The effects of acute intraocular pressure elevation on rat retinal glutamate transport


Nigel L. Barnett
School of Biomedical Sciences
University of Queensland
Queensland 4072
Tel: + 61 7 3365 4089
Fax: + 61 7 3365 4522


Purpose:  To investigate the relationship between intraocular pressure (IOP), retinal glutamate transport and retinal hypoxia during acute IOP elevations of varying magnitude.

Methods:  Female Dark Agouti rats were anaesthetized by ketamine/xylazine/acepromazine (10/5/0.5 mg/kg i.p.). The anterior chamber was cannulated with a 30-gauge needle attached to a saline reservoir. The target IOP (20–120 mmHg, in 10 mmHg increments) was obtained by adjusting the reservoir height. After 10 mins of IOP stabilization, 2 μl of the non-endogenous glutamate transporter substrate, D-aspartate, was injected into the vitreous (final concentration 50 μm), and the elevated IOP maintained for a further 60 mins (total duration of IOP elevation was 70 mins). Glutamate transporter function was assessed by the immunohistochemical localization of D-aspartate. Retinal sections were examined for histological integrity. The experiment was repeated substituting the D-aspartate with the cellular hypoxia marker, Hypoxyprobe-1.

Results:  Under control conditions, D-aspartate was preferentially taken up into the glial Müller cells by glutamate/aspartate transporter (GLAST). This function was maintained at pressures ≤ 70 mmHg, whereafter perturbation of function was evidenced by decreased accumulation of D-aspartate by Müller cells. Failure of GLAST activity was coincident with the appearance of Hypoxyprobe-labelled cells in the inner retina and histological damage.

Conclusions:  Glutamate transport does not appear to change linearly with increased IOP. A pressure threshold exists, above which Müller cell GLAST function is compromised. Moreover, ganglion cell glutamate uptake is only apparent at pressures above those that cause GLAST inhibition. The association between IOP, hypoxia, glutamate transporter dysfunction and subsequent retinal cell death may have important implications for the pathogenesis of IOP/ischaemia-related neuropathy and neuroprotective strategies.


Elevated intraocular pressure (IOP) is a major risk factor for the development and progression of glaucoma, and its reduction remains the mainstay of current therapy (Leske et al. 2003, 2004; Friedman et al. 2004). The selective loss of ganglion cells in glaucoma implies that these neurones are most susceptible to an increase in IOP. Furthermore, this susceptibility has been demonstrated during acute IOP elevations in rats. By assessing retinal function with electroretinography, mild elevations in IOP ≤ 50 mmHg selectively inhibit ganglion cell responses, whereas IOPs > 50 mmHg induce non-specific functional changes in the retina (Bui et al. 2005). The underlying cause of the ganglion cell’s susceptibility to elevated IOP remains unclear.

Ganglion cell susceptibility may be mediated by the effects of IOP at the optic nerve head (ONH). The ONH is a site of amplified IOP-related mechanical stress (Greene 1980). It displays elastic deformation during experimentally elevated IOPs in humans and in animal models (Levy & Crapps 1984; Coleman et al. 1991). Such deformation may alter the alignment of pores in the lamellar plates and induce direct mechanical damage to the ganglion cell axons (Fechtner & Weinreb 1994). Moreover, moderate IOP rises (30–50 mmHg) in monkeys are sufficient to partially block axonal transport (Quigley & Anderson 1977). The blockade of trophic factors such as brain-derived neurotrophic factor (BDNF) at the level of the lamina cribrosa has been hypothesized as a possible cause of IOP-related ganglion cell death (Pease et al. 2000; Quigley et al. 2000). However, chronological studies have shown that retinal ganglion cell apoptosis precedes neurotrophin deprivation, suggesting that factors other than neurotrophin deprivation contribute to the loss of ganglion cells associated with increased IOP (Johnson et al. 2000).

The possibility exists that disrupted glutamate homeostasis contributes to IOP-related retinal dysfunction and neuropathy. However, the involvement of glutamate excitotoxicity in IOP-related ganglion cell dysfunction is controversial. Although it was initially claimed that glutamate levels in the vitreous were elevated in glaucomatous eyes (Dreyer et al. 1996), this has not been confirmed in humans or in animal glaucoma models (Levkovitch-Verbin et al. 2002). However, a large body of evidence supports the idea that glutamatergic mechanisms within the retina contribute to neuronal death. This includes the findings that:

  • 1NMDA glutamate receptor antagonists offer protection to retinal ganglion cells in animal glaucoma models (Hare et al. 2001, 2004; Lipton 2003); consequently, these compounds are currently involved in clinical trials;
  • 2the expression of the endogenous glutamate receptor antagonist, kynurenic acid, is suppressed in experimental glaucoma (Rejdak et al. 2004), and
  • 3elevated IOP differentially regulates metabotropic glutamate receptors in the mouse retina (Dyka et al. 2004).

Furthermore, there is evidence to suggest that glutamate transport mechanisms are perturbed in glaucoma. Extracellular glutamate concentrations are maintained within physiological limits by tightly regulated energy-dependent amino acid transporters (EAATs). Molecular cloning and molecular biological studies have identified five subtypes of EAATs in the retina with the possibility of a sixth, as yet unidentified, transporter (Sarthy et al. 2005). Glutamate/aspartate transporter (GLAST) is present on Müller cells and dominates the uptake of glutamate. Glutamate transporter-1 (GLT-1) is associated with cone photoreceptors and cone bipolar cells, EAAC1 is found on horizontal cells, some amacrine cells and ganglion cells, EAAT4 is found on astrocytes and EAAT5 is associated with photoreceptors and bipolar cells (Rauen & Kanner 1994; Derouiche & Rauen 1995; Rauen et al. 1996, 1998; Barnett & Pow 2000; Pow & Barnett 2000; Ward et al. 2004). Whereas pharmacological or antisense depression of GLAST and GLT-1 induces ganglion cell death (Vorwerk et al. 2000), the effects of glaucoma upon glutamate transporter expression are less clear. For example, it has been reported that GLAST and GLT-1 immunoreactivity are increased in monkey and rat glaucoma models (Carter-Dawson et al. 2002; Woldemussie et al. 2004; Sullivan et al. 2006). Conversely, decreased GLAST and GLT-1 expression have been reported in a different rat glaucoma model (Martin et al. 2002). In human glaucomatous eyes, EAAC1 (EAAT3) immunoreactivity appears to be reduced (Naskar et al. 2000). However, although much evidence exists to implicate glutamatergic mechanisms in glaucomatous neuropathy, their role in primary or secondary pressure-induced neuronal death remains unknown.

Despite the selective effects of elevated IOP on ganglion cell function and the apparent perturbation in glutamate homeostasis during elevated IOP, the direct relationship between IOP elevation and glutamate transport has not been investigated. We sought to clarify this relationship by examining glutamate transport during transient graded elevations of intraocular pressure in rats.

Materials and Methods

All experiments were conducted in accordance with the Association for Research in Vision and Ophthalmology statement for the Use of Animals in Ophthalmic and Vision Research, the Australian Code of Practice for the Care and Use of Animals for Scientific Purposes and the tenets of the Declaration of Helsinki. Adult female Dark Agouti rats (aged 12 weeks) were housed in a 12-hour light/12-hour dark cycle (light on at 06.00 hours). Food and water were available ad libitum.


Acute elevation of IOP

Rats were anaesthetized by intraperitoneal injection of ketamine/xylazine/acepromazine (10/5/0.5 mg/kg) and body temperature maintained with an electric animal heating blanket. The animal’s head was immobilized by resting the front teeth over a horizontal stabilizing bar and securing the skull with adjustable rods inserted in the bony external ear canal. The anterior chamber was cannulated with a 30-gauge needle attached to a reservoir containing 0.9% NaCl. A micromanipulator was used to insert the needle vertically into the anterior chamber at the 12 o’clock position. The target IOP (20–120 mmHg, in 10 mmHg increments) was obtained by adjusting the reservoir height.

Glutamate transporter activity

After 10 mins of IOP stabilization, 2 μl of the non-endogenous glutamate transporter substrate, D-aspartate (750 μm), was injected with a 30-G Hamilton syringe into the vitreous (final concentration 50 μm), and the elevated IOP maintained for a further 60 mins (total duration of IOP elevation was 70 mins, n = 4 for each pressure). A micromanipulator held the injection needle in situ during the experiment to prevent fluid leakage and to maintain the elevated IOP. Elevation of IOP was confirmed with a factory-calibrated TonoLab rodent tonometer (Colonial Medical Supply Inc., Franconia, NH, USA). At the end of the 70 mins, the rats were killed immediately with sodium pentobarbital (100 mg/kg i.p.) and the retinas fixed in 2.5% glutaraldehyde in 0.1 m phosphate buffer (pH 7.4) for 30 mins. Retinas were washed twice in phosphate-buffered saline for 10 mins each, dehydrated in an ethanol series (50%, 70%, 95%, 100%) and embedded in epoxy resin (ProSciTech, Thuringowa, QLD, Australia). Semi-thin (0.5 μm) transverse sections were cut on an Ultracut R ultramicrotome (Leica Microsystems, North Ryde, NSW, Australia) and dried onto glass slides. The sections were etched in 11% sodium hydroxide/ethanol. Glutamate transporter function was assessed by the immunohistochemical localization of D-aspartate. Labelling was detected with a peroxidase-based Vectorstain Elite ABC Kit (Vector Laboratories, Inc., Burlingame, CA, USA) with 3,3-diaminobenzidine (DAB) as the chromogen (Pow & Crook 1993; Barnett et al. 2001). Semi-thin retinal sections were stained with toluidine blue for histological analysis. Retinal sections were observed with a Zeiss Axioskop microscope and photographed with a Spot RT digital camera (Diagnostic Instruments, Sterling Heights, MI, USA).

Hypoxyprobe-1 labelling

To detect cellular hypoxia, the experiment was repeated substituting the D-aspartate with Hypoxyprobe-1 (pimonidazole hydrochloride; Millipore, North Ryde, NSW, Australia). Immunohistochemical detection of hypoxia is appealing as hypoxic regions can be directly visualized in situ and compared with the underlying cellular structure. Pimonidazole hydrochloride, a 2-nitroimidazole compound, forms covalent adducts with cells that have an oxygen partial pressure < 10 mmHg (Arteel et al. 1995). Pimonidazole binding is not dependent on the presence of specialized redox enzymes and wide variations in reduced nicotinamide adenine dinucleotide (NADH) and reduced nicotinamide adenine dinucleotide phosphate (NADPH) levels do not change the oxygen dependence of its binding (Arteel et al. 1998). The subsequent staining of ischaemic tissues with a pimonidazole antibody reveals hypoxic cells within the specimen (Varghese et al. 1976). Hypoxyprobe-1 (2 μl of 5 μm solution diluted in sterile 0.9% NaCl) was injected into the vitreous 10 mins after IOP elevation. The elevated IOP was maintained for a further 60 mins (= 4 for each pressure). After removal of the anterior chamber needle to restore normal IOP, the animals were allowed to recover for 3 hours as per the manufacturer’s instructions and then administered an overdose of sodium pentobarbital (100 mg/kg i.p.). The retinas were fixed with 4% paraformaldehyde in 0.1 m phosphate buffer (pH 7.4) for 60 mins and cryoprotected in 30% sucrose. Monoclonal Hypoxyprobe-1 MAb1 (1:50) was applied to frozen sections (10 μm) for immunolocalization of the Hypoxyprobe. Labelling was detected with a peroxidase-based Vectorstain Elite ABC kit using DAB as the chromogen.


Glutamate transporter activity

D-aspartate immunoreactivity in normotensive eyes (12.8 ± 1.3 mmHg, mean ± standard error of the mean [SEM], = 4) was localized exclusively to Müller cells (Fig. 1A). Intense labelling was present in the Müller cell somata, the transverse processes, the end feet and the outer limiting membrane. Neuronal D-aspartate immunoreactivity was not observed. This localization pattern reflects the activity and distribution of GLAST (Pow & Barnett 1999; Barnett & Pow 2000). The normal pattern of D-aspartate immunoreactivity in the Müller cells persisted as the IOP was elevated to 70 mmHg (Fig. 1B). The accumulation of D-aspartate by Müller cells was impaired at IOPs > 70 mmHg. At 80 mmHg, retinal neurones began to accumulate the glutamate analogue, as evidenced by weak D-aspartate immunoreactivity in photoreceptor inner segments, bipolar cells and ganglion cells (Fig. 1C). At 90 mmHg, Müller cell accumulation of D-aspartate was no longer evident, whereas the intensity of D-aspartate labelling in ganglion cells was enhanced (Fig. 1D). At IOPs > 100 mmHg, a reduction in the intensity of ganglion cell staining became evident (Fig. 1E). All retinal D-aspartate labelling was lost at an IOP of 110 mmHg (Fig. 1F).

Figure 1.

 Immunohistochemical localization of D-aspartate uptake in the rat retina. (A) Under control conditions (intraocular pressure [IOP] 12.8 ± 1.3 mmHg, mean ± standard error of the mean [SEM], = 4), exogenous D-aspartate is accumulated predominantly by the Müller cells. (B) Normal Müller cell uptake of the glutamate analogue is maintained at IOPs ≤ 70 mmHg. (C) At IOPs above a threshold of 70 mmHg, Müller cell uptake of D-aspartate is greatly reduced. Weak D-aspartate immunoreactivity is evident in retinal neurones, including bipolar and ganglion cells, at 80 mmHg. (D) At 90 mmHg there is no evidence of Müller cell D-aspartate uptake. The accumulation of D-aspartate by ganglion cells is maintained. (E) Ganglion cell uptake of d-aspartate is still evident at IOP of 100 mmHg. (F) Retinal uptake of D-aspartate is suppressed by IOP of 110 mmHg. M = Müller cell; GC = ganglion cell; BP = bipolar cell; ONL = outer nuclear layer; OPL = outer plexiform layer; INL = inner nuclear layer; IPL = inner plexiform layer; GCL = ganglion cell layer.


Normal retinal histology is presented in Fig. 2A. Following 60 mins of elevated IOP to 70 mmHg, the histological appearance of the retina was not significantly different from that of the control retina (Fig. 2B). However, at IOPs > 70 mmHg, histological damage to the retina was observed. At 80 mmHg, perforations were present in the inner nuclear layer (INL) (Fig. 2C). Increasing the IOP to 110 mmHg induced significant morphological changes to the retina. Necrotic cells of the INL displayed the characteristic ‘bull’s eye’ appearance and condensed chromatin (Fig. 2D). Furthermore, dying neurones and damage in the ganglion cell layer were clearly evident.

Figure 2.

 Semi-thin (0.5 μm) sections of rat retina stained with toluidine blue showing the effect of elevated intraocular pressure (IOP) on retinal histology. (A) Control retina. (B) Normal retinal integrity is maintained at IOPs ≤ 70 mmHg. (C) An IOP of 80 mmHg induces vacuole formation in the outer plexiform layer and inner nuclear layer (arrows). (D) Gross histological damage to the retina is evident following the elevation of IOP to 110 mmHg. Dying neurones displaying cellular swelling and condensed chromatin are present in the inner nuclear layer and ganglion cell layer (arrows). ONL = outer nuclear layer; OPL = outer plexiform layer; INL = inner nuclear layer; IPL = inner plexiform layer; GCL = ganglion cell layer.

Retinal hypoxia

No Hypoxyprobe-1 labelling was apparent in either control retinas (Fig. 3A) or in retinas subjected to elevated IOP ≤ 70 mmHg (Fig. 3B). The first appearance of Hypoxyprobe-1 immunoreactivity occurred at 80 mmHg, with weak labelling in the INL and the ganglion cells (Fig. 3C). Intense Hypoxyprobe-1 labelling was observed in the INL and ganglion cells at 110 mmHg, together with the appearance of positive Hypoxyprobe-1 staining of the inner plexiform layer (Fig. 3D). No Hypoxyprobe-1 labelling was observed in the outer retina following the elevation of IOP.

Figure 3.

 Hypoxyprobe-1 labelling of rat retinal sections following a graded increase in intraocular pressure (IOP). (A) No Hypoxyprobe-1-labelled cells are present in the control retina. (B) There is no evidence of cellular hypoxia in the retina during elevation of IOP to 70 mmHg. (C) At 80 mmHg, Hypoxyprobe-1-positive ganglion cells appear. (D) The intensity of the Hypoxyprobe-1 labelling of ganglion cells is increased at 110 mmHg. At this IOP, the inner retina (OPL, INL, IPL and GCL) becomes hypoxic. ONL = outer nuclear layer; OPL = outer plexiform layer; INL = inner nuclear layer; IPL = inner plexiform layer; GCL = ganglion cell layer; GC = ganglion cell.


Our present findings suggest that glutamate transport does not change linearly with increased IOP, but rather that a pressure threshold exists, above which GLAST function is compromised. Under physiological conditions, retinal glutamate transport is dominated by the Müller cell glutamate transporter GLAST (Rauen et al. 1998; Barnett & Pow 2000). The specific accumulation of exogenous D-aspartate by Müller cells in eyes with IOPs elevated ≤ 70 mmHg (Fig. 1A, B) is consistent with the maintenance of GLAST activity. The qualitative nature of immunohistochemical labelling does not preclude the possibility that GLAST is functioning at reduced capacity at lower levels of hypertension. However, at pressures > 70 mmHg GLAST function is perturbed, as evidenced by the decreased Müller cell uptake of D-aspartate and the appearance of D-aspartate immunoreactivity in retinal neurones (Fig. 1C–E). Furthermore, our results show that the failure of GLAST coincides with histological damage to the retina (Fig. 2C, D). A similar correlation between GLAST failure and histological damage has been shown in models of retinal ischaemia, confirming that energy-dependent GLAST activity is integral to the regulation of extracellular glutamate concentration and the prevention of excitotoxicity (Harada et al. 1998; Barnett et al. 2001). Hypoxyprobe-1 labelling of cells revealed that GLAST dysfunction correlates with inner retinal hypoxia (PO2 < 10 mmHg [Arteel et al. 1995]) at IOPs > 70 mmHg (Figs 1C–F and 3C, D). Microelectrode detection of oxygen tension in the rat retina during retinal vasculature occlusion demonstrates relative anoxia of the inner retina compared to the outer retina due to perfusion of the outer retina by the choroidal vasculature (Yu & Cringle 2001). This is in agreement with our results, which show that hypoxic labelling is limited to the inner retina, with the ganglion cells being particularly heavily stained at high IOPs of 80–110 mmHg (Fig. 3C, D). Paradoxically, this apparent ganglion cell hypoxia is associated with an uptake of D-aspartate into these neurones. Hypoxia-induced glutamate transporter expression, increased trafficking of the transporter to the membrane or post-translational modification of transporter protein may account for this uptake as it has recently been reported that GLT-1 expression is stimulated by hypoxia (Pow et al. 2004). Furthermore, glaucoma also stimulates GLT-1c (a GLT-1 splice variant) expression by ganglion cells, possibly to protect themselves from glutamate toxicity (Sullivan et al. 2006). It is also apparent from our study that increased transport of glutamate into ganglion cells at higher IOPs may protect these neurones from the type of excititoxic damage evident in the INL (Fig. 2C).

Importantly, we observe that Hypoxyprobe-1 labelling is not present in the inner retina until the IOP is raised to 80 mmHg, suggesting that the elevated IOP could preferentially compromise blood flow to the inner retina without affecting choroidal flow. Moreover, the absence of Hypoxyprobe-1 labelling in the outer nuclear layer (ONL) after hypoxia is observed in the inner retina strongly supports the conclusion that the choroidal perfusion is resistant to increased IOP. Similarly, microelectrode detection of oxygen tension in the rat retina during retinal vasculature occlusion demonstrates relative anoxia of the inner retina compared with the outer retina as a result of perfusion of the outer retina by the choroidal vasculature (Yu & Cringle 2001). Retinal vasculature autoregulation maintains ocular perfusion over a broad range of perfusion pressures (mean arterial pressure minus IOP). In rats, humans and other mammals, ocular perfusion is maintained at perfusion pressures above ∼ 30 mmHg. Decreases in perfusion pressure to < 30 mmHg induce an almost linear decline in ocular perfusion (Kiel & Shepherd 1992; Shonat et al. 1992; Kiel & van Heuven 1995; Riva 1998). As the mean arterial pressure in rats is ∼ 100 mmHg, it is anticipated that ocular perfusion is maintained by vascular autoregulation for IOPs up to ∼ 70 mmHg (a resultant perfusion pressure of ∼ 30 mmHg; Davis & Johns 1995; Mayorov et al. 2001). Our Hypoxyprobe-1 results support this predicted IOP threshold of ∼ 70 mmHg for the maintenance of retinal perfusion. Moreover, the lack of a correlation between GLAST dysfunction and IOP (< 70 mmHg) and the detection of retinal hypoxia coincident with GLAST failure suggest that GLAST function is not affected by IOP per se. Rather, GLAST activity is maintained during IOP elevation until the transporter’s function is compromised by an inadequate vascular supply and the consequent hypoxia. Clearly, the high IOPs required in this experiment to induce ischaemic-related GLAST dysfunction are not associated with chronic glaucoma. However, a growing body of evidence exists to implicate vascular insufficiency in the pathogenesis of glaucoma (Findl et al. 2000; Piltz-Seymour et al. 2001; Hafez et al. 2003; Lam et al. 2005). Furthermore, there is direct evidence that hypoxic tissue stress is present in glaucomatous eyes. Immunoreactivity of hypoxia-inducible factor-1α (HIF-1α), an oxygen-regulated transcriptional factor, is increased in the ONH and retina of glaucomatous eyes (Tezel & Wax 2004).

Our results do not rule out the possibility that disturbed glutamate transport contributes to ganglion cell death in glaucoma; rather, they imply that disturbed glutamate homeostasis does not contribute to the functional deficit observed during acute sub-ischaemic increases in IOP (Bui et al. 2005). Thus, it is likely that mechanisms other than compromised GLAST activity mediate the graded functional changes that occur with acute IOP elevation.


This research was supported by a National Health and Medical Research Council (Australia) grant to NLB and GAG. None of the authors have any commercial or proprietorial interests. We thank Professor David Pow (University of Newcastle, Australia) for the kind donation of D-aspartate antibody and Rowan Tweedale for her careful reading of the manuscript.