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The Journal of Neuroscience, March 1, 2000, 20(5):1809-1821
Mechanisms of Glutamate Metabolic Signaling in Retinal Glial
(Müller) Cells
Serge
Poitry1,
Carol
Poitry-Yamate1,
Joern
Ueberfeld2,
Peter R.
MacLeish3, and
Marcos
Tsacopoulos1
1 Department of Physiology, University of Geneva
Medical School, 1211 Geneva 4, Switzerland, 2 Department of
Applied Physics, University of Geneva, 1211 Geneva 4, Switzerland, and
3 Neuroscience Institute, Morehouse School of Medicine,
Atlanta, Georgia 30310-1495
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ABSTRACT |
Retinal Müller (glial) cells metabolize glucose to lactate,
which is preferentially taken up by photoreceptor neurons as fuel for
their oxidative metabolism. We explored whether lactate supply to
neurons is a glial function controlled by neuronal signals. For this,
we used subcellular fluorescence imaging and either amperometric or
optical biosensors to monitor metabolic responses simultaneously from
mitochondrial and cytosolic compartments of individual Müller
cells from salamander retina. Our results demonstrate that lactate
production and release is controlled by the combined action of
glutamate and NH4+, both at micromolar
concentrations. Transport of glutamate by a high-affinity carrier can
produce in Müller cells a rapid rise of glutamate concentration.
In our isolated Müller cells, glutamine synthetase (GS) converted
transported glutamate to glutamine that was released. This reaction,
predominant when enough NH4+ is
available, was limited at micromolar concentrations of
NH4+, and more glutamate entered then as
substrate into the mitochondrial tricarboxylic acid cycle (TCA).
Increased production of glutamine by GS leads to increased utilization
of ATP, some of which is generated glycolytically. Methionine
sulfoximine, a specific inhibitor of GS, suppressed the stimulatory
effect of glutamate and NH4+ on
glycolysis and induced massive entry of glutamate into the TCA cycle.
The rate of glutamine production also determined the amount of pyruvate
transaminated by glutamate to alanine. Lactate, alanine, and glutamine
can be taken up and metabolized by photoreceptor neurons. We conclude
that a major function of Müller glial cells is to nourish retinal
neurons and to metabolize the neurotoxic ammonia and glutamate.
Key words:
ammonia; glutamine synthetase; biosensor; NADH; fluorescence imaging; lactate; retina
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INTRODUCTION |
There is experimental evidence
suggesting the existence of lactate trafficking from astrocytes to
neurons (for review, see Tsacopoulos and Magistretti, 1996
). For
example, lactate transport across the cytoplasmic membrane of
astrocytes and neurons was shown to be mediated by cell type-specific
monocarboxylate transporters (Bröer et al., 1997; Pellerin et
al., 1998), but probably the most convincing evidence to date for such
trafficking and utilization of lactate as fuel by neurons was provided
by quantitative studies on the metabolic fate of glucose in retinal
cells. Indeed, Poitry-Yamate et al. (1995) found that acutely isolated
Müller cells metabolize glucose to synthesize and release large
amounts of lactate. This lactate, in turn, is taken up by photoreceptor
neurons and serves as the preferred fuel for oxidative metabolism.
Moreover, because the metabolism of glucose and glycogen in
Müller cells was modulated by light absorbed by the visual
pigment of the photoreceptors, it appears that these two cells function
as metabolically coupled units and that there is a signal controlling
the transfer of substrates from glial cells to neurons.
In the honeybee retina, the exact signaling mechanism has been recently
elucidated (Tsacopoulos et al., 1997a,b): functioning photoreceptor
neurons produce and release, in a stimulus-dependent manner,
NH4+ and glutamate which,
in turn, are transported into glia. A stimulus-induced transient rise
in the intraglial concentration of
NH4+ and glutamate caused
a net activation of glycolysis by a direct and synchronous action on
three enzymatic reactions. These clear findings are possibly of
interest to vertebrate studies. Indeed, glutamate is the predominant
excitatory neurotransmitter in the mammalian retina and brain, and it
has been shown that glutamate stimulates the uptake and phosphorylation
of glucose and increases the amount of lactate released by cultured
astrocytes (Pellerin and Magistretti, 1994). However, the exact
metabolic signaling mechanism of glutamate has not been established in
vertebrates because glutamate transported into astrocytes is known to
participate as a substrate in several reactions that occur in the
cytosol or in the mitochondria (Cooper and Plum, 1987; for review, see Schousboe et al., 1997), and there was no model of extreme metabolic compartmentation equivalent to the honeybee retina. We explored metabolic signaling in acutely isolated Müller glial cells from the salamander retina because they present this high degree of metabolic compartmentation. Indeed, mitochondria are found almost exclusively in the distal processes, located in the retina near the
glutamatergic synaptic terminal of photoreceptors (Tsacopoulos et al.,
1998, their Fig. 5b). This feature permitted the use of novel techniques to monitor metabolic responses from metabolically different regions of solitary Müller cells. Here we show that glutamate, in coordination with
NH4+, exerts a strong
activation of glycolysis via the glutamine synthetase reaction. As a
result, the production and release of lactate, glutamine, and alanine
by these glial cells are modulated to meet, presumably, the
requirements of the functioning neurons. We believe that this evidence
provides a basis for interpreting the observed coupling between brain
activity and metabolism (Magistretti et al., 1999).
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MATERIALS AND METHODS |
Materials. Papain was purchased from Worthington
(Freehold, NJ), and neutral glutaminase from Applied Enzyme Technology
(Leeds, UK). 2,7-Biscarboxyethyl-5(6)-carboxyfluorescein-AM (BCECF-AM) and 5-carboxy-4',5'-dimethylfluorescein (CDMF) were obtained from Molecular Probes (Leiden, The Netherlands). Amytal (amobarbital) sodium
was from Serva Feinbiochemica (Heidelberg, Germany). All other
chemicals were purchased from Sigma (Buchs, Switzerland).
Preparation of Müller cells isolated from the salamander
retina. Tiger salamanders (Ambystoma tigrinum) were
purchased from Charles Sullivan (Nashville, TN) and were kept in a tank
filled with tap water maintained at 10°C. The procedure for cell
isolation followed that of MacLeish and Townes-Anderson (1988) with
some modifications. Salamanders were decapitated and pithed. Eyes were enucleated and transsected, and the retinas were removed and placed in
an enzyme solution that contained (in mM): 112 NaCl, 3 KCl, 0.1 CaCl2, 0.1 MgCl2, 0.5 NaH2PO4, 1 NaHCO3, 0.1 choline-Cl, 16 glucose, 10 HEPES, and
5 cysteine-HCl, and 0.001% phenol red and ~15 U/ml papain, pH 6.4. The activity of the enzyme solution was routinely checked before use by
measuring the rate of acid production from
benzoyl-L-arginine ethyl ester (see Worthington
Manual). The 16 mM glucose concentration in the
enzyme solution was to provide the retina with ample substrate during
incubation. Retinas were incubated at 22-25°C with gentle agitation
for 1 hr (or more, depending on the activity of the enzyme) and then
rinsed several times with a slightly hyperosmotic solution of similar
composition, but without cysteine and papain, and with 32 mM sucrose added. Retinas were then gently
triturated using a fire-polished Pasteur pipette previously treated
with bovine serum albumin (BSA; 0.1 mg/ml). Aliquots of the cell
suspension were then placed onto a glass coverslip coated with
salamander-specific antibody (Sal-1) serving as adhesion substrate for
the cells (MacLeish et al., 1983; MacLeish and Townes-Anderson, 1988).
Coating of the coverslip was obtained by applying overnight 100 µl of
goat affinity-purified antibody to mouse IgG (G
M; Cappel Research
Products, Durham, NC) over the thoroughly cleaned glass surface and by
adding then for 1 hr before cell isolation, a 100 µl sample of salt
solution containing the Sal-1 antibody. Cells were left 30 min to
adhere to the coverslip and were then covered with salamander Ringer's solution with the following composition (in mM):
112 NaCl, 3 KCl, 1.8 CaCl2, 0.5 MgCl2, 0.5 MgSO4, 0.5 NaH2PO4, 1 NaHCO3, 0.1 choline-Cl, 0.5 glucose, and 10 HEPES
and 0.001% phenol red, pH 7.4. This concentration of glucose in normal
Ringer's solution is close to the extracellular concentration measured
in rat striatum (Lowry et al., 1998) and was intended to mimic the
extracellular concentration in the retina, which has not yet been
measured. To maintain cells as long as possible, and because larval
tiger salamanders are poikilotherms living in cool water, dissociated
cells were maintained at 17°C, before and during the experiment.
Purified Müller cell islands. The procedure was
similar to the general procedure described above, with a few
modifications. First, only droplets of GM coating (see above) were
deposited onto the surface of the glass coverslip; when dried, these
produced islands 300-800 µm in diameter that were then covered with
100 µl of Sal-1 solution. Müller cells dissociated as described
above were purified by gravity sedimentation through a column of salt solution carrying BSA (500 µl of cell suspension on 500 µl of 10%
BSA-Ringer's solution, in an Eppendorf tube). After 20 min of
sedimentation, a 300 µl band of purified Müller cells was located and removed. These were allowed to settle on the island for
~10 min and were then rinsed several times with salamander Ringer's
solution. Cellular debris remaining after purification and not adhering
to the antibody were then washed away with medium. To obtain complete
purification, in some experiments, cellular debris adhering to the
Sal-1 antibody was removed by aspiration into a micropipette.
Fluorescence imaging of individual cells. Fluorescence was
measured on an inverted Axiovert 135TV epifluorescence microscope (Carl
Zeiss AG, Feldbach, Switzerland) equipped with Zeiss Pan-Neofluar 63×
oil immersion lens (1.25 numerical aperture) and fitted with a xenon
arc lamp, interference and neutral density filters, and an
electromagnetic shutter. Selected cells were positioned in the middle
of the optical field, and puffer pipettes (tip diameter, ~4 µm)
containing the test solution were advanced to within 50 µm from the
central portion of the cell and remained in place for the duration of
the experiment. Images, obtained by collecting fluorescence for 200 msec, were captured every 3 sec with a video CCD camera (Photonic
Science, Robertsbridge, UK) and processed by Ion Vision Software
(ImproVision, Coventry, UK) on a Power Macintosh 8100/100AV computer.
Some data were obtained on another setup, equipped with an Eclipse
TE300-DV microscope (Nikon AG, Küsnacht, Switzerland), a Nikon
Plan Fluor 100× oil immersion lens (1.3 numerical aperture), a
monochromator (T.I.L.L. Photonics, Planegg, Germany), a cooled
intensified CCD camera (Photonic Science), Openlab software
(ImproVision), and a Power Macintosh 9600/350 computer. The NAD(P)H
traces presented in Figure 4 were recorded with another cooled CCD
camera (Photometrics Ltd, Tuscon, AZ), as described in Tsacopoulos et
al. (1998). For pHi measurements, cells were
exposed for 5 min to BCECF-AM (2.5 µg/ml). After washout of
extracellular dye, pHi was measured from the
ratio of light emitted near 550 nm on excitation at 490 and at 440 nm.
For NAD(P)H fluorescence, excitation was at 360 nm, and light emitted
between 450 and 490 nm was collected; we refer to this fluorescence as "NAD(P)H fluorescence", meaning "the sum of NADH and NADPH
fluorescence", because the fluorescence of NADH cannot be
distinguished from that of NADPH (Barbour et al., 1993). For FAD
fluorescence, excitation was at 440 nm, and light emitted between 515 and 565 nm was collected. Fluorescence of different compounds was
measured sequentially (e.g., first NAD(P)H imaging, then BCECF loading
and imaging). A slight drift of the baseline was observed on the
recordings of NAD(P)H or FAD fluorescence. This drift, which was
probably attributable to bleaching of the fluorophore, caused on
average a signal decrease of 1.7 ± 0.3% per min (estimation on
78 traces from 53 cells). To facilitate comparison between individual
responses, all the data were corrected for baseline drift. For this, we
calculated for each trace the average slope of the signal during the 30 sec preceding stimulation and then subtracted the extrapolated baseline from the whole signal trace.
Lactate measurements. Lactate-sensitive microelectrodes (see
Fig. 7A) consisted in glass-insulated platinum (Pt)
microelectrodes onto which a polymer layer immobilizing some lactate
oxidase was deposited. The procedure for fabricating such
microelectrodes has been described in detail (Cosnier et al., 1997;
Poitry et al., 1997). Briefly, the tip of a glass micropipette was
broken with a diamond cutter to an inner diameter of 20-40 µm. A Pt
wire [Pt 90%, iridium (Ir) 10%; 125 µm diameter] was
etched by electrolysis in saturated KCN solution until its tip diameter
was only 2-5 µm. This Pt wire was then introduced into the
micropipette and advanced until its tip protruded from the opening of
the micropipette. Insulation of the Pt wire was obtained by gently
melting the tip of the micropipette to produce a glass sleeve around
the tip of the Pt wire. The excess Pt wire still protruding from the
tip of the micropipette was then etched by electrolysis under a thin jet of 2 M NaCl solution. Finally, a polypyrrole
film in which lactate oxidase was immobilized was deposited onto the
exposed Pt surface by electropolymerization. Lactate oxidase from
Pedioccocus oxidizes L-lactate (but
not D-lactate; Eichel and Rem, 1962; Naka, 1993)
to pyruvate and hydrogen peroxide. The pH dependence of the reaction is
weak, with an optimal pH of ~7.5 (Clark et al., 1984; Raba and
Motola, 1994), and the Km of the
enzyme for L-lactate is 700 µM (Sigma datasheet). The amperometric
detection of L-lactate was achieved by holding
the potential of the microelectrode at +0.6 V relative to an Ag-AgCl
reference electrode to detect the hydrogen peroxide produced in the
reaction. We previously used the same technique to make
glutamate-sensitive microelectrodes based on the entrapment of
glutamate oxidase, and the performance of this type of microelectrode
has been described in detail (Cosnier et al., 1997; Poitry et al.,
1997). The response of the microelectrodes used in the present study
was linear in the range 0-50 µM
L-lactate, and it was not affected by 200 µM alanine, glutamine, glutamate, or
NH4Cl (Fig. 1); it
was also not affected by possible pH changes induced by lactate
released from the cells: the 10 mM HEPES buffer in our Ringer's solution was largely in excess of the concentration of
lactate released, so that pH changes could not have exceeded 0.01 pH
unit in this study. The response times were always <10 sec.
Microelectrodes kept at 4°C retained their sensitivity for up to 3 weeks. This type of microelectrode cannot be used for accurate
measurements in cells and tissues releasing catecholamines or ascorbic
acid (i.e., molecules that can be oxidized directly on the Pt surface),
unless these interfering signals are subtracted. Control experiments
with the same Pt microelectrode as used for lactate measurements, but
without deposited enzyme, showed us that Müller cells do not
release detectable amounts of directly oxidizable compounds. For this
test, the old enzyme layer was removed from the Pt surface by briefly
dipping the microelectrode in 1 M HCl, followed
by rinsing in distilled water and rapid electrolysis of the Pt surface
in 2 M NaCl solution.
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Figure 1.
Current recording showing the sensitivity and the
selectivity of the lactate sensor. Initially, the sensor was in
Ringer's solution, and arrows indicate the times at
which the sensor was exposed to new solution. R,
Ringer's solution; +AA, + alanine + glutamine + glutamate + NH4Cl (200 µM each); 30
(or 15), Ringer's solution + 30 (or 15) µM
lactate.
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Ammonia measurements. The ammonia sensor was made of a
micropipette whose tip was filled with a droplet of fluorescent pH indicator covered with a gas-permeable membrane. This sensor is a
miniaturized version of the sensor described by Kar and Arnold (1992).
The method for fabricating this sensor was similar to that described by
Elamari et al. (1997), with some modifications. A glass micropipette
with short stem was made, using a vertical homemade puller, and its tip
was broken with a diamond cutter to an inner diameter of 50 µm and
fire-polished. A Teflon membrane (standard 65 N; DuPont de Nemours) was
stretched and fitted over the opening of a 10 mm glass tube and then
heated for 10 min at 300°C. Then, the fire-polished micropipette was
pushed through the heated membrane at its thinnest part, so that a
membrane patch remained attached over the tip of the micropipette, thus
closing the opening. This membrane was impermeable to water and ions, but permeable to gases. Only the stem of the micropipette, situated proximally to the Teflon-covered tip, was back-filled with a droplet of
solution containing 115 mM NaCl and 0.5 mM CDMF
indicator, pH 5.4. Unprotonated CDMF is fluorescent (peak excitation,
~505 nm; peak emission, 537 nm), but the protonated CDMF is not.
Because its pKa is close to 7, CDMF can be used
as a fluorescent pH indicator from approximately pH 5-9. An optical
fiber (120 µm diameter; CeramOptech, Bonn, Germany) was then
introduced into the micropipette by means of a specially constructed
holder, so that the extremity of the fiber was positioned in the
droplet as close as possible to the Teflon membrane. Thus, the volume
of the solution trapped between the fiber end and the Teflon membrane
was typically <1 nl, and indicator fluorescence was monitored only in
this minute volume. The other end of the optical fiber was connected to
the collecting end of a multimode optical fiber coupler (Y type,
GI 100/140 µm, 0.29 numerical aperture; Corning, Corning, NY).
One arm of the optical fiber coupler received the light beam from a
green LED (Sibalco, Basel, Switzerland) that passed through a 509 nm
interference filter; this light, traveling down the optical fiber,
excited the fluorescent solution trapped inside the sensor tip. The
emitted fluorescence was collected by the same optical fiber and
measured through the second arm of the optical fiber coupler; this
emitted light passed through a 537 nm interference filter and was
directed, via an optical fiber pigtail, to a single-photon counting
module (SPCM-100 PQ; E G & G, Vaudreuil, Canada) connected to a
timer/counter board of data acquisition (CTM-05/A; Keithley Instruments
S.A., Dübendorf, Switzerland). In principle, the sensor detects
pH-changing gas molecules such as NH3 or
CO2. However, the signals generated by these
molecules are opposite: NH3 causes alkalinization
(and thus increased fluorescence) of the indicator, whereas
CO2 produces acidification of the indicator. In
addition, the detection efficiency of the sensor is much higher for
NH3 than for CO2: because
the pH of the indicator solution is 5.4 and the
pKa of ammonia is 9.15, 99.98% of
NH3 molecules entering the sensor react with
H+ in the indicator solution and are
converted to NH4+; in
contrast, the pKa of
CO2-bicarbonate is 6.17 and therefore only 14.5%
of the CO2 molecules entering the sensor are
converted to bicarbonate and generate H+.
To increase further the detection efficiency for
NH3 in our experiments, all ammonia measurements
were performed in HEPES-buffered Ringer's solution, with no added
bicarbonate, and at pH 7.8: thus, only 2.3% of
CO2-bicarbonate released by the cells was found
as CO2 in the Ringer's, whereas 4.3% of ammonia
released by the cells was found as NH3.
Typically, the sensor signal measured for 5 µM
NH4Cl in Ringer's at pH 7.8 corresponded to an
increase of 0.3 pH unit in indicator pH, close to the Teflon membrane.
The integrity of the Teflon membrane, which is essential for proper functioning of the sensor, was continuously monitored via the sensor
fluorescence signal: any leak through the membrane resulted immediately
in indicator dilution and, therefore, in irreversible decrease of fluorescence.
Chromatographic analysis. HPLC analysis was performed in the
laboratory of Dr. Kim Q. Do (Brain Research Institute, University of
Zurich) using a Hewlett-Packard chromatograph (model 1090) combined
with a variable autoinjector, a cooled (4°C) autosampler, a
programmable fluorescence detector (HP 1046A), and analytical Hyperchrome HPLC columns, 250 × 4.6 mm (Bischoff, Leonberg,
Germany) packed with Hypersil ODS 5 µm (Shandon, Cheshire,
UK). For further detail, see Do et al. (1995).
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RESULTS |
Glutamate transport and signaling
Experiments in which the transport of glutamate is monitored as
transmembrane current (Brew and Attwell, 1987) or as intracellular pH
change (Bouvier et al., 1992) indicate that the uptake has an apparent
Km of ~20 µM
and is fast enough to cause a rapid rise in the intracellular
concentration of glutamate (Barbour et al., 1993). As previously shown
(Brew and Attwell, 1987; Tsacopoulos et al., 1998), the magnitude of
the glutamate-induced inward current in salamander Müller cells
is largest at the level of the distal processes and at the cell soma
(Fig. 2A). In the
intact retina, these are regions extending from the glutamatergic
synapses between photoreceptors and bipolar cells, to near the
glutamatergic synapses between bipolar and ganglion cells (Brew and
Attwell, 1987, their diagram in Fig. 4). These are also regions where
the analog of the excitatory amino acid transporter subtype EAAT1 is
found in salamander Müller cells (Eliasof et al., 1998; see also
Rauen et al., 1998, for similar results on the rat retina). Thus, after a puff application of 40-200 µM glutamate, the
change in the intracellular concentration of glutamate is expected to
be as large in the distal processes, where most mitochondria are found
(Tsacopoulos et al., 1998, their Fig. 5B), as in the
mitochondria-poor soma of the cell. The similarity in the time course
of the glutamate-induced 
pHi measured in the
distal processes and soma (Fig. 2B) is indeed consistent with this reasoning.
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Figure 2.
Effects on pHi and NAD(P)H
fluorescence of pressure-ejecting glutamate (200 µM) onto
the surface of a Müller cell. A, Image of a
freshly isolated salamander Müller cell plated in a Petri dish,
showing the major specialized regions of the cell. In
situ, the distal processes are in contact with synaptic
terminals of photoreceptors. On the left and out of
focus is shown a puffer pipette of the type we used in fluorescence
measurements. B, Glutamate-induced transient
acidification (bottom panel) in a Müller
cell loaded with BCECF-AM. This acidification (Bouvier et al., 1992),
occurs in the distal processes (trace 1), as well as in
the soma (trace 2). In contrast, the glutamate-induced
rise of NAD(P)H (top panel) occurs predominantly
in the distal processes, but in parallel with that of
pHi. Note the different distribution of the fluorescence
of NAD(P)H (top right) as compared to that of the mostly
cytosolic dye BCECF (bottom right): NAD(P)H fluorescence
is much higher in the distal processes (region 1) than in the soma
(region 2) of the cell. These measurements were performed sequentially:
NAD(P)H fluorescence was measured first, and then the cell was loaded
with BCECF-AM and pHi was measured.
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Because the metabolism of glutamate through the tricarboxylic acid
(TCA) cycle is associated with the control of the cytosolic reoxidation
of NADH (see Fig. 12, reaction 3; Dennis and Clark, 1978), we developed
a technique for imaging NAD(P)H fluorescence (see Materials and
Methods) at the subcellular level in single acutely isolated
Müller cells. As shown in Figures 2B and
3, basal NAD(P)H fluorescence was much
more intense in the distal processes than in either the soma or the
endfoot, in sharp contrast to the distribution of the mostly cytosolic
dye BCECF (Fig. 2B, pictures). This distribution of
NAD(P)H fluorescence is consistent with the distribution of
mitochondria: estimates based on Figure 5B from Tsacopoulos
et al. (1998) indicate that the number of mitochondria is 5-10 times
higher in the distal processes than in the rest of the cell, and it is
well known that, although the total concentration of pyridine
nucleotides is at least as high in mitochondria as in the cytosol, the
redox ratio NADH/NAD+ is much higher in
mitochondria than in cytosol (Sies, 1982). To test whether NAD(P)H
fluorescence in the distal processes was indeed predominantly of
mitochondrial origin, we puffed 3 mM of Amytal, a
well established inhibitor of mitochondrial respiration (Ernster et
al., 1955; Chance and Hess, 1959; Slater, 1967). As shown in Figure 3,
C and D, exposure of the cell for 30 sec to Amytal caused a rapid and large (>30%) increase of fluorescence restricted to the distal processes, in accordance with the hypothesis that the recorded fluorescence is predominantly mitochondrial and is
mostly that of NADH (Sies, 1982, their Fig. 3B, Table
1).
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Figure 3.
Relation of NAD(P)H fluorescence and FAD
fluorescence to mitochondrial metabolism. A, Both types
of fluorescence are observed predominantly in the distal processes of
freshly isolated Müller cells, where mitochondria are packed
(Tsacopoulos et al., 1998). Peak wavelengths of fluorescence excitation
and of measured emission are given in parentheses.
Compartmentation of fluorescence is more apparent for NAD(P)H than for
FAD mainly because the fluorescence intensity of FAD is much lower
(Masters and Chance, 1993). B, Schematic representation
showing how the mitochondrial redox potential is linked to the
functioning of the tricarboxylic acid cycle (TCA) and to
respiration. Boxes indicate which member of each redox
couple is fluorescent. C, Pressure-ejecting the
inhibitor of mitochondrial respiration Amytal (3 mM;
solid bar) onto a Müller cell induced a rapid rise
of NAD(P)H fluorescence only in region 1 (top left
panel), which corresponds to the distal processes of the
cell. a.u., Arbitrary units. D, Same
recordings as in C, but represented as relative changes,
to show that the absence of NAD(P)H response from regions 2 and 3 is
not attributable to the lower level of fluorescence.
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Measurements performed on purified suspensions of acutely isolated
Müller cells from the guinea pig retina have shown that the
intracellular concentration of glutamate (~1 mM;
Poitry-Yamate et al., 1995) is much lower than in the whole retina.
Under similar conditions, i.e., acutely isolated Müller cells
bathed in a simple Ringer's solution, we tested whether glutamate
modulates NAD(P)H fluorescence. Puffing either 40 or 200 µM of glutamate for 30 sec onto a Müller cell
caused a transient monophasic increase of this fluorescence (54 cells),
of similar amplitude for both concentrations, occurring mostly in the
distal processes (Fig. 2B). These results are similar
to those obtained by Barbour et al. (1993) who dialyzed salamander
Müller cells with patch-clamp pipettes (see Discussion). In the
first 2 hr after cell isolation, we sometimes recorded a small biphasic
response (18 cells) or a small monophasic decrease (26 cells) of
NAD(P)H fluorescence in response to glutamate application. The same
variability in response shape was seen when the cells were bathed in 16 mM glucose. Therefore, to ensure that the cells
tested were all in a similar metabolic state, we decided in the
following experiments to test cells 2-5 hr after their acute
isolation, i.e., when they all respond to glutamate by an increase in
NAD(P)H fluorescence.
This glutamate-induced increase of mitochondrial NADH appears to be a
typically glial response. Indeed, in the experiment shown in Figure
4, we made a simultaneous recording of
NAD(P)H fluorescence from a glial Müller cell and from the
ellipsoid of a closely located rod (the ellipsoid of a rod is packed
with mitochondria; Townes-Anderson et al., 1985). Glutamate induced in
the rod a small transient decrease of the fluorescence, whereas in the
Müller cell the response was a large rise. Because salamander rod
photoreceptors can also take up glutamate (Grant and Werblin, 1996),
probably via transporters present near their synaptic terminal (Eliasof
et al., 1998), this suggests that, in the same nervous tissue, the
mitochondria in a neuron metabolize glutamate differently from those in
a glial cell.
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Figure 4.
Comparison of the NAD(P)H fluorescence change
induced by glutamate (200 µM) in a freshly isolated
Müller cell and in a rod. Although pressure ejection of glutamate
induced a monophasic rise of NAD(P)H fluorescence in the distal
processes of a Müller cell, the same pressure ejection induced a
small transient decrease of NAD(P)H fluorescence in a neighboring rod
inner segment, also packed with mitochondria. This indicates that the
response of mitochondria to glutamate depends on their environment. The
30 sec period during which the cell was exposed to glutamate is shown
by the thick solid line.
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It has been shown that glutamate transporters transport also
L- and D-aspartate, and by the same ionic
mechanism (Kanai and Hediger, 1992; Barbour et al., 1993). To test
whether the glutamate-induced NAD(P)H response in the Müller cell
was attributable to a specific action of glutamate or to the ionic
current accompanying transport, we compared the effect of puffing
glutamate to that of puffing L- or D-aspartate
at the same concentration. As shown in Figure 5, the nonmetabolizable
D-aspartate exerted a small effect, opposite that of
L-glutamate on the same cells; L-aspartate also
exerted an effect opposite that of glutamate, inducing a small decrease of NAD(P)H in all cells tested. In the experiment of Barbour et al.
(1993), exposure of patch-dialyzed Müller cells to L-
or D-aspartate induced an inward current, but had no effect
on NAD(P)H. Taken together, these results strongly suggest that the
NAD(P)H response to glutamate was attributable predominantly to a
specific action of glutamate on the mitochondrial metabolism of the
Müller cell. Therefore, we explored further the mechanisms
underlying this effect.
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Figure 5.
The glutamate-induced NAD(P)H response does not
simply reflect the activity of the glutamate transporter. Comparison of
the effects of glutamate (200 µM) and either
D- or L-aspartate (200 µM), which
both enter Müller cells by the same transporter as glutamate
(Barbour et al., 1993), on NAD(P)H fluorescence in the distal processes
of Müller cells. The histogram gives the amplitude of the NAD(P)H
response at the end (white columns; ±SE), and 30 sec
after the end (shaded columns; ±SE), of the 30 sec
period during which the test solution was pressure-ejected. The number
of cells in which response pairs were recorded is given in
parentheses.
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Biochemical data obtained initially by Haslam and Krebs (1963) and
extended later by Dennis and Clark (1977, 1978) show that, in
mitochondria of probably mostly astrocytic origin and at millimolar concentrations of glutamate, >70% of the glutamate entering the TCA
cycle is oxidized through transamination with oxaloacetate. Aspartate
and -ketoglutarate are released from these mitochondria into the
cytosol where aspartate can be converted back to oxaloacetate and then
malate, which reenters the mitochondria (see Fig. 12). This
malate-aspartate shuttle, which contributes to the continual reoxidation of cytosolic NADH produced by glycolysis, occurs probably also in the Müller cell, because acutely isolated Müller
cells metabolizing 14C(U)-glucose
produce large amounts of 14C-aspartate
(Poitry-Yamate et al., 1995), and also
14C-malate and
14C--ketoglutarate (Poitry-Yamate,
1994). However, because aspartate and -ketoglutarate are also
released from Müller cells, the TCA cycle in these cells must be
supplied with new intermediates through an anaplerotic process, to
avoid depletion. Among such processes are the deamination of glutamate
to -ketoglutarate by mitochondrial glutamate dehydrogenase (GDH), or
the carboxylation of pyruvate to oxaloacetate by the glial enzyme
pyruvate carboxylase (see Fig. 12; for review, see Schousboe et al.,
1997). Of these, only the process catalyzed by GDH directly produces
NADH (see reaction 4 in Discussion). In contrast, pyruvate carboxylase
hydrolyzes ATP and may therefore indirectly cause a decrease in
mitochondrial NADH by increasing mitochondrial respiration.
To probe the relative importance of transamination in the reactions
occurring in an intact isolated Müller cell, we performed experiments using either a high concentration of pyruvate or the well
known inhibitor of transaminases, aminooxyacetate (AOAA; Westergaard et
al., 1996). Haslam and Krebs (1963) found that 10 mM of
pyruvate strongly inhibited oxaloacetate transamination in brain
homogenates, and Dennis and Clark (1978) observed the same in their
nonsynaptic mitochondria. Our results in acutely isolated Müller
cells show that, after 15 min of exposure to 10 mM
pyruvate, a puff of glutamate did not induce more of an increase of
NAD(P)H fluorescence (Fig.
6A). In contrast,
exposure of a Müller cell for 1 hr to a high concentration of
AOAA (5 mM), had a negligible effect on the basal
NAD(P)H fluorescence and caused a slight increase in the
glutamate-induced rise of NAD(P)H fluorescence (Fig.
6B). Our interpretation of these results is that
GDH-catalyzed deamination of glutamate is the major TCA anaplerotic
process in Müller cells maintained in simple Ringer's solution,
as is also the case in cultured astrocytes (Westergaard et al., 1996).
Thus, this process appears to be a major source of mitochondrial NADH.
Probably, pyruvate at high concentration was metabolized through the
pyruvate carboxylase reaction, leading to a massive increase of
oxaloacetate and, as a consequence, to a massive increase of
-ketoglutarate. This, in turn, inhibited both transamination and
deamination of glutamate in the mitochondria (see Fig. 12, diagram and
Discussion) and thereby suppressed the NAD(P)H response. AOAA, on the
other hand, probably blocked the transamination of glutamate and
thereby favored its deamination, leading to an increased NAD(P)H
response.
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Figure 6.
Effects of a high concentration of pyruvate or of
AOAA on the NAD(P)H response elicited by pressure ejection of 200 µM glutamate. A, Histogram of the response
amplitude at the end (white columns; ±SE), and 30 sec
after the end (shaded columns; ±SE), of the 30 sec
pressure-ejecting period in normal Ringer's solution
(left) and after 15-40 min in 10 mM
pyruvate-Ringer's solution (right). Response pairs
were collected in three cells. B, Effect of exposing the
cells to 5 mM AOAA. Histogram of the amplitude at the end
(white columns; ±SE) and 30 sec after the end
(shaded columns; ±SE) of the 30 sec pressure-ejecting
period. Response pairs were collected in three cells, and the duration
of exposure to AOAA ranged from 30 to 70 min.
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Because pyruvate is the end product of aerobic glycolysis, the results
of Figure 6A suggest a link between the rate of
glycolysis and the mode of oxidation of transported glutamate.
Therefore, we determined whether activation of glycolysis affected
mitochondrial NADH. We tested the effect of
NH4+, first because
biochemical evidence shows that
NH4+ at 1 mM is an allosteric activator of
phosphofructokinase (PFK), which is one of the enzymes controlling the
glycolytic flux (Lowry and Passonneau, 1966), and second because
stimulation of the isolated amphibian retina has been found to cause an
increase in the concentration of ammonia released in the extracellular
fluid (Kar and Arnold, 1992). In addition, deamination of glutamate by
mitochondrial GDH would produce
NH4+.
Ammonia transport and signaling
In the honeybee glial cells (Tsacopoulos et al., 1997a,b; Marcaggi
et al., 1999), cultured mouse astrocytes (Brookes, 1997), and kidney
cells (Kikeri et al., 1989), evidence based on measurements of
intracellular pH indicates that
NH4+ ions, rather than
NH3, enter these cells. Exposure of a
Müller cell to 50-200 µM
NH4Cl had small (
0.1 pH unit) and variable effects on intracellular pH, but exerted a strong metabolic effect (see
below). This suggests that ammonia could enter a Müller cell as
NH3 and
NH4+ in a proportion that
depends on the metabolic state of the cell, but in a sufficient amount
to specifically affect cell metabolism.
We explored these metabolic effects of
NH4+ on acutely isolated
Müller cells. For this, we made purified islands of acutely isolated Müller cells (Fig.
7A) and constructed
enzyme-based lactate-sensitive microelectrodes (see Materials and
Methods; Poitry et al., 1997) to measure net lactate release by these
cells. The cells were bathed for 3-5 hr in a stagnant Ringer's
solution carrying 0.5 mM glucose (physiological
concentration in the mammalian brain; Lowry et al., 1998). The lactate
concentration measured was virtually zero far from the cells, but when
the lactate electrode was positioned near the cells of the island (Fig.
7A), the measured value was in the range of 2-8
µM (Fig. 7B), depending on the size of the island. This result indicates a continuous production and release of lactate by Müller cells. In the presence of 100 µM glutamate in the bathing solution, exposure
of the cells to 100 µM
NH4Cl induced a large increase in lactate release
(Fig. 7C). All islands tested (n = 10)
showed this NH4+-induced
lactate release (range, 10-50 µM at the peak).
In the absence of added glutamate,
NH4+ had no measurable
effect (Fig. 7D). Calculations indicated that even if
Müller cells had released their entire content of lactate under
the combined effect of
NH4+ and glutamate, the
concentration of lactate in the bathing solution near the cells would
never reach such a high concentration without a concomitant strong
stimulation of the production of lactate.
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Figure 7.
Cooperative effect of
NH4+ and glutamate on lactate production
and release. A, Picture showing a portion of a
microisland made of ~250 freshly isolated Müller cells. Also
shown is a lactate microsensor with its tip positioned close to the
cells. Scale bar, 100 µm. B, When the microsensor was
approaching the cells from the bath, the lactate signal increased,
indicating the existence of a lactate concentration gradient.
C, Exposure of the cells to 100 µM
glutamate (solid bar) had no effect on the lactate
signal, but addition of 100 µM NH4Cl
(gray bar) in the presence of 100 µM glutamate induced a large increase in lactate
concentration. Withdrawal of the microprobe from the cells into the
bath caused the lactate signal to decrease close to zero (zero level is
indicated by the dotted line). The interruption in the
trace corresponds to a 30 min period. D, Exposure of the
cells to 100 µM NH4Cl was by itself
insufficient to induce a detectable increase of lactate concentration.
Only after further addition of 100 µM glutamate was a
lactate response recorded. E, Lactate responses elicited
by the cooperative action of glutamate and NH4Cl (100 µM each) were transient, but could be repeated by further
addition of glutamate and NH4Cl (bath concentration
increased to 200 µM for each, indicated by thickening of
the solid bars). The measurement in E was
done on the island shown in A, which comprised
three to four times more cells than those used in the other
measurements.
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We checked that the signal of the lactate-sensitive microelectrode was
not affected by the presence of 200 µM glutamate,
NH4Cl, glutamine, and alanine (the two latter
compounds are released by Müller cells, as shown below). However,
there was still the possibility that the microelectrode sensed another
compound directly oxidized on the Pt electrode. This was tested by
using the same Pt electrode, but the lactate oxidase normally
immobilized in the polymer was omitted. With such an electrode,
polarized at +600 mV, we recorded neither a gradient nor an effect of
NH4+ and glutamate on the
measured current. These results suggest, first, that this enzyme-based
electrode recorded specifically lactate; second, that acutely isolated
Müller cells do not release measurable quantities of molecules
oxidizable at this voltage (e.g., catecholamines, vitamins); and third,
that NH4+ in the presence
of glutamate, or vice versa, strongly increases the
production and the release of lactate.
As shown in Figure 7E, the increased production of lactate
was transient, in spite of the continuous presence of
NH4+ in the bathing
solution, and it could be elicited again only after further addition of
glutamate and NH4+.
Bathing the cells in a Ringer's solution carrying a 10 times higher
concentration of glucose did not prolong the duration of this lactate
increase (result not shown), indicating that the reaction was not
limited by glucose availability. Therefore, the result of Figure
7E suggests that the increase in lactate production is
linked to a reaction shared by the metabolism of both glutamate and
NH4+, and that this
reaction may have been limited in our experiments by the availability
of either one of the reactants. Indeed, to allow detection of released
lactate, the bath was left stagnant after the addition of glutamate and
NH4+; thus, after the
first addition of glutamate and
NH4+, a high uptake rate
of either glutamate or
NH4+ may have caused
their concentration to decrease much below bath value, in the vicinity
of the cells, so that the reaction became diffusion-limited. After
further addition of glutamate and
NH4+, the solution was
mixed, and the concentration near the cells probably increased
transiently to the bath value, allowing lactate production to increase
transiently again. This interpretation is in agreement with the large
decrease in [NH4+]
measured with our ammonia sensor (see below and Fig.
8B). We show below that
the reaction shared by glutamate and
NH4+ is the synthesis of
glutamine.
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Figure 8.
Glutamine is produced and released by Müller
cells exposed simultaneously to glutamate and NH4Cl.
A, B, Measurements of
[NH4+] by a fiber-optic ammonia
sensor. To optimize the detection of ammonia, the pH of the bathing
solution in this type of experiment was adjusted to 7.8. A, Control experiment. When the sensor was in the bath,
addition of neutral glutaminase (100 µl, in Ringer's solution at pH
7.8, added to 2 ml bath; final activity: 0.01 U/ml) did not produce an
increase in the [NH4+] signal, but a
decrease caused by dilution, indicating that the glutaminase solution
contained much less than 10 µM
NH4+. B, When the sensor
was positioned close to freshly isolated Müller cells forming a
microisland (Fig. 7A), and after cells were exposed to
glutamate and NH4+, addition of
glutaminase caused a large increase in
[NH4+]. Twenty minutes before the
beginning of this record, 5 µM NH4Cl was
added to the Ringer's solution bathing the cells. As shown, further
addition of 50 µM glutamate caused a rapid drop of
[NH4+] near the cells, indicating
substantial uptake of ammonia by the cells. The time constant of this
drop was similar to the time constant of the sensor response. After 10 min of exposing the cells to glutamate and
NH4+, glutaminase was added to the bath,
causing the rapid hydrolysis of glutamine present in the bath and a
concomitant increase of [NH4+].
C, Amounts of glutamine (Gln, filled
triangles) and alanine (Ala, open circles)
released into the bath by a microisland of ~850 Müller cells
after 30 min incubating periods in normal Ringer's solution (all bath
collections except third) or in Ringer's solution containing glutamate
and NH4Cl (50 µM each, third bath
collection). Exposure to glutamate and NH4Cl caused a
sizable increase in the production and release of glutamine and
alanine. Amino acid content of bath collections was measured by HPLC,
using ortho-phthalaldehyde derivatization.
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Glutamine synthesis and release
The ammonia content of isolated glial cells decreases with time
after isolation, in solution containing no added ammonia (Tsacopoulos et al., 1997a,b). In cultured cortical astrocytes, Waniewski (1992) showed that, at very low concentrations of ammonia in the bathing medium, the production and release of glutamine were increased when the
cells were exposed to added
NH4+, at a concentration
as low as 50 µM. The enzyme that catalyzes the formation
of glutamine from glutamate and
NH4+ at the expense of
ATP is glutamine synthetase (GS). Using classical immunocytochemistry,
we found that in the salamander retina GS activity is localized
throughout the cytosol of Müller cells, consistent with previous
findings in the mammalian retina and brain (Derouiche and Rauen, 1995;
Derouiche et al., 1996). It is particularly interesting that this
activity is preserved in acutely isolated cells maintained under the
conditions described above (results not shown). Consequently, we
expected that exposure of these cells to micromolar concentrations of
NH4+ would elicit an
increase of glutamine synthesis and release.
We tested this expectation by monitoring the release of glutamine by
Müller cells, using a technique that combines a fiber-optic ammonia sensor with the extracellular application of partially purified
glutaminase diluted in the bathing solution, at pH 7.8. At this pH, the
"neutral" enzyme we used has its maximum activity, and ~4% of
the ammonia produced is in gaseous form and thus sensed by the probe.
Under these conditions, and because the added glutaminase solution had
the same pH as the bathing solution and did not contain ammonia (Fig.
8A), any increase in ammonia detected after addition of glutaminase should result only from the hydrolysis of glutamine by
glutaminase. To detect glutamine produced by the cells, a purified Müller cell island was bathed with Ringer's solution containing 5 µM
NH4+. The ammonia sensor
was advanced in the bath, and its tip was positioned close to the
cells. No gradient of
NH4+ concentration was
detected. The addition of 50 µM glutamate
induced then a rapid decrease of
NH4+ in the bath near the
cells (Fig. 8B). We suggest that added glutamate was
transported into the Müller cells where it induced an increase of
glutamine synthesis. This is likely because the
Km of brain GS for glutamate is much
higher than for NH4+ (see
Pamiljans et al., 1962; for human brain, Listrom et al., 1997).
Probably, in our experiments, the glutamate-induced increase of
glutamine synthesis used intracellular
NH4+, thus increasing the
transmembrane gradient of
NH4+ concentration. This,
in turn, facilitated the entry of
NH4+ from the
extracellular space and caused the observed decrease of
[NH4+] near the cells.
Demonstration that the glutamate-induced decrease of extracellular
[NH4+] might be
attributable to an increased rate of glutamine synthesis was provided
by showing that the cells had produced and released an equivalent
amount of glutamine. Indeed, as shown in Figure 8B,
addition of 0.02 U of glutaminase to the bathing solution caused a
sharp increase of the concentration of ammonia, because glutaminase
rapidly transformed the glutamine released by the cells to glutamate
and NH4+. Similar results
were obtained in five Müller cell islands with five different
sensors. The concentration of ammonia after glutaminase addition was
higher than before the addition of glutamate probably because some
glutamine synthesis and release occurred before the exposure to glutamate.
These results are confirmed by HPLC analysis of the bathing solution
showing that, before the exposure of Müller cells to 200 µM NH4+ and
glutamate and the consecutive increase in glutamine production, there
was a measurable, but slowly decreasing, release of glutamine into the
bath (Fig. 8C). We conclude that acutely isolated
Müller cells produce and release glutamine and that this
production in the presence of micromolar concentration of ammonia is
strongly stimulated by glutamate.
Link between glutamine synthesis and glycolysis
Because exposure to
NH4+ and glutamate caused
a massive increase in lactate production (Fig. 7), the natural question
then was whether this effect was a direct consequence of the increased rate of glutamine synthesis. This possibility was tested by measuring lactate concentration near Müller cell islands in the presence of
1 or 3 mM methionine sulfoximine (MSO), a well established specific inhibitor of GS (Meister, 1974). Whereas the basal level of
lactate production and release by Müller cells did not change significantly after addition of MSO, the increase in lactate production elicited by NH4+ and
glutamate was irreversibly suppressed within 30 min (Fig. 9B). The irreversibility of
this effect was expected, because MSO has been shown to undergo
phosphorylation after binding to the glutamate site of GS, thereby
permanently inactivating the enzyme (Meister, 1974). However, 1 or 3 mM of methionine sulfone, whose inhibitory action
on GS was reported to be more reversible than MSO (Rowe et al., 1969),
produced the same effects (Fig. 9A), irreversible within 1 hr after application (Fig. 9B), which suggests that these
inhibitors tend to accumulate in the cells.
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Figure 9.
The lactate response is linked to the activity of
glutamine synthetase. A, Lactate measurements showing
that exposure of Müller cells to 1 mM methionine
sulfone inhibits the increase in lactate production induced by
glutamate + NH4+ (100 µM
each). B, Histogram showing the effects of 3 mM methionine sulfoximine (white columns; 2 cells) and 3 mM methionine sulfone (black
columns; 6 cells) on the lactate response induced by glutamate + NH4+. Suppression of the response
during exposure to the inhibitor (central column,
MSO) was not reversed 1 hr after returning the
preparation to normal Ringer's (right columns,
wash).
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The effects of MSO and methionine sulfone were consistent with the idea
that the increase in lactate production elicited by NH4+ + glutamate was a
consequence of the increased rate of glutamine synthesis. Moreover,
because the inhibition of GS should produce an elevation of the
intracellular level of
NH4+, these effects
excluded a direct role of
NH4+ in the activation of
glycolysis and lactate production. A possible link between glycolysis
and glutamine synthesis is ATP (or its hydrolysis products ADP and
Pi), because the GS reaction consumes ATP and
because the activity of PFK is known to be regulated by the levels of
ADP and Pi (Lowry and Passonneau, 1966). As
mentioned above, GS is also present in the distal processes of the
Müller cell and, therefore, changes in the cytosolic levels of
ADP and Pi caused by increased glutamine
synthesis may also lead to increased mitochondrial respiration. We
explored this possibility by measuring NAD(P)H fluorescence during
exposure of the Müller cell to
NH4+. As shown in Figure
10A, puffing 200 µM NH4Cl for 30 sec
induced a large monophasic and transient decrease of NAD(P)H, a change occurring only in the distal processes of the cell. The same puff caused a parallel, but smaller, increase in flavin adenine dinucleotide (FAD) fluorescence (Fig. 10B). FAD is the hydrogen
acceptor of the reaction converting succinate to fumarate in the TCA
cycle and is produced by the respiratory chain (Fig. 3B).
Thus, probably ADP and Pi produced by the
increased flux through the GS reaction stimulated mitochondrial
respiration. As shown in Figure 10, C and D,
exposure to MSO decreased significantly, but did not abolish the effect
of NH4+ on NAD(P)H
fluorescence. This incomplete effect of MSO, even after a rather long
incubation period, indicates that
NH4+ probably decreases
NAD(P)H fluorescence also via another reaction than that catalyzed by
GS, for example by decreasing the flux of glutamate through the
glutamate dehydrogenase reaction (see Fig. 12).
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Figure 10.
Changes in the mitochondrial redox potential of
Müller cells are linked to the activity of glutamine synthetase.
A, B, Exposure of a Müller cell to 200 µM NH4Cl caused a decrease of NAD(P)H
fluorescence and an increase of FAD fluorescence in the distal
processes (thick solid traces, see inset)
where mitochondria are packed, but not in other regions of the cell.
C, The NAD(P)H response elicited by
NH4+ in the distal processes is
decreased by 3 mM MSO, an inhibitor of glutamine
synthetase. The response in MSO-Ringer's (dashed line)
was recorded 50 min after switching solution. D, Effect
of MSO on the NAD(P)H response induced by 200 µM
NH4Cl. Histogram of the amplitude at the end (white
columns; ±SE), and 30 sec after the end (shaded
columns; ±SE), of the 30 sec pressure-ejecting period in normal Ringer's
(left) and after 10-60 min of exposure to 3 mM MSO-Ringer's (right). Response pairs
were collected in three cells. Response amplitudes at the end of the
pressure-ejecting period were significantly lower in MSO
(p < 0.01; paired t
test).
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Then we asked whether inhibition of glutamine synthetase by MSO has as
a major consequence an increase in intraglial glutamate availability to
other metabolic pathways. As shown in Figure
11A, we measured the
mitochondrial NAD(P)H fluorescence as a function of time after exposure
of Müller cells to normal (glutamate-free) Ringer's solution.
The basal NAD(P)H fluorescence decreased steadily with time, possibly
because of a sustained loss of TCA intermediates, but puffs of
glutamate induced large rises of NAD(P)H (Fig. 11A). Then, the cell was exposed to 3 mM MSO, which
caused a rapid 40% rise of the basal NAD(P)H fluorescence in the
distal processes and suppressed the glutamate-induced rise of NAD(P)H
fluorescence (Fig. 11A,B). Our interpretation is
that, in the presence of MSO, glutamate is available to enter the TCA
cycle, thus leading to a nearly maximum turnover rate of the cycle and
consequently to the accumulation of NADH. Under this condition, the
transported glutamate could not increase further mitochondrial
NAD(P)H.
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Figure 11.
The glutamate-induced NAD(P)H response is linked
to the activity of glutamine synthetase. A, Plot of the
basal level of NAD(P)H fluorescence (filled
circles) and of the fluorescence at the peak of
glutamate-induced responses (open triangles) as a
function of time after cell isolation, and effects of exposure to 3 mM MSO (solid bar). The basal fluorescence
decreased slightly with time after cell isolation, but MSO caused it to
increase back toward a maximal value and suppressed the
glutamate-induced increase of fluorescence. Arrows
indicate the times at which the glutamate-induced NAD(P)H responses
shown in insets were recorded. Insets, Time
course of the glutamate-induced NAD(P)H responses recorded at the times
indicated by arrows on the plot. Glutamate (200 µM) was pressure-ejected during 30 sec, as indicated by
the solid bar under the traces. B, Effect
of exposing the cells to 3 mM MSO on the glutamate-induced
NAD(P)H response. Histogram of the amplitude at the end (white
columns; ±SE), and 30 sec after the end (shaded
columns; ±SE), of the 30 sec pressure-ejecting period. The
number of cells in which response pairs were recorded is given in
parentheses.
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DISCUSSION |
Based on the average current generated by the glutamate uptake,
Barbour et al. (1993) calculated that the intracellular concentration of glutamate in Müller cells may rise at a rate of ~0.5
mM/sec. Therefore, exposing these cells for 30 sec to 200 µM glutamate may cause the intracellular concentration to
increase up to ~15 mM, if no transformation of glutamate
occurred. However, the similarity of the time courses of NAD(P)H and
pHi, both measured by optical techniques in
this study (Fig. 2B), clearly demonstrates that transported glutamate activates metabolic reactions within seconds.
Our observations are consistent with the occurrence in a Müller
cell of four biochemical reactions involving glutamate: (1) glutamate + NH4++ ATP
glutamine + ADP + Pi; (2) glutamate + pyruvate alanine + -ketoglutarate; (3) glutamate + oxaloacetate 
-ketoglutarate + aspartate; (4) glutamate + NAD+ -ketoglutarate + NADH + NH3. These reactions,
already well established by previous biochemical work on brain
homogenates, isolated brain mitochondria, and cultured astrocytes (for
review, see Sonnewald et al., 1997), occur in the cytosol (reactions 1, 2, and 3), or in the mitochondria (reactions 3 and 4). Although they
are all related in some way to the NADH redox potential, only reaction
4 is directly linked to it.
The interest of our work resides in the use of subcellular imaging of
NAD(P)H fluorescence in an acutely isolated type of astrocyte in which
there is a clear metabolic compartmentation. Measurements in the
cytosol and in the mitochondria were thus possible, allowing us to
probe simultaneously both cellular compartments and to explore the
relationships between the above-mentioned reactions and the complex
signaling role of glutamate. We found that significant and rapid
glutamate-induced NAD(P)H changes occurred only in the distal processes
of the Müller cell, where most mitochondria are located. The
rapid response is explained by the finding that the mitochondria are
located in a region of the cell where glutamate transport is intense
(Tsacopoulos et al., 1998), so that intracellular glutamate
concentration can increase rapidly. The observation that glutamate did
not induce measurable changes of NAD(P)H in the soma, where glutamate
transport is also intense, is compatible with the idea that metabolism
there, as well as in the endfoot region, is predominantly glycolytic
and that glutamate alone does not cause a rapid activation of
glycolysis (see below and Hertz et al., 1998).
The shape of the glutamate-induced change of NAD(P)H depended strongly
on whether the cell was tested shortly after isolation or a few hours
later. With regard to this observation, we excluded a damage of the
transport carrier because L- and D-aspartate
elicited transport currents and NAD(P)H fluorescence decreases in all
Müller cells, independently of time after isolation. We therefore
believe that the parameter that changed over time after isolation was principally the metabolic state of the cells that was determined by the
intracellular levels of glutamate and
NH4+.
Our data show that reaction 1 catalyzed by GS (Fig.
12), which was found in the cytosol of
isolated salamander Müller cells, was the major route for
fixation of NH4+ and
hence for the transformation of cytosolic glutamate to glutamine. The
demonstration, made by Cooper et al. (1979) in the intact rat brain, of
glutamine synthesis 1-2 sec after a bolus injection of
13N-labeled ammonia into the blood stream,
is consistent with our data. The high affinity of GS for ammonia and
the lower affinity for glutamate, implies that at low, physiological
concentrations of both reactants, ammonia is trapped by GS, and the
production of glutamine is finely controlled by glutamate (Waniewski,
1992). Consequently, when the supply of ammonia is limited, as it was for our isolated Müller cells, production of glutamine continues until almost all available ammonia in the cells is trapped. When the
flux through the GS reaction is limited by lack of sufficient amounts
of ammonia (below 10 µM), then more glutamate is
available to enter reactions 3 and 4 occurring in mitochondria. This is consistent with what occurs in MSO-treated rats (Cooper et al., 1979)
and when cultured astrocytes are exposed to MSO (McKenna et al., 1996;
Westergaard et al., 1996).
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Figure 12.
Model diagrams showing biochemical pathways
involved in the trafficking and metabolism of glutamate
(glu) and NH4+
(A) and metabolite exchanges between cytosol and
mitochondrion (B). Numbers
indicate the reactions catalyzed by glutamine synthetase
(1), alanine aminotransferase
(2), aspartate aminotransferase
(3), glutamate dehydrogenase
(4), and pyruvate carboxylase
(5), or the mitochondrial glutamate-aspartate
exchange carrier (6); the circle
indicates transaminating reactions and exchange carrier, and the
square, other reactions. Also indicated are some
inhibitors: MSO for reaction 1 and AOAA for reaction 3. Even though
most of these reactions are reversible in principle, only the
directions of the reactions considered in the analysis of our results
are indicated. Colored arrows indicate whether NADH is
produced (green) or consumed (red)
in the reaction. Mitochondrial respiration is represented schematically
as a single reaction consuming NADH and transforming oxygen
(O2) into a proton gradient
(µH). The final product of the
metabolism of glucose-6-phosphate (g6P), pyruvate
(pyr), is reduced to lactate (lac)
by the enzyme lactate dehydrogenase. This reaction is essential for the
production of NAD+, subsequently used by glycolysis.
However, close to the mitochondrial surface the malate-aspartate
shuttle (m-a shuttle; the reactions involved in this
shuttle are indicated in light and dark
blue in B) may contribute to the production of
cytosolic NAD+. Glutamate
(glu) and NH4+ are
transported into the glial cell where they are transformed to glutamine (gln),
alanine (ala), and -ketoglutarate
(ket). These products leave the glial cell and are
taken up by neurons. In this way nitrogen homeostasis can be
maintained. asp, Aspartate; citr,
citrate; mal, malate; oxal,
oxaloacetate.
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The metabolic fixation of ammonia utilizes ATP (reaction 1). Because
the Km of GS for ATP is high (2.5 mM; Meister, 1974), the cell must restore rapidly
the ATP level to maintain glutamine synthesis. In Müller cells of
all species examined so far, GS is found to be cytosolic, and the
elegant study of Derouiche and Rauen (1995) demonstrated a
colocalization of GS and of the glutamate transporter in the fine
lateral processes of Müller cells. The uneven distribution of
mitochondria, restricted to the distal processes of salamander
Müller cells, suggests that a major part of the ATP used by GS is
produced through the glycolytic pathway. Indeed, in the absence o
TOP
ABSTRACT
INTRODUCTION
