Enzymatic Synthesis of Folylpolyglutamates · 2001-09-05 · Enzymatic Synthesis of...

Enzymatic Synthesis of Folylpolyglutamates CHARACTERIZATION OF THE REACTION AND ITS PRODUCTS*

(Received for publication. I k e m b e r 26. 1979)

John J. McGuire, Pearl Hsieh, James K. Coward,$ and Joseph R. Bertinog From the Department of Pharmacology, Yale UniL>ersity School of Medicine, Neu, Hatsen, Connwticut 06510

Rat liver folylpolyglutamate synthetase was partially purified and its reaction and products were characterized. The preparation contained no conjugase activity. Gel filtration analysis revealed a molecular weight of 69,000. The synthetase was optimally active at pH 8.4 (37"C), required mercaptoethanol and a monovalent cation, and was highly specific for L-glutamate. Only purine nucleoside triphosphates served as the energy source for the reaction; ATP and dATP gave the best activity. All naturally occurring folates (including 5- methyl-tetrahydrofolic acid) as well as a number of folate analogs (including methotrexate) served as substrates. The unnatural diastereoisomer of at least one folate, 5,6,7,8-tetrahydrofolic acid, was also a substrate. Modifications of the terminal, acceptor glutamate led to loss of substrate activity, as well as loss of binding. High pressure liquid chromatography analysis, conjugase digestion, double radiolabel studies, and amino acid analysis of acid-hydrolyzed product confirmed that folylpoly-y-glutamates were synthesized. High concentrations of (dl)-5,6,7,8-tetrahydrofolic acid favored accumulation of short chain (predominantly diglutamate) products while low concentrations favored accumulation of longer chains (predominantly tetraglutamate). This inverse relationship between concentration and chain length may be of regulatory significance. Synthesis of pentaglutamate forms, the predominant chain length of rat liver folates in uiuo, was detected at low (dl)-5,6,7,8-tetrahydrofolic acid, but hexaglutamate was not detected. Synthetic (1)-5,6,7,8- tetrahydropteroylpentaglutamate was a poor substrate for the synthetase but it inhibited formation of polyglutamates from monoglutamates. These observations indicate that the predominant chain length of folates in rat liver may be determined solely by the substrate specificity of the rat liver synthetase. Inhibition by the pentaglutamate derivative offers a means by which folylpolyglutamates could regulate their own synthesis.

Reduced folates' serve as carriers of I-carbon units and are * This investigation was supported by National Institutes of Health

Research Grants CA08010 and CA23209/28097 from the National Cancer Institute and by both National Institutes of Health Postdoc- toral Fellowship GM06321 and Traineeship 5-T32-CA09085 (to J. J. M.). A preliminary account of this work was presented at the Sixth International Symposium on the Chemistry and Biology of Pteridines, La Jolla, CA, September 1978. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "eduertisernent" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

$ Present address, Department of Chemistry, Rensselaer Polytech- nic Institute, Troy, NY 12181.

5 American Cancer Society Professor of Medicine and Pharmacol-

The term folate is used to refer to the general class of compounds OPT.

essential cofactors in the biosynthesis of purines, thymidylate, methionine, and glycine (1). Almost without exception, when- ever these folates are isolated from natural sources, they are found not as monoglutamate derivatives but as poly(?-glutamyl) conjugates containing two to eight glutamates (2). Within any particular source, a single length usually predominates but a distribution of lengths is found. For example, Esche- richia coli contains predominantly pentaglutamate folates but tri- and tetraglutamate derivatives also occur (3). Yeast, which was previously thought to contain only folylheptaglutamates (4), also contains smaller amounts of hexa- and octaglutamates (5). A variety of mammalian sources have been characterized and pentaglutamates predominate in nearly all, including rat liver (6-8).

Although their universal occurrence, even in species unable to biosynthesize folic acid, suggests their physiological importance, the function of these polyglutamates is unclear. Many studies have demonstrated that polyglutamates serve more efficiently than monoglutamates in folate-dependent reactions in vitro (3,9-11,34,52). Polyglutamyl folates may thus be the active cofactors in uiuo and monoglutamates, as suggested by Rabinowitz (121, are merely convenient in vitro analogs. Other evidence indicates that formation of polyglutamates is required for intracellular retention of folates. A mutant CHO cell line which cannot synthesize folylpolyglutamates has been isolated and found to contain very low levels of intracellular folates (13). Since general folate metabolism, including transport of folates, is normal in this line, the lack of folylpolyglutamates must be responsible for the low folate pools and hence polyglutamylation is necessary for retention of folates.

Despite the evidence indicating the importance of polyglutamates in folate metabolism, few reports on the enzyme(s) responsible for their synthesis, folylpolyglutamate synthetase, have appeared. Most studies have focused on the synthesis of polyglutamates in intact cells or whole organisms; interpre- tation of the results is complicated by the effects of transport and subsequent metabolism of the labeled folates used (5-8). Since the original description of a bacterial folylpolyglutamate synthetase by Griffin and Brown (14), only a few in vitro

derived from folic acid. The symbols ( I ) and ( d ) denote the natural and unnatural diastereoisomers of folates, respectively, which are the result of the asymmetric center introduced at C-6 during reduction, and do not indicate optical rotation. All folates contain L-glutamate unless otherwise indicated. The abbreviations used are: PteGlu, Pter- oylglutamic acid, folic acid; HzPteGlu, 7.8-dihydrofolic acid: H,PteGlu, 5,6,7,8-tetrahydrofolic acid; 5-HCO-, IO-HCO-, 5,10-CH~-, and 5-CH:3-H4PteGlu, 5-formyl-, 10-formyl-, 5,10-methylene-, and 5- methyl-tetrahydrofolic acid, respectively; MTX, methotrexate (2-4- diamino-10-methyl-folic acid); (dl)-5-CH.,-H4-homoPteGlu, (d0-5- methyl-tetrahydrohomofolic acid; poly(y-glutamy1) derivatives of all forms are indicated by a following subscript n, where n is the total number of glutamates, e.g. PteClul is PteClu-y-Gh HPLC, high pressure liquid chromatography; CHO, Chinese hamster ovary: d m - syl, 5-dimethylaminonaphthalene-1-sulfonyl.

". ~- " " " .

5776

Rat Liver Folylpolyglutamate Synthetase 5777

studies of this enzyme have appeared (15-17). Such in vitro studies are essential to understand how polyglutamates are synthesized, and what factors, if any, regulate their biosynthesis and determine the distribution found in vivo. In this article, we report the characterization of rat liver folylpolyglutamate synthetase and a detailed examination of the products of the reaction.

MATERIALS AND METHODS

Male Sprague-Dawley rats (200 to 250 g) were obtained from Charles River Breeding Laboratories and maintained on lab chow until sacrifice. Ammonium sulfate (ultrapure) was obtained from Schwarz/Mann. Whatman phosphocellulose (P-11) and DEAE-cellulose (DE52 and DE23) and Eastman DEAE-cellulose were precy- cled according to the manufacturers' instructions. ATP and ADP were from P-L Biochemicals. TPN, TPNH, DPNH, sodium pyruvate, glucose 6-phosphate, and folic acid were from Sigma. New England Nuclear was the source of ~-[B,B-'H]glutamic acid (24 Ci/mmol) and ["Clformate (55.9 mCi/mmol). ['4C]Formaldehyde (15.6 mCi/mmol) and YL-"C]folic acid were products of Amersham. Oxaloacetate was from BoehringH Mannheim and quaternary aminoethvl Sephadex (QAE-Sephadex) and Sephadex G-150 were from Pharmacia. All other chemicals were reagent grade.

Enzymes and Proteins-Clostridium acdi-urici formyltetrahv- drofolate synthetase, purified according to McGuire and Rabinowitz (18), was the generous gift of Dr. J. C. Kabinowitz. Lactic dehydrogenase (beef heart) and glucose-&phosphate dehydrogenase from Sigma, malate dehydrogenase (pig heart) from Boehringer Mann- heim, and ovalbumin from Worthington were used. Crystalline bovine serum albumin standard was from Armour Pharmaceuticals. Lacto- bacillus casei dihydrofolate reductase was from the New England Enzyme Center. Chicken pancreas conjugase was purified according to Leichter et al. (19).

Enzyme Assays-Folylpolyglutamate synthetase was assayed by a method, developed independently in this laboratory, which is similar to the method of Taylor and Hanna (17). The assay depends on the separation, on DEAE-cellulose columns, of [ 'Hlglutamate which is incorporated into folylpolyglutamates from that which is not incorporated. The columns (0.4 X 6 cm) were packed with DE52 which was extensively washed with I M NaCl and equilibrated with 10 mM Tris-CI (pH 7.5), 110 mM NaCI. Approximately 2 mm of acid-washed cellulose equilibrated with this buffer was layered on top of the DE52. Less than 30 min before use, the columns were washed with 3 ml of this buffer containing 25 mM 2-mercaptoethanol (column buffer). Standard assay mixtures (0.85 ml, pH 8.4) which contained Tris-C1 (0.1 M ) , ATP (5 mM), MgCI:! (10 mM), KC1 (20 mM), 2-mercaptoethanol (100 mM), (dl)-HiPteGlu (35 p ~ ) , 4 mM [ 'H]glutamate (4 x 10" cpm/ pnol). and enzyme were incubated at 37°C. When crude extracts were assayed, maximum activity was obtained when the ATP and MgCI, concentrations were doubled because of ATPase activity. High concentrations of MgATP did not inhibit the partially purified rat liver synthetase. At the end of the incubation period, the reaction mixture was diluted with 1 ml of ice-cold 25 mM 2-mercaptoethanol and transferred to an ice water bath. Each reaction mixture was transferred to its respective DEY2 column with a Pasteur pipette. The assay tube was rinsed with 1 ml of column buffer and the rinse was transferred to the column. The columns were washed with column buffer until a total of 14.75 ml was collected. All polyglutamates were eluted with 3 ml of 0.1 N HCI following the wash.

The washing conditions were established using a variety of standard folyldiglutamates synthesized by chemical and enzymatic methods from synthetic PteGlur (9). It was determined that these washing conditions allowed quantitative retention of the di- (and higher polyglutamate) forms of PteGh, HLPteGlu, H,PteGlu, 5,10-CHr- H,PteGlu, and 5-CHI-H4PteGlu. The diglutamate of 10-HCO- HiPteGlu was quantitatively retained only if the NaCl concentration was lowered to 80 mM. Activity in this assay was always corrected for incorporation that occurred in an appropriate blank lacking (dl) . HJ'teGlu (or other folate). One unit is defined as the net incorporation of 1 pmol of ['H]glutamate/h into the polyglutamate fraction. When samples were to be analyzed by HPLC, the same procedure was followed except that columns were eluted with 50 mM H,PO,, 50 mM 2-mercaptoethanol and a portion was analyzed as described under "High Pressure Liquid Chromatography."

Conjugase activity was measured by the assay of Krumdieck and Baugh (20). C. acidi-urici formyltetrahydrofolate synthetase was

assayed by the method of Habinowitz and Pricer (21). Lactic and malic dehydrogenases were assayed by methods in the Boehringer Mannheim Blochemica Information. The activities of IO-HCO- H,PteGlu deacylase and 5-CH:1-H1PteClu demethylase were assaved under synthetase assay conditions using as substrate lO-["ClHCO- H,PteGlue (5 PM) and 5-["C]CH.,-H,PteGlu., (16 pM), respectively, and a procedure similar to that used to assay conjugase (20). Activity was defined as the picomoles of "C released (not absorbable to charcoal) after correction for the nonenzymatic release. These assays were sensitive enough to detect 3 pmol of deacylation and 100 pmol of demethylation during a 3-h incubation.

Folate Compounds-Synthetic PteGlur (9) was supplied by Dr. J. 0. Knipe. Synthetic (f)-5-CH.r-H4PteGlu was the generous gift of Drs. D. W. Horne and C. Wagner. PteGl~2-['~CC]Glu, the conjugase substrate, was a gift of Dr. C. M. Baugh. Pteroyl-D-glutamate and an analog of methotrexate in which the glutamate was replaced with aspartate were gifts of Lederle. Methotrexate and (d0-5-CH1- H,homoPteGlu were supplied by the National Cancer Institute. (d l ) - HIPteGlu was prepared by hydrogenation of folic acid over platinum oxide in neutral solution (22) and purified by chromatography on DEAE-cellulose (23). (dl)-5,10-CHL-H1PteGlu was prepared by mix- ing formaldehyde with (dl)-HIPteGlu (24). HJPteGlu was prepared by dithionite reduction of folic acid (25) and purified (at 4°C) on DEAE-cellulose in 10 mM Tris-CI (pH 7.5), 5 mM dithiothreitol with a gradient from 0 to 500 mM NaCI. (dl)-5-CHi-HIPteGlu (Sigma) was purified on DEAE-cellulose as above except that the gradient was from 0 to 400 mM NaC1. (dl)-5,10-Methenvl-HIPteGlu was synthesized from (dl)-5-HCO-H4PteGlu (calcium salt) (26) and purified according to Curthoys (27) . (2)-5,10-Methenyl-H,PteGlu was prepared by the method of Curthoys (27). Solutions of (dl)- or (l)-lO-HCO-HIl'teGlu were prepared by dissolving (d l ) - or (l)-5,10-methenyl-HlI'teGlu, respectively, in 0.1 M Tris-CI (pH 8.85), 50 mM 2-mercaptoethanol. After standing a t room temperature for 1 h, the material was completely converted to the 10-formyl derivative. (d)-HiPteClu was isolated as a side product of the (1)-5,10-methenvl-HII'teGlu preparation. Its identity was established by its UV spectrum and its lack of activity with clostridial lO-HCO-H,PteGlu synthetase, which is specific for the (I)-isomer (28). In addition, i t s reaction products with formaldehyde were spectrally identical with those of (l)-H,PteGlu and both were identical with those described in the literature (1). (l)-HII'teGlu was prepared according to the general procedure of Curthoys et al. (29) and purified on 1)EAE-cellulose using a gradient elution from 10 to 200 mM Tris-CI (pH 7.5), 5 mM dithiothreitol. It was identified both by its spectrum and by its substrate activity with clostridial IO-HCO- H,PteGlu synthetase. Pte-wGlu was purified by chromatography on DKAE-cellulose using a gradient of 50 to 1100 mM ammonium acetate. pH 8.0. Pte-D-Glu, identified by its spectra at pH 1, 7, and 13, was lyophilized to remove ammonium acetate. H2PteGluL, (dl)-HIPteCluL, (dZ)-H4Pte-['JC]Glu-y-Glu (9), Idl)-5,10-CH~-H~PteGlu2, and (0-10- ['TlHCO-H,PteGlu, (1.1 X 10* cpm/pmol) were prepared analo- gously to the monoglutamates. (rf1)-5-["C)CHI-HII'teGlu~ was prepared by reduction of PteClu, to HII'teCluL with sodium borohydride, addition of ["Clformaldehyde to form 5,10-CHL-H.iPteGlur, and reduction with sodium borohydride (30, 34). (l)-HII'teGlu:> and (1) -[2- "C]H,PteClu were synthesized by direct reduction' of PteClw (9) and [2"'Clfolic acid, respectively. with dihydrofolate reductase from L. cusei. Both compounds were purified on QAE-Sephadex (31).

High Pressure Ltquid Chromatography-HPLC analysis of folylpolyglutamates was performed on a microparticulate anion exchanger (Whatman Partisil-SAX; 10pm) by the procedure of Cashmore et al. (3') except that the initial composition o f the gradient was 0.1 M sodium phosphate, pH 3.3, which resolved mono-, di-, and triglutamates while tetra- and pentaglutamates co-eluted. To simplify analysis and presentation of data (Figs. 2 and 3). the combined tetra-/ pentaglutamate peak was quantitated as if radioactivity were present only in the tetraglutamate. The maximum error introduced by this simplification was determined by resolving the tetra- and pentaglu-

the initial composition was 0.2 M sodium phosphate, pH 3.3) when tamate components (HI'LC conditions as described above except t.hat

pentaglutamate synthesis was maximum (Fig. 4). The amount o f product calculated, assuming that the radioactivity in both peaks was tetraglutamate, was only 10% higher than the sum of the actual values for the tetra- and pentaglutamates. Thus, even the maximum error introduced by this assumption was small and did not affect the conclusions. Sufficient chemically synthesized. oxidized fol.ylpolyglu-

' E. J . Pastore, Y. Gaumont, and K. i. Kisliuk, personal cornmu- " ""

nication.

5778 Rat Liver Folylpolyglutamate Synthetase

tamates to be detected by UV absorbance were co-injected with radiolabeled samples to provide an internal standard. Where possible, authentic standards were synthesized to verify the elution position of the polyglutamate products when reduced/l-carbon-substituted fo- lylmonoglutamates were used as substrates.

Mono-, di-, and trinucleotides are also resolved in this HPLC system and nucleotidase activity was determined from the relative areas of the respective peaks in control and enzyme-treated samples.

Conjugase Treatment-A reaction mixture (1.75 ml) containing only 5 pM (dl)-H,PteGlu but otherwise standard assay components was incubated under N2 for 4.5 h at 37°C. At that time, two samples (0.25 d) were removed, processed as in a standard assay, and eluted as for HPLC analysis. These samples determined the polyglutamate distribution before conjugase treatment. Samples (0.56 ml each) of the remainder were transferred to two separate tubes. To one tube was added 0.1 ml of 0.01 M Tris-C1, pH 8.0, and to the other was added 0.1 ml (12 pg) of chicken pancreas conjugase (19). which was exhaustively dialyzed against 0.01 M Tris-C1, pH 8.0. When assayed at pH 8.3 (37°C) with 2500 pmol of PteGl~~-[ '~C]Glu in 0.5 ml, 2 pl of this conjugase preparation released 130 pmol of ['4C)glutamate in 10 min. The buffer and conjugase-treated samples were incubated a further 1 h at 37"C, after which duplicate 275-pl aliquots were removed from each and processed as above. Polyglutamate products were analyzed by HPLC as described in that section. A control lacking (dl)-H4PteGlu showed no ['H]glutamate incorporation a t 4.5 h.

Identification of Radiolabeled Moiety Incorporated into Prod- uct-Product was isolated by combining eight standard reaction mixtures containing 35 p~ (l)-H4PteGlu after incubation for 10 h under N2 in the dark. Following the usual loading and buffer wash procedures, a 5-ml HzO wash eluted most of the salts but no radioactivity. Product was eluted with 0.1 N HCl and the fractions were pooled and divided into two equal portions (125,OoO cpm; 33,000 pmol of "H product each). All further operations were carried out in duplicate. The samples were reduced to dryness on a rotary evapo- rator, dissolved in 1 ml of 6 N HCI, sealed under vacuum, and hydrolyzed for 20 h at 110'C. The resulting hydrolysate was dried under vacuum a t high temperature, redissolved, and analyzed by dansylation and 2-dimensional chromatography (35) and chromatography on an amino acid analyzer. A portion of the hydrolysate was redissolved in 100 p1 of 0.1 M NaHCOs (final pH 7), lyophilized, redissolved in 100pl of 0.1 M NaHCO:, (final pH -8.5), and dansylated by incubating (37"C, 15 min) with 50 pl of freshly prepared dansyl- chloride (5 mg/ml in acetone). Samples (1 p1) were chromatographed on one side of a polyamide plate (Pierce; 5 X 5 cm) in either solvents 1 and 2, or 1, 2, and 3 of Weiner et al. (35). A mixture of dansylated (35) standard amino acids (Beckman Instruments) co-chromatographed on the reverse side of the plate allowed identification of amino acids in the unknown. Radioactive spots were detected by fluorography (36). Dansyl amino acids were detected by fluorescence under long wave UV light.

Amino acid analysis of the hydrolysate was kindly performed by Dr. K. Williams (Department of Molecular Biochemistry and Bio- physics, Yale University) on a Beckman model 121M analyzer equipped with an Autolab System AA computing integrator. The single microbore column was filled with Durmm DC-4A resin and was sequentially eluted with three sodium citrate buffers (pH 3.25, 4.2, and 7.0, respectively) followed by a 0.2 N NaOH wash. Fractions (160 pl) were collected by directing the column effluent to a fraction collector rather than to the ninhydrin reaction bath. Each fraction was mixed with 0.5 ml of water and 5.5 ml of Aquasol (New England Nuclear) and counted. Authentic [2,3-"H]glutamate was analyzed separately to serve as a standard.

Analysis of Double Labeled Product on DEAE-Cellulose-Stan- d a d assay Conditions were used except that 78 pg (1340 units) of partially purified rat liver folylpolyglutamate synthetase, 40 mM KCI, 5.6 p~ (l)-[2-14C]H4PteGlu (12.5 x 10" cpm/pmol), and 4 mM ~-[2,3- ,'3H]glutamate (3.8 x 10" cpm/pmol) were included. Specific radioac- tivities were determined under the same counting conditions used for all column fractions. Controls lacked either labeled HAPteGlu or labeled glutamate. All assays were incubated for 11 h at 37°C in the dark. Individual assays were diluted with 1 ml of 50 mM 2-mercaptoethanol and applied at room temperature to a DE52 column (0.4 X 6 cm) which was equilibrated with 5 m~ potassium phosphate (pH 7.0). 50 mM 2-mercaptoethanol. The assay tube was rinsed with 1.35 ml of this buffer and the rinse was applied to the column. Elution was carried out with a linear gradient of 5 to 500 mM potassium phosphate (pH 7.0), 50 mM 2-mercaptoethanol in a total Volume Of 60 ml (17). From each fraction (0.95 ml), 0.9 ml was removed, mixed with 1.1 ml

of water and 12 ml of Aqueous Counting Scintillant (Amersham/ Searle), and counted. Counting efficiency was constant throughout the gradient.

Miscellaneous Procedures-A modified phenol reagent method was used to determine protein concentration in samples that had been precipitated with 5% (w/v) trichloroacetic acid (21). All ultra- violet spectra were recorded on a Cary model 15. A Beckman LS230 scintillation counting spectrometer was used to count aqueous samples (1 m l ) mixed with 10 ml of scintillation fluid (33). The tritium counting efficiency was 24% and "C efficiency was 708. Fractions collected from HPLC gradients (0.8 ml) were diluted with 2 ml of water and mixed with 10 ml of either Aquasol-2 (New England Nuclear) or Insta-Gel (Packard). Tris-ATP, for use in monovalent cation studies, was prepared by treating NarATP with excess Dowex 50 and neutralizing the free acid with Tris base.

RESULTS

Enzyme Purification-Folylpolyglutamate synthetase was purified3 approximately %-fold from rat liver (Table I); this preparation was stable for several weeks at -20°C. Polyacryl- amide gel electrophoresis in sodium dodecyl sulfate (not shown) revealed several bands of protein indicating a low degree of purity. Conjugase (pteroylpolyglutamate hydrolase) activity, however, was nearly undetectable in the final preparation. Two lines of evidence indicated that the conjugase activity remaining was insignificant. First, when conjugase was measured (20) under synthetase assay conditions with polyglutamyl substrate levels approximating the amount synthesized during a routine synthetase assay, the hydrolysis of substrate was less than 2%. Also, it was possible to nearly quantitatively convert a monoglutamate substrate to tetra- and pentaglutamate forms (see Fig. 13s) which would not be possible if conjugase activity were significant. Attempts at further purification, for example by chromatography on DEAE-cellulose or hydroxyapatite, were unsuccessful because of extreme enzyme instability a t 4°C.

General Properties-The substrate requirements of both the crude and partially purified enzyme were determined (Table 11). Although crude extracts showed substantial apparent activity in the absence of either (dZ)-H4PteGlu or ATP, maximum product formation occurred in the presence of both these substrates. HPLC analyses of the reaction products synthesized by the crude enzyme (see Fig. 3s) demonstrated that only in the presence of (dl)-H,PteGlu and ATP did a peak of radioactivity appear at the same elution position as authentic (dZ)-H4PteG1u2. No requirement for KC1 was evident for the crude extract. Partially purified enzyme displayed an absolute requirement for both (dl)-H,PteGlu and ATP. Although KC1 dependence could not be determined here because this enzyme preparation contained high levels of KCI, a dependence was established in other experiments (see Fig. 8s). Under standard assay conditions, the partially purified enzyme (40 pg) catalyzed the incorporation of ['HJglutamate into polyglutamates linearly for 3 h and then continued incorporation for at least 7 h longer a t a steadily diminishing rate. Incorporation of ['Hlglutamate was linear with respect to enzyme concentration up to 60 pg, the highest concentration tested, during 3-h incubations. This partially purified folylpolyglutamate synthetase was used in the characterization described below.

Table I1 also illustrates the low background and sensitivity obtainable with this assay. Although approximately 4 X IO6

' The purification of the enzyme as well as much of the supporting evidence, including Figs. 1s to 13s and Tables 1s and 2S, is presented as a miniprint supplement immediately following this paper. Mini- print is easily read with the aid of a standard magnifying glass. Full size photocopies are available from the Journal of Biological Chem- istry, 9650 RockviLle Pike, Bethesda, MD 20014. Request Document No. 79M-2608, cite author(s), and include a check or money order for $3.60 per set of photocopies.


TABLE I Purification of folylpolyglutamate synthetase from rat liver

Activity

Fraction

mg units/mg 9;; Crude Extract 6,100 320 100

Phosphocellulose 310 2,900 46 Sephadex G-150 40 17,200 35

(NH,),SO,, 0 to 35% 1,610 820 67

TABLE I1 Substrate requirements of rat liver folylpolyglutamate synthetase

Standard assay conditions were used except that single components were omitted as noted. Assays of crude extracts (10 mM ATP; 20 mM MgC12; 0.56 mg of protein) showed linear incorporation of ["Hlgluta- mate for 1 h only.

Omission Crude G-150 enzyme pmol f 3HJGlu incorporated/h

None 245 755 Enzyme 85 ATP

60 115 60

(dl)-H,PteGlu 140 60 KC1 230 ND"

' I ND, Not determined.

cpm of [:'H]glutamate are included in each assay point, less than 350 cpm appear as "product" if no enzyme is added. This low background, coupled with the high specific radioactivity of the [,"H]glutamate, allowed quantitation of as little as 25 pmol of product.

Temperature Effect-Determination of the effect of temperature on this reaction was important from two practical standpoints. First, the synthetase activity, even with partially purified enzyme, was relatively low so that any increased activity at another temperature during a practical incubation period (3 h) would be helpful. Second, termination of the reaction was dependent on dilution and lowering the temperature to 4"C, and it was necessary to validate this procedure. Experimentally, the activity at or near 37°C was the highest achievable (see Table IS). Increasing the temperature to 50°C or decreasing it to 22°C substantially decreased the activity. Incubations at 0°C showed little synthesis over 3 h; thus, dilution with ice-cold 2-mercaptoethanol was effective in ter- minating the reaction.

p H Optimum-The pH optimum of rat liver folylpolyglutamate synthetase was between pH 8.2 and 8.6 at 37°C (see Fig. 4s). Activity fell off rapidly at higher or lower pH so the rate was reduced by 50% a t approximately pH 9 or 7.7. The buffering species had no effect on activity. Tris, diethanola- mine, and ethanolamine worked equally well in the optimum region. Tris was the buffer of choice, however, because its blank value (radioactivity as product in the absence of enzyme) was approximately 50% of that obtained with the other buffers. The pH of choice was 8.4 because in crude extracts the apparent activity in the absence of (dl)-H,PteGlu dimin- ished with increasing pH.

Since other folate substrates could display different pH optima either through intrinsic properties of the folate or because they are substrates for a second synthetase that co- purified in our preparation, activity of several other folates as a function of pH was investigated (see Fig. 5s). The pH optima of folic acid, (Z)-lO-HCO-&PteGlu, (Z)-H,PteGlu, and (I)-5-CH:1-H4PteGlu were all near pH 8.4 (37°C) and these substrates showed the same general behavior as (d l ) - H,PteGlu with increasing or decreasing pH. An exception to this pattern of behavior was (dZ)-H4PteGlu2. This substrate

displayed the same pattern of decreasing activity above pH 8.4 but the activity remained relatively constant between pH 8.4 and 7.4. Below pH 7.4, the activity began to decline.

Native Molecular Weight-The native molecular weight of rat liver folylpolyglutamate synthetase was determined on a calibrated (39) Sephadex G-150 column (see Fig. 6s). The synthetase eluted a t a position corresponding to M, = 69,000 -+ 9,OOO. The stated error assumed a one-fraction variation in the elution of the synthetase.

Thiol Requirement-The rat liver enzyme, unlike bacterial synthetases, utilizes oxidized substrates (see below) which are stable in the absence of thiols; thus, the thiol requirement of this enzyme could be determined. Using an oxidized substrate, it was possible to demonstrate a saturable requirement for mercaptoethanol with optimal activity achieved at or above 100 mM (see Fig. 7s).

Activation by Monovalent Cations-Rat liver folylpolyglutamate synthetase was virtually inactive in the absence of monovalent cations. KC1 was the most effective activator with maximum stimulation achieved at 10 to 20 mM (see Figs. 8s and 9s). Activity decreased as the concentration was raised beyond 20 mM. Thus, KC1 was absolutely required for enzyme activity, and activity was stimulated in a concentration-dependent manner.

The pattern of activation by other monovalent cations was determined (see Fig. 9s). Both NHs' and Rb' activated the synthetase although neither was as effective as K'. No activity was detected if Cs+, Na', or Li' were substituted for K'. This pattern of specificity and the molar concentration required for activation are the same as are seen for a wide variety of potassium-activated enzymes (41).

Amino Acid Substrate-L-Glutamate displayed apparent Michaelis-Menten kinetics with the synthetase with a K,, of 0.65 2 0.09 mM (see Fig. 10s). A replot of the data gave a Hill coefficient near 1, indicating no cooperativity. D-Glutamate, L-glutamine, L-aspartate, glycine, p-aminobenzoic acid, L-methionine-DL-sulfoximine, a-aminoadipic acid (the homolog of glutamic acid containing 1 additional methylene carbon), glu- tathione (y-glutamyl-cysteinyl-glycine), and y-L-glutamyl-t- glutamate did not decrease incorporation of subsaturating levels of ["Hlglutamate into product and therefore are neither substrates nor inhibitors of the synthetase (data not shown). Under these conditions, a-ketoglutarate inhibited weakly while unlabeled L-glutamate strongly decreased labeled product formation. The synthetase thus displayed rigid specificity for the amino acid. The addition of glutamates one a t a time was suggested since y-L-glutamyl-L-glutamate did not inhibit the reaction.

Kinetics with ATP and Nucleotide Specificity-Prior to experiments with nucleotides, it was demonstrated, using a HPLC analysis system, that the enzyme preparation contained negligible (<5% hydrolysis) nucleotidase activity under standard assay conditions. Kinetic analysis with ATP as var- iable substrate revealed that its apparent K , was less than 70 PM (see Fig. 11s). An exact value was not obtainable because of nonlinearity of activity with respect to time at low ATP, even at incubation times as short as 20 min (see Figure 11s).

Other nucleotides were tested for their ability to replace ATP in this reaction. dATP, GTP, and dGTP were able to substitute for ATP to varying degrees while both UTP and TTP were inactive (see Table 2s). Activity was detectable when 5 mM ADP replaced ATP; however, it was demonstrated that contaminating adenylate kinase synthesized enough ATP during the incubation to account for this activity. ADP was therefore probably not a substrate. AMP was completely inactive as a substrate.

Folate Specificity-In addition to lacking conjugase activ-

5 780 Rat Liver Folylpolyglutamate Synthetase

ity, sensitive assays demonstrated that neither 10-HCO- H4PteGlu deacylase nor any 5-CH,?-H4PteGlu demethylating activity was present in the partially purified preparation. The absence of interconverting/degrading enzymes allowed the determination of substrate activity of most common folates (Table 111). All of the naturally occurring folates tested were utilized as substrates by the rat liver folylpolyglutamate synthetase, although with different efficiencies. Reduction of the pyrazine ring was not essential (folic acid was a substrate); however, substrate activity increased as the degree of reduction increased. At the lower (more physiological) concentration, H4PteGlu and 10-HCO-H4PteGlu were the best substrates, suggesting that in vivo these forms may serve as the substrates for at least the first step in the polyglutamylation reaction. It is significant that 5-CH:1-H4PteGlu, the main serum folate in mammals, was a substrate. In addition, (l)-5- CH:,-H4PteGlu was active, which argued against the possibility that the activity with the (dl)-mixture was the result of slow metabolism of the (d)-isomer only (43).

The apparent lack of specificity of the synthetase for the pteridine moiety was confirmed by studies with several folate analogs. The potent chemotherapeutic agent MTX (2,4-diamino-10-methyl-folic acid) was also a substrate for the rat liver synthetase (see Fig. 12s). At concentrations greater than 35 p ~ , its activity was higher than the highest level obtainable with (dl)-H,PteGlu, at 35 PM. Since MTX differs from folic acid, a relatively poor substrate, by only two substitutions, two other analogs were tested to determine which features were important for the increased activity. 4-Amino-folic acid (aminopterin) was nearly as good a substrate as MTX while 10-methyl folic acid, like folic acid, was a poor substrate. Thus, the 4-amino substituent was responsible for the increased activity of MTX. The lack of pteridine specificity was further demonstrated by the substrate activity of 5-CH:c-H4-homofo- late (Table III), which contains an extra methylene group between C-9 and N-10, as well as the substrate activity4 of the 5,s-deaza analog of folic acid (44).

Stringent substrate specificity was again evident, however, when folate analogs containing an amino acid other than L- glutamate were tested. Neither the analog of MTX containing aspartate nor the analog of folic acid containing D-glutamate were substrates. Of greater significance was that neither analog a t 17 PM inhibited incorporation of [:'H]glutamate into 5 PM (dl)-H,PteGlu. Apparently, a terminal L-glutamate is absolutely required for binding to the enzyme.

Isomeric Specificity-Purified (d ) - and (l)-H4PteGlu and their racemic mixtures were prepared and tested for substrate activity (Fig. 1). Racemic mixtures of (d ) - and (I)-H4PteGlu, either synthesized by catalytic hydrogenation or equimolar mixtures of pure (d ) - and (I)-isomers, behaved similarly. Activity rose to a maximum at 35 PM total isomer concentration, and decreased activity was evident at high substrate concentrations. The pure (I)-isomer achieved higher incorporation than did the optimum concentration of the racemic mixture and no decrease in incorporation was evident a t high (I)-H4PteGlu concentration. These two observations suggested that the (d)-isomer was inhibitory, but this conclusion was inconsistent with the lower incorporation by the pure (I)- isomer compared with the racemic mixture at (I)-H4PteGlu concentrations of 5 PM or less (Fig. 1). This discrepancy was partially understandable when the pure (d)-isomer was ex- amined (Fig. 1). (d)-H4PteGlu, which is generally considered to be biologically inactive, was a substrate for rat liver synthetase. AlthoEgh its maximum incorporation was only 50% of that of (l)-H4PteGiu, a similar concentration of each isomer

' P. Hsieh, J. J. McGuire, J. B. Hynes, and J. It. Bertino, unpub- lished observation.

TABLE 111 Folate specificity of rat liver folylpolyglutarnate synthetase

Assays were performed under standard conditions but each folate was present at either 5 or 35 p ~ , except for (I)-5-CH.,-HoPteGlu, which was present at 2.5 and 17.5 PM. Results presented are the means of duplicate determinations. When PteGlu and H2PteGlu were used as substrates, assays also contained 1 p~ 2,4-diamino-5-methyl-6- [(3,4,5-trimethoxyanilino)methyl]quinazoline, an inhibitor of dihydrofolate reductase (42), to prevent formation of H,PteGlu. In separate experiments, this quinazoline was shown to be neither a substrate nor an inhibitor of rat liver folylpolyglutamate synthetase. Activity of (dl)-5-HCO-H4PteGlu was determined in a separate experiment in which activity of (dl)-H,PteGlu was redetermined as a control. Values presented are the computed activities relative to H'PteGlu after normalization.

I 5 PM

None 0 PteGlu 270 HzPteGlu 960 (d2)-H4PteGlu 1700 (dl)-CH,-H,PteGlu 1150 (dl)-10-HCO-H4PteGlu 1500 (dl)-5-HCO-H4PteGlu 300 (dl)-5-CH.~-HtPteGlu 330 (1)-5-CH,1-H~PteGlu 170 (dl)-5-CH.r-H4homoPteGlu 70 i

0 16 56

100 68 88 18 19 10 4 __

35 PM

>mol ['HI- Glu incor- ~ ~ ~ i ~ , ~ ~ porated/3 Elative to ,PteClu h

2150

1640 1380 690

1225 48 1075 42 380 15

1 0 10 20 30 40 50

[ H4PleGlu] pM

FIG. 1. Comparison of (I)-H4PteGlu, (d)-RPteGlu, and (dl)- RPteGlu as substrates for rat liver folylpolyglutamate synthetase. Compounds were prepared as described under "Materials and Methods." Standard assay conditions were used with (1)- H4PteGlu (O), (d)-H,PteGlu (O), (dl)-H4PteGlu (m), or an equimolar mixture of pure (1)- and (d)-H,PteClu to simulate the synthetic racemic mixture (0). Concentrations of both ( d l ) and the mixture of ( d ) and ( I ) are given in terms of the (1)-isomer only, i.e. their total concentration is 2 times the indicated concentration.

was required to reach half-maximum incorporation. Compar- ison of (d)-H4PteGlu activity (relative to (dl)-H,PteGlu) and the values for other folates (Table 111) revealed that ( 4 - H,PteGlu served at least as well as most folates, including ( I ) - 5-CHs-H4PteGlu. No decrease in incorporation was evident at high concentrations of (d)-H,PteGlu. HPLC analysis (not shown) of the products obtained at 17.5 PM (d)-H4PteGlu confirmed that polyglutamates were synthesized and demonstrated that products up to at least the tetraglutamate could be synthesized. Substrate activity of the (d)-isomer was not the result of racemization of the (dl- into the (I)-isomer during the incubation. Analysis of reaction mixtures with formyltetrahydrofolate synthetase, which is specific for ( I ) -


H,PteGlu, and not inhibited by the (dbisomer (28), demonstrated that (l)-&PteGlu was quantitatively recovered after 3-h incubations when 40 p~ (I)-isomer was initially present, but that none was detectable if 40 p~ (d)-isomer was initially present. Conversion of less than 2% of the (d)-isomer to ( I ) - isomer would have been detectable. As a result of the substrate activity of the (d)-isomer, the activity of the (dl)-mixture at low concentrations is greater than that expected on the basis of the concentration of the (I)-isomer alone.

I t is not clear why inhibition occurred at high (dl)-H,PteGlu concentrations. If two substrates are utilized in Michaelis- Menten manner by a single enzyme with the same K , but different V,,,, then the kinetics with both substrates present is still Michaelis-Menten. The K,,, will be unchanged but the V,,, will depend on the relative concentrations of the two substrates (45). However, this kinetic analysis may not be valid in the present case because the Michaelis-Menten as- sumptions may not be valid.

The lack of isomeric specificity may not be limited to H,PteGlu. Comparison of the concentration dependence of (dl)- and (l)-10-HCO-H4PteGlu substrate activity showed a pattern similar to the unsubstituted tetrahydrofolates (data not shown). At low, equal (with respect to the (Z)-isomer) concentrations, the (dl)-mixture gave slightly higher activity than the pure (I)-isomer (as in Fig. 1 at 5 p~ or less). The level of incorporation by the ([)-isomer eventually exceeded the highest achieved by the (dl)-mixture. Decreased incorporation was again evident a t higher concentrations of the (dl)-mixture. Thus, although the (d)-isomer was not directly tested, the kinetic pattern suggested that (d)-IO-HCO- H,PteGlu was also a substrate for the rat liver synthet.ase.

Nature of the Glutamate Linkage-Chicken pancreas conjugase specifically hydrolyzes folylpoly-y-glutamates to yield a folyldiglutamate product (19). As shown in Table IV, the products synthesized in vitro by the rat liver folylpolyglutamate synthetase with H4PteGlu as the substrate were sensitive to digestion by a partially purified (19) chicken pancreas conjugase. Extensive synthesis of H,PteGlu,/Glu, occurred prior to treatment while less di- or triglutamate accumulated (Line 1). If buffer alone was added to an aliquot of this reaction mixture (Line 2), during a 1-h incubation a modest increase in the combined tetra/pentaglutamate fractions occurred at the expense of the diglutamate because the synthetase, although diluted, was still active. Treatment of an aliquot for 1 h with conjugase exhaustively dialyzed against this same buffer caused a dramatic decrease in the total amount of ["H]giutamate recovered as product and the decrease occurred in products with three or more glutamates (Line 3). There was a concomitant increase in the diglutamate peak. Nearly quan-

titative recovery of the diglutamate product after this conjugase treatment indicated that only specific degradation OC-

curred. These results demonstrated the presence of a y-glutamyl linkage in the in vitro synthesized product.

Digestion of the higher polyglutamates, particularly the triglutamate, was incomplete. This may be because insuffi- cient time was allowed for digestion or because competition between the synthetase, which was still active under these conditions, and conjugase did not allow complete hydrolysis.

Zdentification of the Incorporated Label as Glutamic Acid-When salt-free product was hydrolyzed, dansylated, and subjected to 2-dimensional thin layer chromatography (35) and fluorography (36), a radioactive spot corresponding to the dansyl glutamic acid standard was observed in two solvent systems. No radioactivity co-chromatographed with the dansyl derivative of any other common amino acid; however, a second radioactive spot appeared at the solvent front of the first dimension in both solvent systems. No dansyl fluorescence coincided with this material. Analysis of the hydrolyzed material on an amino acid analyzer demonstrated a radioactive peak, containing 92% of the injected material, which eluted at the same position as authentic [:lH]glutamic acid. Thus, the dansyl negative, radioactive spot noted above probably represented unreacted glutamic acid since the dansyl reaction is not quantitative in most applications (46). This chromatographic evidence, coupled with the strict glutamic acid specificity noted above, indicated that only labeled glutamic acid was incorporated into the products.

Verification of Folate Nature of Product and Polygluta- mate Length-Assays containing (l)-['4C]H4PteGlu and ["Hlglutamate confirmed that products of the synthetase reaction contained both a folate and glutamate and that they occurred in the correct ratio for a particular polyglutamate. To simplify analysis and assure that sufficient radioactivity was present in product to give meaningful results, the reaction was run under conditions (see below) where t.he presumed pentaglutamate was formed almost exclusively. HPLC characterization of the products confirmed that 91% of the product eluted at the same position as authentic (l)-H4PteGlur, while the remainder eluted as the tetraglutamate. DEAE-cellulose gradient chromatography (17) was then used for quantitation because entire reaction mixtures could be analyzed directly. Assays lacking either ['4C]H4PteGlu or ["Hlglutamate had only single peaks early in the elution profile and radioactivity was background throughout the remainder of the gradient, where polyglutamates elute (17). Chromatography of the complete reaction mixture (see Fig. 13s) showed an additional late eluting peak, containing both labels with a molar ratio :'H/'4C of 3.6 k 0.2 (mean t S.D.), consistent with its identi-

TABLE IV Conjugase sensitivity of product synthesized in I

A 1.75-ml reaction mixture containing standard components and 5 p~ (dl)-H,PteGIu was incubated for 4.5 to allow accumulation of polyglutamates. At that time, aliquots were removed and analyzed immediately or treated for 1 h either with 0.01 M Tris-CI, pH 8.0, or with partially purified (19) chicken pancreas conjugase (12 pg) which had been exhaustively dialyzed against the same buffer. Exact conditions and the HPLC methods used to determine the product distribution are described under "Materials and Methods." Values are

i tro by rat liver folylpolyglutamate synthetase averages of duplicate determinations f S.D. The HPLC system did not resolve tetra- and pentaglutamate derivatives so two values are presented. The top number indicates the picomoles of product if all the radioactivity associated with this combined peak were present as tetraglutamate while the lower value (in parentheses) is the amount if all the radioactivity were present as pentaglutamate. These two values then define the upper and lower limits, respectively, of the amount of product present in the unresolved mixture.

Condition ["HIGlu Incorporation H,PteGlu.. H,PteClu, H4PteGlu,/Glu-, Total product ______ pmol

-

1. 4.5-h incubation 2390 f 60 190 f 30 320 f 10 530+ 10 1040 f 50

2. Buffer (1 h) (980 f 50)

130 f 0 300f0 620 f 10 1070 f 10

3. Conjugase (1 h) (lo00 * 10)

790 f 80 80 f 0 152 5 885 f 85 (15 f 4) (885 k 8 4 )

(470 -t 10)

(550 -t 10) 2580 f 30

1020 & 80

5 782 Rat Liver Folylpolyglutamate Synthetase

Fa’ , , , , , , , , , J OO 2 4 6 8 IO

0

Incubatton Time ( h )

FIG. 2. Product distribution as a function of time at 5 p~ (dl)-H,PteGlu. Standard assay conditions were used at 5 p~ (d l ) - H,PteGlu in a total volume of 2.75 ml. The assay tube was flushed with N2, sealed, and incubated at 37°C. At each time point, duplicate samples (0.25 ml) were diluted into 1 ml of ice-cold 25 mM 2-mercaptoethanol. The assay tube was immediately reflushed and sealed, and the incubation continued. Each sample was then processed, analyzed by HPLC, and quantitated as described under “Materials and Meth- ods.”

37 Total Product A

35s4 1

22 r ’AI7 U Y

Incubation Time ( h )

FIG. 3. Product distribution as a function of time at 35 pM (df)-&PteGlu. Conditions were identical to those described in the legend to Fig. 2 except (dl)-H,PteGlu was initially present at 35 pM.

tication as a folylpentaglutamate. Synthesis of Polyglutamates as a Function of Time with

Tetrahydrofolate as Substrate-The distribution of polyglutamate products was determined through a 10-h time interval at 5 and 35 ~ L M (dl)-H4PteGlu. At 5 p~ substrate (Fig. 21, synthesis of products as long as tetraglutamate was easily discernable even at the shortest time (1 h). At that point, approximately 20% of the substrate was converted to polyglutamyl forms. By 3 h, both di- and triglutamate derivatives reached their maximum accumulation and subsequently both decreased. The levels of these products must have represented

a quasi-steady state in the synthetic system because the combined tetra-/pentaglutamate peak continued to accumu- late product at a nearly linear rate. At 10 h, the total amount of polyglutamate product was approximately 1070 pmol (85% of the original monoglutamate substrate). The tetra-/pentaglutamate peak contained between 510 and 680 pmol (40 and 55% of original monoglutamate) assuming this was all penta- and tetraglutamate, respectively. Thus, at this concentration of (dl)-H,PteGlu, only low levels of short (di- and triglutamate) polyglutamates accumulated, while accumulation of longer (at least tetraglutamate) chains was favored.

At 3 6 p ~ (dl)-H,PteGlu (Fig. 3), the total product and total [‘Hlglutamate incorporated were higher than at 5 p~ for each time point. However, the distribution of products at 35 p~ (dZ)-H,PteGlu was markedly different. The diglutamate pre- dominated throughout the incubation period. Accumulation of triglutamate showed a noticeable lag and then increased, but only approached the level of diglutamate after 10 h. Synthesis of tetra-/pentaglutamates was undetectable until 3 h and then their synthesis was so slow that after 10 h not only was the percentage of conversion to tetra-/pentaglutamate less but their absolute amount was 3-fold less than that observed a t 5 PM (10 h). Thus, at higher concentrations of H,PteGlu, only low levels of longer chains (tetra-/pentaglutamate) were synthesized while high levels of shorter chains, especially diglutamate, accumulated.

Two additional observations on this behavior have been made. First, the concentration dependence of the product distribution is not an artifact of using the racemic mixture of tetrahydrofolate isomers, both of which are substrates (Fig. 1). Five-hour incubations with 5 or 35pM (1)-H,PteGlu showed a similar concentration dependence of product length (data not shown) although the absolute amount of each product was different, presumably because of the different activity of the isomers (Fig. 1). Second, this concentration effect is not peculiar to H,PteGlu. Analysis of the products from 5-h incubations with 2.5 or 25 ~ L M (l)-10-HCO-H4PteGlu demonstrated the product distribution was skewed toward longer polyglutamates at the low concentration.

An additional finding of these experiments was an apparent limit to the length of polyglutamate synthesized. Reanalysis of the products at 5 p~ H,PteGlu after a 10-h incubation was carried out in an HPLC system which increased resolution of higher polyglutamates (Fig. 4). The oxidized pteroyltri-, tetra-, and pentaglutamate standards were resolved from one

N PtcGlu, A

I 1 0 1 2 3 4 5 r e - 0.04 x

D h)

0 m

0.02 5

0

Retention Time (min) FIG. 4. =-LC analysis of reaction products with 5 pM

HPteGlu as substrate after 10-h incubation. Reaction conditions were the same as those described in the legend to Fig. 2. HPLC conditions were the same as those described under “Materials and Methods” except that the initial composition was 0.2 M sodium phosphate, pH 3.3. The absorbance profile resulted from co-injection of synthetic polyglutamate standards whose lengths are noted. Data are presented as raw counts (uncorrected for background radiation) to minimize bias.

Rat Liver Folylpolyglutarnate Synthetase 5783

TABLE V Substrate and inhibitor activity of (Z)-H4PteGlus

purified (l)-H,PteGlu and (2)-H4PteGlu5 were substituted as indicated Assays were performed under standard conditions except that

and incubations were for 4.5 h. Values presented are averages of duplicate determinations.

Folate substrate Net ['Hlglutamate in-

(I)-H,PteGlu (O-H,PteGlu?, corporated

PM pmol 5 2090

35 3360 10 130

5 10 940 35 10 1870

another as well as from higher polyglutamates while resolution of the diglutamate was sacrificed. The radioactivity incorporated into H4PteGlu during the incubation eluted with the tri-, tetra-, and pentaglutamate standards. There was no loss of material in this new system because the amount of triglutamate and the sum of the tetra- and pentaglutamate fractions corresponded to the amounts shown in Fig. 2. This analysis demonstrated that the rat liver synthetase was capable of synthesizing products the length of &PteGlus. A total of 380 and 240 pmol of tetra- and pentaglutamate, respectively, were produced, which represents 30 and 20%, respectively, of the initial 1250 pmol of (dl)-H,PteGlu. Thus, conversion to higher polyglutamates occurred efficiently at low KPteGlu. No radioactive peak eluted at the position corresponding to the hexaglutamate (-22 min). Since the sensitivity of product detection increases with each additional labeled glutamate, very low levels of hexaglutamate synthesis would have been detectable. Thus, rat liver folylpolyglutamate synthetase is capable of synthesizing tetrahydro derivatives only up to the pentaglutamate level or the synthesis of higher derivatives is extremely inefficient.

The accumulation of pentaglutamate derivatives indicated that H4PteGlus itself would either be inactive or poorly utilized as a substrate. The results in Table V confirmed that ( I)-H4PteGlus was a remarkably poor substrate although it is still bound to the enzyme because it inhibited the polyglu- tamylationof (l)-H4PteGlu. At 1 O p ~ (l)-H4PteGlu:,, inhibition of activity with 5 PM (l)-H4PteGlu was only slightly greater than inhibition with 35 ~ L M (l)-H,PteGlu (58 uersus 46%), indicating the inhibition was not of a simple, competitive nature.

DISCUSSION

A partially purified folylpolyglutamate synthetase which could convert folate substrates nearly quantitatively into polyglutamates was obtained from rat liver. The authenticity of the reaction catalyzed by this preparation was supported by several lines of evidence. The reaction was absolutely dependent on the presence of a folate, ATP, and glutamate (Table 11) in agreement with the initial description of this enzyme (14). The products of the reaction co-eluted with chemically synthesized folylpolyglutamates in HPLC. Digestion of these products with chicken pancreas conjugase (Table IV) yielded quantitatively a moiety which co-chromatographed with authentic diglutamate. Since this conjugase is a specific pteroylpolyglutamate hydrolase which digests polyglutamates to the diglutamate level (19), this result indicated that the synthetase product contained y-glutamyl linkages. Hydrolysis of the product and analysis of the resulting material indicated that o d y labeled glutamate was incorporated and no other corn- mon amino acid. In addition, other amino acids did not inhibit the incorporation of label while L-glutamate did. Finally, a

double label experiment (see Fig. 135) demonstrated that the products contained both labeled folate and glutamate, and that they were present in the ratios expected for the respective polyglutamate forms. These results, in sum, indicated that the reaction observed involved ATP-dependent addition of L-glutamate in y linkage to folates to form folylpoly-y-glutamates.

Examination of the general properties reported for the few synthetases investigated thus far indicates some degree of similarity with a few striking differences, particularly between the enzymes from prokaryotes and eukaryotes. In both prokaryotes and eukaryotes, a folate, ATP, and L-glutamate are required in a reaction with a high pH optimum (15-17, 40). Specificity for L-glutamate is absolute but any purine triphos- phate may replace ATP. A number of differences are readily apparent. Specific activity of crude extracts, for example, is 50- to 100-fold lower for eukaryotes (this study; Ref. 17) than for prokaryotes (16). The synthetase from E . coli (16) has a significantly lower molecular weight (43,000) than the rat liver enzyme (69,000). Although the enzymes from both classes require potassium, the optimal levels required by the bacterial synthetase are 10-fold higher. The only exception to this requirement is the CHO synthetase, which is stimulated only 15% by 30 mM potassium, while higher levels are inhibitory (17). The most striking dissimilarity is in folate specificity and the products of the reaction. Partially purified synthetases from E. coli and Corynebacterium have strict folate substrate specificity, only one folate is efficiently utilized, and only a single glutamate is added to any significant extent (16, 40). Since these bacteria contain predominantly penta- and tetraglutamate, respectively (3 , 49), there is a t present a lack of fidelity in these in vitro systems. In contrast, the partially purified rat liver synthetase utilizes all the naturally occurring folates at physiological concentrations although only a few are highly efficient. With a t least one substrate, H,PteGlu, polyglutamates up to the length found in uiuo, pentaglutamate, can be synthesized. Further studies on these two classes of synthetase wiU be instructive as to whether the difference between the properties of the bacterial and mammalian enzymes represent simple species diversity or reflect different physiological roles in prokaryotes and eukaryotes either for the enzyme or its polyglutamate products.

The lack of specificity of the rat liver synthetase for the pteridine moiety contrasts with the stringent specificity for both the terminal glutamate and amino acid to be added. This lack of specificity is not surprising if formation of polyglutamates is essential for cellular retention of folates. Two folates utilized by the rat liver synthetase deserve particular mention. Folic acid had been previously suggested to be inactive with the CHO synthetase (17). This may be in error, however, because folic acid was tested in the absence of a reducing agent, which we have shown is essential for activity of the rat liver synthetase. Several studies, (reviewed in Ref. 47) have demonstrated that ~ - C H : Y H ~ P ~ ~ G ~ U is not converted to polyglutamates in uiuo. This is not inconsistent with our data showing that 5-CHa-H,PteGlu is a substrate because at physiological concentrations its rate of reaction is so slow that its conversion might not be detectable in uiuo.

The substrate activity of the (d)-isomer with the rat liver synthetase was a surprising finding. A few examples are known where an unnatural folate isomer or its polyglutamate form inhibits a folate-dependent reaction (11, 48). However, for all folate enzymes where specificity has been determined, only the naturally occurring isomer is catalytically active. The only exception to this strict specificity is in transport of reduced folates as first described in bacterial (49) and then in mammalian systems (50). In both cases, transport of the two isomers of 5-CH:,-H4PteGlu occurs with similar kinetic con-


stants. The folylpolyglutamate synthetase of L. casei has also been postulated to lack stereospecificity on the basis of indi- rect, in vivo evidence (43), but direct studies on the E , coli enzyme (16) demonstrated that only (Z)-10-HCO-H4PteGlu was active and that the (d)-isomer was slightly inhibitory.

White et al. (50) have stressed that the lack of stereospecificity in transport is important in the current use of high dose methotrexate (anti-folate) chemotherapy followed by reversal of methotrexate toxicity with racemic 5-HCO-H4PteG1u. If (d)-5-HCO-H,PteGlu is transported similarly to (d)-5-CHa- H,PteGlu, its effect on cellular 1-carbon metabolism as well as on methotrexate transport could alter the effectiveness of these protocols. An additional complication is now raised by the knowledge that (d)-H,PteGlu, and probably other (d ) - folates, are substrates for polyglutamylation. In light of these data, the effects on metabolism of both (d)-folates and their polyglutamyl derivatives must now be considered.

Analysis of the products synthesized throughout a 10-h incubation at 5 and 35 WM tetrahydrofolate (Figs. 2 and 3) leads to several conclusions. The lengths of the products are inversely dependent on the initial monoglutamate concentration. High H,PteGlu concentrations lead to synthesis of shorter polyglutamates, principally di- and triglutamates. Low HJPteGlu concentrations favor extensive formation of higher polyglutamates, including pentaglutamates, and they are formed at higher absolute levels even though total product formation is less than at high substrate. A similar concentration dependence has been observed at a single time point with the CHO enzyme (17). This apparent regulatory effect on the synthetase may be responsible for the observation in vivo (37, 38) that law extracellular concentrations of folic acid lead to accumulation of longer polyglutamates in cells while high concentrations lead to shorter polyglutamates. In terms of metabolic efficiency, this is understandable because monoglutamates and short polyglutamates do serve in all mammalian folate reactions, although generally less well than longer polyglutamates. Thus, under conditions of folate excess, formation of long polyglutamates would be wasteful of both ATP and glutamate. In folate-depleted conditions, not only does polyglutamate formation increase retention of folates once transported, but each folate molecule is a more efficient carrier of 1-carbon units.

Rat liver folylpolyglutamate synthetase synthesizes polyglutamates of H,PteClu as long as pentaglutamate in vitro (Fig. 4), the same length which predominates in rat liver in vivo ( 6 4 , and thus can faithfully reproduce synthesis as it occurs in uivo. Synthesis of hexaglutamate and higher derivatives is, at best, extremely slow in uitro, a result of the poor substrate activity, displayed by H4PteGlus (Table V). The pentaglutamate does bind to the synthetase, however, because it inhibits synthesis of polyglutamates from the monoglutamate. These observations with the pentaglutamate derivative may be especially significant in several respects. Since the pentaglutamate binds to the synthetase but is not catalytically active, the data suggest that its length places the free y- carboxyl out of the active site. This, in turn, suggests that there are absolute dimensions for the enzyme active site and that the predominant length of polyglut,amate in rat liver may be determined solely by the active site dimensions and is otherwise unregulated. Inhibition of the polyglutamylation reaction by H4PteGlu5 may be physiologically significant in the rat. This inhibition by the ultimate product of the reaction would offer a means by which folylpolyglutamates could regulate their own synthesis.

Derivatives as long as hexa- and heptaglutamate have been reported by Leslie and Baugh (8) in rat liver following ex- tended periods (28 days) of in vivo labeling. These authors

recently showed their methodology to be inadequate (51) and thus these lengths may be erroneous. If the lengths are correct, it may demonstrate that over long time periods even extremely poor substrates are gradually utilized, or that another enzyme may be involved in the synthesis of longer chain polyglutamates.

We have chosen to approach the problem of folylpolyglutamate function by fust examining the enzyme responsible for its synthesis. The data presented here establish that the preparation, although not homogeneous, can reproduce synthesis as it occurs in uiuo and provide information about possible means of regulating polyglutamate synthesis. In further studies, we hope to explore these regulatory mechanisms more fully and determine the metabolic consequences of this regulation.

Acknowledgments-We wish to thank Ms. Linda D’Ari and Mr. Robert Dreyer for helpful discussions and Mrs. Patricia Kerley for patiently typing the manuscript.

1.

2.

3.

4.

5.

6.

7.

8. 9.

10.

11.

12.

13. 14.

15.

16.

17.

18.

19.

20.

21.

22. 23.

24.

25. 26. 27.

28.

29.

REFERENCES Blakley, R. L. (1968) in Frontiers of Biology (Neuberger, A,, and

Baugh, C. M., and Krumdieck, C. L. (1971) Ann. N . Y. Acad. Sei.

Powers, S. G., and Snell, E. E. (1976) J. Biol. Chem. 251, 3786-

Pfiffner, J. J., Calkins, D. G., Bloom, E. S., and ODell, B. L.

Bassett, R., Weir, R. J., and Scott, J . (1976) J. Gen. Microhiol.

Shin, Y. S., Williams, M. A,, and Stokstad, E. L. R. (1972)

Houlihan, C. M., and Scott, J . M. (1972) Biochem. Biophys. Res.

Leslie, G. I., and Baugh, C. M. (1974) Biochemistry 13,4957-4961 Coward, J . K., Parameswaran, K. N., Cashmore, A. R., and

Baggott, J. E., and Krumdieck, C. L. (1979) Biochemistry 18,

Kisliuk, R. L., Gaumont, Y., and Baugh, C. M. (1974) J . Bid. Chem. 249,4100-4103

Rabinowitz, J. C. (1960) in The Enzymes (Boyer, P. D., Lardy, H., and Myrback, K., eds) 2nd Ed, Vol. 2, pp. 185-252, Academic Press, New York

Tatum, E. L., eds) Vol. 13, North-Holland, Amsterdam

186, 7-28

3793

(1946) J . Am. Chem. Soc. 68, 1392

93, 169-172

Biochem. Biophys. Res. Commun. 47,35-43

Commum. 48, 1675-1681

Bertino, J. R. (1974) Biochemistry 13, 3899-3903

1036-1041

McBurney, M. W., and Whitmore, G. F. (1974) Cell 2, 173-182 Griffin, M. J., and Brown, G. M. (1964) J. Biol. Chem. 239, 310-

Gawthorne, J . M., and Smith, R. M. (1973) Biochem. J. 136,295-

Masurekar, M., and Brown, G . M. (1975) Biochemistry 14,2424-

Taylor, R. T., and Hanna, M. L. (1977) Arch. Biochem. Biophys.

McGuire, J. J., and Rabinowitz, J. C. (1978) J. Biol. Chem. 253,

Leichter, J., Butterworth, C . E., Jr., and Krumdieck, C. L. (1977)

Krumdieck, C. L., and Baugh, C. M. (1970) Anal. Biochem. 35,

Rabinowitz, J . C., and Pricer, W. E., Jr. (1962) J. Biol. Chem.

Blakley, R. L. (1957) Biochem. J . 65, 331-342 Curthoys, N. P., and Rabinowitz, J . C. (1971) J. Biol. Chem. 246,

Kallen, R. G., and Jencks, W. P. (1966) J. Biol. Chem. 241,5851-

Blakley, R. L. (1960) Nature (Lond.) 188, 231-232 Rabinowitz, J . C. (1963) Methods Enzymol. 6,814-815 Curthoys, N. P. (1971) Ph.D. thesis, University of California,

Himes, It. H., and Harmony, J . A. K. (1973) CRC Crit. Reu.

Curthoys, N. P., Scott, J. M., and Rabinowitz, J. C. (1972) J . Biol.

3 I 6

30 1

2430

181,331-344

1079-1085

Proc. Sac. Exp. Biol. Med. 154, 98-101

123-129

237,2898-2902

6942-6952

5863

Berkeley, CA

Biochem. 1, 501-535

Chpm. 247, 1959-1964


30. Blair, J. A,, and Saunders, K. J. (1970) Anal. Biochem. 34, 376-

31. Parker, D. J., Wu, T.-F., and Wood, H. G. (1971) J. Bacteriol.

32. Cashmore, A. R., Dreyer, R. N., Horvath, C., Coward, J. K., Knipe, J. 0.. and Bertino, J. R. (1980) Methods Enzymol., 66,

33. Tierneier, D. C., and Milman, G . (1972) J. B i d . Chem. 247,2272- 2277

34. Coward, J. K., Chello, P. L., Cashmore, A. R., Parameswaran, K. N., DeAngelis, L. M., and Bertino, J. R. (1975) Biochemistry

35. Weiner, A. M., Platt, T., and Weber, K. (1972) J. B i d . Chem. 247,3242-3251

36. Randerath, K., and Randerath, E. (1971) in Procedures in Nucleic Acid Research (Cantoni, G. L., and Davies, D. R., eds) Val. 2, pp. 796-812, Harper & Row, New York

37. Bassett, R., Weir, D., and Scott, J. (1976) Biochem. Soc. Trans.

38. Buehring, K. U., Tamura, T., and Stokstad, E. L. R. (1974) J.

39. Andrews, P. (1965) Biochem. J. 96, 595-606 40. Shane, B., Brody, T., and Stokstad, E. L. R. (1979) in Chemistry

and Biology of Pteridines (Kisliuk, €3. L., and Brown, G. M., eds) pp. 341-346, Elsevier/North Holland Publishing Co. New York

38 1

108,770-776

459-468

14, 1548-1552

4, 500-502

B i d . Chem. 249, 1081-1089

41. Evans, H. J., and Sorger, G. J. (1966) Annu. Rev. Plant Physiol.

42. Bertino, J. R., Sawicki, W. L., Moroson, B. A., Cashmore, A. H., and Elslager, E. F. (1979) Biochem. Pharmacol. 28, 1983-1987

43. Shane, B., and Stokstad, E. L. R. (1977) J . Gen. Microbiol. 103, 249-259

44. Acharya, S. P., and Hynes, J. B. (1975) J. Heterocycl. Chem. 12, 1283-1286

45. Dixon, M., and Webb, E. C. (1958) Enzymes, pp. 91-94, Academic Press, New York

46. Neuhoff, V. (1973) in Micromethods in Molecular Biology (Neu-

Y ork hoff, V., ed) pp. 85-147, Springer-Verlag New York, Inc., New

17,47-76

47. Hoffbrand, A. V. (1975) Prug. Hematol. 9, 85-105 48. Scott, V. F., and Donaldson, K. 0. (1964) Biochem. Biophys. Res.

Cummun. 14,523-526 49. Shane, B., and Stokstad, E. L. R. (1976) J . Biol. Chem. 251,3405-

34 10 50. White, J. C., Bailey, B. D., and Goldman, I. D. (1978) J . Biol.

Chem. 253,242-245 51. Baugh, C. M., May, L., Braverrnan, E.. and Nair, M. G . (1979) in

Chemistry and Biology of Pteridines (Kisliuk, H. L., and Brown, G. M., eds) pp. 219-224. Elsevier/North Holland Puh- lishing Co., New York

52. Dolnick, B. J., and Cheng, Y.-C. (1978) J. Bml. Chern. 253, :3563- 3567

5786 Rat Liver Folylpolyglutamate Synthetase SUPPLEMEIITARY MATERIAL PARTIAL PURIFICATION AND CHARACTERlZATlUN OF FOLYLP0LYGLUTA'I?TE

SYNTHETASE FROM RAT LIVER

John J . M C G U I P B , Pearl Hsleh, James K Coward and Joseph R . B e r t l n o

P a r t l a 1 P u r l f l c a t l o n o f F a l y l p o l y l L u t a m a t e S y n t h e t a s e f r o m R a t Liver

o f amnonlurn s u l f a t e 1 s g lven a s t h e p e r c e n t a g e o f s a t u r a t i o n a t 0' ( l ! ~ The pH was not ad- All operd t l ons were c a r r l e d o u t a t 0-4" unless o t h e r w r e Indicated. The Concentration

J u s t e d a f t e r d d d l t l o n s o f dmnonium S u l f a t e .

Livers were r a p i d r y vemoved, c h l l l e d , and mlnced. Mlnced l i v e r (81 g! was mixed w t h 3 Crude E x t r a c t . Male Spraque-Dawley r a t s ( 7 0 0 - 2 5 0 q l were I a C r l f l C e d by d e c d p l t a t l m

voIumes o f 0 .1 M TPII-CI pH 8.4:U.l M 2-mercaptoethanol and hornoqenlzed by 2 b u r l t l ( l 5 s ! from a Polytron (Bnnkman) homogemier The homogenate was c e n t n f u g e d a t 31,000 x 9 f o r 1 h and decan ted t h rough th ree l aye rs o f cheesec lo th .

Phosphmel lu la re Chromato r a h . The poo led superna tan ts (122 mll *ere d l l u t e d w l t h 74 ml of Buffer A and loaded on% E ~ h o r p h o c e l l u l a r e column (4x48 Cm! p r e - e q u l l l b r a t e d w i t h B u f f e r A. The column was washed w l t h B u f f e r A u n t i l t h e absorbance a t 280 nm w I E l e s s t han 1 and then 11 was e l u t e d w l t h B u f f e r A conta$nlng 400 nM KC1 (Figure I S ) . The s y n t h e t a s e e l u t e d 1" t he 400 mM KC1 wash as a sharp peak wh,ch r l l g h t l y t r a l l e d t h e m a l " protein peak. The large p w t e l n peak wh lch e lu ted w t h Buf fer A conta ined no a c t i v l t y when a s s a y e d a f t e r pool-

d d d l t l o n o f 3 M KCI. '"9 and concentration The KC1 c a n c e n t r a t l o n o f a l l f r a c t l o n s was r a i s e d t o 500 mM by

Frocflon Number

c o l l e c t e d . A C t l Y l t y war d e t e r m n e d ai ~n F l g u k 1s.

Table Is. E f fec t o f Te rnpera twe on Rat L iver Fo ly lpo lyg lu tamate Synthe tase,

Temperature Net t3HlGlutamate Incorporated

I O C l lpmol 1 22

20 610

37 1700 4 1 . 5 1740 50 100

Standard assay Condl t lons e r e used except t h a t t h e pH o f t h e T r l s - C l b u f f e r was v a r i e d S O

o u t f o r 3 h a t t h e i n d i c a t e d temperature. Va lue r pwsen ted are t he average Of duplicate t h a t a t t h e 6 p e c l f l e d t e m p e r a t u r e t h e r e a c t I a n m l x t u l e was pH 8.3. I n c u b a t l o n i Were Car r ied

de te rm ina t ions .

on t he InCOTpOPatlm o f C 3 H J g l u t a m a t e a t a f i x e d l o c u b a t i o n l n t e w a l (3h) and Constant pH

E f f e c t o f Temperature on Fo ly lpo l yg lu tamate S y n t h e t a s e A c t > v > t l . The e f f e c t O f temperature

was determined (Tab le IS). l n c o r p a r a t i a n was h i g h e s t a t 370 and 41 5' A c t l v l t y d e c r e a s e d d r a m a t 7 c a l l y a t lower ( Z P ! or h i g h e r (SO'! temperatures. Only s l l q h t d c t l v l t y was d e t e c t - a b l e a t 0'.

w. Effect o f pH on Foly lpo lyq lutamdte 5ynthetare Act lv l ty . Standard assay cand" t l o n s w e ~ e used except tha t bu f fe rs (100 mM f l n d l c o n c e n t r a t l a n ) o f d i f f e r e n t pH played. The pH I n d i c a t e d i s t h a t o f t he reaction rn lx tu re measured a t 37". (el T F ? s - H - ~ : D~ethmolarn!ne-I iCl; (m) EthdnOlm$ne-IIC1.

246 . . , 4


55-

N

B - 0

E

' L - 2 0

u

O I O 50 90

[ KCI ] mM

2 0

0 8

0 4

' 0 01 0 2 0 3 0 4 0 5 I O

[ATP] m M


O r h e r n u c l e o t l d e i vere t e s t e d f o r t h e l r d b l l l t y t o replace ATP !Table 25). Assays us ing the HPLC demonst ra ted tha t no r p e r l f i c or non- rpec l f lc n u c l e o t l d d i e was presen t . UTP and TTP were essentially I n a c t i v e . GTP and dGTP were l e i 5 a c t i v e t h a n ATP. dATP was s l l g h t l y m o w active than ATP a t t h l s c a n c e n t r a t i o n (in" and a l s o when b o t h wele t e s t e d a t 0.5 mM ( d a t a n o t shown), and t h e r e f o r e t h e \ f o r dATP may d l 3 0 be low.

S u b s t r a t e A c t l v l t x o f M e t h o t r e x a t e . The f o l a t e analog me tho t rexa te (2,4-d~amln(1-10-nlethyl-

h t 50 "M M I X t h e l n c o r p o m t l o n was h i g h e r t h a n t h e maximum ach7evab le w i th ($l)-HIPteGlu. folic ac la ) was 3 good s u b s t r a t e f o r t h e r a t liver f o l y l p o l y g l u t a m a t e s y n t h e t a s e (Flgure 125).

NuCleot lde Net 13H1Glutamate Incorporated

lprnol)

dATP ATP

dGTP GTP

UTP TTP

IS10 1860 710 1160 00 0 2ovi 10

0 0 2 0 40

i ncorpora ted r3Hlq lu tamate and are f a r Of f scale

ANALYSIS OF PRODUCTS

Enzymatic Synthesis of Folylpolyglutamates · 2001-09-05 · Enzymatic Synthesis of...

Documents

Transcript of Enzymatic Synthesis of Folylpolyglutamates · 2001-09-05 · Enzymatic Synthesis of...