(Received for publication, February 27, 1978, and in revised form, January 30, 1980) or galactose, glycolytic activity decreases dramatically, by as

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1 THE JOURNAL OF BOLO(i1CAL CHEMSTRY Vd No. 12. ssue of dune 25. pp , 1980 Prmted m SA. The Pentose Cycle CONTROL AND ESSENTAL FUNCTON N HeLa CELL NUCLEC ACD SYNTHESS* (Received for publication, February 27, 1978, and in revised form, January 30, 1980) Lawrence J. Reitzer,$ Burton M. Wice, and David Kennel1 From the Department of Microbiology and immunology, Division of Biology and Biomedical Sciences, Washington University School of Medicine, St. Louis, Missouri The pentose cycle in HeLa cells, growing on different concentrations of glucose, galactose, or fructose, has been studied with respecto production of NADPH and of pentose-p for nucleic acid synthesis. The flow of hexose carbon through the oxidative arm of the cycle was calculated by measuring the release of %02 from [l- 4C]hexose; complications from randomization and recycling of carbon were shown to be insignificant. This flux was decreased only 2-fold per mass synthesized during growth on 2 m~ fructose, compared to that on 10 mm glucose, even though total hexose was depleted 50X slower. We investigated the basis for control of this flux and observed that the reaction velocities of both dehydro- genases of the oxidative arm are severely limited at the maximum effective cytoplasmic NADP+ concentration and effective NADPH/NADP ratio; however, the effective coenzyme levels did not change significantly during growth on different hexoses. n contrast, the concentrations of the substrates, glucose-6-p and 6-P- function of the pentose cycle in HeLa cells is to generate gluconate, did change severalfold and when introduced ribose-5-p for nucleic acids. into the appropriate rate equations for the pentose two cycle dehydrogenases, could account for the observed changes in the oxidative arm flux. To identify the essential function of the pentose cycle, Cultured HeLa cells metabolize more moles of glucose than the oxidative arm synthesis of ribose-5-p was compared any other organic molecule in the medium. After cell entry, to the rate of nucleic acid (pentose moieties) synthesis. the first glucose intermediate is glucose-6-p, a substrate for The latter was calculated after measuring simultane- glycolysis, glycogen formation, the pentose phosphate cycle, ously the specific activities of cellular UTP at early nucleotide sugar products, and amino sugars. About 80% of times of rh]uridine incorporation and the rate of the glucose carbon is converted into lactic acid via glycolysis counts per min into UMP of RNA. Total RNA synthesis (1-4). A very small fraction ( ~3%) is polymerized to glycogen, was 3 to 4 times net synthesis as a result of the insta- while 7 to 10% of the l-4c is released as Cog. On lower bility of most synthesized RNA; on 10 m~ glucose, total concentrations of sugar or when glucose is replaced by fructose nucleic acid ribose synthesis was about 55% the rate of or galactose, glycolytic activity decreases dramatically, by as ribose-5-p synthesis via the oxidative arm. The actual much as 900-fold in a transition from 10 mm glucose to requirement for ribose-5-p for nucleic acid compounds 2 mm fructose, with only a 2-fold decrease in the exponential growth is a function of the extent of reutilization of the ribose from RNA breakdown. A very large fraction of ribose rate of the cells; most energy (>99% in the latter case) is was found in small nucleotide compounds of the acidsoluble fraction. These compounds could include RNA degradation products, and if the remaining products of decay are reutilized, then on 10 m~ glucose about 34% of the ribose-5-p synthesized by the oxidative arm reactions must be drawn directly into nucleic acids. A small fraction enters protein and the remainder is di- * This research was supported by a special grant for research (BC- 214) from the American Cancer Society, Missouri Division. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked aduertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. $ Some of this work was used to meet partial requirements for the Ph.D. degree from Washington University (L. J. R.). Present address, Department of Biology, Massachusetts nstitute of Technology, Cambridge, Mass. verted through the nonoxidative arm to enter the major flux of sugar carbon to lactate. These conclusions were tested independently by following the fate of exogenous [3H]ribose. On 10 m glucose, about 23% was incorporated into nucleic acids, on 10 mm fructose about 65%, and on 2 mm fructose at least 85%. Thus, in some cases, almost all of the ribose- 5-P, synthesized by the oxidative arm, is utilized for nucleic acid synthesis, and the cellular growth rate can become proportional to the synthetic rate of ribose-5- P. t seems likely that NADPH production is not an essential function of the oxidative arm in HeLa cells. Two different kinds of experiments demonstrated that very little of the synthesized ribose-5-p recycles around the pentose cycle. Without this recycling, the estimated flux rates indicate >80% of the cellular NADPH can be generated by the reaction of malate to pyruvate from glutamine metabolism. Probably the only essential provided by glutamine oxidation (1). n contrast, 14C0, generation from the [l- 4C]hexose varies much less. f this latter parameter is a valid indicator of flux through the oxidative arm of the pentose cycle, then the flowof carbon through these reactions remains relatively invariant while glycolytic activity can practically vanish, suggesting an essential role of this cycle in cell growth. The kinetic basis for this relatively constant flow is investigated here. The function of the pentose cycle in animal cell metabolism has been attributed to two seemingly very dissimilar parts of 5616 metabolism: the regeneration of NADPH foreductive syntheses (5-10), especially that of fatty acid, and for the synthesis of ribose phosphate as a precursor for nucleic acids. From approximate estimations, earlier studies have concluded that the flux through the pentose cycle is in great excess over the need for nucleic acid ribose synthesis. To assess this

2 and Cycle Pentose Nucleic Synthesis Acid 5617 function more accurately, the rate of nucleic acid synthesis was measured in this paper and shown to be much more than is its rate of accumulation. Also, by following the fate of exogenous ribose as a marker, it was possible to correlate the carbon flux through the oxidative arm of the pentose cycle with the flow of pentose phosphate into nucleic acids. Except in the case of high glucose, almost all ribose-5-p derived from the oxidative arm is used in nucleic acid synthesis and in these cases the rate of growth is probably limited by this flux. EXPERMENTAL PROCEDURES Cells and Growth Conditions Cells were maintained as monolayers in 75-cm' T-flasks incubated at 37 C in a Wedco C02 incubator with a 5% Cor-balance air, 100% humidity atmosphere. HeLa strain M cells were grown in T-flasks, as described (l), in Joklik modified minimum essential medium containing antibiotics but lacking NaHCO:, glutamine, and glucose (Kansas City Biological) and supplemented with glutamine to give 4 mm final concentration, plus NaHCO:] (2.5 g/liter) plus 10% (in recent experiments 5%) fetal bovine serum, dialyzed to give <200 PM glucose in the serum (Gibco). Mouse strain L, clone 929, cells were grown under the same conditions except that the medium was Eagle's minimum essential medium minus glucose plus Earle's balanced salt solution for monolayer growth and 10% dialyzed fetal calf serum. Sugar was added as indicated. HeLa cell doubling times in T-flasks were 28 h with 10 mm glucose, 37 h with 10 mm fructose or galactose (1). and 54 h with 2 mm fructose. Enzyme Assays Cell pellets were washed in phosphate-buffered saline (NaC, 0.14 M; KC, 2.7 m ~ Na,HP04, ; 8.1 mm; and KHJ'O,, 1.5 mm) (if stored, they were maintained at -70 C) and extracts prepared by resuspending in 50 mm Tris-HC, ph 7.5, and freeze-thawing twice. Phosphofructokinase (EC ) gave no activity unless the cells were resuspended in the phosphate dilution buffer of Lowry and Passonneau (11) (0.1 M phosphate with K2HP04/NaH2P04 of 4:1, 1 mm EDTA, plus 0.25 m~ AMP). About 50% of the hexokinase (EC ) activity was lost in 4 months at -7O"C, while no loss of phosphofructokinase activity was detected. The other enzymes were assayed from fresh homogenates. Protein contents of homogenates were determined by absorbance at 260 nm and conversion to micrograms of protein from a standard curve of absorbance to protein determined directly by the Fohn reagent (12). An absorbance of 1.0 corresponded to 168 pg of protein/ml in these lysates. The following enzyme assays were performed on a spectrophotometer at 22'C and a wavelength of 340 nm in 1.0 ml with a 1-cm light path. Following the example of Wu and Racker (13), substrate concentrations were saturating and reactions were buffered at ph 7.5 with 50 mm Tris-HC1. Hexokinase-The assay was described by Wu and Racker (13). Phosphofructokinase-Phosphofructokinase was also assayed by the procedure of Wu and Racker (13), except that 2 mm Na,SO, (11) plus 0.05% bovine serum albumin were present to protect the labile activity. Transaldolase-Transaldolase (EC ) was assayed by coupling the formation of glyceraldehyde 3-phosphate to dihydroxyacetone phosphate and then glycerol phosphate with triose phosphate isomerase (present in excess in these extracts) and glycerol-3-phosphate dehydrogenase and following the decrease in NADH in the last reaction. The reaction contained 50 mm Tris (ph 7.5). 2 mm fructose &phosphate, 0.15 m~ NADH, 0.2 mm erythrose 4-phosphate, 20 mm EDTA, ph 7.5, excess coupling enzymes, and lysate. The control with no fructose-6-p gave up to 20% of some observed velocities. This may result from impurities in the commercial erythrose-4-p (85% pure). Omission of erythrose-4-p or lysate gave no activity. The assay mix contained no Mgz+ (and EDTA was added), or ATP, or P, during the freeze-thaw step which eliminated phosphofructokinase or transketolase as activities for triose phosphate formation. Transketolase-Transketoiase (EC ) was assayed by the same coupled reactions used in the transaldolase assay except ribose 5-phosphate and xylulose 5-phosphate were the initial substrates. The reaction included 50 mm Tris (ph 7.5), 5 mm ribose 5-phosphate, 0.2 mm xylulose 5-phosphate, 2 m MgC2, 0.005% thiamin pyrophosphate, and 0.1 mm NADH. The X, for xylulose 5-phosphate with 5 mm ribose 5-phosphate is 0.2 m~ in yeast (14); hence, the observed velocities were about!h V,,,,. The following enzyme assays were conducted at 22 C in 1.0 ml in a Farrand fluorometer A4 using a 7-60 excitation filter and 5-57 plus 3-73 filter for emission. ncreased fluorescence was quantitated by comparison with known standards. Glucose-6-phosphate Dehydrogenase (EC )"The reaction mix contained 50 mm Tris-HC, ph 7.4,5 m~ MgC?, 0.1 mm NADP', 1 mm glucose-6-p, and to 0.02 mg of lysate protein. 6-Phosphogluconate Dehydrogenase (EC.l.1.44)"The assay contained 50 mm Tris-HC, ph 7.4, 0.1 mm NADP', 1 mm 6-Pgluconate, 5 mm MgC12, 0.05% bovine serum albumin, 1 m~ dithioerythritol, plus to 0.08 mg of lysate protein. Cell ntermediates The rapid preparation of perchloric acid cell extracts from a T- flask has been described (1). n all cases the cels had been growing for several generations on the particular sugar at the time of harvest. The assays for cell intermediates were taken directly or with indicated modifications from the text of Lowry and Passonneau (15) which should be consulted for further details and original references. All assays relied upon the generation of oxidized or reduced pyridine nucleotide in a coupled enzyme reaction monitored fluorometrically after amplification of the pyridine nucleotide product by enzymatic cycling (1, 15). Enzymes were from Boehringer Mannheim. Sedoheptulose-7-P was measured by adding 66 p~ glyceraldehyde- 3-P and transaldolase (EC ), 0.2 unit/ml, to the fructose-6-p assay described (15). Transaldolase was centrifuged to remove most of the (NH,),SO,. No other reaction of the pentose cycle will give fructose-6-p since no transketolase (EC ) activity could be detected in the transaldolase preparation. Fructose can and did react with transaldolase to form a complex and generate fructose-6-p by subsequent reaction with glyceraldehyde-3-p; sedoheptulose-7-p could not he measured in fructose-grown cells. Erythrose-4-phosphate was also measured in the fructose-6-p assay (15) with the following additions: 5 mm MgCL, thiamin pyrophosphate, 60 PM xylulose-5-p, and 0.1 unit/ml of transketolase (Sigma). The two-step reaction was modified to remove both glucose- 6-P and fructose-6-p in the fust step. The pentose donates the 2- carbon fragment to form a transketolase.glycolaldehyde complex (16) and erythrose-4-p is the only aldehyde that will produce fructose-6-p from this complex. Our xylulose-5-p contained about 1% ribose-5-p which would generate sedoheptulose-7-p instead of fructose-6-p in the same reaction. Fortunately, our transketolase preparation con- tained no detectable transaldolase activity which would have then formed fructose-6-p as in the sedoheptulose-7-p assay. Transketolase was suspended in 3 M (NHs),S04 and stored at 4 C and centrifuged to remove (NH&SO, before use. Xylulose-5-P was assayed by measuring the formation of glyceraldehyde-3-p by method 1 of Kauffman et al. (17) with addition of ribose-5-p plus transketolase. The glyceraldehyde-3-p formed was followed with glyceraldehyde-p-dehydrogenase (EC ) added to produce NADH. The sedoheptulose-7-p generated did not interfere if high concentrations (50 p ~ of ) ribose-5-p were present. Since transaldolase was not detectable in the transketolase preparations, this should be the only route for glyceraldehyde-3-p formation. NADP+ and NADPH concentrations deserve special comment. t is necessary to distinguish between the total bound and unbound cofactors, as well as to bear in mind that they may be partitioned between cytoplasmic and mitochondrial compartments. For the pur- poses of this paper, our primary concerns are fust, the effective NADPH/NADP+ ratio that modulates the activities of both pentose cycle dehydrogenases and second, the effective concentration of cytoplasmic NADP' that is a substrate in both reactions. Using the approach of Krebs (18). the former value is estimated from the steady state concentrations of the other reactants in two NADPH/NADP+ cytoplasmic reactions that are near equilibrium: the isocitrate dehydrogenase and malic enzyme reactions. The former gives a ratio close to 20 on al sugars in spite of changing reactant concentrations ("Results"), i.e. the reaction is in equilibrium. The malic enzyme reaction gives a value close to this when glucose is the supporting sugar. However, on fructose, galactose, or during sugar starvation, the ratio increases to 100 or more. Krebs and Veech (18) observed similar results in liver from normal versus starved rats. These authors rea- soned that starvation for carbohydrate decreased the pool of pyruvate to create a disequilibrium in the malic reaction of malate u pfvate + CO,. Such a situation would also be expected in fructose- or galactose-grown HeLa cells in which glycolytic activity, and from it pyruvate, are greatly reduced (1). Presumably, the levels of reactants

3 56 18 Pentose Cycle and Nucleic Acid Synthesis in the isocitrate dehydrogenase reaction are maintained largely from the metabolism of glutamine which is relatively constitutive on these different sugars (1). The concentration of NADP is derived from the total cell NADPH measured, after destruction of NADP (l), by enzyme cycling (15) and the NADPH/NADP ratio mentioned above. The total NADPH includes any fraction that is bound as well as mitochondrial NADPH. Deriving the effective cytoplasmic NADP concentration from the total NADPH would give a maximal possible concentration for the free NADP. f the actual unbound concentration of NADPH in the cytoplasm were somewhat lower, the effective NADP concentration would be lower proportionally. However, this would not change any conclusions in this paper ( Discussion ). Measurement of the Rate of Nucleic Acid Synthesis micromoles of UMP incorporated into micromoles of RNA ribose incorporated. The pools of nucleoside triphosphates are provided by de novo Net RNA synthesis per mg of protein was calculated directly from synthesis from hexose and other carbon compounds in the medium, the accumulation rate of RNA (dr/dt) which equals kr where k = from reutilization of degradation products from RNA turnover, and n 2/doubling time and from the RNA/protein ratio of The base from any exogenous labeled compound, e.g. uridine, that may be used ratios of ribosomal RNA (23) were used to convert milligrams of as a marker. For this reason, it is necessary to measure the specific RNA/min/mg of protein into micromoles of ribose/rnin/mg of proactivity of one of the pool compounds in order to translate counts per tein. rnin incorporated into micromoles of nucleic acid polymerized, an n both cases, the rate of DNA synthesis was added to the rate of approach used successfully with bacteria years ago (e.g. Ref. 19). t is also necessary to make this measurement at early times of exposure to a labeled precursor. At later times, when a significant fraction of the unstable RNA has been labeled, the net rate of counts per min RNA synthesis, taking an RNA/DNA ratio of 1.5 (from HeLa DNA per cell (24)) and assuming most DNA synthesized is relatively stable. The Fate of Exogenous r3hjribose incorporated is less than the total rate of synthesis by an amount equal to the rate of degradation (whether or not the products are reutilized). Several 25-cm T-flasks were inoculated with HeLa cells. A day later, the medium in each flask was replaced with 2 ml (rather than 5 ml) of fresh medium and the flask incubated another 3 h before addition of [5,6- H]uridine (to 250 pci/ml and 6.3 nmol/ml). At a specified time, the medium was removed from a given T-flask by aspiration and without washing the cells covered with 0.3 N perchloric acid at 0 C. A rubber policeman was used to remove cell remains from the bottom and the flask contents and a wash were centrifuged at 15,000 X g for 15 min. The supernatant was applied to a 0.5-ml Norit-Celite column in a 5-ml syringe barrel, pre-equilibrated with 0.3 N perchloric acid. The column was washed with about 10 ml of 0.3 N perchloric acid to remove non-nucleotide compounds. After washing with 15 ml of water to remove perchloric acid, the nucleotides were eluted in ethanol: NH,OH (50%:28). After evaporation to dryness, the residue was resuspended in 0.5 ml of water and 5 p applied to a polyethyleneimine cellulose sheet (EM Laboratories, nc., Elmsford, N. Y.) and chromatographed in an ascending system of 2 N formic acid:2 M LiCl (1:l) (20). [5-, H]UTP was added to a control-extracted sample (no [ Hluri- dine) and chromatographed simultaneously in another lane to define acid have been described (1). All chemicals were reagent grade. the position of UTP (RF = 0.2). The thin layer chromatography sheet Radioactive compounds were purchased from New England Nuclear was dipped in 2-methylnaphthalene with 0.4% 2,5-diphenyloxazole Corp. ([5,6-: H]uridine, 40 Ci/mmol; and [5- Hluridine 5 triphos- (PPO) (21) and placed inside a cassette to expose Kodak XR-5 film phate, 20 Ci/mmol) or Amersham/Searle ([l- 4C]fructose. 57 mci/ at -80 C and developed about 24 h later. The UTP spot was cut out mmol; [l- C]glucose, 60 mci/mmol; and [5- H]ribose, 9.6 Ci/mmol). and counted. The micromoles of UTP were determined fluorometrically from an aliquot of the same sample that was chromatographed. The UTP was RESULTS reacted with an excess of glucose--p and UDP-Glu pyrophosphoryl- Problems in Pentose Cycle Flux Measurement-The 1- ase (EC ). The resulting UDP-Glu generated was reacted carbon of an exogenous hexose is released as C02 when it is further with NAD and UDP-Glu dehydrogenase (EC ) to NADH and UDP glucuronate. The reaction (in 30 pl) contained Trisoxidized in the reactions of glucose-6-p to pentose-p (Fig. 1). HC, 50 mm; MgC, 2 mm; NAD, 0.1 mm; UDP-Glu dehydrogenase, Since we wished to compare the flow of carbon through the 0.27 milliunit; glucose-1-p, 0.25 m~; extract, 5 pl; and UDP-Glu oxidative arm to biosynthetic processes, we considered the pyrophosphorylase, 15 milliunits, which was omitted in duplicate possible inaccuracies, discussed earlier (9, 10, 25), in using 1- reactions to measure the amount of UDP-Glu in the extracts (about 14 C evolution for its measurement. First, the only other reac- 20% of the UTP value). tions that would release the 1-carbon as COn would be those The perchloric acid precipitate was washed in 0.3 N perchloric acid of the citrate cycle during the second round of cycling when and then resuspended in 0.3 N KOH and incubated at 23 C for 15 to 20 h to hydrolyze the RNA to mononucleotides. Perchloric acid (6 both the 1- and 6-carbons are first released. Since 4CO2 from N) was added to give neutrality and the K perchlorate precipitate [l- 4C]glucose is >20x that released from [6-4C]glucose(l), removed by centrifugation. The nucleotides were separated on poly- it can be concluded that >95% of the 4CO2 from [ l- 4C]glucose ethyleneinhe cellulose layers using 0.75% formic acid followed, after drying, by 0.25 M Li formate, ph 3, in the same direction (modified from Ref. 22), running each solvent front for 13 cm. The UMP spot was counted. The micromoles of UMP in RNA and the protein per flask were measured in separate parallel flasks in triplicate. These cells were washed twice at 4 C in buffered saline to remove all serum. The preceding labeled cels were not washed since that procedure could have depleted the cellular UTP. The perchloric acid precipitate was hydrolyzed in KOH, an aliquot removed for protein determination (12), and the remainder brought to 0.2 N perchloric acid (0 C) and centrifuged. The ADX was measured on the supernatant and translated into milligrams of RNA using the base ratios of ribosomal RNA (23) and the extinction coefficients of each nucleotide. The micrograms total of RNA could be converted to micrograms of protein using a measured RNA/protein ratio of 0.11 in these cells. Results of a typical experiment are shown in Fig. 2. The UTP pool is approaching maximum specific activity in 10 min. At this time, the slope of the UMP curve is constant and gives the incorporation rate. The base ratios of the heteronuclear RNA (23) were used to convert Subconfluent HeLa cultures growing on the indicated sugar in at least four small (25 cm2) T-flasks were exposed to [5- H]ribose (to 25 pci/ml and 2.6 nmol/ml). Since ribose enters the cells very slowly, the counts per min into lactic acid and nucleic acids per ribose utilized could be determined most accurately from the media of cultures exposed for longer times (12 to 20 h). [5- H]Ribose and [ Hllactic acid in the media were determined by separation on Gelman TLC-SA sheets (20 X 20 cm) using 1-butano1:glacial acetic acid (505) in ascending chromatography. Ribose has an RF of about 0.75 and lactic acid about [ HNucleic acid was determined by precipitating the cells in the T-flask with 5% trichloroacetic acid at 0 C after removal of the medium by aspiration. The acid precipitate was washed on a filter with cold trichloroacetic acid and then 70% ethanol. The filter was exposed to Protosol (0.5 ml) (New England Nuclear) overnight in a 20-ml glass scintillation vial before addition of 10 ml of scintillation mix and 0.05 ml of glacial acetic acid for counting. n separate samples, the acid precipitate was heated at 90 C for 30 rnin to solubilize nucleic acids; this eliminated 85% of the, H from the precipitate. Assay for Major Components The measurement of cell protein, glucose, fructose, or galactose in the medium, and the accumulated COS, lactic acid, and [ 4C]lactic does come from the reactions of the oxidative arm of the pentose cycle. Conversely, the actual flux could be even higher than that estimated by C-1 release if unlabeled carbon entered the C-1 of 6-P-gluconate by any one of the following reactions: 1) a back reaction of ribdose-5-p plus atmospheric (unlabeled)

4 Pentose Cycle and Nucleic Acid Synthesis 5619 GALACTOSE GLUCOSE FRUCTOSE t P-Llpld FG. 1. Outline of the major HeLa cell metabolic pathways considered in this paper. Fructose is phosphorylated by hexokinase (EC ), to fructose-6-p (rather than by fructokinase (EC ), to fructose--p) by HeLa lysates (1). The superscript on the CO, refers to the carbon number on the hexose. Gal--P, galactose--p; Glc--P, glucose-1-p Glc-6-P, glucose-6-p; HK, hexokinase; Glc-6-P COz to 6-P-gluconate; and 2) randomization of the 2- and 3- carbons to positions 1, 2, or 3 of the regenerated glucose-6-p in the reactions of the nonoxidative arm (9, 10, 25) (Fig. 1). Upon recycling, some original 2- and 3-carbon would become the 1-carbon of the sugar-p. The extent of all of these processes can be assessed using glucose with specific carbon labels. 1) f there were a significant fixation of Con to give 6-P-gluconate, then the specific activity of the C-1 of 6-P-gluconate would be lower than is that of the C-1 of glucose-6-p in cells growing on [ l-14c]hexose since the further back reaction of 6-P-gluconate does not occur (26). 2) When cells are growing on [2-L4C]hexose, any recycling and randomization would label the 1-carbons of both of these sugar phosphates. The specific activities of the 1-carbons were measured by trapping the l4co2 released from cellular glucose-6-p or 6-Pgluconate when these intermediates were reacted to ribulose- 5-P plus COz in vitro (Table ). First, in [l-l4c]glucose-grown cells there was no detectable difference in the specific activi- ties of the 1-carbons of the two sugar phosphates (3.84 versus 3.89); this eliminates any significant in vivo condensation of atmospheric Con to 6-P-gluconate which, as indicated, would lead to a decreased specific activity of the 1-carbon of only 6- P-gluconate. Second, there were no detectable counts released in oztro from the 1-carbons of glucose-6-p plus 6-P-gluconate when cells had been grown on [2-'4C]glucose. The resolution of this experiment was not sufficient to eliminate completely any recycling of pentose cycle carbon. However, such recycling, if it does occur, is probably insignificant. Thus, these experiments demonstrated that 4CO2 release from [ l-'4c]glucose is a valid measure of the carbon flux through the oxidative arm of the pentose cycle. The absence of recycling of glucose carbon was only demonstrable because [2-'*C]glucose was available. Since glycolysis virtually disappears on 2 mm fructose, if any hexose-p and triose-p were formed in the nonoxidative arm reactions from ribose-5-p, they might recycle up the oxidative arm Lactate 1 FATTY ACDS C TRATE CYCLE DH, glucose-6-p dehydrogenase; 6-P-gluconate OH, 6-P-gluconate dehydrogenase; Fru-6-P, fructose-6-p Fru-Z,6-diP, fructose 1,6-bisphosphate; GAP, glyceraldehyde-3-p; DHAP, dihydroxyacetone-p; X-5-P, xylulose-5-p; Sed-7-P, sedoheptulose-7-p; PRPP, a-d-5-p-ribosyl-p2; PDH, pyruvate dehydrogenase. rather than be "swept" down glycolysis as is the case on 10 mm glucose (Fig. 3). However, since [2-L4C]fructose is not available, an in vivo experiment was conceived that could measure any recycling on fructose. The 1-carbon of the hexose is released as Con in the initial round of the oxidative arm reactions. Additional recycling of the hexose carbons would release other carbons as COz. These other carbons would be measured as 4COz from [U-'4C]fructose but would be undetected from [l-'4c]fructose. Thus, if there were no recycling, the percentage of input ['4C]fructose converted to 4COz would be 6 times higher from the [1-14C] compared to the [U- 14 Clfructose and less than 6 if there were recycling (to a minimum of one). Note that this experiment relies on the fact that >90% of the 2 mm fructose carbon passes through the oxidative arm reactions (1). Since a significant amount of 4CO2 is recovered from [3,4-14C]glucose(10 rrm) in the reaction of pyruvate to CoASAc + COZ, this particular experiment would be difficult to interpret in cells growing on glucose. However, even on 2 mm fructose there may be some Con released from this reaction so that the extent of recycling estimated below could be even lower. The results of such an experiment with cells grown on 2 mm fructose are shown in Fig. 4. The mean of the ratio of slopes in three such experiments was 1.27/6 rt- 0.12/6 standard error of the mean. Each additional round of recycling would produce only about one-half as much Con as the preceding round since it can be calculated that at least one-thud of the ribose-5-p must be withdrawn directly for nucleic acid synthesis, e.g. one round of recycling would give 1.50/6. Thus, if there is recycling, it is significantly less than one round equivalent. Carbon Flux through the Oxidative Arm of the Pentose Cycle-The fraction of [1-14C]hexose released as COP is a function both of the supporting sugar and its concentration: about 8% on 10 mm glucose, 50% on 10 mm fructose, and >90% on 2 mm fructose (1). However, because sugar is depleted progressively slower in this sequence from 10 m~ glucose to

5 5620 Cycle Pentose and Nucleic Acid Synthesis TABLE The specific activities of the 1-carbons of glucose-6-p and 6-P-gluc :onate in HeLa cells growing on either [-'4CJ- or (2-'4CJglu~~~e The incubations were performed with subconfluent growing HeLa specific activity of the 1-carbon of 6-P-gluconate was determined by cultures in 75-cm' T-flasks from which the old medium had been measuring the l4c0' trapped after completely reacting the 6-P-gluremoved by aspiration and the cells then washed twice with 10 ml of conate in HeLa cell extracts with excess 6-P-gluconate dehydrogenase medium minus glucose. The fresh medium contained either [2-"C]- (1); the resulting counts per min were divided by the concentration of glucose (2.7 x O' cpm/ml and 2.2 mm) or [-'4CC]glucose(1.72 x 10' cpm/ml and 2.2 KM) in 2.5 ml. The counts per min shown for the [- 14C]glucose experiment have been normalized to the specific activity of the [2-'4C]glucose. The cells were incubated for 34 to 38 min with gentle rocking to ensure continuous contact of cells and medium. Media samples were frozen at -7O'C; cells were extracted and the neutralized extract frozen at -70 C before analyses (1). From 65% to 69% of the glucose was depleted in three such experiments. The 6-P-Gluconate Glucose-6-P 6-P-gluconate in the same extrxt. The glucose-6-p plus 6-P-gluconate was determined by complete reaction of both in a two-step reaction. The first reaction was for 90 min in open test tubes using the glucose- 6-P dehydrogenase assay mix (1) and terminated by boiling for 3 min to convert the gluconolactone to 6-P-gluconate and expel to any l4co) in the medium. The second reaction was in a completely sealed scintillation vial to collect 14C0, evolved in the 6-P-gluconate dehydrogenase reaction (1).-, not measured. Glucose-6-P + 6-P-gluconate [''ClCarbon of input glucose CO2 (cpm) from extract" (1 ml) Not detectable Concentration in extract (pmol/ml) Specific activity (cpm/pmol) Not detectable ' The efficiency of the assay was estimated to be 83% and 87% in two experiments by comparing the counts per min of ''Cor recovered from the [l-'4c]glucose in the medium. The reaction of glucose included the two dehydrogenase steps as described above, preceded by conversion of glucose to glucose-6-p in the hexokinase reaction. [-"C]Glucose was counted in the same 1 N NaOH solution as was the "C02. The same assay of the [2-'4C]glucose stock showed a maximum impurity of 0.06% of '"C in the 1-carbon of glucose. TABLE 1 Pentosephosphate cycle activity in HeLa cells measured by CO, production CO, (micromoles) is calculated from accumulated "TO, (counts per min) and the specific activity of the l-14c-sugar (counts per min/ pmol) precursor. This value measures the activity of the oxidative arm of the cycle (see text). Values f the standard error of the mean are given for the averages when two or more experiments were performed. -, not measured. (pmol/mg of new protein) Fructose (2 mm) < mm fructose (Table ), the actual amounts of carbon flowing through this pathway are not so different; when normalized to the respective rates of protein synthesis, there was only a 2.4-fold difference in flux rates per protein synthesized between cells growing on 10 rn glucose compared to 2 mm fructose (Table 11, line 4). Fate of Pentose Cycle Carbon-Pentose cycle carbon must exit from the cycle since, as shown, very little of it recycles after a complete round from glucose-6-p to fructose-6-p plus glyceraldehyde-3-p. Glucose-6-P itself could be diverted to glycogen; however, only about 1% of glucose carbon is used in glycogen synthesis (even less during growth on fructose or galactose). Furthermore, most glucose-6-p could be converted directly to glycogen without passing through the pentose cycle. The major products of pentose carbon are phosphoribosyl- Pp, the direct precursor of the ribose moiety of nucleic acids, and lactic acid that is derived from the diversion of ribose carbon via the nonoxidative arm into glycolysis (Fig. 1). The relative flow of carbon through these pathways from ribose-5- P cannot be assessed by following labeled sugar carbon since a large fraction of the sugar enters glycolysis directly. However, it is possible to do so by following the fate of labeled ribose itself. While exogenous ribose cannot support the TABLE 111 Fate of exogenous ribose in HeLa cells Cells were labeled with [S-'H]ribose during 12 to 20 h of growth in medium containing the indicated sugar. Two independent experiments are given in (a) and (b). The values give the percentages of [3H]ribose counts per min in the medium at the start of the experiment that are recovered in the indicated fraction. The medium was chromatographed to determine ["Hllactic acid production and [5- "Hlribose utilization. The value in column 2 gives the difference between the total counts per min precipitable in 5% trichloroacetic acid at 0 C and counts per min remaining precipitable in 5% trichloroacetic acid after 30 min at 90 C (column 4). The values of the acidsoluble (at 0 C) fraction (column 3) were the same for extractions in 0.3 N perchloric acid plus 1 m~ EDTA as in 5% trichloroacetic acid or when cells had been grown in [-"Clribose rather than in [5-'HJ- ribose. The values in column 6 give the percentages of ribose utilized that were found in nucleic acid compounds (columns (2 + 3)/( )). 1, mm Glucose (a) (b) mm Fructose (a) 5.54 (b) mm Fructose (a) j (b) 4.88 i Lactic acid < Percentage to nucleic acid compounds > growth of HeLa cells, small amounts of it enter the cells to be phosphorylated to ribose-5-p by ribokinase.' Most RNA synthesized is subsequently degraded (see below) and in attempting to account for the ribose released by RNA degradation, a surprisingly large amount of ribose was found in the small molecules of the acid-soluble fraction after a long period of labeling (Table 111). The compounds have yet to be identified, but they included an insignificant amount of free ribose, were negatively charged at neutral ph, most was retained by acid-activated Norit, and the content of uracil relative to uracil in RNA was about the same as the corresponding ratio of ribose in the soluble to precipitable fraction. They have characteristics of nucleotide-containing compounds. The compounds were also unstable. As a consequence ' R. Chait, L. J. Reitzer, and D. Kennel], unpublished observations.

6 Pentose Cycle Synthesis and Nut cleic Acid 5621 of this latter quality, the mass of the fraction relative to the doubling time to give 0.13 nmol of RNA ribose/min/mg of stable nucleic acids could only be estimated after a sufficiently protein (Table V). Even with DNA synthesis added, this long period of labeling to ensure a nearly uniform specific figure only becomes 0.22 nmol of ribose + deoxyribose/min/ activity of all nucleic acid components. After incorporation of mg of protein. From the Con data of Table 11, the flux of [l-l4c]ribose for one cell doubling, the radioactivity in the soluble fraction was about 65% of that in the precipitable, and as expected, it fell to about 32% after 4 generations of labeling carbon through the oxidative arm gives 0.9 nmol of ribose carbon/min/mg of protein during growth on glucose. Thus, the flux through the oxidative arm appears to be in excess of (Table 111). This corresponds to a mass of 60% to 65% of the the need for ribose in nucleic acids (Table V). This kind of acid-precipitable RNA ribose. Chemical assay for ribose in calculation might have led to the impression that outflow cells growing in the absence of exogenous ribose gave a ratio of ribose-reactive material in cold trichloroacetic acid-soluble/ from the cycle to nucleic acids is insignificant compared to the flux within the cycle (10, 25). However, because of very hot trichloroacetic acid-soluble fractions of about 0.6. The extensive turnover, total RNA synthesis at any instant is 3800 PM ATP in the cells (1) would account for about 15% of the ribose mass in the cold-soluble fraction. much greater than is net synthesis. n cells growing on 10 mm glucose, about 0.42 nmol of ribose The accumulations of label from ['Hlribose into nucleic is being incorporated into RNA/min/mg of protein (Table acids, protein, and lactic acid were measured. n normal V). This is about 3.2 times the net accumulation rate, i.e. growth media with 10 mm glucose, close to 75% of the ribose- about 70% of the RNA synthesized is unstable (or does not 5-P was actually diverted into glycolysis with the remainder incorporated into nucleic acids (Table 111). On 10 mm fructose, only about 27% went to lactate. When grown on 2 m~ fructose, accumulate). t is interesting that this is not very different only 4%of the exogenous ribose was converted to lactate; almost all of it was diverted to nucleic acid components. The incorporation into protein could reflect label in alanine, glycine, and serine from pyruvate and 3-P-glycerate as well as the sugars of glycoproteins and also includes any label in most lipids. The Rate of Nucleic Acid Synthesis-The rate of nucleic ::v acid synthesis can be measured and compared to the flux of 1 - carbon through the oxidative arm in order to assess directly the importance of this flow for meeting the requirements of 3,,, this biosynthetic pathway. Measurement of nucleic acid synthesis in cells is complicated by RNA turnover. The major component of synthesis is the heteronuclear RNA because it TME (Mm) has a very high rate of turnover and still is a very large fraction, e.g. it has an estimated 23-min half-life and includes as much as 7% of the total RNA in L cells (27). An estimate based only on net RNA increase ignores its contribution and is thus an underestimate. Net RNA synthesis can be calculated directly (28) from the RNA/ml of cells and the cell TABLE V Pentose cycle flux and rates of nucleic acid synthesis in HeLa cells growing on 10 mm glucose All values are pmol of ribose/min/mg of protein. Pentose cycle flux through the oxidative arm is from Table 11. Total ribose incorporation into RNA was measured by the instantaneous rate of [%]- UTP incorporation into ['HUMP in RNA at early times of cell labeling with ['Hluridine, the specific activity of the cellular UTP at that time, and the base ratios of newly synthesized HeLa cell RNA. Net ribose incorporation into RNA was derived from the RNA/ protein ratio, the base ratios of HeLa ribosomal RNA, and the cell doubling time. DNA synthesis is estimated from an RNA/DNA of 1.5. Total RNA synthesis was calculated from data in the kind of experiment shown in Fig. 2. The total requirement of ribose-5-p synthesized de nouo in the pentose cycle is a function of the extent of reutilization of any ribose derived from RNA degradation. The two extreme cases are shown in the last column. The value for complete reutilization would be with 0.8 the contribution of the acid-soluble fraction, while with no reutilization of products, the cell would require ( ) X pmol of ribose/min/mg of protein (more if the soluble fraction is a primary product of synthesis). Averages of two or more experiments the standard errors of the mean are given. Total nucleic acid (% of ventose cvcle) '~~~~ Net RNA Total RNA DNA Reutilization of RNA-ribose Complete None (34%) 5.1 (57%)?V,,, FG. 2. The specific activity of the intracellular [3H]UTP and the incorporation of [3H]UMP into RNA at early times after exposure of HeLa cells, growing on 10 mm glucose, to [5,6-3H]uridine (41 Ci/mmol) to give 100 pci/ml. Cells were growing in 25-cm' T-flasks with 2 dures"). ml of media (see "Experimental Proce- t PRPP-NUCLEC ACDS FATTY ACDS FG. 3. Diagrammatic representation of the relative net flux of sugar carbon per unit of protein synthesized in the major pathways of metabolism in HeLa cells growing on 10 m~ glucose (shaded) or 2 mm fructose (solid)(see Fig. 1 for more detail of the reactions). The relative flux is approximately proportional to the width of the path, e.g. with glucose to glucose-6-p defined as loo%, the flow of glucose carbon up the oxidative arm of the pentose cycle is lo%, to PRPP is 576, and to lactate, 80%. The flux of fructose per new cell mass is about 5% relative to total glucose flow per new mass and almost all is accounted for by flow to nucleic acids. The values shown are all from this paper or earlier measurements (l), except that the flux from dihydroxyacetone (DHAP) to P-lipid has been assumed to be about 576, and indirect evidence suggests that 4% flux of pyruvate to the citrate cycle returns from citrate to CoASAc for fatty acid synthesis (1). Abbreviations are the same as in Fig. 1.

7 5622 Pentose Cycle and Nucleic Synthesis Acid TABLE V Levels of enzymes important for glycolytic and pentose phosphate cycle activity in cultured mouse L and human HeLa cells grown on different sugars Cells were grown for at least 2 weeks on the given sugar (10 mm). linear with respect to homogenate concentration and time. Each The vdues give nanomoles/min/mg of cell protein. The numbers in enzyme was assayed in optimal known conditions ( Experimental parentheses give the number of different extracts assayed. -, the Procedures ) so that in most cases relative activities between enzymes activity was not determined. Each assay was performed in triplicate could be very different in uiuo. and the values without substrate subtracted. AU enzyme assays were Glucose Galactose HeLa Fructose (10 mm) (2 mm) L Glucose Fructose Mannose Hexokinase (2) 28 (3) (2) - (2) (1) 66 (1) Phosphofructokinase 75 (1) 80 (1) 65 (1) - 62 (1) 46 (1) 79 (1) Glucose-6-P dehydrogenase 12.4 (2) 113 (2) 120 (2) 88 (2) 108 (1) 91 (1) 6-P-gluconate dehydrogenase (3) (2) 20.5 (2) 19 (2) 24 (1) 18 (1) Transaldolase (2) (3) Transketolase (4) (1) (2) - - TABLE vr Concentrations of intermediates in thepentosephosphate cycle and the cytoplasmic NADPH/NADP+ ratio in HeLa cells as a function of exogenous sugar All values are micromolar. The cells had been growing in T-flasks on a given sugar for more than 2 weeks. Each value is the average of determinations on extracts from six cultures (five for galactose) with the standard error of the mean given. -, not determined. Glucose (10 mm) Galactose (10 mm) Fructose (10 mm) Fructose (2 mm) Glucose-6-P 234 f f f Phosphogluconate 75 f f f 4.7 Xylulose-5-P f Sedoheptulose-7-P 70 f f 6 - Erythrose-4-P 16 t 2.6 <2 7.3 f NADPH/NADPih 20.5 t t f NADP The value of sedoheptulose-7-p could not be measured in cells grown on fructose (see Experimental Procedures ). The NADPH/NADP ratio in the cytoplasm is estimated from the isocitrate/a-ketoglutarate ratio and published equilibrium constant (18) for the isocitrate dehydrogenase reaction. The equilibrium constant was determined in conditions as close as possible to physiological. Of the other NADPH-NADP reactions in the cell, the glucose 6-phosphate dehydrogenase and 6-phosphogluconate dehydrogenase reactions are not at equilibrium, and other reactions are difficult to measure, but the agreement in NADPH/NADP ratio from the isocitrate dehydrogenase and malic enzyme reactions in glucose-grown cells (see Experimental Procedures ) and in normal liver (18) suggests that the ratio of their products to substrates is a reasonable measure of the NADPH/NADP ratio. The NADP+ value is derived from the total measured NADPH in the cell (bound plus unbound) and theffective cytoplasmic NADPH/ NADP+ ratio shown here. Thus. it is an estimate of the maximum effective cytoplasmic concentration. The actual NADP cytoplasmic concentration could be higher or lower. from the value in Escherichia coli (reviewed in Ref. 28). The total rate of nucleic acid ribose synthesis is 0.51 nmol/min/ metabolism from 10 mm glucose or 2 mm fructose are approximated schematically in Fig. 3. mg of protein and this value is about 55% of the rate ofribose- Enzymes of the Pentose Cycle-Glucose-6-P dehydrogen- 5-P synthesis from the oxidative arm of the pentose cycle ase, the first enzyme for entry into the pentose cycle (Fig. l), (Table V). Less ribose-5-p would be needed for nucleic acid has a large but unrealized capacity with its activity severely synthesis if part of the ribose derived from RNA breakdown were reutilized, thus sparing the ribose-5-p synthesized de novo. t is not clear what fraction is reutilized, but as noted, limited (about 99% in rat liver) by the ratio of NADPH/ a very large pool of ribose compounds accumulates in the small molecular weight fraction (Table 111). f these compounds represent breakdown products of RNA, then the minimum need of ribose-5-p for nucleic acids can be calculated by assuming that all of the remaining RNA ribose from decay is reutilized. Even in this case, about 34% of the ribose-5-p synthesized de novo from the oxidative arm flux would be needed for nucleic acids (Table V). This compares with the independently derived value of about 23% based on the fate of exogenous ribose (Table 111). f there were no reutilization of RNA decay products, both values would be somewhat higher; 57% of the ribose-5-p would then be required for nucleic acids (Table V). However, we have not been able to detect any fraction other than those noted in Table 11 either in the medium or in the cells. Thus, probably most of the remaining breakdown products are reutilized. The relative flux rates for the major pathways of hexose NADP in the cell (29, 30). n liver, the amounts of this enzyme, as well as 6-P-gluconate dehydrogenase, can change 5- to 10-fold with changes in diet (18, 31-33). However, in HeLa cells, the amounts of these oxidative arm enzymes did not change as a function of supporting sugar (Table V). The levels of transaldolase and transketolase of the nonoxidative arm as well as hexokinase and phosphofructokinase were also invariant. The Concentrations of Pentose Cycle Zntermediates-Pentose cycle intermediates did change with supporting sugar but not necessarily in parallel (Table V). The level of glucose-6- P decreased about 9-fold from 10 m~ glucose to 10 mm fructose and 25-fold on 2 m~ fructose growth, while 6-Pgluconate changed only 3- to 4-fold and 7-fold, respectively. Xylulose-5-P, which is assumed to be in equilibrium and at about equal concentration with ribose-5-p (34), changed only 2- to 3-fold which is also the ratiof oxidative arm flow rates on the two sugars (Table 11). Thus, there is a gradient of concentration ratios whereby a large difference in the starting substrate, glucose-6-p, is damped down to a much smaller

8 difference in pentose-p concentrations. (t is not clear why the erythrose-4-p level is so low in galactose-grown cells. Galactose or galactose-1-p, the only compounds known to be higher in these cells: does not inhibit the enzyme reaction.) As the fructose concentration was increased above 10 mm, levels of glucose-6-p, fructose-6-p, and 6-P-gluconate rose; furthermore, the fraction of [ l-14c]fructose released as 4CO2 declined with a concomitant increase in [l4c]1actate production. At 100 mm fructose, these values were close to those measured in cultures growing on 10 m~ glucose (data not shown). Within the limitations of measurement (see "Experimental Procedures"), the effective level of cytoplasmic NADP' and cytoplasmic NADPH/NADP' did not change significantly as a function of sugar despite different flux rates. These pyridine nucleotides cannot account for the differential reaction rates. DSCUSSON Control of the Pentose-P Cycle-The rate of hexose depletion can vary 50- to 100-fold without a major change in growth rate of HeLa cells (1). This remarkable range in sugar utilization rates reflects the fact that sugar metabolism is not necessary to provide energy. The major source of energy is always from glutamine oxidation which becomes the only source of energy with the very low glycolytic rate on 2 mm fructose (1). However, over this same range of utilization rates, the flux of sugar carbon through the oxidative arm of the pentose cycle changes only 4- to 5-fold. This relative constancy suggests that this pathway is the only essential one for sugar metabolism and it must be maintained in the face of decreasing levels of available hexose in order to sustain continued growth. What factors determine flux the through the oxidative arm? Previous studies have concluded that it is controlled at the glucose-6-p dehydrogenase reaction with the velocity limited severely by the limiting NADP' and inhibitory NADPH (30, 35-37) while the second enzyme reaction is in equilibrium (36, 37). Our data also indicate that this first reaction is not in equilibrium since the ratiof (6-P-gluconate)(NADPH)/(glucose-g-P)(NADP+) is not constant; it increases progressively from 10 mm glucose +. (10 mm -+ 2 m ~ fructose ) (Table V). Our data are insufficient to prove whether or not the second reaction is in equilibrium in HeLa cells during growth on these sugars. f the epimerase reaction of xylulose-5-p and ribulose- 5-P is in equilibrium as it is in brain tissue (17), then the measured values of the former (Table V) would give a ratio of (ribulose-5-p)(nadph)/(6-p-gluconate)(nadp') that is approximately the same 10 on m~ glucose and 10 m~ fructose; this is consistent with an equilibrium reaction. At the cytoplasmic NADPH/NADP' ratio estimated here (Table V), glucose-6-p dehydrogenase was inhibited 80 to 85% (data not shown). However, this severe damping of the flow cannot account for the changes in flux observed with growth on different sugars since the maximum effective level of free cytoplasmic NADP' and the effective NADPH/ NADP' ratio did not change significantly. As noted, the value of NADP' in Table V is the maximum effective (as enzyme substrate) cytoplasmic concentration. f the total measured NADPH includes fractions in the mitochondria or bound molecules that are not contributing to the actual mass action concentration for these enzymes, then the effective NADP' concentration for these enzymes must be correspondingly lower. t is likely indeed that the effective NADP' level is lower. However, none of the conclusions derived below would be changed if (keeping the measured effective NADPH/ ' B. M. Wice, L. J. Reitzer, and D. Kennell, unpublished observations. Pentose Cycle and Nucleic Acid Synthesis 5623 NADP' ratio constant) the NADP' were 10 times or even 100 times lower. We have estimated the velocities of the reactions as a function of the concentrations of glucose-6-p and 6-P-gluconate shown in Table V. The glucose-6-p dehydrogenase reaction is a sequential ordered one (38,39) with NADP' (H) binding to free enzyme. The rate equation for this reaction mechanism was formulated by Cleland (40). where V,,, is the maximum velocity, K, and Kb the Michaelis constants for NADP+ and glucose-6-p, respectively, and K,, and K,, the E. NADP' and E. NADPH dissociation constants for NADP' and NADPH, respectively. A, B, and Q are the concentrations of NADP', glucose-6-p, and NADPH, respectively. We have used the constants reported by Kanji et al. (39) for pig liver glucose-6-p dehydrogenase: K, = 4.8 p ~ K, h = 36 p ~ K,,, = 9,UM, and K,, = 9 VM. They fall within the range of values reported for various mammalian cells and tissues (41, 42), any of which when used in Equation 1 would lead to the same qualitative conclusion. When the concentrations of NADP', NADPH, and glucose-6-p in Table V are used in Equation 1, the relative velocities on 10 mm ghcose/lo mm fructose are 2.3 to 1. The levels of NADP' and NADPH were not measured in cells growing on 2 mm fructose, but using the same values as measured in cells growing on 10 m~ fructose or glucose, the further reduced concentration of glucose-6-p would give a velocity 6.5 times lower than that on 10 mm glucose. These values are remarkably similar to the observed in vivo flux ratios derived from C02 evolution in Table 1 (2.2 for 10 mm glucose/lo mm fructose and 4.5 for 10 m~ glucose/ 2 mm fructose). Procsal and Holten (43) reported that the kinetics of the 6- phosphogluconate dehydrogenase reaction was consistent with a sequential but random mechanism, with the complexing of one substrate not affecting the binding of the other. n this case, the following equation applies (44). The symbols are defined as in Equation 1. We have used average values for constants measured in various mammalian cells (43, 45, 46): K, = 10,UM, Kt, = 20 PM, and Kt, = 20 p ~. With the concentrations of NADP', NADPH, and 6-P-gluconate given in Table V, the relative velocities are: 10 mm glucose/lo mm fructose, 1.4 and 10 m~ glucose/2 m~ fructose, 2.2. Although not as close to the in vivo ratios of rates as were the values calculated for the first reaction, these values show the dependence of velocity on the sugar-p concentration. The low ratio of 2.2 on 10 mm glucose/2 mm fructose compared to the in vivo ratio of 4.5 may reflect the uncertainty associated with the very low concentration of 6-P-gluconate in 2 mm fructose cells with a standard error of the mean close to k50%. Eggleston and Krebs (30) concluded that physiological levels of oxidized glutathione are required to counteract the NADPH inhibition of liver glucose-6-p dehydrogenase. We could not observe such stimulation of the HeLa enzyme in vitro. While these are crude estimations based on kinetic constants that can only approximate those in.the specific environment inside a cell, it can be seen that the observed ranges of sugar-p concentrations by themselves are sufficient to account for the range of flux values seen in uiuo without invoking the participation of any other factors or molecules. As indicated at the start, of physiological interest is not that

9 5624 and Cycle Pentose the flux through the oxidative ann varies but that it varies so little under conditions in which glycolysis, which starts from the same common substrate, glucose-6-p, is varying several hundred-fold. The reason for this relative constancy can be seen from inspection of Equations 1 and 2. n both of them, the largest terms in the velocity expression at these reactant concentrations are the ones containing the factor K,/A (the first term in Equation 1 and the second and fourth ones in Equation 2). This is the case because the level of NADP is so low compared to its Michaelis constant for each dehydrogenase. Thus, the effect of a very large difference in the starting sugar-p substrate (25-fold between 10 mm glucose and 2 mm fructose) is damped (to give only a 4.5-fold difference in flux (Table 11)). n contrast, its effect on the flow through glycolysis is greatly amplified. This is probably a result of control at the complex phosphofructokinase reaction. Part of this amplification could be explained by the fact that the reaction product, fructose 1,6-bisphosphate, is itself an activator of phosphofructokinase (11) and decreases by a greater factor than does the substrate, fructose-6-p (in equilibrium with glucose-6-p (47)). Functions of the Pentose Cycle-t has been assumed that the primary functions of the pentose phosphate cycle in animal cells are to supply pentose-p for synthesis of nucleotides and to regenerate reducing equivalents of NADPH for biosynthetic reactions (especially those in fatty acid synthesis (5)) which specifically require NADPH, rather than NADH (48, 49). There is some, but not perfect, correlation between the rates of lipogenesis and pentose cycle oxidation in liver (reviewed in Ref. 6). n this tissue, the activities of the dehydrogenases and specific lipogenic enzymes increase in response to insulin (50). Elaborate studies of the fate of specifically labeled [ C- or [: H]glucose in adipose tissues have demonstrated the transfer of [H from NADPH to fatty acid (7-9, 51). However, during rapid lipogenesis, the rate of NADPH synthesis increases but is insufficient to meet the biosynthetic demands (7,8), and the remaining NADPH is produced from NADH in a coupled reaction (8, 52). While there may be some correlation between oxidative arm activity and lipogenesis, this says nothing about a causal relationship or the essentiality of one process for the other. For example, conditions which increase lipogenic activity could also favor nucleic acid synthesis (turnover in nongrowing cells) and with it an increased oxidative arm activity in order to provide the needed ribose-p. For that matter, increased activity of the oxidative arm may itself be the result Nucleic Synthesis Acid it would be 0.18 lactate from each glutamine times 1 molecule produced times 0.7 glutamine used per 1 glucose, to give 44% from glutamine. On 2 mm fructose, only 17% would now be from the oxidative arm reactions and 83% from the reaction of malate to pyruvate; approximately the same total amount of NADPH would be produced on the two sugars but the amount from the pentose cycle would decrease and the amount from glutamine metabolism would increase in the transition to 2 mm fructose. t was shown that very little fructose carbon recycles in HeLa cells (Fig. 4). Even if there were maximal recycling, the NADPH produced would only be doubled since close to one-third of the ribose-5-p would have to be withdrawn for nucleic acid synthesis on each round. The malic enzyme reaction would still provide 71% of the NADPH rather than 83%. t seems unlikely that NADPH generation is an essential function of the pentose cycle in HeLa cells. The relative efficiency of the pentose cycle for the production of NADPH as opposed to ribose-p is a function of the patterns of carbon flow. The most efficient production of NADPH would result if the cycle operated as a closed system with a continuing recycling to yield many rounds of reduction per hexose-p. On the other hand, the most effective production of ribose-p would occur if both the oxidative and nonox- idative arms operated in the same direction to convert hexose- P to ribose-5-p. The latter pattern was supported by studies of Hiatt (56, 57) who followed the fate of C-1- and C-2-labeled glucose into specific carbons of RNA ribose and concluded that most ribose-5-p is formed from the nonoxidative arm in HeLa cells. Gumaa and McLean (36) concluded that it accounts for 80% of ribose-5-p synthesis in ascites cells. Horecker (58) suggested that the pathway does not operate a cycle as in animals but rather as two parallel sequences for production of ribose-5-p. However, Katz and Wood (25) argued that these earlier conclusions did not account for exchange of carbons in the reversible transketolase and transaldolase reactions, and that the net flow was from pentose-p to hexose-p and triose-p in E. coli, in rat liver and muscle, and in HeLa cells (10). Katz and Rognstad (10) went a step further and contended that the net flux through the oxidative arm alone was in great excess over any needs for production of ribose for nucleic acids and that the major role of the pentose cycle is to produce NADPH. The preceding studies were all limited to glucose-supported metabolism. The results reported here for glucose-supported HeLa cells agree with some of these conclusions and disagree with others. First, it is necessary to measure the rate of nucleic of an increased synthesis of NADP from an increased rate of acid synthesis in order to assess the requirements of ribose-5- fatty acid synthesis rather than the converse (53). Finally, the P from pentose cycle metabolism. A large fraction of the RNA production of NADPH from the oxidative reactions may not synthesized in animal cells is the unstable heteronuclear RNA be essential for survival as evidenced by the direct observation and total synthesis at any instant is not measured by the net that Drosophila mutants lacking glucose-6-p dehydrogenase accumulation of RNA with time. Most of the RNA being grow and reproduce (54, 55). Presumably, in this case the synthesized in HeLa cells is unstable (Table V). The actual requirement of ribose-5-p for nucleic acids is met by the demands for ribose-5-p for RNA synthesis are also a function nonoxidative arm. At the same time, the requirement for of the size of the acid-soluble pool of ribose compounds in the NADPH must now be met by other enzyme reactions. Of cell and the extent to which ribose derived from RNA breakcourse, in specialized tissues, such as adipose and liver, down is reutilized. The former was found to be very significant, NADPH production from the pentose cycle may be regulated about 65% of the amount of acid-precipitable RNA. by the degree of lipogenic activity (50). On glucose, this total synthetic need of ribose-p for nucleic The relative amounts of NADPH produced from the oxi- acids is at least one-third the rate of ribose-5-p synthesis by dative arm compared to the malic enzyme reaction can be the oxidative arm of the pentose cycle. The fate of the riboseestimated in HeLa cells using data from Ref. 1. f all of the 5-P was followed independently by following the fate of exglutamine that was converted to lactic acid passed through ogenous [3H]ribose. t should end either in nucleic acids or in the malic enzyme reaction, the NADPH produced in glucose- lactate via the nonoxidative arm. This follows since it was grown HeLa cells would be comparable to that produced in shown that the back reaction to 6-P-gluconate is not signifithe two reactions of the pentose cycle. The NADPH from cant (Table ) and flow down the nonoxidative arm would add glucose would be the product of 0.08 through the oxidative to the fructose-6-p and triose-p pools which are a part of the arm, 2 molecules of NADPH produced, while from glutamine major flux to lactate (80% of total glucose) (Fig. 3). ndeed,

10 Pentose Synthesis Acid Cycle Nucleic and 5625 close to one-fourth of this marker ribose ended in nucleic was met by appropriate supplements to the medium. n this acids. These independent results agree and are consistent with case, glucose-6-p and the pentose cycle (and glycolysis too, of a flux through the oxidative arm that is about 3-fold in excess course) might become virtually immeasurable. This would of the actual needs for nucleic acid synthesis. The surplus show directly that the only essential function of sugar metabribose-5-p is diverted down the nonoxidative arm and through olism is to provide carbon in the pentose cycle and, further, glycolysis, into lactate. that, except possibly for specialized tissues, this cycle is only These conclusions suggest that Katz and Wood (25) were essential for nucleic acid synthesis. partially correct; the earlier studies (e.g. Refs. 36, 56, 57) suggesting a net flow up the nonoxidative arm to ribose-5-p Acknowledgments-We benefited from advice and comments of resulted from randomization of labeled carbons and the net flow was really down to glyceraldehyde-3-p plus fructose-6-p, as they concluded. However, our results disagree with their major premise that there is considerable recycling of carbop around the pentose cycle with the major function of these reactions to generate NADPH and with only a small outflow needed for nucleic acid synthesis. As the level of glucose-6-p declines as a result of growth on a different hexose and/or reducing the concentration of exogenous hexose (starvation of an animal), then a greater fraction of the ribose-5-p is used for nucleic acid synthesis. On 2 mm fructose, virtually all of the ribose-5-p is channeled to nucleic acids; there is no surplus to divert into glycolysis. t seems likely that this limitation of ribose-5-p for nucleic acids now limits the growth rate of the cells. These observations argue that the only essential role of sugar for growth is to produce ribose-5-p for nucleic acids, and that this function probably does limit the growth rate unless excess ribose-5-p is produced such as is the case on high glucose. ndeed, HeLa cells should grow in the complete absence of sugar if this requirement for nucleic acid precursors - 02 i 0 5u/ 1 O 1 Ol 0 30 O 20 TME (hrr) FG. 4. The release of COZ from [l- 4CJfructose and from (U-14C]fructose in HeLa cells growing on 2 rn fructose. The cells had been growing on 2 mm fructose for several generations and were distributed to 25-cm2 T-flasks with canted necks (Corning). About 16 h later, the media were removed by aspiration, and without washing, 3 ml of fresh medium was added to each flask. The medium contained either [l- 4C]fructose (1 pci) or [U- 4C]fructose (2 pci) per flask with 2 mm fructose. The flasks were purged with 5% CO2 and 95% air just before inserting the air-tight CO, traps (1). The 14C in each input medium was determined by counting an aliquot in 1 ml of 10% NaOH (the solution that was used to trap Con in each flask), and the fraction of input C released as 4CO2 is plotted versus the time of flask sampling. The more rapid evolution of 4CO2 during the fmt 3 to 4 h must represent an initial disequilibrium situation upon addition of fresh medium with a resultant transient surge of hexose through the pentose cycle. 10 Dr. Oliver H. Lowry and discussions with Dr. Vincent J. 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