Download Invulnerability of retinal ganglion cells to NMDA excitotoxicity

www.elsevier.com/locate/ymcne
Mol. Cell. Neurosci. 26 (2004) 544 – 557
Invulnerability of retinal ganglion cells to NMDA excitotoxicity
E.M. Ullian, a,*,1 W.B. Barkis, a,1 S. Chen, b J.S. Diamond, b and B.A. Barres a
a
b
Department of Neurobiology, Stanford University School of Medicine, Stanford, CA 94305-5125, USA
NIH/NINDS/SPU, Bethesda, MD 20892-4066, USA
Received 15 August 2003; revised 22 March 2004; accepted 10 May 2004
NMDA excitotoxicity has been proposed to mediate the death of retinal
ganglion cells (RGCs) in glaucoma and ischemia. Here, we reexamine
the effects of glutamate and NMDA on rat RGCs in vitro and in situ.
We show that highly purified RGCs express NR1 and NR2 receptor
subunits by Western blotting and immunostaining, and functional
NMDA receptor channels by whole-cell patch-clamp recording.
Nevertheless, high concentrations of glutamate or NMDA failed to
induce the death of purified RGCs, even after prolonged exposure for
24 h. RGCs co-cultured together with ephrins, astrocytes, or mixed
retinal cells were similarly invulnerable to glutamate and NMDA,
though their NMDA currents were 4-fold larger. In contrast, even a
short exposure to glutamate or NMDA induced the rapid and profound
excitotoxic death of most hippocampal neurons in culture. To
determine whether RGCs in an intact retina are vulnerable to
excitotoxicity, we retrogradely labeled RGCs in vivo using fluorogold
and exposed acutely isolated intact retinas to high concentrations of
glutamate or NMDA. This produced a substantial and rapid loss of
amacrine cells; however, RGCs were not affected. Nonetheless, RGCs
expressed NMDA currents in situ that were larger than those reported
for amacrine cells. Interestingly, the NMDA receptors expressed by
RGCs were extrasynaptically localized both in vitro and in situ. These
results indicate that RGCs in vitro and in situ are relatively
invulnerable to glutamate and NMDA excitotoxicity compared to
amacrine cells, and indicate that important, as yet unidentified,
determinants downstream of NMDA receptors control vulnerability
to excitotoxicity.
D 2004 Elsevier Inc. All rights reserved.
Introduction
Although glutamate is the major excitatory neurotransmitter in
the CNS, it is sometimes able to kill neurons (Choi, 1992; Rothman and Olney, 1987). Excitotoxic neuronal death is thought to be
an important contributor to neuron death caused by brain and
spinal cord injuries as well as by many neurological diseases. An
unsolved mystery, however, is why only some types of neurons are
* Corresponding author. Department of Neurobiology, Stanford University School of Medicine, Fairchild Science Building D235, 299 Campus
Drive, Stanford, CA 94305-5125. Fax: +1-650-725-3958.
E-mail address: [email protected] (E.M. Ullian).
1
These authors contributed equally to this paper.
Available online on ScienceDirect (www.sciencedirect.com.)
1044-7431/$ - see front matter D 2004 Elsevier Inc. All rights reserved.
doi:10.1016/j.mcn.2004.05.002
vulnerable to glutamate excitotoxicity. The ionotropic glutamate
receptors are ligand-gated ion channels that are grouped into three
pharmacologically defined classes: NMDA, AMPA, and kainate
receptors. These receptors are encoded by at least six gene families:
a single family for AMPA receptors (GluR1, 2, 3, 4), two for
kainate (GluR5, 6, 7 and KA1, 2), and three for NMDA (NR1,
NR2A, B, C, D, and NR3A, B) (Dingledine et al., 1999). These
subunits combine into multimeric complexes to form functional
receptor channels. To form an NMDA receptor, for instance, two
NR1 subunits must combine with at least two NR2 subunits.
NMDA receptors have been linked to excitotoxic death that occurs
in less than 1 h of exposure, whereas much longer exposures to
kainate or AMPA produce excitotoxicity (Choi, 1992).
We have focused on the effects of glutamate on retinal
ganglion cells (RGCs) in the developing rat retina. RGC death
is a leading cause of blindness resulting from glaucoma and retinal
ischemia, and NMDA receptor-mediated excitotoxicity is presently thought to be an important contributor to RGC death in these
conditions (Ritch, 2000; Wax and Tezel, 2002). The expression of
glutamate receptors by retinal ganglion cells in developing and
adult rats has been extensively studied. Based on in situ hybridization and immunohistochemical studies of rat retinal sections,
RGCs have been shown to express NMDA, AMPA, and kainate
receptors including NR1, NR2A, B, C, D, GluR2, GluR7, and
KA2 (Brandstatter et al., 1994; Grunder et al., 2000a,b; Watanabe
et al., 1994). Electrophysiological recordings of rat retinal ganglion cells in situ clearly indicate that they express functional
NMDA and non-NMDA receptor-activated channels (Chen and
Diamond, 2002). NMDA, AMPA, and kainate-activated currents
can all be observed in RGCs in mixed retinal cultures over the 1day period in which these cells remained viable without added
neurotrophic support (Aizenman et al., 1988; Taschenberger et al.,
1995). In vitro and in vivo, rodent RGCs have been repetitively
shown to be vulnerable to NMDA-mediated excitotoxicity
(reviewed by Lipton, 2001; Sucher et al., 1997). For instance,
both glutamate and NMDA induce the death of RGCs in postnatal
and adult retinas (Izumi et al., 1995; Li et al., 1999; Mosinger et
al., 1991; Sabel et al., 1995). Although kainate-elicited excitotoxicity of RGCs in culture has not generally been observed, one
group reported that kainate killed rat RGCs in purified cultures
(Otori et al., 1998); however, kainate-induced excitotoxicity in
intact retinas appears to be mediated by release of glutamate
which then activates NMDA receptors (Lipton, 2001; Sucher et
al., 1997). Taken together, these studies indicate that the postnatal
E.M. Ullian et al. / Mol. Cell. Neurosci. 26 (2004) 544–557
and adult RGCs express functional NMDA and non-NMDA
receptors, and are vulnerable to NMDA-mediated excitotoxicity.
Because of their expected vulnerability to glutamate, we
initially took measures to avoid exposing RGCs to glutamate
in our studies involving their purification and culture (Barres et
al., 1988). But eventually we were surprised to find that
glutamate exposure actually enhanced the survival of purified
RGCs in culture (Meyer-Franke et al., 1995). We found that
depolarization enhanced RGC survival by enhancing neurotrophic responsiveness (Goldberg and Barres, 2000; Goldberg et
al., 2002; Meyer-Franke et al., 1998; Shen et al., 1999), but we
remained puzzled by the invulnerability of the purified RGCs to
excitotoxicity.
Here, we have investigated the glutamate and NMDA invulnerability of purified RGCs in vitro. By culturing highly purified
RGCs in serum-free conditions, we have found that RGCs in vitro
continue to express the same glutamate receptor subunits that they
do in vivo, yet are invulnerable to glutamate and NMDA excitotoxicity. We also found that in vitro, as in situ (Chen and Diamond,
2002), NMDA receptors are localized extrasynaptically and not
synaptically. To our surprise, we found that RGCs in intact
developing and adult retinas are completely invulnerable to glutamate and NMDA excitotoxicity. Glutamate and NMDA did kill
many cells in the inner nuclear layer (INL) and ganglion cell layer,
but these cells were amacrine cells. Taken together, these studies
cast doubt on the relevance of excitotoxicity to RGC death in
glaucoma and retinal ischemia and strikingly illustrate that the
presence of functional NMDA receptors is insufficient to account
for excitotoxicity.
Results
Expression of NMDA receptor proteins by purified RGCs in culture
As an initial survey of glutamate receptor subunit expression
by RGCs in culture, we first examined mRNA extracted from
purified RGCs after 3 days of culture by Affymetrix gene chip
analysis and found that mRNAs for NR1, NR2, GluR2, and KA2
subunits, but not KA1 or other AMPA subunits, were expressed
(data not shown) closely mirroring the previously determined
RGC pattern of expression in vivo (Brandstatter et al., 1994;
Grunder et al., 2000a,b). We had previously reported that RGCs
in vitro express functional non-NMDA glutamate receptors that
545
are blocked by CNQX and that GluR2 specific antiserum stain
synaptic puncta in RGC cultures (Pfrieger and Barres, 1997;
Ullian et al., 2001), and that in situ, RGCs express GluR2 (J.S.D.,
unpublished observations). However, we had not yet examined
their NMDA receptor protein expression.
To determine whether NMDA receptor proteins were present,
we examined extracts from purified P6 RGCs by Western
blotting with subunit specific antibodies. NR1, NR2A, and
NR2B were all easily detected in RGC extracts prepared after
3 (Fig. 1A) and 7 (data not shown) days in culture. To examine
the localization of these proteins, we performed immunostaining
on RGCs after 7 days in culture using NMDA receptor-specific
antibodies. Bright immunoreactivity for the NR1, NR2A, and
NR2B subunits was distributed throughout the soma and dendritic processes of all RGCs (Figs. 1B, C; labeling for NR2A is
not shown but is identical to that shown for NR2B). We could
not discern whether these subunits were concentrated at synapses
because high levels of staining precluded detection of discrete
synaptic puncta. These data indicate that purified RGCs continue
to express NR1, NR2A, NR2B subunit proteins after 7 days of
culture and appropriately target them to their dendrites.
Expression of functional NMDA channels by purified RGCs in
culture
Although RGCs in vitro express the appropriate subunits to
form NMDARs, we did not know if the receptors are actually
functionally expressed by the neurons. To determine if RGCs
express functional NMDARs, we performed whole-cell patchclamp recordings on purified P6 RGCs cultured for 7 days. In
response to applied NMDA (100 AM), we recorded inward currents
that were blocked by the NMDA antagonist APV (50 AM; Fig.
2A). On average, these currents were small, about 50 pA (Fig. 2C,
N = 6 per condition). These findings demonstrate that purified
RGCs do express functional NMDARs.
We wondered how the size of NMDA currents in our RGC
cultures would compare to those of hippocampal neurons in culture.
We prepared hippocampal cultures from E18 hippocampus and after
2 – 3 weeks of culture and analyzed their currents by whole-cell
patch clamping. In response to applied NMDA (100 AM), we once
again recorded inward currents that were blocked by APV (Fig.
2B). On average, these currents were significantly larger than those
we recorded from RGCs, averaging about 1396 F 177 pA (mean F
SEM, n = 5).
Fig. 1. Purified retinal ganglion cells express NMDA receptor proteins in vitro. (A) Western blots performed on samples harvested from purified P5 – P6 RGCs
after 3 days in culture (DIC) show the presence of NR1, NR2A, and NR2B subunit proteins. (B, C) Immunocytochemistry affirms the presence of NR1 and
NR2B subunit proteins in these cultures. Scale bar, 20 Am.
546
E.M. Ullian et al. / Mol. Cell. Neurosci. 26 (2004) 544–557
Fig. 2. RGCs in vitro have functional NMDA currents. RGCs and
hippocampal neurons were cultured for 3 weeks in defined, serum-free
conditions. Whole-cell patch-clamp recordings reveal that NMDA (100 AM)
results in a sustained, APV (50 AM)sensitive currents in both RGCs (A) and
hippocampal neurons (B). The average whole-cell current amplitudes are
greater in hippocampal neurons compared to RGCs. Example of NMDA
current trace from RGCs treated with control FC antibody (C) or ephrin (D)
shows increased current amplitude in response to ephrin. RGC NMDA
current amplitudes change with culture conditions (E). RGCs cultured under
conditions that increase synapse formation such as with astrocytes (RGC +
Astro) or with mixed retina (RGC + Mixed Retina) show an increase in
NMDA current. Addition of ephrin (RGC + Eph) also increases current
amplitude both in the presence of astrocytes (RGC + Eph + Astro) and in the
absence of astrocytes (RGC + Eph; P < 0.001 Kruskal – Wallis ANOVA, N =
6 per condition).
Effects of glutamate and its agonists on RGCs in vitro
We next investigated whether RGCs exposed to plateau levels
of glutamate or its receptor agonists would undergo excitotoxic
death. We cultured purified P6 RGCs for 3 days and then exposed
them to glutamate (500 AM), NMDA (500 AM, 10 mM), AMPA
(500 AM), or kainate (KA; 500 AM) for 1 h at 37jC in 10% CO2.
We then measured the percentage of cells surviving using a calcein
AM/ethidium homodimer-1 live/dead assay as well as by visualizing cellular and nuclear morphology under Nomarski differential
interference contrast (DIC) optics and DAPI fluorescence (Figs.
3A – F; see Experimental methods). Surprisingly, RGCs were not
killed by glutamate or its agonists including NMDA and kainate
(Figs. 3A – F, 4). Indeed, 100% of the purified RGCs survived after
exposure to these agonists for 1 h (Fig. 4) or even after 1 week
(data not shown). Because it has been found that in some cases
excitotoxicity is enhanced at low cell density (Tezel et al., 1999), in
the presence of serum (Erdo et al., 1990), in the presence of
chronic depolarization (Ha et al., 2002) or high levels of extracellular calcium (Hahn et al., 1988), or after ageing in culture for
several weeks (Choi, 1992), we repeated these studies culturing the
RGCs at low density (500 cells/cm2), in serum-containing media,
in high concentrations of K+ (40 mM) or Ca2+ (10 mM), or for
2 weeks of maturation before glutamate agonist exposure or when
RGCs were acutely platted into excitatory amino acids. In all cases,
greater than 90% of the RGCs survived upon exposure to glutamate, kainate, or NMDA, compared with control RGC cultures
lacking glutamate or its agonists. Thus, despite expression of
functional NMDA and non-NMDA glutamate currents, purified
RGCs in culture are invulnerable to glutamate and NMDA-mediated excitotoxicity.
In contrast to these results, it has been reported that even a 5-min
exposure to NMDA (200 AM) or glutamate (500 AM) produces
widespread excitotoxic death in cultured cortical, hippocampal, and
spinal neurons (Buisson et al., 1996; Choi, 1985). In particular,
cultured hippocampal neurons are reported to be extremely vulnerable to NMDA-mediated excitotoxicity. (Peterson et al., 1989).
Thus, to directly compare the effects of NMDA on RGCs and
hippocampal neurons, to provide a positive control that our agonists
were active and that we could elicit and detect excitotoxicity, we next
applied NMDA to hippocampal neurons in culture at the same
concentration (10 mM) and exposure time (1 h at 37jC in 10% CO2)
used for the RGCs. In contrast to RGCs, virtually all of the
hippocampal neurons were rapidly destroyed by NMDA exposure
as assayed by the fluorescent live/dead assay (Figs. 3I, L) and by
visualizing cellular and nuclear morphology under DIC optics and
DAPI fluorescence (Figs. 3G – H, J – K). These results show that, as
expected, cultured hippocampal neurons are highly vulnerable to
NMDA-mediated excitotoxicity whereas cultured RGCs are not.
Effects of ephrins, astrocytes, and synapse formation on RGC
excitotoxicity
One obvious difference between RGCs and hippocampal neurons is that the RGCs have far smaller NMDA currents. Ephrin
signaling significantly increases NMDAR-dependent calcium
influx into cultured cortical neurons (Dalva et al., 2000; Takasu
et al., 2002). Because NMDA-induced calcium influx is thought to
contribute to its excitotoxicity to RGCs (Hahn et al., 1988) and
RGCs are well described to express ephrin receptors, we examined
whether ephrin stimulation would increase NMDA currents in
RGCs. We clustered ephrinB1 and ephrinB2 using antihuman Fc
(50 ng/ml) at 500 ng/ml and activated the Eph receptors on RGCs
with these clustered, multimeric ligands (Dalva et al., 2000; see
Experimental methods). We found that ephrin treatment induced a
4-fold increase in NMDA currents in RGCs (Figs. 2C – E). This
increase was not accounted for by a change in synapse number per
RGC, which was not affected by ephrin (average number of
E.M. Ullian et al. / Mol. Cell. Neurosci. 26 (2004) 544–557
547
Fig. 3. Purified retinal ganglion cells, but not hippocampal neurons, are invulnerable to NMDA-mediated excitotoxicity in culture. Purified RGCs (A – F) and
hippocampal neurons (G – L) were cultured for 3 weeks in defined, serum-free conditions and were then treated for 1 h with NMDA (10 mM) (D – F; J – L) or
control Neurobasal-Sato (A – C; G – I). Side-by-side Nomarski and DAPI-labeled fluorescent microscopy reveal the cellular and nuclear morphology of healthy
RGCs in both control (A and B) and NMDA-treated (10 mM) (D and E) cultures. Additionally, the fluorescent live/dead assay of calcein AM (live cells; green)
and ethidium homodimer-1 (dead cells, nuclear label; red) affirms the lack of RGC death in the same control (C) and NMDA-treated (F) cultures. In contrast,
panels G – I show healthy hippocampal neurons in the control condition, while J – L show dead hippocampal neurons treated with NMDA (10 mM). Scale bars,
20 Am (A, B, D, E, G, H, J, K), 50 Am (C, F, I, L), N = 27 fields.
synapses 6 F 1 synapses per cell with eprhin or control, N = 6).
Despite the increased NMDA currents, however, the RGCs
remained invulnerable to NMDA excitotoxicity (control survival
100 F 3%, NMDA 102 F 3%, GLUT 98 F 3%; P = 0.63, N = 46
fields per condition).
Another obvious difference between the RGC and hippocampal
cultures was that few synapses are present in our purified RGC
cultures (Ullian et al., 2001), whereas many synapses are present in
the hippocampal cultures which contain astrocytes. To determine if
synapse number or astrocytes affect the levels of NMDA currents
or the vulnerability of RGCs to NMDA, we increased the levels of
synapse formation in RGCs by co-culturing purified, retrogradely
labeled RGCs with a feeding layer of astrocytes. RGCs exposed to
astrocytes have nearly 50 synapses per neuron, which is about 7fold more synapses than RGCs not exposed to astrocytes (Nagler
et al., 2001; Ullian et al., 2001). Co-culture with astrocytes
significantly enhanced the average NMDA receptor currents in
RGCs by almost 3-fold (Fig. 2C). Despite their increased NMDA
currents and synapse number, RGCs co-cultured with astrocytes
were still invulnerable to NMDA-mediated excitotoxicity (control
survival 100 F 2%; NMDA 99 F 2%; P = 0.782 by paired t test).
These results indicate that synapse number does not affect RGC
vulnerability to excitotoxicity in vitro.
Finally, another difference between the RGC and hippocampal
neuronal cultures is that multiple types of neurons are present in
the hippocampal but not the purified RGC cultures. To test if other
548
E.M. Ullian et al. / Mol. Cell. Neurosci. 26 (2004) 544–557
in the mixed retinal cultures. After 7 days, the cultures were
exposed to glutamate (500 AM) or NMDA (500 AM) for 1 h at
37jC. Once again, the RGCs were invulnerable to NMDA-mediated excitotoxicity (control survival 100 F 2%, NMDA 105 F
1%). Taken together, these results indicate that increasing synapse
number, adding other retinal cell types, and enhancing the size of
NMDA currents are all insufficient to enhance RGC vulnerability
to glutamate or NMDA.
Localization of NMDARs
Fig. 4. Quantification of glutamate-mediated excitotoxicity in purified
RGCs in culture. Graph of the percent of RGC survival determined by an
MTT assay after 1-h exposure to glutamate (500 AM), NMDA (500 AM),
AMPA (500 AM), and kainate (KA; 500 AM). No statistically significant
difference was found among conditions (P = 0.202 by one-way ANOVA,
N = 46 fields per condition).
cells within the retina can induce RGC susceptibility to excitotoxicity, we cultured fluorogold retrogradely labeled RGCs with
mixed retina (see Experimental methods). RGCs cultured with
other neuronal types in mixed retinal cultures had a 4-fold increase
in NMDA currents (Fig. 2C). To determine if RGCs had become
susceptible to excitotoxicity, we next examined their vulnerability
Another difference between RGCs and hippocampal neurons is
that hippocampal neurons have both extrasynaptic and synaptic
NMDARs (Sattler et al., 2000; Tovar and Westbrook, 1999)
whereas RGCs in vivo have NMDA receptors primarily at
extrasynaptic sites (Chen and Diamond, 2002; Taylor et al.,
1995). To see if RGCs in vitro also express NMDARs at
extrasynaptic sites, we recorded from RGCs cultured under lowdensity autaptic culture conditions where they are forced to form
synapses onto themselves (Ullian et al., 2001). We used patch
clamping to record spontaneous miniature postsynaptic currents
(mEPSCs) from the autaptic RGCs cultured with astrocyte feeding
layers. We found that all of the mEPSCs are sensitive to CNQX
(Fig. 5A). We were unable to observe an NMDA-mediated
component to the mEPSCs although we recorded under conditions
that allow maximal activation of NMDARs by the synaptically
released glutamate (Fig. 5A, 0 Mg2+ and 20 AM glycine). The
mEPSCs decays were best fit by a single exponential with a fast
Fig. 5. RGCs in vitro do not have an NMDA component to the spontaneous miniature currents (mEPSCs), but do have an NMDA component to the evoked
response. (A) Spontaneous mEPSCs recorded in the presence of glycine (20 AM) and the absence of Mg2+. All events were blocked by CNQX (A, CNQX). (B)
Average of mEPSCs taken from (A) is best fit by a single exponential of 1.1 ms time constant. No obvious slow component to the mEPSCs is present. (C).
Evoked responses of the same cell recorded in (A) and (B) show a slow, APV-sensitive component to the evoked response. The slow component is insensitive
to CNQX treatment (C). Similar results were found for all RGCs (N = 5).
E.M. Ullian et al. / Mol. Cell. Neurosci. 26 (2004) 544–557
549
mediate a component of the EPSC indicating that the NMDARs
expressed by RGCs in vitro are extrasynaptic (Chen and Diamond,
2002; Clark and Cull-Candy, 2002).
Size of NMDA currents in RGCs in intact retinas
Given the relatively small size of the NMDA currents that we
observed in RGCs in vitro and their lack of vulnerability to
excitotoxicity, we next investigated whether the reported vulnerability of RGCs in vivo might reflect the presence of larger NMDA
currents in vivo. To examine the NMDA current in intact retina, we
recorded from RGCs using whole-cell patch clamping in P21
retinal slices as previously described (Chen and Diamond, 2002).
First, we determined the size of currents evoked by direct application of NMDA (100 AM) at a holding potential of +40 mV. The
NMDA was puffed directly onto RGC dendrites in the inner
plexiform layer. On average, this elicited a current of about 300
pA (Figs. 6A, C), surprisingly close to the average size of currents
we recorded from RGCs in mixed retinal cultures. We also
determined the size of NMDA currents induced by synaptic
stimulations using whole-field illumination (Figs. 6B, C). The
synaptic NMDA responses were on average about 200 pA (Fig.
6C). Thus, the NMDA currents elicited in RGCs in vitro and in situ
are comparably small.
Effects of glutamate and NMDA on RGC excitotoxicity in intact
retinas
Fig. 6. RGCs in retinal slices have an NMDA component to synaptic
responses and small responses to applied NMDA. All experiments were
performed at 35jC at a holding potential of +40 mV. (A) Responses to 100
AM NMDA puffed onto the IPL. Glycine (10 mM) and strychnine (10 AM)
were included in the puff solution. (B) EPSCs evoked by full-field light
stimuli. All experiments in (A) were performed in slices from P21 animals.
(C) Summary of responses for the population of neurons (error bars indicate
SEM, N = 8 puff, N = 10 light evoked).
time constant (Fig. 5B, s = 1.1 F 0.17 ms, mean F SD, N = 5).
This fast single-exponential time constant is consistent with the
currents being mediated by AMPARs, but not also by NMDARs
(Chen and Diamond, 2002; Gomperts et al., 2000; O’Brien et al.,
1997; Taylor et al., 1995).
In contrast to the spontaneous mEPSCs, evoked postsynaptic
currents (EPSCs) in the autaptic RGCs showed both AMPA and
NMDA components to the synaptic response. For instance, evoked
current from the same neuron used to record the representative
mEPSCs shown in Fig. 5B showed a slow decay component to the
EPSC (Fig. 5C, Control). This slow component was eliminated by
the NMDAR antagonist APV (Fig. 5C, APV) whereas only the fast
component was eliminated by the AMPAR antagonist CNQX (Fig.
5C, CNQX). Thus, NMDARs do not mediate mEPSCs but do
Because of the invulnerability of RGCs in vitro to excitotoxicity,
but the similarity of RGC glutamate and NMDA responses in vitro
and in situ, we decided to reexamine the vulnerability of RGCs in
intact retinas to excitotoxicity. Rather than studying the effects of
glutamate injection in vivo, we chose the intact isolated (ex vivo) rat
retina (Izumi et al., 1995; Romano et al., 1998). This preparation
offers important advantages in that glutamate (or agonist) levels can
be retained at a stable high concentration without rapidly diffusing
away as would occur in vivo, and furthermore allows analysis of
excitotoxicity immediately after isolation of the retina so that
glutamate receptor expression in the retinal cells matches that in
vivo. It has been shown that the effects of glutamate excitotoxicity
on the ex vivo retinas match those that occur after injection in vivo
with extensive swelling and death of retinal cells in the inner nuclear
layer and retinal ganglion cell layers.
The RGC layer is composed of about equal numbers of RGCs
and displaced amacrines (Linden and Esberard, 1987). Therefore,
to ensure accurate identification of RGCs, we first retrogradely
labeled the RGCs in P6 rats with fluorogold allowing us to
selectively label greater than 99% of the RGCs (see Experimental
methods). After 1 – 14 days, we sacrificed the pups, carefully
dissected out their retinas, and immediately studied the effects of
glutamate and NMDA on RGC survival in the intact retinas. We
examined a large number of retinas over a large range of developmental ages. In a series of experiments, we examined the
following age rats: P6 – P7 (25 retinas per condition), P10 (15
retinas per condition), P14 (5 retinas per condition), and P20 (10
retinas per condition). After carefully dissecting out the retinas, we
immediately exposed them to either glutamate (500 AM or 10 mM)
or NMDA (500 AM or 10 mM) in DPBS for 1 h at 37jC in 10%
CO2. We then labeled the dead cells with propidium iodide, fixed
the retinas, and finally either whole-mounted or sectioned the
retinas for analysis (see Experimental Methods).
550
E.M. Ullian et al. / Mol. Cell. Neurosci. 26 (2004) 544–557
Fig. 7. Retinal ganglion cells are invulnerable to NMDA-mediated excitotoxicity in the intact retina. P20 retinas were dissected, immediately exposed to
agonist, and then fixed and whole-mounted. RGCs retrogradely labeled with fluorogold appear in green; propidium iodide-labeled nuclei, dead cells appear in
red. In the whole mounts, both control (A) or NMDA (10 mM) (C) conditions show healthy RGCs in the ganglion cell layer (GCL) with almost exclusive nondouble-labeling of fluorogold (green) and propidium iodide-labeled nuclei (red), indicating that the affected cells are not RGCs but displaced amacrine cells.
Cross-sections of P7 retinas from control (B) and NMDA-treated (10 mM; D) retina further illustrate the absence of RGC death. In sectioned, NMDA-treated
retinas (D), a significant number of the propidium iodide-labeled nuclei (arrows) are in the inner-half of the inner nuclear layer (INL), suggesting that the
affected cells are amacrine cells. In (C), the affected displaced amacrine cells are swollen and have primarily PI-labeled cytoplasm, whereas in (D) the affected
amacrine cells have PI-labeled nuclei indicating that they are dead. Scale bar, 20 Am (A, C), 30 Am (B, D).
In every retina that we examined, exposure to NMDA (Fig. 7)
or glutamate (data not shown) produced no excitotoxic RGC death,
as evidenced by counting the number of green fluorogold-labeled
RGCs in the ganglion cell layer (GCL) that co-localized with red
propidium iodide labeling of nuclei in whole mount P5 – P20
retinas. For instance, in P20 retinas, NMDA incubation did not
induce propidium iodide labeling of RGC nuclei (Figs. 7A, C).
Rare sporadic cells showed labeled nuclei in both control and test
retinas presumably due to injury incurred during the retinal
dissection. Similarly, retinal sections of NMDA-treated P7 retinas
(Figs. 7B, D) show a nearly complete absence of co-labeled RGCs
and propidium iodide-labeled nuclei in the GCL. However, in the
same retinas, significant cell death occurs within the inner half of
the inner nuclear layer (INL) in NMDA-treated retinas (Fig. 7D),
as indicated by bright propidium iodide labeling of amacrine
nuclei, indicating that amacrine cells are highly vulnerable to
NMDA-mediated excitotoxicity. In addition, within the GCL layer,
many non-fluorogold labeled neurons underwent dramatic swelling
in response to glutamate or NMDA exposure and became strongly
labeled with propidium iodide that leaked into their cytoplasm. As
our fluorogold labeling procedure labels at least 99% of RGCs
(Barres et al., 1988), these PI-labeled cells in the GCL are
displaced amacrine cells, which constitute 40 – 50% of cells in
the GCL (Figs. 7C, D). These results were found for retinas of all
ages tested from P5 to P20. These findings indicate that in both
developing and mature intact retinas, RGCs are invulnerable to
glutamate and NMDA excitotoxicity whereas amacrine cells are
highly vulnerable.
Discussion
Purified RGCs express functional extrasynaptic NMDARs and
NMDA current density is greatly enhanced by other cell types
RGCs in mixed retinal cultures have previously been shown to
express NMDA currents (Aizenman et al., 1988; Taschenberger et
al., 1995), but we investigated whether highly purified RGCs in
culture similarly expressed NMDA currents because of their
invulnerability to NMDA excitotoxicity. The present findings show
that purified RGCs express the NMDA receptor subunits NR1,
NR2A, and NR2B, as measured by immunostaining and Western
blotting, and that these subunits are assembled into functional
NMDARs.
We found that the size of the NMDA currents in RGCs was
significantly increased by ephrin signaling, but that this did not
induce NMDA excitotoxicity. The ability of ephrin signaling to
enhance NMDA-dependent calcium influx in cortical neurons is
E.M. Ullian et al. / Mol. Cell. Neurosci. 26 (2004) 544–557
thought to be mediated by a posttranslational modification by
NMDAR tyrosine phosphorylation (Dalva et al., 2000; Takasu et
al., 2002). In these previous studies, however, the effects of ephrin
signaling on NMDA-induced current size were not examined. Such
a modification could increase NMDA current either by increasing
the number of NMDA receptors on a neuron’s surface, their
probability of opening, or their open times. Any of these changes
would explain the increase in NMDA-induced calcium current. In
these earlier studies, the possibility was raised that the increase in
calcium influx was accounted for by an increased number of
synapses per neuron. Our results show, however, that ephrin
signaling can profoundly increase NMDA-induced currents without altering synapse number.
The NMDA currents in RGCs were also increased severalfold by cell – cell interactions, including astrocytes and mixed
retinal cell types containing astrocytes and Muller glia. Because
astrocytes increase the number of synapses per RGC by 7-fold
(Ullian et al., 2001), the increase in NMDA current could
represent an increase in surface NMDAR expression following
synapse formation. Alternatively, ephrins secreted by astrocytes
could enhance the RGC NMDA currents (Murai et al., 2003).
These results provide evidence that NMDA-current amplitudes
in RGCs can be strongly regulated by cell – cell signaling
interactions.
We also investigated the localization of the functional NMDA
receptors in RGCs. In hippocampal neurons, NMDA receptors are
found both synaptically and extrasynaptically (Tovar and Westbrook, 1999). We found that the NMDA receptors expressed by
purified RGCs in culture were predominantly extrasynaptic, even
in the presence of astrocytes, as has been previously found for
RGCs in vivo (Chen and Diamond, 2002; Matsui et al., 1998, but
see Fletcher et al., 2000). It is not known how localization of
NMDA receptors is controlled. Because in our purified cultures,
RGCs synapse upon each other, whereas in vivo cone bipolar
neurons synapse upon RGCs, it is likely that RGCs play the
dominant role in determining NMDAR localization by directing
them to extrasynaptic sites.
RGCs are invulnerable to glutamate and NMDA excitotoxicity, in
vitro and in intact retinas, whereas amacrine cells are highly
vulnerable
We have been unable to induce acute glutamate or NMDAmediated excitotoxicity in RGCs in intact rat retinas or in vitro,
even under conditions where the NMDA currents are relatively
large. In contrast, we observed that many amacrine cells in the
inner nuclear layer were rapidly and dramatically injured or
killed by glutamate and NMDA and that displaced amacrine
cells in the GCL became swollen. Amacrine cells are well
reported to express NMDAR proteins and currents in the rodent
retina (Brandstatter et al., 1994; Grunder et al., 2000a,b;
Hartveit and Veruki, 1997; Matsui et al., 2001). These studies
suggest that AII (rod) amacrine cells, cholinergic amacrine cells,
and displaced amacrine cells in particular express NMDARs.
Remarkably, despite the presence of NMDA currents in these
subsets of amacrine cells, electrophysiological recordings reveal
only small NMDA currents on average of about 50 pA at a +40
mV holding potential in response to applied NMDA or evoked
excitation (Hartveit and Veruki, 1997; Matsui et al., 2001). The
toxic effects of NMDA and glutamate on amacrines in the intact
retinas were manifested after only 1 h of exposure, which was
551
the earliest time point we examined. It is likely that this damage
may have occurred even within minutes of exposure to glutamate and NMDA, as occurs with hippocampal neurons. Thus,
our findings show that RGCs are invulnerable to glutamate and
NMDA excitotoxicity, whereas amacrine cells are highly vulnerable yet have smaller NMDA currents.
In contrast, a large previous body of literature has reported
that RGCs are highly vulnerable to glutamate and NMDA
excitotoxicity in vitro (Caprioli et al., 1996; Dreyer et al.,
1994; Kawasaki et al., 2000, 2002; Kitano et al., 1996; Otori
et al., 1998; Sucher et al., 1991a,b) and in intact ex vivo retinas
or in vivo (Izumi et al., 1995; Kido et al., 2000; Li et al., 1999;
Lucas and Newhouse, 1957; Moncaster et al., 2002; Moore et
al., 2001; Mosinger et al., 1991; Olney, 1969; Romano et al.,
1998; Sabel et al., 1995; Silprandi et al., 1992; Sisk and
Kuwabara, 1985; Sun et al., 2000; Vorwerk et al., 1996).
How can the discrepancy between our findings and these
previous studies be explained? Lucas and Newhouse (1957),
Olney (1969), and Sisk and Kuwabara (1985) were the earliest
investigators to report that glutamate and NMDA agonists
produced a rapid and severe destruction of the inner retinal
layers including the retinal ganglion cell layer. As nearly half of
the cells in the rodent ganglion cell layer are displaced amacrine
cells, these observations were consistent either with amacrine
cells being primarily affected or with both amacrine and retinal
ganglion cells being affected. Similarly, the majority of later
studies investigating ex vivo and in vivo studies of retinal
glutamate exposure concluding that RGCs were affected failed
to take any measure to distinguish RGCs from displaced
amacrine cells, for instance by retrograde labeling or antigenic
identification. Thus, whereas all studies agree that many retinal
cells in the INL and RGC layers are vulnerable to excitotoxicity, the difference between the present study and these
previous studies may be more of a matter of mistaken previous
interpretations that RGCs were among the affected cells. The
present findings, however, reveal that it is amacrine cells and
not RGCs that are subject to rapid glutamate and NMDA
excitotoxicity in intact retinas and in vivo.
An important point is that our studies examined retinas
acutely after a 1-h exposure to glutamate or NMDA. In many
previous studies, retinas have been examined 1 – 7 days after
intraocular injections of these drugs and a significant thinning
of the GCL was found. In only two of these studies were the
RGCs identified by using retrograde injection of a fluorescent
dye (Manabe and Lipton, 2003; Sun et al., 2000). These
investigators clearly identified death of some RGCs by apoptosis in response to NMDA and, interestingly, this death took at
least 24 h to begin. Delayed apoptosis in the GCL several days
after NMDA injection into rat retina was also observed by Lam
et al. (1999), Kido et al. (2000), and Moore et al. (2001).
Because we did not observe death of RGCs in culture in
response to prolonged exposure of glutamate or NMDA, the
simplest explanation of our findings taken together with these
previous investigators is that NMDA induces rapid loss of
amacrine cells and that apoptotic death of some RGCs follows
because of trophic deprivation from loss of amacrine signals
rather than delayed excitotoxicity.
How can we account for the discrepancy between our culture
studies and those of others who reported that RGCs in vitro are
vulnerable to glutamate and NMDA excitotoxicity? RGC survival strongly depends on target-derived trophic signals and thus
552
E.M. Ullian et al. / Mol. Cell. Neurosci. 26 (2004) 544–557
RGCs in culture rapidly undergo cell death unless appropriate
neurotrophic support is specifically added to the culture medium. As appropriate trophic signals sufficient to support longterm RGC survival were not available before 1995, most of the
early studies of RGC excitotoxicity involved short-term culture
studies of 1 or 2 days at most, during which time the RGCs
were rapidly dying even without any glutamate exposure. In
contrast, our study involved use of RGCs highly purified using
immunopanning that were provided with strong neurotrophic
signals such as BDNF, CNTF, IGF-1, and laminin that promote
their survival (Meyer-Franke et al., 1995).
Otori et al. (1998), however, also used purified cultures of
RGCs under conditions somewhat similar to those we used, yet
observed severe kainate excitotoxicity that we did not observe.
One difference is that IGF-1 was omitted from their culture
conditions; thus, it is likely that RGC viabilities in their cultures
were substantially lower than under the conditions we used
(unfortunately they did not report their raw viabilities). Kainate
activates both AMPA and kainate classes of glutamate receptors,
at least some of which are calcium permeable in RGCs
(Leinders-Zufall et al., 1994), and thus could be toxic particularly if viabilities were low. Nonetheless, Otori et al. (1998) is
one of the few reports of kainate toxicity of RGCs in culture;
most other reports of RGC excitotoxicity in culture have
focused on NMDA toxicity (Aizenman et al., 1988; Lipton,
2001; Pang et al., 1999). Moreover, any toxicity of kainate in
mixed retinal cultures and in intact retinas is thought to be due
to release of glutamate, which then acts upon NMDA receptors
on RGCs (Sucher et al., 1991a, 1997). Finally, another important variable increasingly coming to light is that the cell type
affected by excitotoxicity, as well as the degree of excitotoxicity, may depend on the species and even the strain of the
experimental animal used (Bakalash et al., 2002; Luo et al.,
2001; Schori et al., 2002). We cannot exclude the possibility of
some other variable between ours and previous studies that may
account for why we do not observe excitotoxicity of RGCs in
vitro. Whatever the explanation for the differences between our
findings and previous findings of others, our studies show
clearly that the purified RGCs in our cultures are behaving
exactly the way they behave in intact retinas in that they are
invulnerable to glutamate and NMDA excitotoxicity.
Glutamate excitotoxicity probably does not contribute substantially
to RGC death in retinal ischemia and glaucoma
It is presently thought that glutamate and NMDA excitotoxicity contributes to loss of RGCs in retinal ischemia and glaucoma (Dreyer et al., 1996; Lipton, 2001; Naskar and Dreyer, 2001;
Osborne et al., 1999). Our findings raise some doubts. Glutamate
concentrations become acutely elevated in ischemia and are
reported to be elevated chronically in glaucoma (Dreyer et al.,
1996; Naskar and Dreyer, 2001, but see Dalton, 2001). However,
RGCs die within a few hours after retinal ischemia (Osborne et
al., 1999), whereas only delayed death of RGCs is observed in
response to excitotoxicity (Manabe and Lipton, 2003). Similarly,
chronic elevation of glutamate would be expected to kill amacrine
cells, which are highly vulnerable to excitotoxicity, yet in
glaucoma only loss of RGCs is observed (Quigley, 1999; Wax
and Tezel, 2002). Thus, our observations do not lend support to
an important contribution of glutamate excitotoxicity to retinal
ischemia and glaucoma and in fact suggest that it does not.
Why are RGCs invulnerable to NMDA excitotoxicity whereas
amacrine cells are highly vulnerable?
Why are RGCs not killed by NMDA when amacrine cells
with similarly small NMDA currents are highly vulnerable? One
factor that has been linked to excitotoxicity is the presence of
functional synapses. Embryonic hippocampal neurons are not
vulnerable to NMDA excitotoxicity until they have been cultured
for 3 weeks and synaptogenesis has occurred (Choi et al., 1987).
Perhaps the most striking example of uncoupling the level of
glutamate current to the level of excitotoxicity has been found in
the striatum. Striatal neurons are highly vulnerable to kainate
exposure but are much less sensitive after synapses are removed
by decortication, which does not alter their kainate sensitivity
(Biziere and Coyle, 1978; McGeer and McGeer, 1976). Similarly,
striatal NMDA excitotoxicity is largely eliminated by decortication to remove glutamatergic synapses (Orlando et al., 2001).
However, when we increased the number of synapses that formed
between the RGCs, we were unable to induce glutamate or
NMDA excitotoxicity although we did increase their NMDA
currents 4-fold.
One intriguing difference between RGCs and hippocampal
neurons is that RGCs do not appear to localize NMDARs
directly underneath sites of vesicle release both in situ (Chen
and Diamond, 2002) and in vitro (Taschenberger et al., 1995;
this paper) but instead are extrasynaptic. Could NMDAR
localization play a role in susceptibility to excitotoxicity?
Synaptic NMDARs may have a different subunit composition,
different kinetics, and be associated with different signaling
components (Hardingham et al., 2002; Li et al., 2002; Tovar
and Westbrook, 1999, 2002; Washbourne et al., 2002). Perhaps
when NMDARs are not localized under release sites cells are
intrinsically more resistant to excitotoxicity, both because currents are reduced and because signaling molecules that contribute to cell death are not activated. On the other hand, in cortical
neurons, both synaptic and extrasynaptic NMDA receptors are
able to mediate excitotoxic death (Sattler et al., 2000). Furthermore, in preliminary studies, the NMDA receptors of amacrine
cells appear to be extrasynaptically localized (JSD and Joshua
Singer, unpublished observations). Thus, as amacrine cells are
highly vulnerable, the extrasynaptic localization of NMDA
receptors on RGCs is probably not sufficient to account for
their invulnerability to excitotoxicity.
There are several other apparent differences between retinal
ganglion cells and amacrine cells that may contribute to their
different vulnerabilities to NMDA. First, it has recently been
found that the NR3A subunit of the NMDA receptor is
expressed strongly by RGCs but not by amacrine cells in the
INL (Sucher et al., 2003). Consistent with the known dominant
negative action of NR3 family members (Nishi et al., 2001),
Sucher et al. (2003) found that NMDA-evoked intracellular
calcium responses were significantly greater in NR3A deficient
retinas. Thus, the expression of NR3A by RGCs but not by
amacrine cells in the INL could help to explain a lower
vulnerability to NMDA. Second, neuronal NOS bound to
NMDARs via PSD-95 enhances excitotoxicity (Sattler et al.,
1999) and amacrine cells and displaced amacrines have been
reported to have much higher levels of nNOS (Kim et al., 2000;
Shin et al., 1999; Yamamoto et al., 1993). Whatever the
mechanism, the difference in excitotoxic vulnerability between
RGCs and amacrine cells adds to the growing evidence that
E.M. Ullian et al. / Mol. Cell. Neurosci. 26 (2004) 544–557
NMDA current density is not the sole determinant of susceptibility, but that other downstream signaling mechanisms play
critical roles (Arundine and Tymianski, 2003).
Experimental methods
Cell purification and culture
Step-by-step protocols for all procedures are available on
request from [email protected]. RGCs from postnatal days
5 – 21 (P5 – P21) albino rats (Simonson rats; Simonson Labs,
Gilroy, CA) were purified as previously described (Barres et al.,
1988). Briefly, dissected retinas were enzymatically dissociated in
papain in Dulbecco’s phosphate-buffered saline; Gibco, Carlsbad,
CA) to create a single-cell suspension. RGCs were isolated from
this suspension using sequential immunopanning to greater than
99.5% purity (Barres et al., 1988). Purified RGCs were plated on
12-mm glass coverslips (Carolina Science and Math, Burlington,
NC) in 24-well tissue culture plates (BD Biosciences, San Jose,
CA) at a density of 30,000 cells per coverslip (approximately
26,000 cells/cm2). Coverslips were coated with poly-D-lysine
(PDL, 70 kDa, 10 Ag/ml; Sigma, St. Louis, MO) at room
temperature followed by overnight incubation with mouse laminin
(Sigma).
RGCs were cultured in 500 Al of defined, serum-free medium,
modified from Bottenstein and Sato (1979). Neurobasal media
(Gibco) contained B27, selenium, putrescine, triiodo-thyronine,
transferrin, progesterone, pyruvate (1 mM; Sigma), glutamine (2
mM; Sigma), ciliary neurotrophic factor (CNTF; 10 ng/ml;
Regeneron Pharmaceuticals, Inc., Tarrytown, NY), brain-derived
neurotrophic factor (BDNF; 50 ng/ml; Regeneron Pharmaceuticals, Inc.), insulin (5 Ag/ml; Sigma), and forskolin (10 AM,
Sigma), as defined in Meyer-Franke et al. (1995). Cultures were
maintained at 37jC in a humidified environment of 10% CO2
(Praxair, Danbury, CT). Under these conditions, more than half of
RGCs survive in vitro for at least 1 month (Meyer-Franke et al.,
1995).
Cortical astrocyte cultures were prepared from first- to second-day postnatal (P1 – P2) albino rats as previously described
(McCarthy and de Vellis, 1980). Briefly, the cortex was dissected
and digested in trypsin and plated in tissue culture flasks (BD
Biosciences) in a medium that does not allow survival of neurons
(Dulbecco’s minimum essential medium, fetal bovine serum
(10%), penicillin (100 U/ml), streptomycin (100 mg/ml), glutamine (2 mM), and Na-pyruvate (1 mM). After 4 days in culture,
nonadherent cells were removed from flasks by shaking. Remaining cells were removed from flasks enzymatically and cultured
on PDL (10 Ag/ml; Sigma) coated tissue culture plates (BD
Biosciences).
Purified RGC excitotoxicity experiments
After 3 – 28 days in culture, the RGCs were exposed to
NMDA, glutamate, AMPA, or kainic acid, each at 500 AM or
10 mM for 1 h, followed by a survival assay (see below). Cells
in the 24-h agonist exposure experiment were cultured for 3 days
and then treated with glutamate or NMDA (500 AM) for 24 h.
Neuronal survival was determined using the 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl tetrazolium bromide (MTT; Sigma)
survival assay as described by Mosmann (1983). MTT (500
553
Ag/ml) was added to cultures and incubated at 37jC for 1 h. The
viable and dead cells in each well were counted by bright-field
microscopy.
Alternatively, cells were incubated for 10 min in calcein-AM
(2 AM) and ethidium homodimer-1 (2 AM) following the manufacturer’s protocol (Molecular Probes, Eugene, OR, USA).
Four fields of cells containing approximately 20 cells per field
were counted for viability in each of three wells per condition.
Experiments were repeated three times.
Purified hippocampal neuron excitotoxicity experiments
Hippocampal neurons from E19 Sprague – Dawley rat embryos
were purified according to the methods of Banker and Goslin
(1998) and cultured in Neurobasal plus Sato reagents with B27
(Gibco) in the presence of astrocyte feeder layers.
Astrocyte or mixed retina co-culture excitotoxicity experiments
RGCs were retrogradely labeling with fluorogold (2.5%,
0.5 Al; Fluorochrome, Englewood, CO) bilaterally injected into
the superior collicular brachium. Sixteen to twenty-four hours
postinjection, P5 – P6 animals were sacrificed and their retinas
removed. The labeled RGCs were then purified as described
above and plated at 15,000 RGCs per 12-mm coverslip. Concurrently, a mixed retinal dissociate was done by removing P5 – P6
retinas and dissociating enzymatically with papain to make a
suspension of single cells, essentially as described by Huettner
and Baughman (1986) and plated at 500,000 per well with the
labeled RGCs. In the case of the astrocyte co-cultures, a feeding
layer of astrocytes on a Falcon insert (BD, Franklin Lakes, NJ)
was added after the RGCs had been in culture for 1 day. After
day 3 in culture, the cultures were exposed to NMDA at 500 AM
or 10 mM as described above. Two hundred micromolars of
NMDA is a non-desensitizing dose (Aizenman et al., 1988, Hahn
et al., 1988). The agonist was then gently rinsed off the cells and
the cells were fixed with paraformaldehyde (PFA, 4%). The
coverslips were then mounted on glass slides using Vectashield
mounting and counted by fluorescent microscopy. Cell death was
determined by morphology and live/dead assay on fluorescently
labeled RGCs.
Addition of ephrins
EphrinB1 and ephrinB2 (500 ng/ml; R & D Systems, Minneapolis, MN) were pre-clustered for 1 h at room temperature
using antihuman Fc (50 ng/ml; Jackson Labs). Following clustering, the ligands were added to the RGC cultures for 2 days.
Cultures were then assayed for cell death or NMDA current
amplitude.
RGC excitotoxicity assay in whole mount retina and retinal
sections
Sixteen hours to sixteen days after retrogradely labeling RGCs
with fluorogold, animals were sacrificed and their retinas removed.
The retinas were then placed in a solution of Dulbecco’s phosphatebuffered saline (DPBS) and the experimental condition was treated
with 500 AM or 10 mM NMDA or glutamate for 1 h while the
control was placed only in DPBS. In some experiments, APV (100
AM) or MK-801 (60 AM) was added 15 min before and during the
554
E.M. Ullian et al. / Mol. Cell. Neurosci. 26 (2004) 544–557
hour exposure to glutamate or NMDA. After the 1 h exposure, the
retinas were incubated with propidium iodide for 15 min at 37jC to
label dead cell nuclei. The retinas were then fixed with 4% PFA for
1 h at room temperature and dissected by making four cuts from the
edge into the center of the retina to flatten them (see Shen et al.,
1999). The whole retinas were mounted on glass slides in Vectashield mounting medium containing DAPI (Vector Laboratories
Inc., Burlingame, CA). For sectioning, after fixation, the retinas
were sunk in 30% sucrose overnight at 4jC and mounted in OCT
(Sakura Finetek, Torrance, CA).
Neuronal death was determined for experimental and control
retinas by analyzing the number of propidium iodide-labeled nuclei
and fluorogold-labeled RGCs in the same area of each retina.
NMDAR immunostaining
Cells were fixed for 10 min in 4% PFA, washed three times
in PBS, and blocked in Blocking buffer (50% goat serum, 50%
PBS, 0.1% NP40). Following blocking, primary antibody incubation were done overnight at 4jC. Primary antibodies used
were NR1 1:500 (PharMingen, San Diego, CA) or NR2B 1:50
(Transduction Laboratories, Lexington, KY). Secondary antibodies, goat anti-mouse Alexa 488 or Alexa 594 conjugates
(Molecular Probes) were used at a dilution of 1:200 for 1 h at
room temperature.
Western blot analysis
Standard SDS-PAGE gel electrophoresis was done as previously described (Meyer-Franke et al., 1998). Samples were collected
in boiling 2% SDS and resolved on polyacrylamide gels and
transferred onto polyvinylidene difluoride (Millipore, Bedford,
MA). Blots were probed with monoclonal antibodies against
NR1 (1 Ag/ml; PharMingen), rabbit polyclonal anti-NR2A (1 Ag/
ml; Upstate Biotechnology, Charlottesville, VA), and monoclonal
anti-NR2B (1 Ag/ml; Transduction Labs).
Horseradish peroxidase-conjugated secondary antibodies
(Jackson ImmunoResearch, Westgrove, PA) were used in combination with chemiluminescent horseradish peroxidase substrate
(ECL+Plus; Amersham, Arlington Heights, IL) at 1:30,000 and
SuperSignal WestPico (Pierce, Rockford, IL) at 1:100,000 to
produce the signals on chemiluminescent sensitive film (Scientific Imaging Systems, Eastman Kodak Company, Rochester,
NY).
Electrophysiology
Membrane currents were recorded by whole-cell patch clamping at room temperature (18jC to 22jC) at a holding potential
of 70 mV unless otherwise specified. Patch pipettes (3 – 10 MV)
were pulled from borosilicate capillary glass (WPI). For recordings
of synaptic currents, the bath solution contained (in mM) 120
NaCl, 3 CaCl2, 2 MgCl2, 5 KCl, and 10 Hepes (pH 7.3). For
recording in Mg2+-free conditions, the external bath solution
contained (in mM) 140 NaCl, 3.5 KCl, 10 HEPES, 20 glucose, 3
CaCl2, and 20 AM glycine (pH 7.3). The internal solution
contained (in mM) 100 K-gluconate, 10 KCl, 10 EGTA (Ca2+buffered to 10 6), and 10 Hepes (pH 7.3). For recordings of
autaptic currents, the internal solution contained (in mM) 122.5
K-gluconate, 8 NaCl, 10 Hepes, 0.2 EGTA, 2 Mg-ATP, 0.3 NaGTP, 20 K2-creatine phosphate, and phosphocreatine kinase (50 U/
ml). Currents were recorded using pClamp software for Windows
(Axon Instruments, Foster City, CA). NMDA currents were
expressed as pA as the membrane capacitances of RGCs were
similar under all conditions. Mini-excitatory postsynaptic events
(mEPSCs) were analyzed using Mini Analysis Program (SynaptoSoft, Decatur, GA) and plotted using SigmaPlot or Origin (Microcal, Northampton, MA).
Electrophysiology of retinal slices
Retinal slices were prepared from Sprague – Dawley rats (17 –
22 days) in accordance with the National Institute of Neurological
Disorders and Stroke Animal Care and Use Committee guidelines. Both eyes were removed and immersed in oxygenated
extracellular solution at room temperature. Extracellular solution
contained (in mM): 119 NaCl, 2.5 KCl, 1.3 MgCl2, 2.5 CaCl2,
26.2 NaHCO3, 1 NaH2PO4, 20 glucose, 2 Na pyruvate, and 4 Na
lactate, bubbled with 95% O2 and 5% CO2. The cornea, iris, lens,
and vitreous were removed from one eye with scissors. The retina
was mechanically detached from the eyecup and immersed in 2%
agarose (low-gelling temperature, type VII; Sigma) and cut into
200-Am-thick slices on a vibratome (Leica, Nussloch, Germany).
Slices were prepared and stored in oxygenated extracellular
solution; they were transferred one at a time to the recording
chamber, in which picrotoxin (100 AM) and strychnine (10 AM)
were added to oxygenated extracellular solution to block inhibitory synaptic transmission. The patch pipette solution contained
(in mM): 120 Cs methanesulfonate, 10 EGTA, 20 HEPES, 2
MgATP, and 0.2 NaGTP. All solutions were adjusted to pH 7.4
with NaOH or CsOH and adjusted to 290 – 300 mOsm with
sucrose. Reagents were obtained from Sigma. All recordings were
made from RGCs (identified post recording by filling neurons
and identifying an axonal process) with an Axopatch 1D amplifier (Axon Instruments) in voltage-clamp mode. Patch electrodes
(#0010 glass; World Precision Instruments, Sarasota, FL) had tip
resistances of 4 – 5 MV when filled with internal solution. Access
resistance was 10 – 20 MV and was monitored continuously. Data
acquisition and analysis were performed with custom macros
written in IgorPro (WaveMetrics, Lake Oswego, OR). Data were
filtered at 5 kHz and sampled at 10 kHz. Responses were elicited
by puffing NMDA (100 AM) onto RGC dendrites in the inner
plexiform layer or the distal part of the inner nuclear layer, or by
field illumination.
Acknowledgments
This work was supported by the National Eye Institute (R01
11030 to B.A.B.), and the March of Dimes Birth Defects
Foundation (FY01-0503 to B.A.B.), a Zaffaroni Fellowship
(E.M.U.), a HHMI undergraduate fellowship (W.B), and the
NINDS Intramural Research Program (S.C and J.S.D.). We thank
Regeneron for generously providing BDNF and CNTF.
References
Aizenman, E., Forsch, M.P., Lipton, S.A., 1988. Responses mediated by
excitatory amino acid receptors in solitary retinal ganglion cells from
rat. J. Physiol. 396, 75 – 91.
E.M. Ullian et al. / Mol. Cell. Neurosci. 26 (2004) 544–557
Arundine, M., Tymianski, M., 2003. Molecular mechanisms of calciumdependent neurodegeneration in excitotoxicity. Cell Calcium 34,
325 – 337.
Bakalash, S., Kipnis, J., Yoles, E., Schwartz, M., 2002. Resistance of
retinal ganglion cells to an increase in intraocular pressure is immune-dependent. Invest. Ophthalmol. Visual Sci. 43, 2648 – 2653.
Banker, G., Goslin, K., 1998. Culturing Nerve Cells. Second ed. MIT
Press, Cambridge, MA.
Barres, B.A., Silverstein, B.E., Corey, D.P., Chun, L.L.Y., 1988. Immunological, morphological, and electrophysiological variation among retinal
ganglion cells purified by panning. Neuron 1, 791 – 803.
Biziere, K., Coyle, J.T., 1978. Influence of corticostriatal afferents on striatal kainic acid neurotoxicity. Neurosci. Lett. 8, 303 – 310.
Bottenstein, J.E., Sato, G.H., 1979. . Proc. Natl. Acad. Sci. U. S. A. 76,
514 – 517.
Brandstatter, J.H., Hartveit, E., Sassoe-Pognetto, M., Wassle, H., 1994.
Expression of NMDA and high affinity kainate receptor subunit
mRNAs in the adult rat retina. Eur. J. Neurosci. 6, 1100 – 1112.
Buisson, A., Yu, S.P., Choi, D.W., 1996. DCG-IV selectively attenuates
rapidly triggered NMDA-induced neurotoxicity in cortical neurons. Eur.
J. Neurosci. 8, 138 – 143.
Caprioli, J., Kitano, S., Morgan, J.E., 1996. Hyperthermia and hypoxia
increase tolerance of retinal ganglion cells to anoxia and excitotoxicity.
Invest. Ophthalmol. Visual Sci. 47, 2376 – 2381.
Chen, S., Diamond, J., 2002. Synaptically released glutamate activates
extrasynaptic NMDA receptors on cells in the ganglion cell layer of
the rat retina. J. Neurosci. 22, 2165 – 2173.
Choi, D.W., 1985. Glutamate neurotoxicity in cortical cell culture is calcium dependent. Neurosci. Lett. 58, 293 – 297.
Choi, D.W., 1992. Excitotoxic cell death. J. Neurobiol. 23, 1261 – 1276.
Choi, D.W., Maulucci-Gedde, M.A., Kriegstein, A.R., 1987. Glutamate
neurotoxicity in cortical cell culture. J. Neurosci. 7, 336 – 357.
Clark, B.A., Cull-Candy, S.G., 2002. Activity-dependent recruitment of
extrasynaptic NMDA receptor activation at an AMPA receptor-only
synapse. J. Neurosci. 22, 4428 – 4436.
Dalton, R., 2001. Private investigations. Nature 411, 129 – 130.
Dalva, M.B., Takasu, M.A., Lin, M.Z., Shamah, S.M., Hu, L., Gale, N.W.,
Greenberg, M.E., 2000. EphB receptors interact with NMDA receptors
and regulate excitatory synapse formation. Cell 103, 945 – 956.
Dingledine, R., Borges, K., Bowie, D., Traynelis, S., 1999. The glutamate
receptor ion channels. Pharm. Rev. 51, 7 – 61.
Dreyer, E.B., Pan, Z.H., Storm, S., Lipton, S.A., 1994. Greater sensitivity
of larger retinal ganglion cells to NMDA mediated cell death. Neuroreport 5, 629 – 631.
Dreyer, E.B., Zuracowski, D., Schumer, R.A., Podos, S.M., Lipton, S.A.,
1996. Elevated glutamate levels in the vitreous body of humans and
monkeys with glaucoma. Arch. Ophthalmol. 114, 299 – 305.
Erdo, S.L., Michler, A., Wolff, J.R., Tytko, H., 1990. Lack of excitotoxic
cell death in serum-free cultures of rat cerebral cortex. Brain Res. 526,
328 – 332.
Goldberg, J.L., Barres, B.A., 2000. The relationship between neuronal
survival and regeneration. Annu. Rev. Neurosci. 23, 579 – 612.
Goldberg, J.L., Espinosa, J.S., Xu, Y., Davidson, N., Kovacs, G.T., Barres,
B.A., 2002. Retinal ganglion cells do not extend axons by default:
promotion by neurotrophic signaling and electrical activity. Neuron
33, 689 – 702.
Gomperts, S.N., Carroll, R., Malenka, R.C., Nicoll, R.A., 2000. Distinct
roles for ionotropic and metabotropic glutamate receptors in the maturation of excitatory synapses. J. Neurosci. 15, 2229 – 2237.
Grunder, T., Kohler, K., Guenther, E., 2000a. Distribution and developmental regulation of AMPA receptor subunit proteins in rat retina.
Invest. Ophthalmol. Visual Sci. 41, 3600 – 3606.
Grunder, T., Kohler, K., Kaletta, A., Guenther, E., 2000b. Distribution and
developmental regulation of NMDA receptor subunit proteins in the rat
retina. J. Neurobiol., 333 – 342.
Ha, B.K., Vicini, S., Rogers, R.C., Bresnahan, J.C., Burry, R.W., Beattie,
M.S., 2002. Kainate-induced excitotoxicity is dependent upon extracel-
555
lular potassium concentrations that regulate the activity of AMPA/KA
type glutamate receptors. J. Neurochem. 83, 934 – 945.
Hahn, J.S., Aizenman, E., Lipton, S.A., 1988. Central mammalian neurons normally resistant to glutamate toxicity are made sensitive by
elevated extracellular Ca2+: toxicity is blocked by the N-methyl-Daspartate antagonist MK-801. Proc. Natl. Acad. Sci. U. S. A. 85,
6556 – 6560.
Hardingham, G.E., Fukunaga, Y., Bading, H., 2002. Extrasynaptic
NMDARs oppose synaptic NMDARs by triggering CREB shut-off
and cell death pathways. Nat. Neurosci. 5, 405 – 413.
Hartveit, E., Veruki, M.L., 1997. AII amacrine cells express functional
NMDA receptors. Neuroreport 8, 1219 – 1223.
Huettner, J.E., Baughman, R.W., 1986. Primary culture of identified
neurons from the visual cortex of postnatal rats. Neuroscience 6,
3044 – 3060.
Izumi, Y., Benz, A.M., Kirby, C.O., Labrueyer, J., Zorumski, C.F., Price,
M.T., Olney, J.W., 1995. An ex vivo rat retinal preparation for excitotoxicity studies. J. Neurosci. Methods 60, 219 – 225.
Kawasaki, A., Otori, Y., Barnstable, C.J., 2000. Muller cell protection of rat
retinal ganglion cells from glutamate and nitric oxide neurotoxicity.
Invest. Ophthalmol. Visual Sci. 41, 3444 – 3450.
Kawasaki, A., Han, M., Wei, J., Hirata, K., Otori, Y., Barstable, C.J.,
2002. Protective effects of arachidonic acid on glutamate neurotoxicity in rat retinal ganglion cells. Invest. Ophthalmol. Visual Sci. 43,
1835 – 1842.
Kido, N., Tanihara, H., Honjo, M., Inatani, M., Tatsuno, T., Nakayama, C.,
Honda, Y., 2000. Neuroprotective effects of BDNF in eyes with NMDA
induced neuronal death. Brain Res. 884, 59 – 67.
Kim, K.Y., Ju, W.K., Oh, S.J., Chun, M.H., 2000. The immunocytochemical localization of neuronal nitric oxide synthase in the developing rat
retina. Exp. Brain Res. 133, 419 – 424.
Kitano, S., Morgan, J., Caprioli, J., 1996. Hypoxic and excitotoxic damage
to cultured rat retinal ganglion cells. Exp. Eye Res. 63, 105 – 112.
Lam, T., Abler, A., Kwong, J., Tso, M., 1999. NMDA induced apoptosis in
rat retina. Invest. Ophthalmol. Visual Sci. 40, 2391 – 2397.
Leinders-Zufall, T., Rand, M.N., Waxman, S.G., Kocsis, J.D., 1994. Differential role of two Ca(2+)-permeable non-NMDA glutamate channels
in rat retinal ganglion cells: kainate-induced cytoplasmic and nuclear
Ca2+ signals. J. Neurophysiol. 72, 2503 – 2516.
Li, Y., Schlamp, C.L., Nickells, R.W., 1999. Experimental induction of
retinal ganglion cell death in adult mice. Invest. Ophthalmol. Visual
Sci. 40, 1004 – 1008.
Li, B., Chen, N., Luo, T., Otsu, Y., Murpy, T., Raymond, L.A., 2002.
Differential regulation of synaptic and extrasynaptic NMDA receptors.
Nat. Neurosci. 5, 833 – 834.
Linden, R., Esberard, C.E., 1987. Displaced amacrine cells in the ganglion
cell layer of the hamster retina. Vision Res. 27, 1071 – 1076.
Lipton, S., 2001. Retinal ganglion cells, glaucoma, neuroprotection. Prog.
Brain Res. 131, 712 – 719.
Lucas, D.R., Newhouse, J.P., 1957. The toxic effect of glutamate on the
inner layers of the retina. Arch. Ophthalmol. 158, 193 – 201.
Luo, X., Heidinger, V., Picaud, S., Lambrou, G., Dreyfus, H., Sahel, J.,
Hicks, D., 2001. Selective excitotoxic degeneration of adult pig
retinal ganglion cells in vitro. Invest. Ophthalmol. Visual Sci. 42,
1096 – 1106.
Manabe, S., Lipton, S.A., 2003. Divergent NMDA signals leading to proapoptotic and antiapoptotic pathways in rat retina. Invest. Ophthalmol.
Visual Sci. 44, 385 – 392.
Matsui, K., Hosoi, N., Tachibana, M., 1998. Excitatory synaptic transmission in the inner retina: paired recordings of bipolar cells and neurons in
the ganglion cell layer. J. Neurosci. 18, 4500 – 4510.
Matsui, K., Hasegawa, J., Tachibana, M., 2001. Modulation of excitatory
synaptic transmission by GABA(C) receptor-mediated feedback in the
mouse inner retina. J. Neurophysiol. 86, 2285 – 2298.
McCarthy, K.D., de Vellis, J., 1980. Preparation of separate astroglial
and oligodendroglial cell cultures from rat cerebral tissue. J. Cell
Biol. 85, 890 – 902.
556
E.M. Ullian et al. / Mol. Cell. Neurosci. 26 (2004) 544–557
McGeer, E., McGeer, P., 1976. Duplication of biochemical changes of
Huntington’s chorea by intrastriatal injections of glutamic and kainic
acid. Nature 263, 517 – 519.
Meyer-Franke, A., Kaplan, M.R., Pfrieger, F.W., Barres, B.A., 1995.
Characterization of the signaling interactions that promote the survival
and growth of developing retinal ganglion cells in culture. Neuron 15,
805 – 819.
Meyer-Franke, A., Wilkinson, G.A., Kruttgen, A., Hu, M., Munro, E.,
Hanson, M.G., Reichardt, L.F., Barres, B.A., 1998. Depolarization
and cAMP elevation rapidly recruit TrkB to the plasma membrane of
CNS neurons. Neuron 21, 681 – 693.
Moncaster, J.A., Walsh, D.T., Gentleman, S.M., Jen, L.S., Aruoma, O.I.,
2002. Ergothioneine treatment protects neurons against N-methyl-D-aspartate excitotoxicity in an in vivo rat retinal model. Neurosci. Lett.
328, 55 – 59.
Moore, P., El-sherbeny, A., Roon, P., Schoenlein, P.V., Ganapthy, V., Smith,
S.B., 2001. Apoptotic cell death in the mouse retinal ganglion cell layer
is induced in vivo by the excitatory amino acid homocysteine. Exp. Eye
Res. 73, 45 – 57.
Mosinger, J.L., Price, M.T., Bai, H.Y., Xiao, H., Wozniak, D.F., Olney,
J.W., 1991. Blockade of both NMDA and non-NMDA receptors
is required for optimal protection against ischemic neuronal degeneration in the in vivo adult mammalian retina. Exp. Neurol. 113,
10 – 17.
Mosmann, T., 1983. Rapid colorimetric assay for cellular growth and survival: application to proliferation and cytotoxicity assays. J. Immunol.
Methods 16, 55 – 63.
Murai, K.K., Nguyen, L.N., Irie, F., Yamaguchi, Y., Pasquale, E.B., 2003.
Control of hippocampal dendritic spine morphology through ephrin-A3/
EphA4 signaling. Nat. Neurosci. 6, 153 – 160.
Nagler, K., Mauch, D.H., Pfrieger, F.W., 2001. Glia-derived signals induce
synapse formation in neurones of the rat central nervous system.
J. Physiol. 15, 665 – 679.
Naskar, R., Dreyer, E.B., 2001. New horizons in neuroprotection. Surv.
Ophthalmol. 45, S250 – S255.
Nishi, M., Hinds, H., Lu, H.P., Kawata, M., Hayashi, Y., 2001. Motoneuron-specific expression of NR3B, a novel NMDA-type glutamate receptor subunit that works in a dominant-negative manner. J. Neurosci.
21, RC185.
O’Brien, R.J., Mammen, A.L., Blackshaw, S., Ehlers, M.D., Rothstein,
J.D., Huganir, R.L., 1997. The development of excitatory synapses in
cultured spinal neurons. J. Neurosci. 17, 7339 – 7350.
Olney, J.W., 1969. Glutamate-induced retinal degeneration in neonatal
mice. Electron microscopy of the acutely evolving lesion. J. Neuropathol. Exp. Neurol. 28, 455 – 474.
Orlando, L., Alsdorf, S., Penney, J., Young, A.B., 2001. The role of
group I and group II metabotropic glutamate receptors in modulation
of striatal NMDA and quinolinic acid toxicity. Exp. Neurol. 167,
196 – 204.
Osborne, N.N., Wood, J.P.M., Chidlow, G., Bae, J., Melena, J., Nash, M.S.,
1999. Ganglion cell death in glaucoma: what do we really know? Br. J.
Ophthalmol. 83, 980 – 986.
Otori, Y., Wei, J.Y., Barnstable, C.J., 1998. Neurotoxic effects of low doses
of glutamate on purified rat retinal ganglion cells. Invest. Ophthalmol.
Vis. Sci. 39, 972 – 981.
Pang, I., Wexler, E., Nawy, S., Kapin, M., 1999. Protection by cliprodil
against excitotoxicity in cultured rat retinal ganglion cells. Invest. Ophthalmol. Visual Sci. 40, 1170 – 1176.
Peterson, C., Neal, J.H., Cotman, C.W., 1989. Development of N-methyl-Daspartate excitotoxicity in cultured hippocampal neurons. Brain Res.
Dev. Brain Res. 48, 187 – 195.
Pfrieger, F.W., Barres, B.A., 1997. Synaptic efficacy enhanced by glial cells
in vitro. Science 277, 1684 – 1687.
Quigley, H.A., 1999. Neuronal death in glaucoma. Prog. Retinal Eye Res.
18, 39 – 57.
Ritch, R., 2000. Neuroprotection: is it already applicable to glaucoma
therapy? Curr. Opin. Ophthalmol. 11, 78 – 84.
Romano, C., Chen, Q., Olney, J.W., 1998. The intact isolated (ex vivo)
retina as a model system for the study of excitotoxicity. Prog. Retinal
Eye Res. 17, 465 – 483.
Rothman, S.M., Olney, J.W., 1987. Excitotoxicity and the NMDA receptor.
Trends Neurosci. 10, 299 – 302.
Sabel, B.A., Sautter, J., Stoehr, T., Siliprandi, R., 1995. A behavioral model
of excitotoxicity. Exp. Brain Res. 106, 93 – 105.
Sattler, R., Xiong, Z., Lu, W., Hafner, M., MacDonald, J., Tymianski, M.,
1999. Specific coupling of NMDA receptor activation to nitric oxide
neurotoxicity by PSD-95 protein. Science 284, 1845 – 1848.
Sattler, R., Xiong, Z., Lu, W., MacDonald, J., Tymianski, M., 2000. Distinct roles of synaptic and extrasynaptic NMDA receptors in excitotoxicity. J. Neurosci. 20, 22 – 33.
Schori, H., Yoles, E., Wheeler, L.A., Raveh, T., Kimchi, A., Schwartz, M.,
2002. Immune-related mechanisms participating in resistance and susceptibility to glutamate toxicity. Eur. J. Neurosci. 16, 557 – 564.
Shen, S., Wiemelt, A.P., McMorris, F.A., Barres, B.A., 1999. Retinal
ganglion cells lose trophic responsiveness after axotomy. Neuron 23,
285 – 295.
Shin, D.H., Lee, H.Y., Kim, H.J., Lee, E., Lee, K.H., Lee, W.J., Cho, S.S.,
Baik, S.H., 1999. In situ localization of neuronal nitric oxide synthase
(nNOS) mRNA in the rat retina. Neurosci. Lett. 270, 53 – 55.
Silprandi, R., Canella, R., Carmignoto, G., Schiavo, N., Zanellato, Z.,
Zanoni, R., Vantani, G., 1992. NMDA induced neurotoxicity in the
adult rat retina. Vis. Neurosci. 8, 567 – 573.
Sisk, D.R., Kuwabara, T., 1985. Histological changes in the inner retina of
albino rats following intravitreal injection of glutamate. Graefe’s Arch.
Clin. Exp. Ophthalmol. 223, 250 – 258.
Sucher, N.J., Aizenman, E., Lipton, S.A., 1991a. NMDA antagonists prevent kainate neurotoxicity in rat retinal ganglion cells in vitro. J. Neurosci. 11, 966 – 973.
Sucher, N.J., Lei, S.Z., Lipton, S.A., 1991b. Calcium channel antagonists
attenuate NMDA receptor-mediated neurotoxicity of retinal ganglion
cells in culture. Brain Res. 551, 297 – 302.
Sucher, N.J., Lipton, S.A., Dreyer, E.B., 1997. Molecular basis of
glutamate toxicity in retinal ganglion cells. Vision Res. 37,
3483 – 3493.
Sucher, N.J., Kohler, K., Tenneti, L., Wong, H.K., Grunder, T., Fauser, S.,
Wheeler-Schilling, T., Nakanishi, N., Lipton, S.A., Guenther, E., 2003.
N-methyl-D-aspartate receptor subunit NR3A in the retina: developmental expression, cellular localization, and functional aspects. Invest. Ophthalmol. Visual Sci. 44, 4451 – 4456.
Sun, Q., Oooi, V., Chan, S., 2000. NMDA induced excitotoxicity in the
adult rat retina is antagonized by single systemic injection of MK-801.
Exp. Brain Res. 138, 37 – 45.
Takasu, M.A., Dalva, M.B., Zigmond, R.E., Greenberg, M.E., 2002. Modulation of NMDA receptor dependent calcium influx and gene expression through EphB receptors. Science 295, 491 – 495.
Taschenberger, H., Engert, F., Grantyn, R., 1995. Synaptic current kinetics
in a solely AMPA receptor operated glutamatergic synapse formed by
rat retinal ganglion neurons. J. Neurophysiol. 74, 1123 – 1136.
Taylor, W.R., Chen, E., Copenhagen, D.R., 1995. Characterization of spontaneous excitatory synaptic currents in salamander retinal ganglion
cells. J. Physiol. 486, 207 – 221.
Tezel, G.M., Seigel, G.M., Wax, M.B., 1999. Density-dependent resistance
to apoptosis in retinal cells. Curr. Eye Res. 19, 377 – 388.
Tovar, K.R., Westbrook, G.L., 1999. The incorporation of NMDA receptors with a distinct subunit composition at nascent hippocampal synapses in vitro. J. Neurosci. 19, 4180 – 4188.
Tovar, K.R., Westbrook, G.L., 2002. Mobile NMDA receptors at hippocampal synapses. Neuron 34, 255 – 264.
Ullian, E.M., Sapperstein, S.K., Christopherson, K.S., Barres, B.A., 2001.
Control of synapse number by glia. Science 291, 657 – 661.
Vorwerk, C.K., Lipton, S.A., Zurakowski, D., Hyman, B.T., Sabel, B.A.,
Dreyer, E.B., 1996. Chronic low-dose glutamate is toxic to retinal ganglion cells. Toxicity blocked by memantine. Invest. Ophthalmol. Visual
Sci. 37, 1618 – 1624.
E.M. Ullian et al. / Mol. Cell. Neurosci. 26 (2004) 544–557
Washbourne, P., Bennett, J.E., McAllister, A.K., 2002. Rapid recruitment
of NMDA receptor transport packets to nascent synapses. Nat. Neurosci. 5, 751 – 759.
Watanabe, M., Masayoshi, M., Inone, Y., 1994. Differential distributions of
the NMDA receptor channel subunit mRNAs in the mouse retina. Brain
Res. 634, 328 – 332.
557
Wax, M.B., Tezel, G., 2002. Neurobiology of glaucomatous optic neuropathy: diverse cellular events in neurodegeneration and neuroprotection.
Mol. Neurobiol. 26, 45 – 55.
Yamamoto, R., Bredt, D.S., Snyder, S.H., Stone, R.A., 1993. The localization of nitric oxide synthase in the rat eye and related cranial ganglia.
Neuroscience 54, 189 – 200.