 ,ug/ml, final concentration) was added to an exponentially growing culture of wild-type S. marcescens HY. No alteration of growth rate was noticed in liquid cultures or on minimal agar plates to which the analogue was added. Such a concentration of 2-thiouracil inhibited the growth of E. coli in both cases. Derepression of ATCase. Table 6 is a summary of a series of studies that demonstrate that the formation of ATCase may be derepressed in auxotrophs of S. marcescens HY. Cultures in mid-log growth in SM + uracil (50 jig/ml) were centrifuged at 10,000 x g, and the bacterial cells were resuspended in SM without uracil. The specific activity of ATCase in pyrimidine auxotrophs increased ninefold following incubation for 2 h after removal of uracil. The parental strain produced similar low levels of ATCase in response to various concentrations of uracil or arginine in the growth medium.
DISCUSSION ATCase catalyzes the first biosynthetic reaction unique to the formation of pyrimidine nucleotides and involves the carbamylation of aspartate by carbamyl phosphate. This enzyme is found in procaryotic and eucaryotic organisms, but its structural and regulatory properties extreme variation in relation to pH. Figure 6 show wide diversity (1, 2, 26, 27). The ATCases shows the pH profile for ATCase from S. mar- from different organisms are positively and cescens and the effector responses for ATP, negatively regulated by a variety of nucleotide CTP, and UMP. Tris(hydroxymethyl)amino- effectors. Most of the enteric bacteria are allomethane buffer was used in these studies for sterically inhibited by CTP or UTP. The enpH 10.0 to 7.0. Phosphate buffer was used for zymes from E. coli and S. typhimurium have pH 7.5 to 6.0. In the absence of effectors, the been extensively studied and provide a basis for S. marcescens HY enzyme shows an extremely allosteric effector regulation, regulatory and broad spectrum of activity. This activity is catalytic protomer interactions, and potential greatly enhanced at lower pH upon the addition taxonomic and evolutionary comparisons (12, of either CTP or ATP. It has been reported 15, 27). that the ATCase from a different strain of S. marcescens is inhibited by 0.013 mM UMP or CMP at pH 8.5 (25). We were unable to detect inhibition (greater than 10%) at pH 7.0, 8.5, or 10.0 under similar conditions or with 20-fold higher mononucleotide concentrations in S. marcescens HY. Even at extremely low substrate concentrations no inhibition was observed. This was true for crude extracts and for the partially purified enzyme.
S. marcescens HY possesses a unique ATCase that is not feedback inhibited by pyrimidine or purine nucleotides, nucleosides, or free bases; instead it is activated by a pyrimidine nucleotide, CTP, and by a purine nucleotide, ATP. Neumann and Jones (26), using a different strain of S. marcescens, demonstrated that ATCase was inhibited by UMP and CMP. Extensive attempts have been made to verify this result in strain HY, without success. Preincu-
772
J. BACTERIOL.
WILD, BELSER, AND O'DONOVAN
2.5
2.0 I-
1.5 4
w N 1.0 z w
0.5X e6
1/
r*
0
5
32
MOLANTY OF CARBAMYL PHOSPHATE
48 X 10 4
64
25 20 15 MOLARITY OF CARBAMYL PHOSPHATE x 104 10
FIG. 5. Velocity-substrate saturation curves in the presence of several nucleotides each at 2 mM concentration, pH 7.0. Enzyme activities are expressed as micromoles of carbamyl aspartate producedll5 min per milligram of protein at 30°C. The reaction mixes contained 0.5 mg of protein with 10 mM aspartate and 40 mM potassium phosphate, and varying concentrations of carbamyl phosphate were used to initiate the reaction. Symbols: ATP (a); CTP (0); CMP (*); no effector (E); UMP (-). (Inset) Double-reciprocal plot of carbamyl phosphate saturation for ATCase in the presence of ATP (-), CTP (0), and without effector (O).
bation of the enzyme with effectors for up to 20 min, variation of pH, increased effector concentration to 5 mM, and variation of substrate concentration have failed to establish any conditions in which inhibition could be demonstrated. Neumann and Jones did not report any effect of CTP on Serratia ATCase, but activation of ATCase by CTP was reported in Proteus vulgaris by Martha Bethell (Ph.D. thesis, Brandeis University, Waltham, Mass., 1967). The Proteus extract, however, contained two molecular forms of ATCase, an activatable form with a molecular weight of 300,000 and a "nonresponsive" form of molecular weight 100,000. The native ATCase of S. marcescens HY found in our study has a molecular weight of approximately 300,000.
Although ATP and CTP both function as activators, their mode of activation is different. ATP functions as an activator by altering the [S10.S5 for aspartate at saturating concentrations of carbamyl phosphate. ATP activates the ATCase from E. coli and S. typhimurium in the same manner and competes with CTP for the same binding sites (6). CTP is a competitive inhibitor with aspartate in these systems, but in S. marcescens HY CTP is an uncompetitive activator. Indeed, CTP alters the Vmax of ATCase when aspartate concentration is varied but the same [Sk.5 is maintained. This hitherto unreported observation suggests that there may be a fundamental difference in the mechanism of activation of these two effectors. This observation is in accord with the proposal of
VOL. 128, 1976
S. MARCESCENS ATCase
TABLE 5. Selective destruction of nucleotide activation of ATCasea Treatment and effector
Carbamyl as-
producedb
partate
%
Activityc
Untreated None CTP ATP
27.0 49.0 63.0
100 181 234
Heatd None +CTP +ATP
74.0 73.0 75.0
271 270 273
773
which is considerably higher than the [S6.5 of the E. coli or S. typhimurium enzyme, 2 mM, at approximately the same carbamyl phosphate concentration (26). Thus, a great increase in aspartate level is required for maximal enzymatic activity. Furthermore, this high concentration of aspartate is accompanied by substrate activation of the enzyme. It appears that
2.5
p-CMBe None +CTP +ATP
3.0 2.5
11 9
p-CMBf None 90.0 333 +CTP 328 88.5 +ATP 93.5 346 a ATCase activity is determined in the presence of 2 mM effector, 5 mM L-aspartate, 0.04 mM carbamyl phosphate. b Expressed as nanomoles produced in 15 min at 30°C per milligram of protein. c The activity of ATCase without effectors is set at 100%. d After heat treatment (60°C for 15 min). e After treatment with 2 mM p-CMB. f After treatment with 20 IAM p-CMB.
2.0 C-
1.5 N
z w
1.0
0.5
pH
Wedler and Gasser (29), which suggests that CTP and ATP act by separate mechanisms in affecting aspartate binding inE. coli. ATP may directly affect the rate of aspartate association/ dissociation, whereas CTP induces an intramolecular alteration in the R-C domain of binding (for review, see reference 12). In other closely examined systems, it has been demonstrated that both positive and negative effectors alter the [Sk.5 of the respective ATCases and their effect is overcome by high substrate concentration. Activation studies with CTP and ATP simultaneously present during assay of the S. marcescens ATCase suggest that CTP and ATP compete with one another for activation and act independently of one another. Saturating concentrations of CTP and ATP result in compensatory activation with an intermediate effector response (Table 5). High substrate levels minimize activation by ATP, but activation by CTP is maintained. The kinetics of ATCase activity-substrate plots reveal additional unique characteristics of this enzyme. The concentration of aspartate required for half-maximal saturation is 65 mM,
FIG. 6. Effect of pH on the activation of ATCase by nucleotides. Enzyme activity was determined in the presence of 5 mM aspartate, 0.4 mM carbamyl phosphate, 2 mM effectors. Symbols: CTP (0); ATP (0); UMP or CMP (U); and without effector present (O). Enzyme' activity is expressed as micromoles of carbamyl aspartatel15 min per milligram of protein at 30°C.
TABLE
6. Derepression of ATCasea in wild-type strain HY and mutant pyrEl Strain Growth medium Sp actb SM HY 74.0 HY SM + uracil 63.0 HY SM + arginine 70.0 pyrEl SM + uracil 58.0 pyrEl SM - uracilc 464.0 a Strains were grown in SM supplemented with uracil or arginine (50 ,ug/ml) and harvested in midlog phase (optical density at 660 nm = 0.400 to 0.500). b Expressed as nanomoles of carbamyl aspartate produced per minute per milligram of protein. c Mutant strain pyrEl was grown in the presence of uracil (50 ug/ml), suspended in the absence of uracil, and allowed to incubate for 2 h.
774
WILD, BELSER, AND O'DONOVAN
the saturation curve for carbamyl phosphate is hyperbolic (Fig. 5), but as pointed out by Bethell and co-workers, a more sensitive assay might reveal some cooperative interaction (3). The effects of CTP and ATP are uncompetitive with respect to carbamyl phosphate concentration. The [S]o.5 for carbamyl phosphate, 0.15 mM, is comparable to that ofthe E. coli enzyme (3). ATCase is composed of two types of protein subunits (Table 1), and the purified native enzyme from E. coli has been separated into regulatory and catalytic subunits by p-CMB (15). Similarly, we have selectively dissociated regulatory activity from catalytic activity in S. marcescens HY by heat treatment or by p-CMB. Once dissociated, the catalytic activity was not affected by either ATP or CTP. The desensitized enzyme is shown to be a smaller form, approximately 100,000 daltons, with an S value of 6.0. Thus, the size of the S. marcescens HY ATCase is similar to other type B ATCases before and after dissociation (2). In E. coli and S. typhimurium the subunit dissociation by p-CMB is readily reversible by incubation in 2 mM mercaptoethanol in high ionic strength, 1.0 M (NH4)2SO4, and at pH 9.2 tris(hydroxymethyl)aminomethane buffer. Reconstitution of the S. marcescens HY native enzyme has been achieved from its regulatory and catalytic subunits, and activation by CTP and ATP was restored. Further work on the purification of ATCases from different genera of bacteria is in progress in our laboratory (J. R. Wild, K. F. Foltermann, and G. A. O'Donovan, manuscript in preparation). It is also interesting to speculate on the overall mode of regulation of enzyme activity of pyrimidine biosynthesis in Serratia. The first consideration is that CTP activation of ATCase might be important to increase the flow of pyrimidine intermediates through the pathway. This flow increase might be necessary due to a more sensitive effector response at some other point in the pathway. Carbamyl phosphate synthetase is the most likely target to be considered for such a stringent regulatory role. Aspartate has an extremely high [S]05 for ATCase in S. marcescens (65 mM), and the relative effect of CTP and ATP at this level of aspartate is quite different. At physiological levels of aspartate (5 mM), ATCase is relatively unresponsive to changes in CTP levels, whereas the enzyme is much more responsive to changes in the levels of aspartate and ATP. High endogenous pool levels of CTP may compete with ATP for activation and depress the extent of activation. Therefore, CTP inhibits the activation by
J. BACTERIOL.
ATP and reduces the production of carbamyl aspartate compared to the ATP-activated enzyme without CTP. One further point seems pertinent, namely, that extremely high concentrations of uracil are required to provide maximal growth of pyrimidine auxotrophs of S. marcescens HY in liquid culture (Wild, unpublished data). All of the enzymes in the pathway may be partially derepressed, even in wild-type strains, in order to provide for the unusually high uracil requirement. Thus, the situation may parallel the system for Pseudomonas aeruginosa PAO (20) and P. putida (9), in which very little inhibition was found for ATCase and the entire pathway beyond ATCase seemed to be constitutive. It is possible to elevate levels of ATCase activity in Pseudomonas and Serratia by withdrawal of uracil in pyrimidine auxotrophs, but levels of enzyme cannot be repressed by growth on uracil in prototrophs. The activities of the subsequent enzymes of the de novo pathway in S. marcescens HY vary in starved auxotrophs, but the levels vary only two- to threefold (18). Even so, this variation may be the result of polypeptide interactions in a multifunctional complex rather than "derepression" per se (J. R. Wild and W. L. Belser, Biochem. Genet., in press). These observations suggest that S. marcescens HY might be producing the enzymes involved in pyrimidine biosynthesis at very high levels and might not be capable of conventional "repression" and "derepression." ACKNOWLEDGMENTS This work was supported by research grants from the Office of Naval Research, the Robert A. Welch Foundation, Houston, Tex., and the Texas Agricultural Experiment Station. One of us (J.W.) was a National Science Foundation Predoctoral Fellow. LITERATURE CITED 1. Adair, L. B., and M. E. Jones. 1972. Purification and characteristics of aspartate transcarbamylase from Pseudomonas fluorescens. J. Biol. Chem. 247:23082315. 2. Bethell, M. R., and M. E. Jones. 1969. Molecular size and feedback regulation characteristics of bacterial aspartate transcarbamylases. Arch. Biochem. Biophys. 134:352-365. 3. Bethell, M. R., K. E. Smith, J. S. White, and M. E. Jones. 1968. Carbamyl phosphate: an allosteric substrate for aspartate transcarbamylase of Escherichia coli. Proc. Natl. Acad. Sci. U.S.A. 60:1442-1449. 4. Brenner, D. J., G. R. Fanning, K. E. Johnson, R. V. Citarella, and S. Falkow. 1969. Polynucleotide sequence relationships among members of Enterobacteriaceae. J. Bacteriol. 98:637-650. 5. Cardeilhac, P. T. 1967. A toxic effect of 2-thiouracil on pyrimidine metabolism. Proc. Soc. Exp. Biol. Med. 125:692-696. 6. Changeux, J.-P., J. C. Gerhart, and H. K. Schachman. 1968. Allosteric interactions in aspartate transcarbamylase. I. Binding of specific ligands to the native
VOL. 128, 1976
7.
8. 9.
10. 11.
12. 13.
14.
15.
16.
17.
enzyme and its isolated subunits. Biochemistry 7:531-537. 18. Cocks, G. T., and A. C. Wilson. 1972. Enzyme evolution in the Enterobacteriaceae. J. Bacteriol. 110:793-02. Coleman, M. S., and M. E. Jones. 1971. Aspartate 19. transcarbamylases of Citrobacter freundii. Biochemistry 10:3390-3396. Condon, S., J. K. Collins, and G. A. O'Donovan. 1976. 20. Regulation of arginine and pyrimidine biosynthesis in Pseudomonas putida. J. Gen. Microbiol. 92:375383. 21. Crawford, I. P. 1975. Gene rearrangements in the evolution of the tryptophan synthetic pathway. Bacteriol. Rev. 39:87-120. 22. Endrenzi, L., C. Fazi, and F. H. F. Kwong. 1975. Evaluation of Hill slopes and Hill coefficients when saturation binding or velocity is not known. Eur. J. 23. Biochem. 51:317-328. Gerhart, J. C. 1970. A discussion of the regulatory properties of aspartate transcarbamylase from Esche- 24. richia coli. Curr. Top. Cell. Regul. 2:275-325. Gerhart, J. C., and H. Holoubek. 1967. The purification of aspartate transcarbamylase ofEscherichia coli and separation of its protein subunits. J. Biol. Chem. 25. 242:2886-2892. Gerhart, J. C., and A. B. Pardee. 1962. The enzymology of control by feedback inhibition. J. Biol. Chem. 26. 237:891-896. Gerhart, J. C., and H. K. Schachman. 1965. Distinct subunits for the regulation and catalytic activity of aspartate transcarbamylase. Biochemistry 4:1054- 27. 1062. Gerhart, J. C., and H. K. Schachman. 1968. Allosteric interactions in aspartate transcarbamylase. II. Evi- 28. dence for different conformational states of the protein in the presence and absence of specific ligands. Biochemistry 7:538-552. Goodrich, M. E., and P. T. Cardeilhac. 1970. 2-Thiouri- 29. dine-5'-phosphate and its inhibition of aspartate tmnscarbamylase. Biochim. Biophys. Acta 222:621-
S. MARCESCENS ATCase
775
626.
Hayward, W. S., and W. L. Belser. 1965. Regulation of pyrimidine biosynthesis inSerratia marcescens. Proc. Natl. Acad. Sci. U.S.A. 53:1483-1489. Hutchinson, M. A., and W. L. Belser. 1969. Enzymes of tryptophan biosynthesis in Serratia marcescens. J. Bacteriol. 98:109-115. Isaac, J. H., and B. W. Holloway. 1968. Control of pyrimidine biosynthesis in Pseudomonas aeruginosa. J. Bacteriol. 96:1732-1741. Labrum, E. L., and M. E. Bunting. 1953. Spontaneous and induced color variation of the HY strain of Serratia marcescens. J. Bacteriol. 65:394 404. Lowry, 0. IL, N. J. Rosebrough, A. L. Fan, and R. J. Randall. 1951. Protein measurements with the Folin phenol reagent. J. Biol. Chem. 193:265-275. Marmur, J., S. Falkow, and M. Mandel. 1963. New approaches to bacterial taxonomy. Annu. Rev. Microbiol. 17:329-372. Murphy, T. M., and S. E. Mills. 1969. Immunochemical and enzymatic comparisons ofthe tryptophan synthetase a subunits from five species ofEnterobacteriacae. J. Bacteriol. 97:1310-1320. Neumann, J., and M. E. Jones. 1964. End product inhibition of aspartate transcarbamylase in various species. Arch. Biochem. Biophys. 104:438-447. O'Donovan, G. A., H. Holoubek, and J. C. Gerhart. 1972. Regulatory properties of intergenic hybrids of aspartate transcarbamylase. Nature (London) New Biol. 238:264-266. O'Donovan, G. A., and J. Neuhard. 1970. Pyrimidine metabolism in microorganisms. Bacteriol. Rev. 34:278-343. Rocha, V., I. P. Crawford, and S. E. Mills. 1972. Comparative immunological and enzymatic study of the tryptophan synthetase A subunit in the Enterobacteriaceae. J. Bacteriol. 111:163-168. Wedler, F. C., and F. J. Gasser. 1974. Modes ofmodifier actions in E. coli aspartate transcarbamylase. Arch. Biochem. Biophys. 163:69-78.
JOURNAL OF BACTERIOLOGY, Dec. 1976, p. 766-775 Copyright © 1976 American Society for Microbiology
Unique Aspects of the Regulation of the Aspartate Transcarbamylase of Serratia marcescens JAMES R. WILD,' WILLIAM L. BELSER, AND GERARD A. O'DONOVAN* Department of Biology, University of California, Riverside, California 92502, and Department of Biochemistry and Biophysics, Texas A&M University, College Station, Texas 77843* Received for publication 7 July 1976
Aspartate transcarbamylase (ATCase; EC 2.1.3.2) from Serratia marcescens HY has been purified 134-fold. Its properties are unique. Unlike the ATCase from Escherichia coli and Salmonella typhimurium, the S. marcescens HY enzyme activity is not feedback inhibited by any purine or pyrimidine nucleotide effectors; instead, the enzyme is activated by both cytidine 5'-triphosphate and adenosine 5'-triphosphate. Like the ATCase from E. coli and S. typhimurium, adenosine 5'-triphosphate alters the [S],.5 of the enzyme and, in contrast, cytidine 5'-triphosphate does not alter the [S],.5 but, instead, alters the Vma.. As has been shown for both E. coli and S. typhimurium, effector sensitivity may be selectively dissociated from catalytic activity by treatment with heat, parachloromercuribenzoate, or neohydrin. The dissociated enzyme possesses threefold higher specific activity than the native enzyme. The sedimentation coefficient of the native enzyme is approximately 11.4S, whereas the dissociated enzyme has a value of 6.0S. Whereas it has been possible to reconstitute the E. coli and the S. marcescens ATCase enzymes from their own homologous subunits, it has not been possible to make hybrid enzymes of catalytic and regulatory heterologous subunits from each other. It was not possible to detect repression of ATCase formation after growth of prototrophic strains of S. marcescens HY supplemented with 200 ,tg of uracil per ml, but eightfold derepression was observed after uracil withdrawal in pyrimidine auxotrophs. Aspartate transcarbamylase (ATCase; EC study. Bethell and Jones have classified such 2.1.3.2) is the first enzyme unique to the path- ATCases as "B-type" enzymes (2); these allosway of pyrimidine biosynthesis and therefore terically regulated ATCases are subject to seoccupies an important position in the regula- lective dissociation of effector response and tion of pyrimidine nucleotide formation. Al- characterize all genera of the Enterobacteriathough ATCase is common to all microorga- ceae (2, 25, 27). Serratia marcescens belongs to the Enteronisms possessing a pathway for de novo pyrimidine biosynthesis, the allosteric control of the bacteriaceae (7), but differs from other memenzyme is effected by many nucleotides; the bers of the family in several respects. S. marnegative effectors that have been identified for cescens and Enterobacter (Aerobacter) aerodifferent systems include uridine 5'-triphos- genes represent an extreme in guanine + cytophate (UTP), cytidine 5'-triphosphate (CTP), sine content (58%) among the enterobacteria uridine 5'-monophosphate (UMP), and cytidine (23). The majority of the nucleotide sequences 5'-monophosphate (CMP) (2, 9, 14, 25, 27). AT- of strains of S. marcescens are not related to Case from Escherichia coli has been studied other members of the family by any of the extensively. It contains two nonidentical pro- criteria used by Brenner et al. (4), and thus tein subunits: one, the catalytic subunit, pro- Serratia appears to be the most divergent gevides the active sites of the enzyme, whereas nus of the family. These differences have been the other, the regulatory subunit, provides the supported by the determination of immunologibinding sites for nucleotide effectors (6, 12). cal indexes of dissimilarity of the tryptophan Table 1 summarizes some characteristics of the synthetase a subunit (24) and ,32 subunits (28) ATCase from E. coli that are pertinent to this in comparative studies. The extensive studies in the tryptophan pathway have been recently I Present address: Genetics Section, Texas A&M Uni- reviewed by Crawford (10). In this article we versity, College Station, TX 77843. describe the ATCase from S. marcescens HY 766
VOL. 128, 1976
S. MARCESCENS ATCase
767
TABLE 1. Properties of native ATCase and its catalytic and regulatory subunits from E. colia Property
(2C3) (3R2)
Native enzyme
Mercurials
M o Mercaptoethanol
2C3
Catalytic
Subunit structure ......... ...... (2C3) (3R2) 2(C3) Molecular weight ................ 310,000 2 (100,000) Sedimenation coefficient ......... 11.7S 5.8S Molecular weight of each polypeptide chain 34,000 Shape of saturation curve ........ Sigmoidal Hyperbolic Catalytic activity (binding of substrates) ....................... + + Binding of effectors ........ ...... + a Determined according to O'Donovan and Neuhard, with modifications (27).
and examine some of its unique regulatory properties. MATERIALS AND METHODS
+
3R2
Regulatory
3(R2) 3 (34,000) 2.8S
17,000
+
termination described by Gerhart and Pardee (14). Standard assay mixtures contained: 5 to 10 mM aspartate, pH 7.0; 0.4 mM carbamyl phosphate, pH 7.0; 40 mM potassium phosphate, pH 7.0; and 25 to Bacterial strains. S. marcescens HY wild type 50 ,ul of extract. The volume was adjusted to 0.50 ml and pyrimidine-requiring auxotrophs derived from with distilled water. A final nucleotide concentrathis strain were used in these studies: HYpyrF3 was tion of 2 mM was utilized for all effector studies confirmed by enzyme assay and shown to lack oroti- unless otherwise specified. The reaction mix was dine 5'-phosphate decarboxylase activity (EC incubated at 30°C for 15 min at an extract concentra4.1.1.23); HY pyrEl lacks orotate phosphoribosyl tion such that the final absorbance developed by the transferase (EC 2.4.2.10). colorimetric assay at 560 nm was 0.400 to 0.800. The Growth conditions. The minimal medium used amount of carbamyl aspartate produced was deterwas Serratia minimal medium (SM) defined by La- mined three times during the 15-min reaction to brum and Bunting (21) and contained, per liter of ensure linearity. distilled water: K2HPO4, 8.0 g; ammonium citrate Enzyme hybridization. ATCase was partially dibasic, 5.0 g; MgSO4- 7H2O, 0.5 g; FeCl3, 0.02 g. The purified from a pyrimidine auxotrophic strain wild-type strain HY was grown overnight in 1-liter (pyrF3) of S. marcescens HY by the method of quantities of SM at 30°C. The cells were harvested Gerhart and Holoubek (13), with one modification: by centrifugation during the exponential phase of in the heat step we used a temperature of 58°C ingrowth (optical density at 660 nm = 0.400). The wild stead of 72°C for E. coli. An overall purification of type was used to prepare enzymes for effector stud- 134-fold was achieved, thus giving a protein prepaies. ration homogeneous by gel electrophoresis. Bacterial auxotrophs were grown in SM + uracil ATCase of E. coli K-12 was prepared from a leaky (25 ug/ml). Derepression was achieved by allowing pyrF strain by the method of Gerhart and Holoubek the cells to deplete the available uracil supply at an without modification (13). In both cases the native aggregates were dissooptical density at 660 nm of about 0.250 and continuing starvation for 90 to 120 min in the absence of ciated by p-chloromercuribenzoate (p-CMB) or neouracil. Pyrimidine auxotrophs of S. marcescens re- hydrin treatment (26; M. W. Kirschner, Ph.D. thequired uniquely high levels (50 ,ug/ml) of uracil for sis, University of California, Berkeley, 1971), and maximal yield. the subunits were separated by zone centrifugation Preparation of cell extracts. Cell extracts were on a 5 to 25% sucrose gradient and diethylaminoprepared by sonication of washed bacterial suspen- ethyl-cellulose anion-exchange chromatography, as sions as described by Hutchinson and Belser (19). previously described (16). All extracts were passed over Sephadex G-25 or diChemicals. The chemicals used in these studies alyzed overnight against a 1,000-fold excess of 50 were commercially obtained. 2-Mercaptoethanol, mM potassium phosphate buffer, pH 7.5, containing ethylenediaminetetraacetic acid, dithiothreitol, L2 ,uM ZnSO4. During sonication the extracts were aspartate, and carbamyl phosphate (dilithium salt) maintained below 8°C. The extract was clarified by were purchased from Sigma. p-CMB (sodium salt) centrifugation at 39,000 x g for 1 h. The enzyme and the following nucleotides and nucleosides were preparation was stable for several weeks when obtained from Calbiochem: CTP, cytidine 5'-diphosphate, cytidine 5'-monophosphate (CMP), cytidine, stored at 40C. Analytical methods. Protein was estimated by UTP, uridine 5'-diphosphate, UMP, uridine, guanothe method of Lowry et al. (22), with crystalline sine 5'-triphosphate, guanosine 5'-monophosphate, bovine serum albumin as the standard. The enzy- adenosine 5'-triphosphate (ATP), and adenosine 5'matic activity of ATCase was measured by the de- monophosphate. Catalase was obtained from Wortermination of carbamyl aspartate formed. Carba- thington Biochemicals. Neohydrin was a gift from J. myl aspartate was assayed by the colorimetric de- C. Gerhart.
768
J. BACTERIOL.
WILD, BELSER, AND O'DONOVAN
RESULTS and reassociation. dissociation Enzyme The reconstitution of ATCase from catalytic and regulatory subunits prepared by dissociation withp-CMB is summarized in Table 2. The data are presented as relative enzyme activity (1 unit = 1.0 ,umol of carbamyl aspartic acid produced/min per 25 ,ul of initial sample). Thus, the same concentration of ATCase was assayed for each purified extract before and after dissociation. Separation of the subunits produced a threefold increase in catalytic activity for E. coli (0.220 to 0.661 units) and for S. marcescens (0.440 to 1.240 units). After dissociation from the 3.OS regulatory subunit, the 6.0S catalytic subunit did not possess an effector response to 2 mM CTP for E. coli (0.661 versus 0.665 units) and S. marcescens (1.240 versus 1.257 units). It was possible to reconstitute both the E. coli and S. marcescens native enzymes from their respective homologous subunits and regain the original effector response to CTP, accompanied by a decrease in relative enzyme activity in the absence of effector. The ratio of the percentage of activity in the presence and absence of CTP provides a valuable index for evaluating the reconstitution. The inhibition of ATCase from E. coli and the activation of the ATCase from S. marcescens were virtually identical before dissociation and after reconstitution. We have been unable to reconstitute the heterologous (hybrid) subunits and form the hybrid enzyme that possessed either positive or negative effector response; i.e., the catalytic subunit from S. marcescens mixed with the regulatory subunit from E. coli did not possess any effector response to CTP.
Physical characterization of ATCase. (i) Estimation of sedimentation coefficients. Sedimentation coefficients were determined for the ATCase of S. marcescens HY by sucrose gradient sedimentation. A sample of 0.2 ml of fresh enzyme extract (20 mg/ml) was layered onto a 5-ml, linear 5 to 25% sucrose gradient. The native ATCase was centrifuged in a Beckman Spinco L-2 ultracentrifuge for 13 h at 39,000 rpm at 2°C in an SW39 rotor. The desensitized enzyme (treated with 1 mMp-CMB) was centrifuged for 17 h under the same conditions. Three-drop samples were collected from the bottom of the gradients and diluted with 0.25 ml of 50 mM potassium phosphate buffer, pH 7.5. Each sample was assayed for ATCase activity, and sedimentation profiles are shown in Fig. 1 and 2. Using a value of 11.4S for the catalase marker, sedimentation coefficients were estimated for the enzymes under both conditions (19). The native enzyme has a value of 11.4S and the dissociated enzyme has a value of 6.OS. (ii) Response to nucleotide effectors. The effects of various nucleotides, nucleosides, and
I-
E
5; I-
0.10
TABLE 2. Reconstitution ofATCase from E. coli and S. marcescens HY Relative activity % ActivType of en(+tPI Source - CTP + CTP (+CTP/ zyme 6.0a
E. coli E. coli
0.220 0.661
0.106 0.665
48.1 100.6
3.0b
E. coli
0.233 0.440 1.240
0.096 0.970 1.257
41.1 220 107
0.176
0.334
190
0.350
0.360
102.8
11.7
11.4 6.0a 6.0b
3.0b 6.0b
3.0b
S. marcescens S. marcescens S. marcescens S. marcescens S. marcescens E. coli I
Effector response selectively dissociated by treatment with p-CMB. b Reconstitution of subunits separated by treatment with p-CMB. a
N
I 0
z
z
LUJ
co
005 m
0
5
10 15 20 FRACTION NUMBER
25
FIG. 1. Sucrose gradient sedimentation of native ATCase from S. marcescens HY. A sample of0.4 mg of crude extract was layered onto a 5-ml, linear 5 to 25% sucrose gradient and centrifuged at 39,000 rpm for 13 h in an SW39 rotor at 2°C. Symbols: ATCase (O); catalase (0), used as a marker. The sloping line represents the relative sedimentation values developed by the gradient.
S. MARCESCENS ATCase
VOL. 128, 1976 2.0
E
0.10
§
1.0 w
~~~~~~~z
z
~ ~ ~
0
5
~
10
~
~
~
15
~
20
-0.05
25
FRACTION NUMBER
FIG. 2. Sucrose gradient sedimentation ofp-CMB (2 mM)-treated ATCase from S. marcescens HY. A sample of 0.4 mg oftreated extract was layered onto a 5-ml, linear 5 to 25% sucrose gradient and centrifuged at 39,000 rpm forl3 h in an SW39 rotor at2°C. Symbols: ATCase (a); catalase (0) was used as a marker, measured by absorbance at 405 nm.
free bases on the activity of ATCase in S. marHY wild type were studied. Stimulation by various effectors was determined under standard assay conditions, using an aspartate concentration of 5 mM. No inhibition was observed with any of the effectors (Table 3). Cytosine compounds stimulated enzymatic activity in the order: CTP > cytidine 5'-diphosphate > CMP > cytidine. Deoxycytidine compounds were equally effective as activators as the ribonucleotides. Under similar conditions, ATP was the most effective activator and guanosine 5'-triphosphate showed neither inhibition nor activation. The pH of the reaction mix was then varied in an attempt to identify negative effectors; none were discovered. Various concentrations of aspartate and carbamyl phosphate were included in the assay mix with different potential inhibitors, but still no inhibition was found. (iii) Combinations of effectors. Table 4 presents the effect of various combinations of nucleotides upon the activity of ATCase. The reaction conditions were identical to those used for Table 1 studies, except 2 mM concentrations of each effector were used. The total effector con-
cescens
769
centration was 4 mM. Guanosine 5'-triphosphate, UTP, and UMP had no effect on the activation by CTP. The nucleoside monophosphates in different combinations had no effect on enzymatic activity. The combination of ATP and CTP showed an intermediate response when compared with either effector alone. (iv) Saturation of effectors. The saturation of the nucleotide effectors ATP and CTP is summarized in Fig. 3. At concentrations up to 5 mM, UMP or CMP shows no effect on the activity of ATCase. Both CTP and ATP show saturation of effector activation at 2.5 mM. (v) Kinetics of substrate dependence. Figures 4 and 5 summarize the dependence of ATCase activity of S. marcescens HY on the concentrations of carbamyl phosphate and aspartate. In the absence of effectors, the native enzyme shows a sigmoidal response when the aspartate concentration is varied (Fig. 4) and TABLE 3. Effect of various nucleotide and nucleoside effectors on the activity of S. marcescens HY ATCasea Effector
Enzyme activity (carbamyl as partate produced)"
% Activityc
100 25.2 None 189 47.7 CTP 142 34.8 CDP 108 27.2 CMP 105 26.6 Cytidine 100 25.3 UTP 98 24.5 UDP 97 24.4 UMP 102 25.8 Uridine 101 25.6 dTTP 108 27.2 dTMP 101 25.6 Thymidine 187 47.2 dCTP 110 27.9 dCMP 104 26.2 dUTP 97 24.4 dUMP 104 26.3 GTP 101 25.5 GMP 249 62.9 ATP 96 24.2 AMP a ATCase activity was determined in the presence of 2 mM effector, 5 mM i-aspartate, 0.4 mM carbamyl phosphate, at pH 7.0. Abbreviations: CDP, Cytidine 5'-diphosphate; UDP, uridine 5'-diphosphate; dTTP, deoxythymidine 5'-triphosphate; dTMP,
deoxythymidine 5'-monophosphate; GTP, guanosine 5'-triphosphate; GMP, guanosine 5'-monophosphate; AMP, adenosine 5'-monophosphate. b Expressed as nanomoles of carbamyl aspartate produced in 15 min at 300C per milligram of protein. c
The activity of ATCase without effectors is set
at 100%.
WILD, BELSER, AND O'DONOVAN
J. BACTZRIOL.
TABLz 4. Effect of combinations of nucleotide effectors on the activity of ATCase from S. marcescens Hya
ment, activity was not lost, but actually increased 2.7-fold (Table 5). A high ionic strength 1.0 M (NH4)2S04 and high pH (pH 9.2) provide heat stability to effector response ofthe enzyme and were utilized during partial purification of the enzyme. Enzymatic activity and effector response were determined as described for Table 3. Another method utilized to separate effector response from catalytic activity was treatment with the mercurialp-CMB at 2 mM. Enzymatic activity was greatly reduced (90%) by the initial treatment with this concentration of mercurial. Specific recovery of the enzymatic activity, without effector response, was observed after 3 h of dialysis against 100 volumes of potassium phosphate buffer, pH 7.5, containing 50 mM dithiothreitol and 0.1 mM ethylenediaminetetraacetic acid. Consistent with the report of Gerhart and Pardee for E. coli ATCase (14), the loss of effector response by treatment with p-CMB resulted in a threefold increase in the specific activity of ATCase over the untreated enzyme. Nucleotide activation was also separated from catalytic activity by treatment with neohydrin at 3 mM. (vii) pH studies. Effector responses showed
770
Effector
Carbamyl aspartate produced"
% Activity,
100 25.2 189 47.7 249 62.9 100 25.3 104 26.3 97 24.4 108 27.2 101 25.5 221 55.9 190 48.0 GTP+CTP 178 44.9 UTP+CTP 187 47.2 UMP+CTP 98 24.5 UMP+CMP 105 26.6 GMP+CMP a ATCase activity was determined, as in Table 3, in the presence of 2 mM nucleotide. When the effect of two nucleotides present simultaneously was determined, 2 mM concentrations of each potential effector were present. See Table 3 for abbreviations. b Expressed as nanomoles produced in 15 min at 30°C per milligram of protein. c The activity of ATCase without effectors is set at 100%.
None CTP ATP UTP GTP UMP CMP GMP ATP+CTP
an apparently hyperbolic response to variations in carbamyl phosphate concentration (Fig. 5). The sigmoidal response to aspartate is maintained in the presence of effectors. However, CTP elicits an alteration in V., whereas ATP causes a change in the IS .5. The estimated Hill coefficient (11) in the presence of CTP is similar to that in the absence of effector (napp = 1.8 to 1.9). The value is somewhat lower in the presence of ATP (napp = 1.5 to 1.6). Substrate activation by aspartate is observed in the presence and absence of effectors. The [SL.5 of the native enzyme for aspartate is approximately 65 mM. The [SL.5 for aspartate in the presence of ATP is approximately 20 mM. The hyperbolic response of carbamyl phosphate concentration and velocity is maintained in the presence of nucleotide effectors, and the activation appears to be uncompetitive (see Fig. 5, inset). (vi) Selective destruction of effector response. Selective destruction of activation by both CTP and ATP was obtained by three independent methods. When the cell extract (3 ml) was heated at 60°C for 3 min in 50 mM potassium phosphate buffer, pH 7.5, ATCase could not be activated by 5 mM CTP or ATP, nor could it be inhibited by UMP or CMP. 2-Mercaptoethanol, dithiothreitol, or ethylenediaminetetraacetic acid (1 mM) did not provide any protection to the enzyme. After heat treat-
k-C-) 4
5 4 3 EFFECTORS x 10-3 FIG. 3. Saturation of effectors of S. marcescens HY ATCase. Enzyme activity (micromoles of carbamyl aspartate x 10115 min per milligram of protein at30°C) was determined in the presence of increasing concentrations of nuclotides. Symbols: CTP (0); ATP (-); and UMP or CMP (U) in the standard reaction mix containing 5 mM aspartate, 0.4 mM carbamyl phosphate, and 40 mM phosphate buffer, pH 7.0. 2 1 MOLARITY OF
VOL. 128, 1976
S. MARCESCENS ATCase
P 2.0 w
>.
1.5
w
0
20 40 60 80 MOLARITY OF L-ASPARTATE x10-3
100
FIG. 4. Velocity-substratesaturationcurvesinthe of several nucleotides (at 2 mM concentration) at pH 7.0. Enzyme activities are expressed as micromoles of carbamyl aspartate x 100/15 min per milligram of protein at 30°C. The reaction mixes contained 0.5 mg of protein with 0.4 mM carbamyl phosphate and 40 mM, potassium phosphate, and varying concentrations of aspartate were used to initiate the reaction. Symbols: ATP (a); CTP (0); CMP or UMP (U); no effector (O). presence
771
(viii) Effect of 2-thiouracil. 2-Thiouracil has been shown to be a growth inhibitor when added to exponentially growing cultures of E. coli (5). It has been proposed that 2-thiouracil is converted to 2-thiouridine-5'-monophosphate, which functions as a CTP analogue in inhibiting the catalytic activity of ATCase (17). 2Thiouracil (50 ,ug/ml, final concentration) was added to an exponentially growing culture of wild-type S. marcescens HY. No alteration of growth rate was noticed in liquid cultures or on minimal agar plates to which the analogue was added. Such a concentration of 2-thiouracil inhibited the growth of E. coli in both cases. Derepression of ATCase. Table 6 is a summary of a series of studies that demonstrate that the formation of ATCase may be derepressed in auxotrophs of S. marcescens HY. Cultures in mid-log growth in SM + uracil (50 jig/ml) were centrifuged at 10,000 x g, and the bacterial cells were resuspended in SM without uracil. The specific activity of ATCase in pyrimidine auxotrophs increased ninefold following incubation for 2 h after removal of uracil. The parental strain produced similar low levels of ATCase in response to various concentrations of uracil or arginine in the growth medium.
DISCUSSION ATCase catalyzes the first biosynthetic reaction unique to the formation of pyrimidine nucleotides and involves the carbamylation of aspartate by carbamyl phosphate. This enzyme is found in procaryotic and eucaryotic organisms, but its structural and regulatory properties extreme variation in relation to pH. Figure 6 show wide diversity (1, 2, 26, 27). The ATCases shows the pH profile for ATCase from S. mar- from different organisms are positively and cescens and the effector responses for ATP, negatively regulated by a variety of nucleotide CTP, and UMP. Tris(hydroxymethyl)amino- effectors. Most of the enteric bacteria are allomethane buffer was used in these studies for sterically inhibited by CTP or UTP. The enpH 10.0 to 7.0. Phosphate buffer was used for zymes from E. coli and S. typhimurium have pH 7.5 to 6.0. In the absence of effectors, the been extensively studied and provide a basis for S. marcescens HY enzyme shows an extremely allosteric effector regulation, regulatory and broad spectrum of activity. This activity is catalytic protomer interactions, and potential greatly enhanced at lower pH upon the addition taxonomic and evolutionary comparisons (12, of either CTP or ATP. It has been reported 15, 27). that the ATCase from a different strain of S. marcescens is inhibited by 0.013 mM UMP or CMP at pH 8.5 (25). We were unable to detect inhibition (greater than 10%) at pH 7.0, 8.5, or 10.0 under similar conditions or with 20-fold higher mononucleotide concentrations in S. marcescens HY. Even at extremely low substrate concentrations no inhibition was observed. This was true for crude extracts and for the partially purified enzyme.
S. marcescens HY possesses a unique ATCase that is not feedback inhibited by pyrimidine or purine nucleotides, nucleosides, or free bases; instead it is activated by a pyrimidine nucleotide, CTP, and by a purine nucleotide, ATP. Neumann and Jones (26), using a different strain of S. marcescens, demonstrated that ATCase was inhibited by UMP and CMP. Extensive attempts have been made to verify this result in strain HY, without success. Preincu-
772
J. BACTERIOL.
WILD, BELSER, AND O'DONOVAN
2.5
2.0 I-
1.5 4
w N 1.0 z w
0.5X e6
1/
r*
0
5
32
MOLANTY OF CARBAMYL PHOSPHATE
48 X 10 4
64
25 20 15 MOLARITY OF CARBAMYL PHOSPHATE x 104 10
FIG. 5. Velocity-substrate saturation curves in the presence of several nucleotides each at 2 mM concentration, pH 7.0. Enzyme activities are expressed as micromoles of carbamyl aspartate producedll5 min per milligram of protein at 30°C. The reaction mixes contained 0.5 mg of protein with 10 mM aspartate and 40 mM potassium phosphate, and varying concentrations of carbamyl phosphate were used to initiate the reaction. Symbols: ATP (a); CTP (0); CMP (*); no effector (E); UMP (-). (Inset) Double-reciprocal plot of carbamyl phosphate saturation for ATCase in the presence of ATP (-), CTP (0), and without effector (O).
bation of the enzyme with effectors for up to 20 min, variation of pH, increased effector concentration to 5 mM, and variation of substrate concentration have failed to establish any conditions in which inhibition could be demonstrated. Neumann and Jones did not report any effect of CTP on Serratia ATCase, but activation of ATCase by CTP was reported in Proteus vulgaris by Martha Bethell (Ph.D. thesis, Brandeis University, Waltham, Mass., 1967). The Proteus extract, however, contained two molecular forms of ATCase, an activatable form with a molecular weight of 300,000 and a "nonresponsive" form of molecular weight 100,000. The native ATCase of S. marcescens HY found in our study has a molecular weight of approximately 300,000.
Although ATP and CTP both function as activators, their mode of activation is different. ATP functions as an activator by altering the [S10.S5 for aspartate at saturating concentrations of carbamyl phosphate. ATP activates the ATCase from E. coli and S. typhimurium in the same manner and competes with CTP for the same binding sites (6). CTP is a competitive inhibitor with aspartate in these systems, but in S. marcescens HY CTP is an uncompetitive activator. Indeed, CTP alters the Vmax of ATCase when aspartate concentration is varied but the same [Sk.5 is maintained. This hitherto unreported observation suggests that there may be a fundamental difference in the mechanism of activation of these two effectors. This observation is in accord with the proposal of
VOL. 128, 1976
S. MARCESCENS ATCase
TABLE 5. Selective destruction of nucleotide activation of ATCasea Treatment and effector
Carbamyl as-
producedb
partate
%
Activityc
Untreated None CTP ATP
27.0 49.0 63.0
100 181 234
Heatd None +CTP +ATP
74.0 73.0 75.0
271 270 273
773
which is considerably higher than the [S6.5 of the E. coli or S. typhimurium enzyme, 2 mM, at approximately the same carbamyl phosphate concentration (26). Thus, a great increase in aspartate level is required for maximal enzymatic activity. Furthermore, this high concentration of aspartate is accompanied by substrate activation of the enzyme. It appears that
2.5
p-CMBe None +CTP +ATP
3.0 2.5
11 9
p-CMBf None 90.0 333 +CTP 328 88.5 +ATP 93.5 346 a ATCase activity is determined in the presence of 2 mM effector, 5 mM L-aspartate, 0.04 mM carbamyl phosphate. b Expressed as nanomoles produced in 15 min at 30°C per milligram of protein. c The activity of ATCase without effectors is set at 100%. d After heat treatment (60°C for 15 min). e After treatment with 2 mM p-CMB. f After treatment with 20 IAM p-CMB.
2.0 C-
1.5 N
z w
1.0
0.5
pH
Wedler and Gasser (29), which suggests that CTP and ATP act by separate mechanisms in affecting aspartate binding inE. coli. ATP may directly affect the rate of aspartate association/ dissociation, whereas CTP induces an intramolecular alteration in the R-C domain of binding (for review, see reference 12). In other closely examined systems, it has been demonstrated that both positive and negative effectors alter the [Sk.5 of the respective ATCases and their effect is overcome by high substrate concentration. Activation studies with CTP and ATP simultaneously present during assay of the S. marcescens ATCase suggest that CTP and ATP compete with one another for activation and act independently of one another. Saturating concentrations of CTP and ATP result in compensatory activation with an intermediate effector response (Table 5). High substrate levels minimize activation by ATP, but activation by CTP is maintained. The kinetics of ATCase activity-substrate plots reveal additional unique characteristics of this enzyme. The concentration of aspartate required for half-maximal saturation is 65 mM,
FIG. 6. Effect of pH on the activation of ATCase by nucleotides. Enzyme activity was determined in the presence of 5 mM aspartate, 0.4 mM carbamyl phosphate, 2 mM effectors. Symbols: CTP (0); ATP (0); UMP or CMP (U); and without effector present (O). Enzyme' activity is expressed as micromoles of carbamyl aspartatel15 min per milligram of protein at 30°C.
TABLE
6. Derepression of ATCasea in wild-type strain HY and mutant pyrEl Strain Growth medium Sp actb SM HY 74.0 HY SM + uracil 63.0 HY SM + arginine 70.0 pyrEl SM + uracil 58.0 pyrEl SM - uracilc 464.0 a Strains were grown in SM supplemented with uracil or arginine (50 ,ug/ml) and harvested in midlog phase (optical density at 660 nm = 0.400 to 0.500). b Expressed as nanomoles of carbamyl aspartate produced per minute per milligram of protein. c Mutant strain pyrEl was grown in the presence of uracil (50 ug/ml), suspended in the absence of uracil, and allowed to incubate for 2 h.
774
WILD, BELSER, AND O'DONOVAN
the saturation curve for carbamyl phosphate is hyperbolic (Fig. 5), but as pointed out by Bethell and co-workers, a more sensitive assay might reveal some cooperative interaction (3). The effects of CTP and ATP are uncompetitive with respect to carbamyl phosphate concentration. The [S]o.5 for carbamyl phosphate, 0.15 mM, is comparable to that ofthe E. coli enzyme (3). ATCase is composed of two types of protein subunits (Table 1), and the purified native enzyme from E. coli has been separated into regulatory and catalytic subunits by p-CMB (15). Similarly, we have selectively dissociated regulatory activity from catalytic activity in S. marcescens HY by heat treatment or by p-CMB. Once dissociated, the catalytic activity was not affected by either ATP or CTP. The desensitized enzyme is shown to be a smaller form, approximately 100,000 daltons, with an S value of 6.0. Thus, the size of the S. marcescens HY ATCase is similar to other type B ATCases before and after dissociation (2). In E. coli and S. typhimurium the subunit dissociation by p-CMB is readily reversible by incubation in 2 mM mercaptoethanol in high ionic strength, 1.0 M (NH4)2SO4, and at pH 9.2 tris(hydroxymethyl)aminomethane buffer. Reconstitution of the S. marcescens HY native enzyme has been achieved from its regulatory and catalytic subunits, and activation by CTP and ATP was restored. Further work on the purification of ATCases from different genera of bacteria is in progress in our laboratory (J. R. Wild, K. F. Foltermann, and G. A. O'Donovan, manuscript in preparation). It is also interesting to speculate on the overall mode of regulation of enzyme activity of pyrimidine biosynthesis in Serratia. The first consideration is that CTP activation of ATCase might be important to increase the flow of pyrimidine intermediates through the pathway. This flow increase might be necessary due to a more sensitive effector response at some other point in the pathway. Carbamyl phosphate synthetase is the most likely target to be considered for such a stringent regulatory role. Aspartate has an extremely high [S]05 for ATCase in S. marcescens (65 mM), and the relative effect of CTP and ATP at this level of aspartate is quite different. At physiological levels of aspartate (5 mM), ATCase is relatively unresponsive to changes in CTP levels, whereas the enzyme is much more responsive to changes in the levels of aspartate and ATP. High endogenous pool levels of CTP may compete with ATP for activation and depress the extent of activation. Therefore, CTP inhibits the activation by
J. BACTERIOL.
ATP and reduces the production of carbamyl aspartate compared to the ATP-activated enzyme without CTP. One further point seems pertinent, namely, that extremely high concentrations of uracil are required to provide maximal growth of pyrimidine auxotrophs of S. marcescens HY in liquid culture (Wild, unpublished data). All of the enzymes in the pathway may be partially derepressed, even in wild-type strains, in order to provide for the unusually high uracil requirement. Thus, the situation may parallel the system for Pseudomonas aeruginosa PAO (20) and P. putida (9), in which very little inhibition was found for ATCase and the entire pathway beyond ATCase seemed to be constitutive. It is possible to elevate levels of ATCase activity in Pseudomonas and Serratia by withdrawal of uracil in pyrimidine auxotrophs, but levels of enzyme cannot be repressed by growth on uracil in prototrophs. The activities of the subsequent enzymes of the de novo pathway in S. marcescens HY vary in starved auxotrophs, but the levels vary only two- to threefold (18). Even so, this variation may be the result of polypeptide interactions in a multifunctional complex rather than "derepression" per se (J. R. Wild and W. L. Belser, Biochem. Genet., in press). These observations suggest that S. marcescens HY might be producing the enzymes involved in pyrimidine biosynthesis at very high levels and might not be capable of conventional "repression" and "derepression." ACKNOWLEDGMENTS This work was supported by research grants from the Office of Naval Research, the Robert A. Welch Foundation, Houston, Tex., and the Texas Agricultural Experiment Station. One of us (J.W.) was a National Science Foundation Predoctoral Fellow. LITERATURE CITED 1. Adair, L. B., and M. E. Jones. 1972. Purification and characteristics of aspartate transcarbamylase from Pseudomonas fluorescens. J. Biol. Chem. 247:23082315. 2. Bethell, M. R., and M. E. Jones. 1969. Molecular size and feedback regulation characteristics of bacterial aspartate transcarbamylases. Arch. Biochem. Biophys. 134:352-365. 3. Bethell, M. R., K. E. Smith, J. S. White, and M. E. Jones. 1968. Carbamyl phosphate: an allosteric substrate for aspartate transcarbamylase of Escherichia coli. Proc. Natl. Acad. Sci. U.S.A. 60:1442-1449. 4. Brenner, D. J., G. R. Fanning, K. E. Johnson, R. V. Citarella, and S. Falkow. 1969. Polynucleotide sequence relationships among members of Enterobacteriaceae. J. Bacteriol. 98:637-650. 5. Cardeilhac, P. T. 1967. A toxic effect of 2-thiouracil on pyrimidine metabolism. Proc. Soc. Exp. Biol. Med. 125:692-696. 6. Changeux, J.-P., J. C. Gerhart, and H. K. Schachman. 1968. Allosteric interactions in aspartate transcarbamylase. I. Binding of specific ligands to the native
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7.
8. 9.
10. 11.
12. 13.
14.
15.
16.
17.
enzyme and its isolated subunits. Biochemistry 7:531-537. 18. Cocks, G. T., and A. C. Wilson. 1972. Enzyme evolution in the Enterobacteriaceae. J. Bacteriol. 110:793-02. Coleman, M. S., and M. E. Jones. 1971. Aspartate 19. transcarbamylases of Citrobacter freundii. Biochemistry 10:3390-3396. Condon, S., J. K. Collins, and G. A. O'Donovan. 1976. 20. Regulation of arginine and pyrimidine biosynthesis in Pseudomonas putida. J. Gen. Microbiol. 92:375383. 21. Crawford, I. P. 1975. Gene rearrangements in the evolution of the tryptophan synthetic pathway. Bacteriol. Rev. 39:87-120. 22. Endrenzi, L., C. Fazi, and F. H. F. Kwong. 1975. Evaluation of Hill slopes and Hill coefficients when saturation binding or velocity is not known. Eur. J. 23. Biochem. 51:317-328. Gerhart, J. C. 1970. A discussion of the regulatory properties of aspartate transcarbamylase from Esche- 24. richia coli. Curr. Top. Cell. Regul. 2:275-325. Gerhart, J. C., and H. Holoubek. 1967. The purification of aspartate transcarbamylase ofEscherichia coli and separation of its protein subunits. J. Biol. Chem. 25. 242:2886-2892. Gerhart, J. C., and A. B. Pardee. 1962. The enzymology of control by feedback inhibition. J. Biol. Chem. 26. 237:891-896. Gerhart, J. C., and H. K. Schachman. 1965. Distinct subunits for the regulation and catalytic activity of aspartate transcarbamylase. Biochemistry 4:1054- 27. 1062. Gerhart, J. C., and H. K. Schachman. 1968. Allosteric interactions in aspartate transcarbamylase. II. Evi- 28. dence for different conformational states of the protein in the presence and absence of specific ligands. Biochemistry 7:538-552. Goodrich, M. E., and P. T. Cardeilhac. 1970. 2-Thiouri- 29. dine-5'-phosphate and its inhibition of aspartate tmnscarbamylase. Biochim. Biophys. Acta 222:621-
S. MARCESCENS ATCase
775
626.
Hayward, W. S., and W. L. Belser. 1965. Regulation of pyrimidine biosynthesis inSerratia marcescens. Proc. Natl. Acad. Sci. U.S.A. 53:1483-1489. Hutchinson, M. A., and W. L. Belser. 1969. Enzymes of tryptophan biosynthesis in Serratia marcescens. J. Bacteriol. 98:109-115. Isaac, J. H., and B. W. Holloway. 1968. Control of pyrimidine biosynthesis in Pseudomonas aeruginosa. J. Bacteriol. 96:1732-1741. Labrum, E. L., and M. E. Bunting. 1953. Spontaneous and induced color variation of the HY strain of Serratia marcescens. J. Bacteriol. 65:394 404. Lowry, 0. IL, N. J. Rosebrough, A. L. Fan, and R. J. Randall. 1951. Protein measurements with the Folin phenol reagent. J. Biol. Chem. 193:265-275. Marmur, J., S. Falkow, and M. Mandel. 1963. New approaches to bacterial taxonomy. Annu. Rev. Microbiol. 17:329-372. Murphy, T. M., and S. E. Mills. 1969. Immunochemical and enzymatic comparisons ofthe tryptophan synthetase a subunits from five species ofEnterobacteriacae. J. Bacteriol. 97:1310-1320. Neumann, J., and M. E. Jones. 1964. End product inhibition of aspartate transcarbamylase in various species. Arch. Biochem. Biophys. 104:438-447. O'Donovan, G. A., H. Holoubek, and J. C. Gerhart. 1972. Regulatory properties of intergenic hybrids of aspartate transcarbamylase. Nature (London) New Biol. 238:264-266. O'Donovan, G. A., and J. Neuhard. 1970. Pyrimidine metabolism in microorganisms. Bacteriol. Rev. 34:278-343. Rocha, V., I. P. Crawford, and S. E. Mills. 1972. Comparative immunological and enzymatic study of the tryptophan synthetase A subunit in the Enterobacteriaceae. J. Bacteriol. 111:163-168. Wedler, F. C., and F. J. Gasser. 1974. Modes ofmodifier actions in E. coli aspartate transcarbamylase. Arch. Biochem. Biophys. 163:69-78.
