Intracellular Mg2+ regulates
ADP phosphorylation and adenine nucleotide synthesis in human
erythrocytes
Sarah
Page,
Michael
Salem, and
Maren R.
Laughlin
Departments of Surgery and Physiology, The George Washington
University Medical Center, Washington, District of Columbia 20037
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ABSTRACT |
13C- and
31P-NMR were used in methylene
blue-treated human erythrocytes to determine the dependence on
intracellular Mg2+ concentration
([Mg2+]i)
of the pentose phosphate pathway (PPP), the glycolytic pathway, and
adenine nucleotide synthesis. The PPP flux had an
[Mg2+]i
at half-maximal velocity
([Mg2+]i,0.5)
of 0.02 mM, well below the physiological range (0.2-0.7 mM). Flux
through the PPP was reduced at higher
[Mg2+]i
as flux through phosphofructokinase was increased
([Mg2+]i,0.5 = 0.16 mM).
[Mg2+]i,0.5
of phosphoglycerate kinase flux, which equals net ADP phosphorylation rate, was 0.27 mM, well within the physiological
[Mg2+]i
range. The rate of adenine nucleotide synthesis from
[2-13C]glucose-derived
ribose 5-phosphate and exogenous adenine also exhibited dependence on
[Mg2+]i
but was not saturable up to 1.6 mM. Therefore, net flux through the PPP
and glycolytic pathways in erythrocytes is not strongly dependent on
[Mg2+]i
at physiological ion concentrations, but both ADP phosphorylation and
adenine nucleotide synthesis are likely to be regulated by normal
fluctuations in
[Mg2+]i.
glycolysis; pentose phosphate pathway; adenosine
5'-triphosphate; carbon-13 nuclear magnetic resonance; metabolic
regulation; A-23187
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INTRODUCTION |
SEVERAL STUDIES INDICATE that plasma free ionized
magnesium can fall during severe illness and after cardiac surgery (1, 22, 23) and that clinical administration of
MgSO4 improves patient outcome (3,
10). The intracellular magnesium concentration ([Mg2+]i)
of some cell types, such as lymphocytes (2) and smooth muscle cells
(28), is sensitive to the extracellular magnesium concentration, and
red blood cell
[Mg2+]i
can fall after prolonged magnesium deficiency (21). Metabolic fluxes
may be altered in those pathways that experience regulation via
magnesium concentration, especially those for which MgATP or MgADP is a
substrate. Even though the magnesium requirement for some of the
individual purified enzymes of glycolysis has been reported (6, 17,
20), there are few studies of this requirement in the intact pathways
found in living tissue. It has become clear that metabolic regulation
experienced by a cell in vivo is a distributed phenomenon, which is
sensitive to the fluxes through coupled pathways in addition to
multiple metabolite and effector concentrations, covalent enzyme
modification, gene expression, and protein synthesis (4). Therefore, it
is a poor assumption that the flux through an intact metabolic pathway
will respond as though it were a solution of purified enzymes to
changes in the concentration of a single effector. The erythrocyte
provides a simple model system in which to study the effects of
magnesium on an entire metabolic pathway in the absence of mitochondria or protein synthesis.
A second goal is to determine whether the measured effective
concentration range of the regulating molecule falls within the range
found in the intact cell. Recent findings from
13C- and
31P-NMR kinetic studies of human
erythrocytes consuming
[2-13C]glucose in the
presence of variable magnesium concentration showed that the
[Mg2+]i
at half-maximal velocity
([Mg2+]i,0.5)
for the entire glycolytic pathway is 0.03 mM (14).
[Mg2+]i
in these cells is between 0.2 mM in oxygenated and 0.7 mM in deoxygenated cells (9, 18), and therefore
[Mg2+]i
is unlikely to be regulatory for glycolysis under normal, unstressed conditions. These experiments employed the divalent ionophore A-23187
to allow equilibration between intracellular and extracellular Mg2+, a
31P-NMR assay of ATP-bound
Mg2+ to determine
[Mg2+]i,
and 13C-NMR to monitor metabolism
of the 13C-enriched glucose
substrate (9, 14, 25).
In these earlier experiments the pentose phosphate pathway activity
(PPP) and rate of ADP phosphorylation were both too low to accurately
measure their magnesium dependence. The PPP is important in
erythrocytes to provide the NADPH used for glutathione reduction, which
is in turn used to repair oxidized cellular components (26, 29). This
may be especially important during inflammation, which is associated
with increased release of reactive oxygen species and nitric oxide (12,
24). Therefore, the PPP is likely to be activated under the same
conditions in which plasma magnesium is reduced. In the current study,
the PPP was maximally activated with the redox dye methylene blue. This
dye oxidizes NADPH to produce
NADP+, the substrate for the
rate-limiting enzyme glucose-6-phosphate dehydrogenase (G-6-PDH) (3,
22). The putative sites for magnesium regulation are 6-phosphogluconate
dehydrogenase and transketolase, which contains a tightly bound thiamin
pyrophosphate and requires Mg2+ as
a cofactor (11, 27).
ADP phosphorylation and the synthesis of adenine nucleotides from
exogenous adenine and
[2-13C]glucose were
also activated in the presence of methylene blue, allowing us to
measure the magnesium dependence of these fluxes. In the erythrocyte,
the net rate of ADP phosphorylation can be less than the rate of flux
through the glycolytic pathway, because the 2,3-diphosphoglycerate
(2,3-DPG) shunt bypasses the enzyme phosphoglycerate kinase (PGK) to
produce 2,3-DPG rather than ATP. The rate of
[2-13C]2,3-DPG
appearance can be directly measured in the
13C-NMR experiment, and
subtraction of this rate from net flux through the glycolytic pathway
yields PGK flux, which is equal to the net rate of ADP phosphorylation.
Likewise, production of
[1'-13C]adenine
nucleotide from
[1-13C]ribose
5-phosphate (R-5-P) produced in the
PPP can be directly measured in the
13C-NMR experiment.
These fluxes can be plotted against the measured
[Mg2+]i
to yield saturating curves that are reminiscent of simple
Michaelis-Menten kinetics (velocity = [S] × Vmax/[S] + KM, where
[S] is the substrate concentration,
Vmax is the
maximum velocity, and
KM is the
Michaelis-Menten constant). Clearly, the Michaelis-Menten formalism can
be strictly applied only to well-controlled measurements of the initial
reaction rates of single-substrate reactions catalyzed by purified
enzymes. This is far from the case when the metabolic fluxes in
an intact cell are measured, where alterations of one effector can
result in changes in many substrate and product concentrations, as well as in energy-dependent processes such as membrane transport.
Michaelis-Menten formalism is, however, useful for describing the
general characteristics of control by an effector. Therefore,
much of the following data has been fit to this equation to yield an
overall Vmax.
Rather than a KM,
an analogous value, the intracellular magnesium ion concentration at
half-maximal velocity,
[Mg2+]i,0.5,
will be reported.
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METHODS |
Methods were essentially those found in Refs. 9, 14, and 25. Blood (5 ml) drawn from healthy volunteers who had given informed consent was
centrifuged, and erythrocytes were washed with PBS and incubated at 3%
hematocrit for 60 min at 37°C in a solution containing 110 mM NaCl,
5 mM KCl, 40 mM HEPES, 15 mM Na2HPO4,
5 mM adenine, 10 mM glucose, 6.6 µM methylene blue, pH 7.4, and
MgCl2 between 0 and 1.6 mM.
A-23187 (6 µM), a divalent ionophore, was included to allow
equilibration of magnesium ion across the cell membrane, and 1 mM EGTA
was included to bind residual calcium. This solution was found to
promote maximal glucose use. Magnesium ion concentration in the buffer
was determined by ion-sensitive electrode with an appropriately
calibrated clinical analyzer (NOVA Biomedical, Waltham, MA). Cells were
subsequently washed twice in the same buffer without glucose and
resuspended at 50% hematocrit with 1 mM glucose, oxygenated in 100%
O2 on ice, and placed in a 10-mm
tube for the NMR experiment.
After an initial 20-min 31P-NMR
spectrum (34°C, 60° pulses, 2-s relaxation delays, composite
pulse decoupling), 10 mM
[2-13C]glucose was
added, and glucose, lactate,
PO2, pH,
and Mg2+ were measured with
commercial analyzers from NOVA Biomedical. Fifteen 10-min
13C-NMR spectra were taken (same
parameters), followed by a final two
31P spectra. The
31P-NMR spectra were used to
calculate
[Mg2+]i
from the chemical shift difference between the
- and
-ATP peaks.
The intensities of the following
13C-labeled metabolites were
measured:
[2-13C]glucose,
[2-13C]lactate,
[3-13C]lactate,
[2-13C] 2,3-DPG,
[3-13C]2,3-DPG, and
[1'-13C]adenine
nucleotide.
Cells were frozen in liquid N2,
stored at
50°C, and extracted with perchloric acid.
13C-NMR spectra of extracts were
taken for determination of 13C in
different molecules, and 1H-NMR
spectra were taken for measurement of the enrichment with 13C at C-3 of lactate [1.3
parts per million (ppm) in 1H
spectra] and C-1' in ribose of ATP (6.11 ppm). Enrichment
was calculated as the ratio of the area under the
1H-13C
doublet to the total combined area, including the
1H-12C
central singlet (13). 2,3-DPG was measured in trichloroacetic acid
extracts (Kit 665-PA, Sigma Chemical, St. Louis, MO).
13C-NMR data were analyzed as in
Ref. 14. Because of the spectral broadening that occasionally occurred
at late times in the experiments, [Mg2+]i
was calculated from ATP resonances in the initial
31P-NMR spectrum (9, 14), and all
reported rate data are calculated as the slope at 30 min of
13C-NMR peak areas plotted against
time. All calculations are shown in the
APPENDIX.
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RESULTS |
Glucose consumption was measured as the slope at 30 min of
[2-13C]glucose NMR
peak area plotted vs. time, and the results are plotted against the
initial
[Mg2+]i
for each experiment in Fig.
1. Although these data
measured in a living system show a great deal of scatter, it is clear
that [Mg2+]i
is participating in the regulation of glycolysis flux at the very low
end of the physiological range of 0.2-0.7 mM. Overall Vmax and
[Mg2+]i,0.5
for glucose use are both slightly higher than found in the absence of
methylene blue (Ref. 14, Table 1).

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Fig. 1.
Rate of glucose utilization measured at 30 min of the experiment from
in situ 13C-NMR spectra as
function of intracellular Mg2+
concentration
([Mg2+]i)
in human erythrocytes. , Experiments conducted without ionophore;
, experiments conducted with ionophore. Data are shown with best fit
to the Michaelis-Menten equation, which resulted in maximal velocity
(Vmax) = 0.57 µmol · min 1 · g
hemoglobin 1 and
half-maximal velocity of [Mg2+]i
([Mg2+]i,0.5) = 0.07 mM.
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Figure 2 shows PC, the fractional flux through the PPP,
which falls exponentially as a function of
[Mg2+]i.
PPP activity is shown in Fig. 3. The dotted line is
derived from the functions that represent the best fits in Figs. 1 and 2, and the circles are derived from the individual rates of glucose utilization. The dotted line therefore does not represent a best fit of
the individual data points but does suggest a shape for the magnesium
dependence of the PPP. The estimated
[Mg2+]i,0.5
of 0.02 mM is well below the physiological range. However, it appears
that the flux through the PPP is decreased in the physiological range
between 0.2 and 0.7 mM
[Mg2+]i.
This is unlikely to be due to direct inhibition of this pathway by
magnesium but may be a response to activation of an enzyme in the
competing glycolytic pathway by magnesium in this
range.

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Fig. 2.
PC [fractional pentose phosphate pathway (PPP) activity] as
a function of
[Mg2+]i.
PC was calculated from the ratio of
2,33-13C]diphosphoglycerate
(DPG) to
2,3-[2-13C]DPG in
13C-NMR spectra of perchloric acid
extracts of erythrocytes. Data were fit to exponential shown,
Ae Bx + C, where
A = 0.364, B = 4.53, and C = 0.264. Symbols are the same as for Fig.
1.
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Fig. 3.
Flux through PPP as a function of
[Mg2+]i.
Dotted line represents PPP calculated from fitted curves of Figs. 1 and
2 and Eq. A3. Also shown is PPP
calculated from curve of Fig. 2 and individual data points of Fig. 1
(circles). No attempt was made to fit these data to simple function.
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Figure 4 shows the flux through the portion
of the glycolytic pathway through phosphofructokinase (PFK), i.e.,
between fructose 6-phosphate (F-6-P)
and glyceraldehyde 3-phosphate (GAP). The circles are calculated from
the individual data points in Fig. 1 and the exponential curve in Fig.
2, and the dotted line is the least-squares fit to the Michaelis-Menten
equation (Table 1). Flux through the span that contains
glyceraldehyde-3-phosphate dehydrogenase (GAPDH), GAP-lactate,
expressed in terms of micromole glucose equivalents per minute per gram
hemoglobin, is shown in Fig. 5. The
fractional fluxes through the 2,3-DPG shunt and the enzyme this shunt
bypasses, PGK, were measured from the initial rates of the fitted time
courses of
2,3-[2-13C]DPG and
[2-13C]lactate, and
are also shown in Fig. 5 (14). The behavior of each of these fluxes was
fit to the Michaelis-Menten equation, although this fit required an
offset for
[Mg2+]i
in the case of PGK. Flux through the enzymes of the 2,3-DPG shunt,
2,3-DPG mutase, and 2,3-DPG phosphatase had little or no dependence on
[Mg2+]i
in the physiological range, and the derived kinetic constants are very
similar to those obtained in the absence of methylene blue (Table 1).
On the other hand, PGK had no apparent activity until
[Mg2+]i
was above 0.13 mM, and then had a
Vmax of 0.19 µmol · min
1 · g
hemoglobin
1 and a
[Mg2+]i,0.5
of between 0.2 and 0.3 mM. This indicates that net glycolytic ADP
phosphorylation has a magnesium ion dependence that falls within the
physiological range of concentration.

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Fig. 4.
Phosphofructokinase flux as function of
[Mg2+]i.
Flux through glycolytic span between fructose 6-phosphate
(F-6-P) and glyceraldehyde
3-phosphate (GAP) was calculated from Eq. A4. Circles were calculated as for Fig. 3 and dotted
line is best fit to Michaelis-Menten equation, where
Vmax = 0.45 µmol · min 1 · g
hemoglobin 1 and
[Mg2+]i,0.5 = 0.16 mM.
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Fig. 5.
GAP lactate flux, 2,3-DPG turnover, and phosphoglycerate kinase (PGK)
activity as function of
[Mg2+]i.
For clarity, GAP lactate flux (solid curve) was calculated with
Eq. A5 from 2 ideal curves of Figs. 1
and 2. 2,3-DPG turnover ( ) and PGK flux ( ) were calculated from
GAP lactate flux and fractional fluxes measured from in situ
13C-NMR data. 2,3-DPG shunt has
Vmax = 0.32 µmol · min 1 · g
hemoglobin 1 and
[Mg2+]i,0.5 = 0.04 mM. PGK has maximum activity of 0.19 µmol · min 1 · g
hemoglobin 1 and
[Mg2+]i,0.5 = 0.27 mM.
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There was detectable 13C label in
the C1' position of the ribose in adenine nucleotides, which
resonates at 88 ppm in the in situ experiment and followed a saturating
curve in time (Fig. 6). This indicates that
at least part of the adenine nucleotide pool is in rapid turnover
(synthesis and breakdown rates are equal), because at steady state, net
synthesis would lead to a signal that is constantly increasing with
time. The total 13C in the
C-1' position at the end of the experiment was measured in
extracts and is shown as a function of
[Mg2+]i
in Fig.
7A. The
fractional enrichment with 13C at
this site was measured with 1H-NMR
in extracts and was also a strong exponential function of [Mg2+]i.
[1'-13C]adenine
nucleotide fractional enrichment equals 0.301{1
exp(
6.57 [Mg2+]i)}
0.0012 (data not shown). The rate of formation of
[1'-13C]adenine
nucleotide could be calculated for seven experiments from the initial
slope of the change in time of this signal and is plotted in Fig.
7B.

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Fig. 6.
Corrected
[1'-13C]adenine
nucleotide at 88 ppm over time in representative in situ
13C-NMR experiment. Curve was fit
to Eq. A7.
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Fig. 7.
Total
[1'-13C]adenine
nucleotide and its rate of appearance plotted as function of
[Mg2+]i.
A: total
[1'-13C]adenine
nucleotide was measured in 13C-NMR
spectra of erythrocytes extracted at end of experiments.
B: initial rate of
[1-13C]ribose
5-phosphate incorporation into adenine nucleotides calculated from in
situ 13C-NMR spectra for 7 experiments.
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The fractional enrichment of the
[2-13C]G-6-P
pool was calculated for each experiment from Eq. A8 in the APPENDIX and
is shown in Fig.
8A.
G-6-P is the original substrate of the
PPP, in which the ribose moiety of adenine nucleotide is made. The
calculation does not account for dilution of the
R-5-P pool with unlabeled material and
therefore represents the maximal precursor fractional enrichment. This
fractional enrichment could then be used to calculate the minimum rates
of adenine nucleotide turnover at each
[Mg2+]i,
which are shown in Fig. 8B. Adenine
nucleotide turnover was a strong function of
[Mg2+]i
and was not saturated over the entire concentration range studied. The
least-squares fit to a straight line yields the equation adenine nucleotide turnover equals 0.04 × [Mg2+]i + 0.09, with a correlation of
R2 equal to
0.494.

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Fig. 8.
[2-13C]glucose
6-phosphate (G-6-P) enrichment and
rate of adenine nucleotide turnover.
A:
[2-13C]G-6-P
enrichment was calculated from PC (data and fit of Fig. 2) using
Eq. A8.
B: rate of adenine nucleotide turnover
calculated from Eq. A6 and plotted
against
[Mg2+]i.
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DISCUSSION |
Gupta et al. (9) used 31P to
measure the
[Mg2+]i
in fully oxygenated human erythrocytes (0.25 mM) and in fully
deoxygenated cells (0.67 mM). Under the current experimental
conditions, the [Mg2+]i
in fully oxygenated cells was slightly lower, about 0.17 ± 0.02 mM
(n = 3). The
[Mg2+]i,0.5
for PPP was 0.02 mM, for PGK it was about 0.27 mM, and that for adenine
nucleotide turnover was too high to determine with our experimental
magnesium range. At normal physiological concentrations, then,
[Mg2+]i
is important in the human erythrocyte for the regulation of PGK
activity and adenine nucleotide synthesis from exogenous glucose and
adenine, but is likely to be too high to directly affect PPP activity.
This result compares well with the
KM for
Mg2+ measured in purified PGK from
human erythrocytes, which is 0.3 mM (20). In the erythrocyte, PGK can
be bypassed by the 2,3-DPG shunt and can therefore experience much less
flux than that of glucose through the entire pathway. However, this
step is very important, because PGK flux is equal to the net rate of
ADP phosphorylation (see Fig. 9). These
cells should therefore be able to increase both ADP phosphorylation and
ATP utilization purely as a function of
[Mg2+]i
as they move from the arterial to the venous circulation.

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Fig. 9.
Pathways of glucose utilization in the erythrocyte.
13C-NMR visible molecules are
shown in bold typeface and calculated fluxes are indicated as
1) PPP,
2) glycolytic span between
F-6-P and GAP,
3) glycolytic span between GAP and
lactate, 4) PGK,
5) 2,3-DPG shunt, and
6) adenine nucleotide synthesis. HK,
hexokinase; PFK, phosphofructokinase; GAPDH, glyceraldehyde-3-phosphate
dehydrogenase; PGK, phosphoglycerate kinase; PK, pyruvate kinase;
R-5-P, ribose 5-phosphate. Because 2 ATP-glucose are hydrolyzed
early in pathway and 2 ATP-glucose are produced by PK, flux through PGK
is approximately equal to net rate of ADP phosphorylation.
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It is not clear what the physiological role might be for this
additional capacity for ATP generation in the anaerobic state. Glycolysis is increased in the anaerobic state (7, 25). This may be due
to the fact that the free forms of 2,3-DPG, which inhibit several
glycolytic enzymes including hexokinase (HK), PFK, PGK, and pyruvate
kinase (20), bind tightly to deoxyhemoglobin and are decreased from
62% of total 2,3-DPG in the oxygenated state to 20% in the
deoxygenated cell (18). Increased glycolysis and ATP formation may be
important for ion pumping (16). Certainly the ATP-dependent
Na+/Mg2+
exchange is activated both by artificial elevation of
[Mg2+]i
and in the anaerobic cell, where
[Mg2+]i
is also elevated (5, 9, 18).
The Vmax for
total glucose utilization increased by 30% in the presence of
methylene blue, and its
[Mg2+]i,0.5
may also have been slightly altered (0.03 mM in control vs. 0.07 mM
with methylene blue). The increase in
Vmax was due entirely to increased PPP flux, because the calculated
Vmax of the
F-6-P
GAP span does not change on
addition of a redox stress. This implies that we have been able to
measure the true maximum activity of this portion of the glycolytic
pathway. The apparent magnesium dependence of the
F-6-P
GAP span
([Mg2+]i,0.5)
is, however, elevated in the presence of methylene blue (Table 1). This
could mean that the magnesium dependence of the regulatory enzymes in
this portion of the glycolytic pathway were altered or that the rate
control for glycolysis was no longer found in the
F-6-P
GAP span. It could also mean
that another factor besides
[Mg2+]i
becomes important for setting the flux rate. The last explanation is
likely because of competition between PFK and activated G-6-PDH for the
combined G-6-P and
F-6-P substrate pool.
G-6-P and
F-6-P are thought to be in
equilibrium. The
KM of purified
PFK for F-6-P is about 0.3-0.5 mM
(17, 20). The normal concentration of F-6-P in human erythrocytes is only
10-20 µM (8, 19), and it would likely be decreased by flux of
G-6-P into the activated PPP. PPP flux
should be far less dependent on
G-6-P+F-6-P
concentration, because the
KM of G-6-PDH for
G-6-P is 7.4 µM, well below the cellular concentration of about 40 µM (15, 19).
The PPP also has a much lower dependence on
[Mg2+]i
than the F-6-P
GAP span of
glycolysis under our conditions
([Mg2+]i,0.5
~0.02 mM). The fact that PC falls with increasing
[Mg2+]i
(Fig. 2) means that each individual glucose skeleton turns through the
PPP cycle more times at lower
[Mg2+]i
than it does at higher
[Mg2+]i
when glucose uptake and PFK flux are elevated. This very low requirement of the PPP for Mg2+
supports the work of Thorburn and Kuchel (26), who concluded using
model systems that HK is rate limiting for PPP activity in oxidatively
stressed human erythrocytes. We do not, however, concur with the
literature value of
KM for
Mg2+ of 1.0 mM for purified human
erythrocyte HK, because the rate-limiting step has a
[Mg2+]i,0.5
equal to 0.02 mM whether or not the PPP has been activated (6). Rather,
our results imply that the PPP enzymes that require magnesium
(presumably 6-phosphogluconate dehydrogenase and transketolase, Refs.
11 and 27) have
[Mg2+]i,0.5
values that are much lower than 0.02 mM.
Competition for substrate between PFK and G-6-PDH would
likely depress flux through PFK at low
[Mg2+]i
levels, because of the low
[Mg2+]i,0.5
of the PPP and the low
KM of G-6-PDH for
G-6-P. PPP activation would therefore
cause the observed increase in the measured apparent [Mg2+]i,0.5
for PFK. As
[Mg2+]i
is increased, the flux through the entire glycolytic pathway increases,
perhaps due to activated PGK activity
([Mg2+]i,0.5 = 0.27 mM). This may contribute to the apparent decrease in flux
through the PPP seen in the range of 0.15-0.6 mM
[Mg2+]i.
The rate of turnover of the adenine nucleotide pool continues to
increase with
[Mg2+]i
well after glucose uptake and PPP have reached their plateau. The curve
of adenine nucleotide turnover does not saturate with [Mg2+]i,
and therefore it is not possible to estimate values for
Vmax or
[Mg2+]i,0.5.
There are few measurements because adenine nucleotide pool turnover was
easily detected at high
[Mg2+]i,
but the signal-to-noise ratio of the
13C peak at 88 ppm in the in situ
spectra was often poor at low [Mg2+]i
(see Fig. 7A). The adenine
nucleotide turnover rate was calculated by assuming that the
13C enrichment of the
5-[1-13C]phosphoribosyl-1-pyrophosphate
(PRPP), the immediate precursor of newly synthesized AMP in
erythrocytes, is the same as that of
[2-13C]G-6-P
(shown in Fig. 8A) (20). This is a
reasonable assumption at isotopic equilibrium, because the only point
of label dilution between G-6-P and
PRPP is at R-5-P and is due to
exchange with the ribose moiety of adenine nucleotides. As shown in
Fig. 6, the turnover of the PRPP pool is so rapid that it reached
isotopic equilibrium well before the end of the experiment in all cases where the turnover rate could be measured. At the early time points, dilution of the
[1-13C]PRPP enrichment
would be greater at the higher rates of adenine nucleotide pool
turnover and would tend to make the slope of turnover vs.
[Mg2+]i
even steeper than shown in Fig. 8B.
The rapid turnover of adenine nucleotide pool in these experiments has
an important impact on the calculation of PC. In the prior studies PC
was calculated from the ratio of
[3-13C]lactate/[2-13C]lactate
(14, 25). In the current experiments, this ratio measured in blood
extracts was always lower than the similar ratio, 2,3-[3-13C]DPG/2,3-[2-13C]DPG,
even though 2,3-DPG and lactate have the same precursor pool. This is
because of the turnover of the adenine nucleotides, which exchanges
[1-13C]R-5-P
with the unlabeled ribose moiety and dilutes the enrichment of
[3-13C]lactate at
early times. Label appearing as
[2-13C]lactate is not
similarly diluted, because it does not pass through any unique, large
metabolic pool. This problem of dilution only occurs when the
13C ratio of the growing lactate
pool is used to calculate PC; the 2,3-DPG pool turns over constantly
and quickly reaches a steady-state enrichment. Therefore, an
end-product pool-like lactate that maintains a "history" of all
tracer events during the experiment is not as good an indicator of PPP
activity as a metabolically active pool like 2,3-DPG. The sum of label
in C-1' of adenine nucleotides and
[3-13C]lactate could
be divided by
[2-13C]lactate to
obtain a number similar to the 2,3-DPG ratio (Eq. A2 in APPENDIX).
It is important to point out that although the reported fluxes through
different parts of the glycolytic pathway could be measured somewhat
independently, it is likely that the actual fluxes are very dependent
on each other. For instance, the
Vmax of ADP
phosphorylation (PGK flux) may be dependent on the rate of adenine
nucleotide turnover, which is dependent on adequate PPP activity.
Vmax of PGK is,
in fact, elevated by methylene blue (Table 1). Changes in
[Mg2+]i
per se may also alter fluxes independent of enzyme regulation. The most
obvious manifestation of this is a potentially increased need for ATP
(i.e., PGK flux) to fuel
Na+/Mg2+
exchange (5).
In summary, the magnesium ion dependence of the pathways of glucose
utilization was determined during activation of the PPP by methylene
blue in human erythrocytes. The measured
[Mg2+]i,0.5
values indicate that physiological
[Mg2+]i
is too high to play an important regulatory role for glucose uptake or
PPP activity, although
[Mg2+]i
may be involved in the competition between G-6-PDH (the first committed
step in the PPP) and PFK for substrate. Likewise, 2,3-DPG turnover is
independent of
[Mg2+]i.
Net ADP phosphorylation, equal to PGK activity, has a
[Mg2+]i,0.5
of 0.27 mM and is likely to be regulated by changes in [Mg2+]i
in the physiological range. The rate of adenine nucleotide synthesis
from exogenous adenine and glucose-derived
R-5-P is also highly dependent on
[Mg2+]i
in the physiological range of 0.2-0.7 mM. We have therefore determined that erythrocyte ATP production, both phosphorylation and
adenine nucleotide synthesis, can be regulated by
[Mg2+]i
in its normal physiological range.
 |
APPENDIX |
The calculations are detailed in Refs. 14 and 25 and are presented
briefly here with emphasis only on equations unique to the current
study. Figure 9 shows the pathways of interest, where
13C-NMR visible metabolic pools,
glucose, lactate, 2,3-DPG, and adenine nucleotides are shown in bold.
Fluxes through six segments of the pathway could be estimated and these
are numbered in Fig. 9.
Overall glucose utilization is measured as the change in the area of
the [2-13C]glucose
peak over time (14, 25). PC is defined as the fractional flux through
the PPP
|
(A1)
|
In
other studies (14, 25), PC was determined from the ratio in extracts of
[3-13C]lactate/[2-13C]lactate.
In the current studies we found that where both the 2,3-DPG and lactate
ratios can be measured, the lactate ratio was always less than the
2,3-DPG ratio. This was because some of the carbon skeletons that enter
the PPP are used to produce [1-13C]R-5-P,
which becomes incorporated into adenine nucleotides (resonance found at
88 ppm) under the conditions of high buffer adenine concentration and
elevated PPP activity. This tends to reduce the total amount of
[3-13C]lactate
produced during the entire experiment. 2,3-DPG is a metabolic pool that
reaches a steady-state isotopic enrichment and therefore does not
exhibit this problem. Whenever both could be measured, we found
Eq. A2 to be true
|
(A2)
|
PC was calculated from Eq. A1 or A2, plotted vs.
[Mg2+]i,
and the result was fit using a least-squares algorithm to an
exponential curve. This curve was used in the following equations. PPP
activity is defined as
|
(A3)
|
Two
thirds of the material in the PPP reenters the glycolytic pathway as
F-6-P and proceeds through PFK,
defined as the span between F-6-P and
GAP
|
(A4)
|
On average, one molecule of GAP is produced for every six
G-6-P that enter the PPP, and the flux
of GAP through GAPDH can proceed to lactate through either PGK or the
2,3-DPG shunt
|
(A5)
|
The
initial rates of
[2-13C]lactate and
2,3-[2-13C]DPG
appearance yield flux 4 and
flux 5, respectively.
Flux 6, the rate of synthesis of the
adenine nucleotide pool from PPP-derived
R-5-P and exogenous adenine can be
calculated from Eq. A6 if the
fractional enrichment with 13C of
the R-5-P precursor,
[2-13C]G-6-P,
is known
|
(A6)
|
Appearance of 13C in C-1' of
the ribose moiety in adenine nucleotides (88 ppm) over time could be
fit to Eq. A7. Most adenine and
guanidine nucleotides resonate in this region, although the signal is
predominantly adenine nucleotide. The shape of the curve indicates that
the concentration of 13C reaches a
plateau when the metabolic pool is in isotopic equilibrium. This means
that the rates of synthesis and degradation of the labeled product are
equal. The initial rate of
[1'-13C]adenine
nucleotide appearance was taken as the derivative of Eq. A7 at 10 min after the signal at
88 ppm was observed
|
(A7)
|
The
fractional enrichment of
[2-13C]G-6-P
can be calculated with Eq. A8
|
(A8)
|
The following outlines the derivations of the expressions for
[2-13C]G-6-P
fractional enrichment and the quantity PC. The calculation of PC is
based on the assumption that the fractional enrichment of
[1-13C]F-6-P
leaving the PPP is the same as the
[2-13C]G-6-P
that was its precursor (25). The flow of
G-6-P into the PPP is defined as 3PC × HK (Eq. A3), where HK is
hexokinase flux. The average rate of
F-6-P produced from the PPP is taken to be 2PC × HK, by the following stoichiometry
|
(A9)
|
The combined
G-6-P+F-6-P
sugar 6-phosphate (S-6-P) pool has
two inputs in this model (Fig. 9), where all
[2-13C]S-6-P
arrives through HK and all
[1-13C]S-6-P
arrives from the PPP. If the enrichment of the precursor [2-13C]glucose is 1.0, the enrichment at C-2 of the S-6-P
pool, which gives rise to
[2-13C]lactate and
2,3-[2-13C]DPG,
is
|
(A10)
|
The enrichment of S-6-P at
C-1, which yields
[3-13C]lactate and
2,3-[3-13C]DPG in the
glycolytic pathway, can be calculated by assuming that at steady state
the
[1-13C]F-6-P
newly formed in the PPP has the same enrichment as the precursor
[2-13C]S-6-P
pool
|
(A11)
|
The ratio of the two enrichments yields Eq. A1
|
(A1`)
|
If the enrichment of F-6-P is less
than that of its G-6-P precursor due
to scrambling of label in the PPP, the ratio of Eq. A10 will underestimate PC and therefore the flux
through PPP. If this were the case, we would expect to see a variety of
splitting patterns in 13C-NMR
spectra of the lactate and 2,3-DPG in extracts, as observed in Ref. 25.
This splitting did not appear in the current study. If the adenine
nucleotide pool is large and its turnover is very high, the enrichment
of F-6-P will also be less than that
of G-6-P, but only at early time
points before the nucleotide pools have reached isotopic steady state.
Although every attempt was made to bring the system to metabolic steady
state before measurements were made, this may not have been the case.
If the adenine nucleotide pool expands during the experiment,
[1-13C]R-5-P
would be lost as net adenine nucleotide synthesis, and Eq. A9 could be modified
to
|
(A12)
|
In this case PC underestimates the fractional flux of GAP
from PPP by the quantity x/3. This
would require a correction of PPP flux by about 0.02 µmol · min
1 · g
hemoglobin
1 for each 1 mM
increase of ATP. In the current study, any changes in ATP appeared to
be very small (<<1 mM).
 |
FOOTNOTES |
Address for reprint requests: M. R. Laughlin, Dept. of Surgery and
Physiology, The George Washington Univ. Medical Center, Ross Hall 550, 2300 Eye St., NW, Washington, DC 20037.
Received 24 October 1997; accepted in final form 23 January 1998.
 |
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