Biochem. J. (2013) 454, 91–99 (Printed in Great Britain)
91
doi:10.1042/BJ20130153
Cell cycle regulation of purine synthesis by phosphoribosyl pyrophosphate
and inorganic phosphate
Alla FRIDMAN*1 , Arindam SAHA*1 , Adriano CHAN*, Darren E. CASTEEL*, Renate B. PILZ* and Gerry R. BOSS*2
*Department of Medicine, University of California, San Diego, 9500 Gilman Drive, La Jolla, CA 92093-0652, U.S.A.
Cells must increase synthesis of purine nucleotides/deoxynucleotides before or during S-phase. We found that rates of purine
synthesis via the de novo and salvage pathways increased 5.0- and
3.3-fold respectively, as cells progressed from mid-G1 -phase to
early S-phase. The increased purine synthesis could be attributed
to a 3.2-fold increase in intracellular PRPP (5-phosphoribosylα-1-pyrophosphate), a rate-limiting substrate for de novo and
salvage purine synthesis. PRPP can be produced by the oxidative
and non-oxidative pentose phosphate pathways, and we found a
3.1-fold increase in flow through the non-oxidative pathway, with
no change in oxidative pathway activity. Non-oxidative pentose
phosphate pathway enzymes showed no change in activity, but
PRPP synthetase is regulated by phosphate, and we found that
phosphate uptake and total intracellular phosphate concentration
increased significantly between mid-G1 -phase and early S-phase.
Over the same time period, PRPP synthetase activity increased
2.5-fold when assayed in the absence of added phosphate, making
enzyme activity dependent on cellular phosphate at the time of extraction. We conclude that purine synthesis increases as cells progress from G1 - to S-phase, and that the increase is from heightened
PRPP synthetase activity due to increased intracellular phosphate.
INTRODUCTION
have shown that ribose 5-phosphate, the immediate precursor of
PRPP, can also be rate-limiting to purine nucleotide synthesis
[10–12]. Ribose 5-phosphate is produced by both the oxidative
and non-oxidative branches of the pentose phosphate pathway,
and changes in flow through either pathway can modulate rates
of purine nucleotide synthesis [13–19].
We now show that purine nucleotide synthesis by both the de
novo and salvage pathways increases markedly as cells progress
from G1 into S-phase, and that this increase in purine synthesis
is driven by a corresponding increase in PRPP production. We
found an increase in phosphate uptake, intracellular phosphate
concentration and PRPP synthetase activity, sufficient to increase
PRPP production and purine synthesis.
Purine nucleotides are synthesized as monophosphates, which
are then phosphorylated to the di- and tri-phosphate forms. All
six purine nucleotides, i.e. AMP, ADP, ATP, GMP, GDP and
GTP, are important components of the cell’s metabolome, with
ADP and GDP serving as precursors of purine deoxynucleotides.
The intracellular concentration of purine deoxynucleotides is
small compared with the amount of cellular DNA, and a large
increase in the synthesis of purine deoxynucleotides, and thus
of purine nucleotides, must occur during G1 - and/or S-phase of
the cell cycle. However, purine nucleotide synthesis during the
cell cycle has not been studied specifically [1,2]. Understanding
the regulation of purine nucleotide synthesis during the cell
cycle could lead to better forms of therapy for hyperproliferative
diseases such as cancer and psoriasis.
Cells synthesize purine nucleotides via either the de novo
or salvage pathways. The de novo pathway consists of ten
steps, and work by An et al. [3] showed that the six enzymes
catalysing these reactions cluster into a purinosome in states of
high purine synthesis. The de novo pathway starts with PRPP (5phosphoribosyl-α-1-pyrophosphate) and ends with IMP, which
can be converted into either AMP or GMP. The salvage pathways
combine either adenine or hypoxanthine with PRPP to yield the
corresponding purine nucleotides, i.e. AMP and IMP. Thus both
the de novo and salvage pathways require PRPP, which is a ratelimiting substrate for both pathways [4,5]. PRPP is synthesized
by PRPP synthetase, a highly regulated enzyme that has an
absolute requirement for Pi (inorganic phosphate), and is subject
to feedback inhibition by purine nucleotides, particularly ADP and
GDP [6–8]. The importance of PRPP synthetase to rates of purine
synthesis is underscored by the fact that genetic overactivity of
the enzyme leads to hyperuricaemia and gout [9]. We and others
Key words: cell cycle, pentose phosphate pathway, phosphoribosyl pyrophosphate (PRPP), phosphoribosylpyrophosphate
synthetase, purine nucleotide.
EXPERIMENTAL
Cell culture and cell cycle synchronization
We performed all experiments in the human colon carcinoma cell
line HCT116, which is diploid and commonly used for cell cycle
studies [20]. Key experiments were confirmed in the monkey
fibroblast-like cell line COS-7, making the results generalizable
across species and cell types. HCT116 cells were cultured in
McCoy’s 5A medium supplemented with 10 % (v/v) FBS, and
COS-7 cells were cultured in DMEM (Dulbecco’s modified
Eagle’s medium) supplemented with 10 % (v/v) FBS.
Cells were synchronized using a nocodazole block by
incubating exponentially growing cells with 0.12 μg/ml
nocodazole for 14 h and then recovering cells were arrested in
mitosis by shake-off. The cells were washed twice with PBS and
resuspended in McCoy’s 5A medium (HCT116 cells) or DMEM
(COS-7 cells); both media were supplemented with 10 % (v/v)
FBS that had been dialysed against normal saline. The cells were
Abbreviations used: AICA-riboside, 5-amino-4-imidazolecarboxamide riboside; DMEM, Dulbecco’s modified Eagle’s medium; PRPP, 5-phosphoribosylα-1-pyrophosphate.
1
These authors contributed equally to this work.
2
To whom correspondence should be addressed (email [email protected]).
c The Authors Journal compilation c 2013 Biochemical Society
92
A. Fridman and others
plated in six-well cluster dishes, 25 cm2 flasks or 100 mm tissue
culture plates as noted; cells were incubated for 4, 6, 8 or 10 h,
and then studied as described below.
Cells were also synchronized using a double-thymidine block,
by incubating exponentially growing cells with 2 mM thymidine
for 18 h, releasing cells from the block for 6 h and then re-treating
the cells with 2 mM thymidine for 18 h. After release from the
second block, the cells were studied either immediately or at 4
and 6 h after release.
Overview of biochemical and immunological methods
With the exception of measuring phosphate uptake, all methods
have been described in detail previously [10,11,13,21–23]. Thus
only brief methodological descriptions are provided. In assays
involving incorporation of one of the six different radioactive
precursors used in these studies, ∼ 106 cells were incubated for
1 h with the radioactive compound, and the assays were linear
with time (from 0 to 90 min) and cell number (from 5 × 105 to
1.5 × 106 cells); radioactivity was measured by liquid-scintillation
counting.
Assessment of cell cycle progression
Cells in six-well dishes were harvested at the times indicated,
and extracted under denaturing conditions. Cyclin E and cyclin
A expression were analysed by immunoblotting using mouse
monoclonal antibodies sc-247 and sc-53230 respectively (Santa
Cruz Biotechnology) with α-tubulin as a loading control [24].
DNA synthesis
Cells in six-well dishes were incubated for 1 h with 10 μCi of
[methyl-3 H]thymidine (20 Ci/mmol, final concentration 0.5 μM),
washed with ice-cold PBS and extracted in situ in 10 %
(w/v) trichloroacetic acid [21]. Precipitated DNA was collected
on glass microfibre filters, which were washed with 10 %
(w/v) tricholoroacetic acid, and radioactivity on the filters was
measured.
De novo purine synthesis
Cells in six-well dishes were incubated for 1 h with 10 μCi of
[14 C]formate (54 mCi/mmol, final concentration 185 μM). They
were washed with ice-cold PBS and extracted in situ in 0.4 M
perchloric acid [10,21–23]. The extracts were heated to 100 ◦ C
for 70 min to break the glycosidic bond between the purine base
and ribose group. The samples were centrifuged at 1000 g for
5 min, and supernatants were applied to AG Dowex 50 columns,
which were washed with 0.1 M HCl to remove unreacted formate
and other anions. Purines were eluted in 6 M HCl and radioactivity
was measured.
PRPP content and production
PRPP content. Cells from one 100-mm-diameter dish were
extracted by hypotonic lysis in buffer containing 10 mM EDTA
and phosphatase inhibitors. The extracts were heated at
100 ◦ C for 4 min and cooled, and PRPP in the extracts was
measured by conversion into IMP using [8-14 C]hypoxanthine
(52 mCi/mmol, final concentration, 55 μM) and purified hypoxanthine guanine phosphoribosyltransferase (NovoCIB) [11]. In
control experiments, we recovered 85 % of exogenous PRPP
added to cell extracts.
PRPP production. Cells in six-well dishes were incubated for
1 h with 10 μCi of [8-14 C]adenine (47 mCi/mmol, final concentration 210 μM). They were washed with PBS, collected by
centrifugation at 10 000 g for 1 min, and lysed in water [10,23].
The lysates were applied to 1 cm × 1 cm squares of DE-81 paper,
which were washed in ammonium formate, and radioactivity in
adenylates retained on the squares was measured.
Purine synthesis by the salvage pathway
Cells in six-well dishes were incubated for 1 h with 10 μCi of
[14 C]hypoxanthine (50 mCi/mmol, final concentration 200 μM),
and then processed as described for measuring de novo purine
synthesis [21,23].
AICA-riboside (5-amino-4-imidazolecarboxamide riboside) incorporation
into purines
Cells in six-well dishes were incubated with 10 μM azaserine
and 200 μM AICA-riboside for 1 h before adding 10 μCi of
[14 C]formate (54 mCi/mmol, final concentration 185 μM) [21–
23]. Azaserine inhibits the fourth step of the de novo purine
synthesis pathway, and AICA-riboside enters the pathway at the
last step. The remainder of the protocol was as described for
measuring de novo purine synthesis.
Carbon flow through the oxidative pentose phosphate pathway
Cells in 25 cm2 flasks were transferred to glucose-free,
bicarbonate-free DMEM supplemented with 1 mM glucose,
25 mM Hepes and 10 % (v/v) dialysed FBS (pH 7.4); 5.0 μCi of
[1-14 C]glucose (53 mCi/mmol) was added to a final concentration
of 1.1 mM. The flasks were sealed with rubber stoppers holding
a plastic centre well (Kontes Glass) containing a fluted piece of
filter paper [13,23]. After a 1 h incubation period, 1 M KOH was
injected through the stopper into the wells to saturate the filter
paper; perchloric acid was then injected into the medium to a
final concentration of 0.4 M. The injected acid lysed the cells and
released CO2 from the medium, which was trapped in the basesaturated filter paper. The flasks were kept overnight at room
temperature (22 ◦ C), and the trapped radioactivity bound to the
filter paper was measured.
Intracellular purine nucleotides
Cells in 100-mm-diameter dishes were washed with ice-cold
PBS, extracted in situ with 0.4 N perchloric acid, and the
extracts were centrifuged at 10 000 g for 1 min [10,21–23]. The
supernatants were neutralized using KHCO3 , and precipitated
potassium perchlorate was removed by centrifugation at 10 000 g
for 1 min. The samples were analysed by HPLC on a strong
anion-exchange column monitored by UV absorption at 258 nm
[10,21].
c The Authors Journal compilation c 2013 Biochemical Society
Carbon flow through the non-oxidative pentose phosphate pathway
Cells in six-well dishes were transferred to DMEM containing
1 mM glucose and 10 % (v/v) dialysed FBS; 10 μCi of [114
C]glucose (53 mCi/mmol) was added to a final concentration of
1.2 mM for 1 h [13,23]. The cells were processed as described for
measuring purine nucleotides, except fractions from the HPLC
column corresponding to ATP and GTP were collected, and
radioactivity was measured by liquid-scintillation counting.
PRPP synthetase regulation during the cell cycle
Carbon flow through both pentose phosphate pathways
Cells were treated and processed as described for measuring carbon flow through the non-oxidative pentose phosphate pathway,
except they were incubated with 10 μCi of [6-14 C]glucose
(55 mCi/mmol) [13,23].
Total cellular Pi
Cells in six-well dishes were washed three times with TBS
and extracted in situ in 0.4 M perchloric acid. Extracts were
centrifuged at 10 000 g for 1 min, and supernatants neutralized
with KHCO3 . Pi was measured spectrophotometrically using
Malachite Green in the presence of ammonium molybdate, Tween
20 and sodium citrate by comparison with a standard curve [25].
Phosphate uptake
Cells in six-well dishes were washed twice in TBS and DMEM
containing 0.5 mM KH2 PO4 and 10 % (v/v) dialysed FBS was
added to the cells (these studies had to be done in lowphosphate-containing medium; a low-phosphate or phosphatefree formulation of McCoy’s 5A medium is not available). After
adding 10 μCi of [32 P]Pi (9000 Ci/mmol), the cells were incubated
at 37 ◦ C for 16 min [26]. Cells were washed five times in ice-cold
PBS containing 10 % (w/v) BSA and lysed in 0.5 ml of water, and
450 μl of lysate was subjected to liquid-scintillation counting.
Phosphate uptake was linear from 4 min to at least 32 min, and
from 3 × 105 to 9 × 105 cells.
Enzyme activities
Cells in 100-mm-diameter dishes were washed twice with PBS,
and collected by centrifugation at 10 000 g for 1 min. The cell
pellet was frozen in a dry ice-acetone bath, and stored at − 20 ◦ C
for up to 7 days. The frozen cell pellets were resuspended and
sonicated at 4 ◦ C in 50 mM Tris/HCl (pH 7.6), 5 mM MgCl2 ,
2.5 mM 2-mercaptoethanol, 1 mM EDTA, and protease and
phosphatase inhibitors. The cell lysates were centrifuged at
14 000 g for 10 min, and the supernatants were used immediately
[13].
Activities of the following enzymes were measured spectrophotometrically at 365 nm following either NADP + reduction
(glucose-6-phosphate dehydrogenase and fructose bisphosphatase) or NADH oxidation (aldolase, transaldolase, transketolase
and phosphofructokinase). Substrates and coupling enzymes were
from Sigma–Aldrich. A substrate blank (all assay components
except for the cell extract) and an extract blank (all components
except for the substrate) were included, and had <10 % of
the absorbance change of the full system. Assay time was
10 min, and all assays were linear with time to at least 10 min.
Assays were also linear with protein concentration, which was
measured using the Bradford method [27]. The glucose-6phosphate dehydrogenase assay contained 3.3 mM glucose 6phosphate, and the fructose bisphosphatase assay contained 2 mM
fructose 1,6-bisphosphate and 1 unit each of glucose-6-phosphate
dehydrogenase and glucose phosphoisomerase. The aldolase,
transaldolase, transketolase and phosphofructokinase assays
contained 3 units each of glycerol-3-phosphate dehydrogenase
and triose phosphate isomerase, and the following respective
substrates and reagents: (i) 2 mM fructose 1,6-bisphosphate;
(ii) 1 mM fructose 6-phosphate, 1 mM erythrose 4-phosphate
and 75 mM NaCl; (iii) 1 mM ribose 5-phosphate, 1 mM xylulose
5-phosphate, 1 mM thiamine pyrophosphate and 75 mM NaCl;
and (iv) 5 mM fructose 6-phosphate, 250 nM fructose 2,6bisphosphate, 2.5 mM sodium pyrophosphate and 5 mM ATP.
93
PRPP synthetase activity was measured in cell lysates generated
as described above, except: (i) the cell pellets were lysed by
freeze–thawing instead of sonication; and (ii) the extract buffer
contained either 32 mM NaH2 PO4 when measuring maximal
in vitro activity (V max ) or no added phosphate when assessing
approximate in vivo activity [10,11]. Since enzyme activity was
considerably less in the absence of phosphate, the cells were
extracted at a 2-fold higher density and the assay incubation
time was extended from 10 min to 40 min; with the lower
enzyme activity, the assay remained linear at the longer incubation
time and higher protein concentration. The enzyme assays were
otherwise identical between those performed in the presence or
absence of added phosphate, and contained 5 mM MgCl2 , 2.5 mM
ATP, 2.5 mM ribose 5-phosphate, 1 mM 2-glycerol phosphate,
80 μM [8-14 C]hypoxanthine (50 mCi/mmol) and 1.7 m-units
of hypoxanthine-guanine phosphoribosyltransferase. Substrate
and product were separated on DE-81 paper as described for
measuring PRPP production.
NADP + and NADPH
Cells in six-well dishes were extracted, and NADP + and NADPH
were measured in a fluorimetric assay system (Cell Technology,
Fluoro NADP/NADPH Detection Kit).
Statistical analyses
Statistical analyses between two conditions were assessed by a
two-tailed Student’s t test, and analyses between two or more
conditions by comparison with a control condition were assessed
by a repeated measures ANOVA using a Dunnett’s post-test
analysis. All analyses were performed using GraphPad Prism 5
software. A P value of <0.05 was considered significant.
RESULTS
Purine synthesis de novo and DNA synthesis during phases of the
cell cycle
To synchronize cells at the same phase of the cell cycle, we
arrested HCT116 and COS-7 cells in mitosis using nocodazole
and at the G1 /S interface using a double thymidine block. At 4 h
after release from the nocadozole block, the cells were in mid-G1 phase, by 6 h they were in late G1 -phase, by 8 h they were at the
G1 /S interface, and by 10 h they were in early S-phase, as assessed
by monitoring expression of cyclins A and E (Figure 1a, left-hand
panel; cyclin E migrates as a doublet band at 50 kDa and a singlet
band at 42 kDa due to differential phosphorylation). Within 4 h
after release from the double-thymidine block, cells were well
into S-phase, and by 6 h after release, they were at the S/G2
interface, again as assessed by following expression of cyclins A
and E (Figure 1a, right-hand panel). Throughout the remainder
of the present paper, 4, 6, 8 and 10 h after release from the
nocodazole block are referred to as mid-G1 , late G1 , G1 /S and early
S respectively, and 0, 4 and 6 h after release from the thymidine
block are referred to as G1 /S, mid-S and S/G2 respectively.
Rates of DNA synthesis measured by thymidine incorporation
into acid precipitates increased markedly (>9-fold) as cells
progressed from mid-G1 to early S (Figure 1b, closed bars),
and from G1 /S to mid-S (Figure 1b, open bars). Thymidine
incorporation was measured for 1 h, and thus cells that were at
G1 /S at the start of the incubation period were well within S-phase
at the end of the incubation period; hence the reason for relatively
high rates of DNA synthesis at G1 /S. A small non-significant
c The Authors Journal compilation c 2013 Biochemical Society
94
A. Fridman and others
(Figure 1c, open bars). Overall, a good correlation existed
between rates of DNA synthesis and purine nucleotide synthesis
(Figures 1b and 1c), suggesting the two were closely tied. As
confirmatory evidence for increased purine synthesis during G1 phase, rates of purine synthesis increased 2.1-fold in COS-7 cells
between mid-G1 and G1 /S (Supplementary Figure S1, closed bars,
at http://www.biochemj.org/bj/454/bj4540091add.htm).
Intracellular content of purine nucleotides during mid-G1 and G1 /S
Intracellular purine nucleotides, particularly NMPs (nucleotide
monophosphates), inhibit glutamine amidophosphoribosyltransferase, the first committed and major rate-limiting enzyme of
purine nucleotide synthesis [28]. Moreover, purine nucleotides,
particularly NDPs (nucleotide diphosphates), inhibit PRPP
synthetase, another rate-limiting enzyme of purine nucleotide
synthesis [8]. Thus a decrease in the intracellular concentration
of purine nucleotides, as might occur when DNA synthesis
increases, could explain the increase in purine nucleotide
synthesis we observed as cells progressed from G1 -phase into
S-phase. However, we found no significant difference in the
intracellular concentration of the six purine nucleotides, i.e. AMP,
ADP, ATP, GMP, GDP and GTP, between mid-G1 and G1 /S
(Supplementary Table S1 at http://www.biochemj.org/bj/454/
bj4540091add.htm). Thus the increase in purine synthesis during
cellular progression from G1 -phase into S-phase was not from a
decrease in the intracellular content of purine nucleotides, and the
increased rate of purine synthesis was sufficient to compensate
for the increased demand for purine deoxynucleotides.
Figure 1
De novo purine synthesis and DNA synthesis during the cell cycle
HCT116 cells were arrested in mitosis with nocodazole (left-hand panels) or at G1 /S after a
double-thymidine block (right-hand panels). After release from the blocks, cyclin E and cyclin
A expression were assessed by immunoblotting (a), DNA synthesis was measured following
[methyl -3 H]thymidine incorporation into acid-precipitable material (b) and purine synthesis de
novo was measured following [14 C]formate incorporation into all cellular purines (c). In (a),
times shown are those after release from the blocks, and, in (b) and (c), mid and late G1 , G1 /S
and early S correspond to 4, 6, 8 and 10 h after release from the nocodazole block, and G1 /S,
mid-S and S/G2 correspond to 0, 4 and 6 h after release from the double-thymidine block. In (a),
tubulin was used as a control for protein loading, and in (b) and (c), the data are means +
− S.E.M.
for at least three independent experiments performed in duplicate. *P < 0.05, **P < 0.01 and
***P < 0.005 compared with mid-G1 results for nocodazole block or G1 /S results for thymidine
block. NS, no significant difference between the two cell phases.
increase in DNA synthesis occurred between mid-G1 and late
G1 (Figure 1b, first two closed bars), probably from incomplete
cell synchronization with some cells already in S-phase 4 h after
release from the nocodazole block. Rates of DNA synthesis fell
dramatically as cells left S-phase and entered G2 -phase (Figure 1b,
open bars).
Rates of de novo purine synthesis increased 3.3- and 5.0-fold
as HCT116 cells progressed from mid-G1 to G1 /S and early S
respectively (Figure 1c, closed bars). A small, but significant,
increase in purine synthesis occurred between mid-G1 and late
G1 , probably from incomplete cellular synchronization, but it
also could be from other mechanisms discussed below. Rates
of purine synthesis were similar in cells at G1 /S generated either
by release from the nocodazole block or the double-thymidine
block, indicating that the two methods of cell synchronization
yielded similar results (note the two different y-axis scales). Purine
synthesis peaked during mid-S, approximately 9-fold higher than
in mid-G1 (Figure 1c, compare mid-G1 , closed bar, to mid-S,
open bar), and declined as cells moved from S-phase to G2 -phase
c The Authors Journal compilation c 2013 Biochemical Society
PRPP concentration and production, and purine synthesis via the
salvage pathway during G1 -phase and early S-phase
Another major determinant of rates of purine nucleotide synthesis
is the intracellular concentration and production of PRPP, one of
the two substrates of glutamine amidophosphoribosyltransferase,
the first and rate-limiting enzyme of the de novo purine pathway
[4,5,12,17]. We found that the intracellular content of PRPP, as
measured in cell extracts, increased 3.2-fold (from 5.39 +
− 1.8
6
cells)
as
HCT116
cells
progressed
from
to 17.3 +
1.9
pmol/10
−
mid-G1 to early S (P < 0.05 for comparison between the two
phases). These values translate to PRPP concentrations of 6 and
19 μM respectively, using an intracellular volume of ∼ 0.9 pl
for HTC116 cells [29]. These values are similar to what has
been reported previously for human erythrocytes, cultured human
fibroblasts and mouse liver under various conditions [30], and
both PRPP concentrations are far below the K m of glutamine
amidophosphoribosyltransferase [28].
PRPP production, as measured in intact cells following
conversion of adenine into AMP, increased 3.1-fold over the same
interval (Figure 2a; the inset shows that conversion of adenine
into AMP was linear during the 1 h labelling period). A similar
increase in PRPP content and production reinforces the validity of
the results. Although the increase in PRPP was not as great as the
increase in purine synthesis (compare Figures 2a and 1c), small
changes in the intracellular PRPP concentration can profoundly
influence rates of de novo purine synthesis, because of the high K m
of glutamine amidophosphoribosyltransferase for this substrate
[4,5,12,17,28]. Similar to the de novo pathway, purine synthesis
via the salvage pathway depends on PRPP availability [4,12],
and we found that purine synthesis via the salvage pathway also
increased more than 3-fold as cells progressed from mid-G1 to
early S (Figure 2b).
PRPP synthetase regulation during the cell cycle
95
Figure 3 Oxidative and non-oxidative pentose phosphate pathway activity
from mid-G1 to early S
HCT116 cells were arrested in mitosis using a nocodazole block, with cell cycle phases
corresponding to times after release from the nocodazole block as described in the legend to
Figure 1. Flow through the oxidative pentose phosphate pathway (a), the non-oxidative pentose
phosphate pathway (b), and the oxidative and non-oxidative pentose phosphate pathways (c)
were measured by following [1-14 C]glucose oxidation to CO2 , [1-14 C]glucose incorporation
into ATP and GTP, and [6-14 C]glucose incorporation into ATP and GTP respectively. Results are
means +
− S.E.M. for at least three independent experiments performed in duplicate. *P < 0.05
and ***P < 0.005, compared with mid-G1 results; NS, non-significant.
Figure 2 PRPP production, purine synthesis via the salvage pathway, and
AICA-riboside incorporation into purines from mid-G1 to early S
HCT116 cells were arrested in mitosis using nocodazole, and mid-G1 , G1 /S and early S
correspond to times after release from the nocodazole block as described in the legend to
Figure 1. PRPP production was measured by following [8-14 C]adenine incorporation into
adenylates (a), salvage pathway activity was measured by following [8-14 C]hypoxanthine
incorporation into purine nucleotides (b), and AICA-riboside incorporation into purines was
measured following [14 C]formate incorporation into purine nucleotides in the presence of
azaserine and AICA-riboside (c). The inset in (a) shows that adenine incorporation was linear
over the 1 h labelling period, when assessed at 4 h after release from nocodazole block. Data are
means +
− S.E.M. for at least three independent experiments performed in duplicate. *P < 0.05
and ***P < 0.005, compared with mid-G1 results.
To assess further whether the regulation of de novo purine
synthesis during G1 -phase was at an early step and involved
PRPP, we measured AICA-riboside incorporation into purines.
In this assay, early steps of de novo purine synthesis are inhibited
using azaserine and only the last step of the de novo pathway
is assessed. Moreover, ribose is provided in the form of AICAriboside. This assay is therefore not dependent on PRPP, and we
found that AICA-riboside incorporation into purine nucleotides
showed only a modest (1.5-fold) increase between mid-G1 and
early S (Figure 2c). Thus, although some regulation of purine
synthesis occurs in the late steps of the de novo pathway as cells
move through G1 -phase, the major point of regulation appears to
be at the level of PRPP production.
Oxidative and non-oxidative pentose phosphate pathway activity
during G1 -phase and early S-phase
PRPP can be produced by either the oxidative or non-oxidative
branch of the pentose phosphate pathway (Supplementary Figure
S2 at http://www.biochemj.org/bj/454/bj4540091add.htm), and
which pathway contributes more to PRPP production may be
related to the cell type and the experimental conditions [13–
19]. We found that flow through the oxidative pentose phosphate
pathway did not change as cells progressed from mid-G1 to early
S (Figure 3a). Consistent with no change in oxidative pentose
phosphate pathway activity, we found no change in glucose-6phosphate dehydrogenase activity (Supplementary Table S2 at
http://www.biochemj.org/bj/454/bj4540091add.htm) or the ratio
of NADP + /NADPH between early G1 and G1 /S (the ratio was
0.80 +
− 0.09 and 0.77 +
− 0.1 in early G1 and at G1 /S respectively).
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96
A. Fridman and others
Glucose-6-phosphate dehydrogenase is the first and rate-limiting
enzyme of the oxidative pentose phosphate pathway, and its
activity is strongly regulated by the NADP + /NADPH ratio [31].
Because of the potential for extensive carbon exchange in
the non-oxidative pentose phosphate pathway, it is difficult to
precisely measure flow through this pathway (Supplementary
Figure S2). Since the carbon atom in position 1 of glucose is
lost as glucose transits the oxidative pathway, we and others have
followed [1-14 C]glucose incorporation into purine nucleotides as
a measure of flow through the non-oxidative pathway [13,18,19].
This method is accurate, as long as (i) potential exchange of carbon
atoms at positions 1 and 6 does not occur in the glycolytic pathway
at the level of the triose phosphates, or (ii) little of the trioses are
converted back into glucose via reversal of the glycolytic pathway
or the non-oxidative pentose phosphate pathway. We previously
showed that carbon atom exchange and triose conversion accounts
for <5 % of glucose metabolism in cultured cells, and that
monitoring [1-14 C]glucose incorporation into purine nucleotides
is a valid measure of non-oxidative pentose phosphate pathway
activity [13]. We found that [1-14 C]glucose incorporation into
ATP and GTP increased 3.1-fold between mid-G1 and early S
(Figure 3b), which is very similar to the increase we observed in
PRPP production and content (Figure 2a). Thus increased flow
through the non-oxidative pentose phosphate pathway appeared
to be the basis for increased PRPP production during progression
from G1 -phase to S-phase.
We also measured [6-14 C]glucose incorporation into ATP and
GTP, and found a 2.5-fold increase between mid-G1 and early
S (Figure 3c). The carbon atom at position 6 of glucose can
enter purine nucleotides via either the oxidative or non-oxidative
pentose phosphate pathway and thus this assay measures flow
through both pathways [13,19]. Since no change occurred in
oxidative pentose phosphate pathway activity during progression
from G1 -phase to S-phase, this assay provides further evidence
of increased flow through the non-oxidative pathway during this
time.
To study the mechanism of increased flow through the
non-oxidative pentose phosphate pathway during transition
from G1 -phase to S-phase, we measured activities of the five
enzymes in the pathway, i.e. aldolase, fructose bisphosphatase,
phosphofructokinase, transaldolase and transketolase, under
optimal in vitro conditions, and found no change in the activity of
these enzymes between mid-G1 and G1 /S (Supplementary Table
S2).
PRPP synthetase activity and intracellular Pi during progression
from G1 -phase into early S-phase
We next hypothesized that increased conversion of ribose 5phosphate into PRPP by PRPP synthetase could be the basis
for increased flow through the non-oxidative pentose phosphate
pathway, because reactions in the pathway are largely in
equilibrium [18] and a decrease in concentration of one substrate,
e.g. ribose 5-phosphate, should shift the equilibrium towards
that substrate. We therefore measured PRPP synthetase activity
in vitro, initially under optimal standard assay conditions in
buffer containing 32 mM Pi . Under these conditions, we found no
change in enzyme activity between mid-G1 and G1 /S (Figure 4a,
closed bars). We next measured PRPP synthetase activity under
conditions of no added Pi , where the only phosphate present
in the enzyme assay was that which was in the cells at the
time of extraction. As would be expected, enzyme activity was
considerably less than when the assay was performed with added
phosphate (approximately one-tenth as much; compare open
c The Authors Journal compilation c 2013 Biochemical Society
Figure 4 PRPP synthetase activity, and intracellular phosphate
concentration and uptake of Pi in synchronized cells
(a) HCT116 cells were recovered from a nocodazole block, and 4 and 8 h later, corresponding to
mid-G1 and G1 /S respectively, they were extracted and PRPP synthetase activity was measured
in the extracts. For half of the cells, neither the extract buffer nor the enzyme system contained
Pi (open bars). For the other half of the cells, 32 mM NaH2 PO4 was present in both the extract
buffer and enzyme system (closed bars). (b) Cells were released from nocodazole block, and
placed in standard medium containing 10 % (v/v) dialysed FBS. At 4 and 8 h later, they were
extracted in acid for measuring total intracellular Pi (open bars) or incubated with 32 PO4 for
measuring phosphate uptake as described in the Experimental section (closed bars); for the
latter experiments, the cells were changed to medium containing 0.5 mM KH2 PO4 and 10 %
(v/v) dialysed FBS during the 16 min incubation time with radioactive phosphate. Results are
means +
− S.E.M. for at least three independent experiments performed in duplicate. NS, no
statistical difference. *P < 0.05 and **P < 0.01, for differences between mid-G1 and G1 /S.
and closed bars in Figure 4a, note the different y-axis scales).
Importantly, under these conditions of ‘no phosphate added’, we
found a 2.5-fold increase in enzyme activity as cells progressed
from mid-G1 to G1 /S (Figure 4a, open bars).
These data suggested that Pi increased as cells moved through
G1 -phase, and we found a 43 % increase in total intracellular
content of Pi as HCT116 cells progressed from mid-G1 to G1 /S
(Figure 4b, open bars; P < 0.05 for comparison between the two
cell phases). Phosphate content also increased significantly in
COS-7 cells between mid-G1 and G1 /S (Supplementary Figure
S1, open bars). We hypothesized that the basis for the increased
intracellular phosphate was from increased cellular uptake of
phosphate, and we found that phosphate uptake increased 25 %
as cells moved from mid-G1 to G1 /S (Figure 4b, closed bars;
P < 0.05 for comparison between the two cell phases). Taken
together, these data suggest that the increase in PRPP synthetase
activity between G1 -phase and S-phase can be attributed to an
increase in intracellular phosphate concentration.
DISCUSSION
We are unaware of any other studies that measured purine
synthesis at different phases of the cell cycle, but one group
measured purine synthesis over a 96 h period as rat hepatoma
3924A cells progressed from stationary phase to exponential
growth and back to stationary phase [1]. They found that
PRPP synthetase regulation during the cell cycle
the intracellular PRPP concentration and rates of purine
synthesis increased markedly during exponential growth, and then
returned to baseline as cells ceased growing. Another group of
workers showed that inhibitors of purine synthesis, but not of
pyrimidine synthesis, prevented primary human T-lymphocytes
from progressing from G1 -phase to S-phase [2]. Although both
of these studies are different from the present study, they provide
evidence of the importance of purine synthesis during the cell
cycle.
The small increase in purine nucleotide synthesis we observed
during G1 -phase without a change in purine nucleotide pools was
likely to be from incomplete cell synchronization, with some
cells already in S-phase (as mentioned above). Two additional
explanations are also possible. First, total intracellular purine
nucleotide pools are large relative to newly synthesized purine
nucleotides, and thus a small increase in purine synthesis during
G1 -phase could occur without enlarging the nucleotide pools.
Secondly, intracellular concentrations of dATP and dGTP increase
during G1 -phase [32]; thus purine nucleotides newly synthesized
during G1 -phase could be funnelled towards ribonucleotide
reductase and production of deoxynucleotides [32]. Unlike
purine nucleotides, purine deoxynucleotides exert little feedback
inhibition on purine synthesis, and thus an increase in purine
deoxynucleotides would have a minimal effect on rates of purine
synthesis [33].
The human genome has 3.1 × 109 bp [34]. Assuming purine
bases in DNA are evenly divided between adenine and guanine,
this translates to ∼ 2.6 fmol of adenine and guanine per cell.
Deoxynucleotides for DNA synthesis derive from ADP and GDP,
but the latter are in equilibrium with ATP and GTP; thus the sum
of intracellular ADP and ATP, and GDP and GTP can be viewed as
precursors for DNA. The data in Supplementary Table S1 indicate
that the intracellular pools of ADP and ATP, and GDP and GTP
are equivalent to ∼ 4.6 and 2.9 fmol of adenine and guanine
respectively. As might be expected, a cell must almost double its
purine content to replicate its DNA, and, since the nucleotide pools
are large compared with newly synthesized purines, a severalfold
increase in purine synthetic rates would be needed. Thus the 5fold increase in rates of purine synthesis we observed during the
transition from G1 -phase to S-phase would be compatible with
the increase required to meet the cell’s need for DNA replication.
Both the oxidative and non-oxidative branches of the pentose
phosphate pathway can produce ribose 5-phosphate and therefore
PRPP (Supplementary Figure S2). Which pathway predominates
in producing ribose 5-phosphate/PRPP may depend on a variety
of factors, including cell type, cell cycle phase and availability of
nutrients and growth factors [13–19]. However, most workers,
including Becker and colleagues studying normal human
fibroblasts, have found that the non-oxidative pathway supplies
most of the ribose 5-phosphate for purine nucleotides [13–
16]. Consistent with these data, we now found in synchronized
HCT116 cells that the non-oxidative branch was the major
provider of ribose 5-phosphate and PRPP between mid-G1
and S-phase. The oxidative branch may not produce much
ribose 5-phosphate because it terminates in ribulose 5-phosphate
(Supplementary Figure S1). The intracellular concentration
of ribulose 5-phosphate is well below the K m of either
phosphoriboisomerase or ribulose-5-phosphate epimerase for this
substrate, with the K m of the former enzyme approximately 4-fold
higher than that of the latter enzyme [13]. Hence, under normal
conditions, ribulose 5-phosphate produced by the oxidative
pathway would be more likely to be converted into xylulose
5-phosphate than into ribose 5-phosphate. An increase in the
intracellular xylulose 5-phosphate concentration could actually
decrease the intracellular ribose 5-phosphate concentration
97
through the action of transketolase, which catalyses a fully
reversible reaction that achieves equilibrium between substrates
and products [35]. Thus the oxidative pentose phosphate pathway
may provide primarily reduced NADPH, whereas the nonoxidative pathway provides ribose 5-phosphate and PRPP, and
much data support this viewpoint [13–16,36–38]. Consistent with
our finding that the non-oxidative pathway is the major provider
of PRPP during progression through the cell cycle, inhibition
of the non-oxidative pathway more potently inhibits cell cycle
progression and cell proliferation than inhibition of the oxidative
pathway [39].
Vizán et al. [40] assessed activity of the pentose phosphate
pathway during the cell cycle. Similar to the present study,
they found an increase in pentose phosphate pathway activity,
as assessed by an increase in the intracellular concentration
of pentose phosphates in HT29 colon carcinoma cells during
progression from G1 -phase to S-phase. However, in contrast with
the present study, they found a small increase in the activity of
glucose-6-phosphate dehydrogenase and transketolase between
G1 - and S/G2 -enriched populations obtained by cell sorting.
The data presented by this group are for a single experiment,
and enzyme activities in synchronized non-sorted cells showed
no significant difference. Our results in HCT116 cells differ
from the previous work in two ways. First, HT29 cells are
polyploid, making cell cycle analysis and sorting of S/G2 -enriched
populations difficult. Secondly, the HT29 cells were synchronized
by growth to a confluent state and then serum-starved for 24 h;
this method generally does not synchronize cells as efficiently as
a nocodazole or double-thymidine block [41].
PRPP synthetase self-associates into large complexes, with
little activity in complexes of fewer than 16 or 32 subunits
[6–8]. Phosphate is required for subunit association induced by
magnesium or MgATP, making enzyme activity dependent on the
Pi concentration [8]. Three highly homologous isoforms of PRPP
synthetase exist, i.e. PRSI, PRSII and PRSIII; PRSI and PRSII are
expressed in nearly all tissues, whereas PRSIII is limited to testis
[42,43]. The K a of human PRSI and PRSII for phosphate is ∼ 0.8
and 2.1 mM respectively [44,45]. Using the previously noted
intracellular volume of ∼ 0.9 pl for HTC116 cells [29], the data
from Figure 4(b) would suggest a total Pi concentration in mid-G1
of 6.3 mM. However, it is well recognized that measuring total
Pi in cell extracts overestimates the free intracellular phosphate
concentration severalfold [46,47], suggesting a free phosphate
concentration of <2 mM. Since this concentration is near the K a of
PRPP synthetase for phosphate, even a small change in phosphate
concentration could have a profound effect on enzyme activity.
Thus the 43 % increase in total Pi we observed between mid-G1
and G1 /S could substantially increase PRPP synthetase activity,
and we found a 2.5-fold increase in enzyme activity when the
assay was performed in the absence of added phosphate. This
increase in enzyme activity cannot be attributed to an increase
in enzyme amount between mid-G1 and G1 /S, because enzyme
activity was the same when assessed under optimal conditions, i.e.
at a high Pi concentration. The increased enzyme activity probably
explains the 3-fold increase in PRPP production and content that
we observed as cells moved from G1 -phase to S-phase. Other
studies have shown in cultured cells and rat liver slices that PRPP
synthetase activity, the intracellular PRPP concentration and rates
of de novo purine synthesis are dependent on the phosphate
concentration in the medium [17,48,49].
The basis for the increase in cellular phosphate between mid-G1
and G1 /S could be attributed, at least partially, to a 25 % increase in
the rate of phosphate uptake. Although this increase in phosphate
uptake may appear modest, a considerable increase in cellular
phosphate could occur over several hours. We cannot rule out other
c The Authors Journal compilation c 2013 Biochemical Society
98
A. Fridman and others
possible sources of phosphate contributing to increased cellular
phosphate, such as breakdown of intracellular polyphosphates
[50], but determining the precise basis of the increase in phosphate
between G1 -phase and S-phase was beyond the scope of the
present study.
We conclude that the increase in PRPP synthetase activity
between G1 -phase and S-phase can be attributed to an increase
in intracellular phosphate concentration, and allows for increased
purine synthesis during cell cycle progression. Since the reactions
of the non-oxidative pentose phosphate pathway are reversible,
increased PRPP synthetase activity would be expected to shift
the equilibrium towards ribose 5-phosphate production. Thus
phosphate activation of PRPP synthetase would increase flux
through the non-oxidative pentose phosphate pathway, as we
observed. Our data suggest that agents inhibiting the nonoxidative pentose phosphate pathway or phosphate transport
would impair cell cycle progression, and previous studies have
shown that inhibitors of the non-oxidative pentose phosphate
pathway inhibit growth of cultured pancreatic adenocarcinoma
cells and Ehrlich ascites tumour cells in mice [16,39,51]. Such
agents could serve as novel drugs for treating hyperproliferative
diseases.
AUTHOR CONTRIBUTION
Alla Fridman, Arindam Saha, Adriano Chan and Darren Casteel performed the experiments.
Renate Pilz helped to plan experiments, analysed data and assisted in writing the paper.
Gerry Boss conceived the idea, provided overall direction and wrote much of the paper.
FUNDING
This work was supported, in part, by the Department of Medicine, University of California,
San Diego.
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Received 28 January 2013/3 June 2013; accepted 4 June 2013
Published as BJ Immediate Publication 4 June 2013, doi:10.1042/BJ20130153
c The Authors Journal compilation c 2013 Biochemical Society
Biochem. J. (2013) 454, 91–99 (Printed in Great Britain)
doi:10.1042/BJ20130153
SUPPLEMENTARY ONLINE DATA
Cell cycle regulation of purine synthesis by phosphoribosyl pyrophosphate
and inorganic phosphate
Alla FRIDMAN*1 , Arindam SAHA*1 , Adriano CHAN*, Darren E. CASTEEL*, Renate B. PILZ* and Gerry R. BOSS*2
*Department of Medicine, University of California, San Diego, 9500 Gilman Drive, La Jolla, CA 92093-0652, U.S.A.
Table S2 Activities of non-oxidative pentose phosphate pathway enzymes
and glucose-6-phosphate dehydrogenase at mid-G1 and G1 /S
Cells were released from a nocodazole block and 4 and 8 h later, when they were in mid-G1
or at G1 /S respectively, they were extracted under non-denaturing conditions and enzyme
activities were measured as described in the Experimental section of the main text. Results
are means +
− S.E.M. for three independent experiments performed in duplicate, expressed in
nmol/min per mg of protein. None of the differences in enzyme activities were statistically
significant between the two cell cycle phases.
Figure S1 De novo purine synthesis and intracellular phosphate
concentration in COS-7 cells
Enzyme
Mid-G1
G1 /S
Glucose-6-phosphate dehydrogenase
Aldolase
Fructose bisphosphatase
Phosphofructokinase
Transaldolase
Transketolase
774 +
− 109
18.8 +
− 5.35
5.62 +
− 2.68
235 +
− 87.9
0.97 +
− 0.43
1.23 +
− 0.56
678 +
− 109
14.8 +
− 5.38
3.90 +
− 2.50
230 +
− 109
0.91 +
− 0.39
1.17 +
− 0.62
COS-7 cells were arrested in mitosis with nocodazole as described in the Experimental section
of the main text. At 4 and 8 h after release from the block (mid-G1 and G1 /S respectively), de novo
purine synthesis was measured following [14 C]formate incorporation into all cellular purines
(closed bars), and total intracellular Pi was measured using Malachite Green as described in
the Experimental section of the main text (open bars). Results are means +
− S.E.M., with each
experiment conducted twice on duplicate samples.
Table S1
G1 /S
Intracellular concentration of purine nucleotides at mid-G1 and
Cells were released from a nocodazole block, and 4 and 8 h later, when they were in mid-G1 or at
G1 /S respectively, they were extracted in acid and intracellular purine nucleotides were measured
by HPLC as described in the Experimental section of the main text. Results are means +
− S.E.M.
for three independent experiments performed in duplicate, expressed in nmol/106 cells. None
of the differences for any of the nucleotides were statistically significant between the two cell
cycle phases.
Purine nucleotide
Mid-G1
G1 /S
AMP
ADP
ATP
GMP
GDP
GTP
0.03 +
− 0.005
0.36 +
− 0.07
4.2 +
− 0.64
0.02 +
− 0.006
0.23 +
− 0.03
2.75 +
− 0.15
0.03 +
− 0.008
0.40 +
− 0.08
4.41 +
− 0.68
0.04 +
− 0.007
0.22 +
− 0.04
2.71 +
− 0.25
1
2
These authors contributed equally to this work.
To whom correspondence should be addressed (email [email protected]).
c The Authors Journal compilation c 2013 Biochemical Society
A. Fridman and others
Figure S2
Oxidative and non-oxidative pentose phosphate pathways showing PRPP production
Reactions of the oxidative pathway are shown on the left-hand side, and reactions of the non-oxidative pathway are shown on the right-hand side, with early steps of glycolysis shown in the middle.
PRPP can be produced from either the oxidative or non-oxidative pentose phosphate pathway, and is a limiting substrate for purine synthesis via either the de novo or salvage pathways. -P, phosphate;
-P2 , bisphosphate.
Received 28 January 2013/3 June 2013; accepted 4 June 2013
Published as BJ Immediate Publication 4 June 2013, doi:10.1042/BJ20130153
c The Authors Journal compilation c 2013 Biochemical Society