Address correspondence and reprint requests to Dr Michael B. Robinson, 502 N Abramson Pediatric Research Building, 3615, Civic Center Blvd., Philadelphia, PA 19104–4318, USA. E-mail: Robinson@Pharm.med.upenn.edu
Protein kinase C (PKC) regulates the activity and/or cell surface expression of several different neurotransmitter transporters, including subtypes of glutamate transporters. In the present study, the effects of pharmacological inhibitors of PKC were studied in primary astrocyte cultures that express the glutamate aspartate transporter (GLAST) subtype of glutamate transporter. We found that general inhibitors of PKC, bisindolylmaleimide I (Bis I), bisindolylmaleimide II (Bis II), staurosporine and an inhibitor of classical PKCs, Gö6976, had no effect on Na+-dependent glutamate transport activity. However, rottlerin, a putative specific inhibitor of PKCδ, decreased transport activity with an IC50 value (less than 10 µm) that is comparable to that reported for inhibition of PKCδ. The effect of rottlerin was very rapid (maximal effect within 5 min) and was due to a decrease in the capacity (Vmax) for transport. Rottlerin also caused a drastic loss of GLAST immunoreactivity within 5 min, suggesting that rottlerin accelerates GLAST degradation/proteolysis. Rottlerin had no effect on cell surface or total expression of the transferrin receptor, providing evidence that the effect on GLAST cannot be attributed to a non-specific internalization/degradation of plasma membrane proteins. Down-regulation of PKCδ with chronic phorbol ester treatment did not block rottlerin-mediated inhibition of transport activity. These results suggest a novel mechanism for regulation of the GLAST subtype of glutamate transporter and indicate that there is a rottlerin target that is capable of controlling the levels of GLAST by controlling the rate of degradation or limited proteolysis. It appears that the target for rottlerin may not be PKCδ.
Sodium-dependent high affinity glutamate transport systems clear the excitatory neurotransmitter, glutamate, into neurons and glial cells, thereby terminating glutamatergic neurotransmission and preventing glutamate toxicity. A family of Na+-dependent glutamate transporters mediate this activity, including glutamate aspartate transporter (GLAST) and GLT-1 (glutamate transporter 1: generally expressed by glia), EAAC1 (excitatory amino acid carrier 1) and EAAT4 (excitatory amino acid transporter 4: generally expressed by neurons) and EAAT5 (retinal) (for reviews, see Sims and Robinson 1999; Danbolt 2001). Understanding the mechanisms that regulate these transporters has the potential to impact on both the physiology and pathology of glutamate in the CNS.
Several groups have recently shown that the activity of many neurotransmitter transporters (dopamine, serotonin, γ-aminobutyric acid, norepinephrine) can be regulated by protein kinase C (PKC) (for reviews, see Beckman and Quick 2000; Blakely and Bauman 2000; Robinson 2002). The effects of PKC occur within minutes and are frequently associated with a redistribution of transporter between intracellular vesicles and the plasma membrane. Similar mechanisms appear to regulate some of the glutamate transporter subtypes. Activation of PKC increases the activity and cell surface expression of EAAC1, endogenously expressed in C6 glioma (Dowd and Robinson 1996; Davis et al. 1998), but decreases EAAC1-mediated activity in MDCK cells and in Xenopus oocytes (Trotti et al. 2001). Initially, PKC activation was shown to stimulate phosphorylation of GLT-1 and increase GLT-1-mediated activity when introduced into HeLa cells with vaccinia virus (Casado et al. 1993). In stably transfected HeLa cells, PKC has no effect on GLT-1-mediated transport activity (Tan et al. 1999). In Y-79 human retinoblastoma cells, PKC activation decreases dihydrokainate-sensitive, presumably GLT-1-mediated, glutamate transport activity by increasing the Km value (Ganel and Crosson 1998). In primary cell cultures that endogenously express GLT-1 or when GLT-1 is introduced by transfection into MDCK or C6 glioma cells, PKC decreases GLT-1-mediated activity and cell surface expression (Carrick and Dunlop 1999; Kalandadze et al. 2002). Activation of PKC also has varied effects on the activity of GLAST. When GLAST is expressed in Xenopus oocytes or human embryonic kidney cells, activation of PKC decreases activity by a mechanism that is apparently independent of a change in number of transporters at the plasma membrane (Conradt and Stoffel 1997). In cultured Müller cells and in Bergman glia, PKC activation causes a rapid decrease in transport activity and loss of GLAST protein after long-term treatment with phorbol ester (González and Ortega 1997; González et al. 1999). In primary glial cell cultures that presumably only express GLAST, phorbol ester treatment increases glutamate transport activity (Casado et al. 1991; Susarla et al. 2001).
It seems likely that some of these differences are related to cellular milieu, such as the expression of different proteins required for trafficking or different PKC subtypes. PKC activity is mediated by three groups of proteins. The classical subtypes (α, β1, βII and γ) are activated by phorbol esters and are Ca2+-dependent. The novel subtypes (δ, ε, η and θ) are activated by phorbol esters but are Ca2+-independent. The atypical PKCs (ζ and λ) are not activated by phorbol esters (for review, see Mellor and Parker 1998; Kazanietz et al. 2000). It is possible that various PKC subtypes have differential effects on activity and/or cell surface expression of the various glutamate transporters. In fact, recent studies from our laboratory have suggested that two different subtypes of PKC regulate the activity of the EAAC1 subtype of glutamate transporter. The first, PKCα, appears to increase activity by causing a redistribution of transporter from a subcellular compartment to the cell surface. The second, PKCε, appears to increase activity by a mechanism that is independent of a change in the number of transporters at the plasma membrane (González et al. 2002). Recent studies have demonstrated that chelerythrine, a general inhibitor of PKC, and rottlerin, a putative inhibitor of PKCδ, decrease GLAST-mediated glutamate transport activity in retinal preparations without changing GLAST immunoreactivity. This effect was attributed to inhibition of PKCδ (Bull and Barnett 2002). These studies suggest that pharmacological approaches will be used to examine the role of specific PKC subtypes in the regulation of various neurotransmitter transporters.
In the present study, the effects of general and specific inhibitors of PKC on glutamate transport activity were examined using astrocyte cultures that selectively express the GLAST subtype of glutamate transporter. We found that rottlerin caused a concentration-dependent decrease in transport activity in this system. The decrease in transport was caused by a decrease in the capacity for transport (Vmax) and was associated with a decrease in GLAST protein. We provide evidence that this is not a generalized effect on membrane proteins and that this effect may be independent of PKCδ. Our data suggest that rottlerin accelerates degradation or proteolysis of GLAST by a PKC-independent mechanism.
Materials and methods
l-[3H]Glutamate was obtained from Dupont (Boston, MA, USA). Phorbol 12-myristate 13-acetate (PMA), l-glutamate and anti-actin antibody were purchased from Sigma (St Louis, MO, USA). Sulfo-NHS-biotin and Immunopure immobilized monomeric avidin were obtained from Pierce (Rockford, IL, USA). Except for fetal bovine serum, which was obtained from Hyclone (Logan, UT, USA), all other tissue culture reagents were purchased from Gibco BRL (Gaithersburg, MD, USA). Tissue culture plates were from Corning (Corning, NY, USA). Bisindolylmaleimide I (Bis I), bisindolylmaleimide II (Bis II), staurosporine, Gö6976 and rottlerin were purchased from Calbiochem (La Jolla, CA, USA). Antibodies for PKCα and PKCβ were purchased from Transduction Laboratories (Lexington, KY, USA) and antibodies for PKCγ, PKCδ, PKCε, PKCη and PKCθ were obtained from Santa Cruz Biotechnology, Inc. (Santa Cruz, CA, USA). Phospho-PKCδ (Ser643) antibody was obtained from Cell Signalling Technology, Inc. (Beverly, MA, USA). Chemiluminescence kits were purchased from Amersham (Arlington Heights, IL, USA). Immobilon-P membrane was from Millipore (Bedford, MA, USA). Adult Sprague-Dawley rats were purchased from Charles River Laboratories in Kingston and were bred at the Children's Hospital of Philadelphia. The studies involving animals were reviewed and approved by the institutional animal care use committee of the Children's Hospital of Philadelphia.
Astrocyte cultures were prepared from neonatal (1–3 day old) rat brain cortices according to the protocol described earlier (Garlin et al. 1995). In brief, isolated cortical tissue devoid of meninges and blood vessels was incubated with trypsin–EDTA (0.05%-0.2 mm) for 20 min at 37°C. Tissue was triturated and suspended in Dulbecco's modified eagle medium containing heat inactivated fetal bovine serum (10%), Ham's F12 (10%) and penicillin/streptomycin (0.24%). The suspension was diluted to a density of 2 × 105 cells/mL and 0.5 mL or 10 mL was plated onto 12-well plates or 10-cm dishes, respectively. After 3 days, the media in 12-well plates was replaced with 1 mL of fresh media. Cells were fed with a complete media change twice a week and maintained in a 5% CO2 incubator at 37°C. Based on immunocytochemical criteria, these cultures contain > 95% astrocytes (Garlin et al. 1995). The cultures were used after 14–18 days in vitro. For all experiments, cells were incubated in Dulbecco's modified Eagle's medium containing 0.5% bovine serum albumin for 2 h, followed by two rinses with plain Dulbecco's modified Eagle's medium. Drugs were added directly to fresh Dulbecco's modified Eagle's medium.
Measurement of Na+-dependent l-[3H]glutamate transport activity
After treatment, cells were rinsed twice with either Na+- or choline-containing buffer and transport activity was measured as described earlier (Garlin et al. 1995). Na+-dependent glutamate transport activity was calculated as the difference in the amount of radioactivity accumulated in the presence and absence of Na+ by substituting equimolar concentrations of choline for Na+ in the assay buffer. Cells were solubilized in 1 mL of 0.1 n NaOH; aliquots of this suspension were used for measurement of radioactivity and for analysis of protein (Lowry et al. 1951).
Analyses of cell surface expression
Biotinylation of cell surface proteins was performed as described earlier (Davis et al. 1998). Astrocytes were washed twice with ice-cold phosphate-buffered saline containing CaCl2 (0.1 mm) and MgCl2 (1.0 mm) followed by an incubation with sulfo-NHS-biotin (1 mg/mL of phosphate-buffered saline) for 20 min. The plates were washed and incubated for 20 min with phosphate-buffered saline containing glycine (100 mm) to quench the unreacted biotin. Cells were lysed in radioimmunoprecipitation (RIPA) buffer for 30 min and an aliquot was mixed with an equal volume of sodium dodecyl sulfate-containing sample buffer and stored at −20°C; this fraction is called ‘cell lysate’. An aliquot of avidin beads (300 µL) was centrifuged at 16 500 g and after removing the supernatant, beads were mixed with 300 µL of cell lysate; this slurry was incubated overnight at 4°C on a rotating mixer. The next day, the supernatant containing non-biotinylated proteins (intracellular) was collected by centrifugation and mixed with an equal volume of sodium dodecyl sulfate-containing sample buffer. After washing the beads four times with RIPA buffer, biotinylated proteins were eluted with 600 µL of sodium dodecyl sulfate-containing sample buffer. In this way, the amounts of immunoreactivity in the non-biotinylated and biotinylated fractions should sum to amounts observed in the lysate if the yield of the extraction from the beads is 100%. Protein levels were measured in cell lysates after acid precipitation (Yeang et al. 1998).
Western analyses were performed as described previously (Davis et al. 1998). Twenty-five micrograms of protein from the cell lysates and the same volume of non-biotinylated (intracellular) and biotinylated (cell surface) fractions were loaded onto each gel. After separation of proteins on a sodium dodecyl sulfate/8% polyacrylamide gel and transfer to Immobilon-P, the blots were probed with antibodies against actin, both amino termini of GLAST (1 : 1000) and the carboxyl terminus of GLAST (1 : 75 (Rothstein et al. 1994; Furuta et al. 1997) or with transferrin receptor antibody (Zymed laboratories Inc., San Francisco, CA, USA). Immunoreactive bands were visualized using enhanced chemiluminescence and were quantified using Image software (National Institutes of Health).
All the data are presented as the mean ± standard error of independent experiments. The concentration-dependence of the effects of rottlerin was assessed by non-linear regression fitting to a single population of sites using GraphPad Prism software (version 2.0 GraphPad Software Inc., San Diego, CA, USA). Statistical comparisons were performed using Instat 2.03 (GraphPad Software, Inc.). For the comparisons of kinetic constants, the Km and Vmax values were log transformed because the standard deviations of the untransformed data were significantly different. Comparisons between two groups were made using an unpaired Student's t-test; comparisons between multiple groups were by anova with Bonferroni post hoc analysis.
In the present study, the effects of PKC antagonists on glutamate transporter activity were examined in astrocyte-enriched cultures prepared from rat cortex. In earlier studies, we have demonstrated that these cultures express a robust GLAST immunoreactive band, a faint GLT-1 band and no EAAC1 or EAAT4 immunoreactivity (Schlag et al. 1998). An identical expression pattern was observed in the present study (data not shown, n = 3). Unlike the other transporters, GLAST-mediated activity is essentially insensitive to dihydrokainate (for review, see Robinson and Dowd 1997). As was previously observed (Garlin et al. 1995), 1 mm dihydrokainate had essentially no effect on Na+-dependent glutamate transport activity (94 ± 4% of control, n = 4 observations). These observations strongly suggest that GLAST mediates the bulk of Na+-dependent glutamate transport in the astrocytes used for these studies. In these same cultures, the expression patterns of the classical and novel subtypes of PKC were examined using subtype specific antibodies; brain homogenates were used as a positive control for these studies. The molecular weights of the bands detected were comparable to those reported in earlier studies (Kazanietz et al. 1993; Goodnight et al. 1995). Of the classical subtypes, these cultures expressed PKCα, PKCβ, but not PKCγ. Of the novel subtypes, PKCδ and PKCε were observed, but PKCθ or PKCη were not detected (data not shown, n = 2 independent experiments). The expression pattern of PKC isozymes in these cultures was similar to that previously reported (Gott et al. 1994; Chen et al. 1995a; Slepko et al. 1999).
The effects of three different types of PKC antagonists on Na+-dependent glutamate transport activity were examined in this system. Bis I, Bis II and staurosporine, non-selective inhibitors of both classical and novel PKC subtypes (Toullec et al. 1991; Nixon et al. 1992; Gschwendt et al. 1994a; deVente et al. 1996; Ishii et al. 1996; Fujii et al. 2000), had no effect on transport activity after a 30-min pre-incubation (Fig. 1a). Gö6976, a selective inhibitor of classical PKC subtypes at low µm concentrations (Way et al. 2000), also had no effect on activity. In contrast, rottlerin, a putative selective inhibitor of PKCδ (Gschwendt et al. 1994b), decreased glutamate uptake. The effect of rottlerin was concentration-dependent with the IC50 value of 8.1 ± 2.2 µm when measured at a low concentration (0.5 µm) of l-glutamate (Fig. 1b). This IC50 value is comparable to the concentrations of rottlerin used in several studies to inhibit PKCδ (Gschwendt et al. 1994b; Keenan et al. 1997; Kontny et al. 2000; Soltoff 2001). To examine the time-dependence of this effect, astrocytes were treated with 10 µm rottlerin for different periods of time. The decrease in transport activity was maximal within 5 min (Fig. 2a) with no further decrease up to 1 h. To simplify timing, all subsequent experiments were performed using a 30-min pre-incubation with rottlerin.
The effects of rottlerin on the concentration-dependence of Na+-dependent glutamate transport activity were examined. Rottlerin (10 µm) significantly decreased the Vmax of glutamate transport to ∼10% of control (14.7 ± 4.9 nmol/mg protein per min in vehicle-treated cells compared to 1.1 ± 0.3 nmol/mg protein per min in rottlerin-treated cells, p < 0.001) (Fig. 2b). This change was accompanied by a significant decrease in the Km value for transport activity (69 ± 26 µm in vehicle-treated cells compared to 17 ± 3 µm in rottlerin-treated cells, p < 0.05). The fact that the Km value is not increased strongly suggests that rottlerin is not a competitive inhibitor of glutamate transport activity. The observed decrease in the capacity for transport suggests that rottlerin decreases either the catalytic efficiency (turnover number) of GLAST or changes the number of transporters at the plasma membrane.
To determine if the effect of rottlerin is related to a change in the number of transporters at the cell surface, astrocytes were treated with rottlerin (10 µm) and cell surface proteins were batch extracted after labeling with a membrane impermeant biotinylation reagent. As was observed with the measurement of transport activity, rottlerin decreased GLAST immunoreactivity very rapidly. Within 5 min, there was a 30–40% decrease in immunoreactivity at the cell surface and after 30 min there was approximately a 50% decrease in immunoreactivity. This effect was observed using antibodies directed against the amino or the carboxyl termini (Figs 3a–d). Rottlerin never caused GLAST immunoreactivity to appear in the non-biotinylated (intracellular) fraction, suggesting that the transporter is either subject to rapid proteolysis at the plasma membrane or is rapidly targeted for intracellular degradation. Consistent with this suggestion, rottlerin also decreased total transporter immunoreactivity in the cells (lysate). As was observed with the measurements of transport activity, other PKC antagonists, Bis II and Gö6976, had no effect on GLAST immunoreactivity on the cell surface (Fig. 3e) or in the cell lysate (data not shown) using an antibody directed against the carboxyl terminus. Although it is difficult to imagine that inhibition of transcription or translation could decrease GLAST levels within minutes, the effects of the protein synthesis inhibitor, cycloheximide (10 µg/mL), on GLAST protein levels were examined. A 24-h treatment with cycloheximide only reduces GLAST to 80 ± 2% of control levels (data not shown, n = 3). This suggests that the rapid loss of GLAST is not related to a decrease in the rate of GLAST synthesis, but is instead related to an increase in the rate of GLAST degradation.
Two different studies were conducted to determine if rottlerin affects the activity or expression of other membrane proteins. The effects of rottlerin on Na+-dependent glycine transport were examined to determine if rottlerin is specific for Na+-dependent glutamate transport. As shown in Fig. 4, pre-treatment with rottlerin decreased Na+-dependent glycine transport with a concentration-dependence similar to that observed for inhibition of glutamate transport activity. The effects of rottlerin on transferrin receptor expression were examined to determine if the loss of GLAST might be related to a generalized increase in the rate of degradation of membrane proteins. As would be predicted for the transferrin receptor, which is continuously recycled between the plasma membrane and cytosplasmic vesicles, we found that approximately 50% of the transferrin immunoreactivity (∼100 kDa band) was in the biotinylated (cell surface) fraction and approximately 50% was in the non-biotinylated (intracellular) fraction. At concentrations that caused nearly a 50% loss in GLAST immunoreactivity, rottlerin had no effect on the cell surface expression or total expression of transferrin receptor (Fig. 5).
Rottlerin has been widely used as an inhibitor of PKCδ to define the specific contribution of this subtype of PKC to various biological phenomenon and is thought to directly interact with the ATP binding site on PKC (Gschwendt et al. 1994b). Several groups have shown that chronic incubation with phorbol esters down-regulate the levels of PKC within different cell types (Chen et al. 1995b). To determine if the effects observed in the present study are influenced by down-regulation of PKCδ, astrocytes were chronically treated with PMA prior to treatment with rottlerin. We found that this chronic incubation decreased total PKCδ immunoreactivity to approximately 25% of control levels (Fig. 6a). In some systems, auto-phosphorylation of PKCδ at serine-643 is required for catalytic activity (Li et al. 1997). Therefore, the levels of phospho PKCδ (Ser643) immunoreactivity were examined after treatment of astrocytes with vehicle or PMA for 24 h. The levels of phospho PKCδ (Ser643) were reduced even further than the levels of total PKCδ (Fig. 6b). In these astrocytes that express significantly less PKCδ and phospho PKCδ, rottlerin still reduced transport activity to the same extent as that observed in cells that were not chronically treated with phorbol ester (Fig. 6c). We found that these treatments also down-regulated PKCα and PKCε to the same extent as that observed with PKCδ and completely eliminated expression of PKCβ (data not shown, n = 3). Together with the observation that Bis II, Bis I and staurosporine (general inhibitors of PKC) did not have the same effect as rottlerin, these data would suggest that the inhibition/loss of GLAST is likely to be independent of PKCδ or any other PKC subtypes.
There is also evidence that rottlerin can inhibit calmodulin-dependent protein kinase III (CaM Kinase III) activation at low micromolar (1–5 µm) concentrations (Gschwendt et al. 1994a; Parmer et al. 1997). Therefore, we examined the effects of inhibition of CaM Kinase III (elongation factor-2 kinase) with the calmodulin antagonist, trifluoroperazine. Although the concentration of trifluoroperazine used to inhibit CaM Kinase III activation has varied in different studies (Buss et al. 1994; Vega et al. 1998), 10 µm causes maximal inhibition in lysates from NIH3T3 cells (Vega et al. 1998). Trifluoroperazine (10 µm) had no effect on Na+-dependent glutamate transport activity (110 ± 4% of control, n = 6). Higher concentrations of trifluoroperazine (30 and 100 µm) were toxic to astrocytes. These data would suggest that the effects of rottlerin can not be attributed to inhibition of CaM Kinase III.
Recent studies have demonstrated that PKC regulates the activity and cell surface expression of many different neurotransmitter transporters (for reviews, see Beckman and Quick 2000; Blakely and Bauman 2000; Robinson 2002). Several groups, including ours, have begun to determine if specific subtypes of PKC are responsible for these phenomena (Bull and Barnett 2002; Doolen and Zahniser 2002; González et al. 2002). Although other strategies have been used to define the roles of specific PKC subtypes in biological systems, there is generally a reliance on the use of subtype specific pharmacological tools for an initial assignment of the types of PKC that may be involved.
In the present study, the effects of three types of PKC antagonists on Na+-dependent glutamate transport activity were compared in primary astrocyte cultures. We found that general inhibitors of the phorbol ester activated PKC subtypes (Bis I, Bis II and staurosporine) and a selective inhibitor of the classical subtypes (Gö6976) had no effect on glutamate transport activity, but rottlerin potently inhibited Na+-dependent glutamate transport activity. As was previously observed (Schlag et al. 1998), only two glutamate transporters were detected in these cultures, GLAST and GLT-1, but the immunoreactive band for GLT-1 was very faint compared to levels observed in brain tissue. To determine if GLT-1 contributes significantly to activity in these cultures, the sensitivity to the GLT-1 selective inhibitor, dihydrokainate, was examined. The fact that transport activity was essentially insensitive to inhibition by dihydrokainate suggests that most of the activity is mediated by GLAST. Therefore, the effects of rottlerin on transport activity are presumably caused by either direct or indirect inhibition of GLAST.
This rottlerin-induced inhibition was first examined at a low concentration of glutamate (0.5 µm) relative to the Km value for transport which is ∼60–70 µm. Under these conditions, the effect of rottlerin was concentration-dependent and was consistent with an IC50 value of ∼8 µm. The subsequent analyses of the kinetics of glutamate transport revealed that rottlerin (10 µm) decreased the Vmax to ∼10% of control. This would suggest that the IC50 value for rottlerin-induced inhibition of GLAST-mediated transport activity is lower than that obtained using a single low concentration of glutamate. If one assumes that this inhibition is caused by a simple bimolecular interaction, this reduction in Vmax would indicate that the IC50 value is approximately 1 µm. Treatment with rottlerin also decreased the Km value for transport. It seems likely that the effect of rottlerin on Km explains the difference in IC50 values (1 µm predicted from the decrease in Vmax vs. 8 µm obtained by evaluating the concentration-dependence at a low concentration of glutamate). At a low concentration of glutamate, the decrease in Vmax would be partially counterbalanced by an increase in the apparent affinity of the transporter for glutamate. At present, it is unclear how rottlerin decreases the Km value. In earlier studies, we found that changes in the capacity for transport in this system correlate with changes in Km values (Schlag et al. 1998). This effect may be related to a phenomenon described originally as ‘unstirred layers’ (see Garthwaite 1985 for original discussion). In a high capacity system, substrate can be cleared very rapidly in the local environment of the transporters so that the concentration of glutamate required to occupy 50% of the transporters in the bulk medium is higher than the intrinsic Km value. It is possible that by non-competitively inhibiting GLAST-mediated transport, rottlerin slows the overall clearance of glutamate and reduces the impact of ‘unstirred layers’. Although we suspect that the change in Km is simply related to this phenomenon, we cannot rule out the possibility that rottlerin has an effect on the Km of the transporter for glutamate.
Although changes in protein levels could theoretically be related to changes in transcription/translation, it is highly likely that the loss of GLAST immunoreactivity is independent of a change in protein synthesis. Using antibodies directed against either the amino or carboxyl termini of GLAST, there is 30–40% loss of GLAST protein within 5 min and approximately a 50% loss with 30 min of rottlerin treatment. In order for inhibition of protein synthesis to result in such a large loss of protein, the half-life of newly synthesized transporters would need to be less than 30 min. We found that inhibition of protein synthesis resulted in only a 20% loss of protein after 24 h, suggesting that the half-life of GLAST in this system is greater than 48 h (assuming that degradation follows first order kinetics). Based on these observations, it seems likely that rottlerin either accelerates intracellular (lysosomal or proteosomal) degradation of GLAST or causes limited proteolysis of both termini. In fact, Beckstrom and colleagues (Beckstrom et al. 1999) have demonstrated that in postmortem tissue the loss of GLT-1 immunoreactivity using antibodies directed against either the amino or the carboxyl termini is faster that the loss of immunoreactivity directed against epitopes reacting with middle portion of GLT-1. In addition, there was a loss of GLAST immunoreactivity using antibodies directed against carboxyl terminus. In this study, transport activity measured in a reconstituted system was essentially as stable as the middle part of GLT-1, suggesting that this limited proteolysis did not inactivate the transporter. At present, we cannot distinguish between these two possibilities (limited proteolysis and intracellular degradation). Regardless of the mechanism, unlike the limited proteolysis of GLT-1, the rottlerin-induced loss of GLAST-mediated activity occurs as quickly as the loss of immunoreactivity.
Since rottlerin also inhibited Na+-dependent glycine transport, we were somewhat concerned that rottlerin might have a non-specific effect on the rate of degradation of other membrane proteins. The observation that rottlerin has no effect on the levels of transferrin receptor in these same cells would rule out this type of non-specific effect. These data may also suggest that a common ‘rottlerin-target’, influences both glutamate and glycine transport.
In a recent study, rottlerin was shown to inhibit the accumulation of d-aspartate in retinal preparations (Bull and Barnett 2002). In contrast to the present study, the effect on uptake was not accompanied by a decrease in GLAST immunoreactivity. At present, the reason for this difference is not clear. It is possible that rottlerin has different effects on GLAST in retina and in astrocytes. The effects of rottlerin observed in the present study appear to be independent of PKCδ for the following reasons. Although we observed essentially complete inhibition of glutamate transport activity with rottlerin, relatively high concentrations of the general inhibitors of both classical and novel PKCs, Bis I, Bis II and staurosporine had no effect on transport activity. The concentrations of staurosporine used in the present study (1 µm) are much higher than those required to inhibit PKCδ in several different systems (Gschwendt et al. 1994a; deVente et al. 1996; Ishii et al. 1996). Similarly, the concentrations of Bis I used in the present study are sufficient to inhibit the effects of PKCδ in cellular systems (Fujii et al. 2000; deVente et al. 1996). The effects of down-regulation of PKC were also examined. Although chronic incubations with PMA reduced PKCδ levels to ∼25% of control, rottlerin still inhibited transport activity under these conditions. Chronic incubation with PMA had an even more dramatic effect on the levels of phospho PKCδ (Ser643) which was essentially undetectable under these conditions. Originally it was thought that auto-phosphorylation of PKCδ is required for catalytic activity and therefore levels of immunoreactivity might serve as a useful surrogate marker of active PKCδ (Li et al. 1997). A more recent study confirmed that this was a site of auto-phosphorylation but indicated that mutation of this site did not reduce kinase activity (Stempka et al. 1999). Although these differences are still not resolved, it appears that this serine residue is phosphorylated by PKCδ and may still serve as an indirect marker for kinase activity.
Since it is our strong suspicion that the effects of rottlerin are independent of PKCδ, we considered other potential targets. Rottlerin inhibits CaM Kinase III with an IC50 value comparable to that for inhibition of PKCδ (Gschwendt et al. 1994a); we found that an inhibitor of CaM Kinase III had no effect on transport activity. There is some evidence to suggest that rottlerin may inhibit other signaling molecules, but these have not been identified (Leitges et al. 2001; Soltoff 2001; Zhao et al. 2002). Rottlerin is a structural analog of ATP and is thought to interact with the ATP binding site of PKCδ. Since rottlerin treatment also decreased Na+-dependent glycine transport, we considered the possibility that rottlerin inhibits Na+-dependent transport systems by inhibiting the Na+/K+ ATPase. We found that a pre-incubation with ouabain (1 mm) followed by a rinse into transport buffer, as was performed with rottlerin, had no effect on transport activity (96 ± 5% of control, n = 5 observations). This suggests that astrocytes can very rapidly re-establish the electrochemical gradients required to maintain transport and that the effect of rottlerin on activity is not likely to be related to inhibition of the Na+/K+ ATPase.
Together, these data suggest that there is a novel target for rottlerin that regulates GLAST transport activity by controlling the rate of GLAST degradation or proteolysis. Many membrane proteins undergo a recycling between the plasma membrane and internal vesicles. The rate of recycling would need to be very fast in order for rottlerin to inhibit activity to ∼50% of control after only a few minutes. Although it is theoretically possible that the transporters recycle this quickly, one might expect the presence of an intracellular pool of transporter if the turnover was this quick and we have never observed any evidence of a non-biotinylated (intracellular) pool of GLAST. It seems more likely that rottlerin has an effect on the steps that control endocytosis of GLAST. Rottlerin may either inhibit a process that normally prevents GLAST from being internalized and targeted for degradation or rottlerin may activate a process that targets GLAST for degradation. The identification of this target could be important for understanding the regulation of GLAST levels under physiologic and pathologic conditions. In fact, it has been reported that traumatic brain injury causes a loss of GLAST within 2 h of the insult (Rao et al. 1998; Landeghem et al. 2001). If the half-life of GLAST in vivo is comparable to that observed in astrocyte cultures, this would suggest that these insults are accelerating degradation or proteolysis of GLAST.
In summary, we have shown that rottlerin rapidly inhibits GLAST-mediated activity in astrocyte cultures. This effect is associated with a dramatic decrease in the capacity for transport activity and with a loss of GLAST immunoreactivity. Evidence is provided that this loss of GLAST is caused by an increase in the rate of GLAST degradation or proteolysis, but it is not associated with an increased rate of degradation of another membrane protein (transferrin receptor). These studies also suggest that the effects of rottlerin are independent of inhibition of PKCδ and may be mediated by an interaction with an unidentified target that controls the rate of GLAST degradation. Although beyond the scope of the current study, it may be possible to develop strategies to identify this target using rottlerin as a tool.
The authors would like to thank Keith Fournier, Dr Marco Gonzaléz, Dr Marcelo Katanietz and Dr Avtandil Kalandadze for the helpful suggestions during the preparation of this manuscript. The authors would also like to thank Dr Jeffrey Rothstein for providing the anti-transporter antibodies. This work was supported by the NIH grant NS29868 and NS36465.